CHAPTER 1

Synthesis of Conjugated Linoleic Acid in Ruminants and Humans

K. J. Shingfield*ab and R. J. Wallacec
a MTT Agrifood Research Finland, Animal Production Research, Jokioinen, FI-31600, Finland. E-mail: kes14@aber.ac.uk
b Institute of Biological, Environmental and Rural Sciences, Aberystwyth University, Aberystwyth, Ceredigion, SY23 3EB, UK.
c Rowett Institute of Nutrition and Health, University of Aberdeen, Bucksburn, Aberdeen, AB21 9SB, UK.


Biomedical studies have provided evidence that one or more isomers of conjugated linoleic acid (CLA) may have beneficial effects in the prevention of human chronic disease. Ruminant meat and milk are the principle source of CLA in the human diet. These foods contain numerous positional and geometric isomers of CLA with cis-9,trans-11 as the major isomer. The majority of CLA isomers in meat and milk originate from the isomerization of 18:2n-6 and 18:3n-3 in the rumen, reactions that proceed via different mechanisms catalysed by bacterial enzymes. Diet is the principal determinant of the amount and distribution of CLA isomers formed in the rumen. However, trans-7, cis-9 18:2 and cis-9, trans-11 18:2 are also formed in ruminant tissues and the mammary glands via the action of stearoyl CoA desaturase on trans-7 18:1 and trans-11 18:1, respectively. Virtually all trans-7, cis-9 18:2 and the majority of cis-9, trans-11 18:2 CLA in ruminant lipids and milk fat are synthesized endogenously. Diet is also the major factor determining the synthesis of CLA precursors in the rumen. Even though diet is the major source of CLA in humans, cis-9,trans-11 CLA is also synthesized endogenously with some evidence that CLA isomers may also be formed via the activity of microbial populations in the hindgut. The present chapter provides a comprehensive evaluation of the most recent evidence on the biochemical, microbial, nutritional and physiological factors influencing the biosynthesis of CLA isomers in ruminants and humans.


1.1 Introduction

There is increasing evidence that nutrition plays an important role in the development of human chronic diseases including cancer, cardiovascular disease, insulin resistance and obesity. Developing foods and diets that promote human health is central to public health initiatives for preventing and lowering the economic and social impact of chronic disease.1 Direct and indirect costs of cardiovascular disease (CVD) have been estimated at $445 billion in the United States2 and €200 billion within the European Union.3 Global costs of CVD in 2010 totalled US$863 billion.4 These costs are projected to increase several-fold by 2030, reaching unsustainable levels due to people living longer and the rapid increase in obesity in developed and developing countries.2,5

Following the identification of the anti-mutagenic properties of conjugated linoleic acid (CLA) isomers in cooked beef,6–8 numerous studies have investigated the biological activity of CLA isomers in cell culture and animal models (http://fri.wisc.edu/cla.php). Much of the research has focused on the effects of cis-9, trans-11 18:2 (trivial name rumenic acid) or trans-10, cis-12 18:2 due to the cost and availability in a range of mammalian and avian species. In addition to the inhibition of mutagenesis, specific isomers of CLA have been demonstrated to modulate energy metabolism, immunity, inflammation, insulin resistance and bone metabolism in several animal models.9–21 However, evidence that the physiological effects described in vitro or in other mammalian species are also replicated in humans remains inconclusive.22–28

The optimal intake of one or more isomers of CLA in humans remains to be established. Direct or exponential extrapolation of data from studies in the rat model of carcinogenesis implicate intakes of cis-9, trans-11 18:2 between 95 and 3500 mg per day being required for significant decreases in cancer risk in human populations.29,30 Estimates of cis-9, trans-11 18:2 consumption in human populations vary between 15 and 1500 mg per day29,31–39 depending on the methodology used to estimate dietary intakes, with marked differences between countries, gender and socio-economic groups. Isomers of CLA are present in a wide range of foods including milk, beef and lamb, and to a much lesser extent in pork, poultry, fish and eggs, with trace amounts in some vegetable sources.32,35–43 Milk and dairy products are the major source of CLA in the human diet contributing to between 66 and 80% of total intake.31,35–39 Typically concentrations of CLA in pork, chicken and fish are lower than 0.1 g per 100 g lipid,44 whereas the consumption of lamb, beef and other ruminant meat products account for 15–32% of average daily CLA intakes in developed countries.34–36,38,44

The CLA status of humans can be increased using oral supplements or fortification of foods with synthetic sources, which typically contain equal amounts of cis-9, trans-11 18:2 and trans-10, cis-12 18:2, or from a higher consumption of ruminant-derived foods. In contrast to synthetic sources, meat and milk from ruminants contain numerous positional and geometric isomers of CLA with conjugated double bonds at positions 6,8 through to 13,15, with cis-9, trans-11 18:2 being the major isomer, and trans-7, cis-9 18:2 or trans-11, cis-13 18:2 as the second most abundant.44–47

Producing ruminant-derived foods containing higher amounts of CLA offers the opportunity to increase the consumption of CLA, principally cis-9, trans-11 18:2, without requiring major changes in the habitual diet or eating habits. For this reason, a considerable amount of research has been dedicated to understanding the nutritional, physiological and genetic factors influencing CLA concentrations in meat and milk. The present chapter provides a comprehensive review of the most recent evidence on the biochemical, microbial, nutritional and physiological factors influencing the biosynthesis of CLA isomers in ruminants and humans.

