Prr.hec.gov.pk



SEROLOGICAL AND MOLECULAR INVESTIGATIONS ON COXIELLOSIS AND ITS RELATIONSHIP WITH REPRODUCTIVE DISORDERS IN SMALL RUMINANTS AT LIVESTOCK FARMS OF PUNJAB, PAKISTAN 200406022415500BYQUDRAT ULLAHM. Phil.Regd. No. 2006-ag-1137A thesis submitted in partial fulfillment of the requirements for the degree ofDOCTOR OF PHILOSOPHYINTHERIOGENOLOGYDEPARTMENT OF THERIOGENOLOGY,FACULTY OF VETERINARY SCIENCE,UNIVERSITY OF AGRICULTURE,FAISALABAD, PAKISTAN2018DedicatedDedicatedTo MyLOVING PARENTSLOVING PARENTSMy Beloved WifeMy Kids (Zeeshan and Farhan)Brother and Sisters And Family members especially My Nieces and NephewsBy the virtue of their prayers, I have been able to reach at this position and their hands were always raised for prayer for my well-being, even at this moment of time.ACKNOWLEDGEMENTSI bend myself modestly in front of Almighty ALLAH, the Lord of the worlds, the Omnipotent, the Beneficent, the Merciful, and the Gracious and thanking Him for everything I have been blessed with in my life. Peace and blessings of Almighty Allah be upon Holy Prophet Hazrat Muhammad (Peace be Upon Him), the Apostle of Allah, the greatest social reformer, and the forever source of knowledge. A deep sense of gratitude is due to my Supervisor, Assistant Prof. Dr. Huma Jamil, Department of Theriogenology for her dynamic supervision and fruitful guidance during my course work, and research. I would like to thank Prof. Dr. Zafar Iqbal Qureshi, Chairman, Department of Theriogenology, University of Agriculture, Faisalabad, whose support and encouragement helped me a lot throughout the course. I am also thankful to Assistant Prof. Dr. Muhammad Saqib, Department of Clinical Medicine and Surgery, University of Agriculture, Faisalabad for his guidance in planning my research work, technical support and assistance during the research work. He deserves appreciation more than any other member of my supervisory committee, as he was the driving engine behind the study from its conception to write up stage.I am feeling dearth of words to express my gratitude and appreciation to Prof. Dr. Heinrich Neubauer, Director Institute for Bacterial Infections and Zoonoses (IBIZ), Friedrich Loeffler Institute (FLI), Jena, Germany for providing valuable suggestions, technical support in the form of diagnostic/ laboratory equipments and boosting up my morale during conduct of this study. I have no words to pay thanks for his skilful and ever inspiring intelligent guidance, and absolute friendly atmosphere during my lab work at FLI. I am also very thankful to Dr. Klaus Henning, Dr. Lisa D. Sprague, Dr. Hosny El Adawy, Dr. K. Mertens, Dr. M. Elschner, Dr. F. Melzer, Dr. Gamal Wareth, Tariq Jamil, Roswitha Wehr, Thomas Gkouletsos, Omid Parvizi, MAA Molina and Chrsitopher Sust for their guidance, cooperation and assistance in my research work. Sincere thanks to Prof. Dr. Laeeq Akbar Lodhi, Prof. Dr. Nazir Ahmad, Prof. Dr. Maqbool Ahmad, Prof. Dr. Ijaz Ahmad and Dr. Shujait Ali Rajpoot for their guidance, kindness and moral support during my study. I express my profound sense of appreciation to Associate Prof. Dr. Iahtasham Khan, CVAS, Jhang, who technically guided me about collection, preservation and transport of serum samples to FLI, Germany. I express my profound sense of appreciation to Faisal Shahzad (Berlin, Germany) and members of Pakistani Students Association (PSA) Jena, Germany, especially M. Irfan Badar, Moazzam Bilal, Faraz Rana, Waqas Khan, Inam Khan yusufzai, Nowsherwan Khan, Fawad Khan, Rafiq Malik, Rayan Aziz, Hasan Rahman, Dr. Aurangzaib Jahtool, Dr. Rizwan Mumtaz, Dr. Ali Baig, Faisal Ghafoor Khan, Adnan Nadeem and Majid Bashir for their nice hospitality and care.I am sincerely and earnestly indebted to my Loving parents, Beloved wife, My Kids, brother and sisters, family members and friends, especially Dr. Rifat Ullah Khan and Dr. Mujeeb-Ur-Rehman Nasir, who have always wished to see me glittering high on the skies of success.I am very much thankful to the Higher Education Commission of Pakistan for financial support for this study under HEC Indigenous M.Phil~Ph.D. fellowship for 5000 scholars (Scholar PIN No. 112-34911-2AV1-119). Diagnostic/ laboratory support from Institute for Bacterial Infections and Zoonoses (IBIZ), FLI, Germany is highly appreciated. I am also thankful to the Livestock and Dairy Development Department of Punjab, and the local veterinarians (especially Dr. Muhammad Arif, Dr. M. Usman Mazhar (NIAB), Dr. Aftab Shoukat, Dr. Shabaz Bashir, Dr. Ali Raza, Dr. Mazhar Abbas and Dr. Imran Haider) and farmers for facilitating me in serum sample and ticks collection from small ruminants maintained at livestock farms. Special thanks to Dr. Qudrat Ullah awar, Kifayat Ullah Bettani, Zia-ud-Din Mehsud, Tariq Azam Wazir, Azmat Ullah Dawar, Inam Ullah Wazir, Amin Ullah Bajouri, Haider Khan Sulemankhel, Farooq Shah, Past Gul, Muhammad Haroon, Muhammad Farooq, Syed Bilal, Abidullah Wazir, Roidad Khan, Arsalan said, Ishaq, Aziz Ullah Wazir, Waqar Ali Khan and Anwar Shoaib Wazir for their assistance and support in my research work especially in serum sampling, ticks collection, and their subsequent processing in the laboratories at the University of Agriculture, Faisalabad.(Qudrat Ullah)LIST OF TABLESTable No. Title Page No.2.1World-wide prevalence of Q fever/ Coxiellosis183.1Studied farms and no. of samples collected from each farm433.2Reagents and their volumes present in Q Fever-Indirect ELISA kit453.3Interpretation of Q fever-Indirect ELISA results473.4Contents/ Reagents in High Pure PCR Template Preparation kit483.5Time required for analysis of whole blood and cell culture493.6Working solutions preparation and their use, storage and stability503.7Adjustment of sample volume for DNA extraction503.8Primers and probe sequences used for isocitrate dehydrogenase (icd) real-time qPCR 553.9Conditions for real-time qPCR performed on serum pools553.10Primers and probe sequences used for IS1111 real-time qPCR563.11Conditions for real-time qPCR performed on tick pools563.12Questionnaire for Coxiellosis-Individual Animal Data594.1Farm-wise sero-prevalence of Coxiellosis in small ruminants624.2Farm-wise sero-prevalence of Coxiellosis in goats644.3Farm-wise sero-prevalence of Coxiellosis in sheep664.4District-wise sero-prevalence of Coxiellosis in small ruminants684.5District-wise sero-prevalence of Coxiellosis in goats704.6District-wise sero-prevalence of Coxiellosis in sheep724.7Breed-wise sero-prevalence of Coxiellosis in small ruminants74Table No. Title Page No.4.8Breed-wise sero-prevalence of Coxiellosis in goats764.9Breed-wise sero-prevalence of Coxiellosis in sheep774.10Association of seropositivity against C. burnetii antibodies in small ruminants with history of reproductive disorders794.11Association of seropositivity against C. burnetii antibodies in goats with history of reproductive disorders804.12Association of seropositivity against C. burnetii antibodies in sheep with history of reproductive disorders814.13Association of sero-positivity against C. burnetii antibodies in small ruminants with presence of ticks on sheep and goats834.14Species-wise sero-prevalence of Coxiellosis in small ruminants844.15Association of seropositivity against C. burnetii antibodies with poor body conditions in small ruminants864.16Association of seropositivity against C. burnetii antibodies with poor body conditions in goats874.17Association of seropositivity against C. burnetii antibodies with poor body conditions in sheep884.18Age-wise sero-prevalence of Coxiellosis in small ruminants904.19Age-wise sero-prevalence of Coxiellosis in goats914.20Age-wise sero-prevalence of Coxiellosis in sheep924.21Parity-wise sero-prevalence of Coxiellosis in small ruminants944.22Parity-wise sero-prevalence of Coxiellosis in goats954.23Parity-wise sero-prevalence of Coxiellosis in sheep964.24Sex-wise sero-prevalence of Coxiellosis in small ruminants984.25Sex-wise sero-prevalence of Coxiellosis in goats 994.26Sex-wise sero-prevalence of Coxiellosis in sheep100Table No. Title Page No.4.27Sero-prevalence of Coxiellosis in pregnant and non-pregnant small ruminants1024.28Sero-prevalence of Coxiellosis in pregnant and non-pregnant goats1034.29Sero-prevalence of Coxiellosis in pregnant and non-pregnant sheep1044.30Sero-prevalence of Coxiellosis in lactating and non-lactating small ruminants1064.31Sero-prevalence of Coxiellosis in lactating and non-lactating goats1074.32Sero-prevalence of Coxiellosis in lactating and non-lactating sheep1084.33Real-time qPCR-based prevalence of Coxiellosis in pooled serum samples1104.34Real-time qPCR-based prevalence of Coxiellosis in pools of ticks collected from small ruminants113 LIST OF FIGURESFig. No.TitlePage No.1.1Consequences of C. burnetii infection after invading the gravid uterus33.1Map of Pakistan. Green area on the map showing Punjab province of Pakistan393.2Red dots on the map showing districts of Punjab province from which samples were collected423.3High pure PCR template preparation kit workflow (Roche kit, version 20, Roche Diagnostics GmbH, Germany)523.4Standard curve based on a reference line generated from decimal dilutions of the positive control for qPCR performed on serum pools563.5Standard curve based on a reference line generated from decimal dilutions of the positive control for qPCR performed on tick pools574.1Farm-wise sero-prevalence of Coxiellosis in small ruminants634.2Farm-wise sero-prevalence of Coxiellosis in goats644.3Farm-wise sero-prevalence of Coxiellosis in sheep664.4District-wise sero-prevalence of Coxiellosis in small ruminants694.5District-wise sero-prevalence of Coxiellosis in goats704.6District-wise sero-prevalence of Coxiellosis in sheep724.7Breed-wise sero-prevalence of Coxiellosis in small ruminants754.8Breed-wise sero-prevalence of Coxiellosis in goats764.9Breed-wise sero-prevalence of Coxiellosis in sheep77Fig. No.TitlePage No.4.10Association of seropositivity against C. burnetii antibodies in small ruminants with history of reproductive disorders794.11Association of seropositivity against C. burnetii antibodies in goats with history of reproductive disorders804.12Association of seropositivity against C. burnetii antibodies in sheep with history of reproductive disorders814.13Association of sero-positivity against C. burnetii antibodies in small ruminants with presence of ticks on sheep and goats834.14Species-wise sero-prevalence of Coxiellosis in small ruminants844.15Association of antibodies against C. burnetii with poor body conditions in small ruminants 864.16Association of antibodies against C. burnetii with poor body conditions in goats 874.17Association of antibodies against C. burnetii with poor body conditions in sheep884.18Age-wise sero-prevalence of Coxiellosis in small ruminants904.19Age-wise sero-prevalence of Coxiellosis in goats914.20Age-wise sero-prevalence of Coxiellosis in sheep924.21Parity-wise sero-prevalence of Coxiellosis in small ruminants944.22Parity-wise sero-prevalence of Coxiellosis in goats954.23Parity-wise sero-prevalence of Coxiellosis in sheep964.24Sex-wise sero-prevalence of Coxiellosis in small ruminants984.25Sex-wise sero-prevalence of Coxiellosis in goats 994.26Sex-wise sero-prevalence of Coxiellosis in sheep1004.27Sero-prevalence of Coxiellosis in pregnant and non-pregnant small ruminants102Fig. No.TitlePage No.4.28Sero-prevalence of Coxiellosis in pregnant and non-pregnant goats1034.29Sero-prevalence of Coxiellosis in pregnant and non-pregnant sheep1044.30Sero-prevalence of Coxiellosis in lactating and non-lactating small ruminants1064.31Sero-prevalence of Coxiellosis in lactating and non-lactating goats1074.32Sero-prevalence of Coxiellosis in lactating and non-lactating sheep1084.33Real-time qPCR-based prevalence of Coxiellosis in pooled serum samples1104.34Amplification plots showing fluorescence data acquired during annealing and extension phase of qPCR performed on serum pools1114.35Real-time qPCR-based prevalence of Coxiellosis in pools of ticks collected from small ruminants1134.36Amplification plots showing fluorescence data acquired during annealing and extension phase of qPCR performed on tick pools114ABSTRACTLivestock raising is an important occupation for livelihood of rural poor in Pakistan and plays a vital role in poverty reduction. Coxiellosis is a disease caused by Coxiella burnetii and acts as a major trade barricade adversely affecting the productive and reproductive capabilities of the animal, and hinders with commercialization of animal products at local and international market level. This study was planned to conduct serological and molecular investigations on Coxiellosis and to identify any association of seropositivity against C. burnetii antibodies with sex, age, breed, species, parity, farm, district, lactational status, reproductive status (pregnant or non-pregnant), tick infestation, body condition and reproductive disorders in sheep and goats. A structured questionnaire was used to collect information about individual animal and general farm management. The sampling design was formulated considering an expected prevalence of 50%, confidence interval of 95%, and 5% desired absolute precision. A total of 1000 sera sample (500 from goats and 500 from sheep) were collected from animals maintained at nine different government livestock farms of Punjab. Firstly, all these samples were analyzed through Indirect-ELISA (IDEXX Q Fever, Coxiella burnetii, Antibody Test Kit) and then pools of seropositive, and suspected serum samples (29 pools) were investigated through real-time qPCR, using single copy isocitrate dehydrogenase (icd) gene, for detection of C. burnetii DNA. Additionally, 55 tick pools were also investigated through real-time qPCR, using multicopy IS1111 insertion element, for genomic detection of C. burnetii in these pools. The diagnostic work was carried out at National Reference Laboratory (NRL) for Q fever, Friedrich Loeffler Institute, Jena, Germany. Serological analysis revealed a prevalence of 15.6% (95% CI: 12.5-19.1) and 15.0% (95% CI: 12.0-18.4) in sheep and goats, respectively. Statistically, prevalence of C. burnetii antibodies in serum was non-significantly different (p=0.792, degree of freedom (df)=1, Chi-square (χ2) =0.069) between the two species. A significant association was found between seropositivity against C. burnetii antibodies and different variables like farm (p=0.000, df=8, χ2=141.869), district (p=0.000, df=6, χ22=49.689), breed (p=0.000, df=9, χ2=60.954), lactational status (p= 0.000, df=1, χ22=24.691), reproductive status (p= 0.008, df=1, χ2=7.023), ticks infestation (p=0.000, df=1, χ2=301.914), body condition (p=0.000, df=3, χ2=124.868) and reproductive disorders (p=0.000, df=4, χ2=133.984). However, seropositivity against C. burnetii infection was non-significantly associated with age (p=0.063, df=3, χ2=7.281), parity (p=0.838, df=2, χ2=0.353) and sex (p=0.302, df=1, χ2=1.064) of animal. Univariate analysis revealed a significant (p=0.031, df=1, χ2=4.668) difference in prevalence of C. burnetii DNA in tick pools of sheep and goats, however prevalence of C. burnetii DNA in serum pools was non-significantly (p=0.564, df=1, χ2=0.333) different between the two species. These findings revealed that C. burnetii infection is prevalent in small ruminants maintained at studied livestock farms, as well as in ticks. Further in-depth studies are required to explore its epidemiology more precisely in humans, ticks and various animal species. Chapter-1 INTRODUCTIONLivestock play an important role in food security, poverty reduction and national development. Approximately 8 million Pakistani families are linked with livestock raising obtaining more than 35% earnings from livestock production activities (Anonymous, 2017). According to the Anonymous (2017), the population of sheep and goats in the country is estimated to be 30.1 and 72.2 million heads, respectively. Livestock contributed about 11.4% to national Gross Domestic Production (GDP) and 58.3% to agricultural value added during 2016-17. Gross value addition of domestic animals has increased at a constant factor cost of 2005-06 from Rs. (Rupees) 1288 billion in 2015-16 to Rs. 1333 billion in 2016-17, indicating an increase of 3.4 percent during one year (Anonymous, 2017). Q (Query) fever is a highly infectious disease of bacterial origin, caused by Coxiella burnetii, a Gram-negative coccobacillus and obligate intracellular bacterium which is extremely hardy and ubiquitous in nature. Coxiellosis is a disease of zoonotic importance and is prevalent throughout the world. It has wide range of hosts including cattle, sheep, goats, camels, wild animals and arthropods. Small ruminants act as a reservoir for this pathogen. C. burnetii has also been isolated from ticks. Q fever occurs as a subclinical infection in ruminants (Angelakis and Raoult, 2011; Hadush et al., 2016). Coxiellosis is a zoonotic disease of economic importance and is characterized by reproductive losses including abortion, stillbirth, premature delivery and birth of weak calves. It also adversely affects the succeeding pregnancies (Angelakis and Raoult, 2010). The primary mode of transmission to?humans is?through inhalation?of?C. burnetii contaminated dust or aerosols. Transmission may also occur through intake of uncooked milk and milk products. Birth?products?(including fetuses, placental membranes,?amniotic?and?allantoic?fluids),?milk and excreta of animals are the main sources of infection (Reusken et al., 2011).Q fever is also an occupational disease associated with veterinary doctors, paraveterinary staff, animal researchers, livestock farmers and people working in slaughter houses. It mostly occurs as asymptomatic illness in humans. Sometimes it occurs as an acute febrile illness (usually with self-limiting flu-like symptoms, inflammation of lungs and hepatitis), while chronic form of infection is characterized by severe hepatitis and chronic-fatigue syndrome. A very few of the infected people (~5%) may develop chronic symptoms of the disease mostly in the form of valvular lesions and endocarditis (DeRooij et al., 2012).Antigenic variation is a unique characteristic of C. burnetii and is known as phase variation. It exists in two different antigenic forms i.e., phase I and phase II. Phase I form of C. burnetii is extremely virulent and can be isolated from acutely diseased animals, arthropods and human beings, while Phase II is obtained after several passages of Phase I bacteria in cell culture or embryonated chicken eggs. Avirulent Phase II form of C. burnetii is remarkably different from phase I virulent form in surface protein composition, cell density and surface charge. Phase I has a smooth full length Lipopolysaccharide (LPS) molecule while phase II bacteria contain an incomplete rough LPS molecule (Mertens and Samuel, 2007). The target cells of C. burnetii in the host body are monocytes/macrophages. This micro-organism enters into the target cells through endocytosis/phagocytosis. An integrin avb3 mediates attachment of phase I bacteria to the target cells, while in case of phase II both avb3 and complement receptor CR3 are involved in the attachment to the target cells. C. burnetii is capable of maintaining its intracellular life in the acidic environment of phagolysosomes. The acidic pH of the phagolysosome permits entry of essential nutrients for metabolism of C. burnetii and protects it from antibiotics by disrupting their mechanism of action. It multiplies rapidly within the acidic phagolysosome of host cells with a generation time of 20-45 hours (Mertens and Samuel, 2007; Angelakis and Raoult, 2010).C. burnetii has marked phylogenetic resemblance with bacteria from order Legionellales. It exists in two diverse morphological forms i.e., large cell variant (LCV) and small cell variant (SCV). The SCV is small in size with a compact rod shape possesing high electron-dense center of abridged nucleoid filaments, while LCV is larger in size with less electron-dense center. This LCV is a metabolically active form of C. burnetii present within the cell. It undergoes sporogenic differentiation to yield highly resistant spore-like forms known as small cell variant. These SCV are released upon cell lysis and can exist for long periods of time in harsh environments. They are extremely resistant to heat, drying, chemicals and ultraviolet light (Angelakis and Raoult, 2010; Drozd et al., 2014) Different types of tests can be used for diagnosis of C. burnetii?infection. The disease can be diagnosed by detecting IgG and IgM antibodies against phase I and phase II antigens using an indirect immunofluorescent assay (IFA), or by detecting C. burnetii DNA in blood sample through PCR technique (Anonymous, 2013). The latter is a highly sensitive and specific technique that can be used for detection of C. burnetii DNA both in cell cultures and biological samples (Jones et al., 2010). A positive PCR assay conducted on body secretions like vaginal mucus, milk sample, urine, feces and aborted fetus indicates that animal is infected with C. burnetii. A quantitative?real‐time?PCR may help in analyzing the quantity of organisms responsible for causing adverse pregnancy events (Hazlett et al., 2010). Epidemiological investigation on Coxiellosis based on serum or whole blood samples obtained from different animal species using different diagnostic techniques are necessary for effective risk assessment and preventive measures for humans and animal health. ELISA test is more sensitive for serum samples if antigen is obtained from ruminant isolate as compared to antigen obtained from ticks. So, antigen capture ELISA based on small ruminants isolates is recommended (Touratier et al., 2012). The outcomes of pregnant uterus infected with C. burnetii pathogen can occur in the form of APSW complex (abortion, premature delivery, stillbirth and weak offspring), besides the offspring that are clinically normal and may or may not be congenitally infected. Fig. 1.1 shows the pathways through which C. burnetii infection leads to different pathological conditions after invading pregnant uterus and their localization in placenta (Agerholm, 2013). Figure 1.1: Consequences of C. burnetii infection after invading the gravid uterusData from previous studies indicated that C. burnetii infection may follow one of the two pathways after preliminary localization of bacteria in placenta. Green arrows in Fig. 1.1 (latent infection) indicate that the infection either remains limited to the fetal membranes or it may spread to the fetus. It is probably the most common consequence of this infection, especially in cattle. This latent infection is characterized by normal offspring (though it may or may not be infected congenitally) and presence of bacteria in vaginal secretions during parturition and in postpartum period (Agerholm, 2013). In an active infection (red arrows), there are three possibilities after intrauterine invasion: 1) The infection may remain restricted to fetal membranes, 2) There may be transmission of infection to the fetus through blood (haematogenous route) or 3) The infection may spread to the fetus through amniotic-oral route. These haematogenous and amniotic-oral transmissions of infection will compromise the fetus and results in abortion, premature birth, stillbirth and birth of weak calves (APSW complex), although normal but probably congenitally infected offspring may also be found (Agerholm, 2013). In case of ruminants, after invading the pregnant uterus C. burnetii targets the trophoblastic cells present in the allantochorion of fetal membranes. There is little knowledge on how trophoblastic cells become initially infected. After invasion, more trophoblastic cells are gradually infected and fetal membranes show severe signs of inflammation. However, the trophoblastic cells covering the cotyledonary villi, responsible for nutrient supply and gaseous exchange, are not infected by C. burnetii. The fetal death may occur shortly before or during inflammation induced abortion, or there may be a live birth (Roest et al., 2012; van den Brom et al., 2015). Even though C. burnetii causes disease in many animal species, the infection is mostly asymptomatic. During acute phase of infection, the organism can be found in blood, lungs, spleen and liver of small ruminants. It is not obvious whether Coxiellosis affects the function of organs other than placenta in animals. However, in sheep and goat it results in mild lesions only (Sa?nchez et al., 2006; Roest et al., 2012). In non-pregnant animals, Coxiellosis is asymptomatic. In Pakistan, Coxiellosis/Q fever is among highly neglected diseases both in humans and animal species. So far only five studies (1955- 2016) have reported the prevalence of C. burnetii infection in humans and animal species of Pakistan. Based upon these studies, the prevalence of Coxiellosis ranges from 4.6 to 40% in all livestock species and 10.19 to 26.8% in human beings in the country (Kaplan and Bertagna, 1955; Ahmed, 1987; Ayaz et al., 1993; Zahid et al., 2016; Shabbir et al., 2016). In view of steady increase in prevalence of disease all over the world, the present study was designed to conduct serological, and molecular investigation on Coxiellosis and its relationship with some important risk factors in small ruminants maintained at livestock farms of Punjab, Pakistan. OBJECTIVESThe objectives of this study are summarized below:To conduct serological and molecular investigations on Coxiellosis in small ruminants kept at different livestock farms of Punjab, PakistanTo study association between seropositivity against C. burnetii antibodies and reproductive disorders in small ruminantsAlso to study any relationship between seropositivity against C. burnetii antibodies and variables like sex, age, species, breed, parity, farm, locality of farm, ticks infestation, poor body condition, lactational and reproductive status of the animal Chapter-2REVIEW OF LITERATURE History and BackgroundQ fever was first reported in abattoir workers by Derrick in 1935 in Brisbane, Australia, during an outbreak of a febrile illness of unknown origin, with flu-like symptoms. Later, a rickettsia-like organism was isolated from infected mice by Burnet and Freeman in 1937. They named it as Rickettsia burnetii. Then American and Australian researchers started working together on this bacterium and revealed that the Nine Mile agent and Australian Q fever agent, the zoonotic agent, were in fact the isolates of similar microorganism known as Rickettsia burnetii (Burnet and Freeman, 1937). This bacterium was then renamed as ‘Coxiella burnetii’ in the honour of Cox and Burnet, as pioneers of Q fever (Burnet and Freeman, 1937; Derrick, 1939; Philip, 1948; Maurin and Raoult, 1999)The name Q fever or Query fever was first suggested by Derrick in 1937 for C. burnetii infection because the etiopathogensis of this disease was unknown. The term ‘Q fever’ is used for C. burnetii infection in humans, while in animals, the disease is named as ‘Coxiellosis’ (Mori et al., 2017). Several other synonyms are used for this disease such as Coxiellosis, Australian Q fever, abattoir fever, Balkan influenza, Nine-mile fever, and Pneumorickettsiosis (Vellema and Van den Brom, 2014; Hadush et al., 2016). Influenza like infection called “Balkangrippe” was reported in soldiers from Balkan region (1940) and in German, and American troops during the Second World War (1944-45), which was later identified as Q fever (Georgiev et al., 2013; Vellema and Van den Brom, 2014).Coxiella burnetii is present almost everywhere in the world (Hagenaars et al., 2014). It is strictly an intracellular bacterium and has wide range of hosts, including ticks, fish, reptiles, birds, warm blooded animals such as ruminants and human beings etc. (Cutler et al., 2007). Ruminants, especially sheep and goats, are considered as major reservoir and potential risk for human infection. Therefore, for proper control and prevention of infection in humans, animals and environment, it is important to understand the disease in ruminants (Abdel-Moein and Hamza, 2017). Aerosol is the major route of Q fever transmission. It occurs mainly through inhalation of