Antibiotic-Resistant Enterococci in Seawater and Sediments from a Coastal Fish Farm

Abstract

The aim of this study was to detect and characterize antibiotic-resistant enterococci in seawater and sediment from a Mediterranean aquaculture site where no antibiotics are used. Colonies (650) grown on Slanetz-Bartley (SB) agar were amplified on antibiotic-supplemented SB, and erythromycin (ERY), tetracycline (TET), and ampicillin (AMP) MICs were determined. Of 75 resistant isolates (17 to TET, 5 to ERY, and 45 to AMP), 5 Enterococcus faecalis, 25 E. faecium, 5 E. casseliflavus, 1 E. gallinarum, 1 E. durans, and 23 Enterococcus spp. were identified by genus- and species-specific polymerase chain reaction (PCR). tet(M), tet(O), tet(L), tet(K), erm(B), erm(A), erm(C), mef, msr, blaZ, and int(Tn916) were sought by PCR, including an improved multiplex PCR assay targeting tet(M), tet(L), and erm(B). Tet(M) was the most frequent TET resistance gene; msr(C) was the sole ERY resistance gene detected. blaZ was found in 29/45 AMP-resistant isolates; however, no β-lactamase production was detected. Antibiotic-resistant enterococci were recovered 2 km off the coast despite the absence of selective pressure exerted by antibiotic use. The occurrence of resistant strains in the absence of the tested genes may indicate the presence of less common resistance determinants. This first evidence of resistant enterococci at a Mediterranean aquaculture site suggests the existence of a marine reservoir of antibiotic resistances potentially transmissible to virulent strains that could be affected by mariculture in an antibiotic-independent manner.

Introduction

Enterococci are among the most important agents of nosocomial infections, and increased mortality from enterococcal infections has been described over the past few years.^10,40,43 Enterococcus faecalis and Enterococcus faecium are the species most commonly involved; an increase in the number of infections caused by E. faecium is partly due to its proneness to acquire resistance traits,⁴³ also resulting in the emergence of difficult to eradicate multidrug-resistant (MDR) strains.^5,40 The widespread release of antibiotics in the environment due to their use in human and animal medicine, agriculture, and industrial activities has led to increased selection for and spread of resistant enterococci in different compartments, such as water, soil, food, and wastewater.^9,27,32 On the other hand, the vast majority of antibiotic resistance genes have been found to have an environmental origin.^4,31 For example, more than half of clinical enterococcal isolates are resistant to tetracycline (TET)²⁴ even though TET is not routinely used to treat enterococcal infections, indicating selection in a different environment.²⁷

In enterococci, TET resistance is prevalently mediated by tet(M)—a ribosomal protection gene, mostly carried by Tn916-like elements—and to a lesser extent by tet(O), or by tet(L) and tet(K), which code for the active efflux of the drug across the cell membrane²⁴; macrolide resistance is generally mediated by erm genes, principally erm(B) and less frequently erm(A).²⁴ The efflux gene mef, which codes for the transport of macrolides out of the cell, and msr, which causes resistance to macrolides through the ATP-binding transporter protein, are also involved, although less frequently. Ampicillin (AMP) resistance is caused by overproduction or mutation of penicillin binding protein (PBP) 5, mutation of PBP 4, or by blaZ- encoded β-lactamase production.^5,24

Enterococci are not indigenous to water and are among fecal indicator bacteria (FIB)²⁸; both gastrointestinal and respiratory diseases have been associated with exposure to contaminated water or sediments.¹⁹ Since seawater and sediment are possible vehicles of virulent strains transmissible to humans, it is useful to gain insights into the presence and distribution of enterococci and into their antibiotic resistance mechanisms in marine ecosystems, particularly in sediments, which are a source of organic nutrients and exert a protective effect on microorganisms.^28,37

Aquaculture is spreading throughout the world¹⁶ and is believed to have contributed to the maintenance and spread of antibiotic resistance genes in marine environments through the extensive decade-long use of antibiotics as growth promoters and prophylactic treatments.⁹ It may thus be hypothesized that enterococci in marine environments impacted by mariculture sites may undergo selective pressure, which coupled to their ability to persist and grow in different environmental conditions may have contributed to the evolution and spread of resistant strains.

