Elucidation of a sialic acid metabolism pathway in mucus-foraging Ruminococcus gnavus unravels mechanisms of bacterial adaptation to the gut

Abstract

Sialic acid (N-acetylneuraminic acid (Neu5Ac)) is commonly found in the terminal location of colonic mucin glycans where it is a much-coveted nutrient for gut bacteria, including Ruminococcus gnavus. R. gnavus is part of the healthy gut microbiota in humans, but it is disproportionately represented in diseases. There is therefore a need to understand the molecular mechanisms that underpin the adaptation of R. gnavus to the gut. Previous in vitro research has demonstrated that the mucin-glycan-foraging strategy of R. gnavus is strain dependent and is associated with the expression of an intramolecular trans-sialidase, which releases 2,7-anhydro-Neu5Ac, rather than Neu5Ac, from mucins. Here, we unravelled the metabolism pathway of 2,7-anhydro-Neu5Ac in R. gnavus that is underpinned by the exquisite specificity of the sialic transporter for 2,7-anhydro-Neu5Ac and by the action of an oxidoreductase that converts 2,7-anhydro-Neu5Ac into Neu5Ac, which then becomes a substrate of a Neu5Ac-specific aldolase. Having generated an R. gnavus nan-cluster deletion mutant that lost the ability to grow on sialylated substrates, we showed that—in gnotobiotic mice colonized with R. gnavus wild-type (WT) and mutant strains—the fitness of the nan mutant was significantly impaired, with a reduced ability to colonize the mucus layer. Overall, we revealed a unique sialic acid pathway in bacteria that has important implications for the spatial adaptation of mucin-foraging gut symbionts in health and disease.

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Fig. 1: The nan operon of R. gnavus ATCC 29149.
Fig. 2: Steady-state fluorescence analysis of ligand binding to RgSBP.
Fig. 3: Biophysical analysis of ligand binding to RgSBP.
Fig. 4: Enzymic reaction of R. gnavus sialic acid aldolase.
Fig. 5: RgNanOx catalyses the conversion of 2,7-anhydro-Neu5Ac to Neu5Ac.
Fig. 6: Colonization of germ-free C57BL/6J mice with R. gnavus ATCC 29149 WT or nan mutant strains.

Data availability

Genome and protein sequences are available from NCBI and referenced within the text or the Supplementary Information. Accession numbers for all of the genomes used for multigene alignments are provided in Supplementary Table 5. Raw FASTQ files for the RNA-seq libraries were deposited to the NCBI Sequence Read Archive (SRA) under BioProject accession number PRJNA559470. The crystal structures described in this paper have been deposited in the protein data bank (PDB) under the following identifiers: 6RAB (WT), 6RB7 (K167A) and 6RD1 (RgNanA K167A–Neu5Ac complex). All other data are available from the corresponding author on reasonable request.

Code availability

All code used for statistical analysis is available from the corresponding author on reasonable request.

References

  1. 1.

    Donaldson, G. P., Lee, S. M. & Mazmanian, S. K. Gut biogeography of the bacterial microbiota. Nat. Rev. Microbiol. 14, 20–32 (2016).

  2. 2.

    Johansson, M. E. V., Larsson, J. M. H. & Hansson, G. C. The two mucus layers of colon are organized by the MUC2 mucin, whereas the outer layer is a legislator of host-microbial interactions. Proc. Natl Acad. Sci. USA 108, 4659–4665 (2011).

  3. 3.

    Etienne-Mesmin, L. et al. Experimental models to study intestinal microbes-mucus interactions in health and disease. FEMS Microbiol. Rev. 43, 457–489 (2019).

  4. 4.

    Jensen, P. H., Kolarich, D. & Packer, N. H. Mucin-type O-glycosylation—putting the pieces together. FEBS J. 277, 81–94 (2010).

  5. 5.

    Ndeh, D. & Gilbert, H. J. Biochemistry of complex glycan depolymerisation by the human gut microbiota. FEMS Microbiol. Rev. 42, 146–164 (2018).

  6. 6.

    Tailford, L. E., Crost, E. H., Kavanaugh, D. & Juge, N. Mucin glycan foraging in the human gut microbiome. Front. Genet. 6, 81 (2015).

  7. 7.

