Aeromonas salmonicida, freshwater wetland rhizobacterium, fish pathogen, and a potential environmental indicator.

The objective of this study was the development of potentially pathogenic bacteria as environmental indicators for freshwater wetlands. Aeromonas spp. was selected as a potential environmental indicator because these water-borne bacteria are both fish and emerging human pathogens. Selected bacterial isolates were analyzed using classical microbiological methods, API20E, API20NE, 16S rRNA sequencing, and susceptibility to antibiotics. The selected Aeromonas spp. isolates were identified by 16S rRNA sequencing as Aeromonas salmonicida. The rate of antibiotic resistance (< 50 [micro]g/ml) was high for ampicillin yet low (>10 to 20 [micro]g/ml) for kanamycin, tetracycline, and chloramphenicol for all strains tested. The rate of resistance for ampicillin suggests acquisition by gene transfer. However, the rate of resistance for kanamycin, tetracycline, and chloramphenicol suggests intrinsic antibiotic resistance. Because increased incidence of resistance to ampicillin in freshwater wetlands reflects responses to the increased exposure to antimicrobial compounds (especially ampicillin) over the past several decades, monitoring the response of antibiotic resistant A. salmonicida to environmental changes rather than entire bacterial assemblage represents a potentially productive approach for biomonitoring of natural systems. Utilizing the distribution and antibiotic resistance of Aeromonas spp. in the environment as warning of possible contamination and as index of water and soil/sediment quality deterioration should significantly contribute to the long-term protection of freshwater wetlands.

Keywords: Aeromonas spp., 16S rRNA, antibiotic resistance, environmental indicators

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Bacteria are potentially useful indicators of water and soil/sediment quality because of their species diversity and ability to respond rapidly to changing environmental conditions (Lemke et al., 1997). Previous data have shown that monitoring the response of antibiotic resistant enteric bacteria, rather than the entire assemblage, is a potentially productive approach to the examination of the responses of natural populations of bacteria to anthropogenic disturbances (Halda-Alija et al., 2000; Halda-Alija et al., 2001). Over 12 million kg of antibiotics are produced annually in the United States for use in man (Mazel and Davis, 1998) and almost as much for animal use. Increased introduction of antimicrobial agents into the environment via medical therapy, agriculture, and animal husbandry has resulted in new selective pressures on bacterial populations (Col and O'Connor, 1987). Despite the acquisition and transfer of antibiotic resistance genes in different environments worldwide (Alonso et al., 2001; Lederberg et al., 1992), resistant determinants persist in bacterial genomes over hundreds of generations, even in the absence of antibiotics as selective agents (Jabes et al., 1989). Consequently, there are two general categories of antibiotic resistance traits displayed by bacteria: (i) those that allow bacteria to withstand relatively high levels of a specific antimicrobial agent and conferred by mutations in genes responsible for antibiotic uptake or binding sites, as well as those gained by acquisition of genes on mobile elements (Davis, 1996); and (ii) those provided by genes conferring nonspecific low-level resistance (background levels of resistance) (Davis, 1996). Most investigations of antibiotic resistance in the aquatic habitat have addressed bacteria of fecal origin because they are used as pollution indicators and may be associated with infectious diseases. However, fecal bacteria are of little numerical significance in many freshwater systems, despite the fact that they are discharged into almost all inland waters (Jones et al., 1986). Thus, if the environmental pool of resistance is to be measured, bacteria other than those of fecal origin must be considered (Goni-Urriza et al., 2000).

Aeromonads are ubiquitous, gram-negative bacilli which are autochthonous to aquatic environments (Austin et al., 1996). The widespread use of antimicrobial agents for treating bacterial diseases in aquaculture has been associated with increased antibiotic resistance in A. hydrophila and A. salmonicida (fish pathogen) (Goni-Urriza et al., 2000; Schmidt et al., 2001). When antibiotics are used in aquaculture, they remain in the open environment or may eventually flow into open waterways with unknown consequences with regard to overall rates of antimicrobial resistance acquisition. Our understanding of the ecological aspects of antibiotic resistance in this regard is rudimentary.

