A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae)

Home , Aegyptianella, Argas persicus, Argas walkerae, Dermacentor albipictus, Dermacentor reticulatus, Ehrlichiaceae

By Evert E. Lindquist Terry D. Galloway Harvey Artsob L. Robbin Lindsay Michael Drebot Heidi Wood Richard G. Robbins

With illustrations by King Wan Wu and Barry Flahey Maps by Tom Naughten

All illustrations in this book are copyright by individual authors, and their unauthorized use is prohibited. For further information contact the Biological Survey of Canada.

The Biological Survey of Canada is an incorporated not-for-profit group devoted to promoting biodiversity science in Canada. It develops and coordinates national initiatives in systematics, biodiversity surveys, and publication of knowledge products.

The monograph series of the Biological Survey of Canada comprises invited, fully reviewed publications relevant to the biodiversity of Canada.

ISBN: 978-0-9689321-8-6 doi: 10.3752/9780968932186 A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) i

This handbook is dedicated to John D. (”Jack”) Gregson (17 June 1910–29 October 2006): father of Canadian tick taxonomy and ecology, and lifelong advocate for Canada’s natural heritage. Photograph courtesy of Petrina Gregson. ii E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

List of Contributors

This handbook was assembled through the efforts of seven authors and three illustrators, each of whom contributed to different components of the handbook. Author Bios: Evert E. Lindquist is an honorary research associate with Agriculture and Agri-Food Canada (AAFC), in Ottawa. His research encompasses the systematics of diverse groups of mites, symbiotic relationships between mites, insects, nematodes and fungi, and biodiversity of targeted mite taxa in selected ecosystems. As distinguished visiting lecturer, he taught the agricultural acarology course at Ohio State University for 18 years. Terry D. Galloway is Professor Emeritus in the Department of Entomology, Faculty of Agricultural and Food Sciences at the University of Manitoba in Winnipeg. His research career has concentrated on veterinary and aquatic entomology, with recent focus on arthropods parasitic on vertebrates in Canada. Harvey Artsob is the Former Director of the Zoonotic Diseases and Special Pathogens programme at the National Microbiology Laboratory of the Public Health Agency of Canada in Winnipeg. Much of his research has focused on surveillance for the occurrence of various zoonotic disease agents. His particular interests include arboviruses (California serogroup viruses, Powassan, West Nile and dengue), hantaviruses, and Lyme disease. L. Robbin Lindsay is a research scientist at the Public Health Agency of Canada’s National Microbiology Laboratory in Winnipeg, Manitoba. Robbin has been involved in ticks and tick-associated pathogen surveillance, diagnostic testing and research since 1989 and began his career studying the factors that limit blacklegged ticks on the Long Point peninsula in southern Ontario. Michael A. Drebot is Director of Zoonotic Diseases and Special Pathogens at the Public Health Agency of Canada’s National Microbiology Laboratory in Winnipeg, Manitoba. Michael’s research interests include the study of emerging mosquito and tick-borne pathogens such as West Nile and Powassan viruses. Heidi Wood is Chief of the Rabies, Rickettsia and Related Zoonotic Diseases section at the Public Health Agency of Canada’s National Microbiology Laboratory in Winnipeg. Heidi’s section focuses on arthropod-borne rickettsioses, Q fever and rabies virus. Richard G. Robbins is a recently retired Senior Civilian Entomologist at the Armed Forces Pest Management Board, Office of the Assistant Secretary of Defense for Energy, Installations and Environment, Washington, D.C. He has authored or coauthored over 100 peer-reviewed scientific papers on ticks and tick-borne diseases, as well as five books and several book chapters, and served for seven years as an instructor at the Acarology Summer Program, Ohio State University.

Authors contributing to each chapter of the handbook are as follows: Introduction – E.E. Lindquist and T.D. Galloway; General natural history – E.E. Lindquist; External structures – E.E. Lindquist; Collecting, preserving and studying ticks – R.G. Robbins; Medical and veterinary importance of ticks in North America – H. Artsob, M. Drebot, H. Wood, L.R. Lindsay, and T.D. Galloway; Identification and natural history of the ticks of Canada – E.E. Lindquist, T.D. Galloway and L.R. Lindsay; Illustrations – K.W. Wu and B. Flahey; Maps – T. Naughten. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) iii

Table of Contents List of Contributors...... ii Introduction to the ticks of Canada...... 1 General natural history...... 3 External structures and terms for describing and identifying ticks...... 5 Collecting, preserving, and studying ticks...... 32 Medical and veterinary importance of ticks in North America...... 36 Diseases of microbial etiology...... 37 Bacterial diseases...... 37 Lyme borreliosis...... 37 Southern tick-associated rash illness (STARI)...... 38 Avian borreliosis...... 38 Tick-borne relapsing fever...... 39 Ehrlichioses...... 40 Human granulocytic anaplasmosis (HGA)...... 41 Bovine anaplasmosis...... 42 Anaplasma ovis...... 43 Canine infectious cyclic thrombocytopenia...... 43 Mycoplasma haemocanis...... 43 Q fever...... 43 Tularaemia...... 44 Rocky Mountain spotted fever (RMSF)...... 45 Rickettsia parkeri rickettsiosis...... 48 Potential tick-borne bacterial pathogens...... 48 Other Rickettsia species...... 48 Bartonella species...... 49 Borrelia miyamotoi...... 49 Coinfections acquired from Ixodes ticks...... 50 Protozoan parasitic diseases...... 50 Cytauxzoonosis...... 50 Babesiosis...... 51 Viral diseases...... 51 Colorado tick fever ...... 51 Powassan encephalitis...... 52 Diseases of non-microbial etiology...... 53 Anaemia ...... 53 Tick paralysis...... 54 Skin lesions...... 56 Hypersensitivity reactions...... 56 iv E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Dermacentor albipictus and moose...... 57 Tick resistance...... 57 Immunosuppression caused by tick infestations...... 58 Identification and natural history of the ticks of Canada...... 58 Key to recognition of adults and immature instars of ticks...... 58 Keys to families and genera of ticks in Canada...... 59 Family Ixodidae...... 61 Genus Amblyomma Koch...... 61 Key to species of the genus Amblyomma in Canada...... 64 Amblyomma americanum (Linnaeus)...... 64 Amblyomma maculatum Koch...... 66 Genus Dermacentor Koch...... 69 Key to species of the genus Dermacentor in Canada...... 70 Dermacentor albipictus (Packard)...... 71 Dermacentor andersoni Stiles...... 76 Dermacentor variabilis (Say)...... 83 Genus Haemaphysalis Koch...... 88 Key to Species of the Genus Haemaphysalis in Canada...... 88 Haemaphysalis chordeilis (Packard)...... 88 Haemaphysalis leporispalustris (Packard)...... 91 Genus Ixodes Latreille...... 95 Key to species of Ixodes in Canada – females...... 95 Key to species of Ixodes in Canada – males...... 99 Key to species of Ixodes in Canada – nymphs...... 102 Key to species of Ixodes in Canada – larvae...... 105 Ixodes (Ceratixodes) uriae White...... 109 Ixodes (Trichotoixodes) brunneus Koch...... 113 Ixodes (Ixodes) dentatus Marx...... 118 Ixodes (Ixodes) muris Bishopp and Smith...... 122 Ixodes (Ixodes) pacificus Cooley and Kohls...... 127 Ixodes (Ixodes) scapularis Say...... 134 Ixodes (Ixodes) spinipalpis Hadwen and Nuttall...... 141 Ixodes (Ixodiopsis) angustus Neumann...... 144 Ixodes (Ixodiopsis) ochotonae Gregson...... 148 Ixodes (Ixodiopsis) soricis Gregson...... 152 Ixodes (Multidentatus) auritulus auritulus Neumann...... 155 Ixodes (Pholeoixodes) baergi Cooley and Kohls...... 160 Ixodes (Pholeoixodes) banksi Bishopp...... 164 Ixodes (Pholeoixodes) cookei Packard...... 167 A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) v

Ixodes (Pholeoixodes) hearlei Gregson...... 172 Ixodes (Pholeoixodes) kingi Bishopp...... 176 Ixodes (Pholeoixodes) marmotae Cooley and Kohls...... 181 Ixodes (Pholeoixodes) marxi Banks...... 185 Ixodes (Pholeoixodes) rugosus Bishopp...... 190 Ixodes (Pholeoixodes) sculptus Neumann...... 194 Ixodes (Pholeoixodes) texanus Banks...... 198 Ixodes (Pholeoixodes) gregsoni Lindquist, Wu, and Redner...... 202 Ixodes (Scaphixodes) howelli Cooley and Kohls...... 205 Ixodes (Scaphixodes) signatus Birula...... 209 Genus Rhipicephalus Koch...... 213 Rhipicephalus (Rhipicephalus) sanguineus (Latreille)...... 213 Family Argasidae...... 216 Genus Argas Latreille...... 217 Key to the species of Argas in Canada...... 217 Argas (Argas) cooleyi Kohls and Hoogstraal...... 217 Argas (Persicargus) persicus (Oken)...... 219 Genus Carios Latreille...... 226 Key to the species of Carios in Canada...... 227 Carios concanensis (Cooley and Kohls)...... 227 Carios kelleyi (Cooley and Kohls)...... 229 Genus and subgenus Ornithodoros (Pavlovskyella) Pospelova-Shtrom...... 234 Key to the species of Ornithodoros in Canada...... 234 Ornithodoros (Pavlovskyella) hermsi Wheeler, Herms, and Meyer...... 234 Ornithodoros (Pavlovskyella) parkeri Cooley...... 239 Genus Otobius Banks...... 240 Key to the species of Otobius in Canada...... 240 Otobius lagophilus Cooley and Kohls...... 240 Otobius megnini (Dugès)...... 242 Acknowledgements...... 250 References...... 251 Appendix 1. Host-tick index...... 284 Appendix 2. Tick-host index...... 300 Index...... 311 A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 1

Introduction to the ticks of Canada

It is unlikely that ticks would rank among the top 10 favourite animals for most people. Their surreptitious movements over the body of their hosts, especially upon humans, can result in a surprising discovery, and the presence of an engorged female ixodid is enough to elicit an expression of revulsion from even the most stalwart back country traveller. Even among entomologists, who generally are more complacent about the attractiveness or repulsiveness of terrestrial arthropods, arachnophobia is prevalent, but with ticks more disliked than spiders (Vetter 2013). Ticks are a highly distinctive group of relatively large mites (body length 1–5 mm as unfed adults but up to 20 mm when fully engorged), among which they are distinguished by a set of unique specialisations of their mouthparts and body for seeking and feeding obligately on the blood of amphibians, reptiles, birds, and mammals. All postembryonic instars – larva, nymph, and adult – of most tick species are capable of harming their hosts through exsanguination or secondary infection at sites of attachment. Recent lists of ticks of the world include 867–907 species (Horak et al. 2002; Barker and Murrell 2008; Guglielmone et al. 2010). Although most diverse in tropical and subtropical regions, ticks are also significant pests of wild and domesticated animals and humans in temperate and boreal forest and prairie habitats, and even in harsher tundra of the far north. In addition to causing exsanguination and paralysis, ticks of some species serve as reservoirs and vectors for a remarkable variety of pathogens, including viruses, bacteria, and protozoans. In Canada, these pathogens include the causative agents of Lyme borreliosis, relapsing fever, tularaemia, Rocky Mountain spotted fever, Q fever, Colorado tick fever, Powassan encephalitis, and perhaps others. Ixodid ticks of the genus Dermacentor can cause a motor paralysis in humans and other mammals following extended periods of engorgement. Tick paralysis may result in death if the feeding ticks are allowed to remain attached to the host. Ticks are classified in their own order, Ixodida (sometimes referred to as Metastigmata), in the Acari, or acarines, a subclass of Arachnida (Lindquist et al. 2009). Of the three families of Ixodida generally recognised, two, the Argasidae and Ixodidae, are represented in Canada, where 40 species in 10 genera are known to occur. Argasid ticks, known as “soft ticks” in lacking a hard dorsal shield on their bodies, are generally nocturnal and are rapid feeders, resembling bed bugs in this respect. Only eight species, in four genera (following the concepts of Klompen and Oliver (1993) for genera), are known or thought to occur in Canada. Ixodid ticks, known as “hard ticks” in having a hard dorsal shield, or scutum, that covers at least the anterior portion of the body, are mostly diurnal and slow feeders, remaining attached to their hosts for considerable lengths of time if undisturbed. Thirty-two species, in six genera are known or thought to occur in Canada, 23 of which belong to the genus Ixodes. Keys for identification and other information about ticks in Canada were last presented in a monograph by Gregson (1956). Although that work has served well during the interval since its issue, it has become considerably out of date with regard to the species of ticks that are now known to occur in Canada, their distribution, their range of hosts, and the variety of pathogens that they carry. For example, Lyme borreliosis has only been known as such since the mid-1970s, and human babesiosis is a more recently emerging disease in the northeastern United States of America and southeastern Canada. Moreover, there have been no publications for Canada, and few for the continental United States of America (see Webb et al. 1990) that have included keys to the larvae of tick species. As a reservoir and vector of some pathogens, the larva may be just as important as subsequent instars. An accurate identification of a feeding tick to species, whether it is larva, nymph, female or male, is the essential first step in leading to information as to whether there should be 2 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins concern about the tick being a potential vector of a pathogen of one disease or another. Although certain species of ticks have been shown to be competent vectors of disease- causing pathogens, many others either do not harbour pathogens or do not serve as competent vectors of them. As users of other keys to species of ticks available for various regions of North America, we are aware of how difficult some sections of keys are for non-specialists. We have tried to make our keys as “user-friendly” as possible, by selecting relatively easily visible attributes and by including a second attribute for most couplets, in case the first one is damaged or missing on a specimen. For each active instar (larva, nymph, adult female and male), attributes presented by the keys are augmented by diagnoses, further to help in genus and species identification of ticks. Most of the illustrations were selected among line figures available in previous publications, and redrawn (sometimes with modifications) to make them more standard in appearance. Scanning electron microscopy figures were avoided, as they present excessive details often not useful in routine identification, and their appearance differs much from what non- specialists will see, using an ordinary stereomicroscope. Nevertheless, the line drawings are mostly insufficient in showing the scutal ornamentations noted in the diagnoses. This handbook is designed to present a general survey of available information on most species of ticks known or anticipated to occur in Canada, in such a way that viewers may become familiar with their identities and known natural history. The information will be equally useful in Alaska and the northern tier of contiguous states of the United States of America bordering Canada. Unfortunately, with the availability and scope for travel to exotic destinations, people may bring back a great variety of ticks with them from their excursions. In addition, the pet trade is involved with importation of exotic animals from around the world, and tiny, recently attached ticks may escape detection in quarantine facilities. Given this diverse pool of potential introductions of exotic ticks, it would be impossible to include every species in this handbook. Therefore, when people find ticks for which there is a possibility of travel-related introduction, that species may not appear in this handbook, and the tick may require examination by an expert. It is also important to recognise that ticks may be transported great distances on migrating birds. Consequently, additional species of ticks unlikely to become part of the Canadian tick fauna may be discovered in the future (e.g., see Scott and Durden 2015a, 2015b), but these are not included in this handbook. People using the keys are reminded to use caution when examining ticks infesting such hosts. Information on tick distributions and hosts, beyond that given in Gregson’s work, was based initially on data with specimens accumulated in the Canadian National Collection (CNC) of ticks in the care of the Science and Technology Branch, Agriculture and Agri-Food Canada, Ottawa, Ontario, Canada. Gregson’s extensive collection, which is considerably concentrated on western and mid-western Canadian records, has been deposited in the CNC. It is complemented by substantial numbers of ticks and records that have accumulated largely from mid-eastern and eastern Canadian regions, especially associated with various passive surveillance programmes in recent years. Data from other Canadian sources have also been included to provide insight to the overall distribution of these ticks in Canada. No attempt has been made to include all known locality records. There are many gaps in our knowledge of species of ticks in Canada, particularly their life histories, vector potential, and methods of dispersal. In addition, ranges of many species of ticks are changing rapidly (e.g., Dergousoff et al. 2013) and government policies regarding ticks and tick-borne pathogens are being modified. We hope this handbook will serve to stimulate further investigations in these fields. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 3

General natural history

All ticks have an egg stage, a six-legged (hexapod) larval instar, one or more eight-legged (octopod), sexually immature nymphal instars, and an eight-legged, sexually mature adult instar. Ixodid (hard) ticks have one nymphal instar, whereas argasid (soft) ticks may have as many as eight. When comparing developmental stages, homology is determined from changes in structure of the instar, i.e., the form of the tick between moults. The stage, or stadium, itself is the period of time between moults, or from oviposition until the first moult in the case of the egg, and from the final moult until death in the case of the adult. The instar, or stase, is the form of the individual between two moults or, in the case of the first or last instar, between the egg and the first moult and, respectively, the final moult and its end of postembryonic development. In both families, all larval and nymphal instars and adults of both sexes feed on blood (except adults of Otobius), although ixodid males may feed little, if at all. Hard ticks generally take one blood meal per stage, followed by moulting, and adult females oviposit and die following the blood meal. Soft ticks feed intermittently, and adult females may feed and oviposit several times. Ticks have evolved a great variety of host relationships but are commonly classified as “one-host”, “two-host”, “three-host”, or “many-host” species. One-host ticks complete all feeding and moulting on one animal, usually a large-sized, wandering host (sheep, cow, horse, deer, antelope, buffalo, etc.). Adults mate on the same host, after which females drop to the ground to lay one large batch of eggs before dying (males may remain on the host until death). This strict host specificity is uncommon, being seen in only about a dozen tick species, mostly ixodids and including all members of the economically important subgenus, Boophilus and the moose tick, Dermacentor albipictus (Packard). In two-host ticks, the moult from larva to nymph occurs on the first host, which is usually a small mammal or bird, and the engorged nymph drops to the ground, where it moults to the adult. The adult must then find a second, usually much larger host. Again, there are only about a dozen two-host ticks, almost all of them ixodids (chiefly in the genera Hyalomma and Rhipicephalus), including several associated with livestock. No two-host ticks are known to occur in Canada. All other ixodids (about 625 species) are three-host ticks that detach after engorging at each instar, with moults taking place off the animal. A variety of hosts may be used by each instar, increasing the likelihood of acquiring pathogens, but larvae and nymphs usually feed on smaller hosts than adults. Many-host ticks are typically argasids, which feed on a number of different animals during their life cycle, the adults feeding several times. Note that there are evolutionary advantages and disadvantages to varying the number of stages per host; thus, three-host ticks may be less host-specific but face the challenge of finding a new host after each blood meal, whereas one-host ticks are guaranteed a food source for life but may be threatened with extinction if their sole host dies. Copulation takes place following the last moult, after which the female engorges and produces eggs, which are laid on the ground or in some sheltered location. Female hard ticks lay their eggs in one large batch, often numbering several thousand, then die. Female soft ticks lay multiple smaller egg batches, from less than a hundred to a few hundred at a time. Ticks are extremely hardy and can survive long periods of environmental stress. Some species have been known to survive unfed for several years, and all life history instars can tolerate long periods of submersion in water. Ticks have also developed a number of adaptations to limit water loss; soft ticks in particular are resistant to desiccation, often inhabiting xeric environments (Hafez et al. 1970). 4 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Hard ticks, and a few species of soft ticks, seek their hosts by climbing vegetation, either grass or brush, and waiting for a suitable animal to pass. In a behaviour known as questing, the first pair of legs is extended and used to grasp the host when contact is made. The height at which questing takes place determines the size, and therefore to some extent, the species, of host selected. Ticks cannot jump and do not leap upon their hosts from trees or lower vegetation. Questing heights vary among tick species and among the different life instars of any given species. In contrast, most soft ticks take refuge by day in loose soil or cracks and crevices, attacking their host at night, usually while it is asleep. They crawl to the host, engorge in a few minutes or hours, and then return to their hiding place. Ticks orient to potential hosts in response to a number of stimuli, including host odours, sudden changes in light, temperature and vibrations. Carbon dioxide, a product of host respiration, is especially attractive, even from a distance.

Table 1. Ticks collected in Canadian localities

Taxon Canada Outside Canada Argasidae Argas persicus (Oken) BC Cosmopolitan Argas cooleyi Kohls and Hoogstraal BC California, Montana Carios concanensis (Cooley and Kohls)* AB, SK Western USA, Mexico Carios kelleyi (Cooley and Kohls) AB, SK Central and eastern USA Ornithodoros (Pavlovskyella) hermsi Wheeler, BC Western USA Herms, and Meyer Ornithodoros (Pavlovskyella) parkeri Cooley Western USA Otobius megnini (Dugès) BC Western and central USA, Mexico, Central and South America, Africa, India Otobius lagophilus Cooley and Kohls AB Western USA

Ixodidae Amblyomma americanum (Linnaeus) ON, QC Eastern USA Amblyomma maculatum Koch Southern USA Dermacentor albipictus (Packard) Canada North and Central America Dermacentor andersoni Stiles BC, AB, SK Western USA Dermacentor variabilis (Say) Central and Central and eastern USA, plus eastern Canada CA, OR, WA, ID, TX Haemaphysalis chordeilis (Packard) Canada Eastern and southern USA Haemaphysalis leporispalustris (Packard) Canada USA, Mexico, Argentina Ixodes (Ceratixodes) uriae White Coastal BC, NS, NL cosmopolitan, coastal Ixodes (Trichotoixodes) brunneus Koch MB, ON USA, Europe Ixodes (Ixodes) dentatus Marx ON Eastern USA Ixodes (Ixodes) muris Bishopp and Smith Eastern Canada Eastern USA Ixodes (Ixodes) pacificus Cooley and Kohls BC Western USA, Mexico A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 5

Taxon Canada Outside Canada Ixodes (Ixodes) scapularis Say Central and Eastern USA eastern Canada Ixodes (Ixodes) spinipalpis Hadwen and Nuttall BC, AB Western USA Ixodes (Ixodiopsis) angustus Neumann Canada Northern USA, northeastern Asia Ixodes (Ixodiopsis) ochotonae Gregson BC, SK Western USA Ixodes (Ixodiopsis) soricis Gregson BC Western USA Ixodes (Multidentatus) auritulus Neumann BC, ON Cosmopolitan Ixodes (Pholeoixodes) baergi Cooley and Kohls ON Central and southern USA Ixodes (Pholeoixodes) banksi Bishopp ON, MB Eastern and southern USA Ixodes (Pholeoixodes) cookei Packard Eastern Canada Eastern USA Ixodes (Pholeoixodes) hearlei Gregson BC Western and southern USA Ixodes (Pholeoixodes) kingi Bishopp AB, SK Western USA Ixodes (Pholeoixodes) marmotae Cooley and Kohls BC Western USA Ixodes (Pholeoixodes) marxi Banks NS, ON Eastern USA Ixodes (Pholeoixodes) rugosus Bishopp BC West coastal USA Ixodes (Pholeoixodes) sculptus Neumann BC, AB, SK, MB Western and central USA Ixodes (Pholeoixodes) texanus Banks BC, ON North America Ixodes (Pholeoixodes) gregsoni Lindquist, Wu, Eastern Canada Northeastern USA and Redner Ixodes (Scaphixodes) howelli Cooley and Kohls BC AK, CA, TX Ixodes (Scaphixodes) signatus Birula Coastal BC West coastal USA, Japan Rhipicephalus (Rhipicephalus) sanguineus Eastern Canada Cosmopolitan (Latreille)

* In some cases, there is discord on the higher classification of argasid ticks. In this handbook, we have adopted Klompen and Oliver (1993) and their recognition of Carios as a genus. British Columbia (BC), Alberta (AB), Saskatchewan (SK), Manitoba (MB), Ontario (ON), Québec (QC), Nova Scotia (NS), Newfoundland and Labrador (NL), United States of America (USA), Alaska (AK), Washington (WA), Oregon (OR), California (CA), Idaho (ID), Texas (TX).

External structures and terms for describing and identifying ticks

As with their relatives, the other parasitiform mites, the body of ticks consists of an anterior gnathosoma or capitulum, and a posterior idiosoma or “body” (Fig. 1); these are separated by the circumcapitular suture, which is the only articulation of the body other than on the appendages. The gnathosoma is generally positioned to project apically, so it is visible in dorsal aspect, in larval ticks, but it projects ventrally, and is usually concealed from above, in nymphal and adult argasid ticks. This is in contrast to its visible apical position and anterior projection in nymphal and adult ixodid ticks. Gnathosoma. The gnathosoma superficially resembles the head of insects, and it consists of a subcapitulum or basis capituli, a hypostome, and the paired chelicerae and palpi (Fig. 3). The basis capituli (or, simply, basis) is the basal portion of the gnathosoma to which the palpi and mouthparts (hypostome and chelicerae) are attached 6 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

tarsal hump tarsus

tibia

gnathosoma (capitulum) genu femur trochanter coxa scapula lateral carina cervical groove scutum eye

marginal groove

idiosoma scutal emargination

fovea

marginal body fold

festoons

Fig. 1. Diagrammatic attributes of body dorsum of adult female Ixodidae. Redrawn from Gregson (1956). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 7

gnathosoma (capitulum)

scapula lateral carina cervical groove

pseudoscutum lateral groove

marginal body fold

fovea

scutum

parma

Fig. 2. Diagrammatic attributes of body dorsum of adult male Ixodidae. Redrawn from Cooley and Kohls (1945). 8 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

corona palpal tibiotarsus (article 4)

palpal genu digit of chelicera (article 3) hypostome

palpal femur cheliceral sheath (article 2)

shoulder of hypostome anterior spur palpal trochanter posterior spur (article 1) basis capituli porose area cornu auricula transverse sutural line

A B

Fig. 3. Diagrammatic attributes of gnathosoma of female Ixodidae. A. dorsal. B. ventral. Redrawn from Gregson (1956). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 9

hood

eye

coxa

eye

spiracle

dorso-ventral groove

preanal groove median postanal groove

transverse postanal groove

A B

Fig. 4. Diagrammatic attributes of body of adult female Argasidae. A. lateral. B. ventral. Redrawn from Cooley and Kohls (1944). 10 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

hood

discs

mammillae

dorso-ventral groove

Fig. 5. Diagrammatic attributes of body dorsum of adult Argasidae. Redrawn from Cooley and Kohls (1944). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 11

Fig. 6. Diagrammatic attributes of larval Argasidae. A. dorsal. B. ventral. Redrawn from Kohls et al. (1965). 12 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

corona

denticles file 4

file 3

file 2

file 1

lateral file

base

A B

Fig. 7. Diagrammatic attributes of hypostome of female, nymphal and larval Ixodidae. Redrawn from Cooley and Kohls (1945). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 13

crenulation

lateral file

Fig. 8. Diagrammatic attributes of hypostome of male Ixodidae. Redrawn from Cooley and Kohls (1945). 14 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

(Fig. 3). In nymphal and adult argasid ticks, the gnathosoma lies in a depression, the camerostome, which becomes less clearly defined in engorged specimens. Along its sides, the camerostome has a pair of flaps, the cheeks, which may be either fixed or movable. An anterior projection of the dorsal idiosomal wall, the hood (Figs. 4–5), covers the camerostome. In ixodid ticks, the gnathosoma is fully exposed in all active instars, and a variety of its visible structures are useful diagnostically. Dorsally, the basis capituli commonly has a rectangular shape, with more or less parallel sides, but it is hexagonal, with angularly projecting sides, in adults and immatures of Rhipicephalus and in immatures of Haemaphysalis and in some species of Dermacentor. It is triangular, with posteriorly projecting angles, in adults and immatures of some species of Ixodes and in immatures of some other species of Dermacentor. The basis of adult female ixodids bears a pair of porose areas, which are somewhat depressed with pitted floors (Fig. 3), from which antioxidants are produced to coat the eggs (Sonenshine 1991). Porose areas have characteristic shapes among species in some genera, such as Ixodes; they are absent in males, nymphs, and larvae. Ventrally, the anterolateral margins of the basis capituli, which are sometimes called the “shoulders of the hypostome” (Gregson 1956), are more or less characteristically flattened or humped or angled in leading to the base of the hypostome. Posterior to the attachment of the palpi, the lateral margins of the basis sometimes have a pair of projections, the auriculae (Figs. 3B, 32C), the form of which varies among different species; they may be slight protuberances, flattened ridges, or more strongly developed as hornlike processes or retrorse spurs. Near the posteroventral margin of the basis capituli, a transverse sutural line is evident in some species. Posteriorly, the basis may be constricted to a more or less typical degree among some species, or it may remain untapered. Two or three pairs of setae on the ventral face of the basis (Fig. 51I) are notable diagnostically at the species level for larvae and nymphs. These include one or two pairs of posthypostomal setae, inserted near the base of the hypostome, and one pair of postpalpal setae, inserted more laterally behind the bases of the palpi (Fig. 6B). The former are equivalent to the hypostomal or subcapitular setae, and the latter, found only on some argasid ticks, are equivalent to the palpcoxal or capitular setae, of mesostigmatid mites. A singularly specialised characteristic of ticks, in contrast to other mites, is the hypostome, a forward extension from the anterior portion of the basis that is modified as a piercing organ and armed with retrorse teeth or denticles, or sharp-edged ridges, termed crenulations (or, more concisely, crenulae), on its lateral and ventral surfaces (Figs. 7–8). The denticles are usually arranged in parallel longitudinal rows, or files, and the dentition formula indicates the number of files on each side of the midline of the hypostome. For example, 2/2 indicates the presence of two files on each side, whereas “first 3/3 and then 2/2” indicates that, of three files beginning on the distal portion, one file does not continue to the base of the hypostome. The lateralmost row is designated as file 1. The relative size of denticles differs characteristically among species of ticks, such that the number of teeth in each file is a useful diagnostic attribute; the lateral file generally has denticles at least as large as any present elsewhere on the hypostome. Sexual dimorphism is commonly present in the hypostome among adult ticks of the same species. In males the denticles may be altered to transverse or diagonal rows of sharp or blunt elevations or ridges called crenulations or crenulae as noted above. The argasid genus Otobius is unusual in that its adults have a vestigial hypostome, in contrast to a well developed hypostome in the immature instars. The apex of the hypostome may be pointed or rounded or notched, and it is commonly armed with a crown, the corona, of more or less numerous small denticles that do not form readily recognisable files. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 15

Lying dorsal to the hypostome are the paired chelicerae (Fig. 3), which are reduced to two segments in ticks and consist of an extrudable and retractable shaft that extends through the basis capituli and bears apical cutting digits of a uniquely modified form. These digits consist of a fixed inner digit to which a strongly dentate, outer digit is articulated on its external surface. A bifid or serrate dorsal process is also attached to the external side of the inner digit (see fig. 5.12E, p. 150 in Evans 1992). Only the cutting external teeth are generally exposed, and no emphasis is placed on the chelicerae in diagnoses. The palpi are movable, segmented, paired appendages attached to the basis capituli anteriorly on either side flanking the hypostome (Fig. 3). They are hollowed along their inner faces, such that, when apposed, they more or less cover the hypostome and chelicerae; they are spread apart over the surface of the skin during feeding. The palpi generally consist of four, or rarely three, free segments and these are usually named and numbered articles 1–4 in a proximal to distal sequence in descriptive literature on ticks (e.g., Gregson 1956; Clifford et al. 1961; Edwards 1975). However, palpal setal numbers and setation patterns among larval ticks confirm that the four free segments in Ixodida are homologous with the segments termed trochanter, femur, genu, and fused tibiotarsus of other parasitiform Acari, and we follow Klompen (1992) in applying these terms to ticks. The proximal palpal trochanter is small, and sometimes immovably articulated; it may be visible only ventrally, or it is sometimes vestigial, as in the subgenus Boophilus. It sometimes bears one or two projections or flanges, directed diagonally anteriorly or posteriorly, or both, which may be of diagnostic value, especially among species of Ixodes; these are designated as anterior and posterior “horns” or “spurs” in some treatments. Some members of this genus are characterised by having the basal palpal segment not only fused with the basis capituli but also enlarged and often provided with the spur-like processes noted above. This enlarged structure, termed the palpiger, has been used to some extent in the diagnosis of subgenera of Ixodes. The palpal femur and genu are large conspicuous segments that are often more or less fused, with or without a defined suture line between them; the lateral margin profile of these two segments, whether straight or convex, and their combined length relative to their width are of diagnostic use. In the genus Haemaphysalis, the bases of the palpal femora are strongly flared laterally, giving the palps a characteristically conical shape. Excepting some taxa of Argasidae, the palpal tibiotarsus is small, and it is sometimes reduced to an inconspicuous papilla lying in a ventral depression near the apex of the palpal genu. The number and arrangement of setae on the palpal femur, genu, and tibiotarsus of larval ticks are important diagnostic attributes (Fig. 9). Palpal setation characters are not useful for nymphal and adult ticks because of increased numbers and intraspecific variability of setae during ontogeny, which render the homology of setae difficult and unreliable. The notation applied to the larval palpal setae is that proposed by Evans (1963b) for mesostigmatic mites and modified slightly by Klompen (1992) for ticks. Setae are distinguished and denoted by their position on the anterior (ad) and posterior (pd) dorsal faces, anterolateral (al) and posterolateral (pl) faces, and ventral (v) face of the palpal segments, except that the cluster of distal setae on the tibiotarsus are simply counted but not individually denoted. The palpal trochanter consistently lacks setae in larval ticks, as in larval Mesostigmata. A maximum of six setae is present on the palpal femur of larval ticks; these are denoted al, ad, pd-1, pd-2, pl, and v. Larvae of all Argasidae lack seta v, those of some argasid genera also lack one of the two pd setae, and those of Otobius also lack seta al on the palp femur (Klompen 1992). A maximum of seven setae is present on the palpal genu of larval ticks; these are denoted al, ad-1, ad-2, ad-3, pd-1, pd-2, pl. While larvae of Ixodidae have this full complement of setae or lack but one of the two pd setae, 16 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins those of all Argasidae lack seta ad-3 and one of the two pd setae, and one or two other setae (commonly al, and sometimes ad-2) are absent in some argasid genera (Klompen 1992). A pore-like structure is often evident on the dorsal face in the palpal genual area; care should be taken to not mistake this for an alveolus of a detached seta. According to Klompen (1992), setation of the fused palpal tibiotarsus in larval ticks is maximally 12 (Lindquist et al. 1999 observed 15 in larvae of Ixodes gregsoni), including a group of at most four (al, ad, pd, v) on the main body of the segment and a distal cluster of at most eight, which are not denoted as they are not readily homologised; setae al and pd are absent from the main body of the larval tibiotarsus, and one or two setae are absent from the distal cluster in some genera of Argasidae. All of the above reductions in setal numbers from the maximum complements on the palpal segments are considered to be derived character states within the Ixodida relative to more distantly related outgroups of parasitiform mites (Klompen 1992). Idiosoma. The idiosoma, often simply (and erroneously) called the “body”, is the largest region of the body of ticks, and is analogous to the combined thorax and abdomen of insects. The idiosoma of larval ixodid ticks bears a moderately small but well-developed anterior dorsal shield (Fig. 10) that is retained in relatively similar size on the nymph and adult female, and named the scutum (Fig. 1); the scutum is greatly expanded to cover most of the dorsal surface on the adult male (Fig. 2). In contrast, the idiosoma of larval argasid ticks has at most a small mid-dorsal plate, or mesonotal shield, that is lost in postlarval instars. Argasids are commonly referred to as “soft ticks” due to this lack of a well-developed dorsal scutum. However, the dorsal surface of nymphal and adult argasids is typically leathery and wrinkled, granulate or covered with small elevations called mammillae (Fig. 5), and the lateral margins of the dorsal surface are sometimes sharply delineated from those of the ventral surface by a definitesutural line. In addition, small, somewhat raised or depressed discs, or scutellae, of thickened cuticle may be visible as patterns on both dorsal and ventral surfaces of the idiosoma; these represent muscle attachment sites, or sigillae. The shape of the idiosoma is sometimes diagnostic among argasid species; for example, in dorsal view it is abruptly constricted posteriorly in adults of O. megnini, in contrast to O. lagophilus, and in lateral view, its anterior margin is turned up in adults of A. reflexus, in contrast to A. persicus. The scutum of adult Ixodidae bears a pair of anterolateral projections, the scapulae (Figs. 1–2); these may be pointed, rounded, or reduced to small lobes. The scutum of nymphal ixodids bears more weakly developed scapulae, or these may be indicated only as more narrowly rounded anterolateral corners, as on the larvae. The transverse anterior excavation between the scapulae in adult ixodids, the scutal emargination, accommodates upward movement of the closely articulated gnathosoma; it may be characteristically straight or somewhat sinuous. The scutum in all instars of Ixodidae often bears a pair of linear depressions, the cervical grooves (Figs. 1–2), that begin closely behind the scapulae and extend posterolaterally to somewhat beyond the midlevel of the scutum. They are sometimes absent, and when present they are often shallow, so as to be seen best by reflected light on temporarily dried specimens. Lateral to the cervical grooves, there may be linear elevations, the lateral carinae (Figs. 1–2), close to the margins of the scutum anterolaterally. When present, these are evident as more or less well-developed ridges, but they are often difficult to discern and we prefer to avoid referring to their presence or absence as key characters. The region bearing soft, pliable dorsal cuticle lateral and posterior to the scutum in ixodid immatures and females is sometimes termed the alloscutum. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 17

The shape and ornamentation of the scutum are important diagnostic characters of all instars of ixodid ticks (though less so of adult males). The scutum may be inversely subtriangular versus oval, its greatest width may be lesser versus greater than its median length, and the greatest width may be located anterior to, versus about at its midlength. The scutal surface may be partly or entirely marked with minute irregular fissures (crazed), or pitted with puncta of various size and density, or wrinkled (rugose). The nature of scutal ornamentation of ixodid adults and nymphs is visible under a good stereomicroscope, but that of ixodid larvae is often discernible only with a compound microscope, in which case temporary or permanent slide preparations of specimens need to be made. The scutum of some kinds of adult ixodid ticks is termed ornate when it has a pattern of enamel-like colour superimposed over the base colour of some shade of brown. Both colour and colour pattern or location may be useful diagnostically, particularly among species of Dermacentor, though considerable intraspecific variation of these attributes must be accounted for. Scutal setation, or its “hairiness”, including the relative density and length of setae, is sometimes a useful attribute. As with scutal topography, setation is best seen by reflected light on temporarily dried specimens of nymphs and adults. In larvae of ixodid ticks, the nature of scutal setation may be of more critical diagnostic importance, and includes the number, length, and position of setae, as well as pore-like structures (see below) on the scutum when viewed in slide preparations under a compound microscope. In some genera of ixodid ticks, there is a pair of hyaline lens-like structures, the eyes, near the lateral margins of the scutum (Figs. 1, 10). Two pairs of such structures are present on the anterior submarginal region of the idiosoma of certain argasid ticks. However, there are no eyes in other Argasidae, all species of the genus Ixodes, and some other genera of Ixodidae. In species of Dermacentor, the anterior region of the expansive scutum of males may be distinguished by a difference in colour pattern and by the posterior limits of the cervical grooves. This region, which is homologous with the smaller scutum of the nymph and female, is called the pseudoscutum. In males of other ixodid genera, the scutum may have a pair of lateral grooves that extend along the sides of the shield, beginning near the scapulae and extending to its posterior extremity. These grooves correspond to the pair of marginal grooves found on soft cuticle behind the scutum on adult females. Two circular structures, the foveae, are sometimes present on the midregion of the male scutum (Fig. 2); these are on soft cuticle closely posterior to the scutum of conspecific females. Uniform, subrectangular areas separated by grooves, called festoons, may be present along the posterolateral margins of the idiosomal dorsum in adults of both sexes of some ixodid genera, including Dermacentor, Rhipicephalus, and Haemaphysalis (Fig. 1). Festoons are thought to represent external vestiges of ancestral segmental boundaries (Klompen et al. 1996). The unpaired caudal festoon is termed the parma (Fig. 2). Located on the dorsal scutum in males, but on soft cuticle in females, the number of festoons may be a useful diagnostic attribute at the species or genus level. The setation of the idiosomal dorsum of larval ixodid ticks is extremely important in the generic and species diagnoses of larvae. Diagnostic attributes include the number and position of homologous setae on the scutum and soft cuticle and their positional relationship to certain other structures such as gland pores and lyrifissures, as well as the shape and size of setae. The system of notation follows that of Lindquist and Evans (1965) for mesostigmatic mites, as applied by Klompen et al. (1996) for larval Ixodidae. Dorsal idiosomal setae are recognised to form three longitudinal series, named and denoted as the dorsal median j-J series, dorsal mediolateral z-Z series, and dorsal lateral s-S series (an earlier system of chaetotaxy for larval ixodid ticks, developed by Glashchinskaya- Babenko (1949) and used by Clifford and Anastos (1960), named and denoted the setae 18 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

tibiotarsus al pd (article 4) v ad

ad1 pd genu al (article 3) pl ad2 pd1

al pl femur (article 2) pd2 ad

trochanter (article 1)

Fig. 9. Palpal structure and notation for larval Argasidae. Redrawn from Edwards (1975). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 19

Sc1 z2

Sc2 s2 Sc3 j3

Md1 s3 Cd1 j5 Md2 s4

Cd2 J2 Md3 s6

Ss LWG S1 Md4 Md5 S2 Md6 S3 Md7 S4 Md8 S5

Fig. 10. Setal notation alternatives for body dorsum of larval Ixodidae (see also Table 1). Redrawn from Clifford et al. (1961). 20 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins on soft cuticle as the central dorsals Cd, supplementaries S, and marginal dorsals Md; see Table 1 for equivalents). Lower-case letters are applied to setae on the scutum and elsewhere on the podonotal region (e.g., j5), while upper-case letters are applied to setae on the opisthonotal region (e.g., J2). The numbering of setae is designed to arrange them in transverse rows that correspond with hypothesised segments, which lack external delineation in ticks. Thus setae j3 and s3 are thought to be elements of the third segment of the podonotal region, and J2 and S2 are elements of the second segment of the opisthonotal region (Fig. 10). Klompen et al. (1996) also applied a notation to the lyrifissures and gland pores on the scutum and soft cuticle of ixodid larvae, which enables one to recognise hypothetical homologies among these structures and to use their absence or presence and position relative to adjacent structures as diagnostic attributes. In various works previous to Klompen’s, the lyrifissures were referred to as “sensilla auriformia”, the small glands and their pores as “sensilla hastiformia”, and the large glands and their pores as “sensilla sagittiformia”. Because of their prominence and apparently different function, the large glands have been designated as large wax glands (Fig. 10, LWG). The absence or presence of a pair of large glands on the dorsolateral surface of the idiosoma, and their position relative to the marginal dorsal setae, are useful attributes in distinguishing larval ixodids to genus (Clifford and Anastos 1960; Klompen et al. 1996). However, the use of lyrifissures and gland pores is only beginning to be applied diagnostically (e.g., Lindquist et al. 1999), and sufficient data from the diversity of ticks that occur in Canada are not available for their use as an aid in identifying larvae to species in this handbook. Ventrally, the idiosoma bears the genital aperture between the bases of legs II or III in adult females (Fig. 11) and between legs I, II, or III in adult males, and, posteriorly, the anus in all instars (Figs. 11–12). The genital aperture of the female is more obvious as a somewhat larger, transverse slit than that of the male. The anus is surrounded by a circular or oval sclerotised ring and is covered by a pair of eversible anal valves that bear one to several pairs of setae, the number of which is useful diagnostically among species of Ixodes (Fig. 11). The anus is partly enclosed by an anal groove in the soft cuticle of most ixodid ticks; this groove embraces the anus as an anterior arch in the genus Ixodes (Fig. 13), but is contoured posteriorly to the anus in other genera, though it is absent in a few, e.g., the subgenus Boophilus. A set of ventral plates or shields distinguish males of Ixodes from females as well as from males of other genera of Ixodidae which, as found in Canada, either have only a partial set of these plates (Rhipicephalus) or lack them entirely (Dermacentor, Haemaphysalis). A small pregenital plate may be present between the bases of legs I and II, in front of the male genital aperture (Fig. 30E). A more expansive median plate covers the area behind the genital aperture between and somewhat behind the bases of legs III and IV (Fig. 30E); it abuts both the anal plate, which surrounds the anus, and the pair of adanal plates, which flank the anal plate. A pair of epimeral plates, sometimes less well delineated, may flank the adanal plates (Fig. 45A); these are designated accessory plates in genera such as Rhipicephalus. Although subject to some intraspecific variation, the relative size and shape of these plates are useful diagnostically among species of Ixodes. Nymphal and adult argasids often have a series of ventral grooves or folds, which become less obvious as the ticks engorge. The presence or absence of some of these, such as the paired dorsoventral grooves lateral to the bases of legs IV, and the unpaired preanal, transverse postanal, and median postanal grooves (Fig. 4), are used descriptively, though usually not diagnostically, in keys to species of Ornithodoros. A pair of respiratory spiracular plates, also called stigmatal plates, is located just posterior to coxae IV in the nymphs and adults of all ticks (Fig. 11); larval ticks lack these structures (Fig. 12). The plate may be characteristically round, elliptical, oval, or inversely comma-shaped. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 21

In ixodid ticks, its periphery is bordered by one or several rings of pore-like chambers, which surround a field of goblets, small orifices whose relative size, number and arrangement may be useful diagnostically at the species level (Fig. 14). Near the centre of the spiracular plate of ixodid adults lies a larger and more heavily sclerotised hollow, the macula, which is the principal respiratory opening (Fig. 14). In argasid ticks, the spiracular plate lacks pore-like chambers and goblets, and has a simple opening set in a hinged flap (Fig. 4) which passes directly into an atrial cavity (Hinton 1967). As on the dorsum, the setation of the venter of the larval idiosoma has considerable diagnostic importance (Fig. 12). A chaetotactic notation, developed like that for the dorsal setae, recognises three longitudinal series, named and denoted as the ventral medial Jv series, ventral mediolateral Zv series, and ventral lateral Sv series (the earlier system of chaetotaxy, developed by Glashchinskaya-Babenko (1949) and used by Clifford and Anastos (1960), named and denoted these setae the preanals Pa, premarginals Pm, and marginal ventrals Mv; see Table 1 for equivalents). Only upper-case letters are applied to the ventral setae that are useful diagnostically, as all are on the opisthogastric region. Klompen et al. (1996) also applied a notation to the lyrifissures and gland pores on the ventral soft cuticle of ixodid larvae, as was done for the dorsum. Again, however, the use of lyrifissures and gland pores is only beginning to be applied diagnostically, and sufficient data are not available for their use in this handbook. Legs. All the legs of ticks have six segments (named from base to apex coxa, trochanter, femur, genu, tibia, tarsus) plus a pretarsus that bears a pair of claws and, in ixodids, a well- developed soft pad, the pulvillus, between the claws that enables ticks to walk on smooth surfaces (Figs. 1, 16–17). The coxae of ixodid ticks bear some of the most useful diagnostic attributes for identification, including their shape, presence or absence of spurs or flanges, and setation. When coxa I is denoted as “bifid”, as in the genus Dermacentor, it bears two spurs close together, with a deep incision between them. The absence or presence, shape and size of one or two well-separated spurs on the ventral surface of coxa I is critically important in diagnosis of species of Ixodes. The internal spur is on the proximal region of the coxa, closest to the body midline, and the external spur is on the apical region, farthest from the body midline. The absence or presence of spurs or flanges on the ventral surfaces of coxae II–IV, and whether the posterior edges of the coxae are rounded or have a projected edge, or salience, are used descriptively, but usually not diagnostically, among ixodid species. The trochanter of legs I–IV of ixodids may bear a ventral spur of variable shape and size, and among species of Dermacentor a variably developed retrorse spur also occurs on the dorsal surface of trochanter I; the latter process is called the “dorsal horn” in some works. The coxae and trochanters are generally unarmed in argasid ticks. The tarsus of leg I of all ticks possesses a complex of sensory structures in a posterior capsule and an anterior pit or depression, which together form Haller’s organ (Fig. 17) on the dorsal subapical surface of the segment. Details of the microanatomy and fine structure of Haller’s organ, when viewed under a scanning electron microscope, have been used as an aid in distinguishing between species of Ixodes (Homsher and Sonenshine 1975); however, these attributes are generally not visible when examining specimens under more limited magnifications of stereomicroscopes using incident light. In all ticks, tarsi II–IV consist of a shorter basal metatarsus, or basitarsus, which is delineated by a circumsegmental fissure from a longer apical telotarsus. The length and shape of these elements offer useful diagnostic attributes for some taxa of ticks. In lateral view, the telotarsus in some species tapers gradually and has an even profile to the apex; in others, the dorsal and ventral surfaces are parallel proximally but are humped or abruptly sloped dorsoapically. 22 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

coxa I external spur

internal spur coxa II pregenital plate

genital aperture coxa III

coxa IV median plate

spiracular plate

epimeral plate

adanal plate anus anal groove anal plate

Fig. 11. Diagrammatic attributes of body venter of female Ixodidae. Redrawn from Brinton et al. (1965). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 23

Ph1

st1

st2 st3

Pm1 Pa1 Jv3 Zv1

Pa2 Jv5 Pm2 Zv2

Mv1 Pm3 Sv1 Mv2 Zv5 Pm4 Zv3 Sv2 Mv3 Sv3 Mv4 Sv4 Mv5 Sv5

Fig. 12. Setal notation alternatives for body venter of larval Ixodidae (see also Table 1). Redrawn from Clifford et al. (1961). 24 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

anal groove extending anteriorly around anus

anal groove never extending anteriorly around anus

Fig. 13. Diagrammatic attributes of anal region of Ixodidae. A. Anal groove in Ixodes spp. B. Anal groove characteristic of other genera of Ixodidae in Canada. Redrawn from Keirans and Litwak (1989). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 25

length

dorsal prolongation

goblet

pore of goblet width

macula orifice peripheral perforations

frame

Fig. 14. Spiracular plate attributes of adult and nymphal Ixodidae. Redrawn from Cooley (1938). 26 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

In some argasids, this subapical dorsal protuberance is distinguished from the presence or absence of one to several additional dorsal elevations, called dorsal humps, located more proximally on the telotarsus and sometimes on the basitarsus (Fig. 15). In some adult ixodids, e.g., Rhipicephalus, the ventral surface of tarsi II–IV has one or two projections designated as the terminal and subterminal ventral tarsal spurs. Even more so than on the palps, the number and arrangement of setae on some of the segments of the three pairs of legs of larval ticks are important diagnostic characters (e.g., Figs. 65, 84). The notation applied to the leg setae is that proposed by Evans (1963a, 1969) for mesostigmatic mites and applied to larval Ixodidae by Edwards and Evans (1967) and to larval Argasidae by Edwards (1975) and Klompen (1992). Setae are denoted by their position on the anterior (ad) and posterior (pd) dorsal faces, anterolateral (al) and posterolateral (pl) faces, and anteroventral (av) and posteroventral (pv) faces of the segments (Fig. 18). This scheme of notation generally is not applied to the more complex complement of setae and clusters of sensilla of Haller’s organ on tarsus I. It is also not applied to the leg setation of nymphal and adult ticks, as the complements of setae become much higher on segments and do not lend themselves to being readily assigned a notation that implies homologies of structures. Coxae I–III of argasid larvae consistently bear two setae (av and pv) each, but a third ventral seta is generally present on coxa I, and sometimes on coxae II and III, of ixodid larvae. The absence or presence of the third seta on coxae II and III is a useful, though variable, diagnostic larval attribute among species of Ixodes. Specific setal complements for the coxae are given as simple formulas; for example, 3-2-3 for I. muris indicates three setae on coxa I, two on coxa II, three on coxa III. The larval setal patterns for trochanters I–III are uniform among all species of Argasidae and Ixodidae in Canada, so they are not useful diagnostically: trochanter I bears 4 setae (ad, av, pv, pl), trochanter II bears 5 (al, ad, av, pv, pl), and trochanter III bears 4 (al, ad, av, pv). The larval setal patterns for femora I–III are constant in most species of Ixodidae, and include 10 setae each on femora I and II (al, ad-1, ad-2, av-1, av-2, pd-1, pd-2, pv-1, pv-2, pl) and 9 on femur III (pl absent) (Fig. 18). In contrast, larval argasids do not have this full complement of femoral setae, and show a variety of losses that may be useful diagnostically. Larvae of Argas reflexus have the same full complement of 10 setae on femur I, but they have one less ventral seta (pv-2 absent) on femora II and III than in ixodids. Those of other argasid species deviate in a variety of ways from the ixodid pattern, as discussed by Edwards (1975) and Klompen (1992). For example, larval Ornithodoros kelleyi have eight setae on femur I (av-2, pv-2 absent), nine on femur II (pv-2 absent), and seven on femur III (av-2, pv-2 absent). The larval setal patterns for genua I–III are constant in Ixodidae and some Argasidae, and include the same eight setae (al, ad-1, ad-2, av, pd-1, pd-2, pv, pl). Larvae of some argasid taxa diverge from this pattern in having either a lower or rarely a higher number of genual setae than usually is common to all three legs. For examples (Edwards 1975), Otobius megnini and O. lagophilus have six setae on each of genua I–III (ad-2, pd-2 absent), and Ornithodoros kelleyi has five setae on each of genua I–III (ad-2, pd-2, pv-1 absent). The larva of Argas reflexus is unusual in having 10 setae on genu I (al-2, av-2 added) but nine on genua II-III (al-2 added) (Klompen 1992). The larval setation of tibiae I–III offers perhaps the most interesting patterns for diagnosis of species and higher taxa in both Ixodidae and Argasidae. The maximum complement in Ixodidae occurs in the genus Ixodes, and includes the same eight setae as for the genua of legs I–III (al, ad-1, ad-2, av, pd-1, pd-2, pv, pl). Larvae of the ixodid A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 27

subapical dorsal protuberance

telotarsus tarsus basitarsus

dorsal humps

tibia

genu

femur

trochanter

coxa

Fig. 15. Diagrammatic structure of leg I of adult Argasidae. Redrawn from Cooley and Kohls (1944). 28 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

pulvillus

claw

pretarsus

ad1 pd1

al1 pl1 1st setal vertical dorsal lyrifissure av1 pv1

ad2

al2 pl2

2nd setal verticil pd2 av2 pv2

telotarsus

av3 pv3 intercalary sclerite

pl3 al3 3rd setal verticil ad3 pd3

basitarsus

Fig. 16. Diagrammatic structure and setal notation of tarsi II and III of larval Ixodidae, dorsal aspect. Redrawn from Klompen (1992). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 29

pulvillus

claw

pretarsus

al1 pl1 pv1 av1 dm1

al2 pv2 pl2 av2 pd2

ad2 anterior pit Haller’s organ av3 posterior capsule pv3 al3 pd3 pl3 ad3 dm2 pd4 ad4 ventral lyrifissure

Fig. 17. Diagrammatic structure and setal notation for tarsus I of larval Ixodidae, dorsal aspect. Redrawn from Klompen (1992). 30 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

dorsal ad1 pd1 posterior

al1 pl1 anterior av1 pv1

ad2 pd2 ventral

ad pd al2 pl2

dorsal av2 pv2 posterior ventral seta al pl

anterior dorsal seta

anterior lateral seta ventral posterior lateral seta av pv

Fig. 18. Diagram of surfaces, setal positions and notation for a leg segment of larval Ixodidae. Redrawn from Edwards (1975). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 31

genus Haemaphysalis diverge from this pattern in having seven setae on tibiae I–III (pd-2 absent), while those of the ixodid genera Dermacentor, Rhipicephalus, and Amblyomma differ further from this pattern in having seven setae on tibia I (pd-2 absent) and only six setae on tibiae II-III (ad-2 also absent) (Edwards and Evans 1967). The basic setal pattern of the tibiae in larval Argasidae, which occurs in some species of Argas, is the same as the eight setae found on all legs in Ixodes. However, many argasids, including species of Ornithodoros and Otobius, have only six setae (ad-2, pd-2 absent) on tibiae I-III. Rarely, an addition of one or two setae to the basic complement of eight may occur on the tibiae in scattered species of Argasidae; these may involve any of setae ad-2, pd-2, al-2, pl-2, av-2, pv-2, as discussed by Klompen (1992). The larval setation of tarsus I (Fig. 17) is not readily used diagnostically for species of ticks, due to the number and complexity of structures present, to difficulties in discerning some of them readily, without access to scanning electron microscopy, and to lack of a reliable notation for structures, based on homologies. A comparative discussion of attributes of setae and other structures on tarsus I, which notes family and some genus level differences between larval ticks, is given by Klompen (1992). Surface features of Haller’s organ, as viewed with scanning electron microscopy, have been useful as an aid in distinguishing between adults of species of Ixodes, and determining subgeneric levels of classification in this genus (Homsher and Sonenshine 1975; Homsheret al. 1991). The larval setation of tarsus II (Fig. 16) is the same as that of tarsus III, and shows some diagnostic patterns that distinguish Ixodidae and Argasidae. Ancestrally, these patterns include three whorls, or verticils, of up to six setae each. The six setae in a given whorl are denoted al and pl laterally, ad and pd dorsally, and av and pv ventrally. The notation is applied such that the most distal whorl is the first whorl; thus,ad-2 denotes the anterodorsal seta of the second whorl, while pv-3 denotes the posteroventral seta of the most basal, third whorl. Larval Ixodidae display a full complement of three whorls, each with six setae, for a total of 18 setae, while the setal complement in larval Argasidae is generally reduced, variably so among genera (Klompen 1992). The dorsal setation in Ixodidae presents six setae, including a distal pseudosymmetric “pair” ad-1, pd-1, a median “pair” ad-2, pd-2, and a proximal “pair” ad-3, pd-3. In some genera of Argasidae (e.g., Carios, Ornithodoros, Otobius), an unpaired dorsomedial seta, dm, is present mid-dorsally. This seta is not added until the nymphal instar in genera of Ixodidae and others of Argasidae. Dorsal seta ad-2 is absent in larvae of some genera of Argasidae, e.g., Ornithodoros, while setae pd-2 and ad-3 are absent in larvae of Otobius (Klompen 1992). The lateral and ventral patterns of larval setae in Ixodidae also present six setae each, but the proximal “pair” of ventrals, av-3, pv3, is inserted on a small intercalary sclerite in the region of the circumsegmental fissure, instead of on the basitarsus, which bearsal-3, pl-3 as well as ad-3, pd-3. The lateral and ventral setation is reduced in most larval Argasidae, except for some species of Argas, because of the absence of setal “pairs” al-3, pl-3 and av-3, pv-3; however, in a few species (e.g., Argas persicus), an additional ventral seta, denoted avx, is present on the basitarsus (Klompen 1992). Absences of other setae among some taxa of Argasidae that are not found in Canada, as well as variation in relative positions of tarsal setae in Argasidae, are discussed by Klompen (1992). The position of setae ad-2, pd-2 relative to the dorsal lyrifissure on the telotarsus is another diagnostic larval attribute notable among ticks in Canada. This “pair” of setae is inserted proximal to the lyrifissure among all genera of Ixodidae, but only among some of Argasidae, such as Argas. In some other argasid genera, including Carios, these setae are inserted distal to the lyrifissure, while in yet others, includingOrnithodoros , pd-2 is distal to the lyrifissure whilead-2 , as noted above, is absent (Klompen 1992). 32 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Table 2. Comparison of idiosomal setal notation systems for larval ticks. Notations in parentheses indicate setae infrequently expressed.

Idiosomal dorsum Idiosomal venter Klompen Clifford and Anastos Klompen Clifford and Anastos (1996) (1960) (1996) (1960) j3 Sc3 st1, jv2 St1 j5 Cd1 st2, jv3 St2 J2 Cd2 st3, jv4 St3 (J3) --- Jv3 Pa1 (J4) --- Jv5 Pa2 (J5) --- z2 Sc1 Zv1 Pm1 z4 Sc5 Zv2 Pm2 (z6) --- Zv3 Pm3 (Z1) --- Zv4 --- Z2 S1 Zv5 Pm4 (Z3) --- (Z4) --- (Z5) --- s2 Sc2 Sv1 Mv1 s3 Sc4, Md1 Sv2 Mv1, Mv2 s4 Md1, Md2 Sv3 Mv2, Mv3 s5 Md2 Sv4 Mv3, Mv4 s6 Md3 Sv5 Mv4, Mv5 S1 Md4 S2 Md4, Md5 S3 Md5, Md6 S4 Md6, Md7 S5 Md7, Md8

Collecting, preserving, and studying ticks

Whether for biodiversity studies or disease surveillance or general enquiry, there are several standard methods that can be used to determine the number and types of ticks in a given area. These include tick drags, tick flags, tick walks, dry ice (carbon dioxide, CO2) traps, and tick collections from the bodies, burrows, or nests of host animals. Whichever method is chosen, it should be applied consistently if there is need to ensure that different data sets are statistically comparable. Also, different tick species and instars are collected disproportionately by the various methods, so it may be necessary to use more than one method in order to develop a complete assessment of the tick fauna of an area. Tick drags, in which a piece of cloth is passed over or around areas where ticks are questing, collect representative samples of the ixodid ticks present and more or less mirror the actual exposure that a person might experience in a given area. This technique is labour intensive, yielding few ticks in areas of low to moderate tick density. However, it is ideal as a A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 33

quick “spot check” of tick activity and is particularly useful when surveying for adult Ixodes scapularis, which may take up to an entire day to move toward dry-ice traps. A tick drag can be made from a sheet of soft white material, such as muslin or flannel, and is usually about 1 m long by 1 m wide. Stapling a 1.2 m dowel to the leading edge serves to keep the sheet spread open as it is pulled over vegetation and small obstacles. A 2-m cord is attached to both ends of the dowel to form a loop, which is used to pull the drag. A second dowel is sometimes attached to the trailing edge of the drag, but this may become ensnared in dense vegetation or may limit the ability of the fabric to conform to the contours of the area being surveyed. When ticks are questing close to the surface of the ground or in dense vegetation, a tick flag will sometimes produce better results than a drag. A flag is made by attaching a piece of cloth to a stick or dowel so that it resembles a flag. The flag is then waved slowly back and forth under, in and around vegetation or leaf litter, taking advantage of those areas where ticks are most likely to quest for their preferred host. For example, because immature I. scapularis chiefly infest small mammals, they are more likely to be picked up by low flagging than by dragging over the tops of plants. Also, immatureI . scapularis tend to remain in the leaf litter around a dry-ice trap, rather than climb onto the trap itself, and can therefore best be collected by flagging the trap area. Of course, anyone pulling a tick drag is, in effect, engaged in a tick walk. Ticks attaching to a person walking in a prescribed area provide the best estimate of the tick threat to humans (Chapman and Siegle 2000). To conduct a tick walk, wear white, 100% cotton clothing (pants and shirt, or coveralls, and socks) to highlight any ticks encountered. Pants should be bloused into socks or boots. If live ticks are required to test for pathogens, repellent should not be used, but openings in outerwear should be sealed with tape to prevent tick entry and attachment. For many tick species, dry-ice traps yield the most specimens in terms of labour expended.

This technique capitalises on the ability of ticks to sense CO2 and move toward its source.

Sensitivity to CO2 varies among tick species. Thus, Amblyomma americanum is attracted to

CO2 to a greater degree than Dermacentor variabilis, even though more specimens of the latter can be collected by this method than by dragging, per person-hour. Ixodes scapularis is also attracted to CO2, but since 12–24 hours of trapping may be required to attract these relatively slow-moving ticks, dragging or small mammal checks are generally considered better methods for sampling this species. To construct a CO2 trap, simply place some dry ice in a vented, insulated container and set the container in the centre of a sheet on the ground (see Eads et al. 1982). If the trap will not be monitored, tape can be attached, sticky side out, on the perimeter to capture attracted ticks. A half pound (0.23 kg) of dry ice will last about two hours at 27 ºC (80 ºF) in an insulated container. Many variations on this theme have been developed, including traps designed to collect ticks over a seven-day period using a 12-kg block of dry ice, and traps with tubing to sample argasid ticks in burrows or tree cavities. Carbon dioxide gas cylinders with regulated release valves may be used in place of dry ice. Host trapping is perhaps the best method of assessing local tick populations, particularly if host nests can be sampled and their contents extracted with Berlese-Tullgren funnels. In the case of burrowing mammals (e.g., field mice), far more ticks will be found in nests than on the host animals themselves. However, regardless of the collecting method used, live, unengorged ticks that are to be identified using morphologically-based keys should be killed by immersing them in water that has been brought almost to a boil. This technique causes the legs and mouthparts structures to straighten so that they do not obscure other features. Each tick collection (all the ticks taken from a single host at a given locality and time) should be transferred to a labelled, rimless Pyrex® test tube of 70% ethanol (the alcohol is 34 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins changed after one week if the specimens are numerous or engorged) that is then stoppered with cotton, inverted, and placed in a larger (16 oz. or about 0.47 L), wide-mouth glass museum jar, which is also filled with 70% ethanol. To avoid chipping the mouths of the test tubes, line the jar’s bottom with cotton. The cap of the jar should be made of durable plastic, preferably with reeding, and lined with a polyethylene insert to prevent evaporation. Using dichotomous keys, tick nymphs and adults can usually be determined to species simply by removing them from their collection tubes, patting them down with a tissue so that they are moist but not wet (thereby eliminating glare), and viewing them through a stereoscopic microscope capable of magnifying images 60–90 times. As a general rule, only larval ticks are mounted on microscope slides. Because larvae are smaller and routinely examined by compound transmitted light microscopy, which requires a high degree of transparency, their opaque internal tissues are first macerated by immersing the larvae in lactophenol (see Richards (1964) for formula), an acid corrosive that does not weaken the integument as do basic corrosives (e.g., potassium hydroxide, KOH). After several days, cleared tick larvae may be mounted in Hoyer’s medium, or some equivalent aqueous mountant, as follows: Remove specimens to a white porcelain crucible and wash in 3–4 changes of 70% ethanol until the lactophenol-ethanol interface disappears. Using the clean glass rod of a balsam bottle, place a drop of mountant in the center of a 2.6 x 7.6 cm microscope slide. Lift a specimen from the crucible with a fine (0000) camel’s-hair brush and transfer it to one of the tines of a pair of jeweller’s forceps. Touch the tine to the mountant and press the specimen to the bottom of the droplet, arranging it on a vertical axis with the gnathosoma facing the preparator. By convention, larvae are mounted ventral side down (against the slide); however, if several larvae of the same series are available, at least one should be mounted ventral side up. With a clean pair of angular dissecting forceps, pick up a 12-mm, 0-thickness circular coverslip at its rim, apply the opposite edge to the rim of the droplet of mountant, and gently lower the coverslip into place. Final orientation of the specimen may be accomplished by lightly prodding the coverslip surface with a probe. Place the slide in an oven set at 45 ºC for about 1–2 weeks. Heat-treated slides are allowed to cool to room temperature for several hours or days. A ring of glyptal (presently available under the name Red Glpt insulating varnish from GC Electronics, Rockford, Illinois, United States of America) is then used to seal the edges of the coverslip to the slide surface (Travis 1968; Wu 1986). A first coat may be applied using a number 6 sable brush, but to assure an impervious seal, a second coat is applied with a somewhat smaller (number 4 or 5) brush after the first ring has dried. Alternatively, the sealant may be applied with a small, narrow-nosed polyethylene bottle, which has the advantages of quickly producing a thicker, more uniform coating and minimizing both the user’s exposure to the toxic fumes of volatile glyptal and the waste of unused glyptal exposed to air (Wu 1986). At this point, the slides are ready for labelling and incorporation into the collection. Scanning electron microscopy (SEM) is now universally employed in morphological and systematic studies of ticks and other mites (Robbins and Keirans 1992). By virtue of their size and structure, ticks in any life history stage are ideal subjects for SEM, as are most larger, well-sclerotised mites (e.g., Oribatida, many Gamasina). The principal advantages of SEM are increased magnifying power, resolution, and enhanced depth of field, whereas its disadvantages include the much greater cost of the instrument and space needed for housing it, the sacrifice of specimens used for such imaging, and the need and space for special storage of prepared specimens. Though somewhat labour-intensive, the techniques involved in preparing specimens for SEM are relatively easy to learn and fall into four categories: cleaning, drying, mounting, and coating. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 35

Only the best specimens – but not primary types! – should be selected for SEM, and these should be thoroughly cleaned of any dirt adhering to the integument. Though detergents and ultrasound will remove larger particles (Keirans et al. 1976), the most efficient cleaning method presently available is the “glue technique” of Corwin et al. (1979). Ticks to be cleaned are removed one at a time from the 70% ethanol in which they are normally stored, placed on a piece of blotting paper approximately 5 x 10 cm in size, and observed under a binocular dissecting microscope while the integument gradually dries. When working with adult or nymphal ixodid ticks, it is a good idea to remove the two hindmost legs on one side of the body in order to expose one of the spiracular plates. The dry specimen is glued to the blotting paper, ventral side up, using a small amount of multi-purpose glue thinned with a few drops of an appropriate solvent (e.g., acetone). An artist’s brush is then used to coat all capitular, body, and leg surfaces with an even layer of glue. When the glue is almost dry, it is carefully peeled away with two pairs of fine forceps. If the glue becomes overly dry, it may be softened by applying minute quantities of 95% ethanol held between the tines of a forceps. Concentrated ethanol is also used to release the specimen from the blotting paper. Usually the dorsum will be cleaned by the glue used for attachment. However, if the dorsal surface is still dirty, the specimen may be reattached, ventral side down, and the cleaning process repeated. Most specimens should not be cleaned more than twice because excessive handling loosens intersegmental membranes. Weakened structures will later vibrate and “charge” under an electron beam. Cleaned specimens are held overnight in reagent-grade acetone in order to dissolve any remaining glue and to remove internal fluids. However, specimens to be examined in the high vacuum chamber of a scanning electron microscope must be absolutely dehydrated. This is best accomplished using a critical point dryer (CPD). It is well known that liquids will pass to the gaseous phase at a specific temperature and pressure called the critical point. When this point is reached, no phase boundaries exist, the density of the gas equals that of the liquid, and surface tension is zero. Thus, it is possible to completely dehydrate a biological specimen without exposing it to the tremendous surface tension forces that accompany ordinary evaporation. Such forces can easily distort acarine integument. A CPD with side viewing capability (e.g., the Ladd 38000; Ladd Research, Williston, Vermont, United States of America) is especially suitable for acarological work. Specimen baskets may be purchased or homemade from acetone-resistant materials. The transitional fluid used in CPDs is ultrapure, bone-dry liquid CO2, which is sold by most vendors of bottled and compressed gases. Any

CO2 tank that is to be connected to a CPD must be equipped with an internal siphon or dipping tube that will permit liquid CO2 to enter the CPD specimen chamber readily while the tank is in an upright position. A high pressure, high purity liquid-gas analyser filter, such as the Balston type 95S6 (Parker Hannifin Corp., Haverhill, Massachusetts, United States of America), should be installed in the line between the tank and the CPD. Dried specimens may be mounted immediately on aluminum SEM stubs or stored over Drierite (calcium sulfate, CaSO4) in a desiccator. Specimens are best manipulated with one of the microvacuum systems available from most electron microscopy supply houses. To mount larger specimens, a small drop of silver-based conducting paint is placed at the centre of the stub and observed under a binocular dissecting microscope as it begins to congeal. Just as the paint surface jells, one or both of the hind legs of the specimen are pushed into the drop and the specimen is raised to a vertical position where it is held until the paint is almost dry. In the event of an accident, conducting paint can easily be removed with acetone or thinned with n-butyl acetate. Though this process may seem tedious, an upright position permits views of both the dorsal and ventral specimen surfaces and provides an attractive black background for contrast during photography. Very small 36 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins specimens, such as larvae, should be mounted on adhesive transfer tape (3M Company, St. Paul, Minnesota, United States of America) applied directly to the stub surface. Conducting paint may be used to seal and protect the margins of the adhesive. Mounts that have dried are layered with gold or a gold-palladium alloy in a sputter coater. From an electron microscopist’s perspective, most ticks and many other mites are relatively large, and their features will not be obscured by what some would regard as excessively thick layers of gold (> 1000 Å). In fact, extra coating may reduce or eliminate charging. Also, with very large specimens, sputter coaters do not coat evenly; therefore, it may be necessary to coat in cycles, changing specimen orientation between coats. Finished mounts are extremely delicate and should be stored in a specially designed cabinet where they will be protected from dust, moisture, and sudden changes in temperature.

Medical and veterinary importance of ticks in North America Ticks are of medical and veterinary significance not only because of the disease-causing pathogens they transmit, but because they can induce diseases of non-microbial etiology. Ticks have emerged second only to mosquitoes as the most important group of medically associated arthropods transmitting a diverse range of pathogens. Tick-borne pathogens are believed to be responsible for more than 100 000 cases annually of illness in humans throughout the world. Ticks feed on every class of terrestrial vertebrate and are the most important vectors of disease-causing pathogens in domestic and wild animals (de la Fuente et al. 2008). Since ticks comprise one of the groups of parasitic arthropods whose numbers and geographic range are expected to increase in consequence of global warming, there is likely to be an increase in those infections for which ticks act as vectors (Grist 1992). Ticks are capable of transmitting microbes by several routes including salivary secretions, coxal fluids, regurgitation, and faeces. The role of ticks as transmitters of infectious disease- causing agents is of particular significance with about 40 tick species that are of medical and veterinary importance and up to 40 other species that have some involvement in human and animal disease epidemiology (Hoogstraal 1985). Contributing biological factors to the high vector potential of ticks are their life cycles and characteristic properties of their salivary secretions and blood digestion. Hoogstraal (1985) stated that ticks are carriers of more kinds of micro-organisms than any other single arthropod taxon, including mosquitoes. These microorganisms may be of bacterial (spirochaetes, rickettsiae, Francisella Dorofe’ev species), viral (flaviviruses, coltiviruses, Nairovirus), or protozoan (Babesia Starcovici species) etiology. Diseases of non-microbial etiology caused by ticks include anaemias caused by the blood-feeding activities of ticks, tick paralysis caused by salivary secretions of some long- feeding tick species and inflammatory or hypersensitivity reactions and immunosuppression caused by components of the saliva of the tick. An extensive review of all known tick-transmitted diseases is beyond the scope of the manual. This chapter deals exclusively with diseases of human and veterinary health importance encountered in North America, with an emphasis on diseases recognised in Canada. It is worth noting that many new tick-borne zoonoses are emerging with increasing frequency globally and within North America (Vasconcelos and Calisher 2016). Heartland virus (Savage et al. 2013) and Bourbon virus (Kosoy et al. 2015) are recently described human pathogens associated with lone star tick, and Borrelia mayonii Pritt et al. in blacklegged ticks in the Midwest (Pritt et al. 2016) are two examples of this. Clearly readers of this handbook will need to consult the current literature to be aware of new, emerging tick-associated pathogens. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 37

Diseases of microbial etiology

Bacterial diseases Lyme borreliosis Lyme borreliosis, caused by bacteria of the Borrelia burgdorferi Johnson et al. sensu lato complex (Spirochaetales: Spirochaetaceae), is the most common vector-borne disease in North America. Borrelia burgdorferi is transmitted by ixodid ticks which feed on the wildlife reservoir hosts for these bacteria, particularly rodents and birds, as well as hosts that are not reservoirs of B. burgdorferi, such as deer. Ixodes scapularis, the blacklegged tick, is the main vector in eastern and central North America and I. pacificus, the western blacklegged tick, is the main vector in the west. Both tick species are indiscriminate in their choice of host and will feed on humans when the opportunity arises (Ogden et al. 2008c). Natural reservoirs for B. burgdorferi include the white-footed mouse, Peromyscus leucopus (Rafinesque), other small mammals and a variety of ground-foraging birds (Bouchard et al. 2011). The larvae or nymphs of the tick feed on the white-footed mouse and become infected. Adult ticks, or more commonly, nymphs, may then infect humans. Infected nymphs must remain attached for 36–48 hours or longer, and infected adult ticks must remain attached for 48 hours or longer, before the risk of transmission of B. burgdorferi becomes substantial (Bratton and Corey 2005). Diagnosis of Lyme borreliosis in humans can be complex and difficult (Sperling and Sperling 2009; Lindsay et al. 2014). Early Lyme borreliosis is generally characterised by a skin lesion, erythema migrans (EM), which expands further than 5 cm from the tick bite, accompanied by flu-like symptoms, arthralgias, myalgias, and fever. If untreated, the infection can progress to early disseminated Lyme borreliosis with neurological and cardiac involvement. Late disseminated Lyme borreliosis includes central and peripheral neurological manifestations and Lyme arthritis (Ogden et al. 2008c; Hatchette et al. 2014). Lyme disease is treated with oral antibiotics, usually doxycycline or amoxicillin. Late or severe disease, particularly with cardiac or neurologic involvement, requires treatment with intravenous antibiotics. Late disease may be associated with treatment failures, but antibiotic treatment is curative in most cases of early Lyme borreliosis (Bratton and Corey 2005; Hatchette et al. 2014). Approximately 30 000 cases of Lyme borreliosis are reported annually in the United States of America, although the actual number of cases diagnosed each year is believed to be approximately 300 000 (http://www.cdc.gov/lyme/stats/humancases.html). Most of these cases occur in the northeastern and northcentral states, many of which border Canada. From 2004–2006, only 69 potentially endemic cases of Lyme borreliosis were reported in central and eastern Canada (most likely transmitted by I. scapularis), although this is likely an underestimate of the true number of cases (Ogden et al. 2008c). In 2009, Lyme disease became a reportable disease in Canada. From 2009–2012, 813 cases were reported from across Canada, while it is estimated that over 500 cases were reported in 2013 from across the provinces and territories (http://www.phac-aspc.gc.ca/id-mi/lyme/surveillance- eng.php; Hatchette et al. 2014). It appears that established populations of tick vectors and natural endemic cycles of B. burgdorferi have a limited but expanding geographic scope in Canada, accounting for the relatively low reported incidence of Lyme borreliosis in Canada compared to the United States of America. Established populations of B. burgdorferi-infected I. scapularis have been identified in a number of locations in Ontario, including at Long Point peninsula on the north shore of Lake Erie, Point Pelee National Park and Rondeau and Turkey Point Provincial Parks, around Lake Ontario in the vicinity of the Thousand Islands National 38 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Park (Werden et al. 2014) and tick populations appear to be moving inland in much of eastern Ontario (Nelder et al. 2014). Established populations of infected I. scapularis have also been detected in parts of Québec, Nova Scotia, New Brunswick, and southern Manitoba (http:// www.phac-aspc.gc.ca/id-mi/tickinfo-eng.php). Thus, the distribution of known “endemic” areas for B. burgdorferi infection is limited in Canada but appears to be expanding (Leighton et al. 2012). Thus, the true geographic extent of reproducing I. scapularis populations in Canada is uncertain (Ogden et al. 2008d, 2014). However, the number of established populations of I. scapularis is increasing in Canada, the rate of spread is increasing (Ogden et al. 2013) and climate change is likely to accelerate this trend (Ogden et al. 2008c). Reproducing populations of I. pacificus are widespread in British Columbia. However, transmission cycles involving B. burgdorferi-infected I. pacificus are inefficient, and the prevalence of B. burgdorferi infection in host-seeking I. pacificus is usually much lower than that found in I. scapularis. In addition, I. pacificus is less likely to bite humans, and therefore, the risk of humans contracting Lyme borreliosis infections in I. pacificus-endemic regions is usually lower than in regions where I. scapularis populations are present. In 2001, 141 cases of Lyme borreliosis were reported in the coastal United States of America where I. pacificus is endemic. In British Columbia, approximately four cases are reported each year (Ogden et al. 2008c). From results of passive surveillance in Canada, there is a relatively low possibility of exposure for humans to vectors of Lyme borreliosis in all provinces, even in areas where established tick populations are unknown (Koffiet al. 2012). In this instance, exposure may occur from infected adventitious ticks that are carried by birds from endemic regions of the United States of America into populated areas of Canada (Ogden et al. 2008b, 2008d). Numerous clinical syndromes have been documented in domestic animals infected with B. burgdorferi, including limb and joint disease and neurologic, cardiac, and renal abnormalities. In dogs, lameness, fever, anorexia, lethargy, and lymphadenopathy with or without swollen, painful joints are the most common clinical symptoms (Appel 2005).

Southern tick-associated rash illness (STARI) Southern-tick associated rash illness (STARI), also known as Masters disease, is a Lyme disease-like illness characterised by an erythema migrans rash occurring after the bite of Amblyomma americanum. This Lyme-disease-like illness has been reported in the southeastern United States of America since the mid-1980s. The etiologic agent is not known but may be a Borrelia species, provisionally named Borrelia lonestari James et al. (Spirochaetales: Spirochaetaceae). This spirochaete has been detected in lone star ticks removed from humans and has been cultured from A. americanum (Varela et al. 2004); however, attempts to detect this agent by polymerase chain reaction (PCR) or culture from skin biopsies from patients have not been successful, except for detection of B. lonestari by PCR in one patient (James et al. 2001; Wormser et al. 2005), suggesting that other agents may be involved in the etiology of STARI. Although the exact etiologic agent is unclear, Borrelia burgdorferi sensu stricto is not the causative agent (Masters et al. 2008). Serious systemic complications are not currently recognised with STARI but treatment with doxycycline is recommended (Blanton et al. 2008; Masters et al. 2008).

Avian borreliosis Avian borreliosis, caused by Borrelia anserina (Sakharoff) Bergey et al. (Spirochaetales: Spirochaetaceae), affects a variety of birds, including geese, ducks, turkeys, and chickens in many parts of the world including North America. It is transmitted by Argas species, A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 39

especially Argas persicus, the “cosmopolitan” fowl tick, in tropical and warm temperate regions (Hoogstraal 1985). Infected Argas ticks transmit B. anserina both transstadially and transovarially. The disease begins with a high fever after an incubation period of 3–12 days. Symptoms are highly variable, depending on the virulence of the strain, and may be absent. Signs include listlessness, depression, moderate to severe shaking, and increased thirst. The birds may become cyanotic and have yellowish-green diarrhoea. Surviving birds recover after about two weeks and have long-lasting immunity against homologous but not heterologous strains (Bricker 2005). Avian borreliosis is treated with penicillin derivatives, streptomycins, and tetracyclines.

Tick-borne relapsing fever Tick-borne relapsing fever (TBRF) is caused by a group of approximately 21 different closely related Borrelia species of proven or possible pathogenicity, each associated with their own specific vectors of the genus Ornithodoros (Schwan and Piesman 2002). Important TBRF Borrelia in North America include: B. hermsii (Davis) Steinhaus, B. turicatae (Brumpt) Steinhaus, and B. parkeri (Davis) Steinhaus. The primary reservoirs of these Borrelia species are rodents, rabbits, and hares. These spirochaetes cause a systemic infection characterised by recurrent episodes of fever separated by afebrile periods. Influenza-like symptoms, arthralgias, dizziness, nausea, and vomiting are common during the symptomatic period. The treatment of choice is doxycycline; erythromycin may be used as an alternative treatment but can lead to a Jarisch-Herxheimer reaction (generalised malaise, headache, fever, sweating, rigours, seizure, or stroke), especially if given during the late febrile stage (Bratton and Corey 2005). Ornithodoros parkeri transmits B. parkerii, O. hermsi transmits B. hermsii, and O. turicata transmits B. turicatae. Ornithodoros hermsi and O. parkeri are found in the mountainous regions of western North America, whereas O. turicata prefers drier habitats and is found in underground burrows from Kansas to Texas, United States of America (Dworkin et al. 2002). Tick-borne relapsing fever is endemic in the western United States of America and southern British Columbia. From 1977–2000, 450 cases were reported in the western United States of America, with the majority of cases reported from California, Colorado, and Washington in the months of June to September. In Canada, locally acquired TBRF has been reported only in British Columbia, with O. hermsi as the likely vector of B. hermsii (Dworkin et al. 1998). Very few cases of relapsing fever (fewer than 20 in total) have been documented in British Columbia since the first cases were identified in 1933. However, this infection is likely misdiagnosed as Lyme in many patients and underreported due to a lack of awareness on the part of many physicians (Banerjee et al. 1998; Dworkin et al. 1998). Ornithodoros species have nocturnal feeding habits; these ticks feed rapidly and their bites are usually painless. Tick-borne relapsing fever often occurs in the spring and summer months; however, unlike many other tick-borne infections, TBRF also occurs during the winter. The winter occurrence of TBRF results from humans entering tick-infested dwellings and often results in the familial clustering of cases. For instance, when humans enter cabins during the winter, Ornithodoros can be stimulated by the warmth and carbon dioxide produced by the humans to seek out and feed on the occupants. Ornithodoros can survive for extended periods, even years, in the absence of a blood meal (Dworkin et al. 2002). Transovarial transmission of Borrelia is frequent and efficient in Ornithodoros. Therefore, the longevity of these ticks and their transovarial transmission allow persistence of the spirochaetes in an environment for years, despite long periods of absence or low numbers of suitable vertebrate hosts (Barbour 2005). 40 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Ehrlichioses “Ehrlichiosis” is a generic name for infections caused by obligate, intracellular bacteria in the family Anaplasmataceae, chiefly in the genera Ehrlichia Moshkovski and Anaplasma Theiler. Bacteria in this family have long been considered to be of veterinary importance. However, in the last two decades, three species have been implicated in human disease. Ehrlichia chaffeensis Anderson et al. is the agent of human monocytic ehrlichiosis (HME), Ehrlichia ewingii Anderson et al., the agent of human ewingii ehrlichiosis, and Anaplasma phagocytophilum (Foggie) Dumler et al. (formerly the HGE agent, Ehrlichia phagocytophila (Foggie) Dumler et al. and Ehrlichia equi Lewis et al.) is the agent of human granulocytic anaplasmosis (HGA) (Dumler and Bakken 1998). During 2001–2002, 1176 cases of HME and HGA were reported in the United States of America. However, these infections are likely underreported because of the non-specific nature of their clinical signs (Demma et al. 2005a). The obligate intracellular bacteria that cause these infections are maintained through enzootic cycles between ticks and animals. Amblyomma americanum is the major vector for Ehrlichia chaffeensis and Ehrlichia ewingii, while I. scapularis and I. pacificus are the vectors for A. phagocytophilum (see below for details of HGA). Transovarial transmission of these agents in ticks appears to be inefficient, and consequently, mammals are presumed to play a major role in the maintenance and propagation of these bacteria in nature. The white-tailed deer (Odocoileus virginianus (Zimmermann)) is the sole vertebrate species recognised as a complete and sufficient host for maintaining the transmission cycle of E. chaffeensis. White-tailed deer are an important source of blood for adult and immature instars of A. americanum. The greatest influence on the emergence of ehrlichioses in the United States of America has been the explosive growth of white-tailed deer populations (Childs and Paddock 2003). Because white-tailed deer are a preeminent host for all three instars of A. americanum, a parallel increase in the abundance of lone star ticks has accompanied the dramatic rise in white-tailed deer numbers in the United States of America. Lone star ticks are also aggressive nonspecific feeders and all three instars will bite humans. Dogs also serve as hosts for all instars of A. americanum and can serve as a competent reservoir host for E. chaffeensis and E. ewingii. Little is known about potential reservoirs for E. ewingii other than domestic dogs, although it is likely that one or more species of wildlife are involved in the maintenance of this agent. All forms of human ehrlichiosis share certain clinical manifestations, including fever, chills, headache, malaise, and myalgia. Gastrointestinal symptoms including nausea, vomiting, anorexia, and abdominal pain may also occur. Rash is more common in HME than in the other ehrlichioses, but may only be present in up to one-third of patients. Human monocytic ehrlichiosis is a reportable infection in the United States of America with 417 cases reported in 2005 (Demma et al. 2005a). Human monocytic ehrlichiosis is considered the most severe of the ehrlichioses, with up to 60% of patients requiring hospitalisation and a mortality rate of approximately 3% (Demma et al. 2005a). The drug of choice for the treatment of all ehrlichial infections is doxycycline. Cases occur across the southcentral, southeastern, and mid-Atlantic states, corresponding to regions where white-tailed deer and lone star ticks both occur. Ehrlichia ewingii is also transmitted by A. americanum but much less is known about this pathogen, which was originally associated with disease in dogs (known as canine granulocytic ehrlichiosis). Human infections with E. ewingii were first reported in 1999 in the United States of America with fewer than 20 cases reported to date. This infection is most commonly diagnosed in immunosuppressed patients (Buller et al. 1999). There are A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 41

no known cases of ehrlichiosis contracted in Canada, although cases associated with travel to endemic regions of the United States of America have been diagnosed. This is likely because A. americanum is not established in Canada. Canine monocytic ehrlichiosis (CME) is a disease of dogs caused by Ehrlichia canis (Donatien and Lestoquard) Moshkovski (Rickettsiales: Anaplasmataceae). The primary vector of E. canis is Rhipicephalus sanguineus, the brown dog tick. Canine monocytic ehrlichiosis occurs in acute, subclinical, and chronic phases. Acute infection begins 8–20 days following an infective tick bite and can involve fever, anaemia, anorexia, ataxia, conjunctivitis, depression, and vomiting. The subclinical phase follows the acute phase and may last for 40–120 days or even years during which time dogs can remain persistently infected without clinical signs but with mild thrombocytopenia. The chronic stage resembles the acute phase with recurrent clinical symptoms which may be mild or severe. Doxycycline is effective in alleviating the clinical symptoms of CME but does not appear to clear the infection in all dogs successfully (Skotarczak 2003; Schaefer et al. 2007). Infection of dogs in Canada with E. canis is considered relatively rare, with a seroprevalence of 0.37% detected by Gary et al. (2006). However, seroprevalences for E. canis of 2.4%, 2.9%, and 6.4% have been found in Rhode Island, North Carolina, Virginia, Maryland, and Pennsylvania (Suksawat et al. 2000; Hinrichsen et al. 2001; Duncan et al. 2005), so this infection is generally more common in the United States of America, consistent with the geographic distribution of R. sanguineus. A new pathogenic Ehrlichia species has recently been described from patients in Minnesota and Wisconsin, United States of America (Pritt et al. 2011). The etiological agent is most closely related to Ehrlichia muris Wen et al. and because the taxonomic placement of this new agent is uncertain, it has been referred to as the E. muris-like agent. Blacklegged ticks have been found infected with this agent in recent studies (Pritt et al. 2011) and in tick specimens collected historically (Telford et al. 2011), hence this pathogen has been circulating in this region unrecognised for several decades. The disease in humans is similar to that caused by A. phagocytophilum (see next section) with all patients having fever, malaise, headache, and lymphopenia and all recovered fully after treatment with doxycycline (Pritt et al. 2011). Disease as a result of exposure to infected ticks has also been described in dogs (Hegarty et al. 2012). Blacklegged ticks from Manitoba have been screened for this pathogen but infected specimens were not detected; however, a single female Ixodes muris was infected with a variant of the E. muris-like agent (L.R.L., personal observation). Further research is required to define the ecology of this newly recognised species and its variants in Canada and elsewhere in North America.

Human granulocytic anaplasmosis (HGA) Anaplasma phagocytophilum, the causative agent of human granulocytic anaplasmosis (HGA), is transmitted primarily by I. scapularis and I. pacificus. Because these are the same vectors that transmit Borrelia burgdorferi and Babesia microti (Tyzzer) (Piroplasmida: Babesiidae), coinfections with two or all three of these pathogens are possible. The first case of clinically recognised HGA was described in the United States of America in 1990 (Chen et al. 1994). From 1997–2001, 1088 confirmed or probable cases were reported in the United States of America. Most cases were reported from the northeastern and midwestern regions of the United States of America. The HGA agent was initially detected in Canada in I. scapularis collected at Long Point but has subsequently been demonstrated in established populations of I. scapularis and adventitious ticks in other parts of Canada (Drebot et al. 2001; Ogden et al. 2008b; 42 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

L.R.L., personal observation). The first reported human case of HGA in Canada was acquired in Alberta in 2009, presumably due to an adventitiously-introduced tick, as locally reproducing populations of I. scapularis are not known to exist in Alberta (Parkins et al. 2009). Granuolcytic anaplasmosis was also reported in a dog from British Columbia and three dogs from Saskatchewan with fever, lethargy, inappetence, vomiting, diarrhoea, and lameness (Lester et al. 2005; Cockwill et al. 2009). A case of anaplasmosis in a horse from Saskatchewan has also been described (Burgess et al. 2012). Human anaplasmosis appears most commonly as an undifferentiated febrile illness occurring in the spring and summer. Symptoms include high fever, rigours, generalised myalgias, severe headache, and malaise. Anorexia, arthralgia, nausea, and a non-productive cough are also frequent but a rash is rarely reported. Fatal cases have been documented, but the case fatality rate is estimated to be only about 1%. Doxycyline is the treatment of choice for HGA. Like Ehrlichia chaffeensis, the white-footed mouse is considered the most important reservoir for A. phagocytophilum. The bacterium is transmitted transstadially but not transovarially in Ixodes species; therefore, mammals are critical for maintaining this species in nature. The prevalence of nymphs infected with A. phagocytophilum varies widely, but typically is in the range of 3–25% (Goodman 2005). While the white-footed mouse is likely the dominant small-mammal reservoir, infection is usually transient, unlike the chronic infection observed with B. burgdorferi (Goodman 2005). White-tailed deer are important for the maintenance of Ixodes populations, but the role these large mammals play in the life cycle of A. phagocytophilum is unclear (Massung et al. 2005; Tate et al. 2005; Rainwater et al. 2006). White-tailed deer may be seropositive and/or PCR positive for A. phagocytophilum (Bouchard et al. 2013). The pathogen exists as biologically and ecologically distinct subpopulations that are adapted to specific reservoir hosts and tick species with varying capacities to infect and cause disease in humans and domestic animals (Foley et al. 2009; Scharf et al. 2011). In particular, the 16S rRNA gene variant (referred to as Ap-ha) is the only known A. phagocytophilum variant that has been isolated from humans in the eastern United States of America and it appears to be maintained in the P. leucopus–I. scapularis enzootic cycle. Distinct variants not found in humans have been isolated from deer, other ruminants, horses, dogs, and cats, suggesting only a subset of A. phagocytophilum variants cause disease in humans (Lappin et al. 2004; Lester et al. 2005; Parola et al. 2005a; Poitout et al. 2005; Morissette et al. 2009). More than one strain may be present in a region, and the relative proportion of each strain may vary regionally (Krakowetz et al. 2014). Canine anaplasmosis varies from a subclinical infection to an acute febrile illness accompanied by anorexia, lethargy, and potentially central nervous system dysfunction and lameness. Anaplasma phagocytophilum may cause a less severe disease in dogs than does infection with E. canis, E. chaffeensis, or E. ewingii (Lester et al. 2005). Anaplasma phagocytophilum has also been known as the agent of equine granulocytic anaplasmosis in the United States of America since 1969, when it was initially recognised as Ehrlichia equi, before reclassification into the genusAnaplasma . Infected horses develop high fever, depression, anorexia, limb edema, ataxia, and stiffness in gait (Madigan and Pusterla 2000).

Bovine anaplasmosis Bovine anaplasmosis is a haemolytic disease of cattle caused by Anaplasma marginale Theiler. Clinical symptoms include fever, anaemia, weight loss, abortion, lethargy, and often death in animals older than two years. Cattle that survive acute infection develop persistent infection and can serve as reservoirs for A. marginale by providing a source of A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 43

infective blood for ticks. Anaplasmosis was a federally reportable disease in Canada since 1969 after Canada’s first outbreak in Manitoba until 2015. Anaplasmosis has been recorded on several occasions in Canada: Manitoba in 1968, Québec in 1979, Saskatchewan in 1983, Ontario in 1996, and Saskatchewan in 2000, with all cases associated with beef cow-calf operations, with the exception of the 2000 outbreak, which occurred in bison (de la Fuente et al. 2003). Since 2008, anaplasmosis has also been detected in cattle in Saskatchewan and Manitoba (Howden et al. 2010). In the United States of America, A. marginale is enzootic throughout the southeastern Atlantic states, the Gulf Coast states and the midwestern and western states. Dermacentor variabilis and D. andersoni are vectors with intrastadial (Lysyk 2013), but not transovarial or transstadial transmission of A. marginale, and populations of both species in western Canada have been shown to be capable of transmission of this pathogen experimentally (Lankester et al. 2006). However, the most important vector of A. marginale worldwide is Rhipicephalus (Boophilus) microplus Canestrini, the tropical or southern cattle tick. Rhipicephalus microplus has been extirpated from the United States of America but is widespread in Mexico, including along the United States of America- Mexico border, making reintroduction and reestablishment of this tick vector a cause of concern in the United States of America (Guerrero et al. 2006).

Anaplasma ovis Anaplasma ovis Lestoquard (Rickettsiales: Ehrlichiaceae) is an intraerythrocytic pathogen of sheep, goats, and wild ruminants (but does not infect cattle). Ticks of the genus Dermacentor are the vectors of A. ovis in the western United States of America (de la Fuente et al. 2006, 2007).

Canine infectious cyclic thrombocytopenia Anaplasma platys Dumler et al. (Rickettsiales: Ehrlichiaceae), which causes canine infectious cyclic thrombocytopenia was first reported in the United States of America in 1978 (Harvey et al. 1978) and is enzootic in many parts of the United States of America. Anaplasma platys appears to parasitise platelets of dogs only and is thought to be transmitted by the brown dog tick, R. sanguineus (Martin et al. 2005).

Mycoplasma haemocanis Haemotrophic mycoplasmas are pleomorphic bacteria that attach to the surface and grow upon red blood cells in a wide variety of vertebrate animals. Mycoplasma haemocanis Kützing (Mycoplasmatales: Mycoplasmataceae) has been documented worldwide, including the United States of America and Canada (Messick 2003). The acute form of the disease is characterised by a rapidly developing anaemia, most often in immunocompromised or splenectomised dogs. Lethargy, weight loss, fever, and anorexia are also apparent in many infected dogs. The natural mode of transmission of M. haemocanis in the dog is unknown; however, it is believed that R. sanguineus serves as an important reservoir and vector of infection for dogs. Mycoplasma haemocanis has been experimentally transmitted to splenectomised dogs by R. sanguineus, and transstadial and transovarial transmission of M. haemocanis in this tick has been reported (Messick 2003).

Q fever Q fever is a zoonosis with a worldwide distribution caused by Coxiella burnetii (Derrick) Philip (Legionellales: Coxiellaceae). Infection in humans is often asymptomatic; symptomatic acute Q fever manifests primarily as a self-limited febrile illness, atypical 44 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins pneumonia, or granulomatous hepatitis, whereas endocarditis is the most common presentation of chronic Q fever. The incubation period of acute Q fever is typically 2–3 weeks, followed by an abrupt onset of severe fever, headache, fatigue, and chills in symptomatic patients. The aerosol route is the principal mode of acquisition of C. burnetii infection in humans (Maurin and Raoult 1999; McQuiston and Childs 2002; McQuiston et al. 2002). Doxycycline is used to treat both acute and chronic Q fever. The reservoir for C. burnetii includes many wild and domestic mammals (primarily sheep, cattle, and goats), birds, and arthropods, such as ticks. Infected domestic animals are usually asymptomatic but abortions and stillbirths may occur. Of particular relevance, infected domestic animals may shed high numbers of C. burnettii in urine, milk, and birth products. Although C. burnetii has been isolated from ticks, arthropod-borne transmission of Q fever in humans is unlikely to be significant. However, ticks more likely play a significant role in the transmission of C. burnetii among wild vertebrates, such as rodents, lagomorphs, and birds, and among domestic animal species. Over 40 species of ticks are naturally infected with C. burnetii, including Rhipicephalus sanguineus, Dermacentor andersoni, D. occidentalis Marx, Amblyomma americanum, Haemaphysalis leporispalustris, Ixodes dentatus, Ornithodoros coriaceus Koch, and Otobius lagophilus (Maurin and Raoult 1999). There are no published reports on the isolation of C. burnettii from ticks in Canada. However, organisms that reacted by direct fluorescence microscopy with sera produced to C. burnetti were demonstrated in D. variabilis and H. leporispalustris collected in Nova Scotia in 1980 (H.A., personal observation). Coxiella burnetii replicates in cells of the midgut of experimentally-infected D. andersoni. Infected ticks expelled heavy loads of highly infectious C. burnetii in their faeces onto the skin of the animal host at the time of feeding. As well, C. burnetii may infect tick ovaries, resulting in transovarial transmission to the tick progeny (Maurin and Raoult 1999). Q fever is one of the most important zoonotic diseases in Canada. Human infection due to C. burnetti has been reported from all provinces. In summary of reports from laboratories to the Canadian virus report programme from 1980–1992, there were 474 Q fever cases from across Canada during this time, i.e., an average of 36.5 cases per year. The largest numbers of cases were reported from Ontario and the Maritime provinces, principally Nova Scotia, which reported 301 and 103 cases, respectively. However, since the 1990s, reports of Q fever in Nova Scotia have declined to approximately 4–5 reported cases annually (Marrie et al. 2008). The largest number of cases now appear to occur in the province of Québec, consistent with the large number of sheep herds in this area of Canada (Dolce et al. 2003). Because Q fever is not a notifiable disease in Canada, the actual number of Q fever cases diagnosed annually is unknown. In the United States of America, Q fever became nationally reportable in 1999 with 255 Q fever cases reported from 2000–2004. It is presumed that many more cases of Q fever occur annually but likely go unreported because of the nonspecific nature of the clinical symptoms and lack of definitive early diagnostic tests (McQuiston et al. 2006).

