DETECTION OF CRIMEAN-CONGO HAEMORRHAGIC FEVER VIRUS 

AND RICKETTSIA spp. IN TICKS AND DISEASE EXPOSURE OF 

LIVESTOCK HANDLERS IN THE GREATER ACCRA AND UPPER EAST 

REGIONS OF GHANA 

 

 

 

 

BY 

RONALD ESSAH BENTIL 

(10459351) 

 

 

 

This thesis is submitted to the University of Ghana, Legon in partial fulfilment of the 

requirement for the award of MPHIL in APPLIED PARASITOLOGY Degree 

 

 

 

OCTOBER  2022

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 ii 

DECLARATION 

I do hereby declare that this thesis is my own work, undertaken under the supervision of the 

names listed below and has not been previously submitted either partially or wholly for the 

award of a degree in any University. References to the works of other investigators have been 

duly acknowledged.  

                              28th October 2022 

Ronald Essah Bentil       Date 

(Student) 

 

 

__________________                                   28th October 2022 

Daniel Oduro (PhD)                 Date 

(Supervisor) 

 

___________________          28th October 2022 

Samuel Dadzie (PhD)                  Date 

(Co-Supervisor) 

 

_______________________                          28th October 2022 

Shirley C. Nimo-Paintsil (PhD)     Date 

(Co-Supervisor) 

  

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DEDICATION 

This work is dedicated to God almighty, my parents (Mr. Evans Korsah Bentil and Mrs. Mercy 

Welhemina Bentil) and sister (Joanita Abokoma Bentil). I can never thank you enough for 

everything you have done for me. 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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ACKNOWLEDGEMENT 

I am thankful to the Almighty God for His mercy and favour in bringing me thus far. 

I also wish to extend my sincere thanks to the Noguchi Memorial Institute for Medical Research 

for the opportunity to undertake my research in such a renowned and excellent institute; as well 

as the US Naval Medical Research Unit-3 for support in funding my work. 

My deepest gratitude goes to my supervisors, Dr. Daniel Oduro, Dr. Samuel Dadzie and Dr. 

Shirley C. Nimo-Paintsil for the mentorship and guidance throughout my study. 

I am thankful to Dr. Naiki Attram, for the immense support and contribution. God richly bless 

you.  

To Dr. Daniel Kojo Arhinful, I really appreciate your mentorship and encouragement from a 

distance and leading me this path of science. I remain indebted to you.  

I render heartfelt thanks to my colleagues; Seth Addo, Mba-tihssommah Mosore, Janice Tagoe, 

Clara Yeboah, Selassie Kumordjie, Bright Agbodzi, Eric Behene, Julian Adinkrah, Karen 

Ocansey, Victoria Attoh and Rachel Owusu-Abrokwah for their assistance every step of the 

way. 

I thank the faculty of the Department of Animal Biology and Conservation Science, University 

of Ghana for the robust training given me on this program. 

 

 

 

 

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ABSTRACT 

 

Ticks are important blood-sucking arthropod vectors. They are known to spread a wide range 

of diseases that cause severe and life-threatening illnesses in humans and animals all over the 

world through feeding. In population-dense and trade-dominant areas such as the Greater Accra 

and Upper East Regions, the risk of zoonotic infections may be on the rise. This study, 

therefore, sought to identify the circulating tick species, assess the pathogens they carry and 

the risk of exposure of primary animal handlers. 

A total of 705 ticks were collected from cattle (n=188) and horses (n=11). Three tick genera 

(Hyalomma, Amblyomma and Rhipicephalus) were observed in the study with the predominant 

species being Hyalomma rufipes (n=290, 41.13%), followed by Amblyomma variegatum 

(n=157, 22.27%) and the least, Rhipicephalus sanguineus (n=1, 0.14%). It was also observed 

that the preferred point of attachment of all identified ticks was predominantly the Anal region 

(n=469, 68.37%) with the least site being the Abdominal region (n=5, 0.73%) for cattle except 

for Horses that had the least site being the chest (n=1, 5.26%).   

Out of the 705 tick samples, it was observed that in the Upper East Region, higher percentage 

of ticks collected (62.67%) were infected with Rickettsia as compared to ticks from Greater 

Accra (42.13%). Similarly, slightly more ticks were infected with Rickettsia africae in the 

Upper East Region (91.8%) than Greater Accra (91.38%). In addition, although Amblyomma 

variegatum was the second most prevalent species identified, the highest Rickettsia and 

Rickettsia africae infection rates were recorded in this species. The highest CCHFV infection 

rates were observed in Hyalomma rufipes.  

Serology carried out for a total of 120 human sera samples, for Rickettsia using an in-house 

ELISA revealed that 27.50% had been exposed to Spotted Fever Group (SFG) and 4.17% to 

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Typhus Group (TG) and none exposed to the Scrub Typhus group (STG). Furthermore, 

preliminary analysis for antibodies to CCHFV IgG was detected in 42.5% of the human serum 

(n=120) samples pending confirmatory Plaque Reduction Neutralization Test (PRNT).   

This study further reports the first whole-genome sequencing of CCHFV in tick species within 

Ghana with sequence analysis revealing genotype III (Africa III) may be circulating in the 

Upper East Region of Ghana. This suggests the possible importation of the CCHFV virus into 

the country through trade and thus puts livestock and humans who may have primary contact 

with livestock at risk of infection of these diseases.  

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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TABLE OF CONTENTS 

DECLARATION ....................................................................................................................... ii 

DEDICATION .......................................................................................................................... iii 

ACKNOWLEDGEMENT ........................................................................................................ iv 

ABSTRACT .............................................................................................................................. v 

LIST OF TABLES .................................................................................................................... xi 

LIST OF FIGURES ................................................................................................................. xii 

LIST OF ABBREVIATIONS ................................................................................................ xiii 

CHAPTER ONE ........................................................................................................................ 1 

GENERAL INTRODUCTION ................................................................................................. 1 

1.1 BACKGROUND ................................................................................................................. 1 

1.2 PROBLEM STATEMENT .................................................................................................. 5 

1.3 JUSTIFICATION ................................................................................................................ 7 

1.4 GENERAL OBJECTIVE .................................................................................................... 8 

1.4.1 SPECIFIC OBJECTIVES ................................................................................................. 8 

CHAPTER TWO ....................................................................................................................... 9 

LITERATURE REVIEW .......................................................................................................... 9 

2.1 Overview of ticks and their impact ...................................................................................... 9 

2.2 Tick Systematics and Biology ........................................................................................... 10 

2.2.1 Ticks of medical and veterinary importance .................................................................. 11 

2.3 Tick Behaviour .................................................................................................................. 15 

2.3.1 Host seeking ................................................................................................................... 15 

2.3.2. Feeding .......................................................................................................................... 16 

2.3.3 Disease transmission ....................................................................................................... 17 

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2.3.4 Tick-borne Zoonoses ...................................................................................................... 17 

2.3.4.1 Rickettsial Diseases ..................................................................................................... 18 

2.3.4.2 Spotted Fever Group (SFG) ......................................................................................... 18 

2.3.4.3 Typhus Group (TG) ..................................................................................................... 19 

2.3.4.4 Scrub Typhus Group (STG) ........................................................................................ 19 

2.3.4.5 Other Rickettsioses ...................................................................................................... 20 

2.3.4.6 Crimean Congo Haemorrhagic Fever (CCHFV) ......................................................... 20 

2.4 Crimean Congo Haemorrhagic Fever Virus (CCHFV) Genome ...................................... 21 

2.5 Strategies for tick control and Tick-borne Disease (TBD) prevention .............................. 22 

CHAPTER THREE ................................................................................................................. 23 

METHODOLOGY .................................................................................................................. 23 

3.1 Study sites .......................................................................................................................... 23 

3.2 Ethical clearance ................................................................................................................ 24 

3.3 Study Population ................................................................................................................ 24 

3.4 Sample collection .............................................................................................................. 24 

3.4.1 Sampling from Animals .................................................................................................. 24 

3.4.1.1 Inclusion/ Exclusion of Animals ................................................................................. 24 

3.4.1.2 Tick collection from Animals ...................................................................................... 24 

3.4.2 Sampling from Humans .................................................................................................. 25 

3.4.2.1 Inclusion criteria for human sampling ......................................................................... 25 

3.4.2.2 Exclusion criteria for human sampling ........................................................................ 25 

3.4.2.3 Community entry ......................................................................................................... 26 

3.4.2.4 Participant eligibility and informed consent procedure ............................................... 26 

3.4.2.5 Blood Collection from participants ............................................................................. 26 

3.4.2.5.1 Blood Sample Processing ......................................................................................... 27 

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3.5 Laboratory Testing ............................................................................................................ 27 

3.5.1 Morphological Identification of ticks ............................................................................. 27 

3.5.2 Extraction of total nucleic acid from ticks ...................................................................... 27 

3.5.3 Molecular detection of tick-borne pathogens ................................................................. 29 

3.5.3.1 Real-time PCR (RT-PCR) for detection of CCHFV ................................................... 29 

3.5.3.2 Real-time PCR (RT-PCR) for detection of Rickettsia species .................................... 29 

3.5.3.3 Realtime PCR (RT-PCR) for detection of Rickettsia africae ...................................... 30 

3.5.4 Serological analysis for antibodies against tick-borne pathogens .................................. 30 

3.5.4.1 Magpix Immunoassay for detection of CCHFV antigen ............................................. 30 

3.5.4.2 ELISA Assay to detect Group-specific antibodies against Rickettsiae ....................... 32 

3.5.5 Whole Genome Sequencing (WGS) for CCHFV PCR positives ................................... 32 

3.5.5.1 RNA PICO 6000 assay (for quantification and assessing quality of RNA for NGS 
library preparation) .................................................................................................................. 33 

3.5.5.2 First-strand synthesis ................................................................................................... 34 

3.5.5.3 Second strand synthesis ............................................................................................... 35 

3.5.5.4 Qubit quantification of cleaned pre-enriched cDNA ................................................... 35 

3.5.5.5 Tagmentation of Genomic DNA ................................................................................. 36 

3.5.5.6 Library enrichment ...................................................................................................... 37 

3.5.5.7 DNA high sensitivity assay (for Quantification of DNA library) ............................... 39 

3.5.5.8 Library qPCR ............................................................................................................... 40 

3.5.5.9 Library Pooling for loading ......................................................................................... 41 

3.5.5.10 Pool qPCR ................................................................................................................. 41 

3.5.5.11 Loading the Illumina iSeq sequencer ........................................................................ 42 

3.6 Data Analysis ..................................................................................................................... 42 

CHAPTER 4 ............................................................................................................................ 43 

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RESULTS ................................................................................................................................ 43 

4.1 Distribution of Tick species ............................................................................................... 43 

4.2 Preferred part of tick attachment on the animal host ......................................................... 44 

4.3 Prevalence of Rickettsia in tick species ............................................................................. 47 

