Gastrointestinal Tract: Intestinal Fatty Acid Metabolism and Implications for Health

  • Lesley HoylesEmail author
  • R. John Wallace
Living reference work entry
Part of the Handbook of Hydrocarbon and Lipid Microbiology book series (HHLM)


Short-chain fatty acids (SCFA) are formed from the fermentation of sugars and complex carbohydrates by gastrointestinal (GI) bacteria in man. Acetate is the most abundant SCFA, with lower amounts of propionate and butyrate formed. Propionate and butyrate are also formed from the products of carbohydrate fermentation by other bacteria, for example, from lactate, succinate, and acetate. SCFA play a role in regulating transit of digesta through the GI tract, and in health by, for example, decreasing the risk of colon cancer (butyrate), and promoting satiety and reducing cholesterol load (propionate). Major butyrate-producing (Roseburia and Faecalibacterium spp.) and propionate-producing (Negativicutes and Bacteroides spp.) bacteria are among the most abundant microbes present in the large intestine. Metabolism of longer-chain fatty acids occurs mainly by hydration or hydrogenation of unsaturated fatty acids, the pathway depending on the individual. Hydroxystearic acids are formed in the intestine, particularly under disease conditions. Metabolism of linoleic acid results in the formation of conjugated linoleic acids (CLA) by several species, including Roseburia hominis and Roseburia inulinivorans. Enhancement of GI CLA formation, possibly using probiotics, may be useful in preventing or treating inflammatory bowel disease and be protective of key health-promoting bacteria such as Faecalibacterium prausnitzii.

1 Introduction

The human large intestine harbors 1010–1011 bacteria per gram of wet weight contents (Macfarlane et al. 1998). In this milieu, short-chain fatty acids (SCFAs) are formed from dietary fiber, resistant starches, and other poly- and oligosaccharides that escape digestion in the stomach and small intestine (Flint et al. 2008). This metabolism is considered to be generally beneficial to the health of the host, for reasons that will be described below. Longer-chain fatty acids (FAs) are also released from the various dietary, host, and bacterial fats that enter the gastrointestinal (GI) tract (Chapter “Antimicrobial Activity of Essential Oils”). The metabolism of these FA is restricted, as far as we know, mainly to the more reactive mono- and polyunsaturated FA (MUFA and PUFA, respectively), which undergo hydration or hydrogenation or perhaps both. Once again, this FA metabolism has important implications for health. This chapter summarizes what is known about FA metabolism by GI bacteria, particularly where the metabolism may impinge upon human health.

2 SCFA Production

Fermentation of polysaccharides predominates in the ascending colon, where bacterial numbers are highest, and the microbial by-products of carbohydrate fermentation include CO2, CH4, H2, SCFA, bacterial cell mass, and some heat. Polysaccharides entering the large intestine comprise insoluble plant fiber, in the form of plant cell wall polysaccharides, oligosaccharides, storage polysaccharides (e.g., inulin), and resistant starch. Approximately 10% of the human energy balance is derived from fermentation of polysaccharides by bacteria resident in the large intestine (Bergman 1990). These polysaccharides are fermented to SCFA, predominantly acetate, propionate, and butyrate. SCFA concentrations are highest in the cecum (131 mmol/kg digesta). Concentrations fall during passage through the intestine due to absorption of the SCFA across the gut wall (Cummings et al. 1987).

Acetate is the SCFA produced in the greatest quantity, followed by propionate and butyrate (57:22:21 molar ratio; Cummings et al. 1987). Other organic acids, such as lactate, succinate, formate, valerate and caproate, or branched-chain fatty acids (BCFA) generated from amino acids, are found in much smaller amounts in the intestine (Cummings et al. 1987). Ethanol can also be produced in minor amounts as a by-product of carbohydrate fermentation. The overall pathways of SCFA production are summarized in Fig. 1. Only a few features of SCFA production, mainly those most relevant to health, can be discussed here.
Fig. 1

Prokaryotes involved in the formation and conversion of SCFAs in the human large intestine. A simplified diagram of polysaccharide breakdown and the main routes of carbohydrate fermentation in the large intestine are shown. Two distinct cross-feeding mechanisms operate in the GI tract: one due to the consumption of fermentation end products (lactate, acetate, succinate) and the other due to cross-feeding of partial breakdown products from complex substrates (Falony et al. 2006; Belenguer et al. 2007; Reichardt et al. 2014). Both mechanisms contribute to the production of butyrate and propionate. Updated from Hoyles and Wallace (2010) to include propionate formation (Reichardt et al. 2014) and the bifid shunt, which is restricted to Bifidobacterium spp. in the human gut via the action of fructose 6-phosphate phosphoketolase (Pokusaeva et al. 2011). *Akkermansia muciniphila is thought to be produce propionate via the succinate pathway. †Species predicted from sequence analyses to be capable of reductive acetogenesis (Ohashi et al. 2007; Hylemon et al. 2018). (Figure reproduced with permission of Lesley Hoyles (original available from

Absorbed acetate is the principal route by which the body obtains energy from carbohydrates not absorbed in the small intestine (Salminen et al. 1998). Approximately 60% of the acetate produced by GI bacteria is retained in the liver (Pouteau et al. 2003). Animal studies have shown that acetate is secreted by the liver when portal blood concentrations fall below a critical level (Salminen et al. 1998). Acetate is formed by nearly all heterotrophic anaerobic gut bacteria, but up to one-third of all acetate in the large intestine can come from reductive acetogenesis (Flint 2006), whereby species such as Blautia producta form acetate from H2 and CO2 (Ohashi et al. 2007; Rey et al. 2010) (Fig. 1). Acetate formation benefits bacteria because it results in energy generation by substrate-level phosphorylation of ADP to ATP (Macfarlane and Gibson 1997).

Propionate can be formed by several routes in the GI microbiota – directly from sugars by single species or indirectly by cross-feeding from succinate and lactate producers (Fig. 1). There are three metabolic pathways for propionate formation: the propanediol pathway, the succinate pathway, and the acrylate pathway (Reichardt et al. 2014; Louis and Flint 2017). Belenguer et al. (2007) showed the acrylate pathway was the main pathway by which lactate was converted to propionate. Succinate producers are fairly common. Bacteroides cellulosilyticus, Bacteroides fragilis, Bacteroides ovatus, and Bacteroides thetaiotaomicron have been shown to produce acetate and succinate from sugars in pure culture (Robert et al. 2007). Most cultured members of Clostridium cluster IX (e.g., Selenomonas, Megasphaera, and Veillonella spp.) produce propionate via decarboxylation of succinate (Flint 2006). Propionate formation achieves two needs of bacteria, one for the disposal of reducing equivalents, especially NADH, and the other for ATP synthesis. The presence of electron transport chain components such as cytochromes is widespread in these bacteria (Macfarlane and Gibson 1997), and it can be assumed that they produce ATP by electron transport-linked as well as substrate-level phosphorylation.

Butyrate is presently thought to be the most significant of the SCFA in terms of its influence on human health. Like propionate, butyrate can be formed by more than one route. In terms of substrate, butyrate is formed from sugars by many members of Clostridium clusters XIVa and IV, including Eubacterium, Fusobacterium, Roseburia, and Butyrivibrio (Walker et al. 2005; Duncan et al. 2006; Flint 2006). It may also be formed from lactate, probably by the same bacteria. The fermenter studies of Belenguer et al. (2007) suggested that Eubacterium hallii played a major role in the conversion of lactate to butyrate. Bacteria also employ two metabolic routes to form butyrate, one in which the final step is butyrate kinase and the other where butyrate is released from butyryl-CoA by an acyl transferase. The latter route seems to be more common, used by the predominant Roseburia and Faecalibacterium species (Duncan et al. 2002; Louis et al. 2004). The enzymic mechanism facilitates the formation of butyrate from exogenous acetate, an important route of butyrate synthesis (Duncan et al. 2002; Duncan et al. 2004a; Falony et al. 2006).

