Phenotypic and Molecular Characterization of Hard Ticks

(Acari: Ixodidae) Sampled from Wild Herbivores from Lake

Nakuru and Tsavo National Parks in Kenya

Muruthi Wanjira Carolyn

Reg. 156/25663/2011

A Thesis submitted in partial fulfilment of the requirement for the award of the degree of Master of Science (Medical Biochemistry) in the School of Pure and

Applied Sciences of Kenyatta University

September 2015 ii

DECLARATION

I declare that this thesis is my original work and has not been presented for a degree in any other university for any other award.

Signature...... Date......

We confirm that the work reported here was carried out by the candidate under our supervision.

Dr. Joseph N. Makumi

Department of Biochemistry and Biotechnology

Kenyatta University

Signature...... Date......

Dr. Steven M. Runo

Department of Biochemistry and Biotechnology

Kenyatta University

Signature...... Date......

iii

DEDICATION

This thesis is dedicated with love and respect to my loving dad, the late Benson

Muruthi, Mum Susan Muruthi, Sister Zipporah Muruthi, Brother Christopher

Muruthi, and Nephew Mark Muruthi and the late Dr. J.N Makumi for their continued encouragement, patience, love and huge support during the time I was carrying out the project.

ACKNOWLEDGEMENT

I take this opportunity to thank my supervisors the late Dr J. N. Makumi and Dr.

Steven Runo for their advice, encouragement and contribution to this project.

They have tirelessly supported and followed me from the beginning to the end of this work. I also thank Mr. Moses Otiende and Mr. Vincent Obanda (Kenya

Wildlife Services) who co-ordinated my laboratory work. I express my sincere gratitude to Dr. Olivia Wesula Lwande (ICIPE) who introduced me to tick identification techniques and Mutura Kamawe (Institute of Primate Research) who introduced me to molecular techniques.

My special thanks to personnel at the Veterinary Services Department Kenya

Wildlife Service who introduced me to molecular techniques and assisted me in various activities in the laboratory. This includes, Ms Asenath Owour, Ms.

Wambui Maina, Anthony Kipruto Busienei and Samuel Muthemba who provided support while working in the laboratory.

My deepest appreciation goes to my late dad, Benson Muruthi who was always proud of me and was looking forward to the completion of the project. Mum,

Susan Muruthi who has been my inspiration and has supported me in pursuing my dreams. My special thanks to my brother, Christopher Muruthi, sister, Zipporah

Muruthi and nephew, Mark Muruthi for their special support and love throughout the project. v

TABLES OF CONTENTS

DECLARATION ...... i DEDICATION ...... iii ACKNOWLEDGEMENT ...... iv TABLES OF CONTENTS ...... v LIST OF TABLES ...... vii LIST OF FIGURES ...... viii LIST OF PLATES ...... ix ABBREVIATIONS AND ACRONYMNS ...... x ABSTRACT ...... xi CHAPTER ONE ...... 1 INTRODUCTION ...... 1 1.1 Background ...... 1 1.2 Problem Statement and Justification ...... 3 1.3 Hypothesis ...... 4 1.4 Objectives ...... 4 1.4.1 General Objective ...... 4 1.4.2 Specific Objectives ...... 4 CHAPTER TWO ...... 5 LITERATURE REVIEW ...... 5 2.1 Systematics of ticks ...... 5 2.1.1 Morphology of Hard ticks ...... 6 2.2 Biology of Ticks ...... 7 2.3 Economic importance of ticks ...... 9 2.3.1 Impact on the livestock Industry ...... 9 2.3.2 Impact on human beings ...... 11 2.4 Wildlife ticks ...... 13 2.5 Distribution of ticks in Kenya ...... 15 2.6 Tick identification ...... 16 vi

CHAPTER THREE ...... 20 MATERIAL AND METHODS ...... 20 3.1 Study design and sample size ...... 20 3.2 Study sites ...... 21 3.3 Morphological identification of Ticks ...... 21 3.4 Molecular analyses of tick ...... 22 3.4.1 DNA Isolation...... 22 3.4.2 Polymerase Chain Reaction ...... 23 3.4.3 Electrophoresis and purification of PCR products ...... 23 3.4.4 Sequencing and analysis ...... 24 3.5 Congruency test ...... 24 CHAPTER FOUR ...... 25 RESULTS ...... 25 4.1 Morphological identification ...... 25 4.1.1 Unidentified ticks ...... 35 4.2 Identification of Ticks using PCR and gel electrophoresis ...... 36 4.1.2 Phylogenetic analysis ...... 37 4.3 Morphological and molecular characterization of tick samples ...... 46 CHAPTER FIVE ...... 48 DISCUSSION, CONCLUSIONS AND RECOMMENDATIONS ...... 48 5.1 Discussion ...... 48 5.2 Conclusions ...... 53 5.3 Recommendations ...... 54 5.4 Further Studies ...... 54 REFERENCES ...... 55 APPENDICES ...... 62 APPENDIX 1 ...... 62 APPENDIX 2 ...... 63

vii

LIST OF TABLES

Table 4.1: Ticks identified by morphological keys in genus Amblyomma...... 26

Table 4.2: Ticks identified by morphological keys in genus Rhipicephalus...... 26

Table 4.3: Ticks identified by morphological keys in genus Hyalomma...... 26

Table 4.4: Ticks identified by morphological keys in genus Dermacentor...... 26

Table 4.5: Different sizes of ITS2 as depicted in the gel electrophoresis ……..38

Table 4.6: Morphological and Phylogenetic Identification...... 46

viii

LIST OF FIGURES

Figure 4.1: GenBank blast search for R. pulchellus haplotype………………40

Figure 4.2: GenBank blast search for A. variegatum haplotype……………..41

Figure 4.3: GenBank blast search for H. marginatum rufipes haplotype……42

Figure 4.4: GenBank blast search for sample H. dromedarii………………..43

Figure 4.5: Phylogenetic tree………………..……………………………….45

LIST OF PLATES

Plate 1: Dorsal and ventral region of a female R. pulchellus ………………....27 Plate 2: Dorsal and ventral region of a male R. pulchellus...... 27 Plate 3: Dorsal and ventral region of a male A. gemma……………………….28 Plate 4: Dorsal and ventral region of a female A. gemm...... 29 Plate 5: Dorsal and ventral region a male of A. variegatum…………………...29 Plate 6: Dorsal and ventral region of a male A. tholloni……………………….30 Plate 7: Dorsal and ventral region of a female H. dromedarii ………………..30 Plate 8: Dorsal region of a male H. dromedari...... 31 Plate 9: Ventral region of a male H. dromedarii...... 32 Plate 10: Dorsal and ventral region of a male H. albiparmatum...... 32 Plate 11: Dorsal region of a male H. truncatum...... 33

Plate 12: Ventral region of a male H. truncatum...... 33

Plate 13: Dorsal and ventral region of a male H. marginatum rufipe...... 34

Plate 14: Dorsal and ventral region of a male D. rhinocerinus…………………35

Plate 4.2: Sample agarose gel amplification of PCR products of ITS 2 region of ticks...... 36

ABBREVIATIONS AND ACRONYMNS

BLAST...... Basic Local Alignment Search Tool DNA...... Deoxyribonucleic Acid ITS...... Internal Transcribed Spacer ITS 2...... Internal Transcribed Spacer 2 PCR...... Polymerase Chain Reaction rDNA...... ribosomal DNA. Spp...... Species

ABSTRACT

Hard ticks (Acari: Ixodidae) are ubiquitous blood feeding ectoparasites that infest humans and animals and are vectors of pathogenic micro-organism of rickettsial, bacterial, viral and protozoal origin that cause severe infectious diseases in humans and livestock. Wild herbivores support a large population of these vectors and accurate and quick identification of ticks is therefore important. Morphological identification based on phenotypic criteria has been the main approach but the technique is considered inaccurate and may result in misidentification. Molecular techniques using genetic markers have recently been considered to be appropriate approaches for accurate and rapid identification and Internal Transcribed Spacer 2 (ITS 2) has been shown to differentiate genus of hard ticks. Currently, genetic identification of ticks using ITS 2 has not been carried out in Kenya and there is no database of ITS 2 from locally located tick samples. The aim of this study was therefore, to genetically identify ticks and determine their congruence with phenotypic traits. A total of 98 tick samples were collected from L. Nakuru and Tsavo National Parks during scheduled management activities by the Kenya Wildlife Services. Morphologically, tick samples were identified to genus and species using appropriate identification keys. DNA was extracted from the appendages using DNA extraction kit (DNeasy blood and Tissue Kit, QIAGEN) followed by partial amplification of ITS 2 gene. The PCR products were then analyzed by gel electrophoresis and positive PCR products were sequenced. Of the tick samples four genera were identified morphologically; Amblyomma (26%), Hyalomma (38%), Rhipicephalus (30%) and Dermacentor (3%). Of the tick samples identified and compared with the sequences in the GenBank, six and seven samples showed 98-100% homology with A. variegatum and R. pulchellus respectively and they clustered in their respective monophyletic group in the phylogeny tree with a bootstrap of 99%. Two samples showed 92% homology with H. dromedarii and the study sequences clustered with the reference sequence with a bootstrap of 99% while six samples showed 95% homology with H. marginatum rufipes, however, only four of these samples clustered together with the reference sequence in the phylogeny with a bootstrap of 95%. One sample showed 91% homology with A. humerali and did not cluster together in the phylogeny tree. Congruency between both techniques was high with a correlation coefficient of 0.941. The study concluded that morphological and molecular techniques can be used to identify ticks. This is the first report of phenotypic and genotypic traits of tick species in Kenya and the findings will add value to the existing knowledge in identification of ticks. 1