4.4 Prevalence of CCHFV in tick species ............................................................................... 49 

4.5 Whole-genome sequencing and Phylogeny ....................................................................... 50 

4.6 Seroprevalence of Rickettsia and CCHFV in Human blood samples ............................... 52 

CHAPTER 5 ............................................................................................................................ 61 

DISCUSSION .......................................................................................................................... 61 

5.1 Distribution of tick species ................................................................................................ 61 

5.2 Tick-borne pathogens in sampled ticks ............................................................................. 62 

5.3 Phylogeny .......................................................................................................................... 63 

5.4 Seroprevalence of CCHFV and Rickettsia ........................................................................ 65 

5.5 Limitations ......................................................................................................................... 67 

5.6 Conclusions ....................................................................................................................... 67 

5.7 Recommendations ............................................................................................................. 68 

REFERENCES ........................................................................................................................ 69 

APPENDICES ......................................................................................................................... 92 

Appendix I: Flow chart for consenting Eligible Participants .................................................. 92 

Appendix II: Participant Questionnaire ................................................................................... 93 

Appendix III: Checklist for consenting ................................................................................... 96 

Appendix IV: Participant screening form ................................................................................ 96 

Appendix V: Animal vector questionnaire form ..................................................................... 97 

Appendix VI: Animal vector recording sheet .......................................................................... 98 

Appendix VII: Oligonucleotide primer sequences for Realtime PCR .................................... 99 

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LIST OF TABLES 

Table 1: Occurrence of tick species sampled from the various body parts of livestock….........45 

Table 2: Risk factor analysis of Ticks burden of livestock using Generalized mixed 

model………………………………………………………………………………………....46 

Table 3: Prevalence of Rickettsia and Rickettsia africae in sampled tick species……………..49 

Table 4: Prevalence of CCHFV in sampled tick species……………………………………...50 

Table 5: Demographic factors associated with exposure to CCHFV and Rickettsia……….....54 

Table 6: Risk factors associated with exposure to CCHFV and Rickettsia……………………55 

Table 7: Clinical symptoms associated with exposure to CCHFV and Rickettsia…………….58 

 

 

 

 

 

 

 

 

 

 

 

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LIST OF FIGURES 

Figure 1: Life Cycle of ticks of the Ixodidae family (hard ticks)…………………………….11 

Figure 2: Regional Map of Ghana showing study sites……………………………………….23 

Figure 3: Image demonstrating how ticks were taken off livestock…………………………...25 

Figure 4: Distribution of tick species from the study sites…………………………………….43 

Figure 5: Rickettsia detected in tick samples………………………………………………….48 

Figure 6: Rickettsia africae detected in tick samples………………………………………….48 

Figure 7:  Maximum likelihood phylogeny of CCHF viruses from different geographical 

origins based on the complete coding sequence of the S segment…………………………….51 

Figure 8: Seroprevalence of Rickettsia in human samples……………………………………53 

Figure 9: Seroprevalence of CCHFV in human samples……………………………………...53 

 

 

 

 

 

 

 

 

 

 

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LIST OF ABBREVIATIONS 

CCHFV  Crimean Congo Haemorrhagic Fever Virus 

TBDs   Tick-borne diseases 

SSA   Sub-Saharan Africa 

SAT   Saliva-activated transmission 

CDC   Centers for Disease Control and Prevention 

SFG   Spotted Fever Group 

TG   Typhus Group 

STG   Scrub Typhus Group 

RMSF   Rocky Mountain spotted fever 

MSF   Mediterranean Spotted Fever 

TIBOLA  Tick-Borne Lymphoadenopathy 

USSR   Union of Soviet Socialist Republics 

RNA   Ribonucleic Acid 

SSTs   Serum separator tubes 

PCR   Polymerase Chain Reaction 

RT-PCR  Real-time PCR 

PBS   Phosphate buffered saline 

PBST   Phosphate buffered saline with 0.02% Tween-20 

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PBST-SK  Phosphate buffered saline with 0.02% Tween-20 and 5% skim milk 

eBLT   enrichment Bead-Linked Transposomes 

TB1   tagmentation buffer 

ST2   stop tagment buffer 

TWB   tagment wash buffer 

EPM   Enhanced PCR Mix 

RSB   Resuspension Buffer 

EHB 2   Enrichment Hybridization Buffer 2 

SMB   Streptavidin Magnetic Beads 

EEW   Enhanced Enrichment wash 

EE1   Enrichment Elution Buffer 1 

ET2   elute target buffer 

PPC   PCR primer cocktail 

GLMM  Generalized Linear Mixed Model 

 

 

 

 

 

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CHAPTER ONE 

GENERAL INTRODUCTION 

1.1 BACKGROUND 

Ticks are obligate hematophagous Acari that parasitize mammals, birds and reptiles and 

occasionally bite humans (Parola, 2004).  Ticks are important disease-causing arthropod (De 

La Fuente et al., 2008; Liu & Bonnet, 2014) vectors known to spread a wide range of diseases 

such as protozoa, bacteria, and viruses which are known to cause severe and life-threatening 

illnesses in humans and animals all over the world (Dantas-Torres et al., 2012; Rosenberg et 

al., 2018). In warm and humid climates, they are known to be livestock's most important 

ectoparasites and are responsible for significant livestock economic losses (Kasaija et al., 

2021). Ticks are responsible for the majority of losses due to their potential to spread diseases 

of various aetiologies. 

These ectoparasites can have an impact on animal production and health, either directly through 

their bites or indirectly by the infectious agents they transfer. (Jaime et al., 2018). Feeding on 

their hosts involves the tick biting and pushing the hypostoma into the host's skin, causing 

injury to the epidermis and blood vessel rupture. Tick bites cause skin damage comprised of 

oedema, irritation, infiltration of cells, inflammation or hypersensitivity and a large amount of 

blood loss (Jaime et al., 2018). The animals are predisposed to reduce their food absorption 

and lose weight as a result of these injuries. Tick infestations can result in anaemia as a result 

of blood loss, a decrease in the quality of the hides, and severe dermatitis. (Jaime et al., 2018; 

Rajput et al., 2006).  

 

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Further, tick feeding stresses the host and reduces its immunological response, lowering 

productivity and resulting in meat and milk output losses. (L’hostis & Seegers, 2002; Leal et 

al., 2020; Peter et al., 2005).  These morbidities can lead to death in some situations; with the 

carcass's poor appearance diminishing its marketability. (Jaime et al., 2018). Examples of 

diseases in livestock include bovine babesiosis and anaplasmosis, east coast fever and 

cowdriosis (Blood & Radostits, 1989; Dinkisa, 2018; Marcelino et al., 2012). 

Ticks are the second most prevalent vectors of human infectious disease pathogens, after 

mosquitoes. (De La Fuente et al., 2008; Lawrie et al., 2004) and as pathogen vectors for 

animals (De La Fuente et al., 2008). Infection of ticks with microorganisms occur when they 

feed on hosts (either human or animal) that are infected and in turn further introduce these 

microorganisms into another host on a next blood meal (Jongejan & Uilenberg, 2004; Šimo et 

al., 2017). Ticks are thus known to play a major role when it comes to zoonotic disease 

transmission which contributes to high disease and death rates. Human tick-borne diseases 

zoonoses have been detected in both wild and domestic animals (Baneth, 2014).  

Globally, there have been a considerable increase in tick-borne diseases of epidemiologic 

importance (Rochlin & Toledo, 2020). These diseases include those caused by bacteria; 

Rickettsia, Lyme disease, Rocky Mountain spotted fever, Q-fever and by viruses; Crimean-

Congo Haemorrhagic fever (CCHF), Tick-borne encephalitis, and Powassan encephalitis. 

Other tick borne infections includes those caused by protozoans; Babesiosis (Sambri et al., 

2004).  

Although these vectors  are ubiquitous, they are diverse and more prevalent in the world's 

tropical and subtropical areas (Rehman et al., 2017). Ticks are divided into three groups, two 

of which are veterinary and medically important—Ixodidae (hard ticks) and Argasidae (soft 

ticks) (Rajput et al., 2006). The family Ixodidae, which encompasses most tick species, are 

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characterized by their pestiferous nature and the highest rate of transmitting diseases and 

disease-causing agents among the three (Leal et al., 2020). Ixodidae ticks comprise thirteen 

genera, seven of which are veterinary and medically important species. These are 

Amblyomma, subgenus Rhipicephalus (Boophilus), Rhipicephalus, Haemaphysalis, 

Hyalomma, Dermacentor and Ixodes (Wondimu & Bayu, 2021). The genera Hyalomma, 

Boophilus, Rhipicephalus, and Amblyomma are the most economically important ixodid ticks 

in tropical areas. (Rajput et al., 2006). 

Among the different tick-borne illnesses, CCHF was included on both the United States 

National Institutes of Allergy and Infectious Diseases priority A list as threat to national 

security and public health (Geddes, 2021) and the World Health Organization's priority list for 

research and development (World Health Organization, 2021).  CCHF is endemic throughout 

Africa and has a 40% case fatality rate (Shahhosseini et al., 2021). Depending on how the virus 

was contracted, the incubation period ranges between 1 to 13 days. Initial symptoms are minor, 

but they gradually get worse with time (Kaya et al., 2011). Although an effective vaccine is 

not yet available, Ribavirin (an antiviral medication) has been used in therapy and has been 

shown to reduce mortality (Soares-Weiser et al., 2010).   

Rickettsioses on the other hand are focal; with war, famine, overcrowding, and unhygienic 

circumstances being the main drivers of epidemics. (Wallace et al., 2002). This suggests that, 

in addition to some cases documented in advanced countries, it is an issue in underdeveloped 

and developing nations (Wallace et al., 2002). In humans, incubation can last up to two weeks 

with typical symptoms appearing as rashes that later evolve and progress into other 

manifestations (McCollough, 2018). With a fatality rate of 10 to 40 %, mortality has been 

found to rise with age and rarely deadly in children under the age of 10 (Woods, 2013). Despite 

the fact that some antibiotics have been effective at treating the condition, there is currently no 

licensed vaccination for use, which adds to the issue (McCollough, 2018).   

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In general, there are two types of detection methods: direct and indirect (Springer et al., 2021). 

In the former (direct), patient samples are used for the direct detection of diseases carried by 

ticks using techniques like microscopy and culture. Microscopy is less sensitive, despite being 

quick and inexpensive. This method cannot be utilized for all tick-borne infections since it 

depends on the parasitaemia level and makes it difficult to distinguish between various species 

(Ord & Lobo, 2015). Additionally, culturing typically takes a long period, is difficult, requires 

a lot of work, and requires specific biosafety conditions. (Eldin et al., 2019). As a result, nucleic 

acid amplifications using molecular techniques are the most recommended since they have 

higher sensitivity, produce results quickly, and improve diagnostic effectiveness (Korber et al., 

2017). On the other hand, indirect detection procedures include serological testing techniques 

based on immunological targets and markers (Portillo et al., 2017).  

