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Applied and Environmental Microbiology, April 2007, p. 2224-2229, Vol. 73, No. 7
0099-2240/07/$08.00+0     doi:10.1128/AEM.02099-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Novel Method for Rapid Assessment of Antibiotic Resistance in Escherichia coli Isolates from Environmental Waters by Use of a Modified Chromogenic Agar

A. J. Watkinson,^1,2* G. R. Micalizzi,³ J. R. Bates,³ and S. D. Costanzo¹

National Research Centre for Environmental Toxicology, 39 Kessels Road, Coopers Plains, Brisbane, Queensland, Australia 4108,¹ Cooperative Research Centre for Water Quality and Treatment, PMB 3, Salisbury, South Australia, Australia 5108,² Public Health Microbiology, Queensland Health Scientific Services, 39 Kessels Road, Coopers Plains, Queensland, Australia 4108³

Received 5 September 2006/ Accepted 21 January 2007

Top Abstract Introduction Materials and Methods Results Discussion References

 
We validated a novel method for screening Escherichia coli resistance to antibiotics in environmental samples using modified Difco MI agar (Becton Dickinson) impregnated with selected antibiotics (tetracycline, ampicillin, cephalexin, and sulfamethoxazole), termed MI-R. This method combines an existing rapid assessment technique for E. coli enumeration with clinical reference data for breakpoint analysis of antibiotic resistance and was developed to address issues encountered when clinical methods are used with environmental samples. Initial trials conducted using strains of E. coli with resistance to the selected antibiotics showed that this method was reproducible and accurate with respect to antibiotic resistance. Trials using wastewater effluent demonstrated the precision of the method, and the levels of resistance found in effluent were directly comparable to the levels of antibiotic resistance determined using the more traditional CLSI (formerly NCCLS) disk susceptibility test. All wastewater isolates growing on MI-R plates were confirmed to be resistant using the CLSI disk susceptibility test. Bacterial resistance to ampicillin (38% ± 4% overall), sulfamethoxazole, tetracycline (21% ± 3% overall), and ciprofloxacin (6% ± 1%) were found in wastewater effluent. A successful trial was also conducted with water collected from the Brisbane River, Australia. The levels of antibiotic resistance in E. coli ranged from 0 to 47% for ampicillin, from 0 to 24% for tetracycline, from 0 to 63% for sulfamethoxazole, and from 0 to 1% for ciprofloxacin, with the highest incidence of resistance associated with wastewater treatment plant discharges. This method has great potential for rapid and representative assessment of antibiotic resistance in E. coli and could allow increased sample analysis, resulting in greater confidence in spatial analysis in environmental studies.

Top Abstract Introduction Materials and Methods Results Discussion References

 
The increase in the number of resistant and multiresistant strains of bacteria is a major concern of health officials worldwide, particularly with the decline in the number of new antibiotics available for treatment. While much effort has been directed toward management and monitoring of antibiotic use and the prevalence of bacterial resistance within communities, bacterial resistance to antibiotics in the aquatic environment has received comparatively little attention. Bacterial contamination of surface waters, particularly contamination with fecally derived bacteria, has long been a water quality issue due to the potential for disease transmission. Because of this and the potential for antibiotic resistance, there is a new level of risk associated with these bacteria. Recent studies have also identified antibiotics themselves in surface waters (2, 7, 9, 12), and the role of these antibiotics in the development, transfer, and maintenance of resistance is largely unknown.

In a limited number of studies workers have identified antibiotic-resistant bacteria in the aquatic environment. In a study of 16 United States rivers, antibiotic-resistant bacteria were found to be widespread, and the resistance included resistance to chemically modified and synthesized antibiotics (1). Parveen and coworkers (14) showed that the frequency of antibiotic-resistant and multiple-antibiotic-resistant Escherichia coli isolates was higher close to point source discharges. Boon and Cattanach (3) demonstrated that antibiotic resistance was significantly greater in native heterotrophic bacteria than in E. coli in river samples from southeast Australia. However, there have not been enough studies to assess the fate of antibiotic-resistant bacteria in the aquatic environment.

While monitoring and identification of bacterial resistance in clinical environments is a well-established and developed field, little is known about the occurrence and transfer of resistance in the aquatic environment. E. coli has been the foremost indicator of fecal contamination in water quality monitoring for many decades. E. coli has also been shown to be a significant reservoir of genes coding for antimicrobial drug resistance and therefore is a useful indicator for resistance in bacterial communities (6).

