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Journal of Clinical Microbiology, September 2000, p. 3179-3186, Vol. 38, No. 9
Faculté de Pharmacie, Département de
Microbiologie, Université de Paris-Sud, 92296 Châtenay-Malabry Cedex, France,1 and
Unité de Microbiologie, Département de Biologie,
Université Catholique de Louvain, 1200 Brussels,
Belgium2
Received 3 April 2000/Returned for modification 16 May
2000/Accepted 13 June 2000
Phenotypic and genotypic diversity of the flagellin gene
(fliC) of Clostridium difficile was studied in
47 isolates from various origins belonging to the serogroups A, B, C,
D, F, G, H, I, K, X, and S3. Electron microscopy revealed 17 nonflagellated strains and 30 flagellated strains. PCR and reverse
transcription-PCR demonstrated that the flagellin gene was present in
all strains and that the fliC gene was expressed in both
flagellated and nonflagellated strains. Southern blotting showed the
presence of only one copy of the gene and three different hybridization
patterns. DNA sequence analysis of fliC from the strains
belonging to serogroups C, D, and X, representative of each profile,
disclosed great variability in the central domain, whereas the N- and
C-terminal domains were conserved. The variability of the flagellin
gene fliC was further studied in the isolates by
PCR-restriction fragment length polymorphism (RFLP) analysis. Nine
different RFLP groups were identified (I to IX), among which three (I,
VII, and VIII) corresponded to numerous serogroups whereas the six
others (II, III, IV, V, VI, and IX) belonged to a single serogroup.
Flagellin gene RFLP analysis could constitute an additional typing
method employable in conjunction with other typing methods currently available.
Clostridium difficile is
the major etiological agent of pseudomembranous colitis and
antibiotic-associated diarrhea. In addition to the two major toxins, A
and B, which represent the major virulence factors (7), a
number of other putative accessory virulence factors have been
described. These include adhesins mediating adherence to mucosa
(15, 23, 48), fimbriae, and capsule and tissue-degradative
enzymes (6). However, in some bacterial species, flagella
may also be a virulence factor and play a role in colonization of the
gastrointestinal tract. The flagellar structure plays a role in
internalization of Campylobacter jejuni (18), Salmonella enterica serovar Typhi (26), and
Proteus mirabilis (31) into cultured epithelial
cells. Motility is an important factor in the virulence of Vibrio
cholerae (39) and Vibrio anguillarum (30). Flagella are also involved in chemotaxis and have been implicated in mucus-cell adherence and colonization by
Pseudomonas aeruginosa (1, 40),
Helicobacter pylori (14), and Burkholderia pseudomallei (8). Since flagella are believed to
constitute one of the virulence factors of various infectious bacteria,
the flagellin gene could be considered a useful genetic marker for epidemiological and phylogenetic studies (20, 52).
One aspect of C. difficile that we have studied is its
interaction with target cells (15, 23, 48). Adhesion to and
colonization of target tissues by bacteria are frequently important
first steps in establishing infection. It is likely that C. difficile is unable to colonize without attachment and will be
quickly removed by nonspecific host defense mechanisms.
Our laboratory is interested in finding out whether flagella play a
role in C. difficile intestinal colonization. Few studies concerning C. difficile flagella have been performed;
Delmée et al. established that flagella were involved in
cross-reactions of serogroups (11). In a previous study we
characterized the 39-kDa flagellin protein (45). The
flagellin gene was cloned and sequenced, and the recombinant protein
was characterized.
The aim of this work was to study the phenotypic and genotypic
variability of the flagellin gene (fliC) and its correlation with serogroups in C. difficile isolates from different
origins. Strains were investigated by electron microscopy (EM). The
presence of the fliC gene was verified by PCR amplification,
and the expression of the flagellin gene was studied by reverse
transcription (RT)-PCR. In order to investigate the flagellin gene
structure, Southern analysis with serogroup reference strains and
sequencing of fliC genes from three strains were performed.
PCR amplification of flagellin genes combined with restriction fragment
length polymorphism (RFLP) analysis were used in an attempt to study
the variability among C. difficile isolates.
Bacterial strains, media, and growth conditions.