CHAPTER ONE

INTRODUCTION

1.1 Background Ticks are ubiquitous blood feeding ectoparasites of humans and animals and are vectors of a plethora of pathogens that cause severe infectious diseases in humans and livestock (Jongejan and Uilenberg, 2004). Tick infestation and the diseases they transmit result in huge economic losses in livestock production and continue to cripple the industry, especially in sub-Saharan Africa (Horak et al., 1983). In humans, tick-borne pathogens pose significant public health risks and transmit pathogenic micro-organism of rickettsial, bacterial, viral and protozoal origin.

Majority of tick species infect multiple host species, while some are restricted to particular host taxa. The control of tick infestation and spread is a multi-billion global investment and accurate identification is vital in order to make it effective

(Lampo et al., 1997).

Ticks tend to be localized in specific ecosystems but increased speed and movement of people, ecotourism, translocations of wildlife, trade in live wildlife pets and climate change provide risks of spread beyond their natural ranges.

According to De Meneghi (2006) the diversity and number of wildlife species is enormous all over the world and this diversity sustain a large and diverse tick- borne vectors of pathogens (Lanfranchi et al., 2003; Bengis et al., 2004). Wild herbivores are believed to be significant reservoirs of tick-borne pathogens that 2

affect humans and animals (Tonetti et al., 2009) as they support a large populaton of tick species (Peter et al., 1998).

Reliable and quick identification of ticks is therefore important in the control of spread of tick-borne diseases. Morphological characterization based on phenotypic traits has been the traditional method of identifying ticks.

Nevertheless, the approach is laborious, often inaccurate and may result in misidentification. In addition, differentiation of larvae and nymph of closely related species of ticks is difficult and often inaccurate (Abdigoudarzi et al.,

2011). Morphological identification method is further weakened by the fact that traits used to differentiate species tend to overlap between species or vary within species or according to age and size. Additionally, morphological identification of physically damaged ticks due to poor handling and preservation is often inaccurate. The advent of molecular techniques and development of markers that can identify ticks have revolutionized our understanding of other insect vectors and may be useful in studies of tick taxonomy and diversity.

Identification of ticks especially Rhipicephalus and Hyalomma spp. morphologically is difficult even when applying the nomenclature. Apanaskevich and Horak (2006) found it difficult to differentiate Hyalomma anatolicum excavatum and Hyalomma anatolicum anatolicum using morphological criteria which were weak including colour and the size of the scutum. Additionally, morphological identification of Dermacentor spp., which are rare, is similarly difficult. Zahler et al. (1995) used molecular methods by partial sequence of the 3

ITS 2 region to study Dermacentor and Rhipicephalus spp. Murrell et al. (2001a) used DNA amplification while studying the phylogenetic relationship of hard ticks; and by using molecular methods, the researchers were able to different genera and species of hard tick. With this, genetic techniques have contributed to the understanding of tick taxonomy through improved species identification.

As such, the purpose of this study is to genetically identify ticks sample and confirm whether the distinctive phenotypic traits for tick species differentiation are congruent with genetic traits.

1.2 Problem Statement and Justification Ticks are significant obligatory blood feeding ectoparasites that transmit different protozoan, bacterial and viral agents to animals and humans. To comprehend the epidemiology of tick-borne pathogens and develop effective strategies for controlling the diseases, accurate identification of the vector is vital. Currently, the tick diversity or taxonomy in Kenya is based on historical records that were determined by morphological methods (Walker et al., 2013) and yet the method has inherent challenges and weakness that weaken accuracy of species identification.

Due to contemporary changes in land use, movement of people and animals, the distribution and diversity of tick species may have changed over time with possibility of incursion of new tick species from foreign regions. Although, ticks can be identified directly using genetics, it is important to describe its distinctive phenotypic features and test for congruency with genetic classification. Currently, 4

genetic identification techniques have not been effectively used got identification of tick in Kenya. Still, there is no evidence that the phenotypic and genotypic traits of tick species in Kenya are congruent. Therefore, incongruency in identification based on the two techniques will justify the need for taxonomic re- classification. Additionally, the information generated will quicken future identification of tick species.

1.3 Hypothesis Phenotypic and genetic taxonomic traits can be used to identify tick species collected from wild herbivores in Kenya and the two methods of characterization are congruent.

1.4 Objectives

1.4.1 General Objective To phenotypically and genetically identify ticks collected from wild herbivores from selected areas in Kenya

1.4.2 Specific Objectives i. To identify tick species morphologically

ii. To isolate and identify tick species by partial sequence of ITS2 gene iii. To infer congruency of phenotypic and genetic traits

CHAPTER TWO

LITERATURE REVIEW

2.1 Systematics of ticks

Ticks belong to a small taxanomic group comprising about 860 species (Horak et al., 2002). They can easily be separated from the wide phylum Arthropoda into the subphylum Chelicerata due to the presence of anterior chelicerae. As they are closely related to scorpions and mites, they are classified in the class Arachnida

(Soneshine, 1991). They are in the subclass Acari, order Parasitiformes (Anderson

& Magnarelli, 2008). According to Wall & Shearer (2008) the suborder of

Ixodides contains 4 families Ixodidae (hard ticks), Argasidae (soft ticks),

Laelaptidae and Nuttalliellidae. The latter two families have one species each

(monophyletic) and are of minor importance (Anderson & Magnarelli, 2008). The hard ticks are the largest group with approximately 650 species while the soft ticks comprise about 150 species (Wall & Shearer, 2008).

The two major groups can be differentiated using morphological, behaviour and biological criteria (Mehlhorn, 2008). Morphologically, the soft ticks have leathery and smooth cuticle while the hard ticks have relatively hard cuticle. The latter are found out doors, rarely are they seen in human dwellings. On the other hand, the soft ticks are found near human dwellings in the nest of animals. Common genera of soft ticks include Argas, Antricola, Ornithodorus and Otubius. The hard ticks are classified into two groups Prostrata and Melistriata. Prostriata that contain the

Ixodid, which forms that largest genus with approximately 217 known species and 6

the rest of hard ticks fall under the genus Metastriata (Wall & Shearer, 2008). A profound difference between these two groups is the location of the anal groove; in the Prostriata, it is located posterior to the groove while in the Metastriata, the anus is located posterior to the anus (Mehlhorn, 2008). Important groups in the metastriata genera include Amblyomma, Anocentor, Apanomma, Haemaphysalis,

Hyalomma. Rhipicephalus, Boophilus and Dermacentar.

2.1.1 Morphology of Hard ticks Generally, ticks are closely related to mites on the basis of body structure.

However the presence of Haller’s organs (sensory organ) and hypostome

(mouthpart) separate the ticks from the mites (Krantz, 1978). The hard ticks are relatively large and range from 2 to 20 mm in size. An unfed tick has a dorsoventral body and the structure seem similar to that of a mite; divided into two parts, the anterior capitulum (gnathosoma) and the posterior idiosoma. The gnathosoma bears the palp, a four paired segmented sensory organs. The fourth segment of every palp forms a pincer-like structure (Wall & Shearer, 2008).

Between the palps is the chelicera, two-segmented appendages which is highly sclerotised. The chelicerae are used to cut and pierce a host skin during feeding.

The basis capituli, fused coxae of the palp (Wall & Shearer, 2008), differs in shape among different genera. The basis of capituli is extended both ventrally and anteriorly to form the hypostome.

Podosoma is a region in the idiosoma that bears the legs. Each leg is connected to the main body at the coxa. After the coxa are the trochanter, femur, genu, tibia 7

and the tarsus (Wall & Shearer, 2008). The legs of the ticks end with a pulvillus and a pair of claw. On the tarsi, a Haller’s organ is present that bear the chemoreceptor that are used to locate the hosts. The chemoreceptors are also present in the scutum, chelicerae and palps.

