J. Clin. Microbiol.
Heller, Yves Piemont and Edward B. Breitschwerdt
RemyDarren C. Simpson, Carrie M. Hew, Dorsey L. Kordick,
Chao-Chin Chang, Rickie W. Kasten, Bruno B. Chomel,
Coyotes from Central Coastal California
vinsonii subsp. berkhoffii Infection in
Molecular Epidemiology ofBartonella
a Human Pathogenic Bartonella sp.:
Coyotes (Canis latrans) as the Reservoir for
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JOURNAL OF CLINICAL MICROBIOLOGY,
Copyright © 2000, American Society for Microbiology. All Rights Reserved.
Nov. 2000, p. 4193–4200Vol. 38, No. 11
Coyotes (Canis latrans) as the Reservoir for a Human Pathogenic
Bartonella sp.: Molecular Epidemiology of Bartonella vinsonii subsp.
berkhoffii Infection in Coyotes from Central Coastal California
CHAO-CHIN CHANG,1RICKIE W. KASTEN,1BRUNO B. CHOMEL,1* DARREN C. SIMPSON,2
CARRIE M. HEW,1DORSEY L. KORDICK,3REMY HELLER,4YVES PIEMONT,4
AND EDWARD B. BREITSCHWERDT3
Department of Population Health and Reproduction, School of Veterinary Medicine, University of California, Davis,
California 956161; Wildlife Unit, Vector Control Section, Santa Clara County Department of Health Services,
San Jose, California 951262; Department of Clinical Sciences, College of Veterinary Medicine,
North Carolina State University, Raleigh, North Carolina 276063; and Institut de Bacte ´riologie,
Universite ´ L. Pasteur, Ho ˆpitaux Universitaires, 67000 Strasbourg, France4
Received 6 March 2000/Returned for modification 27 July 2000/Accepted 8 September 2000
Bartonella vinsonii subsp. berkhoffii was originally isolated from a dog suffering infectious endocarditis and
was recently identified as a zoonotic agent causing human endocarditis. Following the coyote bite of a child who
developed clinical signs compatible with Bartonella infection in Santa Clara County, Calif., this epidemiological
study was conducted. Among 109 coyotes (Canis latrans) from central coastal California, 31 animals (28%) were
found to be bacteremic with B. vinsonii subsp. berkhoffii and 83 animals (76%) had B. vinsonii subsp. berkhoffii
antibodies. These findings suggest these animals could be the wildlife reservoir of B. vinsonii subsp. berkhoffii.
PCR-restriction fragment length polymorphism (PCR-RFLP) analysis of the gltA and 16S rRNA genes for these
31 isolates yielded similar profiles that were identical to those of B. vinsonii subsp. berkhoffii. Partial sequencing
of the gltA and 16S rRNA genes, respectively, indicated 99.5 and 100% homology between the coyote isolate and
B. vinsonii subsp. berkhoffii (ATCC 51672). PCR-RFLP analysis of the 16S-23S intergenic spacer region showed
the existence of two different strain profiles, as has been reported in dogs. Six (19%) of 31 Bartonella bacteremic
coyotes exhibited the strain profile that was identified in the type strain of a canine endocarditis case (B.
vinsonii subsp. berkhoffii ATCC 51672). The other 25 bacteremic coyotes were infected with a strain that was
similar to the strains isolated from healthy dogs. Based on whole bacterial genome analysis by pulsed-field gel
electrophoresis (PFGE) with SmaI restriction endonuclease, there was more diversity in fingerprints for the
coyote isolates, which had at least 10 major variants compared to the two variants described for domestic dog
isolates from the eastern United States. By PFGE analysis, three Bartonella bacteremic coyotes were infected
by a strain identical to the one isolated from three healthy dog carriers. Further studies are necessary to
elucidate the mode of transmission of B. vinsonii subsp. berkhoffii, especially to identify potential vectors, and
to determine how humans become infected.
Bartonella species are emerging pathogens in human beings
and cause severe diseases in immunocompromised patients. At
least six Bartonella species are known to be pathogenic for
humans: B. bacilliformis, B. quintana, B. henselae, B. elizabe-
thae, B. grahamii, and B. vinsonii subsp. arupensis (2, 27, 50).
Among these six species, B. quintana, B. henselae, and B. eliza-
bethae have been identified as causative agents of human en-
docarditis (2, 18, 19, 48).
Recently, several new Bartonella species have been isolated
from rodents (20, 34; R. J. Birtles, E. Fichet-Calvet, D. Raoult,
and R. W. Ashford, 13th Sesqui-Annu. Meet. Am. Soc. Rick-
ettsiol. abstr. 34, 1997; R. Heller, M. Kubina, G. Delacour, I.
Mahoudeau, F. Lamarque, M. Artois, H. Monteil, B. Jaulhac,
and Y. Piemont, Abstr. 97th Gen. Meet. Am. Soc. Microbiol.
abstr. B-505, p. 115, 1997), carnivores (32; B. B. Chomel, R. W.
Kasten, C. C. Chang, K. Yamamoto, R. Heller, S. Maruyama,
H. Ueno, D. Simpson, S. S. Swift, Y. Piemont, and N. C.
Pedersen, Abstr. Int. Conf. Emerg. Infect. Dis. vol. 1, p. 21.10,
1998), and wild cervids (12; Chomel et al., Abstr. Int. Conf.
Emerg. Infect. Dis., 1998; R. Heller, M. Kubina, G. Delacour,
F. Lamarque, G. Van Laere, R. Kasten, B. Chomel, and Y.
Piemont, Abstr. Int. Conf. Emerg. Infect. Dis. p. 21.18, 1998).
Furthermore, it is likely that other mammals may also serve as
reservoirs for zoonotic Bartonella spp., involving various vec-
tors for transmission. B. vinsonii subsp. vinsonii, isolated from
a Canadian vole (3), has not yet been identified as a pathogen
in either humans or animals. However, B. vinsonii subsp. aru-
pensis was isolated from a cattle rancher with high fever and
neurological symptoms (50). Recently, B. vinsonii subsp. berk-
hoffii was added to the increasing list of zoonotic Bartonella, as
a human case of endocarditis was associated with this infec-
tious agent, based on sequencing of the gltA and 16S rRNA
genes (43). This agent had been shown to cause endocarditis,
arrhythmia, and myocarditis in dogs (9, 10, 32). Kordick and
Breitschwerdt (31) further identified two different digestion
profiles of B. vinsonii subsp. berkhoffii in dogs, based on the
PCR-restriction fragment length polymorphism (PCR-RFLP)
analysis of the 16S-23S intergenic spacer (ITS) region using
HaeIII restriction endonuclease.
