Coxiella burnetii in Humans, Domestic Ruminants, and Ticks in Rural Western Kenya.
ABSTRACT We conducted serological surveys for Coxiella burnetii in archived sera from patients that visited a rural clinic in western Kenya from 2007 to 2008 and in cattle, sheep, and goats from the same area in 2009. We also conducted serological and polymerase chain reaction-based surveillance for the pathogen in 2009-2010, in human patients with acute lower respiratory illness, in ruminants following parturition, and in ticks collected from ruminants and domestic dogs. The IgG antibodies against C. burnetii were detected in 30.9% (N = 246) of archived patient sera and in 28.3% (N = 463) of cattle, 32.0% (N = 378) of goats, and 18.2% (N = 159) of sheep surveyed. Four of 135 (3%) patients with acute lower respiratory illness showed seroconversion to C. burnetii. The pathogen was detected by polymerase chain reaction in specimens collected from three of six small ruminants that gave birth within the preceding 24 hours, and in five of 10 pools (50%) of Haemaphysalis leachi ticks collected from domestic dogs.
- SourceAvailable from: Yen-Hsu Chen[Show abstract] [Hide abstract]
ABSTRACT: The clinical characteristics of Q fever are poorly identified in the tropics. Fever with pneumonia or hepatitis are the dominant presentations of acute Q fever, which exhibits geographic variability. In southern Taiwan, which is located in a tropical region, the role of Q fever in community-acquired pneumonia (CAP) has never been investigated.PLoS ONE 07/2014; 9(7):e102808. · 3.53 Impact Factor
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ABSTRACT: Two hundred fourteen serosamples were collected from four livestock species across five ranches in Laikipia County, Kenya. Serological analysis for Coxiella burnetii (the causative agent for Q fever) showed a distinct seroprevalence gradient: the lowest in cattle, higher in sheep and goats, and the highest in camels. Laikipia-wide aerial counts show a recent increase in the camel population. One hundred fifty-five stakeholder interviews revealed concern among veterinary, medical, ranching, and conservation professionals about Q fever. Local pastoralists and persons employed as livestock keepers, in contrast, revealed no knowledge of the disease. This work raises questions about emerging Q fever risk in Laikipia County and offers a framework for further integrative disease research in East African mixed-use systems.EcoHealth 03/2014; · 2.27 Impact Factor
Am. J. Trop. Med. Hyg., 88(3), 2013, pp. 513–518
Copyright © 2013 by The American Society of Tropical Medicine and Hygiene
Coxiella burnetii in Humans, Domestic Ruminants, and Ticks in Rural Western Kenya
Darryn L. Knobel,* Alice N. Maina, Sally J. Cutler, Eric Ogola, Daniel R. Feikin, Muthoni Junghae, Jo E. B. Halliday,
Allen L. Richards, Robert F. Breiman, Sarah Cleaveland, and M. Kariuki Njenga
Boyd Orr Centre for Population and Ecosystem Health, Institute of Biodiversity, Animal Health and Comparative Medicine,
University of Glasgow, Glasgow, United Kingdom; Department of Veterinary Tropical Diseases, University of Pretoria, Onderstepoort,
South Africa; Jomo Kenyatta University of Agriculture and Technology, Nairobi, Kenya; Viral and Rickettsial Diseases Department,
Naval Medical Research Center, Silver Spring, Maryland; School of Health, Sports and Biosciences, University of East London, London,
United Kingdom; Kenya Medical Research Institute/Centers for Disease Control and Prevention-Kenya, Kisumu, Kenya;
Global Disease Detection Division, CDC-Kenya, Nairobi, Kenya
clinic in western Kenya from 2007 to 2008 and in cattle, sheep, and goats from the same area in 2009. We also conducted
serological and polymerase chain reaction-based surveillance for the pathogen in 2009–2010, in human patients with
acute lower respiratory illness, in ruminants following parturition, and in ticks collected from ruminants and domestic
dogs. Antibodies against C. burnetii were detected in 30.9% (N = 246) of archived patient sera and in 28.3% (N =
463) of cattle, 32.0% (N = 378) of goats, and 18.2% (N = 159) of sheep surveyed. Four of 135 (3%) patients with
acute lower respiratory illness showed seroconversion to C. burnetii. The pathogen was detected by polymerase chain
reaction in specimens collected from three of six small ruminants that gave birth within the preceding 24 hours, and
in five of 10 pools (50%) of Haemaphysalis leachi ticks collected from domestic dogs.
