An Investigation of Enterococcus Species Isolated from the African Buffalo (Syncerus caffer) in Serengeti National Park, Tanzania

Abstract

We isolated Enterococcus species that colonized in the African buffalo (Syncerus caffer) in order to investigate their genetic relatedness and antimicrobial susceptibility. A total of 219 isolates were obtained and a 16S rRNA gene sequence analysis showed they were classified into Enterococcus avium, E. casseliflavus, E. faecalis, E. faecium, E. hirae, or E. mundtii. Multilocus sequence typing of E. faecalis and E. faecium isolates indicated that some of the isolates showed an evolutionary distance that was far from the primary founders. The antimicrobial susceptibility of the enterococcal isolates suggested that the significant transmission of antimicrobial resistance via human intervention had not yet occurred.

Human and animal microbiomes have developed resistance to commonly used antimicrobials; however, most antimicrobial resistance studies have focused on humans and domestic animals, and not on wild animals (3, 26). Limited information is currently available on the gut microbiome of animals living in a wildlife protection ecosystem.

Serengeti National Park is known for its incredible scenery and magnificent wildlife. It covers 14,763 km² (1°28′−3°17′ S, 33°05′−35°20′ E) in northern Tanzania (altitude of 910 to 1820 m a.s.l). The Serengeti ecosystem is a part of Tanzania’s protected area, which encompasses a geographical region of wildebeest migration (10). The African buffalo (Syncerus caffer) is a key species in the Serengeti ecosystem, and these animals roam freely in Serengeti National Park. They graze on various grass species, and their movement within the park depends on water and grass availability (10). They interact with humans and occasionally with livestock via common water sources and grazing areas. Human–wildlife contact has increased concomitantly with increments in human populations and travel (26).

Since enterococci and Escherichia coli occur within the gut microbiome of humans and animals (1, 13, 31) and easily acquire antimicrobial resistance, they may serve as an indicator organism for the emergence of antimicrobial resistant bacteria (29). We selected commensal bacteria, particularly enterococci, colonizing the gastrointestinal tract of African buffaloes to survey antimicrobial resistance in wild ecosystems. Enterococci are non-spore-forming Gram-positive facultative anaerobic bacteria with spherical cocci appearing separately, in pairs, or in short chains (27). Most enterococci are not virulent and are considered to be relatively harmless, with little potential for human infection (27). However, they have occasionally been identified as nosocomial opportunistic pathogens that exhibit increased resistance to antimicrobials (22). Antimicrobial resistance in enterococci may be divided into two classes: intrinsic resistance and acquired resistance. Intrinsic resistance is due to either a lack of target sites for antimicrobials or the insufficient penetration of antimicrobials to the intracellular target. Acquired resistance arises from genetic mutations or the addition of extrinsic genes. Intrinsic and acquired resistance both hinder the treatment of Enterococcus infection (20, 23). Our study was prompted by concerns of the transfer risk of antimicrobial resistance because of human intervention within the wild animal habitat. This study also aimed to increase our understanding of Enterococcus species that colonized in the African buffalo. In order to achieve this, we used molecular techniques and antimicrobial susceptibility testing, with a focus on genetic relatedness among Enterococcus faecalis and E. faecium based on multilocus sequence typing (MLST).

