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Multi-site Analysis Reveals Widespread Antibiotic Resistance in the Marine Pathogen Vibrio vulnificus

Article in Microbial Ecology · January 2009

DOI: 10.1007/s00248-008-9413-8 · Source: PubMed

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ORIGINAL ARTICLE

Multi-site Analysis Reveals Widespread Antibiotic Resistance in the Marine Pathogen Vibrio vulnificus

Craig Baker-Austin & J. V. McArthur & Angela H. Lindell & Meredith S. Wright & R. Cary Tuckfield & Jan Gooch & Liza Warner & James Oliver & Ramunas Stepanauskas

Received: 10 December 2007 /Accepted: 21 May 2008 # Springer Science + Business Media, LLC 2008

Abstract Vibrio vulnificus is a serious opportunistic human pathogen commonly found in subtropical coastal waters, and is the leading cause of seafood-borne mortality in the USA. This taxon does not sustain prolonged presence in clinical or agricultural settings, where it would undergo human-induced selection for antibiotic resistance. There- fore, few studies have verified the effectiveness of commonly prescribed antibiotics in V. vulnificus treatment. Here we screened 151 coastal isolates and 10 primary

septicaemia isolates against 26 antimicrobial agents repre- senting diverse modes of action. The frequency of multiple resistances to antibiotics from all sources was unexpectedly high, particularly during summer months, and a substantial proportion of isolates (17.3%) were resistant to eight or more antimicrobial agents. Numerous isolates demonstrated resistance to antibiotics routinely prescribed for V. vulnifi- cus infections, such as doxycycline, tetracycline, amino- glycosides and cephalosporins. These resistances were detected at similar frequencies in virulent and non-virulent strains (PCR-based virulence typing) and were present in septicaemia isolates, underlying the public health implica- tions of our findings. Among environmental isolates, there were no consistent differences in the frequency of resis- tance between pristine and anthropogenically impacted estuaries, suggesting natural rather than human-derived sources of resistance traits. This report is the first to demonstrate prevalent antibiotic resistance in a human pathogen with no clinical reservoirs, implying the impor- tance of environmental studies in understanding the spread, evolution and public health relevance of antibiotic resis- tance factors.

Introduction

Bacteria of the genus Vibrio are commonly found in coastal and estuarine waters. Select strains of V. cholerae, V. parahaemolyticus, V. vulnificus and V. mimicus are consid- ered serious human pathogens, [44]. V. vulnificus causes food-borne diseases and wound infections. It carries the highest fatality rate of any food-borne pathogen in the US, often exceeding 50% [29, 30, 34], and 95% of all deaths resulting from seafood consumption in the US are caused by this bacterium [29]. About 85% of V. vulnificus infections

Microb Ecol DOI 10.1007/s00248-008-9413-8

Electronic supplementary material The online version of this article (doi:10.1007/s00248-008-9413-8) contains supplementary material, which is available to authorized users.

C. Baker-Austin: J. V. McArthur : A. H. Lindell : M. S. Wright Savannah River Ecology Laboratory, Drawer E, Aiken, SC, USA

R. C. Tuckfield Savannah River National Laboratory, Bldg. 773-42A, Aiken, SC, USA

J. Gooch National Oceanographic and Atmospheric Administration, Charleston, SC, USA

L. Warner : J. Oliver Department of Biology, University of North Carolina at Charlotte, Charlotte, NC, USA

R. Stepanauskas Bigelow Laboratory for Ocean Sciences, West Boothbay Harbor, ME, USA

C. Baker-Austin (*) Centre for Environment, Fisheries and Aquaculture Science, Weymouth Laboratory, Weymouth, Dorset DT4 8UB, UK e-mail: [email protected]

occur between May and October, as this pathogen thrives in warm water (>20°C) [34]. With advances in refrigeration, seafood originating from habitats where V. vulnificus are most abundant may be consumed by at-risk individuals almost anywhere [43]. It is believed that a large number of V. vulnificus infections may go unreported (CDC estimates ~50% reported), which could greatly contribute to under- estimates of the morbidity and mortality burden associated with this pathogen.

