Environmental and epidemiological surveillance of Vibrio cholerae in a cholera-endemic region in India with freshwater environs

Authors


Neelam Taneja, Department of Medical Microbiology, Post Graduate Institute of Medical Education and Research, Chandigarh 160012, India. E-mail: drneelampgi@yahoo.com

Abstract

Aim:  To conduct epidemiological and ecological surveillance of cholera in freshwater environments.

Methods and Results:  A freshwater region of India was surveyed between April 2007 and December 2008. Vibrio cholerae was isolated from 59·5% of water and plankton samples (n = 357) and 35·5% of stool samples (n = 290). Isolation from water was dependent on air (r = 0·44) and water temperatures (r = 0·49) (P < 0·01) but was independent of rainfall (r = 0·15), chlorophyll a (r = 0·18), salinity (r = 0·2) or pH (r = 0·2) (P > 0·05). Isolation from plankton was dependent on temperature of air (r = 0·45), water temperature (r = 0·44), chlorophyll a concentration (r = 0·42), pH (r = 0·23) and salinity (r = 0·39) (P < 0·01). Cholera cases correlated with rainfall (r = 0·82, P < 0·01) and chlorophyll a concentration (r = 0·42, P < 0·05), but not with air temperature (r = 0·3, P = 0·37). Vibrio cholerae O1 possessed ctxB, ctxA, rstR and tcpA (ElTor), toxR, toxT, rtxA, rtxC, mshA and hylA. Among non-O1–non-O139, the distribution of virulence-associated and regulatory protein genes was heterogeneous with – 0·7, 2·2, 94·77, 97·76, 99·25, 100 and 100% isolates being positive for tcpA, toxT, rtxA, rtxC, hylA, toxR and mshA, respectively. Two-thirds of non-O1–non-O139 isolates exhibited antibiotic resistance to various antibiotics that did not correlate with geographical site or time of origin for the isolates. RAPD and AFLP showed V. cholerae to be a diverse bacterium. AFLP demonstrated separate lineages for non-O1–non-O139 and O1 isolates.

Conclusion:  Environmental parameters played a significant role in the emergence and spread of cholera and the abundance of V. cholerae. But based on virulence gene profiling and genetic fingerprinting, the possibility of origin of toxigenic isolates from nontoxigenic environmental isolates seems unlikely in freshwater environs of India.

Significance and Impact of the Study:  This study explains the ecology, epidemiology and seasonality of cholera in freshwater environs.

Introduction

Vibrio cholerae, a facultative human pathogen, can survive and thrive in aquatic environment. More than 200 serogroups (O1–O200) exist (Chatterjee and Chaudhuri 2004), but only toxigenic strains of the serogroups O1 and O139 cause cholera (Nelson et al. 2009). Non-O1–non-O139 serogroups are indigenous members of the aquatic ecosystem and are generally nonpathogenic (Faruque et al. 1998). Biological and environmental factors play a significant role in the dynamics of cholera. Since the seminal work by Colwell et al. (1977), several investigators have traced the potential environmental ecological niche where vibrios persist and flourish. Several biotic and abiotic factors, viz., salinity, temperature, rainfall and plankton, have been proven as important factors in the ecology of V. cholerae that influence the transmission of the disease (Huq et al. 2005; Constantin de Magny et al. 2008).

Vibrios are well adapted to the aquatic environment and can exist as a free-living form in a planktonic phase or as a particle-associated epibiotic form with phytoplanktons and zooplanktons (Janda 1998). Upon exposure to an unfavourable environment, vibrios persist by entering a viable but nonculturable (VBNC) state (Xu et al. 1982). Humans are only incidental and transient hosts, promoting dissemination into water resources in unsanitary conditions. Upon ingestion, pathogenic V. cholerae colonize the small intestine, multiply, secrete cholera toxin (CT) and are released into the environment by diarrhoeal stools of the hosts (Merrell et al. 2002). The pathogenic strains that belong to serogroups O1 and O139 produce CT and the colonization factor known as toxin-coregulated pilus (TCP). In contrast, more than 95% of non-O1–non-O139 serogroups lack these major virulence factors (Leclerc et al. 2002). However, there are other accessory virulence and regulatory genes, including toxR and toxT (genes that encode regulators), hylA (haemolysin) (Rivera et al. 2001), rstR (regulator of phage lysogeny) (Nusrin et al. 2004), rtxA (cytotoxin) and rtxC (acyltransferase) (Lin et al. 1999), which encode for cytotoxic activity that has been demonstrated in Hep-2 cells, mshA (mannose-sensitive haemagglutinin) (Thelin et al. 1996) and NAG-ST (heat-stable enterotoxin) (Arita et al. 1986). Recent studies have shown that these virulence genes are also dispersed among diverse serogroups that constitute the environmental reservoir (Rivera et al. 2001) (Faruque et al. 2004). These environmental strains are believed to be precursors of pathogenic strains. Therefore, environmental monitoring for the presence of V. cholerae strains with pathogenic potential is important to identify the source of strains that cause either epidemics of cholera or sporadic episodes of gastroenteritis.

