Hostname: page-component-669899f699-vbsjw Total loading time: 0 Render date: 2025-05-06T20:00:02.128Z Has data issue: false hasContentIssue false
  • English
  • Français

The glmS ribozyme: use of a small molecule coenzyme by a gene-regulatory RNA

Published online by Cambridge University Press:  08 September 2010

Adrian R. Ferré-D'Amaré
Adrian R. Ferré-D'Amaré*
Affiliation:
Howard Hughes Medical Institute, Fred Hutchinson Cancer Research Center, 1100 Fairview Avenue North, Seattle, WA 98109-1024, USA
*
*Author for correspondence: A. R. Ferré-D'Amaré, Howard Hughes Medical Institute, Fred Hutchinson Cancer Research Center, 1100 Fairview Avenue North, Seattle, WA 98109-1024, USA. Tel.: 206-667-3622; Fax: 206-667-3331; Email: [email protected]
Get access Rights & Permissions [Opens in a new window]

Abstract

The glmS ribozyme is the first known example of a natural ribozyme that has evolved to require binding of an exogenous small molecule for activity. In Gram-positive bacteria, this RNA domain is part of the messenger RNA (mRNA) encoding the essential enzyme that synthesizes glucosamine-6-phosphate (GlcN6P). When present at physiologic concentration, this small molecule binds to the glmS ribozyme and uncovers a latent self-cleavage activity that ultimately leads to degradation of the mRNA. Biochemical and structural studies reveal that the RNA adopts a rigid fold stabilized by three pseudoknots and the packing of a peripheral domain against the ribozyme core. GlcN6P binding to this pre-organized RNA does not induce conformational changes; rather, the small molecule functions as a coenzyme, providing a catalytically essential amine group to the active site. The ribozyme is not a passive player, however. Active site functional groups are essential for catalysis, even in the presence of GlcN6P. In addition to being a superb experimental system with which to analyze how RNA catalysts can exploit small molecule coenzymes to broaden their chemical versatility, the presence of the glmS ribozyme in numerous pathogenic bacteria make this RNA an attractive target for the development of new antibiotics and antibacterial strategies.

Type
Review Article
Information
Quarterly Reviews of Biophysics , Volume 43 , Issue 4 , November 2010 , pp. 423 - 447
Copyright
Copyright © Cambridge University Press 2010

Access options

Get access to the full version of this content by using one of the access options below. (Log in options will check for institutional or personal access. Content may require purchase if you do not have access.)