V. vulnificus infections are characterised by an extremely short time-span between the onset of symptoms and subsequent clinical outcome, and immediate antibiotic therapy for suspected cases is considered critical. If treatment is delayed greater than 72 h, the fatality rate of primary septicaemia-associated V. vulnificus infections is 100% [20]. Interestingly, V. vulnificus is considered to have low levels of antibiotic resistance, and previous studies have shown sensitivity to tetracyclines, aminoglycosides, third- generation cephalosporins, chloramphenicol and newer fluoroquinolones [26, 41, 43]. Paradoxically, the two V. vulnificus strains CMCP6 and YJ016, for which genomes are publicly available, contain enzymatic-modification systems, active drug transporters and permeases suggestive of a genetic basis for antibiotic resistance. The significant morbidity and mortality associated with V. vulnificus infections, combined with the speed with which these infections progress and symptoms develop, underlie the need for a thorough analysis of the antibiotic resistance capabilities of this species.

The occurrence, diversity and public health implications of environmental antibiotic resistance have only recently been appreciated [2, 6, 16, 33, 42]. Considering the proliferation of antibiotic resistance in clinical settings [22], an understanding of the selective pressures maintain- ing environmental reservoirs of resistance may have a direct impact on the treatment of infectious diseases, particularly for pathogens that persist in the environment. Given the considerable spatial and temporal heterogeneity in the distribution of resistance and the complex factors that affect its evolution, dissemination and persistence, antibiotic resistance is being increasingly viewed as an ecological problem [36]. Addressing antibiotic resistance from an environmental standpoint can promote a better understand- ing of the ecology and evolution of antibiotic resistance, and may provide an early detection system for the development of antibiotic resistance mechanisms in clini- cally relevant bacteria [10]. For example, several recent reports have implicated the role of industrial contamination, and in particular metal contamination, as an indirect selective agent for antibiotic resistance [2, 24, 39, 40, 46].

Of the few studies that have addressed antibiotic resistance in this important human pathogen, most have involved only a few V. vulnificus isolates and relatively few

antimicrobials. Morris and Tenney [26] analysed the antibiotic resistance capabilities of 19 V. vulnificus strains against seven antimicrobial agents. Zanetti et al. [48] tested six V. vulnificus isolates taken from Italian coastal waters for sensitivity to 11 antimicrobials, while Ottaviani et al. [31] studied the resistance of eight environmental isolates to 27 antibiotics, including some antibiotics tested in this study. Thus, the aim of our study was to determine the antibiotic resistance capabilities across a much larger library of V. vulnificus strains (151 isolates) derived from three different estuarine environments and to determine the sensitivity of these isolates to a wide range of antimicrobial agents (26 drugs). Two of the sites analysed in this study are extensively contaminated with heavy metals (Shipyard Creek, Charleston, SC, USA and LCP Chemicals, Bruns- wick, GA, USA), whilst the reference site (the ACE Basin, Beaufort, SC, USA) is considered a pristine estuarine habitat. These habitats were chosen to assess the potential contribution of industrial contamination in driving environ- mental antibiotic resistance in this taxon. In addition, a small number of strains isolated from V. vulnificus- associated primary septicaemia clinical cases were also screened. Our results suggest that the diversity and level of antibiotic resistance in V. vulnificus is far greater than previously thought, with potentially significant clinical ramifications.

Materials and Methods

Sample Collection

Sediment and water samples were obtained from an industrially contaminated estuarine site (Shipyard Creek, Charleston, SC, USA), in March, June and October 2005 and an uncontaminated reference site (ACE Basin, SC, USA), in June and October 2005. Sediment and water samples were also obtained from the industrially contam- inated EPA Superfund site (LCP Chemical site, Brunswick, GA, USA) in October 2005. Triplicate 1-L surface water samples were obtained during maximum ebb and flood tides to isolate bacterial strains entering and leaving tidal creeks. During low tide, top 1-cm sediment samples were collected into sterile plastic bags from five equidistant locations along each creek, starting at creek mouths and ending at the upper reach of each creek. Bacterial isolation was performed within 24 h from sample collection, after refrigerated storage in the dark.