2.2 Biology of Ticks Hard ticks undergo four stages of development; eggs, larvae, nymph and adult and most require three different hosts and are not conspecific (Soneshine 1991). In most cases, the larvae and nymph engorge on smaller and medium host while the adult feed on large hosts. It has been noted that some few ixodid have evolved from multiple host to two-host pattern. In a two-host cycle the tick usually survive for two years. The larvae seeks the first host to attach, moults and the nymph continues feeding on the same host, drops off, moults and the resulting adults seek for a new host. After feeding, the female drops off the host and after several days, lays lots of eggs and dies. Examples of hard ticks that have this kind of cycle include the Hyalomma spp. In the one-host cycle the tick remains on the same individual host for all stages (Junquera, 2013); example of such species include the Rhipicephalus annulatus.

In a three host cycle, the larvae, nymph and adult detach from an individual host after every blood meal. The larvae and nymphs each attach to a new host for another blood meal after moulting in the ground. An adult female deposits eggs on the ground and dies. Example of ticks that have this cycle includes the

Amblyomma spp. whose survival may last for 2 to 4 years (Junquera, 2013). To 8

have a full feeding cycle, larval feeding on warm blooded animals requires 3-7 days while a nymph 4-8 days. Adult females feeding on mammals and birds require 7 -12 days while those that feed on reptiles take longer (Oliver, 1989).

Many ticks have incorporated diapauses in their life cycle (Randolph, 2004) so as to ensure that emergence coincides with optimal conditions needed for survival.

Diapause is significant in influencing seasonal adaptation in ticks (Oliver, 1989).

In addition, it enhances the ticks circadiam rhythms for feeding, drop off, oviposition and host seeking. Diapause in ticks has been found to vary according to climatic conditions. For example, in South Africa the Rhipicephalus adult ticks that emerge after July every year do not starting questing until in November the same year (Randolph, 1994). The short day lengths are seen to induce diapause

(Madder et al., 1999), and this results to near absence of every tick stage successively during the year. Randolph (2004) observed that majority of susceptible egg-larval stage happens during the warm and wet season. This conditions favour good survival and rapid development, despite the fact that wet conditions are associated with high mortality.

In Equatorial Africa, most ticks do not exhibit diapause. The ticks feed throughout the year therefore, resulting to overlapping generations. The above diapause pattern has been observed in some species of genus Rhipicephalus. The diapause pattern is not only important for the survival of the tick species but also for the epidemiology of various associated infections (Randolph, 2004). For instance,

Norval et al. (1991) assert that the occurrence of diapause among Rhipicephalus 9

appendiculatus in southern Africa to some extent account for absential of virulent strain of Theileria parva, a causative agent for East Coast Fever in cattle in central and eastern Africa.

2.3 Economic importance of ticks

Ticks are of great economic importance because of their direct and indirect effects on humans and domestic animals (Soneshine, 1991). Directly, ticks transmit a variety of pathogenic protozoa, spirochaetes, viruses, rickettsiae and micro- organism to humans, livestock and companion animals (Jongejan & Uilenberg,

2004). During feeding, ticks concentrate blood nutrients in their gut and get rid of the excess water from their bodies. They use their salivary glands to pump the waste (by-product of the blood meal) into the host. Through this process, the waste expelled from the body have the potential to transmit rickettsial, bacteria, protozoa, fungal and viral pathogens to animal and human hosts. Transmission of pathogen through a tick bite is not an automatic or immediate effect as an infected tick should attach and feed for more than two days for the transmission to occur

(Sonenshine, 1991).

2.3.1 Impact on the livestock Industry

The impact of ticks on animals depends to a large extent on a number of factors mainly, the species involved, local climatic conditions and also on the vulnerability to tick infestation of the livestock in a region (Jongejan & Uilenberg,

2004). Resistance to ticks infenstation is determined genetically particularly where one host species (Boophilus) are implicated. It has been observed that an 10

uncontrolled Boophilus infestation in a favourably climatic conditions affects the

European cattle severely but has little effect on the zebu cattle. Such infestations severely limit the productivity of exotic cattle resulting to the utilization of intensive and expensive chemical tick control, that leads to resistance against acaricides applied (Willadsen et al., 1998).

The damages caused by tick bites diminishes the value of hides and skin for the production of leather. According to Jongejan and Uilenberg (2004) ticks with short hypostome are significant in this respect if they are present in large numbers on the host. Tick species that have massive and long hypostomes are known to cause secondary bacterial infections. In this way, they may cause lameness or loss of teats, depending on the attachment site; loss of teats leads to an increase in calf mortality (Jongejan & Uilenberg, 2004). The saliva of particular tick species has paralysing toxins. For instance, in southern Africa, Ixodes rubicundus induce severe form of paralysis in sheep. ‘Sweating sickness’ is another form of tick toxicosis, an eczema-like condition affecting calves and other livestock in Africa.

Bovine dermatophilosis is a tick associated disease that is not directly transmitted by ticks. The disease is induced by the presence of A. variegatum which hinders the upgrading of local cattle breeds with susceptible breeds in some regions in the tropics (Ambrose et al., 1999). Exotic livestock are highly susceptible to various tick borne diseases particularly if not previously exposed to natural selection.

Theileriosis, East Coast Fever and cowdriosis, are examples of diseases that affect exotic livestock introdced in a disease free area. Infact cowdriosis and theilerioses 11

may make it uneconomical or impossible to keep exotic ruminants unless veterinary and management infrastructures are adequate. This also applies to local breeds that have never been exposed to specific pathogens. For example, the movement of goats from northen to southen Mozambique, an area infested by A. hebraeum, is hindered due to the risk of contacting cowdriosis (Bekker et al.,

2001).

Ticks and tick-borne diseases are significant in tropical and subtropical areas.

Young et al. (1989) considered the control of ticks as an important health and management problem in Africa. According to most researchers, the economic losses in controlling and managing ticks in livestock industry is high (Jongejan &

Uilenberg, 2004). Mukhebi et al. (1999) estimated the national annual costs due to cowdriosis in Zimbabwe to be US$ 5.6 million while Meltzer et al. (1996) estimated a loss of US$ 6.5 million in cattle on large scale production alone in the same country in 1991. In Kenya and Tanzania, Minjauw & McLeod (2003) estimated the annual cost due to East Coast Fever in smallholder dairy system to be US$ 54.4 and 4.41 million respectively while in the traditional system is 129.5 and 34.1 million respectively.

2.3.2 Impact on human beings

Parola & Raoult (2001) published reviews on human tick borne diseases and reported 15 new bacterial pathogens by 1982. The increasing array of tick borne zoonotic diseases emerging in the world poses an increased public health risk

(Cunha, 2000). Recreational activities in rural areas have increased resulting in an 12

increase in human tick bites. In some rural areas in Tanzania, people sleep outdoors in order to avoid bites from the indoor dwelling soft ticks (Talbert et al.,

1998), resulting in tick control by spraying to reduce the occurrence of tick-borne fever.

As studies on ticks continue due to their impact on human health, novel microbial associations have continued to be described (Jasinkas et al., 2007) and new reports on emerging diseases are on the rise (Paddock & Yabsley, 2007; Lwande et al., 2013). Ticks also serve as hosts for a variety of vertically transmitted bacteria, endosymbiotic and Rickettsia-like organism and novel symbionts of tick mitochondria (Jasinkas et al., 2007).

Different species have been identified as vectors for various infectious diseases in humans. Argas monolakensis is a significant argasid tick that feeds on humans in

California, USA and is the vector of Mono Lake virus (Schwan et al., 1992).

Argas neghmei and Argas moreli have been implicated in human parasitism in

Chile and Peru respectively in areas where chicken and humans cohabit (Keirans et al., 1979). Among the ixodidae, Ambylomma americanum is distributed throughout eastern and southern part of USA, south and central America, it is very aggressive and accounts for 83% of ticks that feed on humans in America

(Felz et al., 1996). In Kenya, Lwande et al. (2013) assosciated viruses noted to be mosquito-borne to R. pulchellus and A. gemma; ticks that have been identified to be abundant among wild herbivores in Kenya (Keesing et al., 2013). 13

Human international traveling, one of the largest evolving industries, has an impact on health. Travel is associated with tick borne diseases and is emerging as a common subject of international travel medicine. African-tick bite fever caused by Rickettsia africae that is transmitted by Amblyomma spp. is found in the sub-

African and Carribean regions. The clinical course of the disease is mild but complications may occur and infection is often confused with tropical diseases such as malaria (Jensenius et al., 2003). Crimean-Congo hemorrhagic fever, a zoonotic viral disease and the causative virus is transmitted by ticks, human- human and animal-human contact (Flick & Whitehouse, 2005), is widely distributed in Middle East, Asia and Africa. The principal vector of the disease is said to be the Hyalomma and the virus causes viral haemorrhagic fever outbreaks reported in various countries with a case fatality rate of 9 – 39 per cent (WHO,

2013).