Lactate seldom accumulates in the colon to a significant level, despite many GI bacterial species producing lactate in pure culture (Macfarlane and Gibson 1997). Interspecies cross-feeding on lactate undoubtedly contributes to the low concentrations in the colon, but it is also likely that the specific growth rate of bacteria in situ is low, such that bacteria that form lactate at maximum growth rates form different products at lower growth rates. This behavior is well documented in ruminal bacteria such as Streptococcus bovis and Selenomonas ruminantium (Russell and Wallace 1997).

Nothing is known about the main members of the microbiota responsible for converting amino acids to BCFA (Flint 2006). However, it is known that 2-methylbutyrate, isobutyrate, and isovalerate are formed as products of isoleucine, valine, and leucine fermentation, respectively, by the fecal microbiota (Macfarlane and Gibson 1995). The proportion of BCFA versus total SCFA is higher in the descending colon compared with other FAs, reflecting that protein fermentation exceeds carbohydrate fermentation in this region due to substrate availability and increased pH (Cummings et al. 1987).

3 Implications of SCFA Production for Health

The total concentration of SCFA in the GI tract and concentrations of individual SCFA have implications for human health. High total SCFA concentrations lead to a lowering of pH. A low pH in feces has been associated with a decreased incidence of colorectal cancer in various populations (Malhotra 1982; Walker et al. 1986). In addition, SCFA contribute to the normal function of the large bowel by stimulating colonic blood flow and water and salt uptake (Roediger 1980; Salminen et al. 1998). SCFA also affect gut motility. Via their remote effects, SCFA are involved in the regulation of upper gut motility in a dose-dependent manner (Cherbut 2003). There are two types of contraction involved in gut motility: tonic contractions, which decrease the volume of the GI tract, and peristaltic contractions, which are propagated over long or short distances and are associated with the backward and forward movement of GI contents. Transit time is dependent upon the pattern of occurrence of these two types of contraction (Cherbut 2003). It is well known that nutrients in the proximal and distal small intestinal lumen, particularly SCFA, participate in regulating gastric motility and emptying. SCFA also influence appetite, via effects on leptin, glucagon-like peptide-1 (GLP-1), and peptide YY (PYY), which act as signals of satiety to the human body (Canfora et al. 2015). Failures in leptin signaling are associated with severe obesity, hyperphagia, infertility, and immunological defects. Studies in mice have suggested a role for SCFA (C2–C6 SCFA and C4–C6 branched-chain FA) in leptin production (Xiong et al. 2004), with SCFA (particularly C3 and C5) activating FFAR3 (GPR41). In addition, in vitro and in vivo studies in mice and humans have shown roles for FFAR3 and FFAR2 (GPR43) in GLP-1 and PYY secretion (Canfora et al. 2015).

Among specific SCFA, acetate is an energy source that is not only absorbed from the intestine but also acts as a precursor for butyrate formation in the intestine. Absorbed propionate is largely cleared from the system by the liver (Salminen et al. 1998). A study in which humans were fed propionate supplements has suggested a role for propionate in improving glucose tolerance and insulin sensitivity, mediated by its effect on hepatic carbohydrate metabolism (Venter et al. 1990). Another feeding study, in which subjects consumed propionate-enriched bread, showed that propionate ingestion reduced high-density lipoprotein and increased serum triglyceride concentrations in addition to improving carbohydrate tolerance (Todesco et al. 1991). In addition, the activity of salivary amylase was inhibited by propionate. It was suggested that this was an additional mechanism by which propionate acted on the host, in addition to its proposed inhibitory effect on HMG-CoA, one of the key regulatory enzymes of cholesteryl ester synthesis from cholesterol.

Of the SCFA produced in the large intestine, butyrate has received most attention due to its role in protection from colorectal cancer and colitis (Roediger 1990; Williams et al. 2003). Butyrate is the principal energy source for colonocytes, is essential in maintaining tissue homeostasis in the colonic epithelium, plays roles in suppressing GI inflammation and in lipid metabolism, and exhibits a range of anti-tumorigenic effects on many cancer cell lines (Hamer et al. 2008). It is estimated that butyrate provides between 40% and 70% of the energy required by the colonic mucosa (Roediger 1980). Using microarray analysis of butyrate-regulated genes in colonic epithelial cells, 221 potentially butyrate-responsive genes specifically associated with the processes of proliferation, differentiation, and apoptosis have been identified (Daly and Shirazi-Beechey 2006).

Lactate only accumulates in significant amounts (up to 100 mM) in the feces of individuals suffering from severe malabsorption, or who have undergone gut resections or are suffering from inflammatory bowel disease and proctitis or Crohn’s colitis (Hove et al. 1994; Hove and Mortensen 1995). The accumulation of d-lactate in short bowel syndrome can lead to neurotoxicity and cardiac arrhythmia (Duncan et al. 2004b). Under normal conditions, the amount of lactate present in the feces is low (<5 mM; Duncan et al. 2004b). Secretion of lactate by the mucosa is minimal in healthy individuals but is increased in patients with colonic inflammation: increased fecal lactate concentrations in these patients are due to mucosal secretions and not to bacterial activity (Hove et al. 1995).

Higher (25%) concentrations of isobutyrate and isovalerate formation have been observed in vitro with systems inoculated with feces from individuals with inflammatory bowel disease (five with Crohn’s disease; three with ulcerative colitis) compared with systems inoculated with feces from healthy individuals (van Nuenen et al. 2004). This is undoubtedly linked to the detrimental effects of other products of proteolytic fermentation in the etiology of inflammatory bowel disease (Gibson et al. 1989; Martin and Rhodes 2000), but much more work is required to establish if there is a direct effect of BCFA.

Upon their uptake into systemic circulation, the three main SCFA are potent bioactive molecules and are found at micromolar concentrations in the peripheral blood of healthy individuals (acetate, 22–42 μM; propionate, 0.9–1.2 μM; butyrate, 0.3–1.5 μM) (Hoyles et al. 2018). It is becoming clear SCFA contribute to the gut–brain axis [i.e., gut-microbiota-generated metabolites influence the central nervous system (CNS)]. All three SCFAs activate members of the free fatty acid receptor (FFAR) family of G protein-coupled receptors; acetate, propionate, and butyrate have affinity in the low millimolar to high micromolar range for FFAR2; propionate and butyrate have mid- to low micromolar affinity for FFAR3. FFAR3 is expressed in human and animal brain cells (Hoyles et al. 2018). Propionate stimulates intestinal gluconeogenesis via a gut–brain neural circuit involving FFAR3 in mice (De Vadder et al. 2014), and increased GI propionate has been associated with reward pathway activity in humans (Byrne et al. 2016). In vitro, both propionate and butyrate are protective to the blood–brain barrier, while this is not the case for acetate, suggesting FFAR3 links SCFA to regulation of brain barrier function (Hoyles et al. 2018). Gut-microbiota-generated acetate can be transported across the blood–brain barrier to the hypothalamus, decreasing food intake through appetite suppression (Frost et al. 2014). The link between SCFA and the gut–brain axis warrants further study. Prebiotics may be one way of promoting SCFA production by GI bacteria to influence the gut–brain axis.

FFAR2 and FFAR3 are expressed in the human white adipose tissue, skeletal muscle, and liver. Acetate and propionate activate FFAR2 in vitro to reduce intracellular lipolytic activity of murine adipocytes (Ge et al. 2008). Hydroxycarboxylic acid receptor 2 HCAR2 (GPR109a), which responds to butyrate but not acetate or propionate, is expressed in gut epithelial cells, adipocytes, and immune cells and contributes to intestinal barrier function. Olfactory receptor Olfr78 (Or51e2), expressed in kidney and vascular smooth muscle cells of mice, can modulate blood pressure when stimulated by butyrate and propionate (Canfora et al. 2015; Kim 2018). The interaction of this small number of host receptors with SCFA and our limited understanding of their effects in humans suggest there may be many more host–microbiome receptor interactions to be discovered. Readers are directed to Canfora et al. (2015) and Kim (2018) for reviews of our current knowledge on the interactions between host receptors and SCFA and implications for host health.