Tularaemia Francisella tularensis (McCoy and Chapin) Dorofe’ev (Thiotricales: Francisellaceae), the causative agent of tularaemia, is maintained in the environment principally by various terrestrial and aquatic mammals such as ground squirrels, rabbits, hares, voles, muskrats, beaver, and other rodents (Hopla 1974). Francisella tularensis can be transmitted to humans through a variety of routes, including tick or insect bite, handling of infected animals (e.g., A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 45

rabbits), ingestion of infected water, food or soil, and inhalation of infectious aerosols (Goethert et al. 2004). Tularaemia causes a wide variety of clinical manifestations, usually related to the route of infection. Six primary clinical syndromes of tularaemia have been described: ulceroglandular, glandular, oculoglandular, oropharyngeal, pneumonic, and typhoidal. The ulceroglandular form is the most commonly reported form of tularaemia in North America, and is associated with arthropod bites. The incubation period typically lasts from 3–6 days, followed by the sudden onset of flu-like symptoms, including chills, fever, headache, fatigue, sore throat, abdominal pain, vomiting, diarrhoea, and generalised aches. An ulcer, which can persist for several months, forms at the site of infection. From this site, bacteria are disseminated via the lymphatic system to the regional lymph nodes, causing lymphadenopathy. The treatment of choice for tularaemia is streptomycin but gentamycin is also effective (Ellis et al. 2002). During 1990–2004, an average of 123 cases of tularaemia were reported annually in the United States of America, with the current focus of tularaemia in Arkansas and Missouri (large numbers of cases also occur in other eastern states, including Georgia, Illinois, Indiana, Kentucky, Louisiana, Mississippi, Ohio, Tennessee, and Virginia) (Eisen 2007). Ticks play a prominent role in the transmission of F. tularensis to humans, accounting for more than 50% of all cases, especially west of the Mississippi River (Eisen et al. 2008). The seasonal distribution of tularaemia cases in the United States of America provides evidence for the prominence of ticks as a source of human infection with a peak in cases from April to August. This coincides with a peak in activity of the tick vectors: the lone star tick, A. americanum, and the American dog tick, D. variabilis, in the southeastern and southcentral United States of America and the Rocky Mountain wood tick, D. andersoni, in the west. From 2005–2006, a total of 31 cases of tularaemia were reported in Canada. The disease is endemic in central and western Canada but less frequently reported from eastern provinces. In western Canada from 1939–1946, F. tularensis was isolated from D. andersoni, D. variabilis, and H. leporispalustris as well as from ground squirrel, house mouse (Mus musculus Linnaeus), meadow vole (Microtus pennsylvanicus (Ord)), deer mouse, rabbit, and Franklin’s Gull (Leucophaeus pipixcan (Wagner)) tissues (Humphreys and Campbell 1947; Gordon et al. 1983). In Ontario, F. tularensis has been isolated from Haemaphysalis leporispalustris (Ditchfield et al. 1960) and Dermacentor variabilis (Artsob et al. 1984) and from beavers, muskrats, and cats (Labzoffsky and Sprent 1952; Ditchfield et al. 1960). Francisella tularensis may be transmitted to mammals via tick saliva or faeces. The bacterium is commonly transmitted transstadially in ticks, but transovarial transmission is rare and probably unimportant in maintaining natural cycles of infection (Hayes 2005). In Canada, D. andersoni, D. variabilis, H. leporispalustris, and the grouse tick, H. chordeilis, have all been implicated as carriers for F. tularensis and help maintain enzootic cycles by transmitting the bacteria among lagomorphs and rodents (Gordon et al. 1983).

Rocky Mountain spotted fever (RMSF) Rocky Mountain spotted fever, caused by Rickettsia rickettsii (Wolbach) Brumpt (Rickettsiales: Rickettsiaceae), is the most severe of all tick-borne rickettsioses. Rocky Mountain spotted fever reached its highest incidence ever in the United States of America in 2012 (Drexler et al. 2016), although this infection is likely underreported because of the lack of awareness by many physicians. The number of RMSF cases reported in Canada is much lower. For example, from 1969–1976, there were 33 hospitalised cases of Rocky Mountain 46 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins spotted fever (Statistics Canada 1969–1976). Most cases of Rocky Mountain spotted fever in Canada have been reported from British Columbia, Alberta, and Saskatchewan, but one case has been reported that was contracted near Ottawa, Ontario (Mackenzie et al. 1979). Rocky Mountain spotted fever is not a notifiable disease in Canada and therefore, the true incidence of this infection in Canada is unknown. Ticks are the natural hosts, reservoirs, and vectors for R. rickettsii, although the organism is also infectious for many mammals. Rickettsia rickettsii is maintained in natural cycles involving wild mammals and ixodid ticks. The primary vector of RMSF in the eastern and southern United States of America is the American dog tick, Dermacentor variabilis. This vector is also present throughout eastern Canada, as far west as southeastern Saskatchewan. The Rocky Mountain wood tick, D. andersoni, is the principal vector of RMSF in the western United States of America, Alberta, and British Columbia. Other species of ticks found in the United States of America have been shown to be naturally infected with R. rickettsii or have been demonstrated to be potential vectors in the laboratory, including Haemaphysalis leporispalustris, Ixodes dentatus, Dermacentor occidentalis, D. parumapertus Neumann, Amblyomma americanum, Rhipicephalus sanguineus, and the soft tick, Ornithodoros parkeri. Rickettsia rickettsii has been isolated from D. andersoni collected in Alberta and British Columbia (Humphreys and Campbell 1947). In central and eastern Canada, R. rickettsii has been isolated from D. variabilis collected in southwestern Ontario (Six Nations reserve, Cardoc, Walpole Island) during 1965–1971, from H. leporispalustris collected from the Ottawa region during 1965–1972, and from D. variabilis and H. leporispalustris collected on the south shore of Nova Scotia during 1976–1980 (H.A., personal observation). However, known pathogenic rickettsial species have not been detected in D. variabilis and D. andersoni collected in recent studies from various locations across Canada, including British Columbia, Alberta, Saskatchewan, Manitoba, and Ontario (Dergousoff et al. 2009; Teng et al. 2011; H.W., personal observation). An outbreak of RMSF in Arizona in 2004 was caused by infected R. sanguineus, a known vector of RMSF in Mexico. Rhipicephalus sanguineus had not been previously associated with RMSF in the United States of America but is now recognised as a potential vector for human infections, particularly in areas where there are no Dermacentor species. All patients in the outbreak reported contact with tick-infested dogs, indicating that dogs may serve as important transport hosts by carrying infected ticks close to their owners (Demma et al. 2005b, 2006). Dogs may also serve as sentinels for disease in their human owners as R. rickettsii causes a severe and sometimes fatal illness in these domestic pets. Rhipicephalus sanguineus bites humans infrequently, but nymphs, when present in larger numbers, have been shown to bite humans. Rhipicephalus sanguineus feeds predominately on dogs in peridomestic habitats, as opposed to D. variabilis, which resides in rural and suburban regions with its small-mammal hosts (Dumler and Walker 2005). Therefore, the epidemiologic features of RMSF transmitted by R. sanguineus are potentially different from those of RMSF transmitted by Dermacentor species. The onset of symptoms of RMSF in humans usually begins 5–7 days after an infective tick bite. Common symptoms include generalised malaise, myalgias, fever, frontal headache, nausea, and vomiting. The classic presenting symptoms include sudden onset of headache, fever, and chills accompanied by a rash appearing within the first few days of symptoms. The macular rash appears initially on the wrists and ankles, and spreads to the trunk of the body becoming maculopapular and then petechial in nature. Neurological manifestations may develop in severe cases, as well as acute renal failure and cerebral and A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 47

pulmonary edema, which are life-threatening. With appropriate treatment, mortality for RMSF is less than 5%; however, the mortality can be as high as 25% if treatment is not initiated or is initiated more than five days from onset of symptoms. Doxycycline is the drug of choice for treatment of adults and children with RMSF because of its effectiveness in treating all spotted fever group (SFG) tick-borne rickettsioses, which can be severe and potentially fatal. Antibiotic treatment should be administered on the basis of clinical suspicion, and should not await diagnostic tests results because delayed administration of antibiotics may result in greater likelihood of morbidity and mortality. Rickettsia rickettsii has been isolated from various small mammals, including meadow voles, golden-mantled ground squirrels (Callospermophilus lateralis (Say), rabbits, and chipmunks, which all develop rickettsaemias of sufficient magnitude and duration to infect ticks. These small mammals are highly susceptible to rickettsial infection and are common blood meal sources for immature ticks, thereby acting as amplifying hosts. Other wild and domesticated animals are susceptible to infection with R. rickettsii but produce rickettsaemias too low or too transiently to infect ticks (Parola et al. 2005b). Rickettsia rickettsii initially infects the epithelial cells of the tick midgut, where it multiplies and subsequently enters the haemocoel, and from there invades and multiplies in other tick tissues including the salivary glands and ovaries. Rickettsia rickettsii is transmitted transovarially and transstadially in D. variabilis and D. andersoni; therefore, these ticks serve as both a reservoir and vector for R. rickettsii. In the laboratory, transovarial transmission of R. rickettsii in D. andersoni diminished the survival and reproductive capacity of tick progeny (Nieblyski et al. 1999). Far fewer offspring developed from infected ticks than from uninfected siblings (Demma et al. 2005b). Ticks may become infected by feeding adjacent to an infected feeding tick; this mechanism would allow for the acquisition of R. rickettsii without the presence of infection in the mammalian host. Infected ticks must remain attached to the host for at least 24 hours in order to transmit the bacteria (Amsden et al. 2005). Interference between different species of rickettsia has been identified as a mechanism by which one species may competitively displace another species from a tick reservoir. In the 1980s, Burgdorfer et al. (1981) showed that ticks infected with the SFG rickettsia, R. peacockii (then known as the east side agent), were refractory to infection with and maintenance of R. rickettsii. Interspecies competition between different rickettsial species in the same tick has been confirmed in more recent studies. Over the past century in the Bitterroot Valley (Montana, United States of America; where R. rickettsii was first isolated), R. rickettsii has become limited to the western slopes, where less than 1% of ticks are infected, despite a widespread abundance of vertebrate hosts. During this same time period, human cases of RMSF in the Bitterroot Valley essentially disappeared. It has been hypothesised that R. peacockii, an avirulent species of rickettsia (which is considered an endosymbiont of D. andersoni) has displaced R. rickettsii from ticks in this region through rickettsial interference. In subsequent laboratory studies, two other “non-pathogenic” rickettsiae, R. montanensis (Lackman et al.) Weiss and Moulder and R. rhipicephali (Burgdorfer et al.) Weiss, blocked transovarial transmission of R. rickettsii. These species are also able to block transovarial transmission of each other, demonstrating that interference occurs independently of the pathogenicity of the rickettsiae. It has been suggested that rickettsial infection of tick ovaries may alter the molecular expression of the oocytes and cause interference or blocking of secondary infection. The process of rickettsial interference could have a significant effect on the distribution and frequency of various pathogenic rickettsiae and their corresponding diseases (Parola et al. 2005b). 48 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Rickettsia parkeri rickettsiosis Rickettsia parkeri Lackman et al. was isolated in 1939 from Amblyomma maculatum, the Gulf Coast tick, collected from cows in Texas (Lackman et al. 1949). For over 60 years, R. parkeri was considered a non-pathogenic species of SFG rickettsiae and received relatively little attention. However, in 2002, the first reported case of human infection with R. parkeri was documented. This patient presented with a relatively mild, febrile illness accompanied by multiple eschars and a maculopapular eruption (Paddock et al. 2004). An eschar, or tache noir, caused by local necrosis of the skin at the tick bite site, is rare in RMSF cases and raised the possibility of a distinct SFG rickettsiosis in the United States of America. The presence of a second SFG rickettsial species capable of causing human disease was subsequently confirmed by molecular studies and isolation of R. parkeri in cell culture from a skin biopsy of the patient. Rickettsia parkeri has now been identified as the cause of spotted fever illness in other patients in the United States of America, and its association with tick bites has been firmly established (Whitman et al. 2007). Because R. parkeri is most closely related to Old World, eschar- producing SFG pathogens, such as R. conorii Brumpt, the causative agent of Boutonneuse fever, infections due to R. parkeri have been termed American Boutonneuse fever. The distribution and significance ofR . parkeri in North America is unclear. However, it is now clear that R. rickettsii is not the sole tick-transmitted SFG in the United States of America capable of causing human disease. Amblyomma maculatum is distributed predominately in coastal regions of the southern and southeastern states that border the Gulf of Mexico and Atlantic Ocean. Rickettsia parkeri has been identified in Gulf Coast ticks collected throughout its range in the United States of America (Paddock et al. 2004). It has been suggested that rare observations of eschars in suspected RMSF cases could actually be caused by infections with R. parkeri rather than R. rickettsii. Because of the recognised cross-reactivity of R. rickettsii with antigens of other SFG rickettsiae, serological titres to R. rickettsii in patients with mild, self-limited infections may actually represent infections with other SFG rickettsiae, including R. parkeri, that are less virulent than R. rickettsii (Paddock 2005).

Potential tick-borne bacterial pathogens Other Rickettsia species Ticks collected throughout North America, including many that parasitise humans, are hosts to many distinct SFG rickettsiae. Many of these rickettsiae, including R. canadensis McKiel et al., R. rhipicephali, R. amblyommii Labruna et al., cause mild to severe disease in rodents or other small mammals but have not yet been isolated from humans with disease (Apperson et al. 2008; Wikswo et al. 2008). Of particular interest from a Canadian perspective is the isolation of R. canadensis from H. leporispalustris collected near Richmond, Ontario in 1964 (McKiel et al. 1967). This rickettsia has been implicated as a causative agent of human disease but conclusive evidence is lacking (Bozeman et al. 1970). Rickettsia 364D, a SFG rickettsia closely related to R. rickettsii and commonly found in D. occidentalis, was detected for the first time in a patient from northern California with mild illness and an eschar (Shapiroet al. 2010). It is reasonable to assume that other tick species may be responsible for transmission of SFG rickettsial species in addition to the classically recognised vectors for RMSF. The first confirmed human case of Rickettsia massiliae Beati and Raoult infection was reported in 2006 from a patient in Italy with fever, a maculopapular rash on the palms and soles and an eschar. Rickettsia massiliae was implicated as the cause of an RMSF-like illness in dogs in southern California, and was present in Rhipicephalus sanguineus collected from the area where the dogs lived (Beeler et al. 2011). Rickettsia massiliae has not yet been detected from humans in North America, although the potential for human infection clearly exists. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 49

Rickettsia montanensis has been found in D. variabilis in Manitoba and Saskatchewan (Dergousoff et al. 2009; Yunik et al. 2015a). The first human case of R. montanensis infection was reported in 2012 from a patient in Georgia, United States of America. This patient developed an afebrile rash illness four days after being bitten by a D. variabilis tick that tested positive for R. montanensis (McQuiston 2012). A study in dogs naturally exposed to A. americanum, A. maculatum, and D. variabilis ticks suggested that several SFG rickettsia species may cause infections in canines, including R. montanensis and R. amblyommii (Barrett et al. 2014). The identification of new SFG rickettsia species involved in human disease will require the addition of culture-based and molecular approaches to the panel of diagnostic tests used to evaluate human samples. Cross-reactive serological assays which are SFG-specific and cannot be used to identify rickettsiae specifically are the primary tests offered for the diagnosis of rickettsial infections, thereby limiting the identification of new SFG pathogens of humans and animals. Contemporary serosurveys in the United States of America have identified remarkably high prevalences of antibodies reactive with R. rickettsii among persons who reported very mild illness not considered consistent with classical RMSF, suggesting that some mild infections are caused by SFG rickettsiae other than R. rickettsii (Paddock 2005).

Bartonella species It is possible that Bartonella Strong et al. (Rhizobiales: Bartonellaceae) species may be transmitted by ticks in North America. Bartonella species are the etiological agents of cat scratch fever (Bartonella henselae (Regnery et al.) Brenner et al.), trench fever (Bartonella quintana (Schmincke) Brenner et al.), and oroya fever (Bartonella bacilliformis (Strong et al.) Strong et al.) and their vectors are fleas, lice, and sand flies, respectively. Bartonella DNA has been detected in several species of ticks, including questing adult Ixodes pacificus (Chang et al. 2001; Holden et al. 2006) and questing adult R. sanguineus in California (Wikswo et al. 2008). However, studies demonstrating transstadial and transovarial transmission of Bartonella species in ticks and experimental transmission of these bacteria from ticks to mammals have not yet been performed. Detection of DNA from a human pathogen in ticks does not in itself indicate involvement of the tick as a vector for human infection. However, there is mounting evidence that ticks could potentially serve as vectors for transmission of Bartonella species.

Borrelia miyamotoi Borrelia miyamotoi Fukunaga et al. (Spirochaetales: Spirochaetaceae), was first described in Ixodes persulcatus (Schulze) and in the blood of rodents collected in Japan in the early 1990s (Fukunaga et al. 1995). Subsequently, B. miyamotoi was detected, for the first time in North America, associated with blacklegged ticks, in several states in the northeastern United States of America (Scoles et al. 2001). Infection rates in field-collected nymphalI. scapularis were 1.9–2.5% and unlike the agent of Lyme disease, Borrelia burgdorferi, B. miyamotoi is transmitted vertically from infected female I. scapularis to a variable proportion of larval progeny (Scoles et al. 2001; Rollend et al. 2013). Initially the public health significance of B. miyamotoi was poorly understood; however, recent studies in Russia have demonstrated that Old World strains of B. miyamotoi, transmitted by I. persulcatus, cause an influenza-like illness with relapsing fever (Platonov et al. 2011). In North America, meningoencephalitis was recently described in an elderly immunocompromised patient (Gugliotta et al. 2013) and from results of a serosurvey of patients from southern New England and New York, B. miyamotoi infection can cause a viral-like illness (Krause et al. 2013). These studies 50 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins support the contention that B. miyamotoi is yet another of the guild of pathogens, which includes the agents of Lyme disease, anaplasmosis, babesiosis, Powassan virus, and the Ehrlichia muris-like agent, associated with blacklegged ticks in North America (Chowdri et al. 2013). The discovery of DNA of B. miyamotoi in ticks during a study of B. burgdorferi diversity signaled the possible occurrence of B. miyamotoi in Canada (Ogden et al. 2011). Subsequent studies quantified the prevalence and distribution of this agent in I. scapularis ticks from across Canada, some co-infected with B. burgdorferi (Dibernardo et al. 2014). Ixodes pacificus populations in California have also been found to harbour this pathogen (Fedorova et al. 2014; Padgett et al. 2014) and it is assumed that it is also present in western blacklegged tick populations in British Columbia.

Coinfections acquired from Ixodes ticks Ticks can become infected with multiple pathogens after one blood meal from a coinfected host or by feeding on one infected host during sequential life stages (Swanson et al. 2006). Ixodes scapularis and I. pacificus are involved in the transmission of three separate agents: Borrelia burgdorferi, Anaplasma phagocytophilum, and Babesia microti (discussed in the protozoan section). All three agents are also maintained by the same rodent reservoirs, white-footed mice. Coinfection of ticks with these agents can and does occur, with varying prevalence of infection depending on the geographic location of the ticks. Coinfections in humans with two or all three of these organisms have been reported (Walker et al. 1996; Thompson et al. 2001; Swanson et al. 2006). The clinical manifestations of concurrent Lyme disease and babesiosis tend to be more severe than would be expected if these infections occurred separately. It is less clear if patients with both Lyme disease and HGA have a different clinical outcome than do patients with Lyme disease alone (Thompson et al. 2001).

Protozoan parasitic diseases Cytauxzoonosis Feline cytauxzoonosis is an invariably fatal blood protozoan disease caused by Cytauxzoon felis Kier (Achromatorida: Theileriidae) (Meinkoth et al. 2000; Meinkoth and Kocan 2005). It is found in the south central and southeastern United States of America, notably Texas, Oklahoma, Missouri, Arkansas, Louisiana, Georgia, Mississippi, and Florida. Ticks, probably D. variabilis, are the most likely vector, but other transmission routes are possible including by blood from bite wounds or transfusions. Infected cats develop clinical signs typical of haemolytic crisis such as anorexia, laboured breathing, lethargy, dehydration, depression, icterus, pale mucous membranes, and fever (Meinkoth and Kocan 2005). Death usually occurs 2–3 days after body temperatures peak, with the entire course of clinical illness usually lasting less than seven days. Attempts to treat naturally infected cats have been unsuccessful. Although generally a fatal disease, there have been reports of cats surviving the disease (Meinkoth et al. 2000). Some of these cats showed a persistent erythro-parasitaemia. These cats may have survived the disease because of infection with a less virulent strain or heightened immune-competency. The diagnosis of C. felis is made by demonstrating organisms within infected erythrocytes on a blood smear or histological identification of schizonts in tissues. Differentiation between C. felis and the smaller feline parasites, such as Babesia felis Davis (Piroplasmida: Babesiidae) and Haemobartonella felis (Clark) Kreier and Ristic (Mycoplasmatales: Mycoplasmataceae), as well as Howell-Jolly bodies must be made. PCR can be used to differentiate between these feline parasites. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 51

Babesiosis Babesiosis is caused by organisms of the genus Babesia, which are tick-borne protozoan parasites that live inside red blood cells (Homer et al. 2000; Vannier et al. 2008). Over 100 species of Babesia have been identified in wild and domestic animals (Levine 1988). The complex life cycle of the agent involves asexual reproduction in the erythrocytes of mammalian hosts and sexual reproduction in arthropod vectors. They are usually host specific and transmitted by ixodid ticks, in which they may actually cause adverse effects. They are responsible for infections in many species of animals and can cause symptomatic infection in humans (Levine 1988; Vannier et al. 2008). Human infections in the United States of America are most frequent in the southeast but have been reported in Hawaii, the southwest, midwest, far west, and middle Atlantic states (Vannier et al. 2008). The etiological agent of human disease in the United States of America is predominantly Babesia microti. The primary tick vector of the species is I. scapularis and the main reservoir in the northeastern United States of America is the white-footed mouse (Peromyscus leucopus) (Spielman et al. 1985). Humans are believed to be accidental hosts and while tick bites are the main risk for transmission, there have been several cases of transfusion-acquired babesiosis in the United States of America (Pantanowitz et al. 2002). Both I. scapularis and reservoirs such as P. leucopus have been identified in areas of southern Ontario, Québec, New Brunswick, and Nova Scotia. Ticks infected with Babesia microti were collected for the first time in Manitoba inI . scapularis in 2012 (L.R.L., personal observation). The first documented clinical case in humans in Canada was reported in Manitoba in 2013 (Bullard et al. 2014). Blacklegged ticks infected with B. microti have been most frequently detected at sites in Manitoba, but small numbers of infected ticks have been reported from most other provinces in eastern Canada (O’Brien et al. 2016). Human babesiosis in the United States of America due to B. microti occurs primarily in nonsplenectomised people and is invariably self-limiting or easily managed with combinations of quinine and clindamycin (Hatcher et al. 2001; Vannier et al. 2008). Most cases are of mild to moderate severity with symptoms (e.g., chills, sweats, headache, arthralgia, myalgia, cough) occurring 1–6 weeks after tick feeding. Severe disease may occur in individuals with underlying immunosuppressive conditions and increased age (Hatcher et al. 2001). Complications of infection include acute respiratory, heart, renal, and liver failure. Babesiosis in animals is a severe infection characterised by fever, anaemia, and bloody urine. Dogs and horses may be infected by Babesia canis vogeli Reichenow and Theileria (Babesia) equi (Laveran), respectively. Amicarbalide, imidocarb, and diminazene have been used successfully to treat the infection in animals. Definitive diagnosis of babesial infection is by microscopic identification ofthe organism in blood smears (Vannier et al. 2008). However, less than 1% of erythrocytes are parasitised early in the course of illness, so multiple smears should be examined over several days. PCR is a sensitive procedure that may detect babesial DNA months after infection. Serological techniques, including IgM and IgG immunoflourescent assays, can also be used to indicate active or recent infections.

Viral diseases Colorado tick fever Colorado tick fever is a generally benign disease caused by the Colorado tick fever virus (Romero and Simonsen 2008). It is a member of the genus Coltivirus Francki et al. (Reoviridae) and is transmitted by Dermacentor andersoni and, to a lesser extent, D. occidentalis (Attoui et al. 2005). It occurs throughout the Rocky Mountain and northwestern 52 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins states. The agent is responsible for several hundreds of reported cases a year in endemic areas of the United States of America, making it the second most commonly identified arboviral infection in North America after West Nile virus (Brackney et al. 2010). Rabbits and hares, golden-mantled ground squirrels, chipmunks, deer mice, and other small mammals have all been implicated as reservoir hosts for Colorado tick fever virus (Carey et al. 1980). In Canada, Colorado tick fever virus was isolated from D. andersoni collected in Alberta in 1952–1954 and in British Columbia in 1965–1966 (Brown 1955; Hall et al. 1968; Artsob 2000). Only one symptomatic human infection has been recognised in Canada due to Colorado tick fever virus (Cimolai et al. 1988). The infection occurred in 1986 in a 20-year-old woman from southwestern Alberta. In serosurveys, exposure of British Columbia residents to Colorado tick fever virus has been demonstrated; presence of the virus in Ontario has also been indicated (Newhouse et al. 1964; Kettyls et al. 1972). The majority of Colorado tick fever virus cases occur during May to July and over 50% of patients are 20–49 years of age (Goodpasture et al. 1978). However, almost a quarter of cases also involve children. A high rate of infection among males has also been observed. The clinical manifestations of Colorado tick fever infection in humans are similar to early signs of Rocky Mountain spotted fever but usually without the rash (Wright and Trott 1988). The onset of clinical illness usually occurs after an incubation period of 3–5 days. Symptoms may include fever, chills, headache, back pain, myalgias, generalised weakness, and photophobia with a rash, usually macular or maculopapular, observed in only 5–15% of patients. Approximately half of the cases have recurrence of fever and acute symptoms after an initial period of improvement (“saddleback fever”) (Goodpasture et al. 1978). Neurological manifestations such as aseptic meningitis and encephalitis have been observed in a small number of infections of children. Death is rare among individuals infected. There is no specific treatment for Colorado tick fever; supportive care is the principal intervention. Thus, avoidance of tick bites and reduction of tick vectors are important. Infection by this virus has been diagnosed by a variety of serological procedures including complement fixation, indirect immunofluoresence, IgM capture enzyme-linked immunosorbent assays (ELISAs), and plaque reduction neutralisation (Attoui et al. 1998). Reverse transcription PCR has been used to detect viral RNA in acute samples of blood collected within five days of illness (Lambertet al. 2007). The virus has also been isolated from blood and cerebrospinal fluid by inoculation of suckling mice and cell culture.

Powassan encephalitis Powassan encephalitis is caused by a tick-transmitted flavivirus (family Flaviviridae) known as Powassan virus (Wright and Trott 1988; Romero and Simonsen 2008). The prototypic strain of the virus was isolated from the brain of a five-year-old boy from Powassan, Ontario in 1958 (McLean and Donahue 1959). In North America, the virus has been isolated from four species of Ixodes (I. cookei, I. scapularis, I. marxi, and I. spinipalpis) and from D. andersoni. Evidence of virus infection has been found in 38 species of mammals; however, the principal vertebrate host appears to be the groundhog (Marmota monax Linnaeus) with squirrels and white-footed mice also significant reservoirs (Artsob 1988). From phylogenetic analysis of the genomes of Powassan virus isolates, two lineages of the virus are recognised in North America (Ebel et al. 2001; Kuno et al. 2001). The genetic lineage that includes the prototypic Powassan virus is distinct from a lineage associated with virus isolates from I. scapularis. The two distinct genotypes are believed to have risen due to vector and ecologically based differences (e.g., association of prototypic Powassan with I. cookei and groundhogs, deer tick virus with I. scapularis and white-footed mice). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 53

Viruses from both lineages cause human disease (Kuno et al. 2001; Tavakoli et al. 2009). Evidence for the presence of Powassan virus by virological or serological analyses has been documented in many areas of Canada including New Brunswick, Nova Scotia, Prince Edward Island, Québec, Ontario, Alberta, and British Columbia (Artsob 1988; Fitch and Artsob 1990; Dibernardo et al. 2007). The virus has been isolated from ticks and/or vertebrates caught in Ontario and Prince Edward Island (Artsob 1988; Dibernardo et al. 2007). In the United States of America, the virus has been shown to circulate in California, Colorado, Connecticut, Massachusetts, New York, Maine, Vermont, Wisconsin, South Dakota, and West Virginia (Romero and Simonsen 2008). Symptomatic human infections with Powassan virus are relatively rare. For example, from 2001–2012, 47 cases of Powassan encephalitis were reported to the Centers for Disease Control and Prevention (Atlanta, Georgia, United States of America) (http://www. cdc.gov/powassan/pdf/POWNDbyYear20012012.pdf). In Canada, there have been only 16 human cases reported to-date from New Brunswick (2), Québec (6), and Ontario (8) (Artsob 1996; M.D., personal observation). Two thirds of Powassan cases have occurred in males and the majority involved children (Romero and Simonsen 2008). However, there appears to be a trend towards increasing number of adults with diagnosed infections which may be due to the more recent testing of neurological cases for West Nile virus. Symptomatic infections of domestic animals have not been documented under natural circumstances, but experimentally-inoculated horses developed a fulminating viral encephalitis (Little et al. 1985). Powassan virus has also been isolated from the brain of a sick fox (Vulpes vulpes (Linnaeus)) found in New York and West Virginia (Artsob 1988). Clinical manifestations of Powassan encephalitis in humans include onset of fever and convulsions; prodromata including sore throat, sleepiness, headache, and disorientation; encephalitis characterised by vomiting, respiratory distress, and prolonged, sustained fever; an electroencephalogram showing diffuse slow-wave change indicating a widespread pathological process (Smith et al. 1974; Artsob 1988; Romero and Simonsen 2008). Overall the mortality in patients with virus-associated neuroinvasive disease is 10–15%. The 16 Powassan cases in Canada have resulted in four fatalities, two during the acute phase of the disease and two from sequelae directly related to the disease. Diagnosis of viral infection has been carried out through isolation of virus from postmortem brain tissue (Romero and Simonsen 2008). However, serological testing is the primary means of documenting cases through the detection of Powassan specific immunoglobulin M in sera/cerebrospinal fluid by enzyme-linked immunosorbent assay (ELISA)-based methodologies and the four-fold rise in titre by haemagglutination inhibition and neutralisation assays conducted on acute and convalescent serum samples. No specific treatments or therapy are available for Powassan virus neurological disease (Romero and Simonsen 2008). Treatment is primarily supportive with mechanical ventilation and anticonvulsant therapy employed in most patients. Prevention is the key, decreasing the risk for being bitten by ticks is the primary means to avoid transmission of the pathogen.

Diseases of non-microbial etiology Ticks can damage their hosts without help from their tick-borne microorganisms, and such effects caused directly by ticks are discussed in this section.

Anaemia Most argasid ticks, during their relatively short feeding sessions, increase their weight by approximately 5–12 times. Their blood meals tend to be smaller than those of the ixodid 54 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins ticks, but they are repeated more often. However, the larvae of Argas persicus and some other argasid species feed for relatively long periods of time and increase their weight 15–20 times during engorgement. Ixodid ticks feed only once in each stadium, but they take more blood per meal. Female ixodid ticks, for example, may increase their body weight 50–150 times, e.g., in smaller forms, such as species of Ixodes, Haemaphysalis, or Rhipicephalus, increasing their weight by 100–400 mg, and by more than 1000 mg in larger forms such as species of Amblyomma (Balashov 1972). Such figures do not take into account the relatively large amounts of faecal material, derived from the blood meal, that is lost from the tick while feeding, nor do they take into account the large volume of water extracted from the blood meal and returned to the host through the salivary glands of the tick during the later stages of feeding. Balashov (1972) attempted to assess the blood volume equivalents removed from hosts by female ixodid ticks using the iron content of the engorged ticks, and presented figures such as 0.7 mL for Ixodes persulcatus, 3.7 mL for Dermacentor reticulatus (Fabricius), and 8.9 mL for Hyalomma asiaticum Schulze and Schlottke. Data of this sort make it believable that large numbers of ticks may cause enough blood loss to outweigh the red cell-regenerative powers of some hosts, and so directly cause significant anaemia. In experimental infestations with ticks believed to be free of anaemia-producing microorganisms, laboratory rabbits showed marked anaemia when 80 or more female D. andersoni were placed on an animal at the same time (Jellison and Kohls 1938). Similarly, Glines and Samuel (1989) worked in Alberta with captive moose experimentally infested with D. albipictus. The animals were infested with 31 000 larvae in the fall, an average number of ticks judging from the previous studies of moose examined in the wild in that area, and even though the infested animals were fed an unlimited high-protein diet and their exercise was limited, clinical anaemia was detectable at times when nymphs and adult female ticks were engorging. Under natural conditions, large tick infestations appear to be the direct cause of anaemia in smaller hosts. For example, young fawns of white-tailed deer carrying very heavy A. americanum loads in Oklahoma were shown to be anaemic (Bolte et al. 1970). In the same area, mature deer did not appear to show anaemia from the large numbers of all instars of this species of tick. Anticoagulants have been reported in extracts of salivary glands or other tissues of ticks, and it has been assumed that they may contribute to the blood loss and anaemias incurred by their hosts (Ribeiro 1987; Binnington and Kemp 1988). Salivary gland extracts from ixodid ticks have been shown to prolong the clotting time of whole blood (Foggie 1959) and anticoagulants directed against coagulation factors V and VII have been demonstrated in salivary gland extracts from D. andersoni (Gordon and Allen 1991a). Another anticoagulant has been isolated from extracts of an argasid tick, Ornithodoros moubata (Murray). This peptide inhibits factor Xa of the coagulation cascade (Waxman et al. 1990).

Tick paralysis Tick paralysis, or tick toxicosis, is a flaccid ascending paralysis (Gregson 1973; Edlow and McGillicuddy 2008). This potentially lethal effect is produced in humans and other host animals by ticks of several different argasid and ixodid species. World-wide there are 55 species of ixodids and 14 species of soft ticks which have been reported to cause paralysis (Gothe et al. 1979; Gothe 1999), but few species have been studied in detail (Mans et al. 2004). However, reports of the paralysing abilities of many of these species are anecdotal and require confirmation. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 55

In species that have been studied in detail, the paralytic effect is caused by toxic components produced by the tick’s salivary glands. No paralysis-producing microorganisms have ever been proven to occur in ticks. The onset of paralysis generally occurs after long-feeding ticks have been attached to the host for several days. The type of paralysis caused by these ticks is very similar in all hosts that are affected. The hind legs of the host are the first to become numb, then uncoordinated, weak and paralysed. Paralysis gradually spreads to the forelimbs, the host is no longer able to stand, and finally the host may die from paralysis of the respiratory muscles. It is not clear if the ability to paralyse their hosts provides a distinct evolutionary advantage to these long-feeding ticks. However, it has been suggested that paralysis of the host might increase the chances of ticks completing their blood meals by reducing the reflex grooming activities which normally remove some of the ticks from their hosts (Gothe 1984). Paralysing toxins have been isolated from only a few species. For example, the toxin from Ixodes holocyclus Neumann has been isolated in Australia from the salivary glands of female ticks which have been feeding for 4–5 days (Goodrich and Murray 1978) and similar isolations have been made from salivary glands of female Rhipicephalus evertsi evertsi Neumann, a South African species (Vijoen et al. 1986). Both of these toxins are antigenic and induce a protective immunological response in animals that have recovered from paralysis or have been vaccinated with the toxin. Also, a toxin capable of paralysing day-old chicks has been isolated from extracts of engorged larvae of Argas walkerae Kaiser and Hoogstraal (Vijoen et al. 1990). In Canada, D. andersoni is the species most commonly associated with paralysis in humans, domestic, and wild animals, with most cases occurring in south-central British Columbia. A very labile and short-lived toxin is presumed to be injected into the body of the host in the saliva (Wilkinson 1982) during tick feeding because removal of the offending female ticks from the host allows a rapid and complete recovery from the paralysis. As in paralyses caused by I. holocyclus and R. evertsi evertsi, demonstrable immunological protection against D. andersoni paralysis occurs in some hosts after recovery from paralysis (Lysyk et al. 2009). Prior feeding on cattle previously exposed to paralysing D. andersoni may even inhibit the ability of ticks to induce paralysis (Lysyk et al. 2005). The paralysis from D. andersoni is caused by female ticks which have been attached for about five days or longer (Gregson 1973). If a paralysing female tick is removed from one host and allowed to re-attach to a new host, the second host usually becomes paralysed within a few hours. In humans and dogs, one tick is often sufficient to produce paralysis. Other mammals, such as cattle and sheep appear to require larger numbers of ticks to produce paralysis, and yet other mammals, including the cat and rabbit, are apparently unable to be paralysed at all by this species of tick (Gregson 1973). Paralysis caused by D. andersoni is much more commonly found in certain restricted areas of the distribution of the tick. For example, paralysis occurs frequently in western parts of the dry belt of British Columbia, but very rarely in Canada east of the Rocky Mountain divide. There is even considerable variation in the ability of D. andersoni in central British Columbia to cause paralysis from year to year and in different locations; this has been shown to be a heritable trait (Lysyk and Majak 2003; Lysyk 2010). A hint of the degree of genetic variation in populations of D. andersoni in Canada was described by Patterson et al. (2009) for populations in Saskatchewan and Alberta, and might be very important in understanding differences in behaviour in this tick. Paralysing D. andersoni from British Columbia typically attach to sites on the “underside” of the animals, whereas the non-paralysing ticks from the prairies tend to attach on the topside of animals, notably around the head and between the shoulder blades. 56 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

However, there is no evidence that the site of attachment of the ticks on man or domestic animals affects the outcome of the paralysis in any way (Wilkinson 1985). The mode of action of the toxin has been studied in animals infested with paralysing ticks. Dogs and marmots have been used in electrophysiological experiments. The passage of electrical impulses along nerves and in muscles of the paralysed animals was monitored, and the speed and amplitude of nerve impulses being conducted along nerve fibres were found to be reduced markedly in paralysed animals (Gothe et al. 1979). Also, there was a reduced ability to transmit excitation at neuromuscular junctions. The motor nerves were apparently more seriously affected than the sensory nerves, though sensory nerves were also slightly affected. In humans, reduced velocities and amplitude of motor nerve impulse conduction were also reported in one case, along with electromyographic abnormalities (Swift and Ignacio 1975). Discovery of the exact chemical nature of the paralysing toxin and associated proteins and the precise mode of action in the mammal must await analysis of this complex system. Contrary to what one might expect, a partially-fed female tick may go unnoticed by the human host. The ticks appear to use a local anaesthetic at the attachment site and often attach at inconspicuous sites, under long hair at the nape of the neck, or in the perineal region. Also, if a patient with paralysis has travelled a long distance in the five days prior to tick attachment, medical personnel in an area where tick paralysis is uncommon may not have this in mind as a differential diagnosis, and may not conduct a search for ticks. Dermacentor variabilis is the other species reported to cause tick paralysis in Canada. Horses carrying large numbers of ticks have shown ascending flaccid paralysis and recovery was rapid after removal of the ticks. As with D. andersoni, paralysis caused by D. variabilis occurs in restricted areas of the distribution of this species, particularly in horses in Manitoba and in dogs in Florida. It is unclear whether these cases are comparable in mode of action to other forms of tick paralysis. These cases usually involve infestation with extraordinary numbers of ticks, and there has seldom been information on the fitness of the affected animal prior to infestation. Dermacentor variabilis has also been reported to cause paralysis in humans in the United States of America (Kocan 1988).

Skin lesions The chelicerae and hypostome of a tick are inserted into the skin of the host, and saliva is injected into the skin as the tick attaches and periodically as the tick feeds. Mechanical disruption of the skin by the mouthparts of the tick and several “inflammatory” components of the saliva of the ticks seem to be responsible for causing skin lesions at the attachment sites of the ticks. Salivary gland extracts of D. variabilis contain components that locally activate the complement cascade and help to mediate the production of lesions in the skin of the host (Ward and Sonenshine 1972). Similarly, components of saliva of D. andersoni, when injected into normal skin, cause lesions histologically similar to those produced in primary tick infestations and activate complement factor 5. These and other salivary components appear to be responsible for inflammatory reactions in the skin of hosts undergoing primary infestations (Ribeiro 1987; Gordon and Allen 1991b).

Hypersensitivity reactions There are many components of the saliva of ticks that act as allergens. These molecules are foreign to the host into which they are injected by the tick and cause no response in the host at first. However, after the host has had time to produce an immunological response to these molecules, and has become hypersensitive to them, an allergic reaction will occur when these molecules are next injected by ticks. The allergic reaction which occurs at the A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 57

attachment site of these ticks is, in effect, a much exaggerated inflammatory response with severe itching and increased swelling, and sometimes more dramatic results may occur. Human patients bitten by argasid or ixodid ticks may become sensitised by allergens introduced into them with the saliva of the ticks, sometimes causing dangerous anaphylactic reactions (Klein-Tebbe et al. 2006; Klotz et al. 2009). In much the same way, other animals produce allergic skin reactions to ticks, sometimes with serious results. Delayed anaphylaxis has also been linked to ingestion of red meat in some individuals with a history of bites from lone star ticks. The carbohydrate found on non-primate mammal and tick glycoproteins (galactose-alpha-1,3-galactose) appears to sensitize people bitten by ticks such that anaphylaxis reactions (i.e., hives, lip and tongue swelling and wheezing) can occur after the consumption of mammalian meat products such as beef (Wolver et al. 2013).

Dermacentor albipictus and moose Severe damage may be done to moose in Canada in late winter, apparently by hypersensitivity reactions to salivary antigens of the winter tick, D. albipictus. In Alberta and Ontario, such reactions to ticks have been associated with serious losses of moose in many winters since the 1930s. In Canada, the larvae of this species climb up the vegetation in the late autumn, from September to mid-November. They occur in large numbers gathered together on the vegetation about 1.0–1.5 m from the ground, at a height suitable to be brushed off by large passing hosts. Because of the aggregated distribution of the larvae in the habitat, if a moose picks up any larvae at all, there may be large numbers of them. Moose have been estimated to carry tens of thousands of these ticks quite regularly in midwinter, and often tick loads may exceed 100 000 on one animal (Samuel 2004). Being a one-host tick, the larvae of D. albipictus stay on the host after they have fed and moult to become nymphs usually beginning by November. The nymphs begin feeding in January, and the first begin moulting to become adults in February. Adults typically engorge during March and April and engorged females drop from the animals to lay their eggs. By May, there are few ticks left on their host (Samuel 2004). It is at this time of the year, when the adult ticks are feeding, that skin hypersensitivity reactions to the injected saliva of the ticks can apparently result in death of the moose. Areas of skin harbouring large numbers of ticks show marked hair loss, and the affected moose spend much time grooming and rubbing these affected areas of the skin. The result is loss of hair, which seems unrelated to the normal spring moult of hair experienced by all moose. It is thought that this premature hair loss in the spring is the result of irritation induced by allergic reactions to the saliva of the tick. The hair loss patterns in this condition are characteristic; light tick loads causing hair-loss over the neck and shoulder regions, and more severe cases showing baldness over almost the entire sides and back. The loss of insulation associated with hair loss at this time of year, when very low temperatures are still prevalent, results in increased energy expenditure and loss of body condition at a time the animals are also likely to be losing relatively large amounts of blood because of the heavy feeding activities of the female ticks. Deaths of such moose occur in quite large numbers in years when late winter snow storms make movement and browsing difficult for already weakened animals (Samuel 2004).

Tick resistance Allergic reactions to tick salivary allergens in many host species, if they are localised and controlled, may be beneficial and protective against ixodid tick infestations. 58 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

The first studies on this subject were reported by Trager (1939a, 1939b), who demonstrated that guinea pigs and other hosts may acquire resistance to the feeding activities of ixodid ticks. This has since been confirmed with several ixodid species in several mammalian species. It appears that the acquired resistance is immunologically mediated and depends on the development of small blisters under the attached ticks in hypersensitised animals. Large numbers of basophil leucocytes accumulate in the blisters and then release histamine and other mediators locally in the skin. These mediators cause an itch and reflex grooming activities that help to remove ticks from the attachment sites. They also apparently have direct effects on the ticks attached nearby, causing them to cease feeding and salivating and to detach. These reactions in hypersensitised animals interfere with the normal feeding activities of long-feeding ticks, either ixodids or the larvae of some argasid species, but apparently are too slow in developing to interfere with the feeding of the faster-feeding argasid species. Reactions of this sort occur in cattle and sheep and other hosts, but are not equally evident in all host species, and some species of ixodids appear to have evolved mechanisms of avoiding the acquisition of effective tick resistance by their usual host species.

Immunosuppression caused by tick infestations Apart from toxins, inflammatory agents and allergens, there are apparently also immunosuppressive components of tick saliva. These seem capable of interfering with the normal immune responses of the host and potentially weakening its defenses against both ticks and tick-borne infections. The exact mechanisms of the immunosuppressive effects are unclear, but it seems that tick infestations and tick salivary gland extracts interfere with T-lymphocyte responses.

Identification and natural history of the ticks of Canada The following section is designed to provide a guide to the identification and diagnostic features of all active instars of ticks known to occur in Canada and a summary of the natural history for each species. Keys and diagnoses were prepared by Evert Lindquist; nearly all figures and plates were prepared by King Wan Wu, by re-drafting illustrations available in the literature; Plates I–IV are original and prepared by Barry Flahey (Agriculture and Agri-Food Canada, Ottawa, Ontario, Canada); maps were generated by Tom Naughten (formerly of Parks Canada, Winnipeg, Manitoba, Canada) based on locality data gathered during the course of this study and from selected published literature. All measurements are in millimetres, unless otherwise indicated. Information on the natural history and references were compiled by Terry Galloway and Robbin Lindsay.

Key to recognition of adults and immature instars of ticks

1. Body with 3 pairs of legs; stigmatic openings absent (Fig. 20G)...... larva – Body with 4 pairs of legs; 1 pair of stigmatic openings present lateral to bases of legs II–IV or ventrolaterally just behind bases of legs IV...... nymphs and adults...... 2

2. Genital aperture absent (undeveloped) (Fig. 22C)...... nymph – Genital aperture present, located mid-ventrally in region between bases of legs I–III (Fig. 11).....adult...... 3

Note: In family Argasidae, other than the shape of the genital aperture (resembling a human thumbnail in males, a pair of lips in females, see Plates IV and II, respectively), the sexes A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 59

are generally similar, such that adult females are not distinguished from adult males in keys to the genera and species.

3. Idiosoma with moderately small shield covering anterior region only, leaving remainder of idiosoma capable of extensive enlargement; dorsal surface of basis capituli with pair of porose areas (Figs. 1, 3A)...... adult female Ixodidae – Idiosoma nearly completely covered by large shield, such that body incapable of becoming greatly enlarged; dorsal surface of basis capituli lacking porose areas (Fig. 2) ...... adult male Ixodidae

Keys to families and genera of ticks in Canada

1. All instars: idiosoma with a dorsal scutum (or shield), which covers anterior region behind gnathosoma in larva, nymph, and adult female (Figs. 1, 10, 21H), and entire dorsal surface in adult male (Fig. 2); gnathosoma (capitulum) projecting anteriorly, and fully visible from above except in greatly engorged females (Fig. 1); adult and nymph: idiosomal integument, other than scutum, finely striated, appearing smooth; spiracular plates located closely posterolateral to coxae IV (Fig. 11); leg coxae often with 1 or 2 spurs (Fig. 42C); larva: palpus with apical palpomere greatly reduced, inserted ventro- subterminally, usually not readily visible in dorsal view (Fig. 23D). Ixodidae...... 2 – All instars: idiosoma lacking a dorsal scutum in nymphs and adults (Fig. 5), sometimes with small mid-dorsal plate in larva (Fig. 6A); gnathosoma placed anteroventrally and covered by anterior projection (hood) of idiosoma in nymphs and adults (Fig. 4B), though projecting anteriorly and fully visible from above in larva (Figs. 6A, B); adult and nymph: idiosomal integument leathery and pebbled or spiny (Fig. 5); spiracular plates located lateral to coxae III or IV (Fig. 4B); leg coxae lacking spurs (Fig. 4B); larva: palpus with apical palpomere shorter but similar in shape to other palpomeres, inserted terminally, readily visible in dorsal view (Figs. 83A, B). Argasidae...... 7

2. All instars: anal groove present, completely or incompletely arching anus anteriorly (Fig. 13A); eyes absent (Figs. 15A, D); scutum inornate; idiosoma usually lacking posteromarginal festoons (Fig. 32D); adult male: venter behind legs entirely covered by a set of 6 nearly contiguous plates (Fig. 45A); larva: idiosoma lacking large wax glands (sensilla sagittiformia); gnathosoma with usually 2 pairs of posthypostomal setae (Figs. 57K, L) �� Ixodes – All instars: anal groove absent or if present, contouring anus posteriorly (Fig. 13B); 1 pair of eyes often present on or beside lateral margin of scutum (Figs. 20A, 22A); scutum ornate or not; idiosoma with posteromarginal festoons often present (Fig. 1) (sometimes faint in engorged specimens); adult male: venter behind legs either partly covered by 4 plates or without plates; larva: idiosoma with 4 pairs of large wax glands (sensilla sagittiformia), including 1 pair on soft integument dorsolaterally and three pairs lateral to bases of legs I–III ventrally (Figs. 10, 12); gnathosoma with 1 pair of posthypostomal setae (Fig. 12)...... 3

3. All instars: palpi conical, flared at the base (Figs. 25A, 25D, 25G, 25K); eyes absent; posterior margin of idiosoma with 11 festoons (Fig. 25D) (sometimes faint in engorged specimens); larva: idiosoma with 2 dorsolateral setae anterior to sensillum sagittiforme on each side (Fig. 25J)...... Haemaphysalis 60 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

– All instars: palpi parallel, not flared at the base; 1 pair of eyes present on or beside lateral margin of scutum (Figs. 19A, 19D); posterior margin of idiosoma usually with 11 festoons (Fig. 23B), sometimes with 9 in larva, or rarely lacking them in all instars; larva: idiosoma with usually 3–5 (Fig. 22D), rarely 2, dorsolateral setae anterior to sensillum sagittiforme on each side...... 4

4. Adult, female and male: dorsal shield ornate, enamelled with brightly coloured markings (Figs. 19A, 19D); male adult: venter lacking a set of delineated plates posterior to legs IV (Fig. 23B); adult and nymph: basis capituli subquadrangular or subtriangular in dorsal outline, midlevel of its sides not produced angulately (Fig. 20A) (nymph of Amblyomma maculatum excepted); larva: idiosoma with 2 or 3 dorsolateral setae anterior to sensillum sagittiforme on each side (Fig. 22D); palpus with 4 palpomeres (femur and genu clearly delineated, each longer than well-delineated trochanter) (Fig. 23D)...... 5 – Adult, female and male: dorsal shield inornate, lacking brightly coloured markings (Fig. 64H); adult male: venter with 4 plates (paired adanal and accessory) posterior to legs IV (Fig. 64I); adult and nymph: basis capituli subhexagonal in dorsal outline, midlevel of its sides produced angulately (Figs. 64A, 64H, 64I); larva: idiosoma with 4 or 5 dorsolateral setae anterior to sensillum sagittiforme on each side; palpus with 3 palpomeres (trochanter or femur or genu not clearly delineated) (Fig. 64K)...... 6

5. Adult and nymph: palpal length shorter than, or equal to, basis capituli width, and palpal femur less than twice as long as wide and usually similar in length to palpal genu (Fig. 23A); adult, female and male: coxae I deeply cleft (Fig. 22B); coxae progressively increasing in size from I–IV (Fig. 22B); larva: idiosoma with 3 dorsolateral setae anterior to sensillum sagittiforme on each side (Fig. 23D), and with 9 festoons along posterolateral margin...... Dermacentor – Adult and nymph: palpal length longer than basis capituli width, and palpal femur usually at least twice as long as wide and twice as long as palpal genu (Fig. 19A); adult, female and male: coxae I with 1 or 2 spurs (Figs. 19B, 19E); coxae I–IV similar in size (Fig. 19B); larva: idiosoma with 2 dorsolateral setae anterior to sensillum sagittiforme on each side, and with 11 festoons along posterolateral margin (Fig. 20D)...... Amblyomma

6. Adult and nymph: palpal apices reaching to or slightly beyond apex of hypostome (Fig. 64E); postanal groove distinct (Fig. 64I); idiosoma with 11 festoons along posterolateral margin; adult, female and male: coxae I deeply cleft (Fig. 64E); larva: idiosoma with 4 dorsolateral setae anterior to sensillum sagittiforme on each side, and with 9 festoons along posterolateral margin (Fig. 64J).....Rhipicephalus (Rhipicephalus) – Adult and nymph: palpi compressed, their apices not reaching apex of hypostome; postanal groove indistinct; idiosoma lacking festoons along posterolateral margin; adult, female and male: coxae I with 2 short blunt spurs; larva: idiosoma with 5 dorsolateral setae anterior to sensillum sagittiforme on each side, and without festoons posteriorly...... Rhipicephalus (Boophilus)

7. Adult, female and male: hypostome vestigial (Plate IV); idiosoma constricted behind legs IV (Fig. 80A), its integument granular and pitted (Fig. 78); nymph (the instar most often seen): hypostome well developed (Figs. 82A–B); idiosoma spinose (Figs. 79A–D); larva: unpaired postanal seta absent; tarsi II and III with seta ad-2 inserted distal to dorsal lyrifissure...... Otobius A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 61

– Adult and nymph: hypostome well developed (Figs. 75A–B); idiosoma elliptical or oval, not constricted behind legs IV, its integument leathery, wrinkly, usually coarsely tuberculate with mammillae and discs, without spines (Plates I–III); larva: unpaired postanal seta usually present (Fig. 77B); tarsi II and III with seta ad-2 inserted proximal to dorsal lyrifissure (as in Fig. 16)...... 8

8. Adult and nymph: idiosoma lacking a sutural line between dorsal and ventral surfaces, with integumental pattern of nearly contiguous mammillae and discs continuous over the sides from dorsal to ventral surfaces (Plate III); idiosoma with dorsoventral groove usually evident behind legs IV (Figs. 4, 5); larva: palp trochanter with small dorsal spines (Fig. 75B)...... Ornithodoros – Adult and nymph: idiosoma with a distinct sutural line delineating dorsal from ventral surfaces (Fig. 66), which covered by wrinkles, small mammillae and discs in various patterns (Plates I, II); idiosoma lacking dorsoventral groove; larva: palp trochanter lacking dorsal spines...... 9

9. Adult and nymph: idiosoma with a flattened marginal rim alongside sutural line that is not obliterated even when tick is fully fed, and with dorsal and ventral surfaces covered with wrinkles and radiating rows of discs (Plate I); gnathosoma not flanked by lateral flaps (cheeks) of idiosoma; tarsi II and III with a single dorsal hump mediodistally; larva: tarsi II and III lacking seta pd-3; genu I usually with seta pv-1...... Argas – Adult and nymph: idiosoma lacking a marginal rim alongside sutural line, and with dorsal and ventral surfaces covered with mammillae and discs sometimes arranged linearly but not clearly radially (Plate II); gnathosoma flanked by pair of lateral flaps (cheeks) of idiosoma (Plate II); tarsi II and III lacking a dorsal hump; larva: tarsi II and III with seta pd-3; genu I usually lacking seta pv-1...... Carios

Family Ixodidae Gnathosoma (capitulum) projecting anteriorly in all instars, and fully visible from above except in greatly engorged females (Fig. 1). Idiosoma with a dorsal scutum (or shield), which covers anterior region behind gnathosoma in larva, nymph, and adult female (Figs. 1, 10, 21H), and covers entire dorsal surface in adult male (Fig. 2). Idiosomal integument, other than scutum, finely striated, appearing smooth in all instars. Spiracular plates located closely posterolateral to coxae IV in nymph and adults (Fig. 11). Leg coxae often with 1 or 2 spurs (Fig. 42C). Larva with apical palpomere greatly reduced in size and dissimilar in shape to other palpomeres, inserted ventro-subterminally, usually not readily visible in dorsal view (Fig. 23D).

Genus Amblyomma Koch (Figs. 19–21) Anus of nymphs and adults with postanal groove. Eyes present (Figs. 19A, D). Posteromarginal festoons present. Palpus with 2nd palpomere (femur) nearly 3 times longer than wide, and not produced laterally (Fig 19A). Basis capituli of adults and nymphs in dorsal view subrectangular, roughly parallel-sided. Adult females and males usually with ornate scutum. Foveae present. Adult males without ventral plates. Coxa I with short internal and longer external spurs. Coxae I–IV similar in size (Fig. 19B). Larvae with 1 pair of posthypostomal setae; scutum with 3 pairs of setae; 8 pairs of marginal dorsal setae, including 2 pairs anterior to sensilla sagittiformia; 2 pairs of central dorsal setae; 11 festoons present (Fig. 21D). 62 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

D E F

Figs. 19A–F. Amblyomma americanum. Fig. 19A, female gnathosoma and scutum, dorsal; Fig. 19B, female coxae and trochanters; Fig. 19C, female hypostome, ventral; Fig. 19D, male gnathosoma and scutum, dorsal; Fig. 19E, male coxae and trochanters; Fig. 19F, male hypostome, ventral. Redrawn from Cooley and Kohls (1944). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 63

A B

F G

Figs. 20A–H. Amblyomma americanum. Fig. 20A, nymph gnathosoma and scutum, dorsal; Fig. 20B, nymph hypostome, ventral; Fig. 20C, nymph coxae and trochanters, ventral; Fig. 20D, larva, dorsal; Fig. 20E, larva hypostome, ventral; Fig. 20F, larva coxae and bases of trochanters, ventral; Fig. 20G, larva, ventral. Figs. 20A–C, E, F redrawn from Cooley and Kohls (1944); Figs. 20D, G redrawn from Clifford et al. (1961). 64 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Key to species of the genus Amblyomma in Canada

1. Adult, female and male: palpus with femur about 2.5 times as long as genu (Figs. 19A, 19D); tibiae II–IV lacking spurs; adult female: scutum with conspicuous white spot posteriorly (Fig. 19A); nymph and larva: basis capituli in dorsal view subquadrangular, with posterolateral corners rounded, not extending laterally beyond scapulae (Figs. 20A, D); coxa I with 2 blunt spurs (Fig. 20C)...... Amblyomma americanum – Adult, female and male: palpus with femur about 1.6 times as long as genu (Figs. 21A, 21D); tibiae II–IV each with a pair of long ventral spurs apically; adult female: scutum with light and dark markings not localised posteriorly; nymph and larva: basis capituli in dorsal view triangular, with posterolateral corners acute, extending laterally beyond scapulae (Figs. 21H, 21L); coxa I with only 1 spur (Fig. 21K) ��Amblyomma maculatum

Amblyomma americanum (Linnaeus) Lone star tick (Figs. 19–20, Map 1) Acarus americanus Linnaeus, 1758: 615. Acarus nigua De Geer, 1778: 153. Ixodes nigua (De Geer): Latreille 1804: 52. Rhynchoprion americanum (Linnaeus): Hermann 1804: 71. Ixodes americanus (Linnaeus): Fabricius 1805: 356. Amblyomma americanum (Linnaeus): Koch 1844: 229. Ixodes unipunctata Packard, 1869: 66. Amblyomma foreli Stoll, 1886–1893: 21. Amblyomma americanum (Linnaeus): Cooley and Kohls 1944.

Adult. Female: Body: Length 2.5–3.4 mm unfed, reaching to 11.0 mm engorged. Gnathosoma: Porose areas of basis capituli subcircular to oval, divergent anteriorly, and separated by interval of about twice the width of one porose area (Fig. 19A). Dorsal posterior margin of basis capituli nearly straight, with rounded corners or small, rounded cornua (Fig. 19A). Hypostome slightly clavate, faintly notched apically; dentition 3/3 along apical half of shaft (Fig. 19C). Palpus long, with femur about 2.5 times as long as genu (Fig. 19A). Scutum: Length about 1.8 greater than width, subtriangular, widest well anterior to mid-length (Fig. 19A). Scapulae pointed. Cervical grooves short, deep. Surface evenly covered with numerous puncta, slightly larger in anterior areas. Posterior angle with a conspicuous white spot with red and green tinges; sometimes with other small spots near eyes. Legs: Coxa I with long, bluntly pointed external spur and a short, rounded internal spur (Fig. 19B). Coxae II–IV each with a broad, plate-like spur, more projected on IV. Tibia of legs II–IV lacking pair of ventral terminal spurs.

Adult. Male: Body: Length approximately 3.0 mm; oval, widest slightly behind midlength. Gnathosoma: Basis capituli in dorsal view subquadrate; cornua short (Fig. 19D). Hypostome as in female, with dentition 3/3 along apical half of length (Fig. 19F). Palpus elongate as in female. Scutum: Surface covered with numerous, small puncta; ornamentation limited to symmetrical, isolated white spots, these much smaller or absent in some specimens (Fig. 19D). Cervical grooves short, deep, convergent anteriorly. Marginal grooves indistinct near eyes, becoming distinct near festoons. Festoons all longer than wide, all limited anteriorly by continuation of marginal groove (Fig. 19D). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 65

Legs: Apical and subapical ventral spurs absent on tarsus I, present on tarsi II–IV. Tibiae II–IV without long ventral apical spurs. Coxa I with external spur longer and more pointed than internal spur (Fig. 19E). Coxae II and III each with a single broad, flat spur. Coxa IV with a long, pointed, internal spur (Fig. 19E).

Nymph. Gnathosoma: Basis capituli with dorsal posterior margin nearly straight, with rounded posterolateral corners not extending laterally beyond scapulae (Fig. 20A). Ventral surface of basis lacking spurs. Hypostome slightly clavate, broadly rounded or slightly flattened apically; dentition 2/2 along apical half of shaft (Fig. 20B). Palpus long, with femur about 2.5 times as long as genu. Scutum: Length about 0.8 width; shape subtriangular, widest at level of eyes, slightly anterior to middle, with nearly straight posterolateral margins (Fig. 20A). Scapulae bluntly pointed. Cervical grooves deep anteriorly, shallow and disappearing behind eyes. Surface sparsely punctate. Legs: Coxa I with short, blunt external and internal spurs (Fig. 20C). Coxae II–IV lacking internal spurs, but each with distinct, blunt external spur.

Larva. Body: Length, excluding gnathosoma, 0.49–0.62 mm. Gnathosoma: Dorsal surface subquadrangular, with posterolateral corners rounded; anterolateral margins with deep indentations at palpal articulation (Fig. 20D); without a pair of sensilla hastiformia. Hypostome slightly clavate; dentition 2/2, with 6–8 teeth per file (Fig. 20E). Palpal genu lacking ventral spur. Scutum: Length approximately 0.6 width, widest at level of eyes, midlength (Fig. 20D). Cervical grooves subparallel, long but not reaching posterolateral margins. Eleven festoons, as wide as long. Legs: Coxa I with two broad, blunt spurs, one near median mesial edge, another near posteromedial margin (Fig. 20F). Coxae II and III each with one short, bluntly rounded spur near middle of posterior margin (Figs. 20F–G).

Natural history. This species does not appear to have become established anywhere in Canada; however, there are frequent incursions, either being returned directly on the bodies of travellers and their pets, or on migrating birds (Scott et al. 2001). Its pattern of seasonal occurrence is relatively consistent wherever it has been studied in the United States of America (e.g., Davidson et al. 1994; Jackson et al. 1996; Gerhardt et al. 1998). Larvae reach their period of greatest abundance in late summer to early autumn, with peak activity in August and September. Occasionally, unfed larvae may survive the winter and resume

Map 1. Collection localities for Amblyomma americanum in Canada. 66 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins host-seeking in the spring. Nymphs are typically active from early spring (March) until October, with April until June being the months of greatest numbers. Adults overwinter and become active first thing in the spring (March), reaching their peak abundance in May and June; their numbers gradually diminish, until they are rarely collected during August and thereafter. In many parts of the eastern United States of America, A. americanum is one of the most frequently encountered ticks by humans and their pets. It has long been considered an important pest species. We now know that it is also an important vector for a variety of human pathogens, including those that cause human monocytic ehrlichiosis (HME), Ehrlichia ewingii erhlichiosis, and southern tick-associated rash illness (STARI) (Childs and Paddock 2003; Paddock and Yabsley 2007). Additional microorganisms have been isolated from lone star ticks in various parts of the United States of America (Francisella tularensis, Coxiella burnettii, Rickettsia parkeri, Rickettsia amblyommii, Panola Mountain ehrlichiosis, and lone star tick virus), but the role of lone star tick transmission and its potential to cause human illness require further investigation (Paddock and Yabsley 2007). Because the lone star tick may be translocated and appears in many parts of Canada, it is important to consider these many infectious agents as possible sources of disease. It is also known to cause tick paralysis in humans.

Distribution. The lone star tick is found in the southern United States of America from central Texas to Missouri, and east to the Atlantic states. Despite many references to the occurrence of this tick throughout Central America and South America, we accept the contention of Guglielmone et al. (2003) that it is restricted to the Nearctic Region. It is not known to have become established in Canada, but there are numerous records from Newfoundland to British Columbia from humans as a result of travel to the United States of America. This tick commonly infests ground-dwelling birds, and migratory activities of these hosts result in wide-spread dispersal of A. americanum into eastern and central Canada.

Hosts. This tick is known from a wide variety of mammals and birds. It can be an important pest on domestic animals, including cattle, horses, and dogs, and all instars may be found attacking humans.

Amblyomma maculatum Koch Gulf coast tick (Fig. 21, Map 2) Amblyomma maculatum Koch, 1844: 227. Amblyomma tigrinum Koch, 1844: 227. Amblyomma rubripes Koch, 1844: 227. Amblyomma ovatum Koch, 1844: 228. Amblyomma triste Koch, 1844: 229. Amblyomma complanatum Berlese, 1888: 191. Amblyomma maculatum Koch: Banks 1908: 39. Amblyomma maculatum Koch: Cooley and Kohls 1944.

Adult. Female: Body: Length, 3.7–4.4 mm unfed, reaching to 18.0 mm engorged. Gnathosoma: Porose areas of basis capituli oval, slightly divergent anteriorly, separated by interval of nearly twice the width of one porose area (Fig. 21A). Dorsal posterior margin of basis capituli slightly concave, with narrowly rounded corners or small cornua (Fig. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 67

21A). Hypostome slightly clavate, notched apically; dentition 3/3 or 4/4 along apical half of shaft (Fig. 21C). Palpus moderately long, with femur about 1.6 times as long as genu. Scutum: Length about 1.1 greater than width, subtriangular, widest anterior to midlength (Fig. 21A). Scapulae pointed. Cervical grooves moderately deep anteriorly, fading posteriorly well before reaching posterolateral margins. Surface with puncta numerous, large and small puncta intermingled, large puncta more numerous in areas anterior to eyes; ornate, with light and dark colour pattern contrasting sharply. Legs: Coxa I with a long, pointed external spur and a very short, blunt internal spur (Fig. 21B). Coxae II–IV each with a broad, short, plate-like spur; progressively smaller from II–IV. Tibia of legs II–IV with a pair of long, ventral terminal spurs, directed distally.

Adult. Male: Body: Length from 3.40–5.00 mm; oval, widest at midlength. Gnathosoma: Basis capituli in dorsal view subquadrate, with moderately short cornua (Fig. 21D). Hypostome as in female, notched terminally and with dentition 3/3 along apical half of length (Figs. 21F–G). Palpus moderately elongate as in female. Scutum: Surface with puncta numerous and deep in area within marginal grooves, sparse on festoons; ornamented with numerous, mostly connected linear spots of golden white (Fig. 21D). Cervical grooves deep anteriorly, shallow and disappearing behind eyes. Marginal grooves deep, complete, starting near eyes, and their continuation delimiting festoons. Festoons mostly longer than wide, except posteriormost three as wide as long (Fig. 21D). Legs: Apical and subapical ventral spurs absent on tarsus I, present on tarsi II–IV. Tibiae II–IV each with long ventral apical spur. Coxa I with long, pointed external spur; internal spur short, almost negligible. Coxae II and III each with short, broad, flat spur. Coxa IV with a long, pointed internal spur (Fig. 21E).

Nymph. Gnathosoma: Basis capituli triangular in dorsal view, its posterolateral corners protruding as acute points laterally beyond scapulae (Fig. 21H). Ventral surface of basis with 2 distinct retrograde spurs (Fig. 21J). Hypostome slightly clavate, rounded apically; dentition 2/2 along apical half of length (Fig. 21I). Palpus long, with femur about 2.5 times as long as genu. Scutum: Length slightly less (0.9) than width; shape suboval, widest at level of eyes, slightly posterior to middle, with slightly convex posterolateral margins (Fig. 21H). Scapulae rounded, covered by posterior angles of basis capituli. Cervical grooves moderately deep anteriorly, continuing as shallow depressions nearly to posterolateral margins. Surface sparsely punctate. Legs: Coxa I with short, blunt internal spur, but lacking external spur (Fig. 21K). Coxa II with small, broad, external spur. Coxae III and IV without spurs (Fig. 21K).

Larva. Body: Broad, oval; widest posterior to midlength (Figs. 21L–M). Length, excluding gnathosoma, 0.50–0.63 mm. Gnathosoma: Dorsal surface triangular, with posterolateral corners acute; anterolateral margins sloped over area of palpal articulation (Fig. 21L); with 1 pair sensilla hastiformia. Ventrally with lateral edges rounded, indented slightly below midlength; with ventral ridge. Hypostome slightly clavate; dentition 2/2 with 6–8 teeth per file (Fig. 21N). Palpal genu with small, blunt spur on ventral surface.Scutum: Length approximately two-thirds (0.66) width, widest at level of eyes, posterior to midlength (Fig. 21L). Cervical grooves shallow, diverging posteriorly, not reaching posterolateral margins. Eleven festoons, wider than long (Fig. 21L). Legs: Coxa I with one broad, blunt spur posteromedially (Fig. 21M). Coxae II and III each with single, short, bluntly rounded spur near middle of posterior margin (Fig. 21M). 68 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A B

F H

E G

K N

L M

Figs. 21A–N. Amblyomma maculatum. Fig. 21A, female gnathosoma and scutum, dorsal; Fig. 21B, female coxae and trochanters, ventral; Fig. 21C, female hypostome, ventral; Fig. 21D, male gnathosoma and scutum, dorsal; Fig. 21E, male coxae and trochanters, ventral; Fig. 21F, male hypostome, ventral; Fig. 21G, enlargement of distal tip of male hypostome, ventral; Fig. 21H, nymph gnathosoma and scutum, dorsal; Fig. 21I, nymph hypostome, ventral; Fig. 21J, spurs on venter of basis capituli, nymph; Fig. 21K, nymph coxae and trochanters, ventral; Fig. 21L, larva, dorsal; Fig. 21M, larva, ventral; Fig. 21N, larva hypostome, ventral. Figs. 21A–K, N redrawn from Cooley and Kohls (1944); Figs. 21L, M redrawn from Clifford et al. (1961). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 69

Natural history. This species has been of historical importance as a pest throughout its natural breeding range along the Gulf of Mexico. The adults commonly attack cattle, on which they often feed in greatest numbers in the ears. They cause considerable irritation and lesions at their feeding sites which serve as an avenue for myiasis caused by the primary screwworm fly,Cochliomyia hominivorax (Coquerel) (Diptera: Calliphoridae) (Spicer and Dove 1938). Following eradication of the screwworm fly in the United States of America, this secondary impact is no longer a concern, but adult A. maculatum still are an important pest of cattle. The larvae and nymphs of A. maculatum feed predominately on ground-foraging birds and small mammals. Its predilection for birds is one reason we see this species in Canada (Scott et al. 2001). In the Gulf States, adults appear early in spring and may be present on large mammals until late summer, though their peak in abundance is usually June to early July (Hixson 1940; Semtner and Hair 1973); their abundance may remain high until September in some areas (Bishopp and Hixson 1936). Eggs are laid within two weeks following female engorgement, and larvae hatch about four weeks later. The peak in larval abundance may occur from late July to September (Bishopp and Hixson 1936; Semtner and Hair 1973); in coastal areas of Texas, Teel et al. (1988; 1998) observed greatest numbers of larvae on meadowlarks during November to January. Ground-foraging birds (e.g., meadowlarks and bobwhite quail) and small mammals are important hosts, and larvae attach to the head and neck regions (Hixson 1940). In some parts of its range, nymphs may be collected in every month of the year (Hixson 1940), while elsewhere, the season may be more limited, with peak abundance in late summer or early autumn (Bishopp and Hixson 1936; Semtner and Hair 1973) or during December or February to March (Teel et al. 1988; 1998). Amblyomma maculatum is not widely known for its ability to transmit pathogens. It is a vector of Rickettsia parkeri (Parker et al. 1939; Sumner et al. 2007), an agent that may cause disease in humans (Paddock et al. 2004). It has also been demonstrated as an intermediate host for Hepatozoon americanum Vincent-Johnson et al. (Eucoccidiorida: Hepatozoidae), the agent of American canine hepatozoonosis in dogs (Mathew et al. 1998; Ewing et al. 2002), and as an agent of tick paralysis in humans (Paffenbarger 1950).

Distribution. The Gulf Coast tick is found mainly in the southeastern United States of America bordering the Gulf of Mexico and the Atlantic coast. Its distribution extends through Mexico into northern South America. It is not known to be established in Canada, and Ontario records are likely the result of travel outside of Canada, or dispersal into Canada on migratory birds.

Hosts. Juvenile instars feed primarily on ground-dwelling birds and a variety of small mammals. Adults feed on large mammals, including humans.

Genus Dermacentor Koch (Figs. 22–24) Anus of adults with postanal groove (Figs. 22A, B), sometimes indistinct on nymphs. Eyes present (Figs. 22A–B). Posteromarginal festoons present. Palpus with 2nd palpomere (femur) not appreciably longer than wide, and not produced laterally (Fig. 23A). Basis capituli in dorsal view subrectangular, roughly parallel-sided on adults, but commonly subtriangular, with posteriorly flared sides on nymphs (Fig. 23C). Adult females and males usually with ornate scutum. Foveae present. Adult males without ventral plates. Coxa I 70 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Map 2. Collection localities for Amblyomma maculatum in Canada.

of adults appearing bifid, with similarly elongate and adjacent internal and external spurs (Fig. 22A). Coxae I–IV increasing in size progressively (Fig. 23A). Larvae with 1 pair of posthypostomal setae; scutum with 3 pairs of setae; 8 pairs of marginal dorsal setae, including 3 pairs anterior to sensilla sagittiformia; 2 pairs of central dorsal setae; 9 festoons present (Fig. 23D).

Key to species of the genus Dermacentor in Canada

1. Adult, female and male: spiracular plate with pronounced postero-dorsal prolongation and with numerous goblets, surrounded by 1 or more peripheral rings of smaller pore- like structures (Figs. 23A, 24A); nymph: coxa I with internal spur distinct, tapered (Fig. 24C)); nymph and larva: basis capituli in dorsal view with lateral margins drawn out to points that extend laterally beyond anterior edges of scapulae (Figs. 23C, D); larva: coxa I with spur distinct, tapered (Fig. 23D)...... 2 – Adult, female and male: spiracular plate with slight or no postero-dorsal prolongation and with relatively few goblets (fewer than 100), not surrounded by several peripheral rings of smaller pore-like structures (Figs. 22A, B); nymph: coxa I with internal spur faint, broadly rounded (Fig. 22C); nymph and larva: basis capituli in dorsal view with lateral margins blunt, not drawn out to extend laterally beyond scapulae (Fig. 22C); larva: coxa I with spur faint, broadly rounded (Fig. 22D) ...... Dermacentor albipictus 2. Adult, female and male: spiracular plate with goblets numerous (greater than 300) and nearly as small as pore-like structures in peripheral rings (Figs. 24A (insert), B); nymph: coxa IV without or with faint external spur (Fig. 24C); larva: palpal femur lacking pore- like opening on dorsodistal surface (Fig. 24D); basis capituli in dorsal view with posterior edge of lateral points often slightly concave...... Dermacentor variabilis – Adult, female and male: spiracular plate with goblets moderately numerous (fewer than 200) and over twice as large as pore-like structures in 1 or 2 peripheral rings (Figs. 23A, B); nymph: coxa IV with small but obvious external spur (Fig. 23C); larva: palpal femur with pore-like opening on dorsodistal surface (Fig. 23D); basis capituli in dorsal view with posterior edge of lateral points often slightly convex ...... Dermacentor andersoni A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 71

Dermacentor albipictus (Packard) Moose or winter tick (Fig. 22, Map 3) Ixodes albipictus Packard, 1869: 65. Ixodes nigrolineatus Packard, 1869: 66. Dermacentor variegatus Marx and Neumann in Neumann, 1897: 367. Dermacentor reticulatus Fabricius: Salmon and Stiles 1901: 448. Dermacentor variegatus kamshadalus Neumann, 1908: 73. Dermacentor albipictus Packard: Banks 1908: 44. Dermacentor nigrolineatus Packard: Banks 1908: 48. Dermacentor salmoni Stiles, 1910: 55. Dermacentor albipictus Packard: Bishopp and Wood 1913: 161. Dermacentor albipictus (Packard): Cooley 1938: 59 (all instars described). Dermacentor kamshadalus Neumann, 1908: Apanaskevich 2013: 692.

Adult. Female: Body: Length: 2.8–5.5 mm unfed, reaching to 17.0 mm engorged. Base colour brown, pattern colour silvery-grey. Gnathosoma: Porose areas of basis capituli irregular in shape, moderately depressed. Cornua variable in length, subacute or rounded apically (Fig. 22A). Scutum: Longer than wide, with section posterior to eyes narrower than in other two species (Fig. 22A). Cervical grooves varying from short and rounded to moderately elongate. Surface with scattered puncta not markedly different in size. Extent of pattern colour highly variable, from predominant to nearly absent; base colour sometimes limited to spots around eyes, cervical grooves, and pair of lines parallel and lateral to those grooves. Spiracular plate: Subcircular to oval, with dorsal prolongation absent, or broad and blunt if present (Fig. 22A, insert). Goblet cells relatively few (compared to D. andersoni and D. variabilis), of constant large size, not usually surrounded by peripheral rings of smaller pore-like perforations near frame (Fig. 22A, insert). Legs: Coxa I with elongate internal and external spurs, their opposing margins parallel or slightly divergent; internal spur much broader and slightly shorter than external spur (Fig. 22A). Small but distinct internal spur present on coxae II and III, indistinct or absent on coxa IV. Coxae II–IV each with large, acute external spur.

Adult. Male: Body: Length: 2.8–6.1 mm. Silvery-grey pattern colour variable as in female. Gnathosoma: Cornua prominent (Fig. 22B), slightly longer than in female. Palpal apex blunt, somewhat flattened. Scutum: Cervical grooves short, about twice as long as broad (Fig. 22B). Surface with puncta small, differing little in size, often indistinct. Spiracular plate: Shape and goblet cell content as in female (Fig. 22B). Legs: Coxae usually progressively enlarged from I–IV, IV sometimes disproportionately enlarged (Fig. 22B). Coxae I–IV with spurs as in female.

Nymph. Body: Length: 1.5–2.3 mm. Gnathosoma: Basis capituli in dorsal view subquadrangular, with blunt lateral margins not extending beyond apices of scapulae; posterolateral corners of posterior margin not developed as acute cornua (Fig. 22C). Ventrally, basis with pair of short, blunt spurs posterolaterally. Scutum: Slightly longer than wide, with slightly angled posterior margin (Fig. 22C). Cervical grooves long, narrow. Spiracular plate: Large, subcircular, with relatively few (15–20, occasionally approximately 45) large, distinct goblet cells (Fig. 22C). Legs: Coxa I with blunt, moderate- sized external spur and small, blunt internal spur. Coxae II–IV each with small, blunt external spur, sometimes indistinct on IV (Fig. 22C). Coxae II–IV lacking internal spurs. 72 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A B

C D

Figs. 22A–D. Dermacentor albipictus gnathosoma and idiosoma, dorsal view (left), ventral view (right). Fig. 22A, female; insert, enlarged spiracle; Fig. 22B, male; Fig. 22C, nymph; Fig. 22D, larva. Figs. 22A modified, B–D redrawn from Brinton et al. (1965); insert redrawn from Yunker et al. (1986). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 73

Larva. Body: Length approximately 0.60 mm. Gnathosoma: Basis capituli in dorsal view with lateral margins forming bluntly pointed posterolateral corners not extending beyond apices of scapulae (Fig. 22D). Palpal femur lacking dorsal gland pore. Scutum: Length about three-fourths (0.75) width, widest at level of eyes, slightly posterior to midlength (Fig. 22D). Legs: Coxa I with weak, broadly rounded spur. Coxae II and III each with posteromarginal spur faint or absent (Fig. 22D).

Natural history. Because all active instars of this tick infest primarily moose (Alces alces (Linnaeus) ) (Anderson and Lankester 1974) and elk (Cervus canadensis (Linnaeus)) (Flook and Stenton 1969; Samuel et al. 1991; Smits 1991), during the winter months, D. albipictus is commonly known as the winter tick, or moose tick (also known, however, as the horse tick, or elk tick) (Bishopp and Wood 1913). The density of D. albipictus (ticks/cm2) on moose is greatest on the neck, mane, withers, shoulders, and abdomen compared to other body regions such as the back and sides (Addison et al. 1979; Glines and Samuel 1984). Similar sites for attachment were selected by D. albipictus on elk (Samuel et al. 1991) and black-tailed deer (Cowan 1946). Some populations of this tick from mountain goats and sheep have been recently viewed as a morphologically distinct species, under the reinstated name, Dermacentor kamshadalus Neumann (Apanaskevich 2013). Adults of this form are distinguished from those of typical D. albipictus by several attributes which, however, may vary considerably among populations (e.g., shorter and broader dorsal spur on trochanter I, less dense and shorter setae on idiosoma, usually more ornate scutal ornamentation, shorter and more posterolaterally-directed spurs on coxa I, shorter and more robust cornua, and more numerous large goblet cells on spiracular plates). The male is also distinguished by having more robust legs, and the female by shorter and broader spurs on coxae II and III. While the nymph is distinguished preliminarily by a shorter external spur on coxa I and by smaller, more numerous goblet cells on the spiracular plates than in other D. albipictus, the larva of this form has not been described. Perry (2014) was unable to separate D. kamshadalus from D. albipictus using molecular techniques. In the absence of comprehensive hybridisation and assortative mating studies or molecular evidence distinguishing these two forms, and in view of overlaps in habitat, geographic distribution, and host preferences, we do not include D. kamshadalus as a species distinct from D. albipictus. Our diagnosis is modified to accommodate the two forms. Throughout its range in Canada, D. albipictus completes its life cycle in one year; however, in warmer regions of the country such as coastal British Columbia (Cowan 1946), animals can be free of ticks earlier in the year than animals from inland or more northern localities (Glines and Samuel 1984). Dermacentor albipictus is the only one-host ixodid tick found in Canada. Both larvae and nymphs feed and moult on one individual host (Bishopp and Trembley 1945). In Canada, winter ticks have also been collected from a variety of other mammals including: mule deer, Odocoileus hemionus (Rafinesque) (Samuelet al. 1981; Welch et al. 1991; Wilkinson 1970), white-tailed deer, Odocoileus virginianus (Zimmermann) (Gregson 1956), woodland caribou, Rangifer tarandus caribou (Gmelin), reindeer, Rangifer tarandus tarandus (Linnaeus) (Welch et al. 1990), plains buffalo or bison, Bison bison bison (Linnaeus), wood buffalo or bison, B. b. athabascae Rhoads (Samuel and Welch 1991) and mountain sheep, Ovis canadensis Shaw. Among domestic animals, horses and cattle are occasionally severely infested (Bruce 1927; Cameron and Fulton 1927; Bishopp and Trembley 1945). Although many mammal species are likely suitable as hosts, the engorgement and reproductive success of female winter ticks are greatest when they are fed on moose (Welch et al. 1991). Winter ticks rarely bite people 74 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins and bites usually occur when adult ticks transfer from wild or domestic hosts onto people (Cameron and Fulton 1927; Bequaert 1945; Bishopp and Trembley 1945). The on-host component of the life cycle of D. albipictus begins when larvae contact a suitable host. Following emergence from eggs, host-seeking or “questing” by larvae is delayed for about two weeks (Drew and Samuel 1985); typically larvae begin to quest actively in early to mid-September, and this behaviour continues through most of November, declining rapidly in December (Wilkinson 1967; Drew and Samuel 1985). Larvae usually aggregate in clumps of several thousand individuals on the tips of plants and shrubs (Drew and Samuel 1985) or inanimate objects such as fencing (Wilkinson et al. 1982). Most aggregations of larvae are 1.0–1.5 m above the ground (Drew and Samuel 1985). Snowfall in combination with average temperature below 0°C inactivates larvae that have not secured a host (Drew and Samuel 1985). Thus, larvae present on vegetation in November, December, or later are likely too sluggish to transfer easily to a host and have little chance of successful attachment (Samuel and Welch 1991). Adult D. albipictus can be found on naturally infested hosts from December to May (Cameron and Fulton 1927; Addison et al. 1979; Glines and Samuel 1984). Males precede females by 2–3 weeks and persist much longer, comprising the majority of adult ticks present on hosts in May (Addison and McLaughlin 1988; Drew and Samuel 1989). Mating occurs on the host shortly after female emergence (Cameron and Fulton 1927) and females must be mated in order to complete engorgement (Oliver 1974). Adult females begin to appear in late January, reaching peak abundance by late March and early April (Addison and McLaughlin 1988; Drew and Samuel 1989). Some females can feed in late February; however, most will engorge and drop from their hosts from late March to early May (Glines and Samuel 1984; Addison and McLaughlin 1988; Drew and Samuel 1989; Welch et al. 1991). Virtually all moose examined in the northern part of their range during the winter months are infested with D. albipictus (Addison et al. 1979; Samuel and Barker 1979; Glines and Samuel 1984; Hoeve et al. 1988; Welch et al. 1991; Samuel and Welch 1991). The prevalence of infestation of winter ticks on other hosts varies by geographic location but is generally less than that observed on moose (Samuel and Welch 1991). Typically moose carry larger numbers and more winter ticks/cm2 than other sympatric host species such as elk and white-tailed deer (Samuel and Barker 1979; Samuel et al. 1991; Welch et al. 1991). In south-central Ontario, moose have been collected with an average of 3852 winter ticks per animal (Addison et al. 1979), whereas in eastern Ontario, moose carry 24–1250 winter ticks (Hoeve et al. 1988). In the western provinces of Manitoba, Alberta, and British Columbia, 212 moose sampled all carried > 25 250 ticks and the density of winter ticks exceeded 1.1 ticks/cm2 on every animal (Samuel and Welch 1991). Over 18% of the moose had infestations greater than 50 000, 6% carried more than 80 000 ticks and the maximum number of ticks infesting one moose was 149 916 ticks (Samuel and Welch 1991). The largest infestation of winter ticks reported from a wild moose is 178 000 (Samuel and Barker 1979). In Ontario and Alberta, during some winters, many moose have been found dead with large numbers of ticks (> 100 000) (Samuel and Barker 1979; Addison and Smith 1981; Hoeve et al. 1988). Although large numbers of winter ticks undoubtedly represent a metabolic cost to moose (e.g., irritation, blood and hair loss, disrupted feeding and resting behaviour (Samuel and Welch 1991)), direct evidence that winter ticks cause the death of heavily infested animals is lacking (Lankester 1987). The off-host component of the life cycle of D. albipictus begins when replete females drop from their hosts over a period of 9–10 weeks (Drew and Samuel 1989). Engorged females do not disperse large distances following disengagement (Drew and Samuel 1986), and as a result, eggs are usually deposited under leaf litter or on the soil in the immediate A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 75

vicinity of where they dropped from their hosts. Depending on the type of habitat where females drop from their hosts, 13–37% of engorged females survive to deposit eggs (Drew and Samuel 1986). Survival of engorged females is also correlated with the amount of snowmelt and ambient temperature at the time of disengagement. Under field conditions in Alberta, most females initiate egg laying by the first week in June (Drew and Samuel 1986). The number of eggs deposited by females is positively correlated with the weight of fed females (Addison and Smith 1981; Drummond et al. 1969). Replete females from experimentally infested animals weigh 420–690 mg and produce an average of 3000–5000 eggs (Drummond et al. 1969; Wright 1971; Ernst and Gladney 1975; Drew and Samuel 1986) and occasionally as many as 10 000 eggs (Addison and Smith 1981). The proportion of eggs that produce larvae ranges from 23–59% (Drew and Samuel 1986) and larvae hatch in late August to mid-October depending on the date females detach from hosts (Glines and Samuel 1984; Drew and Samuel 1986). Duration of the incubation period for eggs appears to be controlled by temperature rather than photoperiod (Patrick and Hair 1975; Drew and Samuel 1986). The longevity of unfed larvae is partly determined by habitat type and the associated microclimatic variation among habitats. Drew and Samuel (1986) reported that 6–36% of larvae that hatched from eggs laid during June in central Alberta were still alive in mid-November. Painful swellings can occur at the site of D. albipictus bites (Cameron and Fulton 1927); however, because they bite humans infrequently, winter ticks have little potential to transfer pathogens directly to humans. Borrelia burgdorferi, the pathogen responsible for Lyme borreliosis, has been isolated from a small percentage of D. albipictus removed from white-tailed deer in B. burgdorferi-endemic regions of the United States of America (Magnarelli et al. 1986, Kocan et al. 1992). However, because D. albipictus is a one-host tick that rarely bites people it has limited potential to transmit B. burgdorferi to humans. Dermacentor albipictus has a limited capacity to transmit pathogens to wild and domesticated animals because winter ticks are one-host ticks and the rates of transovarial transmission of pathogens appear to be low. Winter ticks have been shown experimentally to be a vector of Anaplasma marginale (Stiller et al. 1981), the blood parasite responsible for bovine anaplasmosis. This bacterium can be transferred transstadially from larva to nymph (Stiller et al. 1989; Stiller and Coan 1995). Interestingly, A. marginale failed to be

Map 3. Collection localities for Dermacentor albipictus in Canada. 76 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins transferred transovarially in D. albipictus (Bram and Roby 1970; Stiller et al. 1981), thus the role of this tick as a maintenance vector of A. marginale is likely minimal. Cases of bovine anaplasmosis with onset during the winter months, when no other arthropods are active, also provide circumstantial evidence that winter ticks may have a minor role in transmission of A. marginale to cattle (Alderink and Deitrich 1982; Zaugg 1990). Dermacentor albipictus has also been shown experimentally to act as a vector of a related bacterium, Anaplasma ovis, to sheep (Stiller et al. 1987). Stiller et al. (1980) reported that winter ticks can transmit the equine piroplasm, Babesia caballi (Nuttall and Strickland), to horses and that this pathogen is successfully transmitted transovarially in D. albipictus. The bacterium, Erysipelothrix rhusiopathiae (Migula) Buchanan (Erysipelotrichales: Erysipelotrichidae) has been isolated from moose found dead in Ontario; however, the role, if any, played by D. albipictus in transmission is unknown (Campbell et al. 1994). Francisella-like endosymbionts have been detected in D. albipictus in several locations, including Alberta and Minnesota (Scoles 2004; Balbridge et al. 2009), and were shown by Balbridge et al. (2009) to be transovarially transmitted. Balbridge et al. (2009) isolated A. phagocytophilum in D. albipictus in Minnesota, also being transovarially transmitted.

Distribution. The winter tick is one of the most widely distributed ticks in North America. The geographic distribution and to a lesser extent local abundance of D. albipictus are closely linked with the availability of moose as hosts (Samuel 1989; Zarnke et al. 1990). In Canada, D. albipictus occurs from the east coast to the west coast (Bequaert 1945; Gregson 1956; Wilkinson 1967; Kennedy 1986; Kennedy and Newman 1986), though there are no records for Newfoundland. The northernmost records for D. albipictus are Fort Liard and Fort Smith, Northwest Territories, both of which are close to 60°N (Wilkinson 1967). Moose from the Yukon Territory (as far north as 62°N) have been observed with premature winter hair loss, providing circumstantial evidence that winter ticks are established there (Samuel 1989). In the United States of America, winter ticks are found from coast to coast, southward into Texas. Winter ticks have also been reported from northern Mexico (Bishopp and Trembley 1945) and Central America (Yunker et al. 1986). Dermacentor albipictus has on occasion been detected on livestock exported from North America into Europe and is a potential threat for establishment there (Lillehaug et al. 2002).

Hosts. This species can be present in enormous numbers on moose, but it is also found on elk, deer, cattle, bison, mountain goat, mountain sheep, woodland caribou, and horses. The larvae may infest other animals, including humans, which blunder into them in the fall, but established infestations on these atypical hosts are unknown. Records on large carnivores can be expected as a result of their feeding on carcasses of infested ungulates.

Dermacentor andersoni Stiles Rocky Mountain wood tick (Fig. 23, Map 4) Dermacentor venustus Marx, in Neumann 1897: 365. Dermacentor venustus Marx: Banks 1908: 46. Dermacentor andersoni Stiles, 1908: 949. Dermacentor modestus Banks, 1909: 170. Dermacentor andersoni Stiles: Cooley 1938: 31 (all instars described). Dermacentor andersoni Stiles, 1908: International Commission on Zoological Nomenclature 1978: 88. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 77

Adult. Female: Body: Length: 2.8–5.4 mm unfed, reaching to 16.0 mm engorged. Base colour brown, pattern colour silvery grey. Gnathosoma: Porose areas of basis capituli oval or bean-shaped, depressed. Cornua usually shorter than width at base, rounded apically (Fig. 23A). Scutum: Nearly as long as wide, with posterolateral margins slightly concave behind eyes, posterior margin bluntly angular (Fig. 23A). Cervical grooves elongate. Surface with few, large, deep puncta and numerous small, inconspicuous puncta. Extent of pattern colour highly variable, from extensive to nearly absent. Spiracular plate: Oval, with distinct, moderately broad to slender dorsal prolongation. Goblet cells numerous (typically 100–200), surrounded by usually 1 or 2 peripheral rings of smaller pore-like perforations near frame and on dorsal prolongation (Fig. 23A). Legs: Coxa I with elongate internal and external spurs, their opposing margins parallel; internal spur broader and slightly shorter than external spur (Fig. 23A, insert). Small, rounded internal spur present on coxae II and III, indistinct or absent on coxa IV. Coxae II–IV each with short, bluntly pointed external spur, about as broad at base as long, progressively smaller from II–IV.

Adult. Male: Body: Length: 2.5–6.1 mm. Colour ornamentation variable, as in female. Gnathosoma: Cornua moderately small, subacute (Fig. 23B). Palpal apex broadly to narrowly rounded. Scutum: Cervical grooves 3 times as long as broad. Surface with puncta of markedly different, moderate to large sizes. Spiracular plate: Shape and goblet cell content as in female; goblet cells of moderate size around macula, becoming progressively finer toward periphery of plate (Fig. 23B). Legs: Coxae usually progressively enlarged from I–IV, IV sometimes disproportionately enlarged posteriorly (Fig. 23B). Coxae I–IV with spurs formed much as in female, but with internal spur smaller on coxae II and III, and indistinct on coxa IV (Fig. 23B).

Nymph: Body: Length: 1.30–1.45 mm. Gnathosoma: Basis capituli in dorsal view subtriangular, with lateral margins drawn out to acute points that extend beyond apices of scapulae (Fig. 23C). Ventrally, basis with pair of short, bluntly pointed to rounded spurs posterolaterally. Scutum: Subequally as long as wide, with prominently curved posterior margin (Fig. 23C). Cervical grooves long, narrow. Spiracular plate: Large, subcircular. A ring of moderately large goblet cells around macula, remaining goblet cells numerous, smaller (Fig. 23C). Legs: Coxa I with bluntly pointed external spur much larger than small but distinct internal spur (Fig. 23C). Coxae II–IV each with small, blunt external spur, sometimes indistinct on IV. Coxae II–IV lacking internal spurs.

Larva: Body: Length: 0.57–0.66 mm. Gnathosoma: Basis capituli in dorsal view with lateral margins drawn out posterolaterally to acute points that extend beyond apices of scapulae (Fig. 23D). Palpal femur with dorsal gland pore (Fig. 23D). Scutum: Length about two-thirds (0.67) width, widest at level of eyes, well behind midlength (Fig. 23D). Legs: Coxa I with distinct, bluntly triangular internal spur (Fig. 23D). Coxae II and III each with small, broad spur posteromarginally.

Natural history. The distribution of this tick is centred within the Rocky Mountains of western North America (Cooley 1938; Bishopp and Trembley 1945; Kennedy 1986; James et al. 2006) and hence this tick is commonly known as the Rocky Mountain wood tick (RMWT). Throughout its range in Canada, D. andersoni typically requires at least two years to complete its life cycle. Unfed adult ticks which engorge from March to May will lay eggs that develop into host-seeking larvae in June and July. Larvae feed on hosts in July and 78 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A B

C D

Figs. 23A–D. Dermacentor andersoni gnathosoma and idiosoma, dorsal view (left), ventral view (right). Fig. 23A, female; insert, enlarged spiracle; Fig. 23B, male; Fig. 23C, nymph; Fig. 23D, larva. Figs. 23A modified, B–D redrawn from Brinton et al. (1965); insert redrawn from Yunker et al. (1986). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 79

August and moult to the nymphal instar in August and September. Most of these nymphs will overwinter and begin to seek hosts in early spring and infest hosts from March to the end of May. Engorged nymphs moult to the adult instar and adults overwinter prior to becoming active in the following spring (Wilkinson 1967, 1968, 1979). Under experimental field conditions, the life cycle can be completed in one year. That is, larvae and nymphs feed and moult in the same year and adults overwinter prior to seeking hosts (Wilkinson 1968). Under natural conditions, it is unlikely that the life cycle is often completed this rapidly. At localities at high elevation in Alberta or British Columbia and at some open prairie sites in Saskatchewan, cooler climatic conditions may extend the life cycle to three or more years. Likewise, the life cycle will exceed two years in areas where availability of hosts is low (Wilkinson 1968, 1979). In most endemic areas in the United States of America, the life cycle of D. andersoni is similar to that described for Canada. That is, the life cycle is typically completed in two years, can rarely be completed in one year (Eads and Smith 1983) and can be extended to three or more years in some years and localities (Bishopp and King 1913; Burgdorfer 1969, 1977; Furman and Loomis 1984). Dermacentor andersoni is a three-host tick. Unfed adults of D. andersoni overwinter and become active shortly after snow melt, which varies from late February in some areas of British Columbia to late March to early May in southern Alberta and south-central Saskatchewan. Males usually appear before, and disappear sooner than, females (Gregson 1951). Host-seeking populations usually peak in late March in British Columbia (Hearle 1938; Gregson 1951; Wilkinson and Gregson 1985), or late May to early June in Alberta and Saskatchewan (Holland 1940; Brown 1944; Gregson 1956). In certain localities near Kamloops, British Columbia, adults abruptly disappear by mid-May each spring, though in damper areas of the province adults remain active until June (Gregson 1951). In all provinces, adult activity declines to a minimum by mid-July, though small numbers of adults can occasionally be collected in August (Hearle 1938; Brown 1944) and later (Wilkinson 1979). Some unfed adults can survive through a second winter if hosts are not found in the first year of activity (Gregson 1951; Wilkinson 1968). Adults typically climb to the tips of low vegetation in search of hosts (Gregson 1960; Burgdorfer 1969) and often quest in the “head down” position (Holland 1940; Brown 1944). Based on extensive drag sampling in southern Alberta, Holland (1940) reported that adults were most active on dull, warm, humid days, and peak daily activity occurred in the late afternoon or early evening. Adults are also frequently collected at night (Brown 1944). In montane habitats, adults of D. andersoni are frequently collected along clearings such as power lines or logging roads, and can be concentrated in relatively small areas near shrubby, rocky outcrops in open grassy areas (Wilkinson 1967, 1971; Schaalje and Wilkinson 1985). In British Columbia, unfed adults of D. andersoni were associated with habitats having an abundance of saskatoon (Amelanchier alnifolia (Nuttall) Nuttall ex Roemer (Rosales: Rosaceae)) and rose bushes (Rosales: Rosaceae), presumably reflecting use of these microhabitats by hosts for nymphs (Schaalje and Wilkinson 1985; Laurance and Coan 1987). In prairie habitats, adults can be found in open rangelands, but are generally most abundant on sparse vegetation within dried river beds or coulees (Holland 1940). Because of high densities of hosts for immature RMWT, campgrounds can often be foci for tick populations both in Canada (Wilkinson 1967) and the United States of America (McLean et al. 1981). In Canada, adults of Dermacentor andersoni are common on medium to large domestic mammals, such as horses, cattle, sheep, and dogs (Hearle 1938; Gregson 1956; Wilkinson 80 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

1970; Kennedy 1986) and wild mammals, such as mountain sheep and goats (Bruce 1920; Buckell 1934; Kerr and Holmes 1966); grizzly bears (Hewitt 1915); and coyotes, porcupines, and rabbits (Bruce 1920; Brown 1944). Humans are readily bitten by male and female D. andersoni (Gregson 1956; Kennedy and Newman 1986) and surprisingly, this instar has occasionally been removed from various ground-dwelling bird species (Cooley 1938; Kennedy 1986). Mule deer, moose, yellow-bellied marmots and Columbian ground squirrels are occasionally hosts for adults of D. andersoni (Brown 1944; Gregson 1958; Wilkinson 1965, 1970). In the laboratory, adults of D. andersoni have been successfully fed on mice and guinea pigs (Gregson 1960; Scott and Brown 1986), illustrating that this instar lacks host specificity. Upon contacting a host, adults usually attach to feed on the back-line, head, and neck regions (Gregson 1960). For example, Gregson (1952) reported that most adults of D. andersoni attach to the withers or backs of the heads of cattle in British Columbia. Wilkinson (1972) observed that adults from montane habitats in British Columbia tended to attach to the back-line or upper sides of cattle whereas adults from prairie habitats in Saskatchewan and Alberta attached more frequently to the underside (e.g., brisket or dewlap). It was speculated that because ticks from montane habitats attach to hosts about one month earlier than prairie ticks, selection favours survival of ticks in more sheltered locations on hosts in the prairies (Wilkinson and Lawson 1965; Wilkinson 1985). On people, adults of RMWT usually attach to the scalp (Jellison and Gregson 1950), though they occasionally attach to other regions including the shoulders, back, and groin (Mail and Gregson 1938; Scholten 1977). Data on the abundance of adults of D. andersoni on hosts in Canada are scarce, but the average number of ticks infesting cattle ranges from 1–24 at peak abundance (Brown 1944; Wilkinson and Gregson 1985). Based on drag sampling, abundance of adults varies among habitats and years (Holland 1940; Wilkinson 1984; Wilkinson and Gregson 1985). Females of D. andersoni require 5–12 days (average seven days) to feed to repletion (Bruce 1920; Gregson 1935a; Brown 1944; Loomis 1961). Feeding is a prerequisite to mating for both sexes such that females and males must feed for a minimum of one and five days, respectively, prior to mating (Gregson 1947). Males are capable of mating with several females (Brown 1944) and mating occurs on the host (Feldman-Muhsam and Borut 1971). In the absence of males, females may feed for up to two weeks but will not engorge fully unless they are mated (Gregson 1960). Parthenogenesis is rare in D. andersoni and females that are not mated can lay eggs but they are usually sterile (Gregson 1947; Oliver 1974). Under experimental conditions, crosses between D. variabilis females and D. andersoni males produced viable progeny but not reciprocal crosses. Hybridisation of these two species in areas where their ranges overlap (i.e., central Saskatchewan) is presumably rare (Oliver et al. 1972). When engorged, females drop from their hosts and seek protected locations in which to lay their eggs, such as under stones or around the crown of grass plants (Brown 1944). Temperature influences the duration of the pre-oviposition and oviposition period in D. andersoni (Wilkinson 1967); thus, depending on when females are fed, eggs can be deposited from 1–6 weeks after females drop from their hosts. In parts of British Columbia, females usually begin to lay eggs by early May or June (Wilkinson 1968). Eggs may be deposited even later in the year in climatically cooler parts of the range of the tick, such as central Saskatchewan or at higher elevations in Alberta (Wilkinson 1979). Females produce an average of 4000 eggs (range 2500–7000) over a period of 17–51 days and once the eggs are laid, females die (Hewitt 1915; Bruce 1920; Brown 1944). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 81

Larvae hatch from eggs and infest hosts beginning in June or early July (Hearle 1938; Wilkinson 1984) and larvae have a limited capacity for active dispersal (Sonenshine et al. 1976). The dominant hosts for larvae vary among localities; however, small mammals such as squirrels (e.g., yellow-pine chipmunks, Tamias amoenus Allen; yellow-bellied marmots, Marmota flaviventris (Audubon and Bachman); Richardson’s ground squirrels, Urocitellus richardsonii (Sabine); and red squirrels, Tamiasciurus hudsonicus (Erxleben)), mice (e.g., deer mice, Peromyscus maniculatus (Wagner)), rabbits, and other rodents (i.e., bushy- tailed woodrats, Neotoma cinerea (Ord)) typically serve as hosts for larvae of D. andersoni (Bruce 1920; Hearle 1938; Burgess 1955; Gregson 1956; Wilkinson 1984). Because larvae are rarely observed on hosts in the spring (Wilkinson 1984) and most unfed larvae do not survive more than 10 weeks under field conditions (Sonenshineet al. 1976), in most, but not all, endemic areas of Canada, unfed larvae rarely overwinter (Wilkinson 1967, 1968, 1979). Larvae attach mainly to the ears, face, and neck regions of the body of their host (Hearle 1938; Wilkinson 1968) and feeding usually lasts from 2–8 days (Hewitt 1915; Loomis 1961; Scott and Brown 1986). The abundance and prevalence of larvae on small mammal hosts usually peaks during July and August, and declines rapidly in September, though larvae can be collected from May to October in some localities (Hearle 1938; Wilkinson 1967, 1984). Larvae fed on hosts during July and August may moult to the nymphal instar prior to overwintering (Wilkinson 1967, 1968, 1984). Overwintered nymphs begin to seek hosts as soon as the snow melts, typically from March to early May, depending on geographic locality (Wilkinson 1967, 1968, 1984). Nymphs infest the same species of small mammals as larvae (Hearle 1938; Brown 1944; Gregson 1956). Though nymphs have been observed on hosts throughout the active season (i.e., March to September), their abundance usually peaks in spring (i.e., March to May) and summer (i.e., July and August) (Wilkinson 1968, 1984). The spring peak is composed mainly of overwintered nymphs (i.e., larvae fed in the previous year), whereas the summer peak consists of nymphs which fed as larvae in the same year (Wilkinson 1967, 1968, 1984). Few nymphs are observed on hosts after September. Wilkinson (1968) speculated that diapause may prevent nymphs, moulting from larvae late in the year, from feeding prior to overwintering. Similarly, Pound and George (1991) reported that nymphs from prairie strains of D. andersoni entered diapause more readily than montane strains. This behaviour would presumably be of adaptive advantage to minimise exposure to arid climatic conditions and periods of low host availability in the prairies. As with larvae, most nymphs attach to the ears, to the bases of vibrissae, and around the eyes of their hosts (Wilkinson 1968). The feeding period for nymphs averages about seven days (range 4–11 days) (Hewitt 1915; Brown 1944; Loomis 1961; Scott and Brown 1986). Engorged nymphs moult to the adult instar in late summer but most adults remain inactive until the following spring (Wilkinson 1968). Wilkinson (1971) observed that prolonged exposure to low temperatures (i.e., 3 months at 5 °C) is required to break diapause in recently emerged adults. Unfed adults are long-lived and some are capable of surviving through two winters (Gregson 1951; Wilkinson 1968). Adults and nymphs which overwinter do so in rodent burrows, under dead leaves or other litter, and beneath rocks or similar protective places (Mail 1942). Adults of D. andersoni readily bite humans though larvae and nymphs rarely do so (Bishopp and Trembley 1945). The bite of adults does not cause pain, and because these ticks have relatively short mouthparts they are easily removed (Parker et al. 1937; Gregson 1973). The pathogens responsible for tularaemia, Rocky Mountain spotted fever, and Colorado tick fever have been isolated or detected in D. andersoni collected in Canada. 82 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Adults are the primary vectors of these pathogens to humans because immatures rarely feed on people. Antibodies to Powassan have been detected in small mammals (ground squirrels and hares) in serological surveys in British Columbia (McLean et al. 1968, 1970) and Alberta (Hoff et al. 1970; Zarnke and Yuill 1981), but attempts to isolate this virus from D. andersoni collected from some of these animals or from the environment have failed (McLean et al. 1969, 1970). Powassan virus has been isolated from D. andersoni collected in South Dakota (McLean and Quantz 1964; Keirans and Clifford 1983), Colorado (Thomas et al. 1960) and California (Calisher 1994); thus, this agent may be potentially present in D. andersoni in Canada. In the United States of America, the Rocky Mountain wood tick is the biological vector of Anaplasma marginale, the blood parasite responsible for bovine anaplasmosis (Eriks et al. 1993). Dermacentor andersoni has also been shown experimentally to be capable of transmitting a related species, Anaplasma ovis, to sheep (Kocan and Stiller 1992). Tick paralysis caused by the bite of female D. andersoni has been frequently reported in British Columbia since about 1910 (Gregson 1973). The geographic distribution of the disease does not coincide with the range of D. andersoni (Jellison and Gregson 1950). Most cases of paralysis occur in the north-western portion of the range of this tick, namely, interior British Columbia, the northern parts of Montana, Washington, and Idaho, northeastern Oregon, and the mountains of Wyoming and Colorado (Gregson 1973). In Canada, human cases of paralysis have not been reported from Alberta or Saskatchewan.

