Key Points
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Plant polysaccharides of dietary origin are a major energy source for the dense anaerobic microbial communities that inhabit the mammalian large intestine and rumen. Microbial fermentation products, in turn, provide energy and nutrients to the host that would otherwise be unavailable.
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Only a few species of gut microorganism have the ability to act as primary degraders of insoluble substrates, such as plant cell walls, but many others benefit from cross-feeding interactions.
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The organization and spectrum of enzymes, substrate-binding modules and transport systems largely defines the types of substrate that can be exploited by a given gut bacterium, but only two gut anaerobes have so far been the subject of detailed functional studies.
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In the rumen cellulolytic bacterium Ruminococcus flavefaciens (phylum Firmicutes) enzymes that are involved in the degradation of insoluble plant cell walls are organized into a cellulosome complex that is bound to the cell surface.
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By contrast, the sequestration and breakdown of soluble starch molecules is achieved mainly in the periplasm of the Gram-negative human colonic species Bacteroides thetaiotaomicron.
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More information from microbial genomics is needed to reveal the diversity of the systems that are involved in polysaccharide utilization, and thus the extent of the variation in these two paradigms among gut bacteria.
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Gut bacteria are largely untapped sources of enzymes and binding domains that have potential for biotechnological application.
Abstract
The microbiota of the mammalian intestine depend largely on dietary polysaccharides as energy sources. Most of these polymers are not degradable by the host, but herbivores can derive 70% of their energy intake from microbial breakdown — a classic example of mutualism. Moreover, dietary polysaccharides that reach the human large intestine have a major impact on gut microbial ecology and health. Insight into the molecular mechanisms by which different gut bacteria use polysaccharides is, therefore, of fundamental importance. Genomic analyses of the gut microbiota could revolutionize our understanding of these mechanisms and provide new biotechnological tools for the conversion of polysaccharides, including lignocellulosic biomass, into monosaccharides.
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References
Van Soest, P. J. Nutritional Ecology of the Ruminant 2nd edn (Cornell Univ. Press, 1984).
Lynd, L. R., Weimer, P. J., van Zyl., W. H. & Pretorius, I. S. Microbial cellulose utilization: fundamentals and biotechnology. Microbiol. Mol. Biol. Rev. 66, 506–577 (2002).
Bergman, E. N. Energy contributions of volatile fatty acids from the gastrointestinal tract in various species. Physiol. Rev. 70, 567–590 (1990).
McIntyre, A., Gibson, P. R. & Young, G. P. Butyrate production from dietary fiber and protection against large bowel cancer in a rat model. Gut 34, 386–391 (1993).
Pryde, S. E., Duncan, S. H., Hold, G. L., Stewart, C. S. & Flint, H. J. The microbiology of butyrate formation in the human colon. FEMS Microbiol. Lett. 217, 133–139 (2002).
Macfarlane, G. T. & Gibson, G. R. in Gastrointestinal Microbiology Vol. 1 (eds Mackie, R. I. & White, B. A.) 269–318 (Chapman and Hall, London, 1997).
Flint, H. J., Duncan, S. H., Scott, K. P. & Louis, P. Interactions and competition within the microbial community of the human large intestine: links between diet and health. Environ. Microbiol. 9, 1101–1111 (2007).
Duchmann, R. et al. T cell specificity and cross-reactivity towards enterobacteria, Bacteroides, Bifidobacterium and antigens from resident intestinal flora in humans. Gut 44, 812–818 (1999).
Hooper, L. V. et al. Molecular analysis of commensal host–microbial relationships in the intestine. Science 291, 881–884 (2001).
Ley, R. E., Turnbaugh, P. J., Klein, S. & Gordon, J. I. Microbial ecology — human gut microbes associated with obesity. Nature 444, 1022–1023 (2006).
Turnbaugh, P. J. et al. An obesity-associated gut microbiome with increased capacity for energy harvest. Nature 444, 1027–1031 (2006). Proposed that the gut microbiota of genetically obese ( ob/ob ) mice recover energy from the diet more efficiently than the microbiota from lean mice.
Duncan, S. H. et al. Reduced dietary intake of carbohydrate by obese subjects results in decreased concentrations of butyrate and butyrate-producing bacteria in feces. Appl. Environ. Microbiol. 73, 1073–1078 (2007).
