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Laboratory of Microbiology, Wageningen University, Hesselink van Suchtelenweg 4, 6703 CT Wageningen, The Netherlands
Correspondence
Muriel Derrien
muriel.derrien{at}wur.nl
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Published online ahead of print on 14 May 2004 as DOI 10.1099/ijs.0.02873-0.
The GenBank/EMBL/DDBJ accession number for the 16S rRNA gene sequence of Akkermansia muciniphila MucT is AY271254.
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). A mucus layer covers the GI tract, providing a protective barrier for the underlying epithelium against pathogenic micro-organisms, as well as chemical, physical or enzymic damage. Mucus is a viscous gel mainly composed of high-molecular-mass glycoproteins, termed mucins. Mucins are composed of a peptide core rich in serine and threonine residues that is decorated by oligosaccharides linked via O- or N-glycosidic bonds. The oligosaccharides are composed of one or more of four primary sugars (N-acetylglucosamine, N-acetylgalactosamine, galactose and fucose) and are terminated by sialic acids or sulfate groups. This mucus layer is considered to be an ecological niche for intestinal microbiota. However, the association of microbiota with the mucus is not well understood. Mucus can serve as a barrier to protect the underlying epithelium from the attachment of pathogens and also as a source of nutrients for commensal bacteria. Degradation of mucin is regarded as a pathogenicity factor since loss of the protective mucus layer may expose GI tract cells to pathogens (Ruseler-van Embden et al., 1995; Zhou et al., 2001). However, mucin also constitutes a carbon and energy source for intestinal microbiota. It has been estimated that 1 % of colonic microbiota is able to degrade host mucin using enzymes (e.g. glycosidases and sulfatases) that can degrade the oligosaccharide chains (Hoskins & Boulding, 1981). Despite the apparent low level of mucin-degrading bacteria, these species provide nutrients for other resident bacteria, which can use the monosaccharides or amino acids released from mucin degradation. Based on their capacity to grow on mucin-containing media, isolates belonging to the genera Ruminococcus, Bacteroides, Bifidobacterium and Clostridium have been shown to degrade mucin (Salyers et al., 1977). By measuring the release of reducing sugar monomers from the mucin polymer, it was observed that only mixed cultures of faecal bacteria were able to degrade mucin by more than 90 %, whereas pure cultures of Bacteroides fragilis, Bifidobacterium longum and Clostridium perfringens showed only partial degradation (Willis et al., 1996). It is therefore likely that, in vivo, a co-operative process is required to achieve efficient degradation of the complex structure of mucin.
The introduction of high-resolution molecular techniques has improved analyses of complex microbial ecosystems. The most important advance has been the use of the 16S rRNA gene as a molecular fingerprint to analyse microbial diversity. Molecular approaches have indicated that a lack of knowledge regarding cultivation conditions has hampered our view of the intestinal microbiota (Vaughan et al., 2000). As a consequence, a substantial proportion of the microbiota has not yet been cultured or described (Zoetendal et al., 1998; Suau et al., 1999); this may be due mainly to the lack of appropriate cultivation techniques. However, new, alternative and improved cultivation approaches are continuously being developed and recently a number of novel species and genera have been cultured from the GI tract: Roseburia intestinalis (Duncan et al., 2002), Campylobacter hominis (Lawson et al., 2001), Ruminococcus luti (Simmering et al., 2002), Anaerostipes caccae (Schwiertz et al., 2002), Dorea longicatena (Taras et al., 2002) and Victivallis vadensis (Zoetendal et al., 2003).
In the present study, mucin-degrading bacteria from human faeces were enriched using a most probable number (MPN) approach in which the medium contained mucin as the sole carbon and energy source. The enrichments were analysed by denaturing gradient gel electrophoresis (DGGE) of PCR-amplified 16S rRNA gene sequences. A single DGGE type dominated all the positive MPN enrichments. The organism corresponding to the dominant DGGE type was isolated and characterized; it represents a novel intestinal bacterium, strain MucT, that is able to use gastric mucin in pure culture.
