Mississippi Mosquito Blood Meals and Haemosporidians

Mississippi State University Scholars Junction

Theses and Dissertations Theses and Dissertations

1-1-2019

Jessica Aycock

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Mississippi mosquito blood meals and haemosporidians

By TITLE PAGE Jessica Aycock

A Thesis Submitted to the Faculty of Mississippi State University in Partial Fulfillment of the Requirements for the Degree of Master of Science in Biological Sciences in the Department of Biological Sciences

Mississippi State, Mississippi

May 2019

2019

Mississippi mosquito blood meals and haemosporidians

By APPROVAL PAGE Jessica Aycock

Approved:

______Diana C. Outlaw (Major Professor)

______Jerome Goddard (Minor Professor)

______Andrea S. Varela-Stokes (Committee Member)

______Brandon Barton (Committee Member)

______Mark E. Welch (Graduate Coordinator)

______Rick Travis Dean College of Arts & Sciences

Name: Jessica Aycock ABSTRACT Date of Degree: May 3, 2019

Institution: Mississippi State University

Major Field: Biological Sciences

Major Professor: Diana C. Outlaw

Title of Study: Mississippi mosquito blood meals and haemosporidians

Pages in Study 46

Candidate for Degree of Master of Science

Mosquitoes (Culicidae) transmit several parasites and pathogens including the causative agent of malaria, haemosporidians (Haemosporida). Transmission of these agents to and from the mosquito occurs during the collection of a blood meal. Because of this, it is imperative to gather data on current feeding patterns of mosquitoes. Prior to this study there were no published data on current feeding patterns of mosquitoes in

Mississippi. Mosquitoes were captured with CDC light traps at eleven sites in two collection years. Engorged females were analyzed for blood meal content, and the vertebrate host was identified to species level in 72 mosquitoes. Previously published haemosporidian data were gathered to compare potential transmission of haemosporidians to and from the vertebrate and mosquito hosts identified in this study.

DEDICATION

I dedicate this thesis to my mother, Amy Aycock and my fiancé, Matthew Ber. I would not have made it this far without either of them. Thank you for getting me through the tears, anxiety, and doubt during this work.

ACKNOWLEDGEMENTS

I would like to first acknowledge my major professor, Dr. Diana C. Outlaw.

Without her expertise and guidance none of this would have been possible. I would also like to give a special acknowledgement to Dr. Jerome Goddard, my minor professor, committee member, and entomology mentor. If I can accomplish even half of what you have in the entomological field, I will be happy. I would like to also thank my other committee members, Dr. Andrea Varela-Stokes and Dr. Brandon Barton, for bearing through the many shifts and changes these past years. I would also like to acknowledge everyone who helped develop this project and the protocols within in it such David

Larson, Joan Kenney, and Linda Kothera. I would like to give a special thank you to

Haley Bodden for helping me through the toils of graduate school. Lastly, I would like to acknowledge Andrew Mahoney, Sarah Marshall, Elanie Briggs, Skyla Chanel, and Tate

Johnson for their assistance in this project.

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TABLE OF CONTENTS

DEDICATION ...... ii

ACKNOWLEDGEMENTS ...... iii

LIST OF TABLES ...... vi

LIST OF FIGURES ...... vii

CHAPTER

I. INTRODUCTION ...... 1

1.1 Mosquitoes ...... 1 1.1.1 Classification and Taxonomy ...... 1 1.1.2 Anatomy and Life Cycle ...... 2 1.1.3 Feeding and Vertebrate Hosts ...... 4 1.1.4 Indirect and Direct Disease ...... 6 1.2 Haemosporidians ...... 7 1.2.1 History and Expansion ...... 7 1.2.2 Life Cycle ...... 8 1.2.3 Vertebrate and Dipteran Hosts ...... 9 1.2.4 Purpose of Research ...... 11

II. METHODS ...... 14

2.1 Survey Design ...... 14 2.2 Mosquito Collection ...... 15 2.3 Mosquito Identification and Preparation ...... 15 2.4 Blood Meal Analysis ...... 16 2.5 Statistical Methods ...... 17 2.6 Haemosporidian Analysis and Review ...... 18

III. RESULTS ...... 22

3.1 Engorged Mosquitoes Captured ...... 22 3.2 Blood Meal Identification ...... 23 3.3 Haemosporidian Analysis ...... 23 3.4 Statistical Analyses ...... 24

IV. DISCUSSION ...... 30

4.1 Mosquito Capture Rate Effect ...... 30 4.2 Identified Vertebrate Hosts ...... 30 4.3 Feeding Patterns ...... 31 4.4 Haemosporidian Analysis ...... 32 4.5 Limitations ...... 35 4.6 Conclusion ...... 36

REFERENCES ...... 39

LIST OF TABLES

Table 1.1 Haemosporidian Dipteran and Vertebrate Hosts ...... 12

Table 3.1 Blood Meals by Collection Year ...... 25

Table 3.2 Blood-fed Mosquitoes by Collection Year ...... 25

Table 3.3 Initial Blood Meal Matrix for the Correspondence Analysis ...... 26

Table 3.4 Final Blood Meal Matrix for the Correspondence Analysis ...... 26

Table 3.5 Blood Meal Linear Mixed Model’s p-values ...... 27

Table 4.1 Haemosporidian Transmission within Vertebrates ...... 37

Table 4.2 Haemosporidian Transmission within Mosquitoes ...... 38

LIST OF FIGURES

Figure 1.1 General Haemosporidian Life Cycle ...... 13

Figure 2.1 Map of Collection Sites ...... 20

Figure 2.2 Sella Score ...... 21

Figure 3.1 Unimodal Distribution of Data ...... 28

Figure 3.2 Blood Meal Correspondence Analysis Biplot ...... 29

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CHAPTER I

INTRODUCTION

1.1 Mosquitoes

1.1.1 Classification and Taxonomy

Mosquitoes are dipterans belonging to the family Culicidae and are separated into three subfamilies: Toxorhynchitinae, Anophelinae, and Culicinae. This study will focus on the subfamilies Anophelinae and Culicinae, because mosquitoes belonging to

Toxorhynchitinae do not obtain a blood meal. The subfamily Anophelinae includes the genera Anopheles, Chagasia, and Bironella, but currently only Anopheles spp. are found in Mississippi. The subfamily Culicinae is more extensive and contains the most genera of the three subfamilies. The genera that are most likely to be found in Mississippi are

Aedes, Coquillettidia, Culex, Mansonia, Psorophora, Uranotaenia, and Wyeomyia

(Clements 1992; Lehane 2005; Varnado et al. 2014). Until recently, the previous list would have included the genus Ochlerotatus, which was assumed to have diverged from

Aedes. But, a study in 2015 by Wilkerson et al. showed this separation was unnecessary and advised demoting Ochlerotatus from a genus to a subgenus in Aedes (Wilkerson et al. 2015). This study bases its classification on this 2015 designation.

