https://orcid.org/0000-0002-5285-0432
Figures
Abstract
Bacillus cereus biovar anthracis (Bcbva) causes anthrax-like disease in animals, particularly in the non-human primates and great apes of West and Central Africa. Genomic analyses revealed Bcbva as a member of the B. cereus species that carries two plasmids, pBCXO1 and pBCXO2, which have high sequence homology to the B. anthracis toxin and polyglutamate capsule encoding plasmids pXO1 and pXO2, respectively. To date, only a few studies have investigated the effect of variations in Bcbva sporulation, toxin, and capsule synthesis on animal and macrophage pathogenicity compared to B. anthracis, therefore more research is needed to gain a better understanding of the pathogenesis of this emerging infection. Here, we report that Bcbva can multiply and vegetatively survive on nutrient-rich media for a minimum of six days while generating spores. Sporulation of Bcbva occurred faster and more extensively than B. anthracis Ames. Bcbva tended to secrete less protective antigen (PA) than B. anthracis Ames when cultured in growth medium. We found Bcbva produced a substantially higher amount of attached poly-ƴ-D-glutamic acid (PDGA) capsule than B. anthracis Ames when grown in medium supplemented with human serum and CO2. In a phagocytosis assay, Bcbva spores showed reduced internalization by mouse macrophages compared to B. anthracis Ames. Our research demonstrated that Bcbva is more virulent than B. anthracis Ames using two in vivo models, Galleria mellonella larvae and guinea pigs. Following that, the efficacy of the veterinary vaccine Sterne strain 34F2 against anthrax-like disease was assessed in guinea pigs. Sterne vaccinated guinea pigs had significantly increased anti-PA titers compared to the unvaccinated control group. Toxin neutralizing antibody titers in vaccinated guinea pigs correlated with anti-PA titers. This indicates the Sterne vaccine provides adequate protection against Bcbva infection in laboratory animals.
Author summary
Bacillus cereus biovar anthracis (Bcbva) is a Gram-positive, spore-forming bacteria that causes anthrax-like disease in several animal species including non-human primates. It is listed as a select agent by the US Centers for Disease Control and Prevention (CDC). In sub-Saharan African rainforests, Bcbva is considered highly virulent with high mortality rates across multiple animal species. Bcbva shares key features with B. anthracis, the classical cause of anthrax, including spore production and environmental transmission. Our study demonstrates that Bcbva can replicate, sporulate, and persist in growth media for several days. We showed that Bcbva produced a greater amount of attached capsule when grown in culture media supplemented with human serum and CO2. Additionally, Bcbva caused higher mortality in laboratory animals infected with an equal number of spores compared to B. anthracis Ames. Nevertheless, Bcbva infection in animal models was prevented with the global standard Sterne anthrax vaccine. Further study should focus on the Bcbva lifecycle and transmission pathways in nature to elucidate new preventative strategies for controlling disease outbreaks.
Citation: Jiranantasak T, Bluhm AP, Chabot DJ, Friedlander A, Bowen R, McMillan IA, et al. (2024) Toxin and capsule production by Bacillus cereus biovar anthracis influence pathogenicity in macrophages and animal models. PLoS Negl Trop Dis 18(12): e0012779. https://doi.org/10.1371/journal.pntd.0012779
Editor: Georgios Pappas, Institute of Continuing Medical Education of Ioannina, GREECE
Received: April 10, 2024; Accepted: December 11, 2024; Published: December 23, 2024
This is an open access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.
Data Availability: All data is presented in the manuscript and raw data for figures is present in the supporting information.
Funding: This work was supported by the US Defense Threat Reduction Agency Science and Technology New Initiatives contract #HDTRA121C0033 to M.H.N and J.K.B. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. There was no additional external funding received for this study.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Anthrax occurs nearly worldwide in livestock and wildlife with spillover into humans [1–3]. The disease mainly affects wild and domestic herbivores and is characterized by septicemia and sudden death [4]. Ungulates are frequently infected and are thought to acquire infection by ingestion of contaminated anthrax spores from the environment during feeding [4]. The primary routes of anthrax infection are the cutaneous, inhalational or gastrointestinal routes. Animals die from septicemia due to high concentrations of bacteria and robust secretion of anthrax toxins [5]. The carcasses release vegetative cells of B. anthracis in bloody discharge and body fluids emanating from the nostrils, mouth and anus into the environment after the host succumbs [6]. During decomposition, the vegetative cells within the infected carcasses may be exposed to the environment and spore forming conditions by scavenger animals or gas build up that rupture the abdomen [4,6]. Spores contaminate the area and can persist in the soil for years, creating a locally infectious zone [7,8]. Our previous study found that B. anthracis can multiply outside of the soil and persist on environmental substrates including rocks and leaves for at least seven days in controlled experiments [9].
For classical anthrax, the most effective intervention is sustained, preemptive veterinary vaccine programs for high-risk animal populations in enzootic areas [10]. In sub-Saharan Africa where anthrax risk is high, most livestock vaccination rates are extremely low [1]. Many anthrax vaccination programs are typically implemented reactively after an outbreak starts [1,11]. Currently, Sterne strain 34F2 is the most commonly used vaccine strain for livestock [12]. It is an attenuated vaccine based on a B. anthracis strain lacking the pXO2 plasmid but maintaining pXO1, an acapsular toxigenic live spore vaccine [13]. Once vaccinated, animals usually develop humoral immunity within 4 weeks [14]. Outside of livestock around the world, off label Sterne vaccine is used in some wildlife [15,16]. Annual vaccination is recommended as the immunity is not long-lived, yet over the last century, successful vaccination campaigns helped reduce the incidence of anthrax in livestock [12,17,18] and in parallel, human disease [19].
Until recently, non-human primate deaths from anthrax were rarely reported [20–22]. Increased anthrax rates in non-human primates have coincided with the discovery of Bacillus cereus biovar anthracis (Bcbva). Bcbva is the cause of anthrax-like disease in a variety of animal species including chimpanzees (Pan Troglodytes) [21,22], gorillas (Gorilla gorilla) [22], king colobus monkeys (Colobus polykomos) [23], sooty mangabeys (Cercocebus atys) [23], duikers (cephalophus spp.) [24], mongooses (Herpetes spp.) [24], elephants (Loxodonta African) [25], goats (Capra hircus) [25] and porcupines (Atherurus spp.) [24] among other animals in West and Central Africa. Bcbva was first discovered as a cause of chimpanzee mortality in Taї National Park, Côte d’Ivoire in 2001 [21]. Later, there were additional great ape deaths from Bcbva infection in Dja Reserve, Cameroon in 2004 [22]. Subsequently, Bcbva was found to be more widely distributed through the tropical forests of sub-Saharan African as it was detected in a moribund domestic goat in Democratic Republic of the Congo, and in great apes and an elephant in Central African Republic [25]. A recent study revealed more than 38% of the carcasses found in Taï National Park from 2001 to 2015 were associated with Bcbva [24]. The detection of low antibody rates and high anthrax-like mortality in wildlife across multiple species suggested that Bcbva is highly virulent and the causative agent in sub-Saharan African rainforests [26]. While no human anthrax cases caused by Bcbva have been reported nearly 10% of sera from humans living in Taï National Park tested positive for antibodies against a 35-kDa Bcbva-specific secreted protein, pXO2-60 [5,27]. Most study participants had a history of animal contact, but it was not significantly correlated with Bcbva seroprevalence [27]. This differs from B. anthracis exposure, where human cases are associated with handling infected animals or carcasses [28,29]. Whether these differences are the consequence of virulence factor diversity, human exposure, or lack of diagnostic capacity in the outbreak areas remains to be elucidated.
Bacillus cereus biovar anthracis is a Gram-positive, spore forming, rod-shaped bacterium closely related to B. cereus and B. anthracis. It harbors a B. cereus-like chromosomal background but contains two virulence plasmids, pBCXO1 and pBCXO2. The plasmids are homologous to pXO1 and pXO2 of B. anthracis at 99–100% nucleotide identity [25,30]. Plasmid pBCXO1 encodes the three toxin components, lethal factor (LF), edema factor (EF), and protective antigen (PA), whereas pBCXO2 is responsible for production of the poly-ƴ-D-glutamic acid (PDGA) capsule, respectively [20,22,25,30,31]. Besides PDGA capsule, Bcbva produces a hyaluronic acid capsule similar to B. cereus encoded by hasACB on the pBCXO1 plasmid [30,31]. Both PDGA and hyaluronic acid capsule are co-regulated by the global transcriptional regulator, AtxA [31,32]. Conversely, B. anthracis cannot generate the hyaluronic acid capsule due to a frameshift mutation in hasA causing premature termination of translation [31,33,34] and PDGA capsule expression is only activated by AtxA in the presence of CO2 and bicarbonate [31,32]. During anthrax infections, B. anthracis vegetative cells secrete two exotoxins which are required for pathology and mortality. Lethal toxin (LT) is comprised of protective antigen (PA) and lethal factor (LF), while edema toxin (ET) consists of PA and edema factor (EF) [20,30]. Upon toxin secretion, PA binds to the receptor on the host cells and is cleaved into truncated PA monomers by a furin-like protease enabling pre-pore formation. PA heptamers competitively bind with LF and EF to form complexes allowing their translocation into the cell by endocytosis. Once inside of the host cell, the PA heptamers form a pore at the surface of the endosome resulting in release of LF and EF into the host-cell cytoplasm. LT causes host lethality through systemic action of the LF zinc metallopeptidase activity which cleaves mitogen-activated protein kinase-kinases and other peptide hormones leading to shock and death [35,36]. ET causes edema attributed to the activity of EF as a calmodulin-independent adenylate cyclase which raises intracellular cyclic AMP levels [37,38]. Like B. anthracis, previous studies demonstrated Bcbva from both Côte d’Ivoire and Cameroon expressed PA by growing bacteria in bicarbonate medium supplemented with CO2 [20,31]. Phenotypically, both Bcbva and B. anthracis strains are non-hemolytic in contrast to typical B. cereus [20]. Bcbva strains have variable motility characteristics and are resistant to ƴ-phage, while B. anthracis is non-motile and susceptible to ƴ-phage [20]. In addition, Bcbva exhibits a null-plcR genotype like B. anthracis while the plcR transcriptional regulator in B. cereus is functional, affecting the hemolysis phenotype [30]. Bacillus anthracis is usually sensitive to β-lactam antibiotics, such as penicillin, and quinolones including ciprofloxacin or doxycycline whereas most Bcbva show resistance to β-lactam antibiotics and intermediate sensitivity to amoxicillin-clavulanic acid [20,25,27,30]. Beyond these genetic and phenotypic characteristics, few studies have explored the sporulation properties and kinetics, pathogenic mechanisms in infected macrophages, and lethal doses of Bcbva in animal models, necessitating further investigation [31,39].