4 Metabolism of FA by GI Bacteria: Mechanisms

Our knowledge of lipid metabolism in the human colon has for many decades been rudimentary in comparison with the rumen (Harfoot and Hazlewood 1997). Nevertheless, the knowledge gained from rumen studies has helped advances to be made recently in FA metabolism by human GI bacteria. There is no evidence of oxidation or elongation of FA originally from the diet, although such reactions must occur within bacteria for their endogenous FA synthesis. Metabolism of FA in the large intestine occurs predominantly with MUFA and PUFA, via hydration and biohydrogenation of the unsaturated bonds in the aliphatic chain.

It has been known for many years that the composition of FA in human feces differs greatly from the FA composition of foods (James et al. 1961). For the most part, it was unclear to what extent the different composition was due to differential absorption of FA from the GI tract and how much was due to the metabolic activity of bacteria in the intestine. Some hydroxy FAs were present that did not appear in the diet (James et al. 1961), however, indicating a likely involvement of bacteria. James et al. (1961) postulated that hydroxystearic acids (HSA) arose as an intermediary of the oxidation of stearic acid by GI bacteria. Evidence that HSA were produced by GI bacteria was obtained using dogs that had steatorrhea (fecal fat excretion) secondary to experimentally produced GI blind loops. Dogs treated with tetracycline or which had the blind loop excluded overcame the steatorrhea and had lower concentrations of HSA in the feces, with both outcomes apparently resulting from the elimination of bacterial overgrowth in the intestine (Kim and Spritz 1968). This work showed that HSA could be produced in significant amounts only from oleic and linoleic acids when incubated with the feces of a human and a dog with steatorrhea. Stearic acid was not converted to a hydroxy derivative in significant amounts in either of these in vitro systems (Kim and Spritz 1968). Thus, HSA were formed by hydration of the Δ9 double bond in unsaturated FAs rather than by the oxidation of the saturated acid. Thomas (1972) showed that many anaerobic bacteria, including some colonic species, carried out hydration of oleic acid to HSA. Clostridium perfringens was the most active species. Pearson (1973) incubated 228 strains of GI bacteria from 5 genera with oleic acid and found that 103 strains formed HSA. Thus, HSA formation from unsaturated FA is a widespread function among GI bacteria.

Other hydroxy acids may be formed as intermediates in the metabolism of PUFA. Pearson (1973) did not detect unsaturated hydroxy FAs being produced from linoleic acid by 14 strains of fecal bacteria. In contrast, Devillard et al. (2007) found that some Roseburia strains formed a hydroxy FA identified as a 10-hydroxy,cis-12-18:1. Strains of Lactobacillus, Lactococcus, Eubacterium, Propionibacterium, Bifidobacterium, and Faecalibacterium produced the same hydroxy FA, although to a lower extent than most Roseburia strains. The 10-hydroxy,cis-12-18:1 was converted by the mixed GI microbiota transiently to cis-9,trans-11-18:2 and then to trans-11-18:1 (Fig. 2, Devillard et al. 2007). A different metabolic route can be found in some lactic acid bacteria, such as Lactobacillus plantarum, resulting in the formation of conjugated linoleic acid (CLA) via 10-hydroxy,cis-12-18:1 and also 10-oxo,cis-12-18:1 (Kishino et al. 2013); however, although L. plantarum is described as a “representative gut bacterium,” its abundance is many-fold lower than the typical obligate anaerobes (Qin et al. 2010), and the predominant metabolic fate of linoleic acid in the human colon is biohydrogenation (Devillard et al. 2009). Nevertheless, if they are formed, 10-hydroxy,cis-12-18:1 could be beneficial in terms of lowering inflammation in gut tissues (Miyamoto et al. 2015), while 10-oxo,cis-12-18:1 has a variety of anti-obesity effects (Kim et al. 2017).
Fig. 2

LA metabolism by human fecal bacteria. (Adapted from Devillard et al. 2007). Different fill colors of arrows indicate that the reaction is carried out by bacterial different species. CLA, conjugated linoleic acid, of which rumenic acid is one geometric isomer

The main route of FA metabolism, biohydrogenation, was found in mixed GI bacteria from rats (Eyssen and Parmentier 1974) but was not confirmed in man until the work of Howard and Henderson (1999). The same discovery had been made in the rumen 35 years earlier (Polan et al. 1964). Most attention has been paid to the biohydrogenation of linoleic acid (LA, cis-9,cis-12-18:2), which is metabolized mainly by conversion to the conjugated dienoic acid, rumenic acid (RA, cis-9,trans-11-18:1), which is then hydrogenated to vaccenic acid (VA, trans-11-18:1) and then to stearic acid (18:0) (Fig. 2). α-Linolenic acid (LNA, cis-9,cis-12,cis-15-18:3) is also metabolized rapidly by the fecal microbiota, forming a mixture of 18:3 and 18:2 isomers (Howard and Henderson 1999). Since the route of metabolism of LA by fecal bacteria is similar to that of the ruminal microbiota, one might expect that the pattern by which LNA is metabolized, via firstly cis-9,trans-11,cis-15-18:3, to be similar as well (Harfoot and Hazlewood 1997). Butyrivibrio fibrisolvens, Roseburia inulinivorans, and Roseburia hominis produced VA rapidly from LA, presumably via RA. The bacteria responsible for the conversion of vaccenic acid to stearic acid in the human colon are unknown. Identification of these bacteria may be difficult. From work done with ruminal bacteria (Maia et al. 2007; Paillard et al. 2007), it is known that the bacteria responsible [related to Butyrivibrio proteoclasticus (formerly Clostridium proteoclasticum)] are extremely sensitive to the toxic effects of unsaturated FAs. Growth of the bacteria was necessary for stearate formation to occur, but, as LA was toxic at concentrations as low as 5 μg/ml, growth was inhibited by the substrate. The same may be true of human GI bacteria (Devillard et al. 2007). qPCR based on 16S rRNA gene sequences indicated that B. proteoclasticus was present only at very low numbers in human feces (Devillard et al. 2009), indicating that, as with the earlier steps in the pathway, the species responsible for stearate formation in the two gut ecosystems might be different.

To establish the mechanism by which LA was metabolized by human GI bacteria, mixed and pure cultures were incubated with deuterium oxide, and the fate of the deuterium was analyzed (McIntosh et al. 2009). Fecal bacteria from four human donors and six species of human GI bacteria were incubated with LA in deuterium oxide-enriched medium. The FA products were derivatized, separated by GC, and identified by mass spectrometry. The main CLA products in fecal suspensions, RA and trans-9,trans-11-18:2, were labeled at C-13, as were other 9,11 geometric isomers. Traces of trans-10,cis-12-18:2 formed were labeled to a much lower extent. In pure culture, Bifidobacterium breve formed labeled RA and trans-9,trans-11-18:2, while Butyrivibrio fibrisolvens, Roseburia hominis, Roseburia inulinivorans, and Blautia obeum-like strain A2-162 converted LA to VA, labeled in a manner indicating VA was formed via C-13-labelled RA. Propionibacterium freudenreichii subsp. shermanii, a possible probiotic, formed mainly RA with smaller amounts of trans-10,cis-12-18:2 and trans-9,trans-11-18:2, labeled the same as in the mixed microbiota. Ricinoleic acid (12-OH-cis-9-18:1) did not form CLA in the mixed microbiota, in contrast to CLA formation described for Lactobacillus plantarum (Ogawa et al. 2005). Results were similar to those reported for the mixed microbiota of the rumen. Thus, though the bacterial genera and species responsible for biohydrogenation in the rumen and the human intestine differ, and a second route of RA formation via a 10-OH-18:1 is present in the intestine, the overall labeling patterns of different CLA isomers formation are common to both gut ecosystems. A hydrogen-abstraction enzymic mechanism was proposed that may explain the role of a 10-OH-18:1 intermediate in 9,11-CLA formation in pure and mixed cultures (McIntosh et al. 2009).

5 Influence of Fats on, and Metabolism of FA by, GI Bacteria: Health Implications

In general, the gut microbiota has important implications for health. The composition of the diet, including fats, has a profound influence on the GI microbiota (Scott et al. 2013). Humans consuming a high-fat diet possess a different gut microbiota from those consuming a diet with a lower fat content (Turnbaugh et al. 2009), which has prompted proposals that there is a gut microbiome that is associated with, and by implication causes, obesity (Ley et al. 2005; Turnbaugh et al. 2006). Arguments and experiments for and against such a mechanism have been well rehearsed elsewhere (Chapter “Gastrointestinal Tract: Fat Metabolism in the Colon”). In particular, observations such as those by Hildebrandt et al. (2009), who found the same changes in the microbiota of mice (the high-fat diet increased Firmicutes and Proteobacteria and decreased Bacteroidetes) in both lean and obese animals, question the cause-effect relationships involving dietary fat, the microbiota, and obesity.