A faecal sample from a healthy adult volunteer was freshly collected in a polyethylene bag and 0·5 g was diluted into 9 ml sterile anaerobic Ringer's solution containing 0·5 g cysteine l–1. This suspension was thoroughly mixed and serially diluted (10-fold) in Ringer's. Each dilution (1 ml) was inoculated in triplicate into 9 ml bicarbonate-buffered medium. This basal medium contained (l–1): 0·4 g KH2PO4; 0·53 g Na2HPO4; 0·3 g NH4Cl; 0·3 g NaCl; 0·1 g MgCl2.6H2O; 0·11 g CaCl2; 1 ml alkaline trace element solution; 1 ml acid trace element solution; 1 ml vitamin solution; 0·5 mg resazurin; 4 g NaHCO3; 0·25 g Na2S.7–9H2O. The trace element and vitamin solutions were as described previously (Stams et al., 1993). All compounds were autoclaved, except the vitamins, which were filter-sterilized. This basal medium was supplemented with 0·7 % (v/v) clarified, sterile rumen fluid and 0·25 % (v/v) commercial hog gastric mucin (Type III; Sigma), purified by ethanol precipitation as described previously (Miller & Hoskins, 1981). This medium is further referred to as mucin medium. Unless indicated, incubations were done in serum bottles sealed with butyl-rubber stoppers at 37 °C under anaerobic conditions provided by a gas phase of 182 kPa (1·8 atm) N2/CO2 (80 : 20, v/v). Enrichments were done in 30 ml serum bottles with 10 ml liquid volume. Negative controls comprised one series of mucin media that was not inoculated and another series that was inoculated, but not supplemented with mucin. Mucin-degrading bacteria were quantified using the MPN technique (n=3). The soft agar technique was used to isolate a pure culture as follows: the highest dilution where growth was observed was serially diluted in phosphate buffer (pH 7) until 10–9 dilution and the 10–6 to 10–9 dilutions were re-inoculated into the same medium containing 0·75 % agar (agar noble; Difco). Single colonies were picked, grown in mucin medium and re-inoculated in soft agar mucin medium. This step was repeated until purity.
Generation times were determined in mucin medium and growth was analysed in triplicate by measuring absorbance at 600 nm. The optimum pH and temperature were measured in triplicate on brain–heart infusion (BHI; Difco) supplemented with 1 mM Na2S. Temperatures tested were 4–45 °C, at intervals of 5 °C; growth was determined at pH 5–9, at intervals of 0·5 pH units (adjusted with HCl or NaOH) at 37 °C. Cultures were incubated for at least 1 month.
Potential substrates for growth were tested at a concentration of 10 mM in the same liquid basal medium or in basal medium supplemented with peptone, tryptone, casitone and yeast extract at a concentration of 0·5 or 2 g l–1. Cultures were incubated for up to 4 weeks. Human gastric mucin was isolated from HT-29 MTX human intestinal cell lines and this mucin was added to the basal liquid medium at a concentration of 0·05 %. Rich media BHI and Columbia broth (Difco) and 16 g Wilkens–Chalgren broth (WC broth; Oxoid) l–1 were also tested as growth substrates. To test the origin of the nitrogen source, the solution containing NH4Cl was not added to the mucin medium.
Cell morphology, motility and spore formation were investigated using phase-contrast microscopy. The Gram reaction was assessed using Gram staining as described previously (Plugge et al., 2000). To test for the presence of a capsule, an Indian ink suspension was used.