We conducted serological surveys for Coxiella burnetii in archived sera from patients that visited a rural
Q fever is a disease of humans caused by infection with the
obligate intracellular bacterial pathogen, Coxiella burnetii. It
is a zoonotic infection, typically transmitted from animal hosts
to humans through inhalation of contaminated aerosols or
ingestion of infected animal products such as milk or cheese.1
Ticks have also been implicated as vectors, and over 40 species
of ticks have been found naturally infected with the agent.1
Coxiella burnetii has a wide host range, including wild and
domestic mammals, birds, reptiles, and arthropods.2Domestic
ruminants (primarily goats, cattle, and sheep) represent the
most frequent source of human infection,3although transmis-
sion from dogs and cats is also documented.4,5The uterus
and mammary glands of female animals are sites of chronic
C. burnetii infection, and infected females may shed large
amounts of bacteria into the environment during parturition
or spontaneous abortion.2Once shed, the organisms may
remain infective in the environment for several months.1
Infection in humans, usually by inhalation, may be asymp-
tomatic (up to 60% of infected individuals) or may manifest
clinically after an incubation period ranging between 1 and
3 weeks.1,6Clinical signs of acute Q fever include fever of 2–
14 days’ duration, atypical pneumonia, and/or hepatitis.1
Although the disease is typically self-limiting, severe debili-
tating illness requiring hospitalization can occur in a small
proportion (2–5%) of acutely infected cases.6Chronic dis-
ease may develop following infection, particularly in patients
with predisposing conditions such as preexisting cardiac
valvulopathy, pregnancy, or immunosuppression.1Common
manifestations of chronic disease include endocarditis and
vascular infection.1Coxiella burnetii displays antigenic
(phase) variation associated with loss of virulence and muta-
tional variation in the lipopolysaccharide.1High levels of
antibodies to phase I antigens are detected during chronic
Q fever, whereas antibodies to phase II antigens are pro-
duced in acute disease.6
Coxiella burnetii is found worldwide, with the exception of
New Zealand.7A recent large outbreak in the Netherlands
involved at least 3,523 human cases from 2007 through 2009,8
and was characterized by a high rate of hospitalization, with
20% of notified cases admitted to hospital in 2008–2009.9
Pneumonia was the predominant clinical presentation.8The
Netherlands outbreak has been linked to the increase in the
country’s dairy goat population, which more than doubled in
size between 2000 and 2009, and highlights the public health
risks of Q fever epidemics posed by domestic ruminants.
Despite the high-profile nature of some Q fever outbreaks,
and the attention that C. burnetii has received as a potential
bioterrorism agent,10information on the prevalence of infec-
tion in sub-Saharan Africa is scant.11,12In Kenya, serological
evidence of Q fever in patients with acute febrile and respira-
tory illness was shown in the 1950s.13Other studies showed
the prevalence of antibodies to C. burnetii among Kenyans
to range between 10% and 20%.14,15A more recent investi-
gation found that four people (8%) of a group of 50 travelers
to Kenya contracted Q fever,16and in another recent study
investigators diagnosed acute Q fever in 5% of febrile
patients admitted to two hospitals from September 2007 to
August 2008, in neighboring northern Tanzania.17Among
domestic ruminants in Kenya, the prevalence of antibodies
was reported as 7–57% in cattle and 33–34% in goats.14,15,18,19
There are no other recent reports of investigations into dis-
ease prevalence in resident human or livestock populations,
or any information on the relationship between prevalence
in human and animal populations.
