A New Murine Model for Gastrointestinal Anthrax Infection
Citation: Xie T, Sun C, Uslu K, Auth RD, Fang H, et al. (
A New Murine Model for Gastrointestinal Anthrax Infection
Tao Xie 0
Chen Sun 0
Kadriye Uslu 0
Roger D. Auth 0
Hui Fang 0
Weiming Ouyang 0
David M. Frucht 0
Stefan Bereswill, Charite-University Medicine Berlin, Germany
0 Laboratory of Cell Biology, Division of Monoclonal Antibodies, Office of Biotechnology Products, Center for Drug Evaluation and Research, United States Food and Drug Administration , Bethesda, Maryland , United States of America
The scientific community has been restricted by the lack of a practical and informative animal model of gastrointestinal infection with vegetative Bacillus anthracis. We herein report the development of a murine model of gastrointestinal anthrax infection by gavage of vegetative Sterne strain of Bacillus anthracis into the complement-deficient A/J mouse strain. Mice infected in this manner developed lethal infections in a dose-dependent manner and died 30 h-5 d following gavage. Histological findings were consistent with penetration and growth of the bacilli within the intestinal villi, with subsequent dissemination into major organs including the spleen, liver, kidney and lung. Blood cultures confirmed anthrax bacteremia in all moribund animals, with approximately 1/3 showing co-infection with commensal enteric organisms. However, no evidence of immune activation was observed during infection. Time-course experiments revealed early compromise of the intestinal epithelium, characterized by villus blunting and ulceration in the ileum and jejunum. A decrease in body temperature was most predictive of near-term lethality. Antibiotic treatment of infected animals 24 h following high-dose bacterial gavage protected all animals, demonstrating the utility of this animal model in evaluating potential therapeutics.
Recent bioterrorism attacks  have focused research on the
inhalational route of entry, yet there remains scientific utility in
investigating pathogenic mechanisms involved in gastrointestinal
anthrax, as it is widely held that it is primarily the enteric route of
entry that Bacillus anthracis has evolved to exploit [2,3]. Bacillus
anthracis infection is naturally acquired by ruminant herbivores that
are exposed to spores when feeding in contaminated fields [2,3].
Ruminants are considered to be the most susceptible group within
the mammalian class . However, it has not yet been established
when and where spore germination occurs following oral
consumption . We have previously shown that anthrax lethal
toxin (LT), which is produced by vegetative Bacillus anthracis, elicits
rapid breakdown in the gastrointestinal barrier, characterized by
villus blunting, hemorrhage, and ulceration [4,5]. We have since
been eager to develop an informative animal model that could
address the role of anthrax LT, as well as other virulence factors,
in mediating host-pathogen interactions during gastrointestinal
infection in vivo. Unfortunately, the scientific community has
lacked a murine model of gastrointestinal infection that
incorporates administration of vegetative Bacillus anthracis, which could be
used to investigate pathogenicity via this mode of transmission.
Previously reported animal models for anthrax infection have
mainly involved the administration of B. anthracis spores via
inhalational or parenteral routes [6,7,8,9,10,11,12]. Initial studies
with anthrax spores administered via the gastrointestinal route
failed to establish anthrax infection models in various animal
species [13,14,15]. However, there have been recent reports of the
establishment of infections in mice receiving intragastric B.
anthracis spores [16,17]. One group administered 108 spores of
an encapsulated non-toxigenic strain and reported that B. anthracis
expanded in the Peyers patches, eventually disseminating into
various organs. However, this model was not capable of assessing
the roles of LT in promoting virulence during gastrointestinal
infection. Very recently, another model was reported that utilized
intragastric administration of spores embedded in a thiobendazole
paste . Neither of these models assessed administration of
As ruminant animals use bacterial fermentation to facilitate
digestion, we considered the possibility that in the setting of
natural gastrointestinal infection, the upper gastrointestinal tract
would be exposed to large numbers of vegetative bacteria. Bacillus
species have been shown to germinate and thrive in the conditions
present in the rumen . For this reason, it would seem very
likely that B. anthracis spores would germinate and proliferate in the
rumen of infected animals prior to establishing infection. Under
this scenario, exposure of the gastrointestinal barrier to vegetative
bacteria and the toxins they produce would then lead to barrier
penetration and subsequent dissemination.