1.2 Lipid Metabolism in the Rumen

1.2.1 Substrate Supply

Ruminant diets vary in composition depending on species, physiological state, and the cost and availability of feed ingredients. Diets often contain forage species (grasses, legumes or forage maize) of variable maturity and nutritional value, and differ in composition from those containing forages as the sole feed to combinations of forage, cereals and protein supplements. By-products of the food industry or lipid supplements may also be included. A general characteristic of ruminant diets is the relatively high fibre content (>300 g cell wall constituents per kg dry matter (DM)) and low amounts of lipid (<50 g per kg DM).45 Lipid in cereal grains, plant oils, marine lipids and by-products are predominantly in the form of triacylglycerol (TAG). Most of the lipid in grasses and legume forages is present as phospholipids (PL) and glycolipids (GL) located within thylakoid membranes of chloroplasts.48–50 In forages, GL are the major lipid class, with galactolipids (mono- and di-galactosyl diacylglyerol) being the most prevalent. These differ from TAG in that one or more carbohydrate molecules are linked to one position of the glycerol backbone. Forages and oilseeds also contain several PL species, (phosphatidylcholine, phosphatidylglycerol and phosphatidylethanolamine) as structural components of cell membranes. Most PL contain a diacylglycerol covalently bonded to a phosphate group, which is often esterified to a simple organic molecule such as choline. In general, both fatty acid moieties bound to glycerol in glycolipids or PL are unsaturated. Non-esterified fatty acids (NEFA) are minor components of most ruminant feeds, but are the major lipid class in ensiled grasses and forage legumes and in certain proprietary fat supplements. Changes arising during fermentation in silo are characterized by a substantial decrease in the relative abundance of polar membrane lipid and an increase in NEFA, TAG, diacylglycerol (DAG) and monoacylglycerol (MAG) fractions attributable to the activity of plant and microbial lipases.51–54

The amount and composition of constituent fatty acids differs substantially between ingredients in ruminant diets (Table 1.1). For grasses and legume forages fatty acid content is generally lower than 50 g per kg DM with cis-9, cis-12, cis-15 18:3 (α-linolenic acid; 18:3 n-3) as the major fatty acid (>50 g per 100 g fatty acids).55–59 However, conservation of grass by drying results in substantial decreases in lipid content,58,60,61 principally due to the disappearance of cis-9, cis-12 18:2 (linoleic acid; 18:2 n-6) and 18:3 n-3 by oxidation and leaf shatter.49 Oxidation arises from the activity of plant lipoxygenases, which catalyse the incorporation of molecular oxygen in non-esterified 18:2 n-6 and 18:3 n-3, forming 9- and 13-hydroperoxy polyunsaturated fatty acids, respectively, that are highly reactive and rapidly metabolized into a series of oxylipins including volatile leaf aldehydes and alcohols, hydroxy- and epoxy-fatty acids and jasmonates.62,63 In contrast, 18:2 n-6 is the predominant fatty acid in forage maize, whole crop silages and cereal grains.45,57,64 Ruminant diets may also contain up to around 50 g per kg DM of additional lipid in the form of oils or oilseeds. Oils from rapeseeds, high oleic sunflowerseeds, olives and peanuts are a rich source of cis-9 18:1 (oleic acid),65,66 cottonseeds, safflowerseeds, soyabeans and sunflowerseeds are abundant in 18:2 n-6,65,67 whereas linseeds65,68 and camelina69,70 contain relatively high proportions of 18:3 n-3 (Table 1.1). In contrast, coconut oil is rich in 12:0 (lauric acid), whereas 16:0 (palmitic acid) is the major fatty acid in palm oil.65 Ruminant diets may also be supplemented with fish oil containing cis-5, cis-8, cis-11, cis-14, cis-17 20:5 (eicosapentaenoic acid; 20:5 n-3) and cis-4, cis-7, cis-10, cis-13, cis-16, cis-19 22:6 (docosahexaenoic acid; 22:6 n-3)68 or marine algae enriched in 22:6 n-3.71

Table 1.1 Typical lipid content and fatty acid composition of forages, oilseeds, plant and marine lipid supplements used in ruminant diets.
Oil Fatty acid composition (g/100 g fatty acids)
Ingredient (g/kg dry matter) 16:0 18:0 cis-9 18:1 18:2n-6 18:3n-3 20:5n-3 22:6n-3 Referencea
Grass
 3 week growth 25.0 16.1 1.4 1.94 10.9 67.3 Dewhurst et al.55
 6 week growth 15.2 19.4 2.0 2.40 11.9 60.7 Dewhurst et al.55
 Early cut 33.2 15.1 2.3 4.4 18.2 49.9 Vanhatalo et al.56
 Late cut 28.2 14.5 2.2 6.5 28.8 37.6 Vanhatalo et al.56
 Wilted 20.5 19.0 0.3 1.09 3.7 18.0 Shingfield et al.58
Red clover
 Early cut 33.3 21.5 7.1 4.2 17.3 35.6 Vanhatalo et al.56
 Late cut 30.2 19.5 3.8 3.5 21.4 38.8 Vanhatalo et al.56
Silage
 Grass 19.8 20.1 2.1 2.5 14.2 50.4 Shingfield et al.57
 Maize 25.3 17.4 2.2 20.3 44.8 6.6 Shingfield et al.57
 Red clover 30.4 20.6 6.5 3.4 17.4 40.0 Vanhatalo et al.56
 Whole crop wheat 21.0 17.3 1.0 12.2 40.9 23.2 Noci et al.64
Hay
 Grass 8.1 35.0 0.6 2.6 5.4 15.6 Shingfield et al.58
 Lucerne 10.3 22.9 4.1 4.3 17.7 22.3 AbuGhazaleh et al.59
Rape
 Oil 963 6.0 2.3 48.1 27.4 10.3 Givens et al.66
 Whole seeds 408 4.8 2.0 56.8 19.3 8.3 Givens et al.66
Sunflower
 Oil 962 6.1 3.6 26.5 60.4 0.1 Shingfield et al.67
 Whole seeds 400 5.1 4.3 21.6 66.8 0.2 Woods and Fearon65
Linseed
 Oil 953 4.2 2.7 16.5 15.8 57.8 Shingfield et al.68
 Whole seeds 360 6.1 3.4 18.8 16.3 54.4 Woods and Fearon65
Camelina
 Oil 95.4 5.6 2.4 11.6 15.7 37.0 Halmemies-Beauchet-Filleau et al.70
 Whole seeds 37.8 8.3 17.8 27.6 46.3 Hurtaud and Peyraud69
Fish oil 950 15.0 2.6 11.0 1.2 0.9 16.5 10.5 Shingfield et al.68
Marine algae, Schizochytrium sp. 581 26.3 0.9 1.1 0.3 0.2 <0.1 37.8 Boeckaert et al.71
a Numbers refer to citations listed in the reference section.