The difficulty in diagnosis is a characteristic similarity between the two above-mentioned tick-

borne diseases (Stewart & Stewart, 2021; Tezer & Polat, 2015). Serologic assays and 

polymerase chain reaction (PCR) are two recommended diagnostic procedures that are efficient 

(Stewart & Stewart, 2021; Tezer & Polat, 2015). However, in resource-constrained 

environments, the accessibility, price, and skill required to perform these assays continue to be 

a problem (Walker, 2007).  As a result, incorrectly administering unsuitable medication and 

misdiagnosis may occur, which may lead to the persistence of diseases in the general 

population.  

Several approaches have been made to control ticks, the primary of these is chemical control 

with acaricides. However, this has proven futile because ticks have developed resistance 

against a range of acaricides. In addition, besides being expensive, these chemicals are toxic 

and leave residues in animal products which are in turn harmful to humans upon consumption. 

It has, however, become necessary that control methods that are cost-effective, 

environmentally friendly and pose a minimal adverse effect on the health of livestock and 

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human beings be implemented. Therefore this calls for Integrated Tick Management involving 

novel approaches of pest control, active surveillance and improved testing to mitigate the effect 

of the problem posed (George et al., 2004; Nath et al., 2018).  

 

1.2 PROBLEM STATEMENT 

In several countries across Sub-Saharan Africa, epidemiological data on tick-borne diseases 

are scarce, and their prevalence and distribution are largely less studied. The difficulty in 

identifying, managing tick populations,  and treating infections of tick-borne pathogens makes 

efforts to combat these emerging diseases futile (Kobayashi et al., 2017; Madison-Antenucci 

et al., 2020). In addition, similarities in the symptoms of tick-borne infections with other febrile 

illnesses, often leads to misdiagnosis and, an eventual increase in prevalence through 

transmission (Madison-Antenucci et al., 2020).   

Even though there are significant advances in research and diagnostic techniques such as 

molecular or immunological techniques, which could aid in the diagnosis of tick-borne 

diseases, these advancements remain in highly resourced facilities.  Underdiagnosis still remain 

in limited resource facilities, making the extent of the problem difficult to determine, despite 

these improvements.  The identification and selection of effective care and control methods 

include knowledge of the diseases that can be spread by ticks in particular locations. To deal 

with the problem, the capacity to detect, explain, and acknowledge the expanding public health 

hazard must evolve as new tick-transmitted illnesses are identified and appear in new 

geographic areas (Madison-Antenucci et al., 2020). 

Epidemiology and ecology of tick-borne diseases are influenced by dynamic interactions 

between biotic and abiotic components. Further research has confirmed that pathogens of 

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zoonotic nature as well as their vectors live in separate environments, giving rise to the idea of 

nidality in the natural world, or landscape epidemiology of infectious diseases. The distribution 

and number of ticks, as well as the development, revival, and geographic spread of tick-borne 

illnesses, are all influenced by the following factors: tick and tick-borne pathogen demography, 

micro and macro climate variations, travel, human behaviour, politics, land use and 

modification of habitats (for farming, housing, and entertainment purposes), population 

growth, economy and migration. (Baneth, 2014; Dantas-Torres, 2015). Tick-borne diseases are 

becoming increasingly important, and people are becoming more aware of their effects (Wikel, 

2018). 

Furthermore, since the last release of the list of ticks in Ghana and assessment of the 

distribution across various vegetation zones (Ntiamoa-Baidu, Carr-Saunders, Matthews, 

Preston, & Walker, 2004), it is uncertain if these tick species exist, have undergone adaptations, 

or new species have been introduced. Climate change, which has a significant impact on 

arthropod disease vectors and their capacity to spread diseases (Bett et al., 2019), as well as 

trading of livestock (Nimo-Paintsil et al., 2022) may be contributing factors. Additionally, 

given the incidence of disease outbreaks spread by ticks in neighbouring countries (Ehounoud 

et al., 2016; Nordstrand et al., 2007; Ouedraogo et al., 2021), and past research that has 

revealed the presence of certain tick-borne diseases in Ghana (Akuffo et al., 2016; Kobayashi 

et al., 2017; Sothmann et al., 2017), it is crucial that the impact of such events be evaluated in 

order to be able to promptly alleviate the repercussions. 

That said, due to the indiscriminate hematophagous arthropods, and can transmit infections 

when they feed (Gondard et al., 2017), livestock and domestic animals are at risk of infestation. 

In turn, this creates a health concern to people who take care of these animals and come into 

contact with them. As a result, there is a high chance of exposure should the ticks be harbouring 

any pathogens. 

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1.3 JUSTIFICATION 

Many countries in Sub-Saharan Africa lack epidemiological data on infections caused by ticks, 

and their incidence and spread remaining mostly unknown (Kobayashi et al., 2017). Similarly, 

in the few studies that have been performed in Ghana, some gaps have been identified that need 

further investigation. For example, findings from work done in Kumasi by Clarke et al. (2014) 

revealed the existence of tick-borne pathogens in ticks collected from dogs as well as the 

seroprevalence of these pathogens upon assaying sera collected from the domestic animals. In 

a study conducted by Akuffo et al., (2016), ticks were collected off slaughtered animals and 

blood drawn from animal handlers at the Kumasi abattoir. Study findings showed that some 

ticks carried the Crimean Congo Haemorrhagic fever Virus as well as detecting human 

exposure to the virus among the slaughterhouse workers. Another study conducted by 

Sothmann et al. (2017), further revealed Rickettsia felis infection of some febrile children in 

Ghana upon screening their blood samples. However, the source of exposure/ infection was 

not ascertained by the authors in this case. In addition, serological assays performed on blood 

samples collected from livestock in the Volta Region, Johnson et al. (2019) revealed the 

seroprevalence of Q-fever in their findings. 

Furthermore, although taxonomically reliable information on ticks in Ghana is limited, 

previous work done within the same areas and surrounding areas indicate the presence of 

diverse tick species infesting domestic ruminants and wild mammals within different 

vegetation zones in Ghana ( Walker & Koney, 1999; Ntiamoa-Baidu et al., 2004).    

The above information is an indication that ticks identified in the country can host some tick-

borne pathogens and thus humans who especially cater for or live near livestock are likely to 

be exposed. These studies, however, were localized to study sites and thus do not represent a 

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consistent occurrence of the situation in Ghana. Due to the extensive agricultural activities in 

the study areas, there will likely be more human activity, which will favour the spread of 

zoonoses and a rise in exposure to ticks and tick-borne pathogens in these expanding urban 

environments. It is therefore critical, to determine the prevalence of various tick species 

infesting livestock, as well as the risk burden they pose to their owners. This national 

surveillance system is essential for filling gaps in the data set by providing information on the 

prevalence of tick-borne pathogens and the level of exposure to them across the country. 

1.4 GENERAL OBJECTIVE 

The purpose of the study was to determine the presence of tick-borne pathogens in ticks and 

pathogen exposure among livestock handlers within selected study sites. 

1.4.1 SPECIFIC OBJECTIVES 

1. Identify tick species sampled from livestock within the selected ecozones. 

2. Determine the prevalence of tick-borne pathogens in ticks within the selected ecozones. 

3. Assess previous human exposure to CCHFV and Rickettsia through seroprevalence in 

livestock handlers at selected ecozones 

4. Determine by genomic analysis whether tick-borne pathogens are localised or emerging 

new pathogens     

 

 

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CHAPTER TWO 

LITERATURE REVIEW 

2.1 Overview of ticks and their impact 

Hematophagous ectoparasites are efficient vectors of diseases. Ticks fall within this category, 

derive nutrition through blood-feeding on vertebrates and are of medical and veterinary 

importance (Jongejan & Uilenberg, 2004; De la Fuente et al., 2017). Ticks have been 

implicated as the most important vectors of pathogens that cause disease in animals (both 

domestic and wild), and are responsible for more than 100,000 cases of illness in humans 

worldwide (Jose de La Fuente et al., 2008; Yu et al., 2015). They are involved in the 

transmission of a several pathogenic microorganisms such as Theileria spp., Babesia spp., 

Rickettsiae, Borrelia spp., Powassan, Crimean Congo Hemorrhagic fever and Dugbe viruses, 

compared to any other arthropod vector group (De La Fuente et al., 2008). As transmission of 

tick-borne infections to humans and the associated morbidity and mortality pose significant 

public health concerns, transmission in livestock is an additional major challenge in animal 

production especially in tropical and subtropical regions of the world (De La Fuente et al., 

2008; Dantas-Torres et al., 2012).  

In developing countries, tick-borne diseases have the greatest influence on the lives of 

resource-scarce animal husbandry communities (Jongejan & Uilenberg, 2004; Minjauw & 

McLeod, 2003; Perry et al., 2002). This is certainly relevant in parts of Sub-Saharan Africa, 

Asia, and Latin America, where animal products are in high demand. (Thornton, 2010). Ticks 

and tick-borne illnesses have co-evolved with a variety of wild animal hosts, which often 

coexist and serve as reservoir hosts for ticks and tick-borne pathogens in cattle, pets, and 

humans. (Cançado et al., 2013). Domestic livestock may become infected upon contact with 

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ticks that have fallen off these wild hosts, either because humans moved animals into tick-

infested areas, or because humans moved tick-infested cattle into previously uninfested areas.  

Most ticks have a preference for feeding on certain groups of wild animals, with some even 

being host-specific. Consequently, the number of species pertinent to domestic animals and/or 

humans is limited. Relatively, few species of ticks have successfully adapted to livestock or 

feed on a human subject, and these have developed into efficient vectors of a range of 

pathogenic microorganisms, while virtually all human tick-borne diseases are zoonoses (De la 

Fuente et al., 2017; De La Fuente et al., 2008). A tick species is considered a vector for a 

particular pathogen only if it feeds on an infectious vertebrate host; can acquire the pathogen 

during the blood meal; maintains the pathogen through one or more life stages; and further 

transmit to another host on the next blood meal. (De la Fuente et al., 2017; Kahl et al., 2009) 

2.2 Tick Systematics and Biology 

There are about 898 recognized tick species distributed among three families, namely 

Argasidae (soft-bodied ticks), comprising about 194 species; Ixodidae (hard-bodied ticks) 

comprising about 703 species; and monotypic Nuttalliellidae, made up of 1 species 

(Guglielmone, 2010; Estrada-Peña & De la Fuente, 2014). However, only Argasidae and 

Ixodidae are considered to be of veterinary and medical importance (Tahir et al., 2020). The 

family Ixodidae (hard ticks), which encompasses most tick species (about 80%), are pestiferous 

and have the highest rate of transmitting diseases and disease-causing agents among the three 

families (Leal et al., 2020). The vectorial capacity of Ixodid ticks is linked to their long-term 

co-evolution with the pathogens they transmit, their longevity, their high reproductive 

potential, the wide range of host feeding preferences of several species, and their ability to take 

in a substantial quantity of blood over a relatively long period (De la Fuente et al., 2017). 

