Biofilm acts as a microenvironment for plankton-associated Vibrio cholerae in the aquatic environment of Bangladesh.
ABSTRACT The role of biofilm as a microenvironment of plankton-associated Vibrio cholerae was investigated using plexiglass as a bait. A total of 72 biofilm samples were tested using culture, direct fluorescent antibody (DFA) and molecular techniques following standard procedures. Culturable V. cholerae (smooth and rugose variants) were isolated from 33% of the samples. V. cholerae O1 were detected by FA technique throughout the year except April and June. All V. cholerae O1 isolates were positive for tcpA, ctxA and ace genes while V. cholerae non-O1, non-O139 isolates lacked these genes. V. cholerae O1 (both Inaba and Ogawa) strains had identical ribotype pattern (R1), but V. cholerae non-O1, non-O139 had different ribotype patterns. All V. cholerae O1 strains were resistant to vibrio-static compound (O/129). All V. cholerae O1 except one were resistant to trimethoprime-sulphamethoxazole, streptomycin, nalidixic acid and furazolidone but sensitive to ciprofloxacin, and tetracycline. This study indicates that plexiglass can act as a bait to form biofilm, a microenvironment that provides shelter for plankton containing V. cholerae in the aquatic environment of Bangladesh.
Article: Evidence of interspecies O antigen gene cluster transfer between Shigella boydii 15 and Escherichia fergusonii[show abstract] [hide abstract]
ABSTRACT: of interspe-cies O antigen gene cluster transfer between Shigella boydii 15 and Escherichia fergusonii. APMIS 2012. An environmental bacterial isolate, Iso10, previously found to show serological cross-reactivity with type-specific Shigella boydii 15 antisera was subjected to further molecular and serological analyses that revealed interspecies transfer of the O antigen gene cluster. Western blot analysis of Iso10 cell surface extracts and purified lipopolysaccharides demonstrated strong cross-reactivity with S. boydii 15-specific monovalent antisera and a lipopolysaccharide gel banding profile similar to that of S. boydii 15. Bio-chemical and phylogenetic analyses identified the Iso10 isolate as Escherichia fergusonii. O antigen gene cluster analyses of Iso10, carried out by restriction fragment length analysis of the amplified ~10-kb O antigen-encoding gene cluster, revealed a profile highly similar to that of S. boydii 15, confirming the presence of the S. boydii 15 somatic antigen in Iso10. To the best of our knowledge, this is the first report of interspecies transfer of O antigen-encoding genes between S. boydii and E. fergusonii, and it has implications for our understanding of the role of lateral gene transfer in the emergence of novel Shigella serotypes. Escherichia fergusonii, a member of the Entero-bacteriaceae family, is an infrequent but emerging human pathogen (1). E. fergusonii has been isolated from different types of clini-cal specimens, including blood, urine, faeces, and exudates from wound infections (2). In 2008, Savini et al. identified E. fergusonii as a potential enteric pathogen and reported multi-drug resistance patterns of various E. ferguso-nii strains isolated from a clinical specimen of a case of reported acute cystitis (1). E. ferguso-nii is also considered an important veterinary pathogen, with clinical manifestations of abor-tion, diarrhoea, and meningitis in affected sheep and cattle (3). More recently, septicae-mic distribution of E. fergusonii was noted in a goat with a history of wasting and diarrhoea (4). Despite the pathogenic potential of E. fer-gusonii, its habitat, pathogenic properties, and drug resistance mechanism are still unclear (1). DNA hybridization and biochemical analyses of E. fergusonii and Escherichia coli strains indicate a close relationship between the two genera (2). In addition, Fegan et al. (2007) reported the occurrence of the Escherichia coli O157 somatic antigen in E. fergusonii, suggest-ing a possible genetic transfer of the O antigen gene cluster (5). The O antigen can be used to classify differ-ent Gram-negative bacterial serotypes (6).Apmis 01/2012; · 1.99 Impact Factor
Vibrio cholerae O1 causes Asiatic cholera and has
been regarded as a member of a group of organisms
whose major habitats are aquatic ecosystems (7, 21,
22). V. cholerae is currently classified into 206 “O”
serogroups based on the heat stable somatic “O” antigen
(43, 49). Only V. cholerae O1 and O139 have been
found to be associated with epidemic cholera. V.
cholerae non-O1, non-O139 serogroups are ubiquitous
in the aquatic environment and recognized as causative
agents of sporadic and localized outbreaks (6, 42).
When V. cholerae is not wreaking havoc in the human
intestine, it may be found in diverse aquatic environ-
ment such as estuaries, rivers, ponds, etc. (8, 23).
Adhesion to surfaces both in the human intestine and in
the aquatic environment plays an important role in V.
cholerae’s success as a pathogen and an environmental
organism. In the aquatic environment, V. cholerae can
survive either as free-living planktonic organisms in the
water column, attached to a variety of abiotic surfaces or
associated with phytoplankton and zooplankton (14,
Bacteria attach to surfaces of this kind as widely-sep-
arated individuals, small colonial aggregates or conflu-
ent biofilm communities characterized by interactions
between community members and a three dimensional
architecture that provides channels through which nutri-
ents and metabolic by-products circulate. Biofilms that
form in multi-species habitats can be composed either of
a single strain or of multiple strains or species (10).
Biofilm Acts as a Microenvironment for
Plankton-Associated Vibrio cholerae in the Aquatic
Environment of Bangladesh
Mohammad Sirajul Islam*, 1, Mohammad Iqbal Kabir Jahid2, Mohammad Majibur Rahman2,
Mohammed Ziaur Rahman1, Mohammad Shafiqul Islam1, Mohammad Shahidul Kabir1,
David Allen Sack1, and Gary K. Schoolnik3
1International Centre for Diarrhoeal Disease Research, Bangladesh, GPO Box 128, Dhaka 1000, Bangladesh, 2Department of
Microbiology, University of Dhaka, Dhaka 1000, Bangladesh, and 3Department of Medicine, Division of Infectious Diseases
and Geographic Medicine, and Department of Microbiology and Immunology, Stanford University School of Medicine, Stan-
ford, CA 94305, U.S.A.