Forty-seven
isolates belonging to 12 different serogroups (serogroups A1, A10, B,
C, D, F, G, H, I, K, S3, and X) were selected at the Microbiology Unit
of the Catholic University of Louvain, Brussels, Belgium, with care
taken to choose strains isolated from several geographical locations.
The 10 reference strains for specific serogroups were A (ATCC 43594), B
(ATCC 43593), C (ATCC 43596), D (ATCC 43597), F (ATCC 43598), G (ATCC
43599), H (ATCC 43600), I (ATCC 43601), K (ATCC 43602), and X (ATCC
43603). Clostridium sordellii (Institut Pasteur, Paris,
France) was used as a negative control, and C. difficile
79-685 was used as a positive control for the flagellin gene (Table
1).
0095-1137/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Phenotypic and Genotypic Diversity of the Flagellin
Gene (fliC) among Clostridium difficile Isolates
from Different Serogroups
ABSTRACT
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Abstract
Introduction
Materials and Methods
Results
Discussion
References
INTRODUCTION
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Abstract
Introduction
Materials and Methods
Results
Discussion
References
MATERIALS AND METHODS
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Abstract
Introduction
Materials and Methods
Results
Discussion
References
TABLE 1.
C. difficile
isolates studiedc
EM. The strains of C. difficile were grown overnight on agar plates supplemented with 7% horse blood as described above. A bacterial suspension was made in 100 µl of phosphate-buffered saline. Copper grids (Touzard et Matignon, Paris, France) were placed facedown on the cell suspension for 5 min and then negatively stained with a 2% phosphotungstic acid solution (pH 7.2). The grids were air dried and observed under a transmission electron microscope (EM 301; Philips).
DNA extraction and Southern blotting. DNA was extracted from 10 ml of overnight culture according to the protocol provided in the Puregene DNA gram-positive bacteria and yeast DNA extraction kit (Gentra Systems, Minneapolis, Minn.).
Southern blotting was carried out with 5 µg of DNA digested with 10 U of HindIII for 3 h under the conditions recommended by the provider (Life Technologies, Cergy Pontoise, France). Products of digestion were separated by electrophoresis on a 0.8% (wt/vol) agarose gel. The fragments were transferred onto a positively charged nylon membrane (Roche, Mannheim, Germany) using a vacuum blotter (Appligene-Oncor, Illkirch, France). The amplified fliC gene of the C. difficile 79-685 strain (45) was used as a C. difficile flagellin gene-specific probe. The DNA probe was labeled and detected by using the ECL direct nucleic acid labeling and detection system (Amersham-Pharmacia Biotech, Les Ulis, France). Hybridization and washing of membranes were carried out at low stringency (0.5× SSC [1× SSC is 0.15 M NaCl plus 0.015 M sodium citrate] at 50°C).PCR amplification and PCR-RFLP. For amplification of the fliC gene from various C. difficile isolates, the specific primers used were Nter (5'-ATGAGAGTTAATACAAATGTAAGTGC-3') and Cter (5'-CTATCCTAATAATTGTAAAACTCC-3') corresponding to the 5'- and 3'- end sequences of the fliC gene of the C. difficile 79-685 strain. DNA amplification by PCR was performed in a reaction volume of 100 µl consisting of 1 µl of a bacterial suspension washed twice with phosphate buffer, primer Nter (1 mM), primer Cter (1 mM), deoxynucleoside triphosphates (0.2 mM), MgCl2 (2 mM), 1 U of Taq polymerase, and 1× polymerase buffer (Promega, Madison, Wis.). The reaction mixture was overlaid with mineral oil. Initial denaturation was carried out at 94°C for 5 min. Thirty-five cycles of amplification were performed in a Thermocycler 480 (Perkin-Elmer, Norwalk, Conn.). Each cycle consisted of three steps: denaturation at 94°C (30 s), annealing at 55°C (30 s), and extension at 72°C (1 min). An additional step of extension for 10 min at 72°C was performed at the end of the amplification to complete the extension of the primers. Samples (5 µl) of amplified products were digested with the restriction enzymes HindIII, DraI, HpaI, PvuII, HincII, HinfI (Amersham-Pharmacia Biotech), and RsaI (Life Technologies) according to the vendor's recommendations. The digested amplified products were analyzed by electrophoresis in a 1.2% (wt/vol) agarose gel with a 100-bp ladder (Amersham-Pharmacia Biotech) as the molecular size marker.