Cholera is endemic in northern India, which includes Punjab, Haryana and Chandigarh. It has a freshwater environment with salinity close to zero and subtropical climate conditions. This region experiences hot, humid summers and chilly winters with temperatures falling as low as 0°C (32°F). The annual rainfall is moderate (<1000 mm). Cholera occurs in certain pockets of this region in seasonal outbreaks. The disease is on the rise in this part of India, with frequent sporadic cases and outbreaks (Taneja et al. 2003, 2005, 2009, 2010), particularly with a peak only during the monsoon months. The ecology of V. cholerae in freshwater environs is poorly understood, making it difficult to gauge the exact incidence of disease. Therefore, the aim of this study was to investigate the endemic existence of V. cholerae in freshwater environments in northern India during the various seasons. Epidemiological and environmental surveillance was undertaken, and we attempted to correlate the prevalence of V. cholerae in the environment and clinical cholera cases with physical parameters, viz., air and water temperature, rainfall, chlorophyll, pH and salinity of freshwater bodies. The relationship between clinical and environmental isolates was studied by virulence gene profiling and genetic fingerprinting using randomly amplified palindromic DNA (RAPD) and amplified fragment length polymorphism (AFLP). The antibiotic susceptibility patterns of isolates were also compared.

Materials and methods

Study area

Surveillance was conducted in and around Chandigarh, India (Latitude: 30°43′N, Longitude: 76°47′E). Vibrio cholerae were isolated from environmental and clinical samples and divided into the following categories:

  • Category I: Water and plankton were collected biweekly from eight fixed sites (Fig. S1) for a period of 1 year (April 2007–March 2008).

  • Category II: Samples were also collected randomly during the same time period from natural bodies of freshwater, including major rivers in North India (Satluj, Yamuna, Beas and Ganges) and freshwater lakes, namely Gobind Sagar (Punjab) and Karna (Karnal, Haryana).

  • Category III: Samples of water from hand pumps, taps, tube wells, wells and sewage were collected from the cholera outbreak-affected areas from Mohali, Kurali, Balongi and Madanpura (Punjab) and Rally, Panchkula, Ambala and Noorpur (Haryana) between April 2007 and December 2008.

Sample collection

Water samples were collected in sterilized bottles, and 1·0 l was filtered through 0·22-μm filter membranes (Millipore, Bethesda, MD, USA) using vacuum pressure of 15–20 lb in−2. Subsequently, membranes were washed with 6 ml of phosphate-buffered saline (PBS, pH 7·4). Plankton samples were collected by filtering 20·0 l of water through plankton nets (25 μm) that were previously disinfected with 70% ethyl alcohol. Plankton samples were washed with PBS and concentrated in sterile bottles (final volume, 20 ml). A 10-ml sample of the concentrated plankton was preserved by the addition of formaldehyde (4%, v/v) for identification.

The following physical parameters were measured: salinity by using a hand refractometer (Erma, Tokyo, Japan), temperature by using a laboratory thermometer and pH by using a portable pH meter (Fisher Scientific, Ottawa, Canada) in situ. Chlorophyll a was measured by standard spectrophotometric method (Standard Methods 1999). Weekly rainfall and air temperature data for the northern region were taken from the Directorate of Meteorology, Air Force Headquarters, New Delhi, India.

Stool samples in Carry-Blair were collected from peripheral hospitals in Punjab and Haryana from cholera-affected patients between April 2007 and December 2008. This work was approved by the ‘Institute Ethical Review Committee’ (Ref no. MS/354/PhD/11037).

Vibrio cholerae isolation

Concentrate (3·0 ml) from water and plankton samples was added to alkaline peptone water (APW, pH 8·6). After incubation for 6 h at 37°C, subcultures were plated onto selective thiosulfate citrate bile salt sucrose agar. Single flat, circular, yellow, sucrose-fermenting colonies that are typical for V. cholerae were transferred to sheep blood agar (10%). Presumptive isolates growing on blood agar were identified by conventional biochemical reactions (Old 1996) and genotypically confirmed by species-specific PCR for the ompW gene (Nandi et al. 2000). Isolates were serologically tested with O1 and O139 antiserum (Denka Seiken, Tokyo, Japan).

DFA-DVC for VBNC detection

Environmental samples were investigated for the presence of V. cholerae O1 VBNC cells as previously described (Mishra et al. 2011). Briefly, 1·0 ml of sample concentrate was added to 9·0 ml of 10·0% Tryptic Soy broth containing 0·015 μg ml−1 ciprofloxacin. Samples were incubated for 8 h at room temperature, later fixed with 2·0% formaldehyde and then subjected to Cholera O1 DFA kit (New Horizons Diagnostics Corp., Columbia, MD) and examined under fluorescent microscope (Model BX51; Olympus, Tokyo, Japan). The viable bacterium was appreciated as a cell with increased length (≥1·5-fold) with respect to the initial length.

Antibiotic susceptibility testing

The disc diffusion method was employed following CLSI guidelines. The following antimicrobial agents were tested: nalidixic acid (30 μg), norfloxacin (10 μg), ciprofloxacin (10 μg), ofloxacin (5 μg), gentamicin (10 μg), amikacin (30 μg), cotrimoxazole (25 μg), chloramphenicol (30 μg), neomycin (30 μg), cefotaxime (30 μg), ceftriaxone (30 μg), cefoperazone-sulbactum (30 μg), tetracycline (30 μg) and amoxicillin (10 μg) (Oxoid, Hampshire, UK).