Several studies have addressed antibiotic resistance in FIB, also in proximity of aquaculture sites^2,3,11; however, little research has been conducted on the genetic determinants carried by enterococci.^36,39 In this study, to investigate the role of aquaculture in the emergence and selection of antibiotic-resistant bacterial pathogens of fecal origin, enterococci were isolated from seawater and sediment collected at an aquaculture site and analyzed for their resistance to TET, erythromycin (ERY), and AMP. Resistant isolates were further investigated for species determination and detection of resistance and mobile-element-associated genes.

Materials and Methods

Site and sampling strategy

Seawater and sediment were collected in June 2010 at a fish farm lying ca. 2 km off the coast of Manfredonia, Italy (southern Adriatic Sea, Lat. 41°37'16''N, Lon. 15°56'54''E), at a depth of 11 m. The farm has 35 circular cages holding predominantly sea bass and sea bream. At the time of sampling, the owner reported earlier, sporadic use of EU-approved antibiotics to treat diseases, but none over the last two years. Seawater around the cages was collected from a small boat using sterile Niskin bottles (capacity 5 L) at a depth of ca. 3 m and immediately placed in 2 L sterile polypropylene bottles. Sediment directly under the cages was collected in triplicate (S-1, S-2, and S-3) in the central area of the fish farm by scuba divers using sterile plexiglass corers (internal diameter 4.5 cm). The top 2 cm of each sample was immediately and carefully extruded; the rest was placed in sterile containers for transport to the laboratory. Both types of samples were kept at in situ temperature in the dark until processing a few hours later.

Enterococcus spp. isolation and identification

Seawater samples were prefiltered through a 30-μm membrane to remove large particles and metazoa. Aliquots of 250 ml, 100 ml, 10 ml, and 1 ml were then filtered through 0.45-μm filters (Millipore, Billerica, MA), which were placed on Slanetz-Bartley plates (SB; Oxoid, Basingstoke, United Kingdom) and incubated for 48 h at 37°C. Sediment samples were processed as follows: 20 g or 50 g of sediment was suspended in 200 ml (10% w/v) or 100 ml (50% w/v) of saline solution, respectively, and then vigorously shaken and sonicated to detach bacteria from the sediment matrix, as previously described.²⁸ Resuspensions were then prefiltered through a 30-μm membrane; finally aliquots of 10 ml (10% suspension) or 5 ml (50% suspension) from both sample types were filtered, plated, and incubated as described for seawater.

Grown colonies were counted, amplified on SB plates, and incubated for 48 h at 42°C. A loop of each grown culture was streaked on 8 SB plates, each supplemented with a different antibiotic: TET, 10 μg/ml; ERY, 20 μg/ml; AMP, 20 μg/ml; linezolid (LIN), 10 μg/ml; levofloxacin (LEVO), 20 μg/ml; chloramphenicol (CHLOR), 40 μg/ml; gentamicin (GENTA), 250 μg/ml; and streptomycin (STREP), 1000 μg/ml. Plates were incubated for 48 h at 42°C.

Enterococci were identified to the genus level by amplification of the conserved target sequence shared by known Enterococcus spp., located on 16S rRNA.¹³ E. faecalis and E. faecium were identified by multiplex polymerase chain reaction (PCR) targeting the ddl E. faecalis and ddl E. faecium genes as described by Dutka-Malen et al.¹⁷; E. casseliflavus and E. gallinarum were identified by PCR assays targeting the van(C1) and van(C2) genes and E. durans and E. hirae by targeting the ddl genes, as described previously.⁷

Susceptibility testing

Susceptibility testing was performed by MIC determination according to CLSI guidelines¹² and the results were interpreted according to CLSI 2010, M100-S20. β-Lactamase production was investigated using the nitrocefin-based test.¹² TET, ERY, AMP, GENTA, STREP, and CHLOR were purchased from Sigma-Aldrich (Saint Louis, MO); LIN from Pfizer Italia (Rome, Italy); LEVO from GlaxoSmithKline (Verona, Italy); and nitrocefin from Oxoid.