DNA sequencing. PCR products were purified with the QIAquick PCR purification kit (Qiagen, Hilden, Germany). DNA sequencing was carried out with the BigDye terminator DNA sequencing kit (PE Applied Biosystems, Warrington, England). The samples were analyzed with the automated DNA sequencer ABI PRISM 310 genetic analyzer (Perkin-Elmer). The sequencing was initiated on both strands with the primers Nter and Cter and was finished with internal primers designed from the sequences obtained.
RNA extraction and RT-PCR.
RNA was extracted from 10 ml of
8-h C. difficile anaerobic culture. Bacteria were harvested
by centrifugation at 5,000 × g for 5 min at 4°C and
then resuspended in 0.5 ml of cold TE buffer (10 mM HCl, 1 mM EDTA, pH
7.4) and kept on ice. Glass beads (0.6 g; 425 µm < diameter < 600 µm; Sigma Chemical Co., St. Louis, Mo.) were
added in a solution containing 0.17 ml of 4% (wt/vol) Bentone
rheological additive (Rheox Ltd., Livingston, Scotland), 0.5 ml of acid
phenol (Sigma Chemical Co.), and 0.05 ml of 10% (wt/vol) sodium
dodecyl sulfate solution. The solution was mixed three times by
vortexing for 1 min each, interrupted by 1-min pauses. The aqueous
phase was recovered by centrifugation at 12,000 × g at
4°C for 15 min and then extracted three times with a
phenol-chloroform (1:1, vol/vol) solution and precipitated with
ethanol. The RNA pellet was washed with 75% (vol/vol) cold ethanol,
vacuum dried, and resuspended in 50 µl of TE buffer. The RNA was
treated with DNase I (Amersham-Pharmacia Biotech) and stored at
20°C.
Serogrouping and toxigenicity. Serogroups were determined by slide agglutination with rabbit antisera (12) and were confirmed by typing by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (13). Toxin A production was determined by the C. difficile toxin A test (Oxoid). In vitro cytotoxin (toxin B) determination was performed with HeLa cells cultured in minimum Eagle medium with Earle's salts (Life Technologies) supplemented with 10% fetal calf serum (Life Technologies), 1% nonessential amino acids (Life Technologies), and 200 mM L-glutamine (Life Technologies) in microtiter plates (3 × 104 cells per well). Fivefold serial dilutions of filtrates of 48-h TGY liquid cultures of C. difficile were incubated for 18 h with the cells at 37°C in a 5% CO2 atmosphere. After fixation and coloration of the culture cells with methylene blue, cytotoxic effect was observed by inverse microscopy.
Computer analyses. Nucleotide and protein sequence alignments were performed with DNA Strider software and the Multalin program (9).
Nucleotide sequence accession numbers. The nucleotide sequence of the fliC locus of strains 79-685, 545, 3232, and 5036, corresponding to serogroups S3, C, D, and X, respectively, were assigned GenBank numbers AF065259, AF095236, AF095237, and AF095238.
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RESULTS |
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Introduction
Materials and Methods
Results
Discussion
References
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Detection of fliC gene and flagella by EM and fliC gene expression. As observed by EM, 30 out of 47 strains showed visible flagella, whereas 17 were nonflagellated (Table 1). All strains from serogroup A were flagellated; in contrast, no strain from serogroup C carried flagella. In other serogroups both flagellated and nonflagellated strains were observed. The number and length of flagella also varied considerably among strains.
PCR amplification using fliC-specific oligonucleotide primers Nter and Cter was employed to investigate the presence of the gene in C. difficile isolates. The amplification gave a single product in all 47 C. difficile strains studied (not shown), whereas no amplified product was obtained from the negative control C. sordellii strain. An 870-bp fragment was obtained from 46 strains, whereas the serogroup X reference strain revealed an 850-bp amplified fragment. The lack of flagella on the bacterial surface could be due to the absence of transcription of the fliC gene. Therefore, to investigate expression of the flagellin gene in flagellated and nonflagellated strains, DNA transcription was investigated by detecting flagellin mRNA by RT-PCR. The results (Fig. 1) show that a single 870-bp amplified fragment was obtained from all the 17 nonflagellated strains, including four nonflagellated serogroup C, D, H, and I reference strains.