Molecular characterization of isolates

Genomic DNA was extracted as previously described (Ausubel et al. 2002). The following DNA sequences were targeted with previously known primer sequences: ctxB (Morita et al. 2008), rfbO1 and rfb O139 (Hoshino et al. 1998), ctxA and tcpA (Keasler and Hall 1993), toxR, toxT (Mukhopadhyay et al. 2001), sto/stn, hylA (Rivera et al. 2001), rstR (Nusrin et al. 2004), rtxA, rtxC (Chow et al. 2001) and mshA (Jouravleva et al.1998). The reference strains that were used were V. cholerae Classical 569B (Classical), V. cholerae O1 ElTor N16961 (ElTor), V. cholerae O1 hybrid (Hybrid), V. cholerae non-O1–non-O139 NT5394 (NT5394), V. cholerae O139 and V. cholerae O1 MTCC 3906 (MTCC).

Genetic fingerprinting

RAPD was performed using the following 10-mer primers, as previously described (Leal et al. 2004): P1, 5′-AAGAGCCCGT-3′; P2, 5′-GCGGAAATAG-3′; and P3, 5′-CCGCAGCCAA-3′. P3 was the most discriminatory primer and was used for RAPD of all isolates. For AFLP, DNA was double-digested with EcoRI and MseI and selectively amplified using the primers 5′-GACTGCGTACCAATTCG-3′ and 5′-GATGAGTCCTGAGTAACT-3′ as previously described (Vos et al. 1996). Amplified products were separated on a 6% denaturing polyacrylamide gel using SequiGen gel apparatus 38 × 50 × 0.4 cm (Bio-Rad Laboratories Inc., Hercules, CA, USA) and developed by silver staining (Promega, Madison, WI, USA). RAPD and AFLP gels were manually scored for the presence and absence of bands and evaluated with Jaccard coefficient of similarity using NTYsysPC version 2.003e Applied Biostatics Inc. The dendrogram was constructed using upgma (Sneath and Sokal 1973). Simpson’s coefficient of diversity (D) was calculated by using the formula = 1 − {∑[nj(nj − 1)]}/[N (− 1)], where N is the number of strains tested and nj is the number of strains belonging to jth type (Hunter and Gaston 1988).

Statistical analysis

The Spearman’s rank test was performed for correlation analysis. The Mann–Whitney U-test, anova and post hoc test were used to identify statistical differences. Results were accepted as significant at a probability of ≤0·05.

Results

Identification of Vibrio cholerae

A total of 357 environmental (water and plankton) and 290 stool samples from suspected cholera patients were collected and processed. From environmental samples, 219 sucrose-fermenting colonies were obtained, and 147 (67%) were confirmed as V. cholerae, all from unique samples. The combination of results from oxidase, arginine dihydrolase, string test and cholera red reaction, with individual specificity between 18 and 100%, was sufficient for the identification of environmental V. cholerae isolates (Table S1). Only 13 isolates (3·64%) isolated from the samples collected from cholera-affected areas were confirmed as O1. From the stool samples, 99 isolates were confirmed as V. cholerae O1 Ogawa.

Distribution of Vibrio cholerae and correlation with environmental parameters and clinical cholera

Vibrio cholerae non-O1–non-O139 were isolated from 49 and 34·83% of water and plankton samples, respectively, from the fixed sites. Random sites that included large rivers showed comparatively high isolation percentages of 80·5 and 73·33% for non-O1–non-O139 from water and plankton, respectively. In addition, 17·94 and 33·3% of water samples collected from cholera-affected areas were positive for both non-O1–non-O139 and V. cholerae O1 during 2007 and 2008. Table 1 shows the distribution of percentage isolation of V. cholerae from different types of samples. Interestingly, V. cholerae O1 was not isolated from any fresh or random body of water; however, 11·11% of water samples collected from the cholera-affected regions were positive for V. cholerae O1 Ogawa.

Table 1.   Isolation of Vibrio cholerae from water and plankton samples from fixed sites, random sites, cholera-affected areas and patient stool samples
Sample categorySample typeTotal samplesIsolates (%) of V. choleraeO1 Serogroup
Environmental samplesFixed sitesWater100 49 (49)0
Plankton89 31 (34·83)0
Random sitesWater36 29 (80·55)0
Plankton15 11 (73·33)0
Cholera-affected regionsWater (2007)78 14 (17·94)1
Water (2008)39 13 (33·33)12
 357147 (41·1)13
Clinical samplesSporadic/outbreakStool (2007)15251 (33·5)51
Sporadic/outbreakStool (2008)138 48 (34·78)48

The various physical parameters that were monitored during sampling are listed in Table 2. These parameters fell within the following ranges: water temperature (Tw), 9–33°C; air temperature (Ta), 19·1–38°C; pH, 5·25–7·5; salinity, 0–0·3 ppt; chlorophyll a, 3·81–73·46 μg l−1; and rainfall, 2·2–272·2 mm. The number of cholera cases during the different months, variations in physical parameters and percentages of isolation of V. cholerae from fixed sites are shown in Fig. 1. Vibrio cholerae was isolated from the environment significantly more often during the summer (April–June) and monsoon seasons (July–September) compared with the winter season (October–March) (P < 0·05). Isolation during the summer and monsoon seasons was similar and significantly decreased with a drop in temperature to <16°C in winter (P < 0·05).