Isolation and Genotyping of V. vulnificus Strains

A DNA non-radioactive probe (alkaline phosphatase labelled) targeting the species-specific hemolysin–cytolysin

C. Baker-Austin et al.

(vvh-AP) in Vibrio vulnificus was purchased from DNA Technology A/S (DK-8000 Aarhus C, Denmark) and used for identification and confirmation of V. vulnificus colonies on spread plates.

Sediment and water samples were serially diluted in sterile phosphate buffered saline (PBS) [7.65 g NaCl, 0.724 g anhydrous Na2HPO4 (Sigma), 0.21 g KH2PO4 (Sigma)/L, pH 7.4]. Aliquots of each sample were spread- plated onto two types of media for colony isolation. (a) Vibrio vulnificus agar (VVA) [30 g sodium chloride (Sigma), 10 g cellobiose (Sigma), 20 g peptone (Difco), 0.06 g bromthymol blue (Sigma) and 25 g agar (Difco)/L] and (b) CHROMagar Vibrio (CAV; DRG International, Inc.) 74.7 g/L which contains 15 g/L agar, 8 g/L peptone and yeast extracts, 51.4 g/L salts and 0.3 g/L chromogenic mix [14]. The chromogenic media yields presumptive identifications for specific Vibrio species. Turquoise colo- nies are V. vulnificus/Vibrio cholerae in an approximate 50/ 50 ratio. Final species confirmation by the use of molecular tests utilising V. vulnificus gene-specific markers [35] was subsequently performed.

Typical V. vulnificus colonies on VVA plates and turquoise colonies from CAV plates were transferred with sterile toothpicks into individual wells of a 96-well plate containing 100 μL of tryptic soy broth (TSB) plus 2% extra sodium chloride (total of 2.5% salt) and incubated 16 to 18h at 36°C. Cells were plated to VVA plates [5] and tested with the vvh-AP DNA oligo probe for species confirmation. Colony lift, hybridisation and colorimetric detection proce- dure were performed, essentially as previously described [7, 9, 25, 45].

Sediment samples (3 g each) were weighed into 50-mL sterile tubes and mixed with 27 mL 4° C PBS (to make a 10−1 dilution). A 10−3–10−6 dilution series was set up with the sediment samples, inoculated on VVA plates and cultured overnight at 36°C. A number of V. vulnificus isolation methodologies were simultaneously applied for unambiguous species identification. Firstly, an alkaline phosphatase-labelled DNA probe targeting the species- specific hemolysin–cytolysin (vvh-AP) was used for tenta- tive V. vulnificus identification mentioned above [8, 41]. Isolates were then grown overnight in TSB (amended with 2.5% NaCl) at 36°C. DNA from individual isolates was extracted by boiling (5 min, 95°C) 1 mL of culture followed by centrifugation, using the decanted supernatant as crude template [35]. Colonies that positively hybridised with the vvh-AP DNA were later PCR amplified using the V. vulnificus specific vvhA (hemolysin A) gene with forward primer vvhA-F (CGCCGCTCACTGGGG CAGTGGCTG) and reverse primer vvha-R (CCAGCCGT TAACCGAACCACCCGC) [37]. For hemolysin gene PCR, cells were grown overnight at 22°C in heart infusion (HI) broth (Difco; Detroit, MI, USA) and cell lysates