1.2.2 Lipolysis

Following ingestion and mastication, the ester linkages of TAG, PL and GL are rapidly hydrolysed in the rumen.72–74 Hydrolysis of dietary TAG occurs predominantly as a result of microbial lipases.74 Forage plant tissues are also rich in endogenous galacto- and phospholipases, which remain active once ingested into the rumen for several hours, suggesting that senescence of the plant material itself in the rumen may contribute to ruminal lipolysis in grazing animals.75 Dawson et al.74 challenged this idea and concluded that microbial lipases were more important than plant enzymes. These conclusions were drawn using autoclaved grass as a substrate, which the authors noted was not ideal because of the many effects that autoclaving has in addition to enzyme denaturation. This debate has been revisited by Lee et al.,76 who reported increased NEFA and decreased polar lipid abundance after 6 h incubation of fresh ryegrass leaves in buffer, confirming that plant-catalysed lipolysis occurred. Further, Van Ranst et al.77 reported lipolysis of up to 60% after 8 h incubation of fresh red clover leaves. Both studies suggested the observed lipolysis to be due to active plant lipases that could contribute to overall ruminal lipolysis. However, until plant lipase activity is compared directly with that of ruminal micro-organisms, there will remain an uncertainty about their relative importance. Nonetheless, it may be useful to breed forage plants low in lipase activity in order to increase the amount of polyunsaturated fatty acids escaping the rumen. Plant lipids may also be compartmentalized in the ingested plant material, effectively protecting them from both endogenous and microbial hydrolysis.78

Among the various types of ruminal micro-organisms, the bacteria are considered to be most active in lipolysis.48 The most active bacterial species isolated selectively using TAG as a substrate was Anaerovibrio lipolytica,79 with which most research has been carried out. More recently, Jarvis et al.80 isolated two bacteria from red deer that hydrolysed TAG and grew on glycerol. The bacteria, which were highly active against tallow, tripalmitin and olive oil, were most closely related to the genus Propionibacterium and clostridial cluster XIVa. Cirne et al.81 isolated Clostridium lundense sp. nov. from the bovine rumen. Clostridium lundense exhibited lipolytic activity against olive oil, but neither its activity nor its numbers was reported, so it is difficult to assess its likely importance. Thus, a wider range of bacteria than is usually considered may be involved in the lipolysis of TAG in the rumen.

A. lipolyticus lacks the ability to hydrolyse galactolipids and PL and, therefore, other lipolytic species would be expected to predominate in grazing ruminants. The Butyrivibrio spp. appeared to contain all the phospholipase A, phospholipase C, lysophospholipase and phosphodiesterase activities typical of the mixed rumen contents.48 Their lipase activity against TAG varies between different Butyrivibrio and Pseudobutyrivibrio strains, but not in a manner that corresponds to their position in the phylogenetic tree.82 Little is known about how other lipases vary across different strains/species, nor whether other recently recognized species may possess such activities in the rumen of grazing ruminants.

There have been few recent studies that investigate protozoal lipolysis. Wright83 suggested Epidinium spp. to be responsible for 30–40% of the lipolytic activity in the rumen. Epidinium ecaudatum was reported to liberate galactose from galactolipids, suggesting galactosidase activity, although lipase activity per se was not demonstrated.84 Another protozoal species, Entodinium caudatum, was shown to have phospholipase activity,85 but it is possible that this activity was more relevant to the intracellular metabolism of the protozoa than to the digestion of dietary lipids. The earlier studies to determine the contribution of protozoa to the lipolytic activity in the rumen were conducted using fractionated rumen fluid, with the possibility that lipolytic activity in protozoal fractions was more due to the activity of bacteria that the protozoa had ingested than that of the protozoa themselves. Once again, given that protozoa comprise up to half the microbial biomass present in the rumen, their lipolytic properties warrants further investigation.

Much is known about bacterial lipases in general. They comprise eight well-documented families and their modes of action are reasonably well characterized.86 The lipase activity of A. lipolyticus was investigated in some detail by methods available at the time, some four decades ago.87,88 Perhaps surprisingly, no cloning and sequencing studies seem to have been done when the technology became widely available, and the most detailed recent study89 was derived from genomic analysis. Three enzymes from A. lipolyticus were identified from genomic analysis of A. lipolyticus. The alipA, alipB and alipC encoded 492-, 438- and 248-amino acid peptides, respectively. Phylogenetic analysis indicated that alipA and alipB clustered with the GDSL/SGNH family II, and alipC clustered with lipolytic enzymes from family V. Subsequent expression and purification of the enzymes showed that they had esterase activities with substrate specificities favouring the hydrolysis of caprylate-, laurate- and myristate-containing substrates. Genomes are available for several species in the Butyrivibrio group, but to date no similar analysis appears to have been carried out for their lipase activities.

Metagenomic methods will be invaluable in order to understand the full complement of lipolytic enzymes that are present in the rumen. Expression libraries may be useful. Liu et al.90 screened a metagenomic library from the rumen of grazing Holstein cows for lipase activity, using trioleoylglycerol as substrate. Out of 15 360 bacterial artificial chromosome (BAC) clones, only two were found to have high lipase activity, which seems surprisingly small. The likely origin of the genes was investigated, based on the other open reading frames (ORFs) present in the BAC clones. It was impossible to decipher the host for the first gene, Rlip1, but the second, Rlip2, gave most similar homologues from Thermosinus carboxydivorans, which has not previously been associated with the ruminal ecosystem. Nonetheless, by BLASTing their deposited lipase sequences it was discovered that the genes most likely encoded carboxyl esterases. Thus the prevalence of these lipases in the overall community is far from certain. Clearly many more lipase sequences must be analysed from the metagenome, and assignment to lipases assured, in order to understand the true nature of lipolytic enzymes active in the rumen.