The establishment of a population-based infectious disease
surveillance program in western Kenya20provided a valuable
opportunity for generating domestic animal prevalence data
that could be linked with human health outcomes. To assess
the current status of Q fever among humans and infection
prevalence in domestic ruminants, we conducted surveys for
C. burnetii in cattle, goats, sheep, and ticks in this rural agro-
pastoral community, and tested specimens from human
patients presenting to a clinic in the same area for Q fever.
*Address correspondence to Darryn L. Knobel, Department of
Veterinary Tropical Diseases, Faculty of Veterinary Science, Univer-
sity of Pretoria, Onderstepoort, 0110, South Africa. E-mail: darryn
MATERIALS AND METHODS
Study site. AlldatawerecollectedinAsembowithinRarieda
(formerly Bondo) District in western Kenya in 2007–2010.
This rural site on the eastern shore of Lake Victoria falls
within a health and demographic surveillance system (HDSS)
that has been run by the Kenya Medical Research Institute
(KEMRI) and U.S. Centers for Disease Prevention and
Control (CDC) since 2001. The HDSS collects house-
hold demographic and socioeconomic data three times per
year, and includes information on the number of livestock
owned.21Households are clustered into compounds com-
posed of related family units, with most compounds having
between one and five family units.22The primary economic
activity is subsistence smallholder agro-pastoralism and fish-
ing. In this area, 44% of households own cattle (mean number
owned: 1.84) and 43% own at least one sheep or goat (mean
number owned: 2.12; HDSS data for 2008, unpublished).
The Kenyan International Emerging Infections Program
of KEMRI/CDC has conducted population-based infectious
disease surveillance (PBIDS) of people in Asembo since late
2005, with between 23,500 and 25,000 people under surveil-
lance in 33 villages.20Participants enrolled in PBIDS have
household visits to determine health status every 2 weeks,
and receive free medical care for all acute illnesses at a
centrally located clinic, St. Elizabeth Lwak Mission Hospi-
tal (henceforth Lwak Hospital). Specimen collection and
diagnostic testing are focused on determining etiologies for
four infectious disease syndromes, namely acute respiratory
infection, diarrhea, jaundice, and febrile illness. Sera are
collected from patients participating in PBIDS presenting
to Lwak Hospital and meeting case definitions for acute
respiratory illness, jaundice or acute febrile illness,20or
who are hospitalized for any non-traumatic illness. Between
January 2007 and January 2010, 3,948 serum specimens
were collected and archived at −80°C at the KEMRI/CDC
laboratory in Kisumu.
Ethical review. The collection of specimens from humans
was approved by the KEMRI Ethical Review Committee
(protocol no. 932) and CDC Institutional Review Board
(protocol no. 4566), and from animals by the KEMRI
and CDC Animal Care and Use Committees (protocol no.
1191 and 1562BRETBDX, respectively). Written informed
consent was obtained from all patients or animal owners
before specimen collection.
Human serosurvey. To assess previous exposure to
C. burnetii in humans, we retrospectively examined a subset
of archived sera. Sera collected from January 2007 through
February 2008 were identified and 248 specimens randomly
selected from this period (sufficient to attain a 95% confi-
dence interval with an absolute precision of 5% at an expected
prevalence of 20%).23Sera were tested at the KEMRI/CDC
laboratory in Kisumu for IgG antibodies against phase I and
phase II C. burnetii antigens derived from the Nine Mile
strain, using an indirect fluorescence antibody (IFA) assay
(Fuller Laboratories, Fullerton, CA). Serum specimens were
screened at a dilution of 1:32. A positive and a negative con-
trol were included for each assay run, and all slides were read
by a single operator. A sample was considered positive if
bright, sharply defined apple-green fluorescent elementary
bodies were observed against a background of red counter-
stained material at this serum dilution.