    Martens, E. C., Chiang, H. C. & Gordon, J. I. Mucosal glycan foraging enhances fitness and transmission of a saccharolytic human gut bacterial symbiont. Cell Host Microbe 4, 447–457 (2008).

  8. 8.

    Robbe, C., Capon, C., Coddeville, B. & Michalski, J. C. Structural diversity and specific distribution of O-glycans in normal human mucins along the intestinal tract. Biochem. J. 384, 307–316 (2004).

  9. 9.

    Robbe, C. et al. Evidence of regio-specific glycosylation in human intestinal mucins—presence of an acidic gradient along the intestinal tract. J. Biol. Chem. 278, 46337–46348 (2003).

  10. 10.

    Juge, N., Tailford, L. & Owen, C. D. Sialidases from gut bacteria: a mini-review. Biochem. Soc. Trans. 44, 166–175 (2016).

  11. 11.

    Lewis, A. L. & Lewis, W. G. Host sialoglycans and bacterial sialidases: a mucosal perspective. Cell. Microbiol. 14, 1174–1182 (2012).

  12. 12.

    Almagro-Moreno, S. & Boyd, E. F. Insights into the evolution of sialic acid catabolism among bacteria. BMC Evol. Biol. 9, 118 (2009).

  13. 13.

    Plumbridge, J. & Vimr, E. Convergent pathways for utilization of the amino sugars N-acetylglucosamine, N-acetylmannosamine, and N-acetylneuraminic acid by Escherichia coli. J. Bacteriol. 181, 47–54 (1999).

  14. 14.

    Martinez, J., Steenbergen, S. & Vimr, E. Derived structure of the putative sialic acid transporter from Escherichia coli predicts a novel sugar permease domain. J. Bacteriol. 177, 6005–6010 (1995).

  15. 15.

    Brigham, C. et al. Sialic acid (N-acetyl neuraminic acid) utilization by Bacteroides fragilis requires a novel N-acetyl mannosamine epimerase. J. Bacteriol. 191, 3629–3638 (2009).

  16. 16.

    Vimr, E. R., Kalivoda, K. A., Deszo, E. L. & Steenbergen, S. M. Diversity of microbial sialic acid metabolism. Microbiol. Mol. Biol. Rev. 68, 132–153 (2004).

  17. 17.

    Thomas, G. H. Sialic acid acquisition in bacteria—one substrate, many transporters. Biochem. Soc. Trans. 44, 760–765 (2016).

  18. 18.

    Mulligan, C. et al. The substrate-binding protein imposes directionality on an electrochemical sodium gradient-driven TRAP transporter. Proc. Natl Acad. Sci. USA 106, 1778–1783 (2009).

  19. 19.

    Wahlgren, W. Y. et al. Substrate-bound outward-open structure of a Na+-coupled sialic acid symporter reveals a new Na+ site. Nat. Commun. 9, 1753 (2018).

  20. 20.

    Severi, E., Hosie, A. H., Hawkhead, J. A. & Thomas, G. H. Characterization of a novel sialic acid transporter of the sodium solute symporter (SSS) family and in vivo comparison with known bacterial sialic acid transporters. FEMS Microbiol. Lett. 304, 47–54 (2010).

  21. 21.

    Gangi Setty, T., Cho, C., Govindappa, S., Apicella, M. A. & Ramaswamy, S. Bacterial periplasmic sialic acid-binding proteins exhibit a conserved binding site. Acta Crystallogr. D 70, 1801–1811 (2014).

  22. 22.

    Mulligan, C., Leech, A. P., Kelly, D. J. & Thomas, G. H. The membrane proteins SiaQ and SiaM form an essential stoichiometric complex in the sialic acid tripartite ATP-independent periplasmic (TRAP) transporter SiaPQM (VC1777-1779) from Vibrio cholerae. J. Biol. Chem. 287, 3598–3608 (2012).

  23. 23.

    Muller, A. et al. Conservation of structure and mechanism in primary and secondary transporters exemplified by SiaP, a sialic acid binding virulence factor from Haemophilus influenzae. J. Biol. Chem. 281, 22212–22222 (2006).

  24. 24.

    Severi, E. et al. Sialic acid transport in Haemophilus influenzae is essential for lipopolysaccharide sialylation and serum resistance and is dependent on a novel tripartite ATP-independent periplasmic transporter. Mol. Microbiol. 58, 1173–1185 (2005).

