Orthopoxvirus variola infection of Cynomys ludovicianus
(North American Black tailed prairie dog)
Darin S. Carroll, Victoria A. Olson, Scott K. Smith, Zach H. Braden, Nishi Patel, Jason Abel,
Yu Li, Inger K. Damon, Kevin L. Karemn
Centers for Disease Control and Prevention, National Centers for Zoonotic and Vector-Borne and Enteric Diseases, Division of High Consequence Pathogens
and Pathology, Poxvirus Program, Atlanta, GA, USA
a r t i c l e i n f o
Received 27 March 2013
Returned to author for revisions
2 May 2013
Accepted 20 May 2013
Available online 27 June 2013
a b s t r a c t
Since the eradication of Smallpox, researchers have attempted to study Orthopoxvirus pathogenesis and
immunity in animal models in order to correlate results human smallpox. A solely human pathogen,
Orthopoxvirus variola fails to produce authentic smallpox illness in any other animal species tested to
date. In 2003, an outbreak in the USA of Orthopoxvirus monkeypox, revealed the susceptibility of the
North American black-tailed prairie dog (Cynomys ludovicianus) to infection and fulminate disease.
Prairie dogs infected with Orthopoxvirus monkeypox present with a clinical scenario similar to ordinary
smallpox, including prodrome, rash, and high mortality. This study examines if Black-tailed prairie dogs
can become infected with O. variola and serve as a surrogate model for the study of human smallpox
disease. Substantive evidence of infection is found in immunological seroconversion of animals to either
intranasal or intradermal challenges with O. variola, but in the absence of overt illness.
Published by Elsevier Inc.
Homo sapiens are the only known natural or permissive host for
smallpox (Orthopoxvirus variola). Heroic efforts in the 20th century
eradicated smallpox from the human population and led to the
eventual consolidation of O. variola stocks into two WHO sanctioned
smallpox research collaborating programs in the US and Russia.
Historically, although suckling mice succumb to infections with
O. variola, no adult rodent or small mammal models using O. variola
have been successfully developed (Mayr and Herrlich, 1960; Murti
and Shrivastav, 1957). O. variola challenges of various non-human
primate species were conducted in the 20th century (Hahon and
Wilson, 1960; Brinckerhoff and Tyzzer, 1906; Noble and Rich, 1969),
and some cause generalized rash, but rarely was mortality observed.
In recent years, efforts to develop new therapeutics and vaccines
for smallpox have relied on surrogate, non-O. variola Orthopoxvirus
animal infection models and one O. variola model in non-human
primates. Despite significant animal models of Orthopoxvirus infec-
tion (Chapman et al., 2010) to date the only lethal animal model
using O. variola infection is a high dose intravenous, or intravenous/
aerosol challenge of non-human primates (Cynomolgus macaques)
(Jahrling et al., 2004). At highest challenge dose (109pfu), disease
onset is rapid, within 3–4 days post-infection, with some attributes of
hemorrhagic disease and is often lethal by 7 days post infection.
A lower dose (108pfu), results in a more typical rash illness with rash
onset occuring at 3–4 days post exposure and rash progression
similar to smallpox. Studies based on this latter challenge model
have used small number of animals, and overall mortality and
disease progression has been variable (Jahrling et al., 2004). An O.
monkeypox respiratory challenge model in non-human primates may
provide a surrogate model for O. variola since clinical disease
progression in this model is more similar to human smallpox than
an intravenous challenge (Dyall et al., 2011; Goff et al., 2011; Nalca
et al., 2010). However, human smallpox was not commonly asso-
ciated with extensive pulmonary manifestations, as seen in the
intrabronchial respiratory O. monkeypox challenges. Additional sur-
rogate models have been researched using non-O. variola orthopox-
viruses to mimic smallpox like disease in various animal models
(reviewed in (Chapman et al., 2010)).
In 2003 a US outbreak of O. monkeypox resulted in the inadvertent
infection of a North American species, Cynomys ludovicianus (Black-
tailed prairie dog) as well as a substantial human outbreak (Reed et al.