During routine surveillance for foot and mouth disease and Rift Valley fever virus in July 2012, 105 ear swab samples and 104 rectal swab samples were collected from healthy African buffaloes in the Seronera region in the Serengeti, in which various wild fauna are present. Ear and rectal samples were obtained using cotton swabs, which were placed in Amies transport media without charcoal (Yuhan Lab Tech, Seoul, Korea) after sample collection. Samples were kept in a cold container and delivered to the laboratory in the Tanzania Wildlife Research Institute (TAWIRI) according to the rules of the UN recommendations on the transport of dangerous goods category 2 and then maintained at −20°C until transported. Samples were delivered via air freight in a cold container, which took approximately 48 h. Upon arrival, samples were streaked onto Enterococcosel agar (BBL Becton Dickinson, Sparks, MD, USA). After a 48-h incubation at 37°C, black colonies with a different colony morphology from other colonies on the plate were subcultured on a new Enterococcosel agar plate and trypticase soy agar (BBL) with 5% defibrinated sheep blood and incubated for 24 h at 37°C. After cell morphology was examined under a microscope, each isolated colony was stored at −70°C in 20% glycerol and deposited at the Culture Collection of Antimicrobial Resistant Microbes (CCARM, Seoul, Korea). Bacterial cells grown on tryptic soy agar (BBL) for 24 h at 37°C were suspended in sterile saline and then harvested by centrifugation at 14,000×g for 10 min. Genomic DNA was extracted in 30 μL of 10 mM TE buffer (10 mM Tris-Cl, 1 mM EDTA, pH 8.5) using a G-spin genomic extraction kit (Intron Biotechnology, Seoul, Korea) and stored at −20°C until analyzed. PCR of the 16S rRNA gene was performed with 16S rRNA primers (27F, 5′-AGA GTT TGA TCM TGG CTC AG-3′; 1088R, 5′-GCT CGT TGC GGG ACT TAA CC-3′) to amplify the coding region within the 16S rRNA gene (21). DNA fragments were purified using the QIAquick Gel extraction Kit (Qiagen, Valencia, CA, USA) in accordance with the manufacturer’s instructions, and sequence reactions were performed with an ABI 3100 automated sequencer (Applied Biosystems) by Bionix (Seoul, Korea). DNA sequences were analyzed using the online BLAST algorithm at the National Center for Biotechnology Information web server (http://www.ncbi.nlm.nih.gov) and the EzTaxon server (http://www.ezbiocloud.net/identify) (18) based on 16S rRNA gene sequence data.

Minimal inhibitory concentrations (MICs) were assayed on Mueller–Hinton agar (MH, BBL) in accordance with the recommendations of the Clinical Laboratory Standards Institute (6). The following 10 antimicrobials were tested: ampicillin, chloramphenicol, ciprofloxacin, erythromycin, gentamicin, norfloxacin, synercid, teicoplanin, tetracycline, and vancomycin. All chemicals were purchased from Sigma (St. Louis, MO, USA), except for synercid (May & Baker, Guildford, UK) and teicoplanin (Hoechst Marion Roussel, Kansas City, MO, USA). Enterococcus faecalis ATCC 29212 was included in each batch of the agar dilution test, and CLSI-approved MIC quality-control limits for E. faecalis were used. MIC was assessed following the criteria of Enterococcus species by CLSI (7, 8). Vancomycin MIC higher than 8 μg mL⁻¹ by the agar dilution method was confirmed by Etest^® (BioMérieux SA, Marcy l’Etoile, France).

PCR amplification was performed using primers specific to seven housekeeping genes of E. faecalis (gdh for glucose-6-phosphate dehydrogenase, gyd for glyceraldehyde-3-phosphate dehydrogenase, pstS for the phosphate ATP-binding cassette transporter, gki for glucokinase, aroE for shikimate-5-dehydrogenase, xpt for xanthine phosphoribosyltransferase, and yiqL for acetyl-CoA acetyltransferase) according to the methods of Ruiz-Garbajosa et al. (28). Regarding E. faecium, primers specific to seven housekeeping genes (adk for adenylate kinase, atpA for the ATP synthase, alpha subunit, ddl for D-alanine:D-alanine ligase, gyd for glyceraldehyde-3-phosphate dehydrogenase, gdh for glucose-6-phosphate dehydrogenase, purK for the phosphoribosylaminoimidazol carboxylase ATPase subunit, and pstS for the phosphate ATP-binding cassette transporter) were used according to the method of Homan et al. (16). Amplified products were sequenced in both directions using the PCR primers by Bionix. Sequence types (STs) were identified by comparing the sequences with alleles from the E. faecalis and E. faecium MLST database (http://www.mlst.net). New alleles and STs were submitted to the database curator for assignment. MLST datasets were analyzed using the eBURST algorithm to assess evolutionary relatedness among ST variants. Groups of STs sharing five out of seven alleles were defined as belonging to the same eBURST group, as described elsewhere (32). All available STs of E. faecalis and E. faecium in the MLST database were simultaneously analyzed and illustrated as population snapshots using the eBURST algorithm (11).