Human respiratory patients. Under the PBIDS clinic sur-
veillance protocol, blood specimens are collected from
patients who meet the case definition for acute lower respira-
tory infection (ALRI). This is defined in persons 5 years of
age or older as cough, difficulty breathing or chest pain and
either documented axillary temperature ³ 38.0°C or oxygen
saturation < 90%, and in children under 5 years of age
as cough or difficulty breathing with one of the following:
elevated respiratory rate for age (non-severe pneumonia),20
or lower chest wall indrawing, stridor, oxygen saturation
< 90%, maternal report of convulsions, inability to drink or
breastfeed, or vomiting everything, or on exam lethargy or
unconsciousness (severe and very severe pneumonia).20All
patients who meet the case definition have acute-phase sera
are requested to return to the clinic after 4–6 weeks for the
collection of convalescent-phase serum specimens. Paired sera
are testedserologically for a rangeof respiratorypathogens.
To determine the prevalence of C. burnetii infection in these
patients with respiratory illness, we retrospectively identified
paired serum specimens collected from ALRI cases occurring
from July 2009 through January 2010. Convalescent-phase
serum specimens were screened for IgG antibodies against
C. burnetii phase II antigen by indirect enzyme-linked immu-
nosorbent assay (ELISA) (Panbio, Brisbane, Australia) using
the manufacturer’s recommended cutoff. End-point titers of
IgG antibodies in acute and convalescent sera were determined
by IFA assay for all pairs in which the convalescent-phase
tested positive or equivocal on ELISA, with serial dilutions
starting from 1:16. The operator who performed the IFA was
blind to the status (acute or convalescent) of the specimen. A
³ 4-fold increase in IFA titer to C. burnetii phase II antigen
defined acute Q fever, and a titer ³ 1:1,024 to C. burnetii phase
I antigen in either acute- or convalescent-phase sera defined
possible chronic Q fever.
Domestic ruminants cross-sectional study. Data were col-
lected from January through May 2009. The study population
comprised all domestic ruminants (cattle, goats, and sheep)
living within compounds in the 33 villages of the PBIDS. The
sampling frame was compiled from data on the number of
each species of domestic animal within the compounds,
as reported at the most recent HDSS visit for which data
were available. Three hundred livestock-owning compounds
(LOCs) were randomly selected from the sampling frame of
4,528 LOCs, with a LOC defined by ownership of one or more
animals of any of the following species: cattle, sheep, goats,
chickens. The heads of the selected compounds were
approached and offered enrollment. If livestock were no longer
owned, replacement LOCs was then randomly selected from
the same village.
Specimens were collected from a maximum of three ran-
domly selected animals of any age from each of the species of
domestic ruminants present at enrolled compounds. Blood
was collected by venipuncture of the jugular vein into plain
vacutainers (BD, Franklin Lakes, NJ). Vaginal swabs were
collected from adult females only, using nylon flocked swabs
(eSwabs, Copan Innovation, Brescia, Italy) and placed in
universal transport medium (Copan Innovation). Blood col-
lection tubes were kept at ambient temperature for 15–
30 minutes, and then transported to the field laboratory on
ice with the vaginal swabs. At the field laboratory, blood tubes
were centrifuged at 2,000–3,000 revolutions per minute for
KNOBEL AND OTHERS
7–10 minutes, and serum pipetted into cryovials. Sera and
swabs were then transported on ice to the KEMRI-CDC lab-
oratory and stored at −80°C until testing. Ruminant serum
samples were tested for the presence of IgG antibodies
to C. burnetii by indirect ELISA (Chekit Q fever ELISA,
IDEXX), following the manufacturer’s instructions. DNA was
extracted from vaginal swab specimens using QIAamp DNA
Mini Kit (Qiagen, Valencia, CA) in a biosafety level 3 labora-
tory, and tested by quantitative real-time polymerase chain
reaction (qPCR) assay using the IS1111 gene target for C.
burnetii, as previously described.24
Domestic ruminant births/abortions. To further investigate
shedding of C. burnetii by domestic ruminants, we conducted
surveillance for births and abortions in the ruminant popula-
tion in the PBIDS area during 1 week in August 2009, as part
of a pilot study for a larger surveillance program on animal
disease syndromes. A network of animal health reporters
(AHRs) was established in the 33 villages in the PBIDS (1–
2 AHRs per village), through which all livestock owners could
report births or abortions in their ruminant herds. The AHRs
then contacted the study field veterinary team by mobile
phone, who responded to the case as soon as possible. Avail-
able samples were collected by the veterinary team, including
blood and vaginal swabs from dams, placenta, and liver, lungs,
spleen, and brain of aborted fetuses or stillborn animals.
