The gut microbiota, bacterial metabolites and colorectal cancer

Key Points

  • Dietary intake has an important effect on the gut environment, much of which is mediated by the metabolic activities of the intestinal microbiota on dietary compounds. Different microbial metabolites have the potential to promote and protect against colorectal cancer (CRC).

  • Accumulating evidence suggests that microbial-derived short chain fatty acids control inflammation and regulatory T cell populations. This involves the inhibition of host histone deacetylases and interactions with cell surface receptors.

  • Multiple species in the gut microbiota have complex roles in releasing and converting diet-derived phytochemicals and host-derived bile acids and glycoconjugates, all of which influence the overall microbial metabolome.

  • Alterations in the composition of the gut microbiota can be detected both in faecal samples and in tumour-associated communities that are associated with CRC. Although many of these changes may be consequential, some specific pathogens seem to contribute to causation and disease progression.

  • It is unlikely that the aetiology of CRC can be ascribed to the presence and activities of single pathogenic species, and it is proposed that the cumulative effects of microbial metabolites should be considered to better predict and prevent cancer progression.


Accumulating evidence suggests that the human intestinal microbiota contributes to the aetiology of colorectal cancer (CRC), not only via the pro-carcinogenic activities of specific pathogens but also via the influence of the wider microbial community, particularly its metabolome. Recent data have shown that the short-chain fatty acids acetate, propionate and butyrate function in the suppression of inflammation and cancer, whereas other microbial metabolites, such as secondary bile acids, promote carcinogenesis. In this Review, we discuss the relationship between diet, microbial metabolism and CRC and argue that the cumulative effects of microbial metabolites should be considered in order to better predict and prevent cancer progression.

Access options

Rent or Buy article

Get time limited or full article access on ReadCube.


All prices are NET prices.

Figure 1: Pathways that are responsible for the biosynthesis of the major microbial metabolites that result from carbohydrate fermentation and bacterial cross-feeding.
Figure 2: Anti-inflammatory and anti-apoptotic effects of colonic bacteria and their metabolites that are thought to mitigate colorectal carcinogenesis.
Figure 3: Pro-inflammatory and DNA-damaging effects of colonic bacteria and their metabolites that are thought to contribute to colorectal carcinogenesis.
Figure 4: Major microbial metabolites formed from dietary and environmental compounds that are involved in the initiation and/or progression of colorectal cancer.


  1. 1

    Jemal, A. et al. Global cancer statistics. CA Cancer J. Clin. 61, 69–90 (2011).

  2. 2

    Fearon, E. R. & Vogelstein, B. A genetic model for colorectal tumorigenesis. Cell 61, 759–767 (1990).

  3. 3

    Rabeneck, L., Davila, J. A. & El-Serag, H. B. Is there a true 'shift' to the right colon in the incidence of colorectal cancer? Am. J. Gastroenterol. 98, 1400–1409 (2003).

  4. 4

    Gill, C. I. R. & Rowland, I. R. Diet and cancer: assessing the risk. Br. J. Nutr. 88, S73–S87 (2002).

  5. 5

    Burn, J. et al. Long-term effect of aspirin on cancer risk in carriers of hereditary colorectal cancer: an analysis from the CAPP2 randomised controlled trial. Lancet 378, 2081–2087 (2011).

  6. 6

    World Cancer Research Fund and American Institute for Cancer Research. Food, Nutrition, Physical Activity, and the Prevention of Cancer: A Global Perspective. (AICR, 2007).

  7. 7

    Jess, T., Gamborg, M., Matzen, P., Munkholm, P. & Sørensen, T. I. A. Increased risk of intestinal cancer in Crohn's disease: a meta-analysis of population-based cohort studies. Am. J. Gastroenterol. 100, 2724–2729 (2005).

  8. 8

    Danese, S., Malesci, A. & Vetrano, S. Colitis-associated cancer: the dark side of inflammatory bowel disease. Gut 60, 1609–1610 (2011).

  9. 9

    Schwabe, R. F. & Jobin, C. The microbiome and cancer. Nature Rev. Cancer 13, 800–812 (2013).

  10. 10

    Kostic, A. D. et al. Fusobacterium nucleatum potentiates intestinal tumorigenesis and modulates the tumor–immune microenvironment. Cell Host Microbe 14, 207–215 (2013).

  11. 11

    Tjalsma, H., Boleij, A., Marchesi, J. R. & Dutilh, B. E. A bacterial driver–passenger model for colorectal cancer: beyond the usual suspects. Nature Rev. Microbiol. 10, 575–582 (2012).

