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ABSTRACT |
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Cholera and other enteric diseases are particularly noticeable after natural disasters due to contaminated water supplies. The Indian Ocean tsunami of December 2004 devastated parts of Thailand, India, Sri Lanka, Maldives, Malaysia, Myanmar, Somalia, and much of Indonesia, with an estimated 225,000 deaths from the tidal impact alone. These are areas where cholera is endemic. Many additional deaths were possible in the aftermath of the tsunami due to the spread of infectious diseases, like cholera, dysentery, and viral gastro-enteritis. One of the primary sources of cholera is contaminated drinking water, which may arise from damage to infrastructure, including sewage- and drinking water-handling systems.4,6–9 Last year, months of heavy rains triggered a cholera epidemic in West Africa that was associated with flooding latrines and contaminated wells. Wars and civil conflicts, like the recent unrest in Liberia, have also been linked with outbreaks of cholera due to the displacement of large numbers of civilians.10 In the case of natural or man-made disasters, the loss of electricity, laboratory equipment, and supplies can make conventional testing for water-borne pathogens impossible.
Recently, we identified phosphoglucose isomerase with a novel secondary activity as a lysyl aminopeptidase (LysAP) in members of the Vibrionaceae family, but not present in non-Vibrionaceae pathogens.11–13 We developed a rapid, simple, and inexpensive detection technique to screen for the presence of this LysAP activity in bacteria grown on non-selective laboratory media.14 This technique, known as the colony overlay procedure for peptidases (COPP assay), was used to detect a broad array of Vibrionaceae family members, including V. cholerae serogroups O1, O139, and O155, V. parahaemolyticus, and V. vulnificus. In addition, Aeromonas hydrophila, also a member of the Vibrionaceae family, was detected in environmental samples including shellfish, seawater, and sewage.14 Aeromonas hydrophila is an opportunistic pathogen that has been associated with outbreaks of water-borne illness.15–17 Since A. hydrophila is associated with sewage, it is a potential indicator of fecal contamination in drinking water.18
In this article, we evaluate the use of the COPP assay for the rapid and simple detection and quantitation of V. cholerae serogroups O1, O139, and O155, and of A. hydrophila in well water. We also identify a potential application for the COPP technique as a practical method to determine the sanitary quality of well water in tropical climates when power supplies and laboratory facilities are not available.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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. Escherichia coli O157:H7 was used as a negative control. All bacteria were cultured on tryptic soy agar (Becton, Dickinson and Co., Sparks, MD) supplemented with 0.5% added NaCl (TSA-N) and incubated overnight (16–18 hours) at 37°C, unless otherwise specified.
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Water collection and bacteriological screening. Well water was collected in sterile 50-mL tubes during March, August, September, and October from 5 shallow (5–8 meter deep) wells in Dover and Laurel, Delaware. Aerobic plate counts were performed on the water for indigenous bacteria by spreading 100 µL of each sample (within 3 hours of collection) onto TSA-N plates followed by incubation at 37°C for 24 hours and enumeration of the colonies. Water was also aseptically collected and analyzed from 3 fresh-water lakes in Dover, Delaware to compare any differences in results obtained between lake water and well water. Lake water was intentionally collected and assayed during the peak migratory season for geese and ducks in an effort to increase the diversity of potential bacterial loads.
Colony overlay procedure for peptidases (COPP) assay.