We herein report that we have modeled this scenario in A/J
mice through gavage of vegetative bacteria from the Bacillus
anthracis Sterne strain. Mice infected with toxigenic bacteria via
this route develop gastrointestinal disease, which leads to
bacteremia and lethal dissemination. Moreover, we demonstrate
that this animal model can be used to assess the efficacy of
Intragastric Administration of Vegetative B. anthracis
Sterne Strain (BaS) Results in Systemic Anthrax Infection
We hypothesized that we could establish gastrointestinal
infections in mice by infecting mice with vegetative bacteria,
thereby mimicking conditions that we consider likely to be
present during digestion in ruminants, the predominant hosts of
infection in nature. To investigate this possibility, we
administered increasing concentrations of vegetative Bacillus anthracis
Sterne strain (BaS) bacteria via gavage. The percentage of mice
that succumbed to this treatment increased in a dose-dependent
manner (Figure 1A). At the highest dose (2.36109), 9 of 10
mice died within 4 days of administration. In contrast, mice
that received the vehicle alone showed no signs of toxicity and
were blood culture negative. The LD50 for infection via this
route in this experiment was approximately 2.36107 bacteria.
One animal succumbed to infection with the lowest
administered dose of 2.36106 bacteria. Interestingly 6/19 mice that
were blood culture positive for BaS had mixed infections with
commensal bacteria; 4/19 were blood culture positive for
Enterococcus faecalis, whereas 2/19 were blood culture positive for
enteric Staphylococcus species (Figures 1B and 1C). All mice
that died in response to intragastric challenge were confirmed to
be blood culture positive for BaS. To confirm that infections
were due to vegetative bacteria, and not heat-resistant spores,
mice were gavaged with 26108 heat-treated bacteria. None of
these mice (0/5) became infected, indicating that bacterial
spores were not responsible for gastrointestinal infections in our
Murine Gastrointestinal Anthrax has Characteristic
We next characterized the clinical features of mice infected
with anthrax via the gastrointestinal route. As shown in
Figure 2A, a decline in body weight was noted following the
16 h fast that preceded the gavage. Thereafter, body weights in
animals gavaged with Bacillus anthracis generally continued a
slow trend downward, whereas the body weights of those
receiving the vehicle alone returned to baseline levels. In
contrast, a sudden drop in body temperature (Figure 2B)
predicted mortality in anthrax-infected animals within 1224 h.
Similar to the decrease in body weight, a decline in appearance
and activity was also noted in anthrax-infected animals as soon
as 24 h post gavage, yet the time until death following the
decrease in appearance/activity was variable, sometimes
exceeding 24 h (Figure 2C). A combined clinical score that
included these body weight, body temperature, and
appearance/activity distinguished infected mice from non-infected
controls starting 24 h post gavage (Figure 2D).
Murine Gastrointestinal Anthrax is Associated with Gross
Intestinal Hemorrhage and Edema
Mice that became moribund following development of anthrax
infection were euthanized and autopsied. In contrast to control
animals that received intragastric administration of vehicle alone
(Figures 3A and 3B, left panels), mice that received BaS showed
evidence of intestinal hemorrhage (Figures 3A and 3B, right
panels). These pathological findings of gross hemorrhage were
similar to those that we have reported in mice treated with anthrax
LT alone [4,5]. In addition, infected mice showed evidence of
edema within the peritoneal cavity (Figure 3A, right panel).
Anthrax Infection Causes Breakdown of the
Gastrointestinal Barrier and Local Bacterial Invasion
Compared to normal findings in vehicle-treated control animals
(Figure 4A), histological evaluation of intestinal tissues from
moribund animals with infection revealed villus blunting and
ulceration, along with fragmentation and disruption of the normal
villus structures (Figures 4B and 4C). In addition, the intestinal
lumens of infected mice were marked by areas of sloughed
epithelial cells, with scattered areas of intraluminal neutrophils
(Figures 4DF). Brown and Brenn staining of intestinal sections
demonstrated that infected animals were characterized by invasion
of the villi with Gram+ rods (Figures 4H and 4I), a feature not
observed in control animals (Figure 4G). No evidence of bacteria
was observed in the Peyers patches of infected mice (data not
Gastrointestinal Anthrax Infection in Mice is not
Associated with Immune Activation
Other than scattered areas of neutrophils observed in the
intestinal lumen (Figure 4F), there was little evidence that intestinal
anthrax infection was marked by inflammatory immune responses.