Both Anopheles quadrimaculatus and Anopheles crucians are complexes consisting of several species that are morphologically indistinguishable. In Mississippi, there are currently three described species within Anopheles quadrimaculatus complex

and three described species within Anopheles crucians complex (Cornel et al. 1996;

Wilkerson et al. 2004; Varnado et al. 2014).

Another group of indistinguishable mosquitoes includes Aedes tormentor and

Aedes atlanticus. As adults, these two species are close to impossible to differentiate in ideal conditions, with damage during capture making them completely indistinguishable.

In fact, many identification keys will group these two species within a single couplet

(Sither et al. 2013; Varnado et al. 2014).

1.1.2 Anatomy and Life Cycle

The gravid, female mosquito deposits her 50-500 embryonic eggs in stagnant water or in proximity to potential water sources (Horsfall 1963; Lefkovitch & Brust

1968; McDaniel & Horsfall 1963; Muirhead-Thomson 1951). These eggs contain a hardened shell which is often unique to its clade. Within a few days, the embryo develops into a larva, which will hatch from the egg in most species (Clements 1992). This stage can be found in a variety of positions in the water depending on the genus and/or species.

For example, anopheline mosquitoes are usually found on the surface of the water, while most culicine mosquitoes hang down from a breathing tube on the surface. These peculiar positions are due to the larva’s need to remain in stagnant water but to also breath air

(Keilin 1944). Besides air and an aquatic environment, the mosquito larvae require a food source. This is usually found in the form of bacteria, algae, detritus, and diatoms (Ameen

& Iversen 1978; Pucat 1965; Senior-White 1928).

The larva molts four times during one life cycle; the first three molts have little to no change in appearance, but from the last molt emerges morphological features that resemble the adult stage. After the fourth molt, the larva has developed into a pupa, 2

which remains near the water’s surface during its development. This stage retains buoyancy with an air bubble between its appendages and connects its thorax with the surface of the water. While in the pupal cuticle, many of the adult organs replace their larval predecessors, so the adult is ready to emerge after this stage. The pupa rises to the surface where it rests for emergence (Romoser & Nasci 1979).

The adult takes flight from the water once it emerges. During this stage, mosquitoes focus on feeding and mating. Because the female can fertilize several batches of eggs with previous sperm that she has stored, the male immediately inseminates her, whether or not she has already been inseminated (Bullini et al. 1976). After mating, the female feeds, develops her eggs, and searches for an appropriate habitat for her offspring

(Clements 1992).

Adult mosquitoes are small, two-winged, flies with long slender bodies and long legs. Like all other insects, mosquitoes have three body segments: a head, a thorax, and an abdomen. The head of the mosquito is the center of the sensory organs and contains two eyes, a long proboscis, paired antennae, and paired palps (Carpenter 1955). The eyes of the mosquito are large and compound (Land et al. 1997). The proboscis is an elongated organ found between the palps. It contains the mouthparts within a protective sheath, the labrum (Waldbauer 1962).

The thorax of a mosquito is the center of locomotive control, because it contains the legs and the wings. Each mosquito technically has four wings, but the hind pair, termed halteres, are knob-like organs for locomotion and balance. The fore wings are long and contain “veins”, which are long thickenings for structure. Along these veins are

scales of different color and size depending on the genus. Along the edge of each wing there are small, hair-like fibers called setae (Harbach & Knight 1980)

The dorsal portion of the thorax is covered by a protective scutum (Harbach &

Knight 1980). This is usually a distinguishing feature between the species of mosquitoes within a genus, particularly Aedes spp. as it may be covered in light and dark scales that portray a distinctive pattern (Burkett-Cadena 2013; Varnado et al. 2014). Along the lateral side of the thorax, lies the pleuron which contains two sclerites and spiracles. The sclerites are large exoskeletal plates for protection, and the spiracles are openings for respiration. In addition to the legs and wings, the thorax contains the salivary glands. This organ is of particular importance, because it produces the saliva that may contain infective parasites, such as haemosporidians (Snodgrass 1959).

The abdomen is primarily where reproduction and digestion/excretion occur. The abdomen contains 10 segments each with a tergite (the dorsal plate) and a sternite (the ventral plate). The connections between the plates allow for expansion during feeding and egg production. The end of the abdomen has two cerci for reproduction, specifically copulation and egg laying. The inner anatomy of the segment consists primarily of the alimentary canal, the full pathway for the passage of food. The passage begins at the pharyngeal pump, ends at the rectum, and is separated into the foregut, midgut, and hindgut (Snodgrass 1959).

1.1.3 Feeding and Vertebrate Hosts

Male and female mosquitoes feed on nectar from plants for sustenance, while female mosquitoes may also obtain energy from a blood meal (Kevan & Baker 1983; 4

Nayer & Sauerman 1975). Female anophelines and culicines must obtain protein for egg production by consuming the blood from a host (Kogan 1990). The female’s peg sensilla on the maxillary palps alert her of a potential source of blood by detecting carbon dioxide and host odor. In response to these cues, the female flies toward the source (Daykin et al.

1965; Eiras & Jepson 1991).

Carbon dioxide is a common activator, the first stimulant that directs the female mosquito to her host. Other odors emitted by the host have shown to also activate mosquitoes. Different combinations of carbon dioxide, lactic acid, and host odor have shown varied results in activation on different target mosquito species (Daykin et al.

1965; Eiras & Jepson 1991). Several experiments have even used different host odors

(i.e. human vs cow) and have yielded varied results. This demonstrates how preferential feeding could be based on the mosquito species and not just opportunity (Dekker &

Takken 1998).

Once she has landed on the vertebrate, the female will undergo four stages of the blood feeding: exploratory phase, probing phase, imbibing phase, and withdrawal phase

(Clements 1992). During the exploratory phase, the female lands on the host and uses chemosensillar on her labella, labrum, and tarsi to verify a proper food source. The female begins to insert her stylet into the skin of the host leading to the probing phase.

This includes the first penetration of the skin, blood movement to the mosquito’s mouthparts, and injection of saliva into the host. Next, the imbibing phase is where the mosquito begins to withdraw the blood. Lastly, the withdrawal phase is where the female straightens her forelegs and retracts the stylets from the host (Clements 1992; Jones &

Pilitt 1973).

If undisturbed, the female feeds until fully engorged. Once the blood reaches the midgut, the erythrocytes begin to form into a clot. The blood is utilized by breaking down the proteins to build a proteinaceous yolk which will form in the oocyte. Twenty-four hours after the blood meal is taken, the outer erythrocytes have been digested and turn brown or black in color. The innermost cells within the clot are the last erythrocytes to be digested usually two to three days following feeding (Graf & Briegel 1982).