Article purchase

Temporarily unavailable

References

8. References

Aalberts, D. P. & Hodas, N. O. (2005). Asymmetry in RNA pseudoknots: observation and theory. Nucleic Acids Research 33, 2210–2214.CrossRefGoogle ScholarPubMed
Baird, N. J. & Ferré-D'amaré, A. R. (2010). Idiosyncratically tuned switching behavior of riboswitch aptamer domains revealed by comparative small-angle X-ray scattering analysis. RNA 16, 598–609.Google Scholar
Baird, N. J., Kulshina, N. & Ferré-D'amaré, A. R. (2010). Riboswitch function: flipping the switch or tuning the dimmer? RNA Biology 7, 1–5.CrossRefGoogle ScholarPubMed
Barrick, J. E., Corbino, K. A., Winkler, W. C., Nahvi, A., Mandal, M., Collins, J., Lee, M., Roth, A., Sudarsan, N., Jona, I., Wickiser, J. K. & Breaker, R. R. (2004). New RNA motifs suggest an expanded scope for riboswitches in bacterial genetic control. Proceedings of the National Academy of Sciences USA 101, 6421–6426.Google Scholar
Batey, R. T., Gilbert, S. D. & Montange, R. K. (2004). Structure of a natural guanine-responsive riboswitch complexed with the metabolite hypoxanthine. Nature 432, 411–415.CrossRefGoogle ScholarPubMed
Bevilacqua, P. C. (2003). Mechanistic considerations for general acid–base catalysis by RNA: revisiting the mechanism of the hairpin ribozyme. Biochemistry 42, 2259–2265.Google Scholar
Bevilacqua, P. C. & Yajima, R. (2006). Nucleobase catalysis in ribozyme mechanism. Current Opinion in Chemical Biology 10, 455–464.Google Scholar
Blount, K., Puskarz, I., Penchovsky, R. & Breaker, R. (2006). Development and application of a high-throughput assay for glmS riboswitch activators. RNA Biology 3, 77–81.CrossRefGoogle ScholarPubMed
Brooks, K. M. & Hampel, K. J. (2009). A rate-limiting conformational step in the catalytic pathway of the glmS ribozyme. Biochemistry 48, 5669–5678.CrossRefGoogle ScholarPubMed
Burgers, P. M. J. & Eckstein, F. (1979). Diasteromers of 5′-O-adenosyl 3′-O-uridyl phosphorothioate: chemical synthesis and enzymatic properties. Biochemistry 18, 592–596.CrossRefGoogle Scholar
Carson, M. (1997). Ribbons. Methods in Enzymology 277, 493–505.Google Scholar
Cayley, S., Lewis, B. A., Guttman, H. J. & Record, M. T. (1991). Characterization of the cytoplasm of Escherichia coli K-12 as a function of external osmolarity. Implications for protein–DNA interactions in vivo. Journal of Molecular Biology 222, 281–300.CrossRefGoogle ScholarPubMed
Chi, Y.-I., Martick, M., Lares, M., Kim, R., Scott, W. G., Kim, S.-H. & Joyce, G. F. (2008). Capturing hammerhead ribozyme structures in action by modulating general base catalysis. PLoS Biology 6, e234.CrossRefGoogle ScholarPubMed
Cochrane, J. C., Lipchock, S. V., Smith, K. D. & Strobel, S. A. (2009). Structural and chemical basis for glucosamine 6-phosphate binding and activation of the glmS ribozyme. Biochemistry 48, 3239–3246.Google Scholar
Cochrane, J. C., Lipchock, S. V. & Strobel, S. A. (2007). Structural investigation of the GlmS ribozyme bound to its catalytic cofactor. Chemistry and Biology 14, 97–105.Google Scholar
Collins, J. A., Irnov, I., Baker, S. & Winkler, W. C. (2007). Mechanism of mRNA destabilization by the glmS ribozyme. Genes and Development 21, 3356–3368.CrossRefGoogle ScholarPubMed
Cowan, J. A. (1993). Metallobiochemistry of RNA. Co(NH3)63+ as a probe for Mg2+ (aq) binding sites. Journal of Inorganic Biochemistry 49, 171–175.CrossRefGoogle Scholar
Dambach, M. D. & Winkler, W. C. (2009). Expanding roles for metabolite-sensing regulatory RNA. Current Opinion in Microbiology 12, 161–169.CrossRefGoogle Scholar
Damblon, C., Raquet, X., Lian, L. Y., Lamotte-Brasseur, J., Fonze, E., Charlier, P., Roberts, G. C. & Frère, J. M. (1996). The catalytic mechanism of beta-lactamases: NMR titration of an active-site lysine residue of the TEM-1 enzyme. Proceedings of the National Academy of Sciences USA 93, 1747–1752.Google Scholar