Reductions in neutrophils and T cell populations were noted in the
small intestines of infected animals; B cell populations trended
lower, but the change was not statistically significant (Figure S1A).
The levels of these cell populations in colonic tissues were either
reduced (B cells) or unchanged (T cells and neutrophils) compared
to untreated control animals (Figure S1B). Moreover, intestinal
anthrax infection in mice did not lead to activation of CD4+ and
CD8+ T cells (i.e., CD44+CD62L2 and/or CD69+, Figures S2A
and S2B), B220+ B cells (MHCII+ or CD40+, Figures S2C and
S2D) or CD11c+ dendritic cells (MHCII+, CD80+, and/or CD86+,
Figures S2E and S2F) obtained from mesenteric lymph node cells
48 h following gavage of Bacillus anthracis compared to those
obtained from PBS-treated animals. Corresponding with these
results, there was no significant increase pro-inflammatory
cytokine production (MCP-1/CCL2, IL-1b, IL-6, TNF-a, and
IFN-c) in cultured tissue samples from the jejunum or ileum of
infected mice (Figure S3), Moreover, there was no observed
increase in mRNA levels of pro-inflammatory markers in jejunum
tissue cultures obtained from mice 48 h following gavage with
Bacillus anthracis compared to control mice (IL-6, IL-1b, CXCL2,
and Nos2, Figure S4). Taken together, these data indicate
suppression of immune responses during infection.
Infection via the Gastrointestinal Route Leads to
Tissues from major organs were subsequently assessed for signs
of pathology and infection. Lungs from infected mice showed
intra-alveolar hemorrhage, edema, and interstitial infiltrates
(Figure 5A), associated with the presence of numerous Gram+
rods (Figure 5B). Liver sections revealed areas of hemorrhage
and bacterial invasion (Figures 5A and 5B). Widespread damage
was also observed in the spleens of infected animals, characterized
by destruction of the normal architecture of the white pulp and
accompanied by accumulation of Gram+ bacteria in the marginal
zone (Figures 5A and 5B). In addition, the kidneys of infected
mice were marked by infection, with large numbers of bacteria
present in the renal glomeruli and interstitial areas (Figure 5B).
Collectively, histopathology in moribund mice was characterized
by hemorrhage, edema, and vascular congestion in a wide array of
tissues, accompanied by marked white pulp depletion in the
spleen. These pathological changes were similar to the
observaFigure 1. Intragastric challenge of B anthracis Sterne strain causes lethal anthrax infection. (A) Cohorts of ten week old female A/J mice
were gavaged with varying doses of vegetative bacteria or with the PBS vehicle alone as indicated. The percentage of surviving/non-moribund mice
was assessed at 6 h intervals for a total of 10 d as shown [p values were calculated using the log-rank (Mantel-Cox) test, * P,0.05, **P,0.01]. (B) A
photographic image was taken of a BHI agar culture of blood from a representative animal that developed a co-infection following BaS gavage. The
arrowhead indicates a bacterial colony of BaS, whereas the arrow points colonies of Enterococcus faecalis. (C) This table summarizes blood culture and
bacterial identification results from 19 moribund animals.
tions in mice with disseminated infection acquired through
inhalational or s.c. injection routes [12,19,20].
Villus Blunting and Ulceration Occur Prior to
We next investigated the effects of gastrointestinal anthrax
infection in time-course experiments. Mice gavaged with a high
dose of bacteria (109) showed little evidence of intestinal damage
during the first 12 h following administration (data not shown).
However, evidence of intestinal damage (hemorrhage, villus
blunting and ulceration) was detectable in a majority of mice as
early as 24 h following gavage (six out of ten animals).
Moreover, intestinal pathology was observed in an entire cohort
of mice (6/6 animals) that displayed no evidence of
dissemination 40 h following gavage (e.g., normal activity, blood culture
negative, and negative Brown and Brenn staining of tissues).
This pathology was characterized by villus blunting and
ulceration (Figures 6B and 6C, respectively). The villus
architecture in infected animals was disrupted compared to that
of uninfected control animals (Figure 6A), with pathological
features that were very similar to what we observed previously
in anthrax LT-treated mice [4,5]. Taken together, these data
are consistent with the primary site of infection being the
epithelium of the gastrointestinal tract.