Received July 10, 2006; in revised form, November 7, 2006. Accepted January 15, 2007
Abstract: The role of biofilm as a microenvironment of plankton-associated Vibrio cholerae was investigat-
ed using plexiglass as a bait. A total of 72 biofilm samples were tested using culture, direct fluorescent anti-
body (DFA) and molecular techniques following standard procedures. Culturable V. cholerae (smooth
and rugose variants) were isolated from 33% of the samples. V. cholerae O1 were detected by FA technique
throughout the year except April and June. All V. cholerae O1 isolates were positive for tcpA, ctxA and ace
genes while V. cholerae non-O1, non-O139 isolates lacked these genes. V. cholerae O1 (both Inaba and
Ogawa) strains had identical ribotype pattern (R1), but V. cholerae non-O1, non-O139 had different ribo-
type patterns. All V. cholerae O1 strains were resistant to vibrio-static compound (O/129). All V. cholerae
O1 except one were resistant to trimethoprime-sulphamethoxazole, streptomycin, nalidixic acid and fura-
zolidone but sensitive to ciprofloxacin, and tetracycline. This study indicates that plexiglass can act as a
bait to form biofilm, a microenvironment that provides shelter for plankton containing V. cholerae in the
aquatic environment of Bangladesh.
Key words: Biofilm, Plankton, V. cholerae, Bangladesh
Microbiol. Immunol., 51(4), 369–379, 2007
Abbreviations: APW, alkaline peptone water; DFA, direct fluo-
rescent antibody; LB, Luria Bertani; NA, nalidixic acid; PBS,
phosphate-buffered saline; PCR, polymerase chain reaction;
trimethoprim-sulfamethoxazole; TTGA, taurocholate-tellurite-
gelatin agar; VSC, vibrio-static compound; YE, yeast extract.
*Address correspondence to Dr. Mohammad Sirajul Islam,
Environmental Microbiology Laboratory, Laboratory Sciences
Division, ICDDR, B GPO Box-128, Dhaka-1000, Bangladesh.
Biofilms have been the subject of intense interest in
recent years due to the predominance of biofilm-associ-
ated bacteria in natural environments and the increased
antibiotic resistance of biofilm bacteria compared to the
relative sensitivity of planktonic bacteria (4). It has
also been observed that the rugose colony variant of the
El Tor biotype attaches to a variety of abiotic surfaces
and forms biofilm due to the biosynthesis of extracellu-
lar polysaccharide (50). The rugose variants are resis-
tant to the bactericidal action of chlorine and probably
resistant to biocide (34, 39).
In river ecosystems, the surfaces are covered by
structured autotrophic-heterotrophic assemblies embed-
ded within polysaccharidic material forming a biofilm.
This biofilm contributes significantly to carbon cycling
in rivers and streams (31, 46). The close contact
between the algal and the heterotrophic community in
attached biofilms favors the use of algal material by
microorganisms within the biofilm (12, 36). Algal
accumulation and activity enhance the heterotrophic
community’s use of organic matter by increasing the
amount of substrate available for bacteria (11, 40). V.
cholerae attached to biotic surfaces, such as mucilagi-
nous sheaths of algae and chitinous flora, can derive
nutrients from digestion of the surface. By contrast V.
cholerae attached to an abiotic (non-nutritive) surface
can obtain nutrients only from the water column (or
adsorbed from the water column onto the abiotic sur-
face). The concentrations of such nutrients (in the
water column or adsorbed from it onto the surface) are
typically low. However this situation is dramatically
altered if the abiotic surface also contains other species
which are nutrient producers. The most likely nutrient-
producing surface-attached taxa are organisms which
engage in primary production, i.e., which use photosyn-
thesis to produce carbon- and nitrogen-containing com-
pounds that can be utilized by the heterotrophic bacteria
with which they associate on the same surface. These
primary food producing organisms include various
kinds of phytoplankton.
To date very little is known about the phytoplankton-
and zooplankton-associated V. cholerae O1 in the
biofilm community found in the aquatic environment of
Bangladesh. Therefore, the present study was conduct-
ed to describe the role of biofilm as a microenviron-
ment for plankton-associated V. cholerae in the aquatic
environment of Bangladesh.
Materials and Methods
Biofilm sampling device. The biofilm device devel-
oped by the Maryland Sea Grant program, U.S.A. was
used for the present study. It consists of plexiglass
discs strung on monofilament line at predetermined
depths, one end of the line anchored to the bottom, the
other end suspended from a float. Each biofilm sam-
pling device consists of three plexiglass sampling discs,
one located 12 cm from the surface (within the photic
zone), one 12 cm from the bottom (near the benthic
zone) and the third at mid depth. Adjustment of disc
positions in the water column was done when needed
due to changes in the depth of the water column at vari-
Sampling site. A canal of the Meghna Dhonagoda
River at Matlab was selected as the sampling site. A set
of biofilm sampling devices was placed at the site and
the samples were collected every 2 weeks for 1 year.