Table 2.   Physical parameters that were monitored in this study from April 2007 to March 2008
VariablesSeasonNumber of observationsMedian valueMinimum valueMaximum value
Water temperature (Tw°C)Summer18·0 27·9325·5 31·5
Monsoon24·0 26·022·0 33·0
Winter30·0 16·0 9·0 25·0
Air temperature (Ta°C)Summer 3·0 37·636·0 38·0
Monsoon 4·0 33·132·5 34·4
Winter 5·0 22·419·1 31·8
Rainfall (mm)Summer 3·0  6·0 4·8180·2
Monsoon 4·0 98·8 2·2272·2
Winter 5·0  7·6 2·2 13·0
pHSummer18·0  6·5 5·25  7·0
Monsoon24·0  6·25 6·5  7·0
Winter30·0  6·25 6·5  7·0
Chlorophyll a (μg l−1)Summer15 24·7812·26 64·64
Monsoon15 54·4222·43 73·46
Winter26 11·53 3·81 32·76
Salinity (ppt$) all seasons 72·2  0·1 0  0·3
V. cholerae% isolationSummer69·0100 0100
Monsoon46·5 50 0100
Winter15·5 20 0100
Figure 1.

 Prevalence (%) of cholera cases occurring each month between April 2007 and March 2008 and changes observed in water temperature [Tw (°C)], air temperature [Ta (°C)], rainfall (cm), chlorophyll a (μg l−1) and the percentage isolation of Vibrio cholerae from environment. (inline image) Clinical cholera (% cases); (inline image) Ta (°C); (inline image) Tw (°C); (inline image) Rainfall (cm); (inline image) V. cholerae (% isolation) and (inline image) Chlorophyll a (μg l−1).

Correlation coefficients were calculated for each site between the percentage isolation of non-O1–non-O139 from the water and plankton samples and physical parameters over different months. There was a significant correlation between percentage isolation and water (Tw) and air temperatures (Ta) for Saketri (Ta, 0·72; Tw, 0·70), Sukhna Choe (Ta, 0·81; Tw, 0·70) and Ghaggar (Ta, 0·74; Tw, 0·70) (P < 0·05). In addition, there was a significant correlation between percentage isolation and water temperature only for Sukhna Lake (Tw, 0·76) and Derrabasi (Tw, 0·58) (P < 0·05). No non-O1–non-O139 isolates were detected in water or plankton from Jayanti Devi pond during any season of the year. Similarly, for plankton isolation, a significant correlation was observed between percentage isolation and Tw, Ta and chlorophyll a for Saketri (Ta, 0·5; Tw, 0·6; C, 0·8), Sukhna Lake (Ta, 0·71; Tw, 0·63; C, 0·34) and Sukhna Choe (Ta, 0·78; Tw, 0·73; C, 0·44) (P < 0·05). The details of sitewise correlations between percentage isolation of V. cholerae and various physical parameters are shown in Figs S2–S4. Overall, the isolation of non-O1–non-O139 samples from water showed a significant correlation with air (r = 0·44) and water temperatures (r = 0·49) (P < 0·01). In contrast, no significant correlation was observed with rainfall (r = 0·14, P = 0·24), chlorophyll a (r = 0·18, P = 0·19), salinity (r = 0·2, P = 0·083) and water pH (r = −0·36, P = 0·72). Percentage isolation of non-O1–non-O139 from plankton showed a significant correlation with air temperature (r = 0·45), water temperature (r = 0·441), chlorophyll a (r = 0·42), water pH (r = 0·23) and salinity (r = 0·39) (P < 0·01). Upon microscopic examination of plankton samples, a majority of the phytoplankton population was constituted by Anabaena spp. and Microcystis spp. (blue-green algae). Among zooplanktons, the predominant species were Macrocyclops (freshwater copepod), Oithona spp., Rotifers spp. and Euphausia spp. Almost 23·46% of samples were positive for the presence of zooplanktons, and 17% were positive for phytoplanktons. A significant correlation was observed (r = 0·3) between chlorophyll a (plankton bloom) and an increase in the isolation percentage for non-O1–non-O139 from planktons (P < 0·005).

A total of 51 confirmed cholera cases occurred in between April 2007 and 2008, with a peak during the rainy season (July–September). Clinical cholera cases correlated with rainfall (r = 0·82, P < 0·01), chlorophyll a (r = 0·42, P < 0·05) and air temperature (r = 0·3, P = 0·37). A similar incidence curve was observed in 2008 for cholera, with 48 confirmed cholera cases.

Molecular characterization of isolates

The distribution of different virulence and regulatory genes in V. cholerae that are circulating in cholera-endemic areas is summarized in Table 3. Vibrio cholerae O1 isolates from outbreak-affected waters (n = 13) and all clinical O1 isolates (n = 99) were positive for ctxB (classical), ctxA, rfbO1, rstR and tcpA (ElTor type), toxR, toxT, rtxA, rtxC, mshA and hylA. Of the non-O1–non-O139 environmental isolates (n = 134), 0·7, 2·2, 94·77, 97·76, 99·25, 100 and 100% were positive for tcpA (ElTor), toxT, rtxA, rtxC, hylA, toxR and mshA, respectively. None of the O1 or non-O1–non-O139 isolates that were tested were positive for the sto/stn gene.