C. burnetii contaminated dust or air. The recent outbreak of Q fever in the Netherland was linked to wind dispersion from site where infected goats were kept (Hadush et al., 2016). In ruminants, after transmission the organism enters into the blood stream and reaches the predilection sites i.e., placental membranes, supramammary lymph nodes and mammary glands, where its replication occurs (Hadush et al., 2016). The ability of C. burnetii to multiply within the lysosomal acidic vacuole of phagocytic cells, two distinct morphological forms, and variation in LPS molecule of Phase I and Phase II bacteria are some of the unique characteristics which make it distinctive from other bacteria (Mori et al., 2017). It can exit in two different antigenic forms. The first one is metabolically dormant SCV, which is highly resistant form of coxiella, and the second form is LCV which is metabolically active form of C. burnetii present within the host cell (Boden et al., 2014). According to World Health Organization (WHO), Brucellosis, Q fever and Rifat valley fever are neglected zoonotic diseases (NZDs) which have almost been eliminated from the developed countries but are under diagnosed and under reported in the developing countries (WHO, 2015; Kanouté et al., 2017). The Center for Disease Control and Prevention (CDC), classified this bacterium as category B critical biological agent because of its high infectivity and aerogenic transmission to humans (Rotz et al., 2002; Boden et al., 2014). The bacterium can be used as bio-terrorism agent to induce an acute debilitating disease in target population but it will not cause high mortality (Seo et al., 2016).C. burnetii- the etiological agentC. burnetii is a Gram-negative, strictly intracellular, pleomorphic bacterium ranges in size from 0.2-0.5 μm (width 0.2-0.4 μm and length 0.4-1.0 μm in). It belongs to kingdom Bacteria, phylum Proteobacteria, class Gammaproteobacteria, order Legionellales, family Coxiellaceae, genus Coxiella and species C. burnetii (Million and Raoult, 2015; Bielawska-Dro? et al., 2014).The incubation period of disease is quite variable (2-4 weeks or even more) depending upon the inoculation dose, route of infection and antigenic phase of C. burnetii. An important characteristic of this pathogen is the presence of structurally and antigenically distinctive lipopolysaccharide (LPS) molecule in their cell wall. Based upon the structure of LPS, it exists in two distinct antigenic forms which are Phase-I and Phase-II. Phase-I is a virulent form having a full length LPS molecule and is isolated from infected hosts. Phase II is avirulent form of C. burnetii having an incomplete or truncated LPS molecule without terminal O-antigen sugar. Phase II is obtained by repeated passages of Phase I in embryonated eggs or in cell cultures. In C. burnetii genome, the 38 kb region is the site where all LPS coding genes are present. It has been reported that antigenic phase variation occurs due to chromosomal deletions in this region (Shapiro et al., 2015; Kuley et al., 2015; OIE, 2015). Similarly, C. burnetii can exist in two different morphological forms which can be differentiated under electron microscope. One is LCV and the other is SCV. LCV is large, bacilliform and metabolically active form, while SCV is a small, coccoid and metabolically dormant form of C. burnetii. SCV is highly resistant to environmental stress factors like heat, drying and many disinfectants and thus can survive in the environment for extended period of time (Bielawska-Dro? zd et al., 2014; Schleenvoigt et al., 2015; Hadush et al., 2016)The genus Coxiella also possesses some other putative species like C. cheraxi found in cray fish and a novel Coxiella-like organism present in birds and ticks. C. cheraxi is a Coxiella like organism, having highest genetic homology with C. burnetii. Candidatus Coxiella avium is another novel pleomorphic Coxiella-like organism isolated from the birds. It multiplies within the acidic vacuole of host macrophage cells, leading to systemic infection and mortality. Similarly, Coxiella-like endosymbionts (CLE) are also present in ticks. These are closely related with C. burnetii but are genetically different, indicating that some diversity exits within Coxiella genus (Seo et al., 2016; Mori et al., 2017).The virulence of C. burnetii is associated with the type of strain involved in causing the disease. Different strains are having different virulence potency. Four different strains of C. burnetii were found through Multispacer Sequence Typing (MST) technique. Sequence type (ST) 8 from sheep in France, ST 15 from goats in France, ST 16 was isolated from ticks in Montana (USA) and ST 20 from cattle in France (Sidi-Boumedine et al., 2015).Isolation and PropagationIsolation and propagation technique is not commonly used for the routine diagnosis of C. burnetii due to its zoonotic nature as well as time consuming, laborious, biosafety level (BSL) 3 laboratories restrictions and high level of expertise required. This technique is useful when some new clinical presentation or unusual epidemiological situation of the disease is encountered. It is also important for phenotypic and genotypic molecular characterization of C. burnetii,?using different genotyping methods such as multi-locus variable number of tandem repeats analysis (MLVA) and Multispacer Sequence Typing (MST) etc. Similarly, isolation of organism is useful for whole genome sequencing and to find out new strains for future research. Sample quality, condition and concentration of pathogen in a sample are the factors which strongly affect the success rate of isolation and propagation technique (Berri?et al., 2000; Selim and Elhaig, 2016; Mori et al., 2017). Though C. burnetii is highly resistant to the environmental factors and can survive outside the host for longer period, but it requires host cells for multiplication (Bontje et al., 2016). For proper propagation and isolation of C. burnetii, a concentration above 105 bacteria per ml is recommended (Samuel and Hendrix, 2009). Different methods can be used for isolation and propagation of C. burnetii, including inoculation of embryonated chicken eggs, cell culture, laboratory animals and a novel axenic medium (Mertens et al., 2017). 2.3.1. Embryonated egg inoculation Embryonated egg inoculation method had been used traditionally for direct isolation and propagation of C. burnetii, but nowadays, it is not a recommended method. In this technique, 6-7 days old embryonated chicken eggs are properly disinfected so that they are free of specific pathogens. The yolk sac is then located using a candle light and a suspension of C. burnetii is inoculated via the yolk sac route. The egg shell at the site of injection is then sealed and eggs are incubated at 35-37°C till day 21. Embryos that die after 5 days of inoculation are discarded. This embryonic death may occur because of heavy bacterial load. After 10- 15 days of incubation, the yolk sac is harvested. Stained smears of the yolk sac wall are observed under microscope to assess the presence of C. burnetii and absence of bacterial contamination. A typical straw yellow color with white spot patches develops in infected yolk sacs, while uninfected yolk sacs are of orange color with viscous consistency. PCR is a useful tool to detect the presence of C. burnetii DNA in chicken emrbyos samples after inoculation (OIE, 2015; Mori et al., 2017).2.3.2. Cell cultureSince C. burnetii is an intracellular pathogen, standard biological media are not suitable for its growth (Kersh?et al., 2013). Raoult et al., (1990) described a cell culture system, known as the shell vial cell culture, for isolation and propagation of obligate or facultative intracellular bacteria such as C. burnetii. This culture system is commonly used for isolation of viruses. In this technique, a suspension containing C. burnetii infected material is inoculated into human embryonic lung (HEL) fibroblast cells grown within egg shell vial on 1 cm2 cover-slip. The HEL fibroblasts are the most commonly used cells for isolation and propagation of C. burnetii because of their easy maintenance and ability to keep the monolayer integrity during long incubation periods. Various other cell lines like from epithelial lining (Vero E6), macrophages (P388D1, J774, DH82) and fibroblastic cells (L929, HEL) may be used for C. burnetii replication (Maurin and Raoult, 1999; Mediannikov et al., 2010; Santos et al., 2012). After inoculation, centrifugation is performed for 1 hour at 700g, so that the bacteria properly stick to the cells. Three shell vials are used for the same inoculum. At day 3, 10 and 21 post inoculation, C. burnetii vacuoles can be seen under an inverted microscope. After 10 days, proliferating C. burnetii inside the cells are detected directly on the coverslip within egg shell vial by a direct IFA, using polyclonal anti-C. burnetii antibodies and a proper secondary antibody conjugated to fluorescein isothiocyanate (FITC). Cells present in the remaining shell vial are harvested and shifted to another 25 cm2 culture flask. The culture is then incubated with 5% CO2 at a temperature of 37°C for 2 months which may be extended up to 4-5 months. During incubation, change of culture medium once a week and periodical evaluation of bacterial growth using either light or fluorescence microscopy is required. If Cytopathogenic effects (CPE) are observed in the culture medium and Gimenez staining or PCR gives positive results, then sub-cultures are carried out in a 75 cm2 culture flask. Supernatant is inoculated on confluent layers of Vero cells or fibroblasts (L929, mouse) in a 150 cm2 culture flask to obtain new C. burnetii isolates. Though this cell culture method was established for humans it can be used efficiently for animals as well (Maurin and Raoult, 1999; OIE, 2015; Mori et al., 2017).2.3.3. Laboratory animalsIt is useful technique for isolation and propagation of Coxiella obtained from contaminated samples like ticks or those obtained from animals e.g., faeces, milk, vaginal discharges and fetal parts of placenta. Laboratory animals will act as filtration system for such cases. Mice and guinea pigs are commonly used for this purpose. Following intraperitoneal inoculation with a dose of 0.5 ml per animal, body temperature and antibody titer can be examined. This protocol should be carried out in conjunction with serological assays on other laboratory animals (mice and guinea-pigs) that have been inoculated with the same samples. Sera are collected 21 days after inoculation. A positive result reveals the diagnosis of C. burnetii infection. The results can be confirmed through real-time PCR or by microscopy, using impressions and stained samples of collected spleen, liver and lungs. Splenomegaly is a typical sign caused by substantial growth of C. burnetii. Spleen, liver or lung samples are then inoculated into embryonated chicken eggs or in cell cultures for isolation of C. burnetii (Scott et al., 1987; Santos et al., 2012; OIE, 2015; Mori et al., 2017).2.3.4. Axenic mediaAxenic medium is a novel technique used for isolation and growth of C. burnetii, developed by Omsland et al. (2008). This medium is also called Complex Coxiella Medium (CCM). It is improving continuously. Recently used axenic medium is acidified citrate cysteine (ACC) medium. The actual formulation, designated as defined acidified citrate cysteine medium (ACCM-D), allows replication of Coxiella over 14 days with morphological differentiation (SCV/LCV). In this medium, antimicrobial factors such as proteolytic and hydrolytic enzymes, and an acidic environment similar to that present in the acidic phagolysosome of antigen presenting cells is provided. Henec, axenic medium provides an ideal environment for the growth of C. burnetii. This medium is quite useful for genotypic and phenotypic characterization of C. burnetii variants (Omsland et al., 2008; Omsland and Heinzen, 2011; Mori et al., 2017).According to PMA-PCR (Propidium monoazide-PCR) based study, there is no difference in viable cell count obtained through cell free system (axenic medium) and the cell based culturing system. Moreover, axenic medium does not influence the liveability of cell, relative virulence and antigenic phase variation of C. burnetii as compared to cell-based culturing system (Kuley et al., 2015).Genome and genetic characterizationGenetic characterization of a pathogenic organism such C. burnetii is useful for surveillance purpose and for epidemiological investigation of disease outbreaks. It is also a useful tool to investigate the genotypic variation of a pathogen in a geographical area, and to explore interactions between various types and sub-types of the bacterium. Molecular characterization of pathogenic bacteria is carried out to determine the routes of disease transmission and source of infection (Astobiza?et al., 2012; Sulyok et al., 2014; Pi?ero et al., 2015). Genetic informations are also required to establish an epidemiological link between source of disease outbreak and human cases. This helps in planning a control program for potential reservoirs involved in the life cycle of C. burnetii (Klaassen et al., 2009; Roest et al., 2011; Astobiza?et al., 2012). As a major zoonotic problem, gene level studies on C. burnetii infectivity and pathogenicity are obligatory to know its epidemiology more precisely (Brookea et al., 2015). Several techniques including pulsed-field gel electrophoresis (PFGE), sequence analysis or restriction fragment length polymorphism (RFLP) of single genes can be used for molecular typing of C. burnetii. All these techniques have limitations like poor discriminatory power and an inappropriate reproducibility and transferability (Sulyok et al., 2014). However, two recently PCR-based typing techniques i.e., MLVA (multi-locus variable number of tandem repeats analysis) and multispacer sequence typing (MST) are widely used for molecular typing of C. burnetii nowadays. Both typing systems possess high discriminatory power and are easily reproducible. MST is based on variation in 10 short intergenic regions in a DNA sequence and can be conducted directly on DNA extracted from clinical and environmental samples without need for isolation of C. burnetii. As MLVA and MST having high discrimatory power for genome characterization, it allows the identification of up to 36 different genotypes of C. burnetii (Polo et al., 2015; Leuken et al., 2016; Selim and Elhaig, 2016).The first whole genome sequencing from Nine Mile RSA 493 reference strain of C. burnetii, obtained from an infected group of Dermacentor andersoni ticks in 1935, was released in 2003. Random shotgun technique was used for this sequencing targeting sequence spanning 1,995,275 base pairs. In 2007, another genome sequencing report was published using Henzerling RSA 331 strain obtained from blood of an infected patient in Italy in 1945. The genome of C. burnetii is circular in shape, with about 1.9 to 2 Mbp. It consists of one of the following five plasmids: QpDG (51 kb), QpRS (39 kb), QpH1 (36 kb), QpDV (33,5 kb), plasmid of Chinese isolate (56 kb) or a chromosomal integrated plasmid associated sequences (16 kb). The presence of pseudogenes in high numbers in the genome of C. burnetii indicates that the bacterium undergoes genome reduction (Seshadri et al., 2003; Mori et al., 2017; Mertens et al., 2017). Whole genome sequencing (WGS) technique is becoming more affordable; however, data interpretation remains time consuming, as well as requires special expertise, technical skills, bioinformatics knowledge and additional funding. The most communal MLVA genotype A found in animal species is acquired from caprine, ovine, rats and environmental samples. The presence of this highly prevalent genotype A in humans, environment and different animal species complicates the finding of accurate source of infection (de Bruin et al., 2012; Mori et al., 2017). TransmissionThe major route of acquiring C. burnetii infection is aerosol, while intake of contaminated raw food materials is a minor source of disease transmission. Occasionally, the infection may occur through skin, mucosal contact with contaminated products, blood transfusion or mating (Baziaka et al., 2014; Million and Raoult, 2015). In animals, the disease may spread vertically or sexually but this is not the main route of transmission. However, arthropods, especially ticks, may play an active role in the disease transmission in animals (OIE, 2015). Different body excretions and secretions of animals also contain large amount of C. burnetii, which may result in sexual and vertical transmission of the disease. Body secretions like milk and saliva, and excretion materials like parturition products, aborted materials, urine and feces of the infected animal contains the infectious agent. These body discharges can get dried and combine with dust, ultimately leading to human exposure (Miceli et al., 2010; Bontje et al., 2016). Aerogenic transmission of the disease from contaminated sites to human depends upon atmospheric dispersion and impact of environmental factors on deposition and reaerosolisation. Environmental factors like temperature, prevailing weather conditions, landscape, wind direction etc. strongly influence transmission of Q fever (Leuken et al., 2016). Although human to human transmission of Q fever is very rare, it may occur following contact with parturient women. Transplacental transmission, cutaneous inoculation and postpartum spread of Q fever occur in sporadic cases. The disease is also reported in more than 40 species of ticks from family Ixodidae and family Argasidae, and some other arthropods that feed on animals (Miceli et al., 2010; Bontje et al., 2016). Transmission of C. burnetii infection through tick bite has been proposed in animals, though it is not considered as major cause of infection, while in case of humans it is still dubious. However, they can transmit Coxiella both transovarially and transstadially to their offspring, thus acting as a reservoir. Infected ticks excrete large amount of Coxiella in their feces, which contaminate the skin of host animals. So, the ticks are important for environmental spread of Coxiella infection (Toledo?et al., 2009;?Sprong?et al., 2012;?Cong?et al., 2015; Seo et al., 2016).Occurrence of C. burnetii in different body fluids and tissuesShedding of C. burnetii can occur in different body fluids, and tissues like milk, faeces, urine, birth fluids, vaginal secretion and fetal membranes. Particularly, in case of reproductive failure, a large quantity of bacteria is shed through vaginal secretions and birth fluids (Agerholm, 2013; Khaled et al., 2016). It is reported that approximately 1 billion of C. burnetii/ gram of placenta are excreted in birth fluids of an aborted animal (Hadush et al., 2016). Similarly, placenta of seropositive sheep and goats contain more than 109 hamster infective doses of C. burnetii/ gram of tissue, although a single bacterium is enough to develop Q fever infection (Mori et al., 2013; Vellema and van den Brom, 2014; Freick et al., 2016; Hadush et al., 2016). The organism can shed in body fluids for variable period of time, depending upon animal species and shedding routes. Infected cattle can persistently shed pathogen in their milk for several months without any clinical signs or symptoms, while shedding through vaginal mucus or faeces is sporadic or intermittent in nature (Lucchese et al., 2015). Real-time qPCR is a useful technique to determine the load of bacteria in vaginal and milk samples. During acute phase of Q fever, 104-108 C. burnetii were found per vaginal swab, while 102-106 C. burnetii per milk sample were present. Shedding of pathogen declined continuously within two months to less than 104 bacteria per vaginal swab and 102 per milk sample. At the end of this study, a 10 fold increase in bacterial shedding was reported (Guatteo et al., 2007; Sting et al., 2013). Seropositive animals are mostly found to shed C. burnetii in feces, milk or vaginal discharge; however, some seropositive animals may not excrete the organism. Similarly, some apparently healthy animals may shed the organism although they are seronegative for Q fever. The organism can be excreted in large quantities even during normal parturition (Saglam and Sahin, 2016; Mori et al., 2017).Prevalence of Coxiellosis in milk samples obtained from different ruminants like sheep, goats and cattle may vary due to notable differences in their shedding routes. The principal routes of bacterial shedding in sheep are faeces and vaginal fluids, while these are minor routes in cattle. Milk and blood are not common routes of bacterial shedding in sheep. In cattle, milk is considered as major route of bacterial shedding. Goats can excrete organism through vaginal mucus, faeces and milk. However, major routes of bacterial shedding in caprine species are feces and blood, not the milk. Feces contain the highest numbers of bacteria in goats (Vellema and van den Brom, 2014; Mohammed et al., 2014). PathogenesisC. burnetii possesses a distinct characteristic called Phase variation i.e., it has two phases. Phase I bacterium has a complete LPS molecule and is highly virulent. This virulent form of bacteria can be isolated from infected animals, human beigns and ticks. However, Phase II bacterium is avirulent and can be obtained after serial passages of Phase I bacterium in cell culture or embryonated hen eggs. LPS of phase II is rough and severely truncated. The LPS molecule is an integral part of external cellular membrane of gram-negative pathogens and is important for host-pathogen interaction. It also contributes to immunogenicity and pathogenicity of bacteria. Besides LPS, the two antigenic form of C. burnetii also differ from each other in cell density, surface charge and surface protein configuration (Shah et al., 2015; Mertens et al., 2017). There are two different morphological forms of C. burnetii; Large Cell Variant (LCV) and Small Cell Variant (SCV). These two forms of bacteria differs morphologically and functionally from each other. The LCV is larger in size with less electron-dense center, while SCV is a metabolically inactive and less replicating form, with compact rod shape and dense central region. These SCVs are excreted by infected animals, leading to environmental contamination (Selim and Elhaig, 2016). In humans, Q fever is mostly acquired from inhalation of C. burnetii contaminated aerosol and to a lesser extent, by ingestion of contaminated milk and their products. Similarly in animals, the main route for disseminating C. burnetii infection is through aerosols (Woldehiwet, 2004). Once the organism enters into the body, it attaches to the cell membrane of phagocytes. These phagocytic cells (monocytes/macrophages) act as target cells for C. burnetii. Attachment of virulent bacteria to the phagocytic cells is triggered by avb3 integrin, while in case of avirulent bacteria, both avb3 and complement receptor CR3 mediate the attachment. After inoculation, phase I bacteria have the ability to survive within the phagocytic cells, while phase II bacteria are eliminated. Also phase I bacteria are phagocytosed by the host cells in a considerably lower amount than Phase II bacteria (Angelakis and Raoult, 2010). After transmission, the SCVs are phagocytosed by monocyctes and macrophages, and then enters in to the phagolysosome of host cell. Inside the phagolysosome, SCVs fuses with the lysosomal contents of the cell and changes into metabolically active form. This metabolically active form of SCVs can undergo vegetative growth and ultimately transform into LCVs. Normally, both the antigenic forms of C. burnetii are present within this phagolysosomal niche, however, phase II bacteria are quickly eliminated. This acidic environment of phagolysosome is highly conducive for the growth of C. burnetii. Most important is the organism’s ability to propagate and multiply within the acidic phagolysosome and its tendency to develop persistent infection. The entire developmental cycle of a metabolically active phase I bacteria occurs within this acidic niche (Roest, 2013; Van Schaik?et al., 2013; Selim and Elhaig, 2016). (Woldehiwet, 2004). The adjustment of C. burnetii to its intracellular life is associated with lower pH of phagolysosome within host cell. This acidic pH ensures the availability of nutrients essential for growth of C. burnetii and also protects it from the effect of various antibiotics by changing their mode of action (Angelakis and Raoult, 2010). Cellular immunity plays an active role in elimination of pathogen from infected host. However, little is known about the role of host’s immune system in Q fever patients. The goat’s immune response against C. burnetii infection revealed that both IgG and IgM phase II specific antibodies can be found 2 weeks post-infection and it remains elevated in the blood up to 13 weeks. Phase I antibodies develop four weeks after the development of Phase II antibodies. Duration of immune response against C. burnetii is not exactly determined. However, some previous studies revealed that immune response can persist for several months to years (Van den Brom et al., 2015). The metabolically active LCVs are present in different cells within the host body; however, the main target cells are the trophoblasts of placenta (Van den Brom et al., 2015). During acute infection, the organism is also present in blood, liver, spleen and lungs of the host. The disease is mostly asymptomatic in non-pregnant animals, while in pregnant animals most important clinical manifestations are abortion, stillbirth, birth of weak offspring and premature delivery. Incidence of respiratory and digestive problems in apparently healthy kids, rearing in at-risk areas, can be associated with Q fever infection. Although reproductive disorders are not common consequences of Q fever in domestic animals, increased abortion rates up to 90% have been reported in goats (Wouda and Dercksen, 2007; Van den Brom et al., 2015).In human beings, C. burnetii infection appears both as an acute and chronic infection. Acute infection is often self-limiting with mild flu-like symptoms, while chronic Q fever is life threatening with chronic endocarditis in most of cases (Chakrabartty et al., 2016). In abortions due to C. burnetii infection, fetuses usually look fresh and normal, however, sometimes fetuses may be necrotic. Macroscopically, there is inflammation of placenta with purulent yellow-brownish exudate in severely affected inter-cotyledonary spaces. Microscopically, the trophoblastic cells present at the base of villi and in inter-cotyledonary area of allanto-chorion are mostly affected. This inflammation may vary from mild mononuclear infiltration to chronic necrosis with pus like discharge. The epithelial cells present in chorionic membrane at the base of villi often have basophilic intracytoplasmic granulation and a foamy vacuolated cytoplasm. Histopathological examination of some fetuses showed inflammation of liver with mild granulation, however other organs were found apparently normal (Van den Brom et al., 2012; Van den Brom et al., 2015). EpidemiologyInvestigation on epidemiology of Coxiellosis is complicated due to sub-clinical manifestation of the disease on one side and multi-factorial nature of abortion on other side. Moreover, detailed veterinary investigations, including efforts towards laboratory confirmation of the causative agent following a single abortion in a herd or flock, are usually not carried out (Roest et al., 2011). The largest reported Q fever outbreak has occurred in Netherland during 2007-2010 involving approximately 4000 human cases. This outbreak was linked with infected dairy goat farms situated near residential areas, and most likely involved human exposure through windborne route (Schimmer et al., 2009). During this outbreak, an average of 20% (10-80%) of the pregnant goat population aborted, while at two other suspected sheep farms located in the same area, the estimated abortion rate was 5% (Van den Brom and Vellema, 2009). C. burnetii infection has been reported worldwide with exception of New Zealand and French Polynesia (Million and Raoult, 2015; Musso et al., 2014; Maurin and Raoult, 1999). Regarding global distribution, the disease has been reported from all continents including Africa, Asia, Europe, Oceania, North America and South America (Million and Raoult, 2015) with exception of Antartica (Anonymous, 2013). Prevalence of Q fever is high in ruminants compared to humans. Prevalence ranges from 1.1 to 58.4% in sheep, 6.5 to 65.8% in goats and 0.6 to 46.6% in cattle. Old age, female sex, temperature, large dairy herd size and drinking water from same watercourse are considered potential risk factors for this infection (Carbonero et al., 2015). In addition, dry weather, wind direction, low vegetation densities, low ground water level and climate change are also considered risk factors for spread of C. burnetii infection (Klaasen et al., 2014; Bontje et al., 2016). Both from animal and public health view point, prevalence studies are important to support risk assessment. Epidemiological surveys on Q fever will assist the policy makers in decision making processes (Ohlson et al., 2014). The data given in (Table 2.1) reflect the prevalence of C. burnetii infection in different countries of the world. Table 2.1: Global prevalence of Q feverContinentCountrySpecies% PrevalenceReferencesASIA1.AfghanistanHuman63.9Akbarian et al. (2015)Sheep43.4Goat52.7Cattle5.22.AzerbaijanHuman60.2 Aslanova et al. (2009)3.BangladeshSheep9.52Rahman et al. (2016)Goat3.33Cattle3.574.ChinaGoat22El-Mahallawy et al. (2016)Cattle29Human25Pig3Dog0Cat05.Hong KongHuman8 cases reportedChan et al. (2010)6.IndiaCattle4.54Das et al. (2014)Buffalo8.33Dog07.IndonesiaRuminant7.5Setiyono and Subangkit (2014)8.IranSheep33.9Ezatkhah et al. (2015)Goat22.49.IraqCow9.3Abed et al. (2010)Sheep5.810.IsraelHuman88Amitai et al. (2010)11. JapanSheep8.64Giangaspero et al. (2012)12.JordanSheep7. 