Kruse, H. P., Kleessen, B. & Blaut, M. Effects of inulin on faecal bifidobacteria in human subjects. Br. J. Nutr. 82, 375–382 (1999).
Sonnenburg, J. L. et al. Glycan foraging in vivo by an intestine-adapted bacterial symbiont. Science 307, 1955–1959 (2005).
Scott, K. P., Martin, J. C., Campbell, G., Mayer, C. D. & Flint, H. J. Whole-genome transcription profiling reveals genes up-regulated by growth on fucose in the human gut bacterium Roseburia inulinivorans. J. Bacteriol. 188, 4340–4349 (2006).
McWilliam Leitch, E. C., Walker, A. W., Duncan, S. H., Holtrop, G. & Flint, H. J. Selective colonisation of insoluble substrates by human faecal bacteria. Environ. Microbiol. 72, 667–679 (2006). In vitro demonstration, based on 16S rRNA sequences, that primary colonizers from the human colon are highly substrate-specific.
Eberhardt, R. Y., Gilbert, H. J. & Hazlewood, G. P. Primary sequence and enzymatic properties of two modular endoglucanases, Cel5A and Cel45A, from the anaerobic fungus Piromyces equi. Microbiology 146, 1999–2008 (2000).
Steenbakkers, P. J. M. et al. Noncatalytic docking domains of cellulosomes of anaerobic fungi. J. Bacteriol. 183, 5325–5333 (2001).
Devillard, E. et al. Characterisation of XynB, a modular xylanase from the ruminal protozoan Polyplastron multivesiculatum, bearing a family 22 carbohydrate binding module that binds to cellulose. Biochem J. 373, 495–503 (2003).
Flint, H. J. Polysaccharide breakdown by anaerobic micro-organisms inhabiting the mammalian gut. Adv. Appl. Microbiol. 56, 89–120 (2004).
Dethlefsen, L., Eckburg, P. B., Bik, E. M. & Relman, D. A. Assembly of the human intestinal microbiota. Trends Ecol. Evol. 21, 517–523 (2006).
Suau, A. et al. Direct analysis of genes encoding 16S rRNA from complex communities reveals many novel molecular species within the human gut. Appl. Environ. Microbiol. 24, 4799–4807 (1999).
Tajima, K. et al. Rumen bacterial diversity as determined by sequence analysis of 16S rDNA libraries. FEMS Microbiol. Ecol. 29, 159–169 (1999).
Daly, K., Stewart, C. S., Flint, H. J. & Shirazi-Beechey, S. P. Bacterial diversity within the equine large intestine as revealed by molecular analysis of cloned 16S rRNA genes. FEMS Microbiol. Ecol. 38, 141–151 (2001).
Hold, G. L., Pryde, S. E., Russell, V. J., Furrie, E. & Flint, H. J. Assessment of microbial diversity in human colonic samples by 16S rDNA sequence analysis. FEMS Microbiol. Ecol. 39, 33–39 (2002).
Leser, T. D. et al. Culture-independent analysis of gut bacteria: the pig gastrointestinal tract microbiota revisited. Appl. Environ. Microbiol. 68, 673–690 (2002).
Eckburg, P. B. et al. Diversity of the human intestinal microbial flora. Science 308, 1635–1638 (2005). The largest study of 16S rRNA sequence diversity in a gut community, which revealed more than 395 bacterial phylotypes in samples from 3 subjects.
Morris, E. J. & van Gylswyk, N. O. Comparison of the action of rumen bacteria on cell walls from Eragrostis tef. J. Agric. Sci. (Camb.) 95, 313–323 (1980). Demonstration that only cellulolytic bacteria cause extensive solubilization of plant cell-wall constituents in pure culture.
Halliwell, G. & Bryant, M. P. The cellulolytic activity of pure strains of bacteria from the rumen of cattle. J. Gen. Microbiol. 32, 441–448 (1968).
Slavin, J. L., Brauer, P. M. & Martlett, J. A. Neutral detergent fiber, hemicellulose and cellulose digestibility in human subjects. J. Nutr. 111, 287–297 (1981).
Robert, C. & Bernalier-Donadille, A. The cellulolytic microflora of the human colon: evidence of microcrystalline cellulose-degrading bacteria in methane-excreting subjects. FEMS Microbiol. Ecol. 46, 81–89 (2003).