PCR experiments to detect resistance genes and sequencing assays

All PCR assays were performed in a final volume of 50 μl containing 5 μl of bacterial DNA extracted as described by Hynes et al.,²¹ using a final extension step at 72°C for 7 min in a T Personal thermocycler (Biometra, Goettingen, Germany). The assays performed to seek tet(L) in reference strains used 0.5 μmol/L of each primer, 2 mmol/L MgCl₂, 200 μmol/L dNTPs, and 1 U Taq DNA polymerase (DyNAzyme^™II; Finnzymes, Keilaranta, Finland). The PCR cycling program was as follows: 95°C for 4 min, followed by 30 cycles at 94°C for 1 min, 53°C for 1 min, and 72°C for 1 min. Detection of tet(O), tet(K), erm(A), erm(C), and blaZ was as described by Garofalo et al.¹⁸ The presence of mef and msr was investigated using 0.5 μmol/L of each or mef or msr primer, 2 or 2.5 mmol/L MgCl₂ (for amplification of mef or msr, respectively), 200 μmol/L dNTPs, and 1.5 U hot-start Taq DNA polymerase (AmpliTaq Gold; Applied Biosystems, Foster City, CA). The PCR program involved denaturation at 95°C for 10 min followed by 35 cycles at 94°C for 1 min, 54°C (mef) or 52°C (msr) for 1 min, and 72°C for 1 min.

Detection of msr(C) was confirmed by sequencing and BLAST analysis of the PCR product obtained using the primers targeting the msr gene.

Multiple PCR experiments targeting tet(M), tet(L), and erm(B) were performed using 1.2 μmol/L of each tet(M) primer and 0.5 μmol/L of each tet(L) and erm(B) primers, 6.0 mmol/L MgCl₂, 600 μmol/L dNTPs, and 1.25 U of hot-start Taq DNA polymerase (AmpliTaq Gold; Applied Biosystems, Foster City, CA). Cycling parameters were as follows: initial denaturation at 94°C for 4 min followed by 35 cycles at 94°C for 30 s, 53°C for 30 s, and 72°C for 1 min and 30 s. The presence of the integrase gene of Tn916 [int(Tn916)] was investigated as described by Doherty et al.¹⁵

Amplicons to be sequenced were purified using the GenEluTc^™ PCR CleanUp kit (Sigma-Aldrich) and then sequenced bidirectionally using the BMR Genomics service (www.bmr-genomics.it/).

The control strains and primers used are listed in Tables 1 and 2, respectively.

Table 1.

Control Strains

Bacterial strain	Resistance gene(s)	Reference or source
Enterococcus faecium CM 4.2E	tet(M), tet(L)^a, erm(B), msr(C)^a	44, this study
Enterococcus faecalis PM 2.2T	tet(M), tet(L)^a	44, this study
E. faecium CF 2.1E	tet(M), tet(L)^a, erm(B),	44, this study
E. faecium ET 18	tet(K)	Departmental collection^b
Streptococcus pneumoniae Pn8	int(Tn916)	33
Staphylococcus aureus MU 50	erm(A)	ATCC^c
Staphylococcus aureus 29213	blaZ	ATCC^c
Staphylococcus epidermidis SE12	erm(C)	18
Streptococcus pyogenes 7008	tet(O), mef	8

Detected in this study.

From the collection of the Department of Life and Environmental Sciences—Polytechnic University of Marche, Ancona (Italy).

American Type Culture Collection.

Table 2.