Different fats and their constituent FA also have different effects on gut tissues depending on their degree of unsaturation: fish oil had a greater inflammatory effect than lard, for example, in mice (Li et al. 2017).

HSA concentrations in feces increase as a consequence of various clinical conditions (Wiggins et al. 1974). Patients with ileal disease, ileal resections, or small intestinal bacterial colonization all had more than 5% HSA in their feces. The explanation given for the seemingly high levels of HSA in samples from the ileal disease and colonization patients was the simultaneous occurrence of relatively high concentrations of fat, bacteria, and bile salts in the small intestine. High levels of HSA in the ileal resection patients were not due solely to the resections as patients with comparable ileal resections and steatorrhea but who had undergone removal of most of the colon, thus reducing the time in contact with bile salts, fat and bacteria, had normal levels of HSA. Therefore, the authors concluded that the main site of formation of HSA in ileal resection patients was the colon. This was also the case for patients suffering from pancreatic insufficiency. They suggested that finding normal (<6%) concentrations of HSA in feces could not be taken to exclude any diagnosis; however, the finding of more than 5% HSA in an individual consuming a normal diet with mild steatorrhea and without previous surgery would suggest the presence of ileal disease or a stagnant loop syndrome.

Whether HSA has any implications for health other than a consequence of disease or abnormality is less clear. HSA is chemically similar to ricinoleic acid (12-hydroxy-cis-9-octadecenoic acid), the major FA in castor oil, a known cathartic. The presence of HSA in human feces led to the suggestion that it contributes to the diarrhea frequently associated with steatorrhea (James et al. 1961). In a study examining 87 patients and 12 controls, Wiggins et al. (1974) demonstrated that, in general, the percentage of HSA in feces increased as the fecal fat output rose. In individuals without steatorrhea and excreting 20 g fat/day, less than 5% of the fecal fat comprised HSA, while in individuals with steatorrhea (and, consequently, excreting more fat), between 6% and 23% of the fecal fat comprised HSA. However, no correlation was found between HSA levels and steatorrhea in the majority of cases. A positive correlation was found between fecal weight and HSA excretion expressed as an absolute value, and the authors suggested that HSA may have a direct effect on water absorption. However, they found a correlation of similar magnitude between fecal weight and steatorrhea and stated that it was impossible to conclude that there was a specific action of HSA in the GI tract. Tiruppathi et al. (1983), when working with samples from tropical sprue patients, found no correlation between HSA and the diarrhea associated with the condition, instead suggesting that unsaturated FAs may play a role in diarrhea via their inhibition of colonocyte basolateral membrane ATPases (thereby interfering with colonic Na+ and water absorption).

In addition to converting VA to cis-9,trans-11-CLA, Δ9-desaturase in host tissues converts stearic acid to oleic acid (C18:1). Oleic and stearic acids are known to decrease plasma cholesterol concentrations (Bonanome and Grundy 1988), so biohydrogenating C-18 PUFA and MUFA might be considered in some ways beneficial to health. However, it is the MUFA and PUFA themselves that many consider to be more beneficial to health.

CLA and VA, on the other hand, are considered to have possibly potent effects on human health. In vitro and in vivo animal studies have suggested that the usually most abundant CLA, RA (cis-9,trans-11-18:2), has anticarcinogenic, anti-atherosclerotic, and immune-modulating effects, as well as favorable influences on body composition, blood lipids, liver metabolism, and insulin sensitivity (Belury 2002; Wahle et al. 2004; Tricon and Yaqoob 2006). Whether such benefits can be obtained in humans consuming CLA-rich foods remains uncertain, however, partly because the doses used in rodent trials cannot be replicated in man (Fuke and Nornberg 2017). VA may arguably be considered to be functionally equivalent to RA. VA is converted to RA via the host’s Δ9-desaturase, an enzyme present in the intestine and liver (Rhee et al. 1997; Turpeinen et al. 2002; Mosley et al. 2006). VA has been shown to suppress the growth and affect cellular responses of human mammary and colon cancer cell lines through its conversion to RA by Δ9-desaturase in these cells (Miller et al. 2003). Therefore, increasing CLA and VA intake may have potential benefits on health.

A different sort of benefit deriving from biohydrogenation involves the protection of beneficial bacteria. De Weirdt et al. (2013) noted that Lactobacillus reuteri, a probiotic species, was highly sensitive to toxic effects of linoleic acid and that its survival in a gut-simulating fermenter was enhanced by biohydrogenating bacteria colonizing the mucosal layer. Even more directly, De Weirdt et al. (2017) observed a protective effect of biohydrogenating species Roseburia and Pseudobutyrivibrio toward the highly beneficial anaerobe Faecalibacterium prausnitzii.

One possibility to deliver more CLA/VA to the host was thought to be to use the biohydrogenating ability of GI bacteria. CLA formed in the intestine might be absorbed and contribute to systemic CLA. However, experiments with germ-free rats inoculated with a human fecal microbiota and fed a diet enriched with sunflower-seed oil indicated that no benefit accrued in terms of tissue concentrations of CLA (Kamlage et al. 1999). Kamlage et al. (2000) found that glucose inhibited CLA formation by mixed fecal microorganisms and speculated that this may be the reason for the earlier result. It now seems more likely that CLA is not absorbed from the intestine. Druart et al. (2014) found that CLA, and the corresponding CLnA (cis-9,trans-11, cis-15-18:3) formed from linolenic acid (cis-9, cis-12, cis-15-18:3), accumulated in the cecum and colon, but not in the jejunum and ileum. No increase was observed in plasma, suggesting that the conjugated FA were not absorbed. In contrast, Neyrinck et al. (2011) suggested that increased RA content in white adipose tissue was a response to the fermentation of arabinoxylan in the large intestine of obese mice, which would only occur if RA, or perhaps VA – a precursor of rumenic acid in mammalian tissues as described above – were absorbed from the intestine. CLA-producing lactic acid bacteria used as probiotics result in increased tissue levels of CLA in rodents (O’Shea et al. 2012); however, it might be argued that the CLA formation could occur in the small intestine, from which it is known that FA, including CLA, are absorbed.

Nevertheless, even if CLA absorption from the intestine is minimal, there may be in situ benefits from GI CLA production. In mouse models of inflammatory bowel disease, CLA were shown to exhibit anti-inflammatory properties via endoplasmic and nuclear mechanisms (Bassaganya-Riera et al. 2002; Bassaganya-Riera et al. 2004). Further studies demonstrated that CLA exerted anticarcinogenic activity in the rat colon (Nichenametla et al. 2004) and exhibited antiproliferative properties on the growth of human colon cancer cells in vitro (Kemp et al. 2003). Increased abundance of Bacteroidetes/Prevotella and Akkermansia muciniphila occurred in the cecum of mice receiving dietary CLA, which was interpreted as a beneficial effect (Chaplin et al. 2015). Therefore, mechanisms by which CLA might be delivered to and formed in the intestine have important implications for long-term human gut health. The limited evidence we have suggests that different individuals may have different types of gut microbial FA metabolism (Devillard et al. 2009; Hoyles 2009).