Wedekind, K. J., Mansfield, H. R. & Montgomery, L. Enumeration and isolation of cellulolytic and hemicellulolytic bacteria from human feces. Appl. Environ. Microbiol. 54, 1530–1535 (1988).
Coen, J. A. & Dehority, B. A. Degradation and utilization of hemicellulose from intact forages by pure cultures of rumen bacteria. Appl. Microbiol. 20, 362–368 (1970). Describes the synergy between cellulolytic and non-cellulolytic bacteria in the use of hemicellulose breakdown products from grass cell walls.
Osborne, J. M. & Dehority, B. A. Synergism in degradation and utilization of intact forage cellulose, hemicellulose and pectin by three pure cultures of rumen bacteria. Appl. Environ. Microbiol. 55, 2247–2250 (1989).
Belenguer, A. et al. Two routes of metabolic cross-feeding between bifidobacteria and butyrate-producing anaerobes from the human gut. Appl. Environ. Microbiol. 72, 3593–3599 (2006).
Latham, M. J. & Wolin, M. J. Fermentation of cellulose by Ruminococcus flavefaciens in the presence and absence of Methanobacterium ruminantium. Appl. Environ. Microbiol. 34, 297–301 (1977).
Cheng, K. J., Stewart, C. S., Dinsdale, D. & Costerton, J. W. Electron microscopy of bacteria involved in the digestion of plant cell walls. Anim. Feed Sci. Technol. 10, 93–120 (1983).
Walker, A. W. Influence of Substrate and Environmental Factors on Human Gut Microbial Ecology and Metabolism. Thesis, Univ. Aberdeen (2005).
Larue, R., Yu, Z., Parisi, V. A., Egan, A. R. & Morrison, M. Novel microbial diversity adherent to plant biomass in the herbivore gastrointestinal tract, as revealed by ribosomal intergenic spacer analysis and rrs gene sequencing. Environ. Microbiol. 7, 530–543 (2005).
Salyers, A. A., West, S. E. H., Vercellotti, J. R. & Wilkins, T. D. Fermentation of mucins and plant polysaccharides by anaerobic bacteria from the human colon. Appl. Environ. Microbiol. 34, 529–533 (1977).
Krause, D. O., Dalrymple, B. P., Smith, W. J., Mackie, R. I. & McSweeney, C. S. 16S rDNA sequencing of Ruminococcus albus and Ruminococcus flavefaciens: design of a signature probe and its application in adult sheep. Microbiology 145, 1797–1807 (1999).
Antonopoulos, D. A., Russell, W. M. & White, B. A. Phylogenetic reconstruction of Gram-positive organisms based on comparative sequence analysis of molecular chaperones from the ruminal microorganism Ruminococcus flavefaciens FD-1. FEMS Microbiol. Lett. 227, 1–7 (2003).
Julliand, V., de Vaux, A., Millet, L. & Fonty, G. Identification of Ruminococcus flavefaciens as the predominant cellulolytic bacterial species of the equine cecum. Appl. Environ. Microbiol. 65, 3738–3741 (1999).
Nelson, K. E. et al. Phylogenetic analysis of the microbial populations in the wild herbivore gastrointestinal tract: insights into an unexplored niche. Environ. Microbiol. 5, 1212–1220 (2003).
Krause, D. O., Bunch, R. J., Smith, W. J. M. & McSweeney, C. S. Diversity of Ruminococcus strains: a survey of genetic polymorphisms and plant digestibility. J. Appl. Microbiol. 86, 487–495 (1999).
Bayer, E. A., Shimon, L. J., Shoham, Y. & Lamed, R. Cellulosomes-structure and ultrastructure. J. Struct. Biol. 124, 221–234 (1998).
Bayer, E. A., Belaich, J. P., Shoham, Y. & Lamed, R. The cellulosomes: multi-enzyme machines for degradation of plant cell wall polysaccharides. Annu. Rev. Microbiol. 58, 521–554 (2004).
Doi, R. H. & Kosugi, A. Cellulosomes: plant cell wall degrading enzyme complexes. Nature Rev. Microbiol. 2, 541–551 (2004).
Pettipher, G. L. & Latham, M. J. Characteristics of enzymes produced by Ruminococcus flavefaciens that degrade plant cell walls. J. Gen. Microbiol. 110, 21–27 (1979).
Doerner, K. C. & White, B. A. Assessment of the endo-1,4-beta-glucanase components of Ruminococcus flavefaciens FD-1. Appl. Environ. Microbiol. 56, 1844–1850 (1990).