Traditional techniques, such as the CLSI (formerly NCCLS) disk susceptibility method (8), have proven to be impractical and time-consuming. Due to the great financial and personnel resources required, broad spatial assessments of bacterial resistance to antibiotics have not been feasible. Therefore, we developed a novel method for rapid assessment of bacterial resistance in surface waters to alleviate previous difficulties. In our new method a modified selective agar impregnated with antibiotics is used to rapidly assess the resistance of E. coli to antibiotics. Difco MI agar (Becton Dickinson, New Jersey) is a relatively new agar and has been approved for use by the United States Environmental Protection Agency for enumeration of E. coli and total coliforms (TCs) in water (16). This medium simultaneously detects E. coli and TCs in a variety of water types, including drinking water, and has been shown to be sensitive, selective, and specific and provide results that have low false-positive and false-negative rates within 24 h or less (4). The new method has proven to be superior to the current United States Environmental Protection Agency-approved method in terms of (i) recovery of target organisms, (ii) reduction of background or noncoliform organisms, and (iii) recovery of chlorine-damaged and/or nutrient-deprived target organisms (5) and thus was an ideal candidate for our prospective trial.

MI agar involves the use of a fluorogenic component (4-methylumbelliferyl-ß-D-galactopyranoside) and a chromogenic component (indoxyl-ß-D-glucuronide) that identify E. coli and TCs through reactions with endemic enzymes. Specifically, the E. coli enzyme ß-glucuronidase cleaves indoxyl-ß-D-glucuronide, causing E. coli colonies to appear blue (4). Additionally, the coliform enzyme ß-galactosidase cleaves 4-methylumbelliferyl-ß-D-galactopyranoside, which further confirms the presence of positive colonies under long-wavelength (366-nm) UV light and aids in separating E. coli and coliforms (4). In this study, we used this agar impregnated with specific antibiotics (ampicillin, tetracycline, sulfamethoxazole, and ciprofloxacin) at concentrations that define resistance (breakpoint concentrations) based on clinical studies (8). These antibiotics were chosen because they are widely used in Australia (11) and have been found in wastewater discharges in the study region (9).

Top Abstract Introduction Materials and Methods Results Discussion References

 
Preparation of media.
The following four antibiotics were chosen for development of the new method: ampicillin, tetracycline, sulfamethoxazole, and ciprofloxacin. Additionally, cefsulodin was added to plates to inhibit the growth of gram-positive bacteria and some noncoliform gram-negative organisms that can cause false-positive results (4). All antibiotics were obtained from Sigma Chemical Company, St. Louis, MO. Prior to antibiotic addition, all reagents were autoclaved (103 kPa, 121°C, 15 min) and filtered (PALL Acrodisc syringe filter, 0.2-µm Supor membrane). Ampicillin and cefsulodin were dissolved in 0.1 M phosphate buffer (pH 6.0), and sulfamethoxazole was dissolved in 0.5 volume of hot water with a minimal amount of 2.5 M sodium hydroxide (8). Ciprofloxacin and tetracycline were dissolved in water (8). Intermediate antibiotic solutions were prepared in autoclaved (103 kPa, 121°C, 15 min) amber glass jars. Media were prepared by dissolving 36.5 g of Difco MI powder (Becton Dickinson, New Jersey) in 1 liter of sterile water. Each solution was agitated and heated to aid dissolution and then boiled for 1 min prior to autoclaving (103 kPa, 121°C, 15 min). After the medium had cooled to less than 45°C, cefsulodin was added to obtain a final concentration of 5 µg ml^–1. One set of control plates (containing no antibiotics) were prepared along with a set of plates for each of the four antibiotics used. The antibiotics were added individually to obtain the following batch final concentrations: ampicillin, 32 µg ml^–1; tetracycline, 16 µg ml^–1; sulfamethoxazole, 350 µg ml^–1; and ciprofloxacin, 4 µg ml^–1. These concentrations reflected the breakpoint concentrations for testing with E. coli (8). Medium (8 ml) was dispensed into 50-mm plates for testing, and this medium was designated MI-R to distinguish it from the medium used in the original method (MI).