For TEM of strain MucT, cells were fixed with 0·25 % glutaraldehyde. Negative staining was performed on 400 copper mesh grids with glow-discharged parladion carbon-support film. Micrographs were recorded at a magnification of 40 000x on a JEOL 1010 electron microscope operating at 80 kV. For SEM, droplets of strain MucT were put onto poly-L-lysine-coated Nucleopore polycarbonate membranes (Costar). These membranes were fixed for 1 h in 4 % glutaraldehyde in growing medium. Specimens were dehydrated in a graded series of ethanol and critical-point dried with carbon dioxide. The samples were glued onto a sample holder using carbon adhesive tabs. Samples were sputter-coated with 10 nm platinum in a dedicated preparation chamber (CT 1500 HF) and analysed with a field emission SEM (JEOL 6300 F) at 5 kV.
The G+C content of DNA of strain MucT was determined at the DSMZ (Deutsche Sammlung von Mikroorganismen und Zellkulturen, Braunschweig, Germany) by HPLC (Mesbah et al., 1989).
To monitor the dynamics of the human faecal mucin-degrading population, DGGE analysis of 16S rRNA gene amplicons was performed. DNA was extracted from the faecal sample, enrichment cultures and a pure culture isolated from the highest dilution. DNA isolation and the amplification of the V6 to V8 regions of the 16S rRNA gene from these samples were performed as described previously (Zoetendal et al., 1998). PCR fragments were separated by DGGE consisting of 8 % (v/v) polyacrylamide (ratio of acrylamide to bisacrylamide, 37·5 : 1) and 0·5x Tris/acetate/EDTA (pH 8·0) (TAE) buffer; 100 % denaturing acrylamide was defined as 7 M urea and 40 % formamide. Gradients of 38–48 % were used to separate products amplified with universal primers. After migration of the PCR products at 85 V for 16 h, the gels were stained with AgNO3 as described previously (Sanguinetti et al., 1994).
PCR on the 16S rRNA gene of strain MucT was performed with universal primers 11f and 1510r (Lane, 1991). The following PCR programme was used: 94 °C for 5 min; 40 cycles consisting of 94 °C for 1 min 30 s, 48 °C for 30 s, and 68 °C for 1 min 30 s; and finally 68 °C for 7 min. PCR products were purified and concentrated with the Qiaquick PCR purification kit (Qiagen) according to the manufacturer's instructions. The purified 16S rRNA gene product was sequenced on both strands using infrared Dye 41-labelled primers 7f, 342r, 805f, 1100r and 1510r (Lane, 1991), and 968f (Nübel et al., 1996). One extra primer, Muc1 (5'-GGA AAC CCT GAT GGT GCG-3'), which targets a 339 bp specific region of the 16S rRNA gene sequence of strain MucT, was designed to obtain unambiguous results. Sequences were automatically analysed on a LI-COR DNA sequencer 4000L and corrected manually. Pairwise sequence alignment was performed with the program DNASTAR. The 16S rRNA gene sequence was compared to sequences from GenBank using the program BLASTN 2.0, available through the National Centre for Biotechnology Information (NCBI) website (http://www.ncbi.nlm.nih.gov/blast/). The ARB software package was used to align cloned sequences and 16S rRNA gene sequences of nearest relatives (Strunk & Ludwig, 1995). A phylogenetic tree was constructed with ARB using the neighbour-joining method. The distance matrix used in the neighbour-joining method included stretches of sequence corresponding to Escherichia coli positions 63–1491.
Substrates and fermentation product concentrations were determined in the culture before and after growth using HPLC and GC methods as described previously (Stams et al., 1993).
The use of serial dilution in an anaerobic medium containing mucin as energy source led to the isolation of a predominant mucin-degrading bacterium from the human faecal sample. As in all MPN studies, the low-dilution cultures presumably gave rise to the fastest-growing organisms under the given culture conditions, whereas the high-dilution cultures supported growth of the numerically dominant organisms. In our study, a single band generated by PCR-DGGE (Fig. 1) dominated all dilutions, indicating that there was one predominant mucin-degrading bacterium and that this was also the fastest-growing mucin-degrading organism. The MPN of mucin-degrading organisms present in this faecal sample was estimated at 8·3±0·3x109 (g faeces)–1. No growth was observed in the uninoculated mucin medium, indicating the sterility of the mucin, nor in the medium inoculated with faecal dilutions but not supplemented with mucin. This indicates that growth occurred solely due to utilization of mucin by the faecal bacteria.
