Experimental Infection of White-Tailed Deer with Anaplasma phagocytophilum, Etiologic Agent of Human Granulocytic Anaplasmosis

Article · September 2005with16 Reads
DOI: 10.1128/JCM.43.8.3595-3601.2005 · Source: PubMed
Abstract
Serologic and molecular evidence of Anaplasma phagocytophilum has been demonstrated in white-tailed deer (WTD; Odocoileus virginianus), and deer are an important host for the tick vector Ixodes scapularis. In this study, we describe experimental infection of WTD with A. phagocytophilum. We inoculated four WTD with a human isolate of A. phagocytophilum propagated in tick cells. Two additional deer served as negative controls. All inoculated deer developed antibodies (titers, ≥64) to A. phagocytophilum, as determined by an indirect fluorescent antibody test, between 14 and 24 days postinfection [p.i.]), and two deer maintained reciprocal titers of ≥64 through the end of the 66-day study. Although morulae were not observed in granulocytes and A. phagocytophilum was not reisolated via tick cell culture of blood, 16S reverse transcriptase nested PCR (RT-nPCR) results indicated that A. phagocytophilum circulated in peripheral blood of three deer through at least 17 days p.i. and was present in two deer at 38 days p.i. Femoral bone marrow from one deer was RT-nPCR positive for A. phagocytophilum at 66 days p.i. There was no indication of clinical disease. These data confirm that WTD are susceptible to infection with a human isolate of A. phagocytophilum and verify that WTD produce detectable antibodies upon exposure to the organism. Because adults are the predominant life stage of I. scapularis found on deer and because adult I. scapularis ticks do not transmit A. phagocytophilum transovarially, it is unlikely that WTD are a significant source of A. phagocytophilum for immature ticks even though deer have a high probability of natural infection. However, the susceptibility and immunologic response of WTD to A. phagocytophilum render them suitable candidates as natural sentinels for this zoonotic tick-borne organism.
JOURNAL OF CLINICAL MICROBIOLOGY, Aug. 2005, p. 3595–3601 Vol. 43, No. 8
0095-1137/05/$08.000 doi:10.1128/JCM.43.8.3595–3601.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.
Experimental Infection of White-Tailed Deer with Anaplasma
phagocytophilum, Etiologic Agent of Human
Granulocytic Anaplasmosis
Cynthia M. Tate,
1
* Daniel G. Mead,
1
M. Page Luttrell,
1
Elizabeth W. Howerth,
2
Vivien G. Dugan,
1,3
Ulrike G. Munderloh,
4
and William R. Davidson
1,
Southeastern Cooperative Wildlife Disease Study,
1
Department of Pathology, College of Veterinary Medicine,
2
and Warnell School of Forest Resources,
3
The University of Georgia, Athens, Georgia
30602, and Department of Entomology, University of Minnesota,
St. Paul, Minnesota 55108
4
Received 23 September 2004/Returned for modification 20 December 2004/Accepted 17 April 2005
Serologic and molecular evidence of Anaplasma phagocytophilum has been demonstrated in white-tailed deer
(WTD; Odocoileus virginianus), and deer are an important host for the tick vector Ixodes scapularis. In this
study, we describe experimental infection of WTD with A. phagocytophilum. We inoculated four WTD with a
human isolate of A. phagocytophilum propagated in tick cells. Two additional deer served as negative controls.
All inoculated deer developed antibodies (titers, >64) to A. phagocytophilum, as determined by an indirect
fluorescent antibody test, between 14 and 24 days postinfection [p.i.]), and two deer maintained reciprocal
titers of >64 through the end of the 66-day study. Although morulae were not observed in granulocytes and A.
phagocytophilum was not reisolated via tick cell culture of blood, 16S reverse transcriptase nested PCR
(RT-nPCR) results indicated that A. phagocytophilum circulated in peripheral blood of three deer through at
least 17 days p.i. and was present in two deer at 38 days p.i. Femoral bone marrow from one deer was RT-nPCR
positive for A. phagocytophilum at 66 days p.i. There was no indication of clinical disease. These data confirm
that WTD are susceptible to infection with a human isolate of A. phagocytophilum and verify that WTD produce
detectable antibodies upon exposure to the organism. Because adults are the predominant life stage of I.
scapularis found on deer and because adult I. scapularis ticks do not transmit A. phagocytophilum transovarially,
it is unlikely that WTD are a significant source of A. phagocytophilum for immature ticks even though deer have
a high probability of natural infection. However, the susceptibility and immunologic response of WTD to A.
phagocytophilum render them suitable candidates as natural sentinels for this zoonotic tick-borne organism.
Human granulocytic anaplasmosis is an acute, febrile disease
that may be accompanied by headache, myalgia, pancytopenia,
and elevated serum aminotransferase levels (13). Although the
disease is often mild, delayed treatment, misdiagnosis, and/or
immunosuppression may result in a severe or fatal outcome
(28, 31). Approximately 1,220 cases of human granulocytic
anaplasmosis (HGA) have been diagnosed in the United
States since 1994, when the disease was first described (13, 14).
Sporadic HGA cases have been reported in Europe (10).
Previous to its recognition as a human pathogen, the organ-
ism was known in veterinary medicine as Ehrlichia equi (caus-
ing equine granulocytic ehrlichiosis) and Ehrlichia phagocyto-
phila (causing tick-borne fever in sheep, goats, and cattle in
several European countries). Based on recent phylogenetic
analysis, the etiologic agent of HGA and E. equi were synono-
mized with E. phagocytophila. Furthermore, the analysis indi-
cated that this organism should be reassigned to the genus
Anaplasma, thus resulting in the currently accepted designa-
tion, Anaplasma phagocytophilum (20). The marked differences
in host preference, clinical manifestations, and geographical
distribution are attributed to the existence of multiple variant
strains of A. phagocytophilum (20).
North American strains of A. phagocytophilum cause clinical
disease in domestic animals, notably horses and dogs (20).
Experimentally, both equine and human A. phagocytophilum
strains are pathogenic to horses (37) and cross protective (6).
Disease due to A. phagoctyophilum in domestic cattle has not
been reported in North America. Furthermore, two steers
were not susceptible to experimental infection with human and
equine North American strains; although the steers serocon-
verted, blood was negative for A. phagocytophilum by 16S real-
time PCR at all sampling dates (51). Anaplasma phagocytophi-
lum was first isolated from human patients by using a human
promyelocytic leukemia (HL60) cell line (24) and from horses
and dogs by using Ixodes scapularis cell lines (45, 46).
Knowledge of the natural history of A. phagocytophilum re-
mains incomplete. In the eastern and midwestern United
States, the white-footed mouse (Peromyscus leucopus) and the
black-legged tick (I. scapularis) are a competent reservoir host
and principal vector, respectively (19, 54). Although field stud-
ies indicate that numerous species of wild rodents and other
wild mammals may be naturally infected with A. phagocytophi-
lum (22, 23, 32, 40, 48, 59), the relative importance of these
hosts as sources of A. phagocytophilum for ticks has not been
determined.
White-tailed deer (WTD; Odocoileus virginianus) are the
* Corresponding author. Mailing address: Southeastern Coopera-
tive Wildlife Disease Study, Wildlife Health Building, College of Vet-
erinary Medicine, The University of Georgia, Athens, GA 30602-7393.
Phone: (706) 542-1741. Fax: (706) 542-5865. E-mail: ctate@vet
.uga.edu.
3595
principal hosts for adult I. scapularis (30) and therefore pre-
dictably are exposed to A. phagocytophilum. Sequence-con-
firmed 16S rRNA genes identical to those of A. phagocytophi-
lum and/or A. phagocytophilum-reactive antibodies have been
demonstrated in wild WTD from Connecticut, Georgia, Indi-
ana, Maryland, Missouri, South Carolina, and Wisconsin (3, 8,
33, 38, 39, 60); however, clinical disease due to A. phagocyto-
philum has not been reported in WTD. Thus, WTD have been
identified as both a potential sentinel species for A. phagocy-
tophilum (8, 33, 39, 40, 60), as well as a potential reservoir host
of A. phagocytophilum (3, 5, 6, 8, 43); however, these potential
roles have not been analyzed because data on susceptibility to
and course of infection with A. phagocytophilum in WTD have
previously been unavailable. Therefore, we proposed to ana-
lyze these potential roles by experimentally infecting WTD
with A. phagocytophilum.