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2.2.1 Ticks of medical and veterinary importance  

 

Figure 1: Life Cycle of ticks of the Ixodidae family (hard ticks) 

Source: (Varela-Stokes et al., 2009) 

Four stages exist in the life cycle of an ixodid tick, namely egg, larva, nymph and adult. Unlike 

in argasid ticks where mating occurs off the host in the vegetation, ixodid ticks mate while on 

their host (Horak et al., 2002). All post-embryonic stages of the tick require a blood meal to 

survive. Engorged females drop to the ground and lay eggs in the soil, after which larvae hatch 

from the batches of eggs after about four weeks and take about five days to fully engorge with 

blood. Nymphs may also take up to about twenty days to fully engorge with blood. Upon 

engorgement with blood, ticks detach from the host and fall onto the ground. Although male 

ticks feed, they do not engorge to become as large as the females would, which is attributed to 

the limitation in the quantity of blood imbibed due to the presence of the scutum that covers 

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the dorsum. The females, however, have smaller scutum thus become prominently distended 

when engorged (Cupp, 1991; Nyangiwe et al., 2018). 

Depending on the number of host animals they attach to during their life cycle, Ixodid ticks 

can have one, two, or three hosts. (Walker et al., 2003). The larvae and nymphs feed once to 

engorgement after which moulting occurs. For single-host ticks, moulting occurs two times on 

the same host, from the larval stage to nymph stage and then to the adult stage (that is larval, 

nymph and adult stages remain and feed on the same host and drop off only just before laying 

eggs) (Walker et al., 2003; Latif & Walker, 2004; Nyangiwe et al., 2018). Moulting happens 

only once on a host for two-host ticks, from larvae to nymph. After the engorged nymph falls 

off and moults from the host, the adult seeks out a new host, which may not necessarily be of 

the same species as the first. (that is, the larval and nymph stages feed on the same host, and 

adult stage on another) ( Walker et al., 2003; Latif & Walker, 2004; Nyangiwe et al., 2018). 

For three-host ticks, however, moulting does not occur on the host. The larva that is engorged 

falls off and moults into a nymph, which further engorges on a second host animal.  It falls off 

a second time to moult into the adult stage, which then attaches to a third host animal (Walker 

et al., 2003; Latif & Walker, 2004; Nyangiwe et al., 2018). Adult Ixodidae normally carry out 

mating on the host, after which the female feeds till it becomes  engorged, falls off, lays a huge 

number of eggs (about 1500 to 5000 eggs and dies (while males remain on the host for months) 

(Jongejan & Uilenberg, 2004). One-host ticks lay relatively fewer eggs than three-host ticks, 

as the latter are exposed to more risk in the process of finding a new host at each stage in their 

life cycle  (Jongejan & Uilenberg, 2004). Two-host ticks on the other hand are at less risk than 

species that have three hosts but more than one-host ticks, and also have egg batches that are 

intermediate with regards to size. 

The most important genera of hard-bodied ticks are Ixodes, Haemaphysalis, Dermacentor, 

Hyalomma, Rhipicephalus Amblyomma, and Boophilus. However, owing to the close 

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evolutionary and phylogenetic relationship between those of the genus Boophilus  to the genus 

Rhipicephalus, the former (Boophilus)  is used  as a subgenus of the latter (Rhipicephalus) 

(Barker & Murrell, 2002; Horak et al., 2002; Jongejan & Uilenberg, 2004). 

About 129 species of Amblyomma ticks exist. Characteristic of these ticks are mouthparts that 

are long and scuta that are beautifully coloured and ornamented. Although they have eyes, most 

species do not have them contained in sockets. Amblyomma species have  three hosts ticks in 

their life cycle and are common in (sub)tropical regions where they infest diverse mammalian 

hosts as well as reptiles and amphibians (Spickler, 2003). Birds may get infested with some 

undeveloped stages of some tick species and this contributes significantly to how the ticks are 

distributed. On the African continent, A. variegatum is the regarded as  a vital species because 

it has the widest distribution in Tropical Sub-Saharan Africa and is well-suited to domestic 

livestock. (Nabarro et al., 2020; Saari et al., 2019). Another important pest of livestock is A. 

hebraeum, which is found in Africa's south-eastern region (Jongejan et al., 2020). In addition 

to its prevalence, A. variegatum was introduced into the Caribbean region with cattle from 

West Africa, as far back as the 18th or 19th century. A. variegatum is thus, the only Amblyomma 

of African origin to have established itself successfully beyond Africa (Camus & Barré, 1990).     

Boophilus, which are one-host ticks finish their developmental cycles on the host, growing 

from an unfed larva to an engorged female. Boophilus ticks have mouthparts that are short and 

could cause destruction to skins as the preferred feeding sites are often of good leather potential 

(Jongejan & Uilenberg, 2004; Mkwanazi et al., 2021). B. microplus is the most important 

species which originates from the South-East of Asia but has increased significantly throughout 

the tropics in East and Southern Africa,  Australia and South and Central America (Hoogstraal, 

1956; Spickler, 2003). Till this day, other species such as B. geigyi and B. decoloratus continue 

to be restricted to Africa and are typically found on ruminants within West and Central Africa 

(Estrada-Peña et al., 2006).  

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The genus Dermacentor have three hosts in their life cycle (Földvári et al., 2016) and comprises 

33 species. Characteristic of them are the presence of eyes, ornate scuta and palps that are short 

(Cupp, 1991; Walker et al., 2003; Jongejan & Uilenberg, 2004). The distribution of 

Dermacentor is focal (Földvári et al., 2016). Several species such as D. reticulatus D. 

marginatus parasitize farm and other domestic animals in Europe (Rubel et al., 2016) and Asia 

(Fawaz Dabaja et al., 2017; Hosseini-Chegeni, 2019). In North America, species of the genus 

Dermacentor are among the ticks that infest livestock. However, Dermacentor ticks do not 

have adverse effects  on livestock in Africa (Földvári et al., 2016) although some species have 

been identified in the wild (Horak et al., 2017; Walker et al., 2003).  

About 168 species fall under the genus Haemaphysalis. Small ticks with short mouthparts are 

characteristic of ticks within this genus. Some species have lateral extending posterolateral 

angles on the second segment of the palps, which makes them have a triangular appearance 

and distinguishes them from other tick genera (Marchiondo & Endris, 2019). In these three-

host ticks, eyes are absent (Cupp, 1991). They are found on livestock in Asia, Europe and 

Australia (Rubel et al., 2018). In parts of Southern Asia  for example, H. bispinosa is found on 

cattle  (Brahma et al., 2013), while H. longicornis, which is an East Asian species found on 

cattle and other domestic animals, is now identified in Australia (Marendy et al., 2020). In 

Europe however, the most common on ruminants is H. punctate (Hornok et al., 2015). 

Hyalomma species are ticks ranging in size from medium to large that have long mouthparts 

and eyes found in sockets. They infest  wild and domestic mammals as well as birds and are 

abundant in semi-arid climate zones (Sajid et al., 2018). This genus there are 30 species in all, 

the majority of which carry out their life cycle on three hosts. Their ability to undergo either a 

two-host (e.g., H. truncatum and rufipes) or a three-host (e.g., H. dromedarii) cycle is host 

dependent, while others such as H. scupense undergo a one-host cycle (Spengler et al., 2016). 

Adult Hyalomma ticks actively move from their resting sites to pursue hosts when they 

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approach contrasting most other ixodid ticks, that rest on the vegetation in anticipation of a 

host (Jongejan & Uilenberg, 2004).  

Being the genus of hard ticks with the largest number of 241 species, Ixodes have no eyes and 

a scutum that has no ornamentation. Their life cycle occurs on three hosts and many species 

inhabit nests or holes (Walker et al., 2003). Although commonly scattered among shrubs,  

fewer species infest larger animals (Schwarz et al., 2009).  I. persulcatus and I. ricinus  are the 

common species in Asia and Europe (Rumer et al., 2011), whereas the in North America, I. 

scapularis is prevalent (Ginsberg et al., 2021). However, the unselective feeding attitude of I. 

persulcatus and I. Ricinus on a wide range of hosts makes them important vectors of varied 

zoonotic tick-borne diseases (Aliota et al., 2014; Hyesung Yang & Han, 2018).  

Having a three-host life cycle, Rhipicephalus genus has 70 species made up of ticks of average 

sizes with palps that are short and broad and commonly inornate. They have eyes and festoons 

are present (Dantas-Torres, 2010). In Africa, Rhipicephalus are common on mammals 

(Kanduma et al., 2020). They may either be  three-host or two-host ticks (e.g. R. evertsi) 

(Walker et al., 2000). in Southern and Eastern Africa, the brown ear tick, Rhipicephalus 

appendiculatus, is the most prevalent rhipicephalid tick (Jongejan & Uilenberg, 2004), 

infecting both domestic and wild ruminants with a preference for the ears (Walker et al., 2000).  

 

2.3 Tick Behaviour 

2.3.1 Host seeking 

Ticks spend about 90% of their life cycle off-host. (Needham & Teel, 1991; Leal et al., 2020). 

Although oviposition, egg hatching, metamorphosis, and moulting between the stages often 

occur off-host in the environment, these activities and the survival of the many stages within 

the life cycle require nutrition from a blood meal. (Leal et al., 2020). As a result, being able to 

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quickly locate and attach themselves to a host is critical for survival. In finding a host, ticks 

employ questing behaviour. Generally, two strategies are employed by ticks for this, namely 

the ambush and hunter strategies. The ambush strategy involves ticks (typically of the genera 

Rhipicephalus, Haemaphysalis and Ixodes) climbing to the top of rocks or resting on blades of 

vegetation and extending their front legs in anticipation of a host. The hunter strategy however 

involves the tick (typically of the genera Amblyomma and Hyalomma) actively moving to 

pursue a potential host ( Parola & Raoult, 2001; Leal et al., 2020).  

Ticks perceive certain cues in recognizing their hosts and determine whether to attach for 

feeding or to fall off and resume seeking another host. These include radiant heat, odours, 

vibrations or visual images. Odours are major stimuli perceived such as butyric and lactic acids, 

produced during perspiration and from other fluids from the body. Although breath of humans 

also stimulates a response, it is not as strong as that of cattle. In addition, carbon dioxide in the 

breath of the animal and ammonia in urine and other animal wastes are some essential 

attractants released by hosts. Subtle surges in radiant heat stimulate ticks and act synergistically 

with host odours. Hunter ticks also rely on visual cues in distinguishing between shapes that 

are dark against the sky’s background. In that, they respond to shadows, via the extending their 

legs to enhance interaction. Attachment of tick to host will not proceed unless adequate stimuli 

are perceived in a specific order (Osterkamp et al., 1999; Randolph, 2013). 