The precise isomer(s) of CLA that is formed from LA is significant, given the very different biological effects of RA and other isomers, particularly trans-10,cis-12-CLA (Pariza 2004; Bauman et al. 2005). Such information could have particular relevance to patients using the slimming drug tetrahydrolipstatin (Orlistat; Hauptman et al. 2000), which prevents lipid absorption in the human small intestine. Large amounts of lipid reach the large intestine, sometimes with deleterious consequences (Chanoine et al. 2005). An in vitro study conducted in a gut fermentation model (Macfarlane et al. 1998) inoculated with human feces plus olive oil in the presence and absence of Orlistat showed nonsignificant changes in cis CLA levels over time in the presence of Orlistat, but did not examine other CLA isomers (Hoyles 2009). Orlistat did, however, inhibit microbial lipases in a donor-dependent manner, and this is likely to influence isomers of CLA and other FA produced in the large intestine in the presence of the drug (Hoyles 2009). Conversion of LA to RA, the cis-9,trans-12 CLA isomer, under such circumstances might be beneficial, while the formation of 10,12 isomers could be considered potentially detrimental (Bauman et al. 2005; Pariza 2004). Although the formation of the latter was minimal in our in vitro experiments with a limited number of individuals (McIntosh et al. 2009), it is possible, by analogy with the rumen, that the population may flip over from the production of 9,11 CLA isomers and VA to the production of 10,12 CLA isomers and trans-10-18:1. This type of population shift occurs in dairy cows, causing milk fat depression.

In an effort to increase the amount of RA available to humans, probiotic bacteria have been suggested as a possible method for increasing CLA in the human intestine (Coakley et al. 2003; O’Shea et al. 2012). They may also serve to ensure that the correct isomer of CLA is formed. The rationale for this approach is that ingested bacteria could use dietary LA to produce CLA. Lactobacillus, Propionibacterium, and Bifidobacterium species are known to be involved in the formation of CLA from LA (Devillard et al. 2007). In the study of Coakley et al. (2003), strains of Bifidobacterium breve and Bifidobacterium lactis were identified that were able to convert LA to CLA at high levels. Bifidobacteria have long been used as probiotics in human foods, and they have been shown to elicit specific health benefits upon the host, for example, production of vitamins [folate, cobalamin (B12), menaquinone (K2), riboflavin (B2), and thiamine (B1)] and bacteriocins, and prevention of diarrhea (O’Connor et al. 2005; Khedkar and Ouwehand 2006). Therefore, the identification of probiotic bifidobacteria with the ability to synthesize CLA may offer novel opportunities in the rational design of improved health-promoting functional foods (Coakley et al. 2003).

6 Research Needs

Our detailed understanding of gut bacteria responsible for producing SCFA in humans needs to be underpinned with detailed understanding of the mechanisms by which SCFA influence host cells, organs, and systemic health. We still have no knowledge as to the efficiency of electron transport-linked ATP synthesis during propionate and butyrate production, and the species that are most active in BCFA formation. With the long-chain FAs, our knowledge is limited mainly to the metabolism of LA. We need to know more about the metabolism of other PUFA and MUFA. For example, Howard and Henderson (1999) found that arachidonic acid was not metabolized – is this true for other PUFA such as the fish oil FA? In microbial ecology terms, the glaring unknown is that we still do not yet know species that form stearate from LA or VA, nor do we have a true picture of the lipolytic potential of the human gut microbiota.