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Detection and copy number of fliC in C. difficile isolates.
In some bacteria, fliC can be
present in multiple copies on the bacterial chromosome. To assess
whether fliC is present in mono- or multicopy, the amplified
DNA from strain 79-685 was used as a probe in Southern hybridization of
chromosomal DNA of strain 79-685 and the 10 reference serogroups (A, B,
C, D, F, G, H, I, K, and X). Hybridization under low-stringency
conditions showed that DNA of all isolates hybridized with the
fliC-specific probe. Only one copy of the gene was present
in each strain. Some strains carry a HindIII site and
therefore show the presence of two bands (Fig.
2). The presence of a
HindIII site was confirmed by subjecting the amplified
fliC gene product to HindIII digestion. The
digestion with HindIII allows the classification of the
strains in three groups: the first group exhibits a single 1.94-kb
single band (79-685, C, F, and I), and the second group displays two
bands of 1.64 kb and 0.76 kb (A, B, D, G, H, and K), while there is a
single 3.53-kb band for the serogroup X reference strain (Fig. 2).
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DNA sequence analysis.
In order to confirm that the 870-bp
amplified product described above was the flagellin gene, PCR fragments
obtained from the 10 reference serogroup strains were partially
sequenced with the Nter primer. Sequencing revealed an N-terminal
sequence identical to the fliC gene of the C. difficile 79-685 strain in all PCR products (data not shown). To
investigate the conservation of the fliC gene coding region
in strains representing the different profiles obtained by Southern
blotting, the fliC gene of the serogroup C, D, and X
reference strains was amplified by PCR using specific primers (Nter and
Cter) as described in Materials and Methods. DNA and deduced amino acid
sequence analysis revealed an open reading frame composed of 873 nucleotides (290 amino acids) for the 79-685 strain and our C and D
reference strains, while the open reading frame was 846 nucleotides,
corresponding to 281 amino acids, for the serogroup X reference strain.
The latter strain carried a short deletion in the central region of
flagellin (Fig. 3).
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). The alignment was performed
with the MULTALIN program (
-helical conformation (alanine is a helix-forming
residue); this conservation could reflect the functional importance of
the terminal regions in forming the tertiary structure of the flagellin
protein monomers.
RFLP analysis of flagellin genes. As sequence analysis of fliC from three strains suggested significant variability of gene structure, we decided to examine gene structure by PCR-RFLP analysis in all the isolates studied. The amplified fliC gene was digested with HindIII, DraI, HpaI, PvuII, RsaI, HincII, and HinfI. According to the restriction map (not shown), these restriction sites are distinct and therefore can be used to perform RFLP analysis.
The results (Fig. 4) show the different restriction patterns obtained from the C. difficile strains. Three different restriction profiles were obtained with HindIII, DraI, HpaI, PvuII, HincII, and HinfI enzymes, and four restriction profiles (designated a, b, c, and d) were obtained with RsaI endonuclease. As far as the serogroup X reference strain is concerned, the fliC RFLP analysis revealed a unique and different profile with each enzyme: profile c with HindIII, DraI, HpaI, PvuII, HincII, and HinfI restriction enzymes and profile d with RsaI.
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DISCUSSION |
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Materials and Methods
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Discussion
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For numerous pathogens, the capacity to adhere, invade, and destroy the colonic mucosal cells appears to be an essential aspect of the first stage of their pathogenicity. C. difficile is responsible for the most-frequent hospital-acquired infection consequent to antibiotic therapy. It causes diarrheal disease, which can lead to an intense acute response: pseudomembranous colitis. C. difficile releases two exotoxins into the colon which are responsible for the disease: toxin A and toxin B. Toxin A (an enterotoxin) elicits fluid secretion, mucosal damage, and intestinal inflammation; toxin B (a cytotoxin) is completely devoid of enterotoxicity.
Before these events take place, C. difficile entering the gut must find pathways to reach suitable epithelial cells, which are naturally protected by a layer of dense mucus. C. difficile can adhere to the mucous layer (23). Afterwards the bacterium could penetrate the mucous layer with the aid of its proteases (38) and flagella; finally, C. difficile attaches to enterocytes by means of specific adhesins (23, 48); the role of flagella in this process has yet to be defined. At these different steps, the presence of complex and specialized chemotaxis and flagellar systems may play a role.