2.3.2. Feeding 

The majority of tick species inhabit forests, savannahs, grasslands and scrublands, and survive 

long periods until a suitable host to feed on is found. Upon strategically positioning themselves 

(questing) and successfully identifying and climbing onto a host, the ticks assess suitable sites 

of attachment and hold on firmly to the host, boring into the skin with their mouthparts which 

favours the transmission of pathogens they may be carrying from a previous blood meal from 

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another host (Richter et al., 2013). Ticks are pool feeders who inject saliva through feeding 

and finish engorgement through the same channel in a sporadic sequence of salivating and 

sucking (Bonnet et al., 2018). During feeding, ticks remain attached to the host. Earlier during 

attachment, mouthparts some ticks are cemented to the host’s skin, releasing them when most 

bodily fluids and erythrocyte meals have completely been taken up (Sonenshine & Roe, 2013). 

Salivary glands are thus essential in tick feeding and transmission of pathogens (Tahir et al., 

2020). In that, before a transmission, saliva plays a role in reactivating and/or replicating the 

pathogen from the infected tick to a naïve host (Ebel & Kramer, 2004).  

2.3.3 Disease transmission 

Ticks pose several debilitating threats on their hosts and these are attributed to their bites. These 

range from toxicosis and paralysis to irritation and allergic reactions. Some tick species 

commonly found attached to humans have been implicated as competent vectors of viral, 

bacterial and protozoan pathogens (Estrada-Peña & Jongejan, 1999). Molecules within the 

saliva are exploited by infectious agents to promote their transmission hence the term Saliva-

activated transmission (SAT). At the site of attachment of the tick on the skin, the SAT factor 

is among the immunomodulatory proteins that help maintain prolonged feeding of ticks, thus 

fuelling pathogen transmission (Nuttall, 2019; Randolph et al., 1996).  

2.3.4 Tick-borne Zoonoses 

Ixodid ticks serve as vectors of some pathogens that cause diseases transferrable from animals 

to humans. An example is species of the bacterial genus Rickettsia, which typically is the 

African tick typhus agent R. africae in sub-Saharan Africa, and transmitted by the tropical bont 

tick, Amblyomma variegatum. On another hand, particularly Hyalomma spp is known to 

transmit the virus that causes Crimean-Congo haemorrhagic fever in humans.  

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2.3.4.1 Rickettsial Diseases 

Rickettsia are small gram-negative bacilli that completely replicate inside of eukaryotic cells. 

Organisms within this genus are prevalent worldwide and are usually harboured by arthropod 

vectors that suck blood such as ticks, mites, lice and fleas. Despite the problem posed by 

Rickettsia globally, not much attention is given to it with limited resources allocated to face 

the challenge. This could be attributed to the fact that they present symptoms similar to acute 

febrile infections which are endemic to (sub)tropical regions (Parola et al., 2013; Fang et al., 

2017; Blanton, 2019) and thus result in individuals usually being misdiagnosed. In addition, 

the absence of rapid point of care tests make it difficult to detect acute infections, hence 

healthcare providers usually resort to employing serological methods to ascertain past exposure 

(Blanton, 2019). Rickettsia pathogens thus persist and are emerging and remerging throughout 

the world (Fang et al., 2017). According to the CDC, Rickettsia are divided into spotted fever 

group (SFG), typhus group (TG), scrub typhus group (STG) and other Rickettsioses (Centers 

for Diseases Control and Prevention, 2020). Transmission of Rickettsiae occurs through the 

skin of a person. Following inoculation into the skin, dendritic cells phagocytose pathogens, 

after which transportation to lymph nodes takes place. Replication occurs and organisms enter 

the bloodstream and further infect the endothelium of the microcirculation. This results in 

damage and increased vascular permeability which manifests as rashes and in severe cases 

interstitial pneumonia, acute kidney injury, meningoencephalitis, multiorgan failure and even 

death (Blanton, 2019). 

2.3.4.2 Spotted Fever Group (SFG) 

SFG Rickettsiae infect a wide range of tick species worldwide and thus have a wide geographic 

distribution (Parola et al., 2013). Although ticks are vectors, they may also double as reservoir 

hosts by passing pathogens from one stage to the next in the life cycle as well as from infected 

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females to eggs. The most pathogenic species of Rickettsial diseases, Rickettsia rickettsii falls 

within this group. Less pathogenic species such as R. africae, which causes African tick-bite 

fever also belongs in this class (Blanton, 2019; Parola et al., 2013) 

Others, identified and described based on their geographical prevalence include Rocky 

Mountain spotted fever (RMSF), that is caused by R. Rickettsii, R. parkeri and Candidatus R. 

philippi pathogens, is a disease of the Americas. Mediterranean spotted fever (MSF), that is 

also caused by R. conorii, exists predominantly in Asia, Africa and Europe. Tick-borne 

lymphoadenopathy (TIBOLA), also within this class is caused by R. slovaca and R. raoultii in 

Europe and Asia (Socolovschi et al., 2009)  

2.3.4.3 Typhus Group (TG) 

Vector-borne bacteria that cause typhus group rickettsiosis are Rickettsia typhi and R. 

prowazekii. The former has been implicated to cause the less severe endemic murine typhus in 

tropical coastal regions and transmitted by fleas. The latter however which causes severe 

epidemic typhus in temperate and tropical regions is transmitted by body lice (Rauch et al., 

2018).  

2.3.4.4 Scrub Typhus Group (STG) 

This rickettsiosis is confined to the Asia Pacific region specifically distributed in the 

Tsutsugamushi triangle - the soviet union to the north, China, Philippines and tropical Australia 

to the south , Japan to the east and  India, Pakistan through Afghanistan in the west 

(Chakraborty & Sarma, 2017). The pathogen implicated in causing the disease is Orientia 

tsutsugamushi, transmitted by chigger mites (Chakraborty & Sarma, 2017).  

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2.3.4.5 Other Rickettsioses 

In the United States, ehrlichiosis and anaplasmosis are the commonly reported tick-borne 

infections with Ehrlichia chaffeensis and Anaplasma phagocytophilum being implicated in 

infections respectively. In certain parts of Europe, Africa, Asia and South America, cases with 

various Ehrlichia and Anaplasma spp. have been recorded. In Europe and Asia for example, 

Neoehrlichia mikurensis is the causative tick-borne pathogen. Neorickettsia sennetsu, 

responsible for causing Sennetsu fever has also been reported in Japan and Malaysia and this 

is usually contacted via the consumption of raw fish infected with neorickettsia-infected flukes 

(Centers for Diseases Control and Prevention, 2020).   

2.3.4.6 Crimean Congo Haemorrhagic Fever (CCHFV) 

This is described as one of the very severe zoonotic viral diseases of humans, characterized by 

fever and haemorrhage and often fatal (Ergönül, 2006; Belobo et al., 2021). The virus was first 

identified in Crimea, Russia in 1944 and later found to be identical to the Congo virus identified 

in Congo in 1967, hence its current name and has since been detected in more than 50 counties 

across Asia, Europe and Africa (Ergönül, 2006; Belobo et al., 2021).  

It is tick-borne disease caused by CCHFV of the Orthonairovirus genus within the Nairoviridae 

family and the Bunyavirales order (Adams et al., 2017). Transmission is facilitated by 

Hyalomma ticks of the family Ixodidae, which spread the virus to humans as well as animals 

(both domestic and in the wild) (Gargili et al., 2017). Humans get infected via tick bites, contact 

with individuals carrying the virus in the early stages of infection or upon contact with viraemic 

livestock’s blood or tissues (Ergönül, 2006). Geographically, CCHFV is widely distributed and 

causes severe human disease with a high mortality rate, thus making it an important human 

pathogen (Ergönül, 2006). The virus spreads through ticks to animals and migratory birds, and 

genetic analysis can be used to determine its geographical distribution. Africa, Southeast 

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Europe, and Asia are the most common locations for CCHF (Cajimat et al., 2017; Okely et al., 

2020; Wölfel et al., 2007). 

2.4 Crimean Congo Haemorrhagic Fever Virus (CCHFV) Genome  

The CCHFV is sphere-shaped about 80–100 nm in diameter, a lipid envelope of 5–7 nm 

thickness and glycoprotein spikes that are  8–10 nm in length (Wahid et al., 2019). The CCHFV 

has a tripartite RNA genome (small [S], medium [M], and large [L] segments) with high genetic 

diversity and also has a negative-sense. The major nucleocapsid protein is encoded by the small 

portion of the genome (S, 1.7 kb), the amino acids’ precursor (M ~ 1700) is encoded by the 

medium portion (M, 5.3 kb), which results in the assembly of two envelope glycoproteins, Gn 

(37-kDa)  and Gc (75-kDa), whereas an RNA-dependent RNA polymerase (450 kDa- L-RdRp) 

is encoded by the large portion (L, 12.1 kb)  (Cajimat et al., 2017; Papa, 2019; Voorhees et al., 

2018; Wahid et al., 2019). At the nucleotide level, the S segment which is the most conserved 

has sequences that can be grouped into six lineages (Chamberlain et al., 2005). Every genetic 

lineage has been associated with geographic regions in the Middle East, Africa, Europe and 

Asia where Hyalomma spp. ticks are existent (Hoogstraal, 1979). In Africa, genotypes I, II, 

and III have been described; whereas in Asia genotype IV is described  and in Europe, 

genotypes V and VI (Deyde et al., 2006; Monsalve Arteaga et al., 2021). CCHF has a wide 

ecological distribution. In Asia (India, Pakistan, Tajikistan, Kazakhstan, and Russia), the 

Middle East (Iran and Afghanistan), Europe (Turkey, Bulgaria, Greece, Albania, and Kosovo), 

and more than 30 African republics (including Sudan, Mauritania, Kenya, Senegal, and South 

Africa), minor outbreaks of Crimean-Congo haemorrhagic fever virus (CCHFV) have been 

reported since 2000 (Farhadpour et al., 2016). 

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2.5 Strategies for tick control and Tick-borne Disease (TBD) prevention  

Several strategies to prevent and control the abundance of ticks, the prevalence of pathogens 

they carry and the risk of humans being exposed to these pathogens have been outlined in many 

instances. Broadly, the strategies can be classified into adopting personal protective measures 

(which includes protective clothing, use of repellents, etc), managing ticks to reduce their 

contact with humans (through the application of chemical acaricides on domestic animals and 

the environment), and lowering the transmission of tick-borne diseases via reducing the 

prevalence of pathogens being carried (such as landscape and habitat modification, use of 

biological control agents) (Stafford et al., 2017). However, due to the persistence of tick and 

tick-borne diseases, despite the various efforts made to alleviate these challenges, it is evident 

that no one method is effective at completely tackling the problem. It is thus important that 

Integrated Tick Management programs be employed. This includes surveillance, identification, 

routine disease testing, landscape management, and targeted control of adult ticks (De La 

Fuente et al., 2015).  

