Skip to content


  • Research note
  • Open Access

The reporting of a Bacillus anthracis B-clade strain in South Africa after more than 20 years

  • 1, 5Email author,
  • 1,
  • 2,
  • 3,
  • 4,
  • 3 and
  • 1
BMC Research Notes201811:264

  • Received: 20 March 2018
  • Accepted: 20 April 2018
  • Published:



Anthrax is a disease with an age old history in Africa caused by the Gram-positive endospore forming soil bacterium Bacillus anthracis. Epizootics of wild ungulates occur annually in the enzootic region of Pafuri, Kruger National Park (KNP) in the Limpopo Province of South Africa. Rigorous routine surveillance and diagnostics in KNP, has not revealed these rare isolates since the 1990s, despite unabated annual outbreaks. In 2011 a cheetah was diagnosed as anthrax positive from a private game reserve in Limpopo Province and reported to State Veterinary Services for further investigation. Isolation, molecular diagnostics, whole genome sequencing and comparative genomics were carried out for B. anthracis KC2011.


Bacteriological and molecular diagnostics confirmed the isolate as B. anthracis. Subsequent typing and whole genome single nucleotide polymorphisms analysis indicated it clustered alongside B. anthracis SA A0091 in the B.Br.010 SNP branch. Unlike B. anthracis KrugerB strain, KC2011 strain has unique SNPs and represents a new branch in the B-clade. The isolation and genotypic characterisation of KC2011 demonstrates a gap in the reporting of anthrax outbreaks in the greater Limpopo province area. The identification of vulnerable and susceptible cheetah mortalities due to this strain has implications for conservation measures and disease control.


  • Bacillus anthracis
  • Whole genome sequencing (WGS)
  • Single nucleotide polymorphisms (SNPs)


Anthrax is a zoonotic disease with an age old history in Africa [1], causing acute mortalities. It primarily affects ungulates with episodic spill over into humans and carnivores [2, 3]. The causative agent of this disease is the Gram positive, endospore forming, exotoxin producing soil bacterium Bacillus anthracis [4, 5]. Sporulation is triggered by nutrient scarcity, the presence of bicarbonate and oxygen exposure [6, 7]. The virulence factors of this bacterium are on the two plasmids, pXO1 (181 kilobases) and pXO2 (94 kilobases), encoding the toxin and capsule genes respectively [8, 9]. The toxin complex is composed of three components namely, lethal toxin (LF) made up of the protective (PA) and lethal factor (EF) [10, 11]. The capsule consists of a five gene operon (capBCADE) that produces a poly-gamma-d-glutamic acid (PGA) on the bacterial cell membrane that protects the vegetative cells from phagocytosis [12, 13].

Anthrax was widespread across southern Africa in the early 20th Century, until the development and South African state implementation of the Sterne vaccine in 1937 [14, 15]. While livestock cases have since been infrequent due to vaccination practices; outbreaks are still common amongst wildlife in endemic regions of South Africa and game reserves across the continent [2, 16]. Limpopo province covers 125,754 km2 bordering Botswana and Zimbabwe to the north. The northern half of the Kruger National Park (KNP) makes up the entire eastern border of the Limpopo Province bordering Mozambique. The anthrax endemic region in KNP falls between the Limpopo and Luvhuvhu rivers [19]. The distribution of outbreaks and the animal species affected by anthrax have been described extensively in KNP in the last 35 years [1720]. The genetic population structure of B. anthracis consists of the global A, B and C clades [21]. The B clade isolates identified in South Africa were confined to the northern tip of KNP in Limpopo Province due to it being an ideal soil environment for spore persistence [17]. The A clade isolates, in contrast, have a wide distribution, predominating and prevailing over the rarer B clade isolates during outbreaks since the late 1980s.

Host species differ in their susceptibility to B. anthracis. Wild ungulates are more susceptible to the disease than carnivores [2, 3]. Even amongst herbivores, roan antelope (Hippotragus equinus) are far more susceptible to anthrax than other antelope species [20]. Similarly, cheetahs (Acinonyx jubatus) are more susceptible to the toxins produced by the bacterium than other carnivorous felids and canids [2224]. It is hypothesised that this could be due to the immune response dictated by the monomorphic genetic structure of southern African cheetahs [25] or by the feeding habits of free roaming cheetah [22]. In either event, it is of importance to cheetah conservation practices since these felids are considered vulnerable (bordering on endangered) [26].