The studies undertaken here revealed that with EM certain strains of C. difficile are characterized by their inability to produce in vitro visible flagella. We have shown by PCR that the flagellin gene was present in both flagellated and nonflagellated C. difficile strains. The specificity of the amplification was confirmed by the fact that the flagellin gene of C. sordellii, highly flagellated and genetically very close to that of C. difficile, was not amplified with these primers. Gene amplification by PCR has been frequently used as a rapid method for detection and identification of pathogenic bacteria including Clostridium perfringens (16), Campylobacter spp. (36), Listeria monocytogenes (19), and Bordetella bronchiseptica (22). Therefore, amplification of the fliC gene could be used as a rapid method to detect and identify C. difficile.
We showed here that the flagellin gene was expressed in flagellated and nonflagellated C. difficile strains by detection of flagellin mRNA with the one-step method RT-PCR. According to Macnab (27) and Manson (28), the Escherichia coli flagellin gene was only expressed when both the basal body and the hook of the flagellum were fully formed through the membrane of bacterium. From that, the flagellin, the cap protein, and the junction hook-flagellum proteins were synthesized and assembled to achieve flagellar filament formation from the external membrane (5, 42). In nonflagellated C. difficile strains, the cap and/or junction hook-flagellum proteins could be inactivated by mutation, thus preventing transport of flagellin subunits through the bacterial membrane and polymerization. To confirm this hypothesis, it would be interesting to verify the presence of the flagellin protein in nonflagellated strains.
The nonflagellated C. difficile strains possess a cryptic
flagellin gene. We cannot rule out, however, that in vivo all strains could be flagellated, and we intend to study the in vivo expression of
fliC in our mouse model. Cryptic genes have been
characterized in nonmotile bacteria. Indeed, the expression of surface
flagella in some pathogenic bacteria may be induced only by factors
related to a specific biological microenvironment or under certain in vitro growth conditions. Holt and Chaubal (21) showed that
the carbon source, the viscosity of the medium, and the temperature of
incubation can induce the motility of S. enterica Pullorum, thought to be nonmotile and nonflagellated. Shigella spp.
are described as nonflagellated and nonmotile organisms. However, Giron
(17) detected motility and flagella by EM in four strains and two clinical isolates, depending on the culture conditions under
which temperature, salt, glucose, oxygen, or agar concentrations were
modified. Tominaga et al. (46) showed the presence of intact cryptic flagellin genes in nonflagellated Shigella flexneri
and Shigella sonnei strains. These genes produced
normal-type flagella in an E. coli 
fliC strain. Their
results suggested a loss of the expression potential of flagellar
genes, probably by various mutations and/or gene rearrangements. It
would be interesting to investigate the role of mucus as an inducing
factor for flagellal expression.
In order to study the variability of flagellin genes, the fliC gene was sequenced in three strains representing different profiles obtained by Southern blotting. Sequencing showed high conservation in the N-terminal and C-terminal regions and pronounced variability in the central domain of the flagellin protein (Fig. 3). The N- and C-terminal parts are responsible for secretion and polymerization of flagella, whereas the central region constitutes the surface-exposed antigenic part of the flagellar filament as described by Winstanley and Morgan (52), but flagellin may vary considerably among species in both amino acid sequence and size (49, 52). The deletion of amino acids in the variable domain of the serogroup X reference strain suggests, analogous to other bacteria, that the central region plays no role in the structure of flagella since this strain possesses flagella. Mutations of the flagellin gene do not account for the absence of flagella since the flagellin gene of the flagellated 79-685 strain is strictly identical to that of the nonflagellated serogroup C reference strain.