2004). Subsequent research studies identified the susceptibility of
this species to O. monkeypox and revealed a clinical course similar to
ordinary (human) smallpox (Hutson et al., 2009; Reynolds et al. 2006).
Evidence stemming from the 2003 US outbreak of monkeypox and
Contents lists available at SciVerse ScienceDirect
journal homepage: www.elsevier.com/locate/yviro
0042-6822/$-see front matter Published by Elsevier Inc.
E-mail address: firstname.lastname@example.org (K.L. Karem).
Virology 443 (2013) 358–362
subsequent laboratory challenge studies of O. monkeypox in Cynomys
confirm this susceptibility and lethality through intranasal O. Mon-
keypox challenge with relatively low viral doses (103pfu). Similar
clinical features, and timecourse of illness, to that previously observed
in ordinary human smallpox and in human monkeypox was observed
in cynomys infected with O. monkeypox (Fig. 1). High susceptibility
of this new world mammal species with an old world virus
(O. monkeypox) promoted the question of whether or not Cynomys is
susceptible to O. variola. In an effort to test this hypothesis, Black-tailed
prairie dogs were obtained and subjected to infection with O. variola
via two routes, intranasal and intradermal scarification.
To determine if prairie dogs are susceptible to O. variola, animals
were infected by intranasal or intradermal routes with 6.6?106pfu
O. variola as described in materials and methods and monitored daily
for signs of illness. Slight erythema was noted in all intradermally
challenged animals at days 2–4 at the inoculation site but no observed
lesion could be identified at day 7 (Fig. 2). In animals challenged
intranasally, slight redness was noted at day 3 post challenge around
the nares, but by day 7 no signs of infection could be identified (Fig. 2).
Otherwise no signs of illness or infection were detectable upon gross
physical examination. Body weight was taken every three days and
indicates no significant weight loss post challenge for any animals
ELISA testing for Orthopoxvirus specific antibody indicates
seroconversion to O. variola in 9 out of 10 challenged animals at
day 21. Of these 9, four were challenged intranasally, and five
Fig. 1. Summary of clinical pathogenesis and stages of illness of smallpox in
humans, and monkeypox in humans, prairie dogs and rhesus maquaces.
Neutralization capacity of sera at day 21 post challenge. Sera collected at day 21
was analyzed for virus neutralization capacity using the array scan system. Titers
reported are that which neutralized 50% of the virus titer.
Sample50% neutralizing titer
VRV-11 IN contl
VRV-12 ID contl
nNaïve sera and prebleeds from all animals tested o1:40 dilution for 50% viral
aVIG titer giving 50% viral neutralization; R²¼0.9699.
Fig. 2. Photos of inoculation sites day 7 post inoculation (a) intranasally and (b) intradermally.
PD weights VarV challenge
VRV-11 IN contl
VRV-12 SC contl
Days post infection
Fig. 3. Weights of animals (in grams) post challenge, measured every 3 days.
D.S. Carroll et al. / Virology 443 (2013) 358–362
intradermally (Fig. 4). Control animals were negative by ELISA.
Serum viral neutralization titers were observed in 5 of the
challenge animals; two intranasally and 3 challenged intrader-
mally. The highest 50% neutralizing titer (1:154) was observed in
two intradermal challenge animals. Positive control (VIG) resulted
in a 50% neutralizing titer of 1:6103 (Table 1).