Limited studies have been performed on the epidemiology
of this infection in dogs. According to a serosurvey by Pappa-
lardo et al. (38) involving approximately 2,000 sick dogs from
North Carolina and Virginia, 3.6% of the dogs tested were
* Corresponding author. Mailing address: Department of Popula-
tion Health and Reproduction, School of Veterinary Medicine, Uni-
versity of California, Davis, CA 95616. Phone: (530) 752-8112. Fax:
(530) 752-2377. E-mail: email@example.com.
on June 4, 2013 by guest
seropositive for B. vinsonii subsp. berkhoffii. Because of the low
antibody prevalence in domestic dogs, a very small proportion
of dogs should be Bartonella bacteremic. Therefore, dogs are
not likely to be the main reservoir for B. vinsonii subsp. berk-
hoffii. The fact that Bartonella organisms are very fastidious
bacteria and sometimes are difficult to be isolated by routine
laboratory methods (8) may also explain the very small number
of B. vinsonii subsp. berkhoffii isolates from dogs. Furthermore,
a certain proportion of Bartonella-infected dogs may have been
misdiagnosed by blood culture because of sensitive culture
techniques or concurrent antibiotic treatment.
It is still unknown how this infectious agent is transmitted.
It has been suggested that ticks could be potential vectors for
B. vinsonii subsp. berkhoffii transmission in dogs (38). However,
which vectors or reservoirs are involved in B. vinsonii subsp.
berkhoffii transmission are still unknown and need to be ex-
Because coyotes (Canis latrans) are genetically close to do-
mestic dogs, these wild canids have been shown to be suscep-
tible to or used as sentinels for several viruses (16, 17, 22),
bacteria (11, 51), and parasites (39) that infect domestic dogs.
Following the coyote bite of a child who developed clinical
signs compatible with Bartonella infection in Santa Clara
County, Calif., the child and trapped coyotes were serologically
tested for possible Bartonella infection (13). The identification
of Bartonella-seropositive coyotes prompted us to investigate if
coyotes from central coastal California could serve as a poten-
tial reservoir of B. vinsonii subsp. berkhoffii. Molecular ap-
proaches were applied to determine the characteristics of Bar-
tonella isolates from these animals.
MATERIALS AND METHODS
Sample collection and isolation and identification of B. vinsonii subsp. berk-
hoffii. A sample size of 100 coyotes was determined as necessary for a 95%
confidence interval (CI) with a 10% error for a 50% prevalence estimate. Be-
tween June 1997 and October 1998, a total of 109 coyotes were trapped and
euthanized from nine different sites in central coastal California with the help of
the Santa Clara County Department of Health Services, Wildlife Unit, Vector
Control Section (54 coyotes in 1997 and 55 coyotes in 1998). Blood samples were
collected intracardially in plastic 2-ml EDTA tubes (Becton Dickinson, Franklin
Lakes, N.J.) and frozen at ?70°C until plated. The blood samples were cultured
on heart infusion agar containing 5% rabbit blood and incubated in 5% CO2at
35°C for up to 4 weeks. Identification of the isolates was based on morphological
characteristics and growth time on the blood agar plates and then determined by
PCR-RFLP analysis of the citrate synthase (gltA), 16S rRNA, and 16S-23S ITS
genes. The former two genes were used for comparison because they evolved
slowly enough to allow the use of primers to amplify conserved sequences in
different strains of Bartonella organisms, yet these genes have regions of diversity
that allow for Bartonella species comparison (6, 7, 42). The 16S-23S ITS gene was
used for subtyping of B. vinsonii subsp. berkhoffii as previously described (31).
(i) PCR-RFLP procedures. Isolates were analyzed using PCR-RFLP analysis
of the gltA gene (37, 40), the 16S rRNA gene (24), and 16S-23S ITS gene (45),
as previously described. After approximately 2 cm2of confluent growth was
scraped off and suspended in 100 ?l of sterile water, the bacterial suspension was
heated at 100°C for 15 min and then centrifuged at 15,000 ? g for 10 min at 4°C.
Finally, the supernatant diluted 1:10 was used as the DNA template. An approx-
imately 400-bp fragment of the gltA gene, 1,500-bp fragment of the 16S rRNA
gene, and 2,900-bp fragment of the 16S-23S ITS gene were amplified and then
verified by gel electrophoresis. The amplified product of the gltA gene obtained
with the set of primers suggested by Regnery et al. (40) was digested with TaqI
(Promega, Madison, Wis.) and HhaI (new England BioLabs, Beverly, Mass.)
restriction endonucleases. TaqI and MseI (New England BioLabs) restriction
endonucleases were utilized when using the set of primers suggested by Norman
et al. (37). The amplified product of the 16S rRNA gene was digested with DdeI
(Boehringer GmbH, Mannheim, Germany) and MnlI (New England BioLabs)
restriction endonucleases. The digestion conditions used were the ones recom-
mended by the enzymes’ manufacturer. Banding patterns were compared with
those of a domestic dog isolate (American Type Culture Collection [ATCC]
51672) of B. vinsonii subsp. berkhoffii), B. vinsonii subsp. vinsonii (ATCC VR152),
B. henselae (strain U-4; University of California, Davis) and B. clarridgeiae
(ATCC 51734); the last two Bartonella species are usually isolated from domestic
cats. Finally, the amplified product of the 16S-23S ITS gene was digested with
HaeIII restriction endonuclease (Boehringer GmbH) (31).
(ii) DNA sequencing. The PCR products used for DNA sequencing were
purified with Microcon centrifugal filter devices (Millipore Corp., Bedford,
Mass.) and sequenced with a fluorescence-based automated sequencing system
(Davis Sequencing, Davis, Calif.). Primers BhCS.1137n (5?-AATGCAAAAAG
AACAGTAAACA-3?) (37) and Pc1544 (5?-AAGGAGGTGATCCAGCCGCA-
3?) (25) were used for partial sequencing of the gltA and 16S rRNA genes,
IFA test. B. henselae indirect immunofluorescent-antibody (IFA) test was
performed as previously described (14). For IFA slides using B. vinsonii subsp.
berkhoffii as the antigen, the reference strain (ATCC 51672) was cultured for 4
days on heart infusion agar containing 5% rabbit blood. The bacteria were
harvested into 0.5 ml of sterile saline, washed twice in 0.5 ml of sterile saline, and
finally resuspended in 0.5 ml of sterile saline. The bacteria were heat inactivated
at 55°C for 30 min and then rewashed twice in 0.5 ml of sterile saline. The final
pellet was resuspended in 0.5 ml of sterile saline.