  12. 12

    Elinav, E. et al. Inflammation-induced cancer: crosstalk between tumours, immune cells and microorganisms. Nature Rev. Cancer 13, 759–771 (2013).

  13. 13

    Sears, C. L. & Garrett, W. S. Microbes, microbiota and colon cancer. Cell Host Microbe 15, 317–328 (2014).

  14. 14

    Flint, H. J., Scott, K. P., Louis, P. & Duncan, S. H. The role of the gut microbiota in nutrition and health. Nature Rev. Gastroenterol. Hepatol. 9, 577–589 (2012).

  15. 15

    Eckburg, P. B. et al. Microbiology: diversity of the human intestinal microbial flora. Science 308, 1635–1638 (2005).

  16. 16

    Walker, A. W. et al. Dominant and diet-responsive groups of bacteria within the human colonic microbiota. ISME J. 5, 220–230 (2011). This study is a carefully controlled human dietary trial that demonstrates rapid and reversible changes in the relative abundance of specific bacterial groups in response to carbohydrate intake.

  17. 17

    David, L. A. et al. Diet rapidly and reproducibly alters the human gut microbiome. Nature 505, 559–563 (2014).

  18. 18

    Flint, H. J., Scott, K. P., Duncan, S. H., Louis, P. & Forano, E. Microbial degradation of complex carbohydrates in the gut. Gut Microbes 3, 289–306 (2012).

  19. 19

    Salonen, A. et al. Impact of diet and individual variation on intestinal microbiota composition and fermentation products in obese men. ISME J. (2014).

  20. 20

    Duncan, S. H. et al. Reduced dietary intake of carbohydrates by obese subjects results in decreased concentrations of butyrate and butyrate-producing bacteria in feces. Appl. Environ. Microbiol. 73, 1073–1078 (2007).

  21. 21

    Wu, G. D. et al. Linking long-term dietary patterns with gut microbial enterotypes. Science 334, 105–108 (2011).

  22. 22

    De Filippo, C. et al. Impact of diet in shaping gut microbiota revealed by a comparative study in children from Europe and rural Africa. Proc. Natl Acad. Sci. USA 107, 14691–14696 (2010).

  23. 23

    Le Chatelier, E. et al. Richness of human gut microbiome correlates with metabolic markers. Nature 500, 541–546 (2013).

  24. 24

    Cotillard, A. et al. Dietary intervention impact on gut microbial gene richness. Nature 500, 585–588 (2013).

  25. 25

    Aune, D. et al. Dietary fibre, whole grains, and risk of colorectal cancer: systematic review and dose-response meta-analysis of prospective studies. BMJ 343, 1082 (2011).

  26. 26

    Chen, H. -M. et al. Decreased dietary fiber intake and structural alteration of gut microbiota in patients with advanced colorectal adenoma. Am. J. Clin. Nutr. 97, 1044–1052 (2013).

  27. 27

    Ou, J. et al. Diet, microbiota, and microbial metabolites in colon cancer risk in rural Africans and African Americans. Am. J. Clin. Nutr. 98, 111–120 (2013).

  28. 28

    Marcobal, A. et al. A metabolomic view of how the human gut microbiota impacts the host metabolome using humanized and gnotobiotic mice. ISME J. 7, 1933–1943 (2013).

  29. 29

    Macfarlane, G. T. & Gibson, G. R in Gastrointestinal Microbiology Vol. 1 (eds Maie, R. I. & White, B. A.) 269–318 (Chapman and Hall, 1997).

  30. 30

    Sieber, J. R., McInerney, M. J. & Gunsalus, R. P. Genomic insights into syntrophy: the paradigm for anaerobic metabolic cooperation. Annu. Rev. Microbiol. 66, 429–452 (2012).

  31. 31

    Baughn, A. D. & Malamy, M. H. The strict anaerobe Bacteroides fragilis grows in and benefits from nanomolar concentrations of oxygen. Nature 427, 441–444 (2004).

  32. 32

    Khan, M. T. et al. The gut anaerobe Faecalibacterium prausnitzii uses an extracellular electron shuttle to grow at oxic–anoxic interphases. ISME J. 6, 1578–1585 (2012).

  33. 33

    Circu, M. L. & Aw, T. Y. Intestinal redox biology and oxidative stress. Semin. Cell Dev. Biol. 23, 729–737 (2012).

  34. 34

    Nava, G. M., Carbonero, F., Croix, J. A., Greenberg, E. & Gaskins, H. R. Abundance and diversity of mucosa-associated hydrogenotrophic microbes in the healthy human colon. ISME J. 6, 57–70 (2012).