Distribution. In Canada, populations of D. andersoni occur in British Columbia, Alberta, and Saskatchewan (Gregson 1956). In Saskatchewan, populations occur sporadically in central and western regions of the province. The eastern limit of the range of the tick is approximately 105°W longitude (Wilkinson 1967), where it now overlaps with the expanding westward spread in the distribution of D. variabilis (Dergousoff et al. 2013). In Alberta, populations of D. andersoni are most abundant in the southeastern portions of the province (Brown 1944). Populations occur throughout the southern foothills of the Rocky Mountains to north of Jasper, Alberta (Brown and Kohls 1950) and the northerly record is 53°58’N in west-central Alberta (Wilkinson 1967). Dermacentor andersoni is established throughout central British Columbia and the most northern of the westerly records are from Macalister, British Columbia at about 52°30’N (Bruce 1920; Wilkinson

Map 4. Collection localities for Dermacentor andersoni in Canada. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 83

1967). The western limit of the range of this tick appears to be the coastal mountains as it is rare in coastal regions of British Columbia (Gregson 1956; Wilkinson 1967). Records of this tick from ground squirrels in Manitoba (McLeod 1933) and Ontario (Kennedy and Newman 1986) are almost certainly misidentified specimens. Records in other parts of Canada should be examined carefully, and may be misidentifications or the result of travel. In the United States of America, Rocky Mountain wood tick populations are found in the western regions of South Dakota, North Dakota, and Nebraska (Easton 1983; Yunker et al. 1986; Kietzmann 1987), westward to the Cascade Mountains in Washington, Oregon, and California (Easton et al. 1977; Furman and Loomis 1984). As in British Columbia, D. andersoni is absent (or only occasionally introduced from other areas) from coastal regions of western United States of America (Easton et al. 1977; Furman and Loomis 1984). The southern range limit of D. andersoni is northern New Mexico and Arizona (Cooley 1938; Bishopp and Trembley 1945).

Hosts. The immature instars infest primarily small mammals, especially ground-dwelling sciurids, but including mice and voles. Adults will feed on a wide range of larger wild and domestic mammals such as cattle, horses, sheep, deer, bear, coyote, dogs, cats, and badgers. Adults attach and feed readily upon humans as well. There are occasional records on birds, which are almost certainly incidental. Records from eastern Canada are the result of travel to endemic areas, and emphasise the need for physicians to consider travel history in patients presenting unfamiliar symptoms typical of infection by tick-borne agents.

Dermacentor variabilis (Say) American dog tick (Fig. 24, Map 5) Ixodes variabilis Say, 1821: 77. Dermacentor electus Koch, 1844: 235. Ixodes albipictus (Packard, 1869): Neumann 1911: 101 (as synonym in error). Ixodes quinquestriatus + Ixodes robertsoni (Fitch, 1872): Neumann 1911: 101. Dermacentor americanus (Linnaeus, 1758): Neumann 1911: 101. Dermacentor variabilis (Say): Hooker, Bishopp, and Wood 1912: 190. Dermacentor variabilis (Say): Cooley, 1938: 23 (all instars described).

Adult. Female: Body: Length 2.7–5.2 mm unfed, reaching to 15.0 mm engorged. Base colour brown, pattern colour silvery-grey. Gnathosoma: Porose areas of basis capituli oval or bean-shaped. Cornua shorter than width at base, narrowly to bluntly rounded apically (Fig. 24A). Scutum: Slightly longer than wide, with posterolateral margins slightly concave behind eyes, posterior margin bluntly angular (Fig. 24A). Cervical grooves narrow, pit-like anteriorly, elongate, shallow posteriorly, forming hourglass-shape between them. Surface with large puncta less numerous than small, inconspicuous puncta. Extent of pattern colour highly variable, but typically more extensive than base colour. Spiracular plate: Variably oval, with distinct, moderately broad, dorsal prolongation. Goblet cells minute and numerous (greater than 300), nearly as small as surrounding pore-like perforations in several peripheral rings near frame and on dorsal prolongation (Fig. 24A, insert). Legs: Coxa I with elongate internal and external spurs, their opposing margins parallel; internal spur broader and about as long as external spur (Fig. 24A). Small, rounded internal spur present on coxae II and III, indistinct or absent on coxa IV. Coxae II–IV each with short, bluntly pointed external spur, about as broad at base as long, sometimes progressively smaller from II–IV. 84 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Adult. Male: Body: Length 2.3–4.4 mm. Colour ornamentation variable, much as in female but silvery-grey pattern often less extensive than brown on scutum. Gnathosoma: Cornua slightly more prominent than in female (Fig. 24B). Palpal apex subacute. Scutum: Cervical grooves deep, short, with pit-like posterior depression. Surface with widely scattered large and small puncta, much as in female. Spiracular plate: Shape and goblet size as in female; goblet cells minute and constant in size. Legs: Coxae usually progressively enlarged from I– IV (Fig. 24B). Coxae I–IV with spurs as in female, but no vestige of internal spur on coxa IV.

Nymph: Body: Length 0.9–1.5 mm. Gnathosoma: Basis capituli in dorsal view subtriangular, with lateral margins extending beyond apices of scapulae (Fig. 24C). Ventrally, basis with pair of short, bluntly pointed to rounded spurs posterolaterally. Scutum: Slightly longer than wide, with prominently curved posterior margin (Fig. 24C). Cervical grooves long, narrow. Spiracular plate: Large, subcircular to suboval. A ring of moderately small goblet cells usually surrounds macula, remaining goblet cells numerous, minute (Fig. 24C). Legs: Coxa I with semi-acutely pointed external spur much larger than small but distinct internal spur (Fig. 24C). Coxae II–IV each with small, blunt external spur, sometimes indistinct on IV. Coxae II–IV lacking internal spurs.

Larva: Body: Length 0.58–0.66 mm. Gnathosoma: Basis capituli in dorsal view with lateral margins drawn out posterolaterally to extend beyond apices of scapulae (Fig. 24D). Palpal femur lacking dorsal gland pore. Scutum: Length about five-sixths (0.84) width, widest at level of eyes, slightly behind midlength (Fig. 24D). Legs: Coxa I with a distinct, subacutely triangular internal spur (Fig. 24D). Coxae II and III each with broad, indistinct spur posteromarginally.

Natural history. Throughout North America, dogs (Canis lupus familiaris Linnaeus) are important hosts for adults of D. variabilis (Bishopp and Smith 1938; Cooley 1938; Smith et al. 1946; Koch 1982) and hence this tick is commonly known as the American dog tick. Dermacentor variabilis is a three-host tick. Adult dog ticks typically feed upon medium- sized mammals such as dogs, raccoons (Procyon lotor (Linnaeus)), opossums (Didelphis virginiana (Kerr)), porcupines (Erethizon dorsatum (Linnaeus)), and black bears (Ursus americanus (Pallas)) (Dodds et al. 1969, Garvie et al. 1978; Campbell and MacKay 1979; Pung et al. 1994). They also occasionally infest striped skunks (Mephitis mephitis (Schreber)), eastern chipmunks (Tamias striatus (Linnaeus)), weasels (Mustela species), and ground squirrels (Spermophilus species) (Wright 1979; Gkoroba 1980). Larvae and nymphs of D. variabilis usually infest small mammals such as meadow voles (Microtus pennsylvanicus), red-back voles (Myodes gapperi), deer mice (Peromyscus maniculatus), and white-footed mice (Garvie et al. 1978; Gkoroba 1980; Burachynsky and Galloway 1985). In some localities, meadow jumping mice (Zapus hudsonius (Zimmermann)), eastern chipmunks, and snowshoe hares (Lepus americanus Erxleben), can also serve as hosts to a significant proportion of the larval and nymphal population (Campbell and MacKay 1979; Burachynsky and Galloway 1985). Birds are rarely infested by D. variabilis larvae or nymphs (Battaly et al. 1987; Stafford et al. 1995; Nicholls and Callister 1996). Adult dog ticks readily bite humans (Scholten 1977; Schwartz et al. 1993), though larvae and nymphs rarely do (Felz et al. 1996). Throughout its range in Canada, D. variabilis requires at least two years to complete its life cycle. Unfed adults of D. variabilis overwinter and become active shortly after snow melt (e.g., late March to early May). Males usually appear before females (Sonenshine et al. 1966) and adult host-seeking populations usually peak in late May and June (Garvie A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 85

A B

C D

Figs. 24A–D. Dermacentor variabilis gnathosoma and scutum, dorsal view (left), ventral view (right). Fig. 24A, female; insert, enlarged spiracle; Fig. 24B, male; Fig. 24C, nymph; Fig. 24D, larva. Figs. 24A modified, B–D redrawn from Brinton et al. (1965); insert redrawn from Yunker et al. (1986). 86 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins et al. 1978; Burachynsky and Galloway 1985). Thereafter, adult activity declines and few adults are observed by mid-August, though in Nova Scotia (Garvie et al. 1978), Ontario and Manitoba, small numbers of adults are occasionally observed in September or later. Adults are frequently collected along the boundaries between wooded habitats and fields, along roadsides, trails, fence rows, hedge rows, and similar locations or sites frequented by people and animals (Dodds et al. 1969; Sonenshine and Levy 1972; Newhouse 1983; Carroll et al. 1991). The abundance of adult dog ticks varies among habitats and years (Sonenshine et al. 1965; Sonenshine and Stout 1970; Garvie et al. 1978). Female adult dog ticks require 6–21 days to feed to repletion (Sonenshine 1967; Amin 1969a; Gladney and Dawkins 1971). Mating is required before females engorge fully (Sonenshine 1967; Pappas and Oliver 1971, 1972; Obenchain et al. 1980), though some females of D. variabilis can produce eggs without mating (Oliver 1971, 1974). Male D. variabilis must feed prior to mating and therefore mating occurs on the host (Feldman-Muhsam and Borut 1971; Sonenshine 1991). Fed females can disperse as far as 120 cm prior to establishing an oviposition site (Smith et al. 1946). Temperature influences the duration of the pre-oviposition and oviposition period in D. variabilis (Campbell and Harris 1979), thus females which feed during May and June begin to lay eggs in June and July (Garvie et al. 1978; McEnroe 1982). Females typically produce 4000–6000 eggs (Amin 1969b; Sonenshine and Tigner 1969; Drummond et al. 1971; Koch and Sauer 1984) and occasionally as many as 7000 eggs (Campbell and Harris 1979; Nagar 1968). Larvae emerge from eggs during June to September (Campbell 1979) and usually remain inactive, overwinter, and seek hosts the next year (Sonenshine et al. 1965; Conlon and Rockett 1982; Koch 1989). Occasionally, recently emerged larvae infest hosts in late August (Garvie et al. 1978; Smart and Caccamise 1988; Micher and Rockett 1993) or September (Sonenshine et al. 1965, 1966; Zimmerman et al. 1987), although this fall peak in larvae may not occur in all years (Garvie et al. 1978) or at all geographic localities (Jackson and DeFoliart 1975; Burachynsky and Galloway 1985). Overwintered larvae begin to seek hosts from March to May (Garvie et al. 1978; Burachynsky and Galloway 1985). Larvae rarely actively disperse from egg masses (Conlon and Rockett 1982; McEnroe and Sphect 1987), though they may be carried large distances (i.e., > 300 m) on small mammal hosts (Sonenshine 1973). The duration of the feeding period varies among hosts but typically lasts from 3–9 days (Sonenshine and Atwood 1967). As with nymphs, the peak drop-off of larvae occurs in the daylight hours (i.e., 12–18 hours) and correlates with the period of maximum activity of voles (Amin 1970). The abundance and prevalence of larvae on small mammal hosts usually peaks during late April to early June and declines through July and August (Carey et al. 1980; Burachynsky and Galloway 1985). Larvae fed on hosts during April to May moult to nymphs during June and July (Garvie et al. 1978; Micher and Rockett 1993). In Canada, because of insufficient heat units in late summer, most larvae which feed after August moult in the following year (Campbell 1979). Nymphs typically begin to seek hosts during May (Garvie et al. 1978; Gkoroba 1980) or June (Burachynsky and Galloway 1985). The abundance and prevalence of nymphs on small mammal hosts usually peak during June and July and usually decline through July (Garvie et al. 1978; Burachynsky and Galloway 1985). Most unfed nymphs that do not find a host during June and July will die by September (Campbell 1979). Unfed nymphs rarely overwinter, and survival of individuals that do so is less than 5% (Sonenshine 1979a). Similarly, nymphs rarely infest hosts after mid-August, so few overwinter as fed individuals (Campbell 1979). The feeding period for nymphs averages about five days (range 3–11 days) (Sonenshine and Atwood 1967; Brown 1977). Nymphs fed on hosts A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 87

from April to July generally moult to adults during July to September (Garvie et al. 1978; Campbell 1979). Most of these adults will remain inactive until the following spring; however, adults are sometimes observed on hosts in September or later. If cool climatic conditions or low host availability occur, the life cycle can be extended to three years (Garvie et al. 1978). Yunik et al. (2015b) found that up to 20% of unfed adults were able to survive a second winter in Manitoba. Adult dog ticks readily bite humans (Anderson and Magnarelli 1980; Falco and Fish 1988; Slaff and Newton 1993) and bites typically do not cause pain. The pathogens responsible for Lyme disease, tularaemia, and Rocky Mountain spotted fever in humans have been isolated or detected in D. variabilis collected in Canada (Artsob et al. 1984; Lindsay et al. 1991; Artsob 1996); however, the probability of acquiring a tick-borne pathogen from this species is very low. Despite numerous reports in the United States of America (Anonymous 1996), paralysis of humans (or animals) caused by D. variabilis has never been reported in Canada (Gregson 1973). Dermacentor variabilis has been incriminated as the vector for the protozoan, Cytauxzoon felis, which causes a fatal disease in domestic cats in parts of the southeastern United States of America (Blouin et al. 1984; Hoover et al. 1994). Stiller and Coan (1995) reported that American dog ticks can transmit the equine piroplasm, Babesia caballi, to horses and that this pathogen is successfully transmitted transovarially in D. variabilis. American dog ticks also likely have a limited role in epizootics of tularaemia (Jellison 1974) and in transmission of Ehrlichia risticii, the causative agent of Potomac horse fever (Holland et al. 1985; Carroll and Schmidtman 1986; Schmidtman 1989), though the latter has not been detected in wild or domestic animals in Canada.

Distribution. The American dog tick is widespread in eastern North America, with disjunct populations in California, Idaho, Oregon, Washington, and Mexico. In Canada, there are records from Newfoundland, Prince Edward Island, and New Brunswick; it is often very abundant in parts of Nova Scotia, and is locally abundant in southern Québec and Ontario. It is widely distributed and often abundant in southern Manitoba into northwestern Ontario,

Map 5. Collection localities for Dermacentor variabilis in Canada. 88 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins as well as southeastern Saskatchewan. The American dog tick has expanded its range in Manitoba and Saskatchewan, and is established considerably further north and west than 50 years ago (Dergousoff et al. 2013).

Hosts. Larvae and nymphs feed on a great variety of small mammals, especially rodents, and are occasionally found on cats and dogs. Juveniles are rarely recorded from humans. Adults infest many species of medium-sized to large animals, including dogs, badgers, bear, deer, cattle, horses, bison, skunks, opossum, and humans. Records from British Columbia and the Yukon are almost certainly the results of travel to endemic areas.

Genus Haemaphysalis Koch (Figs. 25–26) Anus of nymphs and adults with postanal groove. Eyes absent. Posteromarginal festoons present (Fig. 25D). Palpus with 2nd palpomere (femur) not appreciably longer than wide, but acutely produced laterally, extending beyond margins of basis capituli (Fig. 26A). Basis capituli in dorsal view subrectangular, roughly parallel-sided. Adult females and males with inornate scutum. Foveae present. Adult males without ventral plates. Coxa I with 1 long or short spur. Coxae I–IV similar in size. Larvae with 1 pair of posthypostomal setae (Fig. 25J); scutum with 3 pairs of setae (Figs. 25K, 26J); 8 pairs of marginal dorsal setae, including 2 pairs anterior to sensilla sagittiformia; 2 pairs of central dorsal setae (Fig. 25K).

Key to Species of the Genus Haemaphysalis in Canada

1. All instars: palpi with flared lateral margins more acutely angled and reflexed than in H. chordeilis (Figs. 26A, 26C–D, 26H); adult, female and male: hypostomal dentition 3/3 (Fig. 26B); basis capituli venter without spurs; adult male: coxa IV with spur short, rounded; nymph and larva: basis capituli in dorsal view subquadrangular, with lateral margins roughly parallel, not drawn out to points (Figs. 26D, 26J) ...... Haemaphysalis leporispalustris – All instars: palpi with flared lateral margins less acutely angled and reflexed than in H. leporispalustris (Figs. 25A, 25D, 25G, 25K); adult, female and male: hypostomal dentition 5/5 (Fig. 25C); basis capituli venter with pair of small spurs; adult male: coxa IV with spur long, pointed; nymph and larva: basis capituli in dorsal view subhexagonal, with lateral margins angled, drawn out to points midlength (Figs. 25F, 25K) ...... Haemaphysalis chordeilis

Haemaphysalis chordeilis (Packard) Bird tick or grouse tick (Fig. 25, Map 6) Ixodes chordeilis Packard, 1869: 67. Haemaphysalis punctata cinnabarina Koch: Neumann 1905: 237. Haemaphyhsalis chordeilis (Packard): Banks 1908: 34. Haemaphysalis punctata cinnabarina Koch: Neumann 1911: 108. Haemaphysalis cinnabarina Koch: Nuttall and Warburton 1915: 372 (in part). Haemaphysalis chordeilis (Packard): Cooley 1946c: 37 (all instars described).

Adult. Female: Body: Length 2.8–2.9 mm unfed, reaching to 9.5 mm engorged. Gnathosoma: Porose areas of basis capituli large, oval, depressed, separated by interval A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 89

slightly greater than width of one porose area (Fig. 25A). Dorsal posterior margin of basis capituli nearly straight or slightly sinuous, with rounded corners or small, rounded cornua (Fig. 25A). Ventrally, posterior region of basis without pair of spurs. Hypostome slightly clavate, with rounded corona (Fig. 25C). Hypostome dentition 5/5, with denticles progressively smaller from lateral to middle files, and with 12–14 teeth in each file. Palpi with flared lateral margins less reflexed and acutely angled than inH . leporispalustris. Scutum: Length slightly greater than width, oval, widest somewhat anterior to middle (Fig. 25A). Scapulae large, bluntly pointed. Cervical grooves conspicuous. Surface covered with numerous, conspicuous, evenly distributed puncta. Legs: Coxa I with internal spur moderately long, subacute, and lacking external spur (Fig. 25B). Coxae II–IV each with single, bluntly pointed spur, placed near middle of posterior side on II and III, but slightly longer and placed near base of posterior side on IV. Trochanter I with a conspicuous, pointed dorsal spur; ventral trochanter spurs absent on all legs. Tarsi I–IV each with small terminal ventral spur.

Adult. Male: Body: Total length, tips of palpi to posterior margin, 2.2–2.8 mm. Gnathosoma: Basis capituli in dorsal view with lateral margins nearly parallel, and with posterior margin slightly concave, with small but distinct, blunt cornua (Fig. 25D). Ventrally, posterior region of basis without pair of spurs. Hypostome form as in female; dentition 5/5, with 8–10 teeth in each file (Fig. 25E). Palpi formed as in female, with flared lateral margins less reflexed and acutely angled than inH . leporispalustris. Scutum: Surface covered with numerous, conspicuous, evenly distributed puncta (Fig. 25D). Marginal grooves well formed anteriorly and continuing posteriorly to delimit anteriormost 2 or 3 festoons. Legs: Coxae I–IV with number and form of spurs as in female except spur on coxa IV longer and more acutely pointed (Fig. 25G).

Nymph. Gnathosoma: Basis capituli in dorsal view hexagonal, with angled lateral margins; posterior margin slightly concave, its posterolateral corners not developed into cornua (Fig. 25F). Ventrally, posterior region of basis without pair of spurs. Hypostome slightly clavate; dentition 2/2, with 7 or 8 teeth in each file (Fig. 25H). Palpi as in adults, with flared lateral margins less reflexed and acutely angled than in H. leporispalustris (Fig. 25I). Scutum: Shape nearly circular, widest slightly anterior to midlevel (Fig. 25F). Cervical grooves distinct, moderately deep, long. Surface not conspicuously punctate. Legs: Coxae I–IV with number and form of spurs similar to those of female (Fig. 25I). Trochanter I with a distinct dorsal spur; ventral trochanter spurs absent on all legs, as in adults.

Larva. Body: Length (excluding gnathosoma) 0.47–0.52 mm. Sternal setae short (approximately 0.2 mm), anterior pair not reaching a third of distance to second pair (Fig. 25J). Eleven festoons. Gnathosoma: Basis capituli in dorsal view hexagonal, with angled lateral margins; posterior margin slightly concave, posterolateral corners not developed into cornua (Fig. 25K). Ventrally, basis with 2 large, spur-like auriculae. Hypostome slightly clavate; dentition 2/2, with 6 or 7 teeth in each file (Fig. 25J). Palpi with flared lateral margins less acutely angled than in H. leporispalustris. Scutum: Length approximately 0.7 times of width, widest slightly anterior to midlength (Fig. 25K). Cervical grooves long, nearly parallel, nearly reaching posterolateral margins. Legs: Coxa I with broadly rounded internal spur; coxae II and III each with weakly developed, broadly rounded spur (Fig. 25J).

Natural history. There are scattered records for this species across Canada (Bequaert 1945; Levine and Goble 1947; Gregson 1956), but there is little published information 90 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A B

E G

F H

J sensilla sagittiforme K

Figs. 25A–J. Haemaphysalis chordeilis. Fig. 25A, female gnathosoma and scutum, dorsal; insert. lateral view of palp; Fig. 25B, female coxae and trochanters, ventral; Fig. 25C, female hypostome, ventral; Fig. 25D, male gnathosoma and scutum, dorsal; Fig. 25E, male hypostome, ventral; Fig. 25F, nymph gnathosoma and scutum, dorsal; Fig. 25G, male gnathosoma, coxae and trochanters, ventral; insert, enlarged palpal seta; Fig. 25H, nymph hypostome, ventral; Fig. 25I, nymph gnathosoma, coxae and trochanters, ventral; insert, enlarged palpal seta; Fig. 25J, larva, ventral; Fig. 25K, larva, dorsal. Figs. 25A–I, redrawn from Cooley (1946); Figs. 25J and K redrawn from Clifford et al. (1961). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 91

Map 6. Collection localities for Haemaphysalis chordeilis in Canada.

on its natural history. Dick (1981) has provided the most detailed information. In Manitoba, adults were most abundant in the spring, first appearing on sharp-tailed grouse (Tympanuchus phasianellus (Linnaeus)) in April, and reaching their maximum numbers in May. Larvae reached their greatest numbers in July, and nymphs were most abundant in early August. Although the larvae and nymphs exhibited clear peaks in their activity, they were present in smaller numbers throughout the season, from April to September. Dick (1981) speculated that the dancing grounds for male grouse were important foci for maintenance of H. chordeilis populations. Dick (1981) reported on the severe reaction to tick bites in sharp-tailed grouse. He found the area of attachment was always swollen, and the bite site often apparent long after the tick had detached. Furman and Loomis (1984) reported on the implication of this species in deaths among turkeys (Meleagris gallopavo Linnaeus) in the United States of America. Gregson (1956) suggested that it is a vector for wildlife diseases, but he provided no specific references to support this contention.

Distribution. This tick is widely distributed in North America from California to Texas, to the southeastern United States of America and across western and south central Canada, east into southeastern Ontario.

Hosts. As its common name implies, H. chordeilis primarily infests birds. Gallinaceous birds, especially grouse, are probably the primary hosts, but there are numerous records from mammals, including cattle, horses, marmots, and occasionally humans.

Haemaphysalis leporispalustris (Packard) Rabbit tick (Fig. 26, Map 7) Ixodes leporis-palustris Packard, 1869 : 67 (not Ixodes chordeilis Packard, 1869 as stated by Neumann 1897: 343.) Rhipistoma leporis Osborn, 1896: 261. Gonixodes rostralis Dugès, 1888: 129 (see also Nuttall and Warburton 1915: 387). Haemaphysalis leporis (Packard): Neumann 1897: 343. Haemaphysalis leporis-palustris (Packard): Hunter and Hooker 1907: 53. Haemaphysalis leporis proxima Aragão, 1911: 167. Synonym. Haemaphysalis leporis-palustris (Packard): Cooley 1946c: 31 (all instars described). 92 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Adult. Female: Body: Length 1.7–2.6 mm unfed, reaching to 10.0 mm engorged. Gnathosoma: Porose areas of basis capituli oval, depressed, widely separated by interval of about twice the width of one porose area (Fig. 26A). Dorsal posterior margin of basis capituli nearly straight, with rounded corners or rounded to bluntly pointed cornua. Ventrally, posterior region of basis with pair of small spurs (Fig. 26C). Hypostome slightly clavate, with rounded corona (Fig. 26B). Hypostome dentition 3/3, with denticles subequal in size, and with about 8 teeth in each file. Palpal femur with posterolateral edge more reflexed than inH . chordeilis (Fig. 26C). Scutum: Length slightly greater than width, oval, widest anterior to middle (Fig. 26A). Scapulae large, bluntly pointed. Cervical grooves conspicuous. Surface sparsely covered with numerous, evenly distributed puncta. Legs: Coxa I with internal spur broad, bluntly pointed, larger than small, bluntly pointed external spur (Fig. 26C). Coxae II–IV each with single, short, bluntly pointed spur placed near middle of posterior side, progressively smaller from II–IV. Trochanter I with a conspicuous, acute dorsal spur and a small ventral spur; trochanters II–IV also each with small ventral spur. Tarsi I–IV lacking a terminal ventral spur.

Adult. Male: Body: Total length, tip of palpi to posterior margin, from 1.4–2.1 mm. Gnathosoma: Basis capituli in dorsal view with lateral margins slightly convergent posteriorly, and with posterior margin nearly straight, with well-formed, bluntly-pointed cornua (Fig. 26H). Ventrally, posterior region of basis with pair of small spurs (Fig. 26I). Hypostome form as in female; dentition 3/3, with approximately 8 teeth in each file (Fig. 26F). Palpi formed as in female, with flared lateral margins more reflexed and acutely angled than in H. chordeilis (Fig. 26I). Scutum: Surface covered with numerous, evenly distributed puncta (Fig. 26H). Marginal grooves incomplete anteriorly, continuing posteriorly to delimit only anteriormost 1 festoon. Legs: Coxae I–IV with number and form of spurs as in female (Fig. 26I).

Nymph. Gnathosoma: Basis capituli in dorsal view subquadrangular, with lateral margins slightly convergent posteriorly; posterior margin slightly concave, its posterolateral corners developed into distinct, triangular cornua (Fig. 26D). Ventrally, posterior region of basis with pair of small spurs as in adults (Fig. 26E). Hypostome slightly clavate; dentition 2/2, with 7 or 8 teeth in each file (Fig. 26G). Palpi much as in adults, with flared lateral margins more reflexed and acutely angled than inH . chordeilis (Fig. 26E). Scutum: Nearly circular, widest near midlevel (Fig. 26D). Surface not conspicuously punctate. Legs: Coxae I–IV with number and form of spurs similar to those of female (Fig. 26E). Trochanter I with a dorsal and a ventral spur as in adults.

Larva. Body: Length (excluding gnathosoma) 0.48–0.55 mm. Sternal setae moderately short (approximately 0.3 mm), anterior pair reaching nearly half distance to second pair (Fig. 26K). Eleven festoons. Gnathosoma: Basis capituli in dorsal view subquadrangular, with lateral margins slightly convergent posteriorly; posterior margin slightly concave, with posterolateral corners developed into distinct, triangular cornua directed posteriorly (Fig. 26J). Ventrally, basis with 2 large, spur-like auriculae (Fig. 26K). Hypostome dentition 2/2, with approximately 7 teeth in each file. Palpi with flared lateral margins more acutely angled than in H. chordeilis (Fig. 26J). Scutum: Length approximately 0.7 times width, widest slightly anterior to midlength (Fig. 26J). Cervical grooves distinct, slightly divergent posteriorly, not reaching posterolateral margins. Legs: Coxa I with broadly rounded internal spur; coxae II and III each with conspicuous, bluntly rounded spur (Fig. 26K). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 93

Natural history. This species is a common parasite of rabbits in Canada, where prevalence of infestation can exceed 75% and an animal may be infested with more than 100 ticks at certain times of the year (Campbell et al. 1980). In Minnesota, Green et al. (1943) reported average infestations of more than 3000 ticks on snowshoe hare (Lepus americanus) in late summer, with as many as 16 000 ticks found on one animal. There are numerous published records for H. leporispalustris in Canada (Hewitt 1915; Hearle 1938; Bequaert 1945; Brown and Kohls 1950; Judd 1953; Scholten et al. 1962; Wilson 1967; Dodds et al. 1969; Martell et al. 1969; Ko 1972a; Wright 1979; Jones and Thomas 1980; Gkoroba 1980) with few specifics on the life cycle and natural history. The most detailed studies on its natural history were carried out in Nova Scotia by Campbell and Glines (1979) in the laboratory and by Green et al. (1943) and Campbell et al. (1980) in the field in Minnesota and Nova Scotia, respectively. Adult ticks overwintered and were present on snowshoe hares early in the spring, at their greatest abundance in May and June, and with their numbers gradually falling off until August. Adults mate on the host, females having the capacity to mate with more than one male (Oliver et al. 1974). Engorged ticks generally leave their host during the day, when the rabbit is resting in its nest or form, so eggs are most often laid in close proximity to a location with a high probability of encountering a second rabbit host (George 1971). At 14.7–35.6 ºC, engorged females took an average of 17.7 ± 3.7 days to begin laying eggs, which were laid over 18.1 ± 7.7 days, respectively. The number of eggs laid was dependent on the temperature in that study, but females laid 21 (at 14.7 °C) to 3327 (19.9 °C) eggs. Eggs hatched in 20–119 days at 35.6–14.7 °C, respectively. Larvae may infest rabbits in the form, or if no host is available in the form, they will readily climb surrounding vegetation and may quest at a height in the vegetation appropriate to the size of their rabbit host (Camin and Drenner 1978). There were two discernible peaks in abundance of larvae, the largest occurring from the end of May to early June, and a second smaller peak at the end of August to early September. In Minnesota, the late season peak of larvae was always the largest. Larvae moulted to nymphs in 14.8–58.1 days at 35.6–14.7 °C, respectively. There was one peak in numbers of nymphs in July and August. Nymphs took 18.2–95.9 days to moult to adults at 35.6–14.7 °C, respectively. In Nova Scotia, Campbell et al. (1980) concluded that H. leporispalustris requires two years to complete one generation. This is different from elsewhere in its range where all stages may be present throughout the year (e.g., Kollars and Oliver 2003). Even in Minnesota, very small numbers of ticks were found on snowshoe hares in the winter months. Haemaphysalis

Map 7. Collection localities for Haemaphysalis leporispalustris in Canada. 94 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

D G H

J K

Figs. 26A–L. Haemaphysalis leporispalustris. Fig. 26A, female gnathosoma and scutum, dorsal; Fig. 26B, female hypostome, ventral; Fig. 26C, female gnathosoma, coxae and trochanters, ventral; insert, enlarged palpal seta; Fig. 26D, nymph, gnathosoma and scutum, dorsal; Fig. 26E, nymph gnathosoma, coxae and trochanters, ventral; Fig. 26F, male hypostome, ventral; Fig. 26G, nymph hypostome, ventral; Fig. 26H, male gnathosoma and scutum, dorsal; Fig. 26I, nymph gnathosoma, coxae and trochanters, ventral; insert, enlarged palpal seta; Fig. 26J, larva, dorsal; Fig. 26K, larva, ventral. Figs. 26A–I redrawn from Cooley (1946); Figs. 26J, K redrawn from Clifford et al. (1961). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 95

leporispalustris is the only species of tick in Canada that has been reported as a host for the parasitoid, Ixodiphagus texanus Howard (Hymenoptera: Encrytidae) (Davis and Campbell 1979; see also Green et al. 1943). The role of Haemaphysalis leporispalustris as a vector for pathogenic microorganisms in Canada is not well understood. McKiel et al. (1967) isolated two strains of a new species of rickettsia, Rickettsia canadensis McKiel et al. (Rickettsiales: Rickettsiaceae), from H. leporispalustris near Ottawa, Ontario. Because juveniles of this tick are so often associated with ground-foraging birds (Scott et al. 2001; Scharf 2004) and therefore might be dispersed over great distances, there is concern about its potential to disseminate potential pathogens. This is of particular concern because of its association with birds carrying Ixodes scapularis and I. pacificus, important vectors of Borrelia burgdorferi. (Durden et al. 2001; Slowik and Lane 2001a; Lane et al. 2006; Scott et al. 2012). The role of H. leporispalustris in the epidemiology of borreliosis in North America is unknown, despite the potential for its infection and for transovarial transmission (Lane and Burgdorfer1988). Goethert and Telford (2003b) ruled out the likelihood that H. leporispalustris was a competent vector for the agent for human granulocytic anaplasmosis. An analysis of geographic variation in structure in larvae, males and females of H. leporispalustris was conducted by Thomas (1968).

Distribution. The rabbit tick is perhaps the most widely distributed species of tick in North America, being found from Alaska south to Mexico, and east across the continent. It occurs in every province in Canada. It has also been recorded in Central and South America, into Argentina.

Hosts. There is no doubt that rabbits and hares are the most important hosts of this widespread tick, and heavy infestations are frequently reported. However, it has been recorded from a great variety of hosts, including ground-foraging birds, which are of undoubted importance in its dispersal. It occurs rarely on large mammals and humans.

Genus Ixodes Latreille (Figs. 27–63) Anus of nymphs and adults with anterior groove (Fig. 13A). Eyes absent. Posteromarginal festoons absent. Palpus variable in form, but not produced laterally. Basis capituli of adults in dorsal view subrectangular, roughly parallel-sided. Adult females and males with inornate scutum. Foveae absent. Adult males with 7 non-salient ventral plates (Fig. 42E). Coxa I with or without spurs. Coxae I–IV similar in size. Larvae with usually 2 pairs of posthypostomal setae (Figs. 35L, 35M); scutum with 5 (rarely 4) pairs of setae; 6–10 pairs of marginal dorsal setae; sensilla sagittiformia absent; 2–6 pairs of central dorsal setae (Fig. 35K).

Key to species of Ixodes in Canada – females

1. Anterior margin of basis capituli conspicuously humped on either side of hypostome (Fig. 59B); scutum punctate and with rugose or wrinkly areas anterolaterally (Fig. 59A); primarily on raccoons, mustelids, and squirrels; western to mid-eastern North America…………...... Ixodes texanus – Anterior margin of basis capituli sloped (Fig. 30B), or at most, slightly humped on either side of hypostome; scutum punctate but usually lacking rugose or wrinkly areas anterolaterally (Fig. 32A) (I. baergi and I. rugosus excepted)...... 2 96 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

2. External spur vestigial or absent on all coxae (Fig. 27D); coxa I with or without internal spur...... 3 – External spur present, small to prominent, on all coxae (Fig. 34C); coxa I usually with internal spur (Fig. 36H)...... 6

3. Coxa I with short internal spur and vestigial external spur (Fig. 54E); dorsal posterior margin of basis capituli with small but distinct cornua (Fig. 54A); auriculae small, usually lobe-like, rarely vestigial...... 4 – Coxa I without spurs (Fig. 27D); dorsal posterior margin of basis capituli without cornua (Fig. 44A); auriculae vestigial or absent...... 5

4. Scutum distinctly (1.2–1.3 times) longer than wide (Fig. 54A); hypostome with pointed apex, dentition 3/3 along distal one-third, then 2/2 to base (Fig. 54B); primarily on squirrels; eastern North America...... Ixodes marxi – Scutum nearly as wide as long (Fig. 48A); hypostome with rounded apex, dentition 2/2 along nearly entire length (Fig. 48B); primarily on squirrels; western North America ...... Ixodes hearlei

5. Scutum coarsely punctate, not wrinkled, with rounded scapulae (Fig. 27A); palp with tibiotarsus terminal and visible dorsally (Fig. 27A); hypostomal dentition 2/2 along nearly entire length; body very setose (Fig. 27A); on sea birds; along Atlantic and Pacific coasts...... Ixodes uriae – Scutum rugose, wrinkled laterally, without coarse puncta, with bluntly pointed scapulae (Fig. 44A); palp with tibiotarsus subterminal and visible only ventrally (Fig. 44C); hypostomal dentition 4/4 subapically, then 3/3 along middle third, then 2/2 to base (Fig. 44B); body not noticeably setose; on cliff swallows; mid-eastern and midwestern North America...... Ixodes baergi

6. Coxae I with internal spur about equal in length to or shorter than external spur (Fig. 63D), or vestigial...... 7 – Coxae I with internal spur considerably longer and more slender than external spur, often extending well over coxae II in unengorged specimens (Fig. 30C)...... 14

7. Coxae I with internal spur small or vestigial, less produced than external spur (Fig. 63D); basis capituli with porose areas large, nearly contiguous, occupying most of dorsal face (Fig. 61A)...... 8 – Coxae I with internal spur short but similarly as well produced as external spur (Fig. 42C); basis capituli with porose areas variable in size but clearly separated medially and not occupying most of dorsal face (Fig. 39A)...... 9

8. Scutum oval, usually distinctly (approximately 1.2–1.4 times) longer than wide (Fig. 63A); hypostome sessile, not arising from a median extension of basis capituli (Fig. 63B); basis capituli with auriculae shelf-like lateral extensions continuous with ridge onto dorsolateral surface (Fig. 63D); primarily on sea birds; along Pacific coast, Aleutians, across Bering Strait to Japan...... Ixodes signatus – Scutum rhomboid to subtriangular, about as long or slightly (to approximately 1.1 times) longer than wide (Fig. 61A); hypostome arising from a short smooth median extension of basis capituli (Fig. 61C); basis capituli with auriculae blunt lateral A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 97

protuberances continuous with ridge onto dorsolateral surface (Fig. 61B); primarily on terrestrial birds; western North America...... Ixodes howelli

9. Auriculae horn-like or “angular” (Fig. 28B); trochanters I–III each with a spur (Fig. 28F) ...... 10 – Auriculae ridge-like or absent (Fig. 41E); trochanters I–III without spurs (Fig. 41E) ...... 11

10. Palptrochanter with prominent, anterior spur-like projection (Figs. 42A, C); auriculae horn-like, retrograde (Fig. 42C); coxae II and usually III each with 2 spurs (Fig. 42C); on birds; western North America, primarily along Pacific coast...... Ixodes auritulus – Palptrochanter without spur-like projection (Fig. 28A); auriculae angular or blunted lateral projections (Fig. 28B); coxae II and III each with 1 spur (internal spur absent) (Fig. 28F); on birds; eastern North America to southwestern United States of America. �� Ixodes brunneus

11. Scutum with longitudinal wrinkles or furrows on posterior face (Fig. 41A); ventral faces of coxae I–IV with prominently barbed setae; exceptionally small tick (body length, unfed, approximately 1.2–1.5 mm); on shrews; western North America ...... Ixodes soricis – Scutum without longitudinal wrinkles on posterior face (Fig. 39A); ventral faces of coxae I–IV with smooth or imperceptibly barbed setae; medium-sized ticks (body length, unfed, approximately 2–3 mm)...... 12

12. Palpi club-like, thick, about 2.0–2.5 times as long as wide (Fig. 60A); scutum subtriangular, about as wide as long; coxae I–IV each with small external spur, coxa I with small internal spur (Fig. 60E); apex of hypostome rounded (Fig. 60C); on mustelids; boreal North America...... Ixodes gregsoni – Palpi linear, slender, over 3–4 times as long as wide; scutum oval, 1.3–1.4 longer than wide (Figs. 38A–B); coxae I–IV each with prominent external spur, coxa I with well- developed internal spur (Fig. 38J); apex of hypostome pointed (Fig. 38C)……...... 13

13. Hypostomal dentition 3/3 along distal one-third to one-half, then 2/2 to base, denticles bluntly pointed, not flared (Fig. 39B); primarily on pikas and wood rats; western North America...... Ixodes ochotonae – Hypostomal dentition 3/3 along nearly entire length, denticles sharp, flared (Fig. 38C); primarily on rodents; western to eastern North America...... Ixodes angustus

14. Auriculae present as horn-like or spur-like structures (Fig. 32C)...... 15 – Auriculae present as mild ridges or absent (Fig. 58E)...... 17

15. Scutum about 1.1 times longer than wide, with numerous coarse puncta posteriorly (Fig. 30A); hypostomal dentition 6/6 to 5/5 apically, then 4/4 along mid-length (Fig. 30B); on rabbits; southeastern Canada and eastern United States of America ...... Ixodes dentatus – Scutum about 1.3 times longer than wide, with puncta more evenly sized and distributed (Fig. 37A); hypostomal dentition at most 4/4 apically, then at most 3/3 along mid-length (Fig. 37B)...... 16 98 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

16. Auriculae straight, blunt, spur-like (Fig. 32C); palptrochanter with weak ventrolateral spur (Fig. 32C); porose areas separated by interval less than width of one porose area (32A); on small rodents, occasionally small birds; eastern North America west to Manitoba……...... Ixodes muris – Auriculae curved, pointed, hook-like (Fig. 37D); palptrochanter with prominent blunt ventrolateral spur; porose areas separated by interval greater than width of one porose area (Fig. 37A); on various rodents, shrews; southwestern Canada and northwestern United States of America...... Ixodes spinipalpis

17. Palpi slender, about 3–4 times as long as wide (Fig. 36A)...... 18 – Palpi thick, club-like in dorsal view, about 2.0–2.5 times as long as wide (Fig. 50A) ……...... 20

18. Scutum with lateral margins angular (Fig. 58A); basis capituli with dorsal posterior margin usually markedly sinuous, with prominent cornua directed posteriorly (Fig. 58A); hypostomal dentition 3/3 on apical one-third, then 2/2 to base (Fig. 58B); primarily on ground squirrels and gophers; central and western North America...... Ixodes sculptus – Scutum oval to nearly circular (Fig. 36A); basis capituli with dorsal posterior margin usually concave, with weakly developed cornua (Fig. 36A); hypostomal dentition 4/4 then 3/3 on apical one-third, then 2/2 to base (Fig. 36B)...... 19

19. Dorsal posterior margin of basis capituli with small but definite cornua (Fig. 36A); auriculae formed as small, diagonal bulges laterally (Fig. 36D); scutum nearly circular, widest at mid-level, with puncta larger marginally and posteriorly (Fig. 36A); on large and small mammals, lizards, birds; eastern North America west to Manitoba ...... Ixodes scapularis – Dorsal posterior margin of basis capituli with blunt corners instead of cornua (Fig. 34A); auriculae formed as short, diagonal ridges laterally (Fig. 34C); scutum oval, widest slightly anterior to mid-level, with uniformly distributed small puncta (Fig. 34A); on large and small mammals, lizards, birds; western North America ...... Ixodes pacificus

20. Cornua prominent as rounded, posterior projections (Fig. 50A); palptrochanter with a distinct, narrowly rounded flange projecting ventrolaterally (Fig. 52D)...... 21 – Cornua small, indistinct, or absent (Fig. 56A); palptrochanter with at most a low, broadly- curved flange-like edge or small ridge-like projection ventrolaterally (Fig. 46C)…...... 22

21. Hypostomal dentition 2/2 along nearly entire length, most denticles blunt (Fig. 50B); scutum with lateral carinae distinct, but cervical grooves weak or absent (Fig. 50A); on wide variety of small to mid-sized mammals; widespread in North America ...... Ixodes kingi – Hypostomal dentition 4/4 then 3/3 along apical one-third, then 2/2 to base, most denticles sharp (Fig. 52B); scutum with lateral carinae faint or absent, but cervical grooves usually distinct (Fig. 50A); primarily on marmots, ground squirrels; western North America...... Ixodes marmotae

22. Scutum with large puncta and wrinkles on anterolateral area (Fig. 56A); basis capituli with dorsal posterior margin concave (Fig. 56A); on mustelids and canids; along west A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 99

coastal North America...... Ixodes rugosus – Scutum with puncta but lacking wrinkles on anterolateral area (Fig. 45A); basis capituli with dorsal posterior margin straight or convex (Fig. 45A)...... 23

23. Basis capituli with dorsal posterior margin straight, with short, blunted cornua (Fig. 46A); porose areas separated by interval about equal to width of one porose area (Fig. 46A); apex of hypostome rounded, densely denticulate (Fig. 46C); palptrochanter with a small, broadly rounded flange-like projection ventrolaterally (Fig. 46B); on wide variety of small to mid-sized mammals, often carnivores; eastern to midwestern North America...... Ixodes cookei – Basis capituli with dorsal posterior margin slightly sinuous or convex, with somewhat flared corners instead of cornua (Fig. 45B); porose areas separated by interval less than width of one porose area (Fig. 45B); apex of hypostome flattened, sparsely denticulate (Fig. 45E); palptrochanter with a slight ventral projection (Fig. 45C); on skunks, muskrats, beaver, canids; eastern to midwestern North America...... Ixodes banksi

Key to species of Ixodes in Canada – males (male of I. gregsoni unknown)

1. Hypostome formed as a plate or shaft nearly devoid of residual denticles (Fig. 27C); palp with apex of femorogenu asymmetrical, either bluntly pointed (Fig. 27G) or broadly flattened (Fig. 63I); coxa I with internal spur weakly formed or absent (Fig. 27G)….... 2 – Hypostome formed as a shaft partly to fully covered with denticles or crenulae (Fig. 44D); palp with apex of femorogenu symmetrical, evenly rounded (Fig. 42E); coxa I with internal spur small to well developed, often conspicuous (Fig. 36H) (I. baergi excepted).....4

2. Caudal margins of ventral body plates lobe-like, with terminal fringe of long setae (Fig. 27G); hypostome deeply indented apically (Fig. 27C); coxae I–IV without spurs (Fig. 27G)...... Ixodes uriae – Caudal margins of ventral body plates not lobed, without a setal fringe (Fig. 63H) or with a marginal fringe of short, thick setae (Fig. 61E, insert); hypostome flattened (Fig. 63J), rounded or bluntly pointed apically (Fig. 61G); coxae I–IV each with a blunt external spur (Fig. 61E), and coxa I with or without (Fig. 63H) a small internal spur ...... 3

3. Hypostome truncate apically (Fig. 63J, insert); apex of palp femorogenu broadly, asymmetrically flattened, with tibiotarsus arising terminally (Fig. 63I); coxa I without internal spur (Fig. 63H); caudal margins of ventral plates without a distinctive fringe of seta (Fig. 63H)...... Ixodes signatus – Hypostome bluntly pointed apically (Fig. 61G); apex of palp femorogenu asymmetrically, bluntly pointed, with tibiotarsus arising ventroterminally (Fig. 61F); coxa I with small internal spur (Fig. 61F); caudal margins of ventral plates with a distinctive fringe of short, thick setae (Fig. 61E, insert)...... Ixodes howelli

4. Scutum smooth, with a pair of mid-lateral pit-like depressions (Fig. 42D); hypostome shaft with apical third tapered, with dentition 3/3 there, increasing to 6/6 or 7/7 at midlength, then gradually decreasing basally (Fig. 42F); coxa I with similarly short, blunt internal and external spurs (Fig. 42E)...... Ixodes auritulus 100 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

– Scutum usually discernibly punctate, sometimes rugose, but without a pair of lateral pit-like depressions (Fig. 48F); hypostome shaft parallel-sided, with denticles or crenulae not more numerous along mid-length (Fig. 52C); coxa I with or without distinct internal and external spurs...... 5

5. Coxa I with at most 2 weakly-developed spurs, internal spur small, bluntly rounded or faint or absent, external spur a low ridge or absent (Fig. 44F); coxae II–IV with external spur small, faint or absent (Fig. 44F); hypostome dentition with 5–7 transverse rows each with 2/2 or 3/3 mild crenulae, these indistinct medially (Fig. 44D)...... 6 – Coxa I with 2 well-developed spurs, internal spur bluntly to sharply pointed, sometimes longer than external spur (Fig. 36H); coxae II–IV each with distinct external spur (Fig. 36H); hypostome dentition variable, often with more strongly formed denticles or crenulae (Fig. 36E)...... 9

6. Coxae I–IV without definite spurs (Fig. 44F); palptrochanter without flange-like projection (Fig. 44F); scutum moderately densely and lightly punctate over most of surface (Fig. 44E); median ventral plate about 1.3 times longer than anal plate, both plates with similarly faint, fine puncta (Fig. 44F)...... Ixodes baergi – Coxa I with weak or faint external and internal spur or saliences, coxae II–IV with faint external spur or salience (Fig. 54G); palptrochanger with ventral ridge or small flange (Fig. 54G); scutum with conspicuous large puncta laterally (Fig. 54F)and sometimes medially (Fig. 59F); median ventral plate faintly punctate, at most slightly longer than conspicuously more coarsely punctate anal plate (Fig. 48G)...... 7

7. Scutum conspicuously more coarsely and deeply punctate laterally than medially (Fig. 54F); hypostome with rounded apex and approximately 7 transverse rows of mostly 2/2 crenulae (Fig. 54C)...... Ixodes marxi – Scutum coarsely punctate over most of surface (Fig. 59F); hypostome with notched apex and approximately 6 transverse rows of mostly 3/3 crenulae (Fig. 59C)...... 8

8. Anterior margin of basis capituli conspicuously humped on either side of hypostome (Fig. 59F); coxa I with internal spur a rounded salience similar in form to that representing external spur (Fig. 59C)...... Ixodes texanus – Anterior margin of basis capituli slightly humped on either side of hypostome (Fig. 48F); coxa I with short blunt internal spur, similar in form to external spur (Fig. 48G)………...... Ixodes hearlei

9. Hypostome with 4 or 5 conspicuous, pointed lateral denticles abruptly differing in shape and larger size from median diagonal or transverse rows of denticles or crenulae (Fig. 37C); coxa I with internal spur long, sharply pointed (Fig. 36H)...... 10 – Hypostome with lateral denticles not abruptly differing in shape and size from median denticles or crenulae (Fig. 58C); coxa I with internal spur variable in form...... 12

10. Posterior margin of basis capituli with distinct, bluntly pointed cornua directed posteriorly (Fig. 37G); spiracular plate subcircular (Fig. 37H)...... Ixodes spinipalpis – Posterior margin of basis capituli with obtuse corners instead of cornua (Fig. 36G); spiracular plate oval or elliptical, its anteroposterior axis clearly longer than its dorsoventral axis (Figs. 34E–G)...... 11 A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 101

11. Spiracular plate elongate, more broadly rounded anteriorly than posteriorly (Fig. 36C); median ventral plate with large deep puncta (Fig. 36H)...... Ixodes scapularis – Spiracular plate oval, similarly rounded anteriorly and posteriorly (Fig. 34G); median ventral plate with small shallow puncta (Fig. 34E)...... Ixodes pacificus

12. Trochanters of legs I–IV each with a spur (as in Fig. 28F)...... Ixodes brunneus – Trochanters of legs I–IV without spurs (Fig. 30E) (vestiges may be evident in Ixodes cookei)...... 13

13. Hypostome with 10–12 diagonal rows of rounded to nearly crenulated denticles, and with dentition 6/6 to 5/5 along most of length (Fig. 30F); median ventral plate deeply punctate (Fig. 30E)...... Ixodes dentatus – Hypostome with 4–8 transverse or diagonal rows of denticles or crenulae, and with dentition 4/4 or 3/3 to 2/2 along most of length (Fig. 32F); median ventral plate with inconspicuous puncta (Fig. 45A) (Ixodes muris excepted, Fig. 32E)...... 14

14. Spiracular plate elongate, with anteroposterior axis clearly longer than dorsoventral axis (Fig. 45A)...... 15 – Spiracular plate broadly oval to circular, with anteroposterior and dorsoventral axes similar in length (Fig. 50E)...... 16

15. Hypostome deeply indented apically, dentition with inner files formed as crenulae (Fig. 32F); median ventral plate conspicuously punctate, about twice as long as anal plate (Fig. 32E)...... Ixodes muris – Hypostome rounded or slightly notched apically, dentition with inner files formed as distinct, bluntly rounded teeth (Fig. 45G); median ventral plate inconspicuously punctate, about 1.4 times as long as anal plate (Fig. 45A)...... Ixodes banksi

16. Dorsal posterior margin of basis capituli with distinct cornua directed posterolaterally (Fig. 50D); scutum with definite lateral carinae (Fig. 50D)...... Ixodes kingi – Dorsal posterior margin of basis capituli with cornua indistinct or absent (Fig. 56F); scutum with lateral carinae faint or absent (Fig. 58G)...... 17

17. Coxa I with internal spur clearly longer than external spur, sharply pointed, well- overlapping partial width of coxa II (Fig. 58H); hypostome with inner files formed as definite blunt teeth, not fully crenulated (Fig. 58C)...... 18 – Coxa I with internal spur moderately long, similar in form or more tapered than external spur, blunt, not overlapping partial width of coxa II (Fig. 41H); hypostome with inner files formed as crenulae (Fig. 41C)...... 20

18. Hypostome dentition 3/3 along most of length, inner files rounded, short, not overlapping (Fig. 58C); coxa I with internal spur long, slightly overlapping width of coxa II (Fig. 58H)...... Ixodes sculptus – Hypostome dentition 4/4 along most of length, inner files bluntly pointed, long, overlapping (Fig. 46G); coxa I with internal spur longer, overlapping nearly half width of coxa II (Fig. 46E)...... 19

19. Ventral body plates with puncta faint, similar in appearance on median, anal and adanal 102 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

plates (Fig. 46E); scutum more coarsely and deeply punctate laterally than medially (Fig. 46D)...... Ixodes cookei – Ventral body plates with puncta faint on median plate but conspicuous and larger on anal and adanal plates (Fig. 56G); scutum with conspicuous deep puncta over most of surface (Fig. 56F)...... Ixodes rugosus

20. Palps bulbous, short (approximately 1.6–1.7 times as long as wide) (Fig. 41G); hypostome with 3 or 4 transverse rows of coarsely overlapping crenulae confined to apical third of shaft (Fig. 41C); ventral faces of coxae I–IV with coarsely branched setae (Fig. 41H); small ticks (body length about 1.0 mm)...... Ixodes soricis – Palps of moderate length (approximately twice as long as wide) (Fig. 39D); hypostome with 5–7 transverse rows of crenulae along at least apical half of shaft (Fig. 39F); ventral faces of coxae I–IV with smooth or slightly barbed setae (Fig. 38H); moderately-sized ticks (body length 1.5–2.5 mm)...... 21

21. Hypostome with rounded apex and 5 or 6 irregularly transverse rows of weakly formed crenulae (Fig. 39F); scutum with indistinct, small puncta over most of surface (Fig. 39D); median ventral plate slightly wider than long (Fig. 39E)...... Ixodes ochotonae – Hypostome with flattened or indented apex and 5–7 regularly transverse rows of distinct crenulae (Fig. 52C); scutum with puncta distinct, slightly to conspicuously coarser along lateral margins (Fig. 52E); median ventral plate usually at least slightly longer than wide (Fig. 52F)...... 22

22. Median ventral plate at most slightly longer than anal plate (Fig. 52F); hypostome distinctly indented apically, and with moderately developed, non-overlapping, transverse rows of crenulae (Fig. 52C); scutum moderately densely and lightly punctate over most of surface, puncta slightly larger laterally (Fig. 52E)...... Ixodes marmotae – Median ventral plate clearly (about 1.5) longer than anal plate (Fig. 38H); hypostome flattened apically, and with strongly developed, overlapping transverse rows of crenulae (Fig. 38G); scutum conspicuously more coarsely and deeply punctate laterally than medially (Fig. 38F)...... Ixodes angustus

Key to species of Ixodes in Canada – nymphs

1. External spurs vestigial or absent on all coxae (Fig. 59E); coxa I with (Fig. 55I) or without internal spur...... 2 – External spur present on coxa I, and present or absent on coxae II–IV (Fig. 36J); coxa I usually with internal spur (Fig. 35I)...... 6

2. Scutum widest near its anterior margin and clearly (approximately 1.2 times) longer than wide (Fig. 27E); palp with tibiotarsus terminal and visible dorsally (Fig. 27E); basis capituli with only 1 pair of post-hypostomal setae...... Ixodes uriae – Scutum widest at anterior one-third of its length or near mid-length, and about as wide as long (Fig. 59H); palp with tibiotarsus subterminal and visible only ventrally (Fig. 59E); basis capituli with 2 pairs of post-hypostomal setae...... 3

3. Anterior margin of basis capituli conspicuously humped on either side of hypostome (best seen in ventral view) (Fig. 59E); scutum rugose or wrinkled anterolaterally (Fig. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 103

59H)...... Ixodes texanus – Anterior margin of basis capituli sloped, or at most, slightly humped on either side of hypostome (Fig. 55I); scutum lacking rugose or wrinkly areas (Fig. 54H)...... 4

4. Basis capituli with dorsal posterior margin concave (Fig. 44G); auriculae faint, ridge- like; hypostomal dentition 3/3 on apical third, then 2/2 to base...... Ixodes baergi – Basis capituli with dorsal posterior margin nearly straight (Fig. 48H); auriculae extending as rounded or slightly angled lobes laterally (Fig. 55I); hypostomal dentition 2/2 along nearly entire length (Fig. 54D)...... 5

5. Dorsal posterior margin of basis capituli with distinct cornua (Fig. 54H); auriculae projecting as lateral lobes posterior to mid-level of basis capituli (Fig. 55I); palptrochanter with small ventral projection (Fig. 55I)...... Ixodes marxi – Dorsal posterior margin of basis capituli with cornua indistinct or absent (Fig. 48H); auriculae projecting as lateral lobes anterior to mid-level of basis capituli, almost flanking bases of palpi (Fig. 49I); palptrochanter with short but conspicuous, blunt, ventrolateral flange (Fig. 49I)...... Ixodes hearlei

6. Basis capituli with auriculae strongly produced as rounded or angular or horn-like or shelf-like projections (Fig. 28C)...... 7 – Basis capituli with auriculae formed as moderately conspicuous lateral bulges or lobes, or lateral ridges, or vestigial or absent (Figs. 31G, 45F)...... 16

7. Basis capituli with auriculae produced as shelf-like lateral extensions continuous with ridge onto dorsolateral surface (Fig. 63E); coxa I with internal spur vestigial or absent, external spur distinct (Fig. 63F)...... Ixodes signatus – Basis capituli with auriculae produced as rounded or angular or horn-like projections (Fig. 28C); coxa I with internal spur slightly smaller than, or longer than, external spur (Fig. 29K) ...... 8

8. Basis capituli with auriculae produced as rounded or angular projections directed laterally or posterolaterally (Fig. 62I)...... 9 – Basis capituli with auriculae produced as triangular or horn-like projections directed posteriorly (Fig. 36I) ...... 14

9. Basis capituli with auriculae produced as 2 variably triangular projections on each side (Fig. 28C); hypostome with pointed apex, and with 3 files of denticles along apical half of shaft (Fig. 28B)...... Ixodes brunneus – Basis capituli with auriculae produced as 1 projection on each side (Fig. 53I); hypostome with rounded apex, and with third file of teeth, if present, confined to apical third of shaft (Fig. 51H)...... 10

10. Palpi club-like in dorsal view, moderately short, at most about 2.5 times as long as wide (Fig. 52G); scutum broadly subtriangular or subrhomboid (Fig. 52G)...... 11 – Palpi linear, long, about 3.5 times as long as wide (Fig. 35H); scutum oval to nearly circular (Fig. 35H)...... 13

11. Palptrochanter lacking flanges (Fig. 62I); coxa I with internal spur blunt, smaller than 104 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

external spur (Fig. 62I)...... Ixodes howelli – Palptrochanter with 1 or 2 flanges (Fig. 51H); coxa I with internal spur tapered, longer than external spur (Fig. 51H)...... 12

12. Palptrochanter with a small ridge anteriorly, and a bluntly pointed flange posterolaterally (Fig. 51H)...... Ixodes kingi – Palptrochanter with 2 long, narrowly rounded flanges of similar size, 1 anteriorly, 1 posterolaterally (Fig. 53I)...... Ixodes marmotae

13. Scutum longer (approximately 1.2) than wide (Fig. 37E); basis capituli with auriculae produced as bluntly triangular lateral lobes (Fig. 37F)...... Ixodes spinipalpis – Scutum about, or slightly wider than long (Fig. 35H); basis capituli with auriculae variably produced as shelf-like to bluntly angular lateral lobes (Fig. 35I)....Ixodes pacificus

14. Palptrochanter with an anteriorly-projecting flange (Fig. 43I); coxae I–IV with salient external spurs, that on coxa I larger than internal spur; trochanters I–IV each with a spur (Fig. 43I)...... Ixodes auritulus – Palptrochanter without an anteriorly-projecting flange (Fig. 33I); coxae I–IV with small external spurs, that on coxa I no larger than internal spur (Fig. 33I); trochanters I–IV without spurs (Fig. 33I)...... 15

15. Basis capituli with dorsal posterior margin sinuous, convex medially, with prominent, pointed cornua (Fig. 36I); basis capituli with auriculae formed as small, pointed, posterior projections (Fig. 36J)...... Ixodes scapularis – Basis capituli with dorsal posterior margin straight, with small, blunt cornua (Fig. 33G); basis capituli with auriculae formed as conspicuous, blunt, posterolateral projections (Fig. 33I)...... Ixodes muris

16. Palptrochanter lacking flange-like projections, at most with a small ventral tubercle (Fig. 30C)...... 17 – Palptrochanter with 1 or 2 flange-like projections (Fig. 41F)...... 18

17. Palpi moderately long (about 3.5 times as long as wide), not club-like in dorsal view (Fig. 31G); hypostome dentition mostly 4/4 along apical half, changing to 3/3 then sometimes to 2/2 near base (Fig. 31H); scutum nearly circular (Fig. 31G)...... Ixodes dentatus – Palpi short (about 2.2 times as long as wide), club-like in dorsal view (Fig. 45F); hypostome dentition 3/3 or 2/2 along distal one-third to one-half, then 2/2 to base (Fig. 45J); scutum broadly subrhomboid (Fig. 45I)...... Ixodes banksi

18. Palptrochanter with both anterior and posterior flange-like projections well developed (Fig. 58F)...... 19 – Palptrochanter with only 1 flange-like projection well developed (usually the posterior projection) (Fig. 47K)...... 21

19. Coxa I with internal spur longer and more tapered than external spur (Fig. 58F); basis capituli dorsally with posterior margin sinuous, concave medially, with prominent cornua (Fig. 58I)...... Ixodes sculptus – Coxa I with internal spur blunt and similar in size to external spur (Fig. 38E); basis A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 105

capituli dorsally with posterior margin nearly straight, with moderately developed cornua (Fig. 38D)...... 20

20. Palpi short, thick (little more than twice as long as wide), club-like in dorsal view (Fig. 41I); scutum oval, clearly (1.3–1.4) longer than wide (Fig. 41I); ventral faces of coxae I–IV with prominently barbed setae (Fig. 41F); small ticks (body length, unfed, at most 1 mm)...... Ixodes soricis – Palpi moderately long (at least 3 times as long as wide), not club-like in dorsal view (Fig. 38D); scutum rhomboid, slightly wider than long (Fig. 38D); ventral faces of coxae I–IV with smooth or inconspicuously barbed setae (Fig. 38E); medium-sized ticks (body length, unfed, more than 1.5 mm)...... Ixodes angustus

21. Coxa I with internal spur clearly longer and more tapered than external spur (Fig. 47K) ...... Ixodes cookei – Coxa I with internal spur blunt and similar in size to external spur (Fig. 40I)...... 22

22. Palptrochanter with posterior flange well developed, tapered (Fig. 40I); dorsal posterior margin of basis capituli with prominent cornua (Fig. 40G)...... Ixodes ochotonae – Palptrochanter with posterior flange moderately developed, rounded or angular (Fig. 60E); dorsal posterior margin of basis capituli with cornua poorly defined or absent (Fig. 56E)...... 23

23. Scutum widest near mid-length (Fig. 56E); coxa I with internal spur more narrowly rounded than external spur (Fig. 56H)...... Ixodes rugosus – Scutum widest at anterior one-third of length (Fig. 60I); coxa I with internal spur bluntly rounded like external spur (Fig. 60M)...... Ixodes gregsoni

Key to species of Ixodes in Canada– larvae

Note: Tick larvae should be examined as slide mounts of cleared whole specimens under a compound microscope that is preferably equipped with an ocular micrometer for observing relative lengths and making measurements of structures. Clearing and mounting procedures are presented in the section on collecting, preserving, and studying ticks. Intraspecific variation occurs among some of the setal and structural characteristics used in keys and diagnoses of larvae. As a result, there will be occasional specimens that are impossible to run through any key. Therefore, this key works most reliably when a series of specimens, rather than one, is available for examination. Because of their small size and variability, the problems associated with making adequate slide mounts of them for study, and the considerable number of species in Canada, the larvae of Ixodes ticks may be difficult to identify with certainty. Authoritative identifications should be made by specialists in matters concerning unusual host or distribution records and specimens as potential vectors of pathogenic agents removed from humans or domestic animals.

1. Palptrochanter lacking flange-like projections (Fig. 62J)...... 2 – Palptrochanter with 1 or 2 small to salient flange-like projections (Fig. 51I)...... 12

2. Coxa I–III without spurs (Fig. 27J)...... 3 – Coxa I with 1 or 2 spurs; coxae II–III with or without 1 or 2 spurs (I. banksi with only 1 weakly-defined spur, internally on coxa I) (Fig. 62L)...... 4 106 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

3. Scutum widest near its anterior margin and clearly (approximately 1.2–1.3 times) longer than wide (Fig. 27I); scutum with 4 pairs of setae (Fig. 27I); basis capituli with 1 pair of post-hypostomal setae (Fig. 27J)...... Ixodes uriae – Scutum widest at about anterior one-third its length, and about as wide or slightly wider than long (Fig. 44J); scutum with 5 pairs of setae (Fig. 44J); basis capituli with 2 pairs of post-hypostomal setae (Fig. 44M)...... Ixodes baergi

4. Soft body cuticle lacking supplementary (dorsal mediolateral) setae, and with 4–6 pairs of marginal ventral (ventral lateral) setae (Fig. 45I); femur III with 8 setae ...... Ixodes banksi – Soft body cuticle with 1 or 2 pairs of supplementary (dorsal mediolateral) setae, and with 3 or 4 pairs of marginal ventral (Fig. 62L); femur III with 9 or 10 setae...... 5

5. Scutum with 4 pairs of setae (Fig. 62K); palpi thick, femur+genu length to width ratio at most 2:1 (Fig. 62L)...... 6 – Scutum with 5 pairs of setae (Fig. 37K); palpi slender, femur+genu length to width ratio near or exceeding 3:1 (Fig. 37L)...... 7

6. Soft body cuticle with 8 or 9 pairs of ventral setae in the premarginal (4), preanal (3), and postanal (1 or 2) regions (Fig. 62L); posteriormost pair of marginal ventral and premarginal ventral setae similar in size to their dorsal counterparts; basis capituli with auriculae well developed, angular, projecting posterolaterally (Fig. 62L); ventral apex of palp femorogenu without protuberances (Fig. 62J)…………………..Ixodes howelli – Soft body cuticle with 7 pairs of ventral setae in the premarginal (5), preanal (2), and postanal (0) regions (Fig. 63L); posteriormost pair of marginal ventral and premarginal ventral setae approximately twice as long as their dorsal counterparts; basis capituli with auriculae indistinct (Fig. 63M); ventral apex of palp femorogenu with a salient flange (Fig. 63L)...... Ixodes signatus

7. Soft body cuticle with dorsal setae relatively long, length of anterior marginal dorsal setae about 4 times that of scutal setae (Fig. 29L); ventral soft body cuticle with 3 pairs of preanal setae (Fig. 29M); palptrochanter weakly formed, nearly indiscernible ventrally (Fig. 29N)...... Ixodes brunneus – Soft body cuticle with dorsal setae and ventral setae mostly similar in length, length of anterior marginal dorsal setae at most twice that of scutal setae (Fig. 37K); ventral soft body cuticle with 2 pairs of preanal setae (Fig. 37L); palptrochanter well developed, clearly discernible ventrally (Fig. 37J)...... 8

8. Soft body cuticle with 2 pairs of central dorsal setae (Fig. 37K)...... Ixodes spinipalpis – Soft body cuticle with 3 or 4 pairs of central dorsal setae (Fig. 33J)...... 9

9. Soft body cuticle with 3 pairs of central dorsal setae (Fig. 31J)...... 10 – Soft body cuticle with usually 4 pairs of central dorsal setae (Fig. 35K)...... 11

10. Basis capituli with auriculae angular, projecting laterally or posterolaterally (Fig. 36K); ventral apex of palp femorogenu without protuberances (Fig. 36L); coxae I–III each with small, blunt external spur (Fig. 36L)...... Ixodes scapularis – Basis capituli with auriculae undeveloped or weakly formed as ridges (Fig. 31J); A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 107

ventral apex of palp femorogenu with small tubercle (Fig. 31K); coxae I–III lacking external spurs (Fig. 31K)...... Ixodes dentatus

11. Auriculae formed as angular lateral projections extending beyond lateral walls of basis capituli (Fig. 33K); soft body cuticle with moderately short dorsal setae, anterior pairs of marginal dorsal setae scarcely (at most 1.5 times) longer than scutal setae (Fig. 33J)...... Ixodes muris – Auriculae formed as blunt posterolateral projections usually not extending beyond lateral walls of basis capituli (Fig. 35L); soft body cuticle with moderately long dorsal setae, anterior pairs of marginal dorsal setae 3–4 times longer than scutal setae (Fig. 35K)...... Ixodes pacificus

12. Palptrochanter with both anterior and posterior flange-like projections developed (Fig. 38N) (anterior projection small, bluntly pointed in Ixodes sculptus, I. rugosus, I. gregsoni, posterior projection small, rounded in I. rugosus and I. gregsoni)...... 13 – Palptrochanter with only 1 flange-like projection developed (usually the posterior projection, except in Ixodes auritulus) (Fig. 47M)...... 18

13. Dorsal posterior margin of basis capituli conspicuously sinuous, concave medially, with cornua distinct, directed posterolaterally (Fig. 58K); palpal femorogenu compact, its length to width ratio at most 1.7:1, and its ventral apex with 1 or 2 small tubercles (Fig. 58L)...... Ixodes sculptus – Dorsal posterior margin of basis capituli nearly straight or slightly sinuous, with cornua indistinct or small, usually directed laterally (Fig. 57J); palpal femorogenu less compact, its length to width ratio from 1.8–2.5:1, and its ventral apex lacking tubercles (Fig. 57L)...... 14

14. Palptrochanter with small flange-like projections, anterior projection bluntly pointed, posterior projection rounded (Fig. 57L)...... 15 – Palptrochanter with salient flange-like projections, anterior projection pointed or narrowly rounded, posterior projection narrowly rounded (Fig. 41L)...... 16

15. Coxa I with small, rounded internal spur (Fig. 60O); palptrochanter with bluntly pointed anterior flange slightly more prominent than rounded posterior flange (Fig. 60K); femur III with usually 9 setae...... Ixodes gregsoni – Coxa I with broadly, bluntly angular internal spur (Fig. 57K); palptrochanter with bluntly pointed anterior flange slightly less prominent than rounded posterior flange (Fig. 57L); femur III with usually 8 setae...... Ixodes rugosus

16. Coxae II–III each with 1 or 2 setae conspicuously unilaterally barbed or setulose, having several, thin barbs (Fig. 41L, insert); soft body cuticle with usually 4 pairs of marginal ventral setae (Fig. 41L)...... Ixodes soricis – Coxae II-III each with all setae similarly smooth or with inconspicuous barbs; soft body cuticle with usually 3 pairs of marginal ventral setae (variably 2–5 pairs in western North America) (Fig. 40L)...... 17

17. Scutum widest at anterior third (Fig. 40K); palptrochanter with anterior flange-like projection bluntly to sharply pointed, often notably shorter than narrowly rounded 108 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

posterior projection (Fig. 40J)...... Ixodes ochotonae – Scutum widest at mid-level (Fig. 38M); palptrochanter with bluntly pointed anterior and narrowly rounded posterior flange-like projections similarly prominent (Fig. 38K)...... Ixodes angustus

18. Soft body cuticle with 2 or 3 pairs of supplementary dorsal setae, and 3–6 pairs of central dorsal setae (Fig. 43J); palptrochanter with prominent anterior flange-like projection (Fig. 43L); trochanters I and II each with small spur or tubercle (Fig. 43K) ...... Ixodes auritulus – Soft body cuticle without supplementary dorsal setae, and with 2 or rarely 3 pairs of central dorsal setae (Fig. 51J); palptrochanter without anterior projection, but with small to prominent posterior flange-like projection (Fig. 59I); trochanters I and II without a spur or tubercle (Fig. 51K)...... 19

19. Soft body cuticle with usually 9 pairs (sometimes 8 pairs in I. texanus and I. hearlei) of marginal dorsal setae (Fig. 59K); scutum finely micropunctate and clearly reticulate- imbricate over entire surface; dorsal posterior margin of basis capituli with narrowly rounded corners instead of cornua (Fig. 55J); femur III with usually 9 setae...... 20 – Soft body cuticle with usually 8 pairs of dorsal lateral setae (Fig. 47L); scutum densely micropunctate over entire surface and lineate-reticulate along anterolateral margins; dorsal posterior margin of basis capituli with cornua divergent, directed laterally (Fig. 51J); femur III with usually 8 setae...... 22

20. Scutum widest at mid-level (Fig. 55J); basis capituli with auriculae angular, projecting laterally (Fig. 55K)...... Ixodes marxi – Scutum widest at anterior third (Fig. 47L); basis capituli with auriculae undeveloped or faintly ridge-like (Fig. 47M)...... 21

21. Posterior margin of scutum slightly rounded, convex (Fig. 59K); dorsal posterior margin of basis capituli nearly straight or slightly sinuous, with rounded corners (Fig. 59K); coxa III lacking external bulge (Fig. 59L)...... Ixodes texanus – Posterior margin of scutum flattened or slightly concave (Fig. 49J); dorsal posterior margin of basis capituli sinuous, with blunt, truncated corners; coxa III with external bulge or slight spur (Fig. 49K)...... Ixodes hearlei

22. Soft body cuticle with 3 pairs of marginal ventral setae (Fig. 53L); sternal setae with anterior pair about twice as long as third pair (Fig. 53L); coxae I–III each with rounded bulge or ridge apicoventrally (Fig. 53L)...... Ixodes marmotae – Soft body cuticle with usually 4 pairs of marginal ventral setae; sternal setae similar to one another in length (Fig. 47M); coxae I–III lacking apical (external) prominences (Fig. 51K)...... 23

23. Palptrochanter with a small, bluntly pointed, anterior ridge (not a flange), and with moderately developed posterior flange usually wider than long (Fig. 47M); eastern North America…………...... Ixodes cookei – Palptrochanter without an anterior ridge, and with moderately developed posterior flange narrowly rounded, about as long as wide (Fig. 51K); western North America ……...... Ixodes kingi A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 109

Ixodes (Ceratixodes) uriae White (Fig. 27, Map 8) Ixodes uriae White, 1852: appendix p. ccx. Hyalomma puta Pickard-Cambridge, 1876: 261. Ixodes borealis Kramer and Neumann, 1883: 527. Ixodes hirsutus Birula, 1895: 356. Ixodes putus: Neumann 1899: 125 (male, nymph). Ceratixodes putus: Neumann 1902: 117. Ixodes uriae White: Nuttall 1912: 60 (larva). Ceratixodes uriae: Schulze 1938: 722. Ixodes (Ceratixodes) putus: Anastos 1950: 124; Pomerantzev 1950: 87. Ixodes (Ceratixodes) putus procellariae Schulze, 1930: 123.

Adult. Female: Body: A large tick, body length 3.8–4.3 mm unfed, reaching to nearly 12.0 mm when fully engorged. Numerous long setae present on dorsum and venter (Fig. 27A). Gnathosoma: Porose areas of basis capituli well defined, large, broadly oval, with their longer axes transverse (Fig. 27A). Dorsal posterior margin of basis capituli straight, posterolateral corners rounded, lacking cornua. Auriculae absent (Fig. 27D). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded; dentition 2/2 along nearly entire length, all denticles strong, almost equal and bluntly pointed (Fig. 27B). Palpi widest distally, clavate in dorsal view. Palpal tibiotarsus often prominent from above. Palpal trochanter lacking flanges, visible dorsally as a broad ring. Scutum: About 1.3 times longer than wide; widest near anterior margin, with posterolateral margins nearly straight, or slightly convex (Fig. 27A). Scapulae rounded. Lateral carinae absent. Cervical grooves well defined, diverging, and reaching posterolateral margins. Surface with puncta numerous, unequal, fairly evenly distributed, occasionally confluent laterally. Legs: Coxae I–IV and trochanters I–IV completely lacking spurs (Fig. 27D). Ventral coxal surfaces with scattered short, smooth setae.

Adult. Male: Body: Length 3.2–3.4 mm. Gnathosoma: Basis capituli dorsal posterior margin straight or weakly concave, its rounded corners not produced into cornua; dorsal face without furrows (Fig. 27F). Ventrally, anterior margin of basis capituli not humped on either side of hypostome (Fig. 27G). Apex of hypostome broad, deeply indented; dentition with approximately 3 transverse rows of faint crenulations confined to either side of apical third of shaft (Fig. 27C). Palpi moderately short, convergent but not club-like in dorsal view, with asymmetrically bluntly pointed apices. Dorsally, palpal femur fused with genu. Palpal tibiotarsus arising from ventral face of palp (Fig. 27G, insert), not visible dorsally (Fig 27F). Palpal trochanter large, easily visible dorsally, lacking flanges. Scutum: Scapulae prominent, bluntly pointed, lateral carinae faint (Fig. 27F). Surface somewhat rugose, densely punctate. Surface lacking pair of pit-like depressions midlaterally. Spiracular plate: Subcircular, without distinct longer axis. Body venter: Caudal margins of ventral plates lobe-like, with continuous fringe of long setae (Fig. 27G). Median plate about as long as anal plate. Anal plate straight- sided, not widened posteriorly, where similarly as wide as adanal plates. Ventral plates with distinct puncta of varying sizes. Legs: Legs long, slender. Coxae I–IV and trochanters I–IV completely lacking spurs (Fig. 27G). Ventral coxal surfaces with scattered, short, smooth setae.

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli straight or slightly concave, its posterolateral corners lacking cornua (Fig. 27E). Auriculae absent. Anterior 110 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins margin of basis capituli not humped on either side of hypostome. Apex of hypostome rounded. Hypostome dentition 2/2 along nearly entire length, lateral and inner denticles similar in size, bluntly pointed. One pair of post-hypostomal setae. Palpi not conspicuously club-like in dorsal view (Fig. 27E). Palpal tibiotarsus visible dorsally. Palpal trochanter lacking flanges, visible dorsally as a broad ring. Scutum: Length about 1.2 times width, shape triangular, widest near anterior margin, with posterolateral margins nearly straight, or slightly concave (Fig. 27E). Scapulae rounded. Lateral carinae absent. Cervical grooves well defined, slightly divergent, and reaching posterolateral margins. Surface with numerous fine puncta amidst sparsely scattered large puncta.Legs: Coxae I–IV and trochanters I–IV completely lacking spurs. Ventral coxal surfaces with scattered short, smooth setae.

Larva. Body: Body length 0.7–0.8 mm unengorged, to 2.2 mm engorged. Gnathosoma: Dorsal posterior margin of basis capituli straight, with corners not developed into cornua (Fig. 27I). Ventrally, auriculae absent (Fig. 27J). Apex of hypostome broadly rounded, dentition below coronal area, 2/2 along entire length; lateral file with 6–8 bluntly pointed, overlapping teeth; inner file with 4–6 blunter, smaller teeth (Fig. 27H). One pair of post– hypostomal setae (Fig. 27H). Palpi moderately thick; femur+genu length to width ratio about 1.8–2.0:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, lacking flanges. Scutum: Elongate, length about 1.3 width, widest near anterior margin. Scapulae rounded apically; shield posterior margin nearly or irregularly flat, or slightly convex (Fig. 27I). Surface densely micropunctate, and reticulated anterolaterally and anteriorly. Setae, 4 pairs, lacking posterolateral pair. Idiosomal setation (Figs. 27I–J): Central dorsal setae, 3–5 pairs. Supplemental dorsal setae absent. Marginal dorsal setae, usually 7 pairs. Marginal ventral setae, 3 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length anterior marginal dorsal setae about twice that of scutal setae. Posteriormost pair of marginal ventral and premarginal ventral setae notably elongate, nearly four times as long as any posterior dorsal setae. Sternal setae similar to one another in short length. Legs: Coxae I–III lacking internal and external spurs. Coxae I–II–III setal formula 2–3–2 (coxa I lacking its anterior seta); coxal setae short, smooth. Femur III with 10 setae. Trochanters I–III lacking spurs (Fig. 27J).

Natural history. Ixodes uriae is one of the most remarkable of our hard ticks, by virtue of its nearly world-wide, coastal distribution. Its ability to feed upon a wide variety of sea birds, and its tolerance of a wide range of environmental conditions have allowed it to become established in sea bird colonies on some of the most remote oceanic islands (e.g., Heath 1977; Bergström et al. 1999a, 1999b; Hänel and Heyne 2008; Heath et al. 2011) and the Antarctic Peninsula (Benoit et al. 2007). Ixodes uriae has been frequently reported in Canada (Bequaert 1945; Gregson 1956; Spencer 1960; Main et al. 1973; Eveleigh and Threlfall 1974, 1975; Main et al. 1976b; Fitzpatrick and Threlfall 1977; Ballard and Ring 1979; Morbey 1996; McCoy et al. 2001, 2003a, 2003b; Muzaffar and Jones 2004, 2007). In recent years, there has been important work done on the genetics of different I. uriae populations, including populations in Canada, which have ramifications on its ecology (McCoy and Tirard 2000; McCoy et al. 2003; de Meeûs et al. 2007). Its natural history and host relationships have been studied in many parts of the world, including Canada. The life cycle of Ixodes uriae was described by Eveleigh and Threlfall (1974) associated with sea birds on Green Island and Gull Island, Newfoundland and Labrador. They found larvae during most sample periods, but they were most abundant in the nest materials in burrows during June and early July. Engorged larvae, which required an average of about six A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 111

B E

F G J

Figs. 27A–J. Ixodes uriae. Fig. 27A, female gnathosoma and idiosoma, dorsal; Fig. 27B, female hypostome, ventral; Fig. 27C, male hypostome, ventral; Fig. 27D, female gnathosoma, coxae and trochanters, ventral; Fig. 27E, nymph, gnathosoma and scutum, dorsal; Fig. 27F, male gnathosoma and idiosoma, dorsal; Fig. 27G, male gnathosoma, idiosoma and coxae, ventral; insert, enlarged palp, lateral; Fig. 27H, larva hypostome, palp trochanter, basis capituli, ventral; Fig. 27I, larva, dorsal; Fig. 27J, larva, ventral. Figs. 27A–G redrawn from Cooley and Kohls (1945); Figs. 27H–I redrawn, 27J modified, from Webb et al. (1990). 112 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins days to feed, were collected in greatest abundance at the beginning of July, but small numbers were present until August. Moulting occurs about 10 weeks after dropping from the host. Nymphs were most abundant during June and July, but engorged nymphs, which required an average of 5.6 days to feed, were collected until mid-August. Nymphs moult after at least six weeks, and may require more than 20 weeks under certain conditions. Males and females were most abundant during June and July but were still present in August. Males do not feed, and females required an average of just over seven days to engorge. Females may mate with several males, most often before engorgement, and offspring are of mixed paternity (McCoy and Tirard 2002). Females laid up to 555 eggs, oviposition initiated after 2–8 weeks, depending on temperature. Eggs may hatch in as little as about 60 days (Murray and Vestjens 1967), but may remain viable for more than a year under certain conditions (Eveleigh and Threlfall 1974). Although the entire life cycle can be completed in 407 days under laboratory conditions, Flint and Kostyrko (1967) and Eveleigh and Threlfall (1974) found that in the field, the life cycle required 4–5 years, with all instars being capable of overwintering. Ixodes uriae can be abundant and obvious on sea birds, especially on chicks (see photos in Murray and Vestjens 1967; Bergström et al. 1999a). This species has been shown to reduce growth rate and prolong time to fledging for Cassin’s auklet (Ptychoramphus aleuticus (Pallas)) nestlings in British Columbia (Morbey 1996). Many sea birds nest in dense colonies under conditions that support large populations of ticks. Their impact under such conditions is probably greatest on chicks in the nests. Though the pathological effects of the bite of I. uriae have been studied on an atypical host (Eveleigh et al. 1974), there is little information on the localised damage done by tick feeding. In the Southern Hemisphere, on the other hand, there are reports of impaired reproductive performance (Mangin et al. 2003) and perhaps even deaths (Gauthier-Clerc et al. 1998) in king penguins as a result of heavy infestation with I. uriae. There are a great many pathogenic organisms known to be transmitted to sea birds by I. uriae (Yunker 1975; Chastel 1988; Muzafarr and Jones 2004; Dietrich et al. 2011). Main et al. (1973; 1976a, 1976b) were the first to isolate three different viruses in sea bird colonies in eastern Canada, all most likely being transmitted by I. uriae. There have

Map. 8. Collection localities for Ixodes uriae in Canada. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 113

been numerous isolates of these viruses since that time (Oprandy et al. 1988). Borrelia garinii Baranton et al. is a spirochaete similar to Borrelia burgdorferi and is known to cause neurologic symptoms of Lyme disease in Europe (Rauter and Hartung 2005) and is transmitted in sea bird colonies by I. uriae. This pathogen has been isolated from this tick from island sea bird colonies off the coast of Newfoundland and Labrador (Smith et al. 2006). Because of the potential of I. uriae to disperse over considerable distances (McCoy et al. 2003) and to bite humans, there is a risk among those who visit sea bird colonies to become infected with a variety of potential pathogens.

Distribution. Ixodes uriae is a tick associated with sea birds and their colonies in coastal areas and islands around the world. In Canada, it has been found on the east coast in Newfoundland and Nova Scotia, and on the west coast of British Columbia. It will no doubt be found in other maritime regions of Canada where suitable hosts are found.

Hosts. Sea birds are the primary hosts, and I. uriae is found attached to the birds, or loose in nests and surrounding rubble. Ixodes uriae is known to feed on humans, especially people who are working in sea bird colonies and handling sea birds.

Ixodes (Trichotoixodes) brunneus Koch (Figs. 28–29, Map 9) Ixodes brunneus Koch, 1844: 232. Ixodes avisugus Berlese, 1889: fascicle 55, number 5. Ixodes frontalis Neumann, 1899: 133 (in part). Ixodes californicus Banks, 1904: 369. Ixodes kelloggi Nuttall and Warburton, 1908: 396. Ixodes frontalis brunneus: Schulze 1933: 437. Ixodes brunneus Koch: Anastos and Smith 1957: 535 (male, nymph, larva). Ixodes (Ixodes) brunneus Koch: Clifford et al. 1973: 495. Ixodes (Trichotoixodes) brunneus Koch: Filippova 1977: 248.

Adult. Female: Body: Length from 1.9–2.2 mm unfed, reaching to 9.0 mm engorged. Gnathosoma: Porose areas of basis capituli large, superficial, with indistinct margins and area between them depressed (Fig. 28A). Dorsal posterior margin of basis capituli slightly concave, with posterolateral corners salient or extended into short, bluntly rounded cornua (Fig. 28A). Auriculae formed as large lateral extensions, variable in shape, usually terminating with 2 corners (Fig. 28F). Anterior margin of basis capituli sloped, slightly humped, on either side of hypostome. Hypostome narrow, with pointed apex, dentition 4/4, becoming 3/3 then 2/2 near base (Fig. 28D). Palpi long, slender, with lateral margin profile nearly straight. Palpal tibiotarsus not visible dorsally. Palpal trochanter with a dorsal and ventral ridge, but lacking flanges. Scutum: About 1.3–1.4 times longer than wide, widest slightly anteriad of middle; posterolateral margins slightly convex (Fig. 28A). Scapulae moderate, narrowly rounded. Lateral carinae absent. Cervical grooves extending nearly to posterolateral margins. Surface with small to medium-sized puncta tending to confluence and striate medially and laterally. Legs: Coxa I with stout internal spur slightly longer than external spur or subequal in size (Fig. 28F). Coxae II–IV lacking internal spurs. Coxae II–IV each with small, bluntly pointed external spur, slightly smaller on IV. Ventral coxal surfaces with scattered smooth or slightly barbed setae. Trochanters I–III each with small ventral spur; trochanter IV with vestige of spur (Fig. 28F). 114 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Adult. Male: Body: Length approximately 1.9–2.3 mm. Gnathosoma: Basis capituli dorsal posterior margin straight, with rounded posterior corners instead of cornua; dorsal face slightly furrowed. Ventrally, anterior margin of basis capituli sloped, slightly humped on either side of hypostome. Apex of hypostome truncated or notched (Fig. 28I). Hypostome dentition 3/3 with approximately 6–8 blunt, somewhat crenulated teeth per file. Palpi thick, club-like in dorsal view, with symmetrically rounded apices. Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter lacking flanges. Scutum: Scapulae short, rounded (Fig. 28G). Lateral carinae absent. Surface generally coarsely punctate. Surface lacking pair of pit-like depressions midlaterally, but with paired grooves midlaterally. Spiracular plate: Suboval, with clearly longer axis anteroposterior. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median plate longer than anal plate. Anal plate weakened, not widened, posteriorly. Adanal plates variable in shape, widest anteriorly, tapered posteriorly. Ventral plates moderately punctate. Legs: Coxa I with stout, blunt internal spur slightly longer than external spur; coxae II–IV lacking internal spurs, but II and III with salient posterior corners (Fig. 28J). Coxae I–IV each with blunt external spur. Ventral coxal surfaces with scattered, moderately long, smooth or slightly barbed setae. Trochanters I–III each with small ventral spur; trochanter IV with vestige of spur.

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli concave, with posterolateral corners salient, extended into short, bluntly pointed cornua (Fig. 28C). Auriculae formed as large, lateral extensions, variable in shape, usually terminating with 2 corners (Fig. 28B). Anterior margin of basis capituli sloped, slightly humped, on either side of hypostome. Two pairs of post-hypostomal setae. Apex of hypostome narrow, pointed, dentition 3/3, becoming 2/2 along basal half. Palpi long, slender, with lateral margin profile nearly straight. Palpal tibiotarsus not visible dorsally. Palpal trochanter with a dorsal and ventral ridge, but lacking flanges. Scutum: Length nearly 1.1 greater than width, shape subtriangular or rhomboid, widest slightly anteriad of middle, with slightly convex posterolateral margins (Fig. 28E). Scapulae moderate, narrowly rounded. Lateral carinae absent. Cervical grooves divergent posteriorly nearly to posterolateral margins. Surface with small to medium-sized puncta. Legs: Coxa I with short, pointed internal spur similar in size to more broadly pointed external spur, coxae II–IV lacking internal spurs (Fig. 29K). Coxae II–IV each with small, bluntly pointed external spur, progressively slightly smaller from II–IV. Ventral coxal surfaces with scattered, moderately long, smooth setae. Trochanter spurs not determined.

Larva. Body: Body length, excluding gnathosoma, 0.53–0.56 mm. Gnathosoma: Dorsal posterior margin straight or slightly concave, with blunt lateral corners instead of cornua (Fig. 29L). Ventrally, auriculae formed as large, triangular, lateral projections; lateral margins of basis anterior to auriculae projected laterally, giving impression of 2 auriculae on each side (Fig. 29M). Apex of hypostome bluntly pointed, dentition 3/3 in anterior third, then 2/2 nearly to base; lateral file with 8–10 teeth, second file with 7–9 teeth (Fig. 29N). Two pairs post-hypostomal setae (Fig. 29N). Palpi moderately slender; femur+genu length to width ratio nearly 3 (2.6–2.7):1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter weakly formed, lacking flanges.Scutum: Length about 0.8 times width, widest near mid-level (Fig. 29L). Scapulae weakly developed; shield posterior margin convex. Surface densely micropuncate, and reticulated anterolaterally. Setae, 5 pairs. Idiosomal setation (Figs. 29L–M): Central dorsal setae, 4 to usually 6 pairs. Supplementary dorsal A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 115

B D F

C E

G H I J

Figs. 28A–J. Ixodes brunneus. Fig. 28A, female gnathosoma and scutum, dorsal; insert enlarged cornua; Fig. 28B, nymph gnathosoma, ventral; Fig. 28C, nymph gnathosoma, dorsal; Fig. 28D, female hypostome, ventral; Fig. 28E, nymph scutum; Fig. 28F, female gnathosoma, coxae and trochanters, ventral; Fig. 28G, male scutum; Fig. 28H, male gnathosoma, dorsal; Fig. 28I, male gnathosoma, ventral; Fig. 28J, male coxae, ventral. Figs. 28A, D, F redrawn from Cooley and Kohls (1945); Figs. 28B, C, E, G–J redrawn from Anastos and Smith (1957). 116 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

M N

Figs. 29K–N. Ixodes brunneus. Fig. 29K, nymph coxae, ventral; Fig. 29L, larva, dorsal; Fig. 29M, larva, ventral; Fig. 29N, larva hypostome, palp trochanter, basis capituli, ventral. Figs. 29K–M redrawn from Anastos and Smith (1957); Fig. 29N redrawn from Webb et al. (1990). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 117

setae, 1 pair. Marginal dorsal setae, 7 pairs. Marginal ventral setae, 3 pairs. Premarginal ventral setae, 3–5 pairs. Preanal ventral setae, 3 pairs. Dorsal setae relatively long, length of anterior marginal dorsal setae about 4 times that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in small size. Legs: Coxa I with small triangular internal spur (Fig. 29M). Coxae II and III each with curved ridge posteroventrally. Coxae I and II each with small triangular external spur, this reduced or lacking on coxa III. Coxae I–II–III setal formula 3–2–2; coxal setae moderately short, smooth. Femur III with 9 setae. Trochanters I–III lacking spurs, but sometimes with a small posteroventral tubercle apically.

Natural history. Ixodes brunneus is a parasite of birds, especially ground-foraging species in the eastern and midwestern United States of America. It is not established in Canada, but because of its propensity to attach to migrating birds, it has been recorded frequently in collections from birds during banding efforts (Durden and Keirans 1996; Scott et al. 2001; Morshed et al. 2005). All instars of I. brunneus are most prevalent during September to March (Sonenshine and Stout 1970; Sonenshine 1979b; Kinsey et al. 2000; Kollars and Oliver 2003), and curiously, males have never been collected in the field. This tick is seldom reported in great abundance, though it was the predominant species collected by Sonenshine and Stout (1970) at their study site in North Carolina. They collected 103 specimens from 1070 dark-eyed juncos (Junco hyemalis (Linnaeus)). Although Rickettsia rickettsii has been isolated from I. brunneus, the status of this tick as a potential vector is unknown (Clifford et al. 1969). However, there is good evidence that I. brunneus may cause paralysis in birds (Bishopp and Tremblay 1945; Gregson 1973; Schwab 1987; Luttrell et al. 1996).

Distribution. Ixodes brunneus is widely distributed in the United States of America, with records from Oregon and California in the west, eastwards through Kansas and Wisconsin, to New York and Maryland in the east. It is found southwards into Texas across to Florida. It is not known to be endemic in Canada, but there are records from migratory birds in New Brunswick, Ontario, and Manitoba.

Hosts. This tick infests ground-foraging birds, and has been recorded from a large number of host species, especially passeriform birds.

Map 9. Collection localities for Ixodes brunneus in Canada. 118 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Ixodes (Ixodes) dentatus Marx (Figs. 30–31, Map 10) Ixodes dentatus Marx in Neumann, 1899: 119. Ixodes dentatus Marx: Smith 1940: 17 (male, nymph, larva). Ixodes (Ixodes) dentatus Marx: Clifford et al. 1973: 495.

Adult. Female: Body: Length 1.8–2.0 mm unfed, reaching to 9.0 mm engorged. Gnathosoma: Porose areas of basis capituli variable in shape, from almost circular to clearly triangular, flattened at posterior margin, separated by nearly the width of one porose area (Fig. 30A). Dorsal posterior margin of basis capituli straight, cornua prominent, bluntly pointed (Fig. 30A). Auriculae sharply pointed, projecting ventrally and posteriorly (Fig. 30C). Anterior margin of basis capituli sloped on either side of hypostome. Hypostome lanceolate, tapering from middle to a blunt point, dentition 6/6 near apex, then 5/5, and 4/4, and finally 3/3 at base (Fig. 30B). Palpi moderately long, not club-like (Fig. 30A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with a sharp spur on posteroventral surface. Scutum: Slightly longer than wide. Shape oval, widest slightly anterior to middle, posterolateral margins convex (Fig. 30A). Scapulae bluntly pointed. Lateral carinae long, slightly curved. Cervical grooves faint, meeting lateral carinae posteriorly. Surface with many coarse puncta, especially numerous posteriorly, progressively smaller and irregular anteriorly. Legs: Coxa I with internal spur tapered, pointed, much longer than external spur, coxae I–IV each with short external spur, these similar in size or slightly diminishing from I–IV (Fig. 30C). Ventral coxal surfaces with long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 30C).