MATERIALS AND METHODS
Animals and experimental design. Six WTD fawns (three females and three
males, orphaned in the state of Georgia) were hand raised and housed in
tick-free facilities at the College of Veterinary Medicine, The University of
Georgia. Prior to initiation of the experiment, at approximately 7 months of age,
the deer were screened for various hematotropic microorganisms known to infect
wild WTD in the southeastern United States. The deer were determined to be
seronegative by indirect fluorescence antibody (IFA) test for A. phagocytophilum
and Ehrlichia chaffeensis (8, 17) and PCR negative for A. phagocytophilum,E.
chaffeensis,Ehrlichia ewingii, and an undescribed Anaplasma sp. of WTD (33, 44).
All deer were determined to be culture negative for Trypanosoma cervi, a pro-
tozoan known to interfere with tissue culture of deer blood (35, 36, 44). Two deer
(WTD132 and WTD133) were diagnosed with Theileria cervi infection by obser-
vation of intraerythrocytic protozoans in whole-blood smears.
For all procedures, deer were anesthetized as described previously (16). Deer
blood samples were collected via aseptic jugular venipuncture at 0, 6, 10, 13, 17,
24, 31, 38, 45, 54, and 66 days postinfection (days p.i.). Approximately 9 ml of
whole blood was collected for culture, reverse transcriptase nested PCR (RT-
nPCR) and clinical hematology in three separate EDTA Vacutainer tubes (Bec-
ton Dickinson, Rutherford, NJ) and for serology in a tube containing no additive.
On all sampling dates, Giemsa-stained blood smears were prepared, and whole
blood was submitted to the Clinical Pathology Laboratory, College of Veterinary
Medicine, The University of Georgia, for analysis of the following parameters:
hematocrit, erythrocyte count, hemoglobin levels, platelet count, total and dif-
ferential leukocyte counts, and fibrinogen levels. Sera and duplicate samples of
whole blood were stored at 20°C.
Deer were observed twice daily for visible signs of clinical illness, including
decreased feed intake, depression, and reluctance to move. Complete physical
examination (11) of each deer was performed at each blood collection date. The
two control deer were removed from the study at 31 days p.i. All deer were
euthanized via intravenous sodium pentobarbital overdose and subjected to
complete necropsies.
Preparatory to and concurrently with the experimental deer trial, female
C3H-HeN mice (Harlan, Indianapolis, IN), approximately 6 weeks old, were
used to test animal infectivity of the A. phagocytophilum isolate propagated in
tick cells (46). Mice were anesthetized for all procedures via subcutaneous
injection of 10 mg of xylazine (Phoenix Scientific, Inc., St. Joseph, MO)/kg of
body weight and 100 mg/kg ketamine HCl (Ft. Dodge Labs, Inc., Fort Dodge,
IA). Samples of blood taken 6 days p.i. for RT-nPCR were collected retro-
orbitally from all mice. After euthanasia of mice, blood samples were collected
for culture by intracardiac puncture, and spleen samples were taken for RT-
nPCR. All animals involved in this study were cared for and used in accordance
with guidelines established by the Institutional Animal Care and Use Committee
of The University of Georgia.
Anaplasma phagocytophilum inoculum. An I. scapularis tick cell line (ISE6) was
used to propagate a human isolate (HGE-1) of A. phagocytophilum (46). Stock
ISE6 cultures were maintained as described previously (47). The inoculum con-
sisted of five 12.5-cm
2
Falcon culture flasks (Becton Dickenson, Franklin Lakes,
NJ) of A. phagocytophilum-infected tick cells, in which approximately 35% of the
cells were infected. Monolayers were resuspended in existing media, and sterile
water (Sigma, St. Louis, MO) was added to bring the volume of each flask to 8.0
ml. Then, the contents of all five flasks were combined for a total volume of 40
ml, and the mixture was divided into 2.0-ml aliquots for deer and 0.5-ml aliquots
for mice. Negative-control injection material was prepared in a similar manner,
using uninfected stock ISE6 cultures.
Each of four deer (WTD131, -132, -133, and -134) was injected with 2.0 ml of
the A. phagocytophilum inoculum by each of four routes: intradermal, subcuta-
neous, intravenous, and intraperitoneal, for a total of 8.0 ml of inoculum per
deer. Concurrently with the deer inoculation, each of nine mice was injected
intraperitoneally with 0.5 ml of the A. phagocytophilum inoculum. Two negative
control deer (WTD127 and WTD139) and three negative control mice were
injected in a similar manner with uninfected ISE6 cells. Remaining fractions of
the A. phagocytophilum inoculum and uninfected tick cells were used to prepare
Giemsa-stained cytospins and were tested by RT-nPCR and DNA sequencing
(see below).
Serology. The IFA test was performed as described previously (8). In brief,
sera were screened at a dilution of 1:64 in 0.01 M phosphate-buffered saline on
commercially prepared HGE substrate slides (Focus Technologies [formerly
MRL Diagnostics], Cypress, CA). Fluorescein isothiocyanate-labeled rabbit anti-
deer immunoglobulin G (Kirkegaard & Perry Laboratories, Gaithersburg, MD),
diluted 1:50 in phosphate-buffered saline, was used as a conjugate. When distinct
fluorescent staining of organisms was observed at a 1:64 dilution, serial twofold
dilutions were performed. Serologic results are reported as reciprocals of the
highest dilution at which specific fluorescence was observed.
Cell culture. Reisolation attempts of A. phagocytophilum from mouse and deer
blood were performed as previously described (44, 45) except that resuspended
ISE6 stock cell monolayers and washed WTD buffy coat cells were mixed,
pelleted at 720 gfor 20 min, and then allowed to stand at room temperature
for 30 min before the pellet was resuspended in “ehrlichia medium” (46) and
divided into two flasks. Duplicate flasks were monitored and maintained sepa-
rately using different sets of reagents as a precaution against contamination;
antibiotics were not used at any time. Cultures were monitored by visual obser-
vation for cytopathic effect (CPE) (46), light microscopy of Giemsa-stained cell
spreads, and RT-nPCR of cell culture supernatant. Monolayers were examined
daily for development and progression of CPE. Periodically, samples were pre-
pared from all cultures by centrifugation (720 gfor 20 min) of the entire
volume of spent medium removed during feeding of the culture, followed by
resuspension of pelleted cells in approximately 1.0 ml of spent medium. For cell
spreads, 100 l of each sample was placed in a Cytofuge filter concentrator
(StatSpin; Iris Co., Norwood, MA) and centrifuged at 27 gfor 4 min using a
Cytofuge 2 cytocentrifuge (StatSpin). Slides were air dried, fixed in 100% meth-
anol, and stained in a 4% solution of Giemsa (Karyomax; GIBCO, Grand Island,
NY) in Sorensen buffer, pH 6.5, for 30 min in a 37°C water bath. Stained cells
were examined microscopically (400 to 1,000) for intracytoplasmic organ-
isms. Our cell culture protocol specified a 60-day monitoring period for CPE,
followed by RT-nPCR testing.
RNA extraction, RT-nPCR, and nucleotide sequence analysis. Total RNA was
extracted from fresh deer and mouse blood samples with the RNA Blood Minikit
(QIAGEN, Inc., Valencia, CA) and from cell culture aliquots and postmortem
tissue stored at 70°C with the QiAmp Viral RNA Extraction kit and the
RNEasy Minikit (QIAGEN, Inc.), respectively. All extractions were performed
according to the manufacturer’s instructions and in an RNase-free environment.
RT-nPCR was performed on RNA extracted from deer blood, cell culture, and
tissues in the following manner. Reverse transcription of 16S r-RNA to cDNA
and subsequent primary amplification using primers ECC and ECB were carried
out in a single-tube reaction, followed by secondary amplification as described
previously (44), except that secondary primers GE9F and GA1UR were used to
generate an internal 411-bp fragment (13, 34).
Two additional gene targets (p44 and groESL) for detection of A. phagocyto-
philum RNA were used. For detection of p44 RNA, RT-PCR was performed
using primers MSP3F and MSP3R (65). Reaction mixtures were subjected to
45°C for 20 min, followed by 39 cycles of the following profile: 94°C for 60 s, 55°C
for 45 s, and 72°C for 60 s, using a PTC-100TM Thermal Cycler. For detection
of groESL RNA, RT-nPCR was performed using primers APF1 (5TAGTGAT
GAAGGAGAGTGAC) and APR1 (5CCAGGIGCCTTIACAGCWGCAAC)
in a primary reaction and primers APF10 (5TATGCTACGGTTGTTTGTTC)
and APR11 (5GGCGAAAGATATCCGCGA) in a secondary reaction to gen-
erate a 652-bp product (primers were generously provided by John Sumner,
Centers for Disease Control and Prevention). Primary reaction mixtures were
subjected to 43°C for 15 min, followed by 95°C for 5 min and 39 cycles of the
following profile: 95°C for 30 s, 52°C for 30 s, and 72°C for 60 s; and a final step
of 72°C for 5 min. Secondary reaction mixtures were subjected to 95°C for 5 min;
29 cycles of 95°C for 30 s, 52°C for 30 s, and 72°C for 60 s; and a final step of 72°C
for 5 min.