A 90% confluent tissue culture flask (Felis catus whole fetus) was inoculated
with the resuspended B. vinsonii subsp. berkhoffii, and the flask was incubated for
3 days at 37°C with 5% CO2. After incubation, the tissue cultures were washed
two times with calcium- and magnesium-free phosphate-buffered saline (PBS)
(Gibco-BRL, Gaithersburg, Md.) and trypsinized (Gibco-BRL) for 10 min at
room temperature. The suspended tissue cultures were combined into one tube
and centrifuged at 200 ? g for 10 min. The supernatant was discarded, and the
cells were resuspended in 30 ml of tissue culture growth medium. Forty micro-
liters of the cell culture were spotted onto HTC supercured glass slides (12-well
slides; Cell-Line/Erie Scientific, Co., Newfield, N.J.) and incubated overnight at
37°C with 5% CO2. The slides were then washed twice in PBS (pH 7.4) (Sigma
Chemical, St. Louis, Mo.), set for 20 min in acetone at room temperature, air
dried, and then stored at ?20°C until they were used. Supernatant of the whole
blood collected in the EDTA tubes after centrifugation was used for serological
testing. Samples added to the test wells were initially screened at 1:32 and 1:64
dilutions in PBS with 5% milk. The slides were then incubated for 35 min at 37°C
and were washed for 5 min in PBS twice. Fluorescein-conjugated goat anti-dog
immunoglobulin (whole-molecule immunoglobulin G; Organon Teknika Corp.,
Durham, N.C.) was diluted at 1:1,400 in PBS with 5% milk containing 0.001%
Evan’s blue, and the mixture was applied to each well. The slides were incubated
for 20 min at 37°C and washed again in PBS for 5 min twice prior to being read
with a fluorescence microscope (magnification, ?400). The intensity of bacillus-
specific fluorescence was scored subjectively from 1 to 4, and a fluorescence score
of ?2 at a dilution of 1:64 was reported as a positive result, as previously
described (38). Any sample positive at 1:64 was titrated in serial twofold dilutions
to the end point. A double-blind reading of each slide was performed by the same
two readers. Negative and positive serum control samples were obtained from
two laboratory dogs before and after they were infected with B. vinsonii subsp.
PFGE. Four canine B. vinsonii subsp. berkhoffii strains isolated at North Caro-
lina State University (including the reference strain, ATCC 51672) were included
for comparison with the coyote isolates in the present study. For pulsed-field gel
electrophoresis (PFGE), a single colony pick of each Bartonella isolate was
subcultured confluently on 5% rabbit agar plate at 35°C for 5 to 7 days in a 5%
CO2incubator. The bacteria grown on the agar plates were scraped off, sus-
pended in sterile saline, and washed twice by centrifugation at 15,000 ? g for 5
min at 4°C. The turbidity of the suspension was adjusted to McFarland standard
6. Then, 0.5 ml of the adjusted suspension was mixed gently but thoroughly with
the same amount of 2% ultrapure low-melting-point agarose (Gibco-BRL, Life
Technologies, Inc., Gaithersburg, Md.), and the mixture was solidified in plug
molds at 4°C. The agarose plugs were then transferred into lysozyme solution (10
mM Tris [pH 7.2], 50 mM NaCl, 0.2% sodium deoxycholate, 0.5% sodium lauryl
sarcosine, 1 mg of lysozyme per ml) and incubated at 37°C overnight. The plugs
were rinsed with sterile water and incubated in proteinase K solution (100 mM
EDTA [pH 8.0], 0.2% sodium deoxycholate, 1% sodium lauryl sarcosine, 1 mg of
proteinase K per ml) at 50°C overnight. This procedure was repeated a second
time. Then, the plugs were washed four times in 10 ml of washing buffer (50 mM
EDTA, 20 mM Tris [pH 8.0]) for 1 h at room temperature with gentle agitation.
Proteinase K was inactivated by the addition of 1 mM phenylmethylsulfonyl
fluoride solution during the second wash. The plugs were stored in wash buffer
at 4°C before endonuclease digestion. Before digestion, plugs were transferred to
1.5-ml sterile microcentrifuge tubes with 0.1? washing buffer at 4°C overnight
and they were equilibrated in 1? endonuclease-specific reaction buffer for 1 h.
SmaI and NotI restriction endonucleases (45) (New England BioLabs) were used
for the analysis of the whole Bartonella genome. Bacterial DNA was digested in
reaction buffer with SmaI endonuclease at 28°C and with NotI endonuclease at
37°C overnight. After digestion, plugs were equilibrated in 0.5? TBE (45 mM
Tris-borate, 1 mM EDTA [pH 8.0]) buffer for 30 min. The chromosomal restric-
tion fragments were separated by PFGE in a CHEF-DRIII system (Bio-Rad,
Hercules, Calif.) by a 1.5% (for SmaI digestion) or 1.0% (for NotI digestion)
pulsed-field certified agarose (Bio-Rad) gel in 0.5? TBE buffer. The electro-
phoresis was equilibrated at 14°C for 26 h at a constant voltage of 5.7 V/cm for
the SmaI-digested plugs and for 33 h at a constant voltage of 4.5 V/cm for the
NotI-digested plugs. Separation of the digested genomic DNA was achieved with
pulse times from 3 to 10 s for SmaI-digested plugs and from 5 to 120 s for
NotI-digested plugs, respectively. After electrophoresis, the gel was stained with
0.5 ?g of ethidium bromide per ml for 30 min, destained with distilled water
4194 CHANG ET AL.J. CLIN. MICROBIOL.
on June 4, 2013 by guest
twice for 15 min each time, and photographed. Lambda ladder pulsed-field gel
markers (48.5 to 970 kbp) (Bio-Rad) were used as molecular size standards.
Three B. vinsonii subsp. berkhoffii strains isolated from healthy dogs (NC95-C02,
NC95-C03, and NC95-C04) and ATCC strain 51672 isolated from a dog with
endocarditis were included in PFGE analysis.
(i) Analysis of PFGE profiles. The PFGE profiles were analyzed by the direct
grouping method suggested by Tenover et al. (49). In addition, cluster analysis
with Molecular Analyst Software (Fingerprinting version 1.12; Bio-Rad) was
performed. The images were processed, and then Jaccard coefficients (SJ) of
band-based similarity were calculated as SJ? NAB/(NA? NB? NAB), where NAB
is the number of bands common to A and B, NAis the total number of bands in
A, and NBis the total number of bands in B. Dendrograms based on results of the
matrix of similarity values were created with unweighted-pair group method
using average linkage clustering.
(ii) Statistical analysis. The data were analyzed by Epi-Info version 6.03. The
chi-square test for homogeneity was used to evaluate the association between a
disease status (bacteremia or seropositivity) and a categorized risk factor, and
then P values were calculated using Yates corrected method or two-tailed Fish-
er’s exact test (for analyses with expected numbers of observations of less than
five). Mantel extension test for trend was also applied to evaluate for the exis-
tence of a seasonal trend for bacteremia and antibody prevalence. The associa-
tion between seropositivity and bacteremia for Bartonella was evaluated by Mc-
Nemar’s test for paired analysis.