Viral DNA detection
Real time PCR was performed for O. variola specific sequences
(A36R), and each sample was tested in triplicate. CT values above
40 are considered negative (see methods). As such, all samples
tested were considered negative. A description of the analysis is as
follows. PCR results from control animals were negative in all oral
swab and tissue samples although certain necropsy samples at day
21 did have PCR results which crossed the threshold after cycle 40
(intranasal control: CT442 for both lung and spleen; and intra-
dermal control: CT442 in liver and gonad). Three inoculated
animals also showed PCR results which crossed the threshold after
cycle 40 in oral swabs. At day 3, oral swabs from two intranasally
challenged animals (VRV-04 female, VRV-05 male) and one intra-
dermally challenged animal (VRV-07 female) exhibited CT values
≥42, yet they were not reproducibly seen (o3 of triplicate reactions
crossing the threshold). Day 7 oral swab from the intradermal
challenged animal (VRV 07) had two of triplicate PCR assays cross
the threshold (CT values of 43.53, and 43.77). All other oral swab
samples remained negative throughout the study. Tissues collected
on day 21 post challenge were also tested for the presence of viral
DNA. All tissues did not have PCR assays which crossed the thresh-
old with the exception of three (VRV-07 female, VRV-08 male, VRV-
09 male) intradermally challenged animals in lung, spleen, and
gonad samples (CT values ≥42 in one of three runs per sample in all
tests). The intradermal challenge site (skin) of one animal (VRV-07)
exhibited a CT≥42 in all three replicate reactions from day 21 post
challenge. No viable virus was cultured from oral swabs or tissue
samples (including inoculation site) following two weeks of incuba-
tion on cell culture. Although CT levels above 40 are considered
negative, the analyses of the results are discussed (below).
Historically, laboratory research efforts have tested several animal
species for susceptibility to O. variola. To date, using adult animals,
only non-human primate models have exhibited overt illness in a
manner comparable to smallpox (Hahon and Wilson, 1960; Jahrling
et al., 2004; Noble and Rich, 1969). However, the recent studies
showing the most significant clinical disease require the intravenous
injection of a high infectious dose, (1?108−1?109pfu). The
discovery of a novel, more permissive/representative animal model
system may facilitate the development of next-generation smallpox
vaccines and therapeutics.
The absence of overt illness in Black-tailed prairie dogs
suggests that this host is not permissive for clinical infection by
O. variola. However, seroconversion of 9 out of 10 challenged
animals to O. variola indicates recognition of the virus by the host
immune system and perhaps some level of viral replication. The
highest serologic response was observed in an intradermally
challenged animal (VRV-08), with all intradermally challenged
animals serocoverting, while 4 of 5 intranasally challenged ani-
mals seroconverted (Fig. 4). PCR results are negative for all
samples tested and all samples were negative for viral growth in
tissue culture. PCR values above the threshold (CT440) in control
animals suggests that these are not reflective of viral DNA, but
perhaps some other factor causing low signals. In fact, 9/10
challenged animals seroconverted while unchallenged controls
did not, suggesting that cross contamination of controls is unlikely
and supports CT values above 40 as being negative. Attempts to
correlate seroconversion to sex of animals, challenge route or
evidence of viral replication (PCR or culture) lack support. Immune
induction in 9/10 challenged animals may be considered a mea-
sure of viral replication, or viral gene expression to some degree,
but the ability to establish viral replication in this host remains
speculative since viral DNA was not detected and cultures from
animals were negative. And while cynomys species are extremely
susceptible to some Orthoxpoxviruses, under the challenge con-
ditions presented here they do not appear to be permissive for
overt illness when challenged with O. variola.
Multiple immortalized cell lines are permissive to in vitro
infection with O. variola but in vivo host restriction indicates some
biological barrier to productive infection (Pirsch et al., 1963). In
C. ludivicanus, an increase in challenge dose and/or altered
challenge route may facilitate a more robust infection, as observed
in non-human primate models. However, our attempt was to
develop a model with natural routes of infection and moderate
dosing to mimic human smallpox disease onset and sequelae.
Additional testing of O. variola in this species to develop a model of
human smallpox is not currently being considered. However,
understanding the different host responses to O. variola or O.