9Aldomy et al. (1998)Goat7 .113.KazakhstanSaiga antelope0.07Orynbayev et al. (2016)14.Lao People’s Democratic RepublicBuffalo0Douangngeun et al. (2016)Cattle2.5Goat0Pig015.LebanonHuman19.3Gababedian et al. (1956)16.MalaysiaFarm workers100Rai et al. (2011)Vet staff37.77Animal farms27.217.NepalHuman0.15Thompson et al. (2015)18.North KoreanHuman21 cases reported (2010-11)Bennett et al. (2013)19.OmanHuman9.8Scrimgeour et al. (2003)Goat5220.a. PakistanSheep17.9Shabbir et al. (2016)Goat16.4b. PakistanSmall ruminant30.8Zahid et al. (2016)c. PakistanHuman phase-II antibodies11.8Ayaz et al. (1993)Human phase-I antibodies10.19Sheep phase-II antibodies21.28Sheep phase-I antibodies18.44d. PakistanHuman26.8Ahmed (1987)Buffalo34.5Cows10.4Sheep18.3Goat4.6Rodent18.0e. PakistanCamel40Kaplan and Bertagna (1955)21.Saudi ArabiaCamel14.83Mohammed et al. (2014)Goat16.09Sheep0Cattle24.4422.South KoreaKorean native cattle1.7Lyoo et al. (2017)dairy cattle10.5Dog2.923.Sri LankaHuman1.6Angelakis et al. (2012)24.TaiwanDog13.2Lai et al. (2017)Cat2.2Rat2.8Pig3.1Cattle0.6Goat0.9Chicken3.8Other animals4.125.ThailandHuman6.4Blacksell et al. (2015)26.TurkeyVeterinarian25.9?etinkol et al. (2017)Buthcer20.0Farmer29.4Hunter22.2Sheep2K?l?? et al. (2016)27.United Arab EmiratesCamel7.9Afzal et al. (1994)AFRICA1.AlgeriaSmall ruminant14.1Khaled et al. (2016)2.Burkina FasoHuman13.1Ki-Zerbo et al. (2000)3.Cape VerdeHuman81.6Miorini et al. (1988)domestic animals 84.74.CameroonCattle31.2Scolamacchia et al. (2010)5.Central African RepublicCattle14.3Nakoune et al. (2004)6.ChadHuman1Schelling et al. (2003)orosHuman one case reportedBrouqui et al. (2005)8.Cote d'Ivoire orIvory Coast Sheep9.4Kanouté et al. (2017)Goat12.4Cattle13.99.EgyptSheep0Abdel-Moein et al. (2017)Goat3.4Cattle0Buffalo0Veterinarians and veterinary assistants9.4Farmers30.810.EthiopiaCattle31.6Gumi et al. (2013)Camel90Goat54.211.GambiaSheep18.5Klaasen et al. (2014)Goat24.212.GhanaHuman (children)16.9Kobbe et al. (2008)Human (adult)8.913.KenyaHuman 2.5Wardrop et al. (2016)Cattle10.514.MalawiCattle1.5Staley et al. (1989)15.MaliHuman 40Steinmann et al. (2005)16.MoroccoSheep15.3Benkirane et al. (2015)Goat27.317.NamibiaHuman 26.1Noden et al. (2014)18.NigeriaCattle14.5Tukur et al. (2014)19.SenegalHuman 7.04Mediannikov et al. (2010)Ticks8.7620.South AfricaCattle8Vanderburg et al. (2014)Human0Cattle and sheep abortion history 10021.SudanCattle29.92Hussien et al. (2017)Camel64.522.TanzaniaHuman20.3Crump et al. (2013)23.TogoHuman35.97Dean et al. (2013)Cattle14.5Sheep15Goat6.624.TunisiaHuman8.5Kaabia et al. (2006)25.ZambiaGoat7.5Qiu et al. (2013)Cattle12.8826.ZimbabweHuman37Kelly et al. (1993)Cattle39Goat15Dog10OCEANIA1.American SamoaHuman0Lau et al. (2016)2.AustraliaBeef cattle16.8Cooper et al. (2011)Common northern bandicoot31.4Cooper et al. (2013)Eastern grey kangaroo41.1Agile wallaby60Red kangaroo0Common wallaroo66.6Brushtail possum0Rufous bettong1Black-striped wallaby13.French PolynesiaHuman0Musso et al. (2014)Ticks 04.New ZealandNew Zealand is free from Q feverCutler et al. (2007); Musso et al. (2014)5.TongaGoats0Saville (1996)EUROPE1.Europe (overall)In general population (human)From 2.4% to >30% in some Mediterranean countriesEuropean Centre for Disease Prevention and Control (2011)2.AlbaniaCattle13.51Ylli et al. (2013)Sheep9.79Goat16.433.ArmeniaHuman5.6Tarasevic et al. (1976)Cattle24.9Sheep14.5M. arvalis6.7M. musculus3.24.AustriaHuman0Rapf et al. (2015)5.AzerbaijanHuman60.2Aslanova et al. (2009)6.BelarusTicks0.9Reye et al. (2013)7.BelgiumWoolsorters/Human50.7Wattiau et al. (2011)8.Bosnia HerzegovinaHuman35.2Hamzic et al. (2006)9.BulgariaSheep11.5 Martinov et al. (2007)Goat13.69Cattle8.53Human 15.7810.CroatiaSheep7.1Racic et al. (2014)Goat29.5Cattle2.7Human 3.1Horse42.211.CyprusSheep33Cantas et al. (2011)Goat50Cattle3512.DenmarkBeef cattle3.54Paul et al. (2014)Crossbred cattle2.98Dairy cattle9.4513.FranceSheep20.0Georgiev et al. (2013)Goat88.1Cattle15.0Human 1.0 to 4.014.GermanySheep8.7Georgiev et al. (2013)Goat2.5Cattle19.3Human 2215.GreeceHuman 48.7Vranakis et al. (2012)16.HungaryCattle26Kreizinger et al. (2015)Sheep47.6Goat20Red deer5.55Roe deer0Fallow deer017.IrelandSheep0.7Ryan et al. (2011)Goat0.318.ItalySheep15.5Rizzo et al. (2016)Goat16.2Goats in mixed flock25.7Sheep in mixed flock16.319.MacedoniaCattle7.14Saiti and Memishi (2015)Sheep26.37Goat6.620.MontenegroSheep5.03Lau?evi? et al. (2011)herlandsSheep3.5Georgiev et al. (2013)Goat7.8Cattle21Human 12.2 to 2422.NorwayDairy cattle0Kampen et al. (2012)Beef cattle0Dairy goats0Sheep023.PolandMilkmaids60Chmielewski et al. (2013)Farm workers28Veterinary staff44Incidental visitors12Raw milk consumers1924.PortugalSheep10.53Cumbassá et al. (2015)Goat23.53Cattle20.83Mongooses11.1Antelopes12.5Giraffe10025.RussiaMeat processing and packing plant workers3.6Ignatovich et al. (2003)Tannery workers27.6Cattle breeders25.6Control group026.SerbiaHuman43 cases reportedMedi? et al. (2012)27.SlovakiaSheep58.42 Dorko et al. (2010)28.Spaindomestic sheep12.7FernándezAguilar et al. (2016)domestic goat0Cattle1.1European mouflon6.8red deer2.4fallow deer0roe deer0Southern chamois029.SwedenCattle8.2Ohlson et al. (2014)Sheep0.6Goat0Moose030.SwitzerlandSheep1.8Magouras et al. (2017)Goat3.4Human1.7 and 3.531.UkraineWorkers of meat-packing factories6.9Fedorov et al. (1983)Workers of fur- and wool-treating establishments8.7Stock breeders5.2Large cattle 2.3Small cattle 5.632.Great BritainSheep0.9Lambton et al. (2016)Goat0.8NORTH AMERICA1.Antigua-BarbudaPregnant Women5.3 ± 6.6Wood et al. (2014)2.BermudaPregnant Women0Wood et al. (2014)3.CanadaSheep and goat64.5Meadows et al. (2016)4.DominicaPregnant Women0Wood et al. (2014)5.El SalvadorDairy cattle26Rice et al. (1979)6.GreenlandHumanone case reportedKoch et al. (2010)7.GrenadaSheep0Stone et al. (2012)Goat08.JamaicaPregnant Women2.1 ± 4.1Wood et al. (2014)9.MartiniqueDog0Boni et al. (1998)10.MexicoDairy cattle28Salinas-Meléndez et al. (2002)Beef cattle10Sheep40Goat3511.TrinidadAbattoir workers4.7Adesiyun et al. (2011)Livestock workers4.6Office workers3.412.USAHuman3.1Anderson et al. (2013)Anderson et al. (2009)SOUTH AMERICA1.ArgentinaHuman0Cicuttin et al. (2015)2.BrazilSheep66.6Mares-Guia et al. (2014)Goat50Dog14.28Cat0Horse03.ChileHuman0.089Weitzel et al. (2016)4.ColombiaHumanone case reportedMattar et al. (2014)5.EcuadorDairy and mixed cattle16.2Carbonero et al. (2015)6.French GuianaHuman24.4Eldin et al. (2014)7.PeruHuman5Blair et al. (2004)ANTARTICAQ fever not reportedAnonymous (2013)Clinical signs and symptoms 2.9.1. Human:In humans, the clinical nature of C. burnetii infection is highly variable. It can lead to an acute infection characterized by mild febrile illness, pneumonia and hepatitis, while in rare cases chronic disease may develop in the form of endocarditis, and abortion and stillbirth in pregnant women. The fever due to C. burnetii infection is remittent and usually persists for 9-14 days. It is considered as self-limiting disease (Polo et al., 2015; Chakrabartty et al., 2016). Approximately 60% of the infected people remain asymptomatic while about 40% of the patients show clinical signs. As far as chronic Q fever is concerned, it develops in 3-5% of patients mostly in the form of endocarditis. Patients with valvular disorders, microbial arteritis, vascular implants and immune-compromised persons are more prone to this infection (Isken et al., 2013). The most common manifestations of Q fever are mild flu-like symptoms with sudden increase in body temperature, restlessness, excessive sweating, severe headache, myalgia, arthritis, anorexia, upper respiratory tract problems, persistent cough, pleuritic chest pain, chills, confusion, and GIT problems like nausea and diarrhea. Another important symptom of the disease is Q fever fatigue syndrome (QFS), which is a debilitating condition following an acute Q fever involving main body systems. It occurs in approximately 20% of patients. Although this persistent fatigue is not a life threatening condition it results in serious social and economical consequences in the form of loss of person’s quality of life and inability to work. QFS was thought to be the major cause of Q fever associated economical losses during the Dutch Q fever outbreak (2007-10). In rare cases, an acute Q fever develops into chronic infection. Endocarditis is the main clinical manifestation of chronic form. Other complications such as pericarditis, myocarditis, thyroiditis, osteomyelitis, nephritis, meningoencephalitis, hemolytic anemia, hemophagocytic syndrome, severe cephalgia and retro-orbital pain are rare manifestations of chronic infection (Isken et al., 2013; van Asseldonk et al., 2013; Keijmel et al., 2015; Y?lmaza et al., 2015). During pregnancy, infection is usually asymptomatic; however, some serious obstetrical disorders like placentitis, spontaneous abortion, fetal growth retardation, stillbirth, premature delivery and birth of weak offspring have been reported. Infection during pregnancy may also result in abortions in the subsequent pregnancies. However, a recent study conducted in an area with highest Q fever outbreaks in Netherland found no association between Q fever and adverse pregnancy outcomes (Van der Hoek et al., 2011; Vellema and van den Brom, 2014). C. burnetii infection results in high morbidity and low mortality. Mortality has been reported in 1-11% of the chronic Q fever patients (Hadush et al., 2016). This infection causes serious long-term implications on patient’s health and social life due to long-lasting Q fever fatigue syndrome (Van Asseldonk et al., 2015). The economical losses caused by Netherland Q fever outbreak (2007-2010) were estimated to be approximately € 0.307 billion (Van Asseldonk et al., 2013). 2.9.2. Animals:In animals, Coxiellosis usually occur without any apparent clinical signs and is not considered a veterinary health problem, except in ruminants, where C. burnetii is a well-known cause of abortion. Ruminants, especially sheep and goats are major reservoir of C. burnetii. The historical Q fever outbreak in the Netherland was linked with infected small goat farms present close to residential areas (Duron et al., 2015; Khaled et al., 2016). In animals, especially small ruminants, Coxiellosis usually results in reproductive problems like spontaneous abortion during late pregnancy, premature delivery, stillbirth and birth of weak offspring, as well as infertility in cattle (Freick et al., 2016; Hadush et al., 2016). In cattle, acute Q fever usually appears as subclinical infection, while chronic infection may result in reproductive disorders. There is no reliable evidence which states that this infection causes retention of fetal membranes, subfertility, metritis and endometritis in cows (Freick et al., 2016). Abortion due to C. burnetii infection usually ranges from 3-8% (Van Asseldonk et al., 2013). During the recent Dutch Q fever outbreak, up to 60% abortion rate was recorded in goats during final month of pregnancy with no apparent signs of illness. Endometritis was reported in some goats with previous history of abortion. Full-term kids were emaciated with lower body weight and high mortality rate. Several other apparently healthy kids showed respiratory and digestive tract problems (Ganter, 2015). In dairy animals, C. burnetii infection may lead to subclinical mastitis. The organism resides in mammary glands and pacenta of pregnant dairy animals with fetus and associated structures having the highest amount of C. burnetii. Thus, post parturient shedding of organism in birth fluids is higher in small ruminant, while lower in case of cattle (Shapiro et al., 2015; Freick et al., 2016). DiagnosisDiagnosis of Q fever on the basis of clinical signs and symptoms or post-mortem examination is almost impossible because of non-specific clinical presentation or missing symptoms and lesions of the disease (Niemczuk et al., 2014). So, for accurate diagnosis of C. burnetii infection, laboratory evidence is obligatory. Generally, four categories of diagnostic techniques are available for Q fever diagnosis: i) isolation and propagation of the organism, which requires BSL-3 laboratory using tissue-culture, embryonated chicken eggs or laboratory animals, ii) sero-diagnostic tests including IFA, CFT and enzyme immunoassay, iii) antigen detection assay such as immunohistochemical staining (IHC) and iv) genomic detection assays like PCR. Use of various laboratory tests in combination i.e., ELISA for serology and PCR for nucleic acid detection is highly suggestive for confirmatory Q fever diagnosis (Niemczuk et al., 2014; Bontje et al., 2016). In ruminants, both IFA and ELISA are suitable techniques for serological investigation of Coxiellosis, while in humans, IFA is considered as an ideal technique for Q fever diagnosis because of its high sensitivity and specificity (Meekelenkamp et al., 2012; Muleme et al., 2016).2.10.1. Serological TestsSero-diagnostic techniques are preferably used for diagnosis of Q fever because isolation techniques are very expensive, time consuming, laborious and less sensitive compared to serological techniques. They are best utilized to estimate herd or flock level prevalence of the disease. Serological diagnosis is based on measuring IgM and IgG antibodies concentration in serum against two distinct antigenic phases of C. burnetii. The difference in level of antibodies titer during acute and chronic form of infection is recognizable, which makes serology a tool of choice for diagnosis of Q fever (Saglam and Sahin, 2016; Lucchese et al., 2015). During acute infection, a higher IgG antibodies concentration is present against Phase II antigen only, while during chronic form of infection, both IgG and IgA antibodies are present against both phase I and phase II bacteria. The occurrence of IgG phase I titer more than 1:800 along with IgA phase I titer of more than 1:25 is an indication of chronic Q fever. However, some other studies reported different serological responses during chronic form of the disease. Overlaps of antibody titers in both acute and chronic forms of Q fever are also seen in some studies. This renders the use of antibody titers as a tool for distinguishing the stage of infection as controversial (Shah et al., 2015; Leuken et al., 2016). Other limitations of using serology for diagnosis of Q fever include failure to identify C. burnetii shedding animals, failure to detect antibodies in early stage due to lag in antibody titer development of 7-15 days after the appearance of clinical signs, and persistence of IgM phase II antibodies during endemic and post-epidemic situations of the acute Q fever (Niemczuk et al., 2014; Wielders et al., 2015). A. Immunofluorescence Assay (IFA):?Immunofluorescence Assay is a reference method for detection of antibodies against C. burnetii infection in humans. It is very useful, especially for follow-up of patient’s disease status and to identify patients having risk of developing chronic infection. This assay can easily differentiate between suspected acute or chronic form of Q fever by measuring phase I and phase II antibodies titers in serum. If phase I antibody titer is ≥phase II, it will indicate chronic form of Q fever and if phase II antibody titer is >phase I, the sample is indicative of an acute infection. IgG antibody titer of ≥1:800 against phase I antigen is indicative of Q fever endocarditis. This technique is commonly used for diagnosis of Q fever in humans, while in animals, no commercial Q fever IFA kit is available yet (Herremans et al., 2013; Selim and Elhaig, 2016; Ferraz et al., 2016). Although IFA is considered as gold standard for diagnosis of Q fever in humans but still it has limitations. This technique is not suitable for detection of early acute Q fever because of lag in antibody titer development (7-15 days after the onset of clinical disease). It requires very specific and expensive instruments along with high level of expertise for proper interpretation of results. The species-specific IFA cannot be used for extensive or herd level screening. That is why IFA, which is not used for routine detection of C. burnetii infection in animals (Selim and Elhaig, 2016; Pan et al., 2013). B. Complement Fixation Test (CFT):?Complement Fixation Test is regarded as a reference assay for Q fever diagnosis on OIE reference serological assay list but nowadays, its use is infrequent because of its lower sensitivity. Another drawback of CFT is the presence of anti-complementary activity in a number of samples which hinders with antibody titer estimation even after repeated attempts. Similarly, anti-C. burnetii present in serum samples of sheep and goat cannot be easily detected by IFA antigen (Shapiro et al., 2015; Selim and Elhaig, 2016). C. Enzyme Linked Immunosorbent Assay (ELISA) Enzyme Linked Immunosorbent Assay is more specific and sensitive than any other serological test and is indorsed by European Food Safety Authority (EFSA) for test harmonization. Particularly, in case of animals, it is more preferred than IFA and CFT because of its convenience for herd or flock level screening and its ability to detect C. burnetii antibodies in various animal species. The IDEXX reported 100% sensitivity and specificity of their ELISA kit (Selim and Elhaig, 2016; Mertens et al., 2017). ELISA kit having antigen from ruminant isolates is more sensitive than kit with antigen from ticks. EFSA recommends ELISA containing C. burnetii antigen from ruminants isolates. ELISA can detect antibodies in serum against both antigenic phases of C. burnetii and provides a cumulative interpretation of results as seropositive, suspect or seronegative status (Ohlson et al., 2014; Selim and Elhaig, 2016). StainingIn this technique, stained tissue or vaginal mucus smears are observed under microscope with an oil immersion objective lens for detection of causative agent. C. burnetii is an acid resistant bacterium. Different kinds of stains like Stamp, Gimenez, Macchiavello, Giemsa, modified Ziehl-Neelsen and modified koster can be used for its visualization under microscope. The first three stains give the best results. However, due to lack of specificity, a positive result is only presumptive indication of C. burnetii infection. Therefore, further investigations should be carried out for confirmatory diagnosis (OIE, 2015). Polymerase Chain ReactionPolymerase Chain Reaction is used for molecular detection of C. burnetii infection. It is a rapid, highly specific and sensitive diagnostic technique for detection of Q fever as compared to all other laboratory methods. It has ability to detect minute quantities of bacterial DNA in any sample. This property of PCR to identify and quantify smaller concentration of bacterial DNA has significantly improved diagnostic and research approaches (OIE, 2015; Selim and Elhaig, 2016). PCR can be performed on variety of biological specimens such as fetal membranes, fetal fluids, genital swabs or samples from aborted fetuses (liver, lung or abomasal contents). Blood, serum, milk, urine, anal and throat swab samples are also useful for detection of C. burnetii using qPCR. In case of chronic infection, samples can be obtained from focal tissue of infected organs like valvular material for endocarditis or aneurism, vessel fragments in case of vasculitis and bone biopsy in artheritis. As the antigen shed intermittently in urine, feces, vaginal discharge and milk, so it is preferable to investigate consecutive samples for genomic detection of pathogen (Niemczuk et al., 2014; Mori et al., 2017). PCR targeting the insertion sequence IS1111, a repetitive transposon-like element, of C. burnetii is considered to be highly sensitive and specific for genomic detection. However, IS1111 cannot be used for quantification of C. burnetii DNA because of having multicopies (20 copies per genome), which may result in misidentification with Coxiella-like organisms. Single copy genes like icd and com1 are useful for quantification of C. burnetii DNA. Different pairs of primers targeting various target genes like superoxide dismutase (sodB), isocitrate dehydrogenase (icd), com1, macrophage infectivity potentiator protein (cbmip), heat shock proteins including htpA, and htpB and some plasmid mediated genes like QpRs, QpH1, cbbE can be used for the detection of C. burnetii DNA (Khalili et al., 2015; Selim and Elhaig, 2016). The best time for PCR assay to detect C. burnetii DNA in blood or serum sample is the first two weeks after onset of clinical infection. During this period, there is lag in antibody titer development, so serological tests are useless during this period. However, the PCR can be used successfully to detect C. burnetii DNA in blood or serum samples during this interval. Two weeks after the onset of clinical signs, IgG antibodies titer starts developing and at the same time C. burnetii DNA becomes undetectable in the blood. Hence, serological techniques can be best utilized two weeks after onset of clinical infection (Niemczuk et al., 2014; Wielders et al., 2015). The quantitative real-time PCR can successfully be utilized to detect C. burnetii shedders. Once the sero-positive animals are detected at flock level with serological tests, then PCR is a tool of choice to trace the shedders (Niemczuk et al., 2014). Prevention and ControlDisease surveillance, and implementation of proper preventive and control strategies are necessary to reduce further disease outbreaks in an area. These strategies have economic and public health significance in reducing reproductive losses in livestock industry and potential risk of transmission of infection to human beings. Determination of herd level prevalence of a disease can help in proper planning, and implementation of preventive and control measures (Ganter, 2015; Van Asseldonk et al., 2015; Meadows et al., 2015). Due to the self-limiting nature of this disease, the prevalence of infection usually declines with passage of time without adopting any control strategies. This may be because of natural immunization of hosts against C. burnetii infection (Selim and Elhaig, 2016). Preventive vaccination, manure management including covering and compositing of manure or treating manure with lime, better livestock farm and wool shearing practices, use of isolated calving pens, restrictions on free animal movement, and proper disposal and burial of aborted materials are important measures to prevent C. burnetii infection. Hygienic practices especially calving pen cleanliness, is considered an important measure in preventing this infection. Similarly, disinfection of calving pens, naval cord disinfection, and proper disposal of aborted fetus and fetal membranes, and provision of new bedding during calving are important measures to reduce the risk of disease transmission. Birth products including fetal membranes and dead fetuses should immediately be disposed off to avoid their ingestion by domesticated animals, stray dogs and wild carnivores, which may spread the infection in the environment (Shapiro et al., 2015; OIE, 2015; Meadows et al., 2015; Van den Brom et al., 2015). Quarantine measures should be implemented at livestock farms and animals from infected flocks should not be mixed up with healthy animals at farm. The raw milk from infected dairies should not be used for drinking or any other purpose, because large number of bacteria are shed in the milk of infected animals. The seropositive animals shedding the organism into the environment are important source of disease transmission. It is important to identify such shedders and cull them from the healthy flocks (Lucchese et al., 2015; Hadush et al., 2016). Training and awareness of livestock associated professionals and farmers are important in reducing the risk of disease spread. Individuals working on disease surveillance should adopt personal protective measures like protective clothing (including FF-3 breathing mask), protective gloves and disinfection of sampling materials immediately after use. Similarly, all consumables should be properly discarded after use (Ganter, 2015). The distance between residential areas and livestock farms, especially those containing pregnant ewes, should not be less than 500 meters to avoid the risk of disease transmission. Producers should avoid transporting and marketing the animals, especially periparturient animals, during ongoing abortion outbreaks to prevent farm to farm spread of infection (Boden et al., 2014; DePuy et al., 2014). As Q fever is a cosmopolitan zoonosis, an interdisciplinary cooperation among medical doctors, veterinarians, lab working groups and farmers is required to understand how this pathogen circulates in a geographical area, and plan strategies for its proper control and prevention (Bellini et al., 2014). 