This status has led to more stringent investigations of cheetah mortalities in both captive and free roaming animals. In 2011 a cheetah mortality from a private game reserve in Limpopo Province was investigated by the local veterinarian. The cheetah was observed to have substantial facial oedema with unclotted blood around the kidneys (Fig. 1A, B). Biological samples from the cheetah were submitted for bacteriologic diagnostics (Fig. 1C, D). Colony morphology indicative of B. anthracis then confirmed the initial diagnosis (Fig. 1D). The isolate was then submitted by Skukuza State Veterinary Services for further analysis on behest of the Anthrax Advisory Committee.
Fig. 1
Fig. 1

A Cheetah (Acinonyx jubatus) demonstrating facial oedema in reaction to Bacillus anthracis toxins. B Dark un-clotted blood surrounding the kidney of a cheetah indicating infection by B. anthracis.C Impression smear of oedematous fluid from cheetah with typical square ended B. anthracis bacilli (purple bacilli indicated by the arrow). D Bacteriologic diagnostics on 5% sheep blood agar demonstrating typical B. anthracis colony morphology and structure

Main text


Isolation and identification

A Giemsa smear stain from the oedema fluid was performed for identification of B. anthracis. Pure culture isolation of KC2011 isolate was obtained by culturing biological samples directly onto 5% sheep blood agar after 24 h incubation at 37 °C characteristics including colony and cell morphology, gamma-phage and penicillin sensitivity were evaluated as described in OIE [3]. Capsule formation was observed by visualization using light microscopy of Giemsa stained fluid and colonies.

DNA extractions and qPCR

Genomic extraction of B. anthracis KC2011 was carried out using a DNA Blood Mini Kit (Qiagen, Germany) according to manufacturer’s instructions. The DNA was quantified using Qubit™ fluorometric quantitation (Invitrogen™, USA). Quality of the extracted DNA was visualized on 0.8% agarose gel electrophoresis. The diagnostic qPCR for B. anthracis was performed with 2.5 µL DNA in 1× FastStart™ Taq DNA Polymerase mastermix (Roche®, Germany) and 0.5 μM of each primer small acid soluble proteins (SASP), Bacillus anthracis protective antigen (BAPA), and capsule region (capC) along with 0.2 μM of probe for each chromosomal and plasmid target pairs with fluorescein on the one and LCRed640 on the other (Tib MolBiol GmbH, Germany) in a final volume of 20 µL as described in Turnbull [3]. The PCR conditions on a LightCycler™ Nano (Roche®, Germany) were used as described in OIE [3, 27].

Melt-MAMA assay

The PCR single nucleotide polymorphism (SNP) assay was performed for the oligonucleotide markers A/B.Br.001, B.Br.003, B.Br.004, B.Br.002, B.Br.001, A.Br.007 and A.Br.002 as described in Birdsell et al. [28]. The reaction included 2.5 µL DNA diluted in 1× FastStart DNA Green Master (Roche®, Germany) with an ancestral forward and a derived forward SNP target primer (GC-clamp: no-GC-clamp) and a common reverse primer with a starting concentration of 0.2 µM depending on the ratio indicated which allowed for separation of melt peaks by at least 5 °C. Thermocycling parameters on the LightCycler™ Nano (Roche®, Germany) were conducted as described in Birdsell et al. [28].

Whole genome sequencing and Bioinformatics analysis

Sequence libraries of the DNA isolate was generated using the Nextera DNA Sample Prep Kit (Illumina, USA) protocol. Sequence reads of paired end library was performed on a HiSeq 2500 sequencer (Illumina, USA). Quality of the genome sequenced reads were assessed using FastQC software 0:10.1 [29]. Trimmomatic [30] was used to remove the ambiguous nucleotide reads. De novo assembly of the B. anthracis KC2011 was carried out using the CLC Genomics Workbench version 7.5 (CLC, Denmark). The assembled contigs were aligned with BLASTn [31] using B. anthracis Ames ancestor (GenBank: AE017334.2, AE017336.2, and AE017336.2) as a reference. Mauve tool [32] was used to align and order the assembled contigs using B. anthracis Ames ancestor as a reference. The genome was annotated using PGAAP at NCBI [33].

The trimmed reads of B. anthracis KC2011 were aligned to B. anthracis Ames ancestor using the Burrows-Wheeler Aligner (BWA) [34]. SAMtools [35] was used to sort and index the aligned sequenced reads. Unified genotyper in GATK [36] was used to call for SNPs. In order to construct WGS- SNPs tree, complete and draft genomes from different clades of B. anthracis available in NCBI Genbank ( were included in this study (Additional file 1: Table S1). SNPs positioning sets were deducted from the aligned genomes of B. anthracis Ames ancestor using molecular evolutionary genetics analysis (MEGA) 7 [37]. SNPs with informative sites (core SNPs) in all genome sequences were used for the phylogenetic tree construction using MEGA 7 tool [37].