prepared as follows: 200 μL of the broth culture was centrifuged, resuspended in 200 μL of filtered, autoclaved, deionised water and boiled for 5 min. PCR was conducted using a Genius thermal cycler (Techne; Princeton, NJ, USA). Briefly, cell lysates (5 μL) were added to a master mix consisting of 17.75 μL diethyl pyrocarbonate-treated water, 5 mM (8 μL) dNTPs (Promega; Madison, WI, USA), 20 mM (0.09 μL) of each primer (Bio-Synthesis; Lewis- ville, TX, USA), 5U (0.25 μL) Taq polymerase (Promega), 25 mM (3.2 μL) MgCl2 (Promega) and 10× (4 μL) Mg-free buffer (Promega) for a final reaction volume of 40 μL. The hemolysin gene was amplified using 24bp oligonucleotides that are specific for a 340-bp fragment located within this 1,416-bp gene unique to V. vulnificus. The primers utilised were Vv1 (CGC CGC TCA CTG GGG CAG TGG CTG) and Vv2 (CCA GCC GTT AAC CGA ACC ACC CGC). For all experiments, a negative control containing all PCR reagents and sterile HI broth was employed. The reaction mixture was overlaid with 20 μL sterile mineral oil. For visualisation, gel electrophoresis was performed using a 2% agarose gel (NuSieve 3:1, BioWhittaker Molecular Appli- cations; Rockland, ME, USA) with PCR products that were stained with ethidium bromide (1.25 μg/mL) [32].

Finally, PCR amplification and sequencing of the 16S rRNA gene was performed on a subset of isolates for unambiguous species verification. Positively identified V. vulnificus colonies were coded to conceal their source from the investigators, stored in TSB plus 2.5% salt with 30% glycerol and frozen at −80°C for later use. To putatively ascertain the potential virulence capabilities of V. vulnificus isolates, a previously identified 200 bp randomly amplified polymorphic DNA (PCR) amplicon associated with clinical isolates was chosen for colony PCR screening for E- and C- genotypes [35]. The presence of class 1 integrons was also ascertained by PCR [27, 47]. Ten clinically derived V. vulnificus strains, from primary septicaemia clinical blood isolations and previously described [35], were also analysed to compare the public health implications of antimicrobial resistance in these bacteria.

Antimicrobial Susceptibility Testing

Minimal inhibitory concentrations (MICs) were determined by microdilution and 48-h incubation onto custom dehydrated 96-well MicroScan® panels (Dade Behring, Sacramento, USA) according to the manufacturer’s instructions and using Mueller–Hinton broth amended with 2.5% NaCl [4]. The following antimicrobial agents (concentration ranges in mg/ L) were used in the MicroScan® panels, based on their mode of action, history of use and resistance, and clinical relevance: amikacin, 8–64; amoxicillin, 4–32; ampicillin, 4–32; apramycin, 8–32; azithromycin, 2–8; cefoxitin, 8–32; ceftriaxone, 8–64; cephalexin, 16–128; cephalothin, 16–128;

Antibiotic Resistance in Vibrio vulnificus

chloramphenicol, 8–32; ciprofloxacin, 1–4; erythromycin, 16–128; gentamicin, 2–16; imipenem, 2–16; meropenem, 2–16; moxifloxacin, 0.25–4; nalidixic acid, 4–32; nitrofur- antoin, 16–128; oxytetraycline, 4–32; ofloxacin, 1–8; peni- cillin, 16–128; streptomycin, 16–128; sulfathiazole, 250–500; tetracycline, 4–32; trimethoprim, 2–16; and trimethoprim– sulfamethoxazole, 2/38 and 4/76. Concentration levels within the specified range for each antibiotic were successive doublings from the range minimum to maximum. Resistance to doxycycline (which was not present on the MicroScan® panels) was later determined using both broth microdilution and agar dilution methods as outlined by the Clinical and Laboratory Standards Institute (formerly National Committee for Clinical Laboratory Standards [42]), using the most resistant 25% of the environmentally derived V. vulnificus isolates. ATCC V. vulnificus strain 27562 was used as a con- trol during the screening process. Isolates that could not be revived or grown over the course of the antimicrobial susceptibility testing were omitted from the final analyses. Randomised subsets of coded isolates were also re-screened at a separate laboratory to assess methodological reproducibility (data not shown). Resistance was defined as growth in the presence of the highest concentration of the antibiotic and partial resistance as growth in concentrations of the antibiotic up to, but not exceeding the highest concentrations used.