In the United States, B. anthracis is stringently controlled by the Centers for Disease Control and Prevention (CDC) and is considered an overlap select agent because of its potential threat to human and animal health [40]. High genetic similarity between the virulence plasmids of Bcbva and B. anthracis, and the ability of Bcbva to cause anthrax in animals have led to its inclusion on the US CDC Tier 1 select agent list [41]. Previous studies revealed dormant Bcbva spores persist in animal components such as bones and teeth for several decades [23,24] and, to date, there is no licensed vaccine available for Bcbva infection in animals.
Understanding the pathogenesis of Bcbva allows for development and testing of efficient intervention strategies to prevent anthrax-like disease. The objectives of this study were to compare sporulation, capsule production, and virulence of Bcbva from Côte d’Ivoire and B. anthracis Ames using in vitro and in vivo models. The median lethal dose (LD50) of Bcbva and mortality data for inhalation route in small animals were investigated. Additionally, since the capsule and toxins produced by Bcbva are homologous to those expressed by B. anthracis, we examined whether the Sterne vaccine could provide sufficient protection against anthrax-like disease cause by Bcbva in inhalation-infected animals, which may be an essential consideration for developing any sustained livestock or wildlife anthrax preventive program in Bcbva enzootic areas.
Materials and methods
Ethics statement
Animal challenges were performed at the Rocky Mountain Regional Biosafety Laboratory at Colorado State University under Colorado State University Institutional Animal Care and Use Committee (IACUC) protocol #1561 following ABSL3 practices and procedures.
Bacterial strains
Strains Bcbva UFBc0009 (replicate UFBc0009.1) and B. anthracis Ames UF00738 (replicate UF00738.1) are curated in the Martin E. Hugh-Jones Bacillus anthracis Collection at the Emerging Pathogen Institute, University of Florida. Bcbva UFBc0009 was recovered from mandible tissue and associated teeth from a monkey carcass (Cercocebus atys) in Taï National Park, Côte d’Ivoire between 1993 to 1994 [23]. The original strain of B. anthracis Ames UF00738 was isolated from a heifer which died of anthrax in Jim Hogg County, Texas in 1981 [42,43] and has long been used as a lab reference strain. All bacterial strains were manipulated using biosafety level 3 (BSL3) standard operating procedures following the Biosafety in Microbiological and Biomedical Laboratories 6th edition [44].
Spore production and purification
Starter cultures were inoculated in 3 mL of brain heart infusion (BHI) broth (Difco, BD Life Sciences, Sparks, MD, USA) in sterile 15 mL tubes with 0.22 μm ventilated caps (CELLTREAT Scientific Products, Pepperell, MA, USA). The inoculations were shaken at 220 rpm at 37°C overnight. 200 μl aliquots of culture were spread on Difco sporulation media (DSM) agar [45] with a total of 10 plates for each strain and incubated at 37°C for 6 days. The sporulation efficiency of each plate was measured with wet mount microscopy and was always ≥99% complete prior to collection. Spores from each plate were harvested in 2 ml of ice-cold sterile MilliQ water and combined into a single tube. The spore suspension was pelleted by centrifugation at 4000 x g at 4°C for 10 min. The supernatant was removed, and the pellets were purified through a sodium diatrizoate gradient (MP Biomedicals, LLC., Slon, OH, USA) as previously described [46,47]. The spore pellets were resuspended in 95% ethanol and incubated at room temperature for 1 h with 1 min of vortexing at 15-min intervals to kill any residual vegetative bacilli. The pellets were washed 3 times and resuspended with ice-cold sterile MilliQ water. The spore suspensions were kept at 4°C until further use. The number of viable spores was identified by serial dilution in sterile 1x phosphate buffer saline with 0.05% (v/v) Tween-20 (PBST), plated on tryptic soy agar (TSA) (Research Products Inter, Mount Prospect, IL, USA), and incubated at 37°C overnight.
Sporulation assay
Strains were inoculated and grown to stationary phase in BHI broth shaken at 220 rpm at 37°C overnight. The cultures were adjusted to an OD600 of 1 and subcultured at 1:100 in 3 ml of heart infusion broth (HIB) (Research Products International, Mt. Prospect, IL, USA) in duplicate. The inoculum was incubated at 37°C until processed at 1-, 2-, and 6-day post incubation. At each timepoint, 100 μl aliquots of each culture were serially diluted in PBST, 100 μl from each dilution was plated on TSA. Another 300 μl aliquots of each inoculation were collected and mixed with 700-μl of absolute ethanol to obtain a final concentration of 70% (v/v). The mixtures were incubated at room temperature and vortexed for 1 min at 15 min intervals for 1 hour to ensure homogenous exposure of ethanol throughout the mixtures. Ethanol mixed samples were serially diluted in sterile PBST, and 100 μl from each dilution was plated on a TSA. All plates were incubated at 37°C overnight to enumerate bacterial colony forming units (CFU). The total number of bacterial cells and spores was determined by multiplying CFU by the dilution factor. This assay was conducted in two independent experiments, and three replicate cultures of bacterial strains were grown in each experiment.
Internalization assays
Murine macrophage cell lines RAW264.7 (American Type Culture Collection, ATCC) was seeded in a 96-well CellBIND plate (Corning, Corning, NY, USA) at 25,000 cells/well in Dulbecco’s Modified Eagle Medium (DMEM)-high glucose + L-glutamine (Gibco, Life Technologies, Grand Island, NY, USA) supplemented with 10% (v/v) fetal bovine serum (FBS) (R&D Systems, Flowery Branch, GA, USA). The plate was incubated at 37°C with 5% CO2 overnight to allow monolayer formation. RAW264.7 cells were infected in triplicate with Bcbva UFBc0009 and B. anthracis Ames UF00738 spores at a multiplicity of infection (MOI) of 20 in 50 μl of DMEM without FBS for 45 min to enhance gravity assisted contact between spores and macrophages. Spores do not germinate in DMEM media alone [48–50]. After the infection, DMEM containing 10% FBS, 0.5 mM L-alanine, 1 mM inosine, and 100 μg/ml gentamicin was added and incubated for 15 min. L-alanine and inosine are germinants and play an important role in initiating the germination of B. cereus and B. anthracis endospores [51,52]. In addition, the germination of B. anthracis spores is faster with the addition of serum to the cell culture growth medium [48]. The combination of L-alanine, inosine, and serum can cause B. anthracis and B. cereus spores to germinate within 15 min [48,51,53]. Upon spore germination, the extracellular vegetative cells were killed by gentamycin [50]. Then, the supernatant was removed, and macrophages were incubated for another 2 h in DMEM plus 10% FBS. The cells were washed with phosphate buffer saline (PBS) 3 times between each step. To lyse the cells, 0.1% (v/v) TritonX-100 (Amresco, Solon, OH, USA) was added for 15 min, and supernatants were serially diluted in PBST. All serial dilutions were plated on TSA for counting the number of bacterial cells.
Lactate Dehydrogenase (LDH)-Mediated Cytotoxicity Test
During anthrax infection, the excessive toxin production during bacterial multiplication leads to cellular damage and cell death. Cytotoxicity is commonly determined by measuring the amount of LDH, a stable cytoplasmic enzyme, that rapidly released from damaged cells into the cell culture supernatant [54]. To determine LDH activity, RAW264.7 cells were seeded as described above. Spore suspensions of Bcbva UFBc0009 and B. anthracis Ames UF00738 in DMEM were added to macrophages at an MOI of 20 in six replicates for each strain and incubated for 45 min. After incubation, DMEM supplemented with 10% FBS, 0.5 mM L-alanine, 1 mM inosine, and 100 μg/ml gentamicin was added and incubated for 15 min. The plate was washed with PBS three times between each step. Fresh DMEM containing 10% FBS was added and incubated further for 16 h. For detecting a maximum LDH activity, 10 μl of 10X lysis solution was added into one set of six replicate wells. Next, 10 μl of sterile ultrapure water was added into another set of six replicate wells to measure spontaneous LDH activity. After incubation, 50 μl of supernatant from each well were transferred to a fresh 96-well plate for measurement of LDH by using CytoTox 96 Non-Radioactive Cytotoxicity Assay following the manufacturer ‘s recommendations (Promega Corporation, WI, USA). Briefly, 50 μl of CytoTox 96 reagent was added to each well and incubated for 30 min at room temperature. Then, 50 μl of stop solution was added to each well to stop the reaction. The level of LDH was detected by measuring the absorbance signal at 490 nm in a spectrophotometer (Agilent Biotek Synergy HTX Multi-Mode Microplate Reader, Agilent Technologies, Inc., Santa Clara, CA, USA).
Protective antigen production
Bacterial cultures were grown in BHI broth at 37°C and shaken at 220 rpm overnight after which they were adjusted to an OD600 of 1 and subcultured at 1:100 in 3 ml of HIB broth or BHI broth in duplicates. The common components across these media are heart extract, digastric enzyme, and electrolytes. BHI is supplemented with glucose as a source of sugar. The comparison of cultures grown in HIB and BHI media was to observe how glucose levels could impact PA secretion without elevation of CO2/bicarbonate in Bcbva compared to B. anthracis. Briefly, the cultures were supplemented with cOmplete, EDTA-Free Protease Inhibitor Cocktail (Roche Diagnostics GmbH, Mannheim, Germany), to a final concentration of 1x following the manufacturer’s instruction. At 24 h post incubation, 1 ml of each culture was collected and spun down at 14,000 x g for 10 min. The supernatant was harvested and filtered through low volume 0.45 μm PVDF filters (Millipore Sigma) followed by Spin-X 0.22 μm filters at 10,000 x g for 1 min. The filtered supernatant was collected and stored at -80°C for further PA measurement. The level of PA was quantified using the Anthrax Protective Antigen 83 ELISA Kit according to the manufacturer’s recommendations (Alpha Diagnostic International, San Antonio, TX, USA). The absorbance was read at 450 nm and 630 nm using a microplate reader (Agilent Biotek Epoch 2, Agilent Technologies, Inc., Santa Clara, CA, USA).
Capsule staining
Strains were streaked and grown on nutrient broth-yeast extract (NBY) agar plates supplemented with 0.7% (w/v) sodium bicarbonate (NaHCO3) (Fisher Scientific, Faire Lawn, NJ, USA). The plates were incubated in 20% CO2 at 37°C for 24 h. To visualize the bacterial capsule, India ink staining was performed by adding a drop of India ink (Becton, Dickson and Company, Sparks, MD, USA) and mixing with a loop of culture on a clean glass slide. The slide was covered with a cover slip and the edges sealed with nail polish. The bacterial capsule was examined under the microscope with oil immersion (100x) objectives (Evos XL Core Imaging System, Life Technologies Corporation, Bothell, WA, USA).