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). In the first dilutions, many bands were detected, but one was predominant. This band became more intense with increasing dilution and, in the highest dilution where growth occurred (10–9), this band was almost unique in the profile. The faecal sample profile showed a band at the same position, suggesting that the enriched micro-organism containing this 16S rRNA gene is the same as the one present in the faecal sample and represents at least 1 % of the total intestinal bacterial community (Muyzer et al., 1993). The mucin-degrading bacterium, whose 16S rRNA gene corresponded to the major DGGE band present in the enrichment (lane 10), was cultured from the highest dilution with growth (10–9) using the soft agar technique. After 6 days, the most predominant colony type, white, was grown in the mucin medium. White colonies were picked, diluted in the mucin medium and transferred into soft agar mucin medium. This purification step was repeated twice. Finally, a single type of white colony appeared. Phase-contrast microscopy revealed only one morphotype, and the DGGE profile of the 16S rRNA gene amplicon of the strain isolated showed the presence of a unique band corresponding to the major band present in the enrichment (Fig. 1, lane 1). The pure culture was designated strain MucT.
An almost-complete 16S rRNA gene sequence of strain MucT was determined (1433 bp). The most similar 16S rRNA gene sequences, which were derived from studies of uncultured colonic bacteria [HuCA18 and HuCC13 (Hold et al., 2002) and L10-6 (Salzman et al., 2002)] were 99 % identical to strain MucT. The cultured bacterium most closely related to strain MucT was Verrucomicrobium spinosum, and this was only distantly related (92 %). A phylogenetic dendrogram based on 16S rRNA gene sequences was constructed; it revealed that strain MucT is related to the genera Prosthecobacter and Verrucomicrobium, which are members of the order Verrucomicrobiales. Thus, MucT belongs to the division Verrucomicrobia and the class Verrucomicrobiae (Fig. 2). The majority of the members of this new division are clones and only a few are cultivated bacteria: a single genus, Verrucomicrobium (Schlesner, 1987), after which the division was named; four species of the genus Prosthecobacter (Prosthecobacter debontii, Prosthecobacter dejongeii, Prosthecobacter fusiformis and Prosthecobacter vanneervenii) (Staley et al., 1976; Hedlund et al., 1997); Opitutus terrae (Chin et al., 2001) and other ultramicrobacteria (Janssen et al., 1997); and the recently described species Victivallis vadensis, the first member of the division Verrucomicrobia to be isolated from the GI tract (Zoetendal et al., 2003). Members of the ‘Verrucomicrobium’ group of bacteria have also been identified in low numbers in human faeces-derived 16S rRNA gene libraries (Wilson & Blitchington, 1996; Suau et al., 1999; Hold et al., 2002). The division Verrucomicrobia is composed of five subdivisions (Hugenholtz et al., 1998) and the genera Prosthecobacter and Verrucomicrobium are part of subdivision 1. Bacteria from these two genera were isolated from freshwater habitats and are both Gram-negative, aerobic and heavily fimbriated. Cells of Verrucomicrobium have many prosthecae, whereas cells of Prosthecobacter have only a single prosthecae. Strain MucT shares some common characteristics; for example, it is Gram-negative and can grow without vitamins. However, strain MucT is distinct among the members of subdivision 1 in that it is strictly anaerobic and cells are oval-shaped in contrast to the other members (see Table 1). On the basis of a phylogenetic analysis, strain MucT does not belong to the Verrucomicrobium or Prosthecobacter clusters and should be considered as a separate phylogenetic branch. It is therefore proposed that strain MucT represents a novel species in a new genus belonging to subdivision 1 of the Verrucomicrobia.