The primary objective of this study was to assess sediments for possible pathogen loads and select alternative indicators of water and sediment deterioration. The rate of acquired antibiotic resistance was investigated for water borne Aeromonas spp. for which antibiotic resistance patterns are well known. Additionally, Aeromonas spp. is fish and emerging human pathogen, and thus may be used as environmental indicator. Aeromonas spp. was selected from the rhizosphere of the freshwater wetland plant Juncus effusus L. and sediments (Subgani, 2002). The secondary objective of this study was to assess the rate of the acquisition of antibiotic resistance for Aeromonas spp. obtained from freshwater wetlands.

MATERIALS AND METHODS

Sampling Sites -- Sediment samples were collected at a pristine, forested site located in Stewart county, Tennessee (Halda-Alija and Johnston, 1999) and an agricultural site located in Calloway county, western Kentucky (Halda-Alija et al., 2001) in spring, summer, and fall of 1997. Physical, chemical, and hydrological characteristics of the streams were described in detail elsewhere (Halda-Alija et al., 1999; Hendricks and Rice, 2000). Briefly, the chemistry of the agricutural site was highly variable (Halda-Alija et al., 2001) with high turbidity (14.1 NTU) and high total suspended solids (14.1 mg/L). The average annual surface water temperature was 13.9[degrees]C and the pH was 7 (calculated for 1997). The chemistry of the pristine site was highly variable with low phosphate inputs (0.02-0.06 mg/L), low nitrate inputs (0.03-0.12 mg/L), and low total suspended solids (3.9 mg/L) (Halda-Alija and Johnston, 1999; Halda-Alija et al., 1999; Hendricks and Rice, 2000).

Additionally, wetland sediments and Juncus effusus L. roots were collected at the University of Mississippi Field Station (UMFS) in wetland areas located in northern Mississippi in March and June 2000. UMFS (www.baysprings.olemiss.edu) is located in Lafayette County, northern Mississippi at the headwaters of the Little Tallahatchie River watershed (Figure 1). The soil type is Lexington silt loam (fine silty mixed thermic Typic Paleudalfs), the average annual air temperature is 15.9[degrees]C, and soil pH is 6.5 (Soil Survey of Lafayette County, MS; UMFS). The average winter temperature in Lafayette County is 5.7[degrees]C, spring air temperature 16[degrees]C, and summer 26[degrees]C (Soil Survey of Lafayette County, MS). Juncus effusus L. (soft rush) is one of the most robust and hardy of the native perennial wetland plant species surveyed at UMFS (Holland and Cooper, 1999). Plants were collected from four 4-[m.sup.2] plots and subdivided into planting units, consisting of no less than six shoots and their associated roots. To obtain bacterial isolates, three replicated planting units with associated sediments were collected to a depth of 30 cm at each 4-[m.sup.2] plot. Microbiological analyses (below) were initiated within 24 h of sample collection.

Enumeration and Characterization of Freshwater Wetland Microflora -- Rhizosphere sediments (10.0 g), closely attached to roots, were homogenized in saline and rhizosphere suspension serially diluted as described previously (Halda et al., 1991; Van Elsas and Smalla, 1997; Zuberer, 1994). Rhizosphere is defined as the thin layer of soil or sediment adhering to the root system after loose soil has been removed by shaking (Dandurand and Knudsen, 1997). In this case, rhizosphere extends up to 5 mm (millimeters) from the root surface. Roots were not included in this study. Briefly, rhizosphere suspension was serially diluted in saline until an appropriate dilution has been reached (up to [10.sup.-8]). Using dilution tubes, samples (0.1 ml) were pipetted and spread onto appropriate agar medium. Four replicate spread plates per dilution tube were used. Sediment samples were homogenized in saline and serially diluted as described above. The following culture media were used for bacterial enumeration: Tryptic Soy Agar (TSA, Difco Laboratories, Detroit, MI) for culturable, heterotrophic aerobic bacteria and MacConkey (Difco Laboratories) for gram-negative bacteria. Plates were incubated at 30[degrees]C in the dark; heterotrophic bacteria were enumerated after 48 to 72 h of incubation and gram-negative bacteria were counted after 48 h of incubation. Obtained isolates were purified and initially assessed for gram-negative and gram-positive reaction using 3% KOH method and the Gram staining method (Baron et al., 1994).