The COPP assay was performed as previously described and as shown in Figure 1.14 In essence, 100 µL of water was inoculated onto a TSA-N plate (Fig. 1A), the water was spread with a flamed and cooled metal triangle (Fig. 1B and 1C), and the plate was inverted and incubated overnight at 37°C (Fig. 1D). In the event of an electrical interruption, the plate may be incubated in a drawer at 22°C or higher (Fig. 1E), until colonies are visible (usually within 24 hours). After incubation, each plate was observed for bacterial colonies (Fig. 1F). If bacteria were present, a previously prepared cellulose acetate membrane containing L-Lys-AFC (see previous discussion) was used to overlay the colony or colonies. One 8 x 15 cm membrane can be easily cut into smaller pieces to overlay individual colonies on a plate. Cultures more than 24 hours old should not be overlaid, because the acid produced by the bacteria in old cultures inhibits enzyme activity.14 Membranes were labeled with the sample number or code using a pen or pencil, and prewet for 5 seconds in 20 mM Tris-HCl, pH 9.0 (Fig. 1G). Excess buffer was allowed to drip from the membranes for 3–5 seconds and the membranes were carefully placed onto the bacterial colonies on the agar plates (Fig. 1H). Care was exercised to prevent bubbles from becoming trapped under the wet membranes. Plates were incubated for precisely 10 minutes at 30–37°C without inversion (Fig. 1I). After the brief overlay, each membrane was removed with forceps and placed in an empty Petri dish (Fig. 1J). Each membrane was viewed with a hand-held UV light (Fig. 1K) or on an ultraviolet (UV) light box (Fig. 1L) at a wavelength of 364 nm. This long-wave UV light is the wavelength for simple "black lights". No fluorescence was visible under short-wave (germicidal) UV. Viewing for fluorescent foci was performed in a darkened room. Membranes were observed for bright spots at the point of contact between the bacterial colony and the membrane. The number of fluorescent foci present on the membrane is a quantitative representation of the colony forming units (cfu) of V. cholerae, A. hydrophila, or other Vibrionaceae family members that are present in the inoculum. This number may be recorded for a permanent record of the level of contamination. Membranes may also be photographed to record the results.
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Analysis of pure cultures.
Pure cultures of V. cholerae O1, O139, and O155, A. hydrophila, Escherichia coli, Salmonella enterica subtype Typhimurium, and Shigella sonii (Table 1
) were grown overnight at 37°C, 250 rpm, in tryptic soy broth (Beckton, Dickinson and Co.) supplemented with 0.5% additional NaCl (TSB-N). Groundwater wells that were found to be free of bacterial contamination were re-sampled and the water was aliquotted, inoculated with 10-8 and 10-9 dilutions of V. cholerae O1, O139, and O155, A. hydrophila, E. coli, S. Typhimurium, and Sh. sonii onto TSA-N plates, and the plates were incubated at 37°C overnight. Individual colonies were subjected to the COPP assay using membrane strips to compare the relative fluorescence of the isolates. The E. coli, S. Typhimurium, and Sh. sonii served as negative controls.
Analysis of mixed cultures. Since sewage-contaminated well water would likely contain a mixture of potential pathogens, well water was also inoculated with dilutions of overnight broth cultures of V. cholerae O1, A. hydrophila, and E. coli (negative control) in varying proportions and 100 µL were plated on TSA-N and incubated at 37°C overnight followed by the analysis of colonies on countable plates by the COPP assay.
Incubation temperature and overlay duration. To determine the influence of incubation temperature and the duration of the overlay during the COPP assay, TSA-N plates were inoculated with the 3 serotypes of V. cholerae, and with A. hydrophila and E. coli. The plates were incubated overnight at 22, 30, and 37°C. Membranes were cut into small strips, wet in 20 mM Tris-HCl, pH 9.0, overlaid onto the overnight colonies for 5, 10, 15, 20, and 25 minutes, incubated at 22, 30, and 37°C, and the fluorescence intensities were compared.
Disinfection. Liquid chlorine (sodium hypochlorite) bleach was evaluated for its effectiveness in disinfecting contaminated plates. High-density, overnight cultures of V. cholerae O1 on TSA-N plates were flooded with 5 mL of a 10% solution of chlorine bleach for periods up to 15 minutes. Since the bleach was originally 6% sodium hypochlorite (5.5% available free chlorine), the diluted bleach contained 0.55% free chlorine. After various exposure intervals, 10 µL of the solution were added to tubes containing 6 mL of TSB-N to enrich for residual V. cholerae. The tubes were incubated at 37°C, 250 rpm and observed for turbidity after 24 hours. In addition, the used disinfection solution was also diluted 1:10, 1:100, and 1:1000 in sterile distilled water, and 100 µL was spread plated onto TSA-N plates, incubated at 37°C overnight, and the bacterial colonies were enumerated.
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RESULTS |
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). Vibrio cholerae O1, O139, O155, and A. hydrophila (strips labeled 1–4, respectively) produced strongly fluorescent foci, while the E. coli, S. Typhimurium, and Sh. sonii (strips labeled 5–7, respectively) were non-fluorescent (Fig. 2A). Aerobic plate counts were also performed on lake water and produced 1, 15, and 27 colonies per 100 µL of which 0, 2, and 5 colonies, respectively, produced fluorescent foci on the COPP assay. Approximately one half of the fluorescent foci were from bacteria and the other one half were associated with fungal colonies.