monkeypox infection may enable our understanding of virus host
interactions. The results of this study suggest that Black-tailed
prairie dogs are not readily susceptible to clinically relevant
O. variola infection through traditional and natural routes of
Materials and methods
All work with live O. variola was conducted within the max-
imum containment laboratory (biosafety level 4) under the Terms
of Reference of the WHO Collaborating Center for Smallpox and
Other Poxvirus Infections at the World Health Organization (WHO)
collaborating center in Atlanta, GA USA. The facility is reviewed for
safety and biosecurity practices by independent U.S. and WHO
teams on a frequent basis. The work was pre-approved by, and
presented to, the WHO Advisory Committee on Variola Virus
(6 females (VRV-02, VRV-04, VRV-06, VRV-07, VRV-10, and VRV-
12), 6 males (VRV-01, VRV-03, VRV-05, VRV-08, VRV-09, and VRV-11)
were used in this study. The animals were approximately one year
old. The animals were topically treated with permethrin prior to
transport to the CDC, and were cared for in accordance with the CDC
wild-caught(Colorado) black-tailedprairie dogs
VRV-11 IN contl
VRV-12 ID contl
Absorbance Post-Pre OD450nm
Fig. 4. ELISA of anti-Orthopoxvirus antibody in animal sera. Values are reported as
post challenge (day 21) minus pre challenge (day 0) absorbance at 450 nm.
D.S. Carroll et al. / Virology 443 (2013) 358–362
Institutional Animal Care and Use Committee (IACUC) under an
IACUC approved protocol (♯ 1729KARPRAC-A2). Animal handling and
husbandry was performed by properly trained personnel using
biosafety level 4 personal protective equipment (PPE).
Animals were screened for Yersinia pestis, Francisella tularensis,
Bartonella spp. and Rickettsia spp. by polymerase chain reaction
(PCR) as previously described (Stevenson et al., 2003; Kugeler
et al., 2006; Sackal et al., 2008). Briefly, these assays use primer
sets to target the Bartonella and rickettsial citrate synthase gene
(gltA), the F. tularensis pdpD gene, and the Y. pestis pla gene.
In addition, serum from these animals taken prior to study start
was tested by Enzyme-Linked Immunosorbant Assay (ELISA) to
detect IgG antibodies to Orthopoxvirus species.
The prairie dogs were provided with a commercially available
diet (Exotic Nutrition, Newport News VA) and water; both ad
libitum. The food is a high fiber formula developed within GMP
guidelines to replicate the prairie dogs natural high fiber intake in
the wild and contains vitamin D-3, A and E supplements. One or
more of the following was provided approximately three times a
week for dietary enrichment: either Sugar Beet Treats (Exotic
Nutrition), Fruit Kabobs (Exotic Nutrition) containing, fresh sweet
potatoes, alfalfa or Timothy Hay (Exotic Nutrition). In addition, the
animals were provided with Monkey Biscuits (Exotic Nutrition) as
treats on a daily basis as they are an easily measured indicator of
inappetance if refused by the animals.
Animal infections and sampling
All animal manipulations were performed on animals, which
were sedated using 5% inhalant isoflurane. Control animals (n¼2)
were sham infected for each route of infection and observed
for possible non-specific signs (e.g., inoculation site trauma).
Intranasal infection was performed in (n¼4) animals with
approximately 6.6?106pfu of purified O. variola strain Solamain
(BSH-74-Sol) in total volume of 10 ul (5 ul per nostril) adminis-
tered with a pipette. Intradermal infection was performed via
scarification in (n¼4) animals by administration of 6.6?106pfu in
total volume of 10 ul to the skin on the back between the shoulder
blades followed by ten intradermal sticks with a tuberculin syringe
needle. The animals were observed daily for 21 days for inappe-
tance and general health. For sample collection, animals were
anesthetized using 5% isoflurane gas anesthetic in a chamber with
maintenance of anesthesia via nosecone and kept on heating
blanket to maintain core temperature in accordance with IACUC-
approved protocol. The hind limb/groin area was sprayed with 70%
isopropanol. A 28-gauge needle was used to collect 1 ml of blood
from the saphenous vein. The femoral vein was used only when
blood could not be obtained from saphenous vein. Blood was
distributed to a serum separation tube (Fisher Scientific) for serum
collection. Sera was gamma irradiated at 4.4?106rads (1 kill dose
for Orthopoxvirus) prior to being tested by ELISA and neutralization
testing. Animals were bled for serum pre infection and again at
day 21. Oral swab and weight of animals were taken pre infection
and post challenge on day 3, day 7, day 10, day 14, and day 21. At
the conclusion of the study (day 21), all animals were euthanized
and necropsy performed to collect lung, liver, gonads and spleen
tissues for immunopathology analysis. Animal carcasses were
autoclaved and incinerated for disposal.