DNA was extracted and tested as described previously.
Collection of arthropods. All domestic ruminants from
which blood was collected were also inspected for ticks. In
addition and when present, up to three domestic dogs in each
enrolled compound were also inspected. In each animal, we
examined specific body sites (ruminants: ear, dewlap, shoul-
der, belly, groin, udder, and perineum; dogs: ear, neck, nape,
belly, and groin) and collected 1–2 non-engorged adult ticks
of different genera per site (although only Amblyomma ticks
were tested from ruminants). Ticks were preserved in 70%
ethanol at room temperature until delivery to the laboratory,
and then stored in ethanol at −80°C. Ticks were pooled by
individual host animal and tick species (1–6 ticks per pool).
Ticks of the genus Amblyomma collected from ruminants and
all ticks collected from dogs were identified using published
entomological keys25; after identification, ticks were washed
in molecular-grade water and mechanically disrupted using
a bead mill (Qiagen TissueLyser LT). Genomic DNA was
extracted using QIAamp blood and tissue kits (Qiagen)
according to the manufacturer’s instructions, using a final
elution volume of 100 mL, and tested by IS1111 qPCR.24
Human serosurvey. A total of 248 and 246 archived sera
were tested by IFA for IgG antibodies against C. burnetii
phase I and phase II antigen, respectively. The IgG antibodies
against C. burnetii phase I antigen were detected in 58 speci-
mens (23.4%, exact binomial 95% confidence intervals [CI]:
18.3–29.2%), and IgG antibodies against phase II antigen
were detected in 76 specimens (30.9%, 95% CI: 25.2–37.1%).
All the specimens that reacted with phase I antigen also
reacted with phase II antigen.
Human respiratory patients. Of 2,246 patients who met the
case definition for ALRI from July 2009 through January
2010, 493 (22%) returned for collection of convalescent sera.
Of these, 135 pairs were available for testing. Convalescent-
phase sera were screened for IgG antibodies against C. burnetii
phase II antigen by indirect ELISA. Nineteen patients (14.1%
of ALRI patients, 95% CI: 8.7–21.1%) had positive titers on
ELISA. Of these, 4 (3% of ALRI patients, 95% CI: 0.8–
7.4%) were diagnosed serologically with acute Q fever by
IFA as determined by a ³ 4-fold rise between acute and con-
valescent sera. All four patients were males, 7–31 years of age,
who presented with fever (documented axillary temperatures
of 38.2–39.5°C), chills, cough, and headache (Table 1). The
reported duration of illness ranged from 2 to 10 days. All
four patients were from households that owned one or more
domestic ruminants around the time of illness. One of the
patients (#2 in Table 1) also showed seroconversion to influ-
enza B virus. No patients were diagnosed with possible chronic
Domestic ruminants cross-sectional study. Of the 300 LOCs
identified, 236 owned one or more species of domestic
ruminant. ELISA results were obtained for sera from 463
cattle, 378 goats, and 159 sheep in these compounds. The
seroprevalence of antibodies to C. burnetii, was 28.3%
(95% CI: 24.2–32.6%) in cattle, 32% (95% CI: 27.3–37%)
in goats, and 18.2% (95% CI: 12.6–25.1%) in sheep. The
age-stratified seroprevalence results for the three species
are given in Figure 1. Coxiella burnetii DNA was detected
by IS1111 PCR in vaginal swabs from 5 of 233 (2.1%) cattle,
5 of 222 (2.3%) goats, and 4 of 85 (4.7%) sheep.