Das, S. R. & Piccirilli, J. A. (2005). General acid catalysis by the hepatitis delta virus ribozyme. Nature Chemical Biology 1, 45–52.CrossRefGoogle ScholarPubMed
De La Peña, M., Dufour, D. & Gallego, J. (2009). Three-way RNA junctions with remote tertiary contacts: a recurrent and highly versatile fold. RNA 15, 1949–1964.CrossRefGoogle ScholarPubMed
Dittrich, M., Hayashi, S. & Schulten, K. (2008). ATP Hydrolysis in the βTP and βDP Catalytic Sites of F1-ATPase. Biophysical Journal 87, 2954–2967.Google Scholar
Draper, D. E. (2004). A guide to ions and RNA structure. RNA 10, 335–343.Google Scholar
Edwards, T. E. & Ferré-D'amaré, A. R. (2006). Crystal structures of the thi-box riboswitch bound to thiamine pyrophosphate analogs reveal adaptive RNA-small molecule recognition. Structure 14, 1459–1468.CrossRefGoogle ScholarPubMed
Edwards, T. E., Klein, D. J. & Ferré-D'amaré, A. R. (2007). Riboswitches: small-molecule recognition by gene regulatory RNA. Current Opinion in Structural Biology 17, 273–279.CrossRefGoogle Scholar
Fedor, M. J. (2009). Comparative enzymology and structural biology of RNA self-cleavage. Annual Reviews in Biophysics 38, 271–299.Google Scholar
Ferré-D'amaré, A. R. (2003). RNA-modifying enzymes. Current Opinion in Structural Biology 13, 49–55.CrossRefGoogle ScholarPubMed
Ferré-D'amaré, A. R. (2004). The hairpin ribozyme. Biopolymers 73, 71–78.CrossRefGoogle ScholarPubMed
Ferré-D'amaré, A. R. & Scott, W. G. (2010). Small self-cleaving ribozymes. Cold Spring Harbor Perspectives in Biology 2, 1–10.CrossRefGoogle ScholarPubMed
Ferré-D'amaré, A. R., Zhou, K. & Doudna, J. A. (1998). Crystal structure of a hepatitis delta virus ribozyme. Nature 395, 567–574.Google Scholar
Gilbert, S. D., Rambo, R. P., Van Tyne, D. & Batey, R. T. (2008). Structure of the SAM-II riboswitch bound to S-adenosylmethionine. Nature Structural and Molecular Biology 15, 177–182.CrossRefGoogle ScholarPubMed
Görke, B. & Vogel, J. (2008). Noncoding RNA control of the making and breaking of sugars. Genes and Development 22, 2914–2929.CrossRefGoogle ScholarPubMed
Grundy, F. J. & Henkin, T. M. (1993). tRNA as a positive regulator of transcription antitermination in B. subtilis. Cell 74, 475–482.Google Scholar
Hampel, K. J. & Tinsley, M. M. (2006). Evidence for preorganization of the glmS ribozyme ligand binding pocket. Biochemistry 45, 7861–7871.CrossRefGoogle ScholarPubMed
Henkin, T. (2008). Riboswitch RNA: using RNA to sense cellular metabolism. Genes and Development 22, 3383–3390.Google Scholar
Jacob, F. & Monod, J. (1961). Genetic regulatory mechanisms in the synthesis of proteins. Journal of Molecular Biology 3, 318–356.CrossRefGoogle ScholarPubMed
Jaffe, E. K. & Cohn, M. (1979). Diasteromers of the nucleoside phosphorothioates as probes of the structure of the metal nucleotide substrates and of the nucleotide binding site of yeast hexokinase. Journal of Biological Chemistry 254, 10839–10845.Google Scholar
Klein, D., Edwards, T. & Ferré-D'amaré, A. (2009). Cocrystal structure of a class I preQ1 riboswitch reveals a pseudoknot recognizing an essential hypermodified nucleobase. Nature Structural and Molecular Biology 16, 343–344.CrossRefGoogle Scholar
Klein, D. J., Been, M. D. & Ferré-D'amaré, A. R. (2007a). Essential role of an active-site guanine in glmS ribozyme catalysis. Journal of the American Chemical Society 129, 14858–14859.CrossRefGoogle ScholarPubMed
Klein, D. J. & Ferré-D'amaré, A. R. (2006). Structural basis of glmS ribozyme activation by glucosamine-6-phosphate. Science 313, 1752–1756.Google Scholar
Klein, D. J. & Ferré-D'amaré, A. R. (2009). Crystallization of the glmS ribozyme-riboswitch. Methods in Molecular Biology 540, 129–139.Google Scholar