Flint, H. J., Martin, J., McPherson, C. A., Daniel, A. & Zhang, J. X. A bifunctional enzyme having separate xylanase and beta (1,3-1,4) glucanase domains, encoded by the xynD gene of Ruminococcus flavefaciens. J. Bacteriol. 175, 2943–2951 (1993).
Aurilia, V. et al. Three multidomain esterases belonging to the plant cell wall degrading enzyme system of the rumen cellulolytic bacterium Ruminococcus flavefaciens 17 carry divergent dockerin sequences. Microbiology 146, 1391–1397 (2000).
Kirby, J., Martin, J. C., Daniel, A. S. & Flint, H. J. Dockerin-like sequences in cellulases and xylanases from the rumen cellulolytic bacterium Ruminococcus flavefaciens. FEMS Microbiol. Lett. 149, 213–219 (1997).
Lamed, R., Naimark, J., Morgenstern, E. & Bayer, E. A. Specialized cell surface structures in cellulolytic bacteria. J. Bacteriol. 169, 3792–3800 (1987). Key evidence that cellulosome structures occur widely among phylogenetically diverse cellulolytic bacteria.
Bayer, E. A., Morag, E. & Lamed, R. The cellulosome — a treasure-trove for biotechnology. Trends Biotechnol. 12, 378–386 (1994).
Ding, S. Y. et al. Cellulosomal scaffoldin-like proteins from Ruminococcus flavefaciens. J. Bacteriol. 183, 1945–1953 (2001). The first report of scaffoldins and cellulosome organization in a gut bacterium.
Rincon, M. T. et al. Novel organisation and divergent dockerin specificities in the cellulosome system of Ruminococcus flavefaciens. J. Bacteriol. 185, 703–713 (2003).
Rincon, M. T. et al. Unconventional mode of attachment of the Ruminococcus flavefaciens cellulosome to the cell surface. J. Bacteriol. 187, 7569–7578 (2005).
Rincon, M. T., McCrae, S. I., Kirby, J., Scott, K. P. & Flint, H. J. EndB, a multidomain cellulase from the rumen cellulolytic bacterium Ruminococcus flavefaciens 17, binds cellulose via a novel cellulose binding domain and to a 130 kDa R. flavefaciens protein via a dockerin domain. Appl. Environ. Microbiol. 67, 4426–4431 (2001).
Jindou, S. et al. Conservation and divergence in cellulosome architecture between two strains of Ruminococcus flavefaciens. J. Bacteriol. 188, 7971–7976 (2006).
Rincon, M. T. et al. Novel structural and catalytic elements in the Ruminococcus flavefaciens cellulosome revealed by genome analysis. Reprod. Nutr. Dev. 46 (Suppl. 1), S57 (2006).
Rincón, M. T. et al. ScaC, an adaptor protein carrying a novel cohesin that expands the dockerin-binding repertoire of the Ruminococcus flavefaciens 17 cellulosome. J. Bacteriol. 186, 2576–2585 (2004).
Rincon, M. T. et al. A novel cell-surface anchored cellulose-binding protein encoded by the sca gene cluster of Ruminococcus flavefaciens. J. Bacteriol. 189, 4774–4783 (2007).
Stewart, C. S., Duncan, S. H. & Flint, H. J. The properties of forms of Ruminococcus flavefaciens which differ in their ability to degrade cotton cellulose. FEMS Microbiol. Lett. 72, 47–50 (1990).
Devillard, E. et al. Ruminococcus albus 8 mutants defective in cellulose degradation are deficient in two processive endocellulases, Cel48A and Cel9B, both of which possess a novel modular architecture. J. Bacteriol. 186, 136–145 (2004).
Xu, Q. et al. A novel family of carbohydrate-binding modules identified with Ruminococcus albus proteins. FEBS Lett. 566, 11–16 (2004).
Ohara, H., Karita, S., Kimura, T., Sakka, K. & Ohmiya, K. Characterisation of the cellulolytic complex (cellulosome) from Ruminococcus albus. Biosci. Biotechnol. Biochem. 64, 254–260 (2000).
Pegden, R. S., Larson, M. A., Grant, R. J. & Morrison, M. Adherence of the gram-positive bacterium Ruminococcus albus to cellulose and identification of a novel form of cellulose-binding protein which belongs to the Pil family of proteins. J. Bacteriol. 180, 5921–5927 (1998).