Genbank submission

The genome sequence of B. anthracis KC2011 was deposited in the Genbank genome database under the accession number: NJGK00000000.


Isolation and identification

A Giemsa stained impression smear from the oedema fluid revealed encapsulated, square ended bacilli pathognomonic for B. anthracis (Fig. 1C). The microbiological characteristics confirmed the bacterium to be B. anthracis on blood smear with typical B. anthracis colony morphology and structure on 5% sheep blood agar [38] (Fig. 1D).

qPCR and Melt-MAMA analysis

The B. anthracis KC2011 isolate was confirmed positive for the presence of B. anthracis BAPA, SASP and capC. B. anthracis KC2011 was further typed using Melt-MAMA and amplified the derived markers for A/B.Br.001, B.Br.003 and B.Br.002, whilst amplifying the B.Br.001 ancestral marker. This SNP profile indicated KC2011 to be a B clade (B.Br.001/002), but not the rare KrugerB sub-clade.

Genome assembly and phylogeny

De novo assembly was carried out which resulted in 45 contigs (Table 1). The total genome contigs contributed to a 5.42 MB, with a GC content of 35%. A total of 6 047 coding sequences (CDSs) was determined in the KC2011 strain (Table 1). Comparative genome alignment of the B. anthracis KC2011 genome with Ames ancestor revealed no evidence of novel genes (Additional file 3: Fig. S1). The genome coverage of B. anthracis KC2011 was 99% covered to B. anthracis Ames ancestor (Additional file 1: Table S1; Additional file 2: Table S2). High copy number of pXO1 was observed represented by 725 coverage (725×).
Table 1

Genome features of the Bacillus anthracis KC2011


Bacillus anthracis KC2011 genome



Genome coverage (X)


Avg. length after trim


Genome size (bp)

5 421 567

Number of contigs


Maximum length

500 750

Minimum length


G+C content (%)


Genes (total)

6 047

CDS (coding)


Genes (RNA)


Complete rRNAS

4, 1 (5S, 23S)

Number of tRNA


Number of ncRNAs


Pseudo genes (total)


A high resolution tree of global B. anthracis strains was constructed to visualise the SNP grouping of the B. anthracis KC2011 (Fig. 2). This yielded 3 247 parsimony informative SNPs that were used to construct the phylogram. The B. anthracis KC2011 grouped in the B-clade separate from the KrugerB strain. The closest related branch was SA B. anthracis A0091.
Fig. 2
Fig. 2

Whole genome SNP phylogeny of the global Bacillus anthracis strains indicating the placement of KC2011 strain. Maximum likelihood was constructed using 3 247 parsimony informative SNPs. The rare B. anthracis KC2011 (indicated with higher font) grouped in the B-clade separately from KrugerB strain


Strain KC2011 is the first B. anthracis identified as a B-clade lineage bacterium in over 20 years in South Africa and represents a new subclade in the B-clade. About 90% of the anthrax outbreaks reported world-wide arise from the A-clade, while less than 10% have been reported from the B-clade (KrugerB subclade) [21]. The majority of the B. anthracis strains isolated from the northern part of KNP from 1970 to 1981 grouped in the B-clade (KrugerB subclade) but this clade became rare since the 1990s [17]. Only one B strain reported between 1982 and 1997 [19] to the extent that it was no longer isolated from 1997 until this report in South Africa.

No significant anthrax outbreaks were noted outside the endemic regions of KNP (northern part of KNP) in 2011 according to state veterinary surveillance data. The isolate KC2011 was only identified because of the delicate conservation status of cheetah in Africa and because it was within the context of the broader Limpopo province area (i.e. outside KNP). Cheetahs are especially susceptible to the B. anthracis exotoxins, however, they do respond well to vaccination with the Sterne vaccine [25]. This study highlighted the importance of vaccinating cheetah, where logistically possible, in conservancies across Limpopo province.

The phylogenetic structure of B. anthracis determined in this study (Fig. 2) is similar to other reported WGS-SNP trees [3942]. Past B-clade (1965–1990) isolates from the State Veterinary archival collection have all clustered in the KrugerB lineage. WGS-SNP analysis defined KC2011 strain grouping as a distinctive sub-clade within southern Africa B.Br.010.