In additional studies, we compared the effects of infection on
various regions in the intestinal tract, including the jejunum,
ileum and colon. As summarized in Table 1, mild damage to
the jejunum and ileum was noted as early as 24 h post gavage.
The pathological scores in these regions of the small intestine
increased over time, and nearly all moribund animals had
evidence of damage in both of these areas. In contrast,
pathological effects in the colon were rare in the first 48 h
post anthrax gavage, and moribund mice showed only mild to
moderate evidence of ulceration. Taken together, these data
indicate that gastrointestinal anthrax in this murine model
preferentially affects the small intestine compared to the colon.
Murine Gastrointestinal Infection Model has Utility in
Having established a murine gastrointestinal anthrax model,
we next investigated whether this model could be used to
evaluate potential therapeutics. To this end, we investigated the
efficacy of the combination of intraperitoneal amoxicillin and
gentamicin in preventing disseminated infection and death. We
used the identical dose regimen (16 mg/kg subcutaneous
gentamicin per day and 100 mg/kg subcutaneous amoxicillin
three times per day) that we used previously to prevent enteric
bacterial infections following treatment with anthrax LT .
Although all 8/9 mice that received bacterial gavage alone died
with disseminated infection experiment, all of the mice that
received the antibiotic combination survived (Figure 7). This
proof-of-concept experiment demonstrates the utility of this
murine model in investigating the efficacy of potential anthrax
In the natural setting, anthrax infection is generally a disease of
herbivores, which develop gastrointestinal infection while feeding
in spore-infected fields [2,3]. Presumably, Bacillus anthracis has
evolved pathogenic mechanisms to facilitate a gastroenteric route
of entry, yet the lack of informative animal models that mimic
natural gastrointestinal anthrax has been a hurdle for investigating
these mechanisms directly and comprehensively. The murine
model that we have established will be a useful tool in overcoming
We hypothesized that the exposure of gastrointestinal tissues to
vegetative bacteria would better parallel the situation in ruminants
following digestive fermentation, which, in turn, would lead to
improved efficiency in establishing infection. Supporting this
hypothesis, our dose-response experiments reveal that the LD50 for
vegetative bacteria via this route of entry is approximately
2.36107, and an infectious dose as low as 2.36106 can lead to
lethal infection. These results contrast those involving animal
models infected with toxigenic Bacillus anthracis spores via the
gastrointestinal tract, where a dose of 108 could not establish
infection in guinea pigs, rabbits or non-human primates
[13,14,15]. Some reduction of the LD50 for mice has been
reported when the spores are embedded in a thiabendazole paste
prior to intragastric administration, but whether this manipulation
mimics natural infection is unclear. Moreover, it is clear that
consumption of infected meat from animals leads to infection in
carnivores. This natural mode of transmission almost certainly
involves exposure to vegetative bacteria growing in fresh meat,
which contrasts infection acquired by ingesting infectious spores,
which might occur in other settings (e.g., bioterrorism). Whether
humans acquire gastrointestinal anthrax primarily through spores
or vegetative bacteria would depend upon local customs (i.e.,
whether raw animal products or undercooked animal products are
Interestingly, increased infectivity of vegetative bacteria vs.
spores was also observed in A/J mice challenged with BaS
through subcutaneous injection. Whereas the LD50 of BaS
spores via subcutaneous injection is 1.16103 , the LD50 is
even lower than 102 if vegetative bacteria are administered .
We speculate that factors produced by vegetative BaS may
contribute to the early pathogenesis of anthrax infection
acquired subcutaneously and that these pathogenic factors
might be shared with gastrointestinal infection as well. In this
regard, vegetative bacteria produce anthrax LT, which likely
promotes infectivity. We previously demonstrated that
LTtreated animals develop a compromise in the integrity of the
intestinal barrier, marked by intestinal hemorrhage and
ulceration [4,5]. This breakdown is associated with development
of systemic infections with commensal enteric bacteria .