Adult. Male: Body: Length approximately 1.9. Gnathosoma: Basis capituli dorsal posterior margin straight or slightly convex, with small cornua; dorsal face without furrows (Fig. 30D). Ventrally, anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded at tip, tapering gradually from near centre, dentition armed with approximately 11 diagonal rows of crenulate plates on each side, each with a somewhat free blunt lateral tooth flanking diagonal series of approximately 5 scallops (Fig. 30F). Palpi moderately robust, club-like in dorsal view, with symmetrically rounded apices (Fig. 30D). Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter with a broad ventral prominence. Scutum: Scapulae prominent, bluntly pointed. Lateral carinae absent. Surface with numerous, coarse puncta, puncta deeper posteromedially. Surface lacking pair of pit-like depressions mid-laterally. Spiracular plate: Oval, with clearly longer axis anteroposterior. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae (Fig. 30E). Median plate about twice as long as anal plate. Anal plate not widened posteriorly, similar in width to adanal plates. Ventral plates with numerous coarse puncta. Legs: Coxa I with internal spur tapered, pointed, much longer than external spur, coxae II and III each with spur- like salience on posterior corner, lacking on IV (Fig. 30E). Coxae I–IV each with bluntly pointed external spur, smaller on I. Ventral coxal surfaces with scattered, moderately long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 30E).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli straight, with bluntly pointed posterior corners instead of cornua (Fig. 31G). Auriculae formed as lateral lobes (Fig. 31I). Anterior margin of basis capituli sloped on either side of hypostome. Hypostome slightly lanceolate, tapering from middle to rounded apex, dentition 4/4 near apex, then A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 119

3/3 at mid-level, and finally 2/2 at base (Fig. 31H). Two pairs of post-hypostomal setae. Palpi moderately long, not conspicuously club-like when viewed dorsally (Fig. 31G). Palpal tibiotarsus not visible dorsally. Palpal trochanter with a small ventral projection. Scutum: Length about 0.9 times width (Fig. 31G). Shape subcircular, widest at middle, posterolateral margins convex. Scapulae very small, blunt. Lateral carinae faint. Cervical grooves shallow, slightly divergent posteriorly nearly to posterolateral margins. Surface with sparse puncta, progressively larger posteriorly, lacking wrinkled or roughened areas. Legs: Coxa I with internal spur tapered, pointed, much longer than external spur, coxae II–IV with more or less salient posterior corners suggesting internal spurs (Fig. 31I). Coxae I–IV each with short external spur, these similar in size or slightly diminishing from I–IV. Ventral coxal surfaces with scattered smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 31I).

Larva. Body: Body length, excluding gnathosoma, approximately 0.6 mm. Gnathosoma: Dorsal posterior margin of basis capituli straight or slightly sinuous, with blunt lateral corners instead of cornua (Fig. 31J). Ventrally, auriculae undeveloped or weakly formed as diagonal ridges (Fig. 31K). Apex of hypostome rounded (Fig. 31K). Hypostomal dentition 3/3 on apical half, then 2/2 nearly to base; lateral file with 9–11 teeth, second file with 8–9 teeth, third file with approximately 5 teeth. Two pairs of post-hypostomal setae. Palpi moderately slender; femur+genu length to width ratio approximately 3:1; ventral apex of femorogenu with a small tubercle or flange. Palpal trochanter distinct, with slight ventral ridge. Scutum: Length about 0.8 times width, widest slightly anterior to mid-level (Fig. 31J). Scapulae weakly developed; shield posterior margin convex. Surface densely uniformly micropunctate, and lineate-reticulate along anterolateral margins. Setae, 5 pairs. Idiosomal setation (Figs. 31J, K): Central dorsal setae, 3 pairs. Supplementary dorsal setae, 1 pair. Marginal dorsal setae, 7 pairs. Marginal ventral setae, 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae at least twice that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in small size. Legs: Coxa I with large, bluntly pointed, subtriangular internal spur, coxae II and III each with angular or curved ridge posteroventrally (Fig. 31K). Coxae I–III lacking external spurs. Coxae I–II–III setal formula 3–2 or 3–3; coxal setae smooth or inconspicuously barbed. Femur III with 9 setae. Trochanters I–III lacking spurs (Fig. 31K).

Natural history. Records for this species in Canada are very sparse (Barker et al. 1992; Morshed et al. 2005), and there is no information on its natural history in Canada. Ixodes dentatus actively parasitised rabbits throughout the year in Missouri, where larvae exhibited two peaks of activity in May and September, nymphs were most abundant in spring and summer, and adult numbers peaked in May (Kollars and Oliver 2003). It undoubtedly requires more than one year in most parts of its range to complete a generation (Smith 1945). In Massachusetts, males were never observed to feed, but were commonly observed on the host, where mating occurred. Females required 4–40 days to engorge, and began to lay eggs 7–24 days after engorgement, depending on conditions; 398–3648 eggs were laid over 12–38 days. Larvae engorged in about six days and moulted 34–110 days later. In some cases, where engorged larvae dropped off late in the season, moulting to the nymphal instar occurred in the following spring. Nymphs engorged in about six days and required 35–144 days to moult to become adults. Smith (1945) found that unfed ticks could live for a long time, adults for two years or more, larvae from 300–612 days, and nymphs two years 120 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A C

D E

Figs. 30A–F. Ixodes dentatus. Fig. 30A, female gnathosoma and scutum, dorsal; Fig. 30B, female hypostome, ventral; Fig. 30C, female gnathosoma, coxae and trochanters, ventral; Fig. 30D, male gnathosoma and scutum, dorsal; Fig. 30E, male gnathosoma, idiosoma, coxae, and trochanters, ventral; Fig. 30F, male hypostome, ventral. Redrawn from Cooley and Kohls (1945). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 121

G H I

J K

Figs. 31G–K. Ixodes dentatus. Fig. 31G, nymph gnathosoma and scutum, dorsal; Fig. 31H, nymph hypostome, ventral; Fig. 31I, nymph gnathosoma, coxae and trochanters, ventral; Fig. 31J, larva, dorsal; Fig. 31K, larva, ventral. Figs. 31G–I redrawn from Cooley and Kohls (1945); Figs. 31J, K redrawn from Clifford et al. (1961). 122 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Map 10. Collection localities for Ixodes dentatus in Canada.

or more. In the field, larvae were most abundant on cottontail rabbits in April and May, and in September and October. In New York, Battaly et al. (1987) also observed a bimodal peak in the abundance of larvae on birds. Nymphs were abundant throughout April to October; females were most abundant in April to June, but were present in August (Smith 1945). The propensity of larvae and nymphs to attach to birds has resulted in the number of diverse records reported in eastern Canada by Morshed et al. (2005). Ixodes dentatus may occasionally bite humans (Hall et al. 1991; Anderson et al. 1996), so its potential as a vector of pathogens is of concern. It is a vector of Babesia divergens McFadyean and Stockman (Telford and Spielman 1989; Goethert and Telford 2003a), Anaplasma phagocytophilum (Goethert and Telford 2003b), and strains of Borrelia burgdorferi (Anderson et al. 1989; Oliver et al. 1996; Lin et al. 2001).

Distribution. Ixodes dentatus is found throughout the United States of America east of the Mississippi River, with the exception of northernmost (Minnesota, New Hampshire, Maine) and southernmost (Florida) states. Adventive records for this tick have been reported in Canada only from migratory birds in Ontario and Manitoba.

Hosts. Ixodes dentatus is primarily a parasite of lagomorphs, most often reported on the eastern cottontail rabbit, but juveniles commonly attach to and feed on ground-foraging birds.

Ixodes (Ixodes) muris Bishopp and Smith (Figs. 32–33, Map 11) Ixodes muris Bishopp and Smith, 1937: 133 (all instars). Ixodes (Ixodes) muris Bishopp and Smith: Clifford et al. 1973: 495.

Adult. Female: Body: Length about 1.9 mm unfed, reaching to at least 6.0 mm engorged. Gnathosoma: Porose areas of basis capituli large, subcircular, superficial, and separated by less than diameter of one porose area (Fig. 32A). Dorsal posterior margin of basis capituli straight, salient. Cornua distinct, small, triangular (Fig. 32A). Auriculae bluntly pointed, projecting posterolaterally (Fig. 32C). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome narrow, pointed, with all denticles long, narrow, pointed, arranged 4/4 subapically, then 3/3, and basally 2/2 (Fig. 32B). Palpi long, narrow, nearly straight in lateral margin profile when viewed dorsally (Fig. 32A). Palpal tibiotarsus not A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 123

visible dorsally. Palpal trochanter lacking flanges.Scutum: About 1.2–1.3 times longer than wide; widest slightly anterior of the middle, with posterolateral margins slightly convex (Fig. 32A). Scapulae long, bluntly pointed. Lateral carinae faint, situated near lateral margins. Cervical grooves shallow, not reaching posterolateral margins. Surface with numerous fine puncta. Legs: Coxa I with internal spur tapered, pointed, much longer than external spur. Coxae II–IV lacking internal spurs (Fig. 32C). Coxae I–IV each with short external spur, becoming slightly smaller from I–IV. Ventral coxal surfaces with scattered, moderately long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 32C).

Adult. Male: Body: Length approximately 1.7 mm. Gnathosoma: Basis capituli dorsal posterior margin nearly straight, with small, blunt cornua; dorsal face without furrows (Fig. 32D). Ventrally, anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome conspicuously notched, dentition roughly 4/4, with 7 or 8 diagonal rows, laterals as sharp denticles, inner rows as pointed crenulations (Fig. 32F). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 32D). Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter lacking flanges. Scutum: Scapulae rounded (Fig. 32D). Lateral carinae absent. Surface with numerous small puncta, usually with a few larger puncta marginally. Surface lacking pair of pit-like depressions midlaterally. Spiracular plate: Oval, with clearly longer axis anteroposterior. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae (Fig. 32E). Median plate about twice as long as anal plate. Anal plate widened posteriorly, somewhat wider than adanals there. Ventral plates with numerous, large puncta. Legs: Coxa I with internal spur tapered, blunt, longer than external spur (Fig. 32E). Coxae II–IV lacking internal spurs, but II and III with posterior saliences. Coxae I–IV each with short, blunt external spur, becoming slightly smaller from I–IV. Ventral coxal surfaces with scattered, moderately long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 32E).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli straight, cornua distinct, small, triangular (Fig. 33G). Auriculae formed as conspicuous, bluntly pointed posterolateral projections (Fig. 33I). Anterior margin of basis capituli sloped on either side of hypostome. Two pairs of post-hypostomal setae. Apex of hypostome narrow, pointed, dentition 3/3 along apical half, then 2/2 to base; denticles long, narrow, pointed, inner denticles smaller (Fig. 33H). Palpi long, narrow, not club-like when viewed dorsally (Fig. 33G). Palpal tibiotarsus not visible dorsally. Palpal trochanter lacking flanges. Scutum: Length about 1.2 width. Shape oval, widest slightly anteriad of middle, with posterolateral margins slightly convex (Fig. 33G). Scapulae bluntly pointed. Lateral carinae absent. Cervical grooves moderately long, divergent but not reaching posterolateral margins. Surface with sparse, fine puncta. Legs: Coxa I with internal spur bluntly pointed, longer and more tapered than external spur. Coxae II–IV lacking internal spurs (Fig. 33I). Coxae I–IV each with short external spur, becoming slightly smaller from I–IV. Ventral coxal surfaces with scattered, moderately long, smooth or slightly barbed setae. Trochanters I– IV lacking spurs (Fig. 33I).

Larva. Body: Body length 0.5–0.6 mm unengorged. Gnathosoma: Dorsal posterior margin sinuous, with cornua divergent, bluntly pointed (Fig. 33J). Ventrally, auriculae formed as large, triangular, lateral projections (Fig. 33K). Apex of hypostome narrowly rounded, dentition 3/3 along apical third, then 2/2 nearly to base; lateral file with 9–10 124 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

B A C

D E

Figs. 32A–F. Ixodes muris. Fig. 32A, female gnathosoma and scutum, dorsal; Fig. 32B, female hypostome, ventral; Fig. 32C, female gnathosoma, coxae and trochanters, ventral; Fig. 32D, male gnathosoma and idiosoma, dorsal; Fig. 32E, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 32F, male hypostome, ventral. Redrawn from Cooley and Kohls (1945). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 125

G I

J K

Figs. 33G–K. Ixodes muris. Fig. 33G, nymph gnathosoma and scutum, dorsal; Fig. 33H, nymph hypostome, ventral; Fig. 33I, nymph gnathosoma, coxae and trochanters, ventral; Fig. 33J, larva, dorsal; Fig. 33K, larva, ventral. Figs. 33G–I redrawn from Cooley and Kohls (1945); Figs. 33J, K, redrawn from Clifford et al. (1961). 126 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins teeth, second file with 8–9 teeth, innermost file with 4–5 teeth (Fig. 33K). Two pairs of post-hypostomal setae. Palpi moderately slender; femur+genu length to width ratio nearly 3.0:1; ventral apex of femorogenu lacking a tubercle or flange (Fig. 33J). Palpal trochanter distinct, lacking flanges. Scutum: Length nearly equal (0.9–1.0 times) to width, widest at, or slightly behind, mid-level (Fig. 33J). Scapulae weakly developed, rounded; shield posterior margin nearly flat. Surface finely micropunctate, and fully reticulate, more conspicuously anterolaterally. Setae, 5 pairs. Idiosomal setation (Figs. 33J–K): Central dorsal setae, 4 pairs. Supplemental dorsal setae, 1 pair. Marginal dorsal setae, 7 pairs. Marginal ventral setae, 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length anterior marginal dorsal setae scarcely (at most 1.5 times) longer than those of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in length. Legs: Coxa I with large, bluntly pointed internal spur. Coxae II and III each with curved internal ridge posteroventrally (Fig. 33K). Coxae I and II each with bluntly triangular external spur, smaller on II, this vestigial or absent on III. Coxae I–II–III setal formula 3–2 or 3–3; coxal setae mostly conspicuously unilaterally barbed. Femur III with 9 setae. Trochanters I–III lacking spurs (Fig. 33K).

Natural history. Although there are many records for this mouse tick in eastern Canada, there have been no detailed studies into its natural history in this region (e.g., Martell et al. 1969; Barker et al. 1992). The most complete examination of its natural history was conducted in Massachusetts (Smith 1944), from which the following has been summarised. Larvae became abundant on rodents in mid-summer, though small numbers were present as early as March and were still found on rodents until October and November. They required about four days (2–9 days) to complete engorgement and drop off the host. Larvae that were isolated in tubes in the field, with no access to a host, survived up to 490 days, but did not survive through two winters. Larvae moulted in late summer and fall, and unfed nymphs overwintered, attaining their greatest abundance on mice in the spring. Some unfed nymphs survived 2–3 years. Nymphs required about three days (2–8 days) to engorge. These nymphs could moult to adults in as few as 25 days in the field, but those that fed in the fall overwintered and moulted the following spring. Mating occurred off the host, and males were never observed to feed. There was no distinct peak in the abundance of adults,

Map 11. Collection localities for Ixodes muris in Canada. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 127

which infested mice in every month from March to September. Females engorged in 4–8 days; those that engorged in the spring and summer began to lay eggs within two weeks, while those that engorged late in the season overwintered in that state and laid eggs in the spring. Once oviposition began, females laid all their eggs within about three weeks, and laid up to 1380 eggs. In the field, eggs hatched in 9–11 weeks. Unfed females survived more than two years under field conditions. Although I. muris is predominately a parasite of mice, juveniles have been reported on birds (Scott et al. 2001; Scharf and Walker 2003; Scharf 2004; Morshed 2005) and this certainly provides opportunities for this species to disperse well beyond its established range. There is no definitive evidence that I. muris is an important vector of pathogens to humans, though Spielman et al. (1984) and Goethert et al. (2003) have suggested its involvement in transmission of Babesia microti in Connecticut. Ixodes muris appears to be a relatively poor vector for Borrelia burgdorferi (Dolan et al. 2000).

Distribution. Ixodes muris is known to occur across the northern tier of United States of America east of the Mississippi River, and there are numerous records across southeastern Canada from the maritime provinces to Manitoba.

Hosts. Ixodes muris is a parasite of small mammals, including mice, voles, and shrews, and is known to feed on cats, dogs and humans. The immature instars frequently attach to migratory ground-foraging birds.

Ixodes (Ixodes) pacificus Cooley and Kohls Western blacklegged tick (Figs. 34–35, Map 12) Ixodes californicus Banks, 1908: 24 (not Ixodes californicus Banks, 1904). Ixodes pacificus Cooley and Kohls, 1943: 140 (female, male, nymph). Ixodes pacificus Cooley and Kohls: Allred et al. 1960: 16 (larva). Ixodes (Ixodes) pacificus Cooley and Kohls: Keirans and Clifford 1978: 104.

Adult. Female: Body: Length about 2.6 mm unfed, reaching to 9.0 mm engorged. Gnathosoma: Porose areas of basis capituli oval or subtriangular, separated by slightly less than the width of one porose area (Fig. 34A). Dorsal posterior margin of basis capituli concave, with blunt corners instead of cornua. Auriculae formed as short diagonal ridges (Fig. 34C). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded, dentition 4/4 in distal portion, then 3/3, and 2/2 near base (Fig. 34B). Lateral denticles large, pointed, inner denticles smaller, rounded, progressively smaller medially. Palpi long, lateral profile margin nearly straight in dorsal view (Fig. 34A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with a blunt triangular point laterally. Scutum: Length slightly greater (1.1 times) than width; oval, widest slightly anterior to midlength, with posterolateral margins convex (Fig. 34A). Scapulae bluntly pointed. Lateral carinae faint but traceable in some specimens. Cervical grooves usually visible only in reflected light as shallow depressions that diverge and end before reaching posterolateral margins. Surface with uniformly distributed small puncta and scattered long setae. Legs: Coxa I with internal spur long, tapering, pointed, much longer than external spur (Fig. 34C). Coxae II–IV lacking internal spurs, but II and III with salient posterior corners. Coxae I–IV each with small, bluntly pointed external spur. Ventral coxal surfaces with scattered long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 34C). 128 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Adult. Male: Body: Length 2.0–2.4 mm. Gnathosoma: Basis capituli dorsal posterior margin straight, with obtuse corners instead of cornua; dorsal face without furrows (Fig. 34D). Ventrally, anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome broadly rounded, dentition with a file of prominent lateral teeth bordering 5 or 6 diagonal rows of crenulations medially, these progressively larger towards base (Fig. 34F). Palpi thick, club-like in dorsal view, with symmetrically rounded apices. Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter with small ventral projection. Scutum: Scapulae bluntly rounded. Lateral carinae absent. Surface with numerous, large puncta, fewer and finer anteriorly. Surface lacking pair of pit-like depressions midlaterally. Spiracular plate: Oval, with clearly longer axis anteroposterior (Fig. 34G). Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median plate about twice as long as anal plate (Fig. 34E). Anal plate slightly widened posteriorly, where slightly wider than adanal plates. Ventral plates with numerous small puncta and fine setae.Legs: Coxa I with internal spur tapered, sharply pointed, much longer than external spur (Fig. 34E). Coxae II–IV lacking internal spurs, but II and III with salient posterior corners. Coxae I–IV each with small, bluntly pointed external spur. Ventral coxal surfaces with scattered long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 34E).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli nearly straight, with definite, bluntly pointed cornua (Fig. 35H). Auriculae formed as shelf-like to bluntly angular lateral projections (Fig. 34I). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome narrowly rounded, dentition 3/3 in distal portion, then 2/2 nearly to base; lateral denticles more pointed and larger than inner ones (Fig. 35J). Two pairs of post-hypostomal setae. Palpi long, not club-like, lateral profile margin nearly straight in dorsal view. Palpal tibiotarsus not visible dorsally. Palpal trochanter with a faint, pointed, ventrolateral projection. Scutum: Oval to subcircular, slightly wider than long, widest at about midlength, with posterolateral margins convex (Fig. 35H). Scapulae short, bluntly pointed. Lateral carinae nearly straight. Cervical grooves distinct, divergent posteriorly nearly to posterolateral margins. Surface with sparse puncta and scattered long setae. Legs: Coxa I with internal spur tapered, bluntly pointed, longer than external spur. Coxae II–IV lacking internal spurs, but II and III with salient posterior corners (Fig. 35I). Coxae I–IV each with small, bluntly pointed external spur, these similar in size. Ventral coxal surfaces with scattered long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 35I).

Larva. Body: Body length 0.5–0.6 mm. Gnathosoma: Dorsal posterior margin of basis capituli straight, with blunt corners instead of cornua (Fig. 35K). Ventrally, auriculae well formed as blunt lateral projections (Fig. 35L–M). Apex of hypostome rounded, dentition 3/3 subapically, then 2/2 to base; lateral file with 9–10 pointed, overlapping teeth, inner file with 6–8 smaller, blunter teeth (Fig. 35M). Two pairs of post-hypostomal setae (Fig. 35M). Palpi moderately slender; femur+genu length to width ratio at least 3:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, lacking flanges. Scutum: Length about 0.8 (0.77–0.83 times) width, widest near or slightly anterior to mid- level (Fig. 35K). Scapulae weakly formed; shield posterior margin convex. Surface finely micropunctate, with underlying reticula becoming emergent anterolaterally. Setae, 5 pairs. Idiosomal setation (Figs. 35K, L): Central dorsal setae, 4 pairs. Supplemental dorsal setae, 1 pair. Marginal dorsal setae, 7 pairs. Marginal ventral setae, 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 129

setae 3–4 times that of scutal setae (Fig. 35K). Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in size. Legs: Coxa I with bluntly triangular internal spur (Fig. 35L). Coxae II and III each with curved ridge posteroventrally. Coxae I and II each with small, blunt external spur, this lacking on coxa III. Coxae I–II–III setal formula 3–2–3; coxal setae smooth or inconspicuously barbed. Femur III with 9 setae. Trochanters I–III lacking spurs (Fig. 35L).

Natural history. Ixodes pacificus is a three-host tick with a wide host range in all active instars. Its western distribution and similar appearance to Ixodes scapularis, the blacklegged tick, account for its common name, the western blacklegged tick. In Canada, adults feed primarily on large to medium-sized mammals such as deer, dogs, cats, and sheep (Gregson 1956). Columbian black-tailed deer, Odocoileus hemionus columbianus, are the primary hosts for adults in the United States of America (Westrom et al. 1985; Foley et al. 2004). In a comprehensive review of the host records for this species in California, Castro and Wright (2007) reported adults from 29 species of mammals and two species of birds. In Canada, Gregson (1935b) believed that larvae and nymphs feed primarily on lizards (Gerrhonotus Wiegmann), and only occasionally feed on small mammals or birds. However, Arnason (1992) noted frequent infestation of small mammals, especially deer mice (Peromyscus maniculatus) by I. pacificus nymphs and larvae. Larvae, and to a greater extent nymphs, also infest northern alligator lizards, Elgaria coerulea (Wiegmann) (Arnason 1992). Though larvae and nymphs parasitise 38 different species of mammals in California, two species of lizards are primary hosts for immatures (Castro and Wright 2007). Squirrels may also be important hosts (and pathogen reservoirs) for I. pacificus larvae and nymphs (Nieto and Foley 2008; Salkeld et al. 2008). Ixodes pacificus larvae and nymphs have also been reported from many species of passeriform birds (Castro and Wright 2007), with some species having a prominent role as hosts and reservoirs for pathogens (Eisen et al. 2004; Wright et al. 2006). As noted by Morshed et al. (2005), birds may also disperse I. pacificus immatures to new localities. All active instars of the western blacklegged tick infest people (Castro and Wright 2007) though nymphs and adults are the only epidemiologically relevant life stages. In California, adult I. pacificus (and nymphs) are one of the most common ticks biting humans (Lane 1990), and in British Columbia, adults of this species frequently bite humans (Gregson 1942a). In British Columbia, I. pacificus likely requires at least three years to complete its life cycle as it does in northern California (Padgett and Lane 2001). Because of the temperate climate in coastal British Columbia, adults of I. pacificus can be collected in every month of the year but they are most abundant from February to June, with peak abundance in May (Arnason 1992). In California, males and females actively seek hosts (Lane 1990) and feed upon large mammals in the fall, winter, and early spring (Westrom et al. 1985). Cohorts of adult ticks likely do not overlap, because unfed adults usually die prior to emergence of the new fall cohort (Padgett and Lane 2001). Adults are frequently collected from vegetation along sun-exposed deer trails or hiking paths (Li et al. 2000) and more adults occurred on south-facing slopes than north-facing ones (Arnason 1992). The abundance of adults and nymphs of western blacklegged ticks varies among habitat types; Arnason (1992), observed adults in British Columbia most frequently in coastal Douglas fir (Pseudotsuga menziesii (Mirbel) Franco; Pinaceae) habitats. In California, though adults are present across a range of wooded habitats (Brown et al. 2006; Killilea et al. 2008), they were most frequently collected in sites classified as hardwood-conifer (Eisen et al. 2006). In the laboratory, females require 6–11 days to feed to repletion (Arthur and Snow 1968; Troughton and 130 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A B C

D E G

Figs. 34A–G. Ixodes pacificus. Fig. 34A, female gnathosoma and scutum, dorsal; Fig. 34B, female hypostome, ventral; Fig. 34C, female gnathosoma, coxae and trochanters, ventral; Fig. 34D, male gnathosoma and idiosoma, dorsal; Fig. 34E, male gnathosoma, coxa and trochanters, ventral; Fig. 34F, male hypostome, ventral; Fig. 34G, male spiracular plate. Redrawn from Cooley and Kohls (1945). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 131

H I

K L

Figs. 35H–M. Ixodes pacificus. Fig. 35H, nymph gnathosoma and scutum, dorsal; Fig. 35I, nymph gnathosoma, coxae and trochanters, ventral; Fig. 35J, nymph hypostome ventral; Fig. 35K, larva, dorsal; Fig. 35L, larva, ventral; Fig. 35M, larva hypostome, palp trochanter, basis capituli, ventral. Figs. 35H–J redrawn from Cooley and Kohls (1945); Figs. 35K modified, 35L–M redrawn from Webb et al. (1990). 132 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Levin 2007). Mating is required for females to engorge fully and this typically occurs on the host (Westrom et al. 1985). Temperature influences the duration of the pre-oviposition period in I. pacificus and replete females placed in the field from December to March deposited eggs from February to April (Peavey and Lane 1996; Padgett and Lane 2001). In laboratory studies using western blacklegged ticks from British Columbia, Arthur and Snow (1968) reported that fed females held at 18–20 °C produced 793–1301 eggs. Larvae of I. pacificus typically hatch from mid-July to late August (Peavey and Lane 1996; Padgett and Lane 2001). Temperature influences the rate of larval hatch such that larvae hatch earlier from eggs deposited later in the season than those laid earlier. Unfed larvae remain in behavioural diapause and do not actively seek hosts until late winter or early spring (Padgett and Lane 2001). Survival rates of unfed larvae vary depending upon habitat and larval cohorts do not appear to overlap (Padgett and Lane 2001). Larval abundance on small mammals in British Columbia was greatest in May and June (Arnason 1992), consistent with the pattern observed in the United States of America. The prevalence and abundance of larvae on hosts decline thereafter with relatively few larvae observed on hosts in August. In the laboratory, larvae feed for up to 10 days (Arthur and Snow 1968) but most drop from their hosts on the fourth and fifth days (Troughton and Levin 2007). Timing of the larval moult to nymphs is influenced by timing of feeding, habitat type, and topographic exposure (i.e., earlier for ticks with southern and western exposure); as a rule, most larvae fed in the spring moulted to nymphs by August (Padgett and Lane 2001). In California, nymphs typically start host-seeking by mid-March, reaching a peak in early May and disappear by late July or mid-August (Eisen et al. 2002). The lengths of the periods of maximum nymphal activity were positively correlated with rainfall and negatively with temperature. Unfed nymphs are active throughout the day and can frequently be collected from logs, the trunks of trees (Slowik and Lane 2001b; Lane et al. 2007), and from leaf litter along shaded trails (Li et al. 2000). Nymphs infest hosts throughout late winter to early summer in California and the maximum abundance of nymphs on hosts occurs several weeks after the peak in larval abundance (Lane and Loye 1989; Manweiler et al. 1992), though there is considerable seasonal overlap between these two life stages. In British Columbia, Arnason (1992) observed similar seasonal patterns of infestation (i.e., nymphs on hosts from April to August) and co-infestation of deer mice by larvae and nymphs. Survival of unfed nymphs varies by habitat type (Eisen et al. 2002) and topographic exposure and cohorts of unfed nymphs typically do not overlap (Padgett and Lane 2001). In the laboratory, the feeding period for nymphs is 6–11 days but most complete feeding in 5–7 days (Arthur and Snow 1968; Troughton and Levine 2007). Most fed nymphs moult to adults in late summer, and this new cohort of adults begins host-seeking in the late fall and winter (Padgett and Lane 2001). Overall, the life cycle of I. pacificus is regulated by temperature, relative humidity and day length. In California, rising summer temperatures accelerate most biological processes, such as larval emergence and moulting. However, these conditions also increase mortality such that most ticks, regardless of life stage, die during the summer drought (Padgett and Lane 2001). Similar, but less dramatic influences may regulate I. pacificus population dynamics in British Columbia, as adults were never collected when air temperatures exceeded 23 °C (Arnason 1992) and in the laboratory, most instars experienced high rates of mortality when exposed to low relative humidity (Arthur and Snow 1968). Adult western blacklegged ticks bite humans in Canada (Gregson 1942a; Arnason 1992) and the United States of America (Lane 1990; Merten and Durden 2000). According to Gregson (1942a), feeding by female I. pacificus can be painful and after tick removal, a persistent swelling can form, lasting for days, sometimes accompanied by a slow- A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 133

healing ulcer, which may last for months. In Canada, the agent of Lyme disease, Borrelia burgdorferi, has been detected in I. pacificus and small numbers of human infections are reported annually (Ogden et al. 2008c). The low disease incidence is consistent with patterns of human disease reported in the far western states (Bacon et al. 2008). As has been reported in other parts of its range (Lane et al. 2001; Holden et al. 2003; Lane et al. 2004), the infection rates of B. burgdorferi in host-seeking I. pacificus in British Columbia are very low (Ogden et al. 2008c), likely because immature ticks preferentially feed on lizards, which are incompetent hosts (Lane and Quistad 1998; Casher et al. 2002). Thus in British Columbia, the risk of human infection with B. burgdorferi is very low, partly because of the widespread but focal nature of the established tick populations and the low prevalence of B. burgdorferi in vector ticks (Ogden et al. 2008c). Elsewhere in the far west of North America, I. pacificus transmits Anaplasma phagocytophilum, the agent of human granulocytic anaplasmosis (HGA), to humans (Fritz et al. 2005). Though human cases of HGA have not been reported in British Columbia, a symptomatic case in a dog was reported from Vancouver Island (Lester et al. 2005). Further studies are required to determine whether the agent of HGA is established in western blacklegged tick populations in British Columbia. In rare instances, bites from I. pacificus have caused severe allergic reactions in humans, presumably the result of repeated exposure to tick bites (Van Wye et al. 1991). Antibodies to B. burgdorferi and signs of clinical disease have been reported in dogs from British Columbia (Banerjee et al. 1996). Given the tendency for adult I. pacificus to bite dogs, this result is not surprising. At least one case of HGA in a dog has been reported in British Columbia (Lester et al. 2005), though the burden of HGA in horses and dogs, in other parts of the range of I. pacificus is much greater (Richter et al. 1996; Foley et al. 2001, 2004) than in British Columbia. Bartonella henselae DNA has been amplified from adult western blacklegged ticks in California (Holden et al. 2006); however, proof of vector competency will require experimental transmission studies with known infected ticks (Billeter et al. 2008). In California, the agent of tularaemia has been detected in I. pacificus and this tick has also been implicated in cases of tick paralysis in dogs, cattle, deer, ponies (Lane et al. 1984; Lane 1990), and mice (Botzler et al. 1980).

Map 12. Collection localities for Ixodes pacificus in Canada. 134 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Distribution. Ixodes pacificus is distributed throughout western North America from Fort Smith in the Northwest Territories (probably an extralimital record) and British Columbia south to Baja California, Mexico, east into Nevada, Idaho, Utah, and Arizona. It was first reported in Canada by Gregson (1935b); established populations in Canada are restricted to British Columbia. Records from outside British Columbia are the result of travel to endemic areas. These extralimital records illustrate the importance of determining travel history when dealing with important vectors of pathogenic agents.

Hosts. This tick will feed on a great variety of hosts, including small mammals, large domestic animals, birds and reptiles. It readily attacks humans.

Ixodes (Ixodes) scapularis Say Blacklegged tick (Fig. 36, Map 13) Ixodes scapularis Say, 1821: 78. Ixodes fuscous Say, 1821: 79. Ixodes reduvius Neumann, 1899: 112 (in part — North American records from New Berne, North Carolina; Baltimore, Maryland; “Carolina;” Florida; Kansas; Texas; and Pennsylvania (specific localities not cited for the four latter states); not the record from California). Ixodes pratti Banks, 1908: 25 (in part, includes I. texanus Banks, 1909) (male). Ixodes ricinus var. scapularis: Nuttall and Warburton, 1911: 156. Ixodes scapularis Say: Hooker et al. 1912: 77 (nymph, larva). Ixodes ozarkus Cooley and Kohls, 1944: 287. Ixodes dammini Spielman, Clifford, Piesman, and Corwin, 1979: 218 (all instars). Ixodes scapularis Say: Oliver et al. 1993: 54 (synonymy with Ixodes dammini).

Adult. Female: Body: Length 2.40–2.70 mm unfed, reaching to 10.00 mm engorged. Gnathosoma: Porose areas of basis capituli large, suboval to subtriangular, slightly depressed into surface, separated by slightly less than width of one porose area (Fig. 36A). Dorsal posterior margin of basis capituli slightly concave. Cornua small, but evident. Auriculae formed as short slightly bulged, diagonal ridges (Fig. 36D). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome broadly rounded, dentition 4/4 in distal portion, then 3/3 and 2/2 to base; lateral denticles large, pointed, inner ones smaller, rounded (Fig. 36B). Palpi long, lateral profile margin nearly straight (Fig. 36A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with a slight rounded projection laterally. Scutum: Length slightly greater (1.1–1.2 times) than width; oval, widest at midlength, with posterolateral margins convex (Fig. 36A). Scapulae bluntly pointed. Lateral carinae absent. Cervical grooves faint, fading out where diverging and not reaching posterolateral margins. Surface with small, numerous puncta, larger marginally and posteriorly, and scattered moderately long setae mostly confined to anterior area.Legs: Coxa I with internal spur long, tapering, sharply pointed, much longer than external spur. Coxae II–IV lacking internal spurs, but II and III with salient posterior corners (Fig. 36D). Coxae I–IV each with small, bluntly pointed external spur. Ventral coxal surfaces with scattered long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 36D).

Adult. Male: Body: Length 1.9–2.1 mm. Gnathosoma: Basis capituli dorsal posterior margin nearly straight, with obtuse corners instead of cornua; dorsal face without furrows (Fig. 36G). Ventrally, anterior margin of basis capituli sloped on either side of A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 135

hypostome. Apex of hypostome broadly rounded or slightly flattened, dentition with a file of prominent lateral teeth bordering 6 or 7 diagonal rows of crenulations medially, these progressively larger towards base (Fig. 36E). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 36G). Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter with small ventral projection. Scutum: Scapulae bluntly rounded (Fig. 36G). Lateral carinae absent. Surface with numerous, large puncta, deeper posteromedially and marginally, fewer and smaller anteriorly. Surface lacking pair of pit-like depressions midlaterally. Spiracular plate: Elongate-oval, with clearly longer axis anteroposterior (Fig. 36C). Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae (Fig. 36H). Median plate about 3 times as long as anal plate. Anal plate not widened posteriorly, where similar in width to adanal plates. Median plate with numerous large, deep puncta; other ventral plates with smaller puncta. Legs: Coxa I with internal spur tapered, sharply pointed, much longer than external spur. Coxae II–IV lacking internal spurs, but II and III with salient posterior corners (Fig. 36H). Coxae I–IV each with small, bluntly pointed external spur. Ventral coxal surfaces with scattered long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 36H).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli sinuous, convex medially (Fig. 36I). Cornua prominent, pointed, directed posteriorly. Auriculae formed as small, pointed, posteriorly-directed projections (Fig. 36J). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome narrowly rounded, dentition 3/3 in distal portion, then 2/2 to base; lateral denticles more pointed and larger than inner denticles (Fig. 36F). Two pairs of post-hypostomal setae. Palpi long, not club-like, lateral profile margin nearly straight in dorsal view (Fig. 36I). Palpal tibiotarsus not visible dorsally. Palpal trochanter with a slight, pointed ventrolateral projection. Scutum: Length slightly greater (1.1–1.2 times) than width; oval, widest at midlength, with posterolateral margins convex (Fig. 36I). Scapulae bluntly pointed. Lateral carinae absent. Cervical grooves faint, fading out where diverging and well short of reaching posterolateral margins. Surface sparsely punctate. Legs: Coxa I with internal spur short, bluntly pointed, hardly longer than external spur. Coxae II–IV lacking internal spurs, but II and III with salient posterior corners (Fig. 36J). Coxae I–IV each with small, bluntly pointed external spur, these similar in size. Ventral coxal surfaces with scattered long, smooth, or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 36J).

Larva. Body: Body length from anterior scutal margin to posterior body margin 0.5–0.6 mm. Gnathosoma: Dorsal posterior margin of basis capituli sinuous, with divergent, bluntly pointed cornua (Fig. 36K). Ventrally, auriculae formed as small to conspicuous, subtriangular, lateral projections (Fig. 36L). Apex of hypostome narrowly rounded to bluntly pointed, dentition 3/3 along anterior third, then 2/2 nearly to base; lateral file with 8–10 pointed, overlapping teeth; inner file with 6–7 smaller, slightly blunter teeth (Fig. 36L). Two pairs of post-hypostomal setae. Palpi slender; femur+genu length to width ratio 3–4:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, lacking flanges. Scutum: Length 0.82–0.95 times width, widest slightly anterior to mid-level (Fig. 36K). Scapulae small, bluntly rounded; shield posterior margin slightly convex. Surface densely micropunctate, with faint underlying reticula becoming emergent anterolaterally and anteriorly. Setae, 5 pairs. Idiosomal setation (Figs. 36K, L): Central dorsal setae, 3 pairs. Supplemental dorsal setae, 1 pair. Marginal dorsal setae, 7 pairs. 136 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

C D

A B

I J

G H K L

Figs. 36A–L. Ixodes scapularis. Fig. 36A, female gnathosoma and scutum, dorsal; Fig. 36B, female hypostome, ventral; Fig. 36C, male spiracular plate; Fig. 36D, female gnathosoma, coxae and trochanters, ventral; Fig. 36E, male hypostome, ventral; Fig. 36F, nymph hypostome, ventral; Fig. 36G, male gnathosoma and idiosoma, dorsal; Fig. 36H, male gnathosoma, idiosoma, coxae, ventral; Fig. 36I, nymph gnathosoma and scutum, dorsal; Fig. 36J, nymph gnathosoma, coxae and trochanters, ventral; Fig. 36K, larva, dorsal; Fig. 36L, larva, ventral. Figs. 36A–J redrawn from Cooley and Kohls (1945); Figs. 36K redrawn, L modified from Clifford et al. (1961). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 137

Marginal ventral setae, 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae similar to, or up to 3 times that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in form and size, either simple or unilaterally barbed. Legs: Coxa I with bluntly triangular internal spur. Coxae II and III each with a small, rounded internal spur or ridge (Fig. 36L). Coxa I–III each with a small, blunt external spur, progressively smaller or vestigial on III. Coxa I–II–III setal formula 3–2 or 3–3; coxal setae similar in form, either short, smooth, or longer, unilaterally barbed. Femur III with 9 setae. Trochanters I–III lacking spurs (Fig. 36L).

Natural history. In North America, Lyme disease was initially associated with the blacklegged tick, Ixodes scapularis (Steere et al. 1978). The tick vector involved was subsequently described as a closely-related new species, the northern deer tick, I. dammini (Spielman et al. 1979). However, based primarily on hybridisation and assortative mating studies between I. scapularis and I. dammini, Oliver et al. (1993) concluded that I. dammini should be considered a junior synonym of I. scapularis. Because of the prevalence of adult ticks feeding on deer, I. scapularis is sometimes referred to as the deer tick, but its official common name is the blacklegged tick. Ixodes scapularis is a three-host tick. Adults attach almost exclusively to large mammals (Anderson 1988). White-tailed deer (Odocoileus virginianus) is considered the most important host for adults (Wilson et al. 1990), but raccoon, red fox, opossum, dogs, and cats, and livestock are also infested (Anderson and Magnarelli 1984; Spielman et al. 1985; Schulze et al. 1985, 1986; Fish and Dowler 1989). Larvae and nymphs feed mainly on small mammals and birds (Anderson 1988, 1989). Peromyscus species (white-footed mouse and deer mouse) are considered the most important mammalian hosts for these instars (Levine et al. 1985; Donahue et al. 1987), but Microtus species (voles), sciurids (squirrels and chipmunks) (Mannelli et al. 1993), shrews, and other small mammals can also be infested (Anderson 1989). Medium-sized mammals, such as opossum, groundhogs, skunks, and raccoons can be significant hosts for immature I. scapularis in endemic areas, and may be particularly important in suburban habitats, where Peromyscus are less abundant (Fish and Dowler 1989). A large variety of ground-dwelling birds are infested by I. scapularis larvae or nymphs (Battaly et al. 1987; Stafford et al. 1995; Nicholls and Callister 1996) and these birds serve as one of the primary mechanisms for short-range and long-range dispersal of blacklegged ticks into Canada (Battaly and Fish 1993; Klich et al. 1996; Morshed et al. 1999, 2005; Scott et al. 2001, 2012; Ogden et al. 2008b). All active instars of blacklegged ticks will bite people (Falco and Fish 1988). In non-endemic areas of Canada, adult females are the instar most often reported from people or pets (Morshed et al. 2006; Ogden et al. 2006c). The most detailed study on the life history of I. scapularis in Canada was conducted at Long Point, Ontario by Lindsay (1995). Throughout its range in Canada, I. scapularis likely requires 3–4 years to complete its life cycle (Lindsay et al. 1998). Unlike the other human-biting ticks in Canada, unfed adults of I. scapularis begin active host-seeking in fall rather than spring. Adults first appear in late September and are most active in October and April though they can be collected as late as mid-December and from mid-March to July (Lindsay et al. 1999a). Adults seeking hosts in the fall and subsequent spring represent the same cohort of ticks. Adults that do not acquire a host in the fall enter a quiescent phase when cold temperatures and snow cover inhibit host-seeking behaviour (Duffy and Campbell 1994); these ticks resume host-seeking in the following spring. Quiescence may 138 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins be incomplete and adults can be collected in the winter months in parts of its range, or in certain years, when extended periods of snow cover or sub-zero temperatures are not experienced (Watson and Anderson 1976). Overwinter survival is variable but typically survival rates are highest for fed females followed by unfed females and males (Lindsay et al. 1998). Adult activity declines following the April peak and adult cohorts rarely overlap because unfed adults do not survive later than August (Lindsay et al. 1998). Adults (and nymphs) are frequently collected in deciduous or coniferous woodlands and from ecotonal (i.e., unmaintained edge) habitats (Maupin et al. 1991), though these instars are also collected on lawns and ornamental shrubbery in suburban residential areas (Stafford and Magnarelli 1993). The abundance of adult blacklegged ticks varies among habitats (Guerra et al. 2002; Brownstein et al. 2005) and from year to year (Lindsay et al. 1999a, 1999b). Female adult blacklegged ticks require 6–10 days to feed to repletion (Troughton and Levin 2007). Males also take blood meals which are completed in less than 24 hours (Krinsky 1979). Mating is required before females engorge fully and occurs most frequently on the host, while females are feeding; however, I. scapularis may mate before they find a host (Yuval and Spielman 1990a). Dispersal by fed females following disengagement is minimal though they do seek protected microhabitats in which to lay eggs (Stafford 1992). Temperature influences, in a non-linear fashion, the duration of the pre-oviposition and oviposition period in I. scapularis (Ogden et al. 2004). In field studies on Long Point, Ontario, regardless of whether females fed in the fall or subsequent spring, eggs were laid during late April and early May (Lindsay et al. 1998). The number of eggs produced per female is dependent on engorgement weight and temperature such that females held at 12–28 °C produced 9034 ± 1706 eggs per gram of fed weight (Lindsay 1995). Larvae hatch during mid- to late July (Lindsay et al. 1998) and rarely actively disperse from egg masses (Stafford 1992), though this instar as well as nymphs can be carried great distances on avian hosts (Scott et al. 2001; Morshed et al. 2005; Ogden et al. 2008b). Unfed larvae may feed during the year of hatch or overwinter without feeding and seek hosts the following year (Lindsay et al. 1998). As a result, I. scapularis larvae typically have two peaks of activity, though in some years they are indistinct (Lindsay et al. 1999b). Overwintered larvae tend to be most abundant on hosts from May to June. The prevalence and abundance of larvae decrease in July and then usually increase again in August when the larvae of that year hatch (Lindsay et al. 1999b). Larval ticks are most abundant in wooded habitats (Maupin et al. 1991; Lindsay et al. 1999a) and survival is predicted to occur at more northerly latitudes in the face of climatic change (Lindsay et al. 1995; Ogden et al. 2006a). Larvae feed for up to seven days, but most drop from their hosts on the fourth day (Troughton and Levin 2007). Engorged larvae (and nymphs) tend to detach from their hosts during the day; the resulting nymphs accumulate in the nest of the host, where they have a higher probability of encountering a host and obtaining a second blood meal (Mather and Spielman 1986). Timing of the larval moult to nymphs is dependent primarily on temperature; larvae that feed later in the year (late July onward), delay moulting until the following year (Ogden et al. 2004). However, fewer larvae successfully moult if they overwinter than those that moult before overwintering. Habitat type and its impact on vapour pressure deficit, also influence the proportion of larvae that survive to moult to nymphs (Lindsay et al. 1998). Ixodes scapularis nymphs are active from early May to late September, though they tend to be most abundant in the environment and on hosts during June and July (Lindsay et al. 1999b). Nymphs are the longest lived instar of the blacklegged tick and unfed nymphs may survive under field conditions an average of 3.4 months longer than unfed adults A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 139

(i.e., about 16 months). Overwintering survival of unfed nymphs is influenced by habitat type and varies among years (Lindsay et al. 1998). Nymphs feed for 3–8 days but most complete feeding in four days (Troughton and Levine 2007). Nymphs fed from April to the end of July moulted in the year of feeding, while those fed later moulted in the subsequent year. Diapause is driven by day-length and may affect rates at which nymphs develop into adults (Ogden et al. 2004). Unlike with larvae, the proportion of nymphs that successfully moulted is similar whether or not they overwinter (i.e., 43.5–46.3%). Host-seeking by I. scapularis nymphs on Long Point precedes the emergence of larvae of the year and this inverted seasonal activity is believed to aid amplification of pathogens (Spielman et al. 1985). Co-feeding of larvae and nymphs may be important for transmission of B. burgdorferi from infected nymphs to larvae (Lindsay et al. 1997). Earlier onset of developmental diapause for nymphs and larvae of I. scapularis is the primary factor that extends the life cycle of this tick in southern Ontario (Lindsay et al. 1998) compared to blacklegged tick populations in the northeast (Yuval and Spielman 1990b) and midwestern United States of America (Platt et al. 1992). Adults and nymphs readily bite humans (Falco and Fish 1988) and in some endemic areas in the United States of America, repeated tick bites can be quite common (Nadelman and Wormser 2007). Dermal reactions in people to uninfected blacklegged tick bites are usually unremarkable but development of a tick-associated itch from repeated tick bites has been reported (Burke et al. 2005). A vigourous immune response from multiple bites may lead to severe pruritus which results in prompt recognition and removal of attached ticks and a decrease in the risk of exposure to Lyme disease compared to people without a dermal reaction. Each year “bird-borne” I. scapularis infected with B. burgdorferi and Anaplasma phagocytophilum, the agents of Lyme disease and human anaplasmosis, respectively, are introduced into widely separated localities in Canada (Morshed et al. 2006; Ogden et al. 2006c, 2008d). These introduced ticks represent a persistent risk of pathogen transmission to people and domestic animals over a large portion of Canada. However, compared to the localities with established blacklegged tick populations, the risk is low because nymphs are infrequently encountered, contact with blacklegged ticks is relatively infrequent and the prevalence of pathogens is much lower. Blacklegged tick populations infected with B. burgdorferi and A. phagocytophilum are limited to a small number of localities in Canada (Drebot et al. 2001; Ogden et al. 2006c); however, the range of blacklegged ticks continues to expand and this trend will likely continue, partly aided by climate change (Ogden et al. 2005, 2006b, 2008a, 2008d). The focal nature of established infected blacklegged tick populations is partly responsible for the relatively small number of human cases of Lyme disease reported in Canada (Ogden et al. 2008c) compared to the United States of America (Bacon et al. 2008). Though A. phagocytophilum is established in some blacklegged tick populations in Manitoba, Ontario, and Nova Scotia, only one locally acquired case of human anaplasmosis has been reported in Canada (Parkins 2008). Blacklegged ticks also transmit a number of other human pathogens in the United States of America, including Babesia microti and deer tick virus, a variant of Powassan virus (Anderson et al. 1991; Ebel et al. 2000). Typically the prevalence of these agents is much lower than B. burgdorferi (Varde et al. 1998; Holman et al. 2004; Kogut et al. 2005; Brackney et al. 2008) and these agents may be absent from some blacklegged tick populations. For example, B. microti was not detected in selected tick populations in Maryland (Swanson and Norris 2007) and Michigan (Hamer et al. 2007). Further study is required to determine the extent to which these pathogens are present in blacklegged ticks in Canada, though it is possible that bird-borne I. scapularis, infected with these agents, 140 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins could cause locally acquired infections among Canadians (Ogden et al. 2008c). The first blacklegged tick infected with B. microti was found in Manitoba in 2013. Studies in the United States of America have also demonstrated the presence of other micro-organisms in blacklegged ticks including: Bartonella (Adelson et al. 2004) and Rickettsia (Swanson and Norris 2007); however, some have challenged the role of ticks in transmission (Matsumoto et al. 2008) and a causal relationship has not been well established for detection of some of these agents and human disease (Halperin and Wormser 2001). Dogs, cats, cattle, horses, and possibly sheep can be infected with the agent of Lyme disease (Sonenshine et al. 2002) through the bite of infected blacklegged ticks. In Canada, dogs are often infested by I. scapularis ticks (Ogden et al. 2006c) and thus are at a heightened risk for exposure to B. burgdorferi and A. phagocytophilum. Clinical disease in dogs has been described for both these agents (Dickinson 2008), though a large proportion of dogs exposed to B. burgdorferi are asymptomatic (Littman et al. 2006). Horses appear to be susceptible to disease as a result of exposure to both of these pathogens (Sonenshine et al. 2002; Pusterla et al. 2002). Ixodes scapularis has also been reported to cause tick paralysis in rare cases in dogs (Gregson 1973).

Distribution. Ixodes scapularis is found throughout the eastern United States of America east of the Rocky Mountains and into Mexico. In Canada, established populations are known in Manitoba, Ontario, Québec, New Brunswick, and Nova Scotia, but the propensity of this tick to attach to migratory birds supports its incursion into parts of Canada well beyond its breeding range. There are numerous records of adult I. scapularis from surveillance programmes in Saskatchewan and Alberta, not all of which are indicated on Map 13. Many extralimital records in Canada are also associated with travel to endemic areas.

Hosts. This tick will attach to and feed on a great variety of hosts, including small mammals, domestic animals, birds, and reptiles. There is one interesting record of larvae from northern pocket gophers near Clavet, Saskatchewan (Anstead and Chilton 2011). All active instars feed readily on humans.

Map 13. Collection localities for Ixodes scapularis in Canada. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 141

Ixodes (Ixodes) spinipalpis Hadwen and Nuttall (Fig. 37, Map 14) Ixodes diversifossus Neumann, 1911: 14 (not Ixodes diversifossus Neumann, 1899). Ixodes diversifossus Bishopp, 1912: 30 (not Ixodes diversifossus Neumann, 1899). Ixodes dentatus spinipalpis Hadwen and Nuttall in Nuttall 1916: 301 (female, nymph , larva). Ixodes neotomae Cooley, 1944: 7. Ixodes spinipalpis Hadwen and Nuttall: Cooley and Kohls 1945: 51 (female, male, nymph). Ixodes (Ixodes) spinipalpis Hadwen and Nuttall: Clifford et al. 1973: 495. Ixodes neotomae Cooley: Norris et al. 1997: 702 (synonymy).

Adult. Female: Body: Length about 1.9 mm unfed, reaching to 9.2 mm engorged. Gnathosoma: Porose areas of basis capituli roughly circular, small, separated by interval more than diameter of one porose area (Fig. 37A). Dorsal posterior margin of basis capituli straight, with short, bluntly pointed cornua (Fig. 37A). Auriculae formed as pair of curved, posteriorly-pointed horns (Fig. 37D). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome narrowly rounded, dentition 4/4 in distal portion, then 3/3, and 2/2 to base (Fig. 37B). Lateral denticles large, pointed, inner ones smaller, blunter. Palpi moderately long, lateral profile margin nearly straight (Fig. 37A). Palpal tibiotarsus not visible dorsally. Palpal femorogenu with elongate setae on ventral face of femoral region. Palpal trochanter with a short, pointed ventrolateral spur. Scutum: Length 1.2–1.3 times greater than width, oval, widest at midlength, with posterolateral margins convex (Fig. 37A). Scapulae pointed. Lateral carinae as low rounded ridges, extending from near scapulae nearly to posterolateral margins. Cervical grooves distinct but shallow, not extending to posterolateral margins. Surface with puncta slightly coarser posteriorly. Legs: Coxa I with internal spur long, tapering, pointed, much longer than external spur (Fig. 37D). Coxae II–IV lacking internal spurs. Coxae I–IV each with small, bluntly pointed external spur, these progressively shorter from I–IV, that on IV sometimes scarcely more than a salience. Ventral coxal surfaces with scattered long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 37D).

Adult. Male: Body: Length 1.6–1.8 mm. Gnathosoma: Basis capituli dorsal posterior margin nearly straight, with bluntly pointed cornua directed posteriorly (Fig. 37G). Ventrally, anterior margin of basis capituli sloped, slightly humped on either side of hypostome. Apex of hypostome broad, indented, dentition with a file of prominent lateral teeth bordering approximately 6 diagonal rows of crenulations medially, these progressively larger towards base (Fig. 37C). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 37G). Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter with ventral ridge. Scutum: Scapulae bluntly rounded (Fig. 37G). Lateral carinae absent. Surface with faint anterior puncta grading into numerous large, shallow, puncta posteriorly and marginally. Surface lacking pair of pit-like depressions midlaterally. Spiracular plate: Subcircular, with slightly longer axis anteroposterior. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae (Fig. 37H). Median plate over twice as long as anal plate. Anal plate widened posteriorly, where wider than adanal plates. Ventral plates with numerous large puncta moderate in size and number, these smaller on anal plate. Legs: Coxa I with internal spur tapered, pointed, much longer than external spur (Fig. 37H). Coxae II–IV lacking internal spurs, but II and III with salient posterior corners. Coxae I–IV each with small, bluntly pointed external spur. Ventral coxal surfaces with scattered long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 37H). 142 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

E C D

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Figs. 37A–L. Ixodes spinipalpis. Fig. 37A, female gnathosoma and scutum, dorsal; Fig. 37B, female hypostome, ventral; Fig. 37C, male hypostome, ventral; Fig. 37D, female gnathosoma, coxae and trochanters, ventral; Fig. 37E, nymph gnathosoma and scutum, dorsal; Fig. 37F, nymph gnathosoma, coxae and trochanters, ventral; Fig. 37G, male gnathosoma and idiosoma, dorsal; Fig. 37H, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 37I, nymph hypostome, ventral; Fig. 37J, larva hypostome, palp trochanter, basis capituli, ventral; Fig. 37K, larva, dorsal; Fig. 37L, larva, ventral. Figs. 37A–I redrawn from Cooley and Kohls (1945); Figs. 37J–L redrawn from Webb et al. (1990). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 143

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli straight, with short, bluntly pointed cornua (Fig. 37E). Auriculae formed as bluntly triangular lateral projections (Fig. 37F). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome narrowly rounded, dentition 3/3 in apical third, then 2/2 to base; lateral denticles more pointed and larger than inner ones (Fig. 37I). Two pairs of post-hypostomal denticles. Palpi long, not club-like, lateral profile margin nearly straight in dorsal view (Fig. 37E). Palpal tibiotarsus not visible dorsally. Palpal trochanter with a short, blunt ventrolateral projection. Scutum: Length 1.1-1.2 times greater than width, oval, widest near midlength, with posterolateral margins slightly convex (Fig. 37E). Scapulae pointed. Lateral carinae nearly straight, extending from near scapulae nearly to posterolateral margins. Cervical grooves distinct but shallow, divergent, not extending to posterolateral margins. Surface with sparse puncta and scattered long setae. Legs: Coxa I with internal spur tapered, bluntly pointed, longer than external spur (Fig. 37F). Coxae II–IV lacking internal spurs. Coxae I–IV each with small, bluntly pointed external spur, these progressively shorter from I–IV. Ventral coxal surfaces with scattered long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 37F).

Larva. Body: Body length, slightly engorged, including gnathosoma 0.9 mm. Gnathosoma: Dorsal posterior margin nearly straight, with bluntly pointed lateral corners instead of cornua (Fig. 37K). Ventrally, auriculae formed as blunt lateral projections (Fig. 37J). Apex of hypostome rounded, dentition 3/3 along apical third, then 2/2 to base; lateral file with 9–10 pointed, overlapping teeth; second file with 8–9 smaller, blunter teeth; subapical file with 4–5 yet smaller teeth (Fig. 37J). Two pairs of post-hypostomal setae (Fig. 37J). Palpi slender; femur+genu length to width ratio 3.5–4.0:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, lacking flanges, but with small ventral ridge. Scutum: Length about 0.85–0.90 times width, widest slightly anterior to mid-level (Fig. 37K). Scapulae weakly formed, bluntly rounded; shield posterior margin slightly convex. Surface finely micropunctate, and with emergent reticula anterolaterally. Setae, 5 pairs. Idiosomal setation (Figs. 37K–L): Central dorsal setae, 2 pairs. Supplemental dorsal setae, 1 pair. Marginal dorsal setae, 7 pairs. Marginal ventral setae, 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae nearly three times that of scutal setae. Posteriormost pairs of ventral marginal and premarginal

Map 14. Collection localities for Ixodes spinipalpis in Canada. 144 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins setae similar in size to their dorsal counterparts. Sternal setae similar to one another in moderate length. Legs: Coxa I with a prominent, tapered, bluntly pointed internal spur (Fig. 37L). Coxae II and III each with a curved ridge posteroventrally. Coxae I and II each with a small, blunt external spur, this absent on coxa III. Coxa I–II–III setal formula 3–2–3; coxal setae smooth or inconspicuously barbed. Femur III with 9 setae. Trochanters I–III lacking spurs (Fig. 37L).

Natural history. Ixodes spinipalpis has been called a nest species, despite its occurrence on a wide variety of hosts, including birds and humans (Gregson 1956). It is a frequent parasite of wood rats and pikas, and perhaps occurs in greater numbers in the nests of these hosts. However, it has recently been implicated in transmission of a variety of pathogens in the United States of America, including Borrelia burgdorferi (Burkot et al. 1999), Borrelia bissetti (Schnieder et al. 2000), Babesia microti (Burkot et al. 2001b), and the agent of human granulocytic anaplasmosis (Zeidner et al. 2000). The importance of I. spinipalpis to maintain an enzootic cycle in small mammals was identified by Burkot et al. (2001a) when they found that larvae and nymphs commonly infested sentinel mice a considerable distance from wood rat nests. There is little information on the occurrence and natural history of I. spinipalpis in Canada, but host records provided by Gregson (1956) certainly support the notion that this species may quest for hosts over a considerable distance. Molecular evidence, along with absence of distinct morphological differences between each of the instars and overlap in habitat, geographic distribution, and host preferences, indicates that Ixodes neotomae should be treated as a junior synonym of I. spinipalpis (Norris et al. 1997).

Distribution. Ixodes spinipalpis is a species of coastal and western North America. There are records from British Columbia and southwestern Alberta to southern California, as far east as South Dakota and Texas.

Hosts. This tick feeds primarily on a variety of shrews, small rodents, and lagomorphs, but larvae and nymphs are also known to feed on birds. It has occasionally been recorded from humans.

Ixodes (Ixodiopsis) angustus Neumann (Fig. 38, Map 15) Ixodes angustus Neumann, 1899: 136. Ixodes angustus Neumann: Nuttall and Warburton 1911: 195 (male, nymph, larva). Ixodes (Pholeoixodes) angustus Neumann: Clifford et al. 1973: 498. Ixodes (Ixodiopsis) angustus Neumann: Filippova 1977: 133. Ixodes (Pholeoixodes) angustus Neumann: Keirans and Clifford 1978: 42. Ixodes (Ixodiopsis) angustus Neumann: Robbins and Keirans 1992: 14.

Adult. Female: Body: length 1.6–2.1 mm unfed, reaching to 7.0 mm engorged. Gnathosoma: Porose areas large but variable, ranging from almost oval to triangular, often with indefinite margins, but clearly separated medially (Fig. 38A). Dorsal posterior margin of basis capituli straight to slightly sinuous or concave, with salient corners instead of cornua (Fig. 38A). Auriculae absent (Fig. 38J). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome pointed, dentition 3/3 throughout nearly entire length; denticles pointed, those in lateral and inner files similar in size (Fig. 38C). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 145

Palpi long, lateral margin profile nearly straight (Fig. 38A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with a short triangular projection ventrolaterally. Scutum: Length about 1.4 times longer than wide; oval, widest at mid-length, with posterolateral margins slightly convex (Figs. 38A–B). Scapulae pointed. Lateral carinae variable, usually distinct, occasionally faint. Cervical grooves shallow, faint, not approaching posterolateral margins. Surface sparsely, rather evenly punctate. Legs: Coxa I with internal spur bluntly pointed, short to moderately long, at least slightly longer than external spur (Figs. 38I–J). Coxae II–IV lacking internal spurs. Coxae I–IV each with bluntly triangular external spur, sometimes progressively smaller from I–IV. Ventral coxal surfaces with few, moderately long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 38J).

Adult. Male: Body: Length 1.6–2.3 mm. Gnathosoma: Basis capituli dorsal posterior margin nearly straight, with blunt corners instead of cornua; dorsal face without furrows (Fig. 38F). Ventrally, anterior margin of basis capituli sloped, slightly humped on either side of hypostome. Apex of hypostome slightly flattened, dentition 3/3 plus few irregular medial denticles, arranged in 5–6 transverse, overlapping, crenulated rows (Fig. 38G). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 38F). Dorsally, palpal femur faintly delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter with small ventrolateral ridge. Scutum: Scapulae bluntly pointed (Fig. 38F). Lateral carinae absent. Surface mildly rugose anterolaterally, and finely, moderately punctate medially, but with large, numerous deep puncta laterally. Surface lacking pair of pit-like depressions midlaterally, but with paired grooves there. Spiracular plate: Subcircular; with slightly longer axis dorsoventral. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae (Fig. 38H). Median plate approximately 1.5 longer than anal plate. Anal plate not widened posteriorly, similar in width to adanal plates. Ventral plates with small, shallow puncta. Legs: Coxa I with internal spur tapered, blunt, slightly longer than external spur (Fig. 38H). Coxae II–IV with salient posterior corners instead of internal spurs. Coxae I–IV each with bluntly pointed external spur, these usually similar in size. Ventral coxal surfaces with few, moderately long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 38H).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli straight or slightly concave, with small, bluntly pointed cornua (Fig. 38D). Auriculae formed as mild, lateral bulges (Fig. 38E). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded, dentition 3/3 in distal portion, then 2/2 throughout nearly all of remaining length; denticles bluntly pointed, those in lateral and inner files similar in size (Fig. 38E). Two pairs of post-hypostomal setae. Palpi long, not club-like, lateral margin profile nearly straight in dorsal view (Fig. 38D). Palpal tibiotarsus not visible dorsally. Palpal trochanter with both anterior and posterior flange-like projections well developed, anterior projection narrowly rounded and variable in size, posterior projection bluntly pointed (Fig. 38E). Scutum: Shape rhomboid, slightly wider then long, widest at mid-length, with posterolateral margins nearly straight (Fig. 38D). Scapulae pointed. Lateral carinae faint, nearly straight. Cervical grooves faint, not approaching posterolateral margins. Surface sparsely punctate. Legs: Coxa I with internal spur bluntly pointed, short to moderately long, hardly longer than external spur (Fig. 38E). Coxae II–IV lacking internal spurs. Coxae I–IV each with bluntly triangular external spur, sometimes progressively smaller from I–IV. Ventral coxal surfaces with few, moderately long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 38E). 146 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A B C D E

I J

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M N

Figs. 38A–N. Ixodes angustus. Fig. 38A, B, variations in female gnathosoma and scutum, dorsal; Fig. 38C, female hypostome, ventral; Fig. 38D, nymph gnathosoma and scutum, dorsal; Fig. 38E, nymph gnathosoma, coxae and trochanters, ventral; insert enlarged nymph hypostome, ventral; Fig. 38F, male gnathosoma and idiosoma, dorsal; Fig. 38G, male hypostome, ventral; Fig. 38H, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 38I, J, variations in female gnathosoma, coxae and trochanters, ventral; Fig. 38K, larva hypostome, palp trochanter, basis capituli, ventral; Fig. 38L, larva coxa I, ventral; Fig. 38M, larva, dorsal; Fig. 38N, larva, ventral. Figs. 38A–J, redrawn from Cooley and Kohls (1945); Figs. 38K, L redrawn from Webb et al. (1990); Figs. 38M, N redrawn from Clifford et al. (1961). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 147

Larva. Body: Length (excluding gnathosoma) 0.5–0.8 mm unengorged, 0.6–1.0 mm engorged. Gnathosoma: Dorsal posterior margin straight or slightly concave, with cornua formed as divergent, blunt, lateral projections (Fig. 38M). Ventrally, auriculae not developed. Apex of hypostome rounded, dentition 2/2 throughout; lateral file with 6–7 bluntly pointed, overlapping teeth, inner file with 4–5 smaller, blunter teeth (Fig. 38K). Two pairs of post-hypostomal setae (Fig. 38K). Palpi moderately thick, femur+genu length to width ratio 1.9–2.1:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, with two well-developed flanges of similar size, anterior flange usually more bluntly pointed (Fig. 38K). Scutum: Length about 0.8 times (0.76–0.89 times) width, widest at mid-level. Scapulae weakly formed, blunt; shield posterior margin convex (Fig. 38M). Surface densely micropunctate, with underlying reticula becoming emergent anterolaterally. Setae, 5 pairs. Idiosomal setation (Figs. 38M–N): Central dorsal setae, 2 or 3 pairs. Supplementary dorsal setae absent. Marginal dorsal setae, 7–10 (usually 8) pairs. Marginal ventral setae, 2–5 pairs (usually 3 pairs in eastern North America). Premarginal ventral setae, 3–5 (usually 4) pairs. Preanal ventral setae, 1–3 (usually 2) pairs. Length of anterior marginal dorsal setae about 1.5 that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in size. Legs: Coxa I with bluntly right-triangular internal spur (Fig. 38L). Coxae II and III each with a curved ridge posteroventrally (Fig. 38N). Coxae I–III lacking external spurs. Coxae I–II–III setal formula 3–2 or 3–2; coxal setae smooth or inconspicuously barbed. Femur III with 9 setae. Trochanters I–III lacking spurs (Fig. 38N).

Natural history. The large range and apparent lack of host specificity have resulted in this tick being reported frequently in the literature from many host species, mostly rodents and insectivores (e.g., Bishopp and Trembley 1945; Judd 1950; Gregson 1956; Martell et al. 1969; Whitaker and French 1982; Barker et al. 1992; Durden and Keirans 1996; Nieto et al. 2007; Anstead et al. 2013). Despite being regularly encountered, it is seldom collected in large numbers, and there are limited data from field studies on its natural history. It is most abundant in cool, moist boreal, montane, and riparian habitats (Robbins and Keirans 1992;

Map 15. Collection localities for Ixodes angustus in Canada. 148 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Sorensen and Moses 1998), where it has been reported to exhibit limited host specificity based on factors presumed to influence overlap with its hosts. Sorensen and Moses (1998) and Anstead et al. (2013) found that it was most prevalent on the red-backed vole (Myodes gapperi (Vigors)) in north-central Alberta and Kootenay National Park, British Columbia, respectively, while Murrell et al. (2003) found Peromyscus keeni Merriam to be most heavily infested in Alaska. Juveniles and females of I. angustus were active in north-central Alberta at least from May to the end of August (Sorensen and Moses 1998); Furman and Loomis (1984) reported activity during all months of the year in California. Males are not collected on animals and are presumed not to feed; they are probably largely confined to the nest environment of the host. Ixodes angustus has been reported as a competent vector of the agent that causes Lyme borreliosis, Borrelia burgdorferi (Banerjee et al. 1994; Peavey et al. 2000), though it rarely bites humans. Fay and Rausch (1969) first reported this tick to transmit what was later shown to be a strain of Babesia microti among rodents in Alaska.

Distribution. Ixodes angustus is a Nearctic and eastern Palaearctic species. It is very widely distributed in North America, from Alaska to Newfoundland, south to central California and across to Georgia and South Carolina. In Canada it is distributed throughout the Maritime Provinces, westward through Alberta to coastal British Columbia. We know of no records for Saskatchewan, though it is expected to occur there.

Hosts. This tick is recorded to feed on a great range of insectivores, rodents, and lagomorphs. In Canada, it is also known to feed on cats and dogs, perhaps as a result of their predation of small mammal hosts; it is known to attack humans as well (Cooley 1946a; Gregson 1956; Spencer 1963).

Ixodes (Ixodiopsis) ochotonae Gregson (Figs. 39–40, Map 16) Ixodes ochotonae Gregson, 1941: 224 (all instars). Ixodes holdenreidi Cooley, 1946b: 103. Ixodes (Pholeoixodes) holdenreidi Cooley: Clifford et al. 1973: 498. Ixodes (Pholeoixodes) ochotonae Gregson: Clifford et al. 1973: 498. Ixodes (Ixodiopsis) ochotonae Gregson: Robbins 1989: 291.

Adult. Female: Body: length 2.2–2.4 mm unfed, reaching to nearly 9.0 mm engorged. Gnathosoma: Porose areas of basis capituli large, separated by slightly less than width of one porose area (Fig. 39A). Dorsal posterior margin of basis capituli straight or slightly concave, with salient corners instead of cornua (Fig. 39A). Auriculae absent (Fig. 39C). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome pointed, dentition 3/3 for about distal third, then 2/2 to base; lateral and inner denticles similar in size; lateral denticles pointed, inner denticles more rounded (Fig. 39B). Palpi long, lateral margin profile nearly straight (Fig. 39A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with small ventrolateral projection. Scutum: Length 1.3–1.4 times longer than wide; oval, widest at midlength, with posterolateral margins nearly straight (Fig. 39A). Scapulae pointed. Lateral carinae prominent, extending to posterolateral margins. Cervical grooves shallow, faint. Surface sparsely, evenly punctate, lacking roughened areas. Legs: Coxa I with internal spur short, about as long as external spur (Fig. 39C). Coxa II with trace of an internal spur; coxae III–IV without them. Coxae I–IV each with a distinct external spur, triangular and A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 149

blunt on I–III, rounded and reduced on IV. Ventral coxal surfaces with scattered, moderately short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 39C).

Adult. Male: Body: Length approximately 2.3 mm. Gnathosoma: Basis capituli dorsal posterior margin slightly concave, with blunt corners instead of cornua; dorsal face without furrows (Fig. 39D). Ventrally, anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome broadly rounded, dentition roughly 3/3, arranged in 5 or 6 irregular transverse rows of weak crenulations (Fig. 39F). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 39D). Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter lacking flanges. Scutum: Scapulae bluntly rounded (Fig. 39D). Lateral carinae absent. Surface slightly rugose anterolaterally, with puncta generally shallow, small, and somewhat more conspicuous near margins. Surface lacking pair of pit-like depressions midlaterally. Spiracular plate: Subcircular, without distinct longer axis. Body venter: Caudal margins of ventral plates not lobe-like, lacking flange of long setae. Median plate slightly wider than long, and longer than anal plate (Fig. 39E). Anal plate not widened posteriorly, hardly wider than adanal plates posteriorly. Ventral plates with scattered, faint puncta. Legs: Coxa I with internal spur short, blunt, hardly longer than external spur (Fig. 39E). Coxae II–IV lacking internal spurs, but II and III with salient posterior corners. Coxae I–IV each with blunt external spur, smaller on IV. Ventral coxal surfaces with scattered, moderately short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 39C).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli straight or slightly concave, cornua distinct, divergent, bluntly rounded (Fig. 40G). Auriculae formed as mild, lateral bulges (Fig. 40I). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded, dentition 3/3 in distal portion, then 2/2 nearly to base; lateral and inner denticles similar in size, inner ones less pointed (Fig. 40H). Two pairs of post- hypostomal setae. Palpi moderately short, slightly club-like, but lateral margin profile nearly straight in dorsal view (Fig. 40G). Palpal tibiotarsus not visible dorsally. Palpal trochanter with well-developed, tapered ventrolateral flange (Fig. 40I). Scutum: Length about 1.2 times greater than width, shape oval, widest at midlength, with posterolateral margins nearly straight (Fig. 40G). Scapulae weakly formed, rounded. Lateral carinae

Map 16. Collection localities for Ixodes ochotonae in Canada. 150 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A B C

D E

Figs. 39A–F. Ixodes ochotonae. Fig. 39A, female gnathosoma and scutum, dorsal; Fig. 39B, female hypostome, ventral; Fig. 39C, female gnathosoma, coxae and trochanters, ventral; Fig. 39D, male gnathosoma and idiosoma, dorsal; Fig. 39E, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 39F, male hypostome, ventral. Redrawn from Cooley and Kohls (1945). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 151

G H I J

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Figs. 40G–L. Ixodes ochotonae. Fig. 40G, nymph gnathosoma and scutum, dorsal; Fig. 40H, nymph hypostome, ventral; Fig. 40I, nymph gnathosoma, coxae and trochanters, ventral; Fig. 40J, larva hypostome, palp trochanter, basis capituli, ventral; Fig. 40K, larva, dorsal; Fig. 40L, larva, ventral. Figs. 40G–I redrawn from Cooley and Kohls (1945); Figs. 40J–L redrawn from Webb et al. (1990). 152 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins distinct, extending nearly to posterolateral margins. Cervical grooves faint, divergent, not reaching posterolateral margins. Surface sparsely, unevenly punctate. Legs: Coxa I with internal spur short, tapered, about as long as external spur (Fig. 40I). Coxae II–IV lacking internal spurs. Coxae I–IV each with small, bluntly pointed external spur, these similar in size or more rounded on IV. Ventral coxal surfaces with scattered, moderately short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 40I).

Larva. Body: Body length 0.7–0.8 mm. Gnathosoma: Dorsal posterior margin slightly sinuous, with cornua distinct, blunt, directed laterally or posterolaterally (Fig. 40K). Ventrally, auriculae undeveloped (Fig. 40L). Apex of hypostome broadly rounded, dentition below coronal area 2/2 nearly to base; lateral file with approximately 7 bluntly pointed teeth, inner file with 5 smaller, more rounded teeth (Fig. 40J). Two pairs of post-hypostomal setae (Fig. 40J). Palpi moderately slender; femur+genu length to width ratio about 2.0–2.4:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, with 2 conspicuous flanges, anterior flange shorter, more bluntly pointed than narrowly rounded posterior flange (Fig. 40J). Scutum: Length about 0.85 width, widest at anterior third (Fig. 40K). Scapulae weakly formed; shield posterior margin convex. Surface densely micropunctate, and with underlying reticula emergent anterolaterally. Setae, usually 5, rarely 6, pairs. Idiosomal setation (Figs. 40K, L): Central dorsal setae, 2 pairs. Supplemental dorsal setae absent. Marginal dorsal setae, 7–9 pairs. Marginal ventral setae, 3 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae slightly larger (approximately 1.5 times larger) than that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in size. Legs: Coxa I with bluntly right-triangular internal spur (Fig. 40L). Coxae II and III each with curved ridge posteroventrally. Coxae I–III lacking external spurs. Coxa I–II–III setal formula 3–2 or 3–2; coxal setae inconspicuously or unilaterally barbed. Femur III with 9 setae. Trochanters I–III lacking spurs (Fig. 40L).

Natural history. There are no published accounts on the natural history of this species. Ixodes ochotonae is not known to be the vector of any pathogenic organisms or parasites.

Distribution. Ixodes ochotonae is largely confined to the mountainous areas of western North America from British Columbia, Washington, Idaho, and Montana south to Wyoming and California, though there are records from South Dakota and Val Marie, Saskatchewan.

Hosts. The primary hosts for this species appear to be pikas and woodrats, though associated small mammals and predators are occasionally reported as hosts.

Ixodes (Ixodiopsis) soricis Gregson Shrew tick (Fig. 41, Map 17) Ixodes soricis Gregson, 1942b: 137 (female, nymph, larva). Ixodes soricis Gregson: Gregson and Kohls 1952: 185 (male). Ixodes (Pholeoixodes) soricis Gregson: Clifford et al. 1973: 498. Ixodes (Ixodiopsis) soricis Gregson: Robbins and Keirans 1992: 51.

Adult. Female: Body: Length 1.2–1.6 mm unfed, reaching to nearly 4.0 mm engorged. Gnathosoma: Porose areas of basis capituli suboval, often poorly defined, separated by A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 153

area of nearly or slightly less than diameter of one porose area (Fig. 41A). Dorsal posterior margin of basis capituli nearly straight or slightly concave, cornua little or no more than salient corners of basis capituli (Fig. 41A). Auriculae absent (Fig. 41E). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome pointed, dentition 3/3 throughout length; all denticles sharp, lateral ones only slightly larger than inner ones (Fig. 41B). Palpi moderately long, lateral margin profile slightly convex (Fig. 41A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with a small ventrolateral projection. Scutum: Length 1.3–1.4 longer than wide, widest slightly anterior to midlength, with posterolateral margins convex (Fig. 41A). Scapulae short, pointed. Lateral carinae distinct, extending from scapulae nearly to posterolateral margins. Cervical grooves faint or indiscernible. Surface with large, shallow puncta on posterior third, changing to furrows near posterior margin. Legs: Coxa I with short, bluntly triangular internal spur and slightly smaller, blunt external spur (Fig. 41E). Coxae II–IV lacking internal spurs. Coxae II–IV each with a small, rounded or bluntly pointed external spur. Ventral coxal surfaces with numerous long, prominently branched setae. Trochanters I–IV lacking spurs (Fig. 41E).

Adult. Male: Body: Length 0.9–1.0 mm. Gnathosoma: Basis capituli dorsal posterior margin straight or slightly irregular, with rounded corners instead of cornua; dorsal surface without furrows (Fig. 41G). Ventrally, anterior margin of basis capituli sloped, slightly humped on either side of hypostome. Apex of hypostome rounded, with slight notch, dentition roughly 3/3, with denticles arranged in approximately 4 transverse, partially overlapping rows of crenulations (Fig. 41C). Palpi short, bulbous in dorsal view, with symmetrically rounded apices (Fig. 41G). Dorsally, palpal femur faintly delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter lacking flanges. Scutum: Scapulae bluntly rounded (Fig. 41G). Lateral carinae absent. Surface slightly rugose anterolaterally, with faint anterior puncta grading into numerous large, shallow puncta posteriorly and marginally. Surface lacking pair of pit-like depressions midlaterally. Spiracular plate: Subcircular, without distinctively longer axis. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median plate about 1.5 longer than anal plate (Fig. 41H). Anal plate slightly widened posteriorly, where slightly wider than adanal plates. Ventral plates faintly punctate. Legs: Coxa I with small, pointed internal spur slightly longer than blunt external spur (Fig. 41H). Coxae II–IV lacking internal spurs, but II and III with salient posterior corners. Coxae I–IV each with a small, blunt external spur of similar size. Ventral coxal surfaces with long, prominently branched setae. Trochanters I–IV lacking spurs (Fig. 41H).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli nearly straight, with posterolateral corners extended into small, bluntly pointed cornua (Fig. 41I). Auriculae absent (Fig. 41F). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded, dentition 2/2, until 1/1 near base; lateral denticles more pointed and larger than inner denticles (Fig. 41D). Two pairs of post-hypostomal setae. Palpi short, thick, club-like in dorsal view (Fig. 41I). Palpal tibiotarsus not visible dorsally. Palpal trochanter with both anterior and posterior flange-like projections well developed, bluntly pointed (Fig. 41F). Scutum: Length 1.3–1.4 times greater than width; shape oval, widest near midlength, with posterolateral margins slightly convex (Fig. 41I). Scapulae short, blunt. Lateral carinae absent. Cervical grooves faint, slightly divergent, well short of reaching posterolateral margins. Surface nearly impunctate. Legs: Coxa I with short, bluntly triangular internal spur similar in size and form to external spur (Fig. 41F). Coxae 154 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

B C D

A E

F I

G H

K L

Figs. 41A–L. Ixodes soricis. Fig. 41A, female gnathosoma and scutum, dorsal; Fig. 41B, female hypostome, ventral; Fig. 41C, male hypostome, ventral; Fig. 41D, nymph hypostome, ventral; Fig. 41E, female gnathosoma, coxae and trochanters, ventral; Fig. 41F, nymph gnathosoma, coxae and trochanters, ventral; Fig. 41G, male gnathosoma and idiosoma, dorsal; Fig. 41H, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 41I, nymph gnathosoma and scutum, dorsal; Fig. 41J, larva hypostome, palp trochanter, basis capituli, ventral; Fig. 41K, larva, dorsal; Fig. 41L, larva, ventral; insert, coxa I. Figs. 41A, B, D–F, I redrawn after Cooley and Kohls (1945); Figs. 41C, G, H redrawn after Gregson and Kohls (1952); Figs. 41J, K, L redrawn after Webb et al. (1990). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 155

II–IV lacking internal spurs. Coxae I–IV each with bluntly pointed external spur, these similar in size or shorter from I–IV. Ventral coxal surfaces with numerous long, prominently branched setae (as in Fig. 41L, insert). Trochanters I–IV lacking spurs (Fig. 41F).

Larva. Body: Body length, unengorged, excluding gnathosoma, 0.45–0.65 mm. Gnathosoma: Dorsal posterior margin nearly straight or slightly sinuous, with divergent, bluntly pointed corners forming small cornua (Fig. 41K). Ventrally, auriculae undeveloped (Fig. 41L). Apex of hypostome rounded, dentition below coronal area 2/2 throughout; lateral file with 6–8 bluntly pointed, overlapping teeth, inner file with 4–5 smaller, more rounded teeth (Fig. 41J). Two pairs of post-hypostomal setae (Fig. 41J). Palpi thick; femur+genu length to width ratio about 1.8–2.0:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter with 2 prominent flanges, a bluntly pointed or truncated anterior flange and a somewhat more rounded posteroventral flange (Fig. 41J). Scutum: Length about 0.84–0.92 width, widest slightly anterior to mid-level (Fig. 41K). Scapulae weakly formed, rounded; shield posterior margin convex. Surface densely micropunctate, with underlying reticula emergent anterolaterally. Setae, 5 pairs. Idiosomal setation (Figs. 41K, L): Central dorsal setae, 2 pairs. Supplemental dorsal setae, absent. Marginal dorsal setae, 8 pairs. Marginal ventral setae, usually 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae less than twice that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in small size. Legs: Coxa I with bluntly right-triangular internal spur (Fig. 41L, insert). Coxae II and III each with a curved ridge posteroventrally. Coxae I–III lacking external spurs. Coxae I–II–III setal formula 3–2 or 3–2; some coxal setae conspicuously unilaterally barbed or usually setulose (Fig. 41L, insert). Femur III with 9 setae. Trochanters I–III lacking spurs (Fig. 41L).

Natural history. This tiny Nearctic tick is found almost exclusively on shrews. It has been reported to feed on northern pocket gophers (Thomomys talpoides Richardson) in Washington (Gregson and Kohls 1952) and humans in British Columbia (Spencer 1963), but these appear to be unusual records. In Oregon, larvae were collected from March to

Map 17. Collection localities for Ixodes soricis in Canada. 156 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

August, while nymphs and females were found during September to April (Easton and Goulding 1974). Males were not collected by Easton and Goulding (1974) and may be confined largely to nests of shrews. Ixodes soricis is probably a three-host tick. There is no information on its role as a vector of pathogenic organisms.

Distribution. Ixodes soricis is recorded from western North America from southern British Columbia to California, and from Utah, Colorado, Arizona and New Mexico.

Hosts. This tick is a parasite of shrews, but it occasionally infests other insectivores.

Ixodes (Multidentatus) auritulus auritulus Neumann (Figs. 42–43, Map 18) Ixodes thoracicus Neumann, 1899: 149 (not Ixodes thoracicus Koch, 1844). Ixodes auritulus Neumann, 1904: 450 (nomen novem for I. thoracicus Neumann, 1899). Ixodes auritulus Neumann: Nuttall 1916: 314 (nymph, larva). Ixodes auritulus auritulus Neumann: Kohls and Clifford 1966: 815 (male). Ixodes (Multidentatus) auritulus Neumann: Clifford et al. 1973: 497.

Adult. Female: Body: Length about 3.0 mm unfed, reaching to 10.0 mm engorged. Gnathosoma: Porose areas of basis capituli very large, oblong, without definite edges, narrowly separated medially (Fig. 42A). Dorsal posterior margin of basis capituli straight, salient (Fig. 42A). Cornua large, bluntly pointed (Fig. 42A). Auriculae formed as large, ventrolateral hornlike projections (Fig. 42C). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome wide, broadly rounded, dentition 5/5 subapically, then 4/4 for most of length, and 3/3 at base, denticles fine, pointed, those in lateral files slightly larger (Fig. 42B). Palpi somewhat thickened, club-shaped when viewed dorsally (Fig. 42A). Palpal tibiotarsus not visible dorsally. Palpal trochanter large, triangular dorsally, with a horn-like projection anteriorly (Figs. 42A, C). Scutum: Length about 1.2–1.3 times greater than width, suboval, widest at or slightly anterior to middle, with posterolateral margins slightly convex (Fig. 42A). Scapulae scarcely formed as salient anterolateral corners of scutum. Lateral carinae absent. Cervical grooves distinct, diverging and fading before reaching posterolateral margins. Surface sparsely, finely punctate.Legs: Coxa I with internal spur short but broad, similar in size and bluntly triangular form to external spur (Fig. 42C). Coxae II and III with salient posterior corners instead of internal spurs, these absent on IV. Coxae I–IV each with bluntly triangular external spur, sometimes progressively smaller from I–IV. Ventral coxal surfaces with scattered, smooth or slightly barbed setae. Trochanters I–IV each with distinct, bluntly pointed posteroventral spur (Fig. 42C).

Adult. Male: Body: Length, approximately 2.7 mm. Gnathosoma: Basis capituli dorsal posterior margin nearly straight, with blunt corners instead of cornua (Fig. 42D); dorsal face without furrows. Ventrally, anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome narrowly rounded, slightly indented, dentition 3/3 on anterior third, increasing to 6/6 or 7/7 at mid-length, then gradually decreasing basally; denticles flattened, blunt (Fig. 42F). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 42D). Dorsally, palpal femur and genu fused. Palpal tibiotarsus not visible dorsally. Palpal trochanter lacking flanges. Scutum: Scapulae short. Lateral carinae absent. Surface nearly smooth, with 2 pit-like depressions dorsolaterally (Fig. 42D, sp), otherwise inconspicuously punctate. Spiracular plate: Subcircular, with slightly longer axis A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 157

anteroposterior. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median plate about twice as long as anal plate (Fig. 42E). Anal plate not widened posteriorly, similar in width to adanal plates. Ventral plates nearly smooth, inconspicuously punctate. Legs: Coxa I with internal spur short, blunt, similar in form and size to external spur (Fig. 42E). Coxae II–IV with salient posterior corners instead of internal spurs, and each with a short, bluntly pointed external spur, these similar in size. Ventral coxal surfaces with scattered, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 42E).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli straight, cornua large, divergent, bluntly pointed (Fig. 43G). Auriculae formed as large, lateral, posteriorly- directed hornlike projections (Fig. 43I). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome broadly rounded, dentition 3/3 for most of length, then 2/2 basally, denticles fine, those in lateral files slightly larger and more pointed (Fig. 43H). Two pairs of post-hypostomal setae. Palpi somewhat thickened, club-shaped when viewed dorsally (Fig. 43G). Palpal tibiotarsus not visible dorsally. Palpal trochanter with a large, bluntly-pointed flange anteriorly (Fig. 43I). Scutum: Length slightly greater than width (Fig. 43G). Shape oval, widest at midlength, with posterolateral margins slightly convex. Scapulae reduced as weakly formed anterolateral corners of scutum. Lateral carinae absent. Cervical grooves diverging and fading before reaching posterolateral margins. Surface sparsely, finely punctate. Legs: Coxa I with internal spur short, broad, similar in size and bluntly triangular form to external spur (Fig. 43I). Coxae II and III with salient posterior corners instead of internal spurs, these absent on IV. Coxae I–IV each with bluntly triangular external spur, sometimes progressively smaller from I–IV. Ventral coxal surfaces with scattered, smooth or slightly barbed setae. Trochanters I–IV each with distinct, bluntly pointed posteroventral spur (Fig. 43I).

Larva. Body: Length of body 0.7–0.8 mm. Gnathosoma: Dorsal posterior margin of basis capituli straight, with cornua divergent, bluntly pointed (Fig. 43J). Ventrally, auriculae prominent triangular, lateral projections (Fig. 43L). Apex of hypostome broadly rounded, dentition 3/3 along apical third, then 2/2 nearly to base; lateral file with 8-10 bluntly pointed teeth, inner files with 6–8 smaller, blunter teeth (Fig. 43L). One or usually 2 pairs of post-hypostomal setae (Fig. 43L). Palpi moderately thick, femur+genu length to width ratio about 2.0:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, with 1 bluntly pointed or narrowly rounded, anterior flange (Fig. 43L). Scutum: Length about 0.8 times (0.75–0.85 times) width, widest at mid-level (Fig. 43J). Scapulae reduced; shield posterior margin flat or slightly convex. Surface densely, uniformly micropunctate, with underlying or emergent reticula, clearly along anterolateral margins. Setae, 5 pairs. Idiosomal setation (Figs. 43J–K): Central dorsal setae, 3–6 pairs. Supplementary dorsal setae, 2 or 3 pairs. Marginal dorsal setae, 8 pairs. Marginal ventral setae, 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 or 3 pairs, plus 2 or 3 pairs in anal region. Length of anterior marginal dorsal setae up to 3 times that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in small size. Legs: Coxa I with small, bluntly right-triangular internal spur (Fig. 43K). Coxae II and III each with curved ridge posteroventrally. Coxae I–III each with conspicuous, pointed external spur. Coxae I–II–III setal formula 3–2 or 3–2 or 3; coxal setae smooth. Femur III with 9 or 10 setae, a sixth seta sometimes present subapically. Trochanters I and II each with small blunt spur, III lacking spur (Fig. 43K). 158 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A B C

F sp

D E

Figs. 42A–F. Ixodes auritulus. Fig. 42A, female gnathosoma and scutum, dorsal; Fig. 42B, female hypostome, ventral; Fig. 42C, female gnathosoma, coxae and trochanters, ventral; Fig. 42D, male gnathosoma and idiosoma, dorsal; sp – dorsal pit; Fig. 42E, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 42F, male hypostome, ventral. Figs. 42A–C redrawn from Cooley and Kohls (1945); Figs. 42D modified, E–F redrawn from Kohls and Clifford (1966). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 159

G H I

J K

Figs. 43G–L. Ixodes auritulus. Fig. 43G, nymph gnathosoma and scutum, dorsal; Fig. 43H, nymph hypostome, ventral; Fig. 43I, nymph gnathosoma, coxae and trochanters, ventral; Fig. 43J, larva, dorsal; Fig. 43K, larva, ventral; Fig. 43L, larva hypostome, palp trochanter, basis capituli, ventral. Figs. 43G–I redrawn from Cooley and Kohls (1945); Figs. 43J–L redrawn from Webb et al. (1990). 160 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Map 18. Collection localities for Ixodes auritulus in Canada.

Natural history. There are relatively few records of I. auritulus from birds in Canada, though Gregson (1956) maintained it was a common species parasitising grouse on the coast of British Columbia. Scott et al. (2016) recorded this species from sooty grouse (Dendrolagapus fuliginosus (Ridway), Galliformes: Phasianidae) and California quail (Callipepla californica (Shaw), Galliformes: Odontophoridae) on Vancouver Island. Morshed et al. (2005) reported larvae, nymphs, and females from 10 species of passerine birds in British Columbia during the course of their study from June to October. Throughout its range, it has been collected from many avian host species. For example, González- Acuña et al. (2005) reported 51 species of birds in the neotropics to be infested with I. auritulus. There is little information on the seasonal activity of I. auritulus. Eisen et al. (2006) collected numerous specimens by dragging vegetation in a northern coastal area of California, but they were unable to resolve relationships in its life history. Some specimens of I. auritulus were found positive for Borrelia burgdorferi in British Columbia (Morshed et al. 2005).

Distribution. Ixodes auritulus auritulus is found in coastal areas of western North America, Florida, and also into Mexico and Central and South America and Africa. Different subspecies are described from Australia and New Zealand. Nearly all records in Canada are from coastal regions of British Columbia.

Hosts. Birds are the primary hosts for this tick. In North America, it has been found on a variety of ground-foraging birds, while in other parts of the world, sea birds are primary hosts.

Ixodes (Pholeoixodes) baergi Cooley and Kohls (Fig. 44) Ixodes baergi Cooley and Kohls, 1942: 1869 (female, male). Ixodes (Pholeoixodes) baergi Cooley and Kohls: Clifford et al. 1973: 498. Ixodes (Pholeoixodes) baergi Cooley and Kohls: Keirans et al. 1993: 735 (nymph, larva).

Adult. Female: Body: Length about 7.5 mm engorged (unfed size not indicated). Gnathosoma: Porose areas of basis capituli large, nearly circular, depressed, reaching A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 161

almost to posterior margin, separated by slightly less than diameter of one porose area (Fig. 44A). Dorsal posterior margin of basis capituli nearly straight or slightly convex, cornua absent (Fig. 44A). Auriculae indicated by faint lateral swellings (Fig. 44C). Anterior margin of basis capituli slightly humped on either side of hypostome. Hypostome short, with rounded apex, dentition 4/4 subapically, then 3/3 along middle third of length, then 2/2 nearly to base (Fig. 44B). Palpi short, club-like when viewed dorsally (Fig. 44A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with slight ventral projection. Scutum: Length 1.1–1.2 times greater than width, widest anterior to middle, with nearly straight posterolateral margins (Fig. 44A). Scapulae bluntly pointed. Lateral carinae faint or absent. Cervical grooves broad, nearly parallel. Surface nearly impunctate, but with numerous longitudinal ridges or wrinkles, especially in lateral areas. Legs: Coxae I–IV lacking any spurs (Fig. 44C). Ventral coxal surfaces with few small, smooth setae. Trochanters I–IV lacking spurs (Fig. 44C).

Adult. Male: Body: Length approximately 2.4 mm. Gnathosoma: Basis capituli dorsal posterior margin straight or slightly concave, with rounded posterior corners instead of cornua (Fig. 44E); dorsal surface with transverse ridge. Ventrally, anterior margin of basis capituli sloped, slightly humped on either side of hypostome. Apex of hypostome broad, cleft, dentition roughly 3/3, teeth of inner rows somewhat crenulate, smaller than more pointed lateral ones (Fig. 44D). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 44E). Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter lacking flanges.Scutum: Scapulae prominent, bluntly pointed (Fig. 44E). Lateral carinae absent. Surface with numerous, fine puncta, sometimes slightly larger near lateral margins; lacking pair of pit-like depressions midlaterally. Spiracular plate: Subcircular, without distinctively longer axis. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median plate nearly 1.3 times longer than anal plate (Fig. 44F). Anal plate not widened posteriorly, similar in width to adanal plates. Ventral plates finely punctate. Legs: Coxae I–IV lacking any spurs (Fig. 44F). Ventral coxal surfaces with scattered, short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 44F).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli concave, its rounded posterolateral corners hardly formed as cornua (Fig. 44G). Auriculae indicated by faint lateral ridges or swellings. Anterior margin of basis capituli sloped on either side of hypostome. Hypostome short, bluntly rounded apically. Hypostome dentition 3/3 on apical third, then 2/2 nearly to base; lateral denticles more pointed and slightly larger than inner denticles. Two pairs of post-hypostomal setae. Palpi short, club-like when viewed dorsally (Fig. 44G). Palpal tibiotarsus not visible dorsally. Palpal trochanter with slight ventral projection. Scutum: Length subequal to width; shape subtriangular, widest slightly anterior to middle, with nearly straight posterolateral margins, scapulae bluntly pointed (Fig. 44H). Lateral carinae faint or absent. Cervical grooves slightly divergent, reaching nearly to posterolateral margins. Surface nearly impunctate, but finely reticulate. Legs: Coxae I–IV lacking any spurs (Fig. 44I). Ventral coxal surfaces with few small, smooth or slightly barbed setae. Trochanters I–IV lacking spurs.

Larva. Body: Length from scapular apices to posterior body margin 0.57–0.63 mm. Gnathosoma: Dorsal posterior margin slightly concave, with rounded lateral corners instead of cornua (Fig. 44J). Ventrally, auriculae undeveloped or weakly formed as 162 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

B C E

I J

F H

K L M

Figs. 44A–M. Ixodes baergi. Fig. 44A, female gnathosoma and scutum, dorsal; Fig. 44B, female hypostome, ventral; Fig. 44C, female gnathosoma, coxae and trochanters, ventral; Fig. 44D, male hypostome, ventral; Fig. 44E, male gnathosoma and idiosoma, dorsal; Fig. 44F, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 44G, nymph gnathosoma, dorsal; Fig. 44H, nymph scutum; Fig. 44I, nymph coxae, ventral; Fig. 44J, larva gnathosoma and scutum; Fig. 44K, larva hypostome, palp trochanter, basis capituli, ventral; Fig. 44L, larva gnathosoma, dorsal; Fig. 44M, larva gnathosoma and coxae, ventral. Figs. 44A–F redrawn from Cooley and Kohls (1945); Figs. 44G–M, redrawn from Keirans et al. (1993). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 163

transverse ridges midlaterally (Fig. 44M). Apex of hypostome broadly rounded, dentition 3/3 subapically, then 2/2 nearly to base; lateral file with approximately 5 bluntly pointed, overlapping teeth, inner file with 4–5 smaller, blunter teeth (Fig. 44K). Two pairs of post-hypostomal setae (Fig. 44M). Palpi moderately thick, femur+genu length to width ratio approximately 2 (2.0–2.3):1 (Fig. 44L); ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, lacking flanges, but with a small ventral ridge. Scutum: Length slightly shorter (about 0.91– 0.95 times) than width, widest at anterior third, scapulae weakly developed; shield posterior margin convex (Fig. 44J). Surface densely micropunctate, with underlying reticula becoming emergent anterolaterally. Setae, 5 pairs. Idiosomal setation: Central dorsal setae, 2 pairs. Supplementary dorsal setae absent. Marginal dorsal setae, 7 or 8 pairs. Marginal ventral setae, 3 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae at most 1.5 that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in size. Legs: Coxae I–III lacking spurs, II with weak curved ridge posteroventrally (Fig. 44M). Coxa I–II–III setal formula 3–2–2; coxal setae short, smooth or inconspicuously barbed. Femur III with 9 setae. Trochanters I–III lacking spurs.

Natural history. This highly specialised species is found in the nests of the colonial cliff swallow (Petrochelidon pyrrhonota Vieillot) where its life cycle is closely synchronised with that of its host. When the swallows return in the spring, larvae become active and feed on the adult birds. Engorged larvae leave their hosts and moult to the nymphal instar, which may feed on adult birds incubating eggs, or on the first chicks to hatch (Hopla and Loye 1983; George 1987). Engorged nymphs leave their hosts and moult in crevices on the outsides of the nests. Adult females feed on the maturing nestlings, attaching primarily to the head and shoulders of the birds (George 1987; Larimore1987), detaching from their hosts mostly at night (George 1987). Males have not been recorded as feeding and it is assumed they do not require a blood meal. Males mate with engorged females as the latter feed, or after they have detached and moved away from the nests. Pairs may be observed as the females move into cracks and crevices to lay their eggs (Baerg 1944). After about two weeks, females may lay several hundreds of eggs (Baerg 1944; Hopla and Loye 1983; Larimore 1987) in concentrated masses in crevices on the outsides of the nest or below the nests (George 1987). Eggs hatch about 40 days later and larvae form dense aggregations; although they are active, they do not attempt to leave the aggregation, and they remain in these sites until the following spring when the birds return to the colony (Larimore 1987). Hopla and Loye (1983), Loye and Hopla (1983), and Hopla (1992) isolated a bunyavirus from I. baergi in cliff swallow colonies in Oklahoma, with evidence of transovarial transmission.

Distribution. Ixodes baergi is found mainly in the midwest of the United States of America. There are no known records of this tick being established in Canada, though its primary host, the cliff swallow breeds extensively here and T.D.G. (personal observation) has examined hundreds of nests during winter in Manitoba. One female was recorded from a cliff swallow collected in Ontario (Scott et al. 2001).

Hosts. This tick is a parasite of the cliff swallow, Petrochelidon pyrrhonota. 164 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Ixodes (Pholeoixodes) banksi Bishopp (Fig. 45, Map 19) Ixodes banksi Bishopp, 1911: 200. Ixodes banksi Bishopp: Cooley and Kohls 1945: 167 (nymph). Ixodes banksi Bishopp: Hays and Lawrence 1957: 213 (male, larva). Ixodes (Pholeoixodes) banksi Bishopp: Clifford et al. 1973: 498.

Adult. Female: Body: Length about 2.5 mm unfed; engorged size not indicated. Gnathosoma: Porose areas of basis capituli subtriangular, depressed, separated by less than width of one porose area (Fig. 45B). Dorsal posterior margin of basis capituli slightly sinuous or convex, with cornua little or no more than flared corners (Fig. 45B). Auriculae formed as mild lateral shelves (Fig. 45C). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome flattened, dentition 3/3 for a couple of subapical rows, then 2/2 to base; lateral denticles larger than medial denticles (Fig. 45E). Palpi moderately long, broadest at middle, lateral margin profile slightly convex, their inner margins more curved (Fig. 45B). Palpal tibiotarsus not visible dorsally. Palpal trochanter with a slight ventrolateral projection. Scutum: Length slightly (1.1 times) greater than width. Shape subtriangular, widest slightly anterior to middle, with nearly straight posterolateral margins, scapulae small, bluntly pointed (Fig. 45B). Lateral carinae moderate, terminating near posterolateral margins. Cervical grooves faint, slightly divergent. Surface with numerous, large, evenly distributed puncta. Legs: Coxa I with internal spur tapered, pointed, much longer than external spur (Fig. 45C). Coxae II–IV lacking internal spurs. Coxae I–IV each with blunt external spur, all similar in size. Ventral coxal surfaces with numerous, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 45C).