  35. 35

    Carbonero, F., Benefiel, A. C. & Gaskins, H. R. Contributions of the microbial hydrogen economy to colonic homeostasis. Nature Rev. Gastroenterol. Hepatol. 9, 504–518 (2012).

  36. 36

    Lewis, S. & Cochrane, S. Alteration of sulfate and hydrogen metabolism in the human colon by changing intestinal transit rate. Am. J. Gastroenterol. 102, 624–633 (2007).

  37. 37

    Miller, T. L. & Wolin, M. J. Pathways of acetate, propionate, and butyrate formation by the human fecal microbial flora. Appl. Environ. Microbiol. 62, 1589–1592 (1996).

  38. 38

    Belenguer, A. et al. Impact of pH on lactate formation and utilization by human fecal microbial communities. Appl. Environ. Microbiol. 73, 6526–6533 (2007).

  39. 39

    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).

  40. 40

    Reichardt, N. et al. Phylogenetic distribution of three pathways for propionate production within the human gut microbiota. ISME J. 8, 1323–1335 (2014).

  41. 41

    Louis, P. et al. Restricted distribution of the butyrate kinase pathway among butyrate-producing bacteria from the human colon. J. Bacteriol. 186, 2099–2106 (2004).

  42. 42

    Louis, P., Young, P., Holtrop, G. & Flint, H. J. Diversity of human colonic butyrate-producing bacteria revealed by analysis of the butyryl-CoA:acetate CoA-transferase gene. Environ. Microbiol. 12, 304–314 (2010).

  43. 43

    Louis, P. & Flint, H. J. Diversity, metabolism and microbial ecology of butyrate-producing bacteria from the human large intestine. FEMS Microbiol. Lett. 294, 1–8 (2009).

  44. 44

    Barcenilla, A. et al. Phylogenetic relationships of butyrate-producing bacteria from the human gut. Appl. Environ. Microbiol. 66, 1654–1661 (2000).

  45. 45

    Sleeth, M. L., Thompson, E. L., Ford, H. E., Zac-Varghese, S. E. K. & Frost, G. Free fatty acid receptor 2 and nutrient sensing: a proposed role for fibre, fermentable carbohydrates and short-chain fatty acids in appetite regulation. Nutr. Res. Rev. 23, 135–145 (2010).

  46. 46

    Fung, K. Y. C., Cosgrove, L., Lockett, T., Head, R. & Topping, D. L. A review of the potential mechanisms for the lowering of colorectal oncogenesis by butyrate. Br. J. Nutr. 108, 820–831 (2012).

  47. 47

    Wilson, A. J. et al. Apoptotic sensitivity of colon cancer cells to histone deacetylase inhibitors is mediated by an Sp1/Sp3-activated transcriptional program involving immediate-early gene induction. Cancer Res. 70, 609–620 (2010).

  48. 48

    Hamer, H. M. et al. Review article: the role of butyrate on colonic function. Aliment. Pharmacol. Ther. 27, 104–119 (2008).

  49. 49

    Chang, P. V., Hao, L., Offermanns, S. & Medzhitov, R. The microbial metabolite butyrate regulates intestinal macrophage function via histone deacetylase inhibition. Proc. Natl Acad. Sci. USA 111, 2247–2252 (2014). This study provides evidence that the inhibition of HDACs by butyrate is responsible for anti-inflammatory effects in colonic macrophages.

  50. 50

    Smith, P. M. et al. The microbial metabolites, short-chain fatty acids, regulate colonic T reg cell homeostasis. Science 341, 569–573 (2013). This study shows that propionate has an anti-inflammatory effect via the modulation of cT Reg cells.

  51. 51

    Furusawa, Y. et al. Commensal microbe-derived butyrate induces the differentiation of colonic regulatory T cells. Nature 504, 446–450 (2013).

  52. 52

    Atarashi, K. et al. Induction of colonic regulatory T cells by indigenous Clostridium species. Science 331, 337–341 (2011).

  53. 53

    Geuking, M. et al. Intestinal bacterial colonization induces mutualistic regulatory T cell responses. Immunity 34, 794–806 (2011).

  54. 54

    Round, J. L. & Mazmanian, S. K. Inducible Foxp3+ regulatory T-cell development by a commensal bacterium of the intestinal microbiota. Proc. Natl Acad. Sci. USA 107, 12204–12209 (2010).

  55. 55

    O'Mahony, C. et al. Commensal-induced regulatory T cells mediate protection against pathogen-stimulated NF-κB activation. PLoS Pathog. 4, e1000112 (2008).