Indirect ELISA for detecting attached and secreted PDGA capsules
The starter cultures were grown to mid-log phase in BHI broth at 37°C while shaken at 220 rpm. OD600 was adjusted to 1 and subcultured at 1:100 to inoculate 3 ml of DMEM with 10% (v/v) normal human serum (NHS) (MP Biomedical, LLC, Solon, OH, USA) in triplicate and supplemented with 20% CO2 to stimulate the formation of capsules. DMEM supplemented with serum and CO2 can mimic the nutritional milieu found within host’s tissues, providing insights into bacterial virulence under conditions like in vivo settings. The cultures were shaken at 220 rpm and incubated at 37°C for 24 or 48 h. At each time point, 1 ml of each culture was harvested and centrifuge at 5,000 x g for 10 min. The supernatants containing secreted capsule were collected. To harvest the attached capsules, the cell pellets were resuspended in sterile MilliQ water and heated at 100°C for 20 min to lyse cells and release the cell membrane attached capsule. The cell suspensions were centrifuged at 5000 x g for 10 min followed by collecting the supernatant harboring the released attached capsules. All samples were stored at -80°C. To assess the level of attached and secreted PDGA capsule from Bcbva UFBc0009 and B. anthracis Ames UF00738, supernatants at 24- and 48-h post inoculation were analyzed using ELISA. Briefly, Immulon 4HBX flat-bottomed 96-well microtiter plates (Immulon, Thermo Scientific, USA) were coated in duplicate with 100 μl per well of either a 1:50 dilution of supernatant containing secreted capsules or 1:200 dilution of supernatant carrying attached capsules in sterile 1x PBS. A known concentration of purified capsule [55] was two-fold serially diluted and used to coat wells of a 96-well microtiter plate with 100 μl per well in duplicate to create a standard curve. All plates were incubated overnight at 4°C and washed with 300 μl of PBST before proceeding. The antigen-coated plates were blocked with 300 μl of blocking solution (5% skim milk in PBST) for 1 h at room temperature. After blocking, 100 μl of 1:6000 diluted mouse anti-PDGA capsule IgM [56,57] in blocking solution were added to the wells and incubated for 1 h at room temperature. After incubation and washing, goat anti-mouse IgM conjugated with horse radish peroxidase (HRP) (Invitrogen, Camarillo, CA, USA) was diluted to 1:3000 in blocking solution, and 100 μl of diluted secondary antibody solution was added to each well and incubated for 1 h at room temperature. Finally, the antigen and antibody complexes were detected by adding 50 μl of 3,3’,5,5’ tetramethylbenzidine (TMB) (1-Step TMB ELISA Substrate Solutions, Thermo Scientific, IL, USA) and incubating for 30 min at room temperature. The reaction was stopped by adding 50 μl of 1 N hydrochloric acid (EMD Millipore Corporation, Burlington, MA, USA) and the absorbance was measured at 450 nm using microplate reader (Agilent Biotek Epoch 2, Agilent Technologies, Inc., Santa Clara, CA, USA). The standard curve from a known concentration of purified capsules was generated, and the concentrations of secreted and attached capsules in each sample were calculated.
Galleria mellonella infection
Galleria mellonella larvae are commonly used as surrogate hosts for infectious disease models including Burkholderia spp., Listeria spp., Pseudomonas spp., Staphylococcus spp., Mycobacterium spp., B. anthracis, and pathogenic B. cereus strains [47,58–64]. Wax moth larvae have several benefits as they are easy to use, and the presence of an innate immune system mimics that of vertebrates [59,65,66]. To investigate the virulence of Bcbva UFBc0009 and B. anthracis Ames UF00738 in G. mellonella, larvae worms weighed between 100–250 mg were used in this study. Worms were purchased from Carolina Biological Supply Company (Burlington, North Carolina, USA) and used within 5 days of receipt. Bcbva UFBc0009 and B. anthracis Ames UF00738 spores were generated and purified as described above. Purified spores were diluted, plated, and enumerated before diluting to the desired inoculum of 107 spores/ml in PBS. Inoculum were serially diluted to 103 spores/ml and kept on ice until use. A total number of 120 worms were randomly assigned into six groups with 20 worms per group. A Hamilton 50 μl microsyringe with a 27-gauge needle was used for injections. Worms were briefly placed on ice to induce torpidity and reduce movement before injection. Each larva was injected with 10 μl of inoculum into the third distal left proleg as desribed previously [47]. Larvae were monitored at room temperature for 30 min before placing at 37°C. The survival of worms was checked at 24-, 48-, and 72 h post-infection by observing melanin pigment formation and no sign of movement when touched.
Animal studies
Guinea pigs were chosen to determine Bcbva virulence because these animals have been widely used for investigation of anthrax pathogenesis and therapeutic efficacy [67–70]. Inhalation anthrax in guinea pigs shows similar infection mechanisms to those seen in non-human primates and humans [67,71]. Although death in guinea pigs often occurs within 2–4 days post infection, protection can be achieved through optimal vaccination [67,72]. Six-to eight-week-old male and female Dunkin-Hartley guinea pigs (Elm Hill Labs, Chelmsford, MA) were provided food and water ad libitum. In the LD50 studies, an equal number of eight-week-old female and male guinea pigs (n = 10 per group) were weighed then anesthetized with intraperitoneal ketamine-xylazine injections and challenged intranasally with spores in 0.5 ml of PBS. For the vaccine challenge experiments, six-week-old guinea pigs (n = 8 per group) were vaccinated subcutaneously with 1x106 spores of the Sterne 34F2 veterinary vaccine (Colorado Serum Company, Denver, CO, USA). Four weeks later the animals were anesthetized, and their blood was collected for further analysis. The anti-PA titers from vaccinated and unvaccinated animals were quantified by using a guinea pig anti-anthrax protective antigen 83 (PA83) ELISA kit according to the manufacturer’s recommendations (Alpha Diagnostic International, San Antonio, TX, USA). Serum from unvaccinated animals served as a control. Animals were then intranasally challenged with 100x LD50 of B. anthracis Ames spores (LD50 = 3.61x105 spores determined for this study) or 100x LD50 of Bcbva spores (LD50 = 3.49x104 spores) and followed for survival for 14 days. Early intervention endpoints were determined for all studies, and guinea pigs were humanely euthanized when moribund or at the end of study. Lungs and spleens were collected and processed to determine bacterial loads in organs.
Anthrax lethal toxin neutralization assay
RAW264.7 cells were seeded as mentioned above and cultured in DMEM supplemented with 10% FBS and 10 mM HEPES (Fisher Scientific, Fai Lawn, NJ, USA) overnight. The toxin neutralization assay was performed as previously described [73]. Pooled serum samples from vaccinated guinea pigs before challenging with Bcbva UFBc0009 and B. anthracis Ames UF00738 spores were prepared at 200-, 1,200-, and 12,000-fold dilutions in DMEM supplemented with 10% FBS and 10 mM HEPES. Pooled serum samples from unvaccinated animals served as a control. The diluted serum samples were then incubated with 50 ng/ml PA (List Labs, Campbell, CA, USA) and 160 ng/ml LF (List Labs, Campbell, CA, USA) for 30 min before adding to RAW264.7 cells. The mixtures of cells, serum, and toxin were incubated for 6 h. After incubation, the supernatant was collected and further used for detection of cytolytic activity by using CytoTox 96 Non-Radioactive Cytotoxicity Assay following the manufacturer ‘s recommendations (Promega Corporation, WI, USA).
Statistical analysis
The sporulation efficiency of Bcbva UFBc0009 and B. anthracis Ames UF00738 spores are presented as means with standard error of the mean (SEM) and are displayed in bar graphs. The Shapiro-Wilk test was used to analyze the sample distribution. The unpaired t-test was performed to determine significance difference of sporulation, cytotoxicity, macrophage internalization, and attached or secreted PDGA capsules in DMEM supplemented with 10% (v/v) NHS in 20% CO2 between Bcbva UFBc0009 and B. anthracis Ames UF00738. The significant difference of PA production between Bcbva UFBc0009 and B. anthracis Ames UF00738 in HIB and BHI media was determined by using the Mann-Whitney U test. A Kruskal-Wallis test with Dunn’s multiple comparison test was used to identify significant differences of anti-PA titers among vaccinated and unvaccinated groups. The survival of G. mellonella larvae and guinea pigs was plotted as a survival curve. The LD50 values of G. mellonella larvae infected with either Bcbva UFBc0009 and B. anthracis Ames UF00738 was determined and compared using nonlinear regression analysis. The LD50 values of B. anthracis Ames UF00738 infected guinea pigs were calculated using the sigmoidal non-linear regression half max calculations in GraphPad Prism fitting the curve to the log transformed group dosing plotted against survival probability at that dose. The log-rank test was used to identify the significance between vaccinated and unvaccinated guinea pig groups. Graph generation and statistical analyses were done in GraphPad PRISM 10 software (GraphPad Software, San Diego, California, USA) with significant level at ρ < 0.05.
Results
Sporulation, cytotoxicity, and macrophage internalization of Bcbva and B. anthracis spores
Spores of Bcbva and B. anthracis are considered the infectious particles of anthrax. The sporulation efficiency of Bcbva UFBc0009 and B. anthracis Ames UF00738 spores was calculated as the percentage of spores relative to the total number of bacteria on day 1, day 2, and day 6 (Fig 1A). Bcbva UFBc0009 sporulated significantly more than B. anthracis Ames UF00738 from day 1 to day 6. The relative percentage of both Bcbva UFBc0009 and B. anthracis Ames UF00738 spores gradually increased from day 1 to day 6. Comparing these two strains, Bcbva UFBc0009 sporulated more rapidly than B. anthracis Ames UF00738. The LDH mediated cytotoxicity assay revealed the cytolytic activities of RAW264.7 macrophages treated with Bcbva UFBc0009 and B. anthracis Ames UF00738 were 64 and 66%, respectively. The study indicated there was no difference in the level of cytotoxicity between groups (Fig 1B). Following phagocytosis, spores germinated and replicated inside of macrophages, leading to cell death due to the robust secretion of anthrax toxins. This suggests the two strains share similar levels of virulence relative to the toxins linked to cell death released by vegetative cells. Next, RAW264.7 macrophages were infected with Bcbva UFBc0009 and B. anthracis Ames UF00738 spores at an MOI of 20 to identify differences in internalization efficiencies. At 2 hours post-infection, both Bcbva UFBc0009 and B. anthracis Ames UF00738 were internalized by RAW264.7 macrophages, however, the internalization efficiency of Bcbva UFBc0009 spores was significantly lower than B. anthracis Ames UF00738 spores (1.75% for Bcbva UFBc0009 vs 34.80% for B. anthracis Ames UF00738) (Fig 1C).