3596 TATE ET AL. J. CLIN.MICROBIOL.
All RT-PCRs were carried out with a PTC-100TM Thermal Cycler. All re-
agent concentrations were identical to those used for A. phagocytophilum 16S
rRNA RTn-PCR. Amplification products were separated by electrophoresis on
a 2% agarose gel, stained in ethidium bromide, and visualized with UV transil-
lumination.
Quality control measures included negative controls (water) that were ex-
tracted and amplified in parallel with all specimens. To minimize the potential
for DNA contamination, three separate, designated areas were used for extrac-
tion of RNA and preparation of primary and secondary PCRs. Additionally, two
thermal cyclers were used, designated for either primary or secondary amplifi-
cation.
RT-nPCR products from the original inoculum, all deer blood samples, se-
lected mouse blood samples and cell culture aliquots, and one postmortem tissue
sample were subjected to DNA sequencing. Gene fragments were purified by gel
electrophoresis, and bands were extracted and purified with the QIAquick gel
extraction kit (QIAGEN, Inc.). DNA was sequenced in forward and reverse
directions at the Molecular Genetics Instrumentation Facility, The University of
Georgia, with an ABI 3100 automated sequencer (Applied Biosystems, Perkin
Elmer Corp., Foster City, CA). The sequences were assembled and edited using
the Sequencher software package, version 4.1.4 (Gene Codes Corp., Ann Arbor,
MI). A nucleotide-nucleotide BLAST (blastn) search was performed to deter-
mine the most similar sequences of the target genes published in GenBank
(http://www.ncbi.nlm.nih.gov/).
Due to incidental molecular detection of Bartonella spp. in deer blood cultures
during the course of the experiment, cell culture samples were subjected to
RT-PCR for the gltA gene of Bartonella sp. using primers CS140f and
BhCS1137n (9). To increase sensitivity, a heminested RT-PCR was developed
for the detection of Bartonella spp. directly from peripheral blood, using primer
set CS140f and BhCS1137n in the primary reaction and Bh731p and BhCS1137n
(49) in the secondary reaction. Reagent concentrations and reaction conditions
were identical to those used for 16S rRNA A. phagocytophilum RT-nPCR.
Pathology. Tissues collected for RT-nPCR included spleen, prescapular and
prefemoral lymph nodes, bone marrow from the sternum and femoral head, and
lung. The aforementioned tissues as well as heart, liver, kidney, adrenal gland,
brain, bladder, haired skin, reproductive organs, and gastrointestinal tract also
were collected in 10% neutral buffered formalin for histopathologic examination.
RESULTS
Animal infectivity of A. phagocytophilum inoculum. The an-
imal infectivity of the A. phagocytophilum inoculum was con-
firmed via RT-nPCR and DNA sequencing of a subset of
products obtained from blood samples taken 6 days p.i. from
seven of nine inoculated mice by using both the 16S and p44
gene targets (data not shown). Furthermore, rare morulae
were observed in granulocytes of blood taken 6 days p.i. from
RT-nPCR-positive mice. Although postmortem spleen sam-
ples for the previously mentioned seven mice were positive for
A. phagocytophilum by 16S RT-nPCR, the organism was not
reisolated in cell culture from terminal blood pooled from
these mice at 11, 15, and 20 days p.i. Negative RT-nPCR
results of blood and spleen samples were obtained for mice
inoculated with uninfected tick cells.
Serology. All experimental deer developed reciprocal anti-
body titers of 64 to A. phagocytophilum between 14 and 24
days p.i. (Table 1). Two experimental deer (WTD133 and
WTD134) seroconverted by 17 days p.i. and remained sero-
positive throughout the 66-day study. The peak reciprocal titer
of 2,048 was detected in one of these deer (WTD134) on 38
days p.i. The other two experimental deer (WTD131 and
WTD132) had peak reciprocal titers of 128 and were seropos-
itive only through 38 days p.i. The geometric mean (57) of all
titers of 64 was 115. Anaplasma phagocytophilum IFA results
for the control deer were negative (64) on all sample dates.
RT-nPCR and cell culture. Blood from all four experimental
deer was positive for A. phagocytophilum by 16S RT-nPCR on
one to five occasions between 6 and 38 days p.i. (Table 1). All
16S rRNA gene amplicons from deer blood were sequenced
and found to be identical to the sequence of the original
inoculum and 99.5% similar to a published sequence of A.
phagocytophilum (GenBank accession number U02521). Blood
from WTD132 and WTD133 was positive for A. phagocytophi-
lum by groESL RT-nPCR on 13 and 17 days p.i. Sequenced
groESL products were 100% identical to published sequences
of A. phagocytophilum (GenBank accession numbers
AY219849 and U96728). Use of the p44 RT-PCR did not yield
A. phagocytophilum amplicons from deer blood. Blood from
the two negative control deer (WTD127 and -139) was nega-
TABLE 1. Test results for experimental infection of white-tailed deer with Anaplasma phagocytophilum
a
Days p.i.
Results of infection
WTD131 WTD132 WTD133 WTD134
CC PCR
b
IFA CC PCR IFA CC PCR IFA CC PCR IFA
0ND64 ND 64 ND 64 ND 64
6—
c
ND — ND —
c
ND —
c
—ND
10 F ⫹⬍64 F ⫹⬍64 F ⫹⬍64 ND 64
13 ND ⫹⬍64 ND
d
64 ND
d
64 —
c
64
17 —
c
⫹⬍64 —
d
64 —
c
d
64 —
c
—64
24 —
c
128 — 128 —
c
256 —
c
— 1,024
31 —
c
64 — 64 —
c
—64
c
— 1,024
38 —
c
64 — 64 — 256 —
c
2,048
45 F 64 F 64 F 128 F 256
54 64 — 64 —
c
128 —
c
— 256
66 —
c
64 —
c
64 —
c
—64
c
— 128
66 Nx ND NA ND NA ND NA ND NA
a
Tests included cell culture (CC), 16S RT-nPCR of peripheral blood with DNA sequence confirmation, and IFA test, reported as reciprocal titers. Two control deer
(WTD127 and WTD139) were negative on all tests through DPI 30 at which time they were removed from the study. F, tick cells failed to reattach after being admixed
with WTD cells; NA, not applicable; ND, not done; —, negative results.
b
PCR results are for whole blood except for last row. Entries for 66 Nx days p.i. (66 Nx) provide results of 16S RT-nPCR of postmortem tissues; sequence-confirmed
A. phagocytophilum DNA was amplified from femoral bone marrow of WTD133 only.
c
Cultures negative for A. phagocytophilum, but Bartonella sp. isolated.
d
groESL RNA of A. phagocytophilum amplified by RT-nPCR.
VOL. 43, 2005 EXPERIMENTAL A. PHAGOCYTOPHILUM IN WHITE-TAILED DEER 3597
tive for A. phagocytophilum by 16S and groESL RT-nPCR and
by p44 RT-PCR at 6, 10, 13, 17, 24, and 31 days p.i.
Anaplasma phagocytophilum was not reisolated in any of 36
culture attempts from experimental deer (Table 1). All 10 and
45 days p.i. cultures failed when tick cells did not reform a
monolayer in the flask after the deer blood culture procedure
was completed. Of 29 remaining culture attempts of A. phago-
cytophilum-inoculated deer, 23 cultures exhibited CPE, and
intracellular bacteria were observed in Giemsa-stained cyto-
spins. Although RT-nPCR of these cell culture samples with
primer pairs ECC-ECB and GE9F-GA1UR yielded products
of the expected size, DNA sequencing revealed that they were
not 16S rRNA gene fragments of A. phagocytophilum, but
rather of Bartonella sp. (see below).
Control deer blood cultures and experimental deer blood
cultures not exhibiting CPE from 6, 17, 24, 31, and 38 days p.i.
were lost due to a single bacterial contamination event involv-
ing use of a commercially purchased component of the tick cell
media at 45, 34, 27, 20, and 13 days in culture (DIC), respec-
tively. For the same reason, the three mouse blood cultures
from 11, 15 and 20 days p.i. were lost at 40, 36, and 31 DIC,
respectively.
Clinical, hematologic and postmortem findings. Clinical
signs attributable to infection with A. phagocytophilum were
not apparent in any deer throughout the 66-day study. Morulae
were not observed in granulocytes on Giemsa-stained deer
blood smears. Although results of complete blood counts were
not always within normal limits for all fawns on all sampling
dates, no consistent pattern of hematologic abnormalities at-
tributable to infection with A. phagocytophilum was apparent.