Mosquitoes will feed on any carbon dioxide-producing organism if given the chance. Despite this, when collecting a blood meal, mosquitoes exhibit host preference.

This is defined as the tendency for a parasite to frequently infect or infest a host or group of hosts. This does not imply the mosquito has a particular idea of what it wants to feed on but instead suggests there may be variables that enhance the probability of a mosquito feeding on the host (Bruce-Chwatt & Gockel 1960; McClelland & Weitz 1963).

One feeding preference found in mosquitoes is based on host location.

Endophagic mosquitoes tend to feed on hosts within houses and animal shelters while exophagic mosquitoes often feed on hosts outside of infrastructure. This may cause feeding patterns due to host availability in each environment. Another abiotic factor that affects a mosquito’s preference is its circadian rhythm. Circadian rhythm is the physiological pattern of an organism within 24 hours. A mosquito primarily active during the daytime may encounter different vertebrates than a nocturnal mosquito (Clements

1992).

1.1.4 Indirect and Direct Disease

Mosquitoes are known to transmit a large variety of pathogens. In addition to haemosporidians, these dipterans transmit filarial worms and many viruses. The filarial 6

worms transmitted by mosquitoes include Wuchereria bancrofti, Brugia malayi, and B. timori all of which are causative agents of elephantiasis in humans. Mosquitoes also transmit the parasitic worms Dirofilaria immitis and D. repens which cause heartworm disease in cats and dogs as well as humans occasionally (Lehane 2005).

Mosquitoes transmit over 200 viruses to their vertebrate hosts. Many of these viruses affect humans including Zika, dengue, yellow fever, and West Nile. Many of these mosquito-borne viruses cause encephalitis in humans and other animals (Lehane

2005; Turell et al. 2001; Marcondes et al. 2015).

In addition, mosquito saliva contains antigens, which are common irritants once injected into the host’s blood. Once a host is exposed to the saliva, they are sensitized to the antigens and will react with either a Type I, Type II, Type III, or Type IV reaction. If the host is exposed to the antigens in the saliva frequently overtime, they will pass through different stages of reaction until they demonstrate non-reactivity (Lengy & Gold

1966; Oka 1989).

1.2 Haemosporidians

1.2.1 History and Expansion

Malaria is a well-known disease caused by haemosporidians in the phylum

Apicomplexa, order Haemosporida. These parasites are transmitted to and from many vertebrates, including humans, by a dipteran host (Perkins 2014; Garnham 1966). The occurrence of these parasites was first recorded in Ancient Egypt in 1550 B.C. as a written diagnosis of distinguishable symptoms of human malaria. The disease itself was not discovered until 1775 through a temporary cure using cinchona bark. The causative

agents, haemosporidians, were not discovered until Laveran examined the parasite through different stages of its life cycle in 1878 using microscopy (Garnham 1966).

Today, haemosporidians have been found all around the world in different environments. The genera of these parasites circumscribe a more informative map as mammalian haemosporidians are primarily located in Asia, Africa, and America, and avian and reptilian haemosporidians are located globally (Valkiunas 2005; Garnham

1966). There are currently 15 extant, known haemosporidian genera. However, the classification of these parasites is undergoing major changes as new genera and species are discovered. The following genera are the most well-studied within haemosporidian research: Plasmodium, Hepatocystis, Polychromophilus, Haemoproteus,

Parahaemoproteus, and Leucocytozoon (Perkins 2014).

1.2.2 Life Cycle

During its life cycle, a haemosporidian infects two hosts: a vertebrate and a dipteran (See Figure 1.1). While inside the vertebrate, the parasite undergoes asexual reproduction, and therefore this host is labelled the intermediate host. Once in the blood of the intermediate host, haemosporidian sporozoites develop into agamic stages, which in turn begin asexual reproduction in tissue cells (Huff 1951). The term for these stages is either exoerythrocytic meronts or schizonts. Because of this division, a dispersing, asexual stage termed uninuclear merozoite is formed. This replication process may occur several times within the host, which causes a gradual increase of the parasite while allowing an adjustment period for the host (Boyd 1935). After several generations, sexual-reproduction-capable stages develop. These stages are gametocytes and are capable of producing the macrogametocytes or microgametocytes (Davey 1946). 8

While inside the dipteran, haemosporidians undergo sexual reproduction, and therefore this host is the definitive host. When a dipteran takes an infected blood meal from a vertebrate, it ingests the aforementioned merozoites. These will develop into microgametes once inside the midgut, which will then proceed to find a macrogamete for fertilization and development of an ookinete. The ookinete travels and breaks through the midgut wall where it develops into an oocyst. This stage’s nucleus divides continuously, forming several sporoblasts, which also divide their nuclei repeatedly to develop sporozoites. The sporozoites break free and migrate to the salivary glands, where they will remain until the dipteran collects another blood meal (Terzakis et al. 1966).

The above life cycle description is a general depiction and varies between genera.

There are many other differences between the genera of haemosporidians, but these can be misleading (see Borner et al. 2016) due to intraspecies variability and interspecies similarities, and therefore DNA sequence data are most often used to reconstruct the evolutionary history (i.e. phylogenetic tree) of these organisms (Borner et al. 2016).

1.2.3 Vertebrate and Dipteran Hosts

Differences between genera are simplified by comparing the various hosts they infect (see Table 1.1). Haemosporidians are considered host-specific, because they must be transmitted to competent dipteran and vertebrate hosts. Hosts are labelled competent or incompetent in relation to the parasite’s ability to complete its life cycle within an organism (Lehane 2005).

Currently, Plasmodium spp. are the only known haemosporidians transmitted by mosquitoes (Garnham 1966). A majority of these natural hosts are Anopheles spp.; however, there have been several cases where various stages of Plasmodium spp. have 9

been detected in other mosquito species such Culex spp. and Aedes spp. with microscopy and molecular methods (Falk et al. 2015; Klein et al. 1987; Valkiunas 2005). In addition, other haemosporidian genera have been found within mosquitoes using molecular methods (Larson et al. 2017). This does not necessarily imply that these mosquitoes could have transmitted the haemosporidian, but simply shows these mosquitoes acquired these parasites from their vertebrate hosts.

Plasmodium spp. are the most versatile haemosporidians in relation to the vertebrate host. Depending on the species, these haemosporidians may be successful in mammals, birds, or reptiles. These species are also the only know haemosporidians to utilize humans as a natural host. Because of this, Plasmodium spp. are the most well- studied of all haemosporidians, particularly the species which are commonly found in humans (Garnham 1966).