Table 3.   Distribution of genes studied in Vibrio cholerae from cholera-endemic area
OriginSerogroupIsolates
n
Virulence-associated and other genes
ompWctxActxB (C*/E*)tcpA (C/E)Rfb (O1/O139)toxRtoxTrstR (C/E)Sto/stnrtxArtxChylA (E)mshA
  1. *V. cholerae: E, ElTor; C, Classical.

EnvironmentO113+++/−−/++/−++−/+++++
Non-O1–non-O139124++−/−++++
5++−/−+++
1+−/+++−/−++++
1+++−/−++++
1+++−/−++++
1++−/−++
1++−/−+
ClinicalO199+++/−−/++/−++−/+++++

Antibiotic susceptibility testing

A total of 36 antimicrobial resistance profiles were observed among environmental and clinical isolates (Table 4). Of the 147 environmental isolates, 45 were susceptible to all of the antibiotic agents tested and 102 isolates were resistant to one or more antibiotics. The percentages of resistance to the antimicrobial agents amikacin, tetracycline, neomycin, norfloxacin, cefoperazone–sulbactam, nalidixic acid, cotrimoxazole and amoxicillin were 7·45, 8·0, 9·3, 10·5, 21·7, 24·2, 28·5 and 35·4, respectively. Most of the clinical isolates were sensitive to antimicrobials, with the exception of cotrimoxazole and nalidixic acid, to which all isolates were uniformly resistant. In addition, 66·7% of isolates from 2008 were also resistant to tetracycline.

Table 4.   Antibiotic resistance profiles that were identified in Vibrio cholerae*
Pattern No.AMKAMXCFP:SULCIPCHLCOTCTXGENNALNEONOROFXTETNo. of isolates (%)
  1. COT, cotrimoxazole; CTX, cefotaxime; AMX, amoxicillin; CFP:SUL, cefoperazone–sulbactam; GEN, gentamicin; AMK, amikacin; NEO, neomycin; TET, tetracycline; CHL, chloramphenicol; NOR, norfloxacin; CIP, ciprofloxacin; NAL, nalidixic acid; OFX, ofloxacin.

  2. *V. cholerae : E, environmental isolates; C, clinical isolates.

1SSSSSSSSSSSSS45[E] (30·61)
2SSSSSRSSRSSSS69[C] (65·71)
11[E] (7·48)
3SSSSSRSSRSSSR36[C] (34·28)
1[E] (0·68)
4SRSSSSSSSSSSS14[E] (9·52)
5SSSSSRSSSSRSR9[E] (6·12)
6SRRSSSSSSSSSS8[E] (5·44)
7SSSSSSSSRSSSS7[E] (4·7)
8SSSSSRSSSSSSS5[E] (3·4)
9SSRSSSSSSSSSS4[E] (2·72)
10SRSSSRSSRSSSS3[E] (2·0)
11RSSSSRSSRSSSS3[E] (2·0)
12SRRSSRSSRSSSS3[E] (2·0)
13SRSSSSSSRSSSS3[E] (2·0)
14RSSSSSSSSSSSS2[E] (1·36)
15SRRSSSSSRSSSS2[E] (1·36)
16RSSSSRSSRSSSS2[E] (1·36)
17RSSSRSSSSSSSS2[E] (1·36)
18SRSSSSSSSRSSS2[E] (1·36)
19RRRSSSSSSRSSS2[E] (1·36)
20RSSSSSSSRSSSS1[E] (0·68)
21SSRSSSSRSSSSS1[E] (0·68)
22SSSSSSSSRRSSR1[E] (0·68)
23RSRSSSSSSRSSS1[E] (0·68)
24SRRSSSSSSRSSS1[E] (0·68)
25RSSSSRSSSRSSS1[E] (0·68)
26SRSSSSSSRRSSS1[E] (0·68)
27SRRSSSSSRSRSS1[E] (0·68
28SRRSSRSSSRSSS1[E] (0·68)
29RRRSSSSRSSSSS1[E] (0·68)
30SRSRSSSSRSRSS1[E] (0·68)
31SRRSSRRSRSSSS1[E] (0·68)
32RRSSSSSSSSSSS1[E] (0·68)
33SSSSSRSSRSRSS1[E] (0·68)
34SRRRSSRSSRSSS1[E] (0·68)
35SRSSSRSSRRSRR1[E] (0·68)
36SSRRSRSSRRRRR1[E] (0·68)

RAPD Fingerprinting

A total of 251 V. cholerae isolates were subjected to RAPD (99 clinical isolates, 147 environmental isolates and five reference strains). The RAPD pattern differentiated V. cholerae isolates into 46 types, A-T1 (Table 5). The most common pattern was designated type A (149 isolates), followed by type D (13 isolates), type I1 (nine isolates), type H (seven isolates) and type H1 (five isolates). The largest, type A contained 149 isolates: 96 clinical isolates, 49 environmental isolates and four control strains (ElTor, MTCC, hybrid and NT5394). The environmental isolates included 19 from fixed sites, 16 from outbreak sites and 14 from large natural bodies of water. The V. cholerae O1 Classical 569B control strain exhibited a unique pattern (type B) that was not found among the other isolates. Overall, there was no correlation between serotype and geographical origin of the isolates; 115 O1 isolates (102 clinical and 13 environmental) belonged to type A, but 37 non-O1–non-O139 isolates of environmental origin also belonged to this type. Three O1 isolates exhibited a different RAPD pattern and were designated as type C. The variability among the RAPD patterns of environmental isolates was very high, as almost 23 isolates exhibited unique patterns. The Simpson’s index of diversity for clinical and environmental samples was 0·105 and 0·778, respectively.