A total of 219 presumptive Enterococcus isolates were obtained from 105 ear swab samples and 104 fecal swab samples from African buffaloes. Ninety-six colonies grew from ear samples and 123 colonies from fecal samples (Table S1 and S2). Enterococcus colonies were white or yellow in color on trypticase soy agar with 5% defibrinated sheep blood. Bacterial cells of these colonies were found to be cocci when examined under a microscope after Gram staining. The 16S rRNA gene sequences of the isolates were classified into six groups and the phylogenetic tree indicated that they belonged to E. avium, E. casseliflavus, E. faecalis, E. faecium, E. hirae, and E. mundtii respectively, in the genus Enterococcus (Table 1 and Fig. S1)

Table 1 Proportion and minimal inhibitory concentrations of Enterococcus species isolated from ear and rectal swab samples of the African buffalo (Syncerus caffer) in the Serengeti, Tanzania.

Species (no. of isolates)	No. of isolates (%)			MIC (μg mL−1) rangea
	Ear	Fecal	Total	AMP (≥16)	CM (≥32)	CIP (≥4)	EM (≥8)	GM (≥500)	NOR (≥16)	SYN (≥4)	TEI (≥32)	TET (≥16)	VAN (≥32)
E. avium	6 (6.3)	0 (0)	6 (2.7)	≤0.25–1	4–8	1–2	≤0.25	0.5–4	1–4	1–4	0.13–2	≤0.25–1	≤0.5–4
E. casseliflavus	67 (69.8)	43 (35.0)	110 (50.2)	≤0.25–1	4–8	0.5–2	≤0.25–4	1–8	2–16	0.5–8	0.06–4	≤0.25–1	1–8
E. faecalis	12 (12.5)	7 (5.7)	19 (8.7)	≤0.25–2	4–8	0.5–2	≤0.25–1	2–32	1–8	2–16	0.06–4	0.5–1	1–8
E. faecium	1 (10)	15 (12.2)	16 (7.3)	≤0.25–4	4–8	0.5–2	≤0.25–4	2–8	1–8	≤0.25–2	0.25–4	≤0.25–0.5	≤0.5–8
E. hirae	0 (0)	3 (2.4)	3 (1.4)	1–2	4–8	≤0.25–1	≤0.25	2–8	1–4	1–2	0.13–0.5	≤0.25–1	1
E. mundtii	10 (10.4)	55 (44.7)	65 (29.7)	≤0.25–1	4–8	0.5–2	≤0.25–1	1–8	2–16	≤0.25–8	≤0.03–4	≤0.25–1	≤0.5–12
Total	96 (100)	123 (100)	219 (100)	≤0.25–4	4–8	≤0.25–2	≤0.25–4	0.5–32	1–16	≤0.25–8	≤0.03–4	≤0.25–1	≤0.5–12
% Susceptibleb				100	100	100	100	100	99.1c	100	100	100	100

^a  AMP, ampicillin; CM, chloramphenicol; CIP, ciprofloxacin; EM, erythromycin; GM, gentamicin; NOR, norfloxacin; SYN, synercid; TEI, teicoplanin; TET, tetracycline; VAN, vancomycin. Values in parentheses are the criterion limits for resistance according to the Clinical Laboratory Standards Institute (CLSI).

^b  Proportion of isolates that are susceptible to an antimicrobial.

^c  Two isolates E. casseliflavus CCARM 5297 and E. mundtii CCARM 5284 detected from a fecal sample were 16 μg mL⁻¹.

The MICs of Enterococcus species from African buffaloes are shown in Table 1. All enterococci isolates, except for two (E. casseliflavus CCARM 5297 and E. mundtii CCARM 5284), were 100% susceptible to ampicillin, chloramphenicol, ciprofloxacin, erythromycin, teicoplanin, tetracycline, and vancomycin. The MICs of E. casseliflavus CCARM 5297 and E. mundtii CCARM 5284 against norfloxacin were 16 μg mL⁻¹, which indicates resistance according to the CLSI criteria (≥16 μg mL⁻¹).