Polymerase Chain Reaction Primers and Target Genes

Target gene	Primer sequence (5'?3')	Product size (bp)	Reference
tet(M)	1-GTTAAATAGTGTTCTTGGAG	657	1
	2-CTAAGATATGGCTCTAACAA
tet(L)	1-CATTTGGTCTTATTGGATCG	475	1
	2-ATTACACTTCCGATTTCGG
tet(O)	1-AACTTAGGCATTCTGGCTCAC	519	18
	2-TCCCACTGTTCCATATCGTCA
tet(K)	1-TCGATAGGAACAGCAGTA	169	35
	2-CAGCAGATCCTACTCCTT
int(Tn916)	1-GCGTGATTGTATCTCACT	1046	15
	2-GACGCTCCTGTTGCTTCT
erm(B)	1-CATTTAACGACGAAACTGGC	425	23
	2-GGAACATCTGTGGTATGGCG
erm(A)	1-TCTAAAAAGCATGTAAAAGAA	645	41
	2-CTTCGATAGTTTATTAATATTAGT
erm(C)	1-TCAAAACATAATATAGATAAA	642	41
	2-GCTAATATTGTTTAAATCGTC
mef	1-AGTATCATTAATCACTAGTGC	348	41
	2-TTCTTCTGGTACTAAAAGTGG
msz	1-GCAAATGGTGTAGGTAAGACAACT	405	41
	2-ATCATGTGATGTAAACAAAAT
blaZ	1-ACTTCAACACCTGCTGCTTTC	240	30/18
	2-TAGGTTCAGATTGGCCCTTAG
16S rRNA	1-TCAACCGGGGAGGGT	733	13
	2-ATTACTAGCGATTCCGG
ddl E. faecalis	1-ATCAAGTACAGTTAGTCT	941	17
	2-ACGATTCAAAGCTAACTG
ddl E. faecium	1-TAGAGACATTGAATATGCC	550	17
	2-CTAACATCGTGTAAGCT
vanC1	1-GGTATCAAGGAAACCTC	822	7
	2-CTTCCGCCATCATAGCT
vanC2	1-CTCCTACGATTCTCTTG	439	7
	2-CGAGCAAGACCTTTAAG
ddl E. durans	1-TTATGTCCCAGTATTGAAAAA	189	7
	2-TGAATCATATTGGTATGCAGT
ddl E. hirae	1-TTATGTCCCTGTTTTGAAAAA	378	7
	2-TTTTGATAGACCTCTTCCGGT

Data analysis

The distribution of resistance strains was calculated as the ratio of isolates resistant to each antibiotic to the total of resistant enterococci, and the result was multiplied by 100. The percent frequency of resistance genes was calculated as the ratio of strains carrying the gene of interest to the total of enterococci resistant to the specifc antibiotic, and the result was multiplied by 100. The nucleotide sequences were analyzed by BIOEDIT and BLAST softwares.

Results

Recovery of antibiotic-resistant enterococci from seawater and sediment

A total of 650 putative enterococci, 150 from seawater and 500 from sediment, were counted on filters placed on SB agar at 37°C. Of these, 111 isolates from seawater (21 on TET, 13 on ERY, and 77 on AMP) and 45 from sediment (9 on TET, 3 on ERY, and 33 on AMP) were able to grow on antibiotic-containing SB plates. No growth was observed in plates supplemented with LIN, LEVO, CHLOR, GENTA, or STREP. To verify the resistance level of the isolates grown in antibiotic-supplemented SB, the MIC of the antibiotic used for the selection was determined. About 50% of isolates (75/156, 51 from water and 24 from sediment) showed MICs exceeding the resistance breakpoint.