Controlled-spike trial.
An initial trial was conducted using E. coli strains with known antibiotic resistance profiles. E. coli ATCC 25922 was used as a control strain having no resistance to the selected antibiotics. From an existing culture library, we chose E. coli strains having resistance to the selected antibiotics which were isolated from wastewater effluent, and these strains included ampicillin-resistant (LP2ecb), tetracycline-resistant (A11eca), sulfamethoxazole-resistant (FF21ecb), and ciprofloxacin-resistant (FF20ecb) strains. Antibiotic resistance was confirmed using the CLSI disk susceptibility test (8). All five strains were grown overnight in brain heart infusion broth at 35°C, and dilutions of each strain were prepared separately in sterile peptone water up to a final dilution of 10^–7. A 10^–7 dilution had previously been determined to maximize the adequate colony counts per plate in order to satisfy statistical growth requirements (16). One milliliter of each dilution was added to 10 ml of peptone water in a filtration apparatus (Gelman, East Hills, NY) and filtered through separate 0.22-µm membrane filters (Millipore, Bedford, MA). For each E. coli strain, six replicate dilutions were prepared for the control (no antibiotics) and each treatment (four antibiotics). Filters were transferred onto the prepared MI and MI-R plates, and the plates were incubated at 35°C for 24 h. Blue colonies were counted under ambient light, and the results were confirmed under long-wavelength UV light (366 nm). A one-way analysis of variance was used to determine significant differences (P < 0.05) between control plates and individual antibiotic plates.

Confirmed wastewater trial.
An additional trial was conducted using effluent from a regional wastewater treatment plant (WWTP). Antibiotic-resistant bacteria have previously been found in effluent from this plant (9), making it an ideal test site for this method. One liter of effluent was collected from the effluent stream in an autoclaved glass amber jar and transported to the laboratory on ice. Two pseudoreplicate series consisting of 1, 0.1, and 0.01 ml of effluent were filtered with 10 ml of peptone water through 0.22-µm membrane filters, transferred to control and antibiotic plates in triplicate, and incubated at 35°C for 24 h. After incubation, blue colonies were counted under ambient light, and the results were confirmed under long-wavelength UV light (366 nm). For each antibiotic, the percent resistance was calculated by directly comparing the counts on the antibiotic plate with the corresponding counts on the control plate:

(1)

A one-way analysis of variance was used to determine significant differences (P < 0.05) between replicate series for each plate type. E. coli ATCC 25922 was used as a control strain that exhibited no resistance.

To validate the levels of resistance determined with this method, we employed the CLSI disk susceptibility method (8) in conjunction with the MI-R method. This was done by randomly selecting 100 isolates from the control plates, which were picked off, plated onto blood agar, and incubated at 37°C for 24 h. Colonies were then picked off and transferred to a 0.85% NaCl solution to obtain a 0.5 McFarland standard. After vortexing to homogenize each solution, a sterile cotton swab was used to spread an even layer of the solution onto a prepared Mueller-Hinton agar plate. Antibiotic disks were placed onto the Mueller-Hinton agar plates, and the plates were incubated at 37°C for 24 h. Resistance was determined by comparing zones of inhibition with CLSI guidelines using E. coli ATCC 25922 as a reference strain (8). Resistance to ampicillin (10 µg; Oxoid), resistance to tetracycline (30 µg; Oxoid), resistance to sulfafurazole (350 µg; Oxoid), and resistance to ciprofloxacin (5 µg; Oxoid) were tested. Fifty colonies from each of the MI-R plates were picked off and individually tested for resistance to ampicillin (10 µg; Oxoid), tetracycline (30 µg; Oxoid), sulfafurazole (350 µg; Oxoid), trimethoprim (1.25 µg; Oxoid)-sulfamethoxazole (23.75 µg; Oxoid), ciprofloxacin (5 µg; Oxoid), and cephalothin (30 µg; Oxoid,) using the disk susceptibility test as described above.

Environmental trial.
The new method was then employed to investigate the extent of E. coli resistance to the four selected antibiotics in surface waters of a large urbanized river subject to a range of wastewater discharges (Brisbane River, Australia) (Fig. 1). Nine sites along the river were chosen based on major influences in the system. At each site, 1 liter of water was collected in autoclaved glass amber jars and transported back to the lab on ice. A dilution series consisting of 100, 10, 1, 0.1, and 0.01 ml was filtered through 0.22-µm membrane filters (Millipore, Bedford, MA), the last three with 10 ml of peptone, and transferred to each of the five types of plates. A dilution series was prepared for the control and each of the antibiotic treatments. The plates were incubated at 35°C for 24 h, and colonies were counted and results were confirmed as described above. For each antibiotic, the percent resistance was calculated by directly comparing counts on the antibiotic plate with the corresponding counts on the control plate as described above (equation 1).