At four sampling dates between 30 and 55 days p.i.,
WTD133 exhibited low platelet counts (48 10
3
to 168
10
3
/l) relative to its own values before and after that time
period and relative to all platelet counts of the other experi-
mental deer and the negative control deer (mean 685 10
3
/
l). During this time period, WTD133 showed no other signs
consistent with thrombocytopenia.
Gross and histopathologic lesions attributable to infection
with A. phagocytophilum were not apparent in any deer. RT-
nPCR assays of postmortem samples of femoral and sternal
bone marrow, prescapular and prefemoral lymph node, spleen,
and lung of experimental and control deer were negative for A.
phagocytophilum, with the exception of a sequence-confirmed
16S rRNA gene amplicon from femoral bone marrow of
WTD133.
Detection of incidental Bartonella sp. infections. Sequencing
of 16S rRNA gene products amplified with primer sets ECC-
ECB and GE9F-GA1UR from a randomly chosen subset (7 of
24) of 16S RT-nPCR-positive cell culture samples revealed
that the products were 98.6% similar to a sequence of Bartonella
schoenbuchensis (GenBank accession number AJ278190.1), an
intraerythrocytic bacteria first isolated from the blood of wild
roe deer (Capreolus capreolus) in Germany (18). Subsequently,
cell culture samples were subjected to RT-nPCR for the citrate
synthase (gltA) gene of Bartonella spp., and 21 of 24 samples
yielded products of the expected size. Furthermore, hemin-
ested gltA RT-PCR for Bartonella spp. developed in this study
for screening whole blood yielded products of the expected size
from preinoculation blood of two deer, WTD134 and
WTD127. DNA sequencing and alignment indicated that these
two gltA fragments had 98.6% and 94.6% sequence similarity,
respectively, to a sequence of B. schoenbuchensis (GenBank
accession number AJ564633) recently isolated from the midgut
of a deer ked (Lipoptena cervi). One 16S rRNA gene sequence
of the Bartonella sp. amplified from tick cell culture and the
gltA DNA sequences of the Bartonella sp. amplified from the
blood of WTD134 and WTD127 are available under GenBank
accession numbers AY805111, AY805109, and AY805110, re-
spectively.
DISCUSSION
We demonstrated that WTD can support infection with a
human-infective strain of A. phagocytophilum and that infec-
tion of WTD with A. phagocytophilum is accurately reflected by
seroconversion. Circulation of A. phagocytophilum in periph-
eral blood of experimental deer was transient, similar to mice
and horses experimentally infected with various North Amer-
ican isolates of the organism (2, 27, 50, 51). Anaplasma phago-
cytophilum RNA was infrequently detectable in peripheral
blood of experimental deer after development of antibodies,
suggesting that the humoral immune system may have played a
role in bacterial clearance. Although the function of humoral
immunity in host elimination of rickettsiae is not well under-
stood, Winslow et al. (61) demonstrated that antibodies can
affect the course of active infection of E. chaffeensis in SCID
mice. Administration of immune sera 10 and 17 days after
infection resulted in partial clearance of E. chaffeensis from
infected mice; however, the organism eventually recolonized
the liver. This indicated that antibodies failed to mediate com-
plete bacterial clearance. Subsequently, the authors hypothe-
sized that E. chaffeensis may have persisted at low levels in the
liver or emigrated from tissues that were inaccessible to the
antibodies.
In the present study, A. phagocytophilum was detected by
RT-nPCR in the peripheral blood of one deer (WTD131) at 38
days p.i., following a 3-week period of negative RT-nPCR
results. The occurrence of recrudescent rickettsemia has been
documented in a 9-month study of WTD experimentally in-
fected with E. chaffeensis (16). Recrudescent rickettsemia may
result from release of organisms sequestered in tissue, but this
hypothesis requires further evaluation with regard to A. phago-
cytophilum in WTD. The detection of A. phagocytophilum
RNA in femoral bone marrow from WTD133 on 66 days p.i.
suggests a potential site of latent infection.
While A. phagocytophilum was not visualized in granulocytes
of experimental deer, the difficulty of finding and definitively
identifying morulae in blood smears of humans with confirmed
A. phagocytophilum infections is well known (5, 58), particu-
larly in blood samples taken from afebrile patients (1, 4).
Furthermore, studies demonstrate that blood of experimen-
tally infected laboratory mice, although morula negative, was
still infectious to naı¨ve mice and sometimes PCR positive and
culture positive as well (26, 53, 54). Thus, our negative light
microscopy results for the deer are consistent with those of
several previous experimental infection studies of mice.
Demonstration of gene transcription by the use of RT-nPCR
is suggestive of A. phagocytophilum survival and replication
within the deer (21). Therefore, our 16S RT-nPCR data imply
that viable A. phagocytophilum circulated until at least 17 days
3598 TATE ET AL. J. CLIN.MICROBIOL.
p.i. in three of the four deer. Although limited, this time period
is of sufficient duration hypothetically to infect ticks. Nonethe-
less, we contend it is unlikely that WTD play an epidemiolog-
ically significant role as a source of A. phagocytophilum for
ticks. This conclusion is based on the fact that WTD are par-
asitized primarily by the adult forms of I. scapularis (30) and
therefore are most likely to be exposed to A. phagocytophilum
at the end of the tick life cycle. Furthermore, A. phagocytophi-
lum is not known to be maintained transovarially in the tick.
Anaplasma phagocytophilum infection initiated by a single
needle inoculation may differ substantially from infection nat-
urally acquired by the bite of one or more infected ticks over
the course of a season. In addition to the likelihood that wild
WTD experience multiple exposures to A. phagocytophilum,
various immunologically active components of tick saliva may
be important factors influencing the outcome of natural expo-
sures of WTD to A. phagocytophilum (25).
We succeeded in reisolating A. phagocytophilum from a
mouse inoculated with tick cell culture in the development
phase of our research (unpublished data); however, we were
unsuccessful in three attempts to reisolate A. phagoctyophilum
from mice inoculated concurrently with deer. Relatively few
studies have attempted culture of blood from laboratory mice;
some have also reported discrepancies between cell culture
and PCR results (26, 53).
With regard to our attempts to reisolate A. phagocytophilum
from experimental deer, we encountered significant difficulties
related to three separate issues. First, on two culture days (10
and 45 days p.i.), tick cells failed to reattach after the mono-
layer was disrupted and the cells were admixed with deer buffy
coat cells. On these days, cellular debris created during resus-
pension may have resulted in cytotoxicity to the tick cells, as
reported previously (64). Alternatively, the deer buffy coat cells
may have killed the tick cells (U. G. Munderloh, unpublished
data). Second, although the experimental design specified
monitoring all cultures not developing CPE for 60 days, bac-
terial contamination of a commercially purchased component
of the tick cell media resulted in the loss of many cultures
before the end of 60 days. Because all “negative” cultures
initiated previous to the contamination event on 51 days p.i.
were destroyed, “negative” culture results for WTD127, -132,
-133, and -139 are equivocal. Third, for at least 21 isolation
attempts, the presence of Bartonella spp. confounded cell cul-
ture. For example, our best opportunities to reisolate A. phago-
cytophilum would have been days when deer were RT-nPCR
positive, as was the case at 6 and 17 days p.i. (WTD131, -132,
and -133) and on 38 days p.i. (WTD131 and -134); however, of
these eight culture opportunities, Bartonella spp. were isolated
in all but two. Bartonella spp. replicated rapidly in deer blood
cultures; a CPE was apparent to the unaided eye as early as 7
DIC and was nearly 100% between 8 and 23 DIC. Perhaps the
vigorous growth of Bartonella sp. in the tick cell medium re-
sulted in conditions unsuitable for the survival of A. phagocy-
tophilum. If this is the case, future attempts to culture A.
phagocytophilum from the blood of deer coinfected with Bar-
tonella spp. may be facilitated by the use of an antibiotic to
which the former is resistant but to which the latter may be
susceptible, such as erythromycin (7, 29).