Statistical Analysis

The antibiotic response data were both left censored and right censored, that is, there are no response measurements below the minimum or above the maximum concentrations, respectively, found on the plates for each antibiotic tested. Left-censored observations were recorded as (MIN − MIN / 2) where MIN was the concentration range minimum per antibiotic. Right censored observations were recorded as (MAX + MAX / 2) where MAX is the concentration range maximum on the plates. There is no empirical justification for replacing a right censored value by a further doubling beyond the MAX concentration, as there is no empirical justification for replacing left censored values by zero. The only certainty for isolates that were inhibited by our lowest concentration or that grew at the highest concentration is that the true response is <MIN or >MAX, respectively. Our choice of left censored replacement values is widely acknowledged [15, 28]. For consistency, we treated right censored values similarly. Two summary response measures were calculated, the total number of antibiotics for which an isolate was resistant to the MIN concentration or higher (NabRes) and the trimmed mean antibiotic concentration producing a resistance effect and per isolate called the average resistance response metric (eCab Re s). The latter differs from the simple average resistance response con- centration among all 26 antibiotics in that the simple

average uses the same divisor (i.e. 26) for the sum (numerator) whether or not any term in the sum is <MIN. Based on the concentration ranges previously established for each antibiotic, eCab Re s is an average containing only those antibiotics with values ≥MIN per isolate. Thus, the number of antibiotics represented in this sum was “trimmed” to a subset of those with evinced resistances. Because the concentration ranges for the different anti- biotics used varied by five orders of magnitude, all measurements among isolates within an antibiotic were first standardised; i.e. converted to Z-scores. Thereafter, all measurements regardless of antibiotic were on a trans- formed and common unitless scale and contributed equally to the average resistance response metric eCab Re s. Simple linear regression was used to model the relationship between NabRes and eCab Re s from which the standard Pearson’s (r) correlation coefficient was obtained. ANOVA methods were applied to these two summary response measures to examine the effects of sampling site (ACE, LCP and Shipyard Creek), month (March, June and October) and sample type (ebb, flood or sediment) and the respective interactions among these three main effects. Since the LCP site was only sampled in October and no V. vulnificus isolates were observed in March with the exception of SYC sediment samples, a summary ANOVA was performed using only two sites (ACE and SYC) and 2 months (June and October). Contingency tables were used to assess the frequency distribution differences between clinical and environmental genotype V. vulnificus isolates among the three sites. All statistical methods were obtained from Steel and Torrie [38] and analyses were performed using the JMP™ 5.12 software.

Results

A total of 151 V. vulnificus isolates were obtained from the three coastal sites (Table 1). Of them, 29 (14.6%) were C- genotype (clinical) and 122 (85.4%) were E-genotype (environmental) according to the virulence typing method of Rosche et al. [35] described above. A greater proportion of C-genotypes were detected in October (26%) than June (9%). There were no significant differences in the frequen-

Table 1 Number of V. vulnificus strains isolated from the various sampling sites and sampling campaigns

Site March June October

ACE Basin, Beaufort, SC 0a 21 27 Shipyard Creek, Charleston, SC 2 37 27 LCP, Brunswick, GA 0a 0a 36

a ACE Basin was not studied in March and the LCP Chemical site was not studied in March or June

C. Baker-Austin et al.

cy of E- and C-genotypes among the three sampling locations and three sample types (ebb, flood and sediment). On average, each isolate was resistant to 5.88 antibiotics, with a unimodal distribution of the number of resistances per isolate (Fig. 1a). No discernible difference was found between the E- and C-genotypes with regard to either the average number of resistances per isolate (NabRes) or the average concentration at which the isolate was resistant (eCab Re s). Resistances to aminoglycosides, cephams, folate pathway inhibitors and penicillins were the most common, varying between 0.6% (cefriaxone) and over 99% (apra- mycin) (Fig. 2). Conversely, less than 1% of the isolates were resistant to chloramphenicol, quinolones, macrolides and carbapenams, except for nalidixic acid (7.3%) and azithrmycin (3.3%). Of the 26 antimicrobial agents tested, total sensitivity was evident for just three agents: imipenem, erythromycin and ciprofloxacin. In an ad hoc study, 38 isolates with the highest NabRes were screened for resistance to the synthetic tetracycline derivative doxycycline, of which one isolate was found resistant (2 mg/L). Of the entire environmental library (151 strains) 68 (45%) were resistant to three or more structural classes of antibiotic (Table S2).