Lipolysis is considered rate limiting for the biohydrogenation of dietary unsaturated fatty acids in the rumen.48 During ruminal digestion of grasses, mono- and digalactosyl diacylglycerides are released following the rupture of chloroplasts.73 Early studies involving incubations of 14C-labelled substrate with strained rumen fluid indicated that lipolysis conforms to first-order kinetics, occurring at a high rate with a short lag time.91 These observations led Hawke and Silcock91 to conclude that hydrolysis of dietary acyl lipids in the rumen would be sufficiently rapid not to impede biohydrogenation. However, such conclusions were not supported by observations that the profile of products formed during intra-ruminal infusion of esterified or non-esterified 18:2 n-6 in sheep differed.92,93

Much of what is known about the factors influencing ruminal lipolysis is based on incubations of various substrates with rumen fluid. Laboratory-scale experiments have obvious drawbacks with respect to mimicking conditions in vivo. Accepting these limitations and the assumptions therein, a number of factors have been determined to influence the rate and extent of lipolysis in a simulated rumen environment (Figure 1.1). Increases in dietary nitrogen content were found to increase the rate at which triolein was hydrolysed,94 whereas the rate of lipolysis was decreased by replacing fibre with starch95 or when more mature forages were incubated.96 As could be expected, lipolysis increases with incubation time,97,98 but is often decreased at low rumen pH.99,100 The extent of lipolysis is also decreased for lipids with a higher melting point containing relatively high proportions of saturated fatty acids.45 Increasing the amount of substrate incubated also lowers progressively the rate and extent of lipolysis in vitro.97 After 24 h incubations with buffered rumen fluid, between 73.7% and 89.5% of TAG in fish oil, linseed oil or sunflower oil was found to be hydrolysed.98 Similarly, lipolysis of lipid in cocksfoot and red clover after 24 h incubations were reported to vary between 65.0% and 82.0%.101,102

Fig. 1.1 Schematic of the main transformation of dietary lipids in the rumen and factors influencing the relative rates of lipolysis, isomerization and biohydrogenation (adapted from Doreau et al.)353 Following ingestion and mastication, dietary lipids are extensively hydrolysed in the rumen liberating non-esterified unsaturated fatty acids that are sequentially reduced to saturated end-products. Metabolism of unsaturated fatty acids proceeds via cis to trans isomerization, reduction or hydration of double bonds. Biohydrogenation to saturated end products is incomplete and numerous intermediates accumulate and escape the rumen.

Measurements of lipolysis in vivo are scarce. Studies in cattle,54,61 goats103 and sheep104 all affirm that a small fraction of esterified dietary lipid escapes the rumen. In lactating cows fed diets based on grass silage or red clover silage, NEFA, PL, TAG, DAG and MAG fractions were found to account for 80%, 12%, 4.4%, 2.4% and 0.8% of total fatty acids in omasal digesta.54 The majority of 18:3 n-3 entered the omasum in the form of polar lipid (42–46%) originating from plant PL and GL, with smaller amounts as NEFA (22–26%), TAG (17–20%), DAG (8–14%) or MAG (2–3%). Between 85% and 93% of esterified lipid was reported to be hydrolysed in the rumen of lactating cows fed diets based on fresh grass or grass silage (forage:concentrate ratio 60:40), the extent of which was decreased to 80–86% and 70% when grass silage was replaced with grass hay61 or red clover silage,54 respectively. While incomplete, ruminal lipolysis is extensive, resulting in NEFA accounting for more than 80% of the total amount of fatty acids leaving the rumen.48,54,61,103,104

1.2.3 Biohydrogenation

From a historical perspective, the existence and importance of ruminal biohydrogenation have been known for more than 50 years. It has only been in recent years following the discovery of the biological activities of CLA isomers that research on ruminal lipid metabolism has focused more on the biochemical mechanisms involved in the formation of the less abundant fatty acid intermediates. Advances in lipid analysis allowed the marked differences in the fatty acid profile of lipids in the diet (principally cis-9 18:1, 18:2 n-6 and 18:3 n-3) and that of lipids leaving the rumen (mainly 16:0 and 18:0) to be established.,105 These differences were discovered to be the consequence of the process we now call biohydrogenation, which converts unsaturated fatty acids to saturated fatty acids via firstly a cis-trans isomerization to trans fatty acid intermediates followed by hydrogenation of the double bonds (Figure 1.1).48,106,107 Little if any oxidation or elongation of the carbon chain is thought to occur.48

Early evidence of ruminal biohydrogenation was obtained when linseed oil was incubated with sheep ruminal contents.108 The 18:3 n-3 content of the linseed oil decreased from 30% to 5% with an accompanying increase in the concentration of 18:2. Shorland et al.105 also noted that the ruminal metabolism of 18:3 n-3 caused the accumulation of 18:2 and 18:1 intermediates, which were then converted to 18:0. The requirement for a free carboxyl group for hydrogenation established that lipolysis must precede biohydrogenation.91 After these initial studies demonstrated the overall features of biohydrogenation in ruminal contents, various in vivo and in vitro experiments were carried out to examine the pathways involved in ruminal digesta. Among these, the work of Dawson and his co-workers was particularly informative. Wilde and Dawson109 constructed a metabolic scheme for the metabolism of 18:3 n-3 to 18:0 based on incubations of sheep ruminal contents with U-14C labelled 18:3 n-3. The initial step was the isomerization of the cis-12 bond to either the Δ11 or Δ13 position. Thereafter, one of the bonds was hydrogenated to leave an 18:2, followed by hydrogenation of another bond producing an 18:1 intermediate. Hydrogenation of the 18:1 intermediate yielded 18:0 as the final product.