Table 5.   Distribution of Vibrio cholerae isolates in 46 RAPD types (A-T1)
S. no.No. of isolatesTypeOrigin/serogroup
  1. *V. cholerae : E, environmental isolates; C, clinical isolates.

1149AO1 (C* = 96; E* = 13), reference strains (4), non-O1–non-O139 (36)
213Dnon-O1–non-O139
39I1non-O1–non-O139
47Hnon-O1–non-O139
55H1non-O1–non-O139
64N, G1, K1non-O1–non-O139
73C, V, J1O1 (type C), others non-O1–non-O139.
82Q, R, S, W, Y, Z, A1, C1, D1, E1, L, M1non-O1–non-O139
91B, E, F, G, I, J, K, L1, M, O, P, T, U, X, B1, F1, N1, O1, P1, Q1, R1, S, TO1 Classical (type B), others non-O1–non-O139

AFLP fingerprinting

A total of 52 isolates were subjected to AFLP, including clinical (n = 13), environmental (n = 34) and reference (n = 5) strains. These isolates were selected based on their RAPD profiles, with 24 isolates from RAPD type A and representative isolates from other types. A total of 68 polymorphic bands, ranging in size from 650 to 150 bp, were scored. A majority of the isolates were divided into two main subclades, subclades 1 and 2, with a similarity index of 0·58 (Fig. 2). Subclade 1 (n = 26) included environmental non-O1–non-O139 isolates and a single O1 isolate. Vibrio cholerae isolates from different geographical locations intermingled in this cluster. Subclade 2 (n = 18) consisted of all O1 clinical and environmental isolates. Apart from above two subclades, eight isolates exhibited independent singletons. The isolates exhibiting similar RAPD patterns were clearly differentiated by AFLP patterns. Overall, the AFLP analysis showed a high level of diversity among environmental isolates, as 32 of 34 isolates exhibited distinguishable AFLP patterns. The Simpson’s index of diversity for AFLP analysis was 0·99.

Figure 2.

 Dendrogram of 52 Vibrio cholerae isolates after pairwise comparison of AFLP patterns generated using EGMT primer combination.

Discussion

Although studied in detail in coastal areas, the epidemiology of cholera remains enigmatic (Huq et al. 2005; Constantin de Magny et al. 2008). The reasons for seasonal cholera and the influence of specific environmental factors on the emergence of this disease have not been explored in noncoastal areas with freshwater environments. The northern region of India, which is located far away from the sea, has witnessed a recent resurgence of cholera, with outbreaks occurring because of V. cholerae O1 Ogawa infections (Taneja et al. 2003, 2005, 2009, 2010). The aims of this study were to monitor the presence of V. cholerae strains with pathogenic potential in freshwater environments in this region and to understand the environmental factors responsible for seasonality of the disease, which coincides with hot and humid months from June to October. Environmental surveillance was conducted at fixed sites, including ponds, lakes and small rivers around Chandigarh. Major rivers and freshwater lakes of North India were included as controls to remove any bias in the selection of the fixed sites. Samples were also collected from drinking water and sewage supplies of cholera outbreak-affected areas during the same period. Vibrio cholerae was isolated from 59·5% of freshwater samples; however, all V. cholerae (n = 120) that were isolated from fresh bodies of water, viz., fixed and random sites, were non-O1–non-O139. Of 117 samples from cholera outbreak-affected areas, 25% were positive for V. cholerae, of which 11·1% were serogroup O1 Ogawa isolates. All clinical isolates (n = 99) were confirmed as V. cholerae O1 Ogawa. Vibrio cholerae O139 was not isolated from any environmental or clinical samples. Viable but nonculturable V. cholerae O1 were also detected in 2·21 and 40·69% of samples from natural bodies of water and cholera-affected areas, respectively, signifying their presence in these waters (Mishra et al. 2011). The absence of culturable V. cholerae O1 and the high prevalence of non-O1–non-139 isolates in natural waters were in congruence with another ecological study from Vellore, India (Jesudason et al. 2000).