(A) Number of spores from Bcbva UFBc0009 and B. anthracis Ames UF00738 produced in HIB broth at 1-, 2-, and 6 day post incubation. At each time point, the number of total spores from Bcbva UFBc0009 (filled circles) and B. anthracis Ames UF00738 (hollow circle) strains were enumerated, and the sporulation efficiency was calculated as the percentage of spores relative to the total number of bacteria. The data are presented as the mean ± SEM (standard error of the mean). The graphs were the average of 2 independent experiments carried out in triplicate. (B) Cytotoxicity measured in RAW264.7 macrophages infected with Bcbva UFBc0009 and B. anthracis Ames UF00738 spores at 16-hour post infection. No treatment samples were RAW264.7 cells incubated in media only and served as a vehicle control, while maximum lysis samples were RAW264.7 cells treated with 10X lysis buffer and served as a maximum LDH release control. Six replicates for each strain were tested. The data were presented as the mean ± SEM. (C) The internalization efficiency of RAW264.7 macrophages infected with Bcbva UFBc0009 and B. anthracis Ames UF00738 spores. Three replicates per strain were examined. The recovered bacteria were identified and calculated as the relative percentage of the initial inoculum. The data were presented as the mean ± SEM. The significance between Bcbva UFBc0009 and B. anthracis Ames UF00738 strains were determined by unpaired t-test with * = ρ<0.05, ** = ρ<0.01, **** = ρ<0.0001.
Secretion of protective antigen (PA) by Bcbva compared to B. anthracis Ames
Secretion of PA by Bcbva UFBc0009 and B. anthracis Ames UF00738 in stationary phase HIB and BHI cultured supernatants was measured by ELISA. The amount of PA83, the 83-kDa form of PA, produced by Bcbva UFBc0009 and B. anthracis Ames UF00738 in HIB broth after 24 h growth was 37.26 and 60.58 ng/ml, respectively (Fig 2A). The average amount of PA secreted by Bcbva UFBc0009 and B. anthracis Ames UF00738 in BHI broth after 24 h growth was 42.48 and 196.78 ng/ml (Fig 2B). During the stationary phase, the bacterial population size is stable, and the growth rate slows down or even stops owing to nutrient limitation and buildup of waste products, decreasing metabolic activities and synthesis of cellular components. For statistical analysis, we assumed that these data were not normally distributed due to a small sample size (n = 2). The nonparametric test was used to analyze the differences of PA productions between two groups. The results showed that B. anthracis Ames UF00738 tended to secrete more PA83 than Bcbva UFBc0009 in both culture media, but not at a significant difference (ρ<0.333).
The average amount of PA secreted by either Bcbva UFBc0009 or B. anthracis Ames UF00738 in HIB (A) and BHI (B) broths at 24 h post-incubation. Two replicates per strain were tested. The data were presented as the mean ± SEM.
Quantification of attached and secreted PDGA capsule produced by Bcbva
Bcbva UFBc0009 and B. anthracis Ames UF00738 were grown at 37°C on bicarbonate agar in a CO2-enriched environment for 24 and 48 h to induce capsule formation. Twenty four hours post incubation, the colonies of Bcbva UFBc0009 and B. anthracis Ames UF00738 appeared mucoid on the agar plates. Capsule produced from vegetative cells of Bcbva UFBc0009 and B. anthracis Ames UF00738 were visualized by staining with India ink under light microscopy. Both Bcbva UFBc0009 (Fig 3A and 3C) and B. anthracis Ames UF00738 (Fig 3B and 3D) were encapsulated and showed highly refractile clear zones surrounding bacterial cells against a dark background due to exclusion of India ink by the bacterial capsule. Bcbva UFBc0009 had short-chain rod shaped vegetative cells, while B. anthracis Ames UF00738 showed long rod-shaped with squared ends arranging in a chain of cells. Qualitatively, Bcbva UFBc0009 produced thicker capsule on the surface of vegetative cells than B. anthracis Ames UF00738. The amount of attached and secreted PDGA capsule produced by Bcbva UFBc0009 and B. anthracis Ames UF00738 was quantified by ELISA using anti-capsule IgM. After 24 hours, Bcbva UFBc0009 produced 8,436 μg/ml of attached capsule, while B. anthracis Ames UF00738 produced a higher amount at 11,460 μg/ml (Fig 4A). At 48-h post incubation, the level of attached capsule from Bcbva UFBc0009 increased to 11,845 μg/ml, whereas B. anthracis Ames UF00738 decreased to 5,105 μg/ml (Fig 4A). The average amount of attached PDGA capsule produced by B. anthracis Ames UF00738 was significantly higher than Bcbva UFBc0009 at the 24 h timepoint (ρ<0.01). At 48 h, the trend was reversed: Bcbva UFBc0009 had significantly higher amounts of attached PDGA capsule than B. anthracis Ames UF00738 (ρ<0.0001).
Bacteria were grown on NBY plates supplemented with 0.7% (v/v) sodium bicarbonate and 20% CO2 at 37°C overnight. Bcbva UFBc0009 (A) and B. anthracis Ames (B) vegetative cells were stained with India ink and examined under 100x oil imersion magnification. Insets magnify Bcbva UFBc0009 (C) and B. anthracis Ames (D) vegetative cells surrounded by capsules.
The average amount of attached (A) and secreted (B) PDGA capsule produced either by Bcbva UFBc0009 or B. anthracis Ames in DMEM+10% (v/v) NHS and 20% CO2 at 24 and 48 h post incubation. The cultures were grown in triplicate, and two replicates per strain were tested with ELISA. Two-independent experiments were performed, and all data were presented as mean ± SEM. The significance between strains were determined by unpaired t-test with * = ρ<0.05, ** = ρ<0.01, **** = ρ<0.0001.
The average amount of secreted PDGA capsule produced at 24 h by Bcbva UFBc0009 (405.9 μg/ml) was significantly higher than B. anthracis Ames UF00738 (351.9 μg/ml) (Fig 4B; ρ<0.05). At the 48 h time point, the levels of secreted capsule from both Bcbva UFBc0009 and B. anthracis Ames UF00738 were higher than the 24 h time point, with B. anthracis Ames UF00738 secreting more capsule (912.7 μg/ml) than Bcbva UFBc0009 (645.4 μg/ml) (Fig 4B). B. anthracis Ames UF00738 secreted significantly more capsule than Bcbva UFBc0009 at 48 h post incubation (ρ<0.01). Additionally, the average level of secreted capsule from both Bcbva UFBc0009 and B. anthracis Ames UF00738 was lower than attached capsule.
Bcbva has a lower lethal dose than B. anthracis in the Galleria mellonella larvae infection model
Our studies revealed G. mellonella larvae did not survive after 24 h when they were infected with 1.17x104 spores of Bcbva UFBc0009, whereas those infected with B. anthracis Ames UF00738 at 1.18x104 spores survived to 72 hours post infection (Fig 5). Bcbva UFBc0009 and B. anthracis Ames UF00738 killed larvae in a dose-dependent manner as shown in the LD50 curves (Fig 5). The 3-day LD50 after hemocoel injection was 9.21x102 spores for G. mellonella larvae infected with Bcbva UFBc0009 spores (Fig 5A) and 2.24x104 spores for B. anthracis Ames UF00738 (Fig 5B). The LD50 of Bcbva UFBc0009 was 24-fold less than that of B. anthracis Ames UF00738 in wax moth worms. The nonlinear fit analysis was used to determine the differences between LD50 curves, and the result showed that the two LD50 values are different as we rejected the null hypothesis that the LD50 values were the same for all data set with a ρ value of 0.0015. These data suggest that Bcbva UFBc0009 is significantly more virulent than B. anthracis Ames UF00738 in the G. mellonella larvae (ρ<0.01). Notably, the worms were found to experience some trauma induced by injection: the survival rate from PBS-injected control group was 80% compared to non-injected control group. The survival rate of those groups either infected with 1.17x101 Bcbva UFBc0009 or 1.18x101 B. anthracis Ames UF00738 spores were found to be like the PBS-injected control group, indicating that the cause of death at these low doses was most likely due to injection injury.
The survival of wax moth worms was monitored at 24-, 48-, and 72 hour post infection and scored by lack of movements when prodding. Spore dosage and survival time of larvae infected with (A) Bcbva UFBc0009 and (B) B. anthracis Ames UF00738 spores are presented as Kaplan-Meier survival curves to visualize the probability of survival versus time.
The LD50 of Bcbva in intranasally challenged guinea pigs and efficiency of the Sterne live spore vaccine
Median lethal dose of Bcbva UFBc0009 was evaluated in guinea pigs via intranasal route of infection, and the results were compared to B. anthracis Ames at the Rocky Mountain Regional Biosafety Laboratory at Colorado State University. The LD50 of B. anthracis determined in this study was consistent with a previous report in guinea pigs following inhalation exposures [74]. Guinea pigs became moribund on day 3, and there was no survival when animals were infected at the highest dose of Ames or Bcbva (Fig 6A and 6B, respectively). The intranasal LD50 of Bcbva UFBc0009 in guinea pigs was 3.49x104 spores which was 10.3-fold lower than found for B. anthracis Ames in this study (LD50 = 3.61x105 spores, as determined by Probit survival calculations, Fig 6A) and 3.4-fold lower than previously reported for B. anthracis Ames (LD50 = 1.2x105 spores) [74]. Next, the efficacy of the Sterne vaccine against Bcbva UFBc0009 was evaluated in the guinea pig model. At 4 weeks post-vaccination, sera from all vaccinated guinea pigs tested positive for PA-specific IgG as measured by ELISA. The average anti-PA titers from vaccinated animals before being challenged with either Bcbva UFBc0009 or B. anthracis Ames UF00738 spores were 102,400 and 83,250 ng/ml, respectively (Fig 7A). The antibody levels to PA in the vaccinated animals were significantly higher when compared to unvaccinated controls but were not significantly different between the two challenge groups. Toxin neutralization activity from vaccinated guinea pigs pre-challenged with Bcbva UFBc0009 ranged from 53.3%-61.7% with a mean of 56.4% (cytotoxicity measurement of 39.3% to 47.7% with a mean of 43.6%) while pre-challenged B. anthracis Ames vaccinated group ranged from 35.0%-42.4% with a mean of 38.2% (cytotoxicity measurement of 58.6% to 65.0% with a mean of 61.8%) at a sera dilution of 1:1,200 (Fig 7B). Pooled serum from unvaccinated guinea pigs diluted 1:1,200 exhibited mean toxin neutralization activity at 13.6% that ranged from 1% to 24.1% (cytotoxicity measurement of 65.9% to 99.0% and mean of 86.4%). The 50% cellular protection (ED50) of vaccinated guinea pigs pre-challenged with Bcbva UFBc0009 and B. anthracis Ames were at 1,756- and 1,136- fold dilutions, respectively. The data indicated that the ability of antibody to neutralize the in vitro cytotoxicity of PA and LF in vaccinated animals was higher than unvaccinated controls. At 14 days post challenge, the results showed 100% and 85% of Sterne vaccinated guinea pigs survived after intranasal spore challenge with 100x LD50 of either Bcbva UFBc0009 or B. anthracis Ames UF00738, respectively (Fig 8A). The survival of unvaccinated animals challenged with B. anthracis Ames UF00738 was significantly different from animals vaccinated then challenged with Bcbva UFBc0009 (ρ<0.0001) and B. anthracis Ames UF00738 (ρ<0.001). Only one vaccinated guinea pig was moribund 5 days post-infection with B. anthracis Ames UF00738. All unvaccinated control guinea pigs were moribund by day 5 post challenge. Sterne vaccinated guinea pigs were completely protected from intranasal Bcbva spore challenge. Spleens of unvaccinated animals contained viable bacteria and ungerminated spores, while infected animals from the vaccinated group demonstrated some levels of organ clearance (Fig 8B).