  25. 25.

    Post, D. M., Mungur, R., Gibson, B. W. & Munson, R. S. Jr. Identification of a novel sialic acid transporter in Haemophilus ducreyi. Infect. Immun. 73, 6727–6735 (2005).

  26. 26.

    North, R. A. et al. The sodium sialic acid symporter from Staphylococcus aureus has altered substrate specificity. Front. Chem. 6, 233 (2018).

  27. 27.

    Hopkins, A. P., Hawkhead, J. A. & Thomas, G. H. Transport and catabolism of the sialic acids N-glycolylneuraminic acid and 3-keto-3-deoxy-d-glycero-d-galactonononic acid by Escherichia coli K-12. FEMS Microbiol. Lett. 347, 14–22 (2013).

  28. 28.

    Sagheddu, V., Patrone, V., Miragoli, F., Puglisi, E. & Morelli, L. Infant early gut colonization by Lachnospiraceae: high frequency of Ruminococcus gnavus. Front. Pediatr. 4, 57 (2016).

  29. 29.

    Qin, J. et al. A human gut microbial gene catalogue established by metagenomic sequencing. Nature 464, 59–65 (2010).

  30. 30.

    Ludwig, W., Schleifer, K.-H. & Whitman, W. B. in Bergey’s Manual of Systematic Bacteriology Vol. 3 (eds De Vos, P. et al.) 1–13 (Springer, 2009).

  31. 31.

    Kraal, L., Abubucker, S., Kota, K., Fischbach, M. A. & Mitreva, M. The prevalence of species and strains in the human microbiome: a resource for experimental efforts. PLoS ONE 9, e97279 (2014).

  32. 32.

    Olbjorn, C. et al. Fecal microbiota profiles in treatment-naive pediatric inflammatory bowel disease—associations with disease phenotype, treatment, and outcome. Clin. Exp. Gastroenterol. 12, 37–49 (2019).

  33. 33.

    Sokol, H. et al. Specificities of the intestinal microbiota in patients with inflammatory bowel disease and Clostridium difficile infection. Gut Microbes 9, 55–60 (2018).

  34. 34.

    Nishino, K. et al. Analysis of endoscopic brush samples identified mucosa-associated dysbiosis in inflammatory bowel disease. J. Gastroenterol. 53, 95–106 (2018).

  35. 35.

    Machiels, K. et al. Specific members of the predominant gut microbiota predict pouchitis following colectomy and IPAA in UC. Gut 66, 79–88 (2017).

  36. 36.

    Hall, A. B. et al. A novel Ruminococcus gnavus clade enriched in inflammatory bowel disease patients. Genome Med. 9, 103 (2017).

  37. 37.

    Fuentes, S. et al. Microbial shifts and signatures of long-term remission in ulcerative colitis after faecal microbiota transplantation. ISME J. 11, 1877–1889 (2017).

  38. 38.

    Joossens, M. et al. Dysbiosis of the faecal microbiota in patients with Crohn’s disease and their unaffected relatives. Gut 60, 631–637 (2011).

  39. 39.

    Willing, B. P. et al. A pyrosequencing study in twins shows that gastrointestinal microbial profiles vary with inflammatory bowel disease phenotypes. Gastroenterology 139, 1844–1854 (2010).

  40. 40.

    Png, C. W. et al. Mucolytic bacteria with increased prevalence in IBD mucosa augment in vitro utilization of mucin by other bacteria. Am. J. Gastroenterol. 105, 2420–2428 (2010).

  41. 41.

    Owen, C. D. et al. Unravelling the specificity and mechanism of sialic acid recognition by the gut symbiont Ruminococcus gnavus. Nat. Commun. 8, 2196 (2017).

  42. 42.

    Monaco, S., Tailford, L. E., Juge, N. & Angulo, J. Differential epitope mapping by STD NMR spectroscopy to reveal the nature of protein-ligand contacts. Angew. Chem. Int. Edn 56, 15289–15293 (2017).

  43. 43.

    Crost, E. H. et al. The mucin-degradation strategy of Ruminococcus gnavus: the importance of intramolecular trans-sialidases. Gut Microbes 7, 302–312 (2016).

  44. 44.