2.4 Wildlife ticks Ticks are of vast significance to the conflict between agricultural interests and wildlife conservation (Anderson et al., 2013). This has become evident given that the diversity and number of wildlife species is enormous all over the world (De

Meneghi, 2006). This variety and abundance is distributed throughout different ecological niches and habitats. Lanfranchi et al. (2003) and Bengis et al. (2004) and noted that these diversity of ecosystems and species may also sustain a large and diverse tick- borne vector pathogens. The current global environmental changes have largely determined the biomodal evolution of the wildlife (De

Meneghi, 2006); and these include increase in human activities, frequency of 14

intercontinental and international travels, disruption of wild habitats and modification of livestock-agriculture production systems.

Wild herbivores have long been believed to be significant in the tick-borne pathogens that are affecting humans and animals, and several wild herbivores species have been identified as major reservoirs for tick borne pathogens (Tonetti et al., 2009); for instance, African buffalo are a reservoir for corridor diseases,

East Coast fever, and heartwater (Young, 1977; Allsopp et al., 1999). Lohr &

Meyer (1973), Carmichael & Hobday (1975), and Peter et al. (1998) demonstrated the transmission of pathogenic microorganisms from the wild herbivores to the livestock.

Ticks are distributed in different environments depending on host availability

(Norval et al., 1994), vegetation structure and type (Tukahirwa 1976; Norval

1982), elevation (Walker, 1974) and climate (Walker 1974; Randolph 1994). All these variables are interrelated; however, their relative importance differs in determining the distribution of ticks. Distribution of ticks in Africa is directly determined by climate effects; such climatic variables are better forecasters of distributions of ticks than vegetation related variables and are not greatly limited by those of their hosts. It can be assumed therefore, that the primary factor that contributes or prevents the expansion of the tick species is the direct effect of climate (Walker, 1974). 15

2.5 Distribution of ticks in Kenya

Kenya is rich in hard ticks that infest both domestic livestock and wild animals.

The diverse climatic conditions in Kenya allow the country to accommodate a variety of tick species. Species under genus Amblyomma are widely distributed in the highland and savannah regions of Kenya. These include the Central, Rift

Valley and coastal regions. In the savannah regions of Tsavo National Park, for instance, the climatic conditions allow the survival of A.hebraeum which requires warmth and moisture bush and brush to survive Most species in genus

Haemaphysalis are also found in Kenya, the favourable climatic conditions include humid and warm conditions making the highlands regions conducive for the survival of these species (Walker et al., 2003).

Species in the genus Hyalomma are well distributed in the drier eastern, northern eastern and coastal regions in the country. The genus is well adapted to extreme dryness and the species include H.dromedarii, H.truncatum and H. marginatum rufipes (Walker, 1974). Genus Rhipicephalus are widely distributed in Kenya; these include R. (Bo.) microplus which are mainly found in the coastal strip of the country, R.appendiculatus and R. evertsi evertsi which occur in the temperate highland region while R.pulchellus is widely distributed in the Rift Valley and the coastal region (Walker, 1974). In their studies Kariuki et al. (2012) noted that

A.gemma, A.lepidium, H. albiparmatum, H.marginatum rufipes. H.truncatum,

R.pulchellus, R.evertsi, R.pravu infested both bufalloes and cattle in Tsavo 16

National Park. They noted new locality records for H.truncatum and

H.impeltatum in the region. Most of the species they identified was in line with

Walker (1974).

2.6 Tick identification

With the growing interest on ticks as vectors of diseases in humans and animals, various reviews on taxonomy have been carried out (Abdigoudarzi, 2004).

Morphological characterization of ticks based on phenotypic criteria has been the traditional method of identifying ticks (Soneshine, 1991). However, various studies have established that there are drawbacks associated with morphological identification. The approach is quite difficult, does not give expected result and may result in misidentification (Anderson et al., 2004). In addition, ticks belonging to different genera are identified using dichotomous key. However, this system requires solid training and an experience in morphology and taxonomy.

Larvae and nymphs are more difficult to identify especially in species that are closely related (Ganjali et al., 2011). In distinguishing characteristics, overlap and high intraspecific variability may occur. Still, morphological identification is not well applicable to damaged ticks. Some morphological traits may vary with size and age and the technique may also represent polymorphism in a single population (Rumer et al., 2011). In their studies Adler & Feldman-Muehsam

(1948) and Hoogstraal & Kaiser (1959) noted that identifiction of Rhipicephalus and Hyalomma species using nomenclatures was difficult. 17

Apanaskevich & Horak (2006) supported this when they tried to discriminate

Hyalomma anatolicum excavatum Koch and Hyalomma anatolicum anatolicum

Koch based on their morphological traits. The identification proved difficult due to the different size and colour of the scutum which differ at various stages of tick growth (Abdigoudarzi et al., 2011). On Rhipicephalus; three species, R. sanguineus Latreille, R. turanicus and R.bursa are quite similar and there is a need to clarify if they are distinct species or subspecies. Using morphological traits in identifying engorged ticks, would be a misleading approach because of changes in body surface after feeding (Abdigoudarzi, 2004).

Subsequently, it is significant to develop more relevant characterization methods in order to differentiate subspecies and species while at the same time offer reliable and convenient approach. Molecular techniques using appropriate markers are more convenient and reliable approaches and have been used to study vectors of parasitic diseases. These techniques offer apparent opportunities as they easily distinguish closely related taxa in ticks (Cruickshank, 2002; Rumer et al., 2011). Pathogenic ticks may be identified by conventional methods such as histological staining techniques, indirect immune-fluorescence or isolation in cell culture. Nevertheless, the advent of PCR has led to improvements of the sensitive and specific detection of pathogen DNA in different species of ticks.

According to Zahler et al. (1995) molecular techniques have been used in clarifying problems arising from species identification. In studying Dermacentor spp. and Rhipicephalus spp., Zahler et al. (1997) used DNA amplification of ITS 18

2 region (Abdigoudarzi, 2004). A study on phylognetic relationships of various subfamilies of hard ticks was carried out by Murrell et al. (2001b) using ITS 2 which provided resolutions of relationships and intrageneric relationship in genera. Studies have been carried out using partial sequences of ITS 2 for both

Boophilus spp and Hyalomma spp.(Murrell et al., 2001a; Rees et al., 2003). After studying phylogenetic relationship of hard ticks using the ITS 2 region Fukunaga et al (2000) and Abdigoudarzi et al. (2011) recommended it as a good marker for identification and analyses of ticks.

Additionally, ITS 2 has been found to be favourable for primer design (Rumer et al., 2011) and is an important tool for genera and species identification even at low taxanomic levels (Robert, 2002). Abdigoudarzi et al. (2011) concluded that

ITS 2-rDNA is useful in studying phylogenetics of Ixodidae at both genus and species levels. Ganajli et al. (2011) noted that ITS 2 represents a region that can be used to identify Hyalomma spp. ; however, further investgation is required so to validate the use of ITS 2 in identifying these species. Similarly, Dergousoff &

Chilton (2007) differentiated three species of Dermacentor; Dermacentor andersoni, Dermacentor albipictus and Dermacentor variabilis using rDNA marker. Rumer et al. (2011) using PCR were able to differentiare medically important tick species; Ixodes hexagonus, dermacentor reticulatus and Ixodes persulcatus in Euro-Asia. They noted that by using PCR approach, ticks at various stages could easily be distingushed including those in unfed and engorged conditions. As such, they concluded that this approach could be useful in large epidemiological studies. 19

In Kenya, identification of ticks is based on historical records that were determined by morphological methods, and yet the method has inherent challenges and weakness that reduce accuracy of species identification. Genetic identification techniques have not been determined in Kenya, and there is no database of ITS 2 from locally collected ticks. The objective was to genetically identify ticks and determine their congruence with phenotypic traits.