  1. Bassaganya-Riera J, Hontecillus R, Beitz DC (2002) Colonic anti-inflammatory mechanisms of conjugated linoleic acid. Clin Nutr 21:451–459CrossRefPubMedPubMedCentralGoogle Scholar
  2. Bassaganya-Riera J, Reynolds K, Martino-Catt S, Cui Y, Hennighausen L, Gonzalez F, Rohrer J, Benninghoff AU, Hontecillas R (2004) Activation of PPAR gamma and delta by conjugated linoleic acid mediates protection from experimental inflammatory bowel disease. Gastroenterology 127:777–791CrossRefPubMedPubMedCentralGoogle Scholar
  3. Bauman DE, Lock AL, Corl BA, Ip C, Salter AM, Parodi PM (2005) Milk fatty acids and human health: potential role of conjugated linoleic acid and trans fatty acids. In: Serjrsen K, Hvelplund T, Nielsen MO (eds) Ruminant physiology: digestion, metabolism and impact of nutrition on gene expression, immunology and stress. Wageningen Academic Publishers, Wageningen, pp 529–561Google Scholar
  4. Belenguer A, Duncan SH, Holtrop G, Anderson SE, Lobley GE, Flint HJ (2007) Impact of pH on lactate formation and utilization by human fecal microbial communities. Appl Environ Microbiol 73:6526–6533CrossRefPubMedPubMedCentralGoogle Scholar
  5. Belury MA (2002) Dietary conjugated linoleic acid in health: physiological effects and mechanisms of action. Annu Rev Nutr 22:505–531CrossRefPubMedGoogle Scholar
  6. Bergman NE (1990) Energy contributions of volatile fatty acids from the gastrointestinal tract in various species. Physiol Rev 70:567–590CrossRefPubMedGoogle Scholar
  7. Bonanome A, Grundy SM (1988) Effect of dietary stearic acid on plasma cholesterol and lipoprotein levels. N Engl J Med 318:1244–1248CrossRefPubMedGoogle Scholar
  8. Byrne CS, Chambers ES, Alhabeeb H, Chhina N, Morrison DJ, Preston T, Tedford C, Fitzpatrick J, Irani C, Busza A, Garcia-Perez I, Fountana S, Holmes E, Goldstone AP, Frost GS (2016) Increased colonic propionate reduces anticipatory reward responses in the human striatum to high-energy foods. Am J Clin Nutr 104:5–14CrossRefPubMedPubMedCentralGoogle Scholar
  9. Canfora EE, Jocken JW, Blaak EE (2015) Short-chain fatty acids in control of body weight and insulin sensitivity. Nat Rev Endocrinol 11:577–591CrossRefPubMedGoogle Scholar
  10. Chanoine JP, Hampl S, Jensen C, Boldrin M, Hauptman J (2005) Effect of orlistat on weight and body composition in obese adolescents – a randomized controlled trial. J Am Med Assoc 293:2873–2883CrossRefGoogle Scholar
  11. Chaplin A, Parra P, Serra F, Palou A (2015) Conjugated linoleic acid supplementation under a high-fat diet modulates stomach protein expression and intestinal microbiota in adult mice. PLoS One 10:e0125091CrossRefPubMedPubMedCentralGoogle Scholar
  12. Cherbut C (2003) Motor effects of short-chain fatty acids and lactate in the gastrointestinal tract. Proc Nutr Soc 62:95–99CrossRefPubMedGoogle Scholar
  13. Coakley M, Ross RP, Nordgren M, Fitzgerald G, Devery R, Stanton C (2003) Conjugated linoleic acid biosynthesis by human-derived Bifidobacterium species. J Appl Microbiol 94:138–145CrossRefPubMedGoogle Scholar
  14. Cummings JH, Pomare EW, Branch WJ, Naylor CP, Macfarlane GT (1987) Short chain fatty acids in the human large intestine, portal, hepatic and venous blood. Gut 28:1221–1227CrossRefPubMedPubMedCentralGoogle Scholar
  15. Daly K, Shirazi-Beechey SP (2006) Microarray analysis of butyrate regulated genes in colonic epithelial cells. DNA Cell Biol 25:49–62CrossRefPubMedGoogle Scholar
  16. De Vadder F, Kovatcheva-Datchary P, Goncalves D, Vinera J, Zitoun C, Duchampt A, Bäckhed F, Mithieux G (2014) Microbiota-generated metabolites promote metabolic benefits via gut-brain neural circuits. Cell 156:84–96CrossRefPubMedGoogle Scholar
  17. De Weirdt R, Coenen E, Vlaeminck B, Fievez V, Van den Abbeele P, Van de Wiele T (2013) A simulated mucus layer protects Lactobacillus reuteri from the inhibitory effects of linoleic acid. Benefic Microbes 4:299–312CrossRefGoogle Scholar
  18. De Weirdt R, Hernandez-Sanabria E, Fievez V, Mees E, Geirnaert A, Van HF, Vilchez-Vargas R, Van den Abbeele P, Jauregui R, Pieper DH, Vlaeminck B, Van de Wiele T (2017) Mucosa-associated biohydrogenating microbes protect the simulated colon microbiome from stress associated with high concentrations of polyunsaturated fat. Environ Microbiol 19:722–739CrossRefPubMedGoogle Scholar
  19. Devillard E, McIntosh FM, Duncan SM, Wallace RJ (2007) Metabolism of linoleic acid by human gut bacteria: different routes for biosynthesis of conjugated linoleic acid. J Bacteriol 189: 2566–2570CrossRefPubMedPubMedCentralGoogle Scholar
  20. Devillard E, McIntosh FM, Paillard D, Thomas NA, Shingfield KJ, Wallace RJ (2009) Differences between human subjects in the composition of the faecal bacterial community and faecal metabolism of linoleic acid. Microbiology 155:513–520CrossRefPubMedGoogle Scholar
  21. Druart C, Neyrinck AM, Vlaeminck B, Fievez V, Cani PD, Delzenne NM (2014) Role of the lower and upper intestine in the production and absorption of gut microbiota-derived PUFA metabolites. PLoS One 9:e87560CrossRefPubMedPubMedCentralGoogle Scholar
  22. Duncan SH, Barcenilla A, Stewart CS, Pryde SE, Flint HJ (2002) Acetate utilization and butyryl coenzyme A (CoA): acetate-CoA transferase in butyrate-producing bacteria from the human large intestine. Appl Environ Microbiol 68:5186–5190CrossRefPubMedPubMedCentralGoogle Scholar
  23. Duncan SH, Holtrop G, Lobley GE, Calder AG, Stewart CS, Flint HJ (2004a) Contribution of acetate to butyrate formation by human faecal bacteria. Br J Nutr 91:915–923CrossRefPubMedGoogle Scholar
  24. Duncan SH, Louis P, Flint HJ (2004b) Lactate-utilizing bacteria, isolated from human feces, that produce butyrate as a major fermentation product. Appl Environ Microbiol 70:5810–5817CrossRefPubMedPubMedCentralGoogle Scholar
  25. Duncan SH, Aminov RI, Scott KP, Louis P, Stanton TB, Flint HJ (2006) 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 56:2437–2441CrossRefPubMedGoogle Scholar
  26. Eyssen H, Parmentier G (1974) Biohydrogenation of sterols and fatty acids by the intestinal microflora. Am J Clin Nutr 27:1329–1340CrossRefPubMedGoogle Scholar
  27. Falony G, Vlachou A, Verbrugghe K, De Vuyst L (2006) Cross-feeding between Bifidobacterium longum BB536 and acetate-converting, butyrate-producing colon bacteria during growth on oligofructose. Appl Environ Microbiol 72:7835–7841CrossRefPubMedPubMedCentralGoogle Scholar
  28. Flint HJ (2006) Prokaryote diversity in the human GI tract. In: Logan NA, Lappin-Scott HM, Oyston PCF (eds) Prokaryotic diversity: mechanisms and significance. Society for General Microbiology symposium no. 66, Warwick. Cambridge University Press, Cambridge, pp 65–90CrossRefGoogle Scholar
  29. Flint HJ, Bayer EA, Rincon MT, Lamed R, White BA (2008) Polysaccharide utilization by gut bacteria: potential for new insights from genomic analysis. Nat Rev Microbiol 6:121–131CrossRefPubMedGoogle Scholar
  30. Frost G, Sleeth ML, Sahuri-Arisoylu M, Lizarbe B, Cerdan S, Brody L, Anastasovska J, Ghourab S, Hankir M, Zhang S, Carling D, Swann JR, Gibson G, Viardot A, Morrison D, Thomas LE, Bell JD (2014) The short-chain fatty acid acetate reduces appetite via a central homeostatic mechanism. Nat Commun 5:3611CrossRefPubMedPubMedCentralGoogle Scholar
  31. Fuke G, Nornberg JL (2017) Systematic evaluation on the effectiveness of conjugated linoleic acid in human health. Crit Rev Food Sci Nutr 57:1–7CrossRefGoogle Scholar
  32. Ge H, Li X, Weiszmann J, Wang P, Baribault H, Chen JL, Tian H, Li Y (2008) Activation of G protein-coupled receptor 43 in adipocytes leads to inhibition of lipolysis and suppression of plasma free fatty acids. Endocrinology 149:4519–4526CrossRefGoogle Scholar
  33. Gibson SAW, Mcfarlan C, Hay S, Macfarlane GT (1989) Significance of microflora in proteolysis in the colon. Appl Environ Microbiol 55:679–683PubMedPubMedCentralGoogle Scholar
  34. Hamer HM, Jonkers D, Venema K, Vanhoutvin S, Troost FJ, Brummer RJ (2008) Review article: the role of butyrate on colonic function. Aliment Pharmacol Ther 27:104–119CrossRefPubMedPubMedCentralGoogle Scholar