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No growth was observed on glucose, cellobiose, lactose, galactose, xylose, fucose, rhamnose, maltose, succinate, acetate, fumarate, butyrate, lactate, casitone (0·5 %), Casamino acids (0·5 %), tryptone (0·5 %), peptone (0·5 %), yeast extract (0·5 %), proline, glycine, aspartate, serine, threonine, glutamate, alanine, N-acetylglucosamine or N-acetylgalactosamine after 4 weeks incubation. Gastric mucin isolated from human intestinal cell lines and adapted on 10–5 M methotrexate (HT-29 MTX) to produce a high amount of mucin (Lesuffleur et al., 1990) resulted in growth of strain MucT to the same density as with hog gastric mucin. Strain MucT could also grow on rich media, Columbia and BHI, but with a final optical density of half that of the mucin medium. No growth was observed on rich WC anaerobe broth. When peptone, yeast extract, tryptone and casitone (each at 2 g l–1) were added to the basal medium, growth was observed only when the sugars N-acetylglucosamine, N-acetylgalactosamine and glucose were added, although bacterial growth was less than a quarter of that on mucin medium. When the solution containing the nitrogen source was not added to the basal medium supplemented with mucin, strain MucT could grow to the same density, indicating that the isolate utilized mucin as both carbon and nitrogen source. Strain MucT did not produce H2, but acetate, propionate and ethanol were formed from mucin fermentation. No sulfides were produced. Sulfates were released during fermentation of mucin (0·71 mM), demonstrating sulfatase activity. It is presumed that the limited ability of strain MucT to grow on the many substrates tested may be due to the complex structure of the mucin, which is composed of both oligosaccharides and amino acids, and that strain MucT requires a combination of all these components to reach a high density. It is likely that strain MucT produces one or more appropriate glycosidases to degrade the N-acetylgalactosamine and N-acetylglucosamine components from mucin, which might be exposed in the terminal part, and to use them as growth substrates.
Cells of strain MucT were oval-shaped (Fig. 3a), showing a different size depending on the medium. In mucin medium, strain MucT was 640 nm in diameter and 690 nm in length and in BHI, strain MucT was 830 nm in diameter and 1 µm in length. Cells stained Gram-negative. Flagella were not seen on negatively stained EM preparations. Spore formation was never observed. In mucin medium, the organism could grow as single cells or in pairs, but rarely in chains; it often formed aggregates in which a translucent layer of material was observed between organisms. In BHI and Columbia media, this material was rarely, if ever, observed and cells occurred singly or in pairs, but rarely in groups. In basal medium, supplemented with N-acetylglucosamine or N-acetylgalactosamine, together with some sources of proteins (a combination of yeast extract, peptone, tryptone and casitone), cells occurred singly and sometimes in pairs. Cells of strain MucT grown in mucin medium could exclude Indian ink, which is characteristic of capsule-possessing bacteria. EM revealed the existence of filamentous structures on cells grown in mucin medium (Fig. 3b). It is assumed that these filaments are capsular polymers that are used to connect cells together. Since this aggregation is mainly observed in mucin medium, this capsule may aid in adhesion and colonization of mucin-secreting epithelia in the GI tract. On soft agar medium, colonies of strain MucT appeared white and were 0·7 mm in diameter.
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Description of Akkermansia gen. nov.
Akkermansia (Ak.ker.man'si.a. N.L. fem. n. Akkermansia derived from Antoon Akkermans, a Dutch microbiologist recognized for his contribution to microbial ecology).
Cells are oval-shaped, non-motile and stain Gram-negative. Strictly anaerobic. Chemo-organotrophic. Mucolytic in pure culture.
The type species is Akkermansia muciniphila.
Description of Akkermansia muciniphila sp. nov.
Akkermansia muciniphila (mu.ci.ni'phi.la. N.L. neut. n. mucinum mucin; Gr. adj. philos loving; N.L. fem. adj. muciniphila mucin-loving).