The other early contribution to delineating biohydrogenation pathways was the seminal work of Tove and his group with the bacterial species, Butyrivibrio fibrisolvens, that had been identified by Polan et al.110 to possess high biohydrogenating activity. When incubated with B. fibrisolvens, 18:2 n-6 was initially isomerized to a conjugated 18:2, thought to be mainly cis-9, trans-11 18:2, but other dienoic isomers were also detected.111The cis-9, trans-11 18:2 intermediate was then hydrogenated to trans-11 18:1 (vaccenic acid) as the final product. However, in mixed ruminal bacteria, the 18:1 intermediate was further hydrogenated to 18:0.112Kepler and Tove113 also incubated B. fibrisolvens with 18:2 n-6 and 18:3 n-3. They confirmed that 18:2 n-6 was first isomerized to cis-9, trans-11 18:2 followed by further hydrogenation to a mixture of trans 18:1 isomers. When 18:3 n-3 was used as a substrate, it was first isomerized to cis-9, trans-11, cis-15 18:3 (conjugated linolenic acid). This product was transient and further hydrogenated to a non-conjugated 18:2 intermediate as the final product. Slightly later, bacteria were isolated that converted monoenoic acids to 18:0 and hydrogenated 18:2 n-6 to 18:0, with traces of the intermediates that had been seen with B. fibrisolvens and mixed ruminal microbes.114,115 The first step of 18:3 n-3 biohydrogenation is analogous to that of 18:2 n-6, namely the formation of a conjugated bond structure, cis-9, trans-11, cis-15 18:3.

Biohydrogenation of cis-6, cis-9, cis-12 18:3 (γ-linolenic acid, 18:3 n-6) during incubations with pure strains of Butyrivibrio and a bacterium identified as Fusocillus babrahamensis has also been investigated.116 The group B bacterium (Fusocillus) hydrogenated 18:3 n-6 to 18:0, whereas incubations with the group A bacterium (Butyrivibrio) yielded cis-6, trans-11 18:2 as an end product. By analogy with the metabolic pathway described for 18:3 n-3, biohydrogenation of 18:3 n-6 is thought to proceed via isomerization of the cis-12 double bond to yield cis-6, cis-9, trans-11 18:3, that is sequentially reduced to cis-6, trans-11 18:2, trans-11 18:1 and 18:0.116 Most feeds fed to ruminants do not contain 18:3 n-6. However, certain oilseeds including evening primrose and borage are relatively abundant in 18:3 n-6, but the effects of these lipids on ruminal lipolysis and biohydrogenation in vivo have not been investigated.

Several recent studies have investigated the metabolism of cis-6, cis-9, cis-12, cis-15 18:4 (stearidonic acid; 18:4 n-3) during incubations with rumen fluid.117,118 Numerous intermediates were found to be formed. Based on the profile of fatty acids detected, the metabolism of 18:4 n-3 was proposed to proceed via the formation of 5,7,11,15 18:4 or 5,8,10,15 18:4.118 Conjugated 18:4 products were reduced to yield 5,11,15 18:3 or 5,10,15 18:3 and hydrogenated yet further to 11,15 18:2 and 5,10 18:2, respectively.118Trans-6 to -12 18:1 were found to accumulate with trans-11 18:1 being the major intermediate formed.117

Much less is known about the fate of longer chain polyenoic fatty acids in the rumen. Detailed analysis of ruminal119,120 and omasal digesta121,122 have indicated that numerous 20- and 22-carbon biohydrogenation intermediates containing one or more trans double bonds are formed during the biohydrogenation of fatty acids in fish oil or marine algae. However, the biochemical pathways responsible are not known. The most recent investigations suggest that the first committed steps of 20:5 n-3 and 22:6 n-3 biohydrogenation in the rumen involve the reduction and/or isomerization of double bonds closest to the carboxyl group.121,122 No conjugated ≥20-carbon fatty acids have been detected.

1.2.4 Metabolic Pathways

Ruminal biohydrogenation of cis-9 18:1, 18:2 n-6 and 18:3 n-3 in vivo varies from 58–87%, 70–95% and 85–100%, respectively.107,123,124 The extent of ruminal fatty acid biohydrogenation is influenced by the composition and amount of lipid in the diet, retention time in the rumen and the characteristics of the microbial population. Biohydrogenation is extensive, with 18:0 being the major fatty acid leaving the rumen on most diets. However, the reduction of unsaturated 18-carbon fatty acids to 18:0 in the rumen is incomplete and numerous 18:1, 18:2 and 18:3 intermediates accumulate. The final reduction of trans 18:1 is thought to be the rate limiting step in the complete biohydrogenation of 18-carbon unsaturated fatty acids.125 Decreases in pH from 6.5 to below 6.0 lowers the extent of cis-9 18:1,126,127 18:2 n-6127–131 and 18:3 n-3127,129,132 biohydrogenation and inhibits the final reduction of trans 18:1 to 18:0127,132in vitro. The effects of pH below normal physiological ranges in the rumen may be related to disruption of bacterial cell membranes resulting in the inactivation of membrane bound isomerases and reductases.113

Characterizing the biohydrogenation process in vivo represents a major challenge. Relatively few studies have reported sufficiently detailed measurements of post-ruminal fatty acid outflows to allow for inferences on metabolic pathways. The situation is further complicated due to oils and oilseeds being used to alter substrate supply rather than pure fatty acids. In this regard incubations of fatty acids with rumen fluid or pure strains of ruminal bacteria have proven invaluable in establishing the profile of possible intermediates formed during the biohydrogenation of unsaturated fatty acids (Table 1.2). However, intermediates formed in vitro may not necessarily parallel biohydrogenation pathways in vivo. Discrepancies may arise from the incubation of excessively high amounts of substrate, methods used to introduce fatty acids into batch cultures, source of ruminal fluid and potential loss of microbial activity over an extended incubation period.107