    Tailford, L. E. et al. Discovery of intramolecular trans-sialidases in human gut microbiota suggests novel mechanisms of mucosal adaptation. Nat. Commun. 6, 7624 (2015).

  45. 45.

    Crost, E. H. et al. Utilisation of mucin glycans by the human gut symbiont Ruminococcus gnavus is strain-dependent. PLoS ONE 8, e76341 (2013).

  46. 46.

    Monestier, M. et al. Membrane-enclosed multienzyme (MEME) synthesis of 2,7-anhydro-sialic acid derivatives. Carbohydr. Res. 451, 110–117 (2017).

  47. 47.

    Xu, G. et al. Three Streptococcus pneumoniae sialidases: three different products. J. Am. Chem. Soc. 133, 1718–1721 (2011).

  48. 48.

    Xu, G. et al. Crystal structure of the NanB sialidase from Streptococcus pneumoniae. J. Mol. Biol. 384, 436–449 (2008).

  49. 49.

    Kumar, J. P., Rao, H., Nayak, V. & Ramaswamy, S. Crystal structures and kinetics of N-acetylneuraminate lyase from Fusobacterium nucleatum. Acta Crystallogr. F 74, 725–732 (2018).

  50. 50.

    Campeotto, I. et al. Pathological macromolecular crystallographic data affected by twinning, partial-disorder and exhibiting multiple lattices for testing of data processing and refinement tools. Sci. Rep. 8, 14876 (2018).

  51. 51.

    North, R. A. et al. Structure and inhibition of N-acetylneuraminate lyase from methicillin-resistant Staphylococcus aureus. FEBS Lett. 590, 4414–4428 (2016).

  52. 52.

    Timms, N. et al. Structural insights into the recovery of aldolase activity in N-acetylneuraminic acid lyase by replacement of the catalytically active lysine with γ-thialysine by using a chemical mutagenesis strategy. ChemBioChem 14, 474–481 (2013).

  53. 53.

    Huynh, N. et al. Structural basis for substrate specificity and mechanism of N-acetyl-d-neuraminic acid lyase from Pasteurella multocida. Biochemistry 52, 8570–8579 (2013).

  54. 54.

    Barbosa, J. A. et al. Active site modulation in the N-acetylneuraminate lyase sub-family as revealed by the structure of the inhibitor-complexed Haemophilus influenzae enzyme. J. Mol. Biol. 303, 405–421 (2000).

  55. 55.

    Daniels, A. D. et al. Reaction mechanism of N-acetylneuraminic acid lyase revealed by a combination of crystallography, QM/MM simulation, and mutagenesis. ACS Chem. Biol. 9, 1025–1032 (2014).

  56. 56.

    Heap, J. T. et al. The ClosTron: mutagenesis in Clostridium refined and streamlined. J. Microbiol. Methods 80, 49–55 (2010).

  57. 57.

    Mandal, C., Schwartz-Albiez, R. & Vlasak, R. in Sialoglyco Chemistry and Biology I: Biosynthesis, Structural Diversity and Sialoglycopathologies Vol. 366 (eds Gerardy-Schahn, R. et al.) 1–30 (Springer, 2015).

  58. 58.

    Vimr, E. R. Unified theory of bacterial sialometabolism: how and why bacteria metabolize host sialic acids. ISRN Microbiol. 2013, 816713 (2013).

  59. 59.

    Robbe-Masselot, C., Maes, E., Rousset, M., Michalski, J. C. & Capon, C. Glycosylation of human fetal mucins: a similar repertoire of O-glycans along the intestinal tract. Glycoconj. J. 26, 397–413 (2009).

  60. 60.

    Pezzicoli, A., Ruggiero, P., Amerighi, F., Telford, J. L. & Soriani, M. Exogenous sialic acid transport contributes to group B Streptococcus infection of mucosal surfaces. J. Infect. Dis. 206, 924–931 (2012).

  61. 61.

    Bidossi, A. et al. A functional genomics approach to establish the complement of carbohydrate transporters in Streptococcus pneumoniae. PLoS ONE 7, e33320 (2012).

  62. 62.

    Marion, C., Burnaugh, A. M., Woodiga, S. A. & King, S. J. Sialic acid transport contributes to pneumococcal colonization. Infect. Immun. 79, 1262–1269 (2011).

  63. 63.