The study was conducted from July 2001 to July 2002
Sample collection and processing. The discs of the
biofilm device were taken out of the water. The samples
were taken by scraping the discs with the edge of a
razor and 3/4 of the sample was preserved in 3.0 ml
phosphate buffered saline (PBS) and transported to the
Environmental Microbiology Laboratory of the Interna-
tional Centre for Diarrhoeal Disease Research,
Bangladesh (ICDDR, B) in a transport box (Jonny Plas-
tic Ice, Pelton Sheperd, Stockton, Calif., U.S.A.) and
processed within 24 hr of collection. One fourth of the
scraped sample was preserved in RNA later and was
sent to Professor Gary Scholnik’s laboratory in Stanford
University, U.S.A., for molecular analysis. From 3 ml
preserved sample in PBS, 1 ml uncrushed sample was
kept in 3.5 ml PBS with 4% formalin for phytoplankton
and zooplankton identification. The rest of the sample
(2 ml) was homogenized using a steadfast stirrer
(Model 300, Fisher Scientific, U.S.A.). One milliliter of
homogenate was enriched in alkaline peptone water
(APW) and incubated for 6 hr at 37 C. Afterwards
enrichment attempts were made to culture vibrios fol-
lowing standard procedures. Isolated vibrios were fur-
ther characterized using PCR (polymerase chain reac-
tion) and ribotyping. From another 1 ml homogenate,
450 µl was taken and supplemented with 0.025% yeast
extract (YE) and 0.002% nalidixic acid (NA) and incu-
bated at room temperature overnight in the dark and
then the samples were preserved with (4%) formalin for
detection of V. cholerae by Direct Fluorescent Antibody
(DFA) technique (30). A flow diagram of sample pro-
cessing is presented in Fig. 1.
Bacteriology and serotyping. From the enrichment
sample in APW, 2 loopfuls were taken and inoculated
onto thiosulphate-citrate-bile-salt-sucrose (TCBS) and
taurocholate-tellurite-gelatin agar (TTGA) plates (33)
and incubated at 37 C for 18–24 hr. Suspected vibrio
colonies were further characterized following the proce-
M.S. ISLAM ET AL
dures described earlier (23). In brief, strains were only
identified as V. cholerae if they fulfilled the following
criteria: Gram negative, oxidase positive, produced acid
from sucrose but not inositol and decarboxylated lysine
and ornithine but not arginine. Strains were serotyped
and biotyped following the procedures described by
Kelly et al. (28).
Detection of V. cholerae O1 by DFA technique. A 5
µl portion of biofilm homogenate was placed into a
well of a PTFE (polytetrafluoroethylene)-coated glass
slide. The fluorescent antibody technique was carried
out following procedures described earlier (25) with
some modification. In brief, the sample was air-dried
and fixed with absolute ethanol (5 µl); then one drop
(10 µl) of Fluorescent iso-thiocyanate conjugated mon-
oclonal antibody (New Horizon Diagnostics Corp.,
U.S.A.) of V. cholerae O1 (cholera DFA) was added to
each well and the slide was incubated at 37 C for 30
min in a moist chamber. The slide was then rinsed with
sterile PBS, gently blotted and air-dried in the dark.
Finally the slide was mounted under a coverslip using
fluorescent mounting media (New Horizons Diagnos-
tics Corp.) and examined under an epifluorescence
microscope (Model BH-2, Olympus Optical Co.,
Tokyo). Cholera-positive control and cholera-negative
control provided by the manufacturer (New Horizon
Diagnostics Corp.) were used as the positive and nega-
tive control respectively. The V. cholerae O1 was
counted following the procedures described earlier (13).
In the viability test, non-culturable but viable V. choler-
ae O1 cells were counted following the procedure
described by Kogure et al. (30). In brief, non-culturable
cells utilize YE (0.025%) as a nutrient and start growing
but due to addition of NA (0.002%), cross wall forma-
tion can not take place for inhibition of DNA gyrase
and termination of DNA polymerization (47). Hence,
the nonculturable but viable cells appear longer than
Multiplex PCR assay for tcpA and ctxA genes. Mul-
tiplex PCR assay was carried out with the boiled tem-
plate of V. cholerae O1 and non-O1, non-O139 strains to
detect the ctxA gene with primers ctxA 1 (5'-CTCA-
GACGGGATTTGTTAGGCACG-3') and ctxA 2 (5'-
TCTATCTCTGTAGCCCCTATTACG-3') which pro-
duced an amplicon of 302 bp (27). For the detection of
tcpA (classical) gene, primers tcpA (classical) 1 (5'-
BIOFILM ASSOCIATED VIBRIO CHOLERAE
Table 1. Presence of V. cholerae O1 and V. cholerae non-O1, non-O139 in 24 samples each from surface, middle and bottom discs
V. cholerae O1
V. cholerae non-O1, non-O139
4 (16.7%)8 (33.3%)
b)Two different clones.
GAATGGAGC-3') were used, which produced an
amplicon of 618 bp (27). For the detection of tcpA (El
Tor) gene, primers tcpA (El Tor) 1 (5'-GAA-
GAAGTTTGTAAAAGAAGAACAC-3') and tcpA (El
Tor) 2 (5'-GAAAGGACCTTCTTTCACGTTG-3')
were used, which produced an amplicon of 472 bp (27).
Amplification was carried out in 50 µl volumes con-
taining 5 µl 10? PCR amplification buffer (500 mM
KCl; 100 mM Tris-HCl, pH 9.0; 0.1% Triton-X), 2 µl
MgCl2 (50 mM), 1 µl of 2.5 mM dNTPs, 20 pmol of
primers specific for ctxA and tcpA (classical/El Tor)
genes, 1 µl Taq polymerase (5 U/ml), 17.25 µl Milli-Q
water and 10 µl DNA template. The cycling condition
included a preincubation at 94 C for 5 min, followed
by a middle step of 40 cycles for 1 min each at 94 C
(denaturation), 56 C (annealing of primer) and 72 C
(DNA polymerase-mediated extension) and a final
extension at 72 C for 10 min for the tcpA and ctxA
genes using an automated thermal cycler (Perkin-
Elmer, Cetus, U.S.A.) as described earlier (24). The
PCR products were analysed by horizontal gel elec-
trophoresis with 1% agarose gel in Tris-borate EDTA
(TBE) buffer (50 mM Tris-borate; 1 mM EDTA; pH
8.2). The gel was stained with ethidium bromide (0.5
M.S. ISLAM ET AL
Fig. 1. Flow chart showing the processing of samples.