  56. 56

    Chung, H. et al. Gut immune maturation depends on colonization with a host-specific microbiota. Cell 149, 1578–1593 (2012).

  57. 57

    Arpaia, N. et al. Metabolites produced by commensal bacteria promote peripheral regulatory T-cell generation. Nature 504, 451–455 (2013). This study shows that propionate and butyrate, but not acetate, promote the generation of anti-inflammatory T Reg cells via the inhibition of HDACs.

  58. 58

    Latham, T. et al. Lactate, a product of glycolytic metabolism, inhibits histone deacetylase activity and promotes changes in gene expression. Nucleic Acids Res. 40, 4794–4803 (2012).

  59. 59

    Ganapathy, V., Thangaraju, M., Prasad, P. D., Martin, P. M. & Singh, N. Transporters and receptors for short-chain fatty acids as the molecular link between colonic bacteria and the host. Curr. Opin. Pharmacol. 13, 869–874 (2013).

  60. 60

    Frank, H. et al. Lactaturia and loss of sodium-dependent lactate uptake in the colon of SLC5A8-deficient mice. J. Biol. Chem. 283, 24729–24737 (2008).

  61. 61

    Brown, A. J. et al. The orphan G protein-coupled receptors GPR41 and GPR43 are activated by propionate and other short chain carboxylic acids. J. Biol. Chem. 278, 11312–11319 (2003).

  62. 62

    Singh, N. et al. Activation of Gpr109a, receptor for niacin and the commensal metabolite butyrate, suppresses colonic inflammation and carcinogenesis. Immunity 40, 128–139 (2014).

  63. 63

    Thangaraju, M. et al. GPFM 09A is a G-protein-coupled receptor for the bacterial fermentation product butyrate and functions as a tumor suppressor in colon. Cancer Res. 69, 2826–2832 (2009).

  64. 64

    Maslowski, K. M. et al. Regulation of inflammatory responses by gut microbiota and chemoattractant receptor GPR43. Nature 461, 1282–1286 (2009).

  65. 65

    Buda, A. et al. Butyrate downregulates a2ß1 integrin: a possible role in the induction of apoptosis in colorectal cancer cell lines. Gut 52, 729–734 (2003).

  66. 66

    Clarke, J. M., Topping, D. L., Bird, A. R., Young, G. P. & Cobiac, L. Effects of high-amylose maize starch and butyrylated high-amylose maize starch on azoxymethane-induced intestinal cancer in rats. Carcinogenesis 29, 2190–2194 (2008).

  67. 67

    Nepelska, M. et al. Butyrate produced by commensal bacteria potentiates phorbol esters induced AP-1 response in human intestinal epithelial cells. PLoS ONE 7, e52869 (2012).

  68. 68

    Belcheva, A. et al. Gut microbial metabolism drives transformation of Msh2-deficient colon epithelial cells. Cell 158, 288–299 (2014).

  69. 69

    Ramos, S. Cancer chemoprevention and chemotherapy: dietary polyphenols and signalling pathways. Mol. Nutr. Food Res. 52, 507–526 (2008).

  70. 70

    Herr, I. & Büchler, M. W. Dietary constituents of broccoli and other cruciferous vegetables: implications for prevention and therapy of cancer. Cancer Treat. Rev. 36, 377–383 (2010).

  71. 71

    Chiva-Blanch, G. & Visioli, F. Polyphenols and health: moving beyond antioxidants. J. Berry Res. 2, 63–71 (2012).

  72. 72

    Bennett, L. L., Rojas, S. & Seefeldt, T. Role of antioxidants in the prevention of cancer. J. Exp. Clin. Med. 4, 215–222 (2012).

  73. 73

    Russell, W. & Duthie, G. Plant secondary metabolites and gut health: the case for phenolic acids. Proc. Nutr. Soc. 70, 389–396 (2011).

  74. 74

    Russell, W. R., Hoyles, L., Flint, H. J. & Dumas, M. -E. Colonic bacterial metabolites and human health. Curr. Opin. Microbiol. 16, 246–254 (2013).

  75. 75

    Cardona, F., Andrés-Lacueva, C., Tulipani, S., Tinahones, F. J. & Queipo-Ortuño, M. I. Benefits of polyphenols on gut microbiota and implications in human health. J. Nutr. Biochem. 24, 1415–1422 (2013).

  76. 76

    Russell, W. R., Labat, A., Scobbie, L. & Duncan, S. H. Availability of blueberry phenolics for microbial metabolism in the colon and the potential inflammatory implications. Mol. Nutr. Food Res. 51, 726–731 (2007).