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shows an example of a COPP assay that was performed on well water that had been spiked with V. cholerae O1, A. hydrophila, and E. coli. Vibrio cholerae O1 were large colonies that produced strongly fluorescent foci. Aeromonas hydrophila colonies also produced strongly fluorescent foci; however, the foci were only about one third the diameter of the V. cholerae, thus allowing their differentiation. The E. coli colonies were about twice the diameter of the A. hydrophila and produced only very weak (negligible) fluorescence (Fig. 2B). Vibrionaceae produce substantially stronger fluorescence than nonvibrios. The analysis of 100 µL of water gave 16 foci from V. cholerae and 14 foci from A. hydrophila (Fig. 2B), representing 160 and 140 bacteria/mL of water, respectively. This demonstrates the quantitative nature of the assay. The E. coli produced only a trace of background fluorescence, which was insufficient to enumerate the foci.
Effects of time and temperature.
We also evaluated the effectiveness of the overlay as a function of time and temperature. Incubation of the A. hydrophila, E. coli, and the 3 subtypes of V. cholerae at 22, 30, and 37°C resulted in the growth of all the bacteria. Colonies were generally smaller when incubated at the lower temperatures. A visual comparison of the fluorescence intensities at various temperatures demonstrated strong fluorescence for V. cholerae O1, O139, and O155 and for A. hydrophila, when overlaid for 10 minutes at 30 and 37°C; however, weaker fluorescence was observed when the overlay was performed at 22°C. At 30 and 37°C, strong fluorescent signal was evident after an overlay for as little as 5 minutes for the 3 V. cholerae with maximum fluorescent signal after only 10 minutes. For A. hydrophila, the signal was comparable between 10 and 25 minutes, indicating that only 10 minutes are required for the overlay of A. hydrophila. In contrast, E. coli produced only a trace of fluorescence after a 10-minute overlay (Fig. 2). This weak signal was previously reported for E. coli and was attributed to the presence of other enzymes that exhibit a weak lysyl aminopeptidase activity in this and other non-Vibrionaceae species.14 Since V. cholerae and A. hydrophila produce substantially stronger signal, it is not likely that their fluorescence would be confused with that seen in E. coli, especially if the overlay time is limited to 10 minutes. A 10-minute overlay at 30–37°C is suitable for general use.
Chlorine disinfection. Disinfection of the TSA-N plates was evaluated using 10% sodium hypochlorite bleach. Results indicated the total elimination of V. cholerae after exposure for only 5 minutes. No growth was obtained on enrichment broth cultures of the bleach-water or on the TSA-N plates inoculated with a 1:10 to 1:1000 dilution of the bleach-water. In areas that may be impacted by the loss of electricity or the lack of an autoclave, our results indicate that the COPP assay plates and membranes may be safely disinfected with chlorine bleach.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Although it has been reported that the infectious dose for V. cholerae may be millions of bacteria, a recent study demonstrated that doses of 104 and 105 frozen V. cholerae O1 were required to elicit weak and moderate illnesses, respectively, in healthy volunteers who had previously ingested sodium bicarbonate to neutralize stomach acids.21 It was also shown that 105 freshly cultured V. cholerae caused infection in volunteers receiving sodium bicarbonate.21 The level of V. cholerae required to illicit infection after a natural disaster, where the population may be in poor health, but where the population is also not receiving sodium bicarbonate treatment, may be similar. Consequently, not all of the pathogens would have to be eliminated from drinking water to make the water relatively safe from V. cholerae. Given that the COPP assay only tests 100 µL of water per plate, that the plate contains the non-selective TSA-N agar that will allow the growth and quantification of virtually all human enteric pathogenic bacteria, and that a single colony of V. cholerae will produce a fluorescent focus on the COPP membrane, the detection threshold of the assay is 1 cfu of V. cholerae per 100 µL or 10 cfu of V. cholerae per ml of water. Even at that level, a person would have to drink 10 L of water to receive a dose of 105 bacteria, but only 1 L to ingest 104 bacteria. In the event greater assay sensitivity is desired, bacteria in the water could be enriched in tubes of TSB-N or in alkaline peptone water for several hours and 100 µL of the broth used to inoculate a plate of TSA-N for overnight incubation and COPP overlay. This would allow the COPP assay to be used to detect lower levels of V. cholerae and A. hydrophila, either of which serve as indicators of fecal pollution and signal the possible presence of other enteric pathogens, including enteric viruses, in the water.