2.11.1. VaccinationVaccination is an active immunological response against potential pathogen present within the body. The use of anthelminthic drugs prior to vaccination is useful to gain an improved immunological response (Lacasta et al., 2015). Currently available inactivated phase I vaccine for animals, containing Nile Mile RSA 493 strain of C. burnetii isolaed from ticks, is recommended by OIE in Q fever endemic areas. This vaccine is reported to casue reduction in abortion rates, decrease in bacterial shedding and lowers the risk of disease transmission to humans. However, it is less effective in outbreak situations compared to regular vaccination (Selim and Elhaig, 2016). Use of inactivated phase I vaccine (Coxevac) in non-infected sheep and goats prior to their first breeding results in reduced abortion rate and bacterial shedding. Some studies reported that use of Coxevac during pregnancy also reduces bacterial shedding, although this vaccine is not approved for use in pregnant animals. Inactivated phase I vaccine was proved to be very effective in small ruminant flocks of Netherland after an extensive vaccination campaign was carried out in 2010. Since then, no C. burnetii borne abortion has been reported from vaccinated flocks and a gradual decline occurred in the number of PCR positive farms based on bulk tank milk (BTM) sampling (Van den Brom et al., 2015). It is recommended to perform repeated annual vaccination in susceptible herds, especially young animals, in at-risk areas (OIE, 2015). Phase I vaccine is equally effective in humans as in case of animals but it is contraindicated in individuals already exposed to C. burnetii infection. Q-vax is the only available Q fever vaccine in Australia for human use. Q-vax is an inactivated phase I whole cell vaccine containing Henzerling RSA 331 strain of C. burnetii isolated from blood of Q fever patient in Italy. In Australia, Q-vax vaccination is done on routine basis in individuals occupationally exposed to Q fever infection (Schoffelen et al., 2013; OIE, 2015). Treatment2.12.1. Treatment in humans: There are two forms of Q fever in humans i.e., acute and chronic. Antibiotic therapy is effective against acute form of the disease, but once the infection proceeds to its chronic form then treatment time is prolonged and recurring of disease is usual, which may lead to high mortality. Duration of antibiotic therapy is established based upon follow-up of serological titers in Q fever patient (Pan et al., 2013; Godinhoa et al., 2015). The antibiotic treatment must be started immediately after the onset of clinical disease, because delayed antibiotic treatment may not be effective (Alvesa et al., 2017). Generally, acute Q fever is self-limiting, however, timely detection and antibiotic administration may decrease the duration of infection and severity of symptoms. The drugs of choice for Q fever are doxycycline and hydroxyl chloroquine. These drugs are mostly used in combination. Other antibiotics, such as erythromycin, rifampin, roxithromycin and clarithromycin, can be used as alternative therapy (Schoffelen et al., 2015; Godinhoa et al., 2015; Hadush et al., 2016). A dose of 100 mg doxycycline two times a day for 2 to 3 weeks is recommended for acute Q fever patients, especially non-pregnant women and adult patients. Hydroxychloroquine can also be used in combination with doxycycline. Hydroxychloroquine is a lysosomotrophic drug which increases the pH of phagolysosome. As hydroxychloroquine elevates the phagolysomal pH, it acts as bacteriostatic because C. burnetii require acidic environment for its multiplication (Hadush et al., 2016). In case of pregnant women and kids < 8 years of age, cotrimoxazole can be used safely for treatment of Q fever (Shah et al., 2015). In case of chronic Q fever, especially native and prosthetic valve endocarditis, similar antibiotics (doxycycline and hydroxychloroquine) can be used at dose rate of 200 mg per day but for a longer period of 18 to 24 months. Combination therapy consisting of doxycycline and hydroxychloroquine is more effective in preventing the development of endocarditis than doxycycline alone. Rifampicin, macrolides and Quinolones are less effective against C. burnetii infection, therefore, they are not usually used as alternative treatment for this disease (Shah et al., 2015; Baziaka et al., 2014). Methotrexate is an important steroid replacement therapy used to suppress vascular inflammation and maintain homeostasis of ascending and thoracic aorta (Baziaka et al., 2014). Follow-up care, such as regular heart beat and eye reflexes examination, is necessary after antibiotic treatment. Photosensitivity may be a problem in some patients after antibiotic use. In advanced stages of chronic Q fever, like severe cardiac failure or abscess formation in heart valve, antibiotics are useless. In these situations cardiac surgery is recommended (Shah et al., 2015; Ferraz et al., 2016). The use of interferon (IFN) and tumor necrosis factor (TNF) for Q fever treatment has also been proven effective (Shah et al., 2015; Hadush et al., 2016). In case of chronic infection, the follow-up of serological response is necessary and the treatment can be stopped when phase I IgG antibodies titer declines by at least four folds. Special attention should be given to individuals more prone to Q fever infection because the infection can lead to high morbidity and mortality if left untreated (Godinhoa et al., 2015; Chieng et al., 2016). 2.12.2. Treatment in animals:Very limited information is available about treatment of Coxiellosis in animals. Extensive data is required to determine the efficacy of antibiotics in preventing the bacterial shedding and reproductive losses in animals due to C. burnetii infection. Usually tetracycline is recommended for treatment of Q fever in animals, but usage of tetracycline in feed of animal during gestation period as a herd-level disease control strategy is not effective because of its less bioavailability during oral administration. Parenteral use of 2 injections of oxytetracycline (long-acting) 20 days apart at 20 mg/kg during ongoing Q fever abortions may be useful in preventing reproductive losses to the animals. However, oral administration of oxytetracycline is not effective in reducing bacterial shedding with birth fluids and in altering the serological status of animal (Anderson et al., 2013). In ruminants, tetracycline administration in pregnant animals from 95th day of gestation till parturition, with an interval of 2-3 weeks, is effective in reducing the risk of abortion due to other pathogens like Chlamydophila abortus (Ganter, 2015). Q fever in PakistanIn Pakistan, Q fever is a highly neglected zoonotic disease. During the period from 1955 to 2016, there are only five studies which reported the prevalence of Q fever in Pakistan. Based upon these studies, prevalence of Q fever in Pakistan ranged from 4.6 to 40% in all livestock species and 10.19-26.8 in humans (Kaplan and Bertagna, 1955; Ahmed, 1987; Ayaz et al., 1993; Zahid et al., 2016; Shabbir et al., 2016). In Pakistan, the disease was first investigated in camels by WHO (1955) which revealed that two out of five camels were seropositive for Q fever through CFT. Another sero-diagnostic study was conducted by Ahmad (1987) in humans, ruminants and commensal rodents of Pakistan. The overall prevalence was found to be 26.8, 34.5, 10.4, 18.3, 4.6 and 18.0% in humans, buffaloes, cattle, caprine, ovine and rodents, respectively through CFT. This investigation showed that Q fever is endemic in humans, animals and rodents of the studied locations of Pakistan.Then another serological survey was carried out by Ayaz et al. (1993) in northern areas of Pakistan. Serum samples collected from humans and sheep were analyzed through microtiter CFT for determination of antibodies against both antigenic phases of C. burnetii. Highest sero-positivity was found in patients with pyrexia of unknown origin (14.5%), followed by veterinary doctors (12.5%) and farmers (12.35%). In Margla region, a high prevalence of phase I antibodies was found in human sera, which revealed that the disease runs a chronic course there (Ayaz et al., 1993).Another cross-sectional study was performed by Shabbir et al. (2016) in Punjab province of Pakistan with the purpose (1) to investigate the prevalence of C. burnetii in soil samples through real-time PCR (RT-PCR) targeting ISIIII gene, (2) to find out association between seropositivity against C. burnetii in soil samples with nutrients (macro and micro nutrients) present in these samples and also with seroconverted small ruminants in areas where its genetic characterization had or had not been elucidated and (3) to determine genetic similarity and variation of the investigated C. burnetii strains with already available gene sequences from other hosts present worldwide. Results obtained from this study revealed that 47 samples (1.94%) obtained from 35 villages included in this study contained C. burnetii DNA. Highest amount of bacterial DNA was found in soil samples collected from Attock (7.11%), followed by Lahore (4.83%), Sahiwal (4.70%), Dera Ghazi Khan (2.33%), Faisalabad (1.35%) and Sheikhupura (0.68%). It was found that odds of detecting C. burnetii DNA in soil samples increased with an increase in sodium ions and organic matter, whereas the odd of detecting bacterial DNA decreased by approximately 1% with an increase in potassium and calcium ions. Higher seroprevalence was recorded in sheep as compared to goats. A significant association was found between detection of C. burnetii DNA in soil samples and Coxiellosis in sheep. Sequence analysis targeting IS1111 element identified a clustering of DNA into different groups with maximum genetic variation. The first group containing sequences from Lahore district clustered with isolates from buffalo and human origin, whereas, the other group containing sequences from other districts of Punjab clustered with isolates from caprine, human and rodent origin.A recent seroprevalence study was carried out by Zahid et al. (2016) in two major livestock farming districts of Punjab, Pakistan. An individual prevalence of 30.8% was recorded in both species. Higher prevalence was found in females (32%) as compared to male animals (21.7%). Similarly, prevalence was higher in nulliparous animals (34.8%) followed by primiparous animals (24.8%) and multiparous animals (32.3%). Univariate analysis showed an association between seropositivity against C. burnetii antibodies with caprine species (p=0.22), female sex (p = 0.10), ticks infestation (p=0.001), previous history of abortion (p=0.143), retaintion of placenta (p=0.35), raising single breed of small ruminanat (p=0.56) and mixed type of feeding practices (p=0.331). ConclusionBased on the literature presented in this chapter, it looks that Q fever is a worldwide zoonosis, except New Zealand and French Polynesia. The disease is highly neglected in many developing and under-developed countries of the world, including Pakistan. C. burnetii is highly resistant to environmental factors and many disinfectants resulting in long-lasting infection risk both for human and animals. As the infection is usually asymptomatic, Q fever mostly remains undiagnosed in animals until and unless adverse pregnancy outcomes occur in a herd. Similarly, in case of human beings, the infection can lead to severe endocarditis and vascular infection if it remains untreated for longer period of time. Limited data is available on molecular epidemiology and evolution of this pathogen, especially in ruminants. Similarly, pathogenesis of Q fever need to be explored through in-depth molecular studies.Chapter-3MATERIALS AND METHODSDescription of study areaThis study was carried out in small ruminants kept at various livestock farms of Punjab province of Pakistan. Punjab is the largest province of Pakistan with highest human and animal population. It is also the second largest province of the country with an area of 205,344 sq. kilometers. Geographically, it is located at 31.17040 N and 72.70970 E in the semiarid lowlands region. Temperature of Punjab ranges from ?2° to 45?°C, but can increase up to 50?°C (122?°F) in summer and can decline up to ?10?°C in winter. Mean annual rainfall in Punjab varies with highest rainfall in northern regions as compared to middle and southern region. The province is bordered North-East side by Kashmir, the Indian provinces of Rajasthan and Punjab towards Eastern side, Sindh province to South side, Baluchistan to South-West, Khyber Pakhtunkhwa (KPK) to the West and Islamabad to its North (Fig. 3.1.1). It has a fertile agriculture zone built on an extensive irrigation network (Khattak and Ali, 2015; Shabbir et al., 2016). Agriculture and livestock are major sources of socio-economic development especially in rural regions of Pakistan. As per livestock census of Pakistan, Punjab dominates the livestock population of the country with 49, 65, 24 and 37% of cattle, buffaloes, sheep and goats, respectively (Pakistan Livestock Census, 2006). We purposively selected these 9 livestock farms located at 7 districts of Punjab for sero-epidemiological survey on Coxiellosis, because no previous studies on C. burnetii infection have been conducted so far at any of these livestock farms and districts which might help us to select a particular geographical area/region to be studied. We selected these livestock farms as these reflect the major livestock production sites of Punjab and there prevails a higher annual incidence of animal and human disease (Directorates of Animal and Human Health, Punjab).4380845028960Indian Ocean00Indian OceanFig 3.1: Map of Pakistan. Green area on the map showing Punjab province of Pakistan (Source:)Estimation of sample size and sample collection (Internet Source)For this study, samples were collected from nine government livestock farms including LES Alladad Jahnia, LES Khushab, LES Fazilpur, LES Jogaitpur, LES Rak Kharewala, LES Bahadarnagar, LES Rakh Ghulaman, GLF Kallurkot and Fine Wool sheep farm-205 TDA. These livestock farms were located at Khanewal, Khushab, Rajanpur, Bhawalpur, Layyah, Okara and Bhakkar districts of Punjab province (Fig. 3.1.2). Blood samples from goats could be collected from only four farms because goats were not kept at other five farms included in this survey, while sheep blood samples were collected from all nine farms. Similarly, goat blood samples could be collected from only four districts, while sheep blood samples were collected from all seven districts included in this survey. Survey Toolbox software was used for the random selection of farms and animals (Cameron, 1999). The survey was based on an unknown prevalence of Q fever in small ruminants in the study areas (Thrusfield, 2007). For the calculation of sample size, the following formula for a 95% confidence interval was applied: n=1.962Pexp (1-Pexp )d2With expected prevalence of 50% and desired absolute precision 5%, the total samples come to be 384, as given below:n=1.962 0.50 (1-0.50)0.052 n=384 However, to be on safe side, 1000 blood samples were collected (500 samples from sheep and goat each). During sampling, individual animal information including species, farm, location (districts), age, sex, breed, parity, lactational status, reproductive status (pregnant or non-pregnant), presence of ticks, body condition, and reproductive disorders and general farm management data were recorded using a structured questionnaire. The parameters/ risk factors included in this study were categorized into different groups. Age-wise the animals were categorized into four groups i.e., up to 1 year, > 1 to 2.5 years, > 2.5 to 4 years and > 4 years. Similarly, sex-wise small ruminants were grouped as male and female. As far as breed was concerned, sera sample were collected from 10 breeds of small ruminants including five breeds of sheep and five breeds of goats. Parity-wise the female animals were grouped as nulliparous, primiparous and multiparous. Similarly, the female animals were classified into lactating/ non-lactating and pregnant/ non-pregnant based on their lactational and reproductive status. Regarding ticks infestation, the animals were classified into two groups i.e., animals with tick infestation (Yes) and animals without tick infestation (No). Based on body score condition, animals were grouped into following two categories i.e., weak animals, animals with weak kid/ lamb delivery. Regarding reproductive disorders, sera samples were collected from animals with previous history of abortion, stillbirth, premature delivery and repeat breeding. Approximately 10 ml of blood was collected from the jugular vein of each animal using disposable needles and evacuated blood collection tubes (Improvacuter, Shanghai International Hamburg Holding, gmbH, Germany). Each tube was properly labeled for its identification and information about its origin, using English alphabets and Arabic numbers. Blood collection tubes were placed vertically in a cool box packed with gel freezing pads, and shipped to the laboratory for further processing. At laboratory, centrifugation was done at 4500 rpm for 10 min. for proper separation of serum from whole blood. Serum was then transferred to disposable screw caped cryovials (Cryo.STM; Greiner Bio-one, GmbH Frickenhausen, Germany) and stored in a deep freezer at -20 ?C for further serological analysis. Fig 3.2: Red dots on the map showing districts of Punjab province from which samples were collected (Modified from source: +punjab&source=lnms&tbm=isch&sa=X&ved=0ahUKEwiogO6HkM7cAhXKfFAKHTULCKMQ_AUICigB&biw=1242&bih=577#imgrc=_uB5leUr7BePSM:)Collection and handling of ticks From the period from January 2016 to June 2016, a total of 163 ticks were collected from small ruminants (sheep and goats) maintained at 9 government livestock farms of Punjab, Pakistan. The ticks were placed in properly-labeled bottles containing 70% ethyl alcohol as a preservative at University of Agriculture Faisalabad. Out of 163 ticks, 85 ticks were collected from sheep while 78 were collected from goats. Later, all these ticks were shipped to FLI, Jena, Germany as per international rules and regulations for transportation of ticks. At FLI, all the 163 ticks were merged into 55 pools including 29 pools for sheep ticks while 26 pools for goat ticks. Each pool was composed of 3 ticks (with only pool 29 containing 1 tick). These pools were then stored in a deep freezer till its use for genomic detection.Study Locales and proportional allocation of samplesThe numbers of samples collected from each farm were computed through Proportional Allocation method, as shown in (Table 3.1)Table 3.1: Studied farms and no. of samples collected from each farmSr. No.Name of FarmTotal Sheep%age of Sheep Pop.Minimum Samples RequiredTotal Goat%age of Goat Pop.Minimum SamplesRequired1LES, AlladadJahnia3830.07940.02810.41203.02LES, Khushab4300.08945.000.000.03LES, Fazilpur900.0199.0600.0943.04LES, Jogaitpur3560.07437.000.000.05LES, RakGhulaman14000.290145.000.000.06LES, RakKharewala7100.14773.02900.42210.07LES, Bahadarnagar8480.17688.0610.0944.08GLF, Kallurkot2140.04422.000.000.09Fine Wool sheep farm, Bhakkar4000.08341.000.000.0Total 48311.000500.06921.00500.0Diagnostic testsTwo types of diagnostic tests were conducted in this study. Firstly, all the serum samples were analyzed through Indirect ELISA (IDEXX Q-Fever (Coxeilla burnetii) Antibody Test Kit). Then pools of some seropositive and suspected serum samples (29 pools) were analyzed through an in-house developed real-time qPCR (FLI standard Jena, LA 190 qPCR, MXPro-MX3000P) using single copy icd gene. Each serum pool was comprised of 6 different serum samples. DNA from serum was extracted using High pure PCR template preparation kit (Roche kit, version 20, Roche Diagnostics GmbH, Germany). All the ELISA-positive samples were not analyzed through qPCR because previous studies have revealed that C. burnetii DNA cannot be detected in serum two weeks after the onset of clinical infection (Schneeberger et al., 2010; Wielders et al., 2013; Niemczuk et al., 2014). Still some pools of ELISA-positive serum samples were investigated through qPCR; but C. burnetii DNA was not detected in any of these seropositive pools. Additionally, 55 ticks pool were also investigated through real-time qPCR (FLI standard Jena, IBIZ, AGr. 180, DNA-Isolation high pure kit, version 2) using multiple copy IS1111 transposase gene. DNA from ticks was extracted through an in-house developed high pure PCR template kit (FLI standard Jena, IBIZ, AGr. 180, DNA-Isolation high pure kit, version 2) following the manufacturer’s protocol for purification of genomic DNA from insects. Prior to DNA extraction, ticks were surface decontaminated by serial passages in 10% and/or 70% ethyl alcohol and then rinsed in sterile water.Q-Fever (Coxiella burnetii) Antibody Test KitName and Intended UseIDEXX Q Fever test provides a sensitive, simple, rapid and specific technique for detecting antibodies against C. burnetii infection in samples of plasma, serum and milk.Description and PrinciplesMicrotiter plates precoated with inactivated antigen are supplied with kit. Sample dilutions to be analyzed were incubated in the ELISA 96-wells plate. Sample dilutions containing antibodies against C. burnetii were bound to the antigen coated on the wells of ELISA plate and an antigen/antibody complex was formed on surface of plate wells. Washing was done to remove unbound material from the wells. After washing, a peroxidase labeled anti-ruminant IgG conjugate was added into the wells, which bound to the antigen-antibodies complexes. To remove unbound conjugate washing was done and then TMB substrate was added to the wells. The intensity of the color that developed was linked with the amount of antibodies specific for C. burnetii found in the sample. The results were recorded by comparing the optical density (OD) value with that of positive control. The following reagents were used for Q fever-Indirect ELISA (Table 3.2).Table 3.2: Reagents and their volumes present in Q Fever- Indirect ELISA kitSr. No.ReagentsVolume1Coxiella burnetii Antigen Coated Plate22Positive Control1x 0.4 ml3Negative Control1x 0.4 ml4Conjugate1x 24 mlATMB Substrate N. 121x 20 mlBStop Solution N. 31x 20 mlCWash Concentrate (10x)2x 100 mlOther Components: Zip lock bag1Storage The reagents are stored at 2-8 oC. Reagents are stable until expiration date, provided they have been stored properly. Materials RequiredPrecision micropipettes or multi-dispensing micropipettesDisposable pipette tipsGraduated cylinder for wash solution 96-well microplate reader (equipped with 450 nm filter)Microplate washer ( manual, semi-automatic or automatic system)Distilled or deionized water for preparation of the reagents Microplate cover (plastic lid, aluminum foil or adhesive) Vortex or equivalent Microplate shakerHumid chamber/ Incubator capable of maintaining a temperature of +37 oC (±3 oC)Preparation of Reagents Wash Solution The wash concentrate (10X) was brought to 18-26 oC temperature and mixed properly to dissolve any precipitated salts. Then 1/10 dilution of wash Concentrate (10X) was done with deionized/ distilled water before use (e.g., Wash Concentrate 30 ml + 270 ml of distilled water/ plate to be analyzed). When prepared under sterile conditions, the Wash Solution was stored for one week at 2-8 oC.Preparation of samples Serum/ plasma samples and controlsSerum/ plasma samples and Positive, and Negative Controls were prediluted 1/400 in a tube using the Wash Solution. Test Procedure All reagents were brought to 18-26 oC before starting analysis. The reagents were mixed by gentle inverting or swirling.Coated plates were obtained and sample position was recorded. 100 ?l of DILUTED Negative Control (NC) was dispended into duplicate wells.100 ?l of DILUTED Positive Control (PC) was dispended into duplicate wells.100 ?l of DILUTED samples was dispended into appropriate wells.The content of the wells were mixed by gently tapping the plate/microplate shaker. The microplate was properly covered and incubated for a period 60 min. (±5 min.) at +37 oC (±3 oC). The plates were tightly sealed or incubated in a humid chamber using plate covers to avoid any evaporation. The solution was then removed and each well was washed with approximately 300 ?l of Wash Solution 3 times. The plate was avoided from drying between the interval of plate washing and before addition of the next reagent. Each plate was tapped onto absorbent material after the final wash to remove any residual wash fluid.100 ?l of the conjugate was dispended into each well.The microplate was covered and incubated for 60 min. (±5 min.) at 37 oC (±3 oC). The plates were tightly sealed or incubated in a humid chamber using plate covers to avoid any evaporation.Step 7 was repeated.100 ?l TMB Substrate N. 12 was dispended into each well.Incubation at 18-26 oC for 15 min. (± 1 min.) away from direct light. 100 ?l of Stop Solution N. 3 was dispended into each well.The results were recorded using a photometer at a wavelength of 450 nm.Following formulas were used for Calculations: Controls NCx?=NC1 A450+NC2 A(450)2PCx?=PC1 A450+PC2 A(450)2Validity criteriaNCx≤0.500PCx≤2.500 PCx- NCx ≥0.300SamplesS/P %=100 X Sample A450-NCx A(450) PCx-NCxBased upon the above calculations, the results were interpreted as shown in (Table 3.3).Table 3.3: Intrepretation of Q fever- Indiect ELISA resultsNegativeS/P % < 30 %Suspect 30 % ≤ S/P % < 40 %Positive S/P % ≥ 40 %DNA Extraction (High Pure PCR Template Preparation kit)For nucleic acid isolation for PCRAll solutions were warmed at a temperature of +15 to +25°C or at +37°C in a water bath until the precipitates were dissolved. The contents needed for extraction of DNA using this kit are shown in (Table 3.4).Table 3.4: Contents/ Reagents in High Pure PCR Template Preparation kit Vial/ CapLabelContents/ Function 1 Tissue Lysis Buffer (TLB) - 20 ml White - [Tris 200 mM, NaCl 20 mM, Urea 4M, pH 7.4 (+25°C), EDTA 200 mM] 2 Binding Buffer - 20 mlGreen - [urea 10 mM, guanidine-HCl 6 M, 10 mM Tris-HCl, pH 4.4 (+25°C) 20% Triton X-100 (v/v)]3 Proteinase k, recom- - act as LyophilizatePink binant PCR grade - For inactivation of endogenous DNase and sample lysis4a Inhibitor Removal Buffer - Total 33 ml+ 20 ml absolute ethanolBlack - [Tris-HCl 20 mM, guanidine-HCl 5 M, With pH 6.6 at (+25°C) final conc. after adding ethanol]4 Wash Buffer - 20 ml+ 80 ml absolute ethanolBlue - [Tris-HCl 2 mM, NaCl 20mM, pH 7.5, at (+25°C) final conc. after adding ethanol]5 Elution Buffer - 40 mlColorless - [Tris-HCl 10 mM, pH 8.5 at (+25°C)] High pure filter tubes 2 bags containing 50 polypropylene tubes with double layered glass fiber fleece, for, 700 μl sample volume usage. Collection tubes 8 bags containing 50 polypropylene tubes of 2ml volume Storage and StabilityThe Kit contents were stored at temperature +15 to +25°C. Contents were assured to be stable till the expiry date mentioned on the label. Inappropriate storage at temperature +2 to +8°C in refrigerator or at 15 to 25°C temperature at freezer has an bad effect on purification of nucleic acid For this reason, the kits are always transported at temperature of +15 to +25°C.Immediately after Proteinase K dissolution, the solution was aliquoted and stockpiled at temperature of 15 to 25°C.As per manufacturer recommendation, the solution remain stable for use up to 1 year at temperature of 15 to 25°C.Additional Equipment and Reagents RequiredThe following additional equipment and reagents were needed for entire isolation procedures:Absolute ethanolAbsolute isopropanolA standard microcentrifuge machine having 13,000 g centrifugal force (e.g., Eppendorf 5415C or similar one)1.5 ml, sterilized Microcentrifuge tubesFor mammalian blood, serum, buffy coat or cultured cells isolation:Phosphate Buffer Saline (PBS)*ApplicationAs per manufacturer recommendation, this kit have the ability to purify DNA from various types of materials like whole blood, cultured cells, and tissue samples. Microorganisms like yeast and bacteria needed a specific prelysis with lyticase or lysozyme. The DNA obtained are ready to be used in subsequent PCR and restriction digestion reactions. Table 3.5 shows the time required for analysis of whole blood and cell culture.Table 3.5: Time required for analysis of whole blood and cell culture Cell culture and whole blood Tissue Total time required Approximately 20 min. Approximately 2hHands-on time Approximately 12 min. Approximately 30 minSample Material 200 – 300 ?l serum or mammalian whole blood.Preparation of Working SolutionsAlthough ready-to-use solutions were provided with kit, some working solutions required some preparation (Table 3.6):Table 3.6: Working solutions preparation and their use, storage and stabilityContentsPreparation/ ReconstitutionStorage and StabilityApplication Proteinase K Proteinase k dissolved in Stored (+15 to +25°C). Sample lysis and DNA (Vial 3; pink 4.5 ml double distilled Stablility 12 months binding protocol step1Cap) water, aliquot sol. Inhibitor Put 20 ml absolute ethanol Stored (+15 to +25°C). Washing and Elution Removal Buf- to inhibitor removal buffer Stablilty: till the protocol step 1 fer(vial 4a, properly label the bottle expiry date Black cap) after adding erhanol mentioned on kit. Wash Buffer pour 80 ml absolute ethanol Stored (+15 to +25°C). Washing and Elution (vial 4a, to washing buffer Stablilty: till the protocol step.2 and 3 Blue cap) properly label bottle expiry date after adding erhanol mentioned on kit Adjustment of Sample VolumeWhen sample volume was less than 200 ?l, PPBS was used to increase the volume up to 200 ?l. Table 3.7 showing adjustment of sample volume.Table 3.7: Adjustment of sample volume for DNA extraction MaterialsActionMammalian whole blood when sample material was < 200 ?l, the volume was increasedor serum sample to 200 ?l using PBS. When sample was > 200 ?l or up to 300 ?l, all other sample Volumes were increased accordingly.Lysis of sample and DNA Binding Procedure for DNA isolation from 300 ?l mammalian blood or serum.Before beginning purification process, the Elution Buffer was warmed to +70°C.Step 1. To 1.5 ml microcentrifuge tube (nuclease-free):200 ?l of sample material was added.200 ?l Binding Buffer was added.40 ?l reconstituted Proteinase K was added. Immediately mixed and incubate at temperature of +70°C for 10 min.Step 2. 