    Gangi Setty, T. et al. Molecular characterization of the interaction of sialic acid with the periplasmic binding protein from Haemophilus ducreyi. J. Biol. Chem. 293, 20073–20084 (2018).

  64. 64.

    Xiao, A. et al. Streptococcus pneumoniae sialidase SpNanB-catalyzed one-pot multienzyme (OPME) synthesis of 2,7-anhydro-sialic acids as selective sialidase inhibitors. J. Org. Chem. 83, 10798–10804 (2018).

  65. 65.

    Duncan, S. H., Hold, G. L., Harmsen, H. J., Stewart, C. S. & Flint, H. J. Growth requirements and fermentation products of Fusobacterium prausnitzii, and a proposal to reclassify it as Faecalibacterium prausnitzii gen. nov., comb. nov. Int. J. Syst. Evol. Microbiol. 52, 2141–2146 (2002).

  66. 66.

    Liu, H. & Naismith, J. H. A simple and efficient expression and purification system using two newly constructed vectors. Protein Expr. Purif. 63, 102–111 (2009).

  67. 67.

    Gerlt, J. A. et al. Enzyme Function Initiative-Enzyme Similarity Tool (EFI-EST): a web tool for generating protein sequence similarity networks. Biochim. Biophys. Acta 1854, 1019–1037 (2015).

  68. 68.

    Shannon, P. et al. Cytoscape: a software environment for integrated models of biomolecular interaction networks. Genome Res. 13, 2498–2504 (2003).

  69. 69.

    Medema, M. H., Takano, E. & Breitling, R. Detecting sequence homology at the gene cluster level with MultiGeneBlast. Mol. Biol. Evol. 30, 1218–1223 (2013).

  70. 70.

    Owen, C. D. et al. Streptococcus pneumoniae NanC: structural insights into the specificity and mechanism of a sialidase that produces a sialidase inhibitor. J. Biol. Chem. 290, 27736–27748 (2015).

  71. 71.

    Perutka, J., Wang, W., Goerlitz, D. & Lambowitz, A. M. Use of computer-designed group II introns to disrupt Escherichia coli DExH/D-box protein and DNA helicase genes. J. Mol. Biol. 336, 421–439 (2004).

  72. 72.

    Mayer, M. & Meyer, B. Characterization of ligand binding by saturation transfer difference NMR spectroscopy. Angew. Chem. Int. Edn 38, 1784–1788 (1999).

  73. 73.

    Hwang, T. L. & Shaka, A. J. Water suppression that works. Excitation sculpting using arbitrary wave-forms and pulsed-field gradients. J. Magn. Reson. A 112, 275–279 (1995).

  74. 74.

    Nepravishta, R., Walpole, S., Tailford, L., Juge, N. & Angulo, J. Deriving ligand orientation in weak protein-ligand complexes by DEEP-STD NMR spectroscopy in the absence of protein chemical-shift assignment. ChemBioChem 20, 340–344 (2019).

  75. 75.

    Mayer, M. & James, T. L. NMR-based characterization of phenothiazines as a RNA binding scaffold. J. Am. Chem. Soc. 126, 4453–4460 (2004).

  76. 76.

    Sanchez-Weatherby, J. et al. VMXi: a fully automated, fully remote, high-flux in situ macromolecular crystallography beamline. J. Synchrotron Radiat. 26, 291–301 (2019).

  77. 77.

    Krissinel, E., Uski, V., Lebedev, A., Winn, M. & Ballard, C. Distributed computing for macromolecular crystallography. Acta Crystallogr. D 74, 143–151 (2018).

  78. 78.

    Vagin, A. & Teplyakov, A. Molecular replacement with MOLREP. Acta Crystallogr. D 66, 22–25 (2010).

  79. 79.

    Keegan, R. M. & Winn, M. D. MrBUMP: an automated pipeline for molecular replacement. Acta Crystallogr. D 64, 119–124 (2008).

  80. 80.

    van Beusekom, B., Joosten, K., Hekkelman, M. L., Joosten, R. P. & Perrakis, A. Homology-based loop modelling yields more complete crystallographic protein structures. IUCr 5, 585–594 (2018).

  81. 81.

    Emsley, P. Tools for ligand validation in Coot. Acta Crystallogr. D 73, 203–210 (2017).

  82. 82.