Identification of resistant isolates

Of the 75 resistant strains 60 were confirmed to belong to the genus Enterococcus. Of these, 5 strains were identified as E. faecalis, 25 as E. faecium, 5 as E. casseliflavus, 1 as E. gallinarum, and 1 as E. durans. There were no E. hirae strains. The remaining 23 resistant isolates were not further analyzed and were generically reported as belonging to Enterococcus spp. To investigate the presence of MDR strains, the 60 Enterococcus isolates were further analyzed for their susceptibility to the two antibiotics not used for selection (TET, ERY, or AMP). Four strains were found to be resistant to ERY and AMP and 3 to TET and AMP (Fig. 1 and Table 3).

FIG. 1.

Distribution of tetracycline, erythromycin, and ampicillin resistance among the 60 antibiotic-resistant enterococci isolated from water (a) and sediment (b). ▮ Total enterococci, □ Enterococcus faecalis, Enterococcus faecium, other Enterococcus species; TET, tetracycline-resistant enterococci; ERY, erythromycin-resistant enterococci; AMP, ampicillin-resistant enterococci; TET-AMP, tetracycline- and ampicillin-resistant enterococci; TET-ERY, tetracycline- and erythromycin-resistant enterococci; ERY-AMP, erythromycin- and ampicillin-resistant enterococci.

Table 3.

Phenotypic and Genotypic Traits of Tetracycline-, Erythromycin-, and Ampicillin-Resistant Enterococci Recovered from Seawater (A) and Sediment (S)

Strain	Phenotype (MIC μg/ml)	Genotype
E. faecium A-1.1.1	AMP 16	–
E. casseliflavus A-1.1.20	TET 32	tet(L)
E. casseliflavus A-1.1.24	TET 16	tet(M)
E. casseliflavus A-1.1.26	TET 32	–
E. faecium A-1.1.55	TET 64	tet(M), tet(K)
E. faecium A-1.2.1	AMP 32	blaZ
Enterococcus spp. A-1.2.2	AMP 32	–
Enterococcus spp. A-1.2.4	AMP 64	blaZ
E. faecalis A-1.2.6	AMP 32	blaZ
E. faecalis A-1.2.7	AMP 32	–
Enterococcus spp. A-1.2.11	AMP 16	blaZ
E. faecium A-1.2.14	AMP 16	blaZ
Enterococcus spp. A-1.2.16	AMP 32	blaZ
E. faecium A-1.2.17	ERY 32, AMP 64	msr(C), blaZ
E. faecium A-1.2.19	AMP 32	blaZ
E. casseliflavus A-1.2.24	TET 16, AMP 16	tet(M), blaZ
Enterococcus spp. A-1.2.29	AMP 16	–
E. faecium A-1.2.33	AMP 16	–
Enterococcus spp. A-1.2.34	AMP 16	–
Enterococcus spp. A-1.2.36	AMP 32	blaZ
Enterococcus spp. A-1.2.37	AMP 64	blaZ
E. faecium A-1.2.39	AMP 16	–
Enterococcus spp. A-1.2.40	AMP 16	blaZ
Enterococcus spp. A-1.2.46	AMP 16	blaZ
E. faecium A-1.2.47	AMP 16	blaZ
E. faecium A-1.2.50	AMP 16	blaZ
Enterococcus spp. A-1.2.51	AMP 16	blaZ
Enterococcus spp. A-1.2.52	AMP 16	–
Enterococcus spp. A-1.2.64	AMP 16	–
Enterococcus spp. A-2.2.3	AMP 16	–
E. faecium A-2.2.8	AMP 32	blaZ
E. faecium A-2.2.14	ERY 128, AMP 16	msr(C), blaZ
E. faecalis A-2.2.16	TET 32, AMP 16,	tet(M), tet(K), blaZ
E. faecium A-3.1.1	TET 64	–
Enterococcus spp. A-3.1.1b	AMP 16	–
E. faecium A-3.1.5	TET 16	–
E. faecium A-3.1.16	TET 32	tet(K)
E. faecium A-3.1.24	TET 32	tet(M)
E. faecium A-3.2.5	AMP 16	–
Enterococcus spp. A-3.2.6	AMP 16	–
E. faecium A-3.2.8	ERY 8, AMP 16	msr(C), blaZ
E. faecium A-3.2.10	ERY 32, AMP 16	blaZ
Enterococcus spp. A-3.2.11	TET 256	tet(L), tet(K)
Enterococcus spp. A-3.2.13	AMP 128	blaZ
E. durans A-3.2.22	AMP 16	blaZ
E. faecium A-3.2.24	TET 64, AMP 256	tet(M), blaZ
E. faecalis S-1.1.2	AMP 16	blaZ
Enterococcus spp. S-1.1.10	AMP 16	–
Enterococcus spp. S-1.1.13	AMP 32	blaZ
Enterococcus spp. S-1.1.15	AMP 16	blaZ
Enterococcus spp. S-1.1.30	TET 32	tet(M), tet(L)
E. faecium S-1.1.31	TET 64	–
E. faecium S-1.2.5	ERY 8	–
Enterococcus spp. S-1.2.6	AMP 16	–
E. casseliflavus S-1.2.13	AMP 256	blaZ
E. faecium S-2.1.9	TET 128	tet(M), tet(L)
E. faecium S-2.1.12	TET 32	–
E. faecalis S-3.1.2	AMP 16	blaZ
E. faecium S-3.1.7	AMP 16	–
E. gallinarum S-3.1.40	TET 32	tet(M)