Epidemiological patterns of B. vinsonii subsp. berkhoffii in-
fection in coyotes. Of 109 coyotes, 31 (28%; 95% CI, 20 to
38%) were positive for Bartonella spp. by blood culture. The
seroprevalence for B. vinsonii subsp. berkhoffii was 76% (83 of
109) (95% CI, 67 to 84%). There was no significant difference
between the prevalence of bacteremia in 1997 (30% [16 of 54])
and in 1998 (27% [15 of 55]) (P ? 0.95). The seroprevalence
was also similar in 1997 and 1998 (72 versus 80% [P ? 0.47]).
Only 10 coyotes less than 1 year old were trapped in this study.
The prevalence of Bartonella bacteremia was significantly
lower in adult coyotes (25%) than in coyotes less than 1 year
old (60%) (Table 1). Conversely, the seroprevalence of Bar-
tonella infection in adult coyotes (91%) was higher than in
coyotes less than 1 year old (60%) (Table 1). There was no
statistically significant prevalence difference for either bacte-
remia or antibodies between male and female coyotes.
Using the blood collection date to investigate seasonal dif-
ferences for the prevalence of Bartonella bacteremia, the low-
est prevalence was observed during winter, followed by sum-
mer and fall. The highest prevalence was seen during spring.
This increasing trend of Bartonella bacteremic prevalence was
statistically significant (P ? 0.05). The trend of seroprevalence
was increased from summer, spring, fall, to winter (P ? 0.05).
The bacteremia and antibody prevalences varied by collection
sites (Table 2). There was no significant association between
age and collection periods or between age and collection sites.
There was no significant association between seropositivity
and bacteremia for Bartonella in coyotes. Twenty-four Bar-
tonella bacteremic coyotes had antibody titers ranging from
1:64 (9 coyotes), 1:128 (5 coyotes), to 1:256 (10 coyotes). How-
ever, seven of the bacteremic coyotes were seronegative (titer
of ?1:32). Of the 78 nonbacteremic coyotes, 59 (75.6%) had B.
vinsonii subsp. berkhoffii antibodies with the following titers:
1:64 (32 coyotes), 1:128 (12 coyotes), 1:256 (13 coyotes), and
1:152 (2 coyotes). Therefore, the positive and negative predic-
tive values of the B. vinsonii subsp. berkhoffii serological assay
for bacteremia status were 29% (24 of 83) and 73% (19 of 26),
respectively. Sixty-three coyotes that were seropositive for B.
vinsonii subsp. berkhoffii were also seropositive for B. henselae
(Table 3). However, thirteen coyotes that were seronegative
for B. vinsonii subsp. berkhoffii were seropositive for B. henselae
PCR-RFLP-based typing. All 31 suspected Bartonella iso-
lates from coyotes were tested by PCR of the gltA gene with
two different sets of primers that are specific for Bartonella
species (37, 40). Before digestion by the restriction endonucle-
ases, a 400-bp band specific for Bartonella spp. was identified
for all 31 coyote isolates by both sets of primers. However,
extra 700- and 190-bp bands were also observed for all isolates
and B. vinsonii subsp. berkhoffii ATCC 51672 with the primers
suggested by Regnery et al. (40) (Fig. 1 and 2). The molecular
pattern of all coyote isolates was identical to that of a domestic
dog isolate of B. vinsonii subsp. berkhoffii (ATCC 51672), based
on PCR-RFLP analysis of the gltA gene with TaqI or HhaI
digestion (Fig. 1) and 16S rRNA gene with DdeI and MnlI
TABLE 1. Prevalence of B. vinsonii subsp. berkhoffii bacteremia
and seropositivity by age, sex, and collection period
for 109 coyotes from central coastal California
?1 yr old
?1 yr old
Spring (Mar. to May)
Summer (June to Aug.)
Fall (Sept. to Nov.)
Winter (Dec. to Feb.)
aPrevalence shown as a percentage, with the number of coyotes with bacte-
remia (or the number of seropositive coyotes) and the total number of coyotes
shown in the parentheses.
bChi-square test for homogeneity.
cNot available for one coyote.
TABLE 2. Bacteremia and antibody prevalences of B. vinsonii
subsp. berkhoffii infection in coyotes from central
coastal California by capture sites
aPrevalence shown as a percentage, with the number of coyotes with bacte-
remia (or the number of seropositive coyotes) and the total number of coyotes
shown in the parentheses.
TABLE 3. Distribution of anti-B. vinsonii subsp. berkhoffii and
anti-B. henselae antibody titers by IFA in coyotes
Anti-B. vinsonii subsp. berkhoffii antibody titer
aAn antibody titer of ?1:64 was reported as seropositive.
VOL. 38, 2000 B. VINSONII SUBSP. BERKHOFFII IN CALIFORNIA COYOTES4195
on June 4, 2013 by guest
digestion (Fig. 3). However, these isolates yielded different
patterns from that of B. vinsonii subsp. vinsonii (ATCC
VR152) when PCR-RFLP analysis of the gltA gene was per-
formed (Fig. 1). The PCR product with the set of primers
suggested by Norman et al. (37) could not be digested by HhaI
restriction endonuclease; however, the TaqI- and MseI-di-
gested profiles were identical for all isolates (Fig. 2). The
partial 16S rRNA gene sequences further showed a 100%
homology between the coyote isolate sequenced and a domes-
tic dog isolate (ATCC 51672). By partial sequencing of the gltA
gene, there was a 99.5% homology between the coyote isolate
and ATCC strain 51672.
By PCR-RFLP analysis of the 16S-23S interspacer region, 6
(19%) of 31 Bartonella bacteremic coyotes were infected with
B. vinsonii subsp. berkhoffii type I strain, similar to the profile
of ATCC strain 51672 (Fig. 4). The other 25 Bartonella bacte-
remic coyotes were all infected with B. vinsonii subsp. berkhoffii
type II strain, which has been isolated from healthy dogs.
There was no major clustering of these two types by capture
PFGE-based typing. Ten coyote isolates were not digested
by NotI restriction enzyme, and only a few high-molecular-
weight bands were observed for the remaining 21 isolates after
digestion. Therefore, PFGE profiles with SmaI digestion were
used for cluster analysis. When a Jaccard coefficient of 70%
was applied for grouping the profiles, the fingerprints with
SmaI identified 11 major patterns for B. vinsonii subsp. berk-
hoffii isolates, including ATCC strain 51672, which had a
unique pattern (Fig. 5). Based on the fingerprints by SmaI
digestion, no clonal distribution according to the place or year
of collection was observed for the coyote isolates (Fig. 5).
Compared to the 16S-23S ITS fingerprints, five of the six coy-
otes infected with type I were grouped with the PFGE pattern
VII and one type I isolate belonged to the PFGE pattern X.