In the present study, 16S RT-nPCR was the most sensitive
assay for detection of A. phagocytophilum RNA in deer blood,
followed by the groESL RT-nPCR assay. Because both of these
assays are nested, we expected them to be more sensitive than
the nonnested p44 RT-PCR. Massung and Slater (41) demon-
strated that specificity and sensitivity vary markedly among the
numerous published PCR assays and primer pairs for A. phago-
cytophilum. With regard to specificity, our cell culture results
illustrate that even in a controlled experimental setting, PCR-
based methods of detection should be confirmed by use of
alternative gene targets or by DNA sequencing. We were
aware of the existence of an undescribed Anaplasma sp. of
WTD that is amplified in 16S PCR assays by using primers
GE9F and GA1UR (34). Additionally, it has been reported
that in blood samples containing a high concentration of
Anaplasma (Ehrlichia)platys DNA, E. equi primers have in-
duced false priming (52). However, we were not aware of a
Bartonella sp. infecting WTD in the southeastern United States
until we sequenced RT-nPCR products amplified with primers
GE9F-GA1UR from tick cell culture of deer blood. Of interest
is the fact that although the deer were apparently coinfected
with A. phagocytophilum and Bartonella sp., we never detected
Bartonella spp. directly from deer blood with the aforemen-
tioned primers. Therefore, we believe that the copy number of
Bartonella spp. circulating in peripheral blood of the deer was
very low. In fact, in retrospective testing of all deer blood
samples collected during this study, we were unable to detect
Bartonella spp. directly in blood by a single-step gltA RT-PCR
assay using primers cited in a standard PCR assay for ampli-
fication of DNA from cervid isolates of Bartonella spp. from
Europe and the western United States (12, 18). Only after
development of a heminested gltA RT-PCR were we able to
detect RNA of Bartonella spp. in preinoculation blood samples
from two of the deer.
Our serologic and molecular findings lend support to the
premise that WTD should be suitable sentinels for human risk
of exposure to A. phagocytophilum (8, 33, 39, 40, 60). Because
two experimental deer maintained detectable antibodies (titer,
64) for at least 49 days through the end of the 66-day study,
we suggest that wild WTD repeatedly exposed to A. phagocy-
tophilum may exhibit relatively long-lasting serologic response,
a desirable trait in a potential sentinel species. Together with
existing field data, our experimental findings, including use of
a 1:64 dilution for serologic screening, lay the foundation for
the development and validation of an A. phagocytophilum sen-
tinel system using WTD. Although serologic cross-reactivity
between E. chaffeensis and A. phagocytophilum in humans has
been previously reported (15, 55), recent work suggests that
this phenomenon is not a significant limitation to the use of
WTD as sentinels for A. phagoctyophilum and E. chaffeensis
when surveillance data sets are validated by confirmatory tests
such as immunoblotting, PCR, and culture (60, 63).
Recently, Massung et al. (42) reported a genetic variant of
A. phagocytophilum (AP-variant 1) from wild WTD in Wiscon-
sin and Maryland and from I. scapularis in Rhode Island and
Connecticut. Because this genovariant was not infectious for
mice, the authors hypothesized that AP-variant 1 may be spe-
cific to WTD and may cycle independently of the human-
infective strain of A. phagocytophilum (AP-ha) that is main-
tained in white-footed mice. If proven, this hypothesis would
have implications for the use of WTD as A. phagocytophilum
sentinels. Our serologic and RT-nPCR findings confirm that
VOL. 43, 2005 EXPERIMENTAL A. PHAGOCYTOPHILUM IN WHITE-TAILED DEER 3599
WTD are susceptible to a human-infective strain of A. phago-
cytophilum by needle inoculation, suggesting that both genetic
variants, AP-ha and AP-variant 1, could be present in wild
WTD. Future research related to A. phagoctyophilum infection
among WTD should include identifying the genovariants
present in deer on a broad geographic scale and determining
the infection dynamics of simultaneous or sequential A. phago-
cytophilum genovariants in WTD, as recently reported for E.
chaffeensis (56, 62).
ACKNOWLEDGMENTS
This work was supported primarily by the National Institutes of
Allergy and Infectious Diseases (5 R01 AI044235-02). Further support
was provided by the Federal Aid to Wildlife Restoration Act (50 Stat.
917) and through sponsorship of the fish and wildlife agencies of
Alabama, Arkansas, Florida, Georgia, Kansas, Kentucky, Louisiana,
Maryland, Mississippi, Missouri, North Carolina, Oklahoma, Puerto
Rico, South Carolina, Tennessee, Virginia, and West Virginia.
We thank Andrea Varela for assistance with acquiring and pre-
screening deer, Andrea Davidson for assistance during mouse inocu-
lation, Chris King and Nat Seney for support regarding animal man-
agement, Jeff Tucker for animal handling expertise, and Molly Murphy
for phylogenetic input during this project.
REFERENCES
1. Aguero-Rosenfeld, M. E., H. W. Horowitz, G. P. Wormser, D. F. McKenna,
J. Nowakowski, J. Munoz, and J. S. Dumler. 1996. Human granulocytic
ehrlichiosis: a case series from a medical center in New York State. Ann.
Intern. Med. 127:89–90.
2. Akkoyunlu, M., and E. Fikrig. 2000. Gamma interferon dominates the mu-
rine cytokine response to the agent of human granulocytic ehrlichiosis and
helps to control the degree of early rickettsemia. Infect. Immun. 68:1827–
1833.
3. Arens, M. Q., A. M. Liddell, G. Buening, M. Gaudreault-Keener, J. W.
Sumner, J. A. Comer, R. S. Buller, and G. A. Storch. 2003. Detection of
Ehrlichia spp. in the blood of wild white-tailed deer in Missouri by PCR assay
and serologic analysis. J. Clin. Microbiol. 41:1263–1265.
4. Bakken, J. S., J. Krueth, C. Wilson-Nordskog, R. L. Tilden, K. Asanovich,
and J. S. Dumler. 1996. Clinical and laboratory characteristics of human
granulocytic ehrlichiosis. JAMA 275:199–205.
5. Bakken, J. S., and J. S. Dumler. 2000. Human granulocytic ehrlichiosis. Clin.
Infect. Dis. 31:554–560.
6. Barlough, J. E., J. E. Madigan, E. DeRock, J. S. Dumler, and J. S. Bakken.
1995. Protection against Ehrlichia equi is conferred by prior infection with
the human granulocytotropic Ehrlichia (HGE agent). J. Clin. Microbiol.
33:3333–3334.
7. Bass, J. W., J. M. Vincent, and D. A. Person. 1997. The expanding spectrum
of Bartonella infections. II. Cat-scratch disease. Pediatr. Infect. Dis. J. 16:
163–179.
8. Belongia, E. A., K. D. Reed, P. D. Mitchell, C. P. Kolbert, D. H. Persing, J. S.
Gill, and J. J. Kazmierczak. 1997. Prevalence of granulocytic Ehrlichia in-
fection among white-tailed deer in Wisconsin. J. Clin. Microbiol. 35:1465–
1468.
9. Birtles, R. J., and D. Raoult. 1996. Comparison of partial citrate synthase
gene (gltA) sequences for phylogenetic analysis of Bartonella species. Int. J.
Syst. Microbiol. 46:891–897.
10. Blanco, J. R., and J. A. Oteo. 2002. Human granulocytic ehrlichiosis in
Europe. Clin. Microbiol. Infect. 8:763–772.
11. Blood, D. C., O. M. Radostits, and J. A. Henderson. 1983. Veterinary med-
icine: a textbook of the diseases of cattle, sheep, pigs, goats and horses, 6th
ed. Bailliere Tindall, Eastbourne, United Kingdom.
12. Chang, 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. Boulois, and Y. Piemont. 2000. Bartonella spp. isolated
from wild and domestic ruminants in North America. Emerg. Infect. Dis.
6:306–311.
13. Chen, S. M., J. S. Dumler, J. S. Bakken, and D. H. Walker. 1994. Identifi-
cation of a granulocytotropic Ehrlichia species as the etiologic agent of
human disease. J. Clin. Microbiol. 32:589–595.
14. Childs, J. E., and C. D. Paddock. 2003. The ascendancy of Amblyomma
americanum as a vector of pathogens affecting humans in the United States.
Annu. Rev. Entomol. 48:307–337.
15. Comer, J. A., W. L. Nicholson, J. G. Olson, and J. E. Childs. 1999. Serologic
testing for human granulocytic ehrlichiosis at a national referral center.
J. Clin. Microbiol. 37:558–564.
16. Davidson, W. R., J. M. Lockhart, D. E. Stallknecht, E. W. Howerth, J. E.
Dawson, and Y. Rechav. 2001. Persistent Ehrlichia chaffeensis infection in
white-tailed deer. J. Wildl. Dis. 37:538–546.
17. Dawson, J. E., D. E. Stallknecht, E. W. Howerth, C. Warner, K. Biggie, W. R.
Davidson, J. M. Lockhart, V. F. Nettles, J. G. Olson, and J. E. Childs. 1994.
Susceptibility of white-tailed deer (Odocoileus virginianus) to infection with
Ehrlichia chaffeensis, the etiologic agent of human ehrlichiosis. J. Clin. Mi-
crobiol. 32:2725–2728.