Morrison, M. & Miron, J. Adhesion to cellulose by Ruminococcus albus: a combination of cellulosomes and Pil-proteins? FEMS Microbiol. Lett. 185, 109–115 (2000).
Rakotoarivinina, H. et al. The Ruminococcus albus pilA1–pilA2 locus: expression and putative role of two adjacent pil genes in pilus formation and bacterial adhesion to cellulose. Microbiology 151, 1291–1299 (2005).
Shinkai, T. & Kobayashi, Y. Localization of ruminal cellulolytic bacteria on plant fibrous material as determined by fluorescence in situ hybridisation and real time PCR. Appl. Environ. Microbiol. 73, 1646–1652 (2006).
Nelson, K. E. et al. The Fibrobacter succinogenes S85 genome sequencing project. Reprod. Nutr. Dev. 42 (Suppl.1), S44 (2002).
Jun, H. S., Qi, M., Ha, J. K. & Forsberg, C. W. Fibrobacter succinogenes, a dominant fibrolytic ruminal bacterium: transition to the post genomic era. Asian–australas. J. Anim. Sci. 20, 802–810 (2007).
Jun, H. S., Qi, M., Gong, J., Egbosimba, E. E. & Forsberg, C. W. Outer membrane proteins of Fibrobacter succinogenes with potential roles in adhesion to cellulose and in cellulose digestion. J. Bacteriol. 189, 6806–6815 (2007).
Maglione, G. J., Russell, J. B. & Wilson, D. B. Kinetics of cellulose digestion by Fibrobacter succinogenes. Appl. Environ. Microbiol. 63, 665–669 (1997).
Dongowski, G., Lorenz, A. & Anger, H. Degradation of pectins with different degrees of esterification by Bacteroides thetaiotaomicron isolated from the human gut flora. Appl. Environ. Microbiol. 66, 1321–1327 (2000).
Hooper, L. V., Midvedt, T. & Gordon, J. I. How host–microbial interactions shape the nutrient environment of the mammalian intestine. Annu. Rev. Nutr. 22, 283–307 (2002).
Chassard, C., Gaillard-Martinie, B. & Bernalier-Donadille, C. Interaction between H2-producing and non-H2-producing cellulolytic bacteria from the human colon. FEMS Microbiol. Lett. 242, 339–344 (2005).
Chassard, C., Goumy, V., Leclerc, M., Del'homme, C. & Bernalier-Donadille, A. Characterization of the xylan-degrading microbial community from human faeces. FEMS Microbiol. Ecol. 61, 121–131 (2007).
Xu, J. et al. A genomic view of the human–Bacteroides thetaiotaomicron symbiosis. Science 299, 2074–2076 (2003). Description of the genome sequence of B. thetaiotaomicron , which revealed 163 susCD paralogues and the largest number of genes related to carbohydrate utilization of any bacterium so far studied.
Anderson, K. L. & Salyers, A. A. Biochemical evidence that starch breakdown by Bacteroides thetaiotaomicron involves outer-membrane starch-binding sites and periplasmic starch-degrading enzymes. J. Bacteriol. 171, 3192–3198 (1989).
Reeves, A. R., D'Elia, J. N., Frias, J. & Salyers, A. A. A Bacteroides thetaiotaomicron outer membrane protein that is essential for utilization of malto-oligosaccharides and starch. J. Bacteriol. 178, 823–830 (1996).
Reeves, A. R., Wang, G. R. & Salyers, A. A. Characterization of four outer membrane proteins that play a role in utilization of starch by Bacteroides thetaiotaomicron. J. Bacteriol. 179, 643–649 (1997). A key paper that reports the structure of the sus gene cluster and, together with reference 82, explains the essential roles of outer-membrane starch-binding proteins in starch utilization by B. thetaiotaomicron.
Cho, K. H. & Salyers, A. A. Biochemical analysis of interactions between outer membrane proteins that contribute to starch utilization by Bacteroides thetaiotaomicron. J. Bacteriol. 183, 7224–7230 (2001).
Shipman, J. A., Berleman, J. E. & Salyers, A. A. Characterization of four outer membrane proteins involved in binding starch to the cell surface of Bacteroides thetaiotaomicron. J. Bacteriol. 182, 5365–5372 (2000).