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FIG. 1. Map of the Brisbane River showing the study sites used for the environmental trial. The sites where there is WWTP discharge are indicated.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Controlled-spike trial.
Our results indicate that the new technique successfully replicated results obtained with traditional techniques (Table 1). The control strain (ATCC 25922) was not observed to grow on any of the antibiotic plates but exhibited the expected growth on the control plate. Colonies of the ampicillin-resistant strain grew on the ampicillin plate at levels comparable to the levels on the control plate (P > 0.05); however, it did not grow on any of the other antibiotic plates (tetracycline, sulfamethoxazole, and ciprofloxacin plates). Similar patterns were observed for the remaining resistant strains; the growth of each of these strains was comparable to the growth of the control on the corresponding antibiotic plate, and the strain did not grow on the remaining plates.

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TABLE 1. Average numbers of CFU of selected antibiotic-resistant strains on control and antibiotic plates

Confirmed wastewater effluent trial.
The E. coli counts in wastewater were comparable for the two psuedoreplicates (48,000 ± 6,500 E. coli cells ml^–1 and 45,600 ± 6,300 E. coli cells ml^–1) (Table 2). The levels of resistance to the four antibiotics investigated were also comparable for the two pseudoreplicates (Table 2). The highest level of antibiotic resistance in E. coli was the resistance to ampicillin (41% ± 5% and 36% ± 3% for the two psuedoreplicates). This was followed by resistance to sulfamethoxazole (37% ± 5% and 34 ± 3%) and resistance to tetracycline (23% ± 4% and 19 ± 3%), and the lowest level of resistance was the resistance to ciprofloxacin (7% ± 1% and 5% ± 1%). There was no significant difference between the two separate dilution pseudoreplicates (P > 0.05). The observed levels of resistance in the MI-R trials were comparable to the levels determined with the CLSI disk susceptibility test for all of the antibiotics investigated (Table 2). The level of resistance to ampicillin (38% ± 4% overall) determined by the MI-R method was slightly greater than the level determined by the CLSI disk susceptibility test (34%), and these results were paralleled in the sulfamethoxazole trial (34% ± 2% overall versus 30%). The overall levels of tetracycline and ciprofloxacin resistance in the MI trial (21% ± 3 and 6 ± 1%, respectively) were slight lower than corresponding estimates obtained using the disk susceptibility test (22 and 9%, respectively); however, all the differences were within acceptable limits.

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TABLE 2. Comparison between MI-R method and CLSI disk susceptibility test for detection of antibiotic-resistant E. coli in wastewater effluent

All E. coli isolates picked off the ampicillin plate and tested using the CLSI disk susceptibility test were confirmed to be ampicillin resistant (Table 3). Identical results were obtained for all of the antibiotics tested, and resistance to each of the antibiotics tested was confirmed. All of the isolates tested exhibited resistance to other antibiotics tested. A high incidence of multiple drug resistance in E. coli was also observed for isolates obtained from the antibiotic plates tested using the CLSI disk susceptibility test (Table 3). Overall, isolates exhibited the highest incidence of multiple resistance to four drugs (36%), followed by multiple resistance to five drugs (24%) and three and two drugs (13%); 9% of all bacteria isolated exhibited resistance to all six antibiotics tested. The levels of resistance varied between isolates for each of the antibiotics tested, as indicated by different zones of inhibition; this included isolates with zones of inhibition close to the breakpoint zone (Table 4).

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TABLE 3. Antibiotic resistance profiles for E. coli isolated from antibiotic-impregnated MI-R plates and occurrence of multiple antibiotic resistance

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TABLE 4. Levels of resistance of MI-R isolates as demonstrated by measured zones of inhibition using the CLSI disk susceptibility test

Environmental trial.
The E. coli concentrations ranged from 3,333 E. coli CFU 100 ml^–1 at site 1 to 5 E. coli CFU 100 ml^–1 at site 9 (Table 5). The highest E. coli counts were obtained for sites with WWTP influence and no disinfection; the counts were 3,333 E. coli CFU 100 ml^–1 at site 1 and 364 E. coli CFU 100 ml^–1 at site 2. The E. coli concentrations at urban sites were lower than the E. coli concentrations at WWTP sites; the concentrations were 300 E. coli CFU 100 ml^–1 at site 3 and 197 E. coli CFU 100 ml^–1 at site 4. At two sites influenced by chlorinated WWTP effluent (sites 5 and 6) the E. coli counts were highly variable; however, at both of these sites the counts were much lower than the counts at nondisinfected WWTP-influenced sites (113 E. coli cells ml^–1 at site 5 and 11 E. coli CFU 100 ml^–1 at site 6). The agriculturally influenced sites, sites 7, 8, and 9, exhibited the lowest overall E. coli concentrations (55, 40, and 5 E. coli CFU 100 ml^–1, respectively).