Kaplan-Meier Survival curves of guinea pigs challenged with the indicated doses of purified Bcbva spores. The number of purified spores were back titrated from the inoculum and used to determine the Ames (A) and Bcbva (B) LD50 in guinea pigs. The intranasal 14-day LD50 was calculated as 3.61 x105 spores for Ames and 3.49x104 spores for Bcbva UFBc0009.
At 4 week post-vaccination, anti-PA titers (A) and toxin neutralization antibody (B) from vaccinated guinea pigs before being challenged with Bcbva UFBc0009 and B. anthracis Ames UF00738 spores were quantified. The anti-PA levels were presented as the mean ± SEM. The significance between groups were determined by a Kruskal-Wallis test with Dunn’s multiple comparison test with ** = ρ<0.01, **** = ρ<0.0001. Toxin neutralizing activities were presented as the mean ± SEM and graphed on a log scale.
(A) Survival of vaccinated and unvaccinated guinea pigs following intranasal challenge with 100x LD50 spores of indicated strains. Sterne vaccinated guinea pigs infected with Bcbva UFBc0009 spores (green triangle and line) and B. anthracis Ames UF00738 spores (red square and line) compared to unvaccinated guinea pigs infected with B. anthracis Ames UF00738 spores (blue dot and line). The log-rank (Mantel–Cox) test was used to determine the significance between vaccinated and unvaccinated groups. (B) The spleens and lungs were collected at 14 days post challenge or at moribundity. Each dot represents data from a single guinea pig with mean ± SD. The limit of detection was shown in the graph (red line).
Discussion
Bacillus cereus biovar anthracis is a genetic near neighbor of B. anthracis and has been a cause of anthrax-like disease in livestock and wildlife across sub-Saharan Africa for decades [24]. Bcbva evolved from B. cereus by acquiring virulence plasmids almost identical to those found in B. anthracis [30]. The ability of Bcbva to cause disease is attributed to the production of secreted anthrax toxins and the PDGA capsule [75,76]. Here, we demonstrated Bcbva UFBc0009 sporulated more rapidly and to a higher magnitude compared to B. anthracis Ames UF00738. This finding is consistent with a recent study that compared Bcbva isolated from chimpanzees in Cameroon and Côte d’Ivoire to B. anthracis strains [39]. A previous study showed wild, low passage B. anthracis strains sporulate faster than laboratory B. anthracis strains which have been repeatedly sub-cultured due to the limitation of nutrients [77]. One reason for this is laboratory strains consume nutrients slower than wild strains, leading to a delay in sporulation time [77]. Therefore, it is possible the wild, low passage Bcbva used in this study can assimilate available nutrients faster than the laboratory B. anthracis Ames UF00738 strain resulting in rapid sporulation.
Furthermore, our data provide additional understanding of sporulation in Bcbva compared to B. anthracis. Bacillus anthracis spores persist in the locally infectious zone (LIZ) for extended periods of time; recent work suggests approximately 10 years where LIZs are undisturbed and decay naturally [78]. The rapidity of spore formation and quantity of spores may serve to efficiently transmit Bcbva in its natural conditions. Studies also demonstrate Bcbva is distributed and transmitted by necrophagous flies [24,79]. This parallels a report of B. anthracis spores being mechanically spread from anthrax carcasses by flies [80], such as in Texas and elsewhere where this occurs during outbreaks. Spore contaminated carcasses are highly valuable sources of nutrients for necrophagous fly replication. Following the case multiplier hypothesis, necrophagous flies can spread viable bacteria from a carcass through the surrounding environment forming a wide area LIZ beyond the immediate carcass site, especially in the first days after host death [80,81]. A previous study estimated a single blow fly can distribute B. anthracis spores from a carcass via fly deposits of up to 8.62 x 105 spores per day [9]. This process leads to increased exposure risk for other animals and contributes to wildlife mortality during an outbreak. Currently, the source of wildlife infection with Bcbva in Africa remains unclear. In Taï National Park, 5% of randomly caught carrion flies were PCR positive for plasmid (pag and capB) and chromosomal (genomic island IV) Bcbva markers, and many of these flies carried viable Bcbva spores [24]. A recent study showed flies associated with Bcbva-infected mangabey monkeys (Cercocebus atys atys) in Taї National Park contained Bcbva DNA, and Bcbva were recovered from the PCR-positive flies [79]. It is conceivable that flies may transport Bcbva as spores or vegetative cells from externally contaminated mouthparts and legs, or from ingesting spores and/or vegetative cells. Further studies are needed to examine the relationship between Bcbva and flies in Taї National Park.
The host-pathogen interaction between macrophages and B. anthracis is initiated by recognition followed by elimination of the pathogen [82]. During the initial stage of infection, macrophages play an essential role in the pathogenesis of B. anthracis infection as they are responsible for phagocytosis, while simultaneously functioning as a vehicle to transport B. anthracis spores to the draining lymph nodes [83]. Macrophages cannot directly kill the B. anthracis spores but rather kill the vegetative bacilli once spores germinate [84]. Upon pulmonary instillation, alveolar macrophages engulf and transport the spores to tracheobronchial lymph nodes where spores germinate and form vegetative cells within the phagolysosome of phagocytic cells [85]. Some vegetative cells are killed, while others remain viable and escape from phagolysosome into the cytoplasm [85]. The bacilli multiply rapidly and secrete toxins resulting in death of macrophages. When the bacillary replication exceeds the capacity of regional lymph nodes, the bacteria enter the bloodstream where they grow extracellularly to levels leading to systemic infection [86]. In advanced anthrax infection, the number of vegetative cells is as high as 108 bacteria/ml of blood [5] suggesting the innate immune responses is overwhelmed. Our data demonstrated that the cytolytic activities of RAW264.7 macrophages infected with Bcbva UFBc0009 and B. anthracis Ames UF00738 spores at 16 hours post infection were comparable. This pattern coincided with the secreted PA levels in both HIB and BHI broths at 24 h post inoculation. The amount of PA generated by Bcbva UFBc0009 and B. anthracis Ames UF00738 did not differ substantially between medium types. Cytotoxicity levels indicate toxin release, as B. anthracis vegetative cells produce toxins and proteases during replication, resulting in tissue lysis further facilitating the dissemination of bacteria to deeper tissue layers [48,58]. The specific mechanisms by which Bcbva and B. anthracis spores survive, and multiply within macrophages are still unclear. The kinetics of bacterial multiplication inside macrophages have yet to be determined. To compare the efficiency of spore uptake by macrophages, we demonstrated Bcbva spores could be internalized and survive within mouse macrophages at the initial stage of infection like B. anthracis. Nevertheless, B. anthracis Ames spores had significantly higher internalization efficiency than Bcbva spores. The low level of recovered bacteria in macrophages infected with Bcbva UFBc0009 is possibly due to differences in components and functions of the exosporium, the hairy-like outermost layer of the spores. In addition, the exosporium of B. anthracis appears to play an essential role in protecting the spores from macrophage-mediated killing [84]. The exosporium regulates intracellular survival and influences spore germination in B. anthracis infected macrophages [82]. Upon phagocytosis and spore germination, the kinetics of capsule production inside phagocytic cells is still not clear. The differences in the encapsulation of Bcbva and B. anthracis could affect the intracellular survival inside macrophages [87]. The additional production of hyaluronic acid to PDGA capsules in Bcbva may give the bacteria a survival advantage by inhibiting phagocytosis and enabling evasion from the innate immune system. Like B. anthracis, the initial interaction of Bcbva spores with the infected macrophages may play role in survival of Bcbva spores from the macrophage killing mechanisms. However, the ability of the exosporium and capsules to promote intracellular survival of Bcbva spores within infected macrophages has yet to be elucidated fully. Additionally, it is important to note the limitations of cytotoxicity and internalization assays in this study. The experiments were designed based on previously published literature [48–51,53]. Further microscopy and molecular studies to directly assess spore germination and replication kinetics inside macrophages are necessary to move past the limits of cytotoxicity and internalization assays seen here. The current study demonstrated that the production of PA by Bcbva UFBc0009 at the stationary phase appeared to be lower than B. anthracis Ames, but not significant differences. This finding is consistent with the study comparing B. anthracis and Bcbva isolates grown in bicarbonate/CO2 conditions, in which the levels of PA secreted by Bcbva were somewhat lower than that of B. anthracis [31]. PA is an important virulence factor of B. anthracis and acts as the major protective immunogen during vaccination [88]. One feature of anthrax vaccination is the production of PA can induce protective immunity against both lethal and edema toxins. In fact, PA is the major immunogenic constituent of the current available human vaccines for anthrax [14,89]. A previous study showed mice vaccinated with formaldehyde-inactivated B. anthracis spores and recombinant PA vaccine were protected against Bcbva spores via subcutaneous inoculation [31]. As a PA-producing bacterium, it is feasible that PA secreted by Bcbva can be used as a target for new therapeutic approaches against anthrax in livestock and wildlife in Africa.
When grown on NBY medium supplemented with bicarbonate in 20% CO2 at 37°C, Bcbva UFBc0009 vegetative cells synthesized capsule on the cell surface like B. anthracis Ames UF00738 vegetative cells. Bcbva UFBc0009 expressed slightly higher amounts of capsule than B. anthracis Ames UF00738 when observed under the light microscope. These findings are consistent with a previous study that found Bcbva expresses both hyaluronic acid and PDGA capsules [31]. By measuring the level of PDGA capsule production, we showed Bcbva UFBc0009 produced significantly more attached PDGA capsule than B. anthracis Ames UF00738 once bacterial cultures were grown in DMEM supplemented with 10% (v/v) NHS and 20% CO2. PDGA capsule is a poor immunogen, is resistant to degradation, and has comparable properties to T cell-independent polysaccharides [86] while hyaluronic acid is a poor immunogen due to antigen mimicry of the mammalian extracellular matrix. During infection, PDGA capsule produced by bacteria inhibits complement deposition and opsonization by macrophages [90,91]. Purified capsule can inhibit the bactericidal activity of alpha defensin in neutrophils in vitro [92]. Taken together, it is possible Bcbva can survive better than B. anthracis in phagocytic cells. However, capsule dependent virulence is unclear, as a study showed that a capsule negative Bcbva cured of pBCXO2, while retaining pBCXO1 and still capable of producing hyaluronic acid capsule, remained highly virulent in mice challenged intranasally when compared to an acapsular, virulent B. anthracis Sterne (pXO2 negative) strain [31]. Previous literature suggested hyaluronic acid capsule encoded by the pBCXO1 plasmid may contribute to a more successful infection by providing higher chance for survival of bacterium in mammalian hosts thus favoring the dispersal of pathogens [31,93]. For B. cereus G9241, a pathogenic strain causing anthrax-like disease, the presence of hyaluronic acid capsule may facilitate in vivo infection [93].