Thirteen STs were obtained from 19 isolates of E. faecalis. Among these, two STs were found in the existing database, while 11 STs were new STs (Table 2). The 11 new STs were assigned in the E. faecalis PubMLST database (http://pubmlst.org/efaecalis/). Five out of 11 STs (ST-648, ST-650, ST-653, ST-654, and ST-667) were new combinations of previously registered alleles. Six STs (ST-649, ST-651, ST-652, ST-665, ST-666, and ST-668) were new combinations of previously registered alleles with the new allele sequences. Nine out of 13 new STs were represented by a single isolate. Two existing STs (ST-273 and ST-436) were identical to the STs assigned to fecal isolates from non-hospitalized individuals in Germany and France, respectively (http://pubmlst.org/efaecalis). Sixteen isolates of E. faecium yielded four novel STs (Table 2). These STs were submitted to the MLST database (http://pubmlst.org/efaeium/) and designated as ST-1081, ST-1082, ST-1083, and ST-1084. In accordance with the same eBURST group defined as groups of STs sharing five out of seven alleles, 74 eBURST groups and 136 singletons of the entire E. faecalis STs were obtained, and the 13 STs from 19 E. faecalis isolates belonged to seven eBURST groups and three singletons (Table 2). Sixteen groups and 49 singletons were produced from all E. faecium STs. ST-1084 belonged to eBURST group 1, ST-1081 and ST-1082 belonged to eBURST group 8, and ST-1083 was a singleton. E. faecalis and E. faecium MLST databases are displayed as a single eBURST diagram by setting the group definition to zero of seven shared alleles (Fig. S2 and S3).

Table 2 Multilocus sequence types of 19 Enterococcus faecalis and 16 E. faecium isolates from the African buffalo (Syncerus caffer) in the Serengeti, Tanzania.

E. faecalis isolates (n=19)
STa	Alleleb							No. of isolates	eBURST groupc
	gdh	gyd	pstS	gki	aroE	xpt	yiqL
ST-273	4	8	12	13	40	46	22	1	26
ST-436	12	7	3	17	6	20	5	1	1
ST-648	12	10	3	17	31	2	5	3	1
ST-649	15	2	80	1	3	15	11	3	4
ST-650	17	7	16	5	75	1	9	1	singleton
ST-651	79	5	79	81	85	73	79	2	50
ST-652	34	1	81	82	3	74	79	1	singleton
ST-653	4	28	17	37	29	23	17	1	20
ST-654	20	1	14	15	3	14	9	2	52
ST-665	80	6	7	16	11	11	11	1	16
ST-666	12	5	79	81	85	73	79	1	50
ST-667	14	8	12	13	31	46	22	1	26
ST-668	81	1	17	6	85	14	28	1	singleton
E. faecium isolates (n=16)
ST	Allele							No. of isolates	eBURST group
	atpA	ddl	gdh	purK	gyd	pstS	adk
ST-1081	2	9	72	7	22	100	1	1	8
ST-1082	2	9	1	7	22	100	1	8	8
ST-1083	20	77	6	81	1	101	1	1	singleton
ST-1084	62	9	1	7	1	1	1	6	1

^a  ST, sequence type. New STs identified in this study are in boldface.

^b  New alleles identified in this study are written in italics and boldface.

^c  STs sharing five out of seven alleles are defined as the same eBURST group.

Advanced molecular technologies have led to considerable changes in the taxonomy of the genus Enterococcus, which currently consists of 35 recognized species, with eight more species to be added (27). Enterococci may reside in the gut of animals, and play an important role in nutrition and health (29). Human and animal gastrointestinal tracts represent reservoirs for enterococci (27). A wide variety of Enterococcus spp. have been recovered from humans and various animals, and differ greatly from each other (17). In general, the predominant species in fecal samples of healthy humans and animals are E. faecium followed by E. faecalis. In human infections, E. faecalis was the most frequently detected Enterococcus species until the mid-1990s. E. faecium has increased, possibly because of the spread of antimicrobial resistance, which particularly affects the incidence of nosocomial infection (22, 25, 31). For example, E. faecium has become a vanA-type (vancomycin-resistant, teicoplanin-susceptible) strain with the increased use of the antimicrobial vancomycin, and has better survival than E. faecalis. Other species, including E. durans, E. gallinarum, and E. casseliflavus, are less common isolates (31). E. avium, E. durans, E. gallinarum, E. casseliflavus, E. hirae, E. mundtii, and E. raffinosus have also been isolated from human infections at low frequencies (14, 31). In isolates from plants, E. casseliflavus, E. mundtii, and E. sulfureus (9, 22) with a yellow pigment have commonly been detected (2). Since the African buffalo is a herbivore, the high frequencies of E. casseliflavus and E. mundtii detected in this study may be related to their diet. In comparison with these species, E. faecalis (8.7%) and E. faecium (7.3%) showed markedly lower isolation rates from the African buffalo than E. casseliflavus and E. mundtii, which is inconsistent with previous findings (24, 30, 31). We presumed that the distributions of enterococci vary in different animals according to the species, geographical region, and environmental conditions, such as diet.