Shipman J. A., Cho, K. H., Siegel, H. A. & Salyers, A. A. Physiological characterization of SusG, an outer membrane protein essential for starch utilization by Bacteroides thetaiotaomicron. J. Bacteriol. 181, 7206–7211 (1999).
D'Elia, J. N. & Salyers, A. A. Contribution of a neopullulanase, a pullulanase and an alpha-glucosidase to growth of Bacteroides thetaiotaomicron on starch. J. Bacteriol. 178, 7173–7179 (1996).
Spence, C., Wells, W. G. & Smith, C. J. Characterization of the primary starch utilization operon in the obligate anaerobe Bacteroides fragilis: regulation by carbon source and oxygen. J. Bacteriol. 188, 4663–4672 (2006).
Xu, J. & Gordon, J. I. Honor thy symbionts. Proc. Natl Acad. Sci. USA 100, 10452–10459 (2003).
Sonnenburg, E. D. et al. A hybrid two component system of a prominent human gut symbiont couples glycan sensing in vivo to carbohydrate metabolism. Proc. Natl Acad. Sci. USA 103, 8834–8839 (2006).
McCarthy, R. E., Kotarsky, S. F. & Salyers, A. A. Location and characteristics of enzymes involved in the breakdown of polygalacturonic acid by Bacteroides thetaiotaomicron. J. Bacteriol. 161, 493–499 (1985).
Cheng, Q., Yu, M. C., Reeves, A. R. & Salyers, A. A. Identification and characterization of a Bacteroides gene, csuF, which encodes an outer membrane protein that is essential for growth on chondroitin sulphate. J. Bacteriol. 177, 3721–3727 (2001).
Weaver, J., Whitehead, T. R., Cotta, M. A., Valentine, P. C. & Salyers, A. A. Genetic analysis of a locus on the Bacteroides ovatus chromosome which contains xylan utilization genes. Appl. Environ. Microbiol. 58, 2764–2770 (1992).
Gasparic, A., Daniel, A., Martin, J. & Flint, H. J. A xylan hydrolase gene cluster from Prevotella ruminicola: sequence relationships, oxygen sensitivity and synergistic interactions of a novel exoxylanase. Appl. Environ. Microbiol. 61, 2958–2964 (1995).
Ramsak, A. et al. Unravelling the genetic diversity of ruminal bacteria belonging to the CFB phylum. FEMS Microbiol. Ecol. 33, 69–79 (2000).
Miyazaki, K. et al. Involvement of the two component regulatory protein XynR in positive control of xylanase gene expression in the ruminal anaerobe Prevotella bryantii B14. J. Bacteriol. 185, 2219–2226 (2003).
Miyazaki, K., Hirase, T., Kojima, Y. & Flint, H. J. Medium to large sized xylo-oligosaccharides are responsible for xylanase induction in Prevotella bryantii B14. Microbiology 151, 4121–4125 (2005).
Miyazaki, K., Martin, J. C., Marinsek-Logar, R. & Flint, H. J. Degradation and utilization of xylans by the rumen anaerobe Prevotella bryantii (formerly P. ruminicola subsp. brevis B14). Anaerobe 3, 373–381 (1997).
Bourne, Y. & Henrissat, B. Glycoside hydrolases and glycosyltransferases: families and functional modules. Curr. Opin. Struct. Biol. 11, 593–600 (2001).
Devillard, E. A. et al. Characterization of XynB, a modular xylanase produced by the anaerobic protozoan Polyplastron multivesiculatum shows close similarity to family 11 xylanases from Gram-positive bacteria. FEMS Microbiol. Lett. 181, 145–152 (1999).
Gilbert, H. J., Hazlewood, G. P., Laurie, J. I., Orpin, C. G. & Xue, G. P. Homologous catalytic domains in a rumen fungal xylanase: evidence for gene duplication and prokaryotic origin. Mol. Microbiol. 6, 2065–2072 (1992).
Ricard, G. et al. Horizontal gene transfer from bacteria to rumen ciliates indicates adaptation to their anaerobic, carbohydrate-rich environment. BMC Genomics 7, 22 (2006).
Kacser, H. & Beeby, R. Evolution of catalytic proteins, or on the origin of enzyme species by means of natural selection. J. Mol. Evol. 20, 38–51 (1984).