100 ?l isopropanol was added and mixed properly.Step 3. A Highly Pure Filter Tube was assembled in one Collection Tube.The sample material was pippted into the upper buffer reservoir of the Filter Tube.The whole Filter Tube assembly was poured into a standard table- top centrifuge.Centrifugation was carried out for 1 min. at 8,000g.Step 4. Proceeded to washing and elution, as described in the flow chartFig 3.3: High pure PCR template preparation kit workflow (Roche kit, version 20, Roche Diagnostics GmbH, Germany)DNA Isolation from TicksNote: Prior to DNA extraction, ticks were surface decontaminated by serial passages in 10% and 70% alcohol, and rinsed in sterile water before starting DNA extraction. As the ticks were preserved in alcohol for a longer period of time, they were triturated in vials containing 1ml PBS (Phosphate Buffer Saline) with 3-4 beads per tube, prior to the addition of TLB (Tissue Lysis Buffer). Then these vials were centrifuged at 1000g for 10 min. After centrifugation, PBS was removed and sediment was collected which was then triturated with 200 ?l TLB.Step-wise Procedure:1. Homogenization of ticksThe tick/ticks were placed in 2 ml Eppendorf tube 3-4 beads were added in each tube 200 ?l Tissue Lysis Buffer (TLB) was added2. TriturationThe tubes were placed evenly over both adapters of TrituratorTriturated for 10 min. at 30 Hz The ticks were properly homogenized (Repeated if necessary)Then centrifuged for 1min. at 1000g3. Lysis and Application40 ?l Proteinase K-Lsg was addedMixed properly (Vortexer)Then incubated at 55°C for ≥ 1hour When incubation was completed, then centrifugion was done for a short time200 ?l Binding Buffer was addedMixed immediately (Vortexer)Again incubated for 10 min. at 70°CWhen incubation was complete, then centrifugion was done for a short time100 ?l Isopropanol was addedThen centrifuged for 1min. at 1000gNew filter tube were taken and labelledFilter tubes were placed in the collection tubeThe volume was poured (from eppendorf tube) into the filter tubeThen centrifuged for 1min. at 800gThe collection tubes were discarded4. Washing stepA)The filter tubes were placed in a new collection tube500 ?l Inhibitor Removal Buffer was addedThen centrifuged for 1min. at 800 gThe collection tubes were discardedB)The filter tubes were placed in a new collection tube500 ?l Inhibitor Removal Buffer was addedThen centrifuged for 1min. at 800 gThe collection tubes were discardedC)The filter tubes were placed in a new collection tube500 ?l Inhibitor Removal Buffer was addedThen centrifuged for 1min. at 800 gThe collections tube were discardedAfter washing, the filter tubes were placed in a new collection tubesCentrifuged maximum up to 10 seconds (for removal of remaining wash buffer).5. ElutionThe filter tubes were placed in a new Eppendorf tubes.The Eppendorf tubes were lablelled.200 ?l Elution Buffer was added (preheated at 70°C)Centrifuged for 1min. at 800 gThe Eppendorf tubes containing DNA sample were stored and the filter tubes were discarded.Thermocycler reactionReal-time qPCR targeting single copy icd gene of C. burnetii was performed in duplicate as demonstrated (serum input volume was 300 ?l while the elution volume was 60 ?l; for reanalyzing, again the serum volume was 300 ?l while the elution volume was 25 ?l). Positive and negative controls were included in each qPCR run to validate the results. DNase- and RNase-free water was used as a negative control for every 10 samples tested. A positive control containing 105 C. burnetii/ ml was delivered with the kit and was used as a quantification standard. Similar procedure was followed for ticks DNA detection through real-time qPCR targeting multicopy IS1111 gene of C. burnetii. Results of qPCR were counted positive when at least one CT value was ≤ 35. Fig. 3.4 and 3.5 showing standard curves based on a reference line generated from decimal dilutions of the positive control for qPCR performed on serum and tick pools. To prevent contamination of qPCR technique with genomic C. burnetii DNA or amplicons from previous PCR reactions, the preparation of the PCR reagents, the isolation of DNA, and the amplification by qPCR (closed system) were performed in three isolated rooms specially designed for those processes. A water control (no-template control [NTC]) and a mock isolation (negative control [NC], i.e., simulated isolation that mimics the process of sample handling) were incorporated in each cycler run. The qPCR assay (Invitrogen?, Life Technologies) was carried out using primers and conditions as shown in (Table 3.8 and 3.9) and (Table 3.10 and 3.11).Table 3.8: Primers and probe sequences used for isocitrate dehydrogenase (icd) real-time qPCR Oligo Name Sequence ( 5′ to 3′ ) Primer icd-439F CGTTATTTTACGGGTGTGCCA icd-439R CAGAATTTTCGCGGAAAATCA Probe Coxburicd-464TM FAM-CATATTCACCTTTTCAGGCGTTTTGACCGT-TAMRA-TTable 3.9: Conditions for real-time qPCR performed on serum poolsCycler: FLI standard MX 3000 Temperature Duration cycles Decontamination 50°C 2 min. 1 Initial denaturation 95°C 10 min. 1 Denaturation 95°C 15 sec. 45 Annealing/Elongation 60°C 30 sec. 45 Table 3.10: Primers and probe sequences used for IS1111 real-time qPCROligo Name Sequence ( 5′ to 3′ ) Primer Cox- R CCCCGAATCTCATTGATCAGC Cox-F GTCTTAAGGTGGGCTGCGTG Probe Cox- TM 6FAM-AGCGAACCATTGGTATCGGACGTTXTATGG-PH Table 3.11: Conditions for real-time qPCR performed on tick poolsCycler: FLI standard MX 3000 Temperature °C Duration cycles Decontamination 50°C 2 min. 1 Initial denaturation 95°C 10 min. 1 Denaturation 95°C 15 sec. 50 Annealing/Elongation 95°C 30 sec. 50 Fig. 3.4: Standard curve based on a reference line generated from decimal dilutions of the positive control for qPCR performed on serum pools Fig. 3.5: Standard curve based on a reference line generated from decimal dilutions of the positive control for qPCR performed on ticks poolsSensitivity and Specificity of an in-house developed real-time qPCR1.??Sensitivity: Less than 10 genome equivalents per reaction were reproducibly detected.2.??Specificity: The following DNA samples from other bacterial species were used as negative controls for PCR: Legionella pneumophila (ATCC 33152, JR32 and 130b), Francisella tularensis, ssp. novicida (ATCC 15482) and ssp. tularensis (Schu4), Bacillus subtilis (DSM 347), Bacillus anthracis (UD III-7), Bacillus cereus (DSM 31), Bacillus thuringiensis (DSM 350), Bacillus megaterium (DSM 90), Bacillus licheniformis (DSM 13), Staphylococcus aureus (DSM 20231), Streptococcus equi (ATCC 9528), Pseudomonas putida (ATCC 12633), Pseudomonas aeruginosa (ATCC 9027), Pseudomonas fluorescens (ATCC 49838), Burkholderia mallei (RR0053), Burkholderia pseudomallei (ATCC 23343), Burkholderia stabilis (CCUG 34168), Burkholderia multivorans (CCUG 37240), Yersinia enterocolitica (O:8 Ye/80), Yersinia pseudotuberculosis (DSM 8992), Yersinia pestis (Kim), Brucella melitensis biotype 1 (16M Weybridge), Brucella abortus biotype 1 (544 Weybridge), Brucella suis biotype 1 (1330 Weybridge), Brucella ovis biotype 1 (63/290 Weybridge), Klebsiella oxytoca (CCUG 15788), Serratia marcescens, Proteus mirabilis, and Escherichia coli (DSM 30083)Statistical AnalysisMinitab17 software was used for calculation of prevalence %age and respective 95% confidence interval. ODD ratio, respective 95% confidence interval for ODD ratio and Chi-square were calculated using IBM SPSS Statistics 13.0 for Windows? (IBM Corporation). For calculation of significance of association (p< 0.05) between seroprevalence and various variables (sex, age, breed, species, parity, farm, district, lactational status, reproductive status, tick infestation, body condition and reproductive disorders) Chi-square test was performed. Table 3.12: Questionnaire for Coxiellosis-Individual Animal Data Farm Name: ___________________ Farm Location: _____________ V.O/Farm Manager Sig & Stamp: __________ Contact: __________S.NAnimalIDSpeciesFarmDistrictAgeSexBreedParityLactation StatusReproductive StatusReproductive DisordersBody ConditionPresence ofTicksIndirect-ELISAqPCRSerum poolsTick pools1234567891011121314151617181920212223… Continue Abbreviations used: Dated: _____________ L=Lactating NL= Non-lactating P= Pregnant NP= Non-PregnantOverall General Farm Data Flock size……...…… Number of samples collected from each flock…..………………… Vaccination history…………………………. Presence of ticks ………………………… Kidding/ Lambing rate ……………………… Twinning rate ………………………………. Management system a) Extensive b) Semi-intensive c) intensive Purpose of farming………………………………………. Feeding system a) Grazing b) stall feeding Watering system………………………………………….. Farm to Farm distance……………………………………. Veternarian/ Paravet/ Farmer awareness of Q fever …………………Qudrat Ullah, PhD Theriogenology, UAF. Q fever Research (+92-346-7761159/ +92-334-7972741)Chapter-4RESULTS4.1: Farm-wise sero-prevalence of Coxiellosis in small ruminants Results obtained from this study revealed a prevalence of 65.9, 26.2, 13.6, 13.3, 11.4, 10.8, 5.8, 5.8 and 4.8% at 205-TDA, RakKharewala, Kallurkot, Khushab, Bahadarnagar, Jogaitpur, Alladad Jahania, Fazilpur, and Rakh Ghulamana farms, respectively (Table 4.1). The overall prevalence of Coxiellosis at nine livestock farms was 15.3%. The highest prevalence (65.9%) of the disease was recorded at 205-TDA farm, while the lowest prevalence (4.8%) was at Rakh Ghulamana farm (Figure 4.1). Statistical analysis revealed that prevalence of C. burnetii antibodies in serum samples obtained from small ruminants was significantly (p=0.000, df=8, χ2=141.87) different among various livestock farms included in this survey.4.2. Farm-wise sero-prevalence of Coxiellosis in goats Results pointed out 24.3, 20.5, 6.97 and 5.9% prevalence of Coxiellosis at RakKharewala, Bahadarnagar, Fazilpur and Alladad Jahania farms, respectively (Table 4.2). Overall sero-prevalence of the disease at goat farms was 15%. Highest C. burnetii seropositive cases (24.3%) were found in goats kept at RakKharewala farm, while seropositivity (5.9%) was lowest in goats at Alladad Jahania farm (Figure 4.2). A statistically significant (p=0.000, df=3, χ2=30.551) difference in prevalence of the disease was observed among various goat farms.4.3. Farm-wise sero-prevalence of Coxiellosis in sheepPrevalence of the disease was recorded as 65.9, 31.5, 13.6, 13.3, 10.8, 6.8, 5.00, 4.8 and 0.00% at 205/TDA, Rakh Kharewala, Kallurkot, Khushab, Jogaitpeer, Bahadurnagar, Alladad Jahania, Rakh Ghulamana and Fazilpur farm, respectively (Table 4.3). The overall prevalence of Coxiellosis at sheep farms was found to be 15.6%. Prevalence of C. burnetii infection was highest at 205/TDA farm (65.9%), while lowest at Fazilpur farm (0.00%) (Figure 4.3). Statistically, the difference in prevalence of the infection among various sheep farms was highly significant (p=0.000, df=8, χ2=116.57). Table 4.1: Farm-wise sero-prevalence of Coxiellosis in small ruminants VariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioFarms205-TDA41271465.949.479.938.0214.03103.0χ2=86.82df=8p=0.000RakKharewala2837420926.221.131.76.983.1215.60Kallurkot2231913.62.934.93.110.7413.07Khushab4563913.35.426.83.030.969.55Bahadarnagar1321511711.46.5 182.531.006.41Jogaitpur3743310.83 25.42.390.668.65Fazilpur523495.83.2 9.51.210.304.85Alladad Jahania243142295.81.2 15.91.210.473.06RakGhulaman14571384.82 9.71--Overall100015384715.312.318.7---Fig. 4.1: Farm-wise sero-prevalence of Coxiellosis in small ruminantsTable 4.2: Farm-wise sero-prevalence of Coxiellosis in goatsVaribaleLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioFarmsRakKharewala2105115924.318.630.74.271.2714.41χ2=30.55df=3p=0.000Bahadarnagar4493520.59.835.33.420.8613.67Fazilpur433406.971.519.10.830.233.11Alladad Jahania203121915.93.110.11--Overall500754251512.0718.33---Fig. 4.2: Farm-wise sero-prevalence of Coxiellosis in goatsTable 4.3: Farm-wise sero-prevalence of Coxiellosis in sheepVariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioFarms205-TDA41271465.8549.479.938.0214.03103.0χ2=116.57df=8p=0.000RakKharewala73235031.5021.143.49.063.6722.43Kallurkot2231913.632.934.93.110.7413.07Khushab4563913.335.126.83.030.969.55Jogaitpur3743310.813.025.42.390.668.64Bahadarnagar886826.812.514.31.440.474.44Alladad Jahania402385.000.616.91.030.215.20RakGhulaman14571384.822.09.713.640.030.18Fazilpur9090.000.028.31--Overall5007842215.612.6218.98---Fig. 4.3: Farm-wise sero-prevalence of Coxiellosis in sheep4.4: District-wise sero-prevalence of Coxiellosis in small ruminants Seroprevalence of Coxiellosis was 26.2, 17.8, 13.3, 11.4, 10.8, 5.8 and 5.8% at Layyah, Bhakkar, Khushab, Okara, Bhawalpur, Khanewal and Rajanpur districts of Punjab, respectively (Table 4.4). Overall prevalence of the disease in these districts was 15.3%. The highest prevalence (26.2%) of the disease was recorded in animals of Layyah district, while the lowest prevalence (5.8%) was in animals of Rajanpur district (Figure 4.4). Statistical analysis showed a significant (p=0.000, df=6, χ2=49.689) difference in prevalence of disease in small ruminants among various districts of Punjab province. 4.5. District-wise sero-prevalence of Coxiellosis in goats Results pointed out a prevalence of 24.2, 20.45, 6.97 and 5.91% at Layyah, Okara, Rajanpur and Khanewal districts of Punjab, respectively (Table 4.5). The overall prevalence of C. burnetii infection was recorded as 15%. Sero-prevalence of the disease was the highest in goats of Rajanpur district (24.2%), while the lowest was in Khanewal district (0.00%) (Figure 4.5). A statistically significant (p=0.000, df=3, χ2=30.551) difference in prevalence of C. burnetii infection in goats was observed among various districts of Punjab province.4.6. District-wise sero-prevalence of Coxiellosis in sheep Results pointed out a prevalence of 31.50, 17.78, 13.33, 10.81, 6.81, 5.00 and 0.00% at Layyah, Bhakkar, Khushab, Bhawalpur, Okara, Khanewal and Rajanpur districts of Punjab, respectively (Table 4.6). Overall disease prevalence in these seven districts was 15.6%. Regarding district-wise prevalence, the highest seropositivity (31.5%) was found in sheep of Layyah district, while seropositivity (0.00%) was the lowest in animals of Rajanpur district (Figure 4.6). A significant (p=0.000, df=6, χ2=25.837) difference in prevalence of ovine Coxiellosis was found among various districts of Punjab province.Table 4.4: District-wise sero-prevalence of Coxiellosis in small ruminantsVariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioDistrictsLayyah2837420926.221.1 31.75.791.7519.11χ2=49.689df=6p=0.000Bhakkar2083717117.812.8 23.73.541.0411.95Khushab4563913.35.1 26.82.520.5910.69Okara1321511711.46.5 182.10.587.56Bhawalpur3743310.83 25.41.980.429.43Khanewal24314495.83.2 9.510.283.61Rajanpur5232295.81.2 15.91--Overall100015384715.312.318.7---Fig. 4.4: District-wise sero-prevalence of Coxiellosis in small ruminantsTable 4.5: District-wise sero-prevalence of Coxiellosis in goatsVariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioDistrictsLayyah2105115924.218.630.74.2771.2714.41χ2=30.551df=3p=0.000Okara4493520.459.835.33.4290.8613.67Rajanpur433406.971.519.13.480.090.95Khanewal 203121915.913.110.11--Overall500754251512.0718.33---Fig. 4.5: District-wise sero-prevalence of Coxiellosis in goatsTable 4.6: District-wise sero-prevalence of Coxiellosis in sheep VariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioDistrictsLayyah73235031.5021.143.48.7401.9439.37χ2=25.837df=6p=0.0053Bhakkar2083717117.7812.823.74.1110.9517.80Khushab4563913.335.126.82.9230.5515.40Bhawalpur3743310.813.025.42.3030.4013.39Okara886826.812.514.31.3900.277.21Khanewal 402385.00.616.96.300.040.70Rajanpur90900.028.31--Overall5007842215.612.6218.98---Fig. 4.6: District-wise sero-prevalence of Coxiellosis in sheep 4.7. Breed-wise sero-prevalence of Coxiellosis in small ruminants Results obtained from this study pointed out a prevalence of 44.11, 28.57, 21.7, 17.1, 13.3, 10.8, 8.7, 7.0, 7.0 and 6.4% in nachi, DDP, thali, teddy, kajli, buchi, mundri, lohi, nukri and beetal breeds of small ruminants, respectively (Table 4.7). The overall prevalence of the infection in various breeds of small ruminants was 15.3%. Prevalence of disease was the highest in nachi breed (44.11%), while the lowest was in beetal (6.4%) (Figure 4.7). Statistical analysis revealed that prevalence of Coxiellosis among various breeds of small ruminants was significantly (p=0.000, df=9, χ2=60.954) different.4.8. Breed-wise sero-prevalence of Coxiellosis in goatsPrevalence of C. burnetii infection was recorded as 44.11, 28.57, 17.07, 6.97 and 6.40% in nachi, DDP, teddy, nukri and beetal breeds of goat, respectively (Table 4.8). Overall prevalence of the disease was 15%. The highest prevalence (44.11%) of the disease was recorded in nachi breed, while the lowest prevalence (6.40%) was in beetal breed (Figure 4.8). Statistically, prevalence of the disease was significantly (p=0.000, df=4, χ2=45.187) different among various breeds of goats.4.9. Breed-wise sero-prevalence of Coxiellosis in sheepResults pointed out a prevalence of 21.70, 13.33, 10.81, 8.69 and 7.01% in thali, kajli, buchi, mundri and lohi breeds of sheep, respectively (Table 4.9). Overall disease prevalence in various breeds of sheep was 15.6%. As far as breed was concerned, the highest seropositivity (21.70%) was found in thali breed, while seropositivity (7.01%) was the lowest in lohi breed of sheep (Figure 4.9). A statistically significant (p=0.003, df=4, χ2=16.168) difference was found in prevalence of Coxiellosis among various breeds of sheep. Table 4.7: Breed-wise sero-prevalence of Coxiellosis in small ruminantsVariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioBreedNachy 34151944.127.262.110.522.7240.79χ2=60.954df=9p=0.000DDP56164028.617.342.25.331.4419.74Thali 2585620221.716.827.23.691.1012.40Teddy 1642813617.111.723.72.740.799.50Kajli 4563913.35.126.82.050.488.78Buchi 3743310.83.025.41.610.347.74Mundri 464428.72.420.81.270.276.03Lohi 114810673.113.41.0060.253.98Nukri 4334071.519.16.320.040.58Beetal 203131906.43.510.71--Overall 100015384715.312.318.7---Fig. 4.7: Breed-wise sero-prevalence of Coxiellosis in small ruminantsTable 4.8: Breed-wise sero-prevalence of Coxiellosis in goatsVariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioBreedNachy 34151944.1127.262.110.520.060.33χ2=45.187df=4p=0.000DDP56164028.5717.342.25.330.291.46Teddy 1642813617.0711.723.72.740.190.80Nukri 433406.971.519.16.30.040.58Beetal 203131906.403.510.71--Overall 500754251512.0718.33---Fig. 4.8: Breed-wise sero-prevalence of Coxiellosis in goatsTable 4.9: Breed-wise sero-prevalence of Coxiellosis in sheep VariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95% StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioBreedThali 2585620221.7016.827.23.6731.697.99χ2=16.168df=4p=0.003Kajli 4563913.335.126.82.0380.666.25Buchi 3743310.813.025.41.6060.455.67Mundri 464428.692.420.81.2620.364.41Lohi 11481067.013.113.41--Overall 5007842215.612.6218.98---Fig. 4.9: Breed-wise sero-prevalence of Coxiellosis in sheep4.10. Association of antibodies against C. burnetii in small ruminants with history of reproductive disorders Sero-prevalence of Coxiellosis was recorded as 51.61, 32.43, 31.81 and 25.00% in animals with history of abortion, premature delivery, stillbirth and repeat breeding, respectively (Table 4.10). The overall prevalence of the disease in small ruminants having history of reproductive disorders was 42.13%. The highest prevalence (51.61%) of the disease was recorded in animals with abortion history, while the lowest prevalence (25.00%) was found in animals with history of repeat breeding (Figure 4.10). Prevalence of C. burnetii infection in animals with history of various reproductive problems was significantly (p=0.000, df=4, χ2=133.984) different. 4.11. Association of antibodies against C. burnetii in goats with history of reproductive disorders Results revealed a prevalence of 59.09, 35.00 and 33.33% in goats with history of abortion, premature delivery and stillbirth, respectively (Table 4.11). Overall prevalence of C. burnetii infection in goats with history of reproductive losses was recorded as 47.56%. Prevalence was the highest in goats with abortion history (59.09%), while the lowest was in animals with history of stillbirth (33.33%) (Figure 4.11). Statistical analysis revealed that sero-prevalence of Coxiellosis was significantly (p=0.000, df=4, χ2=47.530) different in goats with history of various kinds of reproductive problems. 4.12. Association of antibodies against C. burnetii in sheep with history of reproductive disorders Results of the current study pointed out a prevalence of 44.89, 30.76, 29.4 and 25.00% in sheep with history of abortion, stillbirth, premature delivery and repeat breeding, respectively (Table 4.12). An overall prevalence of the disease in sheep with history of reproductive losses was 37.5%. Among reproductive disorders, highest C. burnetii seropositive cases (44.89%) were found in sheep with abortion history, while seropositivity (25.00%) was lowest in sheep with history of repeat breeding (Figure 4.12). Prevalence of the disease was significantly (p=0.000, df=3, χ2=91.483) different in sheep with history of various reproductive problems.Table 4.10: Association of antibodies against C. burnetii in small ruminants with history of reproductive disorders VariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioReproductive disordersAbortion93484551.641 62.110.176.37 16.26χ2=133.984df=4p=0.000Premature delivery37122532.418 49.84.582.21 9.47Stillbirth44143031.818.6 47.64.452.26 8.75Repeat breeder413250.6 80.63.180.33 30.94No822787449.57.6 11.71--Overall100015384715.312.318.7---Fig. 4.10: Association of antibodies against C. burnetii in small ruminants with history of reproductive disorders Table 4.11: Association of antibodies against C. burnetii in goats with history of reproductive disordersVariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower Ratio Upper ratioLower Ratio Upper ratioReproductive disordersAbortion44261859.143.273.715.327.6830.60χ2=91.483df=3p=0.000Premature delivery207133515.459.25.712.1415.23Stillbirth1861233.313.359.05.311.8814.98NO418363828.60.0611.601--Overall500754251512.0718.33---Fig. 4.11: Association of antibodies against C. burnetii in goats with history of reproductive disorders Table 4.12: Association of antibodies against C. burnetii in sheep with history of reproductive disorders VariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioReproductive disordersAbortion49222744.930.759.87.023.6813.42χ2=47.530df=4p=0.000Stillbirth2681830.814.351.83.831.579.35Premature delivery1751229.410.356.03.591.2110.69Repeat breeder413250.680.62.870.2928.25No4044236210.47.613.81--Overall5007842215.612.6218.98---Fig. 4.12: Association of antibodies against C. burnetii in sheep with history of reproductive disorders4.13. Association of antibodies against C. burnetii in small ruminants with presence of ticks on sheep and goats Results revealed that 60.1% of animals with tick infestation and 6.6% of animals without tick infestation were seropositive for coxiellosis (Table 4.13). Based on tick infestation (Yes/ No), overall prevalence of Coxiellosis in small ruminants was 16.3%. Highest odds of seropositivity (60.1%) were recorded in animals with presence of ticks, while lowest (6.6%) in animals without tick infestation (Figure 4.13). A significant (p=0.000, df=1, χ2=301.914) difference was found in prevalence of C. burnetii infection between animals with tick infestation and those without tick infestation.4.14. Species-wise sero-prevalence of Coxiellosis in small ruminants Sero-prevalence of C. burnetii infection was 15.6 and 15% in sheep and goats, respectively (Table 4.14). Overall prevalence of the disease in both species was 15.3%. A slightly higher prevalence of the disease was found in sheep (15.6%) as compared to goats (15%) (Figure 4.14). Prevalence of the disease was non-significantly (p=0.792, df=1, χ2=0.069) different between the two species. Table 4.13: Association of antibodies against C. burnetii in small ruminants with presence of ticks on sheep and goatsVariableLevelsScreened sampleCoxiellosisCI 95%Odd RatiosCI 95%StatisticsSero-positiveSero-negativePrev.(%)LowerRatioUpperRatioLowerRatioUpperRatioPresence of ticksYes163986560.152.267.721.4414.1432.50χ2=301.914df=1p=0.000No837657726.658.51--Overall100016383716.312.318.7---Fig. 4.13: Association of antibodies against C. burnetii in small ruminants with presence of ticks on sheep and goatsTable 4.14: Species-wise sero-prevalence of Coxiellosis in small ruminantsVariablesLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioSpeciesSheep5007842215.612.519.11.0470.7421.478χ2=0.05df=1p=0.821Goat500754251512.018.41--Overall100015384715.312.318.7---Fig. 4.14: Species-wise sero-prevalence of Coxiellosis in small ruminants4.15. Association of antibodies against C. burnetii with poor body conditions in small ruminants Seropositivity against C. burnetii was recorded as 45.71, 44.9 and 37.37% in animals with weak body condition, weak kid delivery and weak lamb delivery, respectively (Table 4.15). The overall prevalence of Coxiellosis in animals with poor body conditions was 43.73%. The highest prevalence (45.71%) of the disease was recorded in small ruminants with weak body condition, while lowest prevalence (37.37%) was found in animals with weak lamb delivery (Figure 4.15). Statistical analysis revealed that prevalence of the disease was significantly (p=0.000, df=3, χ2=124.868) different between weak animals and animals with weak lamb/ kid delivery. 4.16. Association of antibodies against C. burnetii with poor body conditions in goats Results obtained from this study pointed a prevalence of 44.9 and 40.00% in goats with weak kid delivery and weak body condition, respectively (Table 4.16). Overall prevalence of the disease in goats with poor body conditions was 42.69%. A higher prevalence was recorded in weak animals (44.9%) as compared to animals with weak kid delivery (40.00%) (Figure 4.16). A significant (p=0.000, df=2, χ2=65.556) difference in prevalence of C. burnetii infection was found between weak animals and animals with weak kid delivery.4.17. Association of antibodies against C. burnetii with poor body conditions in sheepSero-prevalence of the disease was 53.33% in sheep with weak body condition and 37.73% in ewes with weak lamb delivery. The overall disease prevalence in sheep with poor body conditions was 43.37%. The highest prevalence (53.33%) of the disease was recorded in small ruminants with weak body condition, while lowest prevalence (37.37%) was found in animals with weak lamb delivery (Figure 4.17). Statistical analysis revealed that prevalence of the disease in sheep was significantly (p=0.000, df=2, χ2=61.845) different between weak animals and animals with weak lamb delivery.Table 4.15: Association of antibodies against C. burnetii with poor body conditions in small ruminantsVariablesLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioBody conditionWeak70323845.733.758.17.984.7313.49χ2=124.868df=3p=0.000Weak kid birth49222744.930.759.87.734.2014.20Weak lamb birth53203337.7324.852.15.753.1510.49Apparently Good828797499.57.611.71--Overall100015384715.312.318.7---Fig. 4.15: Association of antibodies against C. burnetii with poor body conditions in small ruminantsTable 4.16: Association of antibodies against C. burnetii with poor body conditions in goatsVariablesLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioBody conditionWeak kid 49222744.930.759.88.2364.2715.88χ2=65.556df=2p=0.000Weak4016244024.956.76.7393.2913.80Apparently Good4113737496.412.21--Overall500754251512.0718.33---Fig. 4.16: Association of antibodies against C. burnetii with poor body conditions in goatsTable 4.17: Association of antibodies against C. burnetii with poor body conditions in sheepVariablesLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioBody conditionWeak30161453.334.371.710.204.6522.37χ2=61.845df=2p=0.000Weak lamb53203337.724.852.15.412.8510.27Apparently Good4174237510.17.413.41--Overall5007842215.612.6218.98---Fig 4.17: Association of antibodies against C. burnetii with poor body conditions in sheep4.18. Age-wise sero-prevalence of Coxiellosis in small ruminantsSero-prevalence of Coxiellosis in different age groups of small ruminants was recorded as 15.78% in up to 1 year age group, 10.16% in >1-2.5 years age group, 19.6% in >2.5-4 years age group and 14.77% in >4years age group (Table 4.18). Overall prevalence of Coxiellosis in different age groups of animals was recorded as 15.3%. Prevalence was highest in >2.5-4 years age group animals (19.6%), while lowest in >1-2.5 years age group (10.6%) (Figure 4.18). Statistical analysis revealed that prevalence of the disease was non-significantly (p=0.063, df=3, χ2=7.281) different among various age groups of small ruminant.4.19. Age-wise sero-prevalence of Coxiellosis in goatsResults pointed out a prevalence of 16.41, 9.19, 16.26 and 18.24% in up to 1 year age group, >1 to 2.5 years age group, >2.5 to 4 years age group and >4 years age group, respectively (Table 4.19). The overall prevalence of C. burnetii infection in different age groups of goats was 15.6%. The highest prevalence (53.33%) of the disease was recorded in >2.5-4 years age group animals, while lowest prevalence (37.37%) was found in >1 to 2.5 years age group (Figure 4.19). Statistically, prevalence of C. burnetii infection among different age groups of goats was non-significantly (p=0.315, df=3, χ2=3.545) different. 