Cells are oval-shaped, non-motile and stain Gram-negative. The long axis of single cells is 0·6–1·0 µm, depending on the substrate used. Cells occur singly, in pairs, in short chains and in aggregates. Growth occurs at 20–40 °C and pH 5·5–8·0, with optimum growth at 37 °C and pH 6·5. Strictly anaerobic. Able to grow on gastric mucin, brain–heart infusion and Columbia media, and on N-acetylglucosamine, N-acetylgalactosamine and glucose when these three sugars are in the presence of (each at 2 g l–1) peptone, yeast extract, casitone and tryptone. Cellobiose, lactose, galactose, xylose, fucose, rhamnose, maltose, succinate, acetate, fumarate, butyrate, lactate, casitone, Casamino acids, tryptone, peptone, yeast extract, proline, glycine, aspartate, serine, threonine and glutamate do not support growth. Capable of using mucin as carbon, energy and nitrogen source. Able to release sulfate in a free form from mucin fermentation. In mucin medium, cells are covered with filaments. Growth occurs without vitamins. Colonies appear white with a diameter of 0·7 mm in soft agar mucin medium.
The type strain is MucT (=ATCC BAA-835T=CIP 107961T), isolated from the human intestinal tract. Its DNA G+C content is 47·6 mol%.
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ACKNOWLEDGEMENTS |
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 P. Ruas-Madiedo, M. Gueimonde, M. Fernandez-Garcia, C. G. de los Reyes-Gavilan, and A. Margolles Mucin Degradation by Bifidobacterium Strains Isolated from the Human Intestinal Microbiota Appl. Envir. Microbiol., March 15, 2008; 74(6): 1936 - 1940. [Abstract] [Full Text] [PDF]
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M. Derrien, M. C. Collado, K. Ben-Amor, S. Salminen, and W. M. de Vos The Mucin Degrader Akkermansia muciniphila Is an Abundant Resident of the Human Intestinal Tract Appl. Envir. Microbiol., March 1, 2008; 74(5): 1646 - 1648. [Abstract] [Full Text] [PDF]
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M. C. Collado, M. Derrien, E. Isolauri, W. M. de Vos, and S. Salminen Intestinal Integrity and Akkermansia muciniphila, a Mucin-Degrading Member of the Intestinal Microbiota Present in Infants, Adults, and the Elderly Appl. Envir. Microbiol., December 1, 2007; 73(23): 7767 - 7770. [Abstract] [Full Text] [PDF]
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J. Yoon, Y. Matsuo, S. Matsuda, K. Adachi, H. Kasai, and A. Yokota Rubritalea spongiae sp. nov. and Rubritalea tangerina sp. nov., two carotenoid- and squalene-producing marine bacteria of the family Verrucomicrobiaceae within the phylum 'Verrucomicrobia', isolated from marine animals Int J Syst Evol Microbiol, October 1, 2007; 57(10): 2337 - 2343. [Abstract] [Full Text] [PDF]
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Y.-J. Choo, K. Lee, J. Song, and J.-C. Cho Puniceicoccus vermicola gen. nov., sp. nov., a novel marine bacterium, and description of Puniceicoccaceae fam. nov., Puniceicoccales ord. nov., Opitutaceae fam. nov., Opitutales ord. nov. and Opitutae classis nov. in the phylum 'Verrucomicrobia' Int J Syst Evol Microbiol, March 1, 2007; 57(3): 532 - 537. [Abstract] [Full Text] [PDF]
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M. Scheuermayer, T. A. M. Gulder, G. Bringmann, and U. Hentschel Rubritalea marina gen. nov., sp. nov., a marine representative of the phylum 'Verrucomicrobia', isolated from a sponge (Porifera). Int J Syst Evol Microbiol, September 1, 2006; 56(Pt 9): 2119 - 2124. [Abstract] [Full Text] [PDF]
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P. Sangwan, S. Kovac, K. E. R. Davis, M. Sait, and P. H. Janssen Detection and Cultivation of Soil Verrucomicrobia Appl. Envir. Microbiol., December 1, 2005; 71(12): 8402 - 8410. [Abstract] [Full Text] [PDF]
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