18. Dehio, C., C. Lanz, R. Pohl, P. Behrens, D. Bermond, Y. Piemont, K. Pelz,
and A. Sander. 2001. Bartonella schoenbuchii sp. nov., isolated from the
blood of wild roe deer. Int. J. Syst. Evol. Microbiol. 51:1557–1565.
19. Des Vignes, F., and D. Fish. 1997. Transmission of the agent of human
granulocytic ehrlichiosis by host-seeking Ixodes scapularis (Acari: Ixodidae)
in southern New York state. J. Med. Entomol. 34:379–382.
20. Dumler, J. S., A. F. Barbet, C. P. Bekker, G. A. Dasch, G. H. Palmer, S. C.
Ray, Y. Rikihisa, and F. R. Rurangirwa. 2001. Reorganization of genera in
the families Rickettsiaceae and Anaplasmataceae in the order Rickettsiales:
unification of some species of Ehrlichia with Anaplasma,Cowdria with Ehr-
lichia and Ehrlichia with Neorickettsia, descriptions of six new species com-
binations and designation of Ehrlichia equi and ‘HGE agent’ as subjective
synonyms of Ehrlichia phagocytophila. Int. J. Syst. Evol. Microbiol. 51:2145–
2165.
21. Felek, S., A. Unver, R. W. Stick, and Y. Rikihisa. 2001. Sensitive detection of
Ehrlichia chaffeensis in cell culture, blood and tick specimens by reverse
transcription-PCR. J. Clin. Microbiol. 39:460–463.
22. Foley, J. E., V. Kramer, and D. Weber. 2002. Experimental infection of
dusky-footed wood rats (Neotoma fuscipes) with Ehrlichia phagocytophila
sensu lato. J. Wildl. Dis. 38:194–195.
23. Goethert, H. K., and S. R. Telford III. 2003. Enzootic transmission of the
agent of human granulocytic ehrlichiosis among cottontail rabbits. Am. J.
Trop. Med. Hyg. 68:633–637.
24. Goodman, J. L., C. Nelson, B. Vitale, J. E. Madigan, J. S. Dumler, T. J.
Kurtti, and U. G. Munderloh. 1996. Direct cultivation of the causative agent
of human granulocytic ehrlichiosis. N. Engl. J. Med. 334:209–215.
25. Hodzic, E., D. L. Borjesson, S. Feng, and S. W. Barthold. 2001. Acquisition
dynamics of Borrelia burgdorferi and the agent of human granulocytic ehrli-
chiosis at the host-vector interface. Vector Borne Zoonotic Dis. 1:149–158.
26. Hodzic, E., J. W. Ijdo, S. Feng, P. Katavolos, W. Sun, C. H. Maretzki, D.
Fish, E. Fikrig, S. R. Telford, and S. W. Barthold. 1998. Granulocytic ehr-
lichiosis in the laboratory mouse. J. Infect. Dis. 177:737–745.
27. Hodzic, E., S. Feng, D. Fish, C. M. Leutenegger, K. J. Freet, and S. W.
Barthold. 2001. Infection of mice with the agent of human granulocytic
ehrlichiosis after different routes of inoculation. J. Infect. Dis. 183:1781–
1786.
28. Jahangir, A., C. Kolbert, W. Edwards, P. Mitchell, J. S. Dumler, and D. H.
Persing. 1998. Fatal pancarditis associated with human granulocytic ehrli-
chiosis in a 44-year-old man. Clin. Infect. Dis. 27:1424–1427.
29. Klein, M. B., C. M. Nelson, and J. L. Goodman. 1997. Antibiotic suscepti-
bility of the newly cultivated agent of human granulocytic ehrlichiosis: prom-
ising activity of quinolones and rifamycins. Antimicrob. Agents Chemother.
41:76–79.
30. Lane, R. S., J. Peisman, and W. Burgdorfer. 1991. Lyme borreliosis: relation
of its causative agent to its vectors and hosts in North America and Europe.
Annu. Rev. Entomol. 36:587–609.
31. Lepidi, H., J. E. Bunnell, M. E. Martin, J. E. Madigan, S. Stuen, and J. S.
Dumler. 2000. Comparative pathology and immunohistology associated with
clinical illness after Ehrlichia phagocytophila-group infections. Am. J. Trop.
Med. Hyg. 62:29–37.
32. Levin, M. L., W. L. Nicholson, R. F. Massung, J. W. Sumner, and D. Fish.
2002. Comparison of the reservoir competence of medium-sized mammals
and Peromyscus leucopus for Anaplasma phagocytophilum in Connecticut.
Vector Borne Zoonotic Dis. 2:125–136.
33. Little, S. E., D. E. Stallknecht, J. M. Lockhart, J. E. Dawson, and W. R.
Davidson. 1998. Natural coinfection of a white-tailed deer (Odocoileus vir-
ginianus) population with three Ehrlichia spp. J. Parasitol. 84:897–901.
34. Little, S. E., J. E. Dawson, J. M. Lockhart, D. E. Stallknecht, C. K. Warner,
and W. R. Davidson. 1997. Development and use of specific polymerase
reaction for the detection of an organism resembling Ehrlichia sp. in white-
tailed deer. J. Wildl. Dis. 33:246–253.
35. Lockhart, J. M., W. R. Davidson, D. E. Stallknecht, J. E. Dawson, and E. W.
Howerth. 1997. Isolation of Ehrlichia chaffeensis from wild white-tailed deer
(Odocoileus virginianus) confirms their role as natural reservoir hosts. J. Clin.
Microbiol. 35:1681–1686.
36. Lockhart, J. M., W. R. Davidson, D. E. Stallknecht, J. E. Dawson, and S. E.
Little. 1997. Natural history of Ehrlichia chaffeensis (Rickettsiales: Ehrli-
chieae) in the piedmont physiographic province of Georgia. J. Parasitol.
83:887–894.
37. Madigan, J. E., P. J. Richter, Jr., R. B. Kimsey, J. E. Barlough, J. S. Bakken,
J. S. Dumler. 1995. Transmission and passage in horses of the agent of
human granulocytic ehrlichiosis. J. Infect. Dis. 172:1141–1144.
38. Magnarelli, L. A., J. W. Ijdo, K. C. Stafford III, and E. Fikrig. 1999. Infec-
3600 TATE ET AL. J. CLIN.MICROBIOL.
tions of granulocytic ehrlichiae and Borrelia burgdorgferi in white-tailed deer
in Connecticut. J. Wildl. Dis. 35:266–274.
39. Magnarelli, L. A., J. W. Ijdo, U. Ramakrishnan, D. W. Henderson, K. C.
Stafford III, and E. Fikrig. 2004. Use of recombinant antigens of Borrelia
burgdorferi and Anaplasma phagocytophilum in enzyme-linked immunosor-
bent assays to detect antibodies in white-tailed deer. J. Wildl. Dis. 40:249–
258.
40. Magnarelli, L. A., K. C. Stafford III, J. W. Ijdo, E. Fikrig, J. H. Oliver, Jr.,
H. J. Hutcheson, and J. L. Boone. 1999. Antibodies to granulocytic ehrlichiae
in white-footed and cotton mice in eastern United States. J. Wildl. Dis.
35:259–265.
41. Massung, R. F., and K. G. Slater. 2003. Comparison of PCR assays for
detection of the agent of human granulocytic ehrlichiosis, Anaplasma phago-
cytophilum. J. Clin. Microbiol. 41:717–722.
42. Massung, R. F., R. A. Priestley, N. J. Miller, T. N. Mather, and M. L. Levin.
2003. Inability of a variant strain of A. phagocytophilum to infect mice.
J. Infect. Dis. 188:1757–1763.
43. McQuiston, J. H., C. L. McCall, and W. L. Nicholson. 2003. Zoonosis
update: ehrlichiosis and related infections. J. Am. Vet. Med. Assoc. 223:
1750–1756.
44. Munderloh, U. G., C. M. Tate, M. J. Lynch, E. W. Howerth, T. J. Kurtti, and
W. R. Davidson. 2003. Isolation of an Anaplasma sp. organism from white-
tailed deer by tick cell culture. J. Clin. Microbiol. 41:4328–4335.
45. Munderloh, U. G., J. E. Madigan, J. S. Dumler, J. L. Goodman, S. F. Hayes,
J. E. Barlough, C. M. Nelson, and T. J. Kurtti. 1996. Isolation of the equine
granulocytic ehrlichiosis agent, Ehrlichia equi, in tick cell culture. J. Clin.
Microbiol. 34:664–670.
46. Munderloh, U. G., S. D. Jauron, V. Fingerle, L. Leitritz, S. F. Hayes, J. M.
Hautman, C. M. Nelson, B. W. Huberty, T. J. Kurtii, G. G. Ahlstrand, B.