  77. 77

    Larrosa, M. et al. Polyphenol metabolites from colonic microbiota exert anti-inflammatory activity on different inflammation models. Mol. Nutr. Food Res. 53, 1044–1054 (2009).

  78. 78

    Etxeberria, U. et al. Impact of polyphenols and polyphenol-rich dietary sources on gut microbiota composition. J. Agr. Food Chem. 61, 9517–9533 (2013).

  79. 79

    Kim, D. -H. & Jin, Y. -H. Intestinal bacterial ß-glucuronidase activity of patients with colon cancer. Arch. Pharm. Res. 24, 564–567 (2001).

  80. 80

    Humblot, C. et al. ß-glucuronidase in human intestinal microbiota is necessary for the colonic genotoxicity of the food-borne carcinogen 2-amino-3-methylimidazo[4,5-f]quinoline in rats. Carcinogenesis 28, 2419–2425 (2007).

  81. 81

    McIntosh, F. M. et al. Phylogenetic distribution of genes encoding β-glucuronidase activity in human colonic bacteria and the impact of diet on faecal glycosidase activities. Environ. Microbiol. 14, 1876–1887 (2012).

  82. 82

    Wallace, B. D. et al. Alleviating cancer drug toxicity by inhibiting a bacterial enzyme. Science 330, 831–835 (2010). This paper describes the design of an inhibitor that specifically targets bacterial β-glucuronidases, which protects against chemotherapy-associated toxicity in a mouse model.

  83. 83

    Russell, W. R. et al. High-protein, reduced-carbohydrate weight-loss diets promote metabolite profiles likely to be detrimental to colonic health. Am. J. Clin. Nutr. 93, 1062–1072 (2011). This human intervention study shows that a diet that is low in carbohydrates and high in protein decreases faecal cancer-protective metabolites and butyrate, whereas levels of potentially harmful N -nitroso compounds increase.

  84. 84

    Windey, K., de Preter, V. & Verbeke, K. Relevance of protein fermentation to gut health. Mol. Nutr. Food Res. 56, 184–196 (2012).

  85. 85

    Russell, W. R. et al. Major phenylpropanoid-derived metabolites in the human gut can arise from microbial fermentation of protein. Mol. Nutr. Food Res. 57, 523–535 (2013).

  86. 86

    Loh, Y. H. et al. N-nitroso compounds and cancer incidence: The European Prospective Investigation into Cancer and Nutrition (EPIC)-Norfolk Study. Am. J. Clin. Nutr. 93, 1053–1061 (2011).

  87. 87

    Hughes, R. & Rowland, I. R. Metabolic activities of the gut microflora in relation to cancer. Microb. Ecol. Health Dis. 12, 179–185 (2000).

  88. 88

    Di Martino, M. L. et al. Polyamines: emerging players in bacteria–host interactions. Int. J. Med. Microbiol. 303, 484–491 (2013).

  89. 89

    Pegg, A. E. Toxicity of polyamines and their metabolic products. Chem. Res. Toxicol. 26, 1782–1800 (2013).

  90. 90

    Hanfrey, C. C. et al. Alternative spermidine biosynthetic route is critical for growth of Campylobacter jejuni and is the dominant polyamine pathway in human gut microbiota. J. Biol. Chem. 286, 43301–43312 (2011).

  91. 91

    Toden, S., Bird, A. R., Topping, D. L. & Conlon, M. A. Resistant starch prevents colonic DNA damage induced by high dietary cooked red meat or casein in rats. Cancer Biol. Ther. 5, 267–272 (2006).

  92. 92

    Windey, K. et al. Modulation of protein fermentation does not affect fecal water toxicity: a randomized cross-over study in healthy subjects. PLoS ONE 7, e52387 (2012).

  93. 93

    Kuhnle, G. G. C. et al. Diet-induced endogenous formation of nitroso compounds in the GI tract. Free Radic. Bio. Med. 43, 1040–1047 (2007).

  94. 94

    Magee, E. A., Richardson, C. J., Hughes, R. & Cummings, J. H. Contribution of dietary protein to sulfide production in the large intestine: an in vitro and a controlled feeding study in humans. Am. J. Clin. Nutr. 72, 1488–1494 (2000).

  95. 95

    Marquet, P., Duncan, S. H., Chassard, C., Bernalier-Donadille, A. & Flint, H. J. Lactate has the potential to promote hydrogen sulphide formation in the human colon. FEMS Microbiol. Lett. 299, 128–134 (2009).

  96. 96

    Roediger, W. E. W., Moore, J. & Babidge, W. Colonic sulfide in pathogenesis and treatment of ulcerative colitis. Dig. Dis. Sci. 42, 1571–1579 (1997).