Natural history experiments reveal that the pathology observed
in murine gastrointestinal anthrax is very similar to that
observed in anthrax LT-treated animals. Our findings are
consistent with a model in which LT-mediated breakdown of
the intestinal barrier leads to a portal of entry for enteric
bacteria, including Bacillus anthracis. Interestingly, nearly 1/3 of
animals that developed systemic infection with Bacillus anthracis
were co-infected with other enteric organisms, similar to the
systemic infections that we reported in anthrax LT-treated mice
. This murine infection model will serve as a useful tool in
investigating the role of anthrax LT directly through
comparison of gastrointestinal BaS infection vs. infection with an
LTdeficient BaS strain.
It should be noted that the BaS strain is deficient in the capsule
virulence factor of wild-type Bacillus anthracis. The mice used in the
study were complement-deficient A/J mice, which are susceptible
to BaS. For this reason, the model would not be amenable to
assessing the role of the capsule virulence factor during
gastrointestinal infection. Nevertheless, an animal model that involves the
Sterne strain and small rodents has practical advantages with
regard to space and biohazard considerations. Moreover, this
represents the first practical model that can directly assess the role
of anthrax LT in an infection acquired through administration of
vegetative Bacillus anthracis.
Our findings are consistent with a model in which Bacillus
anthracis infection leads to intestinal ulceration, likely through the
action of anthrax LT [4,5]. This breakdown in the intestinal
barrier allows a portal of entry for dissemination. Interestingly, we
observed no evidence of amplification of infection in the Peyers
patches, which contrasted previous results derived from a murine
model of gastrointestinal anthrax that utilized non-toxigenic spores
, but is consistent with a model that involved spores imbedded
in a thiobendazole paste . Moreover, infection in our model
did not lead to immune activation, likely due to the well-known
immunosuppressive effects of anthrax LT. Infection with
vegetative bacteria in our model leads to a rapid progression to
hematological dissemination, which would be predicted to result
from the observed destruction of the structural integrity of the
musosal barrier and concomitant suppression of host immune
Figure 5. Murine gastrointestinal anthrax progresses to multi-organ infection. Tissue sections from multiple organs were collected from
control or moribund GI-infected mice, and stained with H&E (A). The left columns show representative sections from untreated control mice, whereas
columns on the right show representative sections from moribund infected mice. The star highlights proteinaceous material in the alveolar spaces,
accompanied by alveolar destruction (lung section); white arrows indicate areas of hemorrhage and/or destruction of the normal tissue architecture
(liver, spleen and kidney sections). Shown in B are sections stained with Brown and Brenn. Arrowheads indicate Gram+ rods, which were present
throughout the interstitial areas of the tissues (lung, liver, spleen, and kidney) of infected animals (right column), but not in uninfected control
animals (left column).
Importantly, we provide data from a proof-of-concept
experiment demonstrating that this murine gastrointestinal infection has
utility in assessing therapeutic efficacy in vivo. As the licensing of
potential anthrax therapeutics will likely depend on efficacy data
from animal models, the development of new animal models is of
importance. This toxigenic murine model of gastrointestinal
anthrax could fill an important niche in the tools available to
assess new therapies for anthrax, especially those that target
anthrax ET and/or LT.
Materials and Methods
Animal experiments were performed in accordance with animal
protocol #WO2011-16, which was approved by the United States
Food and Drug Administration Center for Biologics Evaluation
and Research (CBER) Institutional Animal Care and Use
Committee, in accordance with the U.S. Public Health Service
Policy on Humane Care and Use of Laboratory Animals
Figure 6. Gastrointestinal anthrax is marked by histological damage prior to hematological dissemination. Tissues were collected from
mice 2 h (A) or 40 h (B and C) following intragastric challenge with BaS (1.36109). At neither of these time points were these BaS-challenged mice
bacteremic. Representative images generated from H&E-stained sections from mice in each of these cohorts are shown. Asterisks indicate regions of
villous blunting, whereas block arrows indicate focal regions of ulceration.
(Assurance # A4295-01). The CBER animal program is
accredited by Association for Assessment and Accreditation of
Laboratory Animal Care International. Animal welfare was
assessed at least twice per day.
*The pathological scores of the intestinal specimens were classified into four
grades as follows: no lesions seen (2); mild villous blunting and/or isolated
hemorrhages (+); multifocal areas of moderate mucosal ulceration, villus
blunting, and/or hemorrhage (++); severe mucosal ulceration characterized by
severe structural destruction and abundant bacterial colonies in contact or
within submucosal tissues and/or severe hemorrhage (+++).