Adult. Male: Body: Length approximately 2.5–3.2 mm. Gnathosoma: Basis capituli dorsal posterior margin nearly straight, with rounded posterior corners instead of cornua; dorsal face without furrows. Ventrally, anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded or weakly notched, dentition mostly 3/3, arranged in approximately 6 diagonal rows, with teeth of inner files rounded, smaller than more pointed lateral file (Fig. 45G). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 45A). Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter lacking flanges. Scutum: Scapulae bluntly rounded (Fig. 45D). Lateral carinae absent. Surface with numerous, small puncta, more densely near margins. Surface lacking pair of pit-like depressions midlaterally. Spiracular plate: Ovoid, slightly thicker anteriorly, with clearly longer axis anteroposterior. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median plate about 1.4 times longer than anal plate (Fig. 45A). Anal plate not widened posteriorly, similar in width to adanal plates posteriorly. Ventral plates with numerous fine puncta. Legs: Coxa I with internal spur tapered, pointed, much longer than external spur (Fig. 45A). Coxae II–IV with salient posterior corners, sometimes formed into small, internal spurs. Coxae I–IV each with bluntly pointed external spur, reduced on IV. Ventral coxal surfaces with scattered, moderately long, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 45A).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli nearly straight, its bluntly pointed posterolateral corners not formed into cornua (Fig. 45D). Auriculae formed as faint lateral ridges (Fig. 45F). Anterior margin of basis capituli sloped on either side of A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 165

hypostome. Apex of hypostome rounded, dentition 2/2 below apical corona; lateral denticles larger and more pointed than inner ones (Fig. 45J). Two pairs of post-hypostomal setae. Palpi moderately short, club-like when viewed dorsally (Fig. 45D). Palpal tibiotarsus not visible dorsally. Palpal trochanter with small ventrolateral projection. Scutum: Length slightly less (0.9 times) than width. Shape subtriangular to broadly rhomboid, widest slightly anterior to middle, with nearly straight posterolateral margins (Fig. 45D). Scapulae small, bluntly pointed. Lateral carinae absent. Cervical grooves long, slightly divergent, not reaching posterolateral margins. Surface ornamentation not described. Legs: Coxa I with internal spur tapered, pointed, much longer than external spur (Fig. 45F). Coxae II–IV lacking internal spurs. Coxae I–IV each with short, blunt external spur, progressively smaller from I–IV. Ventral coxal surfaces setation undescribed. Trochanters I–IV lacking spurs (Fig. 45F).

Larva. Body: Body length, including gnathosoma, 0.66–0.72 mm. Gnathosoma: Dorsal posterior margin sinuous or nearly straight, with blunt lateral corners instead of cornua. Ventrally, auriculae undeveloped. Apex of hypostome broadly rounded or slightly indented, dentition below coronal area 2/2; lateral file with 5–6 blunt, overlapping teeth, inner file with 4–5 smaller, blunter teeth (Fig. 45H). Two pairs of post-hypostomal setae (Fig. 45H). Palpi moderately thick, femur+genu length to width ratio approximately 2 (2.0–2.3):1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, without flanges. Scutum: Length about 0.8 times (0.80–0.82 times) width, widest near mid-level, scapulae small to conspicuous, bluntly rounded; shield posterior margin convex (Fig. 45I). Surface finely micropunctate, and with underlying reticula over much of area, emergent anterolaterally. Setae, 5 pairs. Idiosomal setation (Fig. 45I): Central dorsal setae, 2 pairs. Supplementary dorsal setae absent. Marginal dorsal setae, 8 pairs. Marginal ventral setae, 4–6 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Soft cuticle with setae relatively long, but length of anterior marginal dorsal setae not conspicuously longer than that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in moderate length. Legs: Coxa I with small, rounded or somewhat larger, bluntly angular internal spur (Fig. 45I). Coxae II and sometimes III with curved ridge posteroventrally. Coxae I–III lacking external spurs. Coxal I–II–III setal formula 3–2 or 3–2; coxal setae moderately short, smooth or inconspicuously barbed. Femur III with 8 setae. Trochanters I–III lacking spurs (Fig. 45I).

Map 19. Collection localities for Ixodes banksi in Canada. 166 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

D E

F J

H I

Figs. 45A–J. Ixodes banksi. Fig. 45A, male, dorsal gnathosoma and idiosoma, left, ventral gnathosoma, idiosoma, coxae and trochanters, right; Fig. 45B, female gnathosoma and scutum, dorsal; Fig. 45C, female gnathosoma, coxae and trochanters, ventral; Fig. 45D, nymph gnathosoma and scutum, dorsal; Fig. 45E, female hypostome, ventral; Fig. 45F, nymph gnathosoma, coxae and trochanters, ventral; Fig. 45G, male gnathosoma, ventral; Fig. 45H, larva gnathosoma, ventral; Fig. 45I, larva, dorsal left, ventral right; Fig. 45J, nymph hypostome, ventral. Figs. 45A, G–H redrawn, I modified from Hays and Lawrence (1957); Figs. 45B–F, J redrawn from Cooley and Kohls (1945). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 167

Natural history. This species seems to be a parasite primarily of the beaver, though there are records for it infesting muskrat and it is known occasionally to bite humans (Durden and Keirans 1996). It inhabits the host lodges, where it can be found with greater frequency than what has been reported in the literature on their hosts (Lawrence et al. 1956). Records most often come from hosts which have been taken by trappers (Judd 1954; Lawrence et al. 1956), and the ticks are usually found deeply in the outer meatus of the ear (Lawrence et al. 1956; Kollars et al. 1995), though they may be found attached to other parts of the body. Ixodes banksi has been shown to transmit the bacteria that cause tularaemia, Francisella tularensis, to susceptible mice in the laboratory (Lawrence et al. 1956).

Distribution. Records for I. banksi are scattered through Nova Scotia, Ontario, and southeastern Manitoba and into the eastern United States of America as far south as Alabama. Undoubtedly, additional collections will occur in other eastern provinces.

Hosts. The beaver, Castor canadensis Kuhl, seems to be the primary host for this tick, despite the type host being muskrat, Ondatra zibethicus (Linnaeus); all records in Canada are from beaver.

Ixodes (Pholeoixodes) cookei Packard (Figs. 46–47, Map 20) Ixodes cookei Packard, 1869: 67. Ixodes cruciarius Fitch, 1872: 366. Ixodes hexagonus Neumann, 1899: 129. Ixodes hexagonus longispinosus Neumann, 1901: 283. Ixodes cookei Packard: Banks 1908: 2. Ixodes hexagonus cookei: Nuttall and Warburton 1911: 183. Ixodes (Pholeoixodes) cookei Packard: Clifford et al. 1973: 498.

Adult. Female: Body: Length about 2.6 mm unfed, reaching to 9.0 mm engorged. Gnathosoma: Porose areas of basis capituli subcircular to subtriangular, separated by area about the diameter of one porose area (Fig. 46A). Dorsal posterior margin of basis capituli nearly straight. Cornua short, bluntly rounded (Fig. 46A). Auriculae faintly formed as small lateral ridges or projections (Fig. 46B). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded, dentition 4/4 subapically, then 3/3 along apical third, then 2/2 along basal two-thirds (Fig. 46C). Palpi thick, club-like when viewed dorsally (Fig. 46A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with small, broadly rounded flange-like projection ventrolaterally. Scutum: Length about equal to, or slightly greater than, width. Shape oval or slightly subtriangular, widest slightly anteriad of middle, with nearly straight posterolateral margins, scapulae prominent, pointed (Fig. 46A). Lateral carinae present. Cervical grooves faint, diverging posteriorly nearly to posterolateral margins. Surface with moderately numerous, conspicuous puncta, lacking wrinkled or roughened areas. Legs: Coxa I with internal spur tapered, pointed, much longer than external spur (Fig. 46B). Coxae II–IV lacking internal spurs. Coxae I–IV each with small, rounded external spur, progressively smaller from I–IV. Ventral coxal surfaces with scattered, short, smooth or slightly barbed setae. Trochanters I–IV each with small, rounded posteroventral spur (shown only on trochanter III in Fig. 46B). 168 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Adult. Male: Body: Length 2.9–3.5 mm. Gnathosoma: Basis capituli dorsal posterior margin nearly straight, with blunt corners instead of cornua; dorsal face without furrows (Fig. 46D). Ventrally, anterior margin of basis capituli sloped, slightly humped on either side of hypostome (Fig. 46E). Apex of hypostome broadly rounded or slightly indented, dentition roughly 4/4, arranged in approximately 6 diagonal overlapping rows, teeth of inner rows somewhat crenulate, smaller than lateral row (Fig. 46G). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 47I). Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter lacking flanges. Scutum: Scapulae bluntly rounded (Fig. 46D). Lateral carinae absent. Surface with numerous, moderately large puncta, more densely punctate along margins. Surface lacking pair of pit-like depressions midlaterally. Spiracular plate: Subcircular or irregularly oval, without distinctively longer axis. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median plate slightly longer than anal plate (Fig. 46E). Anal plate not widened posteriorly, similar in width to adanal plates posteriorly. Ventral plates with numerous small puncta. Legs: Coxa I with internal spur tapered, pointed, much longer than external spur (Fig. 46E). Coxae II–IV lacking internal spurs, but II–III with salient posterior corners. Coxae I–IV each with small, bluntly pointed external spur, these similar in size. Ventral coxal surfaces with scattered, short, smooth or slightly barbed setae. Trochanters I–IV sometimes each with vestige of rounded posteroventral spur (shown only on trochanter III in Fig. 46E).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli sinuous, cornua prominent, bluntly pointed (Fig. 47J). Auriculae formed as lateral ridges (Fig. 47K). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome broadly rounded, dentition 3/3 subapically, then 2/2 to base (Fig. 46F). Two pairs of post-hypostomal setae. Palpi thick, club-like when viewed dorsally. Palpal tibiotarsus not visible dorsally. Palpal trochanter with short, narrowly rounded anterior flange and long, narrowly rounded posterior flange (Fig. 47K). Scutum: Length about 0.8 times width. Shape subtriangular, widest slightly anteriad of middle, with nearly straight posterolateral margins, scapulae prominent, bluntly pointed (Fig. 47J). Lateral carinae present. Cervical grooves distinct, divergent posteriorly nearly to posterolateral margins. Surface with sparse, small puncta over most of area, lacking wrinkled or roughened areas. Legs: Coxa I with internal spur tapered, pointed, longer than external spur (Fig. 47K). Coxae II–IV lacking internal spurs. Coxae I–IV each with small, bluntly pointed external spur, progressively slightly smaller from I–IV. Ventral coxal surfaces with scattered, short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 47K).

Larva. Body: Body length 0.6–0.7 mm unengorged, excluding gnathosoma. Gnathosoma: Dorsal posterior margin of basis capituli sinuous, with cornua weakly developed, bluntly pointed (Fig. 47L). Ventrally, auriculae faintly formed as diagnonal ridges midlaterally (Fig. 47M). Apex of hypostome broadly rounded or slightly indented, dentition 3/3 subapically, then 2/2 nearly to base; lateral file with 5–7 bluntly pointed, overlapping teeth, inner file with 4–5 smaller, blunter teeth. Two pairs of post-hypostomal setae (Fig. 47M). Palpi moderately thick; femur+genu length to width ratio about 2:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, with a small, blunt ridge anterodorsally and a moderately well-developed, rounded flange posteroventrally.Scutum: Length 0.8–0.9 times width, widest at anterior third, scapulae well developed, rounded apically; shield posterior margin convex (Fig. 47L). Surface densely, uniformly micropunctate, sometimes with underlying reticula, and lineate-reticulate along anterolateral margins. Setae, 4 or 5 pairs, most A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 169

anterolateral pair sometimes off scutum. Idiosomal setation (Figs. 47L–M): Central dorsal setae, 2 pairs. Supplementary dorsal setae absent. Marginal dorsal setae, 8 pairs. Marginal ventral setae, 3 or 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae about twice that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in size. Legs: Coxa I with large, bluntly subtriangular internal spur (Fig. 47M). Coxae II and III each with weakly developed, curved ridge posteroventrally. Coxae I–III lacking external spurs. Coxae I–II–III setal formula 3–2 or 3–2 (Fig. 47M errs in showing only one seta on coxa II); coxal setae smooth or inconspicuously barbed. Femur III with 8 or 9 setae. Trochanters I–III lacking spurs (Fig. 47M).

Natural history. In Canada, Ixodes cookei is primarily associated with the woodchuck, Marmota monax, where prevalence has been reported from 29–50% (McLean et al. 1966; Ko 1972a; Artsob et al. 1984; Farkas and Surgeoner 1990), though Martell et al. (1969) found three of seven porcupines infested in Nova Scotia. In southern parts of its range, medium- sized mammals, especially raccoons and skunks, are important hosts (e.g., Sonenshine 1979b; Kollars and Oliver 2003). All instars may be present in Ontario from April to August, with larvae appearing in the spring slightly later than nymphs and adults (Ko 1972a; Farkas and Surgeoner 1990). As is typical of nest-inhabiting Ixodes, males do not feed. Mating takes place in the host burrow or on the host. Larvae, nymphs, and females required averages of 6.4, 7.9, and 7.3 days to engorge and subsequently larvae and nymphs moulted to the next instar 19.8 and 18.1 days, respectively (Farkas and Surgeoner 1991). There is a positive correlation between engorgement weight and fecundity in females, with a maximum observed number of 2064 eggs by Farkas and Surgeoner (1991). Females may feed and partially engorge when not mated, but they lay fewer eggs which have not been found to hatch. At 29 °C, females began laying eggs 4–5 days after detachment, and continued to lay eggs for an average of about 20 days, most being laid during the first six days (Farkas and Surgeoner 1991). Eggs first hatched an average of 33 days after oviposition at 29 °C; no eggs hatched at 10 °C, and the threshold for development was estimated to be 11.2 °C (Farkas and Surgeoner 1991).

Map 20. Collection localities for Ixodes cookei in Canada. 170 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A B

F G

D E

Figs. 46A–H. Ixodes cookei. Fig. 46A, female gnathosoma and scutum, dorsal; insert, enlarged cornu; Fig. 46B, female gnathosoma, coxae and trochanters, ventral; Fig. 46C, female hypostome, ventral; Fig. 46D, male gnathosoma and idiosoma, dorsal; Fig. 46E, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 46F, nymph hypostome, ventral; Fig. 46G, male hypostome, ventral; Fig. 46H, male gnathosoma, ventral. Figures A-H redrawn, and B, E modified from Cooley and Kohls (1945). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 171

I K

L M

Figs. 47I–M. Ixodes cookei. Fig. 47I, male hypostome, ventral; Fig. 47J, nymph gnathosoma and scutum, dorsal; Fig. 47K, nymph gnathosoma, coxae and trochanters, ventral; Fig. 47L, larva, dorsal; Fig. 47M, larva, ventral. Figures I-K redrawn from Cooley and Kohls (1945). Figures L, M redrawn, M modified from Clifford et al. (1961). 172 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Unfed larvae lived up to 101 weeks at 15 °C, while unfed females survived for 18 months at room temperature at high humidity (Farkas and Surgeoner 1991). Ixodes cookei feeds predominantly on rodents and medium-sized carnivores, though it is known to bite humans as well. Therefore, it may be of importance to human health. Its role in transmission of Powassan virus is well known (Main et al. 1979), and as many as 23% of woodchucks in Ontario were shown to be seropositive (Artsob et al. 1984). This species has not proven to be a good vector of the bacterium that causes Lyme borreliosis (Barker et al. 1993), but there are indications that it may be a vector of other potential pathogens elsewhere in its range (Magnarelli and Swihart 1991). There are relatively few records of ixodid ticks which transmit nematodes, so it is interesting that I. cookei is the vector for a filarial nematode, Ackertia marmotae Webster (Nematoda: Onchocercidae), which parasitises the woodchuck (Ko 1972b).

Distribution. Ixodes cookei is widely distributed in eastern North America from eastern Canada to Texas and Florida. In Canada, it has been recorded in every province from Newfoundland to southeastern Manitoba.

Hosts. This tick is most often found on medium-sized mammals, including woodchuck, skunks, racoon, and porcupine. Carnivores, such as marten, fisher, fox, dogs, and cats are frequently infested, and humans may serve as hosts.

Ixodes (Pholeoixodes) hearlei Gregson (Figs. 48, 49, Map 21) Ixodes hearlei Gregson, 1941: 220 (female, male, nymph). Ixodes (Pholeoixodes) hearlei Gregson: Clifford et al. 1973: 498.

Adult. Female: Body: Length about 2.5 mm unfed, reaching to 6.0 mm engorged. Gnathosoma: Porose areas of basis capituli small, oval to circular, mildly depressed, separated by more than the width of one porose area (Fig. 48A). Dorsal posterior margin of basis capituli slightly concave, salient, cornua short, narrowly rounded (Fig. 48A). Auriculae weakly formed as bluntly triangular lateral lobes or indistinct (Fig. 48D). Anterior margin of basis capituli sloped on either side of hypostome. Apex of relatively small hypostome rounded, dentition 2/2 except for coronal area, with lateral files larger than inner files (Fig. 48B). Palpi moderately long, with lateral profile margin nearly straight in dorsal view, though widest at mid-length so as to appear somewhat club-like (Fig. 48A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with small, pointed ventrolateral projection. Scutum: Length about equal to, or slightly greater than, width. Shape subtriangular, widest slightly anterior to middle, with straight posterolateral margins (Fig. 48A). Lateral carinae absent or sometimes faintly visible. Cervical grooves distinct, starting near scapulae, fading before reaching posterolateral margins. Surface sparsely, coarsely punctate and irregularly mildly rugose or wrinkled along anterolateral margins. Legs: Coxa I with internal spur rounded, short, but larger than faint external spur (Fig. 48D). Coxae II–IV lacking internal spurs. Coxae I–IV with external spurs indicated only by faint bulges. Ventral coxal surfaces with scattered short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 48D).

Adult. Male: Body: Length approximately 2.5 mm. Gnathosoma: Basis capituli dorsal posterior margin straight, with blunt corners instead of cornua; dorsal face without furrows A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 173

(Fig. 48F). Ventrally, anterior margin of basis capituli sloped, slightly humped on either side of hypostome. Apex of hypostome rounded and slightly indented, dentition weak, arranged in approximately six transverse rows of mild, mostly 3/3 crenulations (Fig. 48C). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 48F). Dorsally, palpal femur and genu fused. Palpal tibiotarsus not visible dorsally. Palpal trochanter with a small, blunt ventral flange. Scutum: Scapulae bluntly rounded (Fig. 48F). Lateral carinae absent. Surface with conspicuous, large, shallow puncta and fewer small puncta. Surface lacking pair of pit-like depressions midlaterally. Spiracular plate: Subcircular, without distinctively longer axis. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median plate slightly longer than anal plate (Fig. 48G). Anal plate widened posteriorly, wider than adanal plates there. Median plate with fine, inconspicuous puncta; other ventral plates with mixed large and small puncta as on scutum. Legs: Coxa I with internal spur short, blunt, similar in size to external spur (Fig. 48G). Coxae II–IV lacking internal spurs. Coxae I–IV each with small, blunt external spur. Ventral coxal surfaces with scattered, short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 48G).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli straight, its bluntly rounded or truncated posterolateral corners not formed into cornua (Fig. 48H). Auriculae formed as bluntly triangular lateral lobes anterior to mid-level of basis capituli (Fig. 49I). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded, dentition 2/2 except for coronal area, with lateral files slightly larger and more pointed than inner files (Fig. 48E). Two pairs of post-hypostomal setae. Palpi thick, club-like when viewed dorsally. Palpal tibiotarsus not visible dorsally. Palpal trochanter with short but conspicuous, bluntly pointed ventrolateral flange. Scutum: Length subequal to width. Shape subtriangular, widest slightly anterior to middle, with nearly straight posterolateral margins (Fig. 48H). Scapulae short, rounded. Lateral carinae absent. Cervical grooves distinct, starting near scapulae, divergent and nearly reaching posterolateral margins. Surface sparsely, weakly punctate. Legs: Coxa I with internal spur short, rounded, but larger than vestigial external spur (Fig. 49I). Coxae II–IV lacking internal spurs. Coxae I–IV with external spurs indicated only by faint, blunt bulges. Ventral coxal surfaces with scattered short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 49I).

Larva. Body: Body length 0.5–0.6 mm. Gnathosoma: Dorsal posterior margin of basis capituli sinuous, with bluntly rounded or truncated corners instead of cornua. Ventrally, auriculae faintly formed as diagonal ridges midlaterally. Apex of hypostome nearly flat, slightly indented. Hypostome dentition below coronal area 2/2 nearly to base; lateral file with 5-7 bluntly pointed teeth, inner file with 4–5 smaller, more rounded teeth. Two pairs of post-hypostomal setae. Palpi thick, femur+genu length to width ratio about 1.7– 2.0:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, with a short, broadly to bluntly rounded posteroventral flange (Fig. 49L). Scutum: Length about 0.8 times (0.80–0.83 times) width, widest at anterior third (Fig. 49J); posterior margin flattened or slightly concave. Scapulae rounded; shield posterior margin flattened or slightly concave. Surface micropunctate, and fully reticulated. Setae, 5 pairs. Idiosomal setation: Central dorsal setae, 2 pairs. Supplemental dorsal setae absent. Marginal dorsal setae, 8 or usually 9 pairs. Marginal ventral setae, 3 or 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae less than twice that 174 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

B E

A C D

F G

Figs. 48A–H. Ixodes hearlei. Fig. 48A, female gnathosoma and scutum, dorsal; Fig. 48B, female hypostome, ventral; Fig. 48C, male hypostome, ventral; Fig. 48D, female gnathosoma, coxae and trochanters, ventral; Fig. 48E, nymph hypostome, ventral; Fig. 48F, male gnathosoma and idiosoma, dorsal; Fig. 48G, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 48H, nymph gnathosoma and scutum, dorsal. Redrawn from Cooley and Kohls (1945). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 175

I K

Figs. 49I–L. Ixodes hearlei. Fig. 49I, nymph gnathosoma, coxae and trochanters, ventral; Fig. 49J, larva, outline of scutum, dorsal; Fig. 49K, larva gnathosoma, coxae, ventral; Fig. 49L, larva gnathosoma, ventral. Figs. 49I, K modified from Gregson (1941); J, L are original. 176 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Map 21. Collection localities for Ixodes hearlei in Canada.

of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in size. Legs: Coxa I with small, broadly subtriangular internal spur (Fig. 49K). Coxae II and III with weakly defined, curved ridge posteroventrally. Coxae I and II lacking external spurs, but coxa III with external bulge or slight spur (Fig. 49K). Coxae I–II–III setal formula 3–2–2; coxal setae smooth or inconspicuously barbed. Femur III with 9 setae. Trochanters I–III lacking spurs.

Natural history. Other than host associations, there is no published information on the natural history of this species. There are no known associations of I. hearlei with pathogenic organisms. Foley et al. (2007) examined northern flying squirrels (Glaucomys sabrinus (Shaw)) in California for infections with Anaplasma phagocytophilum and Rickettsia prowazekii, in association with possible arthropod vectors. At the location where prevalence of A. phagocytophilum was 25%, one specimen of I. hearlei was collected, but this tick was not tested.

Distribution. The distribution of I. hearlei is limited to western North America, from British Columbia to Texas. In Canada, it has been recorded only from the interior of British Columbia.

Hosts. In Canada, the only host known is Streator’s tree squirrel, Tamiasciurus hudsonicus streatori (Allen), though it has also been reported from northern flying squirrels and raccoons in the United States of America.

Ixodes (Pholeoixodes) kingi Bishopp Rotund tick (Figs. 50, 51, Map 22) Ixodes pratti Banks, 1908: 27 (in part). Ixodes kingi Bishopp, 1911: 20l (female, male). Ixodes kingi Bishopp: Hooker et al. 1912: 82 (nymph, larva). Ixodes (Pholeoixodes) kingi Bishopp: Clifford et al. 1973: 498. Ixodes (Pholeoixodes) kingi Bishopp: Keirans and Clifford 1978: 81. . Adult. Female: Body: Length about 2.0 mm unfed, reaching to 16.0 mm engorged. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 177

Gnathosoma: Porose areas of basis capituli deep, nearly circular, separated by slightly more than the width of one porose area (Fig. 50A). Dorsal posterior margin of basis capituli nearly straight or slightly concave, cornua prominent, bluntly rounded (Fig. 50A). Auriculae absent or faintly formed as lateral ridges or angled projections (Fig. 50C). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded, dentition 3/3 for a couple of subapical rows, then 2/2 nearly to base, lateral denticles larger, more pointed than inner ones (Fig. 50B). Palpi thickened, club-shaped when viewed dorsally, widest at mid-length (Fig. 50A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with ventrolateral, flange-like projection.Scutum: Length slightly greater (about 1.1 times) than width. Shape subtriangular, widest slightly anterior to middle, with posterolateral margins nearly straight (Fig. 50A). Scapulae prominent, pointed. Lateral carinae distinct, parallel with anterolateral margins. Cervical grooves faint or absent. Surface covered with dense, coarse puncta, especially mesad of lateral carinae. Legs: Coxa I with internal spur tapered, bluntly pointed, longer than external spur (Fig. 50C). Coxa II and III each with somewhat salient posterior corner instead of internal spur, this absent on coxa IV. Coxae I–IV each with short, bluntly pointed external spur. Ventral coxal surfaces with few, short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 50C).

Adult. Male: Body: Length 2.3–2.6 mm. Gnathosoma: Basis capituli dorsal posterior margin straight, with cornua directed posterolaterally (Fig. 50D); dorsal face without furrows. Ventrally, anterior margin of basis capituli sloped, slightly humped on either side of hypostome. Apex of hypostome flattened or slightly indented, dentition 3/3 along apical third, then 2/2 to base, arranged in approximately 6 diagonal rows; denticles discrete, inner denticles slightly smaller, blunter (Fig. 50F). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 50D). Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter with small, blunt ventral flange.Scutum: Scapulae prominent, bluntly rounded (Fig. 50D). Lateral carinae distinct, emphasised by densely punctate depressions on their inner sides. Surface with puncta of intermingled sizes anteriorly, grading to fewer and smaller puncta posteriorly. Surface lacking pair of pit-like depressions midlaterally. Spiracular plate: Broadly oval, with slightly longer axis anteroposterior. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median plate about 1.5 longer than anal plate (Fig. 50E). Anal plate not widened posteriorly, similar in width to adanal plates. Ventral plates with scattered small puncta. Legs: Coxa I with internal spur tapered, bluntly pointed, longer than external spur (Fig. 50E). Coxae II–IV lacking internal spurs, but II and III with salient posterior corners. Coxae I–IV each with short, blunt external spur. Ventral coxal surfaces with few, short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 50E).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli nearly straight or slightly concave, cornua bluntly pointed (Fig. 51G). Auriculae formed as salient, angular lateral lobes (Fig. 51H). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded. Hypostome dentition 2/2 nearly to base. Lateral denticles larger than inner ones. Palpi short, thickened, club-shaped when viewed dorsally (Fig. 51G). Palpal tibiotarsus not visible dorsally. Palpal trochanter with a bluntly pointed, ventrolateral flange (Fig. 51H). Scutum: Length subequal to width. Shape subtriangular, widest slightly anterior to middle, with posterolateral margins straight or slightly concave (Fig. 51G). Scapulae bluntly pointed. Lateral carinae distinct, parallel with anterolateral margins. Cervical grooves faint, divergent, not approaching posterolateral margins. 178 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A C

D E

Figs. 50A–F. Ixodes kingi. Fig. 50A, female gnathosoma and scutum, dorsal; Fig. 50B, female hypostome, ventral; Fig. 50C, female gnathosoma, coxae and trochanters, ventral; Fig. 50D, male gnathosoma and idiosoma, dorsal; Fig. 50E, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 50F, male hypostome, ventral. Redrawn from Cooley and Kohls (1945). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 179

G H I

J K

Figs. 51G–K. Ixodes kingi. Fig. 51G, nymph gnathosoma and scutum, dorsal; Fig. 51H, nymph, gnathosoma, coxae and trochanters, ventral; Fig. 51I, larva hypostome, palp trochanter, basis capituli, ventral; Fig. 51J, larva, dorsal; Fig. 51K, larva, ventral. Figs. 51G, H redrawn from Cooley and Kohls (1945); Figs. 51I–K modified from Webb et al. (1990). 180 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Surface with scattered, sparse puncta. Legs: Coxa I with internal spur tapered, bluntly pointed, somewhat longer than external spur (Fig. 51H). Coxa II–IV lacking internal spurs. Coxae I–IV each with short, bluntly pointed external spur, progressively smaller from I–IV. Ventral coxal surfaces with few, short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 51H).

Larva. Body: Body length 0.7–0.8 mm. Gnathosoma: Dorsal posterior margin of basis capituli sinuous, with cornua formed as divergent, blunt, lateral projections (Fig. 51J). Ventrally, auriculae formed as diagonal ridges or lateral bulges. Apex of hypostome broadly rounded or slightly indented, dentition below coronal area 2/2 along nearly entire length; lateral file with 7–9 bluntly pointed, overlapping teeth, inner file with 6–8 smaller, blunter teeth (Fig. 51I). Two pairs of post-hypostomal setae (Figs. 51I, K). Palpi moderately thick, femur+genu length to width ratio about 2.0:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, with 1 moderately developed, narrowly rounded, posteroventral flange (Fig. 51I). Scutum: Length nearly (0.88–0.97 times) equal to width, widest at anterior third (Fig. 51J). Scapulae weakly formed, rounded; shield posterior margin convex. Surface finely micropunctate, with underlying reticula emergent anterolaterally. Setae, 5 pairs. Idiosomal setation (Figs. 51J, K): Central dorsal setae, 2 pairs. Supplemental dorsal setae absent. Marginal dorsal setae, 8 pairs. Marginal ventral setae, 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae about twice that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in size. Legs: Coxa I with short, bluntly triangular internal spur (Fig. 51K). Coxae II and III each with ridge posteroventrally (Fig. 51K). Coxae I–III lacking external spurs. Coxae I–II–III setal formula 3–2–2; coxal setae smooth or inconspicuously barbed. Femur III with 8 or 9 setae. Trochanters I–III lacking spurs (Fig. 51K).

Natural history. Although the distribution of I. kingi is reasonably well defined in Canada (Brown and Kohls 1950; Gregson 1971), there is little information on its natural history. While Brown collected 161 adults and juveniles in Alberta, dates of collection were not reported, and no comments were made about seasonal patterns of occurrence. Salkeld et al. (2006) found that I. kingi was present in Colorado when they began trapping

Map 22. Collection localities for Ixodes kingi in Canada. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 181

small mammals in May, mainly infesting thirteen-lined ground squirrels (Ictidomys tridecimlineatus (Mitchill)). Peak infestations with nymphs occurred in July and August. Anstead and Chilton (2014) and Anstead et al. (2014) found I. kingi infesting Richardson’s ground squirrel and northern pocket gopher in Saskatchewan. Ixodes kingi has not been identified as a primary vector for pathogens, though Francisella tularensis (Thorpe et al. 1965) and Coxiella burnetii (Sidwell et al.1965) have been isolated from this tick in Utah.

Distribution. Ixodes kingi is widely distributed in North America. It appears to be more common in western and central regions, though it has been recorded from Maryland. In Canada, records are limited to Manitoba and provinces to the west.

Hosts. A variety of hosts are infested by this tick, and in Canada, ground squirrels and their predators, including dogs and cats, are most often reported east of the Rocky Mountains. Gregson (1971) reported pocket gophers, kangaroo rats, and mice (Peromyscus) to be more common hosts west of the continental divide. See Salkeld et al. (2006) for a summary of hosts recorded in North America.

Ixodes (Pholeoixodes) marmotae Cooley and Kohls (Figs. 52–53, Map 23) Ixodes marmotae Cooley and Kohls, 1938b: 2174 (female, male, nymph). Ixodes marmotae Cooley and Kohls: Allred et al. 1960: 12 (larva). Ixodes (Pholeoixodes) marmotae Cooley and Kohls: Clifford et al. 1973: 498.

Adult. Female: Body: Length about 2.7 mm unfed. Gnathosoma: Porose areas of basis capituli subcircular, depressed, separated by about the width of one porose area (Fig. 52A). Dorsal posterior margin of basis capituli nearly straight or slightly sinuous, cornua large, rounded, or bluntly pointed (Fig. 52A). Auriculae absent or weakly formed as lateral projections (Fig. 52D). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded, dentition 4/4 in distal portion, then 3/3, changing to 2/2 at mid-length to base, denticles pointed, those in lateral files slightly larger (Fig. 52B). Palpi somewhat thickened, club-shaped in dorsal view, widest at mid-level (Fig. 52A). Palpal trochanter with rounded ventrolateral flange (Fig. 52D). Scutum: Length slightly greater than width, subtriangular, widest slightly anterior to middle, with posterolateral margins nearly straight but sometimes either slightly convex or concave (Fig. 52A). Scapulae moderate in size, pointed. Lateral carinae faint or absent. Cervical grooves usually distinct, reaching nearly to posterolateral margins. Surface with numerous puncta, more abundant in median area than laterally. Legs: Coxa I with internal spur moderately long, pointed, longer than external spur (Fig. 52D). Coxae II–IV lacking internal spurs. Coxae I–IV each with short, blunt external spur, sometimes progressively smaller from I–IV. Ventral coxal surfaces with few smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 52D).

Adult. Male: Body: Length 2.6–2.7 mm. Gnathosoma: Basis capituli dorsal posterior margin nearly straight, with blunt corners instead of cornua; dorsal surface without furrows (Fig. 52E). Ventrally, anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome broad, clearly indented, dentition roughly 3/3 or 4/4, with approximately 6 transverse rows of non-overlapping crenulations or mild teeth (Fig. 52C). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 52E). 182 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A C D

E F

Figs. 52A–G. Ixodes marmotae. Fig. 52A, female gnathosoma and scutum, dorsal; Fig. 52B, female hypostome, ventral; Fig. 52C, male hypostome, ventral; Fig. 52D, female gnathosoma, coxae and trochanters, ventral; Fig. 52E, male gnathosoma and idiosoma, dorsal; Fig. 52F, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 52G, nymph gnathosoma and scutum, dorsal. Redrawn from Cooley and Kohls (1945). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 183

H I J

K L

Figs. 53H–L. Ixodes marmotae. Fig. 53H, nymph hypostome, ventral; Fig. 53I, nymph gnathosoma, coxae and trochanters, ventral. Fig. 53J, larva gnathosoma, ventral; Fig. 53K, larva, dorsal; Fig. 53L, larva, ventral. Figs. 53H, I redrawn from Cooley and Kohls (1945); Figs. 53J–L redrawn, K, L modified from Allred et al. (1960). 184 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter with a small ventral projection. Scutum: Scapulae short, rounded (Fig. 52E). Lateral carinae absent. Surface with numerous small puncta, somewhat more distinct and larger marginally. Surface lacking pair of pit-like depressions midlaterally, but sometimes with paired furrows in that area. Spiracular plate: Suboval to subtriangular, with slightly longer axis either dorsoventral or anteroposterior. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median plate equal to or usually slightly longer than anal plate (Fig. 52F). Anal plate not widened posteriorly, similar in width to adanal plates posteriorly. Ventral plates all with numerous, fine puncta. Legs: Coxa I with internal spur tapered, blunt, slightly longer than external spur (Fig. 52F). Coxae II–IV lacking internal spurs. Coxae I–IV each with short, blunt external spur, sometimes progressively smaller from I–IV. Ventral coxal surfaces with scattered, short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 52F).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli nearly straight or slightly sinuous, cornua distinct, bluntly pointed (Fig. 52G). Auriculae formed as distinct, rounded, or slightly angular lateral lobes (Fig. 53I). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome broadly rounded, dentition 3/3 in distal fourth, then 2/2 nearly to base; lateral denticles larger and more pointed than inner denticles (Fig. 53H). Two pairs of post-hypostomal setae. Palpi somewhat thickened, club-shaped in dorsal view, widest at mid-level (Fig. 52G). Palpal tibiotarsus not visible dorsally. Palpal trochanter with long, narrowly rounded anterior flange and long, narrowly rounded posterior flange (Fig. 53I). Scutum: Length subequal to width. Shape subtriangular, widest well anterior to middle, with straight or slightly concave posterolateral margins (Fig. 52G). Scapulae moderate in size, bluntly pointed. Lateral carinae short, parallel with anterolateral margins. Cervical grooves usually distinct, divergent posteriorly nearly to posterolateral margins. Surface with sparse, small puncta. Legs: Coxa I with internal spur moderately tapered, pointed, longer than external spur (Fig. 53I). Coxae II–IV lacking internal spurs. Coxae I–IV each with short, blunt external spur, sometimes progressively smaller from I–IV. Ventral coxal surfaces with few smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 53I).

Larva. Body: Body length 0.5–0.6 mm. Gnathosoma: Dorsal posterior margin of basis capituli sinuous, with cornua divergent, bluntly pointed (Fig. 53K). Ventrally, auriculae formed as diagonal bulges laterally. Apex of hypostome broadly rounded, dentition 3/3 subapically, then 2/2 to base; lateral file with 6–8 bluntly pointed teeth, inner file with 5–7 more rounded teeth (Fig. 53J). Two pairs of post-hypostomal setae (Figs. 53J, L). Palpi moderately thick, femur+genu length to width ratio about 1.8–2.0:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, with a blunt anterior bulge and a well-developed, narrowly rounded, posteroventral flange (Fig. 53J). Scutum: Length approximately 0.8 times (0.77–0.88 times) width, widest at anterior third (Fig. 53K). Scapulae well developed, narrowly rounded; shield posterior margin convex. Surface finely micropunctate, and lineate-reticulate along anterolateral margins. Setae, 5 pairs. Idiosomal setation (Figs. 53K–L): Central dorsal setae 2 pairs. Supplemental dorsal setae absent. Marginal dorsal setae, 8 pairs. Marginal ventral setae, 3 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae at most twice that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae with anterior pair about twice as long as third pair (Fig. 53L). Legs: Coxa I with bluntly subtriangular internal spur (Fig. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 185

Map 23. Collection localities for Ixodes marmotae in Canada.

53L). Coxa II with curved ridge posteroventrally, this indistinct on coxa III. Coxae I–III lacking external spurs, but each with rounded bulge or ridge apicoventrally. Coxae I–II–III setal formula 3–2 or 3–2; coxal setae inconspicuously barbed. Femur III with usually 8 setae. Trochanters I–III lacking spurs (Fig. 53L).

Natural history. There is very little information on the natural history of this tick. In the records provided by Gregson (1956), adults were collected in May, June, and July, while juveniles were collected April through July. Wilkinson (1984) collected nymphs on yellow- bellied marmots, Marmota flaviventris, in May and August near Stump Lake, British Columbia. There are no reports of pathogenic organisms being transmitted by I. marmotae.

Distribution. Ixodes marmotae has a relatively limited distribution in western North America, including British Columbia, Washington, Oregon, Idaho, Montana, Wyoming, and Utah.

Hosts. In Canada, the only known hosts are ground-dwelling sciurids (squirrels and marmots) and woodrats.

Ixodes (Pholeoixodes) marxi Banks Squirrel tick (Figs. 54–55, Map 24) Ixodes hexagonus inchoatus Neumann, 1901: 283 (in part). Ixodes marxi Banks, 1908: 32. Ixodes marxi Banks: Cooley and Kohls 1945: 125 (male, nymph). Ixodes marxi Banks: Clifford et al. 1961: 232 (larva). Ixodes (Pholeoixodes) marxi Banks: Clifford et al. 1973: 498.

Adult. Female: Body: Length about 2.1 mm unfed, reaching to 6.3 mm engorged. Gnathosoma: Porose areas of basis capituli small, subcircular, separated by nearly twice the width of one porose area (Fig. 54A). Dorsal posterior margin of basis capituli nearly straight, with small, rounded cornua (Fig. 54A). Auriculae formed as triangular lateral projections (Fig. 54E). Anterior margin of basis capituli sloped on either side of hypostome. Apex of 186 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins hypostome pointed, dentition 3/3 along apical one-third, then 2/2 nearly to base, denticles sharp, those in lateral files larger (Fig. 54B). Palpi with lateral profile margin nearly straight in dorsal view, but somewhat thickened, club-like, widest at mid-length (Fig. 54A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with small ventral projection (Fig. 54E). Scutum: Length about 1.1–1.2 times greater than width, widest slightly anterior to middle; shape subtriangular, with posterolateral margins slightly concave (Fig. 54A). Scapulae bluntly pointed. Lateral carinae faint. Cervical grooves diverging and fading before reaching posterolateral margins. Surface sparsely coarsely punctate, rugo-punctate anterolaterally. Legs: Coxa I with internal spur a short, blunt salience, hardly more developed than apical salience representing external spur (Fig. 54E). Coxae II and III with salient posterior corners instead of internal spurs, these absent on IV. Coxae II, III, sometimes IV, each with apical salience instead of external spur. Ventral coxal surfaces with few, short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 54E).

Adult. Male: Body: Length approximately 2.5 mm Gnathosoma: Basis capituli dorsal posterior margin nearly straight or slightly sinuous, with blunt corners instead of cornua; dorsal face without furrows (Fig. 54F). Ventrally, anterior margin of basis capituli sloped, slightly humped on either side of hypostome. Apex of hypostome slightly rounded, dentition formed as 7 or 8 transverse rows of mild, mostly 2/2 crenulations, becoming indistinct medially (Fig. 54C). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 54F). Dorsally, palpal femur faintly delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter with small, rounded ventral flange (Fig. 54G). Scutum: Scapulae bluntly rounded (Fig. 54F). Lateral carinae absent. Surface with numerous puncta, larger and more dense along margins. Surface lacking pair of pit-like depressions mid-laterally, but sometimes with paired faint grooves there. Spiracular plate: Subcircular, without distinct longer axis. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median and anal plates subequal in length (Fig. 54G). Anal plate widened posteriorly, where somewhat wider than adanal plates. Median plate with fine, inconspicuous puncta; other ventral plates with numerous large puncta. Legs: Coxa I with internal spur a short, blunt salience, similar in form and size to apical salience representing external spur (Fig. 54G). Coxae II and III, with salient posterior corners instead of internal spurs, these absent on IV. Coxae II, III, sometimes IV, each with apical salience instead of external spur. Ventral coxal surfaces with few, short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 54G).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli nearly straight, with salient corners forming small, divergent cornua (Fig. 54H). Auriculae formed as rounded or bluntly angular lateral lobes posterior to mid-level of basis capituli (Fig. 55I). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome narrowly rounded, dentition 3/3 for a couple of subapical rows, then 2/2 nearly to base; lateral denticles larger, more pointed than inner denticles (Fig. 54D). Two pairs of post-hypostomal setae. Palpi somewhat thickened, club-like, in dorsal view, widest at mid-length (Fig. 54H). Palpal tibiotarsus not visible dorsally. Palpal trochanter with small ventral projection (Fig. 55I). Scutum: Length about 1.1–1.2 times width, widest slightly anterior to middle; shape subtriangular, with posterolateral margins straight or slightly concave (Fig. 54H). Scapulae small, bluntly rounded. Lateral carinae absent. Cervical grooves divergent, fading before reaching posterolateral margins. Surface sparsely, finely punctate. Legs: Coxa I with internal spur short, blunt, hardly more developed than apical salience representing external A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 187

spur (Fig. 55I). Coxae II–IV lacking internal spurs and external spurs. Ventral coxal surfaces with few, short, smooth, or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 55I).

Larva. Body: Body length 0.5–0.6 mm. Gnathosoma: Dorsal posterior margin of basis capituli straight or slightly concave, with rounded posterior corners instead of cornua (Fig. 55J). Ventrally, auriculae formed as bluntly angular lateral projections (Fig. 55K). Apex of hypostome broadly rounded, dentition below coronal area 2/2 along entire length; lateral file with 5–6 bluntly pointed teeth, inner file with 4–5 smaller, more rounded teeth (Fig. 55K). Two pairs of post-hypostomal setae (Fig. 55K). Palpi moderately thick, femur+genu length to width ratio about 2.3:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, with 1 small, narrowly rounded posteroventral flange (Fig. 55K). Scutum: Length slightly less (approximately 0.9) than width, widest at mid-level (Fig. 55J). Scapulae well developed, bluntly rounded; shield posterior margin convex. Surface finely micropunctate, and fully imbricated. Setae, 5 pairs. Idiosomal setation (Figs. 55J–K): Central dorsal setae, 2 pairs. Supplemental dorsal setae absent. Marginal dorsal setae, 9 pairs. Marginal ventral setae, 3 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae less than twice that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae usually similar to one another in size. Legs: Coxa I with bluntly subtriangular internal spur (Fig. 55K). Coxae II and sometimes III with a linear ridge posteroventrally. Coxae I–III lacking external spurs. Coxae I–II–III setal formula 3–2 or 3–2; coxal setae smooth or inconspicuously barbed. Femur III with 9 setae. Trochanters I–III lacking spurs (Fig. 55K).

Natural history. There are a number of records for I. marxi in eastern Canada (McLean and Larke 1963; Jones and Thomas 1980; Gyorkos and Hilton 1982; Barker et al. 1992), where it primarily parasitises tree squirrels. These records have mostly been anecdotal in nature, and there is little information on the natural history of this species.

Map 24. Collection localities for Ixodes marxi in Canada. 188 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

E A C

F G

Figs. 54A–H. Ixodes marxi. Fig. 54A, female gnathosoma and scutum, dorsal; Fig. 54B, female hypostome, ventral; Fig. 54C, male hypostome, ventral; Fig. 54D, nymph hypostome, ventral; Fig. 54E, female gnathosoma, coxae and trochanters, ventral; Fig. 54F, male gnathosoma and idiosoma, dorsal; Fig. 54G, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 54H, nymph gnathosoma and scutum, dorsal. Redrawn from Cooley and Kohls (1945). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 189

Figs. 55I–K. Ixodes marxi. Fig. 55I, nymph gnathosoma, coxae and trochanters, ventral; Fig. 55J, larva, dorsal; Fig. 55K, larva, ventral. Fig. 55I redrawn from Kohls (1947); Figs. 55J, K redrawn from Webb et al. (1990). 190 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

McLean and Larke (1963) isolated Powassan virus from I. marxi in Ontario. They suggested that this tick may be important in transmission and maintenance of this virus in red squirrels, but this relationship has not been thoroughly investigated.

Distribution. Ixodes marxi is widely distributed in the United States of America east of the Mississippi River, though it is not abundant in the deep south. In Canada, it has been found in the Maritime Provinces, exclusive of Newfoundland, west to Ontario.

Hosts. Tree squirrels are the most important hosts for this tick, though there are records for snowshoe hare as well as predators of squirrels. There are numerous records for this tick biting cats, dogs and humans.

Ixodes (Pholeoixodes) rugosus Bishopp (Figs. 56, 57, Map 25) Ixodes cookei rugosus Bishopp, 1911: 197 (female, male). Ixodes rugosus Bishopp: Cooley and Kohls 1945: 119 (nymph). Ixodes rugosus Bishopp: Bennett et al. 1989: 262 (larva). Ixodes (Pholeoixodes) rugosus Bishopp: Clifford et al. 1973: 498.

Adult. Female: Body: Length about 2.9 mm unfed, reaching to 7.8 mm engorged. Gnathosoma: Porose areas of basis capituli large, subtriangular, separated by less than the width of one porose area, their posterior edges near posterior margin of basis (Fig. 56A). Dorsal posterior margin of basis capituli slightly concave, with salient corners instead of cornua. Auriculae formed as mild lateral humps. Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded, dentition 2/2 below corona area, denticles bluntly pointed, those in lateral files slightly larger (Fig. 56C). Palpi with lateral profile margin nearly straight in dorsal view, but somewhat thickened, club-like, widest at mid-length (Fig. 56A). Palpal trochanter with small ventrolateral ridge-like projection. Scutum: About as long or slightly longer than wide, widest slightly anterior to midlength (Fig. 56A). Shape subtriangular, with posterolateral margins straight or slightly concave. Scapulae bluntly pointed. Lateral carinae weakly defined, nearly reaching posterolateral margins. Cervical grooves shallow, well short of reaching posterolateral margins. Surface covered with numerous, coarse puncta, sometimes with irregular rugosities intermingled with puncta anterolaterally and posteriorly. Legs: Coxa I with internal spur tapered, pointed, longer than external spur (Fig. 56B). Coxae II–IV lacking internal spurs, but sometimes with salient posterior corners. Coxae I–IV each with small, bluntly pointed external spur. Ventral coxal surfaces with few short, smooth, or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 56B).

Adult. Male: Body: Length 2.7–2.9 mm. Gnathosoma: Basis capituli dorsal posterior margin straight or slightly concave, with bluntly rounded corners instead of cornua; dorsal face without furrows (Fig. 56F). Ventrally, anterior margin of basis capituli gently humped on either side of hypostome. Apex of hypostome slightly indented, dentition roughly 4/4, with a file of prominent, pointed lateral teeth bordering approximately 7 diagonal, overlapping rows of blunt, slightly crenulate teeth (Fig. 56D). Palpi thick, club- like in dorsal view, with symmetrically rounded apices (Fig. 56F). Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter with small ventral ridge. Scutum: Scapulae bluntly rounded (Fig. 56F). Lateral carinae faint. Surface faintly rugose anterolaterally, with numerous, moderately large puncta, denser along A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 191

C D

F G I

Figs. 56A–I. Ixodes rugosus. Fig. 56A, female gnathosoma and scutum, dorsal; Fig. 56B, female gnathosoma, coxae and trochanters, ventral; Fig. 56C, female hypostome, ventral; Fig. 56D, male hypostome, ventral; Fig. 56E, nymph gnathosoma and scutum, dorsal; Fig. 56F, male gnathosoma and idiosoma, dorsal; Fig. 56G, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 56H, nymph gnathosoma, coxae and trochanters, ventral; Fig. 56I, nymph hypostome, ventral. Redrawn from Cooley and Kohls (1945). 192 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

J K

Figs. 57J–L. Ixodes rugosus. Fig. 57J, larva, dorsal; Fig. 57K, larva, ventral; Fig. 57L, larva hypostome, palp trochanter, basis capituli, ventral. Redrawn from Webb et al. (1990). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 193

margins. Surface lacking pair of pit-like depressions mid-laterally, but sometimes with paired, faint grooves there. Spiracular plate: Oval, with slightly longer axis dorsoventral. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median plate nearly 1.5 times longer than anal plate (Fig. 56G). Anal plate slightly widened posteriorly, where slightly wider than adanal plates. Median plate faintly punctate; anal and adanal plates with large puncta, though smaller than on scutum. Legs: Coxa I with internal spur tapered, pointed, much longer than external spur (Fig. 56G). Coxae II–IV lacking internal spurs, but II, III sometimes with salient posterior corners. Coxae I–IV each with small, bluntly pointed external spur. Ventral coxal surfaces with scattered short, smooth, or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 56G).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli slightly concave, with salient corners forming small, divergent cornua (Fig. 56E). Auriculae formed as curved or slightly angular lateral lobes (Fig. 56H). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded, dentition 2/2 below corona area; lateral denticles slightly larger, more pointed than inner ones (Fig. 56I). Two pairs of post- hypostomal setae. Palpi short, thickened, club-like in dorsal view. Palpal tibiotarsus not visible dorsally. Palpal trochanter with salient, rounded posterolateral flange (not well shown in Fig. 56H). Scutum: About as long as wide, widest at about midlength (Fig. 56E). Shape subrhomboid, with posterolateral margins nearly straight. Scapulae bluntly pointed. Lateral carinae distinct, parallel with anterolateral margins. Cervical grooves long, divergent, nearly reaching posterolateral margins. Surface sparsely, weakly punctate. Legs: Coxa I with internal spur narrowly rounded, slightly longer than external spur (Fig. 56H). Coxae II–IV lacking internal spurs. Coxae I–IV each with small, bluntly pointed external spur. Ventral coxal surfaces with few, short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 56H).

Larva. Body: Body length 0.7–0.8 mm. Gnathosoma: Dorsal posterior margin of basis capituli slightly sinuous, with blunt corners projecting laterally or posterolaterally, instead of cornua (Fig. 57J). Ventrally, auriculae formed as diagonal ridges. Apex of hypostome broadly rounded, dentition below coronal area 2/2 along entire length; lateral file with 7–8 pointed teeth, inner file with 5–6 smaller, blunter teeth (Fig. 57L). Two pairs of post- hypostomal setae (Figs. 57K–L). Palpi moderately thick; femur+genu length to width ratio about 2.0–2.2:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, with 2 flanges subequal in small size, a bluntly pointed anterior flange and a more broadly rounded posterior flange (Fig. 57L). Scutum: Length slightly less (approximately 0.95 times) than width, widest near anterior third (Fig. 57J). Scapulae weak, rounded; shield posterior margin convex. Surface densely micropunctate, and lineate-reticulate along anterolateral margins. Setae, 5 pairs. Idiosomal setation (Figs. 57J–K): Central dorsal setae, 2 pairs. Supplemental dorsal setae absent. Marginal dorsal setae, 8 pairs. Marginal ventral setae, 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae at most 1.5 that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in size. Legs: Coxa I with broadly, bluntly triangular internal spur with rounded apex (Fig. 57K). Coxae II and III each with curved ridge posteroventrally. Coxae I–III lacking external spurs. Coxae I–II–III setal formula 3–2 or 3–2; coxal setae smooth or inconspicuously barbed. Femur III with 8 setae. Trochanters I–III lacking spurs (Fig. 57K). 194 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Map 25. Collection localities for Ixodes rugosus in Canada.

Natural history. There is virtually no information on the natural history and occurrence of this species in Canada, though Gregson (1942c, 1956) reported it infesting coyotes. Ixodes rugosus has not been implicated in transmission of any pathogenic organisms or parasites.

Distribution. Ixodes rugosus is restricted to the western regions of North America, from British Columbia south to California.

Hosts. Carnivores, such as skunks, weasels, foxes, and dogs, are overwhelmingly reported as hosts for this tick.

Ixodes (Pholeoixodes) sculptus Neumann (Fig. 58, Map 26) Ixodes sculptus Neumann, 1904: 462. Ixodes aequalis Wherry and Wellman, 1909: 376 (nomen nudum). Ixodes aequalis Banks, 1910: 8. Ixodes sculptus Neumann: Hixson 1932: 36 (male, nymph, larva). Ixodes (Pholeoixodes) sculptus Neumann: Clifford et al. 1973: 498.

Adult. Female: Body: Length 2.2–2.6 mm unfed, reaching to 6.3 mm engorged. Gnathosoma: Porose areas of basis capituli variable in size, shape, from subcircular to subtriangular, separated by width of one porose area (Fig. 58A). Dorsal posterior margin of basis capituli salient, usually markedly sinuous, sometimes concave; cornua large, bluntly pointed, slightly incurved (Fig. 58A). Auriculae faintly formed as lateral bulges or absent. Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded, dentition 3/3 subapically, then 2/2 to base, denticles pointed, those in lateral files slightly larger and sharper (Fig. 58B). Palpi moderately long, lateral profile margin nearly straight in dorsal view, but slightly club-shaped, thicker at mid-length (Fig. 58A). Palpal tibiotarus not visible dorsally. Palpal trochanter with a small ventral projection. Scutum: Length slightly greater (1.1 times) than width. Shape subrhomboidal to subtriangular, widest slightly anterior to middle, with posterolateral margins nearly straight (Fig. 58A). Scapulae bluntly pointed. Lateral carinae usually distinct and reaching to posterolateral A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 195

margins. Cervical grooves usually distinct and reaching nearly to posterolateral margins. Surface entirely covered with coarse puncta, sometimes rugopunctate in anterolateral areas. Legs: Coxa I with internal spur tapered, bluntly pointed, longer than external spur (not well shown in Fig. 58E). Coxae II–IV lacking internal spurs. Coxae I–IV each with bluntly triangular external spur, sometimes progressively smaller from I–IV. Ventral coxal surfaces with few short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 58E).

Adult. Male: Body: Length approximately 2.5 mm. Gnathosoma: Basis capituli dorsal posterior margin straight or slightly concave, with blunt corners instead of cornua; dorsal surface without furrows (Fig. 58G). Ventrally, anterior margin of basis capituli sloped, slightly humped on either side of hypostome. Apex of hypostome broadly rounded, sometimes slightly indented, dentition roughly 3/3, with a file of bluntly pointed lateral teeth bordering approximately 6 diagonal, non-overlapping rows of blunt, slightly crenulate teeth (Fig. 58C). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 58G). Dorsally, palpal femur delineated from genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter with small ventral projection. Scutum: Scapulae rounded, lateral carinae absent (Fig. 58G). Surface with numerous, moderately large puncta, sometimes larger and denser along margins. Surface lacking pair of pit-like depressions mid-laterally, but with paired, elongate grooves there. Spiracular plate: Subcircular, without distinct longer axis. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median plate slightly longer than anal plate (Fig. 58H). Anal plate widened posteriorly, where slightly wider than adanal plates. Median plate with small puncta; anal and adanal plates with larger puncta, much as on scutum. Legs: Coxa I with internal spur tapered, pointed, longer than external spur (Fig. 58H). Coxae II–IV lacking internal spurs, but II and III with salient posterior corners. Coxae I–IV each with bluntly pointed external spur. Ventral coxal surfaces with few short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 58H).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli salient, usually sinuous, sometimes nearly straight, cornua large, bluntly pointed (Fig. 58I). Auriculae formed as conspicuous lateral lobes closely behind bases of palpi (Fig. 58F). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome rounded, dentition 3/3 for a couple of subapical rows, then 2/2 nearly to base; lateral denticles larger, more pointed than inner denticles (Fig. 58D). Two pairs of post-hypostomal setae. Palpi club-shaped in dorsal view (Fig. 58I). Palpal tibiotarus not visible dorsally. Palpal trochanter with conspicuous, narrowly rounded anterior flange and similar, narrowly rounded posterior flange (Fig. 58F). Scutum: About as long as wide. Shape subtriangular, widest slightly anterior to middle, with posterolateral margins nearly straight (Fig. 58I). Scapulae bluntly pointed. Lateral carinae usually distinct and parallel with anterolateral margins. Cervical grooves often faint, divergent, not reaching to posterolateral margins. Surface sparsely covered with puncta. Legs: Coxa I with internal spur tapered, bluntly pointed, somewhat longer than external spur (Fig. 58F). Coxae II–IV lacking internal spurs. Coxae I–IV each with bluntly triangular external spur, sometimes progressively smaller from I–IV. Ventral coxal surfaces with few short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 58F).

Larva. Body: Body length 0.5–0.6 mm. Gnathosoma: Dorsal posterior margin of basis capituli sinuous, with cornua well developed, bluntly pointed, directed posterolaterally (Fig. 58K). Ventrally, auriculae not developed. Apex of hypostome broadly rounded, dentition 196 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A F

C D E

I J

G H

K L

Figs. 58A–L. Ixodes sculptus. Fig. 58A, female gnathosoma and scutum, dorsal; Fig. 58B, female hypostome, ventral; Fig. 58C, male hypostome, ventral; Fig. 58D, nymph hypostome, ventral; Fig. 58E, female gnathosoma, coxae and trochanters, ventral; Fig. 58F, nymph gnathosoma, coxae and trochanters, ventral; Fig. 58G, male gnathosoma and idiosoma, dorsal; Fig. 58H, male, gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 58I, nymph gnathosoma and scutum, dorsal; Fig. 58J, larva hypostome, palp trochanter, basis capituli, ventral; Fig. 58K, larva, dorsal; Fig. 58L, larva, ventral. Figs. 58A–I redrawn from Cooley and Kohls (1945); Figs. 58J–L redrawn and J augmented from Webb et al. (1990). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 197

below coronal area 2/2 along entire length; lateral file with 6–8 bluntly pointed teeth, inner file with 4–6 smaller, blunter teeth (Fig. 58J). Two pairs of post-hypostomal setae (Figs. 58J, L). Palpi thick; femur+genu length to width ratio about 1.7:1; ventral apex of femorogenu with 1 or 2 small tubercles or flanges (Fig. 58J). Palpal trochanter distinct, with two narrowly rounded to bluntly pointed flanges, anterior flange short, rounded, posteroventral flange longer, narrowly rounded (Fig. 58J). Scutum: Length nearly (0.90–0.95 times) equal to width, widest at anterior third (Fig. 58K). Scapulae undeveloped; shield lateral margins slightly incurved at level of second lateral setae; posterior margin convex. Surface densely, uniformly micropunctate, sometimes with underlying reticula emergent anterolaterally and anteriorly. Setae, usually 5 pairs, with anterolateral pair sometimes inserted off margin. Idiosomal setation (Figs. 58K–L): Central dorsal setae, 2 pairs. Supplemental dorsal setae absent. Marginal dorsal setae, usually 8 pairs. Marginal ventral setae, usually 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae about 1.5 longer than that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in size. Legs: Coxa I with broadly, bluntly right-triangular internal spur (Fig. 58L). Coxae II and III each with curved ridge posteroventrally. Coxae I–III lacking external spurs. Coxae I–II–III setal formula 3–2 or 3–2; coxal setae smooth or inconspicuously barbed. Femur III with 8 setae. Trochanters I–III lacking spurs (Fig. 58L).

Natural history. Ixodes sculptus has been reported from Canada by Brown (1944), Brown and Kohls (1950), and Gregson (1956), but despite Brown having collected more than 2500 specimens, there is no published information on its natural history or seasonal occurrence in Canada. Beck et al. (1963) found that there was a peak in activity of nymphs from November to March in Nevada and Salkeld et al. (2006) found nymphs to be most abundant on ground squirrels during June through August (when sampling ended) in northern Colorado. However, the most detailed study on this tick is that of Hixson (1932) in Iowa, where it was a common parasite of thirteen-lined ground squirrels, Ictidomys tridecemlineatus. He reported that all instars were present on their hosts throughout the year. Ixodes sculptus is adapted to life in the burrows of the host, where they may be collected from litter on the burrow floor and unfed ticks in each stage are capable of surviving for more than 100 days under suitable conditions. Larvae required 3–9 days for engorgement, and at room temperature, moulted after 2–3 weeks. Nymphs required 4–9 days to engorge and moulted at room temperature 2–3 weeks later. Females took up to two weeks to engorge and at room temperature, began to lay eggs 6–9 days later. An average of 828 eggs (range 644–1006) was laid over 26–29 days. All engorged instars drop from their hosts at night, when the host is in the burrow. Males were not found on mammal hosts, and probably do not feed. Mating occurs among the litter in the burrow. Prevalence of infestation is often low (e.g., Kietzmann 1987; Lang 1999), but there are reports of ground squirrels being heavily infested (Eddy and Joyce 1942), and more than 70% of ground squirrels at one site in Iowa were infested (Kietzmann and Kietzmann 1987). Ixodes sculptus is not known to be a competent vector of any pathogenic organisms, but Salkeld et al. (2006) emphasised the need to examine this species where it may serve as a silent vector for pathogens maintained primarily by other species of ticks.

Distribution. Ixodes sculptus is found through central regions of North America. Records in Canada are scattered throughout the southern parts of the prairie provinces, from southwestern Manitoba westwards into south central British Columbia. 198 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Map 26. Collection localities for Ixodes sculptus in Canada.

Hosts. The primary hosts of this tick are burrowing mammals, especially ground squirrels, prairie dogs, and pocket gophers, but their predators may become secondarily infested. See Salkeld et al. (2006) for a summary of host records in North America.

Ixodes (Pholeoixodes) texanus Banks (Fig. 59, Map 27) Ixodes pratti Banks, 1908: 27 (in part). Ixodes texanus Banks, 1909: 172. Ixodes texanus Banks: Cooley and Kohls 1945: 78 (male, nymph). Ixodes texanus Banks: Allred et al. 1960: 22 (larva). Ixodes (Pholeoixodes) texanus Banks: Clifford et al. 1973: 498.

Adult. Female: Body: Length reaching to at least 9.0 mm engorged. Gnathosoma: Porose areas of basis capituli deep, irregular to nearly circular, separated by slightly more than the width of one porose area, and surrounded by wrinkled sculpturing (Fi. 59A). Dorsal posterior margin of basis capituli nearly straight or slightly sinuous, salient, with rounded corners or small, rounded cornua (Fig. 59A). Auriculae faintly formed as lateral bulges or ridges, or absent (Fig. 59D). Anterior margin of basis capituli with prominent hump on either side of hypostome (Fig, 59B). Apex of hypostome rounded, dentition 4/4 in distal portion, then briefly 3/3, then 2/2 to base, lateral denticles large, pointed, inner denticles progressively smaller, rounded (Fig. 59B). Palpi thickened, club-shaped when viewed dorsally, widest at mid-length (Fig. 59A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with small ventrolateral projection (Fig. 59D). Scutum: Subequally as wide as long, subtriangular, widest slightly anterior to middle, with posterolateral margins straight or slightly concave, scapulae bluntly pointed (Fig. 59A). Lateral carinae absent. Cervical grooves, if present, usually visible only in mid-region, and often flanked by longitudinal wrinkles laterally. Surface covered with scattered, coarse puncta, and rugose or wrinkled laterally. Legs: Coxa I with internal spur a rounded salience, hardly more developed than apical blunt salience representing external spur (Fig. 59D). Coxa II with salient posterior corner instead of internal spur, this absent on coxae III and IV. Coxae I–III each with blunt apical salience instead of external spur, this absent on coxa IV. Ventral coxal surfaces with few short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 59D). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 199

Adult. Male: Body: Length 1.4–1.5 mm. Gnathosoma: Basis capituli dorsal posterior margin slightly sinuous, with small, blunt cornua (much as in Fig. 59A of female); dorsal face slightly furrowed (Fig. 59F). Ventrally, anterior margin of basis capituli strongly humped on either side of hypostome. Apex of hypostome slightly indented, dentition roughly 3/3, with 6 or 7 rows of faint transverse crenulations, becoming indistinct medially (Fig. 59C). Palpi thick, club-like in dorsal view, with symmetrically rounded apices (Fig. 59F). Dorsally, palpal femur fused with genu. Palpal tibiotarsus not visible dorsally. Palpal trochanter with ventral ridge. Scutum: Scapulae bluntly rounded (Fig. 59F). Lateral carinae absent. Surface slightly rugose anterolaterally, with numerous large, shallow, puncta, denser marginally. Surface lacking pair of pit-like depressions midlaterally. Spiracular plate: Subcircular, with slightly longer axis dorsoventral. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Median plate slightly longer than anal plate (Fig. 59G). Anal plate gradually widened posteriorly, where wider than adanal plates. Median plate with small puncta; anal and adanal plates with larger puncta, similar to scutum. Legs: Coxa I with internal spur a rounded salience, slightly more developed than apical blunt salience representing external spur (Fig. 59G). Coxa II with salient posterior corner instead of internal spur, this absent on coxae III and IV. Coxae I–IV each with blunt apical salience instead of external spur, that on III more spur-like. Ventral coxal surfaces with few short, smooth, or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 59G).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli nearly straight. cornua bluntly pointed, sometimes weakly developed (Fig. 59H). Auriculae faintly formed as lateral ridges. Anterior margin of basis capituli with angular hump on either side of hypostome (Fig. 59E). Apex of hypostome broadly rounded, dentition 3/3 for 1 or 2 rows subapically, then 2/2 nearly to base; lateral denticles large, pointed, inner denticles smaller, rounded (Fig. 59J). Two pairs of post-hypostomal setae. Palpi thickened, club-shaped when viewed dorsally (Fig. 59H). Palpal tibiotarsus not visible dorsally. Palpal trochanter with small, rounded ventrolateral flange (Fig. 59E). Scutum: Length nearly 0.9 times width. Shape subtriangular, widest slightly anterior to middle, with posterolateral margins nearly straight (Fig. 59H). Scapulae narrowly rounded. Lateral carinae absent. Cervical grooves, if present, usually visible only in mid-region, and often flanked by longitudinal wrinkles laterally. Surface with scattered, faint puncta, and rugose or wrinkled laterally. Legs: Coxa I with internal spur short, rounded (Fig. 59E). Coxae II–IV lacking internal spurs. Coxae I–IV with external spurs rudimentary or lacking. Ventral coxal surfaces with few short, smooth, or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 59E).

Larva. Body: Body length 0.46–0.51 mm. Gnathosoma: Dorsal posterior margin of basis capituli straight or slightly concave, with rounded corners instead of cornua (Fig. 59K). Ventrally, auriculae formed on either side as a faint oblique ridge joining medially to form a posteriorly curved ridge. Apex of hypostome broadly rounded or slightly indented, dentition below coronal area 2/2 nearly to base; lateral file with 6–7 bluntly pointed, overlapping teeth, inner file with 5–6 smaller, blunter teeth (Fig. 59I). Two pairs of post-hypostomal setae (Figs. 59I, L). Palpi moderately thick, femur+genu length to width ratio about 1.7–2.0:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, with 1 small, bluntly rounded, posterior spur (Fig. 59I). Scutum: Length about 0.8 width, widest at anterior third (Fig. 59K). Surface finely micropunctate, and fully imbricated. Scapulae weakly developed, rounded; shield posterior margin convex. Setae, 5 pairs. Idiosomal setation (Figs. 59K–L): Central dorsal setae, 2 pairs. Supplemental dorsal setae absent. 200 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A D E

I F G

J L

Figs. 59A–L. Ixodes texanus. Fig. 59A, female gnathosoma and scutum, dorsal; insert enlarged cornua; Fig. 59B, female hypostome, ventral; Fig. 59C, male hypostome, ventral; Fig. 59D, female gnathosoma, coxae and trochanters, ventral; Fig. 59E, nymph gnathosoma, coxae and trochanters, ventral; Fig. 59F, male gnathosoma and idiosoma, dorsal; Fig. 59G, male gnathosoma, idiosoma, coxae and trochanters, ventral; Fig. 59H, nymph gnathosoma and scutum, dorsal; Fig. 59I, larva hypostome, palp trochanter, basis capituli, ventral; Fig. 59J, nymph hypostome, ventral; Fig. 59K, larva, dorsal; Fig. 59L, larva, ventral. Figs. 59A–H, redrawn from Cooley and Kohls (1945); Fig. 59I modified, J redrawn from Clifford et al. (1961); Figs. 59K, L redrawn from Webb et al. (1990). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 201

Marginal dorsal setae, 9 pairs. Marginal ventral setae, 3 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae less than twice that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in size. Legs: Coxa I with weak, rounded or bluntly obtuse-angled internal spur (Fig. 59L). Coxae II and III with curved ridge posteroventrally. Coxae I–III lacking external spurs. Coxae I–II–III setal formula 3–2 or 3–2. Femur III with 9 setae. Trochanters I–III lacking spurs (Fig. 59L).

Natural history. This is a widely distributed tick in North America, but its abundance in Canada is somewhat limited (Gregson 1956; Webster 1966). There is no information on its natural history and life history in Canada. Kollars and Oliver (2003) found that this species was active all year in Missouri, with the greatest number of adults, larvae, and nymphs active in May, October-November, and July, respectively. In North Carolina, females were most abundant on raccoons in summer (absent in fall and winter), larvae were most abundant in fall and winter, and nymphs in winter and spring (Oulette et al. 1997). Males have not been collected on animals. Engorged ticks tend to drop from their hosts in the dark (Darling 1969), perhaps facilitating maintenance in their occupation of the nest. Engorged I. texanus freeze at -5 to -7 ºC (Mail 1942). Evidence for the involvement of I. texanus in transmission of pathogens is largely circumstantial. Anderson et al. (1981) suggested that I. texanus may be the vector for Babesia lotori infection in raccoons. Anderson et al. (1986) isolated Rickettsia rickettsii from a nymph in Connecticut. Dugan et al. (2005) suggested I. texanus as a possible vector for ehrlichiae in raccoons in Georgia.

Distribution. Ixodes texanus is widely distributed throughout the United States of America, but has been recorded in Canada only from Québec, Ontario, and British Columbia, with one disjunct record from Newfoundland. It has recently been recorded from Mexico (Guzmán-Cornejo et al. 2007; Guzmán-Cornejo and Robbins 2010).

Hosts. Throughout the range of this tick, raccoons seem to be the most closely linked host. In Canada, it seems that carnivores, including dogs, are also hosts for this tick. There are records from ground-dwelling and arboreal sciurids as well.

Map 27. Collection localities for Ixodes texanus in Canada. 202 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Ixodes (Pholeoixodes) gregsoni Lindquist, Wu, and Redner (Fig. 60, Map 28) Ixodes (Pholeoixodes) gregsoni Lindquist, Wu and Redner, 1999: 152–170.

Adult. Female: Body: Length unfed not determined, reaching to 9.6 mm engorged. Gnathosoma: Porose areas of basis capituli deep, irregular, or usually subtriangular, separated by furrowed interval slightly less than width of one porose area (Fig. 60A). Dorsal posterior margin of basis capituli nearly straight or slightly sinuous, with rounded corners or small, rounded cornua (Fig. 60A). Auriculae formed as oblique lateral ridges (Fig. 60B). Anterior margin of basis capituli with slight hump on either side of hypostome (Fig. 60C). Apex of hypostome rounded, dentition 3/3 subapically, then 2/2 to base, lateral denticles slightly larger and more pointed than inner denticles (Fig. 60C). Palpi thickened, club-shaped when viewed dorsally, widest at mid-length (Fig. 60A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with small ventrolateral projection. Scutum: Subequally as wide as long, subtriangular, widest slightly anterior to middle, with posterolateral margins straight or slightly concave (Fig. 60D). Scapulae bluntly pointed. Lateral carinae faint. Cervical grooves faint, reaching nearly to posterolateral margins. Surface covered with coarse puncta, lacking rugose or wrinkled areas. Legs: Coxa I with internal spur short, blunt, similar in size to external spur (Fig. 60E). Coxa II–IV lacking internal spurs. Coxae I–IV each with a small, blunt external spur, progressively smaller from I–IV. Ventral coxal surfaces with few, short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs.

Adult male. Unknown

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli slightly concave and sinuous, with salient corners forming small, divergent cornua (Fig. 60F). Auriculae formed as oblique lateral ridges. Anterior margin of basis capituli with slight hump on either side of hypostome. Apex of hypostome rounded, dentition 3/3 for a couple of subapical rows, then 2/2 nearly to base; lateral denticles larger, more pointed than inner ones (Fig. 60H). Two pairs of post-hypostomal setae. Palpi short, thickened, club-shaped when viewed dorsally. Palpal tibiotarsus not visible dorsally. Palpal trochanter with small, blunt anterior flange and moderately developed, rounded posterior flange (Fig. 60G).Scutum: Nearly as long as wide; shape subtriangular, widest slightly anterior to middle, with posterolateral margins straight or slightly concave (Fig. 60I). Scapulae rounded, indistinct. Lateral carinae faint. Cervical grooves faint, slightly divergent, not reaching posterolateral margins. Surface covered with fine reticula and scattered, sparse puncta. Legs: Coxa I with internal spur short, blunt, similar in size and form to external spur (Fig. 60M). Coxa II–IV lacking internal spurs. Coxae I–IV each with a small, bluntly rounded external spur, progressively smaller from I–IV. Ventral coxal surfaces with few short, smooth, or slightly barbed setae. Trochanters I–IV lacking spurs.

Larva. Body: Body length, engorged, 1.25–1.50 mm, excluding gnathosoma. Gnathosoma: Dorsal posterior margin of basis capituli slightly sinuous, with cornua weakly developed, with bluntly rounded apices (Fig. 60J). Ventrally, auriculae formed faintly as oblique midlateral ridges. Apex of hypostome nearly flat or slightly indented, dentition 3/3 subapically, then 2/2 to base; files each with 5 or 6 bluntly pointed teeth, lateral teeth slightly larger, more overlapping than inner teeth (Fig. 60L). Two pairs of post-hypostomal setae. Palpi moderately thick; femur+genu length to width ratio approximately 1.8 (1.77–1.83):1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, with 2 A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 203

A B C

F G H

K L

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P Q

Figs. 60A–Q. Ixodes gregsoni. Fig. 60A, female gnathosoma, dorsal; Fig. 60B, female gnathosoma, ventral; Fig. 60C, female hypostome, ventral; Fig. 60D, female scutum; Fig. 60E, female coxae, ventral; Fig. 60F, nymph gnathosoma, dorsal; Fig. 60G, nymph gnathosoma, ventral; Fig. 60H, nymph hypostome, ventral; Fig. 60I, nymph scutum outline, dorsal; Fig. 60J, larva gnathosoma, dorsal; Fig. 60K, larva palpus, ventral; Fig. 60L, larva hypostome, ventral; Fig. 60M, nymph coxae and trochanters, ventral; Fig. 60N, larva coxae and trochanters, ventral; Fig. 60O, larva coxa I, ventral; Fig. 60P, larva idiosoma, dorsal; Fig. 60Q, larva scutum. Figs. 60A–C, E–G, L, O redrawn, 60P modified from Lindquist et al. (1999); Figs. 69D, H–K, M, N, Q are original. 204 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins flanges, a short, bluntly pointed anterior flange and short, rounded posteroventral flange (Fig. 60K). Scutum: Length about 0.85–0.90 times width, widest at anterior third (Fig. 60Q). Scapulae weakly developed, rounded; shield posterior margin convex. Surface finely micropunctate, and usually lightly reticulated over most of area, more conspicuously so anterolaterally. Setae, five pairs.Idiosomal setation (Fig. 60P): Central dorsal setae, 2 pairs. Supplementary dorsal setae absent. Marginal dorsal setae, 8 pairs. Marginal ventral setae, 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae about 1.5 that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in small size. Legs: Coxa I with small rounded internal spur (Figs. 60N, O). Coxae II and III each with weakly defined, elongate ridge posteroventrally (Fig. 60N). Coxae I–III lacking external spurs. Coxae I–II–III setal formula 3–2 or 3–2; coxal setae inconspicuously barbed. Femur III with 8 or 9 setae. Trochanters I–III lacking spurs (Fig. 60N).

Natural history. Ixodes gregsoni is the most recently described species of tick in Canada (Lindquist et al. 1999), and one of the least well known. The male has not been collected or described, and may be confined largely to the nests of mustelids. All other instars are primarily parasites of mustelids in Canada, and have been collected from these hosts throughout the fall and winter months, October to February. Lindquist et al. (1999) reported its range to be eastern boreal in Canada; their anticipation that it may occur more widely along the northern tier of the contiguous United States of America and Canada, west to Alaska is supported by recent records from Manitoba (T.D.G., personal observation), Maine and Vermont (Lubelczyk et al. 2007). Lubelczyk et al. (2007) reported females collected in April from a domestic cat; it appears that I. gregsoni has the potential to occur in a peridomestic environment where conditions are favourable. Although anticipated to be a three-host tick, no details are available on its life history or capacity to transmit pathogenic organisms.

Distribution. The only records for I. gregsoni in Canada are from Nova Scotia (not shown in map 28), Ontario and Manitoba. Additional collecting at the appropriate time of the year will undoubtedly expand its known range.

Hosts. This tick is known primarily from mustelids: weasels, mink, and marten.

Map 28. Collection localities for Ixodes gregsoni in Canada. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 205

Ixodes (Scaphixodes) howelli Cooley and Kohls (Figs. 61–62) Ixodes howelli Cooley and Kohls, 1938a: 1616. Ixodes howelli Cooley and Kohls: Kohls 1947: 59 (male, nymph, larva). Ixodes (Scaphixodes) howelli Cooley and Kohls: Keirans and Clifford 1978: 76.

Adult. Female: Body: Length about 2.7–2.9 mm unfed, reaching to 7.4 mm engorged. Gnathosoma: Porose areas of basis capituli expansive, narrowly separated by interval of about a third the width of one porose area, their posterior edges near posterior margin of basis (Fig. 61A). Dorsal posterior margin of basis capituli slightly sinuous, with salient corners instead of cornua. Auriculae formed as blunt triangular lateral projections continuous with ridge onto dorsolateral surface (Fig. 61B). Anterior margin of basis capituli slightly bulged along slope on either side of hypostome. Apex of hypostome flattened or notched, dentition 3/3 subapically, then 2/2 to a short, nude, neck at base, lateral denticles larger, more pointed than inner ones (Fig. 61C). Palpi somewhat thickened, club-shaped when viewed dorsally, widest at mid-length (Fig. 61A). Palpal tibiotarsus not visible dorsally. Palpal trochanter with slight ventral projection. Scutum: Length slightly greater than width, subtriangular, widest barely anterior to middle, with posterolateral margins straight or slightly concave (Fig. 61A). Scapulae scarcely formed as salient anterolateral corners of scutum. Lateral carinae absent. Cervical grooves distinct, diverging, and fading before reaching posterolateral margins. Surface sparsely punctate. Legs: Coxa I with internal spur small, blunt, shorter than external spur (Fig. 61B). Coxae II–IV lacking internal spurs. Coxae I–IV each with prominent, bluntly triangular external spur. Ventral coxal surfaces with moderately numerous, smooth setae. Trochanters I–III each with small, bluntly pointed posteroventral spur (Fig. 61B). Legs long, slender.

Adult. Male: Body: Length 3.0–3.5 mm. Gnathosoma: Basis capituli dorsal posterior margin concave, with narrowly rounded corners instead of cornua; dorsal face with a transverse ridge (Fig. 61D). Ventrally, anterior margin of basis capituli sloped on either side of hypostome. Hypostome formed as a plate, apically pointed and with a median ventral carina; dentition rudimentary (Fig. 61G). Palpi somewhat thickened in dorsal view, with asymmetrically bluntly pointed, upturned tips (Fig. 61F). Dorsally, palpal femur and genu fused. Palpal tibiotarsus not visible dorsally. Palpal trochanter indistinctly delineated, lacking flanges. Scutum: Scapulae prominent, blunt, lateral carinae absent (Fig. 61D). Surface slightly rugose on postscapular areas, and with small, scattered puncta. Surface lacking pair of pit-like depressions midlaterally, but with paired grooves there. Spiracular plate: Irregularly oval, without distinctively longer axis. Body venter: Caudal margins of ventral plates not lobe-like, but with dense fringe of short, thick setae extending anterolaterally to spiracular plates (Fig. 61E, insert). Delineation between median and adanal plates poorly defined or absent. Anal plate distinctly delineated, not widened posteriorly. Ventral plates faintly punctate. Legs: Legs long, slender. Coxa I with internal spur weakly formed, slightly smaller than external spur; coxa I also with small spur anterodorsal to external spur (Fig. 61E). Coxae II–IV lacking internal spurs, or II with faint spur or salient posterior corner. Coxae I–IV each with a blunt external spur of similar size. Ventral coxal surfaces with moderately numerous, scattered, short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs (Fig. 61E).

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli slightly sinuous, with salient corners instead of cornua (Fig. 62H). Auriculae formed as large, bluntly triangular, 206 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A B C

D E G

Figs. 61A–G. Ixodes howelli. Fig. 61A, female gnathosoma and scutum, dorsal; Fig. 61B, female gnathosoma, coxae and trochanters, ventral; Fig. 61C, female hypostome, ventral; Fig. 61D, male, gnathosoma and idiosoma, dorsal; Fig. 61E, male, gnathosoma, idiosoma, coxae and trochanters, ventral; insert enlarged setae; Fig. 61F, male palp, lateral; Fig. 61G, male hypostome, ventral. Figs. 61A, B redrawn from Cooley and Kohls (1945); Figs. 61C–G redrawn from Kohls (1947). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 207

H I J

K L

Figs. 62H–L. Ixodes howelli. Fig. 62H, nymph gnathosoma and scutum, dorsal; Fig. 62I, nymph gnathosoma, coxae and trochanters, ventral; Fig. 62J, larva hypostome, palp trochanter, basis capituli, ventral; Fig. 62K, larva, dorsal; Fig. 62L, larva, ventral. Figs. 62H, I redrawn from Kohls (1947); Figs. 62J redrawn, K, L modified from Webb et al. (1990). 208 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins lateral projections (Fig. 62I). Anterior margin of basis capituli slightly bulged along slope on either side of hypostome. Apex of hypostome slightly flattened or notched, dentition below coronal area 2/2 to a short, nude, neck at base; lateral denticles larger, more pointed than inner ones. Two pairs of post-hypostomal setae. Palpi somewhat thickened, club- shaped when viewed dorsally, widest at mid-length (Fig. 62H). Palpal tibiotarsus not visible dorsally. Palpal trochanter lacking flanges. Scutum: Length slightly less than width, broadly oval or subrhomboidal, widest at midlevel, with posterolateral margins nearly straight (Fig. 62H). Scapulae rudimentary as anterolateral corners of scutum. Lateral carinae absent. Cervical grooves diverging and fading before reaching posterolateral margins. Surface sparsely punctate. Legs: Coxa I with internal spur small, blunt, shorter than external spur (Fig. 62I). Coxae II–IV lacking internal spurs (Fig. 62I). Coxae I–IV each with prominent, bluntly triangular external spur, these progressively smaller from I–IV. Ventral coxal surfaces with moderately numerous, smooth setae. Trochanters I–III sometimes each with rudiments of a posteroventral spur (Fig. 62I). Legs long, slender.

Larva. Body: Body length, including gnathosoma, approximately 0.9 mm, slightly engorged. Gnathosoma: Dorsal posterior margin of basis capituli nearly straight, with salient, bluntly pointed corners projecting laterally instead of cornua (Fig. 62K). Ventrally, auriculae formed as large, triangular projections directed posterolaterally (Fig. 62J). Apex of hypostome broadly rounded, dentition below coronal area 2/2 nearly to base; lateral file with approximately 6 overlapping teeth, inner file with 5 teeth; lateral denticles larger, more pointed than inner denticles (Fig. 62J). Two pairs of post-hypostomal setae (Figs. 62J, L). Palpi thick; femur+genu length to width ratio about 1.8–2.0:1; ventral apex of femorogenu lacking a tubercle or flange. Palpal trochanter distinct, lacking flanges (Fig. 62J). Scutum: Length about 0.75 times width, widest at, or slightly behind, mid-level (Fig. 62K). Scapulae undeveloped; shield posterior margin convex. Surface micropunctate, and entirely reticulate. Setae, 4 pairs, lacking posterolateral pair. Idiosomal setation (Figs. 62K, L): Central dorsal setae, 4 or 5 pairs. Supplementary dorsal setae, 2 pairs. Marginal dorsal setae, 7 or 8 pairs. Marginal ventral setae, 3 or 4 pairs. Premarginal ventral setae, 4 pairs. Preanal ventral setae, 3 pairs, sometimes plus 1 or 2 pairs in postanal region. Length of anterior marginal dorsal setae about twice that of scutal setae. Posteriormost pairs of ventral marginal and premarginal setae similar in size to their dorsal counterparts. Sternal setae similar to one another in small size. Legs: Coxae I and II with blunt internal spur (Fig. 62L); coxa III with linear ridge posteroventrally. Coxae I–III each with blunt external spur, similar or slightly smaller in size on I and II to internal spur on coxa I, but smaller, rounder on III. Coxae I–II–III setal formula 3–2–2; coxal setae smooth. Femur III with 10 or 11 setae (i.e., with 6 setae subapically). Trochanters I and II each with small, rounded, posteroventral protuberance apically (Fig. 62L), this lacking on trochanter III.

Natural history. There is very little information on the natural history of I. howelli. Most references are to its distribution and host associations (e.g., Kohls and Ryckman 1962). Kohls (1947) provided the most detailed information available, from a colony of cliff swallows in California. The cliff swallow, Petrochelidon pyrrhonota, may be a primary host, but it has been recorded from other species of birds (Keirans and Clifford 1978; Furman and Loomis 1984). Larvae, nymphs, and females were found feeding mainly on the head and neck region of nestlings, but smaller numbers were found elsewhere on the body. Most males were collected clinging to females; since no males were found attached to a host, it was assumed they do not feed. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 209

Distribution. Ixodes howelli has been recorded in Alaska, Montana, California, and Colorado. There are no known records for it in Canada, though it is likely that it occurs in British Columbia and perhaps in the foothills of Alberta.

Hosts. All recorded hosts for this tick are passeriform and falconiform birds.

Ixodes (Scaphixodes) signatus Birula (Fig. 63, Map 29) Ixodes signatus Birula, 1895: 357–358. Ixodes arcticus Osborn, 1899: 553. Ixodes parvirostris Neumann, 1901: 284. Ixodes eudyptidis signata Neumann, 1904: 451. Ceratixodes signatus (Birula): Banks 1908: 21. Ixodes signatus Birula: Nuttall and Warburton 1911: 263 (nymph, larva). Ixodes signatus Birula: Gregson 1954: 275 (male). Ixodes (Scaphixodes) signatus Birula: Keirans and Clifford 1978: 118.

Adult. Female: Body. Length reaching to 8.0 mm engorged. Gnathosoma: Porose areas of basis capituli expansive, separated only by a narrow median ridge, or sometimes confluent medially, posterior edges of porose areas near posterior margin of basis (Fig. 63A). Dorsal posterior margin of basis capituli slightly sinuous, flared laterally over edges of coxae I, with posterolateral corners instead of cornua (Fig. 63A). Auriculae formed as shelf-like lateral extensions continuous with ridge onto dorsolateral surface (Fig. 63D). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome broadly rounded, dentition 3/3 subapically, then 2/2 to base, lateral denticles larger, more pointed than inner denticles (Fig. 63B). Palpi moderately short, with lateral profile margin slightly convex, thickened, somewhat club-shaped when viewed dorsally, widest at midlength (Fig. 63A). Palpal tibiotarsus not visible dorsally (Fig. 63C). Palpal trochanter with a ventral ridge but no projection. Scutum: Length about 1.25–1.35 times greater than width, oval, widest at about mid-length, with posterolateral margins slightly convex (Fig. 63A). Scapulae scarcely formed as salient anterolateral corners of scutum. Lateral carinae absent. Cervical grooves distinct, diverging and fading before reaching posterolateral margins. Surface ornamentation variable; sometimes nearly smooth with distinct puncta, sometimes with large sparse puncta amidst numerous fine puncta, sometimes with irregular, parallel, or networked rugae. Legs: Coxa I with internal spur reduced to a salient posterior corner, shorter than external spur (Fig. 63D). Coxae II–IV lacking internal spurs, but II, sometimes III, with salient posterior corners. Coxae I–IV each with bluntly triangular external spur, progressively smaller from I–IV. Ventral coxal surfaces with few, short, smooth setae. Trochanters I–IV lacking spurs (Fig. 63D). Legs long, slender.

Adult. Male: Body: Length approximately 3.3 mm. Gnathosoma: Basis capituli dorsal posterior margin slightly sinuous, with obtusely rounded corners instead of cornua; dorsal face without furrows. Ventrally, anterior margin of basis capituli sloped on either side of hypostome. Hypostome formed as a plate, truncate apically and with a median ventral carina; dentition rudimentary, with about 6 files (Fig. 63J, insert). Palpi irregularly thickened in dorsal view, with asymmetrically flattened apices. Dorsally, palpal femur and genu fused, with several stout spine-like setae on femoral area (Fig. 63I, insert). Palpal tibiotarsus arising terminally from palp, slightly visible in dorsal view (Fig. 63J). 210 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

B D E

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Figs. 63A–M. Ixodes signatus. Fig. 63A, female gnathosoma and scutum, dorsal; Fig. 63B, female hypostome, ventral; Fig. 63C, female palp tibiotarsus; Fig. 63D, female gnathosoma, coxae and trochanters, ventral; Fig. 63E, nymph gnathosoma and scutum, dorsal; Fig. 63F, nymph gnathosoma, coxae and trochanters, ventral; Fig. 63G, male gnathosoma and idiosoma, dorsal; Fig. 63H, male, gnathosoma, idiosoma, coxae, ventral; Fig. 63I, male gnathosoma, ventral; insert, palp, lateral; Fig. 63J, male gnathosoma, dorsal; insert, hypostome, ventral; Fig. 63K, larva, dorsal; Fig. 63L, larva, ventral; Fig. 63M, larva hypostome, palp trochanter, basis capituli, ventral. Figs. 63A–F redrawn from Cooley and Kohls (1945); Figs. 63G–J redrawn from Gregson (1956); Fig. 63K–L redrawn, L modified from Webb et al. (1990). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 211

Palpal trochanter weakly delineated, with ventral ridge. Scutum: Scapulae rounded (Fig. 63G). Lateral carinae faint. Surface with numerous puncta, more so on posterior two- thirds. Surface lacking pair of pit-like depressions midlaterally, but with paired grooves there. Spiracular plate: Subcircular, without distinctively longer axis. Body venter: Caudal margins of ventral plates not lobe-like, lacking fringe of long setae. Delineation between median and adanal plates indistinct (Fig. 63H). Anal plate distinctly delineated, not widened posteriorly. Ventral plates faintly punctate anteriorly, becoming increasingly punctate posteriorly. Legs: Legs long, slender. Coxae I–IV lacking internal spurs, but I, II with salient posterior corners (Fig. 63H). Coxae I–IV each with a small, blunt external spur of similar size. Ventral coxal surfaces with few, scattered, short, smooth or slightly barbed setae. Trochanters I–IV lacking spurs.

Nymph. Gnathosoma: Dorsal posterior margin of basis capituli slightly sinuous, flared somewhat laterally over edges of coxae I, with posterolateral corners instead of cornua (Fig. 63E). Auriculae formed as shelf-like lateral extensions continuous with ridge onto dorsolateral surface (Fig. 63F). Anterior margin of basis capituli sloped on either side of hypostome. Apex of hypostome broadly rounded. Hypostome dentition 3/3 for a few subapical rows, then 2/2 nearly to base. Lateral denticles larger, more pointed than inner ones. Two pairs of post-hypostomal setae. Palpi moderately short, thickened, somewhat club-shaped when viewed dorsally, widest at mid-length (Fig. 63E). Palpal tibiotarsus not visible dorsally. Palpal trochanter with a slight ventral ridge or projection. Scutum: Length about 1.3 times greater than width, oval, widest at about mid-length, with posterolateral margins slightly convex (Fig. 63E). Scapulae scarcely formed as salient anterolateral corners of scutum. Lateral carinae absent. Cervical grooves distinct, diverging and fading near posterolateral margins. Surface generally densely covered with fine puncta. Legs: Coxa I with internal spur reduced to a salient posterior corner, shorter than external spur (Fig. 63F). Coxae II–IV lacking internal spurs. Coxae I–IV each with bluntly triangular external spur, progressively smaller from I–IV. Ventral coxal surfaces with few, short, smooth setae. Trochanters I–IV lacking spurs (Fig. 63F). Legs long, slender.

Larva. Body: Body length 0.5–0.6 mm. Gnathosoma: Dorsal posterior margin of basis capituli nearly straight, with bluntly pointed corners instead of cornua (Fig. 63K). Ventrally, auriculae undeveloped or weakly formed as lateral bulges (Fig. 63M). Apex of hypostome broadly rounded or slightly indented, dentition 3/3 on apical third, then 2/2 to base; lateral file with 8–10 teeth, inner file with 6–8; lateral denticles larger, more pointed, and overlapping than inner denticles (Fig. 63M). Two pairs of post-hypostomal setae (Figs. 63L, M). Palpi thick; femur+genu length to width ratio about 1.8–2.0: 1; ventral apex of femorogenu with a salient, seemingly double-edged, tubercle or flange (barely evident in Fig. 63L). Palpal trochanter distinct, lacking flanges (Fig. 63M). Scutum: Length nearly (0.85–0.99 times) equal to width, widest at or slightly anterior to mid-level (Fig. 63K). Scapulae undeveloped; shield posterior margin convex. Surface finely micropunctate, and with underlying reticula becoming emergent anterolaterally. Setae, 4 pairs, lacking second of 2 anterolateral pairs. Idiosomal setation (Figs. 63K, L): Central dorsal setae, 5 pairs. Supplemental dorsal setae, 1 or 2 pairs. Marginal dorsal setae, 7 or 8 pairs. Marginal ventral setae, 3 pairs. Premarginal ventral setae, 5 pairs. Preanal ventral setae, 2 pairs. Length of anterior marginal dorsal setae scarcely longer than scutal setae. Posteriormost pair of marginal ventral and premarginal ventral setae about twice as long as any posterior dorsal setae. Sternal setae similar to one another in small size. Legs: Coxa I with short, 212 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins broadly rounded internal spur or salience (Fig. 63L). Coxae II and III each with curved ridge posteroventrally. Coxae I–III each with blunt external spur, progressively smaller from I–III. Coxae I–II–III setal formula 3–2–2; coxal setae short, smooth. Femur III with 9 setae. Trochanters I–III lacking spurs (Figs. 63L).

Natural history. Information on Ixodes signatus in Canada is quite limited. There are relatively few records and specimens reported in the literature (Gregson 1956; Spencer 1960; Schwan and Kelly 1981; Hoberg and Wehle 1982). Throughout its range, it seems to have a lengthy life cycle, tied in with the short time that its hosts are present on their nests. Each instar of the tick has the opportunity to feed only when the sea birds are nesting. The natural history of I. signatus was described by Asanuma and Fukuda (1957) in nests of black-tailed gulls, Larus crassirostris Vieillot, in Japan, where all instars were present in the nest at all times (as also reported in California by Furman and Loomis (1984)), and each instar fed during the breeding season, to moult subsequent to the fledging of the chicks and departure of the birds from the colony. As a result, it typically required at least three years to complete a generation, though unfed ticks are able to survive long periods without feeding. There are many arboviruses associated with sea bird ticks, and I. signatus is no exception. Two viruses have been isolated from I. signatus in Alaska (Yunker 1975; Chastel 1988). The impact of these viruses on their hosts and the potential for transmission to other animals and humans has not been determined.

Distribution. Being almost entirely restricted to Pacific sea birds as hosts, this species has been recorded in coastal areas of western North America from California to the Aleutian Islands of Alaska, Japan, and the Kamchatka Peninsula of eastern Russia. All records for Canada are from British Columbia.

Hosts. Sea birds, especially cormorants, are the most important hosts for this tick, but there is one unusual record of a passeriform bird being infested (Gregson 1956).

Map 29. Collection localities of Ixodes signatus in Canada. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 213

Genus Rhipicephalus Koch (Fig. 64) Anus of nymphs and adults with postanal groove. Eyes present (Figs. 64A, G, J). Posteromarginal festoons present (Figs. 64H–K). Palpus with 2nd palpomere (femur) not appreciably longer than wide, not acutely produced laterally (Fig. 64E). Basis capituli in dorsal view subhexangular, with angulate lateral sides (Figs. 64E, G). Adult females and males with usually inornate scutum. Foveae present. Adult males with 1 or 2 pairs of ventral plates (Fig. 64I). Coxa I of adults appearing bifid, with similarly elongate and adjacent internal and external spurs (Fig. 64E). Coxae I–IV similar in size. Larvae with 1 pair of posthypostomal setae; scutum with 3 pairs of setae; 8 pairs of marginal dorsal setae, including 4 pairs anterior to sensilla sagittiformia; 2 pairs of central dorsal setae (Fig. 64J).

Rhipicephalus (Rhipicephalus) sanguineus (Latreille) Brown dog tick (Fig. 64, Map 30) Ixodes sanguineus Latreille, 1806: 157. Rhipicephalus sanguineus (Latreille): Koch 1844: 238. Rhipicephalus rutilus Koch, 1844: 235. Rhipicephalus limbatus Koch, 1844: 236. Rhipicephalus siculus Koch, 1844: 239. Rhipicephalus rubicundus Frauenfeld, 1867: 463. Rhipicephalus punctatissimus Gerstaecker, 1873: 481. Rhipicephalus stigmaticus Gerstaecker, 1873: 496. Rhipicephalus becarii Pavesi, 1883: 496. Rhipicephalus sanguineus (Latreille): Neumann 1897: 385. Rhipicephalus texanus Banks, 1908: 34. Rhipicephalus sanguineus (Latreille): Hooker et al. 1912: 102 (all instars described). Rhipicephalus sanguineus (Latreille): Cooley 1946c: 24 (all instars described).

The taxonomic status of R. sanguineus is considered by some to be uncertain (Guglielmone et al. 2014). These authors note that it is not possible to assign a definitive name to any population of this species worldwide. Rhipicephalus sanguineus sensu lato is probably a complex involving a number of species, some of which are cryptic. Here we refer to R. sanguineus in the broadest taxonomic sense.

Adult. Female: Body: Length 2.4–2.7 mm unfed, reaching to 11.5 mm engorged. Nine festoons. Gnathosoma: Porose areas of basis capituli small, oval, separated by interval of at least twice the width of one porose area (Fig. 64A). Basis capituli hexagonal in dorsal view, with triangulate lateral projections; posterior margin nearly straight, with cornua as salient corners (Fig. 64A). Hypostome slightly clavate, rounded apically; dentition 3/3, denticles about equal in size, with 9 or 10 in each file (Fig. 64B). Palpi short, thick, with femur and genu similar in length (Fig. 64A). Palpal trochanter in ventral view with subtriangular projection on inner surface; inner lateral edges of palpal trochanter, genu, and femur fringed with conspicuous, curved, parallel, barbed setae (Fig. 64E, insert). Scutum: Length about 1.1 greater than width, oval, widest at mid-length (Fig. 64A). Scapulae prominent, narrowly rounded. Cervical grooves deep anteriorly, fading posteriorly before reaching posterolateral margins. Lateral carinae faint. Surface 214 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

D E

A C

sensilla sagittiforme

H I K

Figs. 64A–K. Rhipicephalus sanguineus. Fig. 64A, female gnathosoma and scutum, dorsal; Fig. 64B, female hypostome, ventral; Fig. 64C, male hypostome, ventral; Fig. 64D, nymph hypostome, ventral; Fig. 64E, female gnathosoma, coxae and trochanters, ventral; insert enlarged palpal seta; Fig. 64F, nymph gnathosoma, coxae and trochanters, ventral; Fig. 64G, nymph gnathosoma and scutum, dorsal; Fig. 64H, male gnathosoma and idiosoma, dorsal; Fig. 64I, male gnathosoma, idiosoma, coxae and trochanters, ventral; insert enlarged palpal seta; Fig. 64J, larva, dorsal; Fig. 64K, larva, ventral. Figures A–I redrawn from Cooley (1946). Figure J–K redrawn from Clifford et al. (1961). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 215

inornate, with mingled large and small puncta, more numerous in median and posterior areas. Eyes distinct, situated at about mid-length of scutum (Fig. 64A). Legs: Coxa I deeply cleft, with elongate internal and external spurs, with opposing margins slightly divergent; internal spur much broader than pointed external spur (Fig. 64E). Coxae II and III with internal spurs formed as broad, short saliences. Coxae II–IV with short, blunt, external spurs (Fig. 64E).