  97. 97

    Attene-Ramos, M. S. et al. DNA damage and toxicogenomic analyses of hydrogen sulfide in human intestinal epithelial FHs 74 int cells. Environ. Mol. Mutag. 51, 304–314 (2010).

  98. 98

    Attene-Ramos, M. S., Wagner, E. D., Gaskins, H. R. & Plewa, M. J. Hydrogen sulfide induces direct radical-associated DNA damage. Mol. Cancer Res. 5, 455–459 (2007).

  99. 99

    Barrasa, J. I., Olmo, N., Lizarbe, M. A. & Turnay, J. Bile acids in the colon, from healthy to cytotoxic molecules. Toxicol. In Vitro 27, 964–977 (2013).

  100. 100

    Bernstein, H., Bernstein, C., Payne, C. M. & Dvorak, K. Bile acids as endogenous etiologic agents in gastrointestinal cancer. World J. Gastroentero. 15, 3329–3340 (2009).

  101. 101

    Ou, J., DeLany, J. P., Zhang, M., Sharma, S. & O'Keefe, S. J. D. Association between low colonic short-chain fatty acids and high bile acids in high colon cancer risk populations. Nutr. Cancer 64, 34–40 (2012).

  102. 102

    Yoshimoto, S. et al. Obesity-induced gut microbial metabolite promotes liver cancer through senescence secretome. Nature 499, 97–101 (2013).

  103. 103

    Islam, K. B. M. S. et al. Bile acid is a host factor that regulates the composition of the cecal microbiota in rats. Gastroenterology 141, 1773–1781 (2011). This study shows that bile acids have a strong modulatory effect on the gut microbiota, which suggests that they contribute to changes in the microbiota in response to high-fat diets.

  104. 104

    Ridlon, J. M., Kang, D. -J. & Hylemon, P. B. Bile salt biotransformations by human intestinal bacteria. J. Lipid Res. 47, 241–259 (2006).

  105. 105

    Jones, B. V., Begley, M., Hill, C., Gahan, C. G. M. & Marchesi, J. R. Functional and comparative metagenomic analysis of bile salt hydrolase activity in the human gut microbiome. Proc. Natl Acad. Sci. USA 105, 13580–13585 (2008).

  106. 106

    Ridlon, J. M. & Hylemon, P. B. Identification and characterization of two bile acid coenzyme A transferases from Clostridium scindens, a bile acid 7a-dehydroxylating intestinal bacterium. J. Lipid Res. 53, 66–76 (2012).

  107. 107

    Májer, F. et al. New highly toxic bile acids derived from deoxycholic acid, chenodeoxycholic acid and lithocholic acid. Bioorgan. Med. Chem. 22, 256–268 (2014).

  108. 108

    Lee, J. Y. et al. Contribution of the 7β-hydroxysteroid dehydrogenase from Ruminococcus gnavus N53 to ursodeoxycholic acid formation in the human colon. J. Lipid Res. 54, 3062–3069 (2013).

  109. 109

    Devkota, S. et al. Dietary-fat-induced taurocholic acid promotes pathobiont expansion and colitis in Il 10−/− mice. Nature 486, 104–108 (2012).

  110. 110

    Homann, N. Alcohol and upper gastrointestinal tract cancer: the role of local acetaldehyde production. Addict. Biol. 6, 309–323 (2001).

  111. 111

    Hooper, S. J., Wilson, M. J. & Crean, S. J. Exploring the link between microorganisms and oral cancer: a systematic review of the literature. Head Neck 31, 1228–1239 (2009).

  112. 112

    Grivennikov, S. I., Greten, F. R. & Karin, M. Immunity, inflammation, and cancer. Cell 140, 883–899 (2010).

  113. 113

    Schreiber, R. D., Old, L. J. & Smyth, M. J. Cancer immunoediting: integrating immunity's roles in cancer suppression and promotion. Science 331, 1565–1570 (2011).

  114. 114

    Arthur, J. C. et al. Intestinal inflammation targets cancer-inducing activity of the microbiota. Science 338, 120–123 (2012).

  115. 115

    Dove, W. F. et al. Intestinal neoplasia in the ApcMin mouse: independence from the microbial and natural killer (beige locus) status. Cancer Res. 57, 812–814 (1997).

  116. 116

    Fukata, M. et al. Toll-like receptor-4 promotes the development of colitis-associated colorectal tumors. Gastroenterology 133, 1869–1881 (2007).

  117. 117

    Kobayashi, K. S. et al. Nod2-dependent regulation of innate and adaptive immunity in the intestinal tract. Science 307, 731–734 (2005).