In the present study, antimicrobial susceptibility test results revealed no resistant Enterococcus isolates, except for two norfloxacin-resistant isolates with MICs of 16 μg mL⁻¹. Since fluoroquinolones show poor or moderate activity against enterococci and fluoroquinolones are rarely used in the treatment of Enterococcus infection (4, 17), these two isolates do not appear to be important. However, because fluoroquinolone resistance may be transferred to other genera, further studies to characterize the two fluoroquinolone-resistant strains are underway. Fluoroquinolone resistance may occur under the following conditions: a single mutation in the gyrA gene; decreased accumulation in bacteria due to mutations in the outer membrane proteins; and efflux pumps and/or plasmid-mediated resistance qnr gene (4, 19, 33). Since enterococci are facultative anaerobes, they have intrinsic low-level resistance to aminoglycosides via limited drug uptake, which is associated with the proteins involved in electron transport, with MICs ranging between 4 μg mL⁻¹ and 256 μg mL⁻¹ (5, 15, 23). Our results show that the antimicrobial susceptibilities of Enterococcus isolates from the African buffalo were significantly lower than those reported for others (e.g., humans, livestock, companion animals, and wildlife animals) (2, 10, 12, 20). Acquired antimicrobial resistance, which has been reported in human-derived enterococci, was not found in these isolates.

The MLST database for E. faecalis and E. faecium as of December 2016 contains 744 STs and 1275 STs from global collections, respectively. Population snapshots of entire STs in the MLST database showed evolutionary distance among ST variants, and E. faecalis represented a more complicated snapshot than E. faecium (Fig. S2 and S3). A collection of 19 E. faecalis isolates yielded 11 novel STs, with the exception of STs. According to the eBURST analysis, ST-436 was clearly derived from the primary founder ST-6, whereas ST-648 may have descended from ST-6. In both species, 15 novel STs were recovered, and they showed an evolutionary distance far from the primary founders, which is defined as the ST that differs from the largest number of other STs at a single locus only (11, 32). Several genetic elements, such as conjugative plasmids and conjugative transposons, are generally known to be involved in genetic transfer.

In conclusion, the distribution of Enterococcus species in the African buffalo was not similar to that previously reported in humans, livestock, and wildlife (3, 31). Our results showed that the most frequently isolated Enterococcus species in the African buffalo in the Serengeti ecosystem was E. casseliflavus, followed by E. mundtii, which is known to be prevalent in plants. The antimicrobial susceptibilities and molecular characteristics of the Enterococcus isolates in the present study differed greatly from those from different sources and countries (2, 10, 12, 20). These differences may have arisen from the location and environment at which samples were acquired. Our results showed no evidence of antimicrobial resistance transfer in the protected area of the Serengeti ecosystem by human intervention. However, increased human activities within the habitats of wild animals, such as wildlife habitat exploitation, trophy hunting, tourism, livestock keeping, and agriculture, may increase the risk of the transmission of antimicrobial-resistant species and resistance genes. This study is the first to report Enterococcus species in the wild African buffalo in Tanzania with the application of an MLST analysis and antimicrobial resistance assays.

Acknowledgements

The authors would like to thank all staff at the Tanzania Wildlife Research Institute (TAWIRI). We particularly thank Dr. Hansol Park for sampling and sample transport. This work was supported by the National Research Foundation of Korea (NRF) grant funded by the Korea Government (No. 2012K1A3A9A01027507).