CHAPTER THREE

MATERIAL AND METHODS

3.1 Study design and sample size Tick samples were collected randomly during scheduled veterinary management activities within the parks. The animals were immobilized by the Kenya Wildlife

Service veterinarians using a combination of etorphine and xylazine (Norvatis,

PTY, Ltd, South Africa). Ticks were pulled off manually from the animals by hands and with forceps and placed in sterile loosely capped plastic vials and transported to the laboratory in dry ice. In the laboratory the ticks were stored at -

80°C

The sample size was calculated based on the anticipated prevalence and desired degree of precision, derived as; n = P x (100-P) x Z 2/d2 (Daniel, 1999); where (P) is the anticipated prevalence, z the appropriate value from the normal distribution from the desired confidence (0.05), d the desired precision; whereby, the prevalence of ticks with similar genotype and phenotype characteristics ranges between 0 and 5 %. Therefore;

= 0.05 X (1.00-0.05) X (1.96 X 1.96) / (0.05 X 0.05)

= 0.0475 X 1536.64

= 73

From the calculation N= 73, but 98 samples were considered in the study. 21

3.2 Study sites The samples were collected from wild herbivores in Lake Nakuru and Tsavo

National Parks. L. Nakuru National Park was established in 1961 and covers an area of approximately 188 Km2. It lies between longitude 36o 05’ and 0o 24’

South and it is located in Nakuru. The park has four kind of vegetation; forest, woodlands, open waters and open grasslands. Climate conditions in this park vary depending mostly on attitude and ranges from semi-arid, arid and cold. L. Nakuru

National Park is home to over 50 mammal species including rhinoceros, buffaloes, giraffes, elephants and gazelles (KWS, 2002).

Tsavo West National Park is located in the coast and covers an area of approximately 9,065 Km2. It was established in 1948 and lies between longitude

3o 19’30’’ South and 38o8’29’’ East. The savannah ecosystem in the park comprises of scrublands, riverine vegetation, open grasslands, rocky ridges and acacia woodlands. River Galana flows through the park and other waters masses including L. Jipe, Mzima springs and several swamps making it possible for the park to support large mammals including buffaloes, elephants and a home of black rhinos (Wright, 2007).

3.3 Morphological identification of Ticks Sampled ticks were kept at room temperature to thaw, and then washed twice with sterile water to remove excess particulate contamination from animal skin, rinsed once with 70% ethanol. They were mounted on slides and examined using a stereoscope microscope at a magnification of x40, x80, and x100. Identification of the ticks was by sex and species using appropriate identification keys (Walker et 22

al., 2013). The identified ticks were transferred to sterile vials, and stored at -80°C until processing at the KWS Veterinary Laboratory, Nairobi.

3.4 Molecular analyses of tick

3.4.1 DNA Isolation DNA was extracted from each sample using DNA extraction kit (DNeasy blood &

Tissue Kit, QIAGEN (Germany) following the manufacture’s protocol. The stored samples were thawed at room temperature and from each sample tick, the appendages removed and cut into small pieces and placed in labelled microcentrifuge tube. This was followed by addition of 180 µl tissue lysis buffer

ATL, and 20 µl of proteinase K and the mixture was thoroughly mixed by vortexing and then incubated overnight at 56o C. The mixture was vortexed for 15 seconds, then, buffer lysis buffer concentration 216ml (AL) (200 µl) added and mixed thoroughly by vortexing. Immediately, ethanol (200 µl) was added and the mixture again mixed thoroughly by vortexing. The mixture was then pippeted into

DNeasy Mini Spin column in a 2 ml collection tube and centrifuged at 8000 rpm for 1 min, and, the collection tube and flow through discarded. The spin column was placed in a new 2 ml collection tube and 500 µl wash buffer, concentration

242 ml (AWI) added to the mixture and centrifuged at 8000 for 1 min and the flow-through and the collection tube discarded. The washing process was repeated by adding wash buffer 2, concentration 342 ml (AW2) (500 µl) and centrifuged at 14,000 for 3 minutes and the flow-through and collection tube were discarded. The spin column was placed in a 2 ml centrifuge tube and eluted with buffer AE (200 µl), incubated for 1 min at room temperature and then centrifuged 23

at 8000 rpm for 1 minute. The procedure was then repeated so as to increase the overall DNA yield; the DNA samples were stored at -20oC.

3.4.2 Polymerase Chain Reaction Genomic DNA extracted from the 28 ticks’ tissues was amplified. Each species identified morphologically was represented by more than one sample. A pair of degenerate primers designed for ITS2 amplification (Abdigourdarzi et al., 2011) were used; forward 5´-YTGCGARACTTGGTGTGAAT-3´ and reverse 5´-

TATGCTTAARTTYAGSGGGT-3´.

Amplification of DNA was carried out in a final volume of 25 μl containing 15.8

μl of dH20, 2 μl of the genomic DNA, 2 μl of each primer (forward and reverse), 5

μl PCR buffer, 0.2 μl of Taq DNA polymerase. The tubes were then placed into a programmed Applied Biosystems Veriti 96 well thermocycler (Germany) where the reactions mixture was subjected to 2.5 min DNA denaturization at 94°C, 35 cycles of denaturization at 94°C for 30 sec, annealing at 50°C for 1 min and elongation at 72°C for 1 min. The reaction was completed by a further 30 min step at 72°C. The procedure was repeated twice for each sample in order to yield a final volume of 50 μl.

3.4.3 Electrophoresis and purification of PCR products The amplified products were analyzed by electrophoresis on 1% Agarose gel by aliqouting 4 µl of PCR products and 1 kb DNA ladder and run on the gel stained with ethidium bromide for 1 hour at 80 volts. The PCR products were purified using QIAqucik PCR Purification Kit (Germany) following the manufacturer’s 24

protocol. The stored PCR products were thawed and mixed at 25oC, and 46 µl of binding buffer added to 46 µl of the PCR product in the tube, mixed until the mixtures turned yellow. The solutions were then placed in QIAquick purification columns, centrifuged for 1 minute, until the flow through the column, and the flow-through discarded. To the QIAquick column, 750 µl of wash buffer was added, centrifuged for 1 minute and the flow-through discarded. The empty

QIAquick column were centrifuged again so as to remove residual wash buffer, then transferred to a clean 1.5 ml microcentrifuge tube. To elute the DNA, 50 µl of elution buffer were added to the columns and centrifuged for 1 minute, the flow-through discarded and the eluted purified DNA was then stored at -20oC.

3.4.4 Sequencing and analysis Purified DNA products were sent to Biosciences Eastern and Central Africa

(BecA-ILRI) for sequencing. Sequencing was carried out through ABI310 DNA sequencer. Sequences were taken through Bioedit, and aligned using multiple alignment program ClustalX. The default 6.66 and 15 values for gap extension and gap opening respectively were used. Misaligned nucleotides were detected and realigned manually. BLASTN searches were done in the GenBank so as to identify matches to the sample sequences. ITS 2 sequences from the GenBank with 98 per cent or closest similarity to the sample sequence were considered. A table was drawn to compare ticks sample morphologically and genetically.

3.5 Congruency test Correlation statistic were computed to test for congruency between morphological and genetical traits at P= 0.01 25

CHAPTER FOUR

RESULTS

4.1 Morphological identification Tables 4.1-4 show number of tick species that were correctly identified morphologically by genus from Lake Nakuru and Tsavo National Parks. Out of

98 ticks collected from elephants, buffalo and rhinoceros, 80 tick species were correctly identified using morphological keys. Four genera were correctly namely

Amblyomma, Hyalomma, Rhipicephalus and Dermacentor. The species identified in the genus Amblyomma as shown in table 4.1 were A. tholloni (Plate 6), A. gemma (Plates 3 and 4) and A. variegatum Plate 5). In the genus Rhipicephalus, only one species was identified as shown in table 4.2; Rhipicephalus pulchellus

(Plates 1 and 2).

Among the Hyalomma spp. 31 species were identified as shown in table 4.3; namely; H. truncatum (Plates 11 and 12), H. albiparmatum (Plate 10) and H. marginatum rufipes (Plate 13), H. dromedarii (Plates 7, 8 and 9). Table 4.4 shows

3 species which were identified in genus Dermacentor, Dermacentor rhinocerinus

(plate 14). H. truncatum, H. albiparmatum, A. gemma, R. pulchellus were identified from elephants from Tsavo while D. rhinocerinus were identified from rhinoceros in Lake Nakuru National Park. A. variegatum was found in elephants and buffaloes in both National Parks.