  118. 118

    Neal, M. D. et al. Toll-like receptor 4 is expressed on intestinal stem cells and regulates their proliferation and apoptosis via the p53 up-regulated modulator of apoptosis. J. Biol. Chem. 287, 37296–37308 (2012).

  119. 119

    Wu, S. et al. A human colonic commensal promotes colon tumorigenesis via activation of T helper type 17 T cell responses. Nature Med. 15, 1016–1022 (2009).

  120. 120

    Sears, C. L. Enterotoxigenic Bacteroides fragilis: a rogue among symbiotes. Clin. Microbiol. Rev. 22, 349–369 (2009).

  121. 121

    Goodwin, A. C. et al. Polyamine catabolism contributes to enterotoxigenic Bacteroides fragilis-induced colon tumorigenesis. Proc. Natl Acad. Sci. USA 108, 15354–15359 (2011).

  122. 122

    Kostic, A. D. et al. Genomic analysis identifies association of Fusobacterium with colorectal carcinoma. Genome Res. 22, 292–298 (2012).

  123. 123

    Warren, R. L. et al. Co-occurrence of anaerobic bacteria in colorectal carcinomas. Microbiome 1, 16 (2013). This study identifies a potential polymicrobial signature of Gram-negative anaerobic bacteria that is associated with CRC tissue.

  124. 124

    Rubinstein, M. R. et al. Fusobacterium nucleatum promotes colorectal carcinogenesis by modulating E-cadherin/ß-catenin signaling via its FadA adhesin. Cell Host Microbe 14, 195–206 (2013).

  125. 125

    Cuevas-Ramos, G. et al. Escherichia coli induces DNA damage in vivo and triggers genomic instability in mammalian cells. Proc. Natl Acad. Sci. USA 107, 11537–11542 (2010).

  126. 126

    Nougayrède, J. -P. et al. Escherichia coli induces DNA double-strand breaks in eukaryotic cells. Science 313, 848–851 (2006).

  127. 127

    Inaba, Y. et al. Expression of the antimicrobial peptide α-defensin/cryptdins in intestinal crypts decreases at the initial phase of intestinal inflammation in a model of inflammatory bowel disease, IL-10-deficient mice. Inflamm. Bowel Dis. 16, 1488–1495 (2010).

  128. 128

    Schwerbrock, N. M. J. et al. Interleukin 10-deficient mice exhibit defective colonic Muc2 synthesis before and after induction of colitis by commensal bacteria. Inflamm. Bowel Dis. 10, 811–823 (2004).

  129. 129

    Secher, T., Samba-Louaka, A., Oswald, E. & Nougayrède, J. -P. Escherichia coli producing colibactin triggers premature and transmissible senescence in mammalian cells. PLoS ONE 8, e77157 (2013).

  130. 130

    Mukhopadhya, I., Hansen, R., El-Omar, E. M. & Hold, G. L. IBD — what role do Proteobacteria play? Nature Rev. Gastroenterol. Hepatol. 9, 219–230 (2012).

  131. 131

    Grivennikov, S. I. et al. Adenoma-linked barrier defects and microbial products drive IL-23/IL-17-mediated tumour growth. Nature 491, 254–258 (2012). This study demonstrates that defective intestinal barrier function at tumour sites facilitates bacterial translocation, which leads to the activation of myeloid cell-derived cytokine networks (involving IL-17 and IL-23) and the promotion of tumour growth.

  132. 132

    Vogelstein, B. et al. Cancer genome landscapes. Science 340, 1546–1558 (2013).

  133. 133

    McLean, M. H. et al. The inflammatory microenvironment in colorectal neoplasia. PLoS ONE 6, e15366 (2011).

  134. 134

    Lewis, S. J. & Heaton, K. W. Increasing butyrate concentration in the distal colon by accelerating intestinal transit. Gut 41, 245–251 (1997).

  135. 135

    Stephen, A. M., Wiggins, H. S. & Cummings, J. H. Effect of changing transit time on colonic microbial metabolism in man. Gut 28, 601–609 (1987).

  136. 136

    Wichmann, A. et al. Microbial modulation of energy availability in the colon regulates intestinal transit. Cell Host Microbe 14, 582–590 (2013).

  137. 137

    McOrist, A. L. et al. Fecal butyrate levels vary widely among individuals but are usually increased by a diet high in resistant starch. J. Nutr. 141, 883–889 (2011).

  138. 138

    Shen, X. J. et al. Molecular characterization of mucosal adherent bacteria and associations with colorectal adenomas. Gut Microbes 1, 138–147 (2010).