Though other haemosporidian genera are not medically relevant, several of the species may cause either ecological or economical loss. For example, Leucocytozoon spp. are only successful in birds and are commonly transmitted by black flies (Simuliidae).

These parasites are the causative agent of leucocytozoonosis, a disease when left unchecked could result in large economic loss within the poultry industry (Lee et al.

2016). Haemoproteus spp. and Parahaemoproteus spp. are also parasites which affect avian fauna. These parasites are transmitted by louse flies (Hippoboscidae) and biting midges (Ceratopogonidae) respectively (Perkins 2014).

There are many other genera of haemosporidians than the few listed above.

Included in these are haemosporidians which affect other mammals such as bats and non- anthropogenic apes. There are also genera which exclusively affect lizards (Garnham

1966). These haemosporidians are all understudied when compared to the aforementioned genera therefore there is very little data available for these species.

1.2.4 Purpose of Research

Though there has been great progress in the understanding of haemosporidian transmission patterns, there are still many unanswered questions. Previous haemosporidian work in Mississippi saw mosquito acquisition of a previously unknown haemosporidian. Many of these mosquitoes were identified as Culex erraticus, a common mosquito within Mississippi (Larson et al. 2017). In addition, a similar haemosporidian was identified in a survey of the United States. These parasites were identified primarily from white-tailed deer (Martinsen et al. 2016). The transmission pathways and host susceptibility for these cryptic haemosporidians is unclear.

The purpose of this research was to uncover the relationship between the dipteran and vertebrate hosts of haemosporidians in Mississippi. This identified potential transmission pathways for haemosporidians, including the aforementioned previously unknown species. To determine these relationships, we identified blood meals within engorged mosquitoes captured in two surveys: Larson et al.’s survey and a replicate survey of Mississippi (Larson et al. 2017). We also identified any potential feeding patterns of captured mosquitoes. Feeding patterns would allow continuous acquisition of a particular haemosporidian as we see in both Larson et al. and Martinsen et al. (Larson et al. 2017; Martinsen et al. 2016).

Table 1.1 Haemosporidian Dipteran and Vertebrate Hosts

Genus Vertebrate Dipteran Plasmodium Mammals, Birds, Reptiles Culicidae Hepatocystis Mammals Ceratopogonidae Polychromophilus Bats Nycteribiidae Haemoproteus Birds Hippoboscidae Parahaemoproteus Birds Ceratopogonidae Leucocytozoon Birds Simuliidae List of well-studied genera with their vertebrate and dipteran hosts (Perkins 2014; Valkiunas 2005; Garnham 1966)

Figure 1.1 General Haemosporidian Life Cycle

A representative life cycle of an avian Plasmodium sp. (Cole & Friend 1999)

CHAPTER II

METHODS

2.1 Survey Design

Two sampling periods were analyzed: 2013 and 2017. The two sampling periods were identical in collection technique, assay methodology, and data analysis. For each, we identified the mosquito, the vertebrate blood meal, and any infecting haemosporidians within the mosquito. The mosquito and haemosporidian data from the 2013 sampling period were collected previously (Larson et al. 2017). However, the vertebrate hosts had yet to be identified in those mosquitoes. Due to the small sample size of the 2013 collection, mosquitoes were trapped in a second collection period in 2017.

Mosquitoes were captured at the same eleven sites in both the 2013 and 2017 collection periods (see Figure 2.1): Bienville Wildlife Management Area (WMA),

Buccaneer State Park, Caney Creek WMA, Chickasawhay WMA, Legion State Park,

Malmaison WMA, Old River WMA, Pearl River WMA, Stoneville WMA, Upper Sardis

WMA, and Noxubee US National Wildlife Refuge. These sites were randomly selected using ArcGIS® version 10.3 (ESRI 2011.; Larson et al. 2017). The wildlife management area closest to each point was used as a site.

In addition, a random point was selected on the wildlife management area’s map

(Larson et al. 2017). When collecting, traps were placed as close as possible to the random point, though often it was impossible to place the traps directly at the specific

point. For example, the point placed in Noxubee US National Wildlife Refuge was flooded due to heavy rainfall and thus was inadequate for a battery-operated trap.

2.2 Mosquito Collection

Both collection periods used Centers for Disease Control (CDC) Miniature Light

Traps (Sudia & Chamberlain 1962), powered by a six-volt, ten ampere-hour rechargeable, lead-acid battery. For the 2013 collection, three traps were used, while only two traps were used for the 2017 collection due to availability. Each trap was supplemented with approximately 1 kg of dry ice within an insulated, spouted cooler.

Each trap was suspended from either a 64-inch metal garden hook or a tree branch.

Batteries were covered with plastic bags and placed securely on the ground to prevent the clamps from detaching from the battery. Dry ice coolers were suspended in close range to each trap (Larson et al. 2017).

Traps were left over the course of two nights, with samples collected at least every twelve hours, however some sites required more frequent collecting due to a greater number of captured mosquitoes. In addition, mosquitoes were captured at each site with a sweep net and/or an aspirator. The sweep net and aspirator were used for approximately thirty minutes at each site. The 2013 period lasted from 3 July to 8

September (Larson et al. 2017); the 2017 period lasted from 26 June to 4 September.

2.3 Mosquito Identification and Preparation

Blood-fed females were identified based on the presence of blood within the abdomen. Slightly engorged females were also included, as incomplete blood meals often yielded clean sequences of the vertebrate host. Each blood meal was scaled on the Sella

score (see Figure 2.2) which identifies the digestion stage of the blood meal. The females were then identified to species level using two keys: “Identification Guide to Adult

Mosquitoes in Mississippi” and “Mosquitoes of the Southeastern United States”. Due to species similarities, some mosquitoes cannot be identified morphologically (Burkett-

Cadena 2013; Varnado et al. 2014). This survey classified all Aedes atlanticus and A. tormentor as one joined species. In addition, all species within the Anopheles quadrimaculatus complex and A. crucians complex were identified as such.

Mosquitoes were identified using several characteristics found within the aforementioned keys. These features included scale coloration on legs, abdomen, scutum, proboscis, and head; palp length; and setae location (Burkett-Cadena 2013; Varnado et al.

2014). These features were often used in combination to determine the species. Reference species were kept separate and point mounted for confirmation of further specimens.

Identified females were separated into two samples: a thorax and an abdomen and kept at -80 ˚C. Conducting a blood meal analysis on the abdomen alone prevents excess interference of mosquito DNA during the polymerase chain reaction. In addition, it supplements haemosporidian analyses to potentially differentiate acquisition time of the parasite.