Name Sex Number Host Location

A. gemma Male 12 Elephant Tsavo National Park Female 1

Male 7 Buffaloes L. Nakuru National Park A. variegatum Female 0 Male 1 A. thollonia Elephant Tsavo National Park Female 0 Table 4.1: Ticks identified by morphological keys in the genus Amblyomma

Name Sex Number Host Location R. pulchellus Male 19 Elephant Tsavo National Park Female 5 Table 4.2: Ticks identified by morphological keys in the genus Rhipicephalus

Name Sex Numbers Host Location H. marginatum Male 7 Elephant Tsavo National rufipes Female 0 Park H. truncatum Male 15 Elephant Tsavo National Female 2 Park H.dromedarii Male 2 Elephant Tsavo National Female 1 Park H. albiparmatum Male 5 Elephant Tsavo National Female 0 Park

Table 4.3: Ticks identified by morphological keys in the genus Hyalomma

Name Sex Number Host Location D. rhinocerinus Male 3 Rhinos L. Nakuru National Female 0 Park

Table 4.4: Ticks identified by morphological keys in the genus Dermacentor

Species from the genera were identified morphologically and the features used are shown in the plates below. 27

Genus: Rhipicephlus

X80

Plate 1: R. pulchellus (female); A Dorsal region; (1) Ivory white scutum; (2) Sparse punctuation: B Ventral region; (1) U-shaped genital aperture

X80

Plate 2: Rhipicephalus pulchellus (male) A Dorsal region: (1). Flat eyes (2)Punctuation on the lateral grooves (3).Festoons and (4) a caudal appendage at the central festoon (5).Enamel ornamentation- white enamel in a dark background; B (1) Adanal plates that curve posteriorly inwardly

Genus Amblyomma

X80 Plate 3: Amblyomma gemma (male); A Dorsal region: (1) broad posteriomedian stripe; (2) 6 out of 11 festoons enamelled; (3) elongated ornamented mesial region; (4) slightly convex eyes; B Ventral region (1) Genital aperture (2) Anal groove posterior to the anus

X80 Plate 4: Amblyomma gemma (female); A Dorsal region (1) Straight scutum sides (2) small to medium punctuations (3) broad posterior angle of scutum; B Ventral region (1) V-shape posterior lips of genital aperture

X80 Plate 5: Amblyomma variegatum(male) A dorsal region; (1) non-enamelled festoons (2) narrow posteriomedian stripe B ventral region; (1)Anus 30

X80 Plate 6: Amblyomma tholloni (female) A Dorsal region (1) inornate scutum (2) non-enamelled festoons B Ventral region (1) Anus

Genus Hyalomma

X80

Plate 7: Hyalomma dromedarii (female) A Dorsal region: (1) slightly sinous posterior margin of the scutum (2) deep scapular grooves B Ventral region: (1) V-shaped genital aperture

X80

Plate 8: Hyalomma dromedarii male; Dorsal region (1) Eyes (2) Dark and shiny conscutum (3) Legs have pale rings colouration (4) Festoons; paracentral festoons are separated anteriorly and central festoons is pale colured

X80

Plate 9: Hyalomma dromedarii; Ventral region (1) Anus (2) Accessory adanal plates (3) Sub-anal plates do not align with adanal plates (4) Adanal plates have round ends

X80

Plate 10: Hyalomma albiparmatum(male); A Ventral region (1) Squared adanal plates; B Dorsal region (2) central festoon is white forming a distinct parma

X80

Plate 11: Hyalomma truncatum (male); Dorsal region (1) Dark and shiny conscutum (2) Pale rings on the legs (3) Festoons; and caudal depression

X80

Plate 12: Hyalomma truncatum (male);Ventral region (1) Sub-anal plates (2) Adanal plates with squared-ends (3) Anus

X80

Plate 13: Hyalomma marginatum rufipes A Dorsal region (1) Legs with pale rings (2) scutum rugged large punctuations (3) short lateral grooves (4) presence of paracentral festoons; B Ventral region (1) adanal plates, length twice the width and the distance separating the pates is less compared to the width of their ends

Genus Dermacentor

X80 Plate 14: Dermacentor rhinocerinus (male) (A) Dorsal region (1) Festoons (2) shiny ornamented conscutum; B Ventral region (1) Anus

4.1.1 Unidentified ticks A total of 18 tick species could not be identified morphologically as some were damaged and as such lost some parts that could help in identifying the species.

Still, other had morphological changes that had occurred after feeding making it difficult to identify the species correctly.

4.2 Identification of Ticks using PCR and gel electrophoresis DNA was isolated from 28 tick samples and after partial amplification of the PCR products, 22 samples were positive and yielded products of approximately 900-

1200 bp. However, morphologically identified A. tholloni and A. gemma did not give bright bands and as such discarded. Plate 4.2 shows the bands obtained after electrophoresis and the expected fragment sizes.

Plate 4.2: Sample agarose gel amplification of PCR products of ITS 2 region of ticks product size approximately 900-1500 bp; L- Loading Dye (molecular weight marker = 1kb) 37

4.1.2 Phylogenetic analysis The edited sequences were blasted in the GenBank and identified by comparing the sequences with the related sequences of the ITS 2 gene

(http://blast.ncbi.nlm.nih.gov/Blast.cgi). Sequences that showed the highest percentage similarity value were considered as the closest matches to the sequences in this study, and were downloaded from the database for Phylogenetic analyses. The study sequences were assigned names with highest species from the

GenBank. Table 4.5 shows the identified species, base pairs size and similarity with species deposited in the GenBank.

Different species could be discriminated from the ITS 2 fragment size. The fragment size markers of R. pulchellus was estimated at 1100 bp, A. variegatum

1200 bp, H. marginatum 1500bp and H. dromedarii 1200 bp. A comparison of the

ITS 2 sequence identified in this study with the sequence deposited in the

GenBank registered different homology with different genera. From the data, 6 samples (7s, 8s, 47s, 62s, 64s, and 80s) showed 98-100% homology corresponding to R. pulchellus sequences obtained from Australia. Another 7 samples (32s, 37s, 38s, 46s, 54s, 71s and 73s) showed 98% homology with A. variegatum sequences obtained from the U.S.A. Samples 31s and 43s showed

92% homology with H. dromedarii while sample 79s showed 91% homology with ITS2 sequence of A. humerali from Brazil in the GenBank.

Genus Accession Tick species ITS % Reference Country Number fragment Identity sequence Size

Rhipicephalus KM819710 R.pulchellus 1100 100 AF271275 Australia KM819710 R.puchellus 1100 100 AF271275 Australia KM819710 R.pulchellus 1200 98 AF271275 Australia KM819710 R.pulchellus 1200 98 AF271275 Australia KM819710 R.pulchellus 1100 99 AF271275 Australia KM819710 R.pulchellus 1100 99 AF271275 Australia Amblyomma KM819712 A.variegatum 1200 98 HQ856759 U.S.A KM819712 A.variegatum 1200 98 HQ856759 U.S.A KM819712 A.variegatum 1200 98 HQ856759 U.S.A KM819712 A.variegatum 1200 98 HQ856759 U.S.A KM819712 A.variegatum 1200 97 HQ856759 U.S.A KM819712 A.variegatum 1200 98 HQ856759 U.S.A KM819712 A.variegatum 1200 98 HQ856759 U.S.A A.humerali 800 91 AY887111 Brazil Hyalomma KM819713 H. marginatum 1500 95 JQ737104 China rufipes KM819713 H. marginatum 1500 95 JQ737104 China rufipes KM819713 H. marginatum 1500 95 JQ737104 China rufipes KM819713 H. marginatum 1500 95 JQ737104 China rufipes H. marginatum 1600 95 JQ737104 China rufipes H. marginatum 1600 95 JQ737104 China rufipes KM819711 H.dromedarii 1200 95 JQ733570 India KM819711 H.dromedarii 1200 92 JQ733570 India

Table 4.5: Identity of tick species and their bp sizes, and percentage similarity value with the references sequences

Figure 4.1 shows the alignment of the query (R. pulchellus haplotype) and the reference sequence (Sbjct) from the GenBank; the query fully aligned to the reference sequence of R. pulchellus. A. variegatum haplotype (Figure 4.2) showed

98% sequence alignments the reference sample. The alignment had 7 nucleotide mismatches and extra nucleotides indicated by a gap in the sequence. Figure 4.3 shows the alignment of H. marginatum rufipes haplotype and the reference sequence from the GenBank and it had several nucleotide mismatches and nucleotide insertion.

Maximum Score E Value Max. Accession Identification

1024 0.0 100% AF271275

Figure 4.1: GenBank blast search for R. pulchellus haplotype. Query represents haplotype while Sbjct represent R. pulchellus (AF271275). The query fully aligned with thereference sequence. 41

Max Score E. value Max. Accession Identification

1118 0.0 98 HQ856803

Figure 4.2: GenBank blast search for A. variegatum haplotype. Query represents A. variegatum while Sbjct represent A. variegatum (HQ856759). The query nearly fully aligned to the reference sequence; few mismatches (showed with arrows) and insertion can be observed

Maximum Score E Value Max. Accession Identification

776 0.0 95% JQ737104

Figure 4.3: GenBank blast search for H. marginatum rufipes haplotype. Query represents H. marginatum rufipes haplotype while Sbjct represent H. marginatum rufipes (JQ737104). Nucleotide insertions and mismatches can be observed in the alignment (showed with arrows).