An initial three-way ANOVA showed no statistically significant sample type effect (i.e. no differences between sample types ebb, flood or sediment) for either NabRes or eCab Re s. This main effect was dropped and a subsequent two-way ANOVA demonstrated a statistically significant difference (p < .05) in the antibiotic resistance profiles where both NabRes and eCab Re s were higher among ACE Basin isolates than Shipyard Creek isolates. This analysis also revealed a discernible (p < .05) interaction effect in that the difference between ACE and Shipyard isolates for either NabRes or eCab Re s were larger in June than in October (Fig. 1b, Fig. S1). Since sampling was conducted at all three sites in the month of October, a separate ANOVA was performed for this single month’s data and showed that both NabRes and eCab Re s were significantly higher (p < .05) among LCP isolates than either Shipyard Creek or ACE Basin isolates. There was a statistically significant (p <. 05) linear regression relationship between eCab Re s versus NabRes, and the two response measures were positively correlated (r ¼

ffiffiffiffiffi R2

p ¼ :70) (Fig. 3).

For a comparison with the environmental isolates, the same antimicrobial resistance testing procedures were applied on ten V. vulnificus strains isolated from patients suffering from V. vulnificus-associated septicaemia. These strains demonstrated varying levels of resistance, with NabRes = 6.1 (Table 2). All strains were resistant to amikacin, apramycin, cephalexin and streptomycin; and with the exception of strain H3308, gentamicin. One strain (LSU 1606) demonstrated high-level resistance to 12 antimicrobials, including doxycycline (>100 mg/L).

Elevated heavy metal concentrations were confirmed for the SYC and LCP sampling sites using ICP–MS [17] procedures. Compared to ACE, SYC sediments were enriched in Cu, Zn, Sr, Pb and Cd, while LCP sediments were contaminated with Hg (Table S1).

Discussion

This work represents one of the first large-scale surveys of antibiotic resistance and potential virulence in a Vibrio species and provides further evidence for the presence of multi-antibiotic resistance in environmental bacteria. Only a handful of studies have addressed antibiotic resistance in this important human pathogen [13, 26, 41, 43], and these have used a limited range of antimicrobial agents. We found an unexpectedly high frequency and level of resistance to diverse naturally derived as well as synthetic agents, including those often prescribed to treat V. vulnificus infections (tetracycline, gentamicin, ceftriaxone) [3, 19, 26], and even one of the frontline treatment agents, doxycycline (Figs. 1, 2, Table S2). Eleven structural classes of anti- biotics were used in our screening efforts (aminoglycosides,

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Figure 1 Distribution of the number of resistances per isolate (Nab Re s) in the 151 environmental V. vulnificus isolates divided by genotype (a) and by field sampling campaign (b). Since the LCP Chemical site was not sampled in October, only Shipyard Creek and the ACE Basin isolates were included in the seasonal comparison (b)

Antibiotic Resistance in Vibrio vulnificus

carbapenams, cephams, foliate pathway inhibitors, macro- lides, nitrofurans, penicillins, phenicols, quinolones, sul- phonamides and tetracyclines). Of these, 68 isolates (45%) were resistant to agents of three or more structural classes suggestive of widespread multi-antibiotic resistance. Simi- lar resistance patterns were found in V. vulnificus derived from primary septicaemia blood isolates (Table 2). One of the ten analysed clinical isolates (LSU 1606) was resistant to 12 out of 26 antimicrobials. Strikingly, only one V. vulnificus isolate from the 151 environmental and ten clinical strains was susceptible to all 26 antimicrobials (Table S2). These resistances may hinder the treatment of V. vulnificus infections, and could contribute directly to the high rates of mortality associated with these pathogens.