2.4 Blood Meal Analysis

DNA was extracted from each body segment separately with a DNeasy Blood and

Tissue kit using the protocol for arthropod samples. All samples were homogenized with a Fisherbrand™ RNase-Free Disposable Pellet Pestle. Samples were eluted in 50 µl

Buffer AE for the 2013 samples and 200 µl Buffer AE for the 2017 samples. The elution

amount was reduced for the 2013 collection due to DNA degradation from improper, initial storage.

The vertebrate mitochondrial cytochrome b (cyt b) gene was amplified with a polymerase chain reaction. The cyt b gene was used due its success in previous research.

The amplification required the primers H15149[5’-

GCCCCTCAGAATGATATTTGTCCTCA-3’] and L14841[5’-

CCATCCAACATCTCAGCATGATGAAA-3’] with the following thermocycler parameters: 93˚C for 5 minutes; a duration of 93 ˚C for 1 minute, 50 ˚C for 1 minute, and

72 ˚C for 2 minutes for 35 cycles; 72 ˚C for 10 minutes; and held at 4 ˚C (Kent 2009;

Kocher et al. 1989).

Vertebrate DNA presence in the polymerase chain reaction product was tested on a 1.5% agarose gel electrophoresis. Positives were recorded from the gel under UV light.

All positives were cleaned with the DNeasy PCR Purification kit or DNeasy Gel

Extraction kit if double bands were present. Purified PCR product was sequenced with

Sanger sequencing at Arizona State University and sequences were trimmed and edited using Sequencher 5.4.6. (Gene Codes Corporation). Vertebrate species were identified with a BLAST search of the NCBI database (Altschul et al. 1990).

2.5 Statistical Methods

A correspondence analysis was conducted on the contingency table to determine the distance between the two variables: blood meal identity and mosquito species

(McCune & Grace 2002; Paliy & Shankar 2016). The hypothesis tested in the correspondence analysis was that there will be significantly shorter distances between the mosquito species and their hosts. These distances were based on calculations of variance 17

in the relationships between mosquito species and vertebrate host. The null hypothesis was that there would be no significant difference in distance between any mosquito species and host. The correspondence analysis was conducted and summarized in R using the following packages: "FactoMineR", "devtools", "factoextra", "gplots", and "corrplot"

(Kassambara 2015, Kassambara & Mundt 2017, Le et al. 2008; R Core Team 2014;

Warnes et al. 2016; Wei & Simko 2017; Wickham et al. 2018).

In addition, four mosquito species were selected based on larger sample size for further analyses. The mosquito/host relationships were tested individually with a linear mixed model which included a response variable of the number of mosquitoes of each species that had fed on a specified host (i.e. white-tailed deer), two explanatory variables of mosquito species and collection site, and a random effect of year of collection (Munoz et al. 2012).

The hypothesis tested in the models was that collection site and mosquito species will have a significant effect on the vertebrate host identified. The null hypothesis was there would be no significant effect on the vertebrate host by the collection site and mosquito species. Because we did four comparisons, the p-value to reject the null hypothesis must be < 0.0125 (0.05/4). The model was constructed and summarized with the “lme4” package in R (Bates et al. 2015; R Core Team 2014).

2.6 Haemosporidian Analysis and Review

The haemosporidian cyt b gene was amplified with the previous methodology using PerkinsF [5’-TAATGCCTAGACGTATTCCTGATTATCCAG-3’] and PerkinsR

[5’-TGTTTGCTTGGGAGCTGTAATCATAATGTG-3’] primers in an initial amplification using the following thermocycler parameters: 94˚C for 4 minutes; 40 cycles 18

of 94˚C for 20 seconds, 55˚C for 20 seconds, and 68˚C for 1 minute; a prolongation of

68˚C for 10 minutes; and held at 4˚C. The polymerase chain reaction product was then reamplified with primers HaemF [5’-ATGGTGCTTTCGATATATGCATG-3’] and

HaemR2 [5’-GCATTATCTGGATGTGATAATGGT-3’] with the following thermocycler parameters: 94˚C for 3 minutes; 40 cycles of 94˚C for 30 seconds, 51˚C for

30 seconds, and 72˚C for 45 seconds; a prolongation of 72˚C for 10 minutes; and held at

4˚C (Perkins et al. 2002; Perkins et al. 2007; Bensch et al. 2000). The abdomen and thorax were analyzed separately.

Previously reported data on current, haemosporidian transmission patterns were compiled for all blood-fed mosquito species. These transmission rates were compared to the identified feeding patterns from this survey as well as all other blood meal identities which may be due to random occurrence. A literature review was conducted with Google

Scholar and searched for articles with terms such as “sporogony”, “transmission”, and

“gametocytes”. Due to a deficiency in local research, transmission ability of the vertebrate within the same genus was also considered in the review.

Figure 2.1 Map of Collection Sites

A map of all the sites for the 2013 and 2017 collections taken from Larson et al. (2017)

Figure 2.2 Sella Score

A scale of the blood meal/egg ratio in the female mosquito’s abdomen with 1 as an unfed female and 2 as a fully engorged female. Ascending numbers represent blood meal digestion until 7 which represents a gravid female (taken from Detinova 1962)

CHAPTER III

RESULTS

3.1 Engorged Mosquitoes Captured

Of the 151 blood-fed samples from the 2013 collection, forty blood meals were identified: thirty-eight to species level and two to genus level (see Table 3.1). The identified blood meals were found in five mosquito species: Aedes atlanticus/tormentor,

Aedes vexans, Anopheles crucians complex, Anopheles quadrimaculatus complex, and

Culex erraticus (see Table 3.2).

In the 2017 collection, 32 blood meals were identified from 148 engorged females

(see Table 3.1). These blood meals were identified from Aedes canadensis, Aedes vexans,

Anopheles crucians complex, Anopheles quadrimaculatus complex, Coquillettidia perturbans, Culex erraticus, Culex nigripalpus, Psorophora mathesoni, and Uranotaenia sapphirina (see Table 3.2).

There was a large decrease in sample size in both collections. This is largely due to the misidentification of gravid mosquitoes as engorged mosquitoes. It is difficult to differentiate between the two especially for the mosquitoes with darker coloration. In addition, some mosquitoes were identified as partially engorged. Some of these samples may have been mosquitoes which recently released eggs. These mosquitoes would most likely have a slightly enlarged abdomen similar to partially-fed females.

3.2 Blood Meal Identification

The majority of blood meals from the 2013 collection were identified as white- tailed deer with a few samples identified as human, Eastern gray squirrel, Northern grey- headed sparrow, Eastern screech owl, North American beaver, and Northern cardinal.

The two blood-meals identified to genus level were classified as a sparrow and a wood owl. These identities were determined based on the DNA sequence and the geographic location of organisms within the identified genus. A majority of the blood meals identified as white-tailed deer were found within Culex erraticus.