4.20. Age-wise sero-prevalence of Coxiellosis in sheepPrevalence of C. burnetii infection was recorded as 15.15, 11.11, 21.23 and 13.41% in up to 1 year age group, >1 to 2.5 years age group, >2.5 to 4 years age group and >4 years age group, respectively (Table 4.20). An overall prevalence of the disease was recorded as 15% in different age groups of sheep. Prevalence was highest in >2.5-4 years age group of animals (21.23%), while lowest in >1-2.5 years age group (11.11%) (Figure 4.20). Statistical analysis showed that prevalence of the disease was non-significantly (p=0.174, df=3, χ2=4.971) different among various age groups of sheep.Table 4.18: Age-wise sero-prevalence of Coxiellosis in small ruminants VariablesLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioAge (years)Up to 11332112215.81023.10.6530.381.14χ2=7.281df=3p=0.063>1-2.5 1771815910.26.115.61-->2.5-42504920119.614.925.11.0820.631.85>4 4406537514.811.618.41.4060.932.12Overall100015384715.312.318.7---Fig 4.18: Age-wise sero-prevalence of Coxiellosis in small ruminants Table 4.19: Age-wise sero-prevalence of Coxiellosis in goatsVariablesLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioAge (years)Up to 166105615.157.526.11.1520.532.49χ2=4.971df=3p=0.174>1-2.5 90108011.115.519.51-->2.5-4 113248921.2314.129.91.7400.973.13>4 2313120013.419.318.51.580.351.12Overall500754251512.0718.33---Fig. 4.19: Age-wise sero-prevalence of Coxiellosis in goatsTable 4.20: Age-wise sero-prevalence of Coxiellosis in sheepVariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioAge (years)Up to 167115616.418.527.51.0110.482.13χ2=3.545df=3p=0.315>1-2.5 878799.194.117.31-->2.5-4 1372511218.2412.225.71.1490.652.03>4 2093417516.2611.522.01.120.511.56Overall5007842215.612.6218.98---Fig 4.20: Age-wise sero-prevalence of Coxiellosis in sheep4.21. Parity-wise sero-prevalence of Coxiellosis in small ruminantsResults showed a prevalence of 15.6, 14.5 and 13.3% in nulliparous, multiparous and primiparous groups of animals, respectively (Table 4.21). The overall prevalence of the disease was 14.89%. The highest prevalence (15.6%) of the disease was recorded in nulliparous animals, while lowest prevalence (13.3%) was found in primiparous animals (Figure 4.21). Statistical analysis revealed that prevalence of Coxiellosis was non-significantly (p=0.838, df=2, χ2=0.353) different among various parity groups of small ruminants. 4.22. Parity-wise sero-prevalence of Coxiellosis in goats Sero-prevalence of C. burnetii infection was recorded as 15.7, 15.1 and 14.5% in nulliparous, primiparous and multiparous groups of goats, respectively (Table 4.22). Overall prevalence of Coxiellosis in various parity groups of goats was 15.01%. Prevalence was highest in nulliparous goats (15.7%), while lowest in multiparous animals (14.5%) (Figure 4.22). A statistically non-significant (p=0.941, df=2, χ2=0.122) difference in prevalence of the disease was observed among various parity groups of goats.4.23. Parity-wise sero-prevalence of Coxiellosis in sheep Results of the current study revealed a prevalence of 16.3, 15.4 and 11.5% in multiparous, nulliparous and primiparous groups of sheep, respectively (Table 4.23). The overall prevalence of Coxiellosis in different parity groups of sheep was 14.79%. Nulliparous animals showed highest seropositivity (16.3%) against C. burnetii antibodies, while primiparous animals showed lowest seropositivity (11.5%) against the pathogen (Figure 4.23). Statistical analysis of data revealed that prevalence of Coxiellosis was non-significantly (p=0.675, df=2, χ2=0.786) different among various parity groups of sheep. Table 4.21: Parity-wise sero-prevalence of Coxiellosis in small ruminants VariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioParityNulliparous3014725415.611.720.21.20.632.29χ2=0.353df=2p=0.838Multiparous 5949250215.512.718.71.190.652.18Primiparous 105149113.37.521.41--Overall100015384715.312.318.7---Fig. 4.21: Parity-wise sero-prevalence of Coxiellosis in small ruminantsTable 4.22: Parity-wise sero-prevalence of Coxiellosis in goatsVariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioParityNulliparous1652613915.710.622.21.050.442.49χ2=0.122df=2p=0.941Primiparous 5384515.16.727.61.040.412.23Multiparous 2824124114.510.619.21--Overall500754251512.0718.33---Fig. 4.22: Parity-wise sero-prevalence of Coxiellosis in goats Table 4.23: Parity-wise sero-prevalence of Coxiellosis in sheepVariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioParityMultiparous 3125126116.312.420.91.4980.613.69χ2=0.786df=2p=0.675Nulliparous1362111515.49.822.61.4000.533.69Primiparous 5264611.54.423.41--Overall5007842215.612.6218.98---Fig. 4.23: Parity-wise sero-prevalence of Coxiellosis in sheep4.24. Sex-wise sero-prevalence of Coxiellosis in small ruminants Results revealed that prevalence of infection was 18.7 and 14.9% in male and female animals, respectively (Table 4.24). Overall sex-based prevalence of the disease in small ruminants was 15.3%. Prevalence of the disease was higher in males (18.7%) compared to female animals (14.9%) (Figure 4.24). Statistically, prevalence of Coxiellosis was non-significantly (p=0.302, df=1, χ2=1.064) different between the two sexes.4.25. Sex-wise sero-prevalence of Coxiellosis in goats Sero-prevalence of C. burnetii infection was 15.01 and 14.9% in does and bucks, respectively (Table 4.25). The overall prevalence of the disease was 15%. Individual prevalence of Coxiellosis in both male and female goats was almost similar (Figure 4.25). Statistical analysis of data also showed a non-significant (p=0.987, df=1, χ2=0.000) difference in prevalence Coxiellosis between male and female goats. 4.26. Sex-wise sero-prevalence of Coxiellosis in sheepResults showed a prevalence of 35 and 14.8% in ram and ewe, respectively (Table 4.26). Overall prevalence of Coxiellosis in both sexes of sheep was 15.6%. The highest prevalence of the disease was recorded in males (35%), while lowest prevalence was found in female animals (14.8%) (Figure 4.26). Statistically, the prevalence of C. burnetii infection was significantly (p=0.015, df=1, χ2=5.955) different between the two sexes.Table 4.24: Sex-wise sero-prevalence of Coxiellosis in small ruminants VariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioSexMale107208718.711.827.41.310.782.21χ2=1.064df=1p=0.302Female 89313376014.912.617.41--Overall100015384715.312.318.7---Fig. 4.24: Sex-wise sero-prevalence of Coxiellosis in small ruminants Table 4.25: Sex-wise sero-prevalence of Coxiellosis in goats VariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioSexMale4136235115.0111.718.810.521.90χ2=0.000df=1p=0.987Female 87137414.948.224.21--Overall500754251512.0718.33---Fig. 4.25: Sex-wise sero-prevalence of Coxiellosis in goats Table 4.26: Sex-wise sero-prevalence of Coxiellosis in sheep VariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioSexMale207133515.459.23.1021.208.04χ2=5.955df=1p=0.015Female 4807140914.7911.718.31--Overall5007842215.612.6218.98---Fig. 4.26: Sex-wise sero-prevalence of Coxiellosis in sheep 4.27. Sero-prevalence of Coxiellosis in pregnant and non-pregnant small ruminants Prevalence of infection was recorded as 17.24 and 10.65% in pregnant and non-pregnant animals, respectively. The overall prevalence of the disease was 14.89% based on the reproductive status of animal. Prevalence was high (17.24%) in pregnant animals, while low (10.65%) in non-pregnant animals (Figure 4.27). Statistical analysis of data showed a significant (p=0.008, df=2, χ2=7.023) difference in prevalence of Coxiellosis between pregnant and non-pregnant animals. 4.28. Sero-prevalence of Coxiellosis in pregnant and non-pregnant goatsResults of the current study showed a prevalence of 17.87 and 10% in pregnant and non-pregnant goats, respectively (Table 4.28). Overall prevalence of C. burnetii infection was recorded as 15.01%. The highest prevalence (17.87%) of the disease was recorded in pregnant goats, while the lowest prevalence (10%) was found in non-pregnant goats (Figure 4.28). A statistically significant (p=0.031, df=1, χ2=4.638) difference was observed in prevalence of C. burnetii infection between pregnant and non-pregnant goats. 4.29. Sero-prevalence of Coxiellosis in pregnant and non-pregnant sheep Sero-prevalence of the disease was 16.72 and 11.24% in pregnant and non-pregnant sheep, respectively (Table 4.29). An overall prevalence of Coxiellosis was 14.79%. Prevalence was higher in pregnant animals (16.72%), while lower in non-pregnant animals (11.24%) (Figure 4.29). Results obtained from this study showed a non-significant (p=0.106, df=2, χ2=2.607) difference in prevalence of the disease in sheep based on their reproductive status. . Table 4.27: Sero-prevalence of Coxiellosis in pregnant and non-pregnant small ruminants VariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioReproductive statusPregnant5749947517.314.220.61.751.152.65χ2=7.023df=1p=0.008Non-Pregnant3193428510.77.514.61--Overall89313376014.8912.6717.34---Fig. 4.27: Sero-prevalence of Coxiellosis in pregnant and non-pregnant small ruminants Table 4.28: Sero-prevalence of Coxiellosis in pregnant and non-pregnant goatsVariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioReproductive statusPregnant2634721617.8713.423.01.9581.053.64χ2=4.638df=1p=0.031Non-Pregnant15015135105.716.01--Overall4136235115.0111.8118.70---Fig. 4.28: Sero-prevalence of Coxiellosis in pregnant and non-pregnant goatsTable 4.29: Sero-prevalence of Coxiellosis in pregnant and non-pregnant sheepVariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioReproductive statusPregnant3115225916.7212.721.31.5850.902.78χ2=2.607df=1p=0.106Non-Pregnant1691915011.246.917.01--Overall4807140914.7911.8218.18---Fig. 4.29: Sero-prevalence of Coxiellosis in pregnant and non-pregnant sheep4.30. Sero-prevalence of Coxiellosis in lactating and non-lactating small ruminants Results showed a prevalence of 20.20 and 8.3% in lactating and non-lactating animals, respectively (Table 4.30). An overall prevalence of Coxiellosis was 14.89% in small ruminants based on their lactational status. The highest prevalence (20.20%) of the disease was recorded in lactating animals, while lowest prevalence (8.3%) was found in non-lactating animals (Figure 4.30). Statistically, prevalence of the infection was significantly (p=0.000, df=1, χ2=24.691) different between lactating and non-lactating animals. 4.31. Sero-prevalence of Coxiellosis in lactating and non-lactating goatsResults of this study revealed that 20.70% of lactating and 8.1% of non-lactating goats were sero-positive for Coxiellosis (Table 4.31). Overall prevalence of the disease was 15.01%. Prevalence was higher in lactating goats (20.70%) as compared to non-lactating (8.1%) (Figure 4.31). Statistical analysis revealed that prevalence of Coxiellosis was significantly (p=0.000, df=1, χ2=12.803) different between lactating and non-lactating goats.4.32. Sero-prevalence of Coxiellosis in lactating and non-lactating sheepPrevalence of C. burnetii antibodies was recorded as 19.77 and 8.49% in lactating and non-lactating sheep sera, respectively (Table 4.32). The overall prevalence of Coxiellosis in sheep was 14.79% based on their lactational status. Highest seropositivity against C. burnetii antibodies was recorded in lactating (19.77%) as compared to non-lactating sheep (8.49%) (Figure 4.32). A significant (p=0.001, df=2, χ2=11.961) difference was observed in prevalence of the disease between lactating and non-lactating sheep.Table 4.30: Sero-prevalence of Coxiellosis in lactating and non-lactating small ruminants VariableLevelsTotal samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioLactational statusLactating49510039520.216.824.02.8001.844.25χ2=24.691df=1p=0.000Non-lactating398333658.35.811.41--Overall89313376014.8912.6717.34---Fig. 4.30: Sero-prevalence of Coxiellosis in lactating and non-lactating small ruminants Table 4.31: Sero-prevalence of Coxiellosis in lactating and non-lactating goatsVariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioLactational statusLactating2274718020.7015.626.62.9771.605.52χ2=12.803df=1p=0.000Non-lactating186151718.064.613.01--Overall4136235115.0111.8118.70---Fig. 4.31: Sero-prevalence of Coxiellosis in lactating and non-lactating goatsTable 4.32: Sero-prevalence of Coxiellosis in lactating and non-lactating sheepVariableLevelsScreened samplesCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioLactational statusLactating2685321519.7715.225.12.6571.504.69χ2=11.961df=1p=0.001Non-lactating212181948.495.113.11--Overall4807140914.7911.8218.18---Fig. 4.32: Sero-prevalence of Coxiellosis in lactating and non-lactating sheep4.33. Real-time qPCR-based prevalence of Coxiellosis in pooled serum samplesReal-time qPCR based analysis revealed that C. burnetii DNA was not detected in any of the sero-positive (ELISA positive) serum pools, however, C. burnetii DNA was present in 5 pools comprised of suspected serum samples (three from goats and two from sheep). C. burnetii DNA was detected in 21.42 and 13.33% of goats and sheep serum pools, respectively (Table 4.33). Overall prevalence of C. burnetii DNA in serum pools was 17.24%. Prevalence of C. burnetii DNA was higher in serum pools comprised of goat sera (21.42%) compared to that of sheep sera (13.33%) (Figure 4.33). Statistical analysis of the data showed that prevalence of C. burnetii DNA in serum pools was non-significantly (p=0.333, df=1, χ2=0.564) different between the two species. Amplification plots showing fluorescence data acquired during annealing and extension phase of qPCR performed on serum pools are presented in (Fig 4.34).Table 4.33: Real-time qPCR-based prevalence of Coxiellosis in pooled serum samplesVariableLevelsScreened poolsCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveSuspectedPrev.(%)Lower RatioUpper ratioLower RatioUpper ratioSpeciesGoat1431121.44.750.81.610.104.05χ2=0.333df=1p=0.564Sheep1521313.31.740.51--Overall2952417.246.6034.16---Fig. 4.33: Real-time qPCR-based prevalence of Coxiellosis in pooled serum samplesFig 4.34: Amplification plots showing fluorescence data acquired during annealing and extension phase of qPCR performed on serum pools. Positive and negative controls were included in each Q-PCR run to validate the results.4.35. Real-time qPCR-based prevalence of Coxiellosis in pools of ticks collected from small ruminants Results obtained from real-time qPCR based analysis revealed that the prevalence of C. burnetii DNA was 31.03 and 7.69% in sheep and goat tick pools, respectively (Table 4.34). The overall prevalence of C. burnetii DNA in tick pools was 20%. The highest prevalence (31.03%) of C. burnetii DNA was found in sheep tick pools, while lowest prevalence (7.69%) was found in goats tick pools (Figure 4.34). Univariate analysis revealed a significant (p=0.031, df=1, χ2=4.668) difference in prevalence of C. burnetii DNA in tick pools collected from sheep and goats. Amplification plots showing fluorescence data acquired during annealing and extension phase of qPCR performed on tick pools are presented in (Fig 4.36).Table 4.35: Real-time qPCR-based prevalence of Coxiellosis in pools of ticks collected from small ruminantsVariableLevelsScreened poolsCoxiellosisCI 95%Odd RatiosCI 95%StatisticsPositiveNegativePrev.(%)Lower RatioUpper ratioLower RatioUpper ratioTicks InfestationSheep ticks2992031.015.350.84.030.051.22χ2=4.668df=1p=0.031Goat ticks262247.70.925.11--Overall5511442010.9932.10---Fig. 4.35: Real-time qPCR-based prevalence of Coxiellosis in pools of ticks collected from small ruminantsFig. 4.36: Amplification plots showing fluorescence data acquired during annealing and extension phase of qPCR performed on tick poolsChapter-5DISCUSSIONCoxiellosis is a disease of zoonotic importance caused by a bacterium known as C. burnetii. It results in huge economical losses to the livestock industry in the form of reproductive problems. Diagnosis of C. burnetii infection in animals is of great importance not only to identify the infected flocks but also to determine the risk of disease transmission to human beings. It is also necessary for timely treatment of the disease, thereby reducing the chances of developing chronic disease in humans and the occurrence of economical losses to livestock industry (Selim and Elhaig, 2016; OIE, 2015). In Pakistan, the epidemiological information about geographic distribution of C. burnetii infection is very limited both in humans and livestock population. To the best of our information, this is the first molecular and serological survey, investigating the prevalence of Coxiellosis in ticks and small ruminants maintained at different government livestock farms of Punjab, Pakistan. In the current study, the presence of C. burnetii infection in small ruminants has been investigated using both ELISA and real-time qPCR methods, while in ticks, an in-house developed real-time qPCR was used for genomic detection of C. burnetii in pooled samples. Sero-diagnostic techniques are preferably used for screening of infection at herd level and among available serological assays, ELISA is ideal for detection of antibodies against C. burnetii in serum. It is more sensitive and specific than any other serological technique (European Food Safety Authority, 2010; Horigan et al., 2011; Zahid et al., 2016). Selim et al., (2016) and Magouras et al. (2017) reported 100% sensitivity and specificity for IDEXX Q fever Indirect ELISA. Therefore, Q fever-Indirect ELISA was applied to investigate C. burnetii serological status in sheep and goat flocks maintained at different livestock farms. However, use of serological assays for detection of C. burnetii antibodies in early stage of infection could be unproductive due to the time lapse for seroconversion spanning 3-4 weeks after the appearance of the first symptoms. During first two weeks of infection, PCR is the only reliable technique for diagnosis of C. burnetii infection (Howe et al., 2009; Niemczuk et al., 2014).Results of this study revealed an overall sero-prevalence of Coxiellosis in small ruminants as 15.3% (95% CI: 12.3-18.7) which varied from 15% (95% CI: 12.07-18.33) to 15.6% (95% CI: 12.62-18.98) between goats and sheep. The results showed that individual prevalence of the disease was almost similar in both species with a non-significantly (p = 0.792) higher prevalence in sheep as compared to goats. Rizzo et al., 2016 reported almost similar flock level sero-prevalence (14.5%) in sheep and goats. However, a few previous studies reported higher seropositivity in sheep as compared to goats (Hatchette et al., 2002; Vaidya et al., 2010; Kshash, 2012; Anasta?cio et al., 2013). Conversally, other studies pointed out higher prevalence of Coxiellosis in goats than sheep (Klaasen et al., 2014; Zahid et al., 2016; Rizzo et al., 2016). Variations in intrinsic susceptibility to this pathogen among small ruminants have not been described in detail so far (Klaasen et al., 2014). The species-wise variations in the prevalence of disease may be attributed to different farm managemental practices and variation in flock density at sheep and goats farms, which may contribute to the prevalence of the disease (Khaled et al., 2016; Zahid et al., 2016). This survey revealed that small ruminants are important reservoirs of C. burnetii infection in the study area.The prevalence of Coxiellosis varied greatly among seven districts of Punjab province. Results pointed out a very high to low district-wise prevalence of infection ranging from 26.2% (95% CI: 21.1-31.7) in district Layyah to 5.8 % (95% CI: 3.2-9.5) in district Rajanpur. Univariate analysis showed a sigificant (p=0.000) difference in prevalence of the disease among various districts of Punjab province. Ezatkhah et al. (2015) conducted a sero-epidemiological investigation on C. burnetii infection in five counties of Southeast Iranian region. They found that 26.4% of the tested small ruminants had seropositivity against C. burnetii antibodies which ranged from 5% in Sarbaz to 39.2% in Iranshahr. A very high prevalence was reported by Zahid et al. (2016) in Layyah and Muzaffargarh districts of Punjab province which varied from 69.4 to 75.0%, respectively. The higher prevalence of the disease might be due to the prevailing climatic and weather conditions in the study region (Aitken et al., 1987; Van der Hoek et al., 2011; Muema et al., 2017). This variation in prevalence of infection within flock and different geographical areas might be associated with the farm hygienic measures, usual managemental practices and environmental factors such as vegetation, soil moisture and presence of infected animals in the surroundings (Van der Hoek et al., 2011; Paul et al., 2012; Rizzo et al., 2016). Previous studies carried out on Coxiellosis in rural areas have found that poor hygienic and sanitary conditions can also contribute to the transmission of the disease. Similarly, amount of feed, restocking of animals, lambing and kidding rate in sheep and goats, and frequency of visiting people or livestock professionals at a farm also add to the transmission of infection (Lyytika¨inen et al., 1998; Schimmer et al., 2011; Zahid et al., 2016). These managemental and environmental factors might be responsible for higher sero-prevalence of Coxiellosis in these districts.In the current study, the overall prevalence of Coxiellosis at nine livestock farms was 15.3% (95% CI: 12.3-18.7). The highest prevalence 65.9% (95% CI: 49.4-79.9) of the disease was recorded at 205-TDA farm, while the lowest prevalence 4.8% (95% CI: 2.0-9.7) was at Rakh Ghulamana farm. Statistical analysis revealed that prevalence of the disease was significantly (p=0.001) different among various livestock frams included in this survey. A very recent serological investigation carried out by Zahid et al. (2016) in small ruminants of Punjab revealed a very high herd level prevalence (73.1%) of C. burnetii infection, ranging from 58.8% to 94.4%. While a very low herd/ flock level prevalence was recorded by Lambton et al. (2016) in sheep (3%) and goats (10.2%) in Great Britain. Farm-level factors e.g. location of farm, flock density, distance to the nearby livestock farm and the number of visiting peoples, and professional farm workers had a significant association with increased C. burnetii seropositivity. They also reported that comparison between farms belonging to different production systems and geographical locations showed significant differences in disease prevalence and concluded that the intrinsic farm factors are potential risk factors for dissemination of infection (McCaughey et al., 2010; Schimmer et al., 2011; Anastacio et al., 2013; Schimmer et al., 2014). Similarly, contact with other animals and sharing a similar grazing area with other livestock species during seasonal grazing increases the chance of gaining infection from the surroundings (Schimmer et al., 2014; Rizzo et al., 2016).In this study, 10 different breeds (5 each from sheep and goats) were investigated through Q fever-Indirect ELISA for seropositivity against C. burnetii antibodies. An overall breed-wise prevalence of the disease was recorded as 15.3% (95% CI: 12.3-18.7). The highest prevalence 44.1% (95% CI: 27.2-62.1) of the disease was recorded in nachi breed, while the lowest prevalence 6.4 (95% CI: 3.5-10.7) was in beetal breed. The difference in prevalence of disease among these breeds was highly significant (p=0.000). These results are parallel with the findings of previous authors (Paul et al., 2012; McCaughey et al., 2010; Cantas et al., 2011) who also found significant (p<0.05) difference between seropositivity against C. burnetii antibodies and breed-wise prevalence of the disease. However, Muema et al., (2017) did not record any association between breeds and prevalence of disease. The reason for this difference in vulnerability of various breeds to C. burnetii is still not known. Variation in genetic make-up of animals is a possible reason which needs further investigation through in depth studies to find out the possible cause (Paul et al., 2012).Results of the current study revealed that sero-prevalence of Coxiellosis was significantly (p=0.000) different in animal with previous history of reproductive problems including abortion (51.6%), premature delivery (32.4%), stillbirth (31.8%) and repeat breeding (25%). These findings are in agreement with the results of Garcia-Perez et al. (2009) who observed significantly higher prevalence of Coxiellosis in animals associated with history of reproductive problems. However, Ruiz-Fons et al. (2010) and Asadi et al. (2013) did not record any association between prevalence of Coxiellosis and history of reproductive disorders. Seropositivity against C. burnetii antibodies in small ruminants with history of reproductive problems does not means the C. burnetii is the sole cause of reproductive losses in these animals. Pathogens like Brucella melitensis and Toxoplasma gondii may be possible causes other than C. burnetii. The trophoblastic cells of placenta are first target cells of C. burnetii in pregnant animals. Massive multiplication of pathogen occurs in these cells, which may lead to different reproductive problems (Arserim et al., 2011; Saglam et al., 2016; Vaidya et al., 2010; Cantas et al., 2011; Agerholm, 2013). Molecular based studies and larger case series are required to identify the exact cause of reproductive losses in animals, and to explore the mechanism of fetal pathogenesis and pathology (Agerholm, 2013; Zahid et al., 2016). Many previous studies have considered ticks as an important reservoir and vector for transmission of C. burnetii infection. Even the highly virulent Nile Mile reference strain of C. burnetii was isolated from ticks in United States (Derrick, 1937; Psaroulaki et al., 2006; Duron et al., 2015). More than 40 tick species have been found infected with this pathogen, either naturally or experimentally (Lang, 1990; Kovacova and Kazar et al., 2002; Psaroulaki et al., 2006). In the current study, seropositivity was higher in animals with tick infestation 60.1% (as compare to 6.6% in those without ticks. Univariate analysis showed a strong association (p=0.000) between seropositivity against C. burnetii antibodies and presence of ticks on sheep and goats. These results are consistent with the