Table 1.2 Summary of intermediates formed during incubations of 18-carbon unsaturated fatty acids with rumen contents or pure cultures of rumen bacteria.
Substrate Inoculum/Bacterium Intermediates and end-products Referencea
cis-9 18:1 B. proteoclasticus 18:0 McKain et al.134
cis-9 18:1 E. faecalis 10-OH 18:0 Hudson et al.136
cis-9 18:1 P. acnes 10-OH 18:0, 10-0 18:0 McKain et al.134
cis-9 18:1 S. ruminantium 10-OH 18:0 Hudson et al.136
cis-9 18:1 Bovine rumen contents trans-6, -7, -9, -10, -11, -12, -13, -14, -15, -16 18:1, 18:0 Mosley et al.138
cis-9 18:1 Bovine rumen contents trans-6, -7, -9, -10, -11, -12, -13, -14, -15, -16 18:1, 18:0 AbuGhazaleh et al.126
cis-9 18:1 Bovine rumen fluid 10-OH 18:0, 10-0 18:0, 18:0 Jenkins et al.133
trans-9 18:1 Bovine rumen fluid cis-9, -11 18:1, trans-6, -7, -11 18:1, 18:0 Proell et al.139
trans-10 18:1 B. proteoclasticus 18:0 McKain et al.134
trans-10 18:1 P. acnes 10-OH 18:0, 10-0 18:0 McKain et al.134
trans-11 18 :1 B. proteoclasticus 18:0 McKain et al.134
cis-9, cis-12 18:2 B. fibrisolvens trans-11 18:1 Maia et al.207
cis-9, cis-12 18:2 B. fibrisolvens cis-9, cis-11 18:2, cis-9, trans-11 18:2, trans-9, cis-11 18:2, trans-9, trans-11 18:2 Wallace et al.142
cis-9, cis-12 18:2 B. hungatei trans-11 18:1 Maia et al.207
cis-9, cis-12 18:2 B. proteoclasticus cis-9, trans-11 18:2, trans-11 18:1 Maia et al.207
cis-9, cis-12 18:2 B. proteoclasticus cis-9, cis-11 18:2, cis-9, trans-11 18:2, trans-9, cis-11 18:2, trans-9, trans-11 18:2 Wallace et al.142
cis-9, cis-12 18:2 C. aminophilum cis-9 18:1 Maia et al.207
cis-9, cis-12 18:2 E. faecalis 10-OH 18:1, 13-OH 18:1 Hudson et al.144
cis-9, cis-12 18:2 F. succinogenes 16:0 Maia et al.207
cis-9, cis-12 18:2 M. multiacidus cis-9 18:1 Maia et al.207
cis-9, cis-12 18:2 P. acnes cis-10, trans-12 18:2, trans-10, cis-12 18:2, trans-10, trans-12 18:2 Wallace et al.142
cis-9, cis-12 18:2 S. bovis 13-OH 18:1 Hudson et al.144
cis-9, cis-12 18:2 Bovine rumen fluid cis-6, cis-12 18:2, cis-7, cis-12 18:2, cis-8, cis-12 18:2, cis-9, cis-11 18:2, cis-10, cis-12 18:2, cis-9, trans-11 18:2, cis-9, trans-12 18:2, trans-8, cis-10 18:2, trans-8, cis-12 18:2, trans-9, cis-12 18:2, trans-10, cis-12 18:2, trans-9, trans-11 18:2, trans-10, trans-12 18:2, trans-9, trans-12 18:2, trans-6-8, -9, -10, -11, -12, -13-14 18:1, cis-9, -11, -12 18:1, 18:0 Honkanen et al.141
cis-9, cis-12 18:2 Ovine rumen fluid cis-9, cis-11 18:2, cis-9, trans-11 18:2, trans-10, cis-12 18:2, trans-11 18:1 Waşowska et al.146
cis-9, cis-12 18:2 Ovine rumen fluid cis-10, cis-12 18:2, cis-9, trans-11 18:2, cis-9, trans-12 18:2, trans-10, cis-12 18:2, trans-9, trans-11 18:2, trans 4, 5, 6-8, -9, -10, -12 18:1, cis-10, -12, -13 18:1, 18:0 Jouany et al.140
cis-9, cis-12 18:2 Ovine rumen fluid cis-9, cis-11 18:2, cis-10, cis-12 18:2, cis-9, trans-11 18:2, trans-9, cis-11 18:2, trans-10, cis-12 18:2, trans-9, trans-11 18:2 Wallace et al.142
cis-9, trans-11 18:2 B. fibrisolvens trans-11 18:1 McKain et al.134
trans-10, cis-12 18:2 B. fibrisolvens trans-10, -12 18:1, cis-12 18:1 McKain et al.134
trans-9, trans-11 18:2 B. fibrisolvens trans-11 18:1 McKain et al.134
trans-9, trans-11 18:2 B. proteoclasticus trans-9, -11 18:1, cis-11 18:1 McKain et al.134
cis-9, cis-12, cis-15 18:3 Ovine rumen fluid cis-9, trans-11, cis-15 18:3, trans-9, trans-11, cis-15 18:3 trans-11, cis-15 18:2, Waşowska et al.146
cis-9, cis-12, cis-15 18:3 Ovine rumen fluid cis-9, cis-11 18:2, cis-9, cis-15 18:2, cis-9, trans-13 18:2, cis-11, trans-13 18:2, trans-9, cis-12 18:2, trans-11, cis-15 18:2, trans-9, trans-12 18:2, trans-11, trans-13 18:2, trans 6-8,-9, -11, -12, -13-14, -15, -16 18:1, cis-13, -15 18:1, 18:0 Jouany et al.140
cis-9, cis-12, cis-15 18:3 Bovine rumen contentsb cis-9, cis-11 18:2, cis-10, cis-12 18:2, cis-9, trans-11 18:2, trans-9, cis-11 18:2, trans-10, cis-12 18:2, trans-8, trans-10 18:2, trans-9, trans-11 18:2, trans-11, trans-13 18:2 Lee and Thomas148
cis-6, cis-9, cis-12, cis-15 18:4 trans-6-8,-9, -10, -11, -12 18:1, cis-9, -11 18:1, 18:0 Maia et al.117
cis-6, cis-9, cis-12, cis-15 18:4 5,7,11,15 18:4, 5,8,10,15 18:4, 5,10,15 18:3, 5,11,14 18:3, 5,11,15 18:3, trans-5, trans-10 18:2, trans-5, trans-11 18:2 Alves et al.118
a Numbers refer to citations listed in the reference section. b Following 48 h incubations of U-13C cis-9, cis-12, cis-15 18:3 significant 13C enrichment was detected in numerous intermediates including conjugated 18:3 (n=2), non-methylene interupted 18:3 (n=12) and non-methylene-interupted 18:2 (n=5) isomers.