For the 2017 collection, the majority of blood meals were identified as white- tailed deer, Eastern gray squirrel, or Coahuilan box turtle. All Eastern gray squirrel and

Coahuilan box turtle blood meals were sampled from Psorophora mathesoni, while all white-tailed deer blood meals were found in either Culex erraticus or Aedes vexans.

There were also blood meals identified as great blue heron, human, Cape rock thrush, wild boar, swamp rabbit, common box turtle, and Mexican box turtle (see Table 3.1).

3.3 Haemosporidian Analysis

Haemosporidian DNA was only found in two samples. From the 2013 collection,

Plasmodium sp. DNA was detected from the abdomen of a Culex erraticus.

Unfortunately, no vertebrate blood was detected from the abdomen. From the 2017 collection, a Leucocytozoon sp. was detected within the blood-fed abdomen of a

Coquillettidia perturbans. The blood meal within the abdomen was identified as great blue heron.

3.4 Statistical Analyses

Samples were first divided by mosquito species then by blood meal identity, resulting in sixteen different blood meal identities among ten mosquito species (Table

3.3). However, after plotting this ordination, outliers were identified and removed from the data. These outliers were instances of a single sample of a mosquito species and/or blood meal identity and caused the remaining data to gather in one quadrant of the plot.

Once removed, there were five mosquito species and eleven blood meal identities in the matrix (Table 3.4). One requirement for correspondence analyses is the data must be in a unimodal distribution, which was met by the aforementioned contingency table (Figure

3.1). The correspondence analysis resulted in a correlation coefficient of 1.107245 and a chi-square p-value < 0.0005, thus we reject the null hypothesis that there is not variation in the distances between the mosquito species and blood meal identities (Figure 3.2).

The model equation tested the association between the number of an identified blood meal with the mosquito species and the collection site with the collection year as a random effect. The null hypothesis tested the association between the number of an identified blood meal with the collection year as a random effect. The selected four blood meal identities were subjected to the model separately and were tested with an ANOVA with p-values between 0.1061 and 0.4382 (Table 3.5)(R Core Team 2014). None of the p- values were < 0.0125, so we accept the null hypothesis that mosquito identity and site have no effect on the blood meal identity.

Table 3.1 Blood Meals by Collection Year

2013 Collection 2017 Collection American beaver Coahuilan box turtle Eastern gray squirrel* Common box turtle Eastern screech owl Eastern gray squirrel* Eurasian tree sparrow Great blue heron Human* Human* Northern cardinal Mexican box turtle Northern grey-headed Swamp rabbit sparrow White-tailed deer* White-tailed deer* Wood owl Wild boar List of vertebrate species identified from the 2013 and 2017 collections. *denotes vertebrates found within both collection years.

Table 3.2 Blood-fed Mosquitoes by Collection Year

2013 Collection 2017 Collection Aedes atlanticus/tormentor Aedes canadensis Aedes vexans* Aedes vexans* Anopheles crucians* Anopheles crucians* Anopheles Anopheles quadrimaculatus* quadrimaculatus * Culex erraticus* Coquillettidia perturbans Culex erraticus * Culex nigripalpus Psorophora mathesoni Uranotaenia sapphirine List of blood-fed mosquito species identified from the 2013 and 2017 collections. *denotes mosquitoes found within both collection years

Table 3.3 Initial Blood Meal Matrix for the Correspondence Analysis

ABE CRT CBT CXT EGS ESO ETP GBH HUM MBT NCA NGS SRA WTD WBO WOW AQ 0 0 0 0 0 0 0 0 1 1 0 0 0 0 0 0 AC 0 0 0 0 0 0 0 0 0 0 0 1 1 0 0 0 AV 0 0 0 1 0 0 1 0 1 0 0 0 0 8 0 0 AE 0 0 0 0 0 0 0 0 0 0 0 0 0 0 1 0 AA 0 0 0 0 1 0 0 0 0 0 0 0 0 1 0 0 CE 1 1 0 0 0 1 0 0 2 0 1 0 0 26 0 1 CN 0 1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 CP 0 0 0 0 0 0 0 1 0 0 0 0 0 0 0 0 PM 0 0 7 0 8 0 0 0 2 0 0 0 0 0 0 0 US 0 1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Data matrix of blood meal identities among the different mosquito species before outliers were removed. ABE=American beaver, CRT=Cape Rock thrush, CBT=Coahuilan box turtle, CXT=common box turtle, EGS=Eastern gray squirrel, ESO=Eastern screech owl, ETP=Eurasian tree sparrow, GBH=great blue heron, HUM=human, MBT=Mexican box turtle, NCA=northern cardinal, NGS=Northern grey-headed sparrow, SRA=swamp rabbit, WTD=white-tailed deer, WBO=wild boar, WOW=wood owl, AQ= Anopheles quadrimaculatus, AC= Anopheles crucians, AV= Aedes vexans, AE= Aedes canadensis, AA= Aedes atlanticus/tormentor, CE= Culex erraticus, CN= Culex nigripalpus, CP= Coquillettidia perturbans, PM= Psorophora mathesoni, US= Uranotaenia sapphirina

Table 3.4 Final Blood Meal Matrix for the Correspondence Analysis

ABE CRT CBT CXT EGS ESO ETP HUM NCA WTD WOW

AQ 0 0 0 0 0 0 0 1 0 0 0 AV 0 0 0 1 0 0 1 1 0 8 0 AA 0 0 0 0 1 0 0 0 0 1 0 CE 1 1 0 0 0 1 0 2 1 26 1 PM 0 0 7 0 8 0 0 2 0 0 0 Data matrix of blood meal identities among the different mosquito species after outliers were removed. ABE=American beaver, CRT=Cape Rock thrush, CBT=Coahuilan box turtle, CXT=common box turtle, EGS=Eastern gray squirrel, ESO=Eastern screech owl, ETP=Eurasian tree sparrow, HUM=human, NCA=northern cardinal, WTD=white-tailed deer, WOW=wood owl, AQ= Anopheles quadrimaculatus, AV= Aedes vexans, AA= Aedes atlanticus/tormentor, CE= Culex erraticus, PM= Psorophora mathesoni

Table 3.5 Blood Meal Linear Mixed Model’s p-values

Blood Meal Identity ANOVA p-value Coahuilan box turtle 0.3665 Eastern gray squirrel 0.4065 Human 0.4382 White-tailed deer 0.1061 Table of the four selected blood meal identities and their subsequent p-values calculated from an ANOVA between the models of the hypothesis and the null hypothesis.