Reidel, K., Ritter, J. & Bronnenmeier, K. Synergistic interaction of the Clostridium stercorarium cellulases Avicel I (CelZ) and Avicellase II (CelY) in the degradation of microcrystalline cellulose. FEMS Microbiol. Lett. 147, 239–243 (1997).
Robert, C., Chassard, C., Lawson, P. A. & Bernalier-Donadille, A. Bacteroides cellulosilyticus sp. nov., a cellulolytic bacterium from the human gut microbial community. Int. J. Syst. Evol. Microbiol. 57, 1516–1520 (2007).
MacFarlane, G. T. & Englyst, H. N. Starch utilization by the human large intestinal microflora. J. Appl. Bacteriol. 60, 195–201 (1986).
Ramsay, A. G., Scott, K. P., Martin, J. C., Rincon, M. T. & Flint, H. J. Cell associated α-amylases of butyrate-producing Firmicute bacteria from the human colon. Microbiology 152, 3281–3290 (2006).
Ryan, S. N., Fitzgerald, G. F. & van Sinderen, D. Screening for and identification of starch, amylopectin and pullulan-degrading activities in bifidobacterial strains. Appl. Environ. Microbiol. 72, 5289–5296 (2006).
Schell, M. A. et al. The genome sequence of Bifidobacterium longum reflects its adaptation to the human gastrointestinal tract. Proc. Natl Acad. Sci. USA 99, 14422–14427 (2003).
Walker, A. W., Duncan, S. H., McWilliam Leitch, E. C., Child, M. W. & Flint, H. J. pH and peptide supply can radically alter bacterial populations and short-chain fatty acid ratios within microbial communities from the human colon. Appl. Environ. Microbiol. 71, 3692–3700 (2005).
Van der Meulen, R., Makras, L., Verbrugghe, K., Adriany, T. & De Vuyst, L. In vitro kinetic analysis of oligofructose consumption by Bacteroides and Bifidobacterium spp. indicates different degradation mechanisms. Appl. Environ. Microbiol. 72, 1006–1012 (2006).
Demain, A. L., Newcomb, M. & Wu, J. H. D. Cellulase, clostridia and ethanol. Microbiol. Mol. Biol. Rev. 69, 124–154 (2005).
Miller, T. L., Currenti, E. & Wolin, M. J. Anaerobic bioconversion of cellulose by Ruminococcus albus, Methanobrevibacter smithii and Methanosarcina barkeri. Appl. Microbiol. Biotechnol. 54, 494–498 (2000).
Gokarn, R. R., Eiteman, M. A., Martin, S. A. & Eriksson, K. E. L. Production of succinate from glucose, cellobiose and various cellulosic materials by the rumen anaerobic bacteria Fibrobacter succinogenes and Ruminococcus flavefaciens. Appl. Biochem. Biotechnol. 68, 69–80 (1997).
Lissens, G. et al. Advanced anaerobic bioconversion of lignocellulosic waste for bioregenerative life support following thermal water treatment and biodegradation by Fibrobacter succinogenes. Biodegradation 15, 173–183 (2004).
Fierobe, H. P. et al. Action of designer cellulosomes on homogeneous versus complex substrates — controlled incorporation of three distinct enzymes into a defined trifunctional scaffoldin. J. Biol. Chem. 280, 16325–16334 (2005).
Shoseyev, O., Shani, Z. & Levy, I. Carbohydrate binding modules: biochemical properties and novel applications. Microbiol. Mol. Biol. Rev. 70, 283–295 (2006).
van Bielen, J. B. & Li, Z. Enzyme technology: an overview. Curr. Opin. Biotechnol. 13, 338–344 (2002).
Bhat, M. K. Cellulases and related enzymes in biotechnology. Biotechnol. Adv. 18, 355–383 (2000).
Gill, S. R. et al. Metagenomic analysis of the human distal gut microbiome. Science 312, 1355–1359 (2006).
Ferrer, M. et al. Novel hydrolase diversity retrieved from a metagenome library of bovine rumen microflora. Environ. Microbiol. 7, 1966–2010 (2005). Investigation of glycoside hydrolase diversity in the rumen community by functional screening of a metagenome library.
Hayashi, H. et al. Direct cloning of genes encoding novel xylanases from the human gut. Can. J. Microbiol. 51, 251–259 (2005).
Palackai, N. et al. A multifunctional hybrid glycosyl hydrolase discovered in an uncultured microbial consortium from ruminant gut. Appl. Microbiol. Biotechnol. 74, 113–124 (2007).