µg/ml) and visualized with a UV transilluminator and
PCR assay for ace gene. The identified V. cholerae
O1 and non-O1, non-O139 strains were subjected to
simplex PCR for the confirmation of the presence of the
ace gene among the isolates. The primers for the ace
gene, ace-1 (5'-TAAGGATGTGCTTATGAGGACAC-
CC-3') and ace-2 (5'-CGTGATGAATAAAGATACT-
CATAGG-3'), were used, which produced an amplicon
of 289 bp (27). Reset of the procedures was the same as
described for tcpA and ctxA genes above.
Extraction of chromosomal DNA. Chromosomal
DNA was extracted from isolated V. cholerae O1 and
non-O1, non-O139 using the procedure described by
Murray and Thompson (35) with some modifications.
In brief, cells from an 18 hr Luria-Bertani (LB) culture
were harvested and resuspended in TE buffer (10 mM
Tris-HCl; 1 mM EDTA; pH 8.0) treated with 10% SDS
and freshly prepared proteinase K and incubated at 50 C
for 1 hr. After incubation, CTAB/NaCl (10%
cetyltrimethyl ammonium bromide in 0.7 M NaCl) was
added and incubated at 65 C for 15 min. The aqueous
phase was then treated with phenol-chloroform and the
DNA pellet was washed with 70% ethanol. The DNA
was resuspended in TE buffer and treated with RNAse
at 37 C for 1 hr and then preserved for ribotyping.
Ribotyping. Southern blots were prepared for ribo-
typing according to the procedures described by Dals-
gaard et al. (9) with a digoxygenin-labelled rRNA
probe. The 7.5 kb BamHI fragment of plasmid pkk3535
containing the 16S and 23S rRNA genes of Escherichia
coli was used as the rRNA probe (5). Briefly, for
Southern blot, 2 µg of V. cholerae preserved DNA was
digested with BglI restriction enzyme and fragments
were electrophoresed at 70 V through 0.7% agarose gel
in Tris-borate EDTA (TBE) buffer for 2 hr. Gels were
soaked in 0.25 M HCl for 10 min to allow partial
depurination. The gel was then soaked in denaturation
solution for 30 min, followed by neutralisation in 0.5 M
Tris-HCl (pH 7.4) and 1.5 M NaCl for 30 min.
positively charged nylon membrane
(Amersham Pharmacia Biotech, Little Chalfont, Buck-
inghamshire, England) was used with a vacuum pump
unit (Bio-Rad Laboratories) for Southern blotting and
the DNA fragments were fixed to the membrane by
exposure to UV light for 3 min. The membrane was
prehybridized and then hybridized with labelled rRNA
probe. Finally detection of the hybridised bands on the
membrane was carried out according to the manufac-
turer’s instructions (Roche Diagnostic GmbH,
Antimicrobial susceptibility. The susceptibility of V.
cholerae to the antimicrobial agents tested was deter-
mined in vitro by using the modified agar-disc-diffusion
method (2) with commercial antimicrobial discs
(Oxoid, Basingstoke, U.K.). The antibiotic discs used in
this study were trimethoprim-sulfamethoxazole (TMP-
SMZ) (25 µg), streptomycin (S) (10 µg), furazolidone
(FR) (100 µg), NA (30 µg), tetracycline (T) (30 µg)
and ciprofloxacin (CIP) (5 µg). Escherichia coli ATCC
25922 was used as the control strain for susceptibility
studies. Strains were characterized as susceptible or
resistant to antibiotics based on the inhibition zone size
according to the recommendation of WHO (48).
Identification of phytoplankton and zooplankton.
The uncrushed formalin-preserved samples were shaken
gently for proper mixing from which a 1 ml sub-sample
was drawn by pipette and transferred into a Sedgewick-
Rafter counting cell (1). The samples were then
observed under a compound binocular microscope for
identification. The phytoplankton was identified fol-
lowing the procedures described by various authors (1,
15–17, 38, 45). The same procedure was followed for
described elsewhere (3, 32, 37, 45).
Abundance of Culturable V. cholerae in Biofilm Sam-
The overall isolation rate of culturable V. cholerae
was 33% (24/72). The surface disc yielded 4.2% (1/24)
and 16.7% (4/24) of V. cholerae O1 and non-O1, non-
O139 respectively. The abundance of V. cholerae O1
and non-O1, non-O139 in the middle disc was 8.3%
(2/24) and 33.3% (8/24) respectively. The bottom disc
yielded 8.3% (2/24) and 29.16% (7/24) of V. cholerae
O1 and non-O1, non-O139 respectively. Culturable V.
cholerae O1 was isolated in the months of July and
March but V. cholerae non-O1, non-O139 was isolated
many months during the study period (Table 1).
Out of 5 V. cholerae O1 isolates, 3 were rugose vari-
ants whereas out of 10 V. cholerae non-O1, non-O139
isolates, 2 were rugose variants as grown on gelatine
agar (Table 2).
Enumeration of V. cholerae O1 by DFA Technique
The DFA counts of V. cholerae O1 varied from 2.75
to 4.18 log10cells/ml in biofilm homogenate. V. choler-
ae O1 was present almost throughout the year except
April and June (Fig. 2). The V. cholerae O1 (both cul-
turable and viable but non-culturable) was detected in
association with only phytoplankton by DFA technique
(Fig. 3) but no V. cholerae was detected in association
BIOFILM ASSOCIATED VIBRIO CHOLERAE
Detection of tcpA, ctxA and ace Genes by PCR Tech-
The PCR results showed that all the V. cholerae O1
isolates carried the tcpA gene of El Tor biotype, ctxA
and ace but none of the V. cholerae non-O1, non-O139
yielded positive PCR values for the ace, ctxA and tcpA
genes of either classical or El Tor biotypes (Figs. 4 and
Analysis of BglI restriction patterns of conserved
rRNA genes (ribotyping) in the environmental strains
of V. cholerae O1 and non-O1, non-O139 isolated from
biofilm samples revealed that all the V. cholerae O1
(both Inaba and Ogawa) strains were of a single clone
since they produced identical restriction patterns R1
(Fig. 6). The V. cholerae non-O1, non-O139 strains
belonged to 8 different clones (R2, R3, R4, R5, R6, R7,
R8, R9) of which three (VC-5, VC-6, VC-10) showed
identical restriction pattern R2 (Fig. 6 and Table 2).