Figure 3.1 Unimodal Distribution of Data

Distribution of the blood meal occurrences which forms a unimodal pattern

CXT ETP

AV HUM

WTD PM AA CE CBT EGS WOW CRT

NCA ABE ESO

Figure 3.2 Blood Meal Correspondence Analysis Biplot

Plotting of correspondence analysis of blood meal identities and their mosquito counterparts. Red text and arrows represent mosquito samples, while blue text and lines represent vertebrate hosts. ABE=American beaver, CRT=Cape Rock thrush, CBT=Coahuilan box turtle, CXT=common box turtle, EGS=Eastern gray squirrel, ESO=Eastern screech owl, ETP=Eurasian tree sparrow, HUM=human, NCA=northern cardinal, WTD=white-tailed deer, WOW=wood owl, AQ= Anopheles quadrimaculatus, AV= Aedes vexans, AA= Aedes atlanticus/tormentor, CE= Culex erraticus, PM= Psorophora mathesoni

CHAPTER IV

DISCUSSION

4.1 Mosquito Capture Rate Effect

There are several factors that may have affected these data. First, in the 2017 collection, many of the mosquitoes were identified as Psorophora mathesoni. All of these blood-fed mosquitoes were captured in one location, Old River WMA: It is possible that biotic and abiotic factors could have increased the number of blood-fed mosquitoes captured at this site which would create an abundance of the mosquito species within the collection.

In addition, there are several instances where only one vertebrate species was identified in one mosquito species at one collection site. This created a proportion of 1 for that feeding pattern, even though there was only a single occurrence. These examples represent the aforementioned outliers and should be evaluated with caution.

4.2 Identified Vertebrate Hosts

In the 2013 collection, the majority of mosquitoes throughout all sites were identified as Culex erraticus. Despite being found in many different locations, these mosquitoes consistently fed on white-tailed deer. The Culex erraticus mosquitoes from the 2017 collection also consistently fed on white-tailed deer. This recurring relationship supports that the novel haemosporidians identified in Larson et al. and Martinsen et al.

may be the same species (Larson et al. 2017; Martinsen et al. 2016). It is highly possible

Culex erraticus is acquiring this haemosporidian from the natural host, white-tailed deer.

In addition to white-tailed deer, we identified several different vertebrates from the mosquito blood meals. Most of these vertebrates were mammals and birds that are commonly found in Mississippi such as the Northern cardinal and the Eastern gray squirrel (www.iucnredlist.org). However, there were several identifications that are not known in Mississippi or even the United States. For example, the Coahuilan box turtle is an endangered species found in Mexico (www.iucnredlist.org). Other examples of distant blood meal identities are the Cape Rock thrush and the Northern grey-headed sparrow

(www.iucnredlist.org).

4.3 Feeding Patterns

According to the statistical analysis of the blood meal data, the detection of vertebrate hosts was not randomly distributed among mosquito species. First, according to the chi-square statistic p-value < 0.05 and the correlation coefficient > 0, there is a positive association between the two variables, mosquito species and vertebrate host, within the correspondence analysis. In addition, the ordination revealed several outliers within our data. The ordination segregated the mosquito/host relationships which showed an unlikely 1:1 ratio. These perfect ratios were achieved simply due to the extremely low sample size of the mosquito species or blood meal identities, and thus do not properly represent the mosquito/host community within the collection sites.

With the outliers excluded, the plot greatly supported the alternative hypothesis.

Overall, all points representing vertebrates aggregated closer to one or two mosquito species (Figure 3.2). For instance, the Coahuilan box turtle (CBT) and Eastern gray 31

squirrel (EGS) points aggregate extremely closely to Psorophora mathesoni (PM). In fact, these two vertebrate species occupy the same space within the plot. There is a similar occurrence with the many vertebrate points surrounding Culex erraticus (CE).

This means the points have the same variance distance from the mosquito point. They are most closely associated with that mosquito in comparison to the other mosquitoes on the plot.

The vertebrate points occupying the same space were blood meal identities only found within C. erraticus. Instead, the white-tailed deer point (WTD) is plotted between

Culex erraticus and Aedes vexans (AV). This is because white-tailed deer were identified from both of these mosquitoes. The WTD/CE distance is much shorter than the WTD/AV distance. This is due to more of the white-tailed deer samples being identified from C. erraticus than from A. vexans.

Based on the p-values for the linear mixed models, there was no significant effect on the vertebrate identity from the mosquito species and collection site. One explanation for the huge difference between the p-values from the models and correspondence analysis is the representation of the data. The models required each vertebrate be analyzed separately, while the ordination analyzed all data within a single analysis. A single data matrix and analysis may result in a more definitive statistical test.

4.4 Haemosporidian Analysis

Although we did not collect enough haemosporidian data to analyze statistically, we can draw a few conclusions from our two positives. The Plasmodium sp. within the

Culex erraticus in the 2013 collection was not a surprise as mosquitoes are the natural host of this haemosporidian genus (Perkins 2014; Valkiunas 2005; Garnham 1966). 32

However, this Plasmodium spp. is successful in avian species, and due to the large, genetic separation between the avian and mammalian Plasmodium spp. (Borner et al.

2016), we can assume this mosquito had collected the infected blood meal from a bird.

The Leucocytozoon sp. found within the Coquillettidia perturbans was more profound. As stated previously, the only known dipteran host of Leucocytozoon spp. is the black fly (Perkins 2014; Valkiunas 2005; Garnham 1966). Unlike the Culex erraticus, we were able to determine the blood meal, a great blue heron (Ardea herodias), which has been shown to be susceptible to Leucocytozoon spp. (Hsu et al. 1973, Valkiunas

2005). This information and the fact that the haemosporidian DNA was only present in the abdomen, demonstrate the blood meal was most likely infected, not the mosquito.

Despite this, these data show the transmission of a novel parasite to this mosquito, although the parasite may not have been able to reproduce.

The haemosporidian literature review uncovered several connections between the mosquito/vertebrate relationships and the haemosporidians they may transmit (see Table

4.1 and 4.2). Of all the mosquito species captured, Anopheles quadrimaculatus currently transmits the most haemosporidian species, including the agents of human malaria.

However, A. quadrimaculatus has been shown to possess transmission ability of agents of avian malaria including Plasmodium gallinaceum (Garnham 1966; Santiago-Alarcon et al. 2012).

There were only two mosquito species from our collection which currently do not show any transmission ability of haemosporidians: Aedes atlanticus/tormentor and

Psorophora mathesoni. Despite this Aedes atlanticus/tormentor has transmission ability of several viruses (Turell et al. 2013; Watts et al. 1982). Psorophora mathesoni is

currently not known to transmit any disease agents. In addition, only one of the captured mosquito species shows transmission ability of a haemosporidian outside of the

Plasmodium genus. Aedes vexans may transmit Haemoproteus spp. This revelation is due to the discovery of sporozoites within mosquitoes infected with Haemoproteus spp., but this has only been possible in experimental specimens (Santiago-Alarcon et al. 2012).