Klein, D. J., Wilkinson, S. R., Been, M. D. & Ferré-D'amaré, A. R. (2007b). Requirement of helix P2.2 and nucleotide G1 for positioning of the cleavage site and cofactor of the glmS ribozyme. Journal of Molecular Biology 373, 178–189.CrossRefGoogle ScholarPubMed
Kobayashi, K., Ehrlich, S. D., Albertini, A., Amati, G., Andersen, K. K., Arnaud, M., Asai, K., Ashikaga, S., Aymerich, S., Bessieres, P., Boland, F., Brignell, S. C., Bron, S., Bunai, K., Chapuis, J., Christiansen, L. C., Danchin, A., Debarbouille, M., Dervyn, E., Deuerling, E., Devine, K., Devine, S. K., Dreesen, O., Errington, J., Fillinger, S., Foster, S. J., Fujita, Y., Galizzi, A., Gardan, R., Eschevins, C., Fukushima, T., Haga, K., Harwood, C. R., Hecker, M., Hosoya, D., Hullo, M. F., Kakeshita, H., Karamata, D., Kasahara, Y., Kawamura, F., Koga, K., Koski, P., Kuwana, R., Imamura, D., Ishimaru, M., Ishikawa, S., Ishio, I., Le Coq, D., Masson, A., Mauel, C., Meima, R., Mellado, R. P., Moir, A., Moriya, S., Nagakawa, E., Nanamiya, H., Nakai, S., Nygaard, P., Ogura, M., Ohanan, T., O'Reilly, M., O'Rourke, M., Pragai, Z., Pooley, H. M., Rapoport, G., Rawlins, J. P., Rivas, L. A., Rivolta, C., Sadaie, A., Sadaie, Y., Sarvas, M., Sato, T., Saxild, H. H., Scanlan, E., Schumann, W., Seegers, J. F., Sekiguchi, J., Sekowska, A., Seror, S. J., Simon, M., Stragier, P., Studer, R., Takamatsu, H., Tanaka, T., Takeuchi, M., Thomaides, H. B., Vagner, V., Van Dijl, J. M., Watabe, K., Wipat, A., Yamamoto, H., Yamamoto, M., Yamamoto, Y., Yamane, K., Yata, K., Yoshida, K., Yoshikawa, H., Zuber, U. & Ogasawara, N. (2003). Essential Bacillus subtilis genes. Proceedings of the National Academy of Sciences USA 100, 4678–4683.CrossRefGoogle ScholarPubMed
Koonin, E. V. (2009). Intron-Dominated genomes of early ancestors of eukaryotes. Journal of Heredity 100, 618–623.Google Scholar
Kulshina, N., Baird, N. J. & Ferré-D'amaré, A. R. (2009). Recognition of the bacterial second messenger cyclic diguanylate by its cognate riboswitch. Nature Structural and Molecular Biology 16, 1212–1217.CrossRefGoogle ScholarPubMed
Kulshina, N., Edwards, T. E. & Ferre-D'amare, A. R. (2010). Thermodynamic analysis of ligand binding and ligand binding-induced tertiary structure formation by the thiamine pyrophosphate riboswitch. RNA 16, 186–196.Google Scholar
Kuzmin, Y. I., Da Costa, C. P., Cottrell, J. W. & Fedor, M. J. (2005). Role of an active site adenine in hairpin ribozyme catalysis. Journal of Molecular Biology 349, 989–1010.Google Scholar
Lebruska, L. L., Kuzmine, I. I. & Fedor, M. J. (2002). Rescue of an abasic hairpin ribozyme by cationic nucleobases. Evidence for a novel mechanism of RNA catalysis. Chemistry and Biology 9, 465–473.CrossRefGoogle ScholarPubMed
Lilley, D. M. (2004). The Varkud satellite ribozyme. RNA 10, 151–158.CrossRefGoogle ScholarPubMed
Lilley, D. M. J. (1999). Folding of branched RNA species. Biopolymers 48, 101–112.Google Scholar
Lilley, D. M. J. (2003). The origins of RNA catalysis in ribozymes. Trends in Biochemical Science 28, 495–501.CrossRefGoogle ScholarPubMed
Lilley, D. M. J. & Eckstein, F. (2008). Ribozymes and RNA catalysis: introduction and primer. In Ribozymes and RNA Catalysis (eds. Lilley, D. M. J. & Eckstein, F.), pp. 1–10. Cambridge: The Royal Society of Chemistry.Google Scholar
Lim, J., Grove, B. C., Roth, A. & Breaker, R. R. (2006). Characteristics of ligand recognition by a glmS self-cleaving ribozyme. Angewandte Chemie International Edition in English 45, 6689–6693.Google Scholar
Lipfert, J., Ouellet, J., Norman, D., Doniach, S. & Lilley, D. (2008). The complete VS ribozyme in solution studied by small-sngle X-ray scattering. Structure 16, 1357–1367.CrossRefGoogle Scholar