 

 

 

 

 

 

 

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CHAPTER THREE 

METHODOLOGY 

3.1 Study sites 

Samples were collected from January to March 2020. Sampling was carried out within environs 

in Greater Accra and Upper East Region, which fall within the Coastal savannah and Sudan 

Savannah ecological zones respectively in Ghana (Figure 1). These locations also serve as 

study sites for a through a collaboration between the Ghana Armed forces, the Navrongo Health 

Research Centre (NHRC) and the US Naval Medical Research Unit 3 (US NAMRU-3). The 

selected sites included military-owned kraals at Burma camp (5 Infantry Battalion and 3 

Mounted Squadron), Michel camp (1 Infantry Battalion) and Asutuare within Greater Accra; 

and Abattoir, Cow market and Nakong within Navrongo.   

 

Figure 2: Regional Map of Ghana showing study sites. 

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3.2 Ethical clearance 

Ethical clearance for the study was obtained from the Noguchi Memorial Institute for Medical 

Research (NMIMR) (CPN 110/15-16), Ghana Armed Forces (GAF) (IPN: 093/2017), Ghana 

Health Service (GHS) (GHS ERC 004/03/19) and the Naval Medical Research Centre (NMRC) 

(NAMRU3.2016.0010) Institutional Review Boards.  

3.3 Study Population 

Animal Handlers which included livestock rearers and Abattoir workers who resided or worked 

within a 10-kilometre periphery of the study sites were invited to participate in the study. 

3.4 Sample collection 

3.4.1 Sampling from Animals 

3.4.1.1 Inclusion/ Exclusion of Animals 

There was no justification for the exclusion of livestock from being sampled. The study did not 

experiment with animals but only involved collecting ticks off the animals for infection studies. 

At each sampling site, ticks were removed from a convenience sample of animals for tick 

identification and further molecular testing. 

3.4.1.2 Tick collection from Animals 

Ticks were hand-picked either directly or using blunt-tipped forceps. Before tick collection, 

animals were restrained by the owner or handler to provide easy access. Caution was taken not 

to twist or jerk the tick to ensure mouthparts did not break off or remain in the skin. Having 

done these, the tick was pulled up straight until all parts of it were totally out of the animal’s 

skin (Figure 2). The ticks were then placed into labelled tubes containing RNA later and further 

transported to the AFI laboratory at NMIMR for morphological identification.   

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Figure 3: Image demonstrating how ticks were taken off livestock 

Source: (Centers for Disease Control and Prevention, 2021) 

 

 

 

3.4.2 Sampling from Humans 

3.4.2.1 Inclusion criteria for human sampling 

Primary animal handlers above 18 years and who were willing to participate and provide 

informed consent were recruited for the study.  

3.4.2.2 Exclusion criteria for human sampling 

Individuals who did not have primary contact with animals, under 18 years and were unwilling 

to participate and provide informed consent were excluded. 

 

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3.4.2.3 Community entry 

For Navrongo, which was a non-military site, permission was sought from the town elders and 

local chiefs who had jurisdiction over the areas. For military sites within Greater Accra 

however, approval was sought from command.  Before initiation of the study, the target 

population were assembled in groups based on the area of their activities. The study was then 

explained (background, purpose, procedures, time and place for enrolment) to the groups. 

Participants were then individually approached and invited to enrol in the study.  

3.4.2.4 Participant eligibility and informed consent procedure 

Participants were first screened for eligibility. Upon meeting the criteria, the procedure was 

clearly explained to the participants, allowing them to ask questions and to ensure they were 

all comfortable about the procedure. Further, unique identification numbers were assigned to 

each enrolled participant as this information was necessary to enable linking to laboratory 

specimens. Afterwards, informed consent was obtained from participants in duplicate as per 

protocol before blood draw and a copy of informed consent was given to the enrolled 

participant. 

3.4.2.5 Blood Collection from participants 

Serum separator tubes (SSTs) were labelled with the unique participant study ID number and 

date after which a tourniquet was applied proximal to the site of venepuncture upon identifying 

a good-sized vein to ensure engorgement of the vein with blood. The site of venepuncture was 

cleaned with an alcohol swab and a 10 ml syringe with the appropriate needle prepared and 

inserted into the vein. About 10 ml of blood was gently drawn into the syringe after a blood 

flashback in the bevel of the syringe. Immediately, a cotton swab was used to apply direct 

pressure over the puncture site to stem any bleeding. The blood was then transferred from the 

syringe into SST by directly puncturing the tube in the centre and gently injecting blood into 

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the empty tube. Thereafter, the tube was inverted about 10 times to ensure mixing of the sample 

with the anticoagulant inside the tube. The SSTs were then stored in a cool box with ice packs 

while waiting for blood to clot. Band-aids were then affixed to sites of venepuncture to ensure 

bleeding stops after which the specimen logbook with study number and details of patients was 

completed. 

3.4.2.5.1 Blood Sample Processing 

Blood tubes (SSTs) were centrifuged for 10 minutes at 1,000–2,000 x g while ensuring the 

tubes were properly balanced inside of the centrifuge and handled very carefully so as not to 

disturb the separation of layers. Using a sterile pipette, 1 ml serum aliquots for each sample 

were transferred into labelled cryovials. Sera and blood clots were stored in a -80°C mobile 

freezer for transport to the AFI laboratory at Noguchi for analyses. 

3.5 Laboratory Testing 

3.5.1 Morphological Identification of ticks 

Ticks were removed from RNA later onto a clean petri dish and observed under a Motic SMZB 

series dissecting microscope. The various parts (mouthparts, ventral plates, eyes, scutum, etc) 

were observed as outlined in the identification keys by Walker et al., (2003) and ticks grouped 

into the various genera and species. Pictures of identified ticks within the various genera and 

species were taken. Following this, purification of nucleic acid was carried out. 

3.5.2 Extraction of total nucleic acid from ticks 

Extraction of total nucleic acid from each tick was carried out using QIAamp Viral RNA Mini 

Kit (250). Following the protocol of the manufacturer, 0.5g and 0.75g of 0.1mm and 2mm 

beads respectively were added to clean 1.5 ml microcentrifuge tubes. Single ticks were placed 

in each tube and 560µl of prepared Buffer AVL were added. Tubes were then centrifuged at 

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8000rpm for 2minutes to homogenize the ticks.  Four hundred microlitres (400uL) of the 

homogenate was transferred into clean 1.5 ml microcentrifuge tubes after which incubation 

was done at room temperature (between 15-25°C) for 10 minutes. Furthermore, centrifugation 

of the tubes was briefly done to remove drops from inside the lid and 560 µl of absolute ethanol 

(96-100%) added and vortexed for 15 seconds. To remove drops of the mixture from inside the 

caps, the tubes were briefly centrifuged.  

A volume of 600 µl of the solution was carefully pipetted from the previous step into the 

QIAamp Mini spin column without wetting the rim and centrifugation further done at 8000 

rpm for 1 minute. After discarding the filtrate, the QIAamp Mini spin column was placed into 

a clean 2ml collection tube. Further, this step was repeated.  

Following this, Buffer AW1(500 µl) was added to the spin column, and centrifuged at 8000 

rpm for 1 minute. The QIAamp Mini spin column was placed into a clean 2ml collection tube 

and the filtrate was discarded. After carefully opening the QIAamp Mini spin column, 500 µl 

of Buffer AW2 was added and centrifugated at maximum speed (14,000 rpm) for 3 minutes. 

The spin column was then placed in a new 2ml collection tube. An 80 ul volume of elution 

buffer AVE was carefully added to the membrane of the spin column and incubated at room 

temperature for a minute. The nucleic acid was finally eluted by centrifuging the column at 

8000 rpm for 1 minute.  At -20°C, the eluate containing the total nucleic acid was stored at. 

 

 

 

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3.5.3 Molecular detection of tick-borne pathogens 

3.5.3.1 Real-time PCR (RT-PCR) for detection of CCHFV 

In screening for CCHFV, in-house reagents from United States Army Medical Research 

Institute of Infectious Diseases (USAMRIID) were used with primers targeting the S segment 

of the CCHFV genome to screen the nucleic acid extracted from the ticks (Atkinson et al., 

2012) 

The total volume of the reaction was 20 μl, comprising 14.6 μl of prepared master mix, 0.4 μl 

of platinum Taq and 5 μl of the template (nucleic acid extract). Each reaction was ran with a 

positive control, with nuclease-free water as the negative control, simultaneously. The PCR 

was performed on the Applied Biosystems (USA) 7300 Real-time PCR system with cycling 

conditions as follows: 1 cycle of the first hold for 15 minutes at 50°C, 1 cycle of the second 

hold for 5 minutes at 95°C, 45 cycles of the third hold for 1 second at 94°C, 30 seconds at 55°C 

(fluorescence read and data acquisition), for 5 seconds at 68°C and 1 cycle of the fourth hold 

at for 30 seconds at 40° C. No passive reference was chosen, FAM was selected as reporter 

and TAMRA as the quencher. 

3.5.3.2 Real-time PCR (RT-PCR) for detection of Rickettsia species 

For the genus Rickettsia (R17b), in-house reagents from the Vector-borne Diseases division of 

the Naval Medical Research Center (NMRC) were used with primers that targeting the 17kDa 

surface protein to screen the nucleic acid extracted from the ticks for the pathogen (Jiang et al., 

2004). 

Each reaction had a total volume of 20 μl, comprising 14.8 μl of prepared master mix, 0.2 μl 

of platinum Taq and 5 μl of the template (nucleic acid extract). Each reaction was ran with a 

positive control, with nuclease-free water as the negative control, simultaneously. The PCR 

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was performed on the Applied Biosystems (USA) 7300 Real-time PCR system with cycling 

conditions as follows: 1 cycle of the first hold for 2 minutes at 95°C, 45 cycles of the second 

hold at for 5 seconds at 95°C and for 30 seconds at 60°C (fluorescence read and data 

acquisition) and 1 cycle of the third hold for 30 seconds at 40°C. No passive reference was 

chosen, FAM was selected as a reporter and no quencher was selected. 