Interestingly, V. vulnificus antibiotic resistance and genotype composition exhibited a marked temporal varia- tion, with higher NabRes or eCab Re s and higher proportion of E-genotypes in June compared to October (Fig. 1b).

Figure 3 Average resistance response (eCab Re s) as a function of total number of antibiotic resistances per isolate (Nab Re s). Shipyard Creek (SYC), LCP Chemicals (LCP) and the ACE Basin (ACE)

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Figure 2 Frequency of resistance to 26 antibiotics among the 151 environmental V. vulnificus isolates. Resistance was defined as growth in the presence of the highest concentration of the antibiotic and partial resistance as growth in concentrations of the antibiotic up to, but not exceeding, the highest concentrations used. Data for

doxycycline resistance is not shown. Abbreviations used: Amino, aminoglycosides; Pen, penicillins; Carb, carbapenams; Ceph, ceph- ams; FPI, folate pathway inhibitors; Sul, sulfathiazole; Nit, nitro- furantion; Quin, quinolones; Mac, macrolides; Cl, chlorampenicol

C. Baker-Austin et al.

Notably, the highest clinical burden associated with V. vulnificus infections usually takes place during the summer [26]. More extensive studies, encompassing multiple years and a broader geographic scope, should be conducted to assess potential epidemiological implications of such potential seasonal trends.

There were no consistent differences in antibiotic resistance (NabRes and eCab Re s) between V. vulnificus isolated from pristine (ACE Basin) and contaminated (Shipyard Creek and LCP Chemical) sites. The apparent lack of contamination effects on resistance patterns was further supported by the absence of resistance differences among flood, ebb and sediment samples in Shipyard Creek and LCP Chemical sites. This contradicts several prior studies, demonstrating elevated antibiotic resistance in heavy metal- contaminated environments, likely due to resistances to diverse metals and antibiotics being co-selected for [2, 24, 39, 40, 46]. The apparent lack of indirect selection may be due to weak or non-existent genetic linkages between metal and antibiotic resistance in the studied V. vulnificus. Alternatively, although the total concentration of several toxic metals was high in the sediments of Shipyard Creek (Cr and Cd) and the LCP Chemical site (Hg) (Table S1), the bioavailability of these toxic metals may be insufficient to impose significant indirect selection. Some heavy metals are capable of forming stable complexes in seawater [11, 12] reducing bioavailability and thus the potential for

anthropogenic metal emissions to have a significant impact on antibiotic resistance.

Most antibiotics, including many that are clinically relevant, are naturally produced by microorganisms in the environment [5, 18, 36], likely explaining the presence of antibiotic resistance genes and resistant microorganisms in pristine habitats (i.e. the ACE Basin) and in samples that predate the human use of antibiotics [36]. Interestingly, we found a high frequency of resistances to aminoglycosides, cephams and penicillins (Fig. 2), which are often encoded on plasmids in other gram-negative bacteria, including other Vibrio species [44]. However, no class 1 integrons, genetic elements frequently associated with the dissemina- tion of antibiotic resistance genes in clinical isolates, were detected in any of the 151 isolates. Conversely, we did find a positive correlation between plasmid DNA content and antibiotic resistance (data not shown), suggesting that horizontal gene transfer may be partly responsible for the observed resistances, involving mechanisms different from class 1 integron exchange. Furthermore, Vibrio species are known to harbour chromosomal integrons into which mobile gene cassettes potentially conferring antibiotic resistance can be inserted which constitutes an additional potential mechanism contributing to antibiotic resistance in V. vulnificus [23]. It should be emphasised that V. vulnificus does not sustain prolonged presence in clinical settings, where it would undergo human-induced selection for antibiotic resistance, as in the well-known cases of obligate human and agricultural pathogens [21]. Thus, high antimi- crobial resistance found in this study may either reflect innate resistance levels and/or horizontal transfer of resistance factors from anthropogenically derived taxa. Further work is clearly required to determine the precise molecular mechanisms of antibiotic resistance in this taxon.