findings of (Duron et al., 2015; Zahid et al., 2016) who recorded a significant association between ticks infestation and Coxiellosis. On the other hand, Babudieri (1959) did not find ticks as an important vector for C. burnetii infection in livestock. The present study revealed a strong association between Coxiellosis in small ruminants and ticks infestation which reflects the importance of ticks as natural reservoir of C. burnetii. According to Psaroulaki et al. (2006), the circulation of C. burnetii in domestic animals is maintained by the ticks and reported that ticks play an important role in maintaining its viability in nature.In the current study, prevalence of C. burnetii infection varied greatly between lactating (20.2%) and non-lactating (8.3%) animals. The overall prevalence of the disease was recorded as 14.89%. A significantly (p=0.000) higher difference was found in prevalence of the disease between lactating and non-lactating animals. We found only one study about prevalence of Coxiellosis in lactating and non-lactating animals. According to that study, sero-prevalence of the disease was higher in lactating animals as compared to non-lactating animals, although the difference was not statistically significant (Muema et al., 2017). Paul et al. (2012) reported a significantly (p< 0.005) higher prevalence of Coxiellosis in dairy breeds as compared to the beef breeds of cattle. The target sites for proliferation of C. burnetii are placenta and mammary glands of animals (Jung et al., 2014; Shapiro et al., 2015). In ruminants, immediately after inoculation, organism reaches the predilection site through the blood stream and resides in supramammary lymph nodes, mammary glands and placenta of pregnant animals (Hadush et al., 2016).Animals were grouped into two categories based upon their reproductive status i.e. Pregnant and non-pregnant. Higher prevalence 17.3% of Coxiellosis was recorded in pregnant animals as compared to 10.7% in non-pregnant ones. Univariate analysis showed a significant association (p=0.008) between seropositivity against C. burnetii antibodies and reproductive status in small ruminants. Van den Brom et al. (2013) also reported a higher sero-prevalence of the disease in pregnant and periparturient small ruminants as compared to non-pregnant and those in early gestation period. However, Abushahba et al. (2017) observed nearly equal prevalence of C. burnetii infection in sheep and goats based on their reproductive status. They concluded that both groups possess potential risk of disseminating infection to humans living in that area. The trophoblastic cells present in chorioallantoic membrane of placenta are the primary target cells of C. burnetii in pregnant animals (Sa?nchez et al., 2006). During periparturient period, massive multiplication of this pathogen occur within trophoblast cells, causing necrotic suppurative placentitis which ultimately leads to pregnancy failure in the form of abortion, stillbirth, premature delivery and birth of weak offspring (Sa?nchez et al., 2006; Arserim et al., 2011).A significant (p=0.000, df=3, χ2=124.868) difference was found in prevalence of the disease between weak animals and animals with weak lamb/ kid delivery. Highest prevalence (45.7%) of C. burnetii infection was recorded in weak animals followed by weak kids (44.9%) and weak lambs (37.7%). Agerholm (2013) also reported that C. burnetii infective ewes give birth to noticeably small and underweight lambs. Similarly Saegerman et al. (2013) found an association between Coxiellosis and delivery of stillborn or weak calves (OR = 2.14 with 95% CI: 1.05–4.39). Ganter (2015) conducted a study in goats and found that full-term kids delivered by C. burnetii seropositive ewes were emaciated with reduced body weight and high mortality rate. He also found that many of the apparently healthy kids infected with C. burnetii were suffering from digestive and respiratory tract problems.An in-house developed real-time qPCR targeting single copy icd gene was used for genomic detection of C. burnetii DNA in serum pools. The qPCR-based analysis revealed that C. burnetii DNA was not detected in any of the seropositive samples, however, the DNA was detected in 5 pools of suspected serum samples. The prevalence of C. burnetii DNA was recorded as 21.4% (95% CI: 4.7-50.8) in goat serum pools and 13.3% (95% CI: 1.7-40.5) in sheep serum pools. Univariate analysis of the data found no association (p=0.564) between detection of C. burnetii DNA in serum pools and species-wise prevalence of the disease. Schneeberger et al. (2010) also recorded PCR positive results in 98% seronegative samples collected from acute Q fever patients with inconclusive Q fever serology. The presence of C. burnetii DNA in suspected serum pools indicated that infection was in the incubation period or in early acute phase (within 2 weeks post infection), as C. burnetii DNA can easily be detected during this period through qPCR. However, serological assays could be unproductive during this period due to time lapse for seroconversion spanning 3-4 weeks after the onset of clinical infection (Howe et al., 2009; Niemczuk et al., 2014). Vincent et al. (2015) also reported that early serum samples from acute Q fever patients are infectious and valuable source of viable C. burnetii. Apart from serological investiagtion, C. burnetii-specific PCR of serum samples can be an effective tool to diagnose Coxiellosis in early acute phase, but contradictory sensitivities have been reported (Fournier and Raoult, 2003; Schneeberger et al., 2010). Similarly, an in-house developed real-time qPCR targeting multiple copy IS111 transposase gene was used for genomic detection of C. burnetii DNA in tick pools. Real-time qPCR detected C. burnetii DNA in 31.03% (95% CI: 15.3-50.8) and 7.69% (95% CI: 0.9-25.1) of sheep and goats tick pools, respectively. The overall prevalence of C. burnetii DNA in these 55 tick pools was 20%. Univariate analysis revealed a significant (p=0.031, df=1, χ2=4.668) difference in prevalence of C. burnetii DNA in tick pools collected from sheep and goats. Results obtained from the study of Knobel et al. (2013) also revealed C. burnetii DNA prevalence of 2.5 and 20% in ticks feeding on cattle and dogs, respectively. A higher prevalence (25%) of C. burnetii DNA was found in questing ticks collected from domestic animals of Ethopia (Kumsa et al., 2015). While, lower prevalence (0.2%) of C. burnetii DNA was recorded in questing I. ricinus ticks and ticks feeding on wildlife, pets and domestic animals by using a multiplex Q-PCR (Sprong?et al., 2012). Though major pioneering studies have focused on prevalence of C. burnetii in ticks, but still the role of ticks in Q fever epidemiology remains uncertain. Even the highly virulent, Nine Mile, reference strain of C. burnetii was isolated from a guinea pig upon which Dermacentor andersoni ticks had fed (McDade, 1990; Duron et al., 2015). Moreover, several previous studies based on microscopic morphological observations of C. burnetii in ticks revealed that more than 40 tick species carry this pathogen (Babudieri, 1959). The occasional reports of unexpectedly high prevalence of C. burnetii DNA in ticks reflects its role as a vector for Q fever transmission (Duron et al., 2015). ConclusionsBased on findings of the present study, following conclusions can be drawn:This study indicates the presence of anti-C. burnetii antibodies in small ruminants kept at government livestock farms. This epidemiological survey and some previous studies carried out for the detection of C. burnetii infection in different species at different geographical regions of Pakistan indicated that Coxiellosis is endemic in the country. Variables like type and locality of animal farm, presence of ticks, breed, reproductive disorders, animal health status, lactational and reproductive status of the animal have significant role in seropositivity for C. burnetii. Since, there is very limited awareness of the disease (based on information collected during the current survey) among veterinary, para-veterinary and farm workers at the studied livestock farms, an awareness campaign should be launched to educate livestock farmers and professionals about proper preventive and control of Coxiellosis. This survey highlights the importance of clinical herd health management to avoid economical losses in small ruminants as a result of underdiagnosis and underreporting of the disease.Authors hope publication of this study will draw attention of the concerned authorities. Further population based studies at district level are required to clearly understand the epidemiology of the disease. Chapter-6SUMMARYCoxiellosis is one of the major neglected zoonotic diseases, resulting in huge economical losses to the livestock industry in the form of reproductive losses through late term abortion, premature delivery, stillbirth and delivery of weak offsprings. Thus, this disease adversely affects the productive and reproductive capabilities of domestic animals, especially sheep and goats. This study was planned to investigate the epidemiology of Coxiellosis in small ruminants at 9 government livestock farms of Punjab province of Pakistan. Attempts were also made to see if there is any association between seropostivity against C. burnetii antibodies and various factors such as age, sex, parity, breed, species, farms, locational status, reproductive status, lactation, body condition, tick infestation and reproductive disorders. Individual animal and general farm management data were collected using a structured questionnaire. Sample size was estimated using Thrusfield formula with an expected prevalence of 50%, confidence interval (CI) of 95% and 5% desired absolute precision. Accordingly, 1000 blood samples (500 each from sheep and goats) were collected from nine farms. Approximately 10 ml of blood was collected from the jugular vein of each animal using disposable needles and vacutainer tubes. Each tube was properly labeled for its identification and informations about its origin using English alphabets and Arabic numbers. Blood samples collected from small ruminants were placed overnight in inclined position and then centrifugion was done at 1000 rpm for 5 min. for proper separation of serum. Serum was then shifted to disposable screw caped plastic bottles and stored in a deep freezer at -20 ?C till its use for serological analysis. The diagnostic work (Indirect-ELISA and qPCR) was carried out at FLI, Jena, Germany. An Indirect-ELISA was conducted to analyze serum samples for seropositivity against C. burnetii infection. Serological analysis indicated a slightly higher prevalence of C. burnetii antibodies in sheep 15.6% (95% CI: 12.5-19.1) as compare to goats 15% (95% CI: 12.0-18.4). However, the difference in prevalence of disease between the two species was statistically non-significant. A significant association was found between seropositivity against C. burnetii antibodies and different variables like farm (p=0.000, df=8, χ2=141.869), district (p=0.000, df=6, χ2=49.689), breed (p=0.000, df=9, χ2=60.954), lactational status (p= 0.000, df=1, χ2=24.691), reproductive status (p= 0.008, df=1, χ2=7.023), ticks infestation (p=0.000, df=1, χ2=301.914), body condition (p=0.000, df=3, χ2=124.868) and reproductive disorders (p=0.000, df=4, χ2=133.984). However, a non-significant association was found between Coxiellosis and variables like age (p=0.063, df=3, χ2=7.281), parity (p=0.838, df=2, χ2=0.353) and sex (p=0.302, df=1, χ2=1.064) of animal. After serological analysis, a real-time qPCR (FLI standard Jena, LA 190 qPCR, MXPro-MX3000P), based on single copy icd gene, was used for genomic detection of C. burnetii in pools of seropositive and suspected (29 pools) serum samples. The Roche kit was used (Roche kit, version 20, Roche Diagnostics GmbH, Germany) for DNA extraction from serum. Out of 29 sera pools, 14 pools belonged to goats, while 15 pools were from sheep. The qPCR-based analysis did not show the presence of C. burnetii DNA in any of seropositive pools, however, the DNA was detected in 5 pools of suspected serum samples. The prevalence of C. burnetii DNA was recorded as 21.4% (95% CI: 4.7-50.8) in goat serum pools and 13.3% (95% CI: 1.7-40.5) in sheep serum pools. Univariate analysis showed absence of any association between detection of C. burnetii DNA in pooled seum samples and species-wise prevalence of the disease. Additionally, 55 tick pools were investigated through real-time qPCR (FLI standard Jena, LA 190 icd qPCR, MXPro-MX3000P), targeting multiple copy IS1111 transposase gene, for genomic detection of C. burnetii. Prior to DNA extraction, ticks were surface decontaminated by serial passages in 10% and 70% alcohol, and rinsed in sterile water. As the ticks had been preserved in alcohol for a long period of time, they were triturated in vials containing 1ml PBS (Phosphate Buffer Saline) with 3-4 beads per tube, prior to the addition of TLB (Tissue Lysis Buffer). Then these vials were centrifuged at 1000g for 10 min. After centrifugation, PBS was removed and sediment was collected which was then triturated with 200 μl TLB. DNA from ticks was extracted through an in-house developed highly pure PCR template kit (FLI standard Jena, IBIZ, AGr. 180, DNA-Isolation high pure kit, version 2). A total of 163 ticks were investigated through qPCR for the presence of C. burnetii DNA. Out of 163 ticks, 85 ticks were collected from sheep, while 78 were collected from goats. All these 163 ticks were merged into 55 pools, including 29 pools for sheep ticks, while 26 pools for goat ticks. Each pool was comprised of 3 ticks (with only pool 29 containing 1 tick). The qPCR performed on these tick pools detected C. burnetii DNA in 31.03% (95% CI: 15.3-50.8) and 7.69% (95% CI: 0.9-25.1) of sheep and goat tick pools, respectively. While an overall prevalence of C. burnetii DNA in these 55 tick pools was recorded as 20%. Univariate analysis revealed a significant (p=0.031, df=1, χ2=4.668) difference in prevalence of C. burnetii DNA in tick pools collected from sheep and goats. This study highlights the importance of Coxiellosis in small ruminants maintained at government livestock farms of Punjab. Results obtained from this survey showed that C. burnetii infection is endemic in small ruminant population of Pakistan. This study also revealed an active role of ticks as a vector and reservoir of C. burnetii. An awareness campaign should be launched to educate the livestock farmers and livestock professionals about C. burnetii infection, and control of ticks in animals. Further in-depth studies are needed to explore its epidemiology of Q fever more precisely in different species of animals, humans and ticks.LITERATURE CITEDAbdel-Moein KA and Hamza DA, 2017. The burden of Coxiella burnetii among aborted dairy animals in Egypt and its public health implications. Acta Trop 166:92-95.Abed J, Salih AA and Abd-ul-husien A, 2010. Seroprevalence of Coxiella burnetii among cows and sheep in Thi-Qar province Iraq. AL-Qadisiya J Vet Med Sci 9(2):26-30.Abushahba MFN, Abdelbaset AE, Rawy MS, et al., 2017. Cross?sectional study for determining the prevalence of Q fever in small ruminant and humans at El Minya Governorate, Egypt. BMC Res Notes 10:538. DOI 10.1186/s13104-017-2868-2.Adesiyun A, Dookeran S, Stewart-Johnson A, et al., 2011. Frequency of seropositivity for Coxiella burnetii immunoglobulins in livestock and abattoir workers in Trinidad. New Microbiol 34:219-224. Afzal M and Sakkir M, 1994. Survey of antibodies against various infectious disease agents in racing camels in Abu Dhabi, United Arab Emirates. Rev Sci Tech Off Int Epiz 13(3):787-792. Agerholm JS, 2013. Coxiella burnetii associated reproductive disorders in domestic animals- a critical review. Acta Vet Scand 55:13-21.Ahmed IP, 1987. A serological investigation of Q fever in Pakistan. J Pak Med Assoc 37:126-131.Aitken ID, Bogel K, Cracea E, et al., 1987. Q fever in Europe: Current aspects of aetiology, epidemiology, human infection, diagnosis and therapy. Infection 15:323-327.Akbarian Z, Ziay G, Schauwers W, et al., 2015. Brucellosis and Coxiella burnetii infection in householders and their animals in secure villages in Herat province, Afghanistan: A cross-sectional study. PLoS Negl Trop Dis 9(10):1-17.Aldomy FFM, Wilsmore AJ and Safi SH, 1998. Q fever and abortion in sheep and goats in Jordan. Pak Vet J 18(1):43-45.Alvesa J, Almeidaa F, Duroa R, et al., 2017. Presentation and diagnosis of acute Q fever in Portugal- A case series. IDCases 7:34-37.Amitai Z, Bromberg M, Bernstein M, et al., 2010. A large Q fever outbreak in an urban school in Central Israel. Clin Infect Dis 50(11):1433-1438. Anasta?cio S, Tavares N, Carolino N, et al., 2013. Serological evidence of exposure to Coxiella burnetii in sheep and goats in central Portugal. Vet Microbiol 167:500-505.Anderson A, Boyer T, Garvey A, et al., 2013. Prevention and control of Coxiella burnetii infection among humans and animals: Guidance for a coordinated public health and animal health response. National Association of State Public Health Veterinarians (NASPHV), United States of America. pp:1-30.Anderson AD, Kruszon-Moran D, Loftis AD, et al., 2009. Seroprevalence of Q fever in the United States, 2003–2004. Am J Trop Med Hyg 81(4):691-694.Angelakis E and Raoult D, 2010. Q fever.Vet Microbiol 140:297-309.Angelakis E and Raoult D, 2011. Emergence of Q fever. Iran J Pub Health 40:1-18.Angelakis E, Munasinghe A, Yaddehige I, et al., 2012. Short report: detection of Rickettsioses and Q fever in Sri Lanka. Am J Trop Med Hyg 86(4):711-712.Anonymous, 2013. Diagnosis and management of Q fever. Centers for Disease Control and Prevention, United States. Morbidity and Mortality Weekly Report (MMWR) 62(3):1-29. Anonymous, 2017. Economic Survey of Pakistan (2016-17). Economic Advisors Wing, Finance Division, Government of Pakistan, Islamabad, Pakistan.Arserim NB, Yesilmen S, Tel OY, et al., 2011. Seroprevalance of Coxiellosis in cows, sheep, goats and humans in Diyarbakir region of Turkey. Afr J Microbiol Res 5:2041-2043. Asadi J, Kafi K and Khalili M, 2013. Seroprevalence of Q fever in sheep and goat flocks with a history of abortion in Iran between 2011 and 2012. Vet Ital 49:163-168.Aslanova A, Bagirov S, Bakhishova S,?et al., 2009. Identification of an endemic infection: Q fever in rural Azerbaijan. National Assembly of State Animal Health Officials. Proceedings of community-acquired bacterial infections including STDs and Mycobacteria, PP:1-30.Astobiza?I, Tilburg JJHC, Pinero A, et al., 2012. Genotyping of Coxiella burnetii from domestic ruminants in Northern Spain. BMC Vet Res 8(1):241-248.Ayaz M,?Bari A and Humayun A, 1993. Coxiellosis in man and animals in northern parts of Pakistan. Proc Pak Cong Zool 13:425-431.Babudieri B, 1959. Q fever: a zoonosis. Adv Vet Sci 5:81-154.Baziaka F, Karaiskos I, Galani L, et al., 2014. Large vessel vasculitis in a patient with acute Q-fever: A case report. IDCases 1:56-59.Bellini C, Magouras I, Chapuis-Taillard C, et al., 2014. Q fever outbreak in the terraced vineyards of Lavaux, Switzerland. New Microbes New Infect 2(4):93-99.Benkirane A, Essamkaoui S, Idrissi AE, et al., 2015. A sero-survey of major infectious causes of abortion in small ruminant in Morocco. Vet Ital 51(1):25-30.Bennett BW, 2013.The challenge of North Korean biological weapons. Published by the RAND Corporation 1776 Main Street, P.O. Box 2138, Santa Monica, CA 90407-2138 1200 South Hayes Street, Arlington, VA 22202-5050 4570 Fifth Avenue, Suite 600, Pittsburgh, PA 15213-2665 RAND. URL: M, Laroucau K and Rodolakis A, 2000. The detection of Coxiella burnetii from ovine genital swabs, milk and fecal samples by the use of a single touchdown polymerase chain reaction. Vet Microbiol 72:285-293. Bielawska-Dro? zd A, Cies?lik P, Mirski T, et al., 2014. Prevalence of Coxiella burnetii in environmental samples collected from cattle farms in Eastern and Central Poland (2011–2012). Vet Microbiol 174:600-606.Blacksell SD, Kantipong P, Watthanaworawit W, et al., 2015. Underrecognized arthropod-borne and zoonotic pathogens in Northern and Northwestern Thailand: Serological evidence and opportunities for awareness. Vector Borne Zoonotic Dis 15(5):285-290.Blair PJ, Schoeler GB, Moron C, et al., 2004. Evidence of rickettsial and leptospira infections in Andean Northern Peru. Am J Trop Med Hyg 70(4):357-363. Boden K, Brasche S, Straube E, et al., 2014. Specific risk factors for contracting Q fever: Lessons from the outbreak Jena. ?Int J Hyg Environ?Health 217:110-115.Boni M, Davoust B, Tissot-Dupontb H, et al., 1998. Survey of seroprevalence of Q fever in dogs in the southeast of France, French Guyana, Martinique, Senegal and the Ivory Coast. Vet Microbiol 64(1):1-5.Bontje DM, Backer JA, Hogerwerf L, et al., 2016. Analysis of Q fever in Dutch dairy goat herds and assessment of control measures by means of a transmission model. Prev Vet Med 123:71-89.Brookea RJ, Mutters NT, Péter O, et al., 2015. Exposure to low doses of Coxiella burnetii caused high illness attack rates: Insights from combining human challenge and outbreak data. Epidemics 11:1-6.Brouqui P, Rolain JM, Foucault C, et al., 2005. Short report: Q fever and Plasmodium falciparum malaria co-infection in a patient returning from the Comoros Archipelago. Am J Trop Med Hyg 73(6):1028-1030.Burnet FM and Freeman M, 1937. Experimental studies on the virus of “Q” fever. Med J Aust 2:299-305.Cameron A, 1999. Survey Toolbox for Livestcok Diseases: A Practical Manual and Software Package for Active Surveillance in Developing Countries. Australian Center for Internationl Agricultural Research.Cantas H, Muwonge A, Sareyyupoglu B, et al., 2011. Q fever abortions in ruminants and associated on-farm risk factors in northern Cyprus. BMC Vet Res 7(13):2-7.Carbonero A, Guzmán LT, Montano K, et al., 2015. Coxiella burnetii seroprevalence and associated risk factors in dairy and mixed cattle farms from Ecuador: Prev Vet Med 118:427-435.?etinkol Y, Enginyurt O, ?elebi B, et al., 2017. Investigation of zoonotic infections in risk groups in Ordu University Hospital, Turkey. Niger J Clin Pract 20:6-11. Chakrabartty A, Bhattacharjee PK, Sarker RR, et al., 2016. Prevalence of Coxiella burnetii infection in cattle, black Bengal goats and ticks in Bangladesh. Bangl J Vet Med 14(1):65-68.Chan JFW, Tse H, To KKW, et al., 2010. Q fever: under-diagnosed in Hong Kong? Hong Kong Med J 16(1):56-58. Chieng D, Janssen J, Benson S, et al., 2016. 18-FDG PET/ CT scan in the diagnosis and follow-up of fhronic Q fever aortic valve endocarditis. Heart Lung Circ, 25:e17–e20. T and Tylewska-Wierzbanowska S, 2013. Q fever outbreaks in Poland during 2005–2011. Med Sci Monit 19:1073-1079.Cicuttin GL, Degiuseppe JI, Mamianetti A, et al., 2015. Serological evidence of Rickettsia and Coxiella burnetii in humans of Buenos Aires, Argentina. Comp Immunol Microbiol Infect Dis 43:57-60. Cong W, Meng QF, Shan XF, et al., 2015. Coxiella burnetii (Q fever) infection in farmed ruminants in three northeastern provinces and Inner Mongolia autonomous region, China. Vector Borne Zoonotic Dis 15:512-514.Cooper A, Stephens J, Ketheesan K, et al., 2013. Detection of Coxiella burnetii DNA in wildlife and ticks in Northern Queensland, Australia. Vector Borne Zoonotic Dis 13(1):12-16.Cooper A, Hedlefs R, McGowan M, et al., 2011. Serological evidence of Coxiella burnetii infection in beef cattle in Queensland. Aust Vet J 89:260-264. Crump JA, Morrissey AB, Nicholson WL, et al., 2013. Etiology of severe non-malaria febrile illness in Northern Tanzania: A prospective cohort study. PLoS Negl Trop Dis 7(7):1-8. Cumbassá A, Barahonaa MJ, Cunhaa MV, et al., 2015. Coxiella burnetii DNA detected in domestic ruminants and wildlife from Portugal. Vet Microbiol 180:136-141. Cutler SJ, Bouzid M and Cutler RR, 2007. Q fever. J Infect, 54:313-318.Das DP, Malik SVS, Rawool DB, et al., 2014. Isolation of Coxiella burnetii from bovines with history of reproductive disorders in India and phylogenetic inference based on the partial sequencing of IS1111 element. Infect?Genet?Evol?22:67-71. de Bruin A, van Alphen PTW, van der Plaats RQJ, et al., 2012. Molecular typing of Coxiella burnetii from animal and environmental matrices during Q fever epidemics in the Netherlands. BMC Vet Res 8:165-174.Dean AS, Bonfoh B, Kulo AE, et al., 2013. Epidemiology of brucellosis and Q fever is linked with human and animal populations in Northern Togo. PLoS ONE 8(8):1-8. DePuy W, Benka V, Massey A, et al., 2014. Q fever risk across a dynamic, heterogeneous landscape in Laikipia County, Kenya. Ecohealth 11(3):429-433.DeRooij MMT, Schimmer B, Versteeg B, et al., 2012. Risk factors of Coxiella burnetii (Q fever) seropositivity in veterinary medicine students. PLoS One 7(2):e32108.Derrick EH, 1937. “Q” fever, new fever entity: clinical features, diagnosis and laboratory investigation. Med J Aust 2:281-299.Derrick EH, 1939. Rickettsia burneti: cause of Q fever. Med J Aust 1:14.Dorko E, Pilip?inec E, Rimárová K, et al., 2010. Serological study of Q fever in sheep in the Territory of Eastern Slovakia. Ann Agric Environ Med 17:323-325.Douangngeun B, Theppangna W, Soukvilay V, et al., 2016. Seroprevalence of Q fever, Brucellosis, and Bluetongue in selected provinces in Lao People’s Democratic Republic. Am J Trop Med Hyg 95(3):558-561. Drozd AB, Cieslik P, Mirski T, et al., 2014. Prevalence of Coxiella burnetii in environmental samples collected from cattle farms in eastern and central Poland (2011–2012). Vet Microbiol 174:600-606.Duron O, Sidi-Boumedine K, Rousset E, et al., 2015. The importance of ticks in Q fever transmission: what has (and has hot) been demonstrated? Trends Parasitol 31(11):536-552. Eldin C, Mahamat A, Demar M, et al., 2014. Review article: Q fever in French Guiana: Am J Trop Med Hyg 91(4):771-776. El-Mahallawy HS, Kelly P, Zhang J, et al., 2016. Serological and molecular evidence of Coxiella burnetii in samples from humans and animals in China. Ann Agric Environ Med 23(1):87-91.European Centre for Disease Prevention and Control, 2011. Risk assessment on Q fever. http:// ecdc.europa.eu/en/publications/Publications/1005_TER_Risk_Assessment_Q fever.pdf.European Food Safety Authority, 2010. European Food Safety Authority (EFSA). Scientific opinion on Q fever. Panel on Animal Health and Welfare. EFSA J. 8(5):1595 [114 pp.].Ezatkhah M, Alimolaei M, Khalili M, et al., 2015. Seroepidemiological study of Q fever in small ruminant from Southeast Iran. J Infect Public Health 8:170-176. Fedorov EI, 1983. Q fever in the?Ukraine. Zh Mikrobiol Epidemiol Immunobiol 7:109-112.Fernandez?Aguilar X, Cabezón O, Colom?Cadena A, et al., 2016. Serological survey of Coxiella burnetii at the wildlife–livestock interface in the Eastern Pyrenees, Spain. Acta Vet Scand 58(26): 2-5.Ferraz RV, Andrade M, Silva F, et al., 2016. Chronic Q fever: A missed prosthetic valve endocarditis possibly for years. IDCases 6:55-57.Fournier PE and Raoult D, 2003. Comparison of PCR and serology assays for early diagnosis of acute Q fever. J Clin Microbiol 41:5094-5098.Freick M, Enbergs H, Walraph J, et al., 2016. Coxiella burnetii: Serological reactions and bacterial shedding in primiparous dairy cows in an endemically infected herd-impact on milk yield and fertility. Reprod Dom Anim 52:160-169.Gababedian GA, Djanian AY and Johnston EA, 1956. Q fever in Lebanon (Middle East); the presence of complement-fixing antibodies in serum samples obtained from residents of Lebanon. Am J Hyg 63:308-312. Ganter M, 2015. Zoonotic risks from small ruminant. Vet Microbiol 181(1-2):53-65. García-Pérez AL, Astobiza I, Barandika JF, et al., 2009. Investigation of Coxiella burnetii occurrence in dairy sheep flocks by bulk-tank milk analysis and antibody level determination. J Dairy Sci 92:1581-1584. Georgiev M, Afonso A, Neubauer H, et al., 2013. Q fever in humans and farm animals in four European countries, 1982 and 2010. Euro Surveil 18(8):1-13.Giangaspero M, Osawa T, Bonfini B, et al., 2012. Serological screening of Coxiella burnetii (Q fever) and Brucella spp. in sheep flocks in the northern prefectures of Japan in 2007. Vet Ital 48(4):357-365.Godinhoa I, Nogueira EL, Santos