Real time-PCR and tissue infectivity culture of virus
Samples were assayed by RT-PCR using forward primers,
reverse primers and probe complementary to the conserved
O. variolaA36R (a viral envelope protein) gene. The primer/probe
sequences were selected from a conserved region of A36R
(O. variola BSH75_banu, GenBank L22579) with Primer Express
(version 1.5; Applied Biosystems). These included A36R forward
primer (5′–TTC AGA CTA CCA GTT ATT CTC GGA-3′), A36R reverse
primer (5′–AGC CAG GTA GTC AAG ACA TCA GA-3′), and A36R
probe (5′FAM-CA AAT TGC GCC ACA GAA TCA TCA AC-BHQ13′). The
target sequence is identical in all sequenced strains of O. variola
with the exception of a single nucleotide polymorphism within the
forward primer sequence of the O. variola CHN48_horn strain.
Primers and probe were synthesized in the Biotechnology Core
Facility (CDC, Atlanta GA), utilizing standard phosphoramidite
chemistry. The detection probe contained 5′ reporter molecule
(FAM) and 3′ aminomodifier (Glen Research, Sterling, VA). A 3′
quencher molecule, BHQ1 (Molecular Probes, Eugene, OR), was
conjugated to the 3′ amino group after synthesis. PCR assay
conditions were optimized according to standard protocols (pro-
tocol 04304449, Applied Biosystems, Foster City, CA) by adjusting
primer and probe concentrations, and thermal cycling tempera-
tures/duration. Each reaction (30 μL) contained 1?
Universal PCR Master Mix (Applied Biosystems, Foster City, CA),
1 μl of 25 μmol/μL each primer, 1 μl of 25 μmol/μL TaqMan probe,
and 1 μL of template DNA. Thermal cycling conditions for the
ABI7900 (Applied Biosystems, Foster City, CA): one cycle of 95 1C
for 6 min; followed by 45 cycles of 95 1C for 5 s, and 60 1C for 25 s.
PCR amplification is based on fluorescent emission after anneal-
ing/elongation (60 1C). O. variola BSH75_banu was used as a
positive control. Based on positive controls, CT values were
correlated to genome copies with a CT value of 39.3 being
equivalent to 20 genome copies and CT value of 42.6 equivalent
to 2 genome copies. For this assay, CT values above 40 are
considered negative based on reproducibility (among triplicate
testing and in duplicate testing) and absence of viral growth in
tissue culture (BSC40 cells with 10% RPMI media). Any sample
which caused the fluorescent signal to cross the threshold (even if
it was after cycle 40) was evaluated for viability by growth in
Enzyme-linked immunosorbant assays (ELISA)
ELISA was used to screen animal sera for the presence of
orthopoxvirus specific antibodies
(Hutson et al., 2009). Briefly, microtiter plates (Immulon II) were
coated with 100 μl of O. vaccinia (Dryvax purified) at 0.01 μg/well
in carbonate buffer overnight at 4 1C followed by formalin inacti-
vation on the plate. Blocking buffer was used (PBS 0.01 M, pH 7.4
(GibcoBRL cat. ♯93-0223 DK)+0.05% tween-20, 5% dried skim milk
(DIFCO, ♯232100), 2% normal goat serum (CDC ID♯ CP0523), and 2%
bovine serum albumin (Sigma, A-7030)) and test samples were
tested using 2 fold dilutions of 1:100, 1:200, 1:400 and 1:800 in
assay diluent and incubated for 1 h at 37 1C. A 1:30,000 dilution of
Immunopure A/G conjugate (Pierce ImmunoPuresProtein A/G/
Peroxidase conjugated, ♯ 32490) in assay diluent was added for 1 h
at 37 1C. Detection was performed with TMB microwell peroxidase
substrate (KPL ♯50-76-05) 1-component substrate was added
followed by addition of stop solution (KPL ♯50-85-05) Raw
absorbance was determined at OD 450 nm. Values reported
represent the average of duplicate wells of each dilution run with
paired prebleed sample dilution values subtracted. Standard
deviation in raw absorbance values between dublicate wells in
all samples was o0.035. Positive and negative control sera were
used as assay controls.