References

• 1.   Aarestrup,  F.M., and  H.C.  Wegener. 1999. The effects of antibiotic usage in food animals on the development of antimicrobial resistance of importance for humans in Campylobacter and Escherichia coli. Microb Infect. 1:639-644.
• 2.   Aarestrup,  F.M.,  Y.  Agerso,  P.  Gerner-Smidt,  M.  Madsen, and  L.B.  Jensen. 2000. Comparison of antimicrobial resistance phenotypes and resistance genes in Enterococcus faecalis and Enterococcus faecium from humans in the community, broilers, and pigs in Denmark. Diagn Microbiol Infect Dis. 37:127-137.
• 3.   Aarestrup,  F.M.,  H.  Hasman,  L.B.  Jensen,  M.  Moreno,  I.A.  Herrero,  L.  Domínguez,  M.  Finn, and  A.  Franklin. 2002. Antimicrobial resistance among enterococci from pigs in three European countries. Appl Environ Microbiol. 68:4127-4129.
• 4.   Arsène,  S., and  R.  Leclercq. 2007. Role of a qnr-like gene in the intrinsic resistance of Enterococcus faecalis to fluoroquinolones. Antimicrob Agents Chemother. 51:3254-3258.
• 5.   Chow,  J.W. 2000. Aminoglycoside resistance in enterococci. Clin Infect Dis. 31:586-589.
• 6.  Clinical and Laboratory Standards Institute. 2000. Methods for Dilution Antimicrobial Susceptibility Tests for Bacteria that Grow Aerobically; M07-A5 Approved Standard, 5th ed. CLSI, Wayne, PA.
• 7.  Clinical and Laboratory Standards Institute. 2009. Performance Standards for Antimicrobial Disk and Dilution Susceptibility Tests for Bacteria Isolated from Animals; Approved Standard M31-A3, 3rd ed. CLSI, Wayne, PA.
• 8.  Clinical and Laboratory Standards Institute. 2013. Performance Standards for Antimicrobial Susceptibility Testing, 23rd Informational Supplement; M100-S23. CLSI, Wayne, PA.
• 9.   Collins,  M.D.,  J.A.  Farrow, and  D.  Jones. 1986. Enterococcus mundtii sp. nov. Int J Syst Evol Microbiol. 36:8-12.
• 10.   Ernest,  E.M.,  H.  Haanes,  S.  Bitanyi,  R.D.  Fyumagwa,  P.L.  Msoffe,  G.  Bjørnstad, and  K.H.  Røed. 2012. Influence of habitat fragmentation on the genetic structure of large mammals: evidence for increased structuring of African buffalo (Syncerus caffer) within the Serengeti ecosystem. Conserv Genet. 13:381-391.
• 11.   Feil,  E.J.,  B.C.  Li,  D.M.  Aanensen,  W.P.  Hanage, and  B.G.  Spratt. 2004. eBURST: inferring patterns of evolutionary descent among clusters of related bacterial genotypes from multilocus sequence typing data. J Bacteriol. 186:1518-1530.
• 12.   Frei,  A.,  D.  Goldenberger, and  M.  Teuber. 2001. Antimicrobial susceptibility of intestinal bacteria from Swiss poultry flocks before the ban of antimicrobial growth promoters. Syst Appl Microbiol. 24:116-121.
• 13.   Glenn,  L.M.,  M.D.  Englen,  R.L.  Lindsey,  J.F.  Frank,  J.E.  Turpin,  M.E.  Berrang,  R.J.  Meinersmann,  P.J.  Fedorka-Cray, and  J.G.  Frye. 2012. Analysis of antimicrobial resistance genes detected in multipledrug-resistant Escherichia coli isolates from broiler chicken carcasses. Microb Drug Resist. 18:453-463.
• 14.   Hammerum,  A.M. 2012. Enterococci of animal origin and their significance for public health. Clin Microbiol Infect. 18:619-625.
• 15.   Hollenbeck,  B.L., and  L.B.  Rice. 2012. Intrinsic and acquired resistance mechanisms in enterococcus. Virulence. 3:421-569.
• 16.   Homan,  W.L.,  D.  Tribe,  S.  Poznanski,  M.  Li,  G.  Hogg,  E.  Spalburg,  J.D.A.  Embden, and  R.J.  Willems. 2002. Multilocus sequence typing scheme for Enterococcus faecium. J Clin Microbiol. 40:1963-1971.
• 17.   Hoogkamp-Korstanje,  J.A. 1997. In-vitro activities of ciprofloxacin, levofloxacin, lomefloxacin, ofloxacin, pefloxacin, sparfloxacin and trovafloxacin against gram-positive and gram-negative pathogens from respiratory tract infections. J Antimicrob Chemother. 40:427-431.