Martick, M., Horan, L. H., Noller, H. F. & Scott, W. G. (2008). A discontinuous hammerhead ribozyme embedded in a mammalian messenger RNA. Nature 454, 899–902.Google Scholar
Martick, M. & Scott, W. G. (2006). Tertiary contacts distant from the active site prime a ribozyme for catalysis. Cell 126, 309–320.Google Scholar
Mayer, G. & Famulok, M. (2006). High-throughput-compatible assay for glmS riboswitch metabolite dependence. ChemBioChem 7, 602–604.Google Scholar
Mccarthy, T. J., Plog, M. A., Floy, S. A., Jansen, J. A., Soukup, J. K. & Soukup, G. A. (2005). Ligand requirements for glmS ribozyme self-cleavage. Chemistry and Biology 12, 1221–1226.Google Scholar
Milewski, S. (2002). Glucosamine-6-phosphate synthase–the multi-facets enzyme. Biochimica et Biophysica Acta 1597, 173–192.Google Scholar
Mironov, A. S., Gusarov, I., Rafikov, R., Lopez, L. E., Shatalin, K., Kreneva, R. A., Perumov, D. A. & Nudler, E. (2002). Sensing small molecules by nascent RNA: a mechanism to control transcription in bacteria. Cell 111, 747–756.Google Scholar
Murchie, A. I. H., Thomson, J. B., Walter, F. & Lilley, D. M. J. (1998). Folding of the hairpin ribozyme in its natural conformation achieves close physical proximity of the loops. Molecular Cell 1, 873–881.CrossRefGoogle ScholarPubMed
Murray, J. B., Seyhan, A. A., Walter, N. G., Burke, J. M. & Scott, W. G. (1998). The hammerhead, hairpin and VS ribozymes are catalytically proficient in monovalent cations alone. Chemistry and Biology 5, 587–595.CrossRefGoogle ScholarPubMed
Nakano, S.-I., Chadalavada, D. M. & Bevilacqua, P. C. (2000). General acid–base catalysis in the mechanism of a hepatitis delta virus ribozyme. Science 287, 1493–1497.Google Scholar
Nissen, P., Ippolito, J. A., Ban, N., Moore, P. B. & Steitz, T. A. (2001). RNA tertiary interactions in the large ribosomal subunit: the A-minor motif. Proceedings of the National Academy of Sciences USA 98, 4899–4903.CrossRefGoogle ScholarPubMed
Perrotta, A. T., Shih, I. & Been, M. D. (1999). Imidazole rescue of a cytosine mutation in a self-cleaving ribozyme. Science 286, 123–126.Google Scholar
Pley, H. W., Flaherty, K. M. & Mckay, D. B. (1994). Three-dimensional structure of a hammerhead ribozyme. Nature 372, 68–74.CrossRefGoogle ScholarPubMed
Pyle, A. M. (1993). Ribozymes: a distinct class of metalloenzymes. Science 261, 709–714.CrossRefGoogle ScholarPubMed
Raghavan, R. & Minnick, M. F. (2009). Group I introns and inteins: disparate origins but convergent parasitic strategies. Journal of Bacteriology 191, 6193–6202.Google Scholar
Raines, R. T. (1998). Ribonuclease A. Chemical Reviews 98, 1045–1065.CrossRefGoogle ScholarPubMed
Randau, L., Schröder, I. & Söll, D. (2008). Life without RNAe P. Nature 453, 120–123.CrossRefGoogle Scholar
Reichenbach, B., Maes, A., Kalamorz, F., Hajnsdorf, E. & Görke, B. (2008). The small RNA GlmY acts upstream of the sRNA GlmZ in the activation of glmS expression and is subject to regulation by polyadenylation in Escherichia coli. Nucleic Acids Research 36, 2570–2580.Google Scholar
Richards, F. M., Wyckoff, H. W., Carlson, W. D., Allewell, N. M., Lee, B. & Mitsui, Y. (1971). Protein structure, ribonuclease-S, and nucleotide interactions. Cold Spring Harbor Symposium on Quantitative Biology 36, 35–43.Google Scholar
Romani, A. & Scarpa, A. (1992). Regulation of cell magnesium. Archives of Biochemistry and Biophysics 298, 1–12.CrossRefGoogle ScholarPubMed
Roth, A., Nahvi, A., Lee, M., Jona, I. & Breaker, R. R. (2006). Characteristics of the glmS ribozyme suggest only structural roles for divalent metal ions. RNA 12, 607–619.Google Scholar
Rupert, P. B. & Ferré-D'amaré, A. R. (2001). Crystal structure of a hairpin ribozyme-inhibitor complex with implications for catalysis. Nature 410, 780–786.CrossRefGoogle ScholarPubMed