Adult. Male: Body: Length, 1.86–2.70 mm. Nine festoons. Gnathosoma: Basis capituli hexagonal in dorsal view, its posterior margin more concave than in female (Fig. 64H). Hypostome slightly clavate, similar to female but smaller; dentition 3/3, with 7 or 8 teeth in each file (Fig. 64C). Palpus formed as in female, its trochanter in ventral view with a distinct, inner plate-like projection; inner lateral edges of palpal trochanter, genu, and femur lined with curved, barbed setae as in female (Fig. 64I, insert). Scutum: Scapulae long, blunt, cervical grooves deep, short, not reaching to level of eyes (Fig. 64H). Marginal grooves starting near eyes, terminating after delimiting anteriormost pair of festoons. Festoons about as long as wide. Surface covered with small puncta intermingled with larger pits. Body venter: Adanal plates distinct in flanking anal plate medially and accessory (epimeral) plates laterally, variable in form but slightly widened posteriorly where more salient than adjacent plates (Fig. 64I). Legs: Coxa I deeply cleft, with internal and external spurs formed as in female (Fig. 64I). Coxae II and III with salient inner corners, but coxa IV with a short, blunt internal spur. Coxae II–IV with short, blunt external spurs as in female.

Nymph. Body: Length, unengorged, tip of palpi to posterior margin, 1.14–1.30 mm. Nine festoons. Gnathosoma: Basis capituli hexagonal in dorsal view with triangulate lateral projections as in adult; posterior margin nearly straight, with salient corners (Fig. 64G). Hypostome slightly clavate, rounded apically; dentition 2/2, with 6 or 7 teeth in each file (Fig. 64D). Palpal apex narrowly rounded apically. Palpal trochanter visible in ventral view as rounded lobe bearing two long barbed setae on inner side. Scutum: Nearly (0.91–0.94 times) as long as broad, shape suboval, widest slightly behind midlevel (Fig. 64G). Scapulae prominent, narrowly rounded. Surface sparsely, inconspicuously punctate. Legs: Coxa I with 2 short, separated spurs, external spur slightly longer, more pointed than internal spur (Fig. 64F). Coxae II–IV lacking internal spurs. Coxae II and III each with a short, blunt external spur, decreasing in size from II–III; coxa IV with external spur rudimentary or absent (Fig. 64F).

Larva. Body: Nine festoons (Fig. 64J). Gnathosoma: Basis capituli in dorsal view subhexagonal, with bluntly triangular lateral margins; posterior margin lacking cornua (Fig. 64J). Ventrally, auriculae absent (Fig. 64K). Apex of hypostome slightly rounded. Hypostome dentition 2/2, with 5–6 similar-sized teeth per file. Palpi with bluntly flared lateral margins, tapering distally. Palpal genu ventrally with small, blunt spur, and long, barbed setae on inner side. Scutum: Length about two-thirds (0.65 times) width, widest at level of large eyes, well posterior to midlength (Fig. 64J). Scapulae rounded; shield posterior margin broadly convex. Surface sparsely, inconspicuously punctate. Idiosomal setation (Figs. 64J–K): Marginal dorsal setae, 8 pairs, including 4 pairs anterior to sensilla sagittiformia. Marginal ventral setae, 5 pairs. Legs: Coxa I with short, broadly rounded spur (Fig. 64K). Coxae II and III each with small rounded spur on posteromedial margin; coxae I–III lacking external spurs (Fig. 64K). 216 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Map 30. Collection records for Rhipicephalus sanguineus in Canada.

Natural history. It is unlikely that this tick may become established in the wild in Canada, but there are numerous reports of it having become established in sheltered environments, including dog kennels and even inside people’s homes. It readily feeds on humans as well as dogs. Rhipicephalus sanguineus is a pest of major importance worldwide. It is perhaps the most widely distributed species of tick (but see below). Apart from damage and irritation caused by its bite, it is a vector of numerous pathogens, some of the most important of which are canine ehrlichiosis (Ehrlichia canis), canine babesiosis (Babesia canis vogeli), and canine hepatozoonosis (Hepatozoon canis (James)) in dogs, and tick bite fever (Rickettsia conorii) in humans. No pathogens have been isolated yet from R. sanguineus in Canada.

Distribution. Rhipicephalus sanguineus is an Old World tick thought to have originated in Africa, but now found throughout the world, having successfully dispersed along with humans and their dogs. It is not currently possible to assign a definitive name to any population of this species because R. sanguineus, as conventionally known, may in fact be a complex involving Rhipicephalus turanicus Pomerantsev and probably various other cryptic species (Guglielmone et al. 2014). Most records in Canada are the result of people returning from visits abroad, so it may be possible to find this tick in any region of Canada.

Hosts. Dogs are almost exclusively recorded as hosts for this tick, but humans or other animals that are closely associated with dogs may accidentally acquire them.

Family Argasidae Idiosoma lacking a dorsal scutum in nymphs and adults (Fig. 5), but sometimes with a small mid-dorsal shield in larva (Fig. 6A); gnathosoma (capitulum) placed anteroventrally and covered by anterior margin of idiosoma in nymphs and adults (Figs. 4A, B), though projecting anteriorly and fully visible from above in larva (Figs. 6A, B); idiosomal integument of adult and nymph leathery and pebbled or spiny (Figs. 5, 81), though finely striated and smooth in larva; spiracular plates located lateral to coxae III or IV in nymph and adults (Figs. 4A, B); leg coxae lacking spurs. Larva with apical palpomere shorter but similar in cylindrical shape to other segments, inserted terminally and readily visible in dorsal view (Figs. 72A, B). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 217

Genus Argas Latreille (Figs. 65–69, Plate I) Gnathosoma: Adult and nymphs: Hypostome well developed, denticulate, about as long as palpi (Figs. 67A–D). Larva: Palpal femur with 5 setae, including pd-1.

Idiosoma: Adult and nymphs: Body margin with a suture delineating dorsal and ventral surfaces (Figs. 66A–D); unengorged body margin thin and acute; body integument wrinkled and mammillate, without spines (Plate I); body oval or elliptical in dorsal view, sides not constricted; anterior margin of body narrowly rounded, somewhat produced. Larva: Dorsal setae, 23–36 pairs centrally and marginally (Figs. 68A, B). Anal valves surrounded by 2 or 3 pairs of setae, 1 of which preanal. Pair of respiratory organs discernible between bases of coxae I and II.

Legs: Adult and nymphs: Dorsal surfaces of leg I tibia and tarsus smooth, without humps other than subapical tarsal hump. Larva: Genu I with 8 or 9 setae, including 4 dorsal, 2 anterolateral, 2 ventral (Fig. 65). Tibia I with 8 setae, of which 4 dorsal.

Key to the Species of Argas in Canada

1. Adults and nymphs: dorsal integument with peripheral ridge of small rounded tubercles with scattered setae arising from indistinct pits or interstices (Figs. 66A, C); marginal rim of idiosoma with compressed structure giving coarsely striated appearance; postpalpal setae absent (Figs. 67A, B); larva: dorsum of idiosoma with one pair of central setae anterior and lateral to mid-length of mid-dorsal plate (Fig. 68A); tarsus I with a trumpet-shaped sensillum extending posteriorly within capsule of Haller’s organ (Fig. 69A); femora I and II each with 8 setae, each lacking av-2; genu I with 8 setae, lacking al-2...... Argas (Argas) cooleyi – Adults and nymphs: dorsal integument with peripheral ridge of large, rectangular cells, each with a distinct pit from which arises a seta (Figs. 66B, D); marginal rim of idiosoma with uncompressed structure giving quadrangular appearance; pair of long, postpalpal setae present (Figs. 67C, D); larva: dorsum of idiosoma with three or four pairs of central setae anterior and lateral to midlength of mid-dorsal plate (Fig. 68B); tarsus I without a trumpet-shaped sensillum extending posteriorly within Haller’s organ (Fig. 69B); femora I and II each with 9 setae, including av-2 (Fig. 65); genu I with 9 setae, including al-2 (Fig. 65) ...... Argas (Persicargus) persicus

Argas (Argas) cooleyi Kohls and Hoogstraal (Figs. 66A, C, 67A, B, 68A, 69A, Map 31) Argas reflexus: Gregson 1956: 24 (not Fabricius 1794). Argas (Argas) cooleyi Kohls and Hoogstraal, 1960: 625 (all instars).

Adult. Female and male: Body: Female length, unengorged, 4.3–8.2 mm; male length, 3.7–6.2 mm. Gnathosoma: Apex of hypostome slightly flattened, dentition 2/2 along apical half, with approximately 3 or 4 prominent teeth per file, then 3/3 increasing to 5/5 along basal half, with small, weakly formed teeth (Fig. 67A). Postpalpal setal pair absent (Fig. 67A). Body integument: Dorsal integument with peripheral row of small rounded tubercles with scattered setae arising from indistinct pits or interstices (Figs. 66A, C). Dorsal integument 218 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins with irregularly patterned mammillae; ridges smooth, forming a convergent pattern among discs anteriorly; discs moderately large, arranged in radiating rows. Ventral integument similar to that on dorsal surface. Anus bearing approximately 10 setae on each valve. Legs: Tarsi II–IV each with a slightly elevated dorsal subapical protuberance.

Nymph. Similar to adult, but lacking genital aperture (Fig. 67B).

Larva. Body: Length, excluding gnathosoma, unengorged 0.60–0.80 mm. Gnathosoma: Apex of hypostome bluntly rounded or slightly indented. Hypostome dentition 2/2, both rows extending most of shaft length; lateral row with 6 or 7 teeth, inner row with 5 teeth. Basis capituli ventrally with 1 pair of setae, post-hypostomal (postpalpal pair absent). Body integument: Dorsal surface with approximately 32 pairs of setae, including 8 anterolateral pairs, 1 pair beside dorsal plate, 9 postero-mediolateral pairs and 14 posterolateral pairs (Fig. 68A, insert). Legs: Tarsus I with internal, elongate, trumpet-shaped sensillum extending posteriorly from capsule of Haller’s organ (Fig. 69A). Femur I with 8 setae, lacking av-2; genu I with 8 setae, lacking al-2 (compare with fuller set of setae in Fig. 65).

Natural history. Although this species of tick has been recorded in and around the nests of a variety of species of birds, it is most frequently associated with the cliff swallow (Petrochelidon pyrrhonota) (Cook 1972; Howell and Chapman 1976). Cliff swallows are colonial and build clay pots as their nests on cliff faces and various non-natural structures including bridges and buildings, sometimes in enormous numbers. These swallows are present at their colonies for only about three months each year, so A. cooleyi is adapted to the short period of host availability and long periods of their absence. Unfed ticks are negatively phototactic, so they remain hidden in cracks and crevices outside the nests during the day (Howell 1976; George 1987). An aggregation pheromone may play a role in bringing ticks together in harbourages (George 1981) where, in a dense aggregation, they may conserve energy and reduce water loss. When light level falls at night, ticks begin to wander over the nests and respond to various cues to locate their host (Howell 1975). Feeding is completed in about 15 minutes and engorged ticks retreat in the crevices surrounding the nests to digest their blood-meals. George and Cook (1979) and Howell and George (1973) have investigated the influence of various environmental factors on behaviour and physiology of A. cooleyi.

Map 31. Collection record for Argas cooleyi in Canada. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 219

With only one record for A. cooleyi in Canada, and that from an eastern bluebird (Sialia sialis (Linnaeus)), the status of this species as a resident in Canada requires investigation. Argas cooleyi has been reported as a possible vector for a variety of arboviruses of birds in North America (Yunker et al. 1977; Calisher et al. 1988).

Distribution. Argas cooleyi is found primarily in western North America, with one record as far east as South Dakota. In Canada, we have just one record from the interior of British Columbia.

Hosts. This tick is primarily a parasite of the cliff swallow; however, the one Canadian record comes from the nest of an eastern bluebird.

Argas (Persicargus) persicus (Oken) Fowl tick or Persian poultry argas (Figs. 65, 66B, 67C, 68B, 69B, Plate I, Map 32) Rhynchoprion persicum Oken, 1818: 1567. Argas persicus: Fischer von Waldheim, 1823: 282. Argas mauritianus Guérin-Méneville, 1844: 17. Argas chinche Goudot: Gervais, 1844: 462. Argas americana Packard, 1872: 740. Argas persicus (Oken): Kohls et al. 1970: 591 (all instars).

Adult. Female and male: Body: Female length, unengorged, 5.1–9.8 mm; male length 4.7–6.7 mm. Gnathosoma: Apex of hypostome slightly flattened or notched (Fig. 67C). Hypostome dentition 2/2 along apical third, with approximately 3 or 4 prominent teeth per file, followed by several irregular files of minute denticles extending to basal end of shaft. Postpalpal setal pair present, about as long as post-hypostomal pair (Fig. 67C). Body integument: Dorsal integument with peripheral row of approximately 90 relatively large, subrectangular cells, each with conspicuous setiferous pit (Figs. 66B, D); dorsal integument anteriorly with mammillae fewer, smaller, less elevated than those posterior of midlength; ridges forming irregular patterns among discs anteriorly; discs moderately large, arranged in radiating rows (Plate I). Ventral integument similar to that on dorsal surface. Anus bearing approximately 8 setae on each valve. Legs: Tarsi II–IV without dorsal subapical protuberances.

Nymph. Similar to adult, but with fewer setae and lacking genital aperture (Fig. 67D).

Larva. Body: Length, including gnathosoma, unengorged, 0.95–0.98 mm. Gnathosoma: Apex of hypostome bluntly rounded. Hypostome dentition 3/3 on anterior third, then 2/2 to base; lateral file with 9 or 10 teeth, second file with 6–9 teeth. Basis capituli ventrally with 2 pairs of setae, 1 post-hypostomal, 1 postpalpal. Body integument: Dorsal surface with 26–29 pairs of setae, including 7 anterolateral pairs, usually 2 or 3 pairs beside subquadrangular dorsal plate, 9 postero-mediolateral pairs and 8 posterolateral pairs (Fig. 68B). Legs: Tarsus I lacking internal trumpet-shaped sensillum extending from capsule of Haller’s organ (Fig. 69B). Femur I with 9 setae, including seta av-2 (Fig. 65); genu I with 9 setae, including al-2 (Fig. 65).

Natural history. This species is very widely distributed as a result of dispersal by humans, and infests poultry houses where it may become a serious pest. It is found most abundantly in warmer parts of the world, and is limited in occurrence in Canada, where it was Gregson’s (1956) belief that it would be unlikely to become permanently established. 220 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

a1 pd1 p1 a1 ad1 pd1 av1 pv p1 av tibia ad1 pv tibia pd2 ad2 ad1 pd2 al1 ad1 a1 av ad1 p1 p1 pd1 pv av pv al2 ad1 genu

pd2 ad2 pd2 ad2 genu a1 ad1 p1 pd1 av1 p1 av1 pd1 pv ad1 pv femur av2 femur ad2 ad2 pd2 a1 p12 av2 pd2

II I

Fig. 65. Argas. Larva, notation for setae on middle segments of legs I (left) and II (right). Modified from Edwards (1975). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 221

1 mm

B (enlarged)

A D (enlarged)

Figs. 66A–D. Argas adult idiosoma patterns. 66A, C, A. cooleyi. Dorsal integument (A) and peripheral ridge (C). Figs. 66B, D, A. persicus. Dorsal integument (B) and peripheral ridge (D). Figs. 66A, C redrawn from Kohls and Hoogstraal (1960); Figs. 66B, D redrawn from Kohls et al. (1970). 222 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A C postpalpal seta

B D

Figs. 67A–D. Argas ventral gnathosoma attributes. 67A, B, A. cooleyi. A, adult, B, nymph; 67C, D. A. persicus. C, adult, D, nymph. Figs. 67A, B redrawn from Kohls and Hoogstraal (1960); Figs. 67C, D redrawn from Kohls et al. (1970). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 223

A B

Figs. 68A, B. Argas larva dorsal idiosomas; inserts, enlarged marginal seta. A, A. cooleyi. B, A. persicus. Fig. 68A redrawn from Kohls and Hoogstraal (1960); Fig. 68B redrawn from Kohls et al. (1970). 224 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

D A

0.02 mm 0.02 mm

A B

Figs. 69A, B. Argas larvae, sensillum of tarsus I. A, A. cooleyi. B, A. persicus. Initials: D – dorsal; A – anterior. Redrawn from Kaiser et al. (1964). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 225

Plate I. Argas persicus. Upper – dorsal view; lower – ventral view. 226 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Map 32. Collection locality for Argas persicus in Canada.

There is a vast literature on the fowl tick, and a brief summary is provided here. Adult ticks are nocturnal and feed repeatedly from their hosts. Females lay relatively small numbers of eggs following each blood meal, usually in sheltered areas away from the host. Eggs hatch after several weeks, depending on temperature, and the tiny larvae attach to their host and feed up to a week or more. They drop from the host and retreat to cracks or crevices away from the host where they moult to the first instar nymph after a week or more, depending on the temperature. The first-instar nymph repeats the process and moults to the second-instar nymph. This instar feeds, and then moults to the adult instar after a week or more. Adults mate, and females lay eggs, completing a generation in about four months, though under high temperatures and ready host availability, a generation can be completed in as little as five to six weeks. (Lancaster and Meisch 1986). Argas persicus can be present in sufficient numbers to cause death in poultry as a result of blood loss (Hoogstraal 1985), and larvae can cause a form of paralysis in domestic chickens (Rosenstein 1976). They also have the capacity to transmit a number of pathogens, in particular Borrelia anserina, Aegyptianella pullorum Carpano (Rickettsiales: Anaplasmataceae), Mycobacterium avium Chester (Actinomycetales: Mycobacteriaceae), and Pasteurella avicida (Prévot) (Pasteurellales: Pasteurellaceae) (Hoogstraal 1985).

Distribution. Argas persicus is an Old World species that has become widely distributed around the world in association with humans and poultry husbandry. We have only one record of this species in Canada, in British Columbia.

Hosts. This tick feeds primarily on birds, and in some parts of the world may be an important pest of domestic poultry. In Canada, we know of only one record from a passerine bird.

Genus Carios Latreille (Figs. 70–72, Plate II) There is disagreement on the higher classification of argasid ticks, and the genusCarios is not recognised in the latest checklist of the ticks of the world (Guglielmone et al. 2010). We have followed Klompen and Oliver (1993) in their recognition of the Carios group. Carios has been recognised as a subgenus in both the so-called “Soviet school” and “American school” of classifications of argasids, with citations given by Klompen and Oliver (1993). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 227

Gnathosoma: Adult and nymphs: Hypostome well developed, denticulate, shorter to nearly as long as palpi. Larva: Palpal femur with 4 setae, lacking pd-1.

Idiosoma: Adult and nymphs: Body margin with a suture delineating dorsal and ventral surfaces; unengorged body margin thin and acute; body integument wrinkled and mammillated, without spines (Plate II); body oval or elliptical in dorsal view, sides not constricted; anterior margin of body narrowly rounded, somewhat produced (Plate II). Larva: Dorsal setae, 10–15 pairs centrally and marginally. Anal valves surrounded by 2 or 3 pairs of setae, 1 of which preanal; unpaired postanal seta present. Pair of respiratory organs rudimentary between bases of coxae I and II.

Legs: Adult and nymphs: Dorsal surfaces of leg I tibia and tarsus micromammillate, without humps other than subapical tarsal hump. Larva: Genu I with 9 setae, including 4 dorsal, 2 anterolateral, 2 ventral. Tibia I with 8 setae, of which 4 dorsal (as in Fig. 65).

Key to the Species of Carios in Canada

1. Adults and nymphs: hypostome slightly widened distal to midlength (Fig. 70B); palpi with accessory ventral setae on femur (> 4 setae) and genu (> 5 setae) (Fig. 71B); adult cheeks oval, wider anteriorly; larva: hypostome dentition of 3/3 along distal half, then 2/2 to base (Fig. 72B); hypostome arising from short, subtriangular median extension of basis capituli (Fig. 72B)...... Carios concanensis – Adult and nymphs: hypostome sides subparallel, not widened distally (Fig. 70A); palpi without accessory ventral setae on femur (4 setae) and genu (5 setae) (Fig. 71A); adult cheeks reniform, elongate, not wider anteriorly; larva: hypostome dentition of 3/3 limited to distal third (Fig. 72A); hypostome arising abruptly from large, conical, median extension of basis capituli (Fig. 72A)...... Carios kelleyi

Carios concanensis (Cooley and Kohls) (Figs. 70B, 71B, 72B, Map 33) Ornithodoros concanensis Cooley and Kohls, 1941: 910 (adult, nymph). Ornithodoros aguilae Cooley, 1944: 287. Ornithodoros (Alectorobius) concanensis Cooley and Kohls: Kohls et al. 1965: 340 (larva).

Adult. Female and male: Body: Female length, unengorged, approximately 5.5 mm; male length, approximately 4.2 mm. Body suboval, sides nearly parallel. Hood short, elevated apically. Cheeks oval; anterior ends wider and free. Gnathosoma: Hypostome appreciably widened apically beyond midlength, notched apically, dentition of 2/2 limited to distal third, with about 4 teeth per file (as in Fig. 70B for nymph). Palpi hypertrichous, with accessory ventral setae on trochanter, femur, genu (as in Fig. 71B for nymph). Body integument: Mammillae large, close but not crowded; usually with 1–2 circular or crescentic pits on top; some pits with minute seta. Discs depressed on dorsum with elevated edges. Discs on venter linearly arranged in preanal and median postanal grooves. Legs: Surfaces micromammillate. Tarsus I with mild subapical dorsal protuberance; dorsal humps absent on all tarsi.

Nymph (Figs. 70B, 71B). Similar to adult. 228 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Larva. Body: Length, including gnathosoma, unengorged, 0.76–0.80 mm. Gnathosoma: Hypostome: arises directly from a small subtriangular extension of basis capituli, and tapers to pointed apex, dentition 3/3 along distal half of shaft, then 2/2 to base; denticles numerous, fine; lateral file with 18–20 denticles, midlateral file with 16–18 denticles, inner file with 10–12 denticles (Fig. 72B). Body integument: Dorsal plate, pyriform, widest posteriorly, less than one-third as long as idiosoma. Dorsum with 16-19 (typically 17) pairs of setae, including 12–14 lateral pairs and 4 or 5 sub-central pairs.

Natural history. Carios concanensis has rarely been reported in Canada, but Wilkinson et al. (1980) were convinced that it was established in Alberta and Saskatchewan, though they were uncertain whether breeding there or being carried into the area on migrating birds. This species is primarily a parasite in the nests of cliff-dwelling passerines and bats in the western United States of America (Cook 1972; Easton 1983; Knight and Marr 1983), so its prevalence on prairie falcons in Canada is not unexpected where this host nests on cliffs. It also infests cavity-nesting owls in Texas (Proudfoot et al. 2006). Not much has been published on the life cycle of the tick, but it is known that there is a non-feeding first-instar nymph (Davis 1942). There is evidence that these ticks produce a pheromone (George 1981) that allows them to aggregate in cracks in the cliff face, from which they locate their hosts at night using audio cues (Webb et al. 1977). Prairie falcons are sometimes heavily infested (Williams 1947), and Oliphant et al. (1976) attributed the deaths of prairie falcon nestlings to heavy infestations of C. concanensis. Carios concanensis is known to bite humans (Kohls et al. 1965), but not to transmit pathogens to them. It is thought to be a vector for Babesia moshkovskii (Schurenkova) Laird and Lari (Piroplasmida: Babesiidae) among prairie falcons in Wyoming (Croft and Kingston 1975).

Distribution. The distribution of C. concanensis is limited to western North America, from Alberta to California and Texas. We have no records for this tick outside of southcentral Alberta.

Hosts. This tick has been found in bat caves, and a variety of birds have been recorded as hosts, but records from Canada are all from prairie falcons.

Map 33. Collection localities for Carios concanensis in Canada. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 229

Carios kelleyi (Cooley and Kohls) Bat tick (Figs. 70A, 71A, 72B, Plate II, Map 34) Ornithodoros kelleyi Cooley and Kohls, 1941: 912. Ornithodoros kelleyi Cooley and Kohls: Sonenshine 1962: 470 (all instars). Ornithodoros (Alectorobius) kelleyi: Clifford et al. 1964: 433. Carios kelleyi (Cooley and Kohls): Horak et al. 2002: 32.

Adult. Female and male. Body: Female length, unengorged, 5.7–7.4 mm; male length 4.2–5.3 mm. Body suboval, sides nearly parallel. Hood small, developed as small projection of body wall over gnathosoma (Plate II). Cheeks elongate (nearly as long as gnathosoma), reniform with anterior portion free. Gnathosoma: Hypostome with subparallel sides, truncate and notched apically, dentition 2/2 along apical half of shaft, with about 4 teeth per file (as in Fig. 70A for nymph). Palpi not hypertrichous, lacking ventral accessory setae on palpomeres (chaetotaxy 7-12-8-6/7) (as in Fig. 71A for nymph). Body integument: Mammillae moderately large, close, but not crowded. Discs large, conspicuous, concentrated in depressed areas, including a central craterlike cluster on dorsum (Plate II). Discs on venter concentrated in linear arrangement in preanal and transverse postanal grooves and in 3 depressions posterior of transverse postanal groove (Plate II). Legs: Surfaces micromammillate. Tarsus I with subapical dorsal protuberance weakly formed. Legs I–IV lacking dorsal humps on tarsal and other segments.

Nymph (Figs. 70A, 71A). Similar to adult.

Larva. Body: Length, including gnathosoma, unengorged, 1.0–1.1 mm (of which gnathosoma at least half of length, 0.6 mm). Gnathosoma: Hypostome arises abruptly from large, conical, median extension of basis capituli and tapers to pointed apex, dentition 3/3 along distal third, then 2/2 to base; denticles numerous, fine; lateral file with 16–18 denticles, midlateral file with 15–18 denticles, and inner file with 5–8 denticles (Fig. 72A). Body integument: Dorsal plate pyriform, widest posteriorly, prominent, at least half as long as idiosoma. Dorsum with 16 or 17 pairs of setae, including 12 or 13 lateral pairs and 4 more or less sub-central pairs.

Natural history. This tick has been reported in Alberta and Saskatchewan in Canada (Gregson 1956; Wilkinson et al. 1980; Lausen 2005), but there is little information on its natural history in Canada. However, it has been the subject of detailed study in the United States of America (e.g., Sonenshine and Anastos 1960; Sonenshine 1962, 1970; Sonenshine and Gregson 1970). Sonenshine and Anastos (1960) reared C. kelleyi in the laboratory, where a number of details on the life history were determined. Larvae typically feed for an extended period on their hosts, requiring up to about three weeks for engorgement. Larvae moulted to the first-instar nymph 2–6 days after one feeding. There may be two to four nymphal instars, with at least one engorgement between instars, except for the first-instar nymph. The first-instar nymph was not observed to feed, but rather moulted to a second nymphal instar in 8–16 days. Nymphs feed much faster than larvae, usually engorging and dropping off in less than two hours. Second- instar nymphs occasionally moulted directly to the adult, but all were males. Other second-instar nymphs moult to a third-instar nymph, which then feeds before moulting to either an adult or a fourth instar nymph. Fourth-instar nymphs feed and then moult to 230 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A B

Figs. 70A, B. Carios nymph hypostomes, ventral. A, C. kelleyi; B, C. concanensis. Redrawn from Cooley and Kohls (1944).

genu

femur

A B

Figs. 71A, B. Carios nymph hypostome and palps, ventral. A, C. kelleyi; B, C. concanensis. Redrawn from Cooley and Kohls (1944). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 231

Extension of hypostome

A B

Figs. 72A, B. Carios larva gnathosomas, ventral. A, C. kelleyi; B, C. concanensis. Redrawn from Kohls et al. (1965).

al1 pd1 pl1

av1 pv1

ad1 tibia

al1 pd1 pl1

av1 pv1 genu ad1

pd1 al1 av1 pv1 femur ad1

ad2 pd2 av2

III

Fig. 73. Ornithodoros larva, notation for setae on middle segments of leg III. Redrawn from Edwards (1975). 232 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Plate II. Carios kelleyi. Upper – dorsal view; lower – ventral view. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 233

adults, which are always females. Males and females feed repeatedly, usually engorging within two hours. Mating takes place off the host, in the bat roost, and may take up to about an hour. Completion of the life cycle in the laboratory required 54–258 days. Mating was observed before and after feeding, but the pattern in the field is not known. Carios kelleyi is often collected from the roosts of bats, but Lausen (2005) found that prevalence of infestation was highest (about 17%) on big brown bat (Eptesicus fuscus (Palisot de Beauvois)) roosting in rock crevices in southern Alberta from June through September, considerably greater than on western small-footed bat (Myotis ciliolabrum Merriam) (1%), little brown bat (Myotis lucifugus (LeConte)) (1.2%), and long-eared bat (Myotis evotis (Allen)) (0.7%). Intensity of infestation was 4.0 ticks per infested bat, with a maximum of 44 ticks on one host. The presence of ticks on bats late in October in southern Alberta led Lausen (2005) to conclude that this tick overwinters in Canada. Carios kelleyi feeds readily on other small-mammal hosts, which in turn may provide an important alternate source of blood for ticks inhabiting rock crevices in the absence of bats (Lausen 2005). There is evidence from the United States of America that C. kelleyi will feed on humans, sometimes with deleterious consequences (Gill et al. 2004). A number of potential pathogens have been isolated from C. kelleyi, including Rickettsia, Borrelia, and Bartonella (Loftis et al. 2005; Reeves et al. 2006). Schwann et al. (2009) found spirochaetes (Borrelia johnsonii Schwan et al.) in pooled samples of C. kelleyi from Iowa, and in the coxal fluid produced by one male, but the status of this species as a vector for this pathogen has not been demonstrated. Bowles et al. (2013) considered its vector potential for pathogens to humans to be low. No pathogens have been isolated from C. kelleyi in Canada.

Distribution. Carios kelleyi is widely distributed in North America, mostly in the United States of America (Bowles et al. 2013). So far, it has been recorded from Saskatchewan and Alberta in Canada, but its close proximity to the northern tier of states in the United States of America, makes it likely to be found elsewhere on bats.

Hosts. Bats are predominant hosts of this tick, and in Canada, five species of bats have been found to be infested.

Map 34. Collection localities for Carios kelleyi in Canada. 234 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Genus and subgenus Ornithodoros (Pavlovskyella) Pospelova-Shtom (Figs. 74–77, Plate III)

Gnathosoma: Adult and nymphs: Hypostome well developed, denticulate, shorter to nearly as long as palpi (Figs 75A, B). Larva: Palpal femur with 4 setae, lacking pd-1.

Idiosoma: Adult and nymphs: Body margin thick, rounded, without a suture delineating dorsal and ventral surfaces; body integument wrinkled and mammillate, without spines; body oval or elliptical in dorsal view, sides nearly parallel or slightly constricted, and with dorsoventral groove usually evident behind legs IV (Figs. 4A, B, Plate III); anterior margin of body narrowly or broadly rounded, somewhat produced or not. Larva: Dorsal setae, 7–16 pairs centrally and marginally (Figs 76A, B). Anal valves surrounded by 2 or 3 pairs of setae, 1 of which preanal; unpaired postanal seta present or absent (Figs. 77A, B). Pair of respiratory organs absent between bases of coxae I and II.

Legs: Adult and nymphs: Dorsal surfaces of leg I tibia and tarsus mammillate or wrinkled, often with humps in addition to subapical tarsal one (O. hermsi excepted) (Figs. 74A, B). Larva: Genu I with 5 setae, including 2 dorsal, 1 anterolateral, 1 ventral. Tibia I with 6 setae, of which 2 dorsal (compare with fuller set of setae in Fig. 65).

Key to the species of Ornithodoros in Canada

1. Adults and nymph: leg I without dorsal humps on tibia and tarsus (except for subapical dorsal protuberance on tarsus) (Fig. 74A); hypostome slightly notched apically, dentition 2/2 restricted to apical third of length, each file with 3 or 4 teeth of similar size (Fig. 75A); larva: mid-dorsal plate present (Fig. 76A); venter of idiosoma lacking unpaired postanal seta (Fig. 77A); genu III with 5 setae, pv-1 absent...... Ornithodoros hermsi – Adults and nymph: leg I with three or four dorsal humps on each of tibia and tarsus (in addition to subapical dorsal protuberance on tarsus) (Fig. 74B); hypostome truncate apically, dentition 2/2 along apical half of length, each file with 5–7 teeth of similar size (Fig. 75B); larva: mid-dorsal plate absent (Fig. 76B); venter of idiosoma with unpaired postanal seta (Fig. 77B); genu III with 6 setae, pv-1 present (Fig. 77C)...... Ornithodoros parkeri

Ornithodoros (Pavlovskyella) hermsi Wheeler, Herms, and Meyer (Figs. 74A, 75A, 76A, 77A, Plate III, Map 35) Ornithodoros hermsi Wheeler, Herms, and Meyer, 1935: 1290 (all instars). Ornithodoros hermsi: Wheeler 1935: 435 (male, female). Ornithodoros (Pavlovskyella) hermsi Wheeler, Herms, and Meyer: Kohls et al. 1965: 352 (larva).

Adult. Female and male: Body: Female length, 4.5–5.4 mm; male length 3.4–3.7 mm. Body oval, sides nearly parallel. Hood well developed (Plate III). Cheeks absent. Gnathosoma: Hypostome notched apically, dentition of 2/2 limited to distal third of shaft, with 3 or 4 teeth of similar size per file (Fig. 75A). Body integument: Mammillae numerous, moderate in size, elongate, not conical; longer axes of mammillae longitudinal in lateral areas, but tending to be concentric around discs or groups of discs in median areas (Plate III). Discs shallow. Dorsoventral groove weakly extended onto dorsolateral surface (Plate III). Legs: Surfaces smooth, not micromammillate. All tarsi without dorsal humps but each with mild subapical dorsal protuberance (Fig. 74A). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 235

Nymph. Similar to adult.

Larva. Body: Length, including gnathosoma, unengorged, 0.65–0.76 mm. Gnathosoma: Hypostome with sides nearly parallel, not arising from median extension of basis capituli, apically truncate or slightly notched. Hypostome dentition 2/2 restricted to apical third or half of shaft, lateral file with 4 or 5 teeth, inner file with 2–4 teeth of similar size. Post- hypostomal setal pair arising from a prominence. Body integument: Dorsal plate pyriform, widest posteriorly, about one-third as long as idiosoma (Fig. 76A). Dorsum with 13 pairs of setae, including 11 lateral pairs and 2 sub-central pairs. Venter lacking unpaired postanal seta (Fig. 77A). Legs: Leg III genu with 5 setae, lacking seta pv-1, which is shown in Fig. 77C).

Natural history. There is little specific information on the natural history of O. hermsi. Gregson (1956) provided the most relevant information, summarised here. This tick feeds readily on mammals, but it was Gregson’s (1956) opinion that birds were important in its maintenance and dispersal in British Columbia. He found that juvenile instars fed readily on a variety of small mammals including bats; they typically engorged within 30 minutes and dropped off, but some remained attached for a slightly longer period. Larvae moult after their blood-meal, but nymphs took several blood-meals before moulting to the adult. Adults feed repeatedly, the female depositing a small number of eggs after each blood-meal. Despite the importance of small mammals and birds as hosts for this tick, it readily feeds on humans (Furman and Loomis 1984). Gregson (1956) described the response to the bite as sometimes progressing from a typical wheal, to a severe form of transitory shock. In the United States of America, O. hermsi is known to transmit the spirochaete that causes relapsing fever in humans, Borrelia hermsii (Philip and Davis 1940).

Distribution. Ornithodoros hermsi is found widely through the northwestern United States of America, south to California, and has been recorded in many locations in the interior of British Columbia.

Hosts. This tick infests a number of small mammals and passeriform birds and is also known to feed on humans.

Map 35. Collection localities for Ornithodoros hermsi in Canada. 236 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A tibia tarsus

Figs. 74A, B. Ornithodoros adult, apical segments of leg I. A, O. hermsi; B, O. parkeri. Redrawn from Cooley and Kohls (1944).

A B

Figs. 75A, B. Ornithodoros adult hypostomes, ventral. A, O. hermsi; B, O. parkeri. Redrawn from Cooley and Kohls (1944). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 237

A B

Figs. 76A, B. Ornithodoros larva idiosoma, dorsal. A, O. hermsi; B, O. parkeri. Redrawn from Kohls et al. (1965).

A B

Figs. 77A, B. Ornithodoros larva. A, O. hermsi, idiosoma, ventral; B, O. parkeri, idiosoma, ventral. Redrawn from Kohls et al. (1965). 238 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Plate III. Ornithodoros hermsi. Upper – dorsal view; lower – ventral view. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 239

Ornithodoros (Pavlovskyella) parkeri Cooley (Figs. 74B, 75B, 76B, 77B) Ornithodoros parkeri Cooley, 1936: 431 (adults). Ornithodoros wheeleri McIvor, 1937: 365. Ornithodoros (Pavlovskyella) parkeri Cooley: Kohls et al. 1965: 354 (larva).

Adult. Female and male: Body: Female length, 5.8–7.1 mm; male length, 5.0–5.8 mm. Body oval, sides nearly parallel. Hood prominent, not continuous with body wall of dorsum. Cheeks absent. Gnathosoma: Hypostome with sides about parallel, with blunt, flattened apex, dentition 2/2 along distal half of shaft, with 5–7 teeth of similar size per file (Fig. 75B). Body integument: Mammillae numerous, moderately small and dense, not elongate, slightly larger along sides, largest at posterior end. Discs small, depressed. Dorsoventral groove visible on body sides and onto dorsolateral surface (as in Fig. 5). Legs: Surfaces smooth, not micromammillate or granulate. Tarsi I–III each with moderate subapical dorsal protuberance, small or absent on tarsus IV. Leg I tarsus and tibia each with three or four dorsal humps (Fig. 74B).

Nymph. Similar to adult.

Larva. Body: Length, including gnathosoma, unengorged, 0.78–0.86 mm. Gnathosoma: Hypostome with sides nearly parallel, not arising from median extension of basis capituli, apically truncate or slightly notched, dentition 2/2 restricted to apical third or half of shaft, lateral file with 5–7 teeth, inner file with 4 or 5, lateral file slightly larger. Post-hypostomal setal pair not arising from a prominence. Body integument: Dorsal plate absent (Fig. 76B). Dorsum with 13 pairs of setae, including 10–12 (usually 11) lateral pairs and 2 sub- central pairs. Venter with unpaired postanal seta (Fig. 77B). Legs: Leg III genu with 6 setae, pv-1 present (Fig. 77C).

Natural history. The natural history of O. parkeri has been described by Davis (1941) in the United States of America under laboratory conditions. All instars feed relatively quickly on a variety of small mammals, larvae moulting to the first nymphal instar in 1–4 weeks. There may be as many as five nymphal instars, each requiring engorgement before moulting to the next instar. Occasionally, second-instar nymphs moult to adults, but all turn out to be males. Third- instar and fourth-instar nymphs may moult to become either males or females, or a small number of fourth-instar nymphs may moult to become females. Mating takes place off the host, and requires an average of 28 minutes. Davis (1941) found that unfed late-instar nymphs and adults could survive in the absence of a host for a long period of time, up to four years. The spirochaete Borrelia parkeri has been isolated from O. parkeri in the United States of America (Davis 1939; Gage et al. 2001), but its relationship to human disease has not been established.

Distribution. Ornithodoros parkeri is widely distributed in the western United States of America, from Idaho and Utah to California. It has not yet been found in Canada, but is anticipated to occur in southern parts of British Columbia and Alberta where its common hosts occur.

Hosts. This tick is associated with burrowing mammals (there are also records from the burrowing owl) and lagomorphs. It is also known to feed on humans. 240 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Genus Otobius Banks (Figs. 78–84, Plate IV)

Gnathosoma: Hypostome of adult vestigial, without denticles, clearly shorter than palpi (Plate IV); hypostome of nymphs well developed, denticulate, nearly as long as palpi (Figs. 82A, B). Larva: Palpal femur with 3 setae, lacking al and pd-1.

Idiosoma: Adult and nymphs: Body margin thick, rounded, without a suture delineating dorsal and ventral surfaces. Body integument of adult granular, pitted, without spines (Plate IV); that of nymphs spinose, finely striated (Figs. 79A–D). Body pear-shaped or violin-shaped in dorsal view (adults less so than nymphs), sides somewhat constricted (Fig. 80A); anterior margin of body broadly rounded, not produced. Larva: Dorsal setae, 7–11 pairs centrally and marginally (Figs. 83A, B). Anal valves flanked by only 1 pair of setae, none preanal in position; unpaired postanal seta absent. Pair of respiratory organs rudimentary between bases of coxae I and II.

Legs: Adult and nymphs: Dorsal surfaces of leg I tibia and tarsus smooth, without humps other than subapical tarsal hump. Larva: Genu I with 6 setae, including 2 dorsal, 1 anterolateral, 2 ventral. Tibia I with 6 setae, of which 2 dorsal (compare with fuller set of setae in Fig. 65).

Key to the species of Otobius in Canada

1. Adults: dorsal idiosomal integument with pits separated by two or more times their diameters (Fig. 78A); idiosoma constricted at level of legs IV (Fig. 80A, arrow); nymphs: dorsal idiosomal integument with spines thicker anteriorly than posteriorly (Figs. 79A, B); idiosoma at level of legs III–IV strongly constricted posteriorly (Fig. 81A); hypostome dentition mostly 4/4 (Fig. 82A, arrow); larva: idiosoma with two pairs of eyes dorsolaterally (Fig. 83A); mid-dorsal face of tarsus I with row of four adjacent “parascapular” setae (two short setae flanked by a longer seta on either side) inserted on proximal edge of Haller’s organ (Fig. 84A)...... Otobius megnini – Adults: dorsal idiosomal integument with pits separated by no more than their diameters (Fig. 78B); idiosoma indistinctly constricted at level of legs IV (Fig. 80B); nymphs: dorsal idiosomal integument with spines similar in size anteriorly and posteriorly (Figs. 79C, D); idiosoma at level of legs III–IV slightly constricted posteriorly (Fig. 81B); hypostome dentition mostly 3/3 (Fig. 82B, arrow); larva: idiosoma without eyes (Fig. 83B); mid-dorsal face of tarsus I with row of three adjacent “parascapular” setae (one short seta flanked by a longer seta on either side) inserted on proximal edge of Haller’s organ (Fig. 84B)...... Otobius lagophilus

Otobius lagophilus Cooley and Kohls (Figs. 78B, 79C, D, 80B, 81B, 82B, 83B, 84B, Map 36) Otobius lagophilus Cooley and Kohls, 1940: 928 (female, male, nymph). Otobius lagophilus Cooley and Kohls: Kohls et al. 1965: 361 (larva). Ornithodoros megnini (Dugès, 1883): Hadwen 1913: 80; Hewitt 1915: 225; Hearle 1938: 349; Brown 1944: 49 (not Ornithodoros megnini Dugès, 1884).

Adult. Female and male: Body: Female length, 5.4-6.3 mm; male length 4.7–5.0 mm. Body rounded at both ends, indistinctly constricted at level of legs IV (Fig. 80B). Hood and camerostome not apparent. Gnathosoma: Hypostome vestigial, without denticles; sides A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 241

converging towards blunt or notched apex. Body integument: Integument granular with numerous pits, each bearing a minute seta; pits separated by distance subequal to diameter of 1 pit (Fig. 78B); pattern continuous dorsoventrally, but less prominent on venter. Mammillae absent. Legs: Tarsi I–IV each with moderate subapical dorsal protuberance, more pronounced on tarsus IV.

Nymph. Body: Similar to adult in size and shape, but more tapered posteriorly when unfed (Fig. 81B). Hood and eyes present (Fig. 81B). Gnathosoma: Hypostome with lateral margins nearly parallel, apex notched. Hypostome dentition mostly 3/3 amidst some 4/4 along nearly entire length, with 6 or 7 sharp, similar teeth per file (Fig. 82B).Body integument: Surface smooth, shiny, with fine reticulations and transverse striae; discs absent. Slender spines all over body surface; longer and more abundant anteriorly on dorsum, progressively smaller and sparser toward posterior end; less numerous on venter (Figs. 79C, D). Legs: Subapical dorsal protuberance absent or small on tarsi I–III, but distinct on tarsus IV.

Larva. Body: Length, including gnathosoma, unengorged, 0.65–0.71 mm. Eyes absent (Fig. 83B). Gnathosoma: Hypostome prominent, about as long as palpi, with apex flattened (Fig. 83B), dentition 2/2 along entire length of shaft, lateral and inner files each with 10– 12 teeth. Body integument: Dorsal plate with posterior margin nearly as broadly rounded as anterior margin (Fig. 83B). Dorsum with 7–9 pairs of setae, including 5–7 lateral pairs and 2 sub-central pairs (Fig. 83B). Venter lacking both preanal pairs of setae and unpaired postanal seta. Legs: Mid-dorsal face of tarsus I with row of three adjacent “parascapular” setae (1 short seta flanked by a longer seta on either side) inserted on proximal edge of Haller’s organ (Fig. 84B).

Natural history. The first Canadian record for this species appears in Hadwen (1913), in his report as pathologist in charge of the Veterinary Research Laboratory in Agassiz, British Columbia. He briefly described what he calledOtobius megnini occurring on jack rabbits in the Lethbridge area. He noted that they were attached to the muzzle of their hosts, rather than in the ears, the site characteristic for O. megnini. Otobius lagophilus was described much later, in 1940, and today is still only known in Canada from southern Alberta (Brown and Kohls 1950), though its range extends south and west in the United States of America. There has been no research on the natural history of O. lagophilus in Canada, and what is summarised here is drawn from work in the United States of America. The larva crawls on the ground or in low vegetation seeking a suitable host, primarily lagomorphs. Many larvae were found by Loomis (1953) around the entrance of host burrows. When they attain a host, they attach around the muzzle near the vibrissae, on the throat, lower jaw, ears and neck, and occasionally on the back all the way to the base of the tail (Hopla 1955; Bacha 1957; Herrin 1965). The larvae engorge and moult in place to the nymphal instar. Nymphs continue to feed and moult only once to the adult after dropping from the host (Bacha 1957; Herrin and Beck 1965). The adults have no functional mouthparts; females may begin laying eggs two months after they have mated and lay several hundred eggs, up to 655 eggs reported by Bacha (1957). Females lay their eggs in interrupted batches, taking about 7–10 days to complete each batch and they may deposit at least three or four batches (Hopla 1955, temperature not reported). Larvae require about 11 days to hatch (Hopla 1955). In southern parts of its range, O. lagophilus may be found at all times of the year, but there is no information on its seasonal occurrence in Canada. 242 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Map 36. Collection locality for Otobius lagophilus in Canada.

Because O. lagophilus feeds as larvae and nymphs on the same animal, and adults do not feed at all, their potential as vectors of pathogenic organisms is limited, unless a pathogen is transovarially transmitted. Philip et al. (1955) provided evidence of infection with Rickettsia rickettsii in rabbits in the laboratory when they placed ticks collected from wild hares on them. In the same study in Nevada, they found evidence of Colorado tick fever virus in O. lagophilus (Philip et al. 1955). Eklund et al. (1955) and Silva-Goytia and Elizondo (1952) also reported associations of R. rickettsii with this tick in Utah and Mexico, respectively. Despite retaining Francisella tularensis for nearly two years in the laboratory, there was no evidence of transovarial transmission (Furman and Loomis 1984). There are no records of these agents associated with O. lagophilus in Canada.

Distribution. Otobius lagophilus is confined to western North America, from Alberta to Mexico.

Hosts. Jack rabbits are the primary hosts for this tick, though pikas have been recorded as hosts in the United States of America, and predators, such as cats, have also been found infested.

Otobius megnini (Dugès) Spinose ear tick (Figs. 78A–B, 80A, 81A, 82A, 83A, 84A, Plate IV, Map 37) Argas megnini Dugès, 1883: 197. Otobius megnini Banks, 1912: 99. “Argas americana Packard”: Townsend 1893: 50. Rhynchoprium spinosum Marx, 1895: 199. Ornithodoros megnini (Dugès): Neumann 1896: 42. Ornithodoros (Otobius) megnini Dugès: Hoffman 1930: 151. Otobius megnini Banks: Cooley and Kohls 1944: 21 (all instars). Otobius megnini Banks: Kohls et al. 1965: 362 (larva).

Adult. Female and male: Body: Female length, 5.0–8.2 mm; males slightly smaller. Body panduriform, rounded at both ends, constricted just behind leg IV (Plate IV, Fig. 80A). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 243

Hood short, broad. Camerostome lined with numerous fine, long setae. Gnathosoma: Hypostome vestigial, without denticles, with sides converging towards blunt or notched apex. Body integument: Integument granular with numerous pits, each with a minute seta; pits separated by intervals of twice diameter of 1 pit (Fig. 78A); pattern continuous dorsoventrally, but less distinct ventrally (Plate IV). Mammillae absent. Legs: Tarsi I–IV each with subapical dorsal protuberance, inconspicuous on tarsus I, moderate on II–III, more pronounced on IV.

Nymph. Body: Prevalent second instar nymph similar to adult in size and shape, but more narrowed posteriorly when unfed (Fig. 81A); first-instar nymph smaller, more slender than second instar. Hood and camerostome absent. Eyes absent (Fig. 81A). Gnathosoma: Hypostome well developed, with sides converging towards blunt or flattened apex, dentition mostly 4/4 amidst some 3/3 along nearly entire length, with 7 or 8 sharp, similar teeth per file (Fig. 82A). Body integument: Surface smooth, shiny, with fine reticula or striae continuous over depressed areas (counterparts of discs). Surface spinose over nearly entire body except around gnathosoma and on disc–depressions; spines coarser, with thicker bases, on dorsum and venter extending anteriorly from spiracles, progressively slenderer posteriorly (Fig. 79A, B). Legs: Subapical dorsal protuberance absent or small on tarsi I–IV.

Larva. Body: Length, including gnathosoma, unengorged, 0.66–0.72 mm. Two pairs of prominent, well-spaced eyes present (Fig. 83A). Gnathosoma: Hypostome prominent, slightly shorter than palpi, with apex flattened (Fig. 83A). Hypostome dentition 2/2 along entire length of shaft, lateral and inner files each with 10–12 teeth.Body integument: Dorsal plate with posterior margin clearly more narrowly rounded than anterior margin (Fig. 83A). Dorsum with 9–11 pairs of setae, including 6–8 lateral pairs and 3 subcentral pairs (Fig. 83A). Venter lacking both preanal pairs of setae and unpaired postanal seta. Legs: Mid-dorsal face of tarsus I with row of four adjacent “parascapular” setae (2 short setae flanked by a longer seta on either side) inserted on proximal edge of Haller’s organ (Fig. 84A).

Natural history. The spinose ear tick occurs only in British Columbia in Canada where it has been recorded infesting primarily deer, mountain goats, and mountain sheep (Gregson 1956). Elsewhere in Canada, this tick is occasionally encountered as the result of accidental introduction, but it is unlikely that it may become permanently established. In the western United States of America, this species can be a serious pest of domestic cattle, sheep, and horses. Because the parasitic instars attach deep inside the ear canal and feed for long periods of time, they are easily overlooked. This is no doubt part of the reason this tick has been accidentally introduced from its native North America into Central and South America, Africa, and India, where it has become an important pest. Free-living larvae gain access to suitable passing hosts (Rich and Gregson 1968) and then move into the ear canal where they attach and begin to feed. They moult to a first- instar nymph followed by a second-instar nymph, both of which remain imbedded and feeding in the ear canal. When the second instar nymph has finished feeding, it leaves the ear canal and drops to the ground, where it moults to the adult. Adults are not parasitic and do not feed. They mate and females deposit their eggs in sheltered areas on the soil surface or in cracks and crevices in surrounding rocks. The parasitic phase of the life cycle can take up to three months; development from egg to adult was completed in 62–118 days under laboratory conditions (Loomis 1961). 244 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A B

Figs. 78A, B. Otobius adult anterodorsal integuments. A, O. megnini; B, O. lagophilus. Redrawn from Herrin and Beck (1965).

A B

C D

Figs. 79A–D. Otobius nymph dorsal integuments. Figs. 79A, B, O. megnini, A, anterodorsal region; B, posterodorsal region. Figs. 79C, D, O. lagophilus, C, anterodorsal region; D, posterodorsal region. Redrawn from Herrin and Beck (1965). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 245

A B

Figs. 80A, B. Otobius adult idiosomas, dorsal. A. O. megnini, arrow points to idiosomal constriction; B., O. lagophilus. Redrawn from Cooley and Kohls (1944).

A B

Figs. 81A, B. Otobius nymph idiosomas, dorsal. A, O. megnini; B, O. lagophilus. Fig. 81A redrawn from Cooley and Kohls (1944); Fig. 81B redrawn from Herrin and Beck (1965). 246 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

A B

Figs. 82A, B. Otobius nymph gnathosomas, ventral. A, O. megnini; B, O. lagophilus. Redrawn from Herrin and Beck (1965).

A B

Figs. 83A, B. Otobius larva gnathosoma and idiosoma, dorsal. A, O. megnini; B, O. lagophilus. Redrawn from Kohls et al. (1965). A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 247

pdm2 pdm2 absent

A B

Figs. 84A, B. Otobius larvae tarsus I, dorsal. A, O. megnini; B, O. lagophilus. Redrawn from Edwards (1975). 248 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Plate IV. Otobius megnini. Upper – dorsal view; lower – ventral view. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 249

Map 37. Collection localities for Otobius megnini in Canada.

Larvae occasionally attach to humans, and during feeding, can cause considerable irritation and otitis. It is this parasitic instar that physicians detect in the ear canals of recent travellers. Otobius megnini has been implicated in transmission of the agent of Q fever, Coxiella burnetii (Jellison et al. 1948); its role is no doubt limited because it typically feeds on only one host from the time the larva attaches through the nymphal instars. As for Ehrlichia canis in this species (Ewing et al. 1990), there is no evidence for transovarial transmission. A bibliography of information on the spinose ear tick was compiled by Keirans and Pound (2003).

Distribution. Otobius megnini is found throughout western North America, from British Columbia south into Mexico and South America. Because nymphs may remain attached inside the ears of their hosts for a very long time, it is easily transported with infested animals to other parts of the world. Therefore records in Canada outside of British Columbia are not unexpected.

Hosts. This tick prefers to feed on large mammals, including deer, cattle, horses, sheep, and goats. However, it is also occasionally found on smaller mammals, including cats and dogs. It is also known to infest humans. 250 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Acknowledgements

A Handbook to the Ticks of Canada was initiated by members of the Expert Committee on Insect Pests of Animals in 1990–1991. Some of the groundwork for species diagnoses was laid at the time, but the project languished after dissolution of the committee. It was resurrected in 2008; some of the original members of the Expert Committee participated and now see the results of the original commitment to update Gregson’s monograph on the ticks of Canada. We especially thank the Biological Survey of Canada. Without their support and assistance, this project may never have seen the light of day. Patrice Bouchard went beyond the call of duty in serving as our liaison with all parties in the final stages. We thank Tom Naughten (Parks Canada, Winnipeg, retired) who produced the distribution maps; Carol Galloway who spent many hours proof-reading various drafts of the manuscript and modified one map; Petrina Gregson (Kamloops, British Columbia) who provided a variety of photographs of her father, with permission to choose our frontispiece from among them; Owen Lonsdale (Eastern Cereal and Oilseed Research Centre, Canadian National Collection, Agriculture and Agri-Food Canada, Ottawa) who provided logistical support and assistance in preparation of some of the figures in the final stages of publication; Andrew Smith for expert technical editing. We thank Peter G. Mason (Eastern Cereal and Oilseed Research Centre, Canadian National Collection, Agriculture and Agri-Food Canada, Ottawa), one of the members of the Expert Committee in 1990, for his continued support and encouragement. We acknowledge the assistance of Kateryn Rochon as the Microsoft guru for this project, and Donna Giberson for designing the cover. Thanks also to Juliette Nadeau (University of Manitoba Copyright Office) for her advice in obtaining copyright permission for illustrations in the handbook. Copyright permission for reproduction of figures was obtained from the following sources: Western North American Naturalist (Brigham Young University Science Bulletin), The Journal of Vector Ecology, The American Midland Naturalist, The Entomological Society of America (Miscellaneous Publications of the Entomological Society of America, Annals of the Entomological Society of America, Journal of Medical Entomology), and The Journal of Parasitology (Allen Press). Critical financial support was provided by the Biological Survey of Canada, Agriculture and Agri-Food Canada and the Public Health Agency of Canada. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 251

References

Addison, E.M., Johnson, F.J., and Fyvie, A. 1979. Dermacentor albipictus on moose (Alces alces) in Ontario. Journal of Wildlife Diseases, 15: 281–284. Addison, E.M. and McLaughlin, R.F. 1988. Growth and development of winter tick, Dermacentor albipictus, on moose, Alces alces. Journal of Parasitology, 74: 670–678. Addison, E.M. and Smith, L.M. 1981. Productivity of winter ticks (Dermacentor albipictus) collected from moose killed on Ontario roads. Alces, 17: 136–146. Adelson, M.E., Rao, R.S., Tilton, R.C., Cabets, K., Eskow, E., Fein, L., Occi, J.L.,and Mordechsae, E. 2004. Prevalence of Borrelia burgdorferi, Bartonella spp., Babesia microti and Anaplasma phagocytophila in Ixodes scapularis ticks collected in northern New Jersey. Journal of Clinical Microbiology, 42: 2799–2801. Alderink, F.J. and Dietrich, R.A. 1982. Economic and epidemiological implications of anaplasmosis in Texas cattle herds. Annual Meeting of the United States Animal Health Association, 86: 66–75. Allred, D.M., Beck, D.E., and White, L.D. 1960. Ticks of the genus Ixodes in Utah. Brigham Young University Science Bulletin, Biological Series, 1: 1–42. Amin, O.M. 1969a. Growth of the dog tick Dermacentor variabilis Say (Acarina: Ixodidae). I. Growth patterns. Journal of Medical Entomology, 6: 305–316. Amin, O.M. 1969b. Growth of the dog tick Dermacentor variabilis Say (Acarina: Ixodidae). II. The effect of starvation and host species on its growth and fecundity. Journal of Medical Entomology, 6: 321–326. Amin, O.M. 1970. The circadian rhythm of dropping of engorged larvae and nymphs of the American dog tick Dermacentor variabilis Say (Acarina: Ixodidae). Journal of Medical Entomology, 7: 251–255. Amsden, J.R., Warmack, S., and Gubbins, P.O. 2005. Tick-borne bacterial, rickettsial, spirochetal, and protozoal infectious diseases in the United States: a comprehensive review. Pharmacotherapy, 25: 191–210. Anastos, G. 1950. The scutate ticks, or Ixodidae, of Indonesia. Entomologica Americana, 30: 1–144. Anastos, G. and Smith, C.N. 1957. The male, nymph, and larva of Ixodes brunneus Koch (Acarina: Ixodidae). Journal of Parasitology, 43: 535–541. Anderson, J.F. 1988. Mammalian and avian reservoirs for Borrelia burgdorferi. Annals of the New York Academy of Sciences, 539: 180–191. Anderson, J.F. 1989. Epizootiology of Borrelia in Ixodes tick vectors and reservoir hosts. Review of Infectious Diseases, 11: S1451–S1459. Anderson, J.F., Flavell, R.A., Magnarelli, L.A., Barthold, S.W., Kantor, F.S., Wallich, R., et al. 1996. Novel Borrelia burgdorferi from Ixodes scapularis and Ixodes dentatus ticks feeding on humans. Journal of Clinical Microbiology, 34: 524–529. Anderson, J.F. and Magnarelli, L.A. 1980. Vertebrate host relationships and distribution of ixodid ticks (Acari: Ixodidae) in Connecticut, USA. Journal of Medical Entomology, 17: 314–323. Anderson, J.F. and Magnarelli, L.A. 1984. Avian and mammalian hosts for spirochete-infected ticks and insects in a Lyme disease focus in Connecticut. Yale Journal of Biology and Medicine, 57: 627–641. Anderson, J.F., Magnarelli, L.A., LeFebvre, R.B., Andreadis, T.G., McAninch, J.B., Perng, G.-C., and Johnson, R.C. 1989. Antigenically variable Borrelia burgdorferi isolated from cottontail rabbits and Ixodes dentatus in rural and urban areas. Journal of Clinical Microbiology, 27: 13–20. Anderson, J.F., Magnarelli, L.A., Philip, R.N., and Burgdorfer, W. 1986. Rickettsia rickettsii and Rickettsia montana from ixodid ticks in Connecticut. American Journal of Tropical Medicine and Hygiene, 35: 187–191. Anderson, J.F., Magnarelli, L.A., and Sulzer, A.J. 1981. Racoon babesiosis in Connecticut, USA: Babesia lotori sp. n. Journal of Parasitology, 67: 417–425. Anderson J.F., Mintz, E.D., Gadbaw, J.J., and Magnarelli, L.A. 1991. Babesia microti, human babesiosis, and Borrelia burgdorferi in Connecticut. Journal of Clinical Microbiology, 29: 2779–2783. Anderson, R.C. and Lankester, M.W. 1974. Infectious and parasitic diseases and arthropod pests of moose in North America. Naturaliste Canadien (Québec), 101: 23–50. Anonymous. 1996. Tick paralysis – Washington, 1995. Morbidity and Mortality Weekly Report, 45: 325–326. Anstead, C.A. and Chilton, N.B. 2011. Ticks feeding on northern pocket gophers (Thomomys talpoides) in central Saskatchewan and the unexpected detection of Ixodes scapularis larvae. Journal of Vector Ecology, 36: 355–360. Anstead, C.A., Hwang, Y.T., and Chilton, N.B. 2013. Ticks (Acari: Ixodidae) on small mammals in Kootenay National Park, British Columbia, Canada. Journal of Medical Entomology, 50: 1208–1214. 252 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Anstead, C.A. and Chilton, N.B. 2014. Comparison of the DNA sequences and secondary structure of the mitochondrial 16S rRNA gene of Ixodes kingi, Ixodes sculptus and Ixodes angustus. Molecular and Cellular Probes, 28: 155–162. Anstead, C.A., Wallace, S.B., and Chilton, N.B. 2014. Mutation scanning-based identification of larval and nymphal ticks (Acari: Ixodidae) from Richardson’s ground squirrels (Spermophilus richarsonii). Molecular and Cellular Probes, 28: 6–9. Apanaskevich, D.A. 2013. Reinstatement of Dermacentor kamshadalus Neumann (Acari: Ixodidae) as a valid species parasitizing mountain goats and sheep in the United States, Canada, and Russia. Journal of Medical Entomology, 50: 691–700. Appel, M.J. 2005. The Merck veterinary manual, 9th edition. Edited by C.M. Kahne and S. Line. John Wiley & Sons, Whitehouse Station, New Jersey, United States of America. Apperson, C.S., Engber, B., Nicholson, W.L., Mead, D.G., Engel, J., Yabsley, M.J., et al. 2008. Tick-borne diseases in North Carolina: is “Rickettsia amblyommii” a possible cause of rickettsiosis reported as Rocky Mountain spotted fever? Vector-borne and Zoonotic Diseases, 8: 597–606. Aragão, H. de B. 1911. Notas sobre ixódidas brazilleiros. Notes sur les ixodidés du Brésil. Memórias do Instituto Oswaldo Cruz, 3: 145–195, plates 11–12. Arnason, C.S. 1992. Biology of the western black-legged tick, Ixodes pacificus (Cooley and Kohls, 1943): a potential vector of Lyme disease in south coastal British Columbia. M.Sc. thesis. Simon Fraser University, Vancouver, British Columbia, Canada. 64 pp. Available from http://summit.sfu.ca/item/3850 [accessed 29 November 2015]. Arthur, D.R. and Snow, K.R. 1968. Ixodes pacificus Cooley and Kohls, 1943: its life-history and occurrence. Parasitology, 58: 893–906. Artsob, H. 1988. Powassan encephalitis. In The arboviruses: epidemiology and ecology. Edited by T. Monath. Volume IV. CRC Press, Boca Raton, Florida, United States of America. Pp. 29–49. Artsob, H. 1996. Tick-transmitted human disease threats in Canada. Germs and Ideas, 2: 35–39. Artsob, H. 2000. Arthropod-borne disease in Canada: a clinician’s perspective from the “Cold Zone”. Pediatrics & Child Health, 5: 206–212. Artsob, H., Spence, L., Surgeoner, G.A., McCreadie, J., Thorsen, J., Th’ng, C., and Lampotang, V. 1984. Isolation of Francisella tularensis and Powassan virus from ticks (Acari: Ixodidae) in Ontario, Canada. Journal of Medical Entomology, 21: 165–168. Asanuma, K. and Fukuda, S. 1957. On the life history of a tick, Ixodes signatus, infesting a black-tailed gull (Larus crassirostris) in Japan. Japanese Journal of Sanitary Zoology, 8: 147–159. [In Japanese]. Attoui, H., Billoir, F., Bruey, J.-M., de Micco, P., and de Lamballerie, X. 1998. Serological and molecular diagnosis of Colorado tick fever viral infections. American Journal of Tropical Medicine and Hygiene, 59: 763–768. Attoui, H., Jaafar, F.M., de Micco, P., and de Lamballerie, X. 2005. Coltiviruses and seadornaviruses in North America, Europe, and Asia. Emerging Infectious Diseases, 11: 1673–1679. Bacha, W.J. 1957. The life history of Otobius lagophilus. Journal of Parasitology, 43: 560–565. Bacon, R.M., Kugeler, K.J., and P.S. Mead. 2008. Surveillance for Lyme disease – United States, 1992–2006. Morbidity and Mortality Weekly Report, 57(SS10): 1–9. Baerg, W.J. 1944. Ticks and other parasites attacking northern cliff swallows. The Auk, 61: 413–414 + Plate 17. Balashov, Y.S. 1972. Bloodsucking ticks (Ixodoidea) vectors of diseases of man and animals. Miscellaneous Publications of the Entomological Society of America 8: 1–376. Balbridge, G.D., Scoles, G.A., Burkhardt, N.Y., Schloeder, B., Kurtti, T.J., and Munderloh, U.G. 2009. Transovarial transmission of Francisella-like endosymbionts and Anaplasma phagocytophilum variants in Dermacentor albipictus (Acari: Ixodidae). Journal of Medical Entomology, 46: 625–632. Ballard, J.T. and Ring, R.A. 1979. The ectoparasites of some marine birds from Bamfield Marine Station, British Columbia, with particular reference to the common murre, Uria aalge (Pont.). Canadian Journal of Zoology, 57: 1980–1984. Banerjee, S.N., Banerjee, M., Fernando, K., Burgdorfer, W., and Schwan T.G. 1998. Tick-borne relapsing fever in British Columbia, Canada: first isolation ofBorrelia hermsii. Journal of Clinical Microbiology, 36: 3505–3508. Banerjee, S.N., Banerjee, M., Smith, J.A., and Fernando, K. 1994. Lyme disease in British Columbia – an update. British Columbia Medical Journal, 36: 540–541. Banerjee, S.N., Stephen, C., Fernando, K., Coffey, S., and Dong, M. 1996. Evaluation of dogs as sero-indicators of the geographic distribution of Lyme borreliosis in British Columbia. The Canadian Veterinary Journal, 37: 168–169. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 253

Banks, N. 1904. Some Arachnida from California. Proceedings of the California Academy of Sciences, 3: 331–369. Banks, N. 1908. A revision of the Ixodoidea, or ticks, of the United States. United States Department of Agriculture, Bureau of Entomology, Technical Series, 15: 1–61. Banks, N. 1909. Three new ticks from the United States. Proceedings of the Entomological Society of Washington, 10: 170–173. Banks, N. 1910. New American mites (Arachnoidea, Acarina). Proceedings of the Entomological Society of Washington, 12: 2–12. Banks, N. 1912. New American mites. Proceedings of the Entomological Society of Washington, 14: 96–99. Barbour, A.G. 2005. Relapsing fever. In Tick-borne diseases of humans. Edited by J.L. Goodman, D.T. Dennis, and D.E. Sonenshine. American Society for Microbiology Press, Washington, District of Columbia, United States of America. Pp. 268–291. Barker, I.K., Lindsay, L.R., Campbell, G.D., Surgeoner, G.A., and McEwen, S.A. 1993. The groundhog tick Ixodes cookei (Acari: Ixodidae): a poor potential vector of Lyme borreliosis. Journal of Wildlife Diseases, 29: 416–422. Barker, I.K., Surgeoner, G.A., Artsob, H., McEwen, S.A., Elliott, L.A., Campbell, G.D., and Robinson, J.T. 1992. Distribution of the Lyme disease vector, Ixodes dammini (Acari: Ixodidae) and isolation of Borrelia burgdorferi in Ontario, Canada. Journal of Medical Entomology, 29: 1011–1022. Barker, S.C. and Murrell, A. 2008. Systematics and evolution of ticks with a list of valid genus and species names. In Ticks: biology, disease and control. Edited by P.A. Nuttall, and A.S. Bowman. Cambridge University Press, Cambridge, United Kingdom. Pp. 1–39. Barrett, A., Little, S.E., and Shaw, E. 2014. Rickettsia amblyommi and R. montanensis infection in dogs following natural exposure to ticks. Vector-borne and Zoonotic Diseases, 14: 20–25. Battaly, G.R. and Fish, D. 1993. Relative importance of bird species as hosts for immature Ixodes dammini (Acari: Ixodidae) in a suburban residential landscape of southern New York State. Journal of Medical Entomology, 30: 740–747. Battaly, G.R., Fish, D., and Dowler, R.C. 1987. The seasonal occurrence of Ixodes dammini and Ixodes dentatus (Acari: Ixodidae) on birds in a Lyme disease endemic area of southeastern New York State. Journal of the New York Entomological Society, 95: 461–468. Beck, D.E., Allred, D.A., and Brinton, E.P. 1963. Ticks of the Nevada test site. Brigham Young University Science Bulletin, Biology Series, 4: 1–11. Beeler, E., Abramowicz, K.F., Zambrano, M.L., Surgeon, M.M., Khalaf, N., Hu, R., et al. 2011. A focus of dogs and Rickettsia massiliae-infected Rhipicephalus sanguineus in California. American Journal of Tropical Medicine and Hygiene, 84: 244–249. Bennett, S.G., Webb, J.P., and Collins, P.W. 1989. Ixodes rugosus Bishopp (Acari: Ixodidae): description of larvae from Santa Rosa Island, California. Bulletin for the Society for Vector Ecology, 14: 262–265. Benoit, J.B., Yoder, J.A., Lopez-Martinez, G., Elnitsky, M.A., Lee, R.E., and Denlinger, D.L. 2007. Habitat requirements of the sea bird tick, Ixodes uriae (Acari: Ixodidae), from the Antarctic Peninsula in relation to water balance characteristics of eggs, nonfed and engorged stages. Journal of Comparative Biology B, 177: 205–215. Bequaert, J.C. 1945. The ticks, or Ixodoidea, of the northeastern United States and eastern Canada. Entomologica Americana 25: 73–235. Bergström, S., Haemig, P.D., and Olsen, B. 1999a. Distribution and abundance of the tick Ixodes uriae in a diverse subantarctic sea bird community. Journal of Parasitology, 85: 25–27. Bergström, S., Haemig, P.D., and Olsen, B. 1999b. Increased mortality of black-browed albatross chicks at a colony heavily-infested with the tick Ixodes uriae. International Journal of Parasitology, 29: 1359–1361. Berlese, A. 1888 Acari Austro-Americani. Bollettino della Societa Entomologica Italiana, 20: 171–223. Berlese, A. 1889. Acari, Myriapoda et Scorpiones hucusque in Italia reperta. Fascicle 55. Loescher e A. Seeber, Padova, Italy. Billeter, S.A., Levy, M.G., Chomel, B.B., and Breitschwerdt, E.B. 2008. Vector transmission of Bartonella species with emphasis on the potential for tick transmission. Medical and Veterinary Entomology, 22: 1–15. Binnington, K.C. and Kemp, D.H. 1988. Role of tick salivary glands in feeding and disease transmission. Advances in Parasitology, 19: 315–339. Birula, A. 1895. Ixodidae novi vel parum cogniti Musei Zoologici Academiae Caesareae Scientarum Petropolitanae. Izvestia Imperiale Akademie Nauka, serie 5, 2: 353–364. Bishopp, F.C. 1911. Some new North American Ixodidae, with notes on other species. Proceedings of the Biological Society of Washington, 24: 197–208. 254 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Bishopp, F.C. 1912. A new species of Dermacentor, and notes on other North American Ixodidae. Proceedings of the Biological Society of Washington, 25: 29–37. Bishopp, F.C., and Hixson, H. 1936. Biology and economic importance of the Gulf Coast tick. Journal of Economic Entomology, 29: 1068–1076. Bishopp, F.C. and King, W.V. 1913. Additional notes on the biology of the Rocky Mountain spotted-fever tick. Journal of Economic Entomology, 6: 200–211. Bishopp, F.C. and Smith, C.N. 1937. A new species of Ixodes from Massachusetts. Proceedings of the Entomological Society of Washington, 39: 133–138. Bishopp, F.C. and Smith, C.N. 1938. The American dog tick, eastern carrier of Rocky Mountain spotted fever. United States Department of Agriculture Circular, 478: 1–25. Bishopp, F.C. and Trembley, H.L. 1945. Distribution and hosts of certain North American ticks. Journal of Parasitology, 31: 1–54. Bishopp, F.C. and Wood, H.P. 1913. The biology of some North American ticks of the genus Dermacentor. Parasitology, 6: 153–187. Blanton, L., Keith, B., and Brzezinski, W. 2008. Southern tick-associated rash illness: erythema migrans is not always Lyme disease. Southern Medical Journal, 101: 759–760. Blouin, E.F., Kocan, A.A., Glenn, B.L., and Kocan, K.M. 1984. Transmission of Cytauxzoon felis Kier, 1979 from bobcats, Felis rufus (Schreber), to domestic cats by Dermacentor variabilis (Say). Journal of Wildlife Diseases, 20: 241–242. Bolte, J.R., Hair, J.A., and Fletcher, J. 1970. White-tailed deer mortality following tissue destruction induced by lone star ticks. Journal of Wildlife Management, 34: 546–552. Botzler, R.G., Albrecht, J., and Schaefer, T. 1980. Tick paralysis in a western harvest mouse. Journal of Wildlife Diseases, 16: 223–224. Bouchard, C., Beauchamp, G., Nguon, S., Trudel, L., Milord, F., Lindsay, L.R., et al. 2011. Associations between Ixodes scapularis ticks and small mammal hosts in a newly endemic zone in southeastern Canada: implications for Borrelia burgdorferi transmission. Ticks and Tick-borne Diseases, 2: 183–190. Bouchard, C., Leighton, P.A., Beauchamp, G., Nguon, S., Trudel, L., Milord, F., et al. 2013. Harvested white- tailed deer as sentinel hosts for early establishing Ixodes scapularis populations and risk from vector- borne zoonoses in southeastern Canada. Journal of Medical Entomology, 50: 384–393. Bowles, D.E., Robbins, R.G., Harlan, H.J., and Carpenter, T.L. 2013. New Missouri county records and review of the distribution and disease potential of Ornithodoros kelleyi (Arachnida: Ixodida: Argasidae) and Cimex adjunctus (Insecta: Hemiptera: Cimicidae). Proceedings of the Entomological Society of Washington, 115: 117–127. Bozeman, F.M., Elisberg, B.L., Humphries, J.W., Runcik, K., and Palmer, D.B. 1970. Serologic evidence of Rickettsia canada infection of man. Journal of Infectious Diseases, 121: 367–371. Brackney, M.M., Marfin, A.A., Staples, J.E., Stallones, L., Keefe, T., Black, W.C., and Campbell, G.L. 2010. Epidemiology of Colorado tick fever in Montana, Utah, and Wyoming, 1995–2003. Vector-borne and Zoonotic Diseases, 10: 381–385. Brackney, D.E., Nofchissey, R.A., Fitzpatrick, K.A., Brown, I.K., and Ebel, G.D. 2008. Short report: stable prevalence of Powassan virus in Ixodes scapularis in a northern Wisconsin focus. American Journal of Tropical Medicine and Hygiene, 79: 971–973. Bram, R.A. and Roby, T.O. 1970. Attempted hereditary transmission of Anaplasma marginale Theiler with Dermacentor albipictus (Packard). Journal of Parasitology, 56: 620–621. Bratton, R.L. and Corey, R. 2005. Tick-borne disease. American Family Physician, 71: 2323–2330. Bricker, J.M. 2005. The Merck veterinary manual, 9th edition. Merck, Sharp, and Dohme Coporation, Kenilworth, New Jersey, United States of America. Brinton, E.P., Beck, D.E., and Allred, D.M. 1965. Identification of the adults, nymphs and larvae of ticks of the genus Dermacentor Koch (Ixodidae) in the western United States. Brigham Young University Science Bulletin, Biological Series, 5: 1–44. Brown, J.H. 1944. The spotted fever and other Albertan ticks. Canadian Journal of Research, 22: 36–51. Brown, J.H. 1955. Colorado tick fever in Alberta. Canadian Journal of Zoology, 33: 389–390. Brown, J.H. and Kohls, G.M. 1950. The ticks of Alberta with special reference to distribution. Canadian Journal of Research, 28: 197–205. Brown, R.N., Peot, M.A., and Lane, R.S. 2006. Sylvatic maintenance of Borrelia burgdorferi (Spirochaetales) in northern California: untangling the web of transmission. Journal of Medical Entomology, 43: 743–751. Brown, S.J. 1977. Feeding performance, survival and appearance of Dermacentor variabilis nymphs (Acari: Ixodidae), as influenced by host resistance. Journal of Medical Entomology, 14: 497–498. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 255

Brownstein, J.S., Skelly, D.K., Holford, T.R., and Fish, D. 2005. Forest fragmentation predicts local scale heterogeneity of Lyme disease risk. Oecologia, 146: 469–475. Bruce, E.A. 1920. Tick paralysis in British Columbia. Dominion of Canada, Department of Agriculture, Bulletin, 133: 3–7. Bruce, E.A. 1927. Entomological notes of veterinary interest. Proceedings of the British Columbia Entomological Society, 1927: 26–30. Buckell, E.R. 1934. Notes on ticks and insect parasites of game animals in British Columbia. Proceedings of the British Columbia Entomological Society, 1934: 10–16. Bullard, J.M.P., Ahsanuddin, A.N., Perry, A.M., Lindsay, L.R., Iranpour, M.I., Dibernardo, A., and Van Caeseele, P.G. 2014. The first case of locally acquired tick-borneBabesia microti infection in Canada. Canadian Journal of Infectious Diseases and Medical Microbiology, 25: 1–3. Buller, R.S., Arens, M., Hmiel, S.P., Paddock, C.D., Sumner, J.W., Rikhisa, Y., et al. 1999. Ehrlichia ewingii, a newly recognized agent of human ehrlichiosis. New England Journal of Medicine, 341: 148–155. Burachynsky, V.I. and Galloway, T.D. 1985. Seasonal dynamics and distribution of American dog tick, Dermacentor variabilis (Say), larvae and nymphs at Birds Hill Park, Manitoba. Canadian Journal of Zoology, 63: 2748–2755. Burgdorfer, W. 1969. Ecology of tick vectors of American spotted fever. Bulletin of the World Health Organization, 40: 375–381. Burgdorfer, W. 1977. Tick-borne diseases in the United States: Rocky Mountain spotted fever and Colorado tick fever. Acta Tropica, 34: 103–126. Burgdorfer, W., Hayes, S.F., and Mavros, A.J. 1981. Non-pathogenic rickettsiae in Dermacentor andersoni: a limiting factor for the distribution of Rickettsia rickettsii. In Rickettsiae and rickettsial diseases. Edited by W. Burgdorfer and R.L. Anacker. Academic Press, New York, United States of America. Pp. 585–594, 650. Burgess, G.D. 1955. Arthropod ectoparasites of Richardson’s ground squirrel. Journal of Parasitology, 41: 639–640. Burgess, H., Chilton, N.B., Krakowetz, C.N., Williams, C., and Lohmann, K. 2012. Granulocytic anaplasmosis in a horse from Saskatchewan. Canadian Veterinary Journal, 53: 886–888. Burke, G.S., Wikel, S.K., Spielman, A., Telford, S.R., McKay, K., Krause, P.J., and the tick-borne infection study group. 2005. Hypersensitivity to ticks and Lyme disease risk. Emerging Infectious Diseases, 11: 36–41. Burkot, T.R., Clover, J.R., Happ, C.M., DeBess, E., and Maupin, G.O. 1999. Isolation of Borrelia burgdorferi from Neotoma fuscipes, Peromyscus maniculatus, Peromyscus boyleii and Ixodes pacificus in Oregon. American Journal of Tropical Medicine and Hygiene, 60: 453–457. Burkot, T.R., Maupin, G.O., Schneider, B.S., Denatale, C., Happ, C.M., Rutherford, J.A., and Zeidner, N.S. 2001a. Use of a sentinel system to study the questing behaviour of Ixodes spinipalpis and its role in the transmission of Borrelia bissettii, granulocyctic ehrlichiosis, and Babesia microti. American Journal of Tropical Medicine and Hygiene, 65: 293–299. Burkot, T.R., Schneider, B.S., Pieniazek, N.J., Happ, C.M., Rutherford, J.S., Slemenda, S.B., et al. 2001b. Babesia microti and Borrelia bissettii transmission by Ixodes spinipalpis ticks among prairie voles, Microtus ochrogaster, in Colorado. Parasitology, 121: 595–599. Calisher, C.H. 1994. Medically important arboviruses of the United States and Canada. Clinical Microbiology Reviews, 7: 89–116. Calisher, C.H., Schwan, T.G., Lazuick, J.S., Eads, R.B., and Francy, D.B. 1988. Isolation of Mono Lake virus (family Reoviridae, genus Orbivirus, Kemerovo serogroup) from Argas cooleyi (Acari: Argasidae) collected in California. Journal of Medical Entomology, 25: 388–390. Cameron, A.E. and Fulton, J.S. 1927. A local outbreak of the winter or moose tick, Dermacentor albipictus, Pack. (Ixodoidea) in Saskatchewan. Bulletin of Entomological Research, 17: 249–257. Camin, J.H. and Drenner, R.W. 1978. Climbing behavior and host-finding of larval rabbit ticks Haemaphysalis( leporispalustris). Journal of Parasitology, 64: 905–909. Campbell, A. 1979. Ecology of the American dog tick, Dermacentor variabilis in southwestern Nova Scotia. In Recent advances in acarology, volume 2. Edited by J.G. Rodriguez. Academic Press, New York, New York, United States of America. Pp. 135–143. Campbell, A. and Glines, M.V. 1979. Development, survival, and oviposition of the rabbit tick, Haemaphysalis leporispalustris (Packard) (Acari, Ixodidae), at constant temperatures. Journal of Parasitology, 65: 777–781. Campbell, A. and Harris, D.L. 1979. Reproduction of the American dog tick, Dermacentor variabilis, under laboratory and field conditions. Environmental Entomology, 8: 734–739. Campbell, A. and MacKay, P.R. 1979. Distribution of the American dog tick, Dermacentor variabilis (Say), and its small-mammal hosts in relation to vegetative types in a study area in Nova Scotia. Canadian Journal of Zoology, 57: 1950–1959. 256 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Campbell, A., Ward, R.M., and Garvie, M.B. 1980. Seasonal activity and frequency distributions of ticks (Acari, Ixodidae) infesting snowshoe hares in Nova Scotia, Canada. Journal of Medical Entomology, 17: 22–29. Campbell, G.D., Addison, E.M., Barker, I.K., and Rosendal, S. 1994. Erysipelothrix rhusiopathiae, serotype 17, septicemia in moose (Alces alces) from Algonquin Park, Ontario. Journal of Wildlife Diseases, 30: 436–438. Carey, A.B., Krinsky, W.L., and Main, A.J. 1980. Ixodes dammini (Acari: Ixodidae) and associated ixodid ticks in south-central Connecticut, USA. Journal of Medical Entomology, 17: 89–99. Carey, A.B., McLean, R.G., and Maupin, G.O. 1980. The structure of a Colorado tick fever system. Ecological Monographs, 50: 131–151. Carroll, J.F., Russek-Cohen, E., Nichols, J.D., and Hines, J.E.. 1991. Population dynamics of American dog ticks (Acari: Ixodidae) along park trails. Journal of Medical Entomology, 20: 922–929. Carroll, J.F. and Schmidtmann, E.T. 1986. American dog tick (Acari: Ixodidae), summer activity on equine premises enzootic for Potomac horse fever in south-central Maryland. Journal of Economic Entomology, 79: 62–66 Casher, L., Lane, R.S., Barrett, R., and Eisen, L. 2002. Relative importance of lizards and mammals as hosts for ixodid ticks in northern California. Experimental and Applied Acarology, 26: 127–143. Castro, M.B. and Wright, S.A. 2007. Vertebrate hosts of Ixodes pacificus (Acari: Ixodidae) in California. Journal of Vector Ecology, 32: 140–149. Chang, C.C., Chomel, B.B., Kasten, R.W., Romano, V., and Tietze, N. 2001. Molecular evidence of Bartonella spp. in questing adult Ixodes pacificus ticks in California. Journal of Clinical Microbiology, 39: 1221– 1226. Chapman, E.S. and Siegle, T.E. 2000. A novel method for tick retrieval reveals northern migration of Amblyomma americanum into Pike County, IL, USA. International Journal of Acarology, 26: 391–393. Chastel, C.E. 1988. Tick-borne virus infections of marine birds. Advances in Disease Vector Research, 5: 25–60. Chen, S.M., Dumler, J.S., Bakken, J.S., and Walker, D.H. 1994. Identification of a granulocytotropicEhrlichia species as the etiologic agent of human disease. Journal of Clinical Microbiology, 32: 589–595. Childs, J.E. and Paddock, C.D. 2003. The ascendancy of Amblyomma americanum as a vector of pathogens affecting humans in the United States. Annual Review of Entomology, 48: 307–337. Chowdri, H.R., Gugliotta, J.L., Berardi, V.P., Goethert, H.K., Molloy, P.J., Sterling, S.L., and Telford, S.R. 2013. Borrelia miyamotoi infection presenting as human granulocytic anaplasmosis: a case report. Annals of Internal Medicine, 159: 21–27. Cimolai, N., Anand, C.M., Gish, G.J., Calisher, C.H., and Fishbein, D.B. 1988. Human Colorado tick fever in southern Alberta. Canadian Medical Association Journal, 139: 45–46. Clifford, C.M. and Anastos, G. 1960. The use of chaetotaxy in the identification of larval ticks (Acarina: Ixodidae). Journal of Parasitology, 46: 567–578. Clifford, C.M., Anastos, G., and Elbl, A. 1961. The larval ixodid ticks of the eastern United States (Acarina: Ixodidae). Miscellaneous Publications of Entomological Society of America, 2: 213–237. Clifford, C.M., Kohls, G.M., and Sonenshine, D.E. 1964. The systematics of the subfamily Ornithodorinae (Acarina: Argasidae). I. The genera and subgenera. Annals of Entomological Society of America, 57: 429–437. Clifford, C.M., Sonenshine, D.E., Atwood, E.L., Robbins, C.S., and Hughes, L.E. 1969. Tests on ticks from wild birds collected in the eastern United States for rickettsiae and viruses. American Journal of Tropical Medicine and Hygiene, 18: 1057–1061. Clifford, C.M., Yunker, C.E., Easton, E.R., and Kohls, G.M. 1973. Systematics of the subfamily Ixodinae (Acarina: Ixodidae). 1. The subgenera of Ixodes. Annals of the Entomological Society of America, 66: 489–500. Cockwill, K.R., Taylor, S.M., Snead, E.C.R., Dickinson, R., Cosford, K., Malek, S., et al. 2009. Granulocytic anaplasmosis in three dogs from Saskatoon, Saskatchewan. Canadian Veterinary Journal, 50: 835–840. Conlon, J.M. and Rockett, C.L. 1982. Ecological investigations of the American dog tick, Dermacentor variabilis (Say), in northwest Ohio (Acari: Ixodidae). International Journal of Acarology, 8: 125–131. Cook, B. 1972. Hosts of Argas cooleyi and Ornithodoros concanensis (Acarina: Argasidae) in a cliff-face habitat. Journal of Medical Entomology, 9: 315–317. Cooley, R.A. 1936. Ornithodoros parkeri, a new species on rodents. United States Public Health Reports, 51: 431–433. Cooley, R.A. 1938. The genera Dermacentor and Otocentor (Ixodidae) in the United States, with studies in variation. United States National Institute of Health Bulletin, 171: 1–89. Cooley, R.A. 1944. Ixodes ozarkus n. sp. and Ornithodoros aquilae n. sp. with notes on O. talaje and O. kelleyi (Ixodoidea). Journal of Parasitology, 30: 287–294. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 257

Cooley, R.A. 1946a. Note on the tick Ixodes angustus Neumann. Journal of Parasitology, 32: 210. Cooley, R.A. 1946b. Ixodes holdenriedi, a new species of tick from a pocket gopher in California (Acarina, Ixodidae). Pan-Pacific Entomologist,22: 103–104. Cooley, R.A. 1946c. The genera Boophilus, Rhipicephalus, and Haemaphysalis (Ixodidae) of the new world. United States National Institute of Health Bulletin, 187: 1–54. Cooley, R.A. and Kohls, G.M. 1938a. Two new species of ticks (Ixodes) from California (Acarina: Ixodidae). United States Public Health Reports, 53: 1616–1621. Cooley, R.A. and Kohls, G.M. 1938b. Ixodes marmotae – A new species of tick from California. United States Public Health Reports, 53: 2174–2181. Cooley, R.A. and G.M. Kohls. 1940. Two new species of Argasidae (Acarina: Ixodoidea). United States Public Health Reports, 55: 925–933. Cooley, R.A. and Kohls, G.M. 1941. Further new species of Ornithodoros from bats (Acarina: Argasidae). United States Public Health Service Reports, 56: 910–914. Cooley, R.A. and Kohls, G.M. 1942. Ixodes baergi, a new species of tick from Arkansas. United States Public Health Reports, 57: 1869–1872. Cooley, R.A. and Kohls, G.M. 1943. Ixodes californicus Banks, 1904, Ixodes pacificus n. sp. and Ixodes conepati n. sp. (Acarina: Ixodidae). Pan-Pacific Entomologist,19: 139–147. Cooley, R.A. and Kohls, G.M. 1944. The genus Amblyomma (Ixodidae) in the United States. Journal of Parasitology, 30: 77–111. Cooley, R.A. and Kohls, G.M. 1945. The genus Ixodes in North America. United States Public Health Service, Bulletin of the National Institute of Health, 184: 1–246. Corwin, D., Clifford, C.M., and Keirans, J.E. 1979. An improved method for cleaning and preparing ticks for examination with the scanning electron microscope. Journal of Medical Entomology, 16: 352–353. Cowan, I.M. 1946. Parasites, diseases, injuries, and anomalies of the Columbian black-tailed deer, Odocoileus hemionus columbianus (Richardson), in British Columbia. Canadian Journal of Research, 24: 71–103. Croft, R.E. and Kingston, N. 1975. Babesia moshkovskii (Schurenkova, 1938) Laird and Lari, 1957; from the Prairie Falcon, Falco mexicanus, in Wyoming; with comments on other parasites found in this host. Journal of Wildlife Diseases, 11: 229–233. Darling, P.G.E. 1969. Observations on the relation of light to the dropping of the tick Ixodes texanus. Journal of the Entomological Society of British Columbia, 66: 26–28. Davidson, W.R., Siefken, D.A., and Creekmore, L.H. 1994. Seasonal and annual abundance of Amblyomma americanum (Acari: Ixodidae) in central Georgia. Journal of Medical Entomology, 31: 67–71. Davis, A.J. and Campbell, A. 1979. Ixodiphagus texanus Howard (Hymenoptera: Encrytidae) a parasite of the rabbit tick in Nova Scotia. Canadian Journal of Zoology, 57: 1164–1166. Davis, G.E. 1939. Ornithodoros parkeri: distribution and host data; spontaneous infection with the relapsing fever spirochetes. United States Public Health Reports, 54: 2178–2180. Davis, G.E. 1941. Ornithodoros parkeri Cooley: observations on the biology of this tick. Journal of Parasitology, 27: 425–433. Davis, G.E. 1942. Tick vectors and life cycles of ticks. American Association for the Advancement of Science, Publication, 18: 66–76. De Geer, C. 1778 Mémoires pour server à l’histoire des insects. VII. Pierre Hasselberg, Stockholm, Sweden. de la Fuente, J., Atkinson, M.W., Hogg, J.T., Miller, D.S., Naranjo, V., Almazan, C., et al. 2006. Genetic characterization of Anaplasma ovis strains from bighorn sheep in Montana. Journal of Wildlife Diseases, 42: 381–385. de la Fuente, J., Atkinson, M.W., Naranjo, V., Fernandez de Mera, I.G., Mangold, A.J., Keating, K.A., and Kocan, K.M. 2007. Sequence analysis of the msp4 gene of Anaplasma ovis strains. Veterinary Microbiology, 119: 375–381. de la Fuente, J., Estrada-Peña, A., Venzal, J.M., Kocan, K.M., and Sonenshine, D.E. 2008. Overview: ticks as vectors of pathogens that cause disease in humans and animals. Frontiers in Bioscience, 13: 6938–6946. de la Fuente, J., Golsteyn Thomas, E.J., van den Bussche, R.A., Hamilton, R.G., Tanaka, E.E., Druhan, S.E., and Kocan, K.M. 2003. Characterization of Anaplasma marginale isolated from North American bison. Applied Environmental Microbiology, 69: 5001–5005. de Meeûs, T., McCoy, K.D., Prugnolle, F., Chevillon, C., Durand, P., Hurtrez-Boussès, S., and Renaud, F. 2007. Population genetics and molecular epidemiology or how to “débusquer la bête”. Infection, Genetics and Evolution, 7: 308–332. Demma, L.J., Eremeeva, M., Nicholson, W.L., Traeger, M., Blau, D., Paddock, C., et al. 2006. An outbreak of Rocky Mountain spotted fever associated with a novel tick vector, Rhipicephalus sanguineus, in Arizona, 2004: preliminary report. Annals of the New York Academy of Sciences, 1078: 342–343. 258 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Demma, L.J., Holman, R.C., McQuiston, J.H., Krebs, J.W., and Swerdlow, D.L. 2005a. Epidemiology of human ehrlichiosis and anaplasmosis in the United States, 2001–2002. American Journal of Tropical Medicine and Hygiene, 73: 400–409. Demma, L.J., Traeger, M.S., Nicholson, W.L., Paddock, C.D., Blau, D.M., Eremeeva, M.E., et al. 2005b. Rocky Mountain spotted fever from an unexpected tick vector in Arizona. New England Journal of Medicine, 353: 587–594. Dergousoff, S.J., Gajadhar, A.J.A., and Chilton, N.B. 2009. Prevalence of Rickettsia species in Canadian populations of Dermacentor andersoni and D. variabilis. Applied Environmental Microbiology, 75: 1786–1789. Dergousoff, S.J., Galloway, T.D., Lindsay, L.R., Curry, P.S., and Chilton, N.B. 2013. Range extension of Dermacentor variabilis and Dermacentor andersoni (Acari: Ixodidae) near their northern distributional limits. Journal of Medical Entomology, 50: 510–520. Dibernardo, A., Coté, T., Ogden, N.H., and Lindsay, L.R. 2014. The prevalence of Borrelia miyamotoi infection, and co-infections with other Borrelia spp. in Ixodes scapularis ticks collected in Canada. Parasites & Vectors, 7: 183. Dibernardo, A., Lindsay, L.R., Artsob, H., and Drebot, M. 2007. Phylogenetic characterization of a Powassan (POW) virus strain isolated from an Ixodes tick collected in Prince Edward Island (PEI). Association of Medical Microbiology and Infectious Disease Canada – Canadian Association for Clinical Microbiology and Infectious Diseases Annual Conference, Halifax, Nova Scotia, Canada, March 14–18, 2007. Dick, T.A. 1981. Ectoparasites of sharp-tailed grouse, Pediocetes phasianellus. Journal of Wildlife Diseases, 17: 229–235. Dickinson, R. 2008. Anaplasma phagocytophilum infection in three dogs: potential for an emerging infectious disease concern in Saskatchewan? Animal Health Perspectives, 4: 2–3. Dietrich, M., Gómez-Diaz, E., and McCoy, K.D. 2011. Worldwide distribution and diversity of sea bird ticks: implications for the ecology and epidemiology of tick-borne pathogens. Vector-borne and Zoonotic Diseases, 11: 453–470. Ditchfield, J., Meads, E.B., and Julian, R.J. 1960. Tularemia of muskrats in eastern Ontario. Canadian Journal of Public Health, 51: 474–478. Dodds, D. G., Martell, A.M., and Yescott, R.E. 1969. Ecology of the American dog tick, Dermacentor variabilis (Say), in Nova Scotia. Canadian Journal of Zoology, 47: 171–181. Dolan, M.C., Lacombe, E.H., and Piesman, J. 2000. Vector competence of Ixodes muris (Acari: Ixodidae) for Borrelia burgdorferi. Journal of Medical Entomology, 37: 766–768. Dolce, P., Belanger, M.J., Tumanowicz, K., Gauthier, C.P., Jutras, P., Masse, R., et al. 2003. Coxiella burnetii seroprevalence of shepherds and their flocks in the lower Saint-Lawrence River region of Quebec, Canada. Canadian Journal of Infectious Diseases, 14: 97–102. Donahue, J.G., Piesman, J., and Spielman, A. 1987. Reservoir competence of white-footed mice for Lyme disease spirochetes. American Journal of Tropical Medicine and Hygiene, 36: 92–96. Drebot, M.A., Lindsay, L.R., Barker, I.K., and Artsob, H. 2001. Detection and characterization of a human granulocytic (HGE)-like agent from Ixodes scapularis in Ontario, Canada. Emerging Infectious Diseases, 7: 479–480. Drew, M.L. and Samuel, W.M. 1985. Factors affecting transmission of larval moose ticks, Dermacentor albipictus (Packard), to moose, Alces alces L., in Alberta, Canada. Journal of Wildlife Diseases, 21: 274–282. Drew, M.L. and Samuel, W.M. 1986. Reproduction of the winter tick, Dermacentor albipictus, under field conditions in Alberta, Canada. Canadian Journal of Zoology, 64: 714–721. Drew, M.L. and Samuel, W.M. 1989. Instar development and disengagement rate of engorged female winter ticks, Dermacentor albipictus (Acari: Ixodidae), following single- and trickle-exposure of moose (Alces alces). Experimental and Applied Acarology, 6: 189–196. Drexler, N.A., Dahlgren, F.S., Heitman, K.N., Massung, R.F., Paddock, C.D., and Behravesh, C.B. 2016. National surveillance of spotted fever group rickettsioses in the United States, 2008–2012. American Journal of Tropical Medicine and Hygiene, 94: 26–34. Drummond, R.O., Whetstone, T.M., Ernst, S.E., and Gladney, W.J. 1969. Biology and colonization of the winter tick in the laboratory. Journal of Economic Entomology, 62: 235–238. Drummond, R.O., Whetstone, T.M., Ernst, S.E., and Gladney, W.J. 1971. Oviposition of the American dog tick (Acarina: Ixodidae). Annals of the Entomological Society of America, 64: 1305–1309. Duffy, D.C. and Campbell, S.R. 1994. Ambient air temperature as a predictor of activity of adult Ixodes scapularis (Acari: Ixodidae). Journal of Medical Entomology, 31: 178–180. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 259

Dugan, V.G., Gaydos, J.K., Stallknecht, D.E., Little, S.E., Beall, A.D., Mead, D.G., et al. 2005. Detection of Ehrlichia spp. in raccoons (Procyon lotor) from Georgia. Vector-borne and Zoonotic Diseases, 5: 162–171. Dugès, A. 1883. Turicata y garrapata de Guanajuato. La Naturaleza (Mexico), 6: 195–198, plate 4. Dugès, A. 1888. Description d’un nouvel Ixodidé. Bulletin de la Société zoologique de France, 13: 129–133, figures 1–3. Dumler, J.S. and Bakken, J.S. 1998. Human ehrlichioses: newly recognized infections transmitted by ticks. Annual Review of Medicine, 49: 201–213. Dumler, J.S. and Walker, D.H. 2005. Rocky Mountain spotted fever—changing ecology and persisting virulence. New England Journal of Medicine, 353: 551–553. Duncan, A.W., Correa, M.T., Levine, J.F., and Breitschwerdt, E.B. 2005. The dog as a sentinel for human infection: prevalence of Borrelia burgdorferi C6 antibodies in dogs from southeastern and mid-Atlantic States. Vector-borne and Zoonotic Diseases, 5: 101–109. Durden, L.A. and Keirans, J.E. 1996. Nymphs of the genus Ixodes (Acari: Ixodidae) of the United States: taxonomy, identification key, distribution, hosts, and medical/veterinary importance. Thomas Say Publications in Entomology: Monographs 16:, 1–95. Durden, L.A., Oliver, J.H., and Kinsey, A.A. 2001. Ticks (Acari: Ixodidae) and spirochetes (Spirochaetaceae: Spirochaetales) recovered from birds on a Georgia barrier island. Journal of Medical Entomology, 38: 231–236. Dworkin, M.S., Anderson, Jr., D.E., Schwan, T.G., Shoemaker, P.C., Banerjee, S.N., Kassen, B.O., and Burgdorfer, W. 1998. Tick-borne relapsing fever in the northwestern United States and southwestern Canada. Clinical Infectious Diseases, 26: 122–131. Dworkin, M.S., Shoemaker, P.C., Fritz, C.L., Dowell, M.E., and Anderson, Jr., D.E. 2002. The epidemiology of tick-borne relapsing fever in the United States. American Journal of Tropical Medicine and Hygiene, 66: 753–758. Eads, R.B. and Smith, G.C. 1983. Seasonal activity and Colorado tick fever virus infection rates in Rocky Mountain wood ticks, Dermacentor andersoni (Acari: Ixodidae), in north-central Colorado, USA. Journal of Medical Entomology, 20: 49–55.