These strains were identified at different dates and
M.S. ISLAM ET AL
Table 2. Phenotypic, genotypic and antibiogram analysis of V. cholerae isolated from biofilm samples at Matlab in Bangladesh
tcpA ctxA ace
V. cholerae O1, Ogawa (VC-1)
V. cholerae O1, Ogawa (VC-2)
V. cholerae O1, Inaba (VC-3)
V. cholerae O1, Ogawa (VC-4)
V. cholerae non-O1, non-O139 (VC-5)
V. cholerae non-O1, non-O139 (VC-6)
V. cholerae non-O1, non-O139 (VC-7)
V. cholerae non-O1, non-O139 (VC-8)
V. cholerae non-O1, non-O139 (VC-9)
V. cholerae non-O1, non-O139 (VC-10) 5/3/2002 Middle Smooth
V. cholerae O1, Inaba (VC-11)
V. cholerae non-O1, non-O139 (VC-12) 20/3/2002 Middle Rugose
V. cholerae non-O1, non-O139 (VC-13) 8/5/2002 Surface Smooth
V. cholerae non-O1, non-O139 (VC-14) 8/5/2002 Middle Rugose
V. cholerae non-O1, non-O139 (VC-15) 8/5/2002 Bottom Smooth
24/7/2001 Surface Rugose
24/7/2001 Middle Rugose
24/7/2001 Middle Smooth
24/7/2001 Bottom Rugose
23/9/2001 Bottom Smooth
10/1/2002 Bottom Smooth
5/2/2002 Middle Smooth
5/2/2002 Bottom Smooth
5/2/2002 Bottom Smooth
TMP-SMZ, S,NA, FR
TMP-SMZ, S,NA, FR
TMP-SMZ, S,NA, FR
TMP-SMZ, S,NA, FR
TMP-SMZ, S, NA, T
TMP-SMZ, S, NA
TMP-SMZ, S, NA
5/3/2002 Bottom Smooth
a)Ribotypes are based on BglI restriction patterns of the 16S rRNA gene.
b)T, tetracycline; S, streptomycin; TMP-SMZ, trimethoprime-sulfamethoxazole; NA, nalidixic acid; FR, furazolidone; CIP,
c)Susceptible to tetracycline, streptomycin, trimethoprime-sulfamethoxazole, nalidixic acid, furazolidone, ciprofloxacin.
Fig. 2. Counts of V. cholerae O1 by DFA at different depths of biofilm device.
depths (Table 2).
Vibrio-Static Compound and Antimicrobial Susceptibili-
All V. cholerae O1 isolates and 33.3% (3/10) of V.
cholerae non-O1, non-O139 were resistant to vibrio-
static compound O/129 (VSC). All the strains of V.
cholerae O1 were resistant to TMP-SMZ, S, NA and
FR, but sensitive to T and CIP except one strain (VC-
11). This strain (VC-11) was only resistant to TMP-
SMZ and S. Three out of ten V. cholerae non-O1, non-
O139 (VC-9, VC-12, VC-13) were resistant to TMP-
SMZ, S and NA. However, strain VC-9 was also resis-
tant to T (Table 2).
Phytoplankton and Zooplankton Species Associated in
The identified phytoplankton from the uncrushed for-
BIOFILM ASSOCIATED VIBRIO CHOLERAE
Fig. 3. Detection of V. cholerae O1 from the homogenate of plankton by DFA technique.
Fig. 4. Agarose gel electrophoresis of multiplex PCR specific amplicon of ctxA and tcpA (classical and El Tor) of V.
cholerae strains isolated from biofilm samples. Lane-A–D & K: V. cholerae O1 strains (VC-1 to VC-4 and VC-11).
Lane-E–J, L–O: V. cholerae non-O1, non-O139 strains (VC-5 to VC-10, VC-12 to VC-15). Lane-P: V. cholerae O1
569B (classical). Lane-Q: V. cholerae 757 (El Tor). Lane-R: V. cholerae O139. Lane-S: Negative control. Lane-T: mo-
lecular weight marker (HaeIII fragment).
malin-preserved samples was Anabaena spp., Micro-
cystis spp., Oscillatoria spp., Merismopedia spp.. Sti-
geoclonium spp., Fragilaria spp., Pinnularia spp.,
Cymbella spp., Navicula spp., Gomphosphaeria spp.,
Coscinodiscus spp., Ulothrix spp., Trachelomonas spp.,
Cyclotella spp., Synedra spp., Cocconeis spp., Gom-
M.S. ISLAM ET AL
Fig. 5. Agarose gel electrophoresis of PCR amplicon of ace. Lane-A–D & K: V. cholerae O1 strains (VC-1 to VC-4
and VC-11). Lane-E–J, L–O: V. cholerae non-O1, non-O139 strains (VC-5 to VC-10, VC-12 to VC-15). Lane-P: V.
cholerae O1 569B (classical). Lane-Q: V. cholerae 757 (El Tor). Lane-R: V. cholerae O139. Lane-S: Negative control.
Lane-T: molecular weight marker (HaeIII fragment).