Challenge and LD50 experiments were conducted in animal models to characterize the virulence of the earliest known Bcbva isolates. A survival assay was performed to compare the virulence between Bcbva UFBc0009 and B. anthracis Ames UF00738 in G. mellonella larvae and guinea pigs. We demonstrated that G. mellonella results correlate to virulence levels of Bcbva and B. anthracis Ames in small mammal models. We discovered Bcbva UFBc0009 from Taї National Park is significantly more virulent than B. anthracis Ames UF00738. The 3-day LD50 of Bcbva UFBc0009 was 24-fold lower than B. anthracis Ames UF00738 in the G. mellonella model. Akin to G. mellonella infection, intranasal LD50 of Bcbva UFBc0009 in guinea pigs was 3.4-fold lower than reported for B. anthracis Ames [74]. Our findings in guinea pigs suggest Bcbva from Côte d’Ivoire has a lower intranasal LD50 than a neighboring strain in Cameroon [31], however, both had lower LD50 than the B. anthracis strains used for comparison.
Currently, the Sterne vaccine is the most common vaccine used in wildlife and livestock to prevent anthrax [94–98]. Its efficacy in guinea pigs was examined in this study, as they were vaccinated with the Sterne vaccine and later intranasally challenged with spores from Bcbva UFBc0009. The antibody profiles in these vaccinated animals revealed all pre-challenge vaccinated guinea pigs had significantly higher anti-PA levels compared to non-vaccinated animals. The protection was then tested in vitro by the sera’s ability to neutralize the cytotoxic effect of anthrax lethal toxin against RAW264.7 cells. The results showed neutralizing antibody titers correlated with anti-PA titers by ELISA. The increase in anti-PA titers and the lethal toxin neutralization antibodies in vaccinated guinea pigs conferred notable protective immunity to anthrax. A previous PA vaccine study showed protection against Bcbva demonstrated all mice immunized with recombinant PA and formaldehyde-inactivated B. anthracis spores of a genetically detoxified Sterne strain (RPLC 2) survived subcutaneous infection with Bcbva or B. anthracis spores [31]. In the current study, the efficacy of the Sterne vaccine in protecting against Bcbva was complete and comparable to the challenge with B. anthracis Ames UF00738 in the guinea pig model. The vaccine protected against Bcbva UFBc0009 and B. anthracis Ames UF00738 spores, resulting in 8/8 and 7/8 animals surviving, respectively. The Bcbva UFBc0009 group showed a significant increase in survival time of 14 days versus 5 days for the unvaccinated control. However, it should be noted Sterne vaccine did not achieve sterilizing immunity as residual bacteria and ungerminated spores remained in the lungs and spleens at the end of the 14-day study. Several animal models including guinea pigs have shown that a subpopulation of spores may remain ungerminated within the site of infection [72,99–101]. The high level of recovered bacteria in animals challenged with Bcbva UFBc0009 was possibly due to the expression of hyaluronic acid and PDGA capsules which may provide a survival advantage to the bacteria by impeding complement fixation, allowing evasion of host immunity compared to B. anthracis Ames UF00738. The impacts of host immune responses to spore challenge remain to be elucidated. Vaccination is the standard control of anthrax and can break the cycle of transmission [102]. Several studies have demonstrated that vaccine implementation in exposed susceptible animals and proper discarding of infected carcasses impedes the spread of infection and death, especially in endangered species [95,98]. The only licensed animal vaccine available in the US is the Sterne 34F2 live spore vaccine which can be used in livestock including cattle, sheep, goats, pigs, and horses or off-label use in other species [16]; internationally other Sterne-like strains are often used in vaccine production [103]. Further efficacy studies and risk assessments for the Sterne vaccine are required for those additional species, in particular non-human primates of conservation concern.
When comparing the wild Bcbva UFBc0009 isolate and B. anthracis Ames UF00738 strain, we found differences in sporulation efficiencies and PDGA capsule production, specifically Bcbva UFBc0009 sporulated and produced more PDGA capsule than B. anthracis Ames UF00738. B. anthracis Ames UF00738 spores were internalized by mouse macrophages at higher efficiency than Bcbva UFBc0009 spores. However, the cytolytic activity of infected RAW264.7 macrophages and secreted PA levels in minimal media were comparable between the two strains. Although our data demonstrated significant differences between the two strains, these differences could be strain-specific rather than species-specific. More research is required to understand the role of Bcbva strain diversity in pathogenesis.
Besides host-pathogen interaction, future studies should focus on Bcbva lifecycle outside of host cells. These include the kinetics of replication and sporulation of Bcbva on environmental substrates in the LIZ areas may be important as carrion flies can transport viable Bcbva spores and may amplify cases during an outbreak in Africa. The species most affected by Bcbva appear to be non-human primates living in humid tropical forest environments, whereas B. anthracis infection is commonly found in ungulates in savanna habitats. Further investigation into LIZ contamination and transmission pathways may provide new preventative methods to control the recurrence of enzootic outbreaks caused by Bcbva in wildlife and livestock. To prevent the future occurrence of anthrax outbreak caused by Bcbva in West and Central Africa, trials of Sterne vaccine in affected livestock, hoofed wildlife, and several non-human primates should be performed.
Supporting information
S1 Data. The excel workbook contains the raw data used to produce the images in each figure of the paper.
The tabs in the workbook are labeled Figs 1–2 and 4–8 and contain the data points for each of the indicated figures. Fig 3 is a microscopy image figure so there are no data points for this figure.
https://doi.org/10.1371/journal.pntd.0012779.s001
(XLSX)
References
- 1. Carlson CJ, Kracalik IT, Ross N, Alexander KA, Hugh-Jones ME, Fegan M, et al. The global distribution of Bacillus anthracis and associated anthrax risk to humans, livestock and wildlife. Nat Microbiol. 2019 May 13;4(8):1337–43.
- 2. Fasanella A, Galante D, Garofolo G, Jones MH. Anthrax undervalued zoonosis. Vet Microbiol. 2010;140(3–4):318–31. pmid:19747785
- 3. Hugh-Jones M. 1996–97 Global Anthrax Report. J Appl Microbiol. 1999 Aug;87(2):189–91. pmid:10475945
- 4. Hugh-Jones M, Blackburn J. The ecology of Bacillus anthracis. Mol Aspects Med. 2009;30(6):356–67.
- 5. Dupke S, Schubert G, Beudjé F, Barduhn A, Pauly M, Couacy-Hymann E, et al. Serological evidence for human exposure to Bacillus cereus biovar anthracis in the villages around Taï National Park, Côte d’Ivoire. Roeltgen K, editor. PLoS Negl Trop Dis. 2020 May 14;14(5):e0008292.
- 6. Bellan SE, Turnbull PC, Beyer W, Getz WM. Effects of experimental exclusion of scavengers from anthrax-infected herbivore carcasses on Bacillus anthracis sporulation, survival and distribution. Appl Environ Microbiol. 2013;
- 7. Getz WM. Biomass transformation webs provide a unified approach to consumer–resource modelling. Ecol Lett. 2011;14(2):113–24. pmid:21199247
- 8. Turner WC, Kausrud KL, Krishnappa YS, Cromsigt JPGM, Ganz HH, Mapaure I, et al. Fatal attraction: vegetation responses to nutrient inputs attract herbivores to infectious anthrax carcass sites. Proc R Soc B Biol Sci. 2014 Nov 22;281(1795):20141785. pmid:25274365
- 9. Jiranantasak T, Benn JS, Metrailer MC, Sawyer SJ, Burns MQ, Bluhm AP, et al. Characterization of Bacillus anthracis replication and persistence on environmental substrates associated with wildlife anthrax outbreaks. Walter WD, editor. PLOS ONE. 2022 Sep 21;17(9):e0274645.
- 10. Kracalik I, Malania L, Broladze M, Navdarashvili A, Imnadze P, Ryan SJ, et al. Changing livestock vaccination policy alters the epidemiology of human anthrax, Georgia, 2000–2013. Vaccine. 2017 Nov;35(46):6283–9. pmid:28988866
- 11. Metrailer MC, Hoang TTH, Jiranantasak T, Luong T, Hoa LM, Pham QT, et al. Spatial and phylogenetic patterns reveal hidden infection sources of Bacillus anthracis in an anthrax outbreak in Son La province, Vietnam. Infect Genet Evol. 2023;114:105496.
- 12. Turnbull PC. Anthrax vaccines: past, present and future. Vaccine. 1991;9(8):533–9. pmid:1771966
- 13. Sterne M. The use of anthrax vaccines prepared from avirulent (uncapsulated) variants of Bacillus anthracis. Onderstepoort J Vet: Sci & Animal Industry. 1939;13:307–12.
- 14. Ehling-Schulz M, Lereclus D, Koehler TM. The Bacillus cereus Group: Bacillus Species with Pathogenic Potential. Fischetti VA, Novick RP, Ferretti JJ, Portnoy DA, Braunstein M, Rood JI, editors. Microbiol Spectr. 2019 May 31;7(3):7.3.6.
- 15. Blackburn JK, Asher V, Stokke S, Hunter DL, Alexander KA. Dances with Anthrax: Wolves (Canis lupus) Kill Anthrax Bacteremic Plains Bison (Bison bison bison) in Southwestern Montana. J Wildl Dis. 2014;50(2):393–6.
- 16. Sidwa T, Salzer JS, Traxler R, Swaney E, Sims ML, Bradshaw P, et al. Control and Prevention of Anthrax, Texas, USA, 2019. Emerg Infect Dis. 2020 Dec;26(12):2815–24. pmid:33219643
- 17. Hugh-Jones M, De Vos V. Anthrax and wildlife. Rev Sci Tech Int Off Epizoot. 2002;21(2):359. pmid:11974621
- 18.
Turnbull PCB, Nations F and AO of the U, International Office of E, World Health O. Anthrax in humans and animals [Internet]. 4th ed. Geneva, Switzerland: World Health Organization; 2008. Available from: http://lib.myilibrary.com?id=225002
- 19. Kracalik I, Abdullayev R, Asadov K, Ismayilova R, Baghirova M, Ustun N, et al. Changing Patterns of Human Anthrax in Azerbaijan during the Post-Soviet and Preemptive Livestock Vaccination Eras. PLoS Negl Trop Dis. 2014;8(7):e2985. pmid:25032701
- 20. Klee SR, Özel M, Appel B, Boesch C, Ellerbrok H, Jacob D, et al. Characterization of Bacillus anthracis Like Bacteria Isolated from Wild Great Apes from Côte d’Ivoire and Cameroon. J Bacteriol. 2006 Aug;188(15):5333–44.