The detection of A. hydrophila, which appears to be ubiquitous in sewage,18 could serve as an indicator for the possible presence of many other pathogenic bacteria and viruses in ground water supplies. Among these pathogens are Salmonella and Shigella spp., E. coli, and enteric viruses like hepatitis A and E viruses, rotaviruses, and noroviruses. Other media for Vibrio detection, particularly the highly selective thiosulphate citrate bile salts sucrose (TCBS) agar, readily detect many of the Vibrio spp.;22 however, studies in our laboratory showed that TCBS will not support the growth of A. hydrophila (unpublished). In addition, vibrios that are stressed may not grow on TCBS or other selective media, thus reducing the ability to detect these pathogens in well water.
The analysis of seawater by the COPP assay typically produces positive results from the indigenous Vibrionaceae found in the marine environment. The detection of Vibrionaceae could signal instances where fresh well water is contaminated with seawater, giving the impression that the water is contaminated with bacteria of fecal origin. Since seawater is unsafe to drink, a positive COPP assay would signal that the well water may be unsafe to drink, albeit not necessarily from sewage contamination.
We detected COPP-positive colonies in water from 2 of the 3 lakes tested. This was expected since Aeromonas and other Vibrionaceae may be shed in the feces of birds and other wild and domestic animals. We collected the lake water when high numbers of migratory geese, ducks, and other birds were in the area, which likely contributed to some of the organisms isolated. "Natural" contaminants could make the use of the COPP assay inappropriate for V. cholerae screening of lake water. In contrast, groundwater is filtered to some extent by the soil it passes through and may account for the typically negative APCs that we observed for well water. Some carryover of bacteria from fresh water lakes and rivers to nearby shallow wells could occur in areas prone to quick groundwater recharge and, in such areas, the use of the COPP assay for well water testing would have to be further scrutinized.
Restricting the use of well water that tests positive by the COPP assay could reduce or prevent morbidity and mortality from water-borne pathogens. The COPP assay could become a first line of defense in preventing contaminated well water from reaching potential consumers. Restricting the consumption of polluted water could reduce the threat of deadly epidemics. By virtue of the number of COPP-positive colonies detected on TSA-N plates, the assay provides quantitative results, which may be compared over time to determine improving or worsening water quality. In areas where sewage and water infrastructure are being restored, the COPP assay would also allow health authorities to evaluate improvements or failures in water handling systems.
This article shows proof in principle that the COPP assay can detect V. cholerae and A. hydrophila in well water. Further testing is required to determine if the assay is applicable for testing well water in locations around the globe, where different water qualities and microbial flora abound. The simplicity and low cost of the COPP assay will facilitate studies to evaluate the effectiveness of the procedure for regions where cholera is endemic.
Received January 19, 2006. Accepted for publication May 8, 2006.
Disclaimer: Mention of trade names or commercial products in this publication is solely for the purpose of providing specific information and does not imply recommendation or endorsement by the U.S. Department of Agriculture.
* Address correspondence to Gary P. Richards, United States Department of Agriculture, Agricultural Research Service, Microbial Food Safety Research Unit, Delaware State University, Dover, Delaware 19901. E-mail: grichard{at}desu.edu 
Authors’ addresses: Gary P. Richards and Michael A. Watson, USDA, ARS, Delaware State University, James W.W. Baker Center, Dover, Delaware 19901, E-mail: grichards{at}errc.ars.usda.gov.
Reprint requests: Gary P. Richards, USDA, ARS, Delaware State University, James W.W. Baker Center, Dover, DE 19901, E-mail: grichards{at}errc.ars.usda.gov.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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This article has been cited by other articles:
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  G. P. Richards, M. A. Watson, E. J. Crane III, I. G. Burt, and D. Bushek Shewanella and Photobacterium spp. in Oysters and Seawater from the Delaware Bay Appl. Envir. Microbiol., June 1, 2008; 74(11): 3323 - 3327. [Abstract] [Full Text] [PDF]
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