Although, distant in terms of number of SNPs, the isolate A0091 and KC2011 belong to a common albeit broader lineage. The isolate A0091 is recorded as being from South Africa circa 1939, but has no other information to link it to KNP or Limpopo Province. There is also a dearth of information from the countries (Zimbabwe and Mozambique) bordering the South African province across the Limpopo River. Further surveillance of Limpopo Province is required to reveal whether deposits of B-clade isolates are subsisting outside KNP or whether this was an incidental mortality due to circumstances outside the realm of current monitoring criteria. Surveillance in the non-endemic anthrax regions is difficult as there are private game reserves and rural livestock farmers bordering KNP. Moreover the rural subsistence farmers (predominantly cattle farmers) are less likely to report mortalities and/or vaccinate their livestock thus complicating disease monitoring and control. This report of the B. anthracis KC2011 isolated outside the endemic region highlights the necessity of disease surveillance in non-endemic regions.


The identification of KC2011 is significant in that mortalities linked to the B-clade have diminished over several decades and therefore was estimated to have disappeared from the local environment. This suggests a possible gap in surveillance of carcasses submitted for anthrax diagnostics in the Limpopo province especially in the non-endemic regions. It also has implications for the inclusion of the Sterne vaccine as part of the conservation measures to be included for cheetah in sanctuaries and private game reserves across the Limpopo Province in South Africa.


The tree topology of the southern African strains was not well characterised due to the limited number of genomes available. Branch lengths vary according to inclusion of number and variety genomes in the data sets, as this explicates the informative SNPs used to construct the tree. Isolate KC2011 will thus be the basis for further typing of B. anthracis strains for southern Africa.



poly-gamma-d-glutamic acid


lethal toxin




lethal factor


Kruger National Park


whole genome sequencing


single nucleotide polymorphism


melting mismatch amplification mutation analysis


coding sequences


Authors’ contributions

KEL and AH performed the molecular, as well as, in silico analysis and drafting of the manuscript. RL and PR identified the mortality and performed the pathology evaluation. LDKL and HED performed the bacterial and DNA isolations. HvH participated in the design of the study, drafting the manuscript, revising it critically and provided funding. All authors read and approved the final manuscript.


We would like to thank the Anthrax Advisory Committee.

Competing interests

The authors declare that they have no competing interests.

Availability of data and materials

All data generated or analysed during this study are included in this published article.

Consent for publication

Not applicable.

Ethics approval and consent to participate

Not applicable.


This study was funded by the National Research Foundation (NRF) of South Africa.

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (, which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated.

Authors’ Affiliations

Department of Veterinary Tropical Diseases, Faculty of Veterinary Science, University of Pretoria, Onderstepoort, South Africa
Provet Wildlife Services, Raptors Safari Junction, Main Road, Hoedspruit, South Africa
State Veterinary Services, Department of Agriculture, Forestry and Fisheries, Skukuza, South Africa
Vetdiagnostix–Veterinary Pathology Services, 257 Boshoff Street, Pietermaritzburg, South Africa
College of Agriculture and Environmental Sciences, University of South Africa, Christiaan De Wet/Pioneer Dr., Florida, South Africa