- 21. Leendertz FH, Ellerbrok H, Boesch C, Couacy-Hymann E, Mätz-Rensing K, Hakenbeck R, et al. Anthrax kills wild chimpanzees in a tropical rainforest. Nature. 2004 Jul;430(6998):451–2. pmid:15269768
- 22. Leendertz F, Yumlu S, Pauli G, Boesch C, Couacy-Hymann E, Vigilant L, et al. A new Bacillus anthracis found in wild chimpanzees and a gorilla from West and Central Africa. PLoS Pathog. 2006;2(1):e8.
- 23. Norris MH, Zincke D, Daegling DJ, Krigbaum J, McGraw WS, Kirpich A, et al. Genomic and Phylogenetic Analysis of Bacillus cereus Biovar anthracis Isolated from Archival Bone Samples Reveals Earlier Natural History of the Pathogen. Pathogens. 2023 Aug 20;12(8):1065.
- 24. Hoffmann C, Zimmermann F, Biek R, Kuehl H, Nowak K, Mundry R, et al. Persistent anthrax as a major driver of wildlife mortality in a tropical rainforest. Nature. 2017 Aug;548(7665):82–6. pmid:28770842
- 25. Antonation KS, Grützmacher K, Dupke S, Mabon P, Zimmermann F, Lankester F, et al. Bacillus cereus Biovar anthracis Causing Anthrax in Sub-Saharan Africa—Chromosomal Monophyly and Broad Geographic Distribution. Small PLC, editor. PLoS Negl Trop Dis. 2016 Sep 8;10(9):e0004923.
- 26. Zimmermann F, Köhler SM, Nowak K, Dupke S, Barduhn A, Düx A, et al. Low antibody prevalence against Bacillus cereus biovar anthracis in Taï National Park, Côte d’Ivoire, indicates high rate of lethal infections in wildlife. Foley J, editor. PLoS Negl Trop Dis. 2017 Sep 21;11(9):e0005960.
- 27. Dupke S, Barduhn A, Franz T, Leendertz FH, Couacy-Hymann E, Grunow R, et al. Analysis of a newly discovered antigen of Bacillus cereus biovar anthracis for its suitability in specific serological antibody testing. J Appl Microbiol. 2019 Jan;126(1):311–23.
- 28. Beyer W, Turnbull P. Anthrax in animals. Mol Aspects Med. 2009;30(6):481–9. pmid:19723532
- 29. Katani R, Schilling MA, Lyimo B, Eblate E, Martin A, Tonui T, et al. Identification of Bacillus anthracis, Brucella spp., and Coxiella burnetii DNA signatures from bushmeat. Sci Rep. 2021 Jul 21;11(1):14876.
- 30. Klee SR, Brzuszkiewicz EB, Nattermann H, Brüggemann H, Dupke S, Wollherr A, et al. The genome of a Bacillus isolate causing anthrax in chimpanzees combines chromosomal properties of B. cereus with B. anthracis virulence plasmids. PloS One. 2010 Jul 9;5(7):e10986.
- 31. Brézillon C, Haustant M, Dupke S, Corre JP, Lander A, Franz T, et al. Capsules, Toxins and AtxA as Virulence Factors of Emerging Bacillus cereus Biovar anthracis. Small PLC, editor. PLoS Negl Trop Dis. 2015 Apr 1;9(4):e0003455.
- 32. Mock M, Mignot T. Anthrax toxins and the host: a story of intimacy. Cell Microbiol. 2003 Jan;5(1):15–23. pmid:12542467
- 33. Okinaka RT, Cloud K, Hampton O, Hoffmaster AR, Hill KK, Keim P, et al. Sequence and Organization of pXO1, the Large Bacillus anthracis Plasmid Harboring the Anthrax Toxin Genes. J Bacteriol. 1999 Oct 15;181(20):6509–15.
- 34. Oh S, Budzik JM, Garufi G, Schneewind O. Two capsular polysaccharides enable Bacillus cereus G9241 to cause anthrax-like disease. Mol Microbiol. 2011 Apr;80(2):455–70.
- 35. Hammond SE, Hanna PC. Lethal factor active-site mutations affect catalytic activity in vitro. Infect Immun. 1998 May;66(5):2374–8. pmid:9573135
- 36. Klimpel KR, Arora N, Leppla SH. Anthrax toxin lethal factor contains a zinc metalloprotease consensus sequence which is required for lethal toxin activity. Mol Microbiol. 1994 Sep;13(6):1093–100. pmid:7854123
- 37. Leppla SH. Anthrax toxin edema factor: a bacterial adenylate cyclase that increases cyclic AMP concentrations of eukaryotic cells. Proc Natl Acad Sci U S A. 1982 May;79(10):3162–6. pmid:6285339
- 38. Labruyère E, Mock M, Ladant D, Michelson S, Gilles AM, Laoide B, et al. Characterization of ATP and calmodulin-binding properties of a truncated form of Bacillus anthracis adenylate cyclase. Biochemistry. 1990 May 22;29(20):4922–8.
- 39. Gummelt C, Dupke S, Howaldt S, Zimmermann F, Scholz HC, Laue M, et al. Analysis of Sporulation in Bacillus cereus Biovar anthracis Which Contains an Insertion in the Gene for the Sporulation Factor σK. Pathogens. 2023 Dec 13;12(12):1442.
- 40.
PHSBPRA. PUBLIC HEALTH SECURITY AND BIOTERRORISM PREPAREDNESS AND RESPONSE ACT OF 2002 [Internet]. PUBLIC LAW; 2002. Available from: https://www.congress.gov/107/plaws/publ188/PLAW-107publ188.pdf
- 41. Centers for Disease Control and Prevention (CDC), Department of Health and Human Services (HHS). Possession, Use, and Transfer of Select Agents and Toxins—Addition of Bacillus cereus Biovar anthracis to the HHS List of Select Agents and Toxins. Interim final rule and request for comments. Fed Regist. 2016 Sep 14;81(178):63138–43.
- 42. Rasko DA, Worsham PL, Abshire TG, Stanley ST, Bannan JD, Wilson MR, et al. Bacillus anthracis comparative genome analysis in support of the Amerithrax investigation. Proc Natl Acad Sci. 2011;108(12):5027–32.
- 43. Simonson TS, Okinaka RT, Wang B, Easterday WR, Huynh L, U’Ren JM, et al. Bacillus anthracis in China and its relationship to worldwide lineages. BMC Microbiol. 2009;9(1):71.
- 44.
CDC. Biosafety in Microbiological and Biomedical Laboratories (BMBL), 6th Ed. Cent Dis Control Prev [Internet]. 2020; Available from: https://www.cdc.gov/labs/BMBL.html
- 45.
Nicholson W, Setlow P. Molecular Biological Methods for Bacillus, eds. Harwood C. & Cutting S. In New York: John Wiley; 1990. p. 391–450.
- 46. Norris MH, Bluhm AP, Metrailer MC, Jiranantasak T, Kirpich A, Hadfield T, et al. Beyond the spore, the exosporium sugar anthrose impacts vegetative Bacillus anthracis gene regulation in cis and trans. Sci Rep. 2023 Mar 28;13(1):5060.
- 47. Norris M, Kirpich A, Bluhm A, Zincke D, Hadfield T, Ponciano J, et al. Convergent evolution of diverse Bacillus anthracis outbreak strains toward altered surface oligosaccharides that modulate anthrax pathogenesis. PLOS Biol. 2020 Dec 28;18:e3001052.
- 48. Powell JD, Hutchison JR, Hess BM, Straub TM. Bacillus anthracis spores germinate extracellularly at air-liquid interface in an in vitro lung model under serum-free conditions. J Appl Microbiol. 2015 Sep;119(3):711–23.
- 49. Hornstra LM, Van Der Voort M, Wijnands LM, Roubos-van Den Hil PJ, Abee T. Role of Germinant Receptors in Caco-2 Cell-Initiated Germination of Bacillus cereus ATCC 14579 Endospores. Appl Environ Microbiol. 2009 Feb 15;75(4):1201–3.
- 50. Gut IM, Tamilselvam B, Prouty AM, Stojkovic B, Czeschin S, Van Der Donk WA, et al. Bacillus anthracis spore interactions with mammalian cells: Relationship between germination state and the outcome of in vitro. BMC Microbiol. 2011 Dec;11(1):46.
- 51. Ireland JAW, Hanna PC. Amino acid- and purine ribonucleoside-induced germination of Bacillus anthracis DeltaSterne endospores: gerS mediates responses to aromatic ring structures. J Bacteriol. 2002 Mar;184(5):1296–303.
- 52. Clements MO, Moir A. Role of the gerI Operon of Bacillus cereus 569 in the Response of Spores to Germinants. J Bacteriol. 1998 Dec 15;180(24):6729–35.
- 53. Gao X, Swarge BN, Roseboom W, Setlow P, Brul S, Kramer G. Time-Resolved Proteomics of Germinating Spores of Bacillus cereus. Int J Mol Sci. 2022 Nov 6;23(21):13614.
- 54. Kumar P, Nagarajan A, Uchil PD. Analysis of Cell Viability by the Lactate Dehydrogenase Assay. Cold Spring Harb Protoc. 2018 Jun;2018(6):pdb.prot095497. pmid:29858337
- 55. Chabot DJ, Scorpio A, Tobery SA, Little SF, Norris SL, Friedlander AM. Anthrax capsule vaccine protects against experimental infection. Vaccine. 2004 Nov 15;23(1):43–7. pmid:15519706
- 56. De BK, Bragg SL, Sanden GN, Wilson KE, Diem LA, Marston CK, et al. A two-component direct fluorescent-antibody assay for rapid identification of Bacillus anthracis. Emerg Infect Dis. 2002 Oct;8(10):1060–5.
- 57. Ezzell J, Abshire T. Encapsulation of Bacillus anthracis spores and spore identification. Proceeding Int Workshop Anthrax. Salisubury Med Bull. 1999;87:42.
- 58. Kamar R, Gohar M, Jéhanno I, Réjasse A, Kallassy M, Lereclus D, et al. Pathogenic Potential of Bacillus cereus Strains as Revealed by Phenotypic Analysis. J Clin Microbiol. 2013 Jan;51(1):320–3.
- 59. Tsai CJY, Loh JMS, Proft T. Galleria mellonella infection models for the study of bacterial diseases and for antimicrobial drug testing. Virulence. 2016 Apr 2;7(3):214–29.
- 60. Thomas RJ, Hamblin KA, Armstrong SJ, Müller CM, Bokori-Brown M, Goldman S, et al. Galleria mellonella as a model system to test the pharmacokinetics and efficacy of antibiotics against Burkholderia pseudomallei. Int J Antimicrob Agents. 2013 Apr;41(4):330–6.
- 61. Seed KD, Dennis JJ. Development of Galleria mellonella as an Alternative Infection Model for the Burkholderia cepacia Complex. Infect Immun. 2008 Mar;76(3):1267–75.