Domestic ruminant births/abortions. Six births (five in goats
and one in a sheep) and one abortion in a cow were detected
during the week-long surveillance. In all cases, specimens were
collected within 24 hours of the event. Three births in goats
resulted in one stillborn kid in each, along with one to two live
offspring. Samples from the placenta of one of these cases,
and vaginal swabs from a sheep and a goat in the same
compound that gave birth on consecutive days, were positive
for C. burnetii on the qPCR assay for the IS1111 gene target.
Arthropods. A total of 258 adult Amblyomma variegatum
by individual host. Coxiella burnetii DNA was detected by
IS1111 PCR in 4 of 162 (2.5%) pools. In dogs, 102 ticks were
collected from 36 animals, resulting in pools of the following
species: Rhipicephalus sanguineus (15 pools), Rhipicephalus
appendiculatus (9 pools), unspeciated Rhipicephalus (10 pools),
A. variegatum (10 pools), Haemaphysalis leachi (10 pools), and
Clinical features of four patients from rural western Kenya with acute
lower respiratory illness, diagnosed serologically with acute Q fever
COXIELLA BURNETII IN HUMANS, ANIMALS, AND TICKS IN KENYA
Rhipicephalus (Boophilus) decoloratus (5 pools). The number
(and percent) of pools positive using the IS1111 PCR assays
were: Rh. sanguineus 2 (20%), Rh. appendiculatus 1 (11.1%),
unspeciated Rhipicephalus 2 (20%), A. variegatum 2 (20%),
We identified Q fever as a possible cause of acute lower
respiratory illness among rural residents in western Kenya.
In addition, we found a high prevalence of antibodies to
C. burnetii in domestic ruminants in the same area, with high
rates of shedding of the organism shortly after parturition in
a small sample of animals.
The prevalence of antibodies to C. burnetii in patients
attending Lwak Hospital is similar to that reported in several
population-based studies elsewhere in sub-Saharan Africa (17–
37%),12,26,27indicating high levels of exposure to the pathogen.
In addition to the recent study in northern Tanzania in which
acute Q fever was diagnosed in 5.0% of febrile inpatients,17
anotherstudyreportedrecentinfection withC.burnetiiin 9.5%
of febrile patients > 5 years of age in two urban areas in Mali.28
A literature review did not identify any recent investigations
of Q fever as a cause of respiratory illness in Africa, although
in the report of Q fever in travelers returning from Kenya,16
In addition to ALRI, it is likely that Q fever also contrib-
utes to the burden of acute febrile illness in patients in our
study area. Both ALRI and febrile illness are managed empir-
ically in this setting, due largely to a lack of reliable rapid
diagnostic tests for diseases other than malaria. Although
malaria may be responsible for > 50% of febrile illness epi-
sodes in this holoendemic area,29knowledge of the full range
of prevalent etiologies is important to improve the empiric
treatment of this syndrome. Doxycycline is considered the
preferred antibiotic for the treatment of acute Q fever,1and
there is evidence that newer macrolides and fluoroquinilones
are also effective.30Notably, C. burnetii is resistant in vitro to
those antimicrobial agents that are typically used for the
empirical treatment of ALRI in Kenya (penicillin and its
derivatives, and aminoglycosides).1A diagnosis of Q fever
pneumonia should therefore be considered in patients not
responding to first-line antimicrobial therapy, particularly in
those with exposure to known risk factors.
Inhalation of fomites contaminated by the parturient fluids
of infected animals is the main mode of human infection with
C. burnetii.3Despite a very small sample, a high proportion of
small ruminants in our study were found to shed the pathogen
at parturition (2 of 5 goats and 1 of 1 sheep sampled).The
stratified age-seroprevalence curves in ruminants show an
increase in prevalence with age until around 2–3 years, fol-
lowed by a plateau. This pattern has been observed else-
where31and is consistent with cumulative exposure to the
pathogen from a young age, with the highest prevalence of
antibody in reproductively mature animals.
Coxiella burnetii was detected by PCR in all species of ticks
studied. Ticks become infected during the transient bacter-
emia that occurs in the vertebrate host early after infection.