We found 28% of 109 coyotes from central coastal Califor-
nia to be bacteremic with B. vinsonii subsp. berkhoffii, previ-
ously isolated from domestic dogs. The prevalence of bactere-
mia was higher among coyotes ?1 year old than among adult
coyotes, as previously reported for B. henselae infection in cats
(14). The high prevalence (76%) of B. vinsonii subsp. berkhoffii
antibodies in these 109 coyotes was also consistent with the
previous findings of a serosurvey of California coyotes (13).
Seven coyotes that were Bartonella bacteremic were seroneg-
ative for Bartonella, possibly because of recently acquired in-
fection; three coyotes were less than 1 year old, and four
coyotes were adults. Similarly, cats with Bartonella bacteremia
but without detectable antibodies have been previously re-
PCR-RFLP analysis of the gltA and 16S rRNA genes showed
FIG. 1. PCR (lanes 2 to 5) and PCR-RFLP (lanes 6 to 9, TaqI digestion; lanes 10 to 13, HhaI digestion) analyses of the gltA gene of coyote isolates with the set
of primers suggested by Regnery et al. (40). Lanes 1 and 14, standard 100-bp molecular ladder; lanes 2, 6, and 10, coyote isolates; lanes 3, 7, and 11, B. vinsonii subsp.
berkhoffii ATCC 51672; lanes 4, 8, and 12, B. vinsonii ATCC VR152; lanes 5, 9 and 13, B. henselae (strain U-4, University of California, Davis).
FIG. 2. PCR-RFLP analysis (lanes 2 to 11, TaqI digestion; lanes 12 to 21, MseI digestion) of the gltA gene of coyote isolates with the set of primers suggested by
Norman et al. (37). Lanes 1 and 22, standard 100-bp molecular ladder; lanes 2 to 9 and 12 to 19, coyote isolates; lanes 10 and 20, B. vinsonii subsp. berkhoffii ATCC
51672; lanes 11 and 21, B. henselae (strain U-4; University of California, Davis).
4196CHANG ET AL. J. CLIN. MICROBIOL.
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that the PCR-RFLP profiles of all 31 coyote isolates were
identical to that of a domestic dog isolate of B. vinsonii subsp.
berkhoffii (ATCC 51672) but were different from the PCR-RFLP
profiles of the other Bartonella strains tested: B. henselae, B.
clarridgeiae, B. bacilliformis, B. quintana, B. elizabethae, and B.
vinsonii subsp. vinsonii (data not shown). The partial sequenc-
ing of the gltA and 16S rRNA genes, respectively, indicated
99.5 and 100% homology between the coyote isolate and B.
vinsonii subsp. berkhoffii reference strain (ATCC 51672). There
were two different molecular types (designated type I and type
II) of B. vinsonii subsp. berkhoffii using PCR-RFLP analysis of
the 16S-23S ITS region as previously reported for domestic
dogs (31); type I was isolated from a dog with endocarditis, and
type II was isolated from three healthy dogs. Unfortunately,
PCR-RFLP analysis of the 16S-23S ITS was not conducted in
the human endocarditis case caused by B. vinsonii subsp. berk-
hoffii (43), which did not allow comparison with these canine
isolates. In our study, only 19% of 31 Bartonella bacteremic
coyotes were infected with the type I strain, i.e., similar to the
strain isolated from a dog with endocarditis. Further studies
should aimed at determining if the type I strain is specifically
associated with canine and human endocarditis cases.
Because of the large number of B. vinsonii subsp. berkhoffii
isolates obtained from coyotes, we were able to investigate, for
the first time, the molecular diversity of these isolates and
domestic dog isolates by analysis of the whole bacterial ge-
nome using PFGE. PFGE, using SmaI endonuclease, allowed
us to determine 11 variants, including a unique pattern for
ATCC 51672 strain. By PFGE, three coyotes were infected
with a B. vinsonii subsp. berkhoffii strain similar to the strains
isolated from three healthy dogs, but none of the coyotes were
infected with strains identical to the ATCC 51672 strain. The
wide diversity of B. vinsonii subsp. berkhoffii PFGE profiles
could be caused by gene mutation or translocation under nat-
ural circumstances that potentially enhance bacterial transmis-
sibility or pathogenicity. In contrast, NotI endonuclease was
not an appropriate enzyme for differentiating these isolates
because it very infrequently cut B. vinsonii subsp. berkhoffii, as
previously reported by Roux and Raoult (45) for B. henselae
isolates. As has been suggested in other bacterial studies (49),
PFGE analysis appeared to be a more discriminating method
in differentiating B. vinsonii subsp. berkhoffii variants in canids
than PCR-RFLP analysis. In the future, however, it will be of
interest to determine if differences in pathogenicity for canids
will be better identified by PCR-RFLP analysis of the 16S-23S
ITS region or by PFGE fingerprints.
Detection and identification of Bartonella organisms have
been done mainly by PCR-RFLP analysis and/or sequencing
targeting the gltA gene (37, 40), 16S rRNA gene (5, 18, 29, 35,
40, 42), and/or 16S-23S ITS region (31, 36, 44; G. M. Matar,
Letter, J. Clin. Microbiol. 33:3370, 1995). The low rate of
evolutionary sequence divergence of the 16S rRNA gene is
useful for designing primers targeting conserved sequences for
broad-range PCR. Currently, sequencing data of the 16S
rRNA gene for more than 2,000 bacterial species is available in
GenBank. However, it could be difficult to design the genus-
specific primers that are able to specifically amplify one bac-
terial genus, e.g., Bartonella, for rapid diagnosis. It has been
shown that the sequence of the gltA gene is less conserved than
that of the 16S rRNA gene within the genus Bartonella (7),
FIG. 3. PCR-RFLP analysis of the 16S rRNA gene of coyote isolates with
DdeI (A) or MnlI (B) restriction endonuclease. Lanes 1 to 12, coyote isolates;
lane 13, B. vinsonii subsp. berkhoffii ATCC 51672; lane 14, 100-bp molecular
FIG. 4. PCR-RFLP analysis of the 16S-23S ITS region of coyote isolates with HaeIII restriction endonuclease. Lanes 1 and 15, standard 100-bp molecular ladder;
lanes 2 to 10 and 13, coyote isolates (type II); lanes 11 and 12, coyote isolates (type I); lane 14, B. vinsonii subsp. berkhoffii ATCC 51672 (type I).
VOL. 38, 2000 B. VINSONII SUBSP. BERKHOFFII IN CALIFORNIA COYOTES 4197
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facilitating the design of Bartonella-specific primers for PCR
amplification. We suggest using PCR-RFLP analysis and/or
sequencing of the gltA gene as a first step in confirmation and
identification of Bartonella species and to generate phylogenic
trees for the Bartonellaceae family, as done by Kosoy et al. (34).
PCR-RFLP and/or sequencing of the 16S rRNA gene could be
used in a second step for analysis of phylogenic relationships
with the other closely related bacterial genera (41, 42). Finally,
PCR-RFLP and/or sequencing of the 16S-23S ITS gene or
PFGE could be applied for subspecies identification (45, 47).