Eads, R.B., Smith, G.C., and Maupin, G.O. 1982. A CO2 platform trap for taking adult Dermacentor andersoni (Acari: Ixodidae). Proceedings of the Entomological Society of Washington, 84: 342–348. Easton, E.R. 1983. The ticks of South Dakota: an annotated checklist (Acari: Ixodidae). Entomological News, 94: 191–195. Easton, E.R. and Goulding, R.L. 1974. Ectoparasites in two diverse habitats in western Oregon. I. Ixodes (Acarina: Ixodidae). Journal of Medical Entomology, 11: 413–418. Easton, E.R., Keirans, J.E., Gresbrink, R.A., and Clifford, C.M. 1977. The distribution in Oregon of Ixodes pacificus, Dermacentor andersoni, and Dermacentor occidentalis with a note on Dermacentor variabilis (Acarina: Ixodidae). Journal of Medical Entomology, 13: 501–506. Ebel G.D., Campbell, E., Goethert, H.K., Spielman, A., and Telford, S.R. 2000. Enzootic transmission of deer tick virus in New England and Wisconsin sites. American Journal of Tropical Medicine and Hygiene, 63: 36–42. Ebel, G.D., Spielman, A., and Telford III, S.R. 2001. Phylogeny of North American Powassan virus. Journal of General Virology, 82: 1657–1665. Eddy, G.W. and Joyce, C.R. 1942. Ticks collected on the Tama (Iowa) Indian Reservation with notes on other species. Iowa State College Journal of Science, 41: 539–543. Edlow, J.A. and McGillicuddy, D.C. 2008. Tick paralysis. Infectious Disease Clinics of North America, 22: 397–413. Edwards, M.A. 1975. The chaetotaxy of the pedipalps and legs of some larval ticks (Acari: Argasidae). Transactions of the Zoological Society of London, 33: 1–76. Edwards, M.A. and Evans, G.O. 1967. Some observations on the chaetotaxy of the legs of larval Ixodidae (Acari: Metastigmata). Journal of Natural History, 4: 595–601. Eisen, L. 2007. A call for renewed research on tick-borne Francisella tularensis in the Arkansas–Missouri primary national focus of tularemia in humans. Journal of Medical Entomology, 44: 389–397. Eisen, L., Eisen, R.J., and Lane, R.S. 2002. Seasonal activity patterns of Ixodes pacificus nymphs in relation to climatic conditions. Medical and Veterinary Entomology, 16: 235–244. Eisen, L., Eisen, R.J., and Lane, R.S. 2004. The roles of birds, lizards, and rodents as hosts for the western black-legged tick Ixodes pacificus. Journal of Vector Ecology, 29: 295–308. Eisen, L., Eisen, R.J., and Lane, R.S. 2006. Geographical distribution and habitat suitability models for presence of host-seeking ixodid ticks in dense woodlands of Mendocino County, California. Journal of Medical Entomology, 43: 415–427. 260 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Eisen, R.J., Mead, P.S., Meyer, A.M., Pfaff, L.E., Bradley, K.K., and Eisen, L. 2008. Ecoepidemiology of tularemia in the southcentral United States. American Journal of Tropical Medicine and Hygiene, 78: 586–594. Eklund, C.M., Kohls, G.M., and Brennan, J.M. 1955. Distribution of Colorado tick fever and virus-carrying ticks. Journal of the American Medical Association, 157: 335–337. Ellis, J., Oyston, P.C., Green, M., and Titball, R.W. 2002. Tularemia. Clinical Microbiology Reviews, 15: 631–646. Eriks, I.S., Stiller, D., and Palmer, G.H. 1993. Impact of persistent Anaplasma marginale rickettsemia on tick infection and transmission. Journal of Clinical Microbiology, 31: 2091–2096. Ernst, S.E. and Gladney, W.J. 1975. Dermacentor albipictus: hybridization of the two forms of the winter tick. Annals of the Entomological Society of America, 68: 63–67. Evans, G.O. 1963a. Observations on the chaetotaxy of the legs in the free-living Gamasina (Acari: Mesostigmata). Bulletin of the British Museum (Natural History), Zoology, 10: 277–303. Evans, G.O. 1963b. Some observations on the chaetotaxy of the pedipalps in the Mesostigmata (Acari). Annals and Magazine of Natural History, Series 13, 6: 513–527. Evans, G.O. 1969. Observations on the ontogenetic development of the chaetotaxy of the tarsi of legs II–IV in the Mesostigmata (Acari). In Proceedings of the 2nd International Congress of Acarology, Sutton Bonington (England, 9th–25th July 1967). Edited by G.O. Evans. Akademiai Kiado, Budapest, Hungary. Pp. 195–200. Evans, G.O. 1992. Principles of acarology. CAB International, Wallingford, Oxon, United Kingdom. Evans, G.P.A. 1969. Tularemia in Huron County, Ontario. Canadian Journal of Public Health, 60: 447–449. Eveleigh, E.S. and Threlfall, W. 1974. The biology of Ixodes (Ceratixodes) uriae White, 1852 in Newfoundland. Acarologia, 16: 621–635. Eveleigh, E.S. and Threlfall, W. 1975. The bionomics of Ixodes (Ceratixodes) uriae White, 1852 on auks (Alcidae) from Newfoundland. Canadian Journal of Zoology, 53: 82–86. Eveleigh, E.S., Threlfall, W., and Belbeck, L.W. 1974. Histopathological changes associated with the attachment of Ixodes (Ceratixodes) uriae White, 1852. Canadian Journal of Zoology, 52: 1443–1446. Ewing, S.A., DuBois, J.G., Mathew, J.S., and Panciera, R.J. 2002. Larval Gulf Coast ticks (Amblyomma maculatum) [Acari: Ixodidae] as host for Hepatozoon americanum [Apicomplexa: Adeleorina]. Veterinary Parasitology, 103: 43–51. Ewing, S.A., Harkess, J.R., Kocan, K.M., Barker, R.W., Fox, J.C., Tyler, R.D., et al. 1990. Failure to transmit Ehrlichia canis (Rickettsiales: Ehrlichiae) with Otobius megnini (Acari: Argasidae). Journal of Medical Entomology, 27: 803–806. Fabricius, J.C. 1794. Entomologia systematica emendata et aucta. Secundum Classes, Ordines, Genera, Species adjectis synonimis, locis, observationibus, descriptionibus, volume 4. C.G. Proft, Copenhagen, Denmark. Fabricius, J.C. 1805. Systema antliatorum secondum ordines, genera, species adjectis synonimis, locis, observationibus, descriptionibus. C. Reichard, Brunswick, Germany. Falco, R.C. and Fish, D. 1988. Ticks parasitizing humans in a Lyme disease endemic area of southern New York State. American Journal of Epidemiology, 128: 1146–1152. Farkas, M.J. and Surgeoner, G.A. 1990. Incidence of Ixodes cookei (Acari: Ixodidae) on groundhogs, Marmota monax, in southwestern Ontario. Proceedings of the Entomological Society of Ontario, 121: 105–110. Farkas, M.J. and Surgeoner, G.A. 1991. Developmental times and fecundity of Ixodes cookei Packard (Acari: Ixodidae) under laboratory conditions. The Canadian Entomologist, 123: 1–12. Fay, F.H. and Rausch, R.L. 1969. Parasitic organisms in the blood of arvicoline rodents in Alaska. Journal of Parasitology, 55: 1258–1265. Fedorova N., Kleinjan, J.E., James, D., Hui, L.T., Peeters, H., and Lane, R.S. 2014. Remarkable diversity of tick or mammalian-associated Borreliae in the metropolitan San Francisco Bay Area, California. Ticks and Tick-borne Diseases, 5: 951–961. Feldman-Muhsam, B. and Borut, S. 1971. Copulation in ixodid ticks. Journal of Parasitology, 57: 630–634. Felz, M.W., Durden, L.A., and Oliver, J.H. 1996. Ticks parasitizing humans in Georgia and South Carolina. Journal of Parasitology, 82: 505–508. Filippova, N.A. 1977. Ixodid ticks of the subfamily Ixodinae. Fauna SSSR, Paukoobraznye Fauna Series 9, 4: 4–393. [In Russian]. Fischer von Waldheim, G. 1823. Notice sur l’Argas de Perse (Malleh de Mianeh) décrit par les voyageurs sous le nom de punaise venimeuse de Miana. Mémoires de la Société imperiale des naturalistes de Moscou, 6: 269–283, plate 23. Fish, D., and Dowler, R.C. 1989. Host associations of ticks (Acari: Ixodidae) parasitizing medium-sized mammals in a Lyme disease endemic area of southern New York. Journal of Medical Entomology, 26: 200–209. Fitch, A. 1872. Fourteenth report on the noxious, beneficial, and other insects of the State of New York. Transactions of the New York State Agricultural Society (1870), 30: 366. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 261

Fitch, W.M. and Artsob, H. 1990. Powassan encephalitis in New Brunswick. Canadian Family Physician, 36: 1289–1290. Fitzpatrick, C. and Threlfall, W. 1977. The ectoparasites of three species of seabirds from Newfoundland. Canadian Journal of Zoology, 55: 1205–1209. Flint, V.B., and Kostyrko, I.N. 1967. [On the biology of the tick Ixodes putus Pick.] Zoologicheskii Zhurnal, 46: 1253–1256. [In Russian]. Flook, D.R. and Stenton, J.E. 1969. Incidence and abundance of certain parasites in wapiti in the national parks of the Canadian Rockies. Canadian Journal of Zoology, 47: 795–803. Foggie, A. 1959. Studies on the relationship of tick-bite to tick anemia of lambs. Annals of Tropical and Medical Parasitology, 53: 27–34. Foley, J.E., Foley, P., Brown, R.N., Lane, R.S., Dumler, J.S., and Madigan, J.E. 2004. Ecology of Anaplasma phagocytophilum and Borrelia burgdorferi in the western United States. Journal of Vector Ecology, 29: 41–50. Foley, J.E., Foley, P., and Madigan, J.E. 2001. Spatial distribution of seropositivity to the causative agent of granulocytic ehrlichiosis in dogs in California. American Journal of Veterinary Research, 62: 1599–1605. Foley, J.E., Nieto, N.C., Clueit, S.B., Foley, P., Nicholson, W.N., and Brown, R.N. 2007. Survey for zoonotic rickettsial pathogens in northern flying squirrels,Glaucomys sabrinus, in California. Journal of Wildlife Diseases, 43: 684–689. Foley J.E., Nieto, N.C., Massung, R., Barbet, A., Madigan, J., and Brown, R.N. 2009. Distinct ecologically relevant strains of Anaplasma phagocytophilum. Emerging and Infectious Diseases, 15: 842–843. Frauenfeld, G.R. 1867. Zoologische Miscellen. XI. Verhandlungen der Kaiserlich-Königlichen Zoologische- Botanischen Geselschaft in Wien, 17: 425–502. Fritz, C.L., Bronson, L.R., Smith, C.R., Crawford-Miksza, L., Yeh, E., and Schnurr, D. 2005. Clinical, epidemiologic, and environmental surveillance for ehrlichiosis and anaplasmosis in an endemic area of northern California. Journal of Vector Ecology, 30: 4–10. Fukunaga, M., Takahashi, Y., Tsuruta, Y., Matsushita, O., Ralph, D., McClelland, M., and Nakao, M. 1995. Genetic and phenotypic analysis of Borrelia miyamotoi sp. nov., isolated from the ixodid tick Ixodes persulcatus, the vector for Lyme disease in Japan. International Journal of Systematic Bacteriology, 45: 804–810. Furman, D.P. and Loomis, E.C. 1984. The ticks of California (Acari: Ixodidae). Bulletin of the California Insect Survey, 25: viii + 1–239. Gage, K.L., Eggleston, M.E., Gilmore, R.D., Dolan, M.C., Montenieri, J.A., Tanda, D.T., and Piesman, J. 2001. Isolation and characterization of Borrelia parkeri in Ornithodoros parkeri (Ixodida: Argasidae) collected in Colorado. Journal of Medical Entomology, 38: 665–674. Garvie, M.B., McKiel, J.A., Sonenshine, D.E., and Campbell, A. 1978. Seasonal dynamics of American dog tick, Dermacentor variabilis (Say), populations in southwestern Nova Scotia. Canadian Journal of Zoology, 56: 28–39. Gary, A.T., Webb, J.A., Hegarty, B.C., and Breitschwerdt, E.B. 2006. The low seroprevalence of tick- transmitted agents of disease in dogs from southern Ontario and Quebec. Canadian Veterinary Journal, 47: 1194–1200. Gauthier-Clerc, M., Clerquin, Y., and Handrich, Y. 1998. Hyperinfestation by ticks Ixodes uriae: a possible cause of death in adult king penguins, a long-lived sea bird. Colonial Waterbirds, 21: 229–233. George, J.W. 1971. Drop-off rhythms of engorged rabbit ticks, Haemaphysalis leporispalustris (Packard, 1896) (Acari: Ixodidae). Journal of Medical Entomology, 8: 461–479. George, J.E. 1981. The influence of aggregation pheromones on the behavior ofArgas cooleyi and Ornithodoros concanensis (Acari: Ixodoidea: Argasidae). Journal of Medical Entomology, 18: 129–133. George, J.E. 1987. Field observations on the life cycle of Ixodes baergi and some seasonal and daily activity cycles of Oeciacus vicarius (Hemiptera: Cimicidae), Argas cooleyi (Acari: Argasidae), and Ixodes baergi (Acari: Ixodidae). Journal of Medical Entomology, 24: 683–688. George, J.E. and Cook, B. 1979. Exokinetic responses of Argas cooleyi and Ornithodoros concanensis (Acari: Ixodoidea: Argasidae) to various environmental temperatures. Journal of Medical Entomology, 16: 275–285. Gerhardt, R.R., Lohmeyer, K.H., Marsland, E.J., and Paulsen, D.J. 1998. Seasonal abundance of the free-living stages of the lone star tick (Amblyomma americanum) in Cumberland County, Tennessee. Journal of the Tennessee Academy of Science, 73: 100–103. Gerstäcker, A. 1873. Gliederthiere (Insekten, Arachniden, Myriapoden und Isopoden). C.C. von der Decken, Reisen in Ost-Afrika, 3(2): 1–542. Gervais, P. 1844. Acarides. In Histoire naturelle des insectes. Aptères 3. Edited by C.A. Walckenaer. Fain and Thunot, Paris, France. 262 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Gill, J.S., Rowley, W.A., Bush, P.J., Viner, J.P., and Gilchrist, M.R. 2004. Detection of human blood in the bat tick Carios (Ornithodoros) kelleyi (Acari: Argasidae) in Iowa. Journal of Medical Entomology, 41: 1179–1181. Gkoroba, G. 1980. A study of the seasonal population dynamics of ticks Dermacentor variabilis (Say) and Haemaphysalis leporispalustris (Packard) in a marshland habitat. M.Sc. thesis. University of Manitoba, Winnipeg, Manitoba, Canada. Gladney, W.J. and Dawkins, C.C. 1971. Parthenogenetic reproduction by Dermacentor variabilis (Acarina: Ixodidae). Annals of the Entomological Society of America, 64: 1285–1289. Glashchinskaya-Babenko, LV. 1949. Chetotaksiya tela lichinok kleshchei sem. Ixodidae i ee taksonomicheskoe znachenie [Chaetotaxy of the body of larvae of ticks of the family Ixodidae and its taxonomic significance]. Doklady Akademii Nauk SSSR, new series, 65: 245–248. [In Russian]. Glines, M.V. and Samuel, W.M. 1984. The development of the winter tick, Dermacentor albipictus, and its effect on the hair coat of moose, Alces alces, of central Alberta, Canada. In Acarology VI. Edited by D.A. Griffiths and C.E. Bowman, volume 2. Ellis Horwood, Chichester, United Kingdom. Pp. 1208–1214. Glines, M.V. and Samuel, W.M. 1989. Effect of Dermacentor albipictus (Acari: Ixodidae) on blood composition, weight gain, and hair loss of moose, Alces alces. Experimental and Applied Acarology, 6: 197–213. Goethert, H.K., Lubelcyzk, C., LaCombe, E., Holman, M., Rand, P., Smith, R.P., and Telford, S.R. 2003. Enzootic Babesia microti in Maine. Journal of Parasitology, 89: 1069–1071. Goethert, H.K., Shani, I., and Telford, S.R. 2004. Genotypic diversity of Francisella tularensis infecting Dermacentor variabilis ticks on Martha’s Vineyard, Massachusetts. Journal of Clinical Microbiology, 42: 4968–4973. Goethert, H.K. and Telford, S.R. 2003a. Enzootic transmission of Babesia divergens among cottontail rabbits on Nantucket Island, Massachusetts. American Journal of Tropical Medicine and Hygiene, 69: 455–460. Goethert, H.K. and Telford, S.R. 2003b. Enzootic transmission of the agent of human granulocytic ehrlichiosis among cottontail rabbits. American Journal of Tropical Medicine and Hygiene, 69: 633–637. González-Acuña, D., Venzal, J.M., Keirans, J.E., Robbins, R.G., Ippi, S., and Guglielmone, A.A. 2005. New host and locality records for the Ixodes auritulus (Acari: Ixodidae) species group, with a review of host relationships and distribution in the Neotropical Zoogeographical Region. Experimental and Applied Acarology, 37: 147–156. Goodman, J.L. 2005. Human granulocytic anaplamsosis (Ehrlichiosis). In Tick-borne diseases of humans. Edited by J.L. Goodman, D.T. Dennis, and D.E. Sonenshine. ASM Press, Washington, District of Columbia, United States of America. Pp. 218–238. Goodpasture, H.C., Poland, J.D., Francy, D.B., Bowen, G.S., and Horn, K.A. 1978. Colorado tick fever: clinical, epidemiological, and laboratory aspects of 228 cases in Colorado in 1973–1974. Annals of Internal Medicine, 88: 303–310. Goodrich, B.S. and Murray, M.D. 1978. Factors influencing the toxicity of salivary gland extracts ofIxodes holocyclus Neumann. International Journal of Parasitology, 8: 313–320. Gordon, J.R. and Allen, J.R. 1991a. Factors V and VII anticoagulant activities in the salivary glands of feeding Dermacentor andersoni ticks. Journal of Parasitology, 77: 167–170. Gordon, J.R. and Allen, J.R. 1991b. Nonspecific activation of complement factor 5 by isolatedDermacentor andersoni antigens. Journal of Parasitology, 77: 296–301. Gordon, J.R., McLaughlin, B.G., and Nitiuthai, S. 1983. Tularemia transmitted by ticks (Dermacentor andersoni) in Saskatchewan. Canadian Journal of Comparative Medicine, 47: 408–411. Gothe, R. 1984. Tick paralysis: reasons for appearing during ixodid and argasid feeding. In Current topics in vector research. Volume 2. Edited by K.F. Harris. Praeger Scientific, New York, New York, United States of America. Pp. 199–223. Gothe, R. 1999. Zecken Toxikosen. Hieronymus, Munich, Germany. Gothe, R., Kunze, K., and Hoogstraal, H. 1979. The mechanisms of pathogenicity in the tick paralyses. Journal of Medical Entomology, 16: 357–369. Green, R.G., Evans, C.A., and Larson, C.L. 1943. A ten-year population study of the rabbit tick Haemaphysalis leporis-palustris. American Journal of Hygiene, 30: 260–281. Gregson, J.D. 1935a. Tick paralysis. The production of artificial conditions conducive to winter feeding of ticks. Proceedings of the Entomological Society of British Columbia, 31: 6–7. Gregson, J.D. 1935b. A preliminary report of the lizard-tick relationship on the British Columbia coast. Proceedings of the Entomological Society of British Columbia, 31: 17–21. Gregson, J.D. 1941. Two new species of ticks from British Columbia (Ixodidae). The Canadian Entomologist, 73: 220–228. Gregson, J.D. 1942a. The coast tick (Ixodes californicus Banks) problem in British Columbia. The Canadian Entomologist, 74: 3–5. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 263

Gregson, J.D. 1942b. A new species of tick found on shrews (Ixodes soricis n. sp.). The Canadian Entomologist, 74: 137–139. Gregson, J.D. 1942c. Notes on the laboratory reading of some Canadian ticks (Acarina). Proceedings of the Entomological Society of British Columbia, 39: 32–35. Gregson, J.D. 1947. Feeding periods prerequisite to the mating of Dermacentor andersoni (Acarina). Proceedings of the British Columbia Entomological Society, 43: 3–6. Gregson, J.D. 1951. Notes on the spring activity of the Rocky Mountain wood tick, Dermacentor andersoni Stiles (Acarina: Ixodidae). Proceedings of the British Columbia Entomological Society, 47: 4–7. Gregson, J.D. 1952. Further studies on tick paralysis. Proceedings of the Entomological Society of British Columbia, 48: 54–58. Gregson, J.D. 1954. The male of Ixodes signatus Birula (Acarina: Ixodidae). The Canadian Entomologist, 86: 275–277. Gregson, J.D. 1956. The Ixodoidea of Canada. Publication 930. Entomology Division, Science Service, Canada Department of Agriculture, Ottawa, Ontario, Canada. 92 pp. Gregson, J.D. 1958. Host susceptibility to paralysis by the tick Dermacentor andersoni Stiles (Acarina: Ixodidae). The Canadian Entomologist, 90: 421–424. Gregson, J.D. 1960. Morphology and functioning of the mouthparts of Dermacentor andersoni Stiles. Part II. The feeding mechanism in relation to the host. Acta Tropica, 17: 72–79. Gregson, J.D. 1971. Studies on two populations of Ixodes kingi Bishopp (Ixodidae). Canadian Journal of Zoology, 49: 591–587. Gregson, J.D. 1973. Tick paralysis. An appraisal of natural and experimental data. Canada Department of Agriculture Monograph, 9: 1–109. Gregson, J.D. and Kohls, G.M. 1952. The male of Ixodes soricis Gregson (Acarina: Ixodidae). The Canadian Entomologist, 84: 185–188. Grist, N.R. 1992. Ticks are topical. Journal of Infection, 24: 117–121. Guérin-Méneville, F.E. 1829–1839. Iconographie du règne animal de G. Cuvier, ou représentation d’apres nature de l’une des espèces les plus remarkables et souvent non encore figures de chaque genre d’animaux. Arachnides. J.B. Baillière, Paris, France. Guerra, M., Walker, E., Jones, C., Paskewitz, S., Cortinas, M.R., Stancil, A., and Beck., L., Bobo, M., and Kitron, U. 2002. Predicting the risk of Lyme disease: habitat suitability for Ixodes scapularis in north central United States. Emerging Infectious Diseases, 8: 289–297. Guerrero, F.D., Nene, V.M., George, J.E., Barker, S.C., and Willadsen, P. 2006. Sequencing a new target genome: the Boophilus microplus (Acari: Ixodidae) genome project. Journal of Medical Entomology, 43: 9–16. Guglielmone, A.A., Estrada-Peña, A., Keirans, J.E., and Robbins, R.G. 2003. Ticks (Acari: Ixodida) of the Neotropical zoogeographical region. Special Publication, International Consortium on Ticks and Tick- Borne Diseases. Atalanta, Houten, The Netherlands. Guglielmone, A.A., Robbins, R.G., Apanaskevich, D.A., Petney, T.N., Estrada-Peña, A., and Horak, I.G. 2014. The hard ticks of the world (Acari: Ixodida: Ixodidae). Springer, Dordrecht, The Netherlands. Guglielmone, A.A., Robbins, R.G., Apanaskevich, D.A., Petney, T.N., Estrada-Peña, A., Horak, I.G., et al. 2010. The Argasidae, Ixodidae and Nuttalliellidae (Acari: Ixodida) of the world: a list of valid species names. Zootaxa, 2528: 1–28. Gugliotta J.L., Goethert, H.K., Berardi, V.P., and Telford, S.R. 2013. Meningoencephalitis from Borrelia miyamotoi in an immunocompromised patient. New England Journal of Medicine, 368: 240–245. Guzmán-Cornejo, C. and Robbins, R.G. 2010. The genus Ixodes (Acari: Ixodidae) in Mexico: adult identification keys, diagnoses, hosts, and distribution. Revista Mexicana de Biodiversidad,81: 289–298. Guzmán-Cornejo, C., Robbins, R.G., and Pérez, T.M. 2007. The Ixodes (Acari: Ixodidae) of Mexico: parasite- host and host-parasite checklists. Zootaxa, 1553: 47–58. Gyorkos, T.W. and Hilton, D.F.J. 1982. Range extensions for some ectoparasites from rodents of southeastern Quebec. Canadian Journal of Zoology, 60: 486–488. Hadwen, S. 1913. Report of the Veterinary Director General. Department of Agriculture. Appendix, 9: 80. Hafez, M, El-Ziady, S., and Hefnawy, T. 1970. Biochemical and physiological studies of certain ticks (Ixodoidea). Cuticular permeability of Hyalomma (H.) dromedarii Koch (Ixodidae) and Ornithodoros (O.) savignyi (Audouin) (Argasidae). Journal of Parasitology, 56: 154–168. Hall, J.E., Armine, J.W., Gais, R.D., Kolanko, V.P., Hagenbuch, B.E., Gerencser, V.F., and Clark, S.M. 1991. Parasitization of humans in West Virginia by Ixodes cookei (Acari: Ixodidae) a potential vector of Lyme borreliosis. Journal of Medical Entomology, 28: 186–189. Hall, R.R., McKiel, J.A., and Gregson, J.D. 1968. Occurrence of Colorado tick fever virus in Dermacentor andersoni ticks in British Columbia. Canadian Journal of Public Health, 59: 273–275. 264 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Halperin, J.J. and Wormser, G.P. 2001. Of fleas and ticks and cats and mice. Archives of Neurology, 58: 1345–1347. Hamer, S.A., Roy, P.L., Hickling, G.J., Walker, E.D., Foster, E.S., Barber, C.C., and Tsao, J.I. 2007. Zoonotic pathogens in Ixodes scapularis, Michigan. Emerging Infectious Diseases, 13: 1131–1133. Hänel, C. and Heyne, H. 2008. Ticks of the Tristan da Cunha Archipelago (Acarina: Ixodidae: Argasidae). Beiträge zur Entomologie, 58: 121–131. Harvey, J.W., Simpson, C.F., and Gaskin, J.M. 1978. Cyclic thrombocytopenia induced by a Rickettsia-like agent in dogs. Journal of Infectious Diseases, 137: 182–188. Hatcher, J.C., Greenberg, P.D., Antique, J., and Jimenez-Lucho, V.E. 2001. Severe babesiosis in Long Island: review of 34 cases and their complications. Clinical Infectious Diseases, 32: 1117–1125. Hatchette, T.F., Davis, I., and Johnston. B.L. 2014. Lyme disease: clinical diagnosis and treatment. Canadian Communicable Disease Report, 40: 194. Hayes, E.B. 2005. Tularemia. In Tick-borne diseases of humans. Edited by J.L. Goodman, D.T. Dennis, and D.E. Sonenshine. ASM Press, Washington, District of Columbia, United States of America. Pp. 207–217. Hays, K.L. and Lawrence, W.H. 1957. Ixodes banksi Bishopp (Acarina; Ixodidae). I. Description of the male and larva. Journal of Parasitology, 43: 213–216. Hearle, E. 1938. The ticks of British Columbia. Scientific Agriculture,18: 341–354. Heath, A.C.G. 1977. Zoogeography of the New Zealand tick fauna. Tuatara, 23: 26–38. Heath, A.C.G., Palma, R.L., Cane, R.P., and Hardwick, S. 2011. Checklist of New Zealand ticks (Acari: Ixodidae, Argasidae). Zootaxa, 2995: 55–63. Hegarty, B.C., Maggi, R.G., Koskinen, P., Beall, M.J., Eberts, M., Chandrashekar, R., and Breitschwerdt, E.B. 2012. Ehrlichia muris infection in a dog from Minnesota. Journal of Veterinary Internal Medicine, 26: 1217–1220. Hermann, J.-F. 1804 Mémoire aptérologique. F.G. Levrault, Strasbourg, Germany. Herrin, C.S. 1965. Notes on the distribution of Otobius lagophilus. Proceedings of the Utah Academy of Science, 42: 165. Herrin, C.S. and Beck, D.E. 1965. Observations on the biology, anatomy, and morphology of Otobius lagophilus. Brigham Young University Science Bulletin, Biological Series, 6: 1–19. Hewitt, C.G. 1915. A contribution to a knowledge of Canadian ticks. Transactions of the Royal Society of Canada, 9: 225–239. Hinrichsen, V.L., Whitworth, U.G., Breitschwerdt, E.B., Hegarty, B.C., and Mather, T.N. 2001. Assessing the association between the geographic distribution of deer ticks and seropositivity rates to various tick- transmitted disease organisms in dogs. Journal of the American Veterinary Medical Association, 218: 1092–1097. Hinton, H.E. 1967. The structure of the spiracles of the cattle tick, Boophilus microplus. Australian Journal of Zoology, 15: 941–945. Hixson, H. 1932. The life history and habits of Ixodes sculptus Neumann (Ixodidae). Iowa State College Journal of Science, 7: 35–42. Hixson, H. 1940. Field biology and environmental relationships of the Gulf Coast tick in southern Georgia. Journal of Economic Entomology, 33: 179–189. Hoberg, E.P. and Wehle, D.H.S. 1982. Host and geographic records of ectoparasites from Alaskan sea birds (Charadriiformes: Alcidae and Laridae). Canadian Journal of Zoology, 60: 472–475. Hoeve, J., Joachim, D.G., and Addison, E.M. 1988. Parasites of moose (Alces alces) from an agricultural area of eastern Ontario. Journal of Wildlife Diseases, 24: 371–374. Hoff, G.L., Yuill, T.M., Iverson, J.O., and Hanson, R.P. 1970. Selected microbial agents in snowshoe hares and other vertebrates in Alberta. Journal of Wildlife Diseases Proceedings of the Annual Conference, 6: 472–478. Hoffman, C.C. 1930. Monographias para la entomología médica de Mexico. Monografia Núm. 1. Los Argasidos de Mexico. Anales del Instituto Biología, 1: 135–164. Holden, K., Boothby, J.T., Anand, S., and Massung, R.F. 2003. Detection of Borrelia burgdorferi, Ehrlichia chaffeensis, and Anaplasma phagocytophilum in ticks (Acari: Ixodidae) from a coastal region of California. Journal of Medical Entomology, 40: 534–539. Holden, K., Boothby, J.T., Kasten, R.W., and Chomel, B.B. 2006. Co-detection of Bartonella henselae, Borrelia burgdorferi, and Anaplasma phagocytophilum in Ixodes pacificus ticks from California, USA. Vector- borne and Zoonotic Diseases, 6: 99–102. Holland, C.J., Ristic, M., Cole, A.I., Johnson, P., Baker, G., and Goetz, T. 1985. Isolation, experimental transmission, and characterization of causative agent of Potomac horse fever. Science, 227: 522–524. Holland, G.P. 1940. Notes on the ecology of Dermacentor andersoni in southern Alberta. Proceedings of the British Columbia Entomological Society, 36: 8–11. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 265

Holman, M.S., Caporale, D.A., Goldberg, J., Lacombe, E., Lubelczyk, C., Rand, P.W., and Smith, R.P. 2004. Anaplasma phagocytophilum, Babesia microti, and Borrelia burgdorferi in Ixodes scapularis, southern coastal Maine. Emerging Infectious Diseases, 10: 744–746. Homer, M.J., Aguilar-Delfin, I., Telford III, S.R., Krause, P.J., and Persing, D.H. 2000. Babesiosis. Clinical Microbiology Reviews, 13: 451–469. Homsher, P.J., Robbins, R.G., and Keirans, J.E. 1991. Scanning electron microscopy of Haller’s organ for subgeneric systematic studies in the genus Ixodes. In Modern acarology, volume 2. Edited by F. Dusbabek and V. Bukva. Academia, Prague Czech Republic and SPB Academic Publishing, The Hague, The Netherlands. Pp. 335–342. Homsher, P.J. and Sonenshine, D.E. 1975. Scanning electron microscopy of ticks for systematic studies. 2. Structure of Haller’s organ in Ixodes brunneus and Ixodes frontalis. Journal of Medical Entomology, 14: 93–97. Hoogstraal, H. 1985. Argasid and nuttalliellid ticks as parasites and vectors. Advances in Parasitology, 24: 135–238. Hooker, W.A., Bishopp, F.C., and Wood, H.P. 1912. The life history and bionomics of some North American ticks. United States Department of Agriculture, Bureau of Entomology Bulletin, 106: 1–239. Hoover, J.P., Walker, D.B., and Hedges, J.D. 1994. Cytauxzoonosis in cats: eight cases (1985–1992). Journal of the American Veterinary Medical Association, 205: 455–460. Hopla, C.E. 1955. Observations of the life history of a rabbit tick (Otobius lagophilus). Journal of the Kansas Entomological Society, 28: 114–116. Hopla, C.E. 1974. The ecology of tularemia. In Advances in veterinary science and comparative medicine, volume 8. Edited by C.A. Brankley and C.E. Cornelius. Academic Press, New York, United States of America. Pp. 25–53. Hopla, C.E. 1992. Ticks, viruses, and cliff swallows: a study in central Oklahoma. In Proceedings of the First International Conference on Tickborne Pathogens at the Host-Vector Interface: an agenda for research. Edited by U.G. Munderloh and T.J. Kurtti. University of Minnesota, St. Paul, Minnesota, United States of America. Pp. 42–46. Hopla, C.E. and Loye, J.E. 1983. The ectoparasites and microorganisms associated with cliff swallows in west- central Oklahoma. I. Ticks and fleas. Bulletin of the Society of Vector Ecology, 8: 111–121. Horak, I.G., Camicas, J.-L., and Keirans, J.E. 2002. The Argasidae, Ixodidae and Nuttalliellidae (Acari: Ixodida): a world list of valid tick names. Experimental and Applied Acarology, 28: 27–54. Howden, K.J., Geale, D.W., Paré, J., Golsteyn-Thomas, E.J., and Gajadhar, A.A. 2010. An update on bovine anaplasmosis (Anaplasma marginale) in Canada. Canadian Veterinary Journal, 51: 837–840. Howell, F.G. 1975. The roles of host-related stimuli in the behavior of Argas cooleyi (Acarina: Argasidae). Journal of Medical Entomology, 11: 715–723. Howell, F.G. 1976. Influence of the daily light cycle on the behavior ofArgas cooleyi (Acarina: Argasidae). Journal of Medical Entomology, 13: 99–106. Howell, F.G. and Chapman, B.R. 1976. Acarines associated with cliff swallow communities in northwest Texas. The Southwestern Naturalist, 21: 275–280. Howell, F.G. and George, J.E. 1973. Some effects of humidity on the behavior and water balance of Argas cooleyi (Acarina: Argasidae). Journal of Medical Entomology, 10: 459–469. Humphreys, F.A. and Campbell, A.G. 1947. Plague, Rocky Mountain spotted fever, and tularaemia surveys in Canada. Canadian Journal of Public Health, 38: 124–130. Hunter, W.D. and Hooker, W.A. 1907. Information concerning the North American fever tick, with notes on other species. United States Department of Agriculture, Bureau of Entomology Bulletin, 72: 1–87. International Commission of Zoological Nomenclature. 1978. Opinion 1107. Bulletin of Zoological Nomenclature, 35: 88. Jackson, J.O. and DeFoliart, G.R. 1975. Relationships of immature Dermacentor variabilis (Say) (Acari: Ixodidae) with the white-footed mouse, Peromyscus leucopus, in southwestern Wisconsin. Journal of Medical Entomology, 12: 409–412. Jackson, L.K., Gaydon, D.M., and Goddard, J. 1996. Seasonal activity and relative abundance of Amblyomma americanum in Mississippi. Journal of Medical Entomology, 33: 128–131. James, A.M., Freier, J.E., Keirans, J.E., Durden, L.A., Mertins, J.W., and Schlater, J.L. 2006. Distribution, seasonality, and hosts of the Rocky Mountain wood tick in the United States. Journal of Medical Entomology, 43: 17–24. James, A.M., Liveris, D., Wormser, G.P., Schwartz, I., Montecalvo, M.A., and Johnson, B.J. 2001. Borrelia lonestari infection after a bite by an Amblyomma americanum tick. Journal of Infectious Diseases, 183: 1810–1814. 266 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Jellison, W.L. 1974. Tularemia in North America 1930–1974. University of Montana, Missoula, Montana, United States of America. Jellison, W.L., Bell, E.J., Huebner, R.J., Parker, R.R., and Welsh, H.H. 1948. Q fever studies in southern California. IV. Occurrence of Coxiella burneti in the spinose ear tick, Otobius megnini. United States Public Health Reports, 63: 1483–1489. Jellison, W.L. and Gregson, J.D. 1950. Tick paralysis in northwestern United States and British Columbia. Rocky Mountain Medical Journal, 47: 1–28. Jellison, W.L. and Kohls, G.M. 1938. Tick-host anemia: a secondary anemia produced by Dermacentor andersoni Stiles. Journal of Parasitology, 24: 143–154. Jones, G.S. and Thomas, H.H. 1980. Ticks from mammals from Prince Edward Island, New Brunswick, northern Nova Scotia, and Gaspé Peninsula, Quebec. Canadian Journal of Zoology, 58: 1394–1397. Judd, W.W. 1953. Mammal host records of Acarina and Insecta from the vicinity of London, Ontario. Journal of Mammalogy, 34: 137–139. Judd, W.W. 1950. Mammal host records of Acarina and Insecta from the vicinity of Hamilton, Ontario. Journal of Mammalogy, 31: 357–358. Judd, W.W. 1954. Some records of ectoparasitic Acarina and Insecta from mammals in Ontario. Journal of Parasitology, 40: 483–484. Kaiser, M.N., Hoogstraal, H., and Kohls, G.M. 1964. The subgenus Persicargas new subgenus (Ixodoidea, Argasidae, Argas). 1. A. (P.) arboreus, new species, an Egyptian persicus-like parasite of wild birds, with a redefinition of the subgenus Argas. Annals of the Entomological Society of America, 57: 60–69. Keirans, J.E. and Clifford, C.M. 1978. The genus Ixodes in the United States: a scanning electron microscope study and key to the adults. Journal of Medical Entomology, Supplement, 2: 1–149. Keirans, J.E. and Clifford, C.M. 1983. Ixodes (Pholeoixodes) eastoni n. sp. (Acari: Ixodidae), a parasite of rodents and insectivores in the Black Hills of South Dakota, USA. Journal of Medical Entomology, 20: 90–98. Keirans, J.E., Clifford, C.M., and Corwin, D. 1976. Ixodes sigelos, n. sp. (Acarina: Ixodidae), a parasite of rodents in Chile, with a method for preparing ticks for examination by scanning electron microscopy. Acarologia, 18: 217–225. Keirans, J.E., Durden, L.A., and Hopla, C.E. 1993. Description of the immature stages of Ixodes (Pholeoixodes) baergi (Acari: Ixodidae), a parasite of cliff swallows in the United States. Journal of Medical Entomology, 30: 735–739. Keirans, J.E. and Litwak, T.R. 1989. Pictorial key to the adults of hard ticks, family Ixodidae (Ixodida: Ixodoidea), east of the Mississippi River. Journal of Medical Entomology, 26: 435–448. Keirans, J.E. and Pound, J.M. 2003. An annotated bibliography of the spinose ear tick, Otobius megnini (Dugès, 1883) (Acari: Ixodida: Argasidae) 1883–2000. Systematic and Applied Acarology Special Publications, 13: 1–68. Kennedy, M.J. 1986. Synopsis of the parasites of domestic mammals in Canada. Alberta Agriculture, Animal Health Division, Edmonton, Alberta, Canada. Kennedy, M.J. and Newman, R.A. 1986. Synopsis of the parasites of vertebrates in Canada: ectoparasites of terrestrial mammals. Alberta Agriculture, Animal Health Division, Edmonton, Alberta, Canada. Kerr, G.R. and Holmes, J.C. 1966. Parasites of mountain goats in west central Alberta. Journal of Wildlife Management, 30: 786–790. Kettyls, G.D., Verrall, V.M., Wilton, L.D., Clapp, J.B., Clarke, D.A., and Rublee, J.D. 1972. Arbovirus infections in man in British Columbia. Canadian Medical Association Journal, 106: 1175–1179. Kietzmann, G.E. 1987. Ectoparasites of black-tailed prairie dogs (Cynomys ludovicianus) from South Dakota. Journal of Wildlife Diseases, 23: 331–333. Kietzmann, G.E. and Kietzmann, E.A. 1987. Parasites associated with thirteen-lined ground squirrels (Spermophilus tridecemlineatus Mitchill) from Ames, Iowa. Proceedings of the Iowa Academy of Sciences, 94: 22–23. Killilea, M.E., Swei, A., Lane, R., Briggs, C.J., and Ostfeld, R.S. 2008. Spatial dynamics of Lyme disease: a review. EcoHealth, 5: 167–195. Kinsey, A.A., Durden, L.A., and Oliver, J.H. 2000. Tick infestations of birds in coastal Georgia and Alabama. Journal of Parasitology, 86: 251–254. Kleine-Tebbe, J., Heinatz, A., Gräser, I., Dautel, H., Hansen, G., Kespohl, S., et al. 2006. Bites of the European pigeon tick (Argas reflexus): risk of IgE-mediated sensitizations and anaphylactic reactions. Journal of Allergy and Clinical Immunology, 117: 190–195. Klich, M., Lankester, M.W., and Wu, K.W. 1996. Spring migratory birds (Aves) extend the northern occurrence of blacklegged tick (Acari: Ixodidae). Journal of Medical Entomology, 33: 581–585. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 267

Klompen, J.S.H. 1992. Comparative morphology of argasid larvae (Acari: Ixodida: Argasidae), with notes on phylogenetic relationships. Annals of the Entomological Society of America, 85: 541–560. Klompen, J.S.H., Keirans, J.E., Filippova, N.A., and Oliver, Jr., J.H. 1996. Idiosomal lyrifissures, setae, and small glands as taxonomic characters and potential indicators of ancestral segmentation patterns in larval Ixodidae (Acari: Ixodida). International Journal of Acarology, 22: 113–134. Klompen, J.S.H. and Oliver, J.H. 1993. Systematic relationships in the soft ticks (Acari: Ixodida: Argasidae). Systematic Entomology, 18: 313–331. Klotz, J.H., Klotz, S.A., and Pinnas, J.L. 2009. Animal bites and stings with anaphylactic potential. Journal of Emergency Medicine, 36: 148–156. Knight, R.L. and Marr, N.V. 1983. New distributional records of ticks (Ornithodoros concanensis and O. kelleyi) in Washington. Northwest Science, 57: 310–311. Ko, R.C. 1972a. Biology of Ixodes cookei Packard (Ixodidae) of groundhogs (Marmota monax Erxleben). Canadian Journal of Zoology, 50: 433–436. Ko, R.C. 1972b. The transmission of Ackertia marmotae Webster, 1967 (Nematoda: Onchocercidae) of groundhogs (Marmota monax) by Ixodes cookei. Canadian Journal of Zoology, 50: 437–450. Kocan, A.A. 1988. Tick paralysis. Journal of the American Veterinary Medical Association, 192: 1498–1500. Kocan, A.A., Mukolwe, S.W., Murphy, G.L., Barker, R.W., and Kocan, K.M. 1992. Isolation of Borrelia burgdorferi (Spirochaetales: Spirochaetaceae) from Ixodes scapularis and Dermacentor albipictus ticks (Acari: Ixodidae) in Oklahoma. Journal of Medical Entomology, 29: 630–633. Kocan, K.M. and Stiller, D. 1992. Development of Anaplasma ovis (Rickettsiales: Anaplasmataceae) in male Dermacentor andersoni (Acari: Ixodidae) transferred from infected to susceptible sheep. Journal of Medical Entomology, 29: 98–107. Kocan, K.M., Stiller, D., Goff, W.L., Claypool, P.L., Edwards, W., Ewing, S.A., et al. 1992. Development of Anaplasma marginale in male Dermacentor andersoni transferred from parasitemic to susceptible cattle. American Journal of Veterinary Research, 53: 499–507. Koch, C.L. 1844. Systematische Übersicht über die Ordnung der Zecken. Archives für Naturgeschichte, 10: 217–239. Koch, H.G. 1982. Seasonal incidence and attachment sites of ticks (Acari: Ixodidae) on domestic dogs in southeastern Oklahoma and northwestern Arkansas, USA. Journal of Medical Entomology, 19: 293–298. Koch, H.G. 1989. Survival of larvae of three tick species in wooded habitat of southeastern Oklahoma (Acari: Ixodidae). Journal of Agricultural Entomology, 6: 91–99. Koch, H.G. and Sauer, J.R. 1984. Quantity of blood ingested by four species of hard ticks (Acari: Ixodidae) fed on domestic dogs. Annals of the Entomological Society of America, 77: 142–146. Koffi, J.K., Leighton, P.A., Pelcat, Y., Trudel, L., Lindsay, L.R., Milord, F., and Ogden, N.H. 2012. Passive surveillance for I. scapularis ticks: enhanced analysis for early detection of emerging Lyme disease risk. Journal of Medical Entomology, 49: 400–409. Kogut, S.J., Thill, C.D., Prusinski, M.A., Lee, J., Bryon Backenson, P., Coleman, J.L., et al. 2005. Babesia microti, upstate New York. Emerging Infectious Diseases, 11: 476–478. Kohls, G.M. 1947. Notes on the tick Ixodes howelli Cooley and Kohls, with descriptions. Journal of Parasitology, 33: 57–61. Kohls, G.M. and Clifford, C.M. 1966. Three new species of Ixodes from Mexico and description of the male of I. auritulus Neumann, I. conepati Cooley and Kohls, and I. lasallei Mendez and Ortiz (Acarina: Ixodidae). Journal of Parasitology, 52: 810–820. Kohls, G.M. and Hoogstraal, H. 1960. Observations on the subgenus Argas (Ixodoidea, Argasidae, Argas). 2. A. cooleyi, new species from western North American birds. Annals of the Entomological Society of America, 53: 625–631. Kohls, G.M., Hoogstraal, H., Clifford, C.M., and Kaiser, M.N. 1970. The subgenus Persicargas (Ixodoidea, Argasidae, Argas). 9. Redescription and New World records of Argas (P.) persicus (Oken), and resurrection, redescription, and records of A. (P.) radiatus Railliet, A. (P.) sanchezi Dugès, and A. (P.) miniatus Koch, New World ticks misidentified asA. (P.) persicus. Annals of the Entomological Society of America, 63: 590–606. Kohls, G.M. and Ryckman, R.E. 1962. New distributional records of ticks associated with cliff swallows, Petrochelidon spp., in the United States. Journal of Parasitology, 48: 507–508. Kohls, G.M., Sonenshine, D.E., and Clifford, C.M. 1965. The systematics of the sub-family Ornithodorinae (Acarina: Argasidae). II. Identification of the larvae of the Western Hemisphere and descriptions of three new species. Annals of the Entomological Society of America, 58: 331–364. Kollars, T.M., Durden, L.A., Keirans J.E., and Oliver, Jr., J.H 1995. First records of Haemaphysalis (Aboimisalis) chordeilis, Ixodes (Ixodes) brunneus, and Ixodes (Pholeoixodes) banksi (Acari: Ixodidae) from Missouri. Journal of Entomological Science, 30: 511–512. 268 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Kollars, T.M. and Oliver, J.H. 2003. Host associations and seasonal occurrence of Haemaphysalis leporispalustris, Ixodes brunneus, I. cookei, I. dentatus, and I. texanus (Acari: Ixodidae) in southeastern Missouri. Journal of Medical Entomology, 40: 103–107. Kosoy, O.I., Lambert, A.J., Hawkinson, D.J., Pastula, D.M., Goldsmith, C.S., Hunt, D.C., and Staples, J.E. 2015. Novel thogotovirus associated with febrile illness and death, United States, 2014. Emerging Infectious Diseases, 21: 760–764. Krakowetz, C.N., Dibernardo, A., Lindsay, L.R., and Chilton, N.B. 2014. Two Anaplasma phagocytophilum strains in Ixodes scapularis ticks, Canada. Emerging Infectious Diseases, 20: 2064–2067. Kramer, P. and Neumann, C.J. 1883. Acariden, während der Vega-Expedition eingesammelt. Vega-Expeditiones Vetensk (Stockholm), 3: 519–529. Krause P.J., Narasimhan, S., Wormser, G.P., Rollend, L., Fikrig, E., Lepore, T., et al. 2013. Human Borrelia miyamotoi infection in the United States. New England Journal of Medicine, 368: 291–293. Krinsky, W.L. 1979. Development of the tick Ixodes dammini (Acarina: Ixodidae) in the laboratory. Journal of Medical Entomology, 16: 354–355. Kuno, G., Artsob, H., Karabatsos, N., Tsuchiya, K.R., and Chang, G.J.J. 2001. Genomic sequencing of deer tick virus and phylogeny of Powassan-related viruses of North America. American Journal of Tropical Medicine and Hygiene, 65: 671–676. Labzoffsky, N.A. and Sprent, J.A. 1952. Tularemia among beaver and muskrat in Ontario. Canadian Journal of Medical Science, 30: 250–255. Lackman, D.B., Parker, R.R., and Gerloff, R.K. 1949. Serological characteristics of a pathogenic rickettsia occurring in Amblyomma maculatum. United States Public Health Reports, 64: 1342–1349. Lambert A.J., Kosoy, O., Velez, J.O., Russell, B.J., and Lanciotti, R.S. 2007. Detection of Colorado tick fever viral RNA in acute human serum samples by a quantitative real-time RT-PCR assay. Journal of Virological Methods, 140: 43–48. Lancaster, J.L. and Meisch, M.V. 1986. Arthropods in livestock and poultry production. Ellis Horwood, New York, New York, United States of America. Lane, R.S. 1990. Seasonal activity of two human-biting ticks. California Agriculture, 44: 23–25. Lane, R.S. and Burgdorfer, W. 1988. Spirochetes in mammals and ticks (Acari: Ixodidae) from a focus of Lyme borreliosis in California. Journal of Wildlife Diseases, 24: 1–9. Lane, R.S., Foley, J.E., Eisen, L., Lennette, E.T., and Peot, M.A. 2001. Acarologic risk of exposure to emerging tick-borne bacterial pathogens in a semi-rural community in northern California. Vector-borne and Zoonotic Diseases, 1: 197–210. Lane, R.S., Kucera, T.F., Barrett, R.H., Mun, J., Wu, C., and Smith, V.S. 2006. Wild turkey (Meleagris gallopavo) as a host of ixodid ticks, lice, and Lyme disease spirochaetes (Borrelia burgdorferi sensu lato) in California state parks. Journal of Wildlife Diseases, 42: 759–771. Lane, R.S. and Loye, J.E. 1989. Lyme disease in California: interrelationship of Ixodes pacificus (Acari: Ixodidae), the western fence lizard (Sceloporus occidentalis), and Borrelia burgdorferi. Journal of Medical Entomology, 26: 272–278. Lane, R.S., Mun, J., Peribanez, M.A., and Stubbs, H.A. 2007. Host-seeking behavior of Ixodes pacificus (Acari: Ixodidae) nymphs in relation to environmental parameters in dense-woodland and woodland grass habitats. Journal of Vector Ecology, 32: 342–357. Lane, R.S., Peek, J., and Donaghey, P.J. 1984. Tick (Acari: Ixodidae) paralysis in dogs from northern California: acarological and clinical findings. Journal of Medical Entomology, 21: 321–326. Lane, R.S. and Quistad, G.B. 1998. Borreliacidal factor in the blood of the western fence lizard (Sceloporus occidentalis). Journal of Parasitology, 84: 29–34. Lane, R.S., Steinlein, D.B., and Mun, J. 2004. Human behaviors elevating exposure to Ixodes pacificus (Acari: Ixodidae) nymphs and their associated bacterial zoonotic agents in a hardwood forest. Journal of Medical Entomology, 41: 239–248. Lang, J.D. 1999. Ixodid ticks (Acari, Ixodidae) found in San Diego County, California. Journal of Vector Ecology, 24: 61–69. Lankester, M.W. 1987. Pests, parasites and diseases of moose (Alces alces) in North America. In, Swedish wildlife research, supplement 1. Edited by G. Gorunsson and S. Lavsund. Viltrevy, Stockholm, Sweden. Pp. 461–489. Lankester, M.W., Scandrett, W.B., Goldsteyn-Thomas, E.J., Chilton, N.B., and Gajadhar, A.A. 2006. Experimental transmission of bovine anaplasmosis (caused by Anaplasma marginale) by means of Dermacentor variabilis and D. andersoni collected in western Canada. Canadian Journal of Veterinary Research, 71: 271–277. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 269

Lappin, M.R., Breitschwerdt, E.B., Jensen, W.A., Dunnigan, B., Rha, J.Y., Williams, C.R., et al. 2004. Molecular and serologic evidence of Anaplasma phagocytophilum infection in cats in North America. Journal of the American Veterinary Medical Association, 225: 893–896. Larimore, R.W. 1987. Synchrony of cliff swallow nesting and development of the tick, Ixodes baergi. The Southwestern Naturalist, 32: 121–126. Latreille, P.A. 1804 Histoire naturelle, générale et particulière, des crustacés et des insects. VIII. F. Dufart, Paris, France. Latreille, P.A. 1806. Genera cristaceorum et insectorum secundum ordinem naturalem in familias disposita, inconibus exemplisque plurimis explicata, volume 1. Amand Koenig, Paris, France. Laurance, W.F. and Coan, M.E. 1987. Microhabitat and demographic correlates of tick parasitism in a northern Great Basin small mammal community. American Midland Naturalist, 118: 1–9. Lausen, C.L. 2005. First record of hosts for tick Carios kelleyi (Acari: Ixodidae: Argasidae) in Canada and Montana. Journal of Medical Entomology, 42: 497–501. Lawrence, W.H., Fay, L.D., and Graham, S.A. 1956. A report on the beaver die-off in Michigan. Journal of Wildlife Management, 20: 184–187. Leighton, P.A., Koffi, J.K., Pelcat, Y., Lindsay, L.R., and Ogden, N.H. 2012. Predicting the speed of tick invasion: an empirical model of range expansion for the Lyme disease vector Ixodes scapularis in Canada. Journal of Applied Ecology, 49: 457–464. Lester, S.J., Breirschwerdt, E.B., Collis, C.D., and Hegarty, B.C. 2005. Anaplasma phagocytophilum infection (granulocytic anaplasmosis) in a dog from Vancouver Island. The Canadian Veterinary Journal, 46: 825–827. Levine, J.F., Wilson, M.L., and Spielman, A. 1985. Mice as reservoirs of the Lyme disease spirochete. American Journal of Tropical Medicine and Hygiene, 34: 355–360. Levine, N.D. 1988. Progress in taxonomy of the Apicomplexan protozoa. Journal of Protozoology, 35: 518–520. Levine, P.P. and Goble, F.C. 1947. Parasitism and disease. In The ruffed grouse. Life history, propagation and management. Edited by G. Bump, R.W. Darrow, F.C. Edminster, and W.F. Crissey. New York State Conservation Department, Albany, New York, United States of America. Pp. 401–442. Li, X., Peavey, C.A., and Lane, R.S. 2000. Density and spatial distribution of Ixodes pacificus (Acari: Ixodidae) in two recreational areas in north coastal California. American Journal of Tropical Medicine and Hygiene, 62: 415–422. Lillehaug, A., Mehl, R., and Gjerde, B. 2002. Importation of Dermacentor albipictus into Europe. The Veterinary Record, 151: 94–95. Lin, T., Oliver, J.H., Gao, L., Kollars, T.M., and Clark, K.L. 2001. Genetic heterogeneity of Borrelia burgdorferi sensu lato in the southern United States based on restriction length polymorphism and sequence analysis. Journal of Clinical Microbiology, 39: 2500–2507. Lindquist, E.E. and Evans, G.O. 1965. Taxonomic concepts in the Ascidae, with a modified setal nomenclature for the idiosoma of the Gamasina (Acarina: Mesostigmata). Memoirs of the Entomological Society of Canada, 47: 1–64. Lindquist, E.E., Krantz, G.W., and Walter, D.E. 2009. Classification.In A Manual of Acarology, 3rd edition. Edited by G.W. Krantz and D.E. Walter, Texas Tech University Press, Lubbock, Texas, United States of America. Pp. 97–103. Lindquist, E.E., Wu, K.W., and Redner, J.H. 1999. A new species of the tick genus Ixodes (Acari: Ixodidae) parasitic on mustelids (Mammalia: Carnivora) in Canada. The Canadian Entomologist, 131: 151–170. Lindsay, L.R. 1995. Factors limiting the distribution of the blacklegged tick, Ixodes scapularis Say, in Ontario, Canada. Ph.D. Dissertation. University of Guelph, Guelph, Ontario, Canada. Lindsay, L.R., Barker, I.K., Surgeoner, G.A., McEwen, S.A., and Campbell, G.D. 1997. Duration of Borrelia burgdorferi infectivity in white-footed mice for the tick vector Ixodes scapularis under laboratory and field conditions in Ontario. Journal of Wildlife Diseases,33: 766–775. Lindsay, L.R., Barker, I.K., Surgeoner, G.A., McEwen, S.A., Elliott, L.A., and Kolar, J. 1991. Apparent incompetence of Dermacentor variabilis (Acari: Ixodidae) and fleas (Insecta: Siphonaptera) as vectors of Borrelia burgdorferi in an Ixodes dammini endemic area of Ontario, Canada. Journal of Medical Entomology, 28: 750–753. Lindsay, L.R., Barker, I.K., Surgeoner, G.A., McEwen, S.A., Gillespie, T.J., and Addison, E.M. 1998. Survival and development of the different life stages of Ixodes scapularis (Acari: Ixodidae) held within four habitats on Long Point, Ontario, Canada. Journal of Medical Entomology, 35: 189–199. Lindsay, L.R., Barker, I.K., Surgeoner, G.A., McEwen, S.A., Gillespie, T.J., and Robinson, J.T. 1995. Survival and development of Ixodes scapularis (Acari: Ixodidae) under various climatic conditions in Ontario, Canada. Journal of Medical Entomology, 32: 143–152. 270 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Lindsay, L.R., Bernat, K., and Dibernardo, A. 2014. Laboratory diagnostics for Lyme disease. Canadian Communicable Diseases Report, 40: 209. Lindsay, L.R., Mathison, S.W., Barker, I.K., McEwen, S.A., Gillespie, T.J., and Surgeoner, G.A. 1999a. Microclimate and habitat in relation to Ixodes scapularis (Acari: Ixodidae) populations on Long Point, Ontario, Canada. Journal of Medical Entomology, 36: 255–262. Lindsay, L.R., Mathison, S.W., Barker, I.K., McEwen, S.A., and Surgeoner, G.A. 1999b. Abundance of Ixodes scapularis (Acari: Ixodidae) larvae and nymphs in relation to host density and habitat on Long Point, Ontario. Journal of Medical Entomology, 36: 243–254. Linnaeus, C. 1758. Systema naturae. 10th edition, tome 1. Laurentii Salvii, Stockholm, Sweden. Little, P.B., Thorsen, J., Moore, W., and Weninger, N. 1985. Powassan viral encephalitis: a review and experimental studies in the horse and rabbit. Veterinary Pathology, 22: 500–507. Littman, M.P., Goldstein, R.E., Labato, M.A., Lappin, M.R., and Moore, G.E. 2006. ACVIM small animal consensus statement on Lyme disease in dogs: diagnosis, treatment and prevention. Journal of Veterinary Internal Medicine, 20: 422–434. Loftis, A.D., Gill, J.S., Schriefer, M.E., Levin, M.L., Eremeeva, M.E., Gilchrist, M.J.R., and Dasch, G.A. 2005. Detection of Rickettsia, Borrelia, and Bartonella in Carios kelleyi (Acari: Argasidae). Journal of Medical Entomology, 42: 473–480. Loomis, E.C. 1953. A note on Otobius lagophilus (Acarina: Argasidae). The Pan-Pacific Entomologist,29: 198. Loomis, E.C. 1961. Life histories of ticks under laboratory conditions (Acarina: Ixodidae and Argasidae). Journal of Parasitology, 47: 91–99. Loye, J.E. and Hopla, C.E. 1983. Ectoparasites and microorganisms associated with the cliff swallow in west- central Oklahoma. 2. Life history patterns. Bulletin of the Society of Vector Ecologists, 8: 79–84. Lubelczyk, C.B., Hanson, T., Lacombe, E.H., Holman, M.S., and Keirans, J.E. 2007. First U.S. record of the hard tick Ixodes (Pholeoixodes) gregsoni Lindquist, Wu, and Redner. Journal of Parasitology, 93: 718–719. Luttrell, M.P., Creekmore, L.H., and Mertins, J.W. 1996. Avian tick paralysis caused by Ixodes brunneus in the southeastern United States. Journal of Wildlife Diseases, 32: 133–136. Lysyk, T.J. 2010. Tick paraslysis caused by Dermacentor andersoni (Acari: Ixodidae) is a heritable trait. Journal of Medical Entomology 47: 210–214. Lysyk, T.J. 2013. Movement of male Dermacentor andersoni (Acari: Ixodidae) among cattle. Journal of Medical Entomology, 50: 977–985. Lysyk, T.J. and Majak, W. 2003. Increasing the paralyzing ability of a laboratory colony of Dermacentor andersoni Stiles. Journal of Medical Entomology, 40: 185–194. Lysyk, T.J., Majak, W., and Veira, D.M. 2005. Prefeeding Dermacentor andersoni (Acari: Ixodidae) on cattle with prior exposure may inhibit detection of tick paralysis by using hamster bioassay. Journal of Medical Entomology, 42: 376–382. Lysyk, T.J., Veira, D.M., and Majak, W. 2009. Cattle can develop immunity to paralysis caused by Dermacentor andersoni. Journal of Medical Entomology, 46: 358–366. Mackenzie, A.M.R., E. Rossier, E., Polley, J.R., and Corber, S.J. 1979. Rocky Mountain spotted fever – Ontario. Canada Diseases Weekly Report, 5: 130–132. Madigan, J.E. and Pusterla, N. 2000. Ehrlichial diseases. Veterinary Clinics of North America Equine Practice, 16: 487–499, ix. Magnarelli, L.A., Anderson, J.F., Apperson, C.S., Fish, D., Johnson, R.C., and Chappell, W.A. 1986. Spirochetes in ticks and antibodies to Borrelia burgdorferi in white-tailed deer from Connecticut, New York State, and North Carolina. Journal of Wildlife Diseases, 22: 178–188. Magnarelli, L.A. and Swihart, R.K. 1991. Spotted fever group rickettsiae or Borrelia burgdorferi in Ixodes cookei in Connecticut. Journal of Clinical Microbiology, 29: 1520–1522. Mail, G.A. 1942. Lethal temperatures for Dermacentor andersoni Stiles and other ticks in British Columbia. Journal of Economic Entomology, 35: 562–564. Mail, G.A. and Gregson, J.D. 1938. Tick paralysis in British Columbia. Canadian Medical Association Journal, 39: 532–537. Main, A.J., Carey, A.B., and Downs, W.G. 1979. Powassan virus in Ixodes cookei and Mustelidae in New England. Journal of Wildlife Diseases, 15: 585–591. Main, A.J., Downs, W.G., Shope, R.E., and Wallis, R.C. 1973. Great Island and Bauline: two new Kemerovo group orbiviruses from Ixodes uriae in eastern Canada. Journal of Medical Entomology, 10: 229–235. Main, A.J., Downs, W.G., Shope, R.E., and Wallis, R.C. 1976a. Avalon and Clo Mor: two new Sakhalan group viruses from the north Atlantic. Journal of Medical Entomology, 13: 309–315. Main, A.J., Downs, W.G., Shope, R.E., and Wallis, R.C. 1976b. Avian arboviruses of the Witless Bay Seabird Sanctuary, Newfoundland, Canada. Journal of Wildlife Diseases, 12: 182–194. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 271

Mangin, S., Gauthier-Clerc, M., Frenot, Y., Gendner, J.-P., and Le Maho, Y. 2003. Ticks Ixodes uriae and the breeding performance of a colonial seabird, king penguin Aptenodytes patagonicus. Journal of Avian Biology, 34: 30–34. Mannelli, A., Kitron, U., Jones, C.J., and Slajchert, T.L. 1993. Role of eastern chipmunk as a host for immature Ixodes dammini (Acari: Ixodidae) in northwestern Illinois. Journal of Medical Entomology, 30: 87–93. Mans, B.J., Gothe, R., and Neitz, A.W.H. 2004. Biochemical perspectives on paralysis and other forms of toxicoses caused by ticks. Parasitology, 129: S95–S111. Manweiler, S.A., Lane, R.S., and Tempelis, C.H. 1992. The western fence lizard Sceloporus occidentalis: evidence of field exposure toBorrelia burgdorferi in relation to infestation by Ixodes pacificus (Acari: Ixodidae). American Journal of Tropical Medicine and Hygiene, 47: 328–336. Marrie, T.J., Campbell, N., McNeil, S.A., Webster, D., and Hatchette, T.F. 2008. Q fever update, Maritime Canada. Emerging Infectious Diseases, 14: 67–69. Martell, A.M., Yescott, R.E., and Dodds, D.G. 1969. Some records for Ixodidae of Nova Scotia. Canadian Journal of Zoology, 47: 183–184. Martin, A.R., Brown, G.K., Dunstan, R.H., and Roberts, T.K. 2005. Anaplasma platys: an improved PCR for its detection in dogs. Experimental Parasitology, 109: 176–180. Marx, G. 1895. Obituary notice of his work. Proceedings of Entomological Society of Washington, 3: 195–201, 1 plate. Massung, R.F., Courtney, J.W., Hiratzka, S.L., Pitzer, V.E., Smith, G., and Dryden, R.L. 2005. Anaplasma phagocytophilum in white-tailed deer. Emerging Infectious Diseases, 11: 1604–1606. Masters, E.J., Grigery, C.N., and Masters, R.W. 2008. STARI, or Masters disease: lone star tick-vectored Lyme-like illness. Infectious Disease Clinics of North America, 22: 361–376 + viii. Mather, T.N. and Spielman, A. 1986. Diurnal detachment of immature deer ticks (Ixodes dammini) from nocturnal hosts. American Journal of Tropical Medicine and Hygiene, 35: 182–186. Mathew, J.S., Ewing, S.A., Panciera, R.J., and Woods, J.P. 1998. Experimental transmission of Hepatozoon americanum Vincent-Johnson et al., 1997 to dogs by the Gulf Coast tick, Amblyomma maculatum Koch. Veterinary Parasitology, 80: 1–14. Matsumoto, K., Berrada, Z.L., Klinger, E., Goethert, H.K., and Telford, S.R. 2008. Molecular detection of Bartonella schoenbuchensis from ectoparasites of deer in Massachusetts. Vector-borne and Zoonotic Diseases, 8: 549–554. Maupin, G.O., Fish, D., Zultowsky, J., Campos, E.G., and Piesman, J. 1991. Landscape ecology of Lyme disease in a residential area of Westchester County, New York. American Journal of Epidemiology, 133: 1105–1113. Maurin, M. and Raoult, D. 1999. Q fever. Clinical Microbiology Reviews, 12: 518–553. McCoy, K.D., Bouliner, T., Tirard, C., and Michalakis, Y. 2001. Host specificity of a generalist parasite: genetic evidence of sympatric host races in the seabird tick Ixodes uriae. Journal of Evolutionary Biology, 14: 395–405. McCoy, K.D, Boulinier, T., Tirard, C., and Michalakis, Y. 2003a. Host-dependent genetic structure of parasite populations: differential dispersal of seabird tick host races. Evolution, 57: 288–296. McCoy, K.D. and Tirard, C. 2000. Isolation and characterization of microsatellites in the seabird ectoparasite Ixodes uriae. Molecular Ecology, 9: 2213–2214. McCoy, K.D. and Tirard, C. 2002. Reproductive strategies of the seabird tick Ixodes uriae (Acari: Ixodidae). Journal of Parasitology, 88: 813–816. McCoy, K.D., Tirard, C., and Michalakis, Y. 2003b. Spatial genetic structure of the ectoparasite Ixodes uriae within breeding cliffs of its colonial seabird host. Heredity, 91: 422–429. McEnroe, W.D. 1982. Relaxed regulation of Dermacentor variabilis (Say) in Nova Scotia. Canadian Journal of Zoology, 60: 2529–2532. McEnroe, W.D. and Specht, H.B. 1987. Regulation of Dermacentor variabilis by limited dispersion of larvae from the egg mass (Acari: Ixodidae). Folia Parasitologica, 34: 309–310. McIvor, B.C. 1937. A new species of Ornithodoros tick from California (Acarina: Ixodoidea). Journal of Parasitology, 23: 363–367. McKiel, J.A., Bell, E.J., and Lackman, D.B. 1967. Rickettsia canada: a new member of the typhus group of rickettsiae isolated from Haemaphysalis leporispalustris ticks in Canada. Canadian Journal of Microbiology, 13: 503–510. McLean, D.M., Chernesky, M.A., Chernesky, S.J., Goodard, E.J., Ladyman, S.R., Peers, R.R., and Purvin- Good, K.W. 1969. Arbovirus prevalence in the east Kootenay Region, 1968. Canadian Medical Association Journal, 100: 320–326. 272 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

McLean, D.M., Crawford, M.A., Ladyman, S.R., Peers, R.R., and Purvin-Good, K.W. 1970. California encephalitis and Powassan virus activity in British Columbia, 1969. American Journal of Epidemiology, 92: 266–272. McLean, D.M. and Donahue, W.L. 1959. Powassan virus: isolation of virus from a fatal case of encephalitis. Canadian Medical Association Journal, 80: 708–711. McLean, D.M., Ladyman, S.R., and Purvin-Good, K.W. 1968. Westward expansion of Powassan virus prevalence. Canadian Medical Association Journal, 98: 946–949. McLean, D.M. and Larke, R.P.B. 1963. Powassan and Silverwater viruses: ecology of two Ontario arboviruses. Canadian Medical Association Journal, 26: 182–185. McLean, D.M. and Quantz, E.J. 1964. Powassan virus: field investigations during the summer of 1963. American Journal of Tropical Medicine and Hygiene, 13: 747–753. McLean, D.M., Smith, P.A., Livingstone, S.E., Wilson, W.E, and Wilson, A.G. 1966. Powassan virus: summer infection cycle. Canadian Medical Association Journal, 94: 532–536. McLean, R.G., Francy, D.B., Bowen, G.S., Bailey, R.E., Calisher, C.H., and Barnes, A.M. 1981. The ecology of Colorado tick fever in Rocky Mountain National Park in 1974. I. Objectives, study design, and summary of principal findings. American Journal of Tropical Medicine and Hygiene,30: 483–489. McLeod, J.A. 1933. A parasitological survey of the genus Citellus in Manitoba. Canadian Journal of Research, 9: 108–127. McQuiston, J.H. and Childs, J.E. 2002. Q fever in humans and animals in the United States. Vector-borne and Zoonotic Diseases, 2: 179–191. McQuiston, J.H., Childs, J.E., and Thompson, H.A. 2002. Q fever. Journal of the American Veterinary Medical Association, 221: 796–799. McQuiston, J.H., Holman, R.C., McCall, C.L., Childs, J.E., Swerdlow, D.L., and Thompson, H.A. 2006. National surveillance and the epidemiology of human Q fever in the United States, 1978–2004. American Journal of Tropical Medicine and Hygiene, 75: 36–40. McQuiston, J.H., Zemtsova, F., Perniciaro, J., Hutson, M., Singleton, J., Nicholson, W.L., and Levin, M.L. 2012. Afebrile spotted fever group Rickettsia infection after a bite from a Dermacentor variabilis tick infected with Rickettsia montanensis. Vector-borne and Zoonotic Diseases, 12: 1059–1061. Meinkoth, J.H. and Kocan, A.A. 2005. Feline cytauxzoonosis. Veterinary Clinics of North America Small Animal Practice, 35: 89–101. Meinkoth, J.H., Kocan, A.A., Whitworth, L., Murphy, G., Fox, J.C., and Woods, J.P. 2000. Cats surviving natural infection with Cytauxzoon felis: 18 cases (1997–1998). Journal of Veterinary Internal Medicine, 14: 521–525. Merten, H.A. and Durden, L.A. 2000. A state-by-state survey of ticks recorded from humans in the United States. Journal of Vector Ecology, 25: 102–113. Messick, J.B. 2003. New perspectives about hemotrophic mycoplasma (formerly, Haemobartonella and Eperythrozoon species) infections in dogs and cats. Veterinary Clinics of North America Small Animal Practice, 33: 1453–1465. Micher, K.M. and Rockett, C.L. 1993. Field investigations on the American dog tick, Dermacentor variabilis, in northwest Ohio (Acari: Ixodidae). Great Lakes Entomologist, 26: 61–70. Morbey, Y.E. 1996. The abundance and effects of ticks (Ixodes uriae) on nestling Cassin’s auklets (Ptychoramphus aleuticus) at Triangle Island, British Columbia. Canadian Journal of Zoology, 74: 1585–1589. Morissette E., Massung, R.F., Foley, J.E., Alleman, A.R., Foley, P., and Barbet, A.F. 2009. Diversity of Anaplasma phagocytophilum strains, USA. Emerging Infectious Diseases, 15: 928–931. Morshed M.G., Scott, J.D., Banerjee, S.N., Banerjee, M., Fitzgerald, T., Fernando, K., et al. 1999. First isolation of Lyme disease spirochete, Borrelia burgdorferi, from blacklegged tick, Ixodes scapularis, removed from a bird in Nova Scotia, Canada. Canada Communicable Disease Report, 25: 153–155. Morshed, M.G., Scott, J.D., Fernando, K., Beati, L., Mazerolle, D.F., Geddes, G., and Durden, L.A. 2005. Migratory songbirds disperse ticks across Canada, and first isolation of the ymeL disease spirochete, Borrelia burgdorferi, from the avian tick, Ixodes auritulus. Journal of Parasitology, 91: 780–790. Morshed M.G., Scott, J.D., Fernando, K., Geddes, G., McNabb, A., Mak, S., and Durden, L.A. 2006. Distribution and characterization of Borrelia burgdorferi isolates from Ixodes scapularis and presence in mammalian hosts in Ontario, Canada. Journal of Medical Entomology, 43: 762–773. Murray, M.D. and Vestjens, M.J.M. 1967. Studies on the ectoparasites of seals and penguins III. The distribution of the tick Ixodes uriae White and the fleaParapsyllus magellanicus heardi de Meillon on Macquarie Island. Australian Journal of Zoology, 15: 715–725. Murrell, B.P., Durden, L.A., and Cook, J.A. 2003. Host associations of the tick, Ixodes angustus (Acari: Ixodidae), on Alaskan mammals. Journal of Medical Entomology, 40: 682–685. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 273

Muzaffar, S.B. and Jones, I.L. 2004. Parasites and diseases of the auks (Alcidae) of the world and their ecology – a review. Marine Ornithology, 32: 121–146. Muzaffar, S.B., and Jones, I.L. 2007. Activity periods and questing behavior of the seabird tick Ixodes uriae (Acari: Ixodidae) on Gull Island, Newfoundland: the role of puffin chicks. Journal of Parasitology, 93: 258–264. Nadelman, R.B. and Wormser, G.P. 2007. Reinfection in patients with Lyme disease. Clinical Infectious Diseases, 45: 1032–1038. Nagar, S.K. 1968. The value of ovipositional ability in tick taxonomy. Acarologia, 10: 614– 620. Nelder, M.P., Russell, C., Lindsay, L.R., Dhar, B., Patel, S.N., Johnson, S., et al. 2014. Population-based passive tick surveillance and detection of expanding foci of blacklegged ticks Ixodes scapularis and the Lyme disease agent Borrelia burgdorferi in Ontario, Canada. Public Library of Science One 9:e105358. doi: 10.1371/journal.pone.0105358. Neumann, I.G. 1896. Révision de la famille des ixodidés. I. Argasines. Mémoires de la Société de zoologiques de France, 9: 1–44. Neumann, I.G. 1897. Révision de la famille des ixodidés (2e mémoire). Mémoires de la Société de zoologiques de France, 10: 324–420. Neumann, L.G. 1899. Révision de la famille des ixodidés. Mémoires de la Société Zoologique de France, 12: 107–294. Neumann, L.G. 1901. Révision de la famille des ixodides. Mémoires de la Société Zoologique de France, 14: 249–372. Neumann, I.G. 1902. Notes sur les ixodidés. Archives de Parasitologie, 6: 109–128. Neumann, L.G. 1904. Notes sur les ixodidés. II. Archives de Parasitologie, 8: 444–464. Neumann, L.G. 1905. Notes sur les ixodidés. III. Archives de Parasitologie, 9: 225–241. Neumann, I.G. 1908. Notes sur les ixodidés. VII. Notes from the Leyden Museum, 30: 73–91. Neumann, L.G. 1911. Ixodidae. Das Tierreich, 26: 1–169. Newhouse, V.F. 1983. Variations in population density, movement, and rickettsial infection rates in a local population of Dermacentor variabilis (Acarina: Ixodidae) ticks in the piedmont of Georgia. Environmental Entomology, 12: 1737–1746. Newhouse, V.F., McKiel, J.A., and Burgdorfer, W. 1964. California encephalitis, Colorado tick fever and Rocky Mountain spotted fever in eastern Canada. Serological evidence. Canadian Journal of Public Health, 55: 257–261. Nicholls, T.H. and Callister, S.M. 1996. Lyme disease spirochetes in ticks collected from birds in midwestern United States. Journal of Medical Entomology, 33: 379–384. Niebylski, M.L., Peacock, M.G., and Schwan, T.G. 1999. Lethal effect of Rickettsia rickettsii on its tick vector (Dermacentor andersoni). Applied Environmental Microbiology, 65: 773–778. Nieto, N.C., Dabritz, H., Foley, P., Draznovich, N., Calder, L., Adjemian, J., et al. 2007. Ectoparasite diversity and exposure to vector-borne disease agents in wild rodents in central coastal California. Journal of Medical Entomology, 44: 328–335. Nieto, N.C. and Foley, J.E. 2008. Evaluation of squirrels (Rodentia: Sciuridae) as ecologically significant hosts for Anaplasma phagocytophilum in California. Journal of Medical Entomology, 45: 763–769. Norris, D.E., Klompen, J.S.H., Keirans, J.E., Lane, R.S., Piesman, J., and Black, W.C. 1997. Taxonomic status of Ixodes neotomae and I. spinipalpis (Acari: Ixodidae) based on mitochondrial DNA evidence. Journal of Medical Entomology, 34: 696–703. Nuttall, G.H.F. 1912. Notes on ticks. II. (1) New species (Amblyomma, Haemaphysalis) (2) Ixodes putus: description of hitherto unknown larval stage. Parasitology, 5: 50–60. Nuttall, G.H.F. 1916. Notes on ticks, IV. Relating to the genus Ixodes and including a description of three new species. Parasitology, 8: 294–337. Nuttall, G.H.F. and Warburton, C. 1908. On a new genus of the Ixodoidea, together with a description of eleven new species of ticks. Proceedings of the Cambridge Philosophical Society, 14: 392–416. Nuttall, G.H.F. and Warburton, C. 1911. Ticks: a monograph of the Ixodoidea, part 2. Cambridge University Press, London, United Kingdom. Nuttall, G.H.F. and Warburton, C. 1915. Ticks: a monograph of the Ixodoidea, part 3. The genus Haemaphysalis. Cambridge University Press. London, United Kingdom. Pp. 349–550. Obenchain, F.D., Leahy, M.G., and Oliver, J.H. 1980. Implications of tick size on the quantification of engorgement in female Dermacentor variabilis. Journal of Parasitology, 66: 282–286. O’Brien, S.F., Delage, G., Scalia, V., Lindsay, R., Bernier, F., Dubuc, S., Germain, M., Pilot, G., Yi, Q.-L., and Fearon, M.A. 2016. Seroprevalence of Babesia microti infection in Canadian blood donors. Transfusion, 56: 237–243. 274 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Ogden, N.H., Barker, I.K., Beauchamp, G., Brazeau, S., Charron, D.F., Maarouf, A., et al. 2006a. Investigation of ground level and remote-sensed data for habitat classification and prediction of survival ofIxodes scapularis in habitats in southeastern Ontario. 2006. Journal of Medical Entomology, 43: 403–414. Ogden, N.H., Bigras-Poulin, M., Barker, I.K., Lindsay, L.R., , Maarouf, A., O’Callaghan, C.J., et al. 2006b. Climate change and the potential for range expansion of the Lyme disease vector Ixodes scapularis in Canada. International Journal for Parasitology, 36: 63–70. Ogden, N H., Bigras-Poulin, M., O’Callaghan, J.O., Barker, I.K., Kurtenbach, K., Lindsay, L.R., and Charron, D.F. 2008a. Vector seasonality, host infection dynamics and fitness of pathogens transmitted by the tick Ixodes scapularis. Parasitology, 134: 209–227. Ogden, N.H., Bigras-Poulin, M., O’Callaghan, C.J., Barker, I.K., Lindsay, L.R., Maarouf, A., et al. 2005. A dynamic population model to investigate effects of climate on geographic range and seasonality of the tick Ixodes scapularis. International Journal for Parasitology, 35: 375–389. Ogden, N.H., Koffi, J.K., Pelcat, Y., and Lindsay, L.R. 2014. Environmental risk from Lyme disease in central and eastern Canada: a summary of recent surveillance information. Canadian Communicable Disease Report, 40: 74–82. Ogden, N.H., Lindsay, L.R., Charron, D.F., Beauchamp, G., Maarouf, A., O’Callaghan, C.J., et al. 2004. Investigation of the relationships between temperature and development rates of the tick Ixodes scapularis (Acari: Ixodidae) in the laboratory and field. Journal of Medical Entomology, 41: 622–633. Ogden, N.H., Lindsay, L.R., Hanincova, K., Barker, I.K., Bigras-Poulin, M., Charron, D.F., et al. 2008b. The role of migratory birds in introduction and range expansion of Ixodes scapularis ticks, and Borrelia burgdorferi and Anaplasma phagocytophilum in Canada. Applied Environmental Microbiology, 74: 1780–1790. Ogden, N.H., Lindsay, L.R., and Leighton, P.A. 2013. Predicting the rate of invasion of the agent of Lyme disease Borrelia burgdorferi. Journal of Applied Ecology, 50: 510–518. Ogden, N.H., Lindsay, L.R., Morshed, M., Sockett, P.N., and Artsob, H. 2008c. The rising challenge of Lyme borreliosis in Canada. Canada Communicable Disease Report, 34: 1–19. Ogden, N.H., Margos, G., Aanensen, D.M., Drebot, M.A., Feil, E.J., Hanincová, K., et al. 2011. Investigation of genotypes of Borrelia burgdorferi in Ixodes scapularis ticks collected during surveillance in Canada. Applied Environmental Microbiology, 77: 3244–3254. Ogden, N.H., St-Onge, L., Barker, I.K., Brazeau, S., Bigras-Poulin, M., Charron, D.F., et al. 2008d. Risk maps for range expansion of the Lyme disease vector, Ixodes scapularis, in Canada now and with climate change. International Journal of Health Geographics, 7: 24–39. Ogden, N.H., Trudel, L., Artsob, H., Barker, I.K., Beauchamp, G., Charron, D.F., et al. 2006c. Ixodes scapularis ticks collected by passive surveillance in Canada: analysis of geographic distribution and infection with Lyme borreliosis agent Borrelia burgdorferi. Journal of Medical Entomology, 43: 600–609. Oken, L. 1818. Sogenannte giftige Wanze in Persien. Isis, 7: 1567–1570. Oliphant, L.W., Thompson, W.J.P., Donald, T., and Rafuse, R. 1976. Present status of the prairie falcon in Saskatchewan. Canadian Field-Naturalist, 90: 365–368. Oliver, J.H. 1971. Parthenogenesis in mites and ticks (Arachnida: Acari). American Zoologist, 11: 283–299. Oliver, J.H. 1974. Symposium on reproduction of arthropods of medical and veterinary importance. IV. Reproduction in ticks (Ixodoidea). Journal of Medical Entomology, 11: 26–34. Oliver, J.H., Al-Ahmadi, Z., and Osburn, R.L. 1974. Reproduction in ticks (Acari: Ixodoidea). 3. Copulation in Dermacentor occidentalis Marx and Haemaphysalis leporispalustris (Packard) (Ixodidae). Journal of Parasitology, 60: 499–506. Oliver, J.H., Chandler, F.W., James, A.M., Huey, L.O., Vogel, G.N., and Sanders, F.H. 1996. Unusual strain of Borrelia burgdorferi isolated from Ixodes dentatus in central Georgia. Journal of Parasitology, 82: 936–940. Oliver, J.H., Owsley, M.R., Hutcheson, H.J., James, A.M., Chen, C.S., Irby, W.S., Dotson, E.M., and McLain, D.K. 1993. Conspecificity of the ticksIxodes scapularis and Ixodes dammini (Acari: Ixodidae). Journal of Medical Entomology, 30: 54–63. Oliver, J.H., Wilkinson, P.R., and Kohls, G.M. 1972. Observations on hybridization of three species of North American Dermacentor ticks. Journal of Parasitology, 58: 380–384. Oprandy, J.J., Schwan, T.G., and Main, A.J. 1988. Tick-borne Kemerovo group orbiviruses in a Newfoundland seabird colony. Canadian Journal of Microbiology, 34: 782–786. Osborn, H. 1896. Insects affecting domestic animals: an account of the species of importance in North America, with mention of related forms occurring on other animals. Bulletin of the Division of Entomology, United States Department of Agriculture, n.s., 5. 302 Pp. Osborn, H. 1899. List of insects hitherto known from the Pribilof Islands. IX. Acarina. In Fur seals and fur-seal islands of the north Pacific Ocean, part 3.Edited by D.S. Jordan, L. Stejneger, F.A. Lucas, J.F. Moser, C.H. Townsend, G.A. Clark, and J. Murray. Washington, District of Columbia, United States of America. Pp. 553–554. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 275

Oudemans, A.C. 1936. Kritisch historisch overzicht der acarologie. Derde gedeelte, 1805–1850. E.J. Brill, Leiden, The Netherlands. Packard, A.S. 1869. List of hymenopterous and lepidopterous insects collected by the Smithsonian expedition to South America under Prof. James Orton. (Appendix to the report on Articulates). Annual Report to the Trustees of the Peabody Academy of Sciences, 1: 56–69. Packard, A.S. 1872. Arachnida. Report, United States Geological Survey of the Territories, 6: 1–740. Paddock, C.D. 2005. Rickettsia parkeri as a paradigm for multiple causes of tick-borne spotted fever in the Western Hemisphere. Annals of the New York Academy of Science, 1063: 315–326. Paddock, C.D., Sumner, J.W., Comer, J.A., Zaki, S.R., Goldsmith, C.S., Goddard, J., et al. 2004. Rickettsia parkeri: a newly recognized cause of spotted fever rickettsiosis in the United States. Clinical Infectious Diseases, 38: 805–811. Paddock, C.D. and Yabsley, M.J. 2007. Ecological havoc, the rise of white-tailed deer, and the emergence of Amblyomma americanum-associated zoonoses in the United States. Current Topics in Microbiology and Evolutionary Immunology, 315: 289–324. Padgett K., Bonilla, D., Kjemtrup, A., Vilcins, I.M., Yoshimizu, M.H., Hui, L., et al. 2014. Large scale spatial risk and comparative prevalence of Borrelia miyamotoi and Borrelia burgdorferi sensu lato in Ixodes pacificus. Public Library of Science One. 9:e110853. Padgett, K.A. and Lane, R.S. 2001. Life cycle of Ixodes pacificus (Acari: Ixodidae): timing of developmental processes under field and laboratory conditions. Journal of Medical Entomology, 38: 684–693. Paffenbarger, R.S. 1950. Tick paralysis: implicating Amblyomma maculatum. New Orleans Medical and Surgical Journal, 103: 329–332. Pantanowitz, L., Telford, S.R., and Cannon, M.E. 2002. The impact of babesiosis on transfusion medicine. Transfusion Medical Review, 16: 131–143. Pappas, P.J. and Oliver, J.H 1971. Mating necessary for complete feeding of female Dermacentor variabilis (Acari: Ixodidae). Journal of the Georgia Entomological Society, 6: 122–124. Parker, R.R., Kohls, G.M., Cox, G.W., and Davis, G.E. 1939. Observations on an infectious agent from Amblyomma maculatum. United States Public Health Reports, 54: 1482–1484. Parker, R.R., Philip, C.B., Davis, G.E., and Cooley, R.A. 1937. Ticks in the United States in relation to disease in man. Journal of Economic Entomology, 30: 51–69. Parkins, M. 2008. Anaplasmosis, human granulocytic – Canada: First report, (Alberta) [online]. ProMed Posting Archives No. 20080731.2352. Available from http://promedmail.org/post/20080731.2352 {accessed 29 November 2015]. Parkins, M.D., Church, D.L., Jiang, X.Y., and Gregson, D.B. 2009. Human granulocytic anaplasmosis: first reported case in Canada. Canadian Journal of Infectious Diseases and Medical Microbiology, 20: 100–102. Parola, P., Davoust, B., and Raoult, D. 2005a. Tick- and flea-borne rickettsial emerging zoonoses. Veterinary Research, 36: 469–492. Parola, P., Paddock, C.D., and Raoult, D. 2005b. Tick-borne rickettsioses around the world: emerging diseases challenging old concepts. Clinical Microbiology Reviews, 18: 719–756. Patrick, C.D. and Hair, J.A. 1975. Ecological observations on Dermacentor albipictus (Packard) in eastern Oklahoma (Acarina: Ixodidae). Journal of Medical Entomology, 12: 393–394. Patterson, E.I., Dergousoff, S.J., and Chilton, N.B. 2009. Genetic variation in the 16S mitochondrial DNA gene of two Canadian populations of Dermacentor andersoni (Acari: Ixodidae). Journal of Medical Entomology, 46: 475–481. Pavesi, P. 1883. Considerazioni sull’aracnofauna dell’Abissinia. Nota preventive. Reconditi dell’Instituto Lombardo di Scienze e Lettere, Milan, s. 2, 16: 496–501. Peavey, C.A. and Lane, R.S. 1996. Field and laboratory studies on the timing of oviposition and hatching of the western black-legged tick, Ixodes pacificus (Acari: Ixodidae). Experimental and Applied Acarology, 20: 695–711. Peavey, C.A., Lane, R.S., and Damrow, T. 2000. Vector competence of Ixodes angustus (Acari: Ixodidae) for Borrelia burgdorferi sensu stricto. Experimental and Applied Acarology, 23: 77–84. Perry, K.L. 2014. Molecular phylogenetic relationships of North American Dermacentor ticks using mitochondrial gene sequences. M.Sc. Thesis, Paper 1089. Georgia Southern University, Statesboro, Georgia, United States of America. Electronic Theses & Dissertations. Available from http:// digitalcommons.georgiasouthern.edu/etd/1089/ [accessed 26 November 2015]. Philip, C.B., Bell., J.F., and Larson, C.L. 1955. Evidence of infectious diseases and parasites in a peak population of black-tailed jack rabbits in Nevada. Journal of Wildlife Management, 19: 225–233. Philip, C.B. and Davis, G.E. 1940. Relapsing fever: data implicating Ornithodoros hermsi as a vector in northern Idaho. United States Public Health Reports, 55: 504–507. 276 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Pickard-Cambridge, O. 1876. On a new order and some new genera of Arachnida from Kerguelen’s Island. Proceedings of the Zoological Society of London, 17: 258–265. Platonov A.E., Karan, L.S., Kolyasnikova, N.M., Makhneva, N.A., Toporkova, M.G., Maleev, V.V., et al. 2011. Humans infected with relapsing fever spirochete Borrelia miyamotoi, Russia. Emerging Infectious Diseases, 17: 1816–1823. Platt, K.B., Novak, M.G., and Rowley, W.A. 1992. Studies on the biology of Ixodes dammini in the upper midwest of the United States. Annals of the New York Academy of Sciences, 653: 78–87. Poitout, F.M., Shinozaki, J.K., Stockwell, P.J., Holland, C.J., and Shukla, S.K. 2005. Genetic variants of Anaplasma phagocytophilum infecting dogs in western Washington State. Journal of Clinical Microbiology, 43: 796–801. Pomerantzev, B.I. 1950. Arachnoidea, Ticks (Ixodidae). Fauna SSSR, New series, Number 41, 4: 1–224. [In Russian]. Pound, J.M. and George, J.E. 1991. Differential effects of photoperiod on development and reproduction in “montane” and “prairie” strains of the Rocky Mountain wood tick (Acari: Ixodidae). Journal of Medical Entomology, 28: 174–182. Pritt, B. S., Mead, P.S., Johnson, D.K., Neitzel, D.F., Respicio-Kingry, L.B., Davis, J.P., Schiffman, E., Sloan, L.M., Schriefer, M.E., et al. 2016. Identification of a novel pathogenicBorrelia species causing Lyme borreliosis with unusually high spirochaetaemia: a descriptive study. The Lancet Infectious Disease, DOI: 10.101/Z1473-3099. Pritt, B.S., Sloan, L.M., Johnson, D.K., Munderloh, U.G., Paskewitz, S.M., McElroy, K.M., et al. 2011. Emergence of a new pathogenic Ehrlichia species, Wisconsin and Minnesota, 2009. New England Journal of Medicine, 365: 422–429. Proudfoot, G.A., Teel, P.D., and Mohr, R.M. 2006. Ferruginous pygmy-owl (Glaucidium brasilianus) and eastern screech-owl (Megascops asio): new hosts for Philornis mimicola (Diptera: Muscidae) and Ornithodoros concanensis (Acari: Argasidae). Journal of Wildlife Diseases, 42: 873–876. Pung, O.J., Durden, L.A., Banks, C.W., and Jones, D.N. 1994. Ectoparasites of opossums and raccoons in southeastern Georgia. Journal of Medical Entomology, 31: 915–919. Pusterla, N., Chae, J.S., Kimsey, R.B., Berger Pusterla, J., DeRock, E., Dumler, J.S., and Madigan, J.E. 2002. Transmission of Anaplasma phagocytophila (human granulocytic ehrlichiosis agent) in horses using experimentally infected ticks (Ixodes scapularis). Journal of Veterinary Entomology, 49: 484–488. Rainwater, K.K., Ijdo, J., Capuano, A., Gilchrist, M.J., and Gill, J.S. 2006. Serosurveillance for Anaplasma phagocytophilum antibodies in white-tailed deer (Odocoileus virginianus) in Iowa, USA. Vector-borne and Zoonotic Diseases, 6: 275–282. Rauter, C. and Hartung, T. 2005. Prevalence of Borrelia burgdorferi sensu lato genospecies in Ixodes ricinus ticks in Europe: a metaanalysis. Applied and Environmental Microbiology, 71: 7203–7216. Reeves, W.K., Streicker, D.G., Loftis, A.D., and Dasch, G.A. 2006. Serologic survey of Eptesicus fuscus from Georgia for Rickettsia and Borrelia and laboratory transmission of a Rickettsia by bat ticks. Journal of Vector Ecology, 31: 386–389. Ribeiro, J.M.C. 1987. Role of saliva in the bloodfeeding by arthropods. Annual Review of Entomology, 32: 463–478. Rich, G.B. and Gregson, J.D. 1968. The first discovery of free-living larvae of the ear tick,Otobius megnini (Dugès), in British Columbia. Journal of the Entomological Society of British Columbia, 65: 22–23. Richards, W.R. 1964. A short method for making balsam mounts of aphids and scale insects. The Canadian Entomologist, 96: 963–966. Richter, P.J., Kimsey, R.B., Madigan, J.E., Barlough, J.E., Dumler, J.S., and Brooks, D.L. 1996. Ixodes pacificus (Acari: Ixodidae) as a vector of Ehrlichia equi (Rickettsiales: Ehrlichieae). Journal of Medical Entomology, 33: 1–5. Robbins, R.G. and Keirans, J.E. 1992. Systematics and ecology of the subgenus Ixodiopsis (Acari: Ixodidae: Ixodes). Thomas Say Foundation, volume 14. Entomological Society of America, Lanham, Maryland, United States of America. Rollend, L., Fish, D., and Childs, J.E. 2013. Transovarial transmission of Borrelia spirochetes by Ixodes scapularis: a summary of the literature and recent observations. Ticks and Tick-borne Diseases, 4: 46–51. Romero, J.R. and Simonsen, K.A. 2008. Powassan encephalitis and Colorado tick fever. In Tick-borne diseases part II: other tick-borne diseases. Edited by J. Edlow. Infectious Disease Clinics of North America, 22: 545–559. Rosenstein, M. 1976. Paralysis in chickens caused by larvae of the poultry tick, Argas persicus. Avian Diseases, 20: 407–409. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 277

Salkeld, D.J., Eisen, R.J., Antolin, M.F., Stapp, P., and Eisen, L. 2006. Host usage and seasonal activity patterns of Ixodes kingi and I. sculptus (Acari: Ixodidae) nymphs in a Colorado prairie landscape, with a summary of published North American host records for all life stages. Journal of Vector Ecology, 31: 168–180. Salkeld, D.J., Leonhard, S., Girard, Y.A., Hahn, N., Mun, J., Padgett, K.A., and Lane, R.S. 2008. Identifying the reservoir hosts of the Lyme disease spirochete Borrelia burgdorferi in California: the role of the western gray squirrel (Sciurus griseus). American Journal of Tropical Medicine and Hygiene, 79: 535–540. Salmon, D.E. and Stiles, C.W. 1901. The cattle ticks (Ixodoidea) of the United States. 17th Annual Report of the Bureau of Animal Industry. United States Department of Agriculture, Washington, District of Columbia, United States of America. Pp. 380–488, figures 42–257. Samuel, W.M. 1989. Location of moose in northwestern Canada with hair loss probably caused by the winter tick, Dermacentor albipictus (Acari: Ixodidae). Journal of Wildlife Diseases, 25: 436–439. Samuel, W.M. 2004. White as a ghost. Winter ticks and moose. Federation of Alberta Naturalists. Natural History Series, 1: 1–100. Samuel, W.M. and M.J. Barker. 1979. The winter tick, Dermacentor albipictus (Packard, 1869), on moose, Alces alces (L.), of central Alberta. Proceedings of the North American Moose Conference Workshop, 15: 303–348. Samuel, W.M., Grinnell, E.R., and Kennedy, A. 1981. Ectoparasites (Mallophaga, Anoplura, Acari) on mule deer, Odocoileus hemionus, and white-tailed deer, Odocoileus virginianus, of Alberta, Canada. Journal of Medical Entomology, 17: 15–17. Samuel, W.M. and Welch, D.A. 1991. Winter ticks on moose and other ungulates: factors influencing their population size. Alces, 27: 169–182. Samuel, W.M., Welch, D.A., and Smith, B.L. 1991. Ectoparasites of elk (Cervus elaphus nelsoni) from Wyoming. Journal of Wildlife Diseases, 27: 446–451. Savage, H.M., Godsey, Jr., M.S., Lambert, A., Panella, N.A., Burkhalter, K.L., Harmon, J.R., Lash, R.R., Ashley, D.C., and Nicholson, W.L. 2013. First detection of heartland virus (Bunyaviridae: Phlebovirus) from field collected arthropods. American Journal of Tropical Medicine and Hygiene,89: 445–452. Say, T. 1821. An account of the Arachnides of the United States. Journal of the Academy of National Sciences of Philadelphia, 2: 59–82. Schaalje, G.B. and Wilkinson, P.R. 1985. Discriminant analysis of vegetational and topographical factors associated with the focal distribution of Rocky Mountain wood ticks, Dermacentor andersoni (Acari: Ixodidae), on cattle range. Journal of Medical Entomology, 22: 315–320. Schaefer, J.J., Needham, G.R., Bremer, W.G., Rikihisa, Y., Ewing, S.A., and Stich, R.W. 2007. Tick acquisition of Ehrlichia canis from dogs treated with doxycycline hyclate. Antimicrobial Agents Chemotherapy, 51: 3394–3396. Scharf, W.C. 2004. Immature ticks on birds: temporal abundance and reinfestation. Northeastern Naturalist, 11: 143–150. Scharf, W.C., Schauer, S., Freyburger, F., Petrovec, M., Schaarschmidt-Kiener, D., Liebisch, G., et al. 2011. Distinct host species correlate with Anaplasma phagocytophilum ankA gene clusters. Journal of Clinical Microbiology, 49: 790–796. Scharf, W.C. and Walker, E.D. 2003. New host records for Ixodes muris (Acarina: Ixodidae) from Michigan birds. Great Lakes Entomologist, 35: 113–114. Schmidtman, E.T. 1989. American dog tick: vector of the Potomac fever rickettsia, Ehrlichia risticii? Parasitology Today, 5: 355–356. Scholten, T.H. 1977. Human tick infestations in Ontario: findings at the Toronto Public Health Laboratory, 1967–1977. Canadian Journal of Public Health, 68: 494–496. Scholten, T.H., Ronald, K., and McLean, D.M. 1962. Parasite fauna of the Manitoulin Island region. I. Arthropoda parasitica. Canadian Journal of Zoology, 40: 605–606. Schulze, P. 1930. Erster Beitrag zu einer Zeckenfauna Dänmarks. Sitzungsberichte und Abhandlungen der Naturforschenden Gesellschaft zu Rostock (1927–1929), 2: 120–124. Schulze, P. 1933. Neue und wenig bekannte deutsche Ixodes-Arten. Zeitschrift für Parasitenkunde, 6: 432-437. Schulze, P. 1938. Zur Zeckenfauna Burmas. Zeitshrift für Parasitenkunde, 10: 722–728. Schulze, T.L., Bowen, G.S., Lakat, M.F., Parkin, W.E., and Shisler, J.K. 1985. The role of adult Ixodes dammini (Acari: Ixodidae) in the transmission of Lyme disease in New Jersey, USA. Journal of Medical Entomology, 22: 88–93. Schulze, T.L., Bowen, G.S., Laket, M.F., Parkin, W.E., and Shisler, J.K. 1986. Seasonal abundance and hosts of Ixodes dammini (Acari: Ixodidae) and other ixodid ticks from an endemic Lyme disease focus in New Jersey, USA. Journal of Medical Entomology, 23: 105–109. 278 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Schwab, D. 1987. Partial paralysis in a yellow-rumped warbler may have been caused by a tick. Journal of Field Ornithology, 58: 291–291. Schwan, T.G. and Kelly, P.R. 1981. Ixodes signatus (Ixodoidea: Ixodidae) parasitizing pigeon guillemots on the Channel Islands, California. Journal of Medical Entomology, 18: 171–172. Schwan, T.G. and Piesman, J. 2002. Vector interactions and molecular adaptations of Lyme disease and relapsing fever spirochetes associated with transmission by ticks. Emerging Infectious Diseases, 8: 115–121. Schwan, T.G., Raffel, S.J., Schrumpf, M.E., Gill, J.S., and Piesman, J. 2009. Characterization of a novel relapsing fever spirochete in the midgut, coxal fluid, and salivary glands of the bat tickCarios kelleyi. Vector-borne and Zoonotic Diseases, 9: 643–647. Schwartz, B.S., Nadelman, R.B., Fish, D., Childs, J.E., Forseter, G., and Wormser, G.P. 1993. Entomologic and demographic correlates of anti-tick saliva antibody in a prospective study of tick bite subjects in Westchester County, New York. American Journal of Tropical Medicine and Hygiene, 48: 50–57. Scoles, G.A. 2004. Phylogenetic analysis of the Francisella-like endosymbionts of Dermacentor ticks. Journal of Medical Entomology, 41: 277–286. Scoles G.A., Papero, M., Beati, L., and Fish, D. 2001. A relapsing fever group spirochete transmitted by Ixodes scapularis ticks. Vector-borne and Zoonotic Diseases, 1: 21–34. Scott, J.D., Anderson, J.F., and Durden, D.A. 2012. Widespread dispersal of Borrelia burgdorferi-infected ticks collected from songbirds in Canada. Journal of Parasitology, 98: 49–59. Scott, J.D., Anderson, J.F., Durden, L.A., Smith, M.L., Manord, J.M., and Clark, K.L. 2016. Ticks parasitizing gallinaceous birds in Canada and first record ofBorrelia burgdorferi-infected Ixodes pacificus (Acari: Ixodidae) from California quail. Systematic & Applied Acarology, 21: 1–12. Scott, J.D. and Durden, L.A. 2015a. Songbird-transported tick Ixodes minor (Ixodida: Ixodidae) discovered in Canada. The Canadian Entomologist, 147: 46–50. Scott, J.D. and Durden, L.A. 2015b. First record of Amblyomma rotundatum tick (Acari: Ixodidae) parasitizing a bird collected in Canada. Systematic and Applied Acarology, 20: 155–161. Scott, J.D., Fernando, K., Banerjee, S.N., Durden, L.A., Byrne, S.K., Banerjee, M., et al. 2001. Birds disperse ixodid (Acari: Ixodidae) and Borrelia burgdorferi-infected ticks in Canada. Journal of Medical Entomology, 38: 493–500. Scott, T.W. and Brown, S.J. 1986. Differential attachment and blood-feeding by the tick Dermacentor andersoni (Acari: Ixodidae). Acarologia, 27: 241–245. Semtner, P.J. and Hair, J.A. 1973. Distribution, seasonal abundance, and hosts of the Gulf Coast tick in Oklahoma. Annals of the Entomological Society of America, 66: 1264–1268. Shapiro, M.R., Fritz, C.L., Tait, K., Paddock, C.D., Nicholson, W.L., Abramowicz, K.F., et al. 2010. Rickettsia 364D: a newly recognized cause of eschar-associated illness in California. Clinical Infectious Diseases, 50: 541–548. Sidwell, R.W., Lundgren, D.L., Bushman, J.B., and Thorpe, D.B. 1965. The occurrence of a possible epizootic of Q fever in fauna of the Great Salt Desert of Utah. American Journal of Tropical Medicine and Hygiene, 13: 754–762. Silva-Goytia, R. and Elizondo, A. 1952. Estudios sobre fiebre manchada en Mexico. 2. Parasitos hematofagos encontrados naturalmente infectados. Revista Asociacion Medica Mexicana, 32: 278–282. Skotarczak, B. 2003. Canine ehrlichiosis. Annals of Agricultural and Environmental Medicine, 10: 137–141. Slaff, M. and Newton, N.H. 1993. Location of tick (Acari: Ixodidae) attachment sites on humans in North Carolina. Journal of Medical Entomology, 30: 485–488. Slowik, T.J. and Lane, R.S. 2001a. Birds and their ticks in northwestern California: minimal contribution to Borrelia burgdorferi enzootiology. Journal of Parasitology, 87: 755–761. Slowik, T.J. and Lane, R.S. 2001b. Nymphs of the western black-legged tick (Ixodes pacificus) collected from tree trunks in woodland-grass habitat. Journal of Vector Ecology, 26: 165–171. Smart, D.L. and Caccamise, D.F. 1988. Population dynamics of the American dog tick (Acari: Ixodidae) in relation to small mammal hosts. Journal of Medical Entomology, 25: 515–522. Smith, C.N. 1940. The male, nymph, and larva of Ixodes dentatus Marx (Acarina: Ixodidae). Proceedings of the Entomological Society of Washington, 42: 16–20. Smith, C.N. 1944. Biology of Ixodes muris Bishopp and Smith (Ixodidae). Annals of the Entomological Society of America, 37: 221–234. Smith, C.N. 1945. Biology of Ixodes dentatus Neumann (Ixodidae). Annals of the Entomological Society of America, 38: 223–233. Smith, C.N., Cole, M.M., and Gouck, H.K. 1946. Biology and control of the American dog tick. United States Department of Agriculture Technical Bulletin, 905: 1–74. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 279