Of particular interest is the significant positive correlation between total number of antibiotic resistances (NabRes) and average resistance response (eCab Re s) (Fig. 3). It suggests that as the number of antibiotics to which an isolate is resistant increases, the concentration of the corresponding antibiotics that can be tolerated by the isolate increases as well. This seems to contradict the common assumption that the frequency and rates of antibiotic resistance in bacterial populations are directly related to the antibiotic exposure and inversely related to the cost that resistance imposes on the fitness of bacteria [1]. The positive relationship between NabRes and eCab Re s implies low fitness costs of the maintenance of antibiotic resistance genes in V. vulnificus and/or synergistic effects among the various antibiotic resistance factors. Likewise, a recent study indicates low or non-existent fitness impact imposed on an environmentally derived E. coli strain via carriage of several mobile antibiotic resistance elements [8]. Low

Table 2 Antimicrobial resistance information of the clinical and type strains

Strain Source Genotype Resistance profilea

C7184K2 Clinical C Ak, Apr, Cex, Gm, St LSU 763 Clinical C Ak, Apr, Cex, Gm, St LSU 1009 Clinical C Ak, Apr, Cex, Gm, St LSU 1365 Clinical C Ak, Apr, Cex, Gm, St LSU 1003 Clinical C Ak, Apr, Cex, Gm, St H3308 Clinical C Ak, Apr, Cex, NA, St CMCP6 Clinical C Ak, Apr, Cex, Cfx, Gm, St LSU 1015 Clinical C Ak, Apr, Cex, Gm, St, T LSU 1456 Clinical C Ak, Apr, Cex, Cfx, Gm, NA,

St LSU 1606 Clinical C Ak, Apr, Azi, Cex, E, Fd,

Gm, Otet, St, Sz, TE, Doxy V. vulnificus Type C Ak, Apr, Cex, Cfx, Gm, St ATCC 27562 strain

a Abbreviations used: Ak, amikacin; Am, ampicillin; Amx, amoxicillin; Apr, ampramycin; Azi, azithromycin; C, chloramphenicol; Cax, ceftriaxone; Cex, cephalexin; Cf, cephalothin; Cfx, cefoxitin; Deoxy, doxycycline; Fd, nitrofurantoin; Gm, gentamicin; Mer, meropenem; Mox, moxifloxacin; NA, nalidixic acid; Otet, oxytetracycline; P, penicillin; St, streptomycin; Sz, sulfathiazole, T, trimethoprim; TE, tetracycline

Antibiotic Resistance in Vibrio vulnificus

fitness cost of antibiotic resistance genes may explain slow recovery of antimicrobial sensitivity in obligate pathogens after the discontinuation of antimicrobials [1].

To our knowledge this report constitutes the first to demonstrate prevalent antibiotic resistance in a human pathogen with no clinical reservoirs, implying the impor- tance of environmental studies in understanding the spread, evolution and public health relevance of antibiotic resis- tance factors.

Acknowledgements We thank Blaine West, Charles Zemp, Kirk Kessler, Richard Gregory, and Marc Frischer for help accessing field sample collection sites. Brian Thompson and Brian Robinson are acknowledged for their assistance in isolate preparation. This work was supported by the National Oceanographic and Atmospheric Administration (NOAA) awards NA04OAR4600198 to Stepanauskas and NA05NOS4781244 to Oliver. Additional support was provided from the US Department of Energy Financial Assistance Award DE- FC09-96SR18546 to the University of Georgia Research Foundation.

Disclaimer This report was prepared as an account of work sponsored by an agency of the United States Government. Neither the United States Government nor any agency thereof, nor any of their employees, makes any warranty, express or implied, or assumes any legal liability or responsibility for the accuracy, completeness, or usefulness of any information, apparatus, product, or process disclosed, or represents that its use would not infringe privately owned rights. Reference herein to any specific commercial product, process, or service by trade name, trademark, manufacturer, or otherwise does not necessarily constitute or imply its endorsement, recommendation, or favouring by the United States Government or any agency thereof. The views and opinions of authors expressed herein do not necessarily state or reflect those of the United States Government or any agency thereof.

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Antibiotic Resistance in Vibrio vulnificus