CM, et al., 2015. Chronic Q fever in a renal transplant recipient: A case report. Transplant Proc 47:1045-1047.Guatteo R, Beaudeau F, Joly A, et al., 2007. Coxiella burnetii shedding by dairy cows. Vet Res 38:849-860.Gumi B, Firdessa R, Yamuah L, et al., 2013. Seroprevalence of brucellosis and Q fever in Southeast Ethiopian pastoral livestock. J Vet Sci Med Diagn 2(1):1-11.Hadush A, Kandi V and Pal M, 2016. Epidemiology and public health implications of Q fever. Perspect Med Res 4(3):42-46.Hagenaars JCJP, Wever PC, van Petersen AS, et al., 2014. Estimated prevalence of chronic Q fever among Coxiella burnetii seropositive patients with an abdominal aortic/iliac aneurysm or aorto-iliac reconstruction after a large Dutch Q fever outbreak. J Infect 69:154-160.Hamzic S, Beslagic E and Zvizdi? S, 2006. Serotesting of human Q fever distribution in Bosnia and Herzegovina. ?Ann NY Acad Sci?1078(1):133-136.Hatchette T, Campbell N, Whitney H, et al., 2002. Seroprevalence of Coxiella burnetii in selected populations of domestic ruminants in Newfoundland. Can Vet J 43:363-364.Hazlett?M, Cai H, DeLay J, et al., 2010. The AHSI small ruminant abortion project- an update. AHL?Newsletter,?University?of?Guelph, 14(2):9-18.?Herremans T, Hogema BM, Nabuurs M, et al., 2013. Comparison of the performance of IFA, CFA and ELISA assays for the serodiagnosis of acute Q fever by quality assessment. Diagn Microbiol Infect Dis 75:16-21.Horigan MW, Bell MM, Pollard TR, et al., 2011. Q fever diagnosis in domestic ruminants: comparison between complement fixation and commercial enzyme-linked immunosorbent assays. J Vet Diagn Invest 23:924-931.Howe GB, Loveless BM, Norwood D, et al., 2009. Real-time PCR for the early detection and quantification of Coxiella burnetii as an alternative to the murine bioassay. Mol Cell Probes 23:127-131.Hussien MO, Enan KA, Alfaki SH, et al., 2017. Seroprevalence of Coxiella burnetii in dairy cattle and camel in Sudan. Int J Infect 4(3):e42945: doi: 10.17795/iji.42945.Ignatovich V, Penkina G and Ummova N, 2003. Seroimmunological monitoring of several species of rickettsiaceae and bartonellaceae circulating in the Moscow region. Ann NY Acad Sci 990:419-423.Isken LD, Kraaij-Dirkzwager M, Bondt PEV, et al., 2013. Implementation of a Q fever vaccination program for high-risk patients in the Netherlands. Vaccine 31:2617-2622.Jones?RM,? Twome DF, Hannon S,?et al., 2010. Detection of Coxiella? burnetii ?in ?placenta? and?abortion?samples?from?British?ruminants?using real-time PCR. Vet? Rec, 167(25): 965‐967. Jung BY, Seo MG, Lee SH, et al., 2014. Molecular and serological detection of Coxiella burnetii in native Korean goats (Capra hircus coreanae). Vet Microbiol 173:152-155.Kaabia N, Rolain JM, Khalifa M, et al., 2006. Serologic study of Rickettsioses among acute febrile patients in Central Tunisia. Ann NY Acad Sci 1078:176-179.Kampen AH, Hopp P, Gr?neng GM, et al., 2012. No indication of Coxiella burnetii infection in Norwegian farmed ruminants. BMC Vet Res 8(59):2-6.Kanouté YB, Gragnon BG, Schindler C, et al., 2017. Epidemiology of brucellosis, Q fever and Rift Valley fever at the human and livestock interface in northern C?te d’Ivoire. Acta Trop 165:66-75.Kaplan MM and Bertagna P, 1955. The geographical distribution of Q fever. Bull World Health Organ 13(5):829-860.Keijmel SP, Saxe J, van der Meer JWM, et al., 2015. A comparison of patients with Q fever fatigue syndrome and patients with chronic fatigue syndrome with a focus on inflammatory markers and possible fatigue perpetuating cognitions and behavior. J Psychosom Res 79:295-302.Kelly PJ, Matthewman LA, Mason PR, et al., 1993. Q?fever?in Zimbabwe. A review of the disease and the results of a serosurvey of humans, cattle, goats and dogs. S Afr Med J 83(1):21-25.Kersh GJ, Fitzpatrick KA, Self JS, et al., 2013. Presence and persistence of Coxiella burnetii in the environments of goat farms associated with a Q fever outbreak. Appl Environ Microbiol 79:1697-1703.Khaled H, Sidi-Boumedine K, Merdja S, et al., 2016. Serological and molecular evidence of Q fever among small ruminant flocks in Algeria. Comp Immunol Microbiol Infect Dis 47:19-25. Khalili M, Diali HG, Mirza HN, et al., 2015. Detection of Coxiella burnetii by PCR in bulk tank milk samples from dairy caprine herds in southeast of Iran. Asian Pac J Trop Dis 5(2):119-122.Khattak MS and Ali S, 2015. Assessment of temperature and rainfall trends in Punjab province of Pakistan for the period 1961-2014. J Himalayan Earth Sci (JHES) 48(2):42-61.K?l?? A, Kalender H, Ko? O, et al., 2016. Molecular investigation of Coxiella burnetii infections in aborted sheep in eastern Turkey. Iran J Vet Res 17(1):41-44.Ki-Zerbo GA, Tall F, Nagalo K, et al., 2000. Seroprevalence des rickettsioses et de la fievre Q chez les patients febriles a I’hopital de Bobo-Dioulasso (Burkina Faso). Med Mal Infect 30:270-274.Klaasen M, Roest HJ, Hoek W, et al., 2014. Coxiella burnetii Seroprevalence in small ruminant in the Gambia. PLoS ONE 9(1):1-6. Klaassen CH, Nabuurs-Franssen MH, Tilburg JI, et al., 2009. Multigenotype Q fever outbreak, The Netherlands. Emerg Infect Dis 15:613-614.Knobel DL, Maina AN, Cutler SJ, et al., 2013. Coxiella burnetii in humans, domestic ruminants and ticks in rural western Kenya. Am J Trop Med Hyg 88:513-518.Kobbe R, Kramme S, Kreuels B, et al., 2008. Q fever in young children, Ghana. Emerg Infect Dis 14(2):344-345. Koch A, Svendsen CB, Christensen JJ, et al., 2010. Q fever in Greenland. Emerg Infect Dis 16(3):511-513.Kovacova E and Kazar J, 2002. Q fever-still a query and underestimated infectious disease. Acta Virol 46:193-210.Kreizinger Z, Szeredi L, Bacsadi A, et al., 2015. Occurrence of Coxiella burnetii and Chlamydiales species in abortions of domestic ruminants and in wild ruminants in Hungary, Central Europe. J Vet Diagn Inves 27(2):206-210.Kshash QH, 2012. Prevalence of Q-fever in small ruminant in Al- Qassim city. Basrah J Vet Res 11:342-348.Kuley R, Smith HE, Frangoulidis D, et al., 2015. Cell-free propagation of Coxiella burnetii does not affect its relative virulence. PLoS ONE 10(3):1-16. Kumsa B, Socolovschi C, Almeras L, et al., 2015. Occurrence and genotyping of Coxiella burnetii in ixodid ticks in Oromia, Ethiopia. Am J Trop Med Hyg 93(5):1074-1081.Lacasta D, Ferrer LM, Ramos JJ, et al., 2015. Vaccination schedules in small ruminant farms. Vet Microbiol 181(1-2):34-46. Lai CH, Sun W, Lee CH, et al., 2017. The epidemiology and characteristics of Q fever and co-infections with scrub typhus, murine typhus or leptospirosis in Taiwan: A nationwide database study. Zoonoses Public Health 64(7):517-526.Lambton SL, Smith RP, Gillard K, et al., 2016. Serological survey using ELISA to determine the prevalence of Coxiella burnetii infection (Q fever) in sheep and goats in Great Britain. Epidemiol Infect 144:19-24. Lang GH, 1990. Coxiellosis (Q fever) in animals. In: Marrie TJ (ed.) Q fever. CRC, Boca Raton, FL, pp:23-48.Lau C, Musso D, Fournier P, et al., 2016. Absence of serological evidence of Rickettsia spp., Bartonella spp., Ehrlichia spp. and Coxiella burnetii infections in American Samoa. Ticks Tick Borne Dis 7:703-705. Lau?evi? D, 2011. Prevalence of Coxiellae burnetii antibodies in sheep in the territory of Montenegro. ?Acta veterinaria?51(2-3).Leuken JPGV, Swart AN, Brandsma J, et al., 2016. Human Q fever incidence is associated to spatiotemporal environmental conditions. One Health 2:77-87.Lucchese L, Capello K, Barberio A, et al., 2015. IFAT and ELISA phase I/ phase II as tools for the identification of Q fever chronic milk shedders in cattle. Vet Microbiol 179:102-108.Lyoo K, Kim D, Jang HG, et al., 2017. Prevalence of antibodies against Coxiella burnetii in Korean native cattle, dairy cattle and dogs in South Korea. Vector Borne Zoonotic Dis 17(3):213-216. Lyytika¨inen O, Ziese T and Schwartla¨nder B, 1998. An outbreak of sheep-associated Q fever in a rural community in Germany. Eur J Epidemiol 14:193-199.Magouras I, Hunninghaus J, Scherrer S, et al., 2017. Coxiella burnetii infections in small ruminant and humans in Switzerland. Transbound Emerg Dis 64:204-212.Mares-Guia MAMM, Rozental T, Guterres A, et al., 2014. Molecular identification of the agent of Q fever – Coxiella burnetii – in domestic animals in State of Rio de Janeiro, Brazil. Rev Soc Bras Med Trop 47(2):231-234. Martinov S, 2007. Contemporary state of the problem Q fever in Bulgaria. Biotechnol Biotechnol Equip 21(3):353-361.Mattar S, Contreras V, González M, et al., 2014. Infection by Coxiella burnetii in a patient from a rural area of Monteria, Colombia. Rev Salud Pública 16(6):958-961.Maurin M and Raoult D, 1999. Q fever. Clin Microbiol Rev 12:518-553. McCaughey C, Murray LJ, McKenna JP, et al., 2010. Coxiella burnetii (Q fever) seroprevalence in cattle. Epidemiol Infect 138:21-27.McDade JE, 1990. Historical aspects of Q fever. In Q FEVER-The Disease (Marrie, T.J., Ed.). CRC Press 1:5-22Meadows S, Jones-Bitton A, McEwen S, et al., 2015. Coxiella burnetii seropositivity and associated risk factors in goats in Ontario, Canada. Prev Vet Med?121(3-4):199-205. Meadows S, Jones-Bitton A, McEwen SA, et al., 2016. Coxiella burnetii (Q fever) seropositivity and associated risk factors in sheep and goat farm workers in Ontario, Canada. Vector Borne Zoonotic Dis 16(10):643-649.Meadows S, Jones-Bitton A, McEwen S, et al., 2015. Coxiella burnetii seropositivity and associated risk factors in sheep in Ontario, Canada. Prev Vet Med?122(1-2):129-134. Mediannikov O, Fenollar F, Socolovschi C, et al., 2010. Coxiella burnetii in humans and ticks in rural Senegal. PLoS Negl Trop Dis 4(4):1-8. Medi? S, Kaluski DN, ?eguljev Z, et al., 2012. Q fever outbreak in the village of No?aj, Srem County, Vojvodina province Serbia, January to February 2012. Euro Surveill 17(15):1-4.Meekelenkamp JC, Schneeberger PM, Wever PC, et al., 2012. Comparison of ELISA and indirect immunofluorescent antibody assay detecting Coxiella burnetii IgM phase II for the diagnosis of acute Q fever. Eur J Clin Microbiol Infect Dis 31:1267-1270.Mertens K and Samuel JE, 2007. Bacteriology of Coxiella. In: Raoult D and Parola P (eds.) Rickettsial diseases, London. Informa Healthcare, pp. 257–270.Mertens K, Gerlach C, Neubauer H, et al., 2017. Q fever-An update. Curr Clin Micro Rpt 4:61-70.Miceli MH, Veryser AK, Anderson AD, et al., 2010. A case of person-to-person transmission of Q fever from an active duty serviceman to his spouse. Vector Borne Zoonotic Dis 10:539-541.Million M and Raoult D, 2015. Recent advances in the study of Q fever epidemiology, diagnosis and management. J Infect 71:S2-S9.Miorini T, Brosch R, Buchrieser C,?et al., 1988. Further serological investigations in humans and domestic animals on the?Cape Verde?Islands (Q-fever, brucellosis, listeriosis, shigellosis, campylobacteriosis, yersiniosis, toxoplasmosis and chlamydia of PLT-group). Geogr Med Suppl?1:19-31.Mohammed OB, Jarelnabi AA, Aljumaah RS, et al., 2014. Coxiella burnetii, the causative agent of Q fever in Saudi Arabia: molecular detection from camel and other domestic livestock. Asian Pac J Trop Dis (2014):715-719.Mori M, Mertens K, Cutler SJ, et al., 2017. Critical aspects for detection of Coxiella burnetii. Vector-Borne Zoonotic Dis 17(1):33-41.Mori M, Boarbi S, Michel P, et al., 2013. In vitro and in vivo infectious potential of Coxiella burnetii: A study on Belgian livestock isolates. PLoS ONE 8(6):e67622. Muema J, Thumbi SM, Obonyo M, et al., 2017. Seroprevalence and factors associated with Coxiella burnetii infection in small ruminant in Baringo County, Kenya. Zoonoses Public Health 64:e31–e43.Muleme M, Stenos J, Vincent G, et al., 2016. Bayesian validation of the indirect immunofluorescence assay and its superiority to the enzyme-linked immunosorbent assay and the complement fixation test for detecting antibodies against Coxiella burnetii in goat serum. Clin Vaccine Immunol 23:507-514.Musso D, Broult J, Parola P, et al. et al., 2014. Absence of antibodies to Rickettsia spp., Bartonella spp., Ehrlichia spp. and Coxiella burnetii in Tahiti, French Polynesia. BMC Infect Dis 14(255):1-4. Nakoune E, Debaere O, Koumanda-Kotogne F, et al., 2004. Serological surveillance of brucellosis and Q fever in cattle in the Central African Republic. Acta Trop 92:147-151.Niemczuk K, Szymanska-Czerwinska M, Smietanka K, et al., 2014. Comparison of diagnostic potential of serological, molecular and cell culture methods for detection of Q fever in ruminants. Vet Microbiol 171:147-152. Noden BH, Tshavuka FI, Colf BE, et al., 2014. Exposure and risk factors to Coxiella burnetii, spotted fever group and Typhus group Rickettsiae, and Bartonella henselae among volunteer blood donors in Namibia. PLOS ONE 9(9):1-8. e108674. Ohlson A, Malmsten J, Frossling J, et al., 2014. Surveys on Coxiella burnetii infections in Swedish cattle, sheep, goats and moose. Acta Vet Scand 56:1-9.OIE, 2015. (Office International des Epizootics, Paris, France), OIE Terrestrial Manual 2015, Chapter 1.2.16.-Q fever: pp:1-15.Omsland A and Heinzen RA, 2011. Life on the outside: The rescue of Coxiella burnetii from its host cell. Annu Rev Microbiol 65:111-128.Omsland A, Cockrell DC, Fischer ER, et al., 2008. Sustained axenic metabolic activity by the obligate intracellular bacterium Coxiella burnetii. J Bacteriol 190:3203-3212.Orynbayev MB, Beauvaisb W, Sansyzbaya AR, et al., 2016. Seroprevalence of infectious diseases in saiga antelope (Saiga tataricatatarica) in Kazakhstan 2012–2014. Prev Vet Med 127:100-104. Pakistan Livestock Census, 2006. Statistics Division. Agricultural Census Organization. Government of Pakistan. Lahore, Pakistan. Pan L, Zhang L, Fan D, et al., 2013. Rapid, simple and sensitive detection of Q fever by Loop-Mediated Isothermal Amplification of the htpAB gene. PLoS Negl Trop Dis 7(5):1-6. Paul S, Agger JF, Markussen B, et al., 2012. Factors associated with Coxiella burnetii antibody positivity in Danish dairy cows. Prev Vet Med 107:57-64. Paul S, Agger JF, Agerholm JS, et al., 2014. Prevalence and risk factors of Coxiella burnetii seropositivity in Danish beef and dairy cattle at slaughter adjusted for test uncertainty. Prev Vet Med 113:504-511.Philip CB, 1948. Comments of the name of the Q fever organism. Public Health Rep, 63:58.Pi?ero A, Barandika JF, García-Pérez AL, et al., 2015. Genetic diversity and variation over time of Coxiella burnetii genotypes in dairy cattle and the farm environment. Infect Genet Evol 31:231-235.Polo MF, Mastandrea S, Santoru L, et al., 2015. Pulmonary inflammatory pseudotumor due to Coxiella burnetii. Case report and literature review. Microbes Infect 17(11-12):795-98. Psaroulaki A, Hadjichristodoulou C, Loukaides F, et al., 2006. Epidemiological study of Q fever in humans, ruminant animals and ticks in Cyprus using a geographical information system. Eur J Clin Microbiol Infect Dis 25:576-586.Qiu Y, Nakao R, Namangala B, et al., 2013. Short report: First genetic detection of Coxiella burnetii in Zambian livestock. Am J Trop Med Hyg 89(3):518-519.Racic I, Spicic S, Galov A, et al., 2014. Identification of Coxiella burnetii genotypes in Croatia using multi-locus VNTR analysis. Vet Microbiol 173:340-347.Rahman MA, Alam MM, Islam MA, et al., 2016. Serological and molecular evidence of Q fever in domestic ruminants in Bangladesh. Vet Med Int 2016:1-7. Rai SB, Kamaludin F, Chow TS, et al., 2011. First documented zoonotic case of Q fever in Penang, Malaysia. Organ Studies Innov Rev (OSIR) 4(1):1-5. Raoult D, Vestris G and Enea M, 1990. Isolation of 16 strains of Coxiella burnetii from patients by using a sensitive centrifugation cell culture system and establishment of the strains in HEL cells. J Clin Microbiol 28:2482-2484.Rapf S, 2015. Q fever in Austria. Diploma Thesis. University Clinic for Internal Medicine. Section for Infectiology and Tropical Medicine. Graz Medical University, Austria.Reusken?C,?Plaats RVD, Opsteegh M, et al., 2011. Coxiella burnetii (Query fever) in?Rattus?norvegicus?and?Rattus?rattus?at?livestock?farms?and?urban locations in the Netherlands;?could?Rattus?species?represent?reservoirs?for (re)introduction. Prev?Vet?Med 101:124-130.Reye AL, Stegniy V, Mishaeva NP, et al., 2013. Prevalence of tick-borne pathogens in Ixodes ricinus and Dermacentor reticulatus ticks from different geographical locations in Belarus. Plos One 8(1):1-9.Rice DA and Knoke MAR, 1979. The prevalence of Q fever antibodies in dairy cows in El Salvador. Trop Anim Health Prod ll:50-50.Rizzo F, Vitaleb N, Ballardinia M, et al., 2016. Q fever seroprevalence and risk factors in sheep and goats in northwest Italy. Prev Vet Med 130:10-17. Roest HJ, Gelderen B, Dinkla A, et al., 2012. Q fever in pregnant goats: pathogenesis and excretion of Coxiella burnetii. PLoS One 7(11):1-14. Roest HI, Tilburg JJ, van der Hoek W, et al., 2011. The Q fever epidemic in the Netherlands: history, onset, response and reflection. Epidemiol Infect 139(1):1-12.Roest HJ, 2013. Coxiella burnetii in pregnant goats. Ph.D. Thesis, Department of Bacteriology and TSEs, Central Veterinary Institute of Wageningen UR, Lelystad, The Netherlands.Rotz LD, Khan AS, Lillibridge SR, et al., 2002. Public health assessment of potential biological terrorism agents. Emerg Infect Dis 8 (2):225-230.Ruiz-Fons F, Astobiza I, Barandika JF, et al., 2010. Seroepidemiological study of Q fever in domestic ruminants in semi-extensive grazing systems. BMC Vet Res 6:3. E, Kirby M, Clegg T, et al., 2011. Seroprevalence of Coxiella burnetii antibodies in sheep and goats in the Republic of Ireland. Vet Record 169(280):1-3.Sa?nchez J, Souriauy A, Buend??a AJ, et al., 2006. Experimental Coxiella burnetii infection in pregnant goats: a histopathological and immunohistochemical study. J Comp Path 135:108-115.Saegerman C, Speybroeck N, Pozzo FD, et al., 2013. Clinical indicators of exposure to Coxiella burnetii in dairy herds. Transbound Emerg Dis 62(1):46–54. Saglam AG and Sahin M, 2016. Coxiella burnetii in samples from cattle herds and sheep flocks in the Kars region of Turkey. Vet Med 61(1):17-22.Saiti I and Memishi SH, 2015. The frequency variation of Q-fever and comparison of infection in farm animals according to regions in Western Macedonia. Int J Educ Sci Tech Innov Health Environ 01(04):52-63.Salinas-Meléndez JA, Avalos-Ramírez R, Riojas-Valdez V, et al., 2002. Serologic survey in animals of ‘Q’ fever in Nuevo Leon. Rev Latinoam Microbiol 44(2):75-78. Samuel JE and Hendrix LR, 2009. Laboratory maintenance of Coxiella burnetii. Curr Proto Micriobiol 6C (suppl. 15):1-16.Santos AS, Tilburg JJ, Botelho A, et al., 2012. Genotypic diversity of clinical Coxiella burnetii isolates from Portugal based on MST and MLVA typing. Int J Med Microbiol 302:253-256.Saville P, 1996. The animal health status of Tonga. South pacific commission cataloguing-in-publication data Saville, Peter H. Report on the animal health status of Tonga. Prepared for publication by the SPC Regional Media Centre, Suva and printed by Oceania Printers Ltd, Suva Fiji, Tonga.Schelling E, Diguimbaye C, Daoud S, et al., 2003. Brucellosis and Q-fever seroprevalences of nomadic pastoralists and their livestock in Chad. Prev Vet Med 61:279-293.Schimmer B, Dijkstra F and Vellema P, 2009. Sustained intensive transmission of Q fever in the south of the Netherlands. Euro Surveill 14(19). PMID:19442401.Schimmer B, de Lange MMA, Hautvast JLA, et al., 2014. Coxiella burnetii seroprevalence and risk factors on commercial sheep farms in the Netherlands. Vet Rec 175(1):1-17.Schimmer B, Luttikholt S, Hautvast JLA, et al., 2011. Seroprevalence and risk factors of Q fever in goats on commercial dairy goat farms in the Netherlands, 2009–2010. BMC Vet Res 7:81-85.Schleenvoigt BT, Sprague LD, Mertens K, et al., 2015. Acute Q fever infection in Thuringia, Germany, after burial of roe deer fawn cadavers (Capreolus capreolus): a case report. New Microbe New Infect 8:19-20.Schneeberger PM, Hermans MHA, van Hannen EJ, et al., 2010. Real-time PCR with serum samples is indispensable for early diagnosis of acute Q fever. Clin Vaccine Immunol 17(2):286–290. Schoffelen T, Self JS, Fitzpatrick KA, et al., 2015. Early cytokine and antibody responses against Coxiella burnetii in aerosol infection of BALB/c mice. Diagn Microbiol Infect Dis 81:234-239.Schoffelen T, Herremans T, Sprong T, et al., 2013. Limited humoral and cellular responses to Q fever vaccination in older adults with risk factors for chronic Q fever. J Infect 67:565-573.Scolamacchia F, Handel IG, Fevre EM, et al., 2010. Serological patterns of brucellosis, leptospirosis and Q fever in Bos indicus cattle in Cameroon. PLoS ONE 5(1):1-11. e8623Scott GH, Williams JC and Stephenson EH, 1987. Animal models in Q fever: pathological responses of inbred mice to phase I Coxiella burnetii. J Gen Microbiol 133:691-700.Scrimgeour EM, Al-Ismaily SIN, Rolain JM, et al., 2003. Q fever in human and livestock populations in Oman. Ann NY Acad Sci 990:221-225.Selim A and Elhaig M, 2016. Q fever in domestic small ruminant. Asian J Anim Vet Adv 11(1):1-8.Seo MG, Lee SH, Ouh IO, et al., 2016. Molecular detection and genotyping of Coxiella-like endosymbionts in ticks that infest horses in South Korea. PLoS ONE 11(10):1-9. e0165784. doi:10.1371/journal.Seshadri R, Paulsen IT, Eisen JA, et al., 2003. Complete genome sequence of the Q-fever pathogen Coxiella burnetii. Proc Natl Acad Sci 100(9):5455-5460. Setiyono A and Subangkit M, 2014. Immunohistochemical detection of Coxiella burnetii in ruminants: A case study of Q fever in Indonesia. Global Vet 12(6):865-868. Shabbir MZ, Akram S, Hassan Z, et al., 2016. Evidence of Coxiella burnetii in Punjab province, Pakistan. Acta Trop 163:61-69. Shah SY, Kovacs C, Tan CD, et al., 2015. Delayed diagnosis of Q fever endocarditis in a rheumatoid arthritis patient. IDCases 2:94-96.Shapiro AJ, Bosward KL, Heller J, et al., 2015. Seroprevalence of Coxiella burnetii in domesticated and feral cats in eastern Australia. Vet Microbiol 177:154-161.Sidi-Boumedine K, Adam G, Angen O, et al., 2015. Whole genome PCR scanning (WGPS) of Coxiella burnetii strains from ruminants. Microbes Infect 17(11-12):772-775.Sprong H, Tijsse-Klasen E, Langelaar A, et al., 2012. Prevalence of Coxiella burnetii in ticks after a large outbreak of Q fever. Zoonoses Public Health 59:69-75.Staley GP, Myburgh JG and Chaparro F, 1989. Serological evidence of?Q fever?in cattle in?Malawi. Onderstepoort J Vet Res 56(3):205-206.Steinmann P, Bonfoh B, Peter O, et al., 2005. Seroprevalence of Q-fever in febrile individuals in Mali. Trop Med Int Health 10(6):612–617.Sting R, Molz K, Philipp W, et al., 2013. Quantitative real-time PCR and phase specific serology are mutually supportive in Q fever diagnostics in goats. Vet Microbiol 167:600-608.Stone DM, Kumthekar S, Chikweto A, et al., 2012. Exposure to zoonotic abortifacients among sheep and goats in Grenada. Int J Anim Vet Adv 4(2):113-118.Sulyok KM, Kreizinger Z, Hornstra HM, et al., 2014. Genotyping of Coxiella burnetii from domestic ruminants and human in Hungary: indication of various genotypes. BMC Vet Res 10:107-113.Tarasevic IV, Plotnikova LF, Fetisova NF, et al., 1976. Natural foci of rickettsioses in the Armenian Soviet Socialist Republic. Bull World Health Organ 53:25-30.Thompson CN, Blacksell SD, Paris DH, et al., 2015. Undifferentiated febrile illness in Kathmandu, Nepal. Am J Trop Med Hyg 92(4):875-878. Thrusfield M, 2007. Veterinary Epidemiology, 3rd Ed., Blackwell Sci Ltd, New Jersey, USA. Toledo A, Jado AS, Olmeda MA et al., 2009. Detection of Coxiella burnetii in ticks collected from central Spain. Vector Borne Zoonotic Dis 9:465-468.Touratier A, Baurier F, Beaudeau F, et al., 2012. Comment Faire le Diagnostic d’un ?levage Cliniquement Atteint de Fièvre Q? In Pathologie Infectieuse: Actualités cliniques, diagnostiques et thérapeutiques, Syndromes émergents. Edited by Société Nationale des Groupements Techniques Vétérinaires, Paris. Journées Nationales des GTV 2012:147-155.Tukur HB, Ajogi I, Kabir J, et al., 2014. Seroprevalence of Coxiella burnetti in cattle and its risk factors in Kaduna Metropolis, Kaduna State, Nigeria. IOSR-JAVS 7(2):01-05.Vaidya VM, Malik SVS, Bhilegaonkar KN, et al., 2010. Prevalence of Q fever in domestic animals with reproductive disorders. Comp Immunol Microbiol Infect Dis 33:307-321.Van Asseldonk MAPM, Bontje DM, Backer JA, et al., 2015. Economic aspects of Q fever control in dairy goats. Prev Vet Med 121:-115-122.Van Asseldonk MAPM, Prins J and Bergevoet RHM, 2013. Economic assessment of Q fever in the Netherlands. Prev Vet Med 112:27-34.Van den Brom R and Vellema P, 2009. Q fever outbreaks in small ruminant and people in the Netherland. Small Rumin Res 86:74-83.Van den Brom R, Moll L, van Schaik G, et al., 2013. Demography of Q fever seroprevalence in sheep and goats in The Netherlands in 2008. Prev Vet Med 109:76-82.Van den Brom R, van Engelen E, Luttikholt S, et al., 2012. Coxiella burnetii in bulk tank milk samples from dairy goat and dairy sheep farms in the Netherlands in 2008. Vet Rec 170(12):310.Van den Brom R, van Engelen E, Roest HIJ, et al., 2015. Coxiella burnetii infections in sheep or goats: an opinionated review. Vet Microbiol 181:119-129.Van der Hoek W, Meekelenkamp JC, Leenders AC, et al., 2011. Antibodies against Coxiella burnetii and pregnancy outcome during the 2007–2008 Q feverout breaks in The Netherlands. BMC Infect Dis 11:44-52. 44, S, Rubach MP, Halliday JEB, et al., 2014. Epidemiology of Coxiella burnetii infection in Africa: A one health systematic review. PLoS Negl Trop Dis 8(4):1-10. e2787. Van Schaik EJ, Chen C, Mertens K, et al., 2013. Molecular pathogenesis of the obligate intracellular bacterium Coxiella burnetii. Nat Rev Microbiol 11:561-573.Vellema P and van den Brom R, 2014. The rise and control of the 2007–2012 human Q fever outbreaks in the Netherlands. Small Ruminant Res 118:69-78.Vincent GA, Graves SR, Robson JM, et al., 2015. Isolation of Coxiella burnetii from serum of patients with acute Q fever. J Microbiol Methods 119:74-78.Vranakis I, Kokkini S, Chochlakis D, et al., 2012. Serological survey of Q fever in Crete, southern Greece. Comp Immunol Microbiol Infect Dis 35:123-127. Wardrop NA, Thomas LF, Cook EAJ, et al., 2016. The sero-epidemiology of Coxiella burnetii in humans and cattle, Western Kenya: Evidence from a cross-sectional study. PLoS Negl Trop Dis 10(10):1-17. e0005032. Wattiau P, Boldisova E, Toman R, et al., 2011. Q fever in wool sorters, Belgium. Emerg Infect Dis 17(12):2368-2369.Weitzel T, López J, Acosta-Jamett G, et al., 2016. Absence of convincing evidence of Coxiella burnetii infection in Chile: a cross-sectional serosurvey among healthy adults in four different regions. BMC Infect Dis 16(541):1-6.WHO, 2015. Neglected zoonotic diseases (NZDs). Geneva, Switzerland. (accessed 3.26.15.).Wielders CCH, Wijnbergen PCA, Renders NHM, et al., 2013. High Coxiella burnetii DNA load in serum during acute Q fever is associated with progression to a serologic profile indicative of chronic Q fever. J Clin Microbiol 51(10):3192-3198.Wielders CCH, Teunis PFM, Hermans MHA, et al., 2015. Kinetics of antibody response to Coxiella burnetii infection (Q fever): Estimation of the seroresponse onset from antibody levels. Epidemics 13:37-43.Woldehiwet Z, 2004. Q fever (coxiellosis): epidemiology and pathogenesis. Res Vet Sci 77:93-100. H, Drebot MA, Dewailly E, et al., 2014. Short report: Seroprevalence of seven zoonotic pathogens in pregnant women from the Caribbean. Am J Trop Med Hyg 91(3):642-644.Wouda W and Dercksen DP, 2007. Abortion and stillbirth among dairy goats as a consequence of Coxiella burnetii. Tijdschr Diergeneeskd 132(23):908-11.Y?lmaza G, ?ztürkb B, Memikoglu O, et al., 2015. An unusual manifestation of Q fever: Peritonitis. J Infect Public Health 8:373-376.Ylli B, Ismije S, Arta L, et al., 2013. Serological survey of Q fever in small ruminant and cattle in five regions of Albania: An update. J Anim Vet Adv 12(3):402-405.Zahid MU, Hussain MH, Saqib M, et al., 2016. Sero-prevalence of Q fever (Coxiellosis) in small ruminant of two district in Punjab, Pakistan. Vector Borne Zoonotic Dis 16: 449-454. ................
................

In order to avoid copyright disputes, this page is only a partial summary.

Google Online Preview   Download