Fig. 6. Representative ribotype patterns of BglI-digested DNA of Vibrio cholerae. Lane-A: VC-1, lane-B: VC-2, lane-
C: VC-3, lane-D: VC-4, lane-E: VC-5, lane-F: VC-6, lane-G: VC-7, lane-H: VC-8, lane-I: VC-9, lane-J: VC-10, lane-
K: VC-11, lane-L: VC-12, lane-M: VC-13, lane-N: VC-14, lane-O: VC-15, lane-P: V. cholerae O1 569B (classical),
lane-Q: V. cholerae 757 (El Tor), lane-R: V. cholerae O139. Ribotype patterns: lane-A, lane-B, lane-C, lane-D, lane-K
(R1); lane-E, lane-F, Lane-J (R2); lane-G (R3); lane-H (R4), lane-I (R5), lane-L (R-6), lane-M (R-7), lane-N (R-8),
lane-O (R-9), lane-P: V. cholerae O1 569B (classical), lane-Q: V. cholerae 757 (El Tor), lane-R: V. cholerae O139.
Diploneis spp., and Melosira spp. The identified zoo-
plankton was Monostyla spp., Arcella spp., Philodina
spp., Difflugia spp., Brachionus spp. and Chironomid
spp., Gyrosigma spp., Amphoraspp.,
The present study showed that the algae containing V.
cholerae as well as the free floating V. cholerae from the
water column move to the surface of the plexiglass disc
and form biofilm communities in which a complex
interaction takes place between the algal and V. cholerae
communities in the tributaries of Meghna River in
Bangladesh. It has also been observed that biofilm pro-
vides V. cholerae a microenvironment where rugose
variants can persist. In a laboratory-based experiment,
Yildiz et al. (50) observed that rugose variants can form
biofilm better than smooth variants as the former can
produce extra-cellular polysaccharides. In the present
field-based study, rugose variants of V. cholerae were
also isolated from biofilm samples. The presence of
ctxA, tcpA and ace genes in V. cholerae O1 isolates
indicates that V. cholerae O1 do not loose their viru-
lence properties when they form biofilm in the aquatic
environment. The finding of this study is also consistent
with findings of a previous study (26). It was also
shown that different clones of V. cholerae O1 persist in
the aquatic environment of Bangladesh (26). In the
present study, the isolation of the same clone (same
ribotype pattern) of V. cholerae O1 from the biofilm
samples indicates that a particular clone of V. cholerae
O1 may have advantages to form biofilm in the aquatic
environment of Bangladesh. The repeated isolation of
R2 ribotype of V. cholerae non-O1, non-O139 indicated
that this particular ribotype was more likely to be sur-
face associated than the other ribotypes.
Dramatic increases in resistance of V. cholerae O1
clinical isolates to both T and TMP-SMZ were noted
over the course of 1991 and 1992, rising from 2 to 90%
for T and from 18 to 90% for TMP-SMZ (29). In
another study, Sack et al. (41) also observed that T
resistance among El Tor strains rapidly increased from
1.9% in 1990 to 7.6% in 1991, 61.1% in 1992 and
85.4% in 1993. The present study showed that the V.
cholerae O1 El Tor isolated from biofilm has developed
resistance to TMP-SMZ, S, NA and FR. The resistance
of environmental strains of V. cholerae O1 to TMP-
SMZ is consistent with the previous findings of clinical
strains. It was also found that all El Tor V. cholerae O1
were resistant to VSC which might indicate that biofilm
can act as a reservoir of VSC- and antibiotic-resistant V.
cholerae in the aquatic environment of Bangladesh.
The main limitation of the study is the low rate of
recovery of the V. cholerae O1 in culturable form from
the biofilm sample using the presently available con-
ventional culture technique. Attempts were made to
isolate culturable V. cholerae O1 using enrichment
media and growing it in selective plates including
TCBS and TTGA. However, V. cholerae O1 has been
detected throughout the year except April and June by
DFA technique only in association with algae (Fig. 3)
but V. cholerae O1 were isolated in culturable form
only in the months of July and March. Improved meth-
ods are needed to culture V. cholerae from biofilm.
Therefore, in our future study, we are planning to use
colony blot hybridization by plating the biofilm
homogenate onto non-selective plates, e.g. LB agar, in
an attempt to increase the isolation of culturable V.
The result of the present study revealed that V.
cholerae associated with phytoplankton could form
biofilm in the aquatic environment on abiotic surfaces
such as plexiglass discs. Therefore, it may be concluded
that plexiglass devices may be used as a bait to concen-
trate the phytoplankton to form biofilm which can play
an important role as a microenvironment of phyto-
plankton associated V. cholerae in the aquatic environ-
ment of Bangladesh.
This research was funded by National Institute of Health
(NIH), Grant no. 5 R01 A143422-03 through Stanford Universi-
ty, U.S.A. ICDDR, B acknowledges with gratitude the commit-
ment of National Institute of Health and Stanford University to
the Centre’s research efforts.
1) American Public Health Association (APHA). 1998. Stan-
dard methods for the examination of water and wastewater,
20th ed, America Public Health Association/American
Water Works Association/Water Environment Federation,
Washington, D.C., U.S.A.
2) Bauer, A.W., Kirby, W.M., Sherris, J.C., and Turck, M.
1966. Antibiotic susceptibility testing by a standardized sin-
gle disk method. Am. J. Clin. Pathol. 45: 493–496.
3) Bhouyain, A.M., and Asmat, G.S.M. 1992. Freshwater zoo-
plankton from Bangladesh, Ghazi Publishers, Dhaka,
4) Brooun, A., Liu, S., and Lewis, K. 2000. A dose-response
study of antibiotic resistance in Pseudomonas aeruginosa
biofilms. Antimicrob. Agents Chemother. 44: 640–646.
5) Brosius, J., Ullrich, A., Raker, M.A., Gray, A., Dull, T.J.,
Gutell, R.R., and Noller, H.F. 1981. Construction and fine
mapping of recombinant plasmids containing the rrnB ribo-
somal RNA operon of E. coli. Plasmid 6: 112–118.
6) Chakraborty, S., Garg, P., Ramamurthy, T., Thungapathra,
M., Gautam, J.K., Kumar, C., Maiti, S., Yamasaki, S., Shi-
BIOFILM ASSOCIATED VIBRIO CHOLERAE
mada, T., Takeda, Y., Ghosh, A., and Nair, G.B. 2001.