Mice, Bacterial Strain and Gastrointestinal Challenge
Female A/J mice used in the experiments were obtained from
The Jackson Laboratory (Bar Harbor, ME, USA) and were 912
weeks of age at the time of the experiments. Mice were allowed at
least one week to acclimatize to the animal facilities prior to
experimentation. Mice were housed using standard cages that had
a capacity for up to 5 animals. Food and water were provided ad
libitum, with the exception of the fasting period described below.
Study animals that became moribund were euthanized. Mice were
euthanized either via terminal exanguination under anesthesia
[i.p. administration of ketamine (6070 mg/kg) and xylazine (12
14 mg/kg)], carbon dioxide inhalation in a euthanasia chamber,
or by cervical dislocation by experienced animal handlers (in cases
requiring rapid tissue collection).
Bacillus anthracis (Sterne strain 7702; BaS) was kindly provided by
Dr. Tod Merkel . All experiments with this strain were carried
out using biosafety level 2 procedures. To prepare vegetative BaS
bacteria, 50 mL of frozen stock was incubated in 5 mL BBL Brain
Heart Infusion medium (BHI; Becton, Dickinson and Company,
Franklin Lakes, NJ, BD 221813) at 37uC for 16 h (overnight). The
overnight culture was diluted 1:1 with fresh, pre-warmed BHI
medium and cultured an additional 6 h, at which time the density
was at least 26108. This estimate was based on repeated
experiments with similarly prepared suspensions. However, the
actual challenge dose (colony forming units, CFU) was calculated
post-hoc via dilution and plating on BHI plates. To distinguish
between bacilli and spores in culture, a fraction of the bacterial
culture was heat-treated at 60uC for 30 min to kill vegetative
bacilli. Heat-treated and untreated samples were serially diluted
and plated, and results were recorded as numbers of CFU. Using
the culture conditions described above, we determined that the
concentration of heat-resistant colony forming units (spores)/
vegetative bacteria was 4/108.
Gastrointestinal Infection Experiments
For gastrointestinal challenge experiments, the mice were first
fasted for 16 hours to allow the passage of stomach contents. Mice
were then received 6070 mg/kg intraperitoneal ketamine and
1214 mg/kg xylazine for anesthesia. When the mice were deeply
sedated, 50 ml of 8.5% (W/V) NaHCO3 was administrated, which
was followed immediately by 150 ml of the bacteria suspension
that had been concentrated (4000 g68 min) or diluted in varying
amounts of culture medium (pre-warmed to 37uC) to provide the
desired concentration for administration. As a control to assess a
potential role for bacterial spores, some mice were gavaged with
heat-treated bacteria (60uC for 30 min). Single-use plastic feeding
tubes (20 GA 38 mm, Instech, Plymouth Meeting, PA) were used
for intragastric administration. After the procedure, mice were
returned to barrier housing and received food and water ad libitum.
To eliminate allocation bias in time-course experiments, mice
were pre-segregated into treatment groups corresponding to each
time point to be assessed. A clinical assessment of the mice was
performed by monitoring body weight, rectal temperature,
locomotor activity and general appearance . As described in
the figure legend, these parameters were scored and added
together to provide a combined clinical score.
Fluorescent dyelabeled antibodies to the cell surface markers
CD4 (RM4-5), CD8 (536.7), TCRb (H57-597), CD69 (H1.2F3),
CD62L (MEL-14), CD44 (1M7), B220 (RA3-6B2), MHCII (M5/
114.15.2), CD40 (1C10), CD11c (N418), CD80 (16-10A1) and
CD86 (2F4) were purchased from eBiosciences (San Diego, CA),
along with corresponding isotype control antibodies. Mesenteric
lymph nodes were dissected from euthanized mice and were
digested in collagenase (2 mg/mL; Sigma, St. Louis, MO)
dissolved in DMEM medium (Invitrogen, Grand Island, NY)
containing 5% FBS at 37u for 45 min. After digestion, the cells
were washed twice and resuspended in DMEM medium.
Mesenteric lymph node cells were incubated for 15 min on ice
with these specific antibodies or corresponding isotype control
antibodies using standard methods . All samples were acquired
and analyzed with an LSR II (Becton Dickinson, Franklin Lakes,
NJ) and FlowJo software (TreeStar).