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TABLE 5. Antibiotic resistance profiles for E. coli in the Brisbane River

Top Abstract Introduction Materials and Methods Results Discussion References

 
Laboratory trials conducted in this study demonstrated that the MI-R method is both accurate and reproducible for determination of antibiotic resistance in E. coli (Table 1). The controlled-spike trial showed that E. coli concentrations can be accurately reproduced on both control and antibiotic plates using this new method. This is consistent with previous work which demonstrated the high precision and reproducibility of MI media (4, 5). Here we demonstrated this in the presence of inhibitory levels of antibiotics. The antibiotic-resistant test strains grew comparably on the control plates and the plates containing the antibiotics to which the bacteria were resistant.

Trials with wastewater also confirmed that this new method is highly reproducible, accurate, and representative (Table 2). All values for antibiotic resistance were comparable to values generated with the CLSI disk susceptibility test. This supports the conclusion that this method can be used to accurately determine levels of antibiotic resistance in E. coli. Various zones of inhibition for the isolates indicated that the method can enumerate strains with various levels of resistance, including strains with MICs close to breakpoint concentrations, indicating that the method is suitable for even marginally resistant isolates (Table 4).

All colonies isolated from the antibiotic plates were confirmed to be resistant to the corresponding antibiotics using the CLSI disk susceptibility test (Table 3). This further strengthens the conclusion that this method can be used for assessment of antibiotic resistance in E. coli. Substantial resistance of the isolates to other antibiotics was also demonstrated, and a high incidence of multiple resistance was evident (Table 3). While Grabow et al. (10) showed that nontransferable resistance was more common than transferable resistance in wastewater-borne fecal coliforms (via plasmids, transposons, etc.), the prevalence of multiple-antibiotic-resistant bacteria could indicate that this is no longer the case, and research has demonstrated that there is exchange of plasmids between E. coli and other coliform bacteria in wastewater systems (10).

The final step in the validation process was to evaluate this method using an environmental trial (Table 5). This analysis was successful, and the results reiterated the strengths of the method as an accurate, reproducible, and rapid technique for assessment of E. coli resistance to antibiotics. The rapidity of the method was demonstrated in this environmental trial; the results were available within 24 h of sampling, compared with a minimum of 5 days for the previously used method combining membrane filtration (15) with the CLSI disk susceptibility test. The incidence of antibiotic-resistant bacteria appeared to be higher in locations adjacent to sites of WWTP discharge. Similar results have been obtained in other studies (3, 14), demonstrating the importance of point source discharges for the addition of antibiotic-resistant bacteria to the aquatic environment. The highest incidence of antibiotic-resistant bacteria was at site 5, where the effluent was chlorinated prior to discharge. Despite this, the E. coli concentrations in this effluent were still substantial, and the associated resistance was comparable to that seen in the raw effluent trial. One possible explanation for this anomaly is that organisms that survived the chlorination process also had high levels of antibiotic resistance, a phenomenon that has been demonstrated previously (13). This was not apparent at site 6, where chlorination also occurred, but the phenomenon could be dose dependent given the difference in E. coli concentrations between these two sites.

In conclusion, the method described here for rapid assessment of antibiotic resistance in waterborne E. coli has proven to be extremely successful. Not only is this method highly reproducible, accurate, and precise, but it also provides results within 24 h, greatly reducing the labor and the cost and allowing greater sample analysis and therefore spatial assessment.

 
We thank the staff of the Microbiology Department of Queensland Health Scientific Services for their support of and assistance with this project and Leon Peters for his aid with laboratory work.

This project was supported by an Australian Research Council Linkage Grant (grant LP0453-708) and in part by the Wastewater Program of the Cooperative Research Centre for Water Quality and Treatment (project 666003).

 
* Corresponding author. Mailing address: National Research Centre for Environmental Toxicology, 39 Kessels Rd., Coopers Plains, Brisbane, QLD, Australia 4108. Phone: 61 7 3274 9004. Fax: 61 7 3274 9003. E-mail: a.watkinson{at}uq.edu.au.

Published ahead of print on 2 February 2007.

Top Abstract Introduction Materials and Methods Results Discussion References

 

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Applied and Environmental Microbiology, April 2007, p. 2224-2229, Vol. 73, No. 7
0099-2240/07/$08.00+0     doi:10.1128/AEM.02099-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

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This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Watkinson, A. J. Articles by Costanzo, S. D. Search for Related Content PubMed PubMed Citation Articles by Watkinson, A. J. Articles by Costanzo, S. D. Agricola Articles by Watkinson, A. J. Articles by Costanzo, S. D.