Different methods have been developed to study the epidemiology of C. difficile or to identify or type strains. Analysis of restriction patterns of DNA of clinical isolates has been used for investigations of epidemiology and typing of C. difficile-associated diarrhea (24). Pantosti et al. (37) used the electrophoretic patterns of extracted proteins to characterize C. difficile strains from various sources and showed correlation between certain electrophoretic patterns and virulence. Delmée et al. (13) compared serogrouping of C. difficile by slide agglutination with rabbit antisera and polyacrylamide gel electrophoresis of whole-cell proteins, permitting correlation between the two typing systems and establishment of a single classification. Recently, new molecular techniques have been developed to type C. difficile strains based on DNA polymorphism. DNA pattern profiles have been obtained by PCR amplification of a specific chromosomal region such as the rRNA gene (4) or the 16S-23S rRNA gene intergenic spacer region (44). Another molecular method, based on DNA polymorphism, has been found to be useful to distinguish strains of C. difficile, namely, arbitrary primed PCR, also called random amplified polymorphic DNA analysis. Arbitrary primed PCR or random amplified polymorphic DNA analysis has been used as an efficient discriminative method for investigation of nosocomial outbreaks of C. difficile-associated diarrhea (2, 3, 5; F. Barbut, N. Mario, J. Frottier, and J. C. Petit, Letter, Eur. J. Clin. Microbiol. Infect. Dis. 12:794-795, 1993).
In our study, we have used the PCR-RFLP method to study genetic diversity among C. difficile strains. With this molecular technique, correlation between RFLP groups and serogroups was clear for certain serogroups. Serogroups C and I are represented by a single RFLP group, group I, and serogroups D, F, and G are represented by RFLP groups VIII, II, and VII, respectively. We noticed that serogroup F exclusively possesses pattern II, which is not shared by any other strain in this study. Similar data were shown by Rupnik (41) concerning the toxinotype of strains belonging to serogroup F. Other serogroups (A, B, H, K, S3, and X) were subdivided into several RFLP groups. However, six RFLP groups, corresponding each to a single serogroup, could be used to confirm some strains. Indeed, RFLP groups II, III, IV and V, and VI and IX were correlated to serogroups F, B, A, and X, respectively. Some conclusions could also be drawn concerning the state of flagellation and toxigenesis, but a larger number of strains need to be investigated to draw more-definitive conclusions.
The study of flagellin gene diversity has been also carried out with other bacteria, such as C. jejuni (33, 35, 43), P. aeruginosa (32, 50), S. enterica (10), Vibrio parahaemolyticus (29), H. pylori (34), or Burkholderia cepacia (47, 51). The results have clearly demonstrated the pronounced genetic diversity of the flagellar gene of various bacteria. The PCR-RFLP method has sometimes been used with success to differentiate several bacterial flagellal types from isolates. However, in certain cases, this procedure does not appear sufficient to type bacterial species.
The flagellin genes are excellent biomarkers with which to study strain diversity. The particular structure of the flagellin gene, with terminal conserved regions allowing gene amplification by PCR, allows analysis by RFLP and sequencing to study the variations in the central region. The PCR-RFLP procedure is rapid, highly specific, and reproducible. If a vaccine is to be developed for C. difficile disease based on different proteins, the preparations should include a mixture of flagellin proteins from major RFLP groups to allow the best possible protection. C. difficile flagella could play a role in intestinal colonization during the first stage of pathogenesis. Colonization is induced in response to environmental conditions. It is likely that production of flagella could be under the control of a system which may be turned on or off by various factors in the gut environment. Important questions remain to be explored as to the identity of these factors and what role flagella and motility play in the pathogenic scheme.
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ACKNOWLEDGMENTS |
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This work was supported in part by the FAIR Programme of the European Union (contract CT95-0433); program ACC-SV6 (Actions Concertées Coordonnées des Sciences du Vivant) of the Ministère de l'Education Nationale, de l'Enseignement Supérieur et de la Recherche of France; and a Medical Research Council Programme grant (G9122850).
We thank Michel Lemullois and Danielle Jaillard (Université de Paris-Sud, Orsay, France) for helping us with EM.
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FOOTNOTES |
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* Corresponding author. Mailing address: Faculté de Pharmacie, Département de Microbiologie, Université de Paris-Sud, 5, rue J. B. Clément, 92296 Châtenay-Malabry Cedex, France. Phone: (33)-1 46 83 55 49. Fax: (33)-1 46 83 58 83. E-mail: marie-claude.barc{at}cep.u-psud.fr.
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Materials and Methods
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Discussion
References
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Journal of Clinical Microbiology, September 2000, p. 3179-3186, Vol. 38, No. 9
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