FIG. 5. (A) Fingerprints of B. vinsonii subsp. berkhoffii isolates by PFGE with SmaI digestion. Lane M, molecular size markers; lanes 2 to 11, B. vinsonii subsp.
berkhoffii patterns I to X of the coyote isolates; lane 12, ATCC strain 51672 (pattern XI); lanes 13 to 15, isolates from three healthy dogs (pattern II). (B) Dendrogram
of the fingerprints as determined by unweighted-pair group method using average linkage clustering. NC95-C02, NC95-C03, and NC95-C04 are B. vinsonii subsp.
berkhoffii isolates from three healthy dogs. ID, identification.
4198 CHANG ET AL. J. CLIN. MICROBIOL.
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When preparing IFA slides, it was observed that B. vinsonii
subsp. berkhoffii-infected F. catus whole fetus cells were prone
to clump together, making the preparation of IFA slides dif-
ficult. We found that heat inactivation during antigen prepa-
ration could prevent cell clumping and improve the quality of
IFA tests for detection of B. vinsonii subsp. berkhoffii antibod-
Sixty-three coyotes which were seropositive for B. vinsonii
subsp. berkhoffii were also seropositive for B. henselae, but
thirteen coyotes that were seronegative for B. vinsonii subsp.
berkhoffii were seropositive for B. henselae. These results may
be related to cross-reactivity between various Bartonella spe-
cies, which has been reported in humans and animals (4, 21,
26). Moreover, it is still unknown if canids can be naturally
infected with B. henselae, isolated at present only from felids
and humans, and become seropositive. However, all 31 Bar-
tonella bacteremic coyotes were found to be infected only with
B. vinsonii subsp. berkhoffii, not with B. henselae or any other
Compared to B. vinsonii subsp. berkhoffii infection in domes-
tic dogs (38), the significantly higher bacteremia and antibody
prevalences in coyotes implies that coyotes might be a wildlife
reservoir. The ability to maintain a particular infectious agent
for a long period of time and the high prevalence of infection
with this infectious agent in a given animal species are the
characteristics of animal reservoirs, as illustrated for cats and
B. henselae. The high prevalence of B. vinsonii subsp. berkhoffii
infection found in coyotes also suggests that these animals
could serve as a potential reservoir for B. vinsonii subsp. berk-
hoffii. Repeated isolation of the infectious agent from captured
wild coyotes provides presumptive evidence of reservoir com-
petency, although this capacity was not fully demonstrated by
the present study.
In our laboratory, experimental inoculation of two domestic
dogs by the intradermal route with a coyote isolate led to a
prolonged bacteremia (at least 8 weeks) (B. B. Chomel et al.,
unpublished data). As seen for B. henselae infection in cats,
which can remain bacteremic for periods ranging from several
months to years (28, 30, 46), both dogs had a high level of
Bartonella bacteremia for a few weeks after infection but with-
out fever and clinical signs. These data suggest that, in addition
to the capacity to be infected, canids can maintain levels of
bacteremia for periods long enough to allow possible arthro-
pod transmission. However, this cross-sectional study did not
allow us to determine the duration of B. vinsonii subsp. berk-
hoffii bacteremia in wild coyotes. Coyotes could be a source of
infection for domestic dogs, especially when Bartonella enzo-
otic coyote populations have an overlapping home range with
Modes of transmission between coyotes and domestic dogs
could be either by mechanical means (biting and scratching) or
through arthropod vectors. However, Bartonella spp. are usu-
ally transmitted by arthropod vectors (2). B. bacilliformis, the
agent of Carrio ´n’s disease, mainly found in the Andes moun-
tains, is transmitted by sand flies. B. quintana, which is trans-
mitted by the human body louse, causes trench fever. Cats are
the main reservoir for B. henselae (28), and cat fleas are a
competent vector for the transmission between cats (15). No
direct transmission of B. henselae from cat to cat has been
documented in experimental settings (1, 23). The seasonal
fluctuation of Bartonella bacteremic prevalence in coyotes
from central coastal California observed in this study could
possibly be related to arthropod activity. Based on our previous
study, the clustered distribution of higher Bartonella seropreva-
lence in coyotes from coastal California compared to coyotes
from the inland regions also suggested that the geographical
distribution of this Bartonella infection in coyotes could be
associated with the presence of certain arthropod species, such
as ticks or sand flies (13). We therefore hypothesize that B.
vinsonii subsp. berkhoffii transmission within coyotes and be-
tween coyotes and dogs could be arthropodborne. Until now,
investigations of which vectors are competent for B. vinsonii
subsp. berkhoffii transmission among dogs have been very lim-
ited. Tickborne infection has been suggested for dogs from the
eastern United States (38). To confirm this hypothesis, one
needs to show vector competency by allowing ticks to feed on
experimentally infected dogs and/or coyotes to determine if
ticks are able to acquire B. vinsonii subsp. berkhoffii infection
and further transmit this agent to noninfected canids. Further-
more, testing of field-collected tick samples should be per-
Dogs and possibly humans could become infected during
recreational activities through bites of infected arthropods that
may have fed on Bartonella bacteremic coyotes. The recent
identification of B. vinsonii subsp. berkhoffii in a human endo-
carditis case (43) warrants further investigation to elucidate
the mode of transmission of B. vinsonii subsp. berkhoffii, espe-
cially to identify potential vectors, and to determine how hu-
mans become infected.
We thank Didier Raoult for kindly sharing unpublished data on the
human case of Bartonella vinsonii subsp. berkhoffii endocarditis. We
also thank Ian Gardner (Department of Medicine and Epidemiology,
School of Veterinary Medicine, University of California, Davis) and
Robert S. Lane (Department of Environmental Science, Policy and
Management, College of Natural Resources, University of California,
Berkeley) for suggestions and help in preparing this manuscript.
1. Abbott, R. C., B. B. Chomel, R. W. Kasten, K. A. FloydHawkins, Y. Kikuchi,
J. E. Koehler, and N. C. Pedersen. 1997. Experimental and natural infection
with Bartonella henselae in domestic cats. Comp. Immunol. Microbiol. Infect.
2. Anderson, B. E., and M. A. Neuman. 1997. Bartonella spp. as emerging
human pathogens. Clin. Microbiol. Rev. 10:203–219.
3. Baker, J. A. 1946. A rickettsial infection in Canadian voles. J. Exp. Med.
4. Baneth, G., D. L. Kordick, B. C. Hegarty, and E. B. Breitschwerdt. 1996.
Comparative seroreactivity to Bartonella henselae and Bartonella quintana
among cats from Israel and North Carolina. Vet. Microbiol. 50:95–103.
5. Bergmans, A. M., J. F. Schellekens, J. D. van Embden, and L. M. Schouls.
1996. Predominance of two Bartonella henselae variants among cat-scratch
disease patients in The Netherlands. J. Clin. Microbiol. 34:254–260.