D.S. Carroll et al. / Virology 443 (2013) 358–362
High content screening-green fluorescent protein (HCS-GFP) Download full-text
Neutralizing antibody titers against O. vaccinia were measured
using a previously validated and described GFP-based assay
(Johnson et al., 2008). In brief, Vero E6 Cells were diluted with
10% DMEM from a stock suspension to 1.7?105cells/mL, seeded
into a 96 well flat bottom plate and overnight in a 37 1C CO2
incubator to form a cell monolayer. Serial dilutions of the prairie
dog serum samples were made with 2% DMEM. Tubes containing
180 μL of virus (virus only, no serum), and 12 tubes containing
360 μL of media only (no virus or serum) were used as assay
controls. WR-GFP O. vaccinia was thawed on ice for 1 h followed by
three rounds of sonication and dilution in 2% DMEM. This viral was
added (180 μL) to samples and used as plate viral only control.
Samples were briefly rocked and placed in a 37 1C CO2incubator
for 2 h, and rocked once every 30 min. The media was carefully
removed from the cell monolayer (96 well plate) and replaced
with test samples and controls in triplicate (3 wells per sample).
After a 2 h incubation, the inoculum was aspirated and replaced
with 100 μl of 2% DMEM +46 μg/mL cytosine arabinoside (Ara-C)
and then placed in a 37 1C CO2incubator for 18 h. The cells were
formalin fixed and stained with DAPI (3 mM). Two channel
analysis was performed for DAPI and GFP signals. The plate was
analyzed on the Array Scan HCS Reader with the target acquisition
software from Cellomics (Thermo Scientific: Cellomics, Pennsylvania).
Prism 5.0 (GraphPad) was used to confirm the Gaussian distribution
of our data. Because the data passed the D'Agostino-Pearson omnibus
test, which measures the deviation from a predicted Gaussian
distribution by skewness (asymmetry) and kurtosis (shape), further
statistics were performed assuming a Gaussian “normal” distribution.
The column statistics option was used to determine means, medians,
standard deviations, and standard error from the mean measure-
ments of our data. Additional statistics utilized in this paper include
The authors would like to acknowledge the assistance of the
following individuals in accomplishing this project: Allison
Williams DVM and Eddie Jackson of the Roybal ARB staff for
animal husbandry. This work was funded through program funds
at the poxvirus program in the Centers for Disease Control and
Prevention. The findings and conclusions in this report are those of
the authors and do not necessarily represent the views of the Centers
for Disease Control and Prevention.
Brinckerhoff, W.R., Tyzzer, E.E., 1906. Studies upon the Immunity Reactions of the
Monkey after Inoculation with Vaccine or with Variola Virus: Part III. J. Med.
Res. 14, 321–339.
Chapman, J.L., Nichols, D.K., Martinez, M.J., Raymond, J.W., 2010. Animal models of
orthopoxvirus infection. Vet. Pathol. 47, 852–870.
Dyall, J., Johnson, R.F., Chen, D.Y., Huzella, L., Ragland, D.R., Mollura, D.J., Byrum, R.,
Reba, R.C., Jennings, G., Jahrling, P.B., Blaney, J.E., Paragas, J., 2011. Evaluation of
monkeypox disease progression by molecular imaging. J. Infect. Dis. 204,
Goff, A.J., Chapman, J., Foster, C., Wlazlowski, C., Shamblin, J., Lin, K., Kreiselmeier,
N., Mucker, E., Paragas, J., Lawler, J., Hensley, L., 2011. A novel respiratory model
of infection with monkeypox virus in Cynomolgus macaques. J. Virol. 85,
Hahon, N., Wilson, B.J., 1960. Pathogenesis of variola in Macaca irus monkeys. Am. J.