Comparison of antibiogram, virulence genes, ribotypes and
DNA fingerprint of Vibrio cholerae of matching serogroups
isolated from hospitalised diarrhoea cases and from the
environment during 1997–1998 in Calcutta, India. J. Med.
Microbiol. 50: 879–888.
7) Colwell, R.R., Seidler, R.J., Kaper, J., Joseph, S.W.,
Garges, S., Lockman, H., Maneval, D., Bradford, H.,
Roberts, N., Reemers, E., Huq, I., and Huq, A. 1981.
Occurrence of Vibrio cholerae serotype O1 in Maryland
and Louisiana estuaries. Appl. Environ. Microbiol. 41:
8) Colwell, R.R., and Spira, W.M. 1992. The ecology of Vibrio
cholerae, p. 107–127. In Barua, D., and Greenough, W.B.,
III (eds), Cholera, Plenum Medical Book Company, New
9) Dalsgaard, A., Mortensen, H.F., Molbak, K., Dias, F.,
Serichantalergs, O., and Echeverria, P. 1996. Molecular
characterization of Vibrio cholerae O1 strains isolated dur-
ing cholera outbreaks in Guinea-Bissau. J. Clin. Microbiol.
10) Davey, M.E., and O’Toole, G.A. 2000. Microbial biofilms:
from ecology to molecular genetics. Microbiol. Mol. Biol.
Rev. 64: 847–867.
11) Espeland, E.M., Francoeur, S.N., and Wetzel, R.G. 2001.
Influence of algal photosynthesis on biofilm bacterial pro-
duction and associated glucosidase and xylosidase activi-
ties. Microb. Ecol. 42: 524–530.
12) Haack, T.K., and McFeters, G.A. 1982. Nutritional relation-
ships among microorganisms in an epilithic biofilm com-
munity. Microbiol. Ecol. 8: 115–126.
13) Hasan, J.A., Bernstein, D., Huq, A., Loomis, L., Tamplin,
M.L., and Colwell, R.R. 1994. Cholera DFA: an improved
direct fluorescent monoclonal antibody staining kit for
rapid detection and enumeration of V. cholerae O1. Federa-
tion of European Microbiological Societies 120: 143–148.
14) Huq, A., Small, E.B., West, P.A., Huq, M.I., Rahman, R.,
and Colwell, R.R. 1983. Ecological relationships between
Vibrio cholerae and planktonic crustacean copepods. Appl.
Environ. Microbiol. 45: 275–283.
15) Islam, A.K.M.N., and Khatun, M. 1966. Preliminary studies
on the phytoplankton of the polluted waters. Sci. Res. 3:
16) Islam, A.K.M.N., and Nahar, L. 1967. Preliminary studies
on the phytoplankton of the polluted waters. Sci. Res. 4:
17) Islam, A.K.M.N., Khondker, M., Begum, A., and Akhter, N.
1992. Hydrobiological studies in two habitats at Dhaka. J.
Asiat. Soc. Bangladesh Sci. 18: 47–52.
18) Islam, M.S., and Aziz, K.M.S. 1978. Association of vibrios
with some hydrophytic plants, 52 pp. In Proceedings of the
3rd Annual Science Conference of Bangladesh Association
for the Advancement of Science, Chittagong, Abstract No.
19) Islam, M.S., Drasar, B.S., and Bradley, D.J. 1990. Long-
term persistence of toxigenic Vibrio cholerae O1 in the
mucilaginous sheath of a blue-green alga, Anabaena vari-
abilis. J. Trop. Med. Hyg. 93: 133–139.
20) Islam, M.S., Drasar, B.S., and Bradley, D.J. 1990. Survival
of toxigenic Vibrio cholerae O1 with a common duckweed,
Lemna minor, in artificial aquatic ecosystems. Trans. R.
Soc. Trop Med. Hyg. 84: 422–424.
21) Islam, M.S., Drasar, B.S., and Sack, R.B. 1993. The aquatic
environment as reservoir of Vibrio cholerae: a review. J.
Diarr. Dis. Res. 11: 197–206.
22) Islam, M.S., Drasar, B.S., and Sack, R.B. 1994. Probable
role of blue-green algae in maintaining endemicity and sea-
sonality of cholera in Bangladesh: a hypothesis. J. Diarr.
Dis. Res. 12: 245–256.
23) Islam, M.S., Alam, M.J., and Khan, S.I. 1995. Occurrence
and distribution of culturable Vibrio cholerae O1 in aquatic
environments of Bangladesh. Int. J. Environ. Stud. 47:
24) Islam, M.S., Rahim, Z., Alam, M.J., Begum, S., Moniruzza-
man, S.M., Umeda, A., Amako, K., Albert, M.J., Sack,
R.B., Huq, A., and Colwell, R.R. 1999. Association of Vib-
rio cholerae O1 with the cyanobacterium, Anabaena sp.,
elucidated by polymerase chain reaction and transmission
electron microscopy. Trans. R. Soc. Trop. Med. Hyg. 93:
25) Islam, M.S., Goldar, M.M., Morshed, M.G., Khan, M.N.,
Islam, M.R., and Sack, R.B. 2002. Involvement of the hap
gene (mucinase) in the survival of Vibrio cholerae O1 in
association with the blue-green alga, Anabaena sp. Can. J.
Microbiol. 48: 793–800.
26) Islam, M.S., Talukder, K.A., Khan, M.N.H., Mahmud,
Z.H., Rahman, M.Z., Nair, G.B., Siddique, A.K., Yunus,
M., Sack, D.A., Sack, R.B., Huq, A., and Colwell, R.R.
2004. Variation of toxigenic Vibrio cholerae O1 in the
aquatic environment of Bangladesh and its correlation with
the clinical strains. Microbiol. Immunol. 48: 773–777.