Early studies demonstrated that mixed or pure cultures of ruminal bacteria were capable of hydrogenating cis-9 18:1 to 18:0.48 More recent experiments have indicated that cis-9 18:1 is also hydrated to 10-OH 18:0 (10-hydroxystearic acid), an intermediate that is further oxidized to 10-O 18:0 (10-ketostearic acid) during incubations with ruminal contents.133,134 Hydration of the cis-9 double bond involves the incorporation of one hydrogen atom from water.134 A select number of bacteria isolated from the rumen, including species identified as Fusocillus babrahamensis,135Selenomonas ruminantium,136Enterococcus faecalis,136Streptococcus bovis137 and Propionibacterium acnes134 are capable of hydrating cis-9 18:1. P. acnes is known to catalyse the oxidation of 10-OH 18:0 to 10-O 18:0.134 Incubations of 1-13C cis-9 18:1 with rumen contents have indicated that cis-to trans isomerization of the Δ9 double bond also takes place, resulting in the formation of trans-6, -7, -9, -10, -11, -12, -13, -14, -15, and -16 18:1 intermediates.138 Further investigations of the metabolism of 1-13C cis-9 18:1 by mixed rumen microbes grown in continuous culture at pH 5.5 or pH 6.5 and at different dilution rates indicated that decreases in pH and dilution rate prevented the formation of trans 18:1 intermediates with a double bond beyond the Δ10 position.126 There is also evidence to suggest that trans to cis isomerization may occur, following reports that incubations of trans-9 18:1 with mixed ruminal bacteria resulted in the production of cis-9 18:1 and cis-11 18:1.139 Incubations with pure or mixed cultures of ruminal bacteria have allowed several biochemical pathways of cis-9 18:1 metabolism to be characterized (Figure 1.2).

Fig. 1.2 Pathways of cis-9 18:1 metabolism in the rumen. Formation of intermediates and end products illustrated is based on studies involving the incubation of labelled and unlabelled cis-9 18:1 with mixed and pure cultures of rumen bacteria. Arrows with solid lines highlight the major biohydrogenation pathway while arrows with dashed lines describe the formation of minor fatty acid metabolites.

For many years isomerization of the cis-12 double bond leading to the formation of cis-9, trans-11 CLA has been considered the first committed step of 18:2 n-6 biohydrogenation in the rumen, followed by reduction to trans-11 18:1 and 18:0 (Figure 1.3). More recent studies have shown that geometric isomers of both 9,11 and 10,12 18:2 are formed during incubations of 18:2 n-6 with rumen fluid127,132,140,141 and pure cultures of several ruminal bacteria.142 The amounts of 9,11 and 10,12 CLA isomers formed during the isomerization of 18:2 n-6 also appear to be pH-dependent, but reports are conflicting. Some investigations demonstrated that decreases in pH below 6.0 lower the accumulation of cis-9, trans-11 18:2 and trans-10, cis-12 18:2,127,129 whereas increased formation was reported in others.130,131 Conjugated 18:2 products formed during the initial isomerization of 18:2 n-6 are transient and reduced to yield both trans (-4, -5, 6-8, -9, -10, -11, -12, -13 and -14) and cis (-9, -10, -11, -12 and -13) 18:1 intermediates134,140,141(Table 1.2). Decreases in pH from 6.4 to 5.6 have been shown to promote trans-10 18:1 formation, changes accompanied by a decrease in the accumulation of trans-11 18:1 accumulation.127 Increases in the amounts of 18:2 n-6 incubated have also been shown to increase the ratio of trans-10 18:1/trans-11 18:1.141,143

Fig. 1.3 Pathways of cis-9, cis-12 18:2 metabolism in the rumen. Formation of intermediates and end products illustrated is based on studies involving the incubation of labelled and unlabelled cis-9, cis-12 18:2 with mixed and pure cultures of rumen bacteria. Arrows with solid lines highlight the major biohydrogenation pathway while arrows with dashed lines describe the formation of minor fatty acid metabolites.

Incubations with strains of Enterococcus faecalis isolated from the rumen have demonstrated that 18:2 n-6 may also be hydrated to yield 10-OH, cis-12 18:1 and cis-9, 13-OH 18:1.144 Recent investigations provided the first evidence that biohydrogenation of 18:2 n-6 may also proceed by a mechanism involving the migration of the cis-9 double bond rather than hydrogen abstraction or isomerization of the cis-12 double bond resulting in the formation of cis-6, cis-12 18:2, cis-7, cis-12 18:2 and cis-8, cis-12 18:2.141

Owing to the higher number of double bonds the metabolic pathways responsible for the biohydrogenation of 18:3 n-3 in the rumen are far more complex than for cis-9 18:1 or 18:2 n-6. Most investigations indicate the metabolism of 18:3 n-3 in the rumen involves the formation of a single 18:3 intermediate cis-9, trans-11, cis-15 18:3, that is sequentially reduced to trans-11, cis-15 18:2, followed by trans-11 18:1 with 18:0 as an end product48,109,113,145 (Figure 1.4).