Beloqui, A. et al. Novel polyphenol oxidase mined from a metagenome expression library of bovine rumen. J. Biol. Chem. 281, 22933–22942 (2006).
Handelsman, J., Rondon, M. R., Brady, S. F., Clardy, J. & Goodman, R. M. Molecular biological access to the chemistry of unknown soil microbes: a new frontier for natural products. Chem. Biol. 5, R245–R249 (1988).
Margulies, M. et al. Genome sequencing in microfabricated high-density picolitre reactors. Nature 437, 376–380 (2005).
Brulc, J. M. et al. Comparative metagenomics of the rumen microbiome by random sampling pyrosequencing. Microb. Ecol. Health Dis. 19, 15 (2007).
Duncan, S. H. et al. Proposal of Roseburia faecis sp. nov., Roseburia hominis sp. nov. and Roseburia inulinivorans sp. nov., based on isolates from human faeces. Int. J. Syst. Evol. Microbiol. 56, 2437–2441 (2006).
Ayers, W. A. Phosphorolysis and synthesis of cellobiose by cell extracts of Ruminococcus flavefaciens. J. Biol. Chem. 234, 2819–2822 (1959).
Cerdeno-Tarraga, A. M. et al. Extensive DNA inversions in the B. fragilis genome control variable gene expression. Science 307, 1463–1465 (2005).
Acknowledgements
The Rowett Research Institute receives support from the Scottish Government Rural and Environment Research and Analysis Directorate. M.T.R. is grateful for support from the European Union FP5 project GEMINI.
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Glossary
- Lignocellulose
-
A complex mixture of structural polysaccharides, mainly cellulose and hemicellulose, that has varying amounts of polyphenolic lignins, and is the main constituent of plant cell-wall material.
- Inulin
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A storage polysaccharide that is found in some plants and consists mainly of β-(1,2)-linked D-fructose residues. Inulin, and the shorter fructo-oligosaccharides that are derived from it, is not hydrolysed by mammalian digestive enzymes, but can be used by bacteria in the large intestine.
- Resistant starch
-
The fraction of dietary starch that evades digestion in the upper gut of monogastric animals, notably humans, and thus becomes available for fermentation by the microbial community in the large intestine.
- Hemicellulose
-
A cross-linking glycan that constitutes up to 30% of plant cell walls; the two major hemicelluloses are xyloglucan and glucuronoarabinoxylan.
- Dockerin
-
A calcium-binding, modular component of cellulosomal enzymes (and other components) that is characterized by a duplicated sequence that binds to a complementary cohesin.
- Cohesin
-
A module in the scaffoldin to which cellulosomal dockerin-containing enzymes (and/or other components) are bound. The cohesin–dockerin pairs are grouped according to sequence similarity into phylogenetically different types.
- Cellulosome
-
An extracellular enzyme complex that consists of a scaffoldin (or scaffoldins) and cellulosomal enzymes that are dedicated to the efficient degradation of plant cell walls.
- Sortase
-
A transpeptidase that links peptide units on separate chains of peptidoglycan. Specifically, sortases link the threonine (T) residue of the LPXTG motif (where X denotes any amino acid) to the bacterial cell wall by a transpeptidation reaction.
- Scaffoldin
-
A structural cellulosomal subunit that comprises cohesin modules and binds dockerin-bearing enzymes (and/or other components).
- Neopullulanase
-
An amylase that has the ability to cleave the α-(1,4) linkages in amylose (a linear α-(1,4)-linked glucose polymer), amylopectin (amylose chains joined in a branching structure by α-(1,6) linkages) and pullulan (a linear polymer of α-(1,4)- and α-(1,6)- linked glucose residues).
- Extracytoplasmic function-type sigma factor
-
A class of sigma factor that stimulates the transcription of specific genes by RNA polymerase in response to environmental signals. Each extracytoplasmic function-type sigma factor is typically complexed with a specific membrane-bound anti-sigma factor, and becomes active only when released in response to the environmental stimulus.
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Flint, H., Bayer, E., Rincon, M. et al. Polysaccharide utilization by gut bacteria: potential for new insights from genomic analysis. Nat Rev Microbiol 6, 121–131 (2008). https://doi.org/10.1038/nrmicro1817
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DOI: https://doi.org/10.1038/nrmicro1817
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