6. Birtles, R. J. 1995. Differentiation of Bartonella species using restriction
endonuclease analysis of PCR-amplified 16S rRNA genes. FEMS Microbiol.
7. Birtles, R. J., and D. Raoult. 1996. Comparison of partial citrate synthase
gene (gltA) sequences for phylogenetic analysis of Bartonella species. Int. J.
Syst. Bacteriol. 46:891–897.
8. Breathnach, A. S., J. M. Hoare, and S. J. Eykyn. 1997. Culture-negative
endocarditis: contribution of Bartonella infections. Heart 77:474–476.
9. Breitschwerdt, E. B., C. E. Atkins, T. T. Brown, D. L. Kordick, and P. S.
Snyder. 1999. Bartonella vinsonii subsp. berkhoffii and related members of the
alpha subdivision of the Proteobacteria in dogs with cardiac arrhythmias,
endocarditis, or myocarditis. J. Clin. Microbiol. 37:3618–3626.
10. Breitschwerdt, E. B., D. L. Kordick, D. E. Malarkey, B. Keene, T. L. Had-
field, and K. Wilson. 1995. Endocarditis in a dog due to infection with a
novel Bartonella subspecies. J. Clin. Microbiol. 33:154–160.
11. Burgess, E. C., and L. A. Windberg. 1989. Borrelia sp. infection in coyotes,
black-tailed jack rabbits and desert cottontails in southern Texas. J. Wildl.
12. Chang, C. C., B. B. Chomel, R. W. Kasten, R. Heller, K. M. Kocan, H. Ueno,
K. Yamamoto, V. C. Bleich, B. M. Pierce, B. J. Gonzales, P. K. Swift, W. M.
Boyce, S. S. Jang, H. Boulouis, and Y. Piemont. 2000. Bartonella spp. isolated
from wild and domestic ruminants in North America. Emerg. Infect. Dis.
13. Chang, C. C., K. Yamamoto, B. B. Chomel, R. W. Kasten, D. C. Simpson,
C. R. Smith, and V. L. Kramer. 1999. Seroepidemiology of Bartonella vin-
sonii subsp. berkhoffii infection in California coyotes, 1994–1998. Emerg.
Infect. Dis. 5:711–715.
VOL. 38, 2000 B. VINSONII SUBSP. BERKHOFFII IN CALIFORNIA COYOTES4199
on June 4, 2013 by guest
14. Chomel, B. B., R. C. Abbott, R. W. Kasten, K. A. Floyd-Hawkins, P. H. Kass, Download full-text
C. A. Glaser, N. C. Pedersen, and J. E. Koehler. 1995. Bartonella henselae
prevalence in domestic cats in California: risk factors and association be-
tween bacteremia and antibody titers. J. Clin. Microbiol. 33:2445–2450.
15. Chomel, B. B., R. W. Kasten, K. Floyd-Hawkins, B. Chi, K. Yamamoto, J.
Roberts-Wilson, A. N. Gurfield, R. C. Abbott, N. C. Pedersen, and J. E.
Koehler. 1996. Experimental transmission of Bartonella henselae by the cat
flea. J. Clin. Microbiol. 34:1952–1956.
16. Clark, K. A., S. U. Neill, J. S. Smith, P. J. Wilson, V. W. Whadford, and G. W.
McKirahan. 1994. Epizootic canine rabies transmitted by coyotes in south
Texas. J. Am. Vet. Med. Assoc. 204:536–540.
17. Cypher, B. L., J. H. Scrivner, K. L. Hammer, and T. P. O’Farrell. 1998. Viral
antibodies in coyotes from California. J. Wildl. Dis. 34:259–264.
18. Daly, J. S., M. G. Worthington, D. J. Brenner, C. W. Moss, D. G. Hollis, R. S.
Weyant, A. G. Steigerwalt, R. E. Weaver, M. I. Daneshvar, and S. P.
O’Connor. 1993. Rochalimaea elizabethae sp. nov. isolated from a patient
with endocarditis. J. Clin. Microbiol. 31:872–881.
19. Drancourt, M., J. L. Mainardi, P. Brouqui, F. Vandenesch, A. Carta, F.
Lehnert, J. Etienne, F. Goldstein, J. Acar, and D. Raoult. 1995. Bartonella
(Rochalimaea) quintana endocarditis in three homeless men. N. Engl.
J. Med. 332:419–423.
20. Ellis, B. A., R. L. Regnery, L. Beati, F. Bacellar, M. Rood, G. G. Glass, E.
Marston, T. G. Ksiazek, D. Jones, and J. E. Childs. 1999. Rats of the genus
Rattus are reservoir hosts for pathogenic Bartonella species: an old world
origin for a new world disease? J. Infect. Dis. 180:220–224.
21. Engbaek, K., and C. Koch. 1994. Immunoelectrophoretic characterization
and cross-reactivity of Rochalimaea henselae, Rochalimaea quintana and
Afipia felis. APMIS 102:931–942.
22. Foreyt, W. J., and J. F. Evermann. 1985. Serologic survey of canine corona-
virus in wild coyotes in the western United States. J. Wildl. Dis. 21:428–430.
23. Guptill, L., L. Slater, C. C. Wu, T. L. Lin, L. T. Glickman, D. F. Welch, and
H. HogenEsch. 1997. Experimental infection of young specific pathogen-free
cats with Bartonella henselae. J. Infect. Dis. 176:206–216.
24. Gurfield, A. N., H. J. Boulouis, B. B. Chomel, R. Heller, R. W. Kasten, K.
Yamamoto, and Y. Piemont. 1997. Coinfection with Bartonella clarridgeiae
and Bartonella henselae and with different Bartonella henselae strains in
domestic cats. J. Clin. Microbiol. 35:2120–2123.
25. Heller, R., M. Artois, V. Xemar, D. De Briel, H. Gehin, B. Jaulhac, H.
Monteil, and Y. Piemont. 1997. Prevalence of Bartonella henselae and Bar-
tonella clarridgeiae in stray cats. J. Clin. Microbiol. 35:1327–1331.
26. Jackson, L. A., D. H. Spach, D. A. Kippen, N. K. Sugg, R. L. Regnery, M. H.
Sayers, and W. E. Stamm. 1996. Seroprevalence to Bartonella quintana
among patients at a community clinic in downtown Seattle. J. Infect. Dis.
27. Kerkhoff, F. T., A. M. C. Bergmans, A. Van Der Zee, and A. Rothova. 1999.
Demonstration of Bartonella grahamii DNA in ocular fluids of a patient with
neuroretinitis. J. Clin. Microbiol. 37:4034–4038.
28. Koehler, J. E., C. A. Glaser, and J. W. Tappero. 1994. Rochalimaea henselae
infection—a new zoonosis with the domestic cat as reservoir. J. Am. Med.