3.5.3.3 Realtime PCR (RT-PCR) for detection of Rickettsia africae 

A species-specific assay was chosen due to the incidence and spread of spotted fever 

Rickettsiae on the African continent to test samples that were positive for the Rickettsia genus, 

for Rickettsia africae (Maina et al., 2014). Each reaction had a total volume of 25 μl, 

comprising 12.5 μl of 2X TaqMan Universal PCR master mix (Applied Biosystems, USA), 5 

μl of nuclease-free water, 1.25 μl of forward and reverse primers, 1 μl of the probe, 1 μl of 

MgCl2 and 3μl of the template (nucleic acid extract).  Each reaction was ran with a positive 

control, with nuclease-free water as the negative control, simultaneously. The PCR was 

performed on the Applied Biosystems (USA) 7300 Real-time PCR system with cycling 

conditions as follows: 1 cycle of the first hold for 10 minutes at 95°C, 45 cycles of the second 

hold for 15 seconds at 95°C and for 1 minute at 60°C (fluorescence read and data acquisition 

stage). ROX was chosen as the passive reference, FAM as a reporter and no quencher selected.  

3.5.4 Serological analysis for antibodies against tick-borne pathogens 

3.5.4.1 Magpix Immunoassay for detection of CCHFV antigen 

In a biological safety hood, 2 µl patient samples were diluted in 198 µl PBST-SK (Phosphate 

buffered saline with 0.02% Tween-20 and 5% skimmed milk) in a 1:100 ratio. Negative 

controls were then diluted in PBST-SK in a 1:10 ratio (450 µl PBST-SK added to a negative 

control aliquot tube). Positive control was also diluted in a 1:2 ratio in PBST-SK (100 µl PBST-

SK was added to one positive control aliquot tube). The capture bead mix was vortexed for 10 

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seconds and diluted in PBST (Phosphate buffered saline with 0.02% Tween-20) in a ratio of 

1:250, (ensuring beads are protected from light before and after dilution. Into the desired wells 

of a white 96 well plate 50 µl was added to the wells. Sealing of the plate was done and plate 

further placed on the Luminex Plate Magnet (ensuring that the connection was secure with the 

two metal arms latched on the sides of the plate), covered with foil and the beads allowed to 

collect at the bottom for 60 seconds. After, the Luminex plate magnet was grasped (while still 

latched to the plate) and the buffer was discarded into a sink with a downward motion. 

Following this, the negative and positive controls, as well as the samples, were added to the 

plate and the sample locations recorded. The plate was then sealed and put onto a plate shaker 

to shake for an hour at 400 rpm (making sure the plate was covered with foil during this step). 

After this, the plate was removed from the shaker, plate sealer removed, placed on the Luminex 

plate magnet and covered with foil. The liquid was discarded in the sink while the magnet was 

still latched to the plate as above. Washing of the beads were done by adding 100 µl of PBST 

to the wells of the plate after which the plate was put onto the magnet, sealed, covered with a 

foil and then buffer discarded after a minute. This step was carried out two additional times to 

make a total of three washes. Furthermore, the Human anti-IgG-PE was diluted in PBST-SK 

(1:100) after which 50 µl was added to the wells. The plate was then sealed, covered with a 

foil and placed on the plate shaker for an hour. After, taking the plate off the shaker, and placing 

on the Luminex magnet for a minute, the liquid was discarded, leaving the beads bound to the 

bottom of the plate. Three washes were then done by adding 100 µl PBST to the well, 

incubating on the Luminex magnet for a minute and buffer discarded. After the final wash, 100 

µl of PBST was added to the wells and then proceeded to read the plate on the Luminex 

Magpix. A sample was called positive if the signal to noise (S/N) ratio was greater than 10 (i.e., 

S/N > 10) 

 

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3.5.4.2 ELISA Assay to detect Group-specific antibodies against Rickettsiae  

One-half of 96 well microtiter plates were coated with 100 µl Rickettsia STG (Karp = 1:1000; 

Kato = 1:1000 and Gilliam = 1:1500), SFG (1:2500) and TG (1:2000) antigens diluted in PBS. 

For the other half of the plates, however, 100 µl of PBS only was added with no antigen. Plate 

sealers were used to seal the plates, which were subsequently wrapped in aluminium foil and 

incubated for a minimum of 48 hours at 4°C. Furthermore, wash buffer was used to wash the 

coated plates three times (0.1% Tween in 20 in PBS). After that, 200 µl of blocking buffer (5 

percent skim milk [Difco] in wash buffer) was added to each well to block the plates. Plates 

were covered in aluminium foil and incubated for 1 hour at room temperature, after which wash 

buffer was used three times. After diluting the patient serum in blocking buffer (1:100), 100 µl 

was pipetted into allotted wells for each sample on both the antigen and PBS half of the plate. 

Washing was done three times with wash buffer after an hour of incubation at room 

temperature. 100 µl of peroxidase-conjugated goat antihuman IgG (Kirkegaard & Perry) was 

added to each well after diluting it in blocking buffer (1:2000) and incubating it at room 

temperature for 1 hour. After that, the plates were washed three times in wash buffer. 100 µl 

of ABTS (Kirkegaard & Perry) substrate was mixed in a 1:1 ratio and added to each well of 

the plate, which was then incubated in the dark for 30 minutes. Plates were then scanned at 405 

nm (minus a reference value of 650 nm) with ODs from antigen-free wells used to subtract 

background absorbance. If the net optical density (NET OD) values were more than or equal 

to 1.0 (i.e., NET OD 1.0), the sample was considered positive. 

3.5.5 Whole Genome Sequencing (WGS) for CCHFV PCR positives 

Whole Genome Sequencing was carried out to determine the entire sequence of the pathogen’s 

genome after which genome assembly and phylogeny was carried out. 

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3.5.5.1 RNA PICO 6000 assay (for quantification and assessing quality of RNA for NGS 

library preparation) 

The RNA ladder was stretched down and heat-denatured for 2 minutes at 70°C, after which the 

vial was placed on ice to cool. A total of 90 µl of RNase-free water were added and well mixed. 

The appropriate amount for normal daily use was then prepared in approved 0.5 mL RNase-

free vials and stored at -70°C. Before use, all reagents were brought to room temperature for 

30 minutes. To begin, 550 µl of RNA 6000 Pico gel matrix was deposited in the top receptacle 

of a spin filter, which was centrifuged at 4000 rpm for 10 minutes. After that, 65 µl of filtered 

gel was aliquoted into the provided 0.5 ml RNase-free microcentrifuge tubes. Aliquots were 

kept at 4°C for a month and then used.  

The RNA 6000 Pico dye concentrate was briefly vortexed, span down, and 1 μl pipetted into a 

vial containing the prepared 65 μl aliquoted filtered gel. The tube was capped, thoroughly 

vortexed and visually inspected to ensure gel and dye were properly mixed after which 

centrifugation was done at 14000 rpm for 10 minutes. The dye concentrate was then stored in 

the dark at 4°C to ensure that it was shielded from light.  

A new RNA chip was removed from its sealed package and placed on the chip priming station 

after allowing the gel-dye mix to equilibrate to room temperature for 30 minutes. In addition, 

9 μl of the gel-dye mix was pipetted and placed into the bottom of the indicated well (G). The 

chip priming station was then closed after a 30-second timer was set, ensuring that the plunger 

was positioned at 1 ml. The syringe plunger was then pushed down until it was caught by the 

clip. The plunger was visually inspected until it moved to at least the 0.3 ml mark after 30 

seconds, and then carefully pushed back to the 1 ml position using the clip release mechanism. 

The chip priming station was next opened, and 9.0 μl of the gel-dye mix was added to the two 

remaining wells (G). Pipetting 9 μl of RNA 6000 Pico conditioning solution into the indicated 

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well CS and 5 μl of RNA 6000 Pico marker into the well with a ladder symbol and each of the 

11 sample wells was done. 

In addition, 1 l of diluted RNA 6000 Pico ladder (made as indicated above) was pipetted into 

the ladder symbol-marked well. The samples were then placed in the sample wells (1 l of each 

sample). On an IKA vortex mixer, the chip was vortexed for 60 seconds at 2400 rpm. The chip 

was carefully inserted into the receptacle of the Agilent 2100 Bioanalyzer instrument, the 

proper assay was chosen from the assay menu, and the chip run began. 

3.5.5.2 First-strand synthesis 

The fragmentation and priming reaction mix were made on ice in a nuclease-free tube using 5 

μl of input RNA, 4 μl of NEBNext First Strand Synthesis Reaction Buffer, and 1 μl of Random 

Primers for a total of 10 μl per reaction while ensuring thorough mixing. The samples were 

then placed in a thermocycler and heated to 94 degrees Celsius. The incubation time differed 

depending on whether the RNA was intact or partially degraded, as determined by the 

bioanalyzer (RIN >7 (intact RNA) = 15 minutes; RIN 2-6 (partially degraded RNA) =7-8 

minutes). Following this, the tubes were immediately placed on the ice. 

The first-strand synthesis reaction mix was then put together on ice in a nuclease-free tube with 

10 μl fragmented and primed RNA, 8 μl NEBNext Strand Specificity Reagent, and 2 μl 

NEBNext First-Strand Synthesis Enzyme Mix to form a total of 20 μl per reaction while 

assuring complete mixing. The reaction mixture was then incubated in a preheated 

thermocycler under the following cycling conditions: Step 1: 25°C for 10 minutes, 42°C for 15 

minutes, 70°C for 15 minutes, step 3: 70°C for 15 minutes, and step 4: 4°C hold. 

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3.5.5.3 Second strand synthesis 

The second strand cDNA synthesis reaction was made on ice in a nuclease-free tube with the 

20 l first-strand synthesis product, 8 μl NEBNext Second Strand Synthesis Reaction Buffer, 4 

μl NEBNext Second Strand Synthesis Enzyme Mix, and 48 μl nuclease-free water to make a 

total of 80 μl per reaction while ensuring thorough mixing. The thermocycler was used to 

incubate the samples for 1 hour at 16°C with the heated lid set to a temperature of 40°C. After 

that, the NEBNext Sample Purification Beads were vortexed to resuspend them, and 144 μl of 

the second strand synthesis product (80 μl) was added. This was followed by thorough 

vortexing and a 5-minute incubation period at room temperature. The tube was briefly 

centrifuged before being put on a magnet to separate the beads from the supernatant. The 

supernatant was eliminated when the solution became clear, taking care not to disturb the beads 

(which are bound to DNA). 200 μl of freshly made 80 % ethanol was added while the magnetic 

stand was still in use, and the supernatant was discarded after 30 seconds of incubation at room 

temperature. After two washes, the beads were air-dried for 5 minutes with the tube remaining 

on the rack. After that, the tube was taken from the magnetic stand, and the DNA target was 

eluted from the beads using 53 μl 0.1X TE buffer, which was thoroughly mixed. Brief spinning 

was performed, followed by an incubation period at room temperature. The tube was placed 

back on the magnetic stand, and 50 μl of supernatant was transferred to a clean nuclease-free 

PCR tube when the solution became clear. 