Grieg, M. A. Mellencamp, and J. L. Goodman. 1999. Invasion and intracel-
lular development of the human granulocytic ehrlichiosis agent in tick cell
culture. J. Clin. Microbiol. 37:2518–2524.
47. Munderloh, U. G., Y. Liu, M. Wang, C. Chen, and T. J. Kurtti. 1994.
Establishment, maintenance and description of cell lines from the tick Ixodes
scapularis. J. Parasitol. 80:533–543.
48. Nicholson, W. L., S. Muir, J. W. Sumner, and J. E. Childs. 1998. Serologic
evidence of infection with Ehrlichia spp. in wild rodents (Muridae: Sigmo-
dontidae) in the United States. J. Clin. Microbiol. 36:695–700.
49. 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.
Microbiol. 33:1797–1803.
50. Pusterla, N., J. S. Chae, R. B. Kimsey, J. B. Pusterla, E. DeRock, J. S.
Dumler, and J. E. Madigan. 2002. Transmission of Anaplasma phagocyto-
phila (human granulocytic ehrlichiosis agent) in horses using experimentally
infected ticks (Ixodes scapularis). J. Vet. Med. B Infect. Dis. Vet. Public
Health 49:484–488.
51. Pusterla, N., R. J. Anderson, J. K. House, J. B. Pusterla, E. DeRock, and
J. E. Madigan. 2001. Susceptibility of cattle to infection with Ehrlichia equi
and the agent of human granulocytic ehrlichiosis. J. Am. Vet. Med. Assoc.
218:1160–1162.
52. Suksawat, J., C. Pitulle, C. Arraga-Alvarado, K. Madrigal, S. I. Hancock,
and E. B. Breitschwerdt. 2002. Coinfection with two Ehrlichia species in dogs
from Thailand and Venezuela with emphasis on consideration of 16s ribo-
somal DNA secondary structure. J. Clin. Microbiol. 39:90–93. (Author’s
correction, 40:3887.)
53. Sun, W., J. W. IJdo, S. R. Telford III, E. Hodzic, Y. Zhang, S. W. Barthold,
and E. J. Fikrig. 1997. Immunization against the agent of human granulo-
cytic ehrlichiosis in a murine model. J. Clin. Investig. 100:3014–3018.
54. Telford, S. R., III, J. E. Dawson, P. Katavolos, C. K. Warner, C. P. Kolbert,
D. H. Persing, A. F. Azad, and E. McSweegan. 1996. Perpetuation of the
agent of human granulocytic ehrlichiosis in a deer tick-rodent cycle. Proc.
Natl. Acad. Sci. USA 93:6209–6214.
55. Unver, A., S. Felek, C. D. Paddock, N. Zhi, H. W. Horowitz, G. P. Wormser,
L. C. Cullman, and Y. Rikihisa. 2001. Western blot analysis of sera reactive
to human monocytic ehrlichiosis and human granulocytic ehrlichiosis agents.
J. Clin. Microbiol. 39:3982–3986.
56. Varela, A. S., D. E. Stallknecht, M. J. Yabsley, V. A. Moore, E. W. Howerth,
W. R. Davidson, and S. E. Little. 2005. Primary and secondary infection with
Ehrlichia chaffeensis in white-tailed deer (Odocoileus virginianus). Vector
Borne Zoonotic Dis. 5:48–57.
57. Villegas, P. 1998. Titration of biological suspensions, p. 248–254. In D. E.
Swayne, J. R. Glisson, M. W. Jackwood, J. E. Pearson, and W. M. Reed
(ed.), A laboratory manual for isolation and identification of avian patho-
gens, 4th ed. American Association of Avian Pathologists, Inc., Kennett
Square, Pa.
58. Walker, D. H., and J. S. Dumler. 1997. Human monocytic and granulocytic
ehrlichioses. Discovery and diagnosis of emerging tick-borne infections and
the critical role of the pathologist. Arch. Pathol. Lab. Med. 121:785–791.
59. Walls, J. J., B. Greig, D. F. Neitzel, and J. S. Dumler. 1997. Natural infection
of small mammal species in Minnesota with the agent of human granulocytic
ehrlichiosis. J. Clin. Microbiol. 35:853–855.
60. Walls, J. J., K. M. Asanovich, J. S. Bakken, and J. S. Dumler. 1998. Serologic
evidence of a natural infection of white-tailed deer with the agent of human
granulocytic ehrlichiosis in Wisconsin and Maryland. Clin. Diagn. Lab. Im-
munol. 5:762–765.
61. Winslow, G. M., E. Yager, S. Konstantin, E. Volk, A. Reilly, and F. K. Chu.
2000. Antibody-mediated elimination of the obligate intracellular bacterial
pathogen Ehrlichia chaffeensis during active infection. Infect. Immun. 68:
2187–2195.
62. Yabsley, M. J., S. E. Little, E. J. Sims, V. G. Dugan, D. E. Stallknecht, and
W. R. Davidson. 2003. Molecular variation in the variable-length PCR target
and 120-kilodalton antigen genes of Ehrlichia chaffeensis from white-tailed
deer. J. Clin. Microbiol. 41:5202–5206.
63. Yabsley, M. J., V. G. Dugan, D. E. Stallknecht, S. E. Little, J. M. Lockhart,
J. E. Dawson, and W. R. Davidson. 2003. Evaluation of a prototype Ehrlichia
chaffeensis surveillance system using white-tailed deer (Odocoileus virginia-
nus) as natural sentinels. Vector Borne Zoonotic Dis. 3:195–207.
64. Yunker, C. E., J. Cory, and H. Meibos. 1984. Tick tissue and cell culture:
applications to research in medical and veterinary acarology and vector-
borne disease, p. 1082–1088. In D. A. Griffiths and C. E. Bowman (ed.),
Acarology VI, vol. 2. Ellis Harwood, Chichester, United Kingdom.
65. Zeidner, N. S., T. R. Burkot, R. Massung, W. L. Nicholson, M. C. Dolan, J. S.
Rutherford, B. J. Biggerstaff, and G. O. Maupin. 2000. Transmission of
human granulocytic ehrlichiosis by Ixodes spinipalpis ticks: evidence of an
enzootic cycle of dual infection with Borrelia burgdorferi in northern Colo-
rado. J. Infect. Dis. 182:616–619.
VOL. 43, 2005 EXPERIMENTAL A. PHAGOCYTOPHILUM IN WHITE-TAILED DEER 3601
    • Patients present an undifferentiated illness characterized by high-degree fever, headache, myalgia and/or arthralgia and lymphadenopathy , associated with bicytopenia and characterized by elevated serum levels of liver enzymes (Koebel et al., 2012). In the North of America, two strains of A. phagocytophilum are reported, the " human-active " strain which is pathogenic for humans, but can also infect both ruminants and mice under experimental conditions , and the " Variant 1 " , which is not infectious for humans, but can infect goats and deer, although not mice (Keesing et al., 2014; Massung et al., 2003 Massung et al., , 2005 Massung et al., , 2006 Tate et al., 2005). In contrast, the European variants infect humans, livestock, as well as rodents and are associated with severe disease and high antibody prevalence in ruminants (Foley et al., 2009).
    [Show abstract] [Hide abstract] ABSTRACT: Rhipicephalus bursa is one of 79 species of the genus Rhipicephalus in the family of Ixodidae. In this study, we investigated Anaplasmataceae bacteria associated with R. bursa collected after an epizootic outbreak of ovine anaplasmosis. 76 adult ticks, (60 male and 16 female ticks), were removed from sheep in two farms and all identified as R. bursa, all females were partially engorged. We found that 50% of the ticks were positive in the initial Anaplasmataceae qPCR screening. Bacterial species was identified by analyzing the sequences of amplicons of 23S rRNA, groEL and rpoB genes. 22.4% of ticks contained DNA of Anaplasma phagocytophilum and 7.9% the DNA of Anaplasma ovis. Based on 23S rRNA and groEL genes analysis, we found that 19.7% of ticks contained a potentially new species of Ehrlichia. We propose the status of Candidatus for this uncultured species and we provisionally name it Candidatus Ehrlichia urmitei. No Wolbachia were identified. These results show that R. bursa can be a carrier of Anaplasmataceae bacteria.
    Article · Sep 2016
    • An American strain infectious for horses is not infectious for ruminants[19], while a European variant pathogenic for cattle does not cause any clinical disease in horses[20]. In the USA, the Ap-Variant 1 infects goats and deer, but not mice[21][22][23], whereas the Ap-ha variant can infect both ruminants and mice under experimental conditions[21,22,24]. Taken together, these results suggest that distinct A. phagocytophilum ecotypes with varying host tropisms, circulate in Europe and the USA.