  1. Farrar WE. Anthrax: from mesopotamia to molecular biology. Pharos Alpha Omega Alpha Honor Med Soc. 1995;58(2):35–8.PubMedGoogle Scholar
  2. Hugh-Jones ME, de Vos V. Anthrax and wildlife. Rev Sci Tech. 2002;21(2):359–83.View ArticlePubMedGoogle Scholar
  3. Organization WH. Epizootics IOo: anthrax in humans and animals. Geneva: World Health Organization; 2008.Google Scholar
  4. Leppla SH. Anthrax toxin edema factor: a bacterial adenylate cyclase that increases cyclic AMP concentrations of eukaryotic cells. Proc Natl Acad Sci USA. 1982;79(10):3162–6.View ArticlePubMedPubMed CentralGoogle Scholar
  5. Sterne M. Variation in Bacillus anthracis. Onderstepoort J Vet Sci Anim Ind. 1937;8:271–349.Google Scholar
  6. Koehler TM. Bacillus anthracis physiology and genetics. Mol Aspects Med. 2009;30(6):386–96.View ArticlePubMedPubMed CentralGoogle Scholar
  7. Sterne M. The effects of different carbon dioxide concentrations on the growth of virulent anthrax strains: pathogenicity and immunity tests on Guineapigs and sheep with anthrax variants derived from virulent strains. Onderstepoort J Vet Sci. 1937;9:49–67.Google Scholar
  8. Okinaka R, Cloud K, Hampton O, Hoffmaster A, Hill K, Keim P, Koehler T, Lamke G, Kumano S, Manter D, et al. Sequence, assembly and analysis of pX01 and pX02. J Appl Microbiol. 1999;87(2):261–2.View ArticlePubMedGoogle Scholar
  9. Pannucci J, Okinaka RT, Sabin R, Kuske CR. Bacillus anthracis pXO1 plasmid sequence conservation among closely related bacterial species. J Bacteriol. 2002;184(1):134–41.View ArticlePubMedPubMed CentralGoogle Scholar
  10. Mock M, Fouet A. Anthrax. Annu Rev Microbiol. 2001;55:647–71.View ArticlePubMedGoogle Scholar
  11. Mock M, Mignot T. Anthrax toxins and the host: a story of intimacy. Cell Microbiol. 2003;5(1):15–23.View ArticlePubMedGoogle Scholar
  12. Green BD, Battisti L, Koehler TM, Thorne CB, Ivins BE. Demonstration of a capsule plasmid in Bacillus anthracis. Infect Immun. 1985;49(2):291–7.PubMedPubMed CentralGoogle Scholar
  13. Drysdale M, Bourgogne A, Hilsenbeck SG, Koehler TM. atxA controls Bacillus anthracis capsule synthesis via acpA and a newly discovered regulator, acpB. J Bacteriol. 2004;186(2):307–15.View ArticlePubMedPubMed CentralGoogle Scholar
  14. Viljoen PR. Anthrax in South Africa. In: 13th and 14th reports of the director of veterinary education and research, Onderstepoort, South Africa; 1928.Google Scholar
  15. Sterne M. Distribution and economic importance of anthrax. Fed Proc. 1967;26(5):1493–5.PubMedGoogle Scholar
  16. Pienaar UD. Epidemiology of anthrax in wild animals and the control of anthrax epizootics in the Kruger National Park, South Africa. Fed Proc. 1967;26(5):1496–502.PubMedGoogle Scholar
  17. Smith KL, DeVos V, Bryden H, Price LB, Hugh-Jones ME, Keim P. Bacillus anthracis diversity in Kruger National Park. J Clin Microbiol. 2000;38(10):3780–4.PubMedPubMed CentralGoogle Scholar
  18. De Vos V, Bryden H. Anthrax in the Kruger National Park: temporal and spatial patterns of disease occurrence. Salisbury Med Bull. 1996;87:26–31.Google Scholar
  19. Odendaal MW, Pieterson PM, de Vos V, Botha AD. The antibiotic sensitivity patterns of Bacillus anthracis isolated from the Kruger National Park. Onderstepoort J Vet Res. 1991;58(1):17–9.PubMedGoogle Scholar
  20. De Vos V. The ecology of anthrax in the Kruger National Park, South Africa. Salisbury Med Bull Suppl. 1990;68:19–23.Google Scholar
  21. Van Ert MN, Easterday WR, Huynh LY, Okinaka RT, Hugh-Jones ME, Ravel J, Zanecki SR, Pearson T, Simonson TS, U’Ren JM, et al. Global genetic population structure of Bacillus anthracis. PLoS ONE. 2007;2(5):e461.View ArticlePubMedPubMed CentralGoogle Scholar
  22. Good KM, Houser A, Arntzen L, Turnbull PC. Naturally acquired anthrax antibodies in a cheetah (Acinonyx jubatus) in Botswana. J Wildl Dis. 2008;44(3):721–3.View ArticlePubMedGoogle Scholar
  23. Good KM, Marobela C, Houser AM. A report of anthrax in cheetahs (Acinonyx jubatus) in Botswana. J S Afr Vet Assoc. 2005;76(4):186.View ArticleGoogle Scholar
  24. Jager HG, Booker HH, Hubschle OJ. Anthrax in cheetahs (Acinonyx jubatus) in Namibia. J Wildl Dis. 1990;26(3):423–4.View ArticlePubMedGoogle Scholar