–, no resistance gene found.

TET, tetracycline; ERY, erythromycin; AMP, ampicillin.

Detection of resistance genes

The 60 resistant enterococcal isolates were then screened for the presence of the TET and ERY resistance genes most frequently reported in enterococci, that is, tet(M), tet(O), tet(L), and tet(K), and erm(B), erm(A), erm(C), mef, and msr(C), respectively. The presence of the β-lactamase-encoding blaZ gene was also investigated. A multiplex PCR protocol to detect tet(M), tet(L), and erm(B) was developed and validated using the control strains described previously. The other resistance genes, tet(K), tet(O), erm(A), erm(C) mef, msr, and blaZ, were sought by single PCR assays. Among TET-resistant enterococci (n=17), tet(M) was harbored by 5 strains, tet(O) by no strain, tet(L) by 1, tet(K) by 1, tet(M) and tet(L) by 2, tet(M) and tet(K) by 2, and tet(L) and tet(K) by 1 strain, whereas 5 strains harbored none of the resistance genes investigated. The nine tet(M)-positive strains were further investigated for the presence of Tn916. Four [3 carrying only tet(M) and 1 carrying both tet(M) and tet(K)] were amplified with int(Tn916) specific primers. As regards ERY-resistant isolates (n=5), 3 strains carried msr(C), while erm(B), erm(A), erm(C), and mef were never detected. Among AMP-resistant enterococci (n=45) blaZ was detected in 29 strains; however, β-lactamase production was demonstrated in none of them. Ten strains were found to carry more than one resistance gene (Table 3). The results are summarized in Fig. 2 and Table 3.

FIG. 2.

Frequency (%) of resistance genes detected in enterococci isolated from water (a) and sediment (b). ▮ Total enterococci, □ E. faecalis, Enterococcus faecium E. faecium, other Enterococcus species.

Discussion

Several investigations have focused on isolates from farmed animals^7,9,20,44,45 and from the environment, including seawater and sediment,^26,32,34 to achieve a greater understanding of the origin, spread, and persistence of antibiotic-resistant enterococci. However, not many data are available on antibiotic-resistant enterococci from water and sediment near coastal mariculture sites³⁶ and, to our knowledge, this is the first study carried out in the Adriatic Sea. Sixty enterococcal strains resistant to TET, ERY, and/or AMP were recovered from samples collected at a fish farm lying 2 km from the coast. Resistance to AMP (ca. 75%) was the most prevalent, followed by TET resistance (ca. 28%); ERY resistance was seen in less than 10% of isolates. In a similar study of water, sediment, and fish intestine collected in fish farms in Thailand, Petersen and Dalsgaard found a prevalence of ERY compared with TET resistance.³⁶ Barros et al. also reported a greater frequency of ERY than TET resistance in enterococci from fecal samples of Gilthead sea bream captured in the western Atlantic Ocean.⁶ The same authors described a lower frequency of resistance to AMP than to TET, as also reported by Kimiran-Erdem et al. in isolates from coastal seawater near Istanbul.²⁶