Smith, R., Woodall, J.P., Whitney, E., Deibel, R., Gross, M.A., Smith, V., and Bast, T.F. 1974. Powassan virus infection. A report of three human cases of encephalitis. American Journal of Diseases of Children, 127: 691–693. Smith, R.P., Muzaffar, S.B., Lavers, J., Lacombe, E.H., Cahill, B.K., Lubelczyk, C.B., et al. 2006. Borrelia garinii in sea bird ticks (Ixodes uriae), Atlantic coast, North America. Emerging Infectious Diseases, 12: 1909–1912. Smits, J.E.G. 1991. A brief review of infectious and parasitic diseases of wapiti, with emphasis on western Canada and northwestern United States. Canadian Veterinary Journal, 32: 471–479. Sonenshine, D.E. 1962. External anatomy of the bat tick Ornithodoros kelleyi Cooley and Kohls (Acarina: Argasidae). Journal of Parasitology, 48: 470–485. Sonenshine, D.E. 1967. Feeding time and oviposition of Dermacentor variabilis (Acarina: Ixodidae) as affected by delayed mating. Annals of the Entomological Society of America, 60: 489–490. Sonenshine, D.E. 1970. A contribution to the internal anatomy and histology of the bat tick Ornithodoros kelleyi Cooley and Kohls, 1941. 2. The reproductive, muscular, respiratory, excretory, and nervous systems. Journal of Medical Entomology, 7: 289–312. Sonenshine, D.E. 1973. Ecology of the American dog tick, Dermacentor variabilis, in a study area in Virginia. 3. Studies of dispersal of immature ticks by small-mammal hosts. Annals of the Entomological Society of America, 66: 44–46. Sonenshine, D.E. 1979a. Zoogeography of the American dog tick, Dermacentor variabilis. In Recent advances in acarology, volume 2. Edited by J.G. Rodriguez. Academic Press, New York, USA. Pp. 123–134. Sonenshine, D.E. 1979b. Ticks of Virginia (Acari: Metastigmata). The Insects of Virginia 13. Research Division Bulletin, 139: 1–44. Sonenshine, D.E. 1991. Biology of ticks. Volume 1. Oxford University Press, New York, New York, United States of America. 447 pp. Sonenshine, D.E. and Anastos, G. 1960. Observations on the life history of the bat tick Ornithodoros kelleyi (Acarina: Argasidae). Journal of Parasitology, 46: 449–454. Sonenshine, D.E. and Atwood, E.L. 1967. Dynamics of feeding of the American dog tick, Dermacentor variabilis (Acarina: Ixodidae). Annals of the Entomological Society of America, 60: 362–373. Sonenshine, D.E, Atwood, E.L., and Lamb, J.T 1966. The ecology of ticks transmitting Rocky Mountain spotted fever in a study area in Virginia. Annals of the Entomological Society of America, 59: 1234–1262. Sonenshine, D.E. and Gregson, J.D. 1970. A contribution to the internal anatomy and histology of the bat tick Ornithodoros kelleyi part 1. The alimentary system with notes on the food channel in Ornithodoros denmarki. Journal of Medical Entomology, 7: 46–64. Sonenshine, D.E, Lamb, J.T., and Anastos, G. 1965. The distribution, hosts and seasonal activity of Virginia ticks. Virginia Journal of Science, 16: 26–91. Sonenshine, D.E., Lane, R.S., and Nicholson, W.L. 2002. Ticks (Ixodida). In Medical and veterinary entomology. Edited by G.R. Mullen and L.A. Durden. Academic Press, Boston, Massachesetts, United States of America. Pp. 518–551. Sonenshine, D.E. and Levy, G.F. 1972. Ecology of the American dog tick, Dermacentor variabilis, in a study area in Virginia. 2. Distribution in relation to vegetative types. Annals of the Entomological Society of America, 65: 1175–1182. Sonenshine, D.E. and Stout, I.J. 1970. A contribution to the ecology of ticks infesting wild birds and rabbits in the Virginia-North Carolina piedmont (Acarina: Ixodidae). Journal of Medical Entomology, 7: 645–654. Sonenshine, D.E. and Tigner, J.A. 1969. Oviposition and hatching in two species of ticks in relation to moisture deficit. Annals of the Entomological Society of America,62: 628–640. Sonenshine, D.E., Yunker, C.E., Clifford, C.M., Clark, G.M., and Rudbach, J.A. 1976. Contributions to the ecology of Colorado tick fever virus. 2. Population dynamics and host utilization of immature stages of the Rocky Mountain wood tick, Dermacentor andersoni. Journal of Medical Entomology, 12: 651–656. Sorensen, T.C. and Moses, R.A. 1998. Host preferences and temporal trends of the tick Ixodes angustus in north-central Alberta. Journal of Parasitology, 84: 902–906. Spencer, G.J. 1960. Two records of Ixodes signatus Birula and Ixodes uriae White, marine bird ticks. Proceedings of the Entomological Society of British Columbia, 57: 46. Spencer, G.J. 1963. Attacks on humans by Ixodes angustus Neumann, the coast squirrel tick, and Ixodes soricis Gregson, the shrew tick. Proceedings of the Entomological Society of British Columbia, 60: 40. Sperling, J.L.H. and Sperling, F.A.H. 2009. Lyme borreliosis in Canada: biological diversity and diagnostic complexity from an entomological perspective. The Canadian Entomologist, 141: 521–549. Spicer, W.J., and Dove, W.E. 1938. The screwworm and the Gulf Coast tick in southern Texas. Journal of Economic Entomology, 31: 642–646. 280 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Spielman, A., Clifford, C.M., Piesman, J., and Corwin, M.D. 1979. Human babesiosis on Nantucket Island, U.S.A.: description of the vector, Ixodes (Ixodes) dammini, n. sp. (Acarina: Ixodidae). Journal of Medical Entomology, 15: 218–234. Spielman, A., Levine, J.F., and Wilson, M.L. 1984. Vectorial capacity of North American Ixodes ticks. Yale Journal of Biology and Medicine, 57: 507–513. Spielman, A., Wilson, M.L., Levine, J.F., and Piesman, J. 1985. Ecology of Ixodes dammini-borne human babesiosis and Lyme disease. Annual Review of Entomology, 30: 439–460. Stafford, K.C. 1992. Oviposition and larval dispersal of Ixodes dammini (Acari: Ixodidae). Journal of Medical Entomology, 29: 129–132. Stafford, K.C., Bladen, V.C., and Magnarelli, L.A. 1995. Ticks (Acari: Ixodidae) infesting wild birds (Aves) and white-footed mice in Lyme, CT. Journal of Medical Entomology, 32: 453–466. Stafford, K.C. and Magnarelli, L.A. 1993. Spatial and temporal patterns of Ixodes scapularis (Acari: Ixodidae) in southeastern Connecticut. Journal of Medical Entomology, 30: 762–771. Statistics Canada. 1969–1976. Hospital morbidity database. In Statistics Canada (Rocky Mountain spotted fever section, ICDA-8 code number 082.0 (spotted fevers). P. 51. Steere, A.C., Broderick, T.F., and Malawista, S.E. 1978. Erythema chronicum migrans and Lyme arthritis: epidemiologic evidence for a tick vector. American Journal of Epidemiology, 108: 312–321. Stiles, C.W. 1908. The common tick (Dermacentor andersoni) in the Bitter Root Valley. United States Public Health Report, 23: 949. Stiles, C.W. 1910. The taxonomic value of the microscopic structure of the stigmal plates in the tick genus Dermacentor. United States Public Health and Marine Hospital Service, Hygienic Laboratory Bulletin 62: 1–72, plates I–XLIII. Stiller, D. and Coan, M.E. 1995. Recent developments in elucidating tick vector relationships for anaplasmosis and equine piroplasmosis. Veterinary Parasitology, 57: 97–108. Stiller, D., Frerichs, W.M., Leatch, G., and Kuttler, K.L. 1980. Transmission of equine babesiosis and bovine anaplasmosis by Dermacentor albipictus (Packard) (Acari: Ixodidae). Journal of the New York Entomological Society, 88: 75–76. Stiller, D., Johnson, L.W., and Kuttler, K.L. 1987. Experimental transmission of Anaplasma ovis to sheep by Dermacentor albipictus and Dermacentor variabilis male ticks. Abstracts, Proceedings of the 8th Annual Western Conference on Food Animal Veterinary Medicine. Boise, Idaho, United States of America. P. 36. Stiller, D., Kocan, K.M., Edwards, W., Ewing, S.A., Hair, J.A., and Barron, S.J. 1989. Detection of colonies of Anaplasma marginale in salivary glands of three Dermacentor spp. infected as nymphs or adults. American Journal of Veterinary Research, 50: 1381–1385. Stiller, D., Leatch, G., and Kuttler, K.L. 1981. Dermacentor albipictus (Packard): an experimental vector of bovine anaplasmosis. Annual Meeting of the United States Animal Health Association, 85: 65–75. Stoll, O. 1886–1893. Arachnida Acaridea. Biologia Centrali-Americana, Zoology. Taylor and Francis, London, United Kingdom. Suksawat, J., Hegarty, B.C., and Breitschwerdt, E.B. 2000. Seroprevalence of Ehrlichia canis, Ehrlichia equi, and Ehrlichia risticii in sick dogs from North Carolina and Virginia. Journal of Veterinary Internal Medicine, 14: 50–55. Sumner, J.W., Durden, L.A., Goddard, J., Stromdahl, E.Y., Clark, K.L., Reeves, W.K., and Paddock, C.D. 2007. Gulf Coast ticks (Amblyomma maculatum) and Rickettsia parkeri, United States. Emerging Infectious Diseases, 13: 751–753. Swanson, K.I. and Norris, D.E. 2007. Co-circulating microorganisms in questing Ixodes scapularis nymphs in Maryland. Journal of Vector Ecology, 32: 243–251. Swanson, S.J., Neitzel, D., Reed, K.D., and Belongia, E.A. 2006. Coinfections acquired from Ixodes ticks. Clinical Microbiology Review, 19: 708–727. Swift, T.R. and Ignacio, O.J. 1975. Tick paralysis: electrophysiologic studies. Neurology, 25: 1130–1133. Tate, C.M., Mead, D.G., Luttrell, M.P., Howerth, E.W., Dugan, V.G., Munderloh, U.G., and Davidson, W.R. 2005. Experimental infection of white-tailed deer with Anaplasma phagocytophilum, etiologic agent of human granulocytic anaplasmosis. Journal of Clinical Microbiology, 43: 3595–3601. Tavakoli, N.P., Wang, H., Dupuis, M., Hull, R., Ebel, G.D., Gilmore, E.J., and Faust, P.L. 2009. Fatal case of deer tick virus encephalitis. New England Journal of Medicine, 360: 2099–2107. Teel, P.D., Fleetwood, S.C., Hopkins, S.W., and Cruz, D. 1988. Ectoparasites of eastern and western meadowlarks from the Rio Grande plains of South Texas. Journal of Medical Entomology, 25: 32–38. Teel, P.D., Hopkins, S.W., Donahue, W.A., and Strey, O.F. 1998. Population dynamics of immature Amblyomma maculatum (Acari: Ixodidae) and other ectoparasites on meadowlarks and northern bobwhite quail resident to the coastal prairie of Texas. Journal of Medical Entomology, 35: 483–488. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 281

Telford, S.R., Goethert, H.K., and J.A. Cunningham. 2011. Prevalence of Ehrlichia muris in Wisconsin deer ticks collected during the mid 1990s. The Open Microbiology Journal, 5: 18–20. Telford, S.R. and Spielman, A. 1989. Competence of a rabbit-feeding Ixodes (Acari: Ixodidae) as a vector of the Lyme disease spirochete. Journal of Medical Entomology, 26: 118–121. Teng, J., Lindsay, L.R., Bartlett, K., Klinkenberg, B., Dibernardo, A., Wood, H., and Morshed, M.G. 2011. Prevalence of tick-borne pathogens in the South Okanagan, British Columbia: active surveillance in ticks (Dermacentor andersoni) and deer mice (Peromyscus maniculatus). British Columbia Medical Journal, 53: 122–127. Thomas, L.A., Kennedy, R.C., and Eklund, C.M. 1960. Isolation of a virus closely related to Powassan virus from Dermacentor andersoni collected along North Cache la Poudre River, Colorado. Proceedings of the Society for Experimental Biology and Medicine, 104: 355–359. Thomas, P.A. 1968. Geographic variation of the rabbit tick, Haemaphysalis leporispalustris, in North America. University of Kansas Science Bulletin, 47: 787–828. Thompson, C., Spielman, A., and Krause, P.J. 2001. Coinfecting deer-associated zoonoses: Lyme disease, babesiosis, and ehrlichiosis. Clinical Infectious Diseases, 33: 676–685. Thorpe, B.D., Sidwell, R.W., Johnson, D.E., Smart, K.L., and Parker, D.D. 1965. Tularemia in wildlife and livestock of the Great Salt Lake region, 1951–1964. American Journal of Tropical Medicine and Hygiene, 14: 622–637. Townsend, C.H.T. 1893. Ticks in the ears of horses. Journal of New York Entomological Society, 1: 49–52. Trager, W. 1939a. Acquired immunity to ticks. Journal of Parasitology, 25: 57–81. Trager, W. 1939b. Observations on acquired immunity to the tick Dermacentor variabilis Say. Journal of Parasitology, 25: 137–141. Travis, B.V. 1968. Glyptal – a useful slide ringing compound. Journal of Medical Entomology, 5: 24. Troughton, D.R. and Levin, M.L. 2007. Life cycles of seven ixodid tick species (Acari: Ixodidae) under standardized laboratory conditions. Journal of Medical Entomology, 44: 732–740. Vannier, E, Gewurz, B.E., and Krause, P.J. 2008. Human babesiosis. Infectious Disease Clinics of North America, 22: 469–488. Van Wye, J.E., Hsu, Y.P., Terr, A.I., Moss, R.B., and Lane, R.S. 1991. Anaphylaxis from a tick bite. New England Journal of Medicine, 324: 777–778. Varde, S.J.B., Beckley, J., and I. Schwartz, I. 1998. Prevalence of tick-borne pathogens in Ixodes scapularis in a rural New Jersey county. Emerging Infectious Diseases, 4: 97–99. Varela, A.S., Luttrell, M.P., Howerth, E.W., Moore, V.A., Davidson, W.R., Stallknecht, D.E., and Little, S.E. 2004. First culture isolation of Borrelia lonestari, putative agent of southern tick-associated rash illness. Journal of Clinical Microbiology, 42: 1163–1169. Vasconcelos, P. F. and Calisher, C.H. 2016. Emergence of human arboviral diseases in the Americas, 2000-2016. Vector-Borne Zoonotic Diseases, DOI:10.1089/vbz.2016.1952. Vetter, R.S. 2013. Arachnophobic entomologists: when two more legs makes a big difference. American Entomologist, 59: 168–175. Vijoen, J.G., Bezuidenhout, J.D., Oberem, P.T., Vermeulen, N.M.J., Visser, L., Gothe, R., and Neitz, A.W.H. 1986. Isolation of a neurotoxin from the salivary glands of female Rhipicephalus evertsi evertsi. Journal of Parasitology, 72: 865–874. Vijoen, J.G., Wyngaardt, S.V., Gothe, R., Visser, L., Bezuidenhout, J.D., and Neitz, A.W.H. 1990. The detection and isolation of a paralysis toxin present in Argas (Persicargas) walkerae. Onderstepoort Journal of Veterinary Research, 57: 163–168. Walker, D.H., Barbour, A.G., Oliver, J.H., Lane, R.S., Dumler, J.S., Dennis, D.T., et al. 1996. Emerging bacterial zoonotic and vector-borne diseases. Ecological and epidemiological factors. Journal of the American Medical Association, 275: 463–469. Ward, P.A. and Sonenshine, D.E. 1972. Tick-bite injury: mediation by a complement-derived chemotactic peptide. Journal of Immunology, 109: 451–456. Watson, T.G. and Anderson, R.C. 1976. Ixodes scapularis Say on white-tailed deer (Odocoileus virginianus) from Long Point, Ontario. Journal of Wildlife Diseases, 12: 66–71. Waxman, L, Smith, D.E., Arcuri, K.E., and Vlasuk, G.P. 1990. Tick anticoagulant peptide (TAP) is a novel inhibitor of blood coagulation factor Xa. Science, 243: 593. Webb, J.P., Bennett, S.G., and Challet, G.L. 1990. The larval ticks of the genus Ixodes Latreille (Acari: Ixodidae) of California. Bulletin of the Society for Vector Ecology, 15: 73–124. Webb, J.P., George, J.E., and Cook, B. 1977. Sound as a host-detection cue for the soft tick Ornithodoros concanensis. Nature, 265: 443–444. Webster, W.A. 1966. Ixodes texanus in eastern Canada. Bulletin of the Wildlife Disease Association, 2: 78–79. 282 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Welch, D.A., Samuel, W.M., and Wilke, C.J. 1990. Dermacentor albipictus (Acari, Ixodidae) on captive reindeer and free-ranging woodland caribou. Journal of Wildlife Diseases, 26: 410–411. Welch, D.A., Samuel, W.M., and Wilke, C.J. 1991. Suitability of moose, elk, mule deer, and white-tailed deer as hosts for winter ticks (Dermacentor albipictus). Canadian Journal of Zoology, 69: 2300–2305. Werden, L., Barker, I.K., Bowman, J., Gonzales, E.K., Leighton, P.A., Lindsay, L.R., and Jardine, C.M. 2014. Geography, deer, and host biodiversity shape the pattern of Lyme disease emergence in the Thousand Islands archipelago of Ontario, Canada. Public Library of Science One 9:e85640. doi: 10.1371/journal. pone.0085640. Westrom, D.R., Lane, R.S., and Anderson, J.R. 1985. Ixodes pacificus (Acari: Ixodidae): population dynamics and distribution on Columbian black-tailed deer (Odocoileus hemionus columbianus). Journal of Medical Entomology, 22: 507–511. Wheeler, C.M. 1935. A new species of tick which is a vector of relapsing fever in California. American Journal of Tropical Medicine, 15: 435–438. Wheeler, C.M., Herms, W.B., and Meyer, K.F. 1935. A new tick vector of relapsing fever in California. Proceedings of the Society for Experimental Biology and Medicine, 32: 1290–1292. Wherry, W.B. and Wellman, F.C. 1909. Ticks on the California ground squirrel. Entomological News, 20: 376. Whitaker J.O. and French, T.W. 1982. Ectoparasites and other associates of some insectivores and rodents from New Brunswick. Canadian Journal of Zoology, 60: 2787–2797. White, A. 1852. Insects and Aptera. In F.C. Sutherland, Journal of a voyage in Baffin’s Bay and Barrow Straite, in the years 1850–1851 performed by H.M. Ships “Lady Franklin” and “Sophia,” under the command of Mr. William Penny, in search of the missing crews of H.M. Ships Erebus and Terror. Volume 2: appendix. Edited by P.C. Sutherland. Longman, Brown, Green, and Longmans, London, United Kingdom. Pp. ccviii–ccxi. Whitman, T.J., Richards, A.L., Paddock, C.D., Tamminga, C.L., Sniezek, P.J., Jiang, J., Byers, D.K., and Sanders, J.W. 2007. Rickettsia parkeri infection after tick bite, Virginia. Emerging Infectious Diseases, 13: 334–336. Wikswo, M.E., Hu, R., Dasch, G.A., Krueger, L., Arugay, A., Jones, K., et al. 2008. Detection and identification of spotted fever group rickettsiae in Dermacentor species from southern California. Journal of Medical Entomology, 45: 509–516. Wilkinson, P.R. 1965. A first record of paralysis of a deer byDermacentor andersoni (Stiles) and notes on the “host-potential” of deer in British Columbia. Proceedings of the Entomological Society of British Columbia, 62: 28–30. Wilkinson, P.R. 1967. The distribution of Dermacentor ticks in Canada in relation to bioclimatic zones. Canadian Journal of Zoology, 45: 517–537. Wilkinson, P.R. 1968. Phenology, behavior, and host-relations of Dermacentor andersoni Stiles in outdoor “rodentaria”, and in nature. Canadian Journal of Zoology, 46: 677–689. Wilkinson, P.R. 1970. Dermacentor ticks on wildlife and new records of paralysis. Journal of the Entomological Society of British Columbia, 67: 24–28. Wilkinson, P.R. 1971. Termination of diapause in laboratory-reared Dermacentor andersoni adults. Proceedings of the 3rd International Congress of Acarology, Prague, Czechoslovakia, 1971. Pp. 803–806. Wilkinson, P.R. 1972. Sites of attachment of “prairie” and “montane” Dermacentor andersoni (Acarina: Ixodidae) on cattle. Journal of Medical Entomology, 9: 133–137. Wilkinson, P.R. 1979. Early achievements, recent advances, and future prospects in the ecology of the Rocky Mountain wood tick. In Recent advances in acarology, volume 2. Edited by J.G. Rodriguez. Academic Press, New York, New York, United States of America. Pp. 105–112. Wilkinson, P.R. 1982. Tick paralysis. In CRC Press handbook series in zoonoses. Edited by J. H. Steele, G. Hillyer, and C. E. Hopla. CRC Press, Boca Raton, Florida, United States of America. Pp. 275–282. Wilkinson, P.R. 1984. Hosts and distribution of Rocky Mountain wood ticks (Dermacentor andersoni) at a tick focus in British Columbia rangeland. Journal of the Entomological Society of British Columbia, 81: 57–71. Wilkinson, P.R. 1985. Differences in the paralyzing ability and sites of attachment to cattle of Rocky Mountain wood ticks (Acari: Ixodidae) from three regions of western Canada. Journal of Medical Entomology, 22: 28–31. Wilkinson, P.R., Abbott, H.R., and Willman, J.N. 1982. Location of larvae of the winter tick, Dermacentor albipictus (Pack.), in Elk Island National Park, Alberta. Canadian Field-Naturalist, 96: 211–213. Wilkinson, P.R., Fyfe, R., and Martin, J.E.H. 1980. Further records of Ornithodoros ticks on prairie falcons and in bat-inhabited buildings in Canada. The Canadian Field-Naturalist, 94: 191–193. Wilkinson, P.R. and Gregson, J.D. 1985. Comparisons of sampling methods for recording the numbers of Rocky Mountain wood ticks (Dermacentor andersoni) on cattle and range vegetation during control experiments. Acarologia, 26: 131–139. A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 283

Wilkinson, P.R. and Lawson, J.E. 1965. Differences of sites of attachment of Dermacentor andersoni Stiles to cattle in southeastern Alberta and in south central British Columbia, in relation to possible existence of genetically different strains of ticks. Canadian Journal of Zoology, 43: 408–411. Williams, R.B. 1947. Infestation of raptorials by Ornithodoros aquilae. The Auk, 64: 185–188, Plate 7. Wilson, M.L., Litwin, T.S., Gavin, T.A., Capkanis, M.C., Maclean, D.C., and Spielman, A. 1990. Host- dependent differences in feeding and reproduction of Ixodes dammini (Acari: Ixodidae). Journal of Medical Entomology, 27: 945–954. Wilson, N. 1967. Ectoparasites of Canadian birds and mammals. Proceedings of the Entomological Society of Washington, 69: 349–353. Wolver, S.E., Sun, D.R., Commins, S.P., and Schwartz, L.B. 2013. A peculiar cause of anaphylaxis: no more steak? The journey to discovery of a newly recognized allergy to galactose-alpha-1,3-galactose found in mammalian meat. Journal of General Internal Medicine, 28: 322–325. Wormser, G.P., Masters, E., Liveris, D., Nowakowski, J., Nadelman, R.B., Holmgren, D., et al. 2005. Microbiologic evaluation of patients from Missouri with erythema migrans. Clinical Infectious Diseases, 40: 423–428. Wright, B. 1979. Mites, ticks, fleas and lice in the Nova Scotia Museum and Acadia University Museum collections. Proceedings of the Nova Scotia Institute of Science, 29: 185–196. Wright, J.E. 1971. Relationship of seasonal activity of Dermacentor albipictus (Acarina: Ixodidae) to photoperiod: a laboratory study. Annals of the Entomological Society of America, 64: 1456–1458. Wright, S.A., Lemenager, D.A., Tucker, J.R., Armijos, M.V., and Yamamoto, S.A. 2006. An avian contribution to the presence of Ixodes pacificus (Acari: Ixodidae) and Borrelia burgdorferi on the Sutter Buttes of California. Journal of Medical Entomology, 43: 368–374. Wright, S.W. and Trott, A.T. 1988. North American tick-borne diseases. Annals of Emerging Medicine, 17: 967–972. Wu, K.W. 1986. Review of the polyethylene bottle applicator technique for sealing microslide preparations of mites. International Journal of Acarology, 12: 87–89. Yunik, M.E.M., Galloway, T.D., and Lindsay, L.R. 2015a. Assessment of prevalence and distribution of spotted fever group rickettsiae in Manitoba, Canada, in the American dog tick, Dermacentor variabilis (Acari: Ixodidae). Vector-borne and Zoonotic Diseases, 15: 103–108. Yunik, M.E.M., Galloway, T.D., and Lindsay, L.R. 2015b. Ability of unfed Dermacentor variabilis (Acari: Ixodidae) to survive a second winter as adults in Manitoba, Canada, near the northern limit of their range. Journal of Medical Entomology, 52: 138–142. Yunker, C.E. 1975. Tick-borne viruses associated with seabirds in North America and related islands. Medical Biology 53: 302–311. Yunker, C.E., Clifford, C.M., Thomas, L.A., Keirans, J.E., Casals, J., George, J.E., and Parker, J.C. 1977. Sunday Canyon virus, a new ungrouped agent from the tick Argas (A.) cooleyi in Texas. Acta Virologica, 21: 36–44 + Plate II. Yunker, C.E., Keirans, J.E., Clifford, C.M., and Easton, E.R. 1986. Dermacentor ticks (Acari: Ixodoidea: Ixodidae) of the New World: a scanning electron microscope atlas. Proceedings of the Entomological Society of Washington, 88: 609–627. Yuval, B. and Spielman, A. 1990a. Sperm precedence in the deer tick Ixodes dammini. Physiological Entomology, 15: 123–128. Yuval, B. and Spielman, A. 1990b. Duration and regulation of the developmental cycle of Ixodes dammini (Acari: Ixodidae). Journal of Medical Entomology, 27: 196–201. Zarnke, R.L., Samuel, W.M., Franzmann, A.W., and Barrett, R. 1990. Factors influencing the potential establishment of the winter tick (Dermacentor albipictus) in Alaska. Journal of Wildlife Diseases, 26: 412–415. Zarnke, R.L. and Yuill, T.M. 1981. Powassan virus infection in snowshoe hares (Lepus americanus). Journal of Wildlife Diseases, 17: 303–310. Zaugg, J.L. 1990. Seasonality of natural transmission of bovine anaplasmosis under desert mountain range conditions. Journal of the American Veterinary Medical Association, 196: 1106–1109. Zeidner, N.S., Burkot, T.R., Massung, T., Nicholson, W.L., Dolan, M.C., Rutherford, J.S., et al. 2000. Transmission of the agent of human granulocytic ehrlichiosis by Ixodes spinipalpis ticks: evidence on an enzootic cycle of dual infection with Borrelia burgdorferi in northern Colorado. Journal of Infectious Diseases, 182: 616–619. Zimmerman, R H., McWherter, G.R., and Bloemer, S.R. 1987. Role of small mammals in population dynamics and dissemination of Amblyomma americanum and Dermacentor variabilis (Acari: Ixodidae) at Land Between the Lakes, Tennessee. Journal of Medical Entomology, 24: 370–375. 284 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Appendix 1. Host-tick index

Class Reptilia

Order Lacertilia, lizards Family Anguidae Northern alligator lizard, Elgaria coerulea principis Baird and Girard Ixodes pacificus Cooley and Kohls

Class Mammalia Order Insectivora, insectivores Family Talpidae Scheffer’s mole, Scapanus orarius schefferi Jackson Ixodes pacificus Cooley and Kohls Ixodes angustus Neumann Hairy-tailed mole, Parascalops breweri (Bachman) Ixodes angustus Neumann Star-nosed mole, Condylura cristata (Linnaeus) Ixodes angustus Neumann

Family Soricidae Vagrant shrew, Sorex vagrans Baird Ixodes soricis Gregson Masked shrew, Sorex cinereus Kerr Dermacentor variabilis (Say) Ixodes soricis Gregson Ixodes angustus Neumann Montane shrew, Sorex monticolus Merriam Ixodes soricis Gregson Smoky shrew, Sorex fumeus Miller Ixodes angustus Neumann Gaspé shrew, Sorex gaspensis Batchelder Ixodes angustus Neumann Water shrew, Sorex palustris Richardson Dermacentor variabilis (Say) Ixodes angustus Neumann Pygmy shrew, Sorex hoyi Baird Ixodes angustus Neumann Shrews, Sorex species Ixodes pacificus Cooley and Kohls Ixodes spinipalpis Hadwen and Nuttall Ixodes angustus Neumann Ixodes soricis Gregson Short-tailed shrew, Blarina brevicauda (Say) Dermacentor variabilis (Say) Ixodes angustus Neumann A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 285

Order Chiroptera, bats Family Vespertilionidae Bats (genus unknown) (or mice) Ornithodoros hermsi Wheeler, Herms, and Meyer Big brown bat, Eptesicus fuscus (Palisot de Beauvois) Carios kelleyi (Cooley and Kohls) Little brown bat, Myotis lucifugus (LeConte) Carios kelleyi (Cooley and Kohls) Long-eared bat, Myotis evotis (Allen) Carios kelleyi (Cooley and Kohls) Long-legged bat, Myotis volans Allen Carios kelleyi (Cooley and Kohls) Small-footed bat, Myotis ciliolabrum Merriam Carios kelleyi (Cooley and Kohls)

Order Primates, primates Family Hominidae Human, Homo sapiens Linnaeus Ornithodoros hermsi Wheeler, Herms, and Meyer Dermacentor andersoni Stiles Dermacentor albipictus (Packard) Dermacentor variabilis (Say) Rhipicephalus sanguineus (Latreille) Haemaphysalis leporispalustris (Packard) Haemaphysalis chordeilis (Packard) Ixodes muris Bishopp and Smith Ixodes pacificus Cooley and Kohls Ixodes scapularis Say Ixodes spinipalpis Hadwen and Nuttall Ixodes angustus Neumann Ixodes cookei Packard Ixodes marxi Banks Ixodes muris Bishopp and Smith Amblyomma americanum (Linnaeus) Amblyomma maculatum Koch

Order Carnivora, carnivores Family Procyonidae Pacific raccoon,Procyon lotor pacificus Merriam Ixodes texanus Banks Eastern raccoon, Procyon lotor lotor Linnaeus Dermacentor variabilis (Say) Ixodes texanus Banks Ixodes cookei Packard

Family Ursidae Black bear, Ursus americanus (Pallas) Dermacentor variabilis (Say) Bear, Ursus species Dermacentor andersoni Stiles Dermacentor albipictus (Packard) 286 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Family Canidae Coyote, Canis latrans Say Dermacentor andersoni Stiles Dermacentor albipictus (Packard) Ixodes rugosus Bishopp Wolf, Canis lupus Linnaeus Dermacentor albipictus (Packard) Domestic dog, Canis lupus familiaris Linnaeus Dermacentor andersoni Stiles Dermacentor variabilis (Say) Rhipicephalus sanguineus (Latreille) Ixodes pacificus Cooley and Kohls Ixodes muris Bishopp and Smith Ixodes scapularis Say Ixodes angustus Neumann Ixodes texanus Banks Ixodes cookei Packard Ixodes kingi Bishopp Ixodes marxi Banks Ixodes gregsoni Lindquist, Wu, and Redner Amblyomma americanum (Linnaeus) Amblyomma maculatum Koch

Family Felidae Domestic cat, Felis silvestris catus Linnaeus Otobius megnini (Dugès) Otobius lagophilus Cooley and Kohls Dermacentor andersoni Stiles Dermacentor variabilis (Say) Haemaphysalis leporispalustris (Packard) Ixodes muris Bishopp and Smith Ixodes pacificus Cooley and Kohls Ixodes scapularis Say Ixodes angustus Neumann Ixodes marxi Banks Ixodes cookei Packard Ixodes kingi Bishopp Amblyomma maculatum Koch

Family Mustelidae Pine marten, Martes americana americana (Turton) Ixodes cookei Packard Ixodes gregsoni Lindquist, Wu, and Redner Fisher, Martes pennanti pennanti (Erxleben) Ixodes cookei Packard Short-tailed weasel, Mustela erminea Linnaeus Dermacentor variabilis (Say) Prairie long-tailed weasel, Mustela frenata longicauda Bonaparte Dermacentor variabilis (Say) Ixodes sculptus Neumann A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 287

Weasels, Mustela species Ixodes texanus Banks Ixodes kingi Bishopp Ixodes sculptus Neumann Ixodes gregsoni Lindquist, Wu, and Redner Mink, Neovison vison (Schreber) Ixodes pacificus Cooley and Kohls Ixodes texanus Banks Ixodes cookei Packard Ixodes gregsoni Lindquist, Wu, and Redner Puget Sound spotted skunk, Spilogale phenax olympica (Elliot) (= Western Spotted Skunk, Spilogale gracilis latifrons (Merriam)) Ixodes rugosus Bishopp Eastern striped skunk, Mephitis mephitis (Schreber) Dermacentor variabilis (Say) Ixodes cookei Packard Badger, Taxidea taxus (Schreber) Dermacentor andersoni Stiles Dermacentor variabilis (Say) Ixodes kingi Bishopp Ixodes sculptus Neumann

Order Lagomorpha Suborder Duplidentata Family Ochotonidae Pika, Ochotona princeps (Richardson) Dermacentor andersoni Stiles Ixodes spinipalpis Hadwen and Nuttall Ixodes angustus Neumann Ixodes ochotonae Gregson

Family Leporidae White-tailed jack rabbit, Lepus townsendii Bachman Otobius lagophilus Cooley and Kohls Dermacentor andersoni Stiles Haemaphysalis leporispalustris (Packard) Ixodes sculptus Neumann Snowshoe hare, Lepus americanus Erxleben Haemaphysalis leporispalustris (Packard) Dermacentor variabilis (Say) Ixodes marxi Banks Rabbits, Lepus species Ixodes spinipalpis Hadwen and Nuttall Ixodes angustus Neumann Eastern cottontail, Sylvilagus floridanus (Allen) Haemaphysalis leporispalustris Blackhills cottontail, Sylvilagus nuttallii grangeri (Allen) Haemaphysalis leporispalustris (Packard) Rabbits (genus unknown) Haemaphysalis leporispalustris (Packard) Ixodes sculptus Neumann 288 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Order Rodentia Family Sciuridae Yellow-bellied marmot, Marmota flaviventris (Audubon and Bachman) Dermacentor andersoni Stiles Haemaphysalis leporispalustris (Packard) Haemaphysalis chordeilis (Packard) Ixodes texanus Banks Ixodes marmotae Cooley and Kohls Woodchuck, Marmota monax (Linnaeus) Dermacentor variabilis (Say) Ixodes cookei Packard Richardson’s ground squirrel, Urocitellus richardsonii (Sabine) Dermacentor andersoni Stiles Haemaphysalis leporispalustris (Packard) Ixodes kingi Bishopp Ixodes sculptus Neumann Columbian ground squirrel, Urocitellus columbianus (Ord) Dermacentor andersoni Stiles Ixodes marmotae Cooley and Kohls Ixodes sculptus Neumann Franklin’s ground squirrel, Poliocitellus franklinii (Sabine) Dermacentor andersoni Stiles Dermacentor variabilis (Say) Ixodes sculptus Neumann Golden-mantled ground squirrel, Callospermophilus lateralis (Say) Dermacentor andersoni Stiles Thirteen-lined ground squirrel, Ictidomys tridecemlineatus (Mitchill) Dermacentor andersoni Stiles Dermacentor variabilis (Say) Ixodes sculptus Neumann Black-tailed prairie-dog, Cynomys ludovicianus ludovicianus (Ord) Dermacentor andersoni Stiles Eastern chipmunk, Tamias striatus (Linnaeus) Dermacentor variabilis (Say) Northwestern chipmunk, Tamias striatus lysteri (Richardson) Ixodes angustus Neumann Lake Superior chipmunk, Eutamias minimus neglectus (Allen) Ixodes angustus Neumann Chipmunks, Eutamias species Dermacentor andersoni Stiles Haemaphysalis leporispalustris (Packard) Ixodes pacificus Cooley and Kohls Ixodes spinipalpis Hadwen and Nuttall Ixodes angustus Neumann Southern red squirrel, Tamiasciurus hudsonicus loquax (Bangs) Haemaphysalis leporispalustris (Packard) Ixodes angustus Neumann Ixodes marxi Banks A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 289

Streator’s red squirrel, Tamiasciurus hudsonicus streatori (Allen) Haemaphysalis leporispalustris (Packard) Haemaphysalis chordeilis (Packard) Ixodes hearlei Gregson Vancouver red squirrel, Tamiasciurus hudsonicus vancouverensis (Allen) Ixodes angustus Neumann Northwestern red-bellied squirrel, Tamiasciurus douglasii mollipilosus (Audubon and Bachman) Ixodes angustus Neumann Tree squirrels, Tamiasciurus species Dermacentor andersoni Stiles Haemaphysalis leporispalustris (Packard) Haemaphysalis chordeilis (Packard) Ixodes pacificus Cooley and Kohls Ixodes angustus Neumann Ixodes texanus Banks Grey squirrel, Sciurus carolinensis Gmelin Dermacentor variabilis (Say) Ixodes marxi Banks Northern flying squirrel, Glaucomys sabrinus (Shaw) Dermacentor variabili (Say) Ixodes angustus Neumann Ixodes marxi Banks

Family Castoridae North American beaver, Castor canadensis Kuhl Ixodes banksi Bishopp

Family Geomyidae Northern pocket gopher, Thomomys talpoides Richardson Dermacentor andersoni Stiles Dermacentor variabilis (Say) Ixodes kingi Bishopp Ixodes sculptus Neumann Ixodes scapularis Say

Family Cricetidae Dusky harvest mouse, Reithrodontomys megalotis nigrescens Howell Ixodes spinipalpis Hadwen and Nuttall Deer mouse, Peromyscus maniculatus (Wagner) Dermacentor andersoni Stiles Dermacentor albipictus (Packard) Dermacentor variabilis (Say) Ixodes scapularis Say Ixodes pacificus Cooley and Kohls Ixodes spinipalpis Hadwen and Nuttall Ixodes angustus Neumann Ixodes ochotonae Gregson 290 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

White-footed mouse, Peromyscus leucopus (Rafinesque) Ixodes scapularis Say Ixodes angustus Neumann Western bushy-tailed woodrat, Neotoma cinerea (Ord) Dermacentor andersoni Stiles Ixodes spinipalpis Hadwen and Nuttall Ixodes angustus Neumann Ixodes ochotonae Gregson Ixodes marmotae Cooley and Kohls Southern bog lemming, Synaptomys cooperi (Baird) Dermacentor variabilis (Say) Ixodes angustus Neumann Northern bog lemming, Synaptomys borealis (Richardson) Ixodes angustus Neumann Red-backed vole, Myodes gapperi (Vigors) Dermacentor variabilis (Say) Haemaphysalis leporispalustris (Packard) Ixodes scapularis Say Ixodes angustus Neumann Red-backed voles, Myodes species Ixodes angustus Neumann Drummond’s meadow mouse, Microtus pennsylvanicus drummondi (Audubon and Bachman) Ixodes angustus Neumann Meadow vole, Microtus pennsylvanicus (Ord) Dermacentor andersoni Stiles Dermacentor variabilis (Say) Haemaphysalis leporispalustris (Packard) Long-tailed vole, Microtus longicaudus (Merriam) Ixodes angustus Neumann Rock vole, Microtus chrotorrhinus (Miller) Ixodes angustus Neumann Meadow voles, Microtus species Dermacentor andersoni Stiles Dermacentor variabilis (Say) Ixodes angustus Neumann Ixodes ochotonae Gregson Ixodes auritulus Neumann Pallid pigmy vole, Lemmiscus curtatus pallidus (Merriam) Dermacentor andersoni Stiles Ixodes kingi Bishopp Heather vole, Phenacomys intermedius (Merriam) Ixodes angustus Neumann Mouse (genus unknown) (or bat) Ornithodoros hermsi Wheeler, Herms, and Meyer Rat, Rattus species Ixodes spinipalpis Hadwen and Nuttall Ixodes angustus Neumann A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 291

Family Dipodidae Western jumping mouse, Zapus princeps minor Preble Dermacentor andersoni Stiles Dermacentor variabilis (Say) Meadow jumping mouse, Zapus hudsonius (Zimmermann) Dermacentor variabilis (Say) Ixodes angustus Neumann Jumping mouse, Zapus species Ixodes angustus Neumann Eastern woodland jumping mouse, Napaeozapus insignis (Miller) Ixodes muris Bishopp and Smith

Family Erethizontidae Porcupine, Erethizon dorsatum (Linnaeus) Dermacentor andersoni Stiles Dermacentor variabilis (Say) Ixodes cookei Packard

ORDER ARTIODACTYLA, even-toed hoofed animals Family Cervidae Rocky Mountain mule deer, Odocoileus hemionus hemionus (Rafinesque) Otobius megnini (Dugès) Dermacentor andersoni Stiles Dermacentor albipictus (Packard) Columbian black-tailed deer, Odocoileus hemionus columbianus (Richardson) Dermacentor albipictus (Packard) Ixodes pacificus Cooley and Kohls White-tailed deer, Odocoileus virginianus (Zimmermann) Dermacentor albipictus (Packard) Dermacentor variabilis (Say) Ixodes scapularis Say Elk, Cervus canadensis (Linnaeus) Dermacentor albipictus (Packard)

Family Bovidae Montana mountain goat, Oreamnos americanus missoulae Allen Otobius megnini (Dugès) Dermacento andersoni Stiles Rocky Mountain bighorn, Ovis canadensis canadensis Shaw Otobius megnini (Dugès) Dermacentor andersoni Stiles Domestic sheep, Ovis aries Linnaeus Dermacentor andersoni Stiles Dermacentor variabilis (Say) Ixodes pacificus Cooley and Kohls Domestic goat, Capra aegagrus hircus (Linnaeus) Dermacentor andersoni Stiles Ixodes pacificus Cooley and Kohls 292 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

American bison, Bison bison bison (Linnaeus) Dermacentor albipictus (Packard) Dermacentor variabilis (Say) Caribou, Rangifer tarandus (Linnaeus) Dermacentor albipictus (Packard) Moose, Alces americanus (Linnaeus) Dermacentor albipictus (Packard) Cow, Bos taurus Linnaeus Dermacentor andersoni Stiles Dermacentor albipictus (Packard) Dermacentor variabilis (Say) Haemaphysalis chordeilis (Packard) Ixodes pacificus Cooley and Kohls Ixodes cookei Packard

Order Perissodactyla, odd-toed hoofed animals Family Equidae Horse, Equus caballus Linnaeus Dermacentor andersoni Stiles Dermacentor albipictus (Packard) Dermacentor variabilis (Say) Haemaphysalis chordeilis (Packard) Ixodes kingi Bishopp

Class Aves Order Suliformes, cormorants and allies Family Phalacorcoracidae Double-crested Cormorant, Phalacrocorax auritus (Lesson) Haemaphysalis leporispalustris (Packard) Cormorants, Phalacrocorax species Ixodes signatus Birula Ixodes uriae White

Order Anseriformes, ducks, geese, and allies Family Anatidae Mallard, Anas platyrhynchos Linnaeus Haemaphysalis leporispalustris (Packard) Northern shoveler, Anas clypeata Linnaeus Haemaphysalis leporispalustris (Packard) Wood duck, Aix sponsa (Linnaeus) Ixodes scapularis

Order Accipitriformes, hawks, eagles, and vultures Family Accipitridae Bald eagle, Haliaeetus leucocephalus (Linnaeus) Haemaphysalis leporispalustris (Packard) Ixodes auritulus Neumann A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 293

Northern harrier, Circus cyaneus (Linnaeus) Haemaphysalis leporispalustris (Packard) Sharp-shinned hawk, Accipiter striatus Vieillot Haemaphysalis leporispalustris (Packard) Cooper’s hawk, Accipiter cooperii (Bonaparte) Haemaphysalis leporispalustris (Packard) Northern goshawk, Accipiter gentilis (Linnaeus) Haemaphysalis leporispalustris (Packard) Broad-winged hawk, Buteo platypterus (Vieillot) Haemaphysalis leporispalustris (Packard) Swainson’s hawk, Buteo swainsoni Bonaparte Dermacentor variabilis (Say) Hawk (genus unknown) Dermacentor andersoni Stiles

Order Falconiformes, falcons Family Falconidae Prairie falcon, Falco mexicanus Schlegel Haemaphysalis leporispalustris (Packard) Carios concanensis (Cooley and Kohls)

Order Galliformes, gallinaceous birds Family Phasianidae Greater sage grouse, Centrocercus urophasianus (Bonaparte) Dermacentor andersoni Stiles Ruffed grouse, Bonasa umbellus (Linnaeus) Haemaphysalis leporispalustris (Packard) Haemaphysalis chordeilis (Packard) Spruce grouse, Falcipennis canadensis (Linnaeus) Haemaphysalis leporispalustris Sharp-tailed grouse, Tympanuchus phasianellus (Linnaeus) Haemaphysalis leporispalustris (Packard) Haemaphysalis chordeilis (Packard) Prairie chicken, Tympanuchus species Haemaphysalis chordeilis (Packard) Sooty grouse, Dendragapus fuliginosus (Ridway) Ixodes auritulus Neumann Grouse (genus unknown) Dermacentor andersoni Stiles Haemaphysalis leporispalustris (Packard) Haemaphysalis chordeilis (Packard) Ixodes pacificus Cooley and Kohls Ixodes auritulus Neumann Ring-necked pheasant, Phasianus colchicus Linnaeus Haemaphysalis leporispalustris (Packard) Haemaphysalis chordeilis (Packard) Pheasant, Phasianus species Haemaphysalis leporispalustris (Packard) Haemaphysalis chordeilis (Packard) 294 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Domestic turkey, Meleagris gallopavo Linnaeus Haemaphysalis leporispalustris (Packard) Haemaphysalis chordeilis (Packard) Domestic chicken, Gallus gallus domesticus (Linnaeus) Haemaphysalis chordeilis (Packard)

Family Odontophoridae California quail, Callipepla californica (Shaw) Ixodes pacificus Cooley and Kohls Ixodes auritulus Neumann Ixodes spinipalpis Hadwen and Nutall

Order Gruiformes, cranes, rails, and allies Family Rallidae American coot, Fulica americana Gmelin Haemaphysalis leporispalustris (Packard) Sora, Porzana carolina (Linnaeus) Haemaphysalis leporispalustris (Packard)

Order Charadriiformes, shore birds, gulls, auks, and allies Family Scolopacidae American woodcock, Scolopax minor Gmelin Haemaphysalis leporispalustris (Packard) Wilson’s snipe, Gallinago delicata Ord Haemaphysalis leporispalustris (Packard)

Family Alcidae Razor-billed auk, Alca torda Linnaeus Ixodes uriae White Common murre, Uria aalge (Pontoppidan) Ixodes uriae White Atlantic puffin,Fratercula arctica (Linnaeus) Ixodes uriae White

Order Cuculiformes Family Cuculidae Black-billed Cuckoo, Coccyzus erythrophthalmus (Wilson) Haemaphysalis leporispalustris (Packard)

Order Strigiformes, owls Family Strigidae Great horned owl, Bubo virginianus (Gmelin) Haemaphysalis leporispalustris (Packard) Dermacentor variabilis (Say) Snowy owl, Bubo scandiacus (Linnaeus) Haemaphysalis leporispalustris (Packard) Ixodes cookei Packard Northern hawk owl, Surnia ulula (Linnaeus) Haemaphysalis leporispalustris (Packard) A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 295

Barred owl, Strix varia Barton Haemaphysalis leporispalustris (Packard) Great grey owl, Strix nebulosa Forster Haemaphysalis leporispalustris (Packard) Long-eared owl, Asio otus (Linnaeus) Haemaphysalis leporispalustris (Packard) Owl, Strix species Haemaphysalis leporispalustris (Packard)

Order Caprimulgiformes, nighthawks and allies Family Caprimulgidae Common nighthawk, Chordeiles minor (Forster) Haemaphysalis leporispalustris (Packard)

Order Piciformes, woodpeckers Family Picidae Northern flicker, Colaptes auratus (Linnaeus) Haemaphysalis leporispalustris (Packard) Woodpecker (genus unknown) Haemaphysalis leporispalustris (Packard)

ORDER PASSERIFORMES, perching birds Family Tyrannidae Eastern wood pewee, Contopus virens (Linnaeus) Haemaphysalis leporispalustris (Packard)

Family Corvidae Steller’s jay, Cyanocitta stelleri (Gmelin) Ixodes auritulus Neumann Blue jay, Cyanocitta cristata (Linnaeus) Ixodes scapularis Say Black-billed magpie, Pica hudsonia Sabine Haemaphysalis leporispalustris (Packard) American crow, Corvus brachyrhynchos Brehm Haemaphysalis leporispalustris (Packard) Ixodes auritulus Neumann Common raven, Corvus corax Linnaeus Haemaphysalis leporispalustris (Packard)

Family Hirundinidae Cliff swallow, Petrochelidon pyrrhonota Vieillot Ixodes baergi Cooley and Kohls Ixodes howelli Cooley and Kohls

Family Sittidae White-breasted nuthatch, Sitta carolinensis Latham Haemaphysalis leporispalustris (Packard) 296 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Family Paridae Black-capped chickadee, Parus atricapillus (Linnaeus) Ixodes scapularis Say

Family Certhiidae Brown creeper, Certhia americana Bonaparte Ixodes auritulus Neumann

Family Troglodytidae Canyon wren, Catherpes mexicanus (Swainson) Haemaphysalis leporispalustris (Packard) Carolina wren, Thryothorus ludovicianus (Latham) Haemaphysalis leporispalustris (Packard) Bewick’s wren, Thyromanes bewickii (Audubon) Ixodes auritulus Neumann House wren, Troglodytes aedon (Vieillot) Haemaphysalis leporispalustris (Packard) Ixodes scapularis Say Winter wren, Troglodytes troglodytes (Linnaeus) Ixodes auritulus Neumann Sedge wren, Cistothorus platensis (Latham) Dermacentor variabilis (Say)

Family Turdidae Eastern bluebird, Sialia sialis (Linnaeus) (nest) Argas cooleyi Kohls and Hoogstraal Ornithodoros hermsi Wheeler, Herms, and Meyer Veery, Catharus fuscescens (Stephens) Haemaphysalis leporispalustris (Packard) Ixodes scapularis Say Ixodes muris Bishopp and Smith Ixodes brunneus Koch Grey-cheeked thrush, Catharus minimus (Lafresnaye) Amblyomma americanum (Linnaeus) Ixodes scapularis Say Swainson’s thrush, Catharus ustulatus (Nuttall) Haemaphysalis leporispalustris (Packard) Ixodes pacificus Cooley and Kohls Ixodes scapularis Say Ixodes auritulus Neumann Ixodes dentatus Marx Amblyomma maculatum Koch Hermit thrush, Catharus guttatus (Pallas) Haemaphysalis leporispalustris (Packard) Ixodes scapularis Say Ixodes spinipalpis Hadwen and Nuttall Ixodes muris Bishopp and Smith A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 297

Ixodes auritulus Neumann Ixodes brunneus Koch American robin, Turdus migratorius Linnaeus Haemaphysalis leporispalustris (Packard) Ixodes scapularis Say Ixodes auritulus Neumann Varied thrush, Ixoreus naevius Gmelin Ixodes spinipalpis Hadwen and Nuttall Ixodes auritulus Neumann

Family Mimidae Grey catbird, Dumetella carolinensis (Linnaeus) Ixodes scapularis Say Ixodes brunneus Koch Brown thrasher, Toxostoma rufum (Linnaeus) Haemaphysalis leporispalustris (Packard)

Parulidae Tennessee warbler, Oreothlypis peregrina (Wilson) Haemaphysalis leporispalustris (Packard) Orange-crowned warbler, Oreothlypis celata (Say) Ixodes scapularis Say Nashville warbler, Oreothlypis ruficapilla (Wilson) Haemaphysalis leporispalustris (Packard) Northern parula, Setophaga americana (Linnaeus) Haemaphysalis leporispalustris (Packard) Yellow warbler, Setophaga petechia (Linnaeus) Haemaphysalis leporispalustris (Packard) Ixodes scapularis Say Magnolia warbler, Setophaga magnolia (Wilson) Haemaphysalis leporispalustris (Packard) Ixodes scapularis Say Yellow-rumped warbler, Setophaga coronata (Linnaeus) Haemaphysalis leporispalustris (Packard) Palm warbler, Setophaga palmarum (Gmelin) Haemaphysalis leporispalustris (Packard) Blackpoll warbler, Setophaga striata (Forster) Haemaphysalis leporispalustris (Packard) American redstart, Setophaga ruticilla (Linnaeus) Haemaphysalis leporispalustris (Packard) Ixodes scapularis Say Black-and-white warbler, Mniotilta varia (Linnaeus) Haemaphysalis leporispalustris (Packard) Ovenbird, Seiurus aurocapillus (Linnaeus) Haemaphysalis leporispalustris (Packard) Northern waterthrush, Seiurus noveboracensis (Gmelin) Haemaphysalis leporispalustris (Packard) 298 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Ixodes scapularis Say Ixodes muris Bishopp and Smith Mourning warbler, Oporornis philadelphia (Wilson) Haemaphysalis leporispalustris (Packard) Ixodes scapularis Say Ixodes muris Bishopp and Smith Wilson’s warbler, Cardellina pusilla (Wilson) Ixodes pacificus Cooley and Kohls Canada warbler, Cardellina canadensis (Linnaeus) Haemaphysalis leporispalustris (Packard) Common yellowthroat, Geothlypis trichas (Linnaeus) Haemaphysalis leporispalustris (Packard) Ixodes auritulus Neumann Ixodes scapularis Say Ixode muris Bishopp and Smith Warbler (genus unknown) Haemaphysalis leporispalustris (Packard)

Family Emberizidae American tree sparrow, Spizella arborea (Wilson) Haemaphysalis leporispalustris (Packard) Chipping sparrow, Spizella passerina (Bechstein) Haemaphysalis leporispalustris (Packard) Haemaphysalis chordeilis (Packard) Ixodes scapularis Say Clay-coloured sparrow, Spizella pallida (Swainson) Haemaphysalis leporispalustris (Packard) Le Conte’s Sparrow, Ammodramus leconteii (Audubon) Haemaphysalis leporispalustris (Packard) Fox sparrow, Passerella iliaca (Merrem) Ixodes auritulus Neumann Ixodes brunneus Koch Song sparrow, Melospiza melodia (Wilson) Haemaphysalis leporispalustris (Packard) Ixodes muris Bishopp and Smith Ixodes auritulus Neumann Ixodes brunneus Koch Lincoln’s sparrow, Melospiza lincolnii (Audubon) Haemaphysalis leporispalustris (Packard) Ixodes muris Bishopp and Smith Ixodes scapularis Say White-throated sparrow, Zonotrichia albicollis (Gmelin) Haemaphysalis leporispalustris (Packard) Ixodes brunneus Koch Ixodes scapularis Say White-crowned sparrow, Zonotrichia leucophrys (Forster) Haemaphysalis leporispalustris (Packard) A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 299

Golden-crowned sparrow, Zonotrichia atricapilla (Gmelin) Argas persicus (Oken) Haemaphysalis leporispalustris (Packard) Ixodes spinipalpis Hadwen and Nuttall Ixodes auritulus Neumann Dark-eyed junco, Junco hyemalis (Linnaeus) Haemaphysalis leporispalustris (Packard) Ixodes brunneus Koch Ixodes auritulus Neumann

Family Cardinalidae Northern cardinal, Cardinalis cardinalis (Linnaeus) Haemaphysalis leporispalustris (Packard) Rose-breasted grosbeak, Pheucticus ludovicianus (Linnaeus) Haemaphysalis leporispalustris (Packard)

Family Icteridae Red-winged blackbird, Agelaius phoeniceus (Linnaeus) Haemaphysalis leporispalustris (Packard) Ixodes scapularis Say Western meadowlark, Sturnella neglecta Audubon Haemaphysalis leporispalustris (Packard) Yellow-headed blackbird, Xanthocephalus xanthocephalus (Bonaparte) Haemaphysalis leporispalustris (Packard) Haemaphysalis chordeilis (Packard) Brewer’s blackbird, Euphagus cyanocephalus (Wagler) Haemaphysalis leporispalustris (Packard) Common grackle, Quiscalus quiscula (Linnaeus) Haemaphysalis leporispalustris (Packard) Ixodes scapularis Say Brown-headed cowbird, Molothrus ater (Boddaert) Haemaphysalis leporispalustris (Packard) Orchard oriole, Icterus spurius (Linnaeus) Haemaphysalis leporispalustris (Packard) Baltimore oriole, Icterus galbula (Linnaeus) Haemaphysalis leporispalustris (Packard)

Family Fringillidae Grey-crowned rosy-finch, Leucosticte tephrocotis (Swainson) Ixodes signatus Birula Purple finch, Haemorhous purpureus (Gmelin) Haemaphysalis leporispalustris (Packard) Pine siskin, Spinus pinus (Wilson) Haemaphysalis leporispalustris (Packard)

Family Passeridae House sparrow, Passer domesticus (Linnaeus) Haemaphysalis leporispalustris (Packard)

300 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

By drag and clothing Amblyomma americanum (Linnaeus) Dermacentor andersoni Stiles Dermacentor albipictus (Packard) Dermacentor variabilis (Say) Haemaphysalis chordeilis (Packard) Ixodes pacificus Cooley and Kohls Ixodes scapularis Say Ixodes spinipalpis Hadwen and Nuttall

Appendix 2. Tick-host index Argasidae Argas cooleyi Kohls and Hoogstraal Bluebird, Sialia sialis (Linnaeus) (nest) Argas persicus (Oken) Golden-crowned sparrow, Zonotrichia atricapilla (Gmelin) Carios concanensis (Cooley and Kohls) Prairie falcon, Falco mexicanus Schlegel Carios kelleyi (Cooley and Kohls) Big brown bat, Eptesicus fuscus (Palisot de Beauvois) Little brown bat, Myotis lucifugus (LeConte) Long-eared bat, Myotis evotis (Allen) Long-legged bat, Myotis volans Allen Small-footed bat, Myotis ciliolabrum Merriam Ornithodoros hermsi Wheeler, Herms, and Meyer Bat or mouse (genus unknown) Human, Homo sapiens Linnaeus Bluebird, Sialia sialis (Linnaeus) (nest) Otobius megnini (Dugès) Domestic cat, Felis silvestris catus Linnaeus Rocky Mountain mule deer, Odocoileus hemionus hemionus (Rafinesque) Montana mountain goat, Oreamnos americanus missoulae Allen Rocky Mountain bighorn, Ovis canadensis canadensis Shaw Otobius lagophilus Cooley and Kohls Domestic cat, Felis silvestris catus Linnaeus White-tailed jack rabbit, Lepus townsendii Bachman

Ixodidae Dermacentor andersoni Stiles Human, Homo sapiens Linnaeus Bear, Ursus species Coyote, Canis latrans Say Dog, Canis lupus familiaris Linnaeus Domestic cat, Felis silvestris catus Linnaeus Badger, Taxidea taxus (Schreber) Pika, Ochotona princeps (Richardson) White-tailed jack rabbit, Lepus townsendii Bachman A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 301

Yellow-bellied marmot, Marmota flaviventris (Audubon and Bachman) Richardson’s ground squirrel, Urocitellus richardsonii (Sabine) Columbian ground squirrel, Callospermophilus columbianus (Ord) Franklin’s ground squirrel, Callospermophilus franklinii (Sabine) Golden-mantled ground squirrel, Callospermophilus lateralis (Say) Thirteen-lined ground squirrel, Ictidomys tridecemlineatus (Mitchill) Black-tailed prairie-dog, Cynomys ludovicianus ludovicianus (Ord) Chipmunk, Eutamias species Tree squirrel, Tamiasciurus species Northern pocket gopher, Thomomys talpoides Richardson Deer mouse, Peromyscus maniculatus (Wagner) Western bushy-tailed woodrat, Neotoma cinerea (Ord) Meadow vole, Microtus pennsylvanicus (Ord) Meadow voles, Microtus species Pallid pygmy vole, Lemmiscus curtatus pallidus (Merriam) Western jumping mouse, Zapus princeps minor Preble Porcupine, Erethizon dorsata (Linnaeus) Rocky Mountain mule deer, Odocoileus hemionus hemionus (Rafinesque) Montana mountain goat, Oreamnos americanus missoulae Allen Rocky Mountain bighorn, Ovis canadensis canadensis Shaw Domestic sheep, Ovis aries Linnaeus Domestic goat, Capra aegagrus hircus (Linnaeus) Cow, Bos taurus Linnaeus Horse, Equus caballus Linnaeus Hawk (genus unknown) Greater sage grouse, Centrocercus urophasianus (Bonaparte) Grouse (genus unknown) Drag or clothing Dermacentor albipictus (Packard) Human, Homo sapiens Linnaeus Bear, Ursus species Coyote, Canis latrans Say Deer mouse, Peromyscus maniculatus (Wagner) Wolf, Canis lupus Linnaeus Rocky Mountain mule deer, Odocoileus hemionus hemionus (Rafinesque) Columbian black-tailed deer, Odocoileus hemionus columbianus (Richardson) White-tailed deer, Odocoileus virginianus (Zimmermann) Elk, Cervus canadensis (Linnaeus) American bison, Bison bison bison (Linnaeus) Reindeer, Rangifer tarandus (Linnaeus) Moose, Alces americanus (Linnaeus) Cow, Bos taurus Linnaeus Horse, Equus caballus Linnaeus Drag or clothing Dermacentor variabilis (Say) Masked shrew, Sorex cinereus Kerr Short-tailed shrew, Blarina brevicauda (Say) Water shrew, Sorex palustris Richardson 302 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Human, Homo sapiens Linnaeus Raccoon, Procyon lotor Linnaeus Black bear, Ursus americanus (Pallas) Dog, Canis lupus familiaris Linnaeus Domestic cat, Felis silvestris catus Linnaeus Eastern striped skunk, Mephitis mephitis (Schreber) Short-tailed weasel, Mustela erminea Linnaeus Prairie long-tailed weasel, Mustela frenata longicauda Bonaparte Badger, Taxidea taxus (Schreber) Woodchuck, Marmota monax (Linnaeus) Red squirrel, Tamiasciurus hudsonicus (Erxleben) Grey squirrel, Sciurus carolinensis Gmelin Franklin’s ground squirrel, Poliocitellus franklinii (Sabine) Eastern chipmunk, Tamias striatus (Linnaeus) Thirteen-lined ground squirrel, Ictidomys tridecemlineatus (Mitchill) Northern flying squirrel, Glaucomys sabrinus (Shaw) Northern pocket gopher, Thomomys talpoides Richardson Deer mouse, Peromyscus maniculatus (Wagner) Southern bog lemming, Synaptomys cooperi (Baird) Red-backed vole, Myodes gapperi (Vigors) Meadow vole, Microtus pennsylvanicus (Ord) Meadow vole, Microtus species Meadow jumping mouse, Zapus hudsonius (Zimmermann) Western jumping mouse, Zapus princeps Preble Porcupine, Erethizon dorsatum (Linnaeus) Snowshoe hare, Lepus americana Erxleben Domestic sheep, Ovis aries Linnaeus American bison, Bison bison bison (Linnaeus) Cow, Bos taurus Linnaeus White-tailed deer, Odocoileus virginianus (Zimmermann) Horse, Equus caballus Linnaeus Swainson’s hawk, Buteo swainsoni Bonaparte Great horned owl, Bubo virginianus (Gmelin) Sedge wren, Cistothorus platensis (Latham) Drag or clothing Rhipicephalus sanguineus (Latreille) Human, Homo sapiens Linnaeus Dog, Canis lupus familiaris Linnaeus Haemaphysalis leporispalustris (Packard) Human, Homo sapiens Linnaeus Domestic cat, Felis silvestris catus Linnaeus White-tailed jack rabbit, Lepus townsendii Bachman Snowshoe hare, Lepus americanus Erxleben Eastern cottontail, Sylvilagus floridanus(Allen) Black Hills cottontail, Sylvilagus nuttallii grangeri (Allen) Rabbits, genus unknown Yellow-bellied marmot, Marmota flaviventris (Audubon and Bachman) Richardson’s ground squirrel, Urocitellus richardsonii (Sabine) A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 303

Chipmunk, Eutamias species Southern red squirrel, Tamiasciurus hudsonicus loquax (Bangs) Streator’s red squirrel, Tamiasciurus hudsonicus streatori (Allen) Tree squirrels, Tamiasciurus species Red-backed vole, Myodes gapperi (Vigors) Meadow vole, Microtus pennsylvanicus (Ord) Double-crested cormorant, Phalacrocorax auritus (Lesson) Mallard, Anas platyrhynchos Linnaeus Northern Shoveler, Anas clypeata Linnaeus Northern harrier, Circus cyaneus (Linnaeus) Sharp-shinned hawk, Accipiter striatus Vieillot Cooper’s hawk, Accipiter cooperii (Bonaparte) Northern goshawk, Accipiter gentilis (Linnaeus) Bald eagle, Haliaeetus leucocephalus (Linnaeus) Broad-winged hawk, Buteo platypterus (Vieillot) Prairie falcon, Falco mexicanus Schlegel Ruffed grouse, Bonasa umbellus (Linnaeus) Spruce grouse, Falcipennis canadensis (Linnaeus) Sharp-tailed grouse, Tympanuchus phasianellus (Linnaeus) Grouse (genus unknown) Ring-necked pheasant, Phasianus colchicus Linnaeus Domestic turkey, Meleagris gallopavo Linnaeus American coot, Fulica americana Gmelin Sora, Porzana carolina (Linnaeus) American woodcock, Scolopax minor Gmelin Wilson’s snipe, Gallinago delicata Ord Black-billed cuckoo, Coccyzus erythrophthalmus (Wilson) Great horned owl, Bubo virginianus (Gmelin) Snowy owl, Bubo scandiacus (Linnaeus) Northern hawk owl, Surnia ulula (Linnaeus) Barred owl, Strix varia Barton Great grey owl, Strix nebulosa Forster Long-eared owl, Asio otus (Linnaeus) Owl, Strix species Common nighthawk, Chordeiles minor (Forster) Northern flicker, Colaptes auratus (Linnaeus) Woodpecker (genus unknown) Eastern wood pewee, Contopus virens (Linnaeus) Black-billed magpie, Pica hudsonia Sabine American crow, Corvus brachyrhynchos Brehm Common raven, Corvus corax Linnaeus White-breasted nuthatch, Sitta carolinensis Latham Canyon wren, Catherpes mexicanus (Swainson) Carolina wren, Thryothorus ludovicianus (Latham) House wren, Troglodytes aedon (Vieillot) Veery, Catharus fuscescens (Stephens) Swainson’s thrush, Catharus ustulatus (Nuttall) Hermit thrush, Catharus guttatus (Pallas) 304 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

American robin, Turdus migratorius Linnaeus Brown thrasher, Toxostoma rufum (Linnaeus) Tennessee warbler, Oreothlypis peregrina (Wilson) Magnolia warbler, Setophaga magnolia (Wilson) Nashville warbler, Oreothlypis ruficapilla(Wilson) Northern parula, Setophaga americana (Linnaeus) Yellow warbler, Setophaga petechia (Linnaeus) Yellow-rumped warbler, Setophaga coronata (Linnaeus) Palm warbler, Setophaga palmarum (Gmelin) American redstart, Setophaga ruticilla (Linnaeus) Blackpoll warbler, Setophaga striata (Forster) Black-and-white warbler, Mniotilta varia (Linnaeus) Ovenbird, Seiurus aurocapillus (Linnaeus) Northern waterthrush, Seiurus noveboracensis (Gmelin) Mourning warbler, Oporornis philadelphia (Wilson) Canada warbler, Cardellina canadensis (Linnaeus) Common yellowthroat, Geothlypis trichas (Linnaeus) Warbler (genus unknown) American tree sparrow, Spizella arborea (Wilson) Chipping sparrow, Spizella passerina (Bechstein) Clay-coloured sparrow, Spizella pallida (Swainson) Le Conte’s Sparrow, Ammodramus leconteii (Audubon) Song sparrow, Melospiza melodia (Wilson) Lincoln’s sparrow, Melospiza lincolnii (Audubon) White-throated sparrow, Zonotrichia albicollis (Gmelin) White-crowned sparrow, Zonotrichia leucophrys (Forster) Golden-crowned sparrow, Zonotrichia atricapilla (Gmelin) Dark-eyed junco, Junco hyemalis (Linnaeus) Northern cardinal, Cardinalis cardinalis (Linnaeus) Rose-breasted grosbeak, Pheucticus ludovicianus (Linnaeus) Red-winged blackbird, Agelaius phoenicius (Linnaeus) Western meadowlark, Sturnella neglecta Audubon Yellow-headed blackbird, Xanthocephalus xanthocephalus (Bonaparte) Brewer’s blackbird, Euphagus cyanocephalus (Wagler) Common grackle, Quiscalus quiscula (Linnaeus) Brown-headed cowbird, Molothrus ater (Boddaert) Orchard oriole, Icterus spurius (Linnaeus) Baltimore oriole, Icterus galbula (Linnaeus) Purple finch, Haemorhous purpureus (Gmelin) Pine siskin, Spinus pinus (Wilson) House sparrow, Passer domesticus (Linnaeus) Haemaphysalis chordeilis (Packard) Human, Homo sapiens Linnaeus Yellow-bellied marmot, Marmota flaviventris (Audubon and Bachman) Streator’s red squirrel, Tamiasciurus hudsonicus streatori (Allen) Tree squirrel, Tamiasciurus species Cow, Bos taurus Linnaeus Horse, Equus caballus Linnaeus Ruffed grouse, Bonasa umbellus (Linnaeus) A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 305

Sharp-tailed grouse, Tympanuchus phasianellus (Linnaeus) Prairie chicken, Tympanuchus species Grouse (genus unknown) Ring-necked pheasant, Phasianus colchicus Linnaeus Pheasant, Phasianus species Domestic turkey, Meleagris gallopavo Linnaeus Domestic chicken, Gallus gallus domesticus (Linnaeus) Chipping sparrow, Spizella passerina (Bechstein) Yellow-headed blackbird, Xanthocephalus xanthocephalus (Bonaparte) Drag or clothing Ixodes pacificus Cooley and Kohls Northern alligator lizard, Elgaria coeruleus principis Baird and Girard Human, Homo sapiens Linnaeus Dog, Canis lupus familiaris Linnaeus Domestic cat, Felis silvestris catus Linnaeus Mink, Neovison vison (Schreber) Scheffer’s mole, Scapanus orarius schefferi Jackson Shrew, Sorex species Chipmunk, Eutamias species Tree squirrel, Tamiasciurus species Deer mouse, Peromyscus maniculatus (Wagner) Columbian black-tailed deer, Odocoileus hemionus columbianus (Richardson) Domestic sheep, Ovis aries Linnaeus Domestic goat, Capra aegagrus hircus (Linnaeus) Cow, Bos taurus Linnaeus California quail, Callipepla californica, (Shaw) Grouse (genus unknown) Swainson’s thrush, Catharus ustulatus (Nuttall) Wilson’s warbler, Cardellina pusilla (Wilson) Drag or clothing Ixodes spinipalpis Hadwen and Nuttall Human, Homo sapiens Linnaeus Shrews, Sorex species Pika, Ochotona princeps (Richardson) Rabbit, Lepus species Chipmunk, Eutamias species Dusky harvest mouse, Reithrodontomys megalotis nigrescens Howell Deer mouse, Peromyscus maniculatus (Wagner) Western bushy-tailed woodrat, Neotoma cinerea (Ord) Rat, Rattus species California quail, Callipepla californica (Shaw) Hermit thrush, Catharus guttatus (Pallas) Varied thrush, Ixoreus naevius Gmelin Golden-crowned sparrow, Zonotrichia atricapilla (Gmelin) Drag or clothing Ixodes scapularis Say Human, Homo sapiens Linnaeus Dog, Canis lupus familiaris Linnaeus 306 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Domestic cat, Felis silvestris catus Linnaeus Northern pocket gopher, Thomomys talpoides Richardson Deer mouse, Peromyscus maniculatus (Wagner) White-footed mouse, Peromyscus leucopus (Rafinesque) Red-backed vole, Myodes gapperi (Vigors) White-tailed deer, Odocoileus virginianus (Zimmermann) Wood Duck, Aix sponsa (Linnaeus) Blue jay, Cyanocitta cristata (Linnaeus) Black-capped chickadee, Parus atricapillus (Linnaeus) House wren, Troglodytes aedon (Vieillot) Veery, Catharus fuscescens (Stephens) Grey-cheeked thrush, Catharus minimus (Lafresnaye) Swainson’s thrush, Catharus ustulatus (Nuttall) Hermit thrush, Catharus guttatus (Pallas) American robin, Turdus migratorius Linnaeus Grey catbird, Dumetella carolinensis (Linnaeus) Orange-crowned warbler, Oreothlypis celata (Say) Yellow warbler, Setophaga petechia (Linnaeus) Magnolia warbler, Setophaga magnolia (Wilson) American redstart, Setophaga ruticilla (Linnaeus) Northern waterthrush, Seiurus noveboracensis (Gmelin) Mourning warbler, Oporornis philadelphia (Wilson) Common yellowthroat, Geothlypis trichas (Linnaeus) Chipping sparrow, Spizella passerina (Bechstein) Lincoln’s sparrow, Melospiza lincolnii (Audubon) White-throated sparrow, Zonotrichia albicollis (Gmelin) Red-winged blackbird, Agelaius phoenicius (Linnaeus) Common grackle, Quiscalus quiscula (Linnaeus) Ixodes angustus Neumann Scheffer’s mole, Scapanus orarius schefferi Jackson Hairy-tailed mole, Parascalops breweri (Bachman) Masked shrew, Sorex cinereus Kerr Smoky shrew, Sorex fumeus Miller Gaspé shrew, Sorex gaspensis Batchelder Water shrew, Sorex palustris Richardson Pygmy shrew, Sorex hoyi Baird Star-nosed mole, Condylura cristata (Linnaeus) Short-tailed shrew, Blarina brevicauda (Say) Shrews, Sorex species Human, Homo sapiens Linnaeus Dog, Canis lupus familiaris Linnaeus Domestic cat, Felis silvestris catus Linnaeus Pika, Ochotona princeps (Richardson) Rabbit, Lepus species Northwestern chipmunk, Tamias striatus lysteri (Richarson) Lake Superior chipmunk, Eutamias minimus neglectus (Allen) Chipmunks, Eutamias species Southern red squirrel, Tamiasciurus hudsonicus loquax (Bangs) A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 307

Vancouver red squirrel, Tamiasciurus hudsonicus vancouverensis (Allen) Northwestern red-bellied squirrel, Tamiasciurus douglasii mollipilosus (Audubon and Bachman) Tree squirrels, Tamiasciurus species Northern flying squirrel, Glaucomys sabrinus (Shaw) Deer mouse, Peromyscus maniculatus (Wagner) White-footed mouse, Peromyscus leucopus (Rafinesque) Western bushy-tailed woodrat, Neotoma cinerea (Ord) Northern bog lemming, Synaptomys borealis (Richardson) Red-backed vole, Myodes gapperi (Vigors) Drummond’s meadow vole, Microtus pennsylvanicus drummondi (Audubon and Bachman) Rock vole, Microtus chrotorrhinus (Miller) Long-tailed vole, Microtus longicaudus (Merriam) Heather vole, Phenacomys intermedius (Merriam) Meadow voles, Microtus species Meadow jumping mouse, Zapus hudsonius (Zimmermann) Jumping mouse, Zapus species Rat, Rattus species Ixodes soricis Gregson Vagrant shrew, Sorex vagrans Baird Masked shrew, Sorex cinereus Kerr Montane or dusky shrew, Sorex monticolus Merriam Shrews, Sorex species Ixodes ochotonae Gregson Pika, Ochotona princeps (Richardson) Deer mouse, Peromyscus maniculatus (Wagner) Western bushy-tailed woodrat, Neotoma cinerea (Ord) Meadow vole, Microtus species Ixodes texanus Banks Pacific raccoon,Procyon lotor pacificusMerriam Eastern raccoon, Procyon 1otor lotor Linnaeus Dog, Canis lupus familiaris Linnaeus Mink, Neovison vison (Schreber) Weasels, Mustela species Yellow-bellied marmot, Marmota flaviventris (Audubon and Bachman) Tree squirrel, Tamiasciurus species Ixodes hearlei Gregson Streator’s red squirrel, Tamiasciurus hudsonicus streatori (Allen) Ixodes marxi Banks Human, Homo sapiens Linnaeus Domestic cat, Felis silvestris catus Linnaeus Dog, Canis lupus familiaris Linnaeus Snowshoe hare, Lepus americanus Erxleben Southern red squirrel, Tamiasciurus hudsonicus loquax (Bangs) Grey squirrel, Sciurus carolinensis Gmelin Northern flying squirrel, Glaucomys sabrinus (Shaw) 308 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Ixodes rugosus Bishopp Coyote, Canis latrans Say Puget Sound spotted skunk, Spilogale phenax olympica (Elliot) (= Western Spotted Skunk, Spilogale gracilis latifrons (Merriam)) Ixodes marmotae Cooley and Kohls Yellow-bellied marmot, Marmota flaviventris (Audubon and Bachman) Columbian ground squirrel, Callospermophilus columbianus (Ord) Western bushy-tailed woodrat, Neotoma cinerea (Ord) Ixodes cookei Packard Human, Homo sapiens Linnaeus Eastern raccoon, Procyon 1otor lotor Linnaeus Dog, Canis lupus familiaris Linnaeus Domestic cat, Felis silvestris catus Linnaeus Pine marten, Martes americana americana (Turton) Fisher, Martes pennanti pennanti (Erxleben) Mink, Neovison vison (Schreber) Eastern striped skunk, Mephitis mephitis (Schreber) Woodchuck, Marmota monax (Linnaeus) Porcupine, Erethizon dorsatum (Linnaeus) Cow, Bos taurus Linnaeus Snowy owl, Bubo scandiacus (Linnaeus) Ixodes kingi Bishopp Dog, Canis lupus familiaris Linnaeus Domestic cat, Felis silvestris catus Linnaeus Weasels, Mustela species Badger, Taxidea taxus (Schreber) Richardson’s ground squirrel, Urocitellus richardsonii (Sabine) Northern pocket gopher, Thomomys talpoides Richardson Pallid pigmy vole, Lemmiscus curtatus pallidus (Merriam) Horse, Equus caballus Linnaeus Ixodes sculptus Neumann Prairie long-tailed weasel, Mustela frenata longicauda Bonaparte Weasels, Mustela species Badger, Taxidea taxus (Schreber) White-tailed jack rabbit, Lepus townsendii Bachman Rabbits (genus not reported) Richardson’s ground squirrel, Urocitellus richardsonii (Sabine) Columbian ground squirrel, Callospermophilus columbianus (Ord) Franklin’s ground squirrel, Callospermophilus franklinii (Sabine) Thirteen-lined ground squirrel, Ictidomys tridecemlineatus (Mitchill) Northern pocket gopher, Thomomys talpoides Richardson Ixodes muris Bishopp and Smith Human, Homo sapiens Linnaeus Dog, Canis lupus familiaris Linnaeus Domestic cat, Felis silvestris catus Linnaeus Eastern woodland jumping mouse, Napaeozapus insignis insignis (Miller) Veery, Catharus fuscescens (Stephens) Hermit thrush, Catharus guttatus (Pallas) A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 309

Northern waterthrush, Seiurus noveboracensis (Gmelin) Mourning warbler, Oporornis philadelphia (Wilson) Common yellowthroat, Geothlypis trichas (Linnaeus) Lincoln’s sparrow, Melospiza lincolnii (Audubon) Song sparrow, Melospiza melodia (Wilson) Ixodes auritulus Neumann Meadow vole, Microtus species Bald eagle, Haliaeetus leucocephalus (Linnaeus) Sooty grouse, Dendragapus fuliginosus (Ridway) California quail, Callipepla californica (Shaw) Grouse (genus unknown) Steller’s jay, Cyanocitta stelleri (Gmelin) American crow, Corvus brachyrhynchos Brehm Brown creeper, Certhia americana Bonaparte Winter wren, Troglodytes troglodytes (Linnaeus) Bewick’s wren, Thyromanes bewickii (Audubon) Swainson’s thrush, Catharus ustulatus (Nuttall) Hermit thrush, Catharus guttatus (Pallas) American robin, Turdus migratorius Linnaeus Varied thrush, Ixoreus naevius Gmelin Common yellowthroat, Geothlypis trichas (Linnaeus) Song sparrow, Melospiza melodia (Wilson) Golden-crowned sparrow, Zonotrichia atricapilla (Gmelin) White-crowned sparrow, Zonotrichia leucophrys (Forster) Fox sparrow, Passerella iliaca (Merrem) Dark-eyed junco, Junco hyemalis (Linnaeus) Ixodes howelli Cooley and Kohls Cliff swallow, Petrochelidon pyrrhonota Vieillot Ixodes signatus Birula Cormorant, Phalacrocorax species Grey-crowned rosy-finch, Leucosticte tephrocotis (Swainson) Ixodes uriae White Cormorant, Phalacrocorax species Razor-billed auk, Alca torda Linnaeus Common murre, Uria aalge (Pontoppidan) Atlantic puffin,Fratercula arctica (Linnaeus) Ixodes banksi Bishopp North American beaver, Castor canadensis Kuhl Ixodes gregsoni Lindquist, Wu, and Redner Domestic dog, Canis lupus familiaris Linnaeus Pine marten, Martes americana (Turton) American mink, Neovison vison (Schreber) Weasel, Mustela species Ixodes brunneus Koch Hermit thrush, Catharus guttatus (Pallas) Veery, Catharus fuscescens (Stephens) Grey catbird, Dumetella carolinensis (Linnaeus) Song sparrow Melospiza melodia (Wilson) 310 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Fox sparrow, Passerella iliaca (Merrem) White-throated sparrow, Zonotrichia albicollis (Gmelin) Dark-eyed junco, Junco hyemalis (Linnaeus) Ixodes baergi Cooley and Kohls Cliff swallow, Petrochelidon pyrrhonota Vieillot Ixodes dentatus Marx Swainson’s thrush, Catharus ustulatus (Nuttall) Amblyomma americanum (Linnaeus) Human, Homo sapiens Linnaeus Dog, Canis lupus familiaris Linnaeus Grey-cheeked thrush, Catharus minimus (Lafresnaye) Amblyomma maculatum Koch Human, Homo sapiens Linnaeus Dog, Canis lupus familiaris Linnaeus Domestic cat, Felis silvestris catus Linnaeus Swainson’s thrush, Catharus ustulatus (Nuttall) A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 311

Index

Acarus 64 aequalis 194 aguilae 227 albipictus 3, 4, 54, 57, 70, 71–76, 83 Alectorobius 227, 229 Amblyomma 4, 31, 33, 38, 40, 44, 46, 48, 54, 60, 61, 64–66, 69, 70 amblyommii 48, 49, 66 Amelanchier alnifolia 79 americana 219, 242, 300 americanum 4, 33, 38, 40, 44–46, 49, 54, 64–66, 69 americanus 64, 83, 84, 93 anaemia 53 Anaplasma 40–43, 50, 75, 76, 82, 122, 139, 176 Anaplasma ovis 43, 76, 82 andersoni 4, 43–47, 51, 52, 54–56, 70, 71, 76–83 angustus 5, 97, 102, 105, 108, 144–148 anserina 38, 39, 226 arcticus 209 Argas 4, 26, 31, 38, 54, 55, 61, 217–219, 226, 242 Argasidae 1, 4, 15, 17, 26, 31, 58, 59, 216 auritulus 5, 97, 99, 104, 107, 108, 156–160 avian borreliosis 38 avisugus 113 Babesia 36, 41, 50, 51, 76, 87, 122, 127, 139, 144, 148, 201, 216, 228 babesiosis 51 bacilliformis 49 badger 83, 88 baergi 5, 95, 96, 99, 100, 103, 106, 160–163 banksi 5, 99, 101, 104–106, 164–167 Bartonella 49, 133, 140, 233 beaver 44, 45, 99, 167 becarii 213 bison 43, 73, 76, 88 Bison bison athabascae 73 Bison bison bison 73 Boophilus 3, 15, 20, 43, 60 borealis 109 Borrelia 37–39, 41, 49, 50, 75, 113, 122, 127, 133, 144, 148, 160, 226, 233, 235, 239 bovine anaplasmosis 42, 75, 76, 82 brunneus 4, 97, 101, 103, 106, 113–117 buffalo 3, 73 burgdorferi 37, 38, 41, 42, 49, 50, 75, 113, 122, 127, 133, 139, 144, 148, 160 burnetii 43, 44, 181, 249 bushy-tailed woodrat 81 caballi 76, 87 californicus 113, 127 312 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Callospermophilus lateralis 47 canadensis 48, 73, 167 canine ehrlichiosis 216 Canine monocytic ehrlichiosis 41 canis 41, 42, 51, 216, 249 Canis lupus familiaris 84 Carios 4, 5, 31, 61, 226, 227, 228, 229, 233 Castor canadensis 167 cat 42, 45, 50, 55, 83, 87, 88, 127, 129, 137, 140, 148, 172, 181, 190, 204, 242, 249 cattle 42–44, 55, 58, 66, 69, 73, 76, 79, 80, 83, 88, 91, 133, 140, 243, 249 Ceratixodes 4, 109, 209 chaffeensis 40, 42 chinche 219 chordeilis 4, 45, 88–91, 92 cinnabarina 88 cliff swallow 96, 163, 208, 218, 219 CME 41 Cochliomyia hominivorax 69 Colorado tick fever 1, 51, 52, 81, 242 Columbian black-tailed deer 129 Columbian ground squirrel 80 complanatum 66 concanensis 4, 227–231 cookei 5, 52, 99, 101, 102, 105, 108, 167–172, 190 cooleyi 4, 217–224 Coxiella 43, 44, 66, 181, 249 coyote 80, 194 cruciarius 167 Cytauxzoon 50, 87 Cytauxzoonosis 50 dammini 134, 137 dark-eyed junco 117 deer mouse 45, 52, 81, 84, 129, 132, 137 dentatus 4, 44, 46, 97, 101, 104, 107, 118–122, 141 Dermacentor 1, 3, 4, 14, 17, 20, 21, 31, 33, 43–46, 51, 54, 56, 57, 60, 69–71, 73, 75, 76, 79, 82, 83, 84, 87 Didelphis virginiana 84 divergens 122 diversifossus 141 dog 38, 40–43, 46, 48, 49, 51, 55, 56, 66, 69, 79, 83, 84, 88, 127, 129, 133, 137, 140, 148, 172, 181, 190, 194, 198, 201, 216 Douglas fir 129 eastern bluebird 219 eastern chipmunk 84 Ehrlichia 40–42, 50, 66, 87, 216, 249 Ehrlichiosis 40 electus 83 Elgaria coerulea 129, 287 A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 313

elk 73, 74, 76 equi 40, 42, 51 Erethizon dorsatum 84 Erysipelothrix rhusiopathiae 76 eudyptidis signata 209 evertsi 55 ewingii 40, 42, 66 felis 50, 87 foreli 64 Francisella 36, 44, 45, 66, 76, 167, 181, 242 frontalis 113 fuscous 134 Gerrhonotus 129 gnathosoma 5 Gonixodes 91 gregsoni 5, 16, 97, 99, 105, 107, 202–204 grizzly bear 80 groundhog 52, 137 Haemaphysalis 4, 14, 15, 17, 20, 31, 44–46, 54, 59, 88, 91, 93, 95 Haemobartonella 50 haemocanis 43 hearlei 5, 96, 100, 103, 108, 172–176 henselae 49, 133 hermsi 4, 39, 234–238 hermsii 39, 235 hexagonus 167, 185 HGA 40, 41, 42, 50, 133 hirsutus 109 HME 40, 66 holdenreidi 148 holocyclus 55 horse 3, 42, 51–53, 56, 66, 73, 76, 79, 83, 87, 88, 91, 133, 140, 243, 249 howelli 5, 97, 99, 104, 106, 205–208, 209 human granulocytic anaplasmosis 40, 41, 95, 133, 144 human 1, 33, 36–57, 66, 69, 75, 76, 80–84, 87, 88, 91, 95, 105, 113, 122, 127, 129, 132–134, 139, 140, 144, 148, 155, 167, 172, 190, 212, 216, 219, 226, 228, 233, 235, 239, 249 Hyalomma 3, 54, 109 hypersensitivity reactions 56 idiosoma 16 immunosuppression 58 Ixodes 1, 4, 5, 14–17, 20, 21, 26, 31, 33, 37, 41, 42, 44, 46, 49, 50, 52, 54, 55, 59, 64, 71, 83, 88, 91, 95–110, 112, 113, 117–119, 122, 126, 127, 129, 133, 134, 137, 138, 140, 141, 143, 144, 148, 152, 156, 160, 163–165, 167, 169, 172, 176, 180, 181, 185, 187, 190, 194, 197, 198, 201, 202, 204, 205, 209, 212, 213 Ixodidae 1, 4, 15, 16, 17, 20, 26, 31, 59, 61 Ixodiopsis 5, 144, 148, 152 Ixodiphagus 95 314 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins

Junco hyemalis 117, 302 kamshadalus 71, 73 kangaroo rat 181 kelleyi 4, 26, 227, 229–233 kelloggi 113 kingi 5, 98, 101, 104, 108, 176–181 lagophilus 4, 16, 26, 44, 240–242, 244–247 leporis 91 leporis-palustris 91 leporispalustris 4, 44–46, 48, 88, 89, 91–95 leporis proxima 91 Lepus americanus 84, 93 limbatus 213 lonestari 38 longispinosus 167 Lyme borreliosis 1, 37, 38, 75, 148, 172 maculatum 4, 48, 49, 60, 64, 66–69, 70 marginale 42, 75, 82 marmotae 5, 98, 102, 104, 108, 172, 181–185 Marmota flaviventris 81, 185 Marmota monax 52, 169 marten 172, 204 marxi 5, 52, 96, 100, 103, 108, 185–189 massiliae 48 mauritianus 219 meadow jumping mouse 84 meadow vole 45, 47, 84 megnini 4, 16, 26, 240, 241, 242–249 Meleagris gallopavo 91 Mephitis mephitis 84 microplus 43 microti 41, 50, 51, 127, 139, 144, 148 Microtus pennsylvanicus 45, 84 mink 204 miyamotoi 49 modestus 76 montanensis 47, 49 moose 3, 54, 57, 73, 74, 80 moshkovskii 228 moubata 54 mountain goat 73, 76, 243 mountain sheep 73, 76, 80, 243 mule deer 80 muris 4, 26, 41, 50, 98, 101, 104, 107, 122–127 muskrat 44, 45, 167 Mycoplasma haemocanis 43 Myodes gapperi 84, 148 Neotoma cinerea 81 A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 315

neotomae 141, 144 nigrolineatus 71 nigua 64 northern alligator lizard 129 northern pocket gopher 140, 155, 181 occidentalis 44, 46, 48, 51 ochotonae 5, 97, 102, 105, 108, 148–152 Odocoileus hemionus columbianus, 129 Odocoileus virginianus 40, 73, 137 Ondatra zibethicus 167 opossum 84, 88, 137 Ornithodoros 4, 20, 26, 31, 39, 44, 46, 54, 61, 227, 229, 234, 235, 239, 240, 242 Otobius 4, 14, 15, 26, 31, 44, 60, 240, 241, 242, 249 ovatum 66 ozarkus 134 pacificus 4, 37, 38, 40, 41, 49, 50, 98, 101, 104, 107, 127–134 parkeri 4, 39, 46, 48, 66, 69, 234, 239 parvirostris 209 Pavlovskyella 4, 234, 239 peacockii 47 Peromyscus leucopus 37, 51 Peromyscus maniculatus 81, 84, 129 Persicargus 217, 219 persicus 4, 16, 31, 39, 54, 217–226 persulcatus 49, 54 Petrochelidon pyrrhonota 163, 208, 218 phagocytophila 40 phagocytophilum 40, 41, 42, 50, 76, 122, 133, 139, 176 Pholeoixodes 5, 144, 148, 152, 160, 164, 167, 172, 176, 181, 185, 190, 194, 198, 202 platys 43 porcupine 80, 84, 169, 172 Powassan encephalitis 1, 52, 53 Powassan virus 50, 52, 53, 82, 139, 172, 190 pratti 134, 176, 198 Pseudotsuga menziesii 129 punctata cinnabarina 88 punctatissimus 213 puta 109 putus procellariae 109 Q fever 1, 43, 44, 249 quinquestriatus 83 quintana 49 rabbit 39, 44, 45, 47, 54, 55, 80, 81, 91, 93, 95, 97, 119, 122, 241, 242 raccoon 84, 95, 137, 169, 176, 201 Rangifer tarandus tarandus 73 red-backed vole 84, 148 reduvius 134 reindeer 73 316 E. E. Lindquist, T. D. Galloway, H. Artsob, L. R. Lindsay, M. Drebot, H. Wood, and R. G. Robbins resistance 57, 58 reticulatus 54, 71 rhipicephali 47, 48 Rhipicephalus 3, 5, 14, 17, 20, 26, 31, 41, 43, 44, 46, 48, 54, 55, 60, 213, 216 Rhipistoma 91 Rhynchoprion 64, 219 Richardson’s ground squirrel 81, 181 ricinus 134 Rickettsia 45–49, 66, 69, 95, 117, 140, 176, 201, 216, 233, 242 rickettsii 45–49, 117, 201, 242 RMSF 45–49 robertsoni 83 Rocky Mountain spotted fever 1, 45, 46, 52, 81, 87 rose 79 rostralis 91 rubicundus 213 rubripes 66 rugosus 5, 95, 99, 102, 105, 107, 190–194 rutilus 213 saddleback fever 52 salmoni 71 sanguineus 5, 41, 43, 44, 46, 48, 49, 213–216 saskatoon 79 Scaphixodes 5, 205, 209 scapularis 5, 33, 37, 38, 40, 41, 42, 49–52, 98, 101, 104, 106, 129, 134–140 sculptus 5, 98, 101, 104, 107, 194–198 sharp-tailed grouse 91 shrew 97, 98, 127, 137, 144, 155, 156 siculus 213 signatus 5, 96, 99, 103, 106, 209–212 skin lesions 56 skunk 84, 88, 99, 137, 169, 172, 194 snowshoe hare 84, 93 soricis 5, 97, 102, 105, 107, 152–156 southern tick-associated rash illness 38, 66 spinipalpis 5, 52, 98, 100, 104, 106, 141–144 STARI 38, 66 stigmaticus 213 streatori 176 Streator’s tree squirrel 176 Tamias amoenus 81 Tamiasciurus hudsonicus 81, 176 Tamias striatus 84 TBRF 39 texanus 5, 95, 100, 103, 108, 134, 198–201, 213 Thomomys talpoides 155 thoracicus 156 tick-borne relapsing fever 39 A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) 317

tick paralysis 1, 36, 54, 56, 66, 69, 82, 133, 140 tigrinum 66 Trichotoixodes 4, 113 triste 66 tularaemia 44, 45, 81, 87, 133, 167 tularensis 44, 45, 66, 167, 181, 242 turanicus 216 turicata 39 turicatae 39 turkey 38, 91 Tympanuchus phasianellus 91 unipunctata 64 uriae 4, 96, 99, 102, 106, 109–113 Urocitellus richardsonii 81 Ursus americanus 84 variabilis 4, 33, 43–47, 49, 50, 56, 70, 71, 80, 82–84, 87 variegatus 71 venustus 76 walkerae 55 weasel 84, 194, 204 western blacklegged tick 129, 132, 133 wheeleri 239 white-footed mouse 37, 42, 50, 51, 84, 137 white-tailed deer 40, 54, 73–75, 137 wood buffalo 73 woodchuck 169, 172 woodland caribou 73, 76 yellow-bellied marmot 80, 81, 185 yellow-pine chipmunk 81 Zapus hudsonius 84 A Handbook to the Ticks of Canada (Ixodida: Ixodidae, Argasidae) Evert E. Lindquist, Terry D. Galloway, Harvey Artsob, L. Robbin Lindsay, Michael Drebot, Heidi Wood, and Richard G. Robbins With illustrations by King Wan Wu and Barry Flahey and Maps by Tom Naughten

Despite their insignificant form and size, ticks are one of the most readily recognized groups of arthropods in Canada. They are repugnant to most people not only because of their blood-feeding habits, but some species may cause paralysis or serve as reservoirs and vectors of a remarkable variety of pathogens, including viruses, rickettsiae, bacteria, protozoans and spirochaetes. In Canada, these pathogens include the causative agents of Lyme borreliosis, relapsing fever, tularaemia, Rocky Mountain spotted fever, Q fever, Colorado tick fever, Powassan encephalitis, and perhaps others, some of which afflict wild and domesticated animals as well as humans, leading to highly pervasive medico-veterinary and economic issues. An important step is correct identification of the offending entity, which with ticks may be any of the active immature or adult forms.

This publication is an identification and information guide to all known active instars of the nine genera and forty species of ticks that occur in Canada. Identification keys and other information about ticks in Canada were last presented by John D. Gregson in a 1956 monograph which gradually became outdated. Previous keys did not account for the larva, which may be just as important as subsequent instars. Also, the distributions of ticks have changed markedly, in view of climatic changes and other factors.

Handbook topics include: general information on tick life history, external structures and terms used for identifying ticks, methods of collecting, rearing and preserving ticks, medical, veterinary and wildlife importance of ticks in Canada, illustrated keys to larvae, nymphs, females and males of genera and species, and for each species, diagnoses, summary life histories, host preferences, general distribution, disease vector potential, and maps of Canadian distribution. Host/tick and tick/host indices are also provided. This handbook will be equally useful in Alaska and the northern tier of contiguous states of the United States of America.

Front cover images: Background figure, Carios kelleyi, by Barry Flahey. Boxed figures from left to right: female American dog tick, Dermacentor variabilis (Photo by G.D. Alpert); engorged nymph of Ixodes gregsoni, a tick which infests mustelids in winter (Photo by Klaus Bolte); picking ticks from a dragcloth (Photo by K. Rochon); active stages of Ixodes scapularis, including a gravid female with eggs (Photo by J.L. Occi); a questing female Ixodes scapularis (Photo by J.L.Occi).

Cover design by Sheila Torgunrun and Donna Giberson. For information on The Biological Survey of Canada, please visit http://biologicalsurvey.ca

ISBN: 978-0-9689321-8-6 doi: DX.DOI.ORG/10.3752/9780968932186