  25. Turnbull PC, Tindall BW, Coetzee JD, Conradie CM, Bull RL, Lindeque PM, Huebschle OJ. Vaccine-induced protection against anthrax in cheetah (Acinonyx jubatus) and black rhinoceros (Diceros bicornis). Vaccine. 2004;22(25–26):3340–7.View ArticlePubMedGoogle Scholar
  26. Durant SM. Predator avoidance, breeding experience and reproductive success in endangered cheetahs, Acinonyx jubatus. Anim Behav. 2000;60(1):121–30.View ArticlePubMedGoogle Scholar
  27. Ellerbrok H, Nattermann H, Ozel M, Beutin L, Appel B, Pauli G. Rapid and sensitive identification of pathogenic and apathogenic Bacillus anthracis by real-time PCR. FEMS Microbiol Lett. 2002;214(1):51–9.View ArticlePubMedGoogle Scholar
  28. Birdsell DN, Pearson T, Price EP, Hornstra HM, Nera RD, Stone N, Gruendike J, Kaufman EL, Pettus AH, Hurbon AN, et al. Melt analysis of mismatch amplification mutation assays (Melt-MAMA): a functional study of a cost-effective SNP genotyping assay in bacterial models. PLoS ONE. 2012;7(3):e32866.View ArticlePubMedPubMed CentralGoogle Scholar
  29. Andrews S. FastQC: a quality control tool for high throughput sequence data. 2010.
  30. Bolger AM, Lohse M, Usadel B. Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics. 2014;30(15):2114–20.View ArticlePubMedPubMed CentralGoogle Scholar
  31. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ. Basic local alignment search tool. J Mol Biol. 1990;215(3):403–10.View ArticlePubMedGoogle Scholar
  32. Darling AC, Mau B, Blattner FR, Perna NT. Mauve: multiple alignment of conserved genomic sequence with rearrangements. Genome Res. 2004;14(7):1394–403.View ArticlePubMedPubMed CentralGoogle Scholar
  33. Pruitt KD, Tatusova T, Klimke W, Maglott DR. NCBI reference sequences: current status, policy and new initiatives. Nucleic Acids Res. 2009;37((Database issue)):D32–6.View ArticlePubMedGoogle Scholar
  34. Li H, Durbin R. Fast and accurate short read alignment with Burrows–Wheeler transform. Bioinformatics. 2009;25(14):1754–60.View ArticlePubMedPubMed CentralGoogle Scholar
  35. Li H, Handsaker B, Wysoker A, Fennell T, Ruan J, Homer N, Marth G, Abecasis G, Durbin R. Genome project data processing S: the sequence alignment/map format and SAMtools. Bioinformatics. 2009;25(16):2078–9.View ArticlePubMedPubMed CentralGoogle Scholar
  36. McKenna A, Hanna M, Banks E, Sivachenko A, Cibulskis K, Kernytsky A, Garimella K, Altshuler D, Gabriel S, Daly M, et al. The genome analysis toolkit: a MapReduce framework for analyzing next-generation DNA sequencing data. Genome Res. 2010;20(9):1297–303.View ArticlePubMedPubMed CentralGoogle Scholar
  37. Kumar S, Stecher G, Tamura K. MEGA7: molecular evolutionary genetics analysis version 7.0 for bigger datasets. Mol Biol Evol. 2016;33(7):1870–4.View ArticlePubMedGoogle Scholar
  38. Butler J, MacCallum I, Kleber M, Shlyakhter IA, Belmonte MK, Lander ES, Nusbaum C, Jaffe DB. ALLPATHS: de novo assembly of whole-genome shotgun microreads. Genome Res. 2008;18(5):810–20.View ArticlePubMedPubMed CentralGoogle Scholar
  39. Pearson T, Okinaka RT, Foster JT, Keim P. Phylogenetic understanding of clonal populations in an era of whole genome sequencing. Infect Genet Evol. 2009;9(5):1010–9.View ArticlePubMedGoogle Scholar
  40. Vergnaud G, Girault G, Thierry S, Pourcel C, Madani N, Blouin Y. Comparison of French and Worldwide Bacillus anthracis strains favors a recent, post-Columbian origin of the predominant North-American Clade. PLoS ONE. 2016;11(2):e0146216.View ArticlePubMedPubMed CentralGoogle Scholar
  41. Derzelle S, Girault G, Kokotovic B, Angen O. Whole genome-sequencing and phylogenetic analysis of a historical collection of bacillus anthracis strains from Danish cattle. PLoS ONE. 2015;10(8):e0134699.View ArticlePubMedPubMed CentralGoogle Scholar
  42. Girault G, Blouin Y, Vergnaud G, Derzelle S. High-throughput sequencing of Bacillus anthracis in France: investigating genome diversity and population structure using whole-genome SNP discovery. BMC Genomics. 2014;15:288.View ArticlePubMedPubMed CentralGoogle Scholar


© The Author(s) 2018