Most of the vertebrates identified from the blood meals have shown transmission ability of at least one haemosporidian species. The exceptions are Coahuilan box turtle,

Cape rock thrush, common box turtle, Mexican box turtle, North American beaver, and wild boar. It should be noted the parasitology of many of these vertebrates is understudied. For example, there is very little molecular work on haemosporidians within turtles. Usually reptilian haemosporidian studies focus on lizard parasites, neglecting other reptiles such as turtles.

As predicted, there were very little data on the identified vertebrates, particularly the local mammals. A majority of the associations were between the haemosporidian and a close relative of the identified vertebrate. The literature review also revealed a majority of Plasmodium transmission across all identified vertebrates (Garnham 1966; Martinsen et al. 2016; Shurulinkov et al. 2003; Tavernier et al. 2005; Valkiunas 2005). However, a few of the avian species have shown the ability to transmit Haemoproteus spp.,

Leucocytozoon spp., and Parahaemoproteus spp. (Hsu et al. 1973; Karadjian et al. 2013;

Valkiunas 2005; Valkiunas et al. 2014). In addition, close relatives of the Eastern gray squirrel may be able to transmit Hepatocystis vassal (Garnham 1966).

4.5 Limitations

Due to unpreventable occurrences, a cautionary note must be included here. First, due to the age of the 2013 collection at the time of mosquito identification, several of the mosquitoes identified as Culex erraticus were ambiguous. Though this was the most likely species, it was difficult to be sure due to the absence of the abdominal scales. It is possible that a few of the Culex erraticus in the 2013 collection were misidentified.

However, a majority of the unfed and engorged mosquitoes captured in 2013 were identified as Culex erraticus, so this was most likely identity of these ambiguous Culex spp. Because the 2017 mosquitoes were identified directly following capture, this problem did not occur.

As mentioned previously, abiotic and biotic factors out of our control may have influenced the amounts of blood-fed mosquitoes captured. This may also have influenced which vertebrates were fed upon or the mosquitoes species captured. In a perfect survey, all environmental factors would be constant at each site, however this is not feasible in a survey of free-living specimens, thus we needed to determine the direct influence of our variables on the blood meal identification.

In addition, these data can only show species supplemented by the NCBI

(National Center for Biotechnology Information) database. It is possible these mosquitoes are consistently feeding within a broader group (i.e. Psorophora mathesoni may feed on several species of box turtles), or the actual vertebrate species may not be available in the

NCBI database.

4.6 Conclusion

In this study, we identified a connection between two potential hosts of a newly discovered haemosporidian: Culex erraticus and white-tailed deer. We showed that Culex erraticus are feeding on white-tailed deer in Mississippi and are potentially acquiring and/or transmitting haemosporidians to this vertebrate host. Due to our findings, it is possible that the novel haemosporidian identified by David et al. and Martinsen et al. represent the same species (Larson et al. 2017; Martinsen et al. 2016).

From our statistical analyses, we determined a significant variation in the relationships between mosquito species and the vertebrate identity. However, we did not see a significant effect of the mosquito identity and collection site on the blood meal identity. Based on these analyses, we can say that certain vertebrate and mosquito relationships may occur more frequently, but they do not necessarily represent a feeding pattern.

Based on the results of this study, there are new, important questions to answer.

First, the haemosporidian prevalence within Mississippi white-tailed deer must be analyzed, particularly for the species identified by Martinsen et al. (2016). Though

Martinsen et al. did a national survey of white-tailed deer, they were unable to sample from Mississippi (Martinsen et al. 2016). Also, we now know Culex erraticus is potentially acquiring this parasite form the natural host, thus host compatibility testing of these mosquitoes may be necessary.

Table 4.1 Haemosporidian Transmission within Vertebrates

EGS ESO ETS GBH HUM NCA NGS WTD WOW *Pla. Pla. Pla. relictum Hae. Pla. vivax Pla. *Pla. relictum **Pla. *Pla. berghei subpraecox herodiadis relictum sp. subpraecox *Hep. *Par. *P. Fal. P. ovale *P. *P. vassali noctuae cathemerium neotropicalis cathemerium gundersi *P. syrnii *P. matutinum Leu. P. malariae *P. matutinum *Hae. Leboeufi syrnii Leu. *P. *L. P. *P. *Leu. danilewskyi circumflexum nycticoraxi falciparum circumflexum danilewskyi Pla. *P. vaughani L. ardeae *P. vaughani *Pla. forresteri forresteri *Hae. *P. rouxi *P. rouxi *Hae. syrnii noctuae *P. elongatum *P. elongatum

Hae. passeris Hae. passeris *Pla. polare *Pla. polare *P. pinottii *P. pinottii *P. *P. octamerium octamerium *P. leanucleus *P. leanucleus List of haemosporidians which may be transmittable by the vertebrate. Identified vertebrates with no evidence of haemosporidian transmission were not included (Bukauskaitė et al. 2015; Garnham 1966; Hsu et al. 1973; Karadjian et al. 2013; Martinsen et al. 2016; Shurulinkov et al. 2003; Tavernier et al. 2005; Valkiunas 2005; Valkiunas et al. 2014). EGS=Eastern gray squirrel, ESO=Eastern screech owl, ETS=Eurasian tree sparrow, GBH=great blue heron, HUM=human, NCA=northern cardinal, NGS=Northern grey-headed sparrow, WTD=white-tailed deer, WOW=wood owl*denotes haemosporidians which are transmittable by a closely related species **According to the phylogenetic tree reconstructed by Martinsen et al. (2016), this haemosporidian does not group closely to other mammalian Plasmodium spp.

Table 4.2 Haemosporidian Transmission within Mosquitoes

AE AV AC AQ CP CE CN Pla. *Hae. lineage Pla. Pla. relictum Pla. polare Pla. Pla. hermani Gallinaceum LIN6h falciparum floridense Pla. P. vivax P. gallinaceum P. circumflexum P. elongatum gallinaceum P. relictum P. relictum P. fallax P. vaughani

P. lophurae P. gallinaceum

P. falciparum

P. vivax

P. cynomolgi

P. gonderi

P. cathemerium List of haemosporidians which may be transmittable by the mosquito. Identified mosquitoes with no evidence of haemosporidian transmission were not included (Falk et al. 2015; Garnham 1966; King 1916; Klein et al. 1987; Santiago-Alarcon et al. 2012; Valkiunas 2005) AE= Aedes canadensis, AV= A. vexans, AC= Anopheles crucians, AQ= A. quadrimaculatus, CP= Coquillettidia perturbans, CE= Culex erraticus, CN= C. nigripalpus **mosquito transmission potential of Haemoproteus spp. has only been demonstrated experimentally

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