The assay to detect cytokine levels in supernatants of cultured
tissues was adopted from a published method . In brief,
longitudinally-cut intestinal biopsies were washed once with PBS
and three times in RPMI 1640 (with antibiotics, pre-warmed to
37uC; Hyclone Laboratory, Inc., Logan, Utah). Five 1 cm sections
of tissue were then placed in 24-well flat-bottom culture plates
containing 1 mL of serum-free RPMI 1640 medium
supplemented with penicillin (100 mg/ml) and streptomycin (100 mg/ml) and
maintained at 37uC. Following culture for 18 h, culture
supernatants were collected and tested for cytokine concentrations.
Cytokine assays were performed in duplicate using a
customerdesigned Procarta multiplex bead-based kit that assessed MCP-1,
IL-1b, IL-6, TNF-a, and IFN-c (Affymetrix, CA). Bead
fluorescence was measured and cytokine levels analyzed using a BioPlex
200 analyzer (Luminex; Bio-Rad) and BioPlex Manager software
Tissue Section Immunofluorescence Staining
Paraffin embedded tissue sections (5 mm) were prepared from
intestine tissues, and sections were de-paraffinized in xylene and
rehydrated in a series of graded alcohols. Antigen retrieval was
then performed in Antigen Unmasking Solution (H-3300, Vector
Burlingame, CA). Slides were incubated at 4uC overnight at a
1:150 dilution with the following primary antibodies: CD3
(NBP172167, Novus, Littleton, CO), B220 (RA3-6B2, eBioscience, San
Diego, CA) or Gr-1 (RB6-8C5, eBioscience, San Diego, CA).
Following three 10 min washes in PBS, slides were incubated with
Alexa Fluor-labeled secondary antibody (1:500) at room
temperature for 90 min. The stained sections were mounted in Prolong
Gold Anti-fade reagent (P36935, Invitrogen) and examined using a
Keyence BZ-9000 All-in-one fluorescence microscope.
Quantitative Reverse Transcription-PCR
RNA was extracted from frozen tissues using a Mini RNeasy
Kit (Qiagen, Gaithersburg, MD) and treated with DNase
(DNAfree, Ambion, Austin, TX) to remove DNA contamination.
Reverse transcription (RT) was performed using the SuperScript
Vilo cDNA Synthesis kit (11754-050, Invitrogen, Grand Island,
NY). Gene expression levels were measured by quantitative
RTPCR using the ABI Prism 7900 Real-time PCR system (Applied
Biosystems, Foster City, CA). PCR reactions (20 ml total volume)
included cDNA, 100 nM primers and 10 ul of SYBR Green
MasterMix (Applied Biosystems, Foster City, CA). To obtain the
relative quantification of the mRNA of the genes, the mRNA
levels of the genes were normalized to b-actin mRNA levels in
each sample, which were determined simultaneously by the same
method. The following primer pairs were used for RT-PCR:
CXCL2, Forward 59-AACATCCAGAGCTTGAGTGTGA-39
and Reverse 59-TTCAGGGTCAAGGCAAACTT-39;
IL-6, Forward 59-ATGGATGCTACCAAACTGGAT-39 and
Reverse 59-TGAAGGACTCTGGCTTTGTCT-39; IL-1b,
Reverse 59- GATCCACACTCTCCAGCTGCA-39; iNOS2,
Forward 59-CAGCTGGGCTGTACAAACCTT-39and Reverse
59CATTGGAAGTGAAGCGTTTCG; and b-actin, Forward
59TTCCATCATGAAGTGTGACGTT-39 and Reverse 59
Antibiotic Efficacy Experiments
Female A/J mice were infected via gavage with 1.26109 CFU
of vegetative BaS. Twenty four hours later, the infected mice were
treated with amoxicillin and gentamicin or PBS alone for 3
consecutive days. The following antibiotic regimen was used:
16 mg/kg subcutaneous gentamicin per day and 100 mg/kg
subcutaneous amoxicillin three times per day. Mice were
monitored at least every 6 h until the termination of the
experiment (10 days post infection). Blood culture results from
all animals that died showed disseminated anthrax infection.
Mice were euthanized when moribund or at specific time points
post-infection as indicated in the figure legends. The intestines and
other organs were dissected and fixed in neutral buffered formalin.