29. Koehler, J. E., F. D. Quinn, T. G. Berger, P. E. Le Boit, and J. W. Tappero.
1992. Isolation of Rochlimaea species from cutaneous and osseous lesions of
bacillary angiomatosis. N. Engl. J. Med. 327:1625–1631.
30. Kordick, D. L., and E. B. Breitschwerdt. 1995. Intraerythrocytic presence of
Bartonella henselae. J. Clin. Microbiol. 33:1655–1656.
31. Kordick, D. L., and E. B. Breitschwerdt. 1998. Persistent infection of pets
within a household with three Bartonella species. Emerg. Infect. Dis. 4:325–
32. Kordick, D. L., B. Swaminathan, C. E. Greene, K. H. Wilson, A. M. Whitney,
S. O’Connor, D. G. Hollis, G. M. Matar, A. G. Steigerwalt, G. B. Malcolm,
P. S. Hayes, T. L. Hadfield, E. B. Breitschwerdt, and D. J. Brenner. 1996.
Bartonella vinsonii subsp. berkhoffii subsp. nov., isolated from dogs; Bar-
tonella vinsonii subsp. vinsonii; and emended description of Bartonella vin-
sonii. Int. J. Syst. Bacteriol. 46:704–709.
33. Kordick, D. L., K. H. Wilson, D. J. Sexton, T. L. Hadfield, H. A. Berkhoff,
and E. B. Breitschwerdt. 1995. Prolonged Bartonella bacteremia in cats
associated with cat-scratch disease patients. J. Clin. Microbiol. 33:3245–3251.
34. Kosoy, M. Y., R. L. Regnery, T. Tzianabos, E. L. Marston, D. C. Jones, D.
Green, G. O. Maupin, J. G. Olson, and J. E. Childs. 1997. Distribution,
diversity, and host specificity of Bartonella in rodents from the southeastern
United States. Am. J. Trop. Med. Hyg. 57:578–588.
35. Lucey, D., M. J. Dolan, C. W. Moss, M. Garcia, D. G. Hollis, and S. Wegner.
1992. Relapsing illness due to Rochalimaea henselae in immunocompetent
host: implication for therapy and new epidemiological associations. Lin.
Infect. Dis. 14:683–688.
36. Matar, G. M., B. Swaminathan, S. B. Hunter, L. N. Slater, and D. F. Welch.
1993. Polymerase chain reaction-based restriction fragment length polymor-
phism analysis of a fragment of the ribosomal operon from Rochalimaea
species for subtyping. J. Clin. Microbiol. 31:1730–1734.
37. Norman, A. F., R. Regnery, P. Jameson, C. Greene, and D. C. Krause. 1995.
Differentiation of Bartonella-like isolates at the species level by PCR-restric-
tion fragment length polymorphism in the citrate synthase gene. J. Clin.
38. Pappalardo, B. L., M. T. Correa, C. C. York, C. Y. Peat, and E. B.
Breitschwerdt. 1997. Epidemiologic evaluation of the risk factors associated
with exposure and seroreactivity to Bartonella vinsonii in dogs. Am. J. Vet.
39. Pappas, L. G., and A. T. Lunzman. 1985. Canine heartworm in the domestic
and wild canids of southeastern Nebraska. J. Parasitol. 71:828–830.
40. Regnery, R. L., B. E. Anderson, J. E. Clarridge III, M. C. Rodriguez-
Barradas, D. C. Jones, and J. H. Carr. 1992. Characterization of a novel
Rochalimaea species, R. henselae sp. nov., isolated from blood of a febrile,
human immunodeficiency virus-positive patient. J. Clin. Microbiol. 30:265–
41. Relman, D. A., P. W. Lepp, K. N. Sadler, and T. M. Schmidt. 1992. Phylo-
genetic relationships among the agent of bacillary angiomatosis, Bartonella
bacilliformis, and other alpha-proteobacteria. Mol. Microbiol. 6:1801–1807.
42. Relman, D. A., J. S. Loutit, T. M. Schmidt, S. Falkow, and L. S. Tompkins.
1990. The agent of bacillary angiomatosis. An approach to the identification
of uncultured pathogens. N. Engl. J. Med. 323:1573–1580.
43. Roux, V., S. J. Eykyn, S. Wyllie, and D. Raoult. 2000. Bartonella vinsonii
subsp. berkhoffii as an agent of afebrile blood culture-negative endocarditis in
a human. J. Clin. Microbiol. 38:1698–1700.
44. Roux, V., and D. Raoult. 1995. The 16S–23S rRNA intergenic spacer region
of Bartonella (Rochalimaea) species is longer than usually described in other
bacteria. Gene 156:107–111.
45. Roux, V., and D. Raoult. 1995. Inter- and intraspecies identification of
Bartonella (Rochalimaea) species. J. Clin. Microbiol. 33:1573–1579.
46. Sander, A., C. Buhler, K. Pelz, E. von Cramm, and W. Bredt. 1997. Detection
and identification of two Bartonella henselae variants in domestic cats in
Germany. J. Clin. Microbiol. 35:584–587.
47. Sander, A., M. Ruess, S. Bereswill, M. Schuppler, and B. Steinbrueckner.
1998. Comparison of different DNA fingerprinting techniques for molecular
typing of Bartonella henselae isolates. J. Clin. Microbiol. 36:2973–2981.
48. Spach, D. H., K. P. Callis, D. S. Paauw, Y. B. Houze, F. D. Schoenknecht,
D. F. Welch, H. Rosen, and D. J. Brenner. 1993. Endocarditis caused by
Rochalimaea quintana in a patient infected with human immunodeficiency
virus. J. Clin. Microbiol. 31:692–694.
49. Tenover, F. C., R. D. Arbeit, R. V. Goering, P. A. Mickelsen, B. E. Murray,
D. H. Persing, and B. Swaminathan. 1995. Interpreting chromosomal DNA
restriction patterns produced by pulsed-field gel electrophoresis: criteria for
bacterial strain typing. J. Clin. Microbiol. 33:2233–2239.
50. Welch, D. F., K. C. Carroll, E. K. Hofmeister, D. H. Persing, D. A. Robison,
A. G. Steigerwalt, and D. J. Brenner. 1999. Isolation of a new subspecies,
Bartonella vinsonii subsp. arupensis, from a cattle rancher: identity with
isolates found in conjunction with Borrelia burgdorferi and Babesia microti
among naturally infected mice. J. Clin. Microbiol. 37:2598–2601.
51. Willeberg, P. W., R. Ruppanner, D. E. Behymer, H. H. Higa, C. E. Franti,
and R. A. Thompson. 1979. Epidemiologic survey of sylvatic plague by
serotesting coyote sentinels with enzyme immunoassay. Am. J. Epidemiol.
4200 CHANG ET AL. J. CLIN. MICROBIOL.
on June 4, 2013 by guest