Isolation and Identification of Aeromonas spp -- MacConkey agar (Difco Laboratories) was used for the selection of gram-negative bacteria. Representative colonies having different colonial and cellular morphology were purified, stored, and characterized as previously described (Halda-Alija and Johnston, 1999; Halda-Alija et al., 2001). Preliminary identification of strains obtained in pure culture was based on Gram staining, respiration-fermentation tests, and oxidase reaction (Baron et al., 1994). Biochemical tests were performed with the API bacterial identification system from Bio-Merieux Inc., Hazelwood, MO. API 20E test kits were used for identification of enteric bacteria and API 20NE for aeromonads. Bacteria were identified according to Krieg and Holt (1984). Final identification of Aeromonas spp. was carried out according to criteria for environmental isolates recommended by Austin et al. (1996). Strains that could not be identified at the genus level were grouped as "unidentified." Identification of representative isolates was confirmed using 16S rRNA sequencing (MIDI Inc., Newark, DE).

[FIGURE 1 OMITTED]

16S rRNA Sequencing -- The 16S rRNA gene was PCR amplified from genomic DNA isolated from pure bacterial colonies (PrepMan[TM] Ultra Sample Preparation Reagent, Applied Biosystems Inc., Foster City, CA) following standard PCR protocols (Halda-Alija and Johnston, 1999) and using primers corresponding to E. coli positions 005 and 531 (500 bp packages). Amplification products were purified from excess primers and dNTPs using Microcon 100 (Millipore Inc., Bedford, MA) molecular weight cut-off membranes, and checked for quality and quantity by running 5 [micro]l of the products on 2.5% NuSieve/SeaKem agarose gel (Biowhittaker Inc., Rockland, ME) at 148 V (Sambrook, 1989).

Cycle sequencing of the 16S rRNA amplification products was carried out using AmpliTaq FS DNA polymerase (Applied Biosystems Inc., Foster City, CA) and dRhodamine dye terminators (Applied Biosystems Inc., Foster City, CA). Excess dye-labeled terminators were removed from the sequencing reactions (Applied Biosystems Inc., Foster City, CA) using a Sephadex G-50 spin column (Sephadex-Amersham Inc., Upsala, Sweden). The products were collected by centrifugation (1000 g for 2 min) at 4[degrees]C, dried under vacuum and frozen at -20[degrees]C until ready to load. Samples were resuspended in a solution of formamide (83% v/v)/blue dextran (0.8% w/v)/5 mM EDTA and denatured prior to loading. The samples were electrophoresed on an ABI Prism 377 DNA sequencer (Applied Biosystems Inc., Foster City, CA). Data were analyzed using Applied Biosystems DNA (Applied Biosystem Inc., Foster City, CA) editing and assembly software and sequence comparisons were obtained using the MicroSeq software (Applied Biosystem Inc., Foster City, CA). Bacterial identifications were based on 16S rRNA gene sequence similarity. Sequence analysis was performed using Applied Biosystems MicroSeq [TM] microbial analysis software and database (Applied Biosystem Inc., Foster City, CA). The top ten alignment matches were presented in a percent genetic distance format. Neighbors joining phylogenetic trees (Saitou and Nei, 1987) were generated using the top ten alignment matches.

Antibiotic Susceptibility Testing -- Antibiotics representing different chemical families were employed. Antibiotic susceptibility was determined by the plate agar and disk diffusion tests (Leboffe and Pierce, 2002). For both tests, Nutrient Agar (NA, Difco Laboratories) medium was used. In the plate agar method, test plates were identical to control plates except that 10, 50, and 100 [micro]g of either ampicillin ([beta]-lactams), kanamycin (aminoglycoside), tetracycline, or chloramphenicol per ml was added. The results obtained by the plate agar method were confirmed with disk diffusion test (Leboffe and Pierce, 2002). The results from three to four replicates were averaged. Statistical tests and the differences in mean values were compared using SYS-TAT v.9.2.1 (SPSS Inc., Chicago, IL).