- 62. Ignasiak K, Maxwell A. Galleria mellonella (greater wax moth) larvae as a model for antibiotic susceptibility testing and acute toxicity trials. BMC Res Notes. 2017 Dec;10(1):428. pmid:28851426
- 63. Wand ME, Müller CM, Titball RW, Michell SL. Macrophage and Galleria mellonella infection models reflect the virulence of naturally occurring isolates of B. pseudomallei, B. thailandensis and B. oklahomensis. BMC Microbiol. 2011 Dec;11(1):11.
- 64. Mukherjee K, Altincicek B, Hain T, Domann E, Vilcinskas A, Chakraborty T. Galleria mellonella as a Model System for Studying Listeria Pathogenesis. Appl Environ Microbiol. 2010 Jan;76(1):310–7.
- 65. Kavanagh K, Reeves EP. Exploiting the potential of insects for in vivo pathogenicity testing of microbial pathogens. FEMS Microbiol Rev. 2004 Feb;28(1):101–12. pmid:14975532
- 66. Mukherjee K, Mraheil MA, Silva S, Müller D, Cemic F, Hemberger J, et al. Anti-Listeria Activities of Galleria mellonella Hemolymph Proteins. Appl Environ Microbiol. 2011 Jun 15;77(12):4237–40.
- 67. Savransky V, Sanford DC, Syar E, Austin JL, Tordoff KP, Anderson MS, et al. Pathology and Pathophysiology of Inhalational Anthrax in a Guinea Pig Model. Pirofski L, editor. Infect Immun. 2013 Apr;81(4):1152–63.
- 68. Perry MR, Ionin B, Barnewall RE, Vassar ML, Reece JJ, Park S, et al. Development of a guinea pig inhalational anthrax model for evaluation of post-exposure prophylaxis efficacy of anthrax vaccines. Vaccine. 2020 Feb;38(10):2307–14. pmid:32029323
- 69. Welkos S, Bozue J, Twenhafel N, Cote C. Animal Models for the Pathogenesis, Treatment, and Prevention of Infection by Bacillus anthracis. Eichenberger P, Driks A, editors. Microbiol Spectr. 2015 Feb 27;3(1):3.1.12.
- 70. Twenhafel NA. Pathology of Inhalational Anthrax Animal Models. Vet Pathol. 2010 Sep;47(5):819–30. pmid:20656900
- 71. Levy H, Weiss S, Altboum Z, Schlomovitz J, Glinert I, Sittner A, et al. Differential Contribution of Bacillus anthracis Toxins to Pathogenicity in Two Animal Models. Blanke SR, editor. Infect Immun. 2012 Aug;80(8):2623–31.
- 72. Altboum Z, Gozes Y, Barnea A, Pass A, White M, Kobiler D. Postexposure prophylaxis against anthrax: evaluation of various treatment regimens in intranasally infected guinea pigs. Infect Immun. 2002 Nov;70(11):6231–41. pmid:12379702
- 73. Ngundi MM, Meade BD, Lin TL, Tang WJ, Burns DL. Comparison of three anthrax toxin neutralization assays. Clin Vaccine Immunol CVI. 2010 Jun;17(6):895–903. pmid:20375243
- 74. Peterson JW, Comer JE, Noffsinger DM, Wenglikowski A, Walberg KG, Chatuev BM, et al. Human monoclonal anti-protective antigen antibody completely protects rabbits and is synergistic with ciprofloxacin in protecting mice and guinea pigs against inhalation anthrax. Infect Immun. 2006 Feb;74(2):1016–24. pmid:16428748
- 75. Green BD, Battisti L, Koehler TM, Thorne CB, Ivins BE. Demonstration of a capsule plasmid in Bacillus anthracis. Infect Immun. 1985 Aug;49(2):291–7.
- 76. Mikesell P, Ivins B, Ristroph J, Vodkin M, Dreier T, MD. AMRIOIDFD. Plasmids, Pasteur, and anthrax. Defense Technical Information Center; 1983.
- 77. Norris M, Zincke D, Leiser O, Kreuzer H, Hadfied T, Blackburn J. Laboratory strains of Bacillus anthracis lose their ability to rapidly grow and sporulate compared to wildlife outbreak strains. PLOS ONE. 2020 Jan 24;15:e0228270.
- 78. Barandongo ZR, Dolfi AC, Bruce SA, Rysava K, Huang YH, Joel H, et al. The persistence of time: the lifespan of Bacillus anthracis spores in environmental reservoirs. Res Microbiol. 2023;104029.
- 79. Gogarten JF, Düx A, Mubemba B, Pléh K, Hoffmann C, Mielke A, et al. Tropical rainforest flies carrying pathogens form stable associations with social nonhuman primates. Mol Ecol. 2019 Sep;28(18):4242–58. pmid:31177585
- 80. Blackburn JK, Mullins JC, Van Ert M, Hadfield T, O’Shea B, Hugh-Jones ME. The necrophagous fly anthrax transmission pathway: Empirical and genetic evidence from a wildlife epizootic in west Texas 2010. Vector-Borne Zoonotic Dis. 2014;14(8):576–83.
- 81. Blackburn JK, Curtis AJ, Hadfield T, Hugh-Jones ME. Spatial and temporal patterns of anthrax in white-tailed deer, Odocoileus virginianus, and hematophagous flies in west Texas during the summertime anthrax risk period. Ann Assoc Am Geogr. 2014;
- 82. Dixon TC, Fadl AA, Koehler TM, Swanson JA, Hanna PC. Early Bacillus anthracis-macrophage interactions: intracellular survival survival and escape. Cell Microbiol. 2000 Dec;2(6):453–63.
- 83. Ross JM. The pathogenesis of anthrax following the administration of spores by the respiratory route. J Pathol Bacteriol. 1957 Apr;73(2):485–94.
- 84. Kang TJ, Fenton MJ, Weiner MA, Hibbs S, Basu S, Baillie L, et al. Murine macrophages kill the vegetative form of Bacillus anthracis. Infect Immun. 2005 Nov;73(11):7495–501.
- 85. Guidi-Rontani C. The alveolar macrophage: the Trojan horse of Bacillus anthracis. Trends Microbiol. 2002 Sep;10(9):405–9.
- 86. Kozel TR, Murphy WJ, Brandt S, Blazar BR, Lovchik JA, Thorkildson P, et al. mAbs to Bacillus anthracis capsular antigen for immunoprotection in anthrax and detection of antigenemia. Proc Natl Acad Sci U S A. 2004 Apr 6;101(14):5042–7.
- 87. Tonello F, Zornetta I. Bacillus anthracis Factors for Phagosomal Escape. Toxins. 2012 Jul 10;4(7):536–53.
- 88. Cote CK, Rossi CA, Kang AS, Morrow PR, Lee JS, Welkos SL. The detection of protective antigen (PA) associated with spores of Bacillus anthracis and the effects of anti-PA antibodies on spore germination and macrophage interactions. Microb Pathog. 2005 May;38(5–6):209–25.
- 89. Lee DY, Chun JH, Ha HJ, Park J, Kim BS, Oh HB, et al. Poly-gamma-d-glutamic acid and protective antigen conjugate vaccines induce functional antibodies against the protective antigen and capsule of Bacillus anthracis in guinea-pigs and rabbits. FEMS Immunol Med Microbiol. 2009 Nov;57(2):165–72.
- 90. Scorpio A, Chabot DJ, Day WA, O’Brien DK, Vietri NJ, Itoh Y, et al. Poly-γ-Glutamate Capsule-Degrading Enzyme Treatment Enhances Phagocytosis and Killing of Encapsulated Bacillus anthracis. Antimicrob Agents Chemother. 2007 Jan;51(1):215–22.
- 91. Sharma S, Bhatnagar R, Gaur D. Bacillus anthracis Poly-γ-D-Glutamate Capsule Inhibits Opsonic Phagocytosis by Impeding Complement Activation. Front Immunol. 2020 Mar 31;11:462.
- 92. O’Brien DK, Ribot WJ, Chabot DJ, Scorpio A, Tobery SA, Jelacic TM, et al. The capsule of Bacillus anthracis protects it from the bactericidal activity of human defensins and other cationic antimicrobial peptides. Blanke SR, editor. PLOS Pathog. 2022 Sep 29;18(9):e1010851.
- 93. Scarff JM, Seldina YI, Vergis JM, Ventura CL, O’Brien AD. Expression and contribution to virulence of each polysaccharide capsule of Bacillus cereus strain G9241. Koehler TM, editor. PLOS ONE. 2018 Aug 22;13(8):e0202701.
- 94. De Vos V, Van Rooyen GL, Kloppers JJ. Anthrax immunization of free-ranging Roan Antelope Hippotragus Equinus in the Kruger National Park. Koedoe. 1973 Jul 28;16(1):11–25.
- 95. Kaitho T, Ndeereh D, Ngoru B. An outbreak of anthrax in endangered Rothschild’s giraffes in Mwea National Reserve, Kenya. Vet Med Res Rep. 2013 Nov;45.
- 96. Peterson MJ, Davis DS, Templeton JW. An Enzyme-linked Immunosorbent Assay for Detecting Anthrax Antibody in White-tailed Deer (Odocoileus virginianus): Evaluation of Anthrax Vaccination and Sera from Free-ranging Deer. J Wildl Dis. 1993 Jan;29(1):130–5.
- 97. Shlyakhov EN, Rubinstein E. Human live anthrax vaccine in the former USSR. Vaccine. 1994 Jan;12(8):727–30. pmid:8091851
- 98. Turnbull PCB, Tindall BW, Coetzee JD, Conradie CM, Bull RL, Lindeque PM, et al. Vaccine-induced protection against anthrax in cheetah (Acinonyx jubatus) and black rhinoceros (Diceros bicornis). Vaccine. 2004 Sep;22(25–26):3340–7.
- 99. Corre JP, Piris-Gimenez A, Moya-Nilges M, Jouvion G, Fouet A, Glomski IJ, et al. In Vivo Germination of Bacillus anthracis Spores During Murine Cutaneous Infection. J Infect Dis. 2013 Feb 1;207(3):450–7.
- 100. Loving CL, Kennett M, Lee GM, Grippe VK, Merkel TJ. Murine Aerosol Challenge Model of Anthrax. Infect Immun. 2007 Jun;75(6):2689–98. pmid:17353290
- 101. Henderson DW, Peacock S, Belton FC. Observations on the prophylaxis of experimental pulmonary anthrax in the monkey. J Hyg (Lond). 1956 Mar;54(1):28–36. pmid:13319688
- 102. Turner A, Galvin J, Rubira R, Condron R, Bradley T. Experiences with vaccination and epidemiological investigations on an anthrax outbreak in Australia in 1997. J Appl Microbiol. 1999;87(2):294–7. pmid:10475972
- 103.
WHO. Anthrax in humans and animals. World Health Organization, International Office of Epizootics; 2008.