27) Keasler, S.P., and Hall, R.H. 1993. Detection and biotyping
of Vibrio cholerae O1 with multiplex polymerase chain
reaction. Lancet 341 (8861): 1661.
28) Kelly, M.T., Hickman-Brenner, F.W., and Farmer, J.J., III.
1992. Vibrio, p. 384–395. In Balows, A., Hausler, W.J., Jr.,
Hermann., K.L., Isenberg, H.D., and Shadomy, H.J. (eds),
Manual of clinical microbiology, 5th ed, ASM Press, Wash-
29) Khan, W.A., Begum, M., Salam, M.A., Bardhan, P.K.,
Islam, M.R., and Mahalanabis, D. 1995. Comparative trial
of five antimicrobial compounds in the treatment of cholera
in adults. Trans. R. Soc. Trop. Med. Hyg. 89: 103–106.
30) Kogure, K., Simidu, U., and Taga, N. 1979. A tentative
direct microscopic method for counting living marine bac-
teria. Can. J. Microbiol. 25: 415–420.
31) Lock, M.A. 1993. Attached microbial communities in
rivers, p. 113-138. In Ford, T.E. (ed), Aquatic microbiology:
an ecological approach, Blackwell Scientific Publications,
32) Mellanby, H. 1971. Animal life in fresh-water (6th ed), A
guide to fresh-water invertebrates, Chapman and Hall, Ltd.,
33) Monsur, K.A. 1961. A highly selective gelatin taurocholate
tellurite medium for isolation of Vibrio cholerae. Trans. R.
Soc. Trop. Med. Hyg. 55: 440–445.
34) Morris, J.G., Jr., Sztein, M.B., Rice, E.W., Nataro, J.P.,
Losonsky, G.A., Panigrahi, P., Tacket, C.O., and Johnson,
M.S. ISLAM ET AL
J.A. 1996. Vibrio cholerae O1 can assume a chlorine-resis-
tant rugose survival form that is virulent for humans. J.
Infect. Dis. 174: 1364–1368.
35) Murray, M.G., and Thompson, W.F. 1980. Rapid isolation
of high molecular weight plant DNA. Nucleic Acids Res. 8:
36) Nakano, S. 1996. Bacterial response to extracellular dis-
solved organic carbon released from healthy and senescent
Fragilaria crotonensis (Bacillariophyceae) in experimental
systems. Hydrobiology 339: 47–55.
37) Needham, J.G., and Needham, P.R. 1962. A guide to the
study of fresh water biology, 5th ed, Holden-Day, Inc., San-
38) Prescott, G.W. 1984. The algae: a review, Bishen Singh
Mahendra Pal Singh, Dehra Dun, India.
39) Rice, E.W., Johnson, C.J., Clark, R.M., Fox, K.R., Reasoner,
D.J., Dunnigan, M.E., Panigrahi, P., Johnson, J.A., and
Morris, J.G., Jr. 1992. Chlorine and survival of “rugose”
Vibrio cholerae. Lancet 340 (8821): 740.
40) Romani, A.M., and Sabater, S. 1999. Effect of primary pro-
ducers on the heterotrophic metabolism of a stream biofilm.
Freshwat. Biol. 41: 729–736.
41) Sack, D.A., Lyke, C., Mclaughlin, C., and Suwanvanichkij,
V. 2001. Antimicrobial resistance in shigellosis, cholera and
campylobacteriosis (WHO/CDS/CSR/DRS/2001.8), World
Health Organization, Geneva, Switzerland.
42) Sanyal, S.C., Alam, K., Neogi, P.K.B., Huq, M.I., and Al-
Mahmud, K.A. 1983. A new cholera toxin. Lancet i: 1337.
43) Shimada, T., Arakawa, E., Itoh, K., Okitsu, T., Matsushima,
A., Asai, Y., Yamai, S., Nakazato, T., Nair, G.B., Albert,
M.J., and Takeda, Y. 1994. Extended serotyping scheme for
Vibrio cholerae. Curr. Microbiol. 28: 175–178.
44) Singh, D.V., Matte, M.H., Matte, G.R., Jiang, S., Sabeena,
F., Shukla, B.N., Sanyal, S.C., Huq, A., and Colwell, R.R.
2001. Molecular analysis of Vibrio cholerae O1, O139,
non-O1, and non-O139 strains: clonal relationships
between clinical and environmental isolates. Appl. Environ.
Microbiol. 67: 910–921.
45) Ward, H.B., and Whipple, G.C. 1959. Freshwater biology,
2nd ed, Edmonson, W.T. (ed), John Wiley & Sons, Inc.,
New York, N.Y.
46) Wetzel, R.G. 1993. Microcommunities and microgradients:
linking nutrient regeneration, microbial mutualism, and
high sustained aquatic primary production. Neth. J. Aquat.
Ecol. 27: 3–9.
47) Wolfson, J.S., and Hooper, D.C. 1985. The fluoro-
quinolones: structure, mechanisms of action and resistance
and spectra of activity in vitro. Antimicrob. Agents Chemo-
ther. 28: 581–586.
48) World Health Organization. 1993. Guidelines for cholera
control. World Health Organization, Geneva.
49) Yamai, S., Okitsu, T., Shimada, T., and Katsube, Y. 1997.
Distribution of serogroups of Vibrio cholerae non-O1 non-
O139 with specific reference to their ability to produce
cholera toxin, and addition of novel serogroups. J. Jpn.
Assoc. Infect. Dis. 71: 1037–1045.
50) Yildiz, F.H., and Schoolnik, G.K. 1999. Vibrio cholerae O1
EI Tor: identification of a gene cluster required for the
rugose colony type, exopolysaccharide production, chlorine
resistance, and biofilm formation. Proc. Natl. Acad. Sci.
U.S.A. 96: 4028–4033.
BIOFILM ASSOCIATED VIBRIO CHOLERAE