The influence of environmental parameters, viz., temperature, pH, salinity, rainfall and chlorophyll, on the abundance of V. cholerae is well established in marine environments (Pardio Sedas 2007). In freshwater, environmental parameters were observed to have a significant effect on the isolation of V. cholerae, which are considered aboriginal members of the aquatic ecosystem (Xu et al. 1982). Among abiotic factors, salinity was constant throughout the year, while other parameters, including air and water temperatures, chlorophyll, rainfall and pH, were variable. A significant difference was observed in the isolation of V. cholerae in different seasons, which peaks during summers (69%) and monsoons (46·5%) and is minimal in winters (15·5%). Thus, temperature directly influences the abundance of the organism in the environment. Isolation of non-O1–non-O139 from water and planktons significantly correlated with air and water temperature but was independent of rainfall. However, there was a significant correlation between the isolation of non-O1–non-O139 from planktons and both pH and chlorophyll concentration. Plankton blooms were measured by an increase in chlorophyll a concentration. Although initial isolation from water showed a parallel increase with chlorophyll a concentration (Fig. 1), there was no significant correlation between the isolation of non-O1–non-O139 from water and plankton blooms. In fact, when the chlorophyll concentration peaked, the isolation of non-O1–non-O139 from water dropped, may be due to the accumulation of toxic algal products. The plankton population was constituted by similar phytoplanktons and zooplanktons in different sites. The phytoplankton included mainly freshwater blue-green algae, which resulted in bright green blooms and a further increase in the zooplankton population, which in turn resulted in an increase in the isolation of V. cholerae. The pH of water significantly increased with plankton blooms, although the actual changes were small (6·5–7·0). Nevertheless, there was a significant correlation between the plankton bloom, which governs pH changes, and an increase in isolation percentage for V. cholerae from planktons. Therefore, environmental parameters influence the isolation and hence the abundance of non-O1–non-O139 in fresh bodies of water.

Clinical cholera cases coincided with elevated rainfall, chlorophyll concentration and air temperature. On multivariate regression analysis, rainfall was found to be an independent predictor for outbreaks of cholera, whereas elevated temperature had a significant effect only if combined with rainfall. Chlorophyll a also exhibited a significant correlation with the occurrence of cholera outbreaks. These findings might therefore explain the seasonal variation of the disease coinciding with the monsoon months. During winter months (November–February), the temperature dropped to below 16°C and the isolation of non-O1–non-O139 dropped significantly. No cases of cholera occurred during the winters. We observed a lag period of 1 month between the appearance of clinical cases of cholera and the increase in temperature, isolation of non-O1–non-O139 from the environment and chlorophyll concentration. We hypothesize that failure to isolate V. cholerae O1 from freshwater samples may be due to either the high prevalence of non-O1–non-O139, which are believed to be indigenous members of the aquatic ecosystem that mask the isolation of pathogenic strains, or the presence of the organism in a VBNC state, thereby escaping detection by culture methods, which we were able to demonstrate (Mishra et al. 2011). Either way, at the onset of favourable conditions of increased temperature and nutrition in the form of planktons, V. cholerae increased in number, leading to the appearance of cholera cases. Later, the rains lead to the runoff of water and the overflow of sewage. These factors lead to increased chances of contamination by seepage of water; indeed, in our study, water from hand pumps and tube wells was found to be contaminated with V. cholerae O1. Later, the outbreak is driven by the contamination of drinking water supplies, with increasing cholera cases leading to more pollution. Rainfall is known to have an effect on the appearance of cholera outbreaks (Pardio Sedas 2007). The positive association between counts of V. cholerae O1, water temperature 2 months earlier and the appearance of cholera cases was demonstrated in a study in Peru (Franco et al. 1997). In addition, the correlation between cholera outbreaks and algal blooms was previously reported (Colwell 1996); however, there is no direct evidence that this correlation leads to an enrichment of V. cholerae O1 strains that are responsible for cholera cases. Similarly, in this study, a correlation between chlorophyll a (an indicator of algal blooms) and cholera cases was observed; however, toxigenic V. cholerae O1 was not isolated from the freshwater environment, with the exception of cholera-affected areas.

All V. cholerae O1, whether isolated from clinical or environmental samples, possessed ctxA, ctxB (classical type), tcpA and rstR (ElTor type). This finding confirmed the presence of ElTor variants (Safa et al. 2010) in our region. Overall, non-O1–non-O139 isolates were free of CTXΦ phage, as both ctxA and rstR were absent. The absence of another horizontal genetic element, VPI, in the majority of non-O1–non-O139 isolates was confirmed; tcpA and toxT, which are components of VPI, were present only in one and three non-O1–non-O139 isolates, respectively. Such tcpA+ strains are believed to be selected in human intestines and become pathogenic upon transduction with CTXΦ lysogenic phage (Faruque et al. 1998). Merrell et al. (2002) proposed that pre-epidemic amplification of ctxA- and tcpA-positive V. cholerae occurs in the human host, which leads to the start of an epidemic cycle of V. cholerae that spreads rapidly through environmental waters that are otherwise free of pathogenic isolates. Among the other accessory genes studied, toxR, rtxA, rtxC and hylA showed an almost equal distribution in environmental and clinical isolates, in agreement with previous studies (Rivera et al. 2001; Faruque et al. 2004) that depicted the environmental repertoire of accessory virulence genes. Similar to our study, previous studies in other parts of the world have shown fairly low or no isolation of V. cholerae O1 (Jesudason et al. 2000; Rivera et al. 2001; Faruque et al. 2006). However, Faruque et al. (2004) demonstrated that, in Bangladesh, strains that were isolated by conventional culture methods were mostly (99·2%) negative for the major virulence genes tcpA and ctxA and were not pathogenic in animal models. In contrast, when water samples were directly enriched in rabbit intestines, O1 strains that were isolated were colonization-competent, and 56·8% of these strains carried genes encoding tcpA alone or both tcpA and ctxA. We demonstrated the presence of O1 strains in a VBNC state by DFA-DVC. Therefore, the freshwater environment of the cholera-endemic region around Chandigarh was shown to harbour pathogenic V. cholerae, although the isolation of V. cholerae O1 from the environment is a rare event because the environment contains relatively abundant nontoxigenic isolates.