Max score E value Max. Accession Identification

693 00 92 JQ733570

Figure 4.4: Figure 4.2 GenBank blast search for sample 43 (H. dromedarii); Query represents sample 43 while Sbjct represents H. dromedarii (JQ733570). Nucleotide insertions and mismatches can be observed in the alignment (showed with arrows).

Sequence from this study and reference sequence obtained from the GenBank were aligned using multiple alignment program Clustal X. Maximum Composite

Likelihood method was used in computing the evolutionary distance. Figure 4.5 shows the Phylogenetic tree generated from partial ITS sequence data. The branch length represents evolutionary changes that have taken place over time and the amount of genetic change is represented by a scale of 0.1. The number of substitution which is related to the clustering together of the taxa as a bootstrap test, is represented by a percentage value; that is, the number of substitution per

100 nucleotide sites and is shown above the branches. The tree was rooted to the genus Ixodes.

From the figure, R. pulchellus and A. variegatum haplotypes clustered together at their respective group with a high bootstrap value of 99%. H. marginatum rufipes also clustered together with the reference sequence while sample 39s and 40s which shared recent ancestor origin with H. anatolicum with a bootstrap support of 79%. H. dromedarii clustered together with the reference H. dromedarii ITS 2 sequence deposited in the GenBank with a high bootstrap (99%) while A. humerali is clustered together with A. tuberculatum than A. humerale with a low bootstrap support of 51%. Oxidi dammini was used as an outgroup making the tree be a rooted one.

Figure 4.5: Sequence from the study and reference sequence obtained from the GenBank aligned using multiple alignment programs Clustal X. Evolutionary distance computed using the Maximum Composite Likelihood. The amount of genetic change is represented by a scale of 0.1. The number of substitution is represented by a percentage value; that is, the number of substitution per 100 nucleotide sites and is shown above the branches. The reference sequences are in bold. The tree was rooted to the genus Ixodes 46

4.3 Morphological and molecular characterization of tick samples The number of ticks identified using each technique were compared. Table 4.6 shows the number of tick samples identified using phenotypic and phylogenetic techniques.

Species No. identified No. identified genetically Morphologically

R. pulchellus 6 6

H.marginatum rufipes 4 6

H.truncatum 1 0

H.dromedarii 2 2

H. albiparmatum 1 0

A. variegatum 7 7

A. humerali 0 1

D. rhinocerinus 1 0

Morphology Genetic

Pearson Correlation 1 0.941**

Morphology Sig. (2-tailed) .000

N 8 8

Pearson Correlation 0.941** 1

Genetic Sig. (2-tailed) .000

N 8 8

**. Correlation is significant at the 0.01 level.

Table 4.6: Number of ticks identified using both techniques and the correlation coefficient 47

The number of R. pulchellus, A. variegatum and H. dromedarii identified both morphologically and genetically were the same; that is, 6, 7 and 22 respectively while 4 species were identified H. marginatum rufipes morphologically and 6 genetically. H. truncatum and H. albiparmatum were each identified morphological but were not identified using genetic techniques. A. humerali was identified genetically while D. rhinocerinus was identified morphologically.

Congruency between morphological and genetic traits was high as depicted by correlation co-efficient of 0.941 (P=0.01)

CHAPTER FIVE

DISCUSSION, CONCLUSIONS AND RECOMMENDATIONS

5.1 Discussion Ticks are ubiquitous blood feeding ectoparasites of humans and animals and therefore vectors of a plethora of pathogens that cause severe infectious diseases in humans and livestock (Lawrence et al., 2004). Various genera are associated with various diseases, for example Rhipicephalus spp. are known vectors of

Crimean-Congo haemorrhagic fever virus while Amblyomma spp. are associated with viruses noted to be mosquito-borne (Lwande et al., 2013). More importantly, the impact of ticks on human health is on the rise and more novel microbial associations have continued to be described (Jasinkas et al., 2007). Wild herbivores are considered to be the most important reservoirs for tick-borne pathogens that affecting humans and animals as they support a large populaton of ticks species (Peter et al., 1998). There is need therefore for accurate identification of ticks in order to develop better control measures.

In this study, ticks collected from wild herbivores in L. Nakuru and Tsavo

National Park were identified both morphologically and genetically.

Morphologically, Amblyomma spp. and R. pulchellus were easily identified using

Walker’s (2013) identification keys. Male R. pulchellus were easily identified by their unique striped enamel on a brown-dark background over their conscutum as 49

described by Beati & Keirans (2001) who observed that male R. pulchellus have characters that are useful for classification and easily distinguished. The stripes clearly distinguished them from R. humeralis that are also characterized by the white spots on the scapulae. Female R. pulchellus were easily identified by their association with the males (Walkers et al., 2013) and through their fine and sparse large punctuation on the scutum and U-shaped genital aperture. Amblyomma spp. were identified by ornamentation, long mouths and festoons in line with Piazark

(2005).

It was difficult to differentiate some Male H. truncatum from male H. marginatum rufipes. The distinctive features that differentiate the two species, the caudal depression present on H. truncatum and absent in H. marginatum rufipes; and dense even punctuation on H. marginatum rufipes. However, some species identified as H. truncatum had morphology of H. marginatum; that is, despite the presence of caudal depression, some species identified as H. truncatum had dense even covering of punctuations on their conscutum, a feature of H. marginatum rufipes. As such, it was difficult using Walker’s identification to distinguish between H. truncatum and H. marginatum rufipes, though the offspring were regarded as H.truncatum. These observations are similar to other studies which have reported difficulties in identifying Hyalomma spp using nomenclatures

(Abdigourdarzi et al., 2011). This is attributed to natural hybridization in the genus Hyalomma (Rees et al., 2003). The authors reported that initial hybridization between H. marginatum rufipes and H. truncatum resulted in a hybrid with more morphological characteristic of H. truncatum. Similar 50

observations were made by Hadani & Cwilich (1963) who noted that hybridization of H. excavatum and H. marginatum resulted to male offpsprings with morphological characterization of both H. margiatum and H. excavatum. In this study, difficulties observed in differentiating Hyalomma spp. may also be attributed to such natural hybridization between the two species in the wild.

Various tick samples could not be identified due to lost body parts and physical damages making it difficult to identify them morphologically. For example, in the genus Amblyomma, some tick samples had partially lost the conscutum tampering with the ornamentation, making it difficult to identify a problem observed by Abdigoudarzi et al. (2011) who noted that morphological changes that occur on ticks’ surface body may result to misleading identification. In their studies, some damaged ticks could not be easily identified morphologically due to decomposition but using phylogenetic analysis, there were able to identify the tick species.

Recent molecular work on tick species has used the ITS 2 as a marker in identifying ticks especially in differentiating closely related genus (Hillis &

Dixon, 1991; Rumer et al. 2011). Similar observations were made by Lv et al.

(2014) who showed that ITS 2 has the highest interspecific divergence and a low intraspecific variation compared to other markers and therefore, concluded that

ITS 2 may be the most useful DNA marker for discriminating tick species.

Amplification of the entire ITS 2 have been done by various reserachers and mixed results were observed including Zahler et al. (1995) and Murell et al. 51

(2001) who attributed the usatisfactory to specific nature of the ITS 2 region in ticks and some repeated fragments in the region. In their study Abdigourdarzi et al. (2011) noted that partial amplification of the region is good in discrimaniting different species of hard ticks; however, the authors asserted that it is important to have a complete sequence of ITS 2 region in order to be able to resolve issues related to partial sequences that were seen in their study.

In this study, partial sequence of the ITS 2 region was used to identify tick samples and different genera were differentiated based on the fragment sizes. The

Rhipicephalus spp. was estimated at 1100 bp and Amblyomma spp., 1200 and

Hyalomma spp., at 1500 bp. The results are similar to findings by Abdigourdarzi et al. (2011) who reported similar sizes for the three genera.

Sampled ticks were compared and identified with the sequences from the

GenBank, and R. pulchellus and A. variegatum had a percentage similarity ranging between 98-100%. This suggested that both species could be regarded according to the genotypic identification given that in the phylogenetic tree, both species fell in their respective monophyletic group, clustering together, with a high bootstrap of 99%. The low percentage similarity value (92-95%) shown by

Hyalomma spp. to the corresponding accession sequences in the GenBank could be due to diverse evolution as a result of geographical separations (Taberlet et al.,

1997). The difference could be attributed to single nuclueotide substitution between both sequences due to individual variation as a result of their wide range of distribution; an observation that is in agreement with other studies which 52

showed that sequence divergence may be due to phylogeographical units in a given species (Avise & Walker, 1999).