    Smart, O. S. et al. Exploiting structure similarity in refinement: automated NCS and target-structure restraints in BUSTER. Acta Crystallogr. D 68, 368–380 (2012).

  83. 83.

    Langer, G., Cohen, S. X., Lamzin, V. S. & Perrakis, A. Automated macromolecular model building for X-ray crystallography using ARP/wARP version 7. Nat. Protoc. 3, 1171–1179 (2008).

  84. 84.

    Winn, M. D., Murshudov, G. N. & Papiz, M. Z. Macromolecular TLS refinement in REFMAC at moderate resolutions. Methods Enzymol. 374, 300–321 (2003).

  85. 85.

    Williams, C. J. et al. MolProbity: more and better reference data for improved all-atom structure validation. Protein Sci. 27, 293–315 (2018).

  86. 86.

    McCoy, A. J. et al. Gyre and gimble: a maximum-likelihood replacement for Patterson correlation refinement. Acta Crystallogr. D 74, 279–289 (2018).

  87. 87.

    Vonrhein, C. et al. Data processing and analysis with the autoPROC toolbox. Acta Crystallogr. D 67, 293–302 (2011).

  88. 88.

    STARANISO (Global Phasing Ltd., 2018); http://staraniso.globalphasing.org/cgi-bin/staraniso.cgi

  89. 89.

    Kim, D. et al. TopHat2: accurate alignment of transcriptomes in the presence of insertions, deletions and gene fusions. Genome Biol. 14, R36 (2013).

  90. 90.

    Anders, S. & Huber, W. Differential expression analysis for sequence count data. Genome Biol. 11, R106 (2010).

  91. 91.

    Schindelin, J. et al. Fiji: an open-source platform for biological-image analysis. Nat. Methods 9, 676–682 (2012).

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Acknowledgements

We thank M. Rejzek (JIC) for help with the purification of 2,7-anhydro-Neu5Ac; N. Minton (University of Nottingham) for access to ClosTron technology; H. Wu, M. Philo and G. Savva at QIB for their help with the SSN, mass spectrometry analysis and statistical analyses, respectively; A. Brion and A. Goldson for their technical help with gnotobiotic mouse experiments; staff at Diamond Light Source beamlines VMXi, I03, I04 and I24 for beamtime and assistance; and R. Keegan of CCP4 and STFC for assistance with phasing the RgNanA crystal structure. We acknowledge the support of the Biotechnology and Biological Sciences Research Council (BBSRC); this research was funded by the BBSRC 42854000B responsive mode grant, the BBSRC Institute Strategic Programme Gut Microbes and Health BB/R012490/1 and its constituent project BBS/E/F/000PR10353 (Theme 1, Determinants of microbe-host responses in the gut across life). A.B. was supported by the BBSRC Norwich Research Park Biosciences Doctoral Training Partnership grant number BB/M011216/1.

Author information

N.J. conceived the study and wrote the manuscript with contribution from the co-authors. A.B. performed bioinformatics analyses, transcriptomics, heterologous expression, site-directed mutagenesis, enzymatic assays, analytical product characterization (HPLC and mass spectrometry), protein–ligand interaction experiments (ITC and fluorescence spectroscopy) and R. gnavus mutagenesis. J.B. supervised the ClosTron mutagenesis. G.H.T. supervised the fluorescence spectroscopy experiments. D.L. developed the HPLC and mass spectrometry analysis protocols. C.D.O. performed the X-ray crystallography experiments under the supervision of M.A.W. L.V. performed the immunohistochemistry experiments. E.C. contributed to the mouse study and RNA-seq analyses. A.B., E.C., L.V. and D.L. worked under the supervision of N.J. R.N. performed the NMR experiments under the supervision of J.A. A.X. and W.L. synthesized the 2,7-anydro-Neu5Ac used in this study under the supervision of X.C. J.C. performed the cluster bioinformatics analyses. All of the authors reviewed and corrected the final manuscript.

Correspondence to Nathalie Juge.

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Bell, A., Brunt, J., Crost, E. et al. Elucidation of a sialic acid metabolism pathway in mucus-foraging Ruminococcus gnavus unravels mechanisms of bacterial adaptation to the gut. Nat Microbiol 4, 2393–2404 (2019). https://doi.org/10.1038/s41564-019-0590-7

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