Overall, the entire spectrum of the V. cholerae population in the environment of a cholera-endemic area was highly heterogeneous. The variability among RAPD patterns of environmental isolates was high, as almost 23 V. cholerae exhibited unique patterns. RAPD fingerprinting did not reveal any correlation within serotype or geographical origin of the isolates. The genetic profiles of toxigenic V. cholerae O1 from clinical cases and the environment were similar. Interestingly, a large number of ctxA, tcpA non-O1–non-O139 isolates exhibited RAPD profiles similar to toxigenic isolates, presumably because of the low discriminatory power of this technique. Recent studies on the genomic characterization of V. cholerae non-O1–non-O139, compared with O1 and O139, have shown that they are divergent and belong to distinct lineages. The evidence for this finding was provided by comparative genomic microarrays and sequencing of non-O1–non-O139 and serogroup O1 isolates, indicating that they are quite divergent (Dziejman et al. 2005; Chun et al. 2009). Therefore, to resolve the overlapping patterns of RAPD, we selected 52 V. cholerae isolates based on their RAPD profiles, 24 of which were RAPD type A, and subjected them to highly discriminatory AFLP fingerprinting. We compared the RAPD and AFLP profiles of these 52 isolates. Vibrio cholerae non-O1–non-O139, which exhibited similar profiles to clinical isolates, were clearly differentiated by AFLP. Non-O1–non-O139 strains were distinguishable from pathogenic O1 stains, as they did not possess any major virulence genes based on PCR analysis and exhibited different AFLP patterns that belong to a separate cluster. The V. cholerae O1 population was not clonal but closely related. Therefore, it appears that the majority of environmental nonpathogenic and pathogenic V. cholerae O1 isolates have distinct lineages, with the exception of one C8 isolate, which is an O1 strain that clustered with non-O1–non-O139 isolates. The possibility that toxigenic strains originate from nontoxigenic, non-O1–non-O139 isolates does not seem to be likely in our region.

Antibiotic resistance patterns fluctuate spatially and temporally and are very important to monitor. Two-thirds of environmental isolates were resistant to one or more (up to 8) antibiotics that were tested. One isolate each was resistant to 8, 6 and 5 antibiotics. Overall, environmental V. cholerae isolates exhibited varied resistant against amikacin, tetracycline, neomycin, norfloxacin, cefoperazone–sulbactam, nalidixic acid, cotrimoxazole and amoxicillin. No correlation was observed between the susceptibility patterns and geographical distribution of isolates. The O1 serogroup isolates were 100% resistant to cotrimoxazole and nalidixic acid, and 66% of the 2008 isolates were also resistant to tetracycline. It will be interesting in future to observe the role of Class 1 integrons and transferable elements, such as conjugative plasmids and transposons, in the carriage and dissemination of antimicrobial resistance among V. cholerae isolates.

This study was aimed at monitoring the influence of environmental parameters on the presence of V. cholerae strains in the environment. No culturable V. cholerae O1 or O139 were found in the environment during the inter-epidemic intervals. During the outbreaks, V. cholerae O1 could be isolated from 11·11% samples. Nonculturable V. cholerae O1 (VBNC) was demonstrated in 2·21 and 40·69% samples from natural water bodies and cholera-affected areas, respectively (Mishra et al. 2011). In comparison, the V. cholerae non-O1–non-O139 was isolated frequently, and environmental parameters, viz. temperature of water and air, chlorophyll and pH, had a significant role in their isolation. The cholera outbreaks also coincided with elevated rainfall, chlorophyll (plankton blooms) and temperature of water and air.

Next, antibiotic susceptibility testing, virulence genes profiling, RAPD and AFLP were carried out to compare V. cholerae isolates. The antibiotic patterns were not comparable and fluctuated irrespective of geographical origin, and non-O1–non-O139 were found to be devoid of main virulence factors –ctxA, ctxB and tcpA, barring one tcpA-positive isolate. The other virulence factors were evenly distributed in pathogenic and nonpathogenic isolates. Genetic fingerprinting analysis established that lineages for pathogenic and nonpathogenic isolates are different. Therefore, the potential of origin of pathogenic isolates from environmental isolates does not seem to be real in the freshwater environments of North India. However, the VBNC state could still be demonstrated in the environment (Mishra et al. 2011). Therefore, when a host comes in contact with V. cholerae O1 in the environment, it results in clinical cholera or sporadic cases. Combined with rainfall and poor sanitation conditions, this will result in further contamination of water sources, amplification of the organism and hence the beginning of an outbreak.

Acknowledgements

The help of Directorate of Meteorology, Air Force Headquarters, New Delhi, India, is acknowledged. The authors wish to thank Dr G.B. Nair, Dr T. Ramamurthy, NICED Kolkata and Dr Sabu Thomas, RGCB, Trivandrum, for providing reference strains. This study was partially financed by DST, Chandigarh, and PGIMER, Chandigarh.

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