    [Show abstract] [Hide abstract] ABSTRACT: Background: Anaplasma phagocytophilum is a zoonotic and obligate intracellular bacterium transmitted by ticks. In domestic ruminants, it is the causative agent of tick-borne fever, which causes significant economic losses in Europe. As A. phagocytophilum is difficult to isolate and cultivate, only nine genome sequences have been published to date, none of which originate from a bovine strain.Our goals were to; 1/ develop a sequencing methodology which efficiently circumvents the difficulties associated with A. phagocytophilum isolation and culture; 2/ describe the first genome of a bovine strain; and 3/ compare it with available genomes, in order to both explore key genomic features at the species level, and to identify candidate genes that could be specific to bovine strains. Results: DNA was extracted from a bovine blood sample infected by A. phagocytophilum. Following a whole genome capture approach, A. phagocytophilum DNA was enriched 197-fold in the sample and then sequenced using Illumina technology. In total, 58.9% of obtained reads corresponded to the A. phagocytophilum genome, covering 85.3% of the HZ genome. Then by performing comparisons with nine previously-sequenced A. phagocytophilum genomes, we determined the core genome of these ten strains. Following analysis, 1281 coding DNA sequences, including 1001 complete sequences, were detected in the A. phagocytophilum bovine genome, of which four appeared to be unique to the bovine isolate. These four coding DNA sequences coded for "hypothetical proteins of unknown function" and require further analysis. We also identified nine proteins common to both European domestic ruminants tested. Conclusion: Using a whole genome capture approach, we have sequenced the first A. phagocytophilum genome isolated from a cow. To the best of our knowledge, this is the first time that this method has been used to selectively enrich pathogenic bacterial DNA from samples also containing host DNA. The four proteins unique to the A. phagocytophilum bovine genome could be involved in host tropism, therefore their functions need to be explored.
    Full-text · Article · Nov 2014
    • Experimental studies support the hypothesis that different epidemiological contexts are associated with considerable strain variation [17,18]: an American strain infectious for horses was not infectious for ruminants [19], whereas a European variant pathogenic for cattle did not cause any clinical disease in horses [20]. In the US, the Ap-ha variant, which is pathogenic for humans, can also infect both ruminants and mice under experimental conditions, whereas the Ap-Variant 1, which is not infectious for humans, can infect goats and deer, but not mice21222324. As A. phagocytophilum is not transmitted transovarially in ticks, it is thought to be maintained in vertebrate reservoir hosts.
    [Show abstract] [Hide abstract] ABSTRACT: Anaplasma phagocytophilum is a tick-borne intragranulocytic alpha-proteobacterium. It is the causative agent of tick-borne fever in ruminants, and of human granulocytic anaplasmosis in humans, two diseases which are becoming increasingly recognized in Europe and the USA. However, while several molecular typing tools have been developed over the last years, few of them are appropriate for in-depth exploration of the epidemiological cycle of this bacterium. Therefore we have developed a Multiple-Locus Variable number tandem repeat (VNTR) Analysis typing technique for A. phagocytophilum. Five VNTRs were selected based on the HZ human-derived strain genome, and were tested on the Webster human-derived strain and on 123 DNA samples: 67 from cattle, 7 from sheep, 15 from roe deer, 4 from red deer, 1 from a reindeer, 2 from horses, 1 from a dog, and 26 from ticks. From these samples, we obtained 84 different profiles, with a diversity index of 0.96 (0.99 for vertebrate samples, i.e. without tick samples). Our technique confirmed that A. phagocytophilum from roe deer or domestic ruminants belong to two different clusters, while A. phagocytophilum from red deer and domestic ruminants locate within the same cluster, questioning the respective roles of roe vs red deer as reservoir hosts for domestic ruminant strains in Europe. As expected, greater diversity was obtained between rather than within cattle herds. Our technique has great potential to provide detailed information on A. phagocytophilum isolates, improving both epidemiological and phylogenic investigations, thereby helping in the development of relevant prevention and control measures.
    Full-text · Article · Sep 2014
    • Experimental studies support the hypothesis that different epidemiological contexts are associated with considerable strain variation [17,18]: an American strain infectious for horses was not infectious for ruminants [19], whereas a European variant pathogenic for cattle did not cause any clinical disease in horses [20]. In the US, the Ap-ha variant, which is pathogenic for humans, can also infect both ruminants and mice under experimental conditions, whereas the Ap-Variant 1, which is not infectious for humans, can infect goats and deer, but not mice21222324. As A. phagocytophilum is not transmitted transovarially in ticks, it is thought to be maintained in vertebrate reservoir hosts.
    [Show abstract] [Hide abstract] ABSTRACT: Background: Anaplasma phagocytophilum is a tick-borne intragranulocytic alpha-proteobacterium. It is the causative agent of tick-borne fever in ruminants, and of human granulocytic anaplasmosis in humans, two diseases which are becoming increasingly recognized in Europe and the USA. However, while several molecular typing tools have been developed over the last years, few of them are appropriate for in-depth exploration of the epidemiological cycle of this bacterium. Therefore we have developed a Multiple-Locus Variable number tandem repeat (VNTR) Analysis typing technique for A. phagocytophilum. Methods: Five VNTRs were selected based on the HZ human-derived strain genome, and were tested on the Webster human-derived strain and on 123 DNA samples: 67 from cattle, 7 from sheep, 15 from roe deer, 4 from red deer, 1 from a reindeer, 2 from horses, 1 from a dog, and 26 from ticks.
    Full-text · Article · Aug 2014
    • In the northern USA, the most important An. phagocytophilum reservoirs in the I. scapularis distribution area are the white-footed mice (Peromyscus leucopus) and white-tailed deer (Odocoileus virginianus) (Ravyn et al., 2001; Tate et al., 2005; Telford III et al., 1996). It has been shown that An. phagocytophilum strains differing in their host tropism can circulate in the same location.
    File · Data · Dec 2012 · Journal of wildlife diseases
    • The Ap-ha strains, however, cause human disease and infect white-footed mice. Although a human isolate infected white-tailed deer experimentally (Tate et al., 2005), there is no evidence that Ap-ha strains infect deer in nature. In this study, we used a human isolate of A. phagocytophilum (NCH-1 strain) to produce whole-cell and recombinant p44 antigens.
    [Show abstract] [Hide abstract] ABSTRACT: Whole-blood samples were obtained from 214 white-tailed deer (Odocoileus virginianus) representing 44 sites in Connecticut (USA) during 1992, 1993, 1996, 1999, and 2000 through 2006. Sera were analyzed for total antibodies to whole-cell or recombinant antigens of Borrelia burgdorferi sensu stricto and Anaplasma phagocytophilum, the respective causative agents of Lyme borreliosis and human granulocytic anaplasmosis. Deer sera contained antibodies to both bacteria during different seasons and throughout the 11-yr study. Of the 224 sera tested, 141 (63%) contained antibodies to whole-cell B. burgdorferi in a polyvalent enzyme-linked immunosorbent assay, whereas 124 (55%) were positive to whole-cell A. phagocytophilum by indirect fluorescent antibody staining. Use of highly specific recombinant antigens (VlsE of B. burgdorferi and protein 44 of A. phagocytophilum) provided strong confirmatory results of past or current infections. There was coexistence of antibodies to whole-cell or recombinant antigens of both agents in 72 (32%) sera. Analyses of 18 sera from eight deer that were marked, released, and recaptured, showed minimal changes in antibody titer over sampling time intervals ranging from 17 days to 5.1 yr. Relatively high antibody prevalences for both bacterial agents in different seasons and years reaffirm that there are well-established foci for both tick-borne infections and probably reflect frequent exposure of deer to infected Ixodes scapularis ticks. November and December is a suitable period to obtain blood samples from deer to conduct serosurveillance for both bacteria.
    Full-text · Article · Jul 2010
Show more
Article
December 1991 · Journal of the American Veterinary Medical Association · Impact Factor: 1.56
    Article
    February 1992 · New York state journal of medicine
      Article
      February 1992 · Journal of the South Carolina Medical Association (1975)
        Based on a survey of hunter-harvested deer, the suspected primary vector of Lyme disease in the Southeast, I. scapularis, is most prevalent in sandhill and coastal plain counties of South Carolina. None of 271 I. scapularis examined were found to be infected with the Lyme disease spirochete. However, many more specimens of I. scapularis, A. americanum, and other tick species must be examined... [Show full abstract]
        Article
        December 1991 · Canada diseases weekly report = Rapport hebdomadaire des maladies au Canada
          Discover more