Rupert, P. B., Massey, A. P., Sigurdsson, S. T. & Ferré-D'amaré, A. R. (2002). Transition state stabilization by a catalytic RNA. Science 298, 1421–1424.CrossRefGoogle ScholarPubMed
Salehi-Ashtiani, K. & Szostak, J. W. (2001). In vitro evolution suggests multiple origins for the hammerhead ribozyme. Nature 414, 82–84.Google Scholar
Scott, W. G., Finch, J. T. & Klug, A. (1995). The crystal structure of an all-RNA hammerhead ribozyme: a proposed mechanism for RNA catalytic cleavage. Cell 81, 991–1002.Google Scholar
Serganov, A. (2009). The long and the short of riboswitches. Current Opinion in Structural Biology 19, 251–259.CrossRefGoogle Scholar
Serganov, A., Huang, L. & Patel, D. (2009). Coenzyme recognition and gene regulation by a flavin mononucleotide riboswitch. Nature 457, 233–237.Google Scholar
Serganov, A., Polonskaia, A., Phan, A. T., Breaker, R. R. & Patel, D. J. (2006). Structural basis for gene regulation by a thiamine pyrophosphate-sensing riboswitch. Nature 441, 1167–1171.Google Scholar
Soukup, G. A. & Breaker, R. R. (1999). Relationship between internucleotide linkage geometry and the stability of RNA. RNA 5, 1308–1325.Google Scholar
Tinsley, R. A., Furchak, J. R. & Walter, N. G. (2007). Trans-acting glmS catalytic riboswitch: locked and loaded. RNA 13, 468–477.Google Scholar
Torelli, A. T., Krucinska, J., & Wedekind, J. E. (2007). A comparison of vanadate to a 2′–5′ linkage at the active site of a small ribozyme suggests a role for water in transition-state stabilization. RNA 13, 1052–1070.Google Scholar
Wang, L., Yu, X., Hu, P., Broyde, S. & Zhang, Y. (2007). A water-mediated and substrate-assisted catalytic mechanism for Sulfolobus solfataricus DNA polymerase IV. Journal of the American Chemical Society 129, 4731–4737.CrossRefGoogle ScholarPubMed
Wang, Y. & Schlick, T. (2008). Quantum mechanics/molecular mechanics investigation of the chemical reaction in Dpo4 reveals water-dependent pathways and requirements for active site reorganization. Journal of the American Chemical Society 130, 13240–13250.Google Scholar
Webb, C.-H. T., Riccitelli, N. J., Ruminski, D. J. & Lupták, A. (2009). Widespread occurrence of self-cleaving ribozymes. Science 326, 953–953.Google Scholar
White, H. B. (1976). Coenzymes as fossils of an earlier metabolic state. Journal of Molecular Evolution 7, 101–104.Google Scholar
Wilkinson, S. R. & Been, M. D. (2005). A pseudoknot in the 3′ non-core region of the glmS ribozyme enhances self-cleavage activity. RNA 11, 1788–1794.CrossRefGoogle ScholarPubMed
Wilson, T. J. & Lilley, D. M. J. (2009). Biochemistry. The evolution of ribozyme chemistry. Science 323, 1436–1438.Google Scholar
Winkler, W., Nahvi, A. & Breaker, R. R. (2002). Thiamine derivatives bind messenger RNA directly to regulate bacterial gene expression. Nature 419, 952–956.CrossRefGoogle ScholarPubMed
Winkler, W. C., Nahvi, A., Roth, A., Collins, J. A. & Breaker, R. R. (2004). Control of gene expression by a natural metabolite-responsive ribozyme. Nature 428, 281–286.Google Scholar
Xiao, H., Edwards, T. E. & Ferré-D'amaré, A. R. (2008a). Structural basis for specific, high-affinity tetracycline binding by an in vitro evolved aptamer and artificial riboswitch. Chemistry and Biology 15, 1125–1137.Google Scholar
Xiao, H., Murakami, H., Suga, H. & Ferré-D'amaré, A. R. (2008b). Structural basis of specific tRNA aminoacylation by a small in vitro selected ribozyme. Nature 454, 358–361.Google Scholar
Xue, Y., Xu, Y., Liu, Y., Ma, Y. & Zhou, P. (2001). Thermoanaerobacter tengcongensis sp. nov., a novel anaerobic, saccharolytic, thermophilic bacterium isolated from a hot spring in Tengcong, China. International Journal of Systematic and Evolutionary Microbiology 51, 1335–1341.Google Scholar