  35. Harfoot CG, Hazlewood GP (1997) Lipid metabolism in the rumen. In: Hobson PN, Stewart CS (eds) The rumen microbial ecosystem. Chapman & Hall, London, pp 382–426CrossRefGoogle Scholar
  36. Hauptman J, Lucas C, Boldrin MN, Collins H, Segal KR (2000) Orlistat in the long-term treatment of obesity in primary care settings. Arch Fam Med 9:160–167CrossRefPubMedPubMedCentralGoogle Scholar
  37. Hildebrandt MA, Hoffmann C, Sherrill-Mix SA, Keilbaugh SA, Hamady M, Chen YY, Knight R, Ahima RS, Bushman F, Wu GD (2009) High-fat diet determines the composition of the murine gut microbiome independently of obesity. Gastroenterology 137:1716–1724CrossRefPubMedPubMedCentralGoogle Scholar
  38. Hove H, Mortensen PB (1995) Influence of intestinal inflammation (IBD) and small and large bowel length on fecal short-chain fatty acids and lactate. Dig Dis Sci 40:1372–1380CrossRefPubMedPubMedCentralGoogle Scholar
  39. Hove H, Nordgaard-Andersen I, Mortensen PB (1994) Faecal dl-lactate concentration in 100 gastrointestinal patients. Scand J Gastroenterol 29:255–259CrossRefGoogle Scholar
  40. Hove H, Holtug K, Jeppesen PB, Mortensen PB (1995) Butyrate absorption and lactate secretion in ulcerative colitis. Dis Colon Rectum 38:519–525CrossRefPubMedGoogle Scholar
  41. Howard FAC, Henderson C (1999) Hydrogenation of polyunsaturated fatty acids by human colonic bacteria. Lett Appl Microbiol 29:193–196CrossRefPubMedGoogle Scholar
  42. Hoyles L (2009) In vitro examination of the effect of Orlistat on the ability of the faecal microbiota to utilize dietary lipids. PhD thesis, University of Reading, United KingdomGoogle Scholar
  43. Hoyles L, Wallace RJ (2010) Gastrointestinal tract: intestinal fatty acid metabolism and implications for health. In: Timmis K (ed) Handbook of hydrocarbon and lipid microbiology. Springer, Berlin, pp 3119–3132CrossRefGoogle Scholar
  44. Hoyles L, Snelling T, Umlai UK, Nicholson JK, Carding SR, Glen RC, McArthur S (2018) Microbiome–host systems interactions: protective effects of propionate upon the blood-brain barrier. Microbiome 6:55CrossRefPubMedPubMedCentralGoogle Scholar
  45. Hylemon PB, Harris SC, Ridlon JM (2018) Metabolism of hydrogen gases and bile acids in the gut microbiome. FEBS Lett. Scholar
  46. James AT, Webb JPW, Kellock TD (1961) The occurrence of unusual fatty acids in faecal lipids from human beings with normal and abnormal fat absorption. Biochem J 78:333–339CrossRefPubMedPubMedCentralGoogle Scholar
  47. Kamlage B, Hartmann L, Gruhl B, Blaut M (1999) Intestinal microorganisms do not supply associated gnotobiotic rats with conjugated linoleic acid. J Nutr 129:2212–2217CrossRefPubMedGoogle Scholar
  48. Kamlage B, Hartmann L, Gruhl B, Blaut M (2000) Linoleic acid conjugation by human intestinal microorganisms is inhibited by glucose and other substrates in vitro and in gnotobiotic rats. J Nutr 130:2036–2039CrossRefPubMedGoogle Scholar
  49. Kemp MQ, Jeffy BD, Romagnolo DF (2003) Conjugated linoleic acid inhibits cell proliferation through a p53-dependent mechanism: effects on the expression of G1-restriction points in breast and colon cancer cells. J Nutr 133:3670–3677CrossRefPubMedGoogle Scholar
  50. Khedkar CD, Ouwehand AC (2006) Modifying the gastrointestinal microbiota with probiotics. In: Ouwehand A, Vaughan EE (eds) Gastrointestinal microbiology. Taylor & Francis Ltd., New York, pp 315–333CrossRefGoogle Scholar
  51. Kim CH (2018) Immune regulation by microbiome metabolites. Immunology. Scholar
  52. Kim YS, Spritz N (1968) Metabolism of hydroxy fatty acids in dogs with steatorrhea secondary to experimentally produced intestinal blind loops. J Lipid Res 9:487–491PubMedGoogle Scholar
  53. Kim M, Furuzono T, Yamakuni K, Li Y, Kim YI, Takahashi H, Ohue-Kitano R, Jheng HF, Takahashi N, Kano Y, Yu R, Kishino S, Ogawa J, Uchida K, Yamazaki J, Tominaga M, Kawada T, Goto T (2017) 10-oxo-12(Z)-octadecenoic acid, a linoleic acid metabolite produced by gut lactic acid bacteria, enhances energy metabolism by activation of TRPV1. FASEB J 31:5036–5048CrossRefPubMedGoogle Scholar
  54. Kishino S, Takeuchi M, Park SB, Hirata A, Kitamura N, Kunisawa J, Kiyono H, Iwamoto R, Isobe Y, Arita M, Arai H, Ueda K, Shima J, Takahashi S, Yokozeki K, Shimizu S, Ogawa J (2013) Polyunsaturated fatty acid saturation by gut lactic acid bacteria affecting host lipid composition. Proc Natl Acad Sci USA 110:17808–17813CrossRefPubMedGoogle Scholar
  55. Ley RE, Backhed F, Turnbaugh P, Lozupone CA, Knight RD, Gordon JI (2005) Obesity alters gut microbial ecology. Proc Natl Acad Sci USA 102:11070–11075CrossRefPubMedGoogle Scholar
  56. Li H, Zhu Y, Zhao F, Song S, Li Y, Xu X, Zhou G, Li C (2017) Fish oil, lard and soybean oil differentially shape gut microbiota of middle-aged rats. Sci Rep 7:826CrossRefPubMedPubMedCentralGoogle Scholar
  57. Louis P, Flint HJ (2017) Formation of propionate and butyrate by the human colonic microbiota. Environ Microbiol 19:29–41CrossRefPubMedGoogle Scholar
  58. Louis P, Duncan SH, McCrae SI, Millar J, Jackson MS, Flint HJ (2004) Restricted distribution of the butyrate kinase pathway among butyrate-producing bacteria from the human colon. J Bacteriol 186:2099–2106CrossRefPubMedPubMedCentralGoogle Scholar
  59. Macfarlane GT, Gibson GR (1995) Microbiological aspects of the production of short-chain fatty acids in the large bowel. In: Cummings JH, Rombeau JL, Sakata T (eds) Physiological and chemical aspects of short-chain fatty acids. Cambridge University Press, Cambridge, pp 87–105Google Scholar
  60. Macfarlane GT, Gibson GR (1997) Carbohydrate fermentation, energy transduction and gas metabolism in the human large intestine. In: Mackie RI, White BA (eds) Gastrointestinal microbiology, vol. 1. Gastrointestinal ecosystems and fermentations. Chapman & Hall, New York, pp 269–318Google Scholar
  61. Macfarlane GT, Macfarlane S, Gibson GR (1998) Validation of a three-stage compound continuous culture system for investigating the effect of retention time on the ecology and metabolism of bacteria in the human colon. Microb Ecol 35:180–187CrossRefPubMedGoogle Scholar
  62. Maia MRG, Chaudhary LC, Figueres L, Wallace RJ (2007) Metabolism of polyunsaturated fatty acids and their toxicity to the microflora of the rumen. Antonie Van Leeuwenhoek 91:303–314CrossRefPubMedGoogle Scholar
  63. Malhotra SL (1982) Faecal urobilinogen levels and pH of stools in population groups with different incidence of cancer of the colon, and their possible role in aetiology. J R Soc Med 75:709–714PubMedPubMedCentralGoogle Scholar
  64. Martin HM, Rhodes JM (2000) Bacteria and inflammatory bowel disease. Curr Opin Inflamm Dis 13:503–509CrossRefGoogle Scholar
  65. McIntosh FM, Shingfield KJ, Devillard E, Russell WR, Wallace RJ (2009) Mechanism of conjugated linoleic acid and vaccenic acid formation in human fecal suspensions and pure cultures of intestinal bacteria. Microbiology 155:285–294CrossRefPubMedGoogle Scholar
  66. Miller A, McGrath E, Stanton C, Devery R (2003) Vaccenic acid (t11-18:1) is converted to c9,t11-CLA in MCF-7 and SW480 cancer cells. Lipids 38:623–632CrossRefPubMedGoogle Scholar
  67. Miyamoto J, Mizukure T, Park SB, Kishino S, Kimura I, Hirano K, Bergamo P, Rossi M, Suzuki T, Arita M, Ogawa J, Tanabe S (2015) A gut microbial metabolite of linoleic acid, 10-hydroxy-cis-12-octadecenoic acid, ameliorates intestinal epithelial barrier impairment partially via GPR40-MEK-ERK pathway. J Biol Chem 290:2902–2918CrossRefPubMedGoogle Scholar
  68. Mosley EE, McGuire MK, Williams JE, McGuire MA (2006) cis-9,trans-11 conjugated linoleic acid is synthesized from vaccenic acid in lactating women. J Nutr 136:2297–2301CrossRefPubMedGoogle Scholar
  69. Neyrinck AM, Possemiers S, Druart C, Van de Wiele T, De BF, Cani PD, Larondelle Y, Delzenne NM (2011) Prebiotic effects of wheat arabinoxylan related to the increase in bifidobacteria, Roseburia and Bacteroides/Prevotella in diet-induced obese mice. PLoS One 6:e20944CrossRefPubMedPubMedCentralGoogle Scholar
  70. Nichenametla SN, South EH, Exon JH (2004) Interaction of conjugated linoleic acid, sphingomyelin, and butyrate on formation of colonic aberrant crypt foci and immune function in rats. J Toxicol Environ Health A 67:469–481CrossRefPubMedPubMedCentralGoogle Scholar