3.5.5.4 Qubit quantification of cleaned pre-enriched cDNA 

Standards and samples were placed in separate assay tubes. 190 μl of Qubit working solution 

was pipetted into the tube for standards, and 10 μl of standards (1 and 2) were pipetted into 

separate tubes. In separate tubes for each sample, 198 μl of Qubit working solution and 2 μl of 

samples were added to the tubes containing the cDNA samples. All mixtures were briefly 

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vortexed and allowed to incubate at room temperature before the concentrations were measured 

(standards first, then samples) using the Qubit (Invitrogen life technologies). 

3.5.5.5 Tagmentation of Genomic DNA 

A 30 μ volume of DNA was poured into the wells of a 96-well PCR plate. 20 μl tagmentation 

master mix containing 11.5 μl enrichment Bead-Linked Transposomes (eBLT) and 11.5 μl 

tagmentation buffer 1 (TB1) was added to each sample, mixed completely, sealed, and shaken 

for 1 minute at 1600 rpm. The plate was placed on a thermal cycler, with the preheat lid set to 

100°C and the reaction volume set to 50 μl, incubation at 55°C for 5 minutes and a hold at 

10°C. After allowing the plate to cool for 2 minutes, 10 μl of stop tagment buffer 2 (ST2) was 

added to the tagmentation reaction, sealed, and shaken for 1 minute at 1600 rpm. After that, 

the plate was placed on a magnetic stand for 3 minutes to allow the liquid to clear.  

After that, 60 μl of the supernatant was removed and discarded before washing. The plate was 

removed from the magnetic stand, and 100 μl of tagment wash buffer (TWB) was added 

directly onto the beads and stirred thoroughly until they were fully resuspended. The plate was 

re-positioned on the magnetic stand until the liquid was clear, and 100 μl of supernatant was 

discarded. For a total of two washes, the wash process was repeated. 

The plate was further taken off the magnetic stand after removing and discarding 100 μl of the 

supernatant. After, 40 μl of PCR master mix, consisting of 23 μl of Enhanced PCR Mix (EPM) 

and 23 μl of Nuclease-free water was added and mixed thoroughly till beads were fully 

resuspended. The index adapter plate was then prepared and briefly centrifuged, after which 

10 μl of pre-paired index 1 (i7) and index 2 (i5) distinct index adapters were added to each 

sample in the wells. This was thoroughly mixed, the plate was sealed, and the preheat lid option 

was selected and set on a thermal cycler to 100°C, and volume set to 50 μl. Cycling conditions 

for the run were as follows: 72°C for 3 minutes for step 1, 98°C for 3 minutes for step 2, 12 

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cycles of 98°C for 20 seconds, 60°C for 30 seconds and 72°C for 1 minute for step 3, 72°C for 

3 minutes for step 4 and a final hold at 10°C.  

The plate was then placed on the magnetic stand till liquid was clear and 45 μl of each sample 

was transferred to a new plate. Further, ensuring AMPure XP beads were brought to room 

temperature, 88 μl was added, followed by the addition of 77 μl of nuclease-free water. This 

was thoroughly combined and allowed to incubate for 5 minutes at room temperature. After 

that, the plate was placed on the magnetic stand until the liquid was clear, and 200 μl 

supernatant from each sample was transferred to a fresh plate. In addition, 20 μl of AMPure 

XP beads were pipetted into the wells containing the supernatant, vigorously mixed, and 

incubated for 5 minutes at room temperature. While on the magnet, the supernatant was 

removed and disposed without harming the beads.  

After that, two washes were done as follows: with the plate on the magnet, 200 μl of freshly 

produced 80 % ethanol was added without mixing, followed by incubation at room temperature 

for 30 seconds and removal and discarding the supernatant. After draining any remaining 

ethanol, the plate was dried for 5 minutes on the magnetic stand. The plate was taken off the 

magnetic stand and 17 μl of resuspension buffer (RSB) was added to the beads in each well, 

which were then incubated at room temperature for 2 minutes before being placed back on the 

magnetic stand for 2 minutes. For library enrichment, 15 l of each sample's supernatant was 

transferred to wells of a fresh plate. 

3.5.5.6 Library enrichment 

Into the wells of a new plate, 9 μl of each sample was transferred, 12 μl of new hybridization 

buffer 1 (NHB 1), 1 μl of probes, 2.5 μl of enrichment hybridization buffer 2 (EHB 2) were 

added, making a total of 25 μl reaction volume. The plate was then placed on a thermocycler 

programmed as follows: denaturation at 95°C for 5 minutes for step 1, 18 cycles of ramping 

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down from 94°C to 58°C (decreasing 2°C every cycle) for 1 minute for step 2, hybridization 

at 58°C for 90 minutes for step 3 and a temperature hold at 58°C for step 5.  

The plate was removed from the thermocycler at the end of the run, 62.5 μl of streptavidin 

magnetic beads (SMB) were added, the plate was sealed and shaken for 4 minutes at 1200 rpm, 

then incubated at 58°C for 15 minutes.  

After that, the plate was removed from the thermocycler and placed on a magnetic stand for 

two minutes. 88 l of the supernatant was collected and discarded once the liquid had become 

clear. The plate was removed from the magnet and washed three times, first with 50 l of pre-

warmed enhanced enrichment wash (EEW), then shaking at 1600 rpm for four minutes, 

incubating at 58°C for five minutes, and then returning to the magnet to collect and discard the 

supernatant. After the washes, 50 μl of pre-warmed EEW was added to each well and shaking 

was done at 1600 rpm for 4 minutes, incubated at 58°C for 5 minutes and then placed on the 

magnet after which 50 μl of supernatant was removed and discarded.  

Thereafter, the plate was taken off the magnet, and 5.75 μl from elution master mix comprising 

7.125 μl enrichment elution buffer 1 (EE1) and 0.375 μl sodium hydroxide was added to each 

well, shaking was done at 1800 rpm for 2 minutes and incubated at room temperature for 2 

minutes. The plate was placed on the magnetic stand, and 5.25 μl of supernatant was transferred 

to a new plate, after which 1 μl of elute target buffer (ET2) was added and thoroughly mixed. 

The enriched library was then amplified by mixing 1.25 μl of PCR primer cocktail (PPC) with 

5 μl of enhanced PCR mix (EPM) in a thermocycler set to the following: 95°C for 5 minutes 

for step 1, 12 cycles of 98°C for 10 seconds, 60°C for 30 seconds and 72°C for 1 minute for 

step 2, 72°C for 5 minutes for step 3 and a temperature hold at 4°C. 

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After the amplification, 22.5 μl of AMPure XP beads were added to samples in each well, 

thorough mixing was done, shaking at 1800 rpm for 2 minutes was carried out and further 

incubated at room temperature for 5 minutes. Further, the plate was placed on the magnetic 

stand for 2 minutes, and 35 μl of the supernatant was discarded. While on the magnet, 200 μl 

of freshly prepared 80% ethanol was added to each well and discarded after 30 seconds. After 

two washes, the plate was kept at room temperature for 5 minutes to evaporate any remaining 

ethanol. After that, the plate was removed from the magnet and 32 μl of RSB was added to 

resuspend it, followed by 2 minutes of shaking at 1800 rpm and 2 minutes of incubation at 

room temperature. After 2 minutes on the magnet, 30 μl of the supernatant (clean enriched 

library) was transferred to the wells of a new plate. 

3.5.5.7 DNA high sensitivity assay (for Quantification of DNA library) 

Before use, all reagents were brought to room temperature for 30 minutes. To begin, 15 μl of 

High Sensitivity dye concentrate was pipetted into the High Sensitivity DNA gel matrix vial 

after brief vortexing. The dye concentrate was stored again at 4 °C in the dark.  

The tube was capped, thoroughly vortexed and visually inspected to ensure gel and dye were 

properly mixed. The gel-dye mix was completely transferred to the top receptacle of a spin 

filter, and the tube was centrifuged at 6000 rpm for 15 minutes at room temperature.  

After allowing the gel-dye mix to equilibrate to room temperature for 30 minutes, a new High 

Sensitivity DNA chip was taken out of its sealed bag. 9 μl of the gel-dye mix was pipetted and 

dispensed into the bottom of the well which was marked (G). The chip priming station was 

then closed after a 60-second timer had been set and the plunger had been set at 1 ml. The 

syringe plunger was pushed down until the clip caught it. The plunger was visually inspected 

until it moved to at least the 0.3 ml mark after 60 seconds, and then carefully pulled back to 

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the 1 ml point using the clip release mechanism. The chip priming station was then opened, 

and each of the additional wells labelled with a 9.0 l gel-dye mix was added (G). 

Further, 5 μl of the DNA marker was pipetted into the well with a ladder symbol and each of 

the 11 sample wells. After, 1 μl of the ladder was pipetted into the well with the ladder symbol. 

The samples (cleaned enriched library) were then put to the sample wells (1 μl of each sample). 

On an IKA vortex mixer, the chip was vortexed for 60 seconds at 2400 rpm. The chip was 

carefully inserted into the receptacle of the Agilent 2100 Bioanalyzer instrument, the proper 

assay was chosen from the assay menu, and the chip run began. For use, the average library 

size (bp) and the Molarity (nM) for each library were recorded. 

3.5.5.8 Library qPCR 

Using the concentrations from the high sensitivity bioanalyzer, qPCR was conducted to further 

quantify libraries to obtain proper concentrations that will help in library loading. The libraries 

were first diluted 1:1000 so that they were within the range of the qPCR standards (20-0.0002 

pm). The total volume of each reaction was 10 μl consisting of 1.8 μl of nuclease-free water, 

2X KAPA SYBR FAST qPCR Master Mix with primers, 0.2 μl of 50X ROX High and 2 μl of 

the template (diluted library).  The samples were run in duplicates. Six standards (1-6) of 

concentrations 20 pM, 2 pM, 0.2 pM, 0.02 pM, 0.002 pM and 0.0002 pM, as well as nuclease-

free water as the negative control, were run simultaneously with each reaction. The PCR was 

run on a 7300 Real-time PCR system (Applied Biosystems, USA) with cycling conditions as 

follows: 1 cycle of the first hold at 95°C for 5 minutes and 35 cycles of the second hold at 95°C 

for 30 seconds and 60°C for 45 minutes (fluorescence read and data acquisition stage). ROX 

was chosen as the passive reference, SYBR as reporter and ROX as the quencher.  

 

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3.5.5.9 Library Pooling for loading 

Upon obtaining concentrations from the qPCR run, the average concentrations from the qPCR 

run were computed together with fragment sizes from high sensitivity bioanalyzer run per 

sample. To ensure an equal representation of each sample, all samples were normalized to the 

same 2 nm concentration. Since the concentration of samples from qPCR was pico-mole, the 

sample concentrations were converted to nano-mole. With the needed volume of resuspension 

buffer, sample concentrations were standardized. The total amount of resuspension buffer 

required to normalize each sample to 2 nm was added to a 1.5 mL microcentrifuge tube, along 

with the equivalent volume for each sample.