RESULTS

Samples collected from sediments and the Juncus effusus L. rhizosphere gave rise to colonies exhibiting disparate morphologies on the TSA and MacConkey media. Occurrence of Enterobacter spp. did not exceed 5.6% (as a percentage of total number of heterotrophic bacteria) in the pristine stream and 8.5% in the Juncus effusus L. rhizosphere (Tables 1 and 2). The occurrence of Enterobacter spp. and Aeromonas spp. decreased in winter. Abundance of culturable enteric bacteria was higher in the agricultural streams (11.9% in fall) than in other environments (Halda-Alija and Johnston, 1999; Halda-Alija et al., 2001; this study). Enterobacteriacae and Aeromonas spp. are of little numerical significance compared to Pseudomonas and Bacillus spp. in freshwater wetlands (Tables 1 and 2; Halda-Alija and Johnston, 1999; Halda-Alija et al., 2001). Because Pseudomonas spp. is characterized by an extraordinary nutritional versatility (Palleroni, 1984), population sizes of Pseudomonas spp. do not reflect environmental differences. Antibiotic resistant pseudomonads are common in wetlands (Halda-Alija, unpublished). Although most strains of Pseudomonas spp. are significantly more resistant to many antimicrobial agents (including [beta]-lactams, tetracycline, chloramphenicol, and fluoroquinolones) than most other gram-negative rods, this resistance has been assumed to be intrinsic resistance (Li et al., 1994).

Despite the polyphasic approach employed, identification to the species level was inconclusive for most of the bacterial strains. Sequencing of 16S rRNA revealed A. hydrophyla and A. salmonicida to be among the antibiotic resistant rhizobacteria (Figure 2).

The resistance to [beta]-lactams was prevalent for aeromonads (Table 3). [beta]-lactams and tetracycline are widely used in human and veterinary practices and isolates resistant to these compounds were the most frequent. Results for chloramphenicol and kanamycin were similar. Chloramphenicol resistance was low; thereby suggesting it is conferred by nonspecific low-level resistance genes (i.e., background level of resistance). This is in agreement with other studies (Goni-Urriza et al., 2000), and probably the result from the restricted use of kanamycin and chloramphenicol. Although kanamycin is an effective antibiotic against a number of pathogenic bacteria, they have side effects that limit their use (Salyers and Whitt, 2002).

DISCUSSION

This study focused on antibiotic resistant Aeromonas spp. and demonstrated the abundance of antibiotic resistance in non-clinical environments. This observation is in agreement with four independent studies of antibiotic resistance in bacteria isolated from European mineral water (Massa et al., 1995; Mary et al., 2000; Papatropoulou et al., 1994; Rosenberg and Hernandez, 1989).

High level of resistance (<50 [micro]g/ml) for ampicillin indicates that ampicillin resistance was acquired through gene transfer. The ampicillin was deployed in 1961 and the resistance developed in 1973 (Lewis et al., 2002). It has been reported that Aeromonas salmonicida has the ability to transfer the antibiotic resistant phenotype to E. coli by R-plasmids (Adams et al., 1998). The hypothesis that antibiotic resistant genes are transferred between enterics and aeromonads in the environment requires further testing.

Our understanding of antibiotic resistance is rudimentary, especially with respect to the ecological aspects (Lemke and Leff, 1999). One study showed an increase in antibiotic resistance in an industrially perturbed stream, which positively correlated with mercury concentrations in the sediments (McArthur and Tuckfield, 2000). Antibiotic resistance genes can be present in replicons that contain other selectable markers and that might explain the predominance of ampicillin resistance in the wetland plant rhizospheres. Consequently, selection for antibiotic resistance determinants might occur in the environment as a function of high levels of antibiotic use and chemical or heavy metal pollution (McArthur and Tuckfield, 2000). Aeromonads play multiple roles in the environment as wetland plant-growth promoters (Subgani, 2002), nitrifiers (DeBoer and Kowalchuk, 2001; Nemergut and Schmidt, 2002), and potential environmental indicators (present study).

Aeromonads present adequate markers because they are ubiquitous in aquatic environments such as wetlands. We propose using aeromonad species, specifically A. hydrophyla and A. salmonicida, as environmental indicators of pollution because Aeromonas is considered an emerging human pathogen (suspected to cause a wide variety of diseases including gastroenteritis, septicaemia, endocarditis, osteomyelitis, myonecrosis, haemolytic uraemic syndrome, meningitis, peritonitis, respiratory tract disease and ocular infections) (Austin et al., 1996). In addition to being pathogenic to humans, Aeromonas spp. are also important pathogens in amphibians, reptiles and fish with the latter resulting in economic problems for the fish farming industry (Austin and Austin, 1993). Aeromonas species have also been isolated from a variety of foods such as vegetables, raw milk, ice cream, meat and seafood (Austin et al., 1996). Therefore, monitoring the presence of environmental aeromonads in order to detect possible contamination and water and soil/sediment quality deterioration will significantly contribute to the long-term protection of human health and the environment.