Paraffin sections were prepared and stained with hematoxylin and
eosin (H&E) and/or Brown and Brenn (B&B) by Histoserv, Inc
(Germantown, MD). The stained slides were scanned using an
Aperio ScanScope (ScanScope, Aperio, CA) and acquired using
206or 406magnification. These digital images were converted
into TIF files for generating figures. The magnification factors are
provided within the figures or figure legends. Histological
assessments and scoring were made by an investigator blinded to
the treatment cohort from which the sections were obtained.
Bacterial Isolation and Identification
Bacterial isolates were cultured on blood agar plates (RO1202,
Remel, Lenexa, KS) using blood samples obtained aseptically from
cardiac puncture. In some cases, blood samples were also cultured
on BHI plates (#221569, Becton Dickinson and Company).
Bacterial identifications were performed using the Omnilog
Microbial Identification System (Biolog Inc., Hayward, CA).
Bacterial colonies were isolated and suspended in a
nutrientdeficient inoculating medium containing tetrazolium redox dyes
provided by Biolog Inc., using the manufacturers recommended
protocol. Suspended bacteria were then incubated in 96 well
MicroPlates provided by Biolog Inc., which were then incubated
in the identification instrument at 33uC for analysis. Identifications
were made through colorimetric analysis of the plates, which
produced a metabolic fingerprint that could be compared with a
software database that included a fingerprint for BaS.
Figure S1 Intestinal anthrax infection is not associated
with inflammatory cell infiltrates in intestinal tissues.
Intestinal tissue sections were prepared from moribund, infected
mice and uninfected control mice. The average numbers of T cells
(positive for CD3), B cells (positive for B220) and neutrophils
(positive for Gr-1) in small intestinal (jejunum) (A) and colonic (B)
sections were generated from six to twelve randomly picked
microscopic fields per animal (206, Keyence BZ-9000
fluorescence microscope). The numbers of analyzed animals are provided
in parentheses on the X-axes. Average values for each cohort are
shown as horizontal bars; statistical significance (p-value) was
determined using the Students t-test.
Figure S2 Bacillus anthracis infection does not activate
T cells, B cell and dendritic cells. Mesenteric lymph node
cells were obtained from mice 48 h following gavage of PBS
(control, C) or BaS (infected, I), stained with relevant antibodies,
and assessed by flow cytometry. (A) T cells were identified through
TCR-b staining (left panels); TCR-b+-gated cells were
subsequently assessed for CD4 and CD8 expression (right panels). (B)
CD4+ and CD8+ subsets, in turn, were assessed for CD44, CD62L
and CD69 expression. Activated T cells are CD44+CD62L2 and/
or CD69+. (C) B cells were identified through B220 staining (left
panels). (D) B220+-gated cells were subsequently assessed for
MHCII and CD40, markers of B cell activation. (E) Dendritic cells
were identified through CD11c staining. (F) CD11c+-gated cells
were subsequently assessed for MHCII, CD80, and CD86
expression, markers of dendritic cell activation.
Figure S3 Intestinal anthrax infection has minimal
effects on inflammatory cytokine production. Samples
from the jejunum and ileum were obtained 48 h following gavage
with BaS (infected, I) or PBS (control, C) and were cultured ex vivo.
Cytokine/chemokine levels in the supernatants of these ex vivo
small intestine cultures (normalized to tissue weight) were assessed
and are shown, with each dot representing the results for one
animal (n = 4 for each treatment group).
Figure S4 Intestinal anthrax infection does not increase
inflammatory gene expression. Using RT-PCR, mRNA
levels of the indicated pro-inflammatory genes were measured in
jejunum samples obtained from mice 48 h following gavage with
PBS (control, open bar,) or BaS (infected, closed bar). mRNA
levels were first normalized to b-actin levels. Mean levels of each
cytokine in PBS controls were arbitrarily assigned a relative level
of 1 (n = 5/group; SEM values are shown).
The authors thank Drs. Wen Jin Wu and Jennifer Swisher for thoughtful
review of the manuscript. We also thank Dr. Tod Merkel for critical
reagents provided for our studies. The information presented in this
manuscript reflects the work of the authors and does not necessarily
represent the views of the U.S. Food and Drug Administration.
Conceived and designed the experiments: DMF TX WO CS. Performed
the experiments: TX RDA KU HF WO CS. Analyzed the data: DMF TX
WO CS. Wrote the paper: DMF TX.
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