In our study, the highest prevalence of infection with
C. burnetii was found in H. leachi (the yellow dog tick), one
of two ticks adapted to feeding on domestic dogs in tropical
and sub-tropical areas. Detection of C. burnetii in this spe-
cies has been reported in Kenya previously32; these findings
suggest that domestic dogs may play a role as reservoir hosts
and sources of human infection with C. burnetii in Kenya, as
has been reported elsewhere.5
Our findings show that Q fever is a significant yet under-
diagnosed cause of human respiratory illness in Kenya.
Kenya from January through May 2009. The three panels show the age seroprevalence in cattle, goats, and sheep, respectively. Within each
panel, bold horizontal bars indicate the mean seroprevalence in each age class and the vertical bars show the 95% confidence intervals for
Age-stratified seroprevalence of IgG antibodies to Coxiella burnetii in sera collected from cattle, goats, and sheep in western
KNOBEL AND OTHERS
Domestic ruminants are sources of human infection, through
direct contact or contamination of the environment during
parturition or abortion. Although not assessed in this study,
consumption of dairy products from infected ruminants is also
likely to pose a risk. Although risk factor studies would be
helpful to identify the principal modes of transmission to
humans in this setting, our results support the likely efficacy
of recognized public health and hygiene measures for mitigat-
ing the risk of transmission from domestic ruminants, such as
confinement of domestic animals during parturition, avoid-
ance of contaminated pastures and contact with placental
material, and boiling or pasteurization of milk before con-
sumption. This study also suggests that investigation of the role
of domestic dogs and tick-borne transmission is warranted to
identify potential sources of infection and risk factors that are
currently not well recognized.
Received March 15, 2012. Accepted for publication December 15, 2012.
Published online February 4, 2013.
Acknowledgments: We thank the IEIP-Z field team (Samuel Asembo,
Michael Otieno, James Oyigo, and Pauline Otieno) for the collection
of ticks and specimens from domestic animals, and Immaculate Amadi
and Catherine Sonye for assistance with laboratory testing.
Financial support: This research was supported by the Wellcome Trust,
UK (grant no. 081828/B/06/Z) and U.S. Centers of Disease Con-
trol and Prevention and Global Emerging Infections Surveillance and
Response System Program (work unit no. 188M.0931.001.A0074).
Disclaimer: The findings and conclusions are those of the authors.
They do not necessarily represent the official policy or position of
the Centers for Disease Control and Prevention, the Department of
the Navy, Department of Defense, or the U.S. Government.
ical Diseases, Faculty of Veterinary Science, University of Pretoria,
Maina and Allen L. Richards, Viral and Rickettsial Diseases Depart-
ment, Naval Medical Research Center, Silver Spring, MD, E-mails:
email@example.com and firstname.lastname@example.org. Sally J.
Cutler, School of Health, Sports and Bioscience, University of East
London, United Kingdom, E-mail: S.Cutler@uel.ac.uk. Eric Ogola
and Muthoni Junghae, Kenya Medical Research Institute, Centers
for Disease Control and Prevention-Kenya, International Emerging
Infections Program, Kenya, E-mails: EOgola@kemricdc.org and
email@example.com. Daniel R. Feikin, Johns Hopkins Bloomberg
School ofPublic Health andCentersforDisease Control and Preven-
tion, Baltimore, MD, E-mail: firstname.lastname@example.org. Jo E. B. Halliday
and Sarah Cleaveland, Institute of Biodiversity, Animal Health
and Comparative Medicine, College of Medical, Veterinary and
Life Sciences, University of Glasgow, United Kingdom, E-mails:
email@example.com and firstname.lastname@example.org.
Robert F. Breiman, International Emerging Infections Program,
Global Disease Detection Division, Centers for Disease Control
and Prevention, Kenya, E-mail: email@example.com. M. Kariuki
Njenga, Integrated Human-Animal Health Program, Global Disease
Detection Division, Centers for Disease Control and Prevention,
Kenya, E-mail: firstname.lastname@example.org.
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