TET-, ERY-, and AMP-resistant isolates were more common among E. faecium than among E. faecalis strains, as previously noted by Petersen and Dalsgaard,³⁶ in line with the propensity of the former to accumulate resistance traits. Our molecular experiments indicated that tet(M) was the most frequently detected TET resistance gene. Similar findings have often been reported in enterococci from terrestrial farm animals^14,22,44 and in isolates from aquaculture sites.^25,42 However, there are no recent data regarding enterococci from mariculture.

The combinations tet(M)–tet(L), tet(M)–tet(K), and tet(L)–tet(K) were found in a small number of strains, but did not result in greater MICs compared with strains carrying a single resistance gene. In enterococci, tet(M) is usually carried by Tn916; however, only 50% of our tet(M)-positive strains were amplified with primers specific for int(Tn916), suggesting the common involvement of other elements in marine isolates.

ERY resistance was rare and, unexpectedly, no erm gene was found. The only ERY resistance gene detected was msr(C), identified in 3 isolates of E. faecium with ERY MICs of 8, 32, and 128 μg/ml, respectively. However, msr(C) does not explain the high MICs of E. faecium A-2.2.14 (128 μg/ml), since it does not cause high resistance levels in enterococci.³⁸ These findings agree with its although nonuniform association with the species E. faecium and its possible involvement in mechanisms other than antibiotic resistance.⁴⁶

Petersen and Dalsgaard described erm(B) in 87% of ERY-resistant enterococcal isolates from water and sediment, chicken manure, and fish intestine samples.³⁶ The discrepancy with our findings may be explained by the different types of reared stock and possibly the different effect of human activities and pollution on the areas sampled in the two studies.

blaZ was detected in more than half of AMP-resistant isolates; however, no strain produced β-lactamase, in line with its scarce production by enterococci. On the other hand high-level AMP resistance in enterococci is prevalent due to either overproduction of PBP5 or its mutation, mainly in species other than E. faecalis.²⁴

No resistance gene was detected in one-third (22/60) of resistant isolates, as also reported by other researchers.^29,44 The presence of other, different determinants seems likely. Overall these results highlight the presence of a marine reservoir of antibiotic-resistant enterococci showing resistance traits only partially overlapping with those typical of clinical strains and characterized by an important contribution from species rarely recovered from humans. Its role in the increase and diffusion of enterococcal antibiotic resistance warrants further research.

Resistant enterococci were recovered from both water and sediment; however, whereas the isolates recovered from the two matrices exhibited a similar frequency of tet(M) and blaZ, this did not apply to tet(L) and, especially, tet(K) and msr(C). It may be concluded that the distribution of resistance genes in water and sediment is only partially overlapping. This finding highlights the need for analyzing both water and sediment if a comprehensive view of antibiotic-resistant enterococci is to be achieved.

Finally, although the farm owner denied using antibiotics before and during the time of the study, the role of mariculture in favoring the spread of antibiotic resistance to enteric bacteria of nonmarine origin requires further investigation, also in order to understand its possible role in the emergence of multiresistant strains in an antibiotic-independent manner.

Footnotes

Acknowledgments

The authors are grateful to Dr. Andrea Brenciani and to Dr. Marina Mingoia for supplying the reference strains Streptococcus pyogenes 7008 and Streptococcus pneumoniae Pn8 and to Dr. Claudio Palmieri for technical assistance.

This work was supported by the Italian Ministry of Research and Education (contract PRIN 2008–I31J10000050001FYXAXL_003).

Disclosure Statement

None of the authors has any conflicts of interest to disclose.

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