[FIGURE 2 OMITTED]

Table 1. Seasonal pattern of the occurrence of Enterobacter spp.,
Aeromonas spp., Pseudomonas spp., and Bacillus spp. in a pristine and an
agricultural stream as the percentage of the total number of
isolates (a).

                                              Genus
Season                   Aeromonas  Enterobacter  Pseudomonas  Bacillus
                         spp.       spp.          spp.         spp.

Winter
Pristine stream (b)      0.6 a       ND (c)       32.7 a        9.6 a
Agricultural stream (b)  0.6 a       3.7 a        29.8 a       10.2 a
Summer
Pristine stream          3.6 b       4.6 a        23.8 b       14.8 b
Agricultural stream      4.6 b      11.4 b        25.1 b       14.3 b
Fall
Pristine stream          3.1 b       5.6 a        24.8 b       13.9 b
Agricultural stream      4.7 b      11.9 b        23.6 b       11.7 b

(a) Values are average of three subsamplings and three replicates.
(b) Data obtained at five stream sites were combined and average is
presented (see Materials and Methods for details).
(c) Not determined.
abc-or each column, means followed by a common letter are not
significantly different according to LSD test (P < 0.05).

Table 2. Seasonal pattern of the occurrence of Enterobacter spp.,
Aeromonas spp., Pseudomonas spp., and Bacillus spp. in wetlands as the
percentage of the total number of isolates (a).

                                          Genus
Season (b)           Aeromonas  Enterobacter  Pseudomonas  Bacillus
                     spp.       spp.          spp.         spp.

Winter
Wetland sediment     0.9 a      4.3 a         17.3 a       16.1 a
Rhizosphere          0.7 a      2.1 a         16.1 a       13.6 a
(Juncus effusus L.)
Spring
Wetland sediment     1.7 b      5.1 a         20.3 b       17.3 a
Rhizosphere          2.2 b      8.5 b         18.7 b       15.3 a
(Juncus effusus L.)
Summer
Wetland sediment     3.4 c      6.3 c         15.6 c       19.2 b
Rhizosphere          3.7 c      8.1 c         13.9 c       18.6 b
(Juncus effusus L.)

(a) Values are average of three subsamplings and three replicates.
(b) Data are not available for fall.
abc-For each column, means followed by a common letter are not
significantly different according to LSD test (P<0.05).

Table 3. Percentage of Aeromonas spp. isolates obtained from different
environments resistant to antibiotics (a).

Environment                   % of resistant Aeromonas spp.
                      Ampicillin
                      ([beta]-lactam)  Tetracycline  Chloramphenicol

Pristine stream        ND (c)            ND            ND
Agricultural stream    ND                ND            ND
Wetland sediment (b)   17.30 a           7.30 a        0.50 a
Rhizosphere (b)        23.50 b           3.40 b        1.10 a
(Juncus effusus L.)

(a) Values are average of three subsamplings and three replicates.
(b) Combined data for spring and summer are presented (no significant
differences were found between spring and summer at p<0.05).
(c) ND = Not determined.
abc-For each column, means followed by a common letter are not
significantly different according to LSD test (P < 0.05).

ACKNOWLEDGMENTS

This study was supported by an American Association for the Advancement of Science/National Science Foundation grant and a grant from the University of Mississippi Office of Research. The authors acknowledge Charles Copper (United States Department of Agriculture) and Marjorie Holland (The University of Mississippi Field Station) for helpful discussions. The authors thank Clay Borden, Emily Garner, and Dinesh Talreja for technical assistance. Map assistance was provided by The University of Mississippi Geoinformatics Center, which is supported by National Aeronautics and Space Administration grant NAG13-00037.

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L. Halda-Alija (1) and R. K. Subgani

Department of Biology, University of Mississippi, University, Mississippi 38677

(1) Author for correspondence (E-mail: [e-mail])