  71. O’Connor EB, Barrett E, Fitzgerald G, Hill C, Stanton C, Ross RP (2005) Production of vitamins, exopolysaccharides and bacteriocins by probiotic bacteria. In: Tamine A (ed) Probiotic dairy products. Blackwell Publishing Ltd., Oxford, pp 167–194Google Scholar
  72. O’Shea EF, Cotter PD, Stanton C, Ross RP, Hill C (2012) Production of bioactive substances by intestinal bacteria as a basis for explaining probiotic mechanisms: bacteriocins and conjugated linoleic acid. Int J Food Microbiol 152:189–205CrossRefPubMedPubMedCentralGoogle Scholar
  73. Ogawa J, Kishino S, Ando A, Sugimoto S, Mihara K, Shimizu S (2005) Production of conjugated fatty acids by lactic acid bacteria. J Biosci Bioeng 100:355–364CrossRefGoogle Scholar
  74. Ohashi Y, Igarashi T, Kumazawa F, Fujisawa T (2007) Analysis of acetogenic bacteria in human feces with formyltetrahydrofolate synthetase sequences. Biosci Microflora 26:37–40CrossRefGoogle Scholar
  75. Paillard D, McKain N, Chaudhary LC, Walker ND, Pizette F, Koppova I, McEwan NR, Kopecny J, Vercoe PE, Louis P, Wallace RJ (2007) Relation between phylogenetic position, lipid metabolism and butyrate production by different Butyrivibrio-like bacteria from the rumen. Antonie Van Leeuwenhoek 91:417–422CrossRefGoogle Scholar
  76. Pariza MW (2004) Perspective on the safety and effectiveness of conjugated linoleic acid. Am J Clin Nutr 79:1132S–1136SCrossRefGoogle Scholar
  77. Pearson JR (1973) Alteration of dietary fat by human intestinal bacteria. Proc Nutr Soc 32:8A–9AGoogle Scholar
  78. Pokusaeva K, Fitzgerald GF, van Sinderen D (2011) Carbohydrate metabolism in bifidobacteria. Genes Nutr 6:285–306CrossRefPubMedPubMedCentralGoogle Scholar
  79. Polan CE, McNeill JJ, Tove SB (1964) Biohydrogenation of unsaturated fatty acids by rumen bacteria. J Bacteriol 88:1056–1064PubMedPubMedCentralGoogle Scholar
  80. Pouteau E, Ngyuen P, Ballèvre O, Krempf M (2003) Production rates and metabolism of short-chain fatty acids in the colon and whole body using stable isotopes. Proc Nutr Soc 62:87–93CrossRefPubMedGoogle Scholar
  81. Qin JJ, Li RQ, Raes J, Arumugam M, Burgdorf KS, Manichanh C, Nielsen T, Pons N, Levenez F, Yamada T, Mende DR, Li JH, Xu JM, Li SC, Li DF, Cao JJ, Wang B, Liang HQ, Zheng HS, Xie YL, Tap J, Lepage P, Bertalan M, Batto JM, Hansen T, Le Paslier D, Linneberg A, Nielsen HB, Pelletier E, Renault P, Sicheritz-Ponten T, Turner K, Zhu HM, Yu C, Li ST, Jian M, Zhou Y, Li YR, Zhang XQ, Li SG, Qin N, Yang HM, Wang J, Brunak S, Dore J, Guarner F, Kristiansen K, Pedersen O, Parkhill J, Weissenbach J, Bork P, Ehrlich SD, Wang J (2010) A human gut microbial gene catalogue established by metagenomic sequencing. Nature 464:59–65CrossRefPubMedPubMedCentralGoogle Scholar
  82. Reichardt N, Duncan SH, Young P, Belenguer A, McWilliam Leitch C, Scott KP, Flint HJ, Louis P (2014) Phylogenetic distribution of three pathways for propionate production within the human gut microbiota. ISME J 8:1323–1335CrossRefPubMedPubMedCentralGoogle Scholar
  83. Rey FE, Faith JJ, Bain J, Muehlbauer MJ, Stevens RD, Newgard CB, Gordon JI (2010) Dissecting the in vivo metabolic potential of two human gut acetogens. J Biol Chem 285:22082–22090CrossRefPubMedPubMedCentralGoogle Scholar
  84. Rhee SK, Kayani AJ, Ciszek A, Brenna JT (1997) Desaturation and interconversion of dietary stearic and palmitic acids in human plasma and lipoproteins. Am J Clin Nutr 65:451–458CrossRefPubMedGoogle Scholar
  85. Robert C, Chassard C, Lawson PA, Bernalier-Donadille A (2007) Bacteroides cellulosilyticus sp. nov., a cellulolytic bacterium from the human gut microbial community. Int J Syst Evol Microbiol 57:1516–1520CrossRefPubMedGoogle Scholar
  86. Roediger WE (1980) Role of anaerobic bacteria in the metabolic welfare of the colonic mucosa in man. Gut 21:793–798CrossRefPubMedPubMedCentralGoogle Scholar
  87. Roediger WE (1990) The starved colon – diminished mucosal nutrition, diminished absorption, and colitis. Dis Colon Rectum 33:858–862CrossRefPubMedGoogle Scholar
  88. Russell JB, Wallace RJ (1997) Energy yielding and consuming reactions. In: Hobson PN, Stewart CS (eds) The rumen microbial ecosystem. Chapman & Hall, London, pp 185–215Google Scholar
  89. Salminen S, Bouley C, Boutron-Ruault MC, Cummings JH, Franck A, Gibson GR, Isolauri E, Moreau MC, Roberfroid M, Rowland I (1998) Functional food science and gastrointestinal physiology and function. Br J Nutr 80(suppl. 1):S147–S171CrossRefPubMedGoogle Scholar
  90. Scott KP, Gratz SW, Sheridan PO, Flint HJ, Duncan SH (2013) The influence of diet on the gut microbiota. Pharmacol Res 69:52–60CrossRefPubMedGoogle Scholar
  91. Thomas PJ (1972) Identification of some enteric bacteria which convert oleic acid to hydroxystearic acid in vitro. Gastroenterology 62:430–435PubMedGoogle Scholar
  92. Tiruppathi K, Balasubramanian KA, Hill PG, Mathan VI (1983) Faecal free fatty acids in tropical sprue and their possible role in the production of diarrhoea by inhibition of ATPases. Gut 24:300–305CrossRefPubMedPubMedCentralGoogle Scholar
  93. Todesco T, Rao AV, Bosello O, Jenkins DJA (1991) Propionate lowers blood glucose and lipid metabolism in healthy subjects. Am J Clin Nutr 54:860–865CrossRefPubMedGoogle Scholar
  94. Tricon S, Yaqoob P (2006) Conjugated linoleic acid and human health: a critical evaluation of the evidence. Curr Opin Clin Nutr Metab Care 9:105–110CrossRefPubMedGoogle Scholar
  95. Turnbaugh PJ, Ley RE, Mahowald MA, Magrini V, Mardis ER, Gordon JI (2006) An obesity-associated gut microbiome with increased capacity for energy harvest. Nature 444:1027–1031CrossRefPubMedPubMedCentralGoogle Scholar
  96. Turnbaugh PJ, Hamady M, Yatsunenko T, Cantarel BL, Duncan A, Ley RE, Sogin ML, Jones WJ, Roe BA, Affourtit JP, Egholm M, Henrissat B, Heath AC, Knight R, Gordon JI (2009) A core gut microbiome in obese and lean twins. Nature 457:480–484CrossRefPubMedPubMedCentralGoogle Scholar
  97. Turpeinen AM, Mutanen M, Aro A, Salminen I, Basu S, Palmquist DL, Griinari JM (2002) Bioconversion of vaccenic acid to conjugated linoleic acid in humans. Am J Clin Nutr 76:504–510CrossRefPubMedGoogle Scholar
  98. van Nuenen MH, Venema K, van der Woude JC, Kuipers EJ (2004) The metabolic activity of fecal microbiota from healthy individuals and patients with inflammatory bowel disease. Dig Dis Sci 49:485–491CrossRefPubMedGoogle Scholar
  99. Venter CS, Vorster HH, Cummings JH (1990) Effects of dietary propionate on carbohydrate and lipid metabolism in healthy volunteers. Am J Gastroenterol 85:549–553PubMedGoogle Scholar
  100. Wahle KWJ, Heys SD, Rotondo D (2004) Conjugated linoleic acids: are they beneficial or detrimental to health. Prog Lipid Res 43:553–557CrossRefPubMedGoogle Scholar
  101. Walker ARP, Walker BF, Walker AJ (1986) Fecal pH, dietary fibre intake, and proneness to colon cancer in four South African populations. Br J Cancer 53:489–495CrossRefPubMedPubMedCentralGoogle Scholar
  102. Walker AW, Duncan SH, McWilliam Leitch EC, Child MW, Flint HJ (2005) 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 71:3692–3700CrossRefPubMedPubMedCentralGoogle Scholar
  103. Wiggins HS, Pearson JR, Walker JG, Russell RI, Kellock TD (1974) Incidence and significance of faecal hydroxystearic acid in alimentary disease. Gut 15:614–621CrossRefPubMedPubMedCentralGoogle Scholar
  104. Williams EA, Coxhead JM, Mathers JC (2003) Anti-cancer effects of butyrate: use of micro-array technology to investigate mechanisms. Proc Nutr Soc 62:107–115CrossRefPubMedGoogle Scholar
  105. Xiong Y, Miyamoto B, Shibata K, Valasek MA, Motoike T, Kedzierski RM, Yanagisawa M (2004) Short-chain fatty acids stimulate leptin production in adipocytes through the G protein-coupled receptor GPR41. Proc Natl Acad Sci USA 101:1045–1050CrossRefPubMedGoogle Scholar

Copyright information

© Springer Nature Switzerland AG 2019

Authors and Affiliations

  1. 1.Department of BioscienceNottingham Trent UniversityNottinghamUK
  2. 2.Rowett InstituteUniversity of AberdeenAberdeenUK

Personalised recommendations