• 18.   Kim,  O.S.,  Y.J.  Cho,  K.  Lee,  S.H.  Yoon,  M.  Kim,  H.  Na, and  S.  Won. 2012. Introducing EzTaxon-e: a prokaryotic 16S rRNA gene sequence database with phylotypes that represent uncultured species. Int J Sys Evol Microbiol. 62:716-721.
• 19.   Korten,  V.,  W.M.  Huang, and  B.E.  Murray. 1994. Analysis by PCR and direct DNA sequencing of gyrA mutations associated with fluoroquinolone resistance in Enterococcus faecalis. Antimicrob Agents Chemother. 38:2091-2094.
• 20.   Kwon,  K.H.,  S.Y.  Hwang,  B.Y.  Moon,  Y.K.  Park,  S.  Shin,  C.Y.  Hwang, and  Y.H.  Park. 2012. Occurrence of antimicrobial resistance and virulence genes, and distribution of enterococcal clonal complex 17 from animals and human beings in Korea. J Vet Diag Invest. 24:924-931.
• 21.   Lane,  D.J. 1991. 16S/23S rRNA sequencing, p.115-175. In  E.  Stackebrandt, and  M.  Goodfellow (ed.), Nucleic Acid Techniques in Bacterial Systematics. John Wiley and Sons, Chichester.
• 22.   Lebreton,  F.,  R.J.L.  Willems, and  M.S.  Gilmore. 2014. Enterococcus diversity, origins in nature, and gut colonization, p.1-24. In  M.S.  Gilmore,  D.B.  Clewell,  Y.  Ike, et al (ed.), Enterococci: From Commensals to Leading Causes of Drug Resistant Infection [Internet]. Massachusetts Eye and Ear Infirmary, Boston.
• 23.   McBride,  S.M.,  V.A.  Fischetti,  D.J.  LeBlanc,  R.C.  Moellering, Jr, and  M.S.  Gilmore. 2007. Genetic diversity among Enterococcus faecalis. PLoS One. 2:e582.
• 24.   Moura,  I.,  H.  Radhouani,  C.  Torres,  P.  Poeta, and  G.  Igrejas. 2010. Detection and genetic characterization of vanA—containing Enterococcus strains in healthy Lusitano horses. Equine Vet J. 42:181-183.
• 25.   Murray,  B.E. 1990. The life and times of the Enterococcus. Clin Microbiol Rev. 3:46-65.
• 26.   Pesapane,  R.,  M.  Ponder, and  K.A.  Alexander. 2013. Tracking pathogen transmission at the human–wildlife interface: Banded mongoose and Escherichia coli. Ecohealth. 10:115-128.
• 27.   Radhouani,  H.,  N.  Silva,  P.  Poeta,  C.  Torres,  S.  Correia, and  G.  Igrejas. 2014. Potential impact of antimicrobial resistance in wildlife, environment and human health. Front Microbiol. 5:23.
• 28.   Ruiz-Garbajosa,  P.,  M.J.  Bonten,  D.A.  Robinson,  J.  Top,  S.R.  Nallapareddy,  C.  Torres, and  R.J.L.  Willems. 2006. Multilocus sequence typing scheme for Enterococcus faecalis reveals hospital-adapted genetic complexes in a background of high rates of recombination. J Clin Microbiol. 44:2220-2228.
• 29.   Sava,  I.G.,  E.  Heikens, and  J.  Huebner. 2010. Pathogenesis and immunity in enterococcal infections. Clin Microbiol Infect. 16:533-540.
• 30.   Silva,  N.,  G.  Igrejas,  N.  Figueiredo,  A.  Gonçalves,  H.  Radhouani,  J.  Rodrigues, and  P.  Poeta. 2010. Molecular characterization of antimicrobial resistance in enterococci and Escherichia coli isolates from European wild rabbit (Oryctolagus cuniculus). Sci Total Environ. 408:4871-4876.
• 31.   Silva,  N.,  G.  Igrejas,  A.  Gonçalves, and  P.  Poeta. 2012. Commensal gut bacteria: distribution of Enterococcus species and prevalence of Escherichia coli phylogenetic groups in animals and humans in Portugal. Ann Microbiol. 62:449-459.
• 32.   Spratt,  B.G.,  W.P.  Hanage,  B.  Li,  D.M.  Aanensen, and  E.J.  Feil. 2004. Displaying the relatedness among isolates of bacterial species—the eBURST approach. FEMS Microbiol Lett. 241:129-134.
• 33.   Tankovic,  J.,  F.  Mahjoubi,  P.  Courvalin,  J.  Duval, and  R.  Leclercq. 1996. Development of fluoroquinolone resistance in Enterococcus faecalis and role of mutations in the DNA gyrase gyrA gene. Antimicrob Agents Chemother. 40:2558-2561.