Hyg. 71, 69–80.
Hutson, C.L., Olson, V.A., Carroll, D.S., Abel, J.A., Hughes, C.M., Braden, Z.H., Weiss, S.,
Self, J., Osorio, J.E., Hudson, P.N., Dillon, M., Karem, K.L., Damon, I.K., Regnery, R.
L., 2009. A prairie dog animal model of systemic orthopoxvirus disease using
West African and Congo Basin strains of monkeypox virus. J. Gen. Virol. 90,
Jahrling, P.B., Hensley, L.E., Martinez, M.J., Leduc, J.W., Rubins, K.H., Relman, D.A.,
Huggins, J.W., 2004. Exploring the potential of variola virus infection of
Cynomolgus macaques as a model for human smallpox. Proc. Natl. Acad. Sci.
USA 101, 15196–15200.
Johnson, M.C., Damon, I.K., Karem, K.L., 2008. A rapid, high-throughput vaccinia
virus neutralization assay for testing smallpox vaccine efficacy based on
detection of green fluorescent protein. J. Virol. Methods 150, 14–20.
Kugeler, K.J., Pappert, R., Zhou, Y., Petersen, J.M., 2006. Real-time PCR for Francisella
tularensis types A and B. Emerg. Infect. Dis. 12, 1799–1801.
Mayr, A., Herrlich, A., 1960. Cultivation of the variola virus in the infantile mouse.
Arch. Gesamte. Virusforsch. 10, 226–235.
Murti, B.R., Shrivastav, J.B., 1957. A study of biological behaviour of variola virus. II.
Experimental inoculation of laboratory animals. Indian J. Med. Sci. 11, 580–587.
Nalca, A., Livingston, V.A., Garza, N.L., Zumbrun, E.E., Frick, O.M., Chapman, J.L.,
Hartings, J.M., 2010. Experimental infection of Cynomolgus macaques (Macaca
fascicularis) with aerosolized monkeypox virus. PloS One 5.
Noble, Jr., J., Rich, J.A., 1969. Transmission of smallpox by contact and by aerosol
routes in Macaca irus. Bull. WHO 40, 279–286.
Pirsch, J.B., Mika, L.A., Purlson, E.H., 1963. Growth characteristics of variola virus in
tissue culture. J. Infect. Dis. 113, 170–178.
Reed, K.D., Melski, J.W., Graham, M.B., Regnery, R.L., Sotir, M.J., Wegner, M.V.,
Kazmierczak, J.J., Stratman, E.J., Li, Y., Fairley, J.A., Swain, G.R., Olson, V.A.,
Sargent, E.K., Kehl, S.C., Frace, M.A., Kline, R., Foldy, S.L., Davis, J.P., Damon, I.K.,
2004. The detection of monkeypox in humans in the Western Hemisphere. N.
Engl. J. Med. 350, 342–350.
Reynolds, M.G., Yorita, K.L., Kuehnert, M.J., Davidson, W.B., Huhn, G.D., Holman, R.C.,
Damon, I.K., 2006. Clinical manifestations of human monkeypox influenced by
route of infection. J. Infect. Dis. 194, 773–780.
Sackal, C., Laudisoit, A., Kosoy, M., Massung, R., Eremeeva, M.E., Karpathy, S.E., Van
Wyk, K., Gabitzsch, E., Zeidner, N.S., 2008. Bartonella spp. and Rickettsia felis in
fleas, Democratic Republic of Congo. Emerg. Infect. Dis. 14, 1972–1974.
Stevenson, H.L., Bai, Y., Kosoy, M.Y., Montenieri, J.A., Lowell, J.L., Chu, M.C., Gage, K.L.,
2003. Detection of novel Bartonella strains and Yersinia pestis in prairie dogs
and their fleas (Siphonaptera: Ceratophyllidae and Pulicidae) using multiplex
polymerase chain reaction. J. Med. Entomol. 40, 329–337.
D.S. Carroll et al. / Virology 443 (2013) 358–362