This was also noted in H. dromedarii, with a low similarity value (92%) while the study sequence clustered together with the reference sequence with a high bootstrap of 99%. Such similarity value could be associated with the cryptic hybridization factor which according to Rees et al. (2003) results to nucleotide substitution. Two samples considered H. marginatum rufipes failed to cluster together with the rest of the species in the phylogenetic tree. Phenotypically, the tick samples were identified as H. truncatum and H. albiparmatum; and therefore, the high dissimilarity value and failure to cluster together with the rest of H. marginatum rufipes could be attributed to lack of corresponding sequences in the

GenBank. This could also be the case in A. humerale which had been identified as

D. rhinocerinus phenotypically.

In this study, congruency between morphology and genetic traits was high as the correlation coefficient was high at 0.942. Congruency was observed in

Rhipicephalus and Amblyomma spp. as they had samples matching between morphological and genetical traits. Similar findings have been reported by Lu et al. (2013) who observed consistency in their morphological and molecular characteristic of identifying tick species in Japan. Inconsistency was observed in genus Hyalomma and this could be attributed to lack of or few correpsonding sequences deposited in the GenBank. It could also have been contributed by 53

several nucleotides mismatches and insertion which could be as a result of geographical separation and hybridization among the species.

5.2 Conclusions

From the findings of the study,

 Rhipicephalus spp. and Amblyomma spp. were easily identified

using morphologically characteristics

 Some H. marginatum rufipes and H. truncatum could not be easily

identified morphologically.

 Partial amplification of ITS 2 was successful in differentiating tick

species

 Rhipicephalus spp. and Amblyomma spp. showed high percentage

similarity value with their corresponding sequences from the

GenBank (98-100%) while Hyalomma spp. showed low percentage

similarity value with the corresponding sequences deposited in the

GenBank (92-95%)

 Congruency between morphological and genetic traits was high as

depicted by correlation coeffienct of 0.941 (P = 0.01)

Therefore, I concluded that morphological and genetic taxonomic traits can be used to identify tick species collected from wild herbivores in Kenya. 54

5.3 Recommendations

From the findings it is recommended that;

 The ITS2 is a good loci for identification od ticks in Kenya and the

sequences deposited in the NCBI could be used

 For correct identication of engorged, damage, closely related species and

ticks in sub-adult stages molecular markers such as ITS should be used.

5.4 Further Studies  A wide collection of Ixodidae from Kenya should be added to contribute

to the existing ITS nucleotide database in order to have sufficient genetic

database to cover tick species

 More studies to be done on cryptic hybridization of genus Hyalomma

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APPENDICES

APPENDIX 1 A work sheet for DNA analysis, partial sequence of ITS 2

Reagents

MyTAq HS DNA Polymerase

5XMyTaq Reaction Buffer

7.73 nmoles C452E

10.9 nmoles C452F

DNA- extracted from ticks

Equipment

0.2 ml PCR tubes- Applied Biosystems

1.5 ml microcentrfuge tubes

Thermocycler (Applied Biosystem verity 96 well thermal cycler)

Amplification Reaction Mix

Reagent Volume dH2O 15.8 µl

5X My Taq reaction Buffer 5.0 µl

C452E (Forward Primer) 1.0 µl

C452F (Reverse Primer) 1.0 µl

My Taq Reaction Buffer 0.2 µl

Template DNA 2.0 µl

Total 25 µl

APPENDIX 2 Key to Genera

1. Scutum with no ornamentation or eyes; absent of festoons, anal groove surrounding the anus anterior...... Ixodes -Festoons present; eyes present; anus with an anal groove posteriorly...... 2 2. Without ornamentation and eyes...... Haemphysalis -Scutum ornamented and eyes presents...... 3 3. Ornamented scutum and basis capitulum hexagonal shape from a dorsal view.....Rhipicephalus -Scutum ornamented including the festoons and basis capitulum subrectangular in shape...... 4 4. Long mouth parts and festoons ornamented and large in size; legs with pale rings...... Amblyomma -Basis capitulum subrectangulared and scutum ornamented or not.....5 5. Short and wide palps and scutum ornamented...... Dermacentor -Scutum, inornamented and loner palps than wider.....Hyalomma

Key to Ixodes females

1. Ixodes canisuga Dorsal region: Shorter palp compared to the width of the basis capitulum; scutum is narrowly rounded at the posterior Ventral region: Genital aperture located between coxae II 2. Ixodes ricinus Dorsal region: Scutum widely rounded at the posterior position Ventral region: Genital aperture is located between coxae IV 3. Ixodes hexagonus Dorsal region: Scutum is hexagonal shaped Ventral region: genital aperture located between coxae III 64

Key to Ixodes male

4. Ixodes canisuga Dorsal region: Coxa I has an internal spur Ventral region: Median plate broad posteriorly but narrow anteriorly; long epimeral and adanal plates 5. Ixodes ricinus Dorsal region: Coxae II-IV with vestigial internal spurs Ventral region: long pregenitial and media plates 6. Ixodes hexagonus Dorsal region: Coxae II-IV with absent internal spurs Ventral region: Long median plate and hexagonal shaped pregenital plate

Key to Rhipicephalus males

1. Rhipicephalus pulchellus Dorsal region: Small species (1.9-2.2mm); fine and large punctuations scattered in the scutum Ventral region: Narrow adanal plates that curve posteriorly inwardly 2. Rhipicephalus bursa Dorsal region: Large species (3-4 mm); fine and numerous punctuations on the scutum Ventral: Large adanal plate which are subtriangular shaped with wide posterior; broad spiracles 3. Rhipicephalus sanguineus Dorsal region: Medium in size (2-3.2mm); fine to large punctuation of the scutum; regular lows of large punctuation Ventral region: triangulared an elongated adanal plates; with wide posterior

Key to Rhipicephalus females 65

1. Rhipicephalus pulchellus Dorsal region: Small species (2-2.4mm); fine and sparse large punctuation on the conscutum Ventral region: U-shaped genital aperture 2. Rhipicephalus bursa Dorsal region: Large species (3-4mm); conscutum with large sparse and fine punctuation Ventral region: V-shaped genital aperture 3. Rhipicephalus sanguineus Dorsal region: medium in size (3-3.5mm); fine and large sparse punctuation on the conscutum Ventral region: U-shaped genital aperture

Key to Hyalomma males

1. Hyalomma marginatum marginatum Dorsal region: smooth scutum with fine and regular and to some extent large punctuations; caudal depression; central browned coloured festoon; lack dense punctuations Ventral region: Adanal plates, length twice the width; the distance separating the plates are equal to the width of the ends 2. Hyalomma marginatum rufipes Dorsal region: Scutum rugged large punctuations; short lateral grooves; presence of paracentral festoons Ventral region: Adanal plates, length twice the width and the distance separating the pates is less compared to the width of their ends 3. Hyalomma truncatum Dorsal region: Dark and shiny conscutum; caudal depression Ventral region: Square ends adanal plates; subanal plate align with adanal plates 4. Hyalomma arbiparmatum 66

Dorsal region: Central festoon as a white parma 5. Hyalomma dromedarii Dorsal region: Central festoons paled coloured Ventral region: Sub-anal plates do not align with adanal plates; Adanal plates have round ends

Key to Hyalomma females

1. Hyalomma marginatum marginatum Ventral region: Broad U-shaped genital aperture 2. Hyalomma marginatum rufipes Dorsal region: Deep scapular grooves; dense punctuation on the scutum Ventral region: Broad V-shaped genital aperture 3. Hyalomma truncatum Ventral region: Broad U shaped genital aperture 4. Hyalomma dromedarii Dorsal region: Slightly sinous posterior margin of the scutum Ventral region: V –shaped posterior lip

Key to Amblyomma Males

1. Amblyomma gemma Dorsal region: 6 out of 11 festoons enamelled; broad posteriomedian; elongated ornamented mesial region; slightly convex eyes 2. Amblyomma variegatum Dorsal region: Convex eyes; no enamel on festoons; narrow posteriomedian does not touch 3. Amblyomma lepidium Dorsal region: Narrow posteromedian stripe; two outermost and central festoons no enamel 4. Amblyomma dromedarii Dorsal region: Dark conscutum 67

Ventral region: Sub-anal are outside in alignment with the adanal plates

Key to Amblyomma females

5. Amblyomma gemma Dorsal region: Straight scutum sides; small to medium punctuations; broad posterior angle of scutum Ventral region: Narrow v-shaped genital aperture 6. Amblyomma variegatum Dorsal region: Convex eyes Ventral region: Wide U-shaped genital aperture 7. Amblyomma lepidium Dorsal region: Large conspicuous punctuation; small mesial region; posterior angle of the scutum is narrow Ventral region: V-shaped genital aperture