  139. 139

    Sanapareddy, N. et al. Increased rectal microbial richness is associated with the presence of colorectal adenomas in humans. ISME J. 6, 1858–1868 (2012).

  140. 140

    Wang, T. et al. Structural segregation of gut microbiota between colorectal cancer patients and healthy volunteers. ISME J. 6, 320–329 (2012). This study finds that the faecal microbiota of patients with CRC and healthy controls differs.

  141. 141

    Zackular, J. P. et al. The gut microbiome modulates colon tumorigenesis. mBio 4, e00692–13 (2013). This study provides evidence that the composition of the gut microbiota influences the development of carcinogen-induced tumours in the intestines of mice.

Download references


P.L. and H.F. acknowledge support from the Scottish Government Food Land and People programme. The authors thank A. Walker and R. Barker for critical reading of the manuscript.

Author information

Correspondence to Harry J. Flint.

Ethics declarations

Competing interests

The authors declare no competing financial interests.

PowerPoint slides


Resistant starch

The fraction of dietary starch that is not digested in the small intestine by host enzymes and reaches the colon.

Non-digestible carbohydrates

Dietary carbohydrates that are not digested by mammalian enzymes in the small intestine and reach the colon, where they may be used as substrates for the resident microbiota.


A high molecular weight glycoprotein that is produced by the gut epithelium and forms the mucus layer that lines the gut wall; it can be used as an energy source by some gut bacteria.

Non-starch polysaccharides

Non-digestible carbohydrates other than resistant starch, including cellulose, arabinoxylans, xyloglucans, pectins and gums.


A term used to describe bacteria that produce acetate from carbon dioxide and hydrogen (or formate) via the Wood–Ljungdahl pathway.


The process by which methane is formed from carbon dioxide and hydrogen (or formate) by methanogenic archaea.

Wood–Ljungdahl pathway

The collection of sequential biochemical reactions that lead to the formation of acetate from carbon dioxide and hydrogen. Several bacteria use this pathway for the generation of energy, and it is also used by certain bacteria and archaea for the assimilation of carbon dioxide into biomass.

Acrylate pathway

The collection of sequential biochemical reactions that lead to the conversion of lactate to propionate by certain Firmicutes bacteria.

Propanediol pathway

The collection of sequential biochemical reactions that lead to the conversion of deoxy-sugars (such as rhamnose and fucose) to propionate by certain gut bacteria.

Histone deacetylases

(HDACs). Enzymes that remove acetyl groups from histones, which are structural proteins that package DNA into structural units. Deacetylation leads to more condensed chromatin, which alters gene expression.

Colonic regulatory T cells

(cTReg cells). A subset of T lymphocytes that are found in the colon and are crucial for the maintenance of immune tolerance.

AP-1 signalling pathway

A pathway that regulates gene expression via the transcription factor activator protein 1 (AP-1).


Organic compounds that contain phenol moieties; they are produced by plants and are involved in many biological processes, including defence against herbivores.


Sulphur-containing natural compounds that are present in Brassica vegetables. They are converted to secondary compounds (including isothiocyanates) that are thought to have health-promoting effects.


Natural compounds that are produced by plants and belong to a range of biochemical classes.


A chemical compound that is not normally found in an organism, such as a drug or environmental pollutant.


Chemical compounds that are bound to a sugar residue via a glycosidic linkage.


Chemical compounds that are bound to a glucuronic acid residue via a glycosidic linkage.


The compounds that are produced following the removal of the sugar or sugar acid (such as glucuronic acid) residue from a glycoside or glucuronide.


The process by which compounds are converted to nitroso group (-N=O)- containing derivatives.

Secondary bile acids

Metabolites that are produced by enteric bacteria from primary bile acids.

Primary bile acids

Steroid acids that are synthesized in the liver and secreted into the gut to promote fat digestion.

Pattern recognition receptors

(PRRs). Innate immune proteins that are essential for recognizing and responding to microorganisms. The most common PRRs include Toll-like receptors, NOD-like receptors, RIGI-like receptors and DNA receptors (that is, cytosolic sensors for DNA).

Microorganism-associated molecular patterns

(MAMPS). Conserved microbial components, such as lipopolysaccharide, peptidoglycan, flagellin and nucleic acids, that are detected by the host innate immune system via pattern recognition receptors.


Inflammation of the colon or large intestine.

Rights and permissions

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Louis, P., Hold, G. & Flint, H. The gut microbiota, bacterial metabolites and colorectal cancer. Nat Rev Microbiol 12, 661–672 (2014).

Download citation

Further reading