Fig. 1.4 Pathways of cis-9, cis-12, cis-15 18:3 metabolism in the rumen. Formation of intermediates and end products illustrated is based on studies involving the incubation of labelled and unlabelled cis-9, cis-12, cis-15 18:3 with mixed and pure cultures of rumen bacteria. Arrows with solid lines highlight the major biohydrogenation pathway, while arrows with dashed lines describe the formation of minor fatty acid metabolites.

Further studies have demonstrated that a diverse range of intermediates are formed during incubations of 18:3 n-3 with ruminal bacteria. In addition to cis-9, trans-11, cis-15 18:3, trans-9, trans-11, cis-15 18:3 is also produced during the initial isomerization of 18:3 n-3.146 Biohydrogenation of 18:3 n-3 has been reported to result in the accumulation of numerous conjugated 18:2 (cis-9, cis-11 18:2, cis-11, trans-13 18:2 and trans-11, trans-13 18:2), non-methylene-interrupted 18:2 (cis-9, cis-15 18:2, cis-9, trans-13 18:2, trans-9, cis-12 18:2, trans-11, cis-15 18:2, trans-9, trans-12 18:2), trans 18:1 (6-8,-9, -11, -12, -13-14, -15 and -16) and cis 18:1 (-13 and -15) intermediates.140 Two strains of B. fibrisolvens have also been identified as capable of converting trans-11,cis-15 18:2 to trans-11, cis-13 18:2.147 The complexity of the reactions responsible for the biohydrogenation of 18:3 n-3 was reinforced following reports that 14 positional and geometric isomers of 18:3 isomers were formed during incubations of U-13C 18:3 n-3 with ruminal contents.148 After 48 h of incubation, 13C enrichment was detected in trans-8, trans-10 18:2, trans-11, trans-13 18:2 and geometric isomers of 9, 11 and 10, 12 18:2. This is the first report that cis-9, trans-11 18:2 may also be formed from 18:3 n-3. In addition to studies in vitro, feeding diets enriched in 18:3 n-3 have been shown to increase cis-9, trans-13, cis-15 18:3, cis-9, trans-11, trans-13 18:3, trans-9, trans-11 18:2, trans-13, trans-15 18:2, 11,13 18:2 and 12,14 18:2 concentrations in bovine milk.149–151 One or more of these fatty acids may originate from the rumen. While the major pathway describing the biohydrogenation of 18:3 n-3 in the rumen is well established, those responsible for the formation of minor intermediates remain incomplete and require further investigation.

1.3 Synthesis of CLA in the Rumen

1.3.1 Mechanisms of CLA Biosynthesis

We are indebted to the early work of Tove and his colleagues for establishing many of the properties of the first step in the metabolism of 18:2 n-6 and 18:3 n-3.113,152,153 Linoleate isomerase (EC 5.2.1.5) is described as an enzyme that catalyses the conversion of 18:2 n-6 to other positional and geometric 18:2 isomers, including conjugated intermediates. The properties of this enzyme activity vary according to the host bacterial species and the product that is formed. Kepler and Tove113 carried out a partial purification of linoleate isomerase activity from B. fibrisolvens and developed a convenient assay method based on the UV absorbance of the conjugated double bond system at 233 nm. The particulate cell-free preparation formed mainly cis-9, trans-11-18:2 from 18:2 n-6; the equilibrium was strongly in favour of cis-9,trans-11 18:2 formation. Perhaps significantly when it comes to understanding the mechanism, the authors failed to isolate a homogeneous preparation that was active. Furthermore, the reaction was short-lived in reaching an endpoint, and the reaction could be restored by the addition of more enzyme, suggesting that a cofactor was required. Isomerization of 18:3 n-3 occurred in a similar manner, but at a faster rate, to cis-9, trans-11, cis-15 18:3. Further refinement of the substrate specificity and mechanism of action followed,152,153 but little further work was done for the following three decades.

In contrast, cloning, crystallization and structural analysis of the linoleate isomerase catalysing the formation of trans-10, cis-12 18:by non-ruminal P. acnes revealed at the atomic level, with a single protein, a mode of action that involves hydride abstraction by enzyme-bound FAD.154,155 Isotope studies with a ruminal P. acnes were consistent with this mode of action.142

In non-ruminal lactic acid bacteria, which produce both isomers of CLA, it has been proposed that cis-9, trans-11 18:2 formation from 18:2 n-6 by lactic acid bacteria involves a hydration-dehydration mechanism via a 10-hydroxy,cis-12 18:1 intermediate.156 Such a mechanism has been eliminated as a possible route of cis-9, trans-11 18:2 synthesis from 18:2 n-6 by B. fibrisolvens, since 10-hydroxy, cis-12 18:1 was not converted to cis-9, trans-11 18:2.142 The Lactobacillus linoleate isomerase seems, like the B. fibrisolvens enzyme, to be a multi-component enzyme,157 one component derived from the membrane fraction and another from the soluble cell content.

Two publications have claimed to have identified a gene encoding a linoleate isomerase that forms cis-9, trans-11 18:2. Park et al.158 published a sequence purporting to be a linoleate isomerase from B. fibrisolvens A-38. The gene sequence of linoleate isomerase in non-ruminal lactic acid bacteria has been published within a patent.159 Our own in silico analysis suggests that both claims may be mistaken. Furthermore, BLASTing the sequences against the recently available genomes of human intestinal species that produce cis-9, trans-11 18:2, including B. fibrisolvens, Roseburia inulinivorans and R. hominis (the last two are cis-9, trans-11 18:2 producers from the human intestine), failed to identify credible isomerase genes. In other preliminary studies, we have attempted to use published data from studies of the CLA reductase of B. fibrisolvens to identify neighbouring genes as possible candidates for linoleate isomerase, based on the assumption that CLA reductase and linoleate isomerase might be coordinately expressed. Fukuda et al.160 isolated a mutant of B. fibrisolvens that formed exceptionally high concentrations of cis-9, trans-11 18:2 from 18:2 n-6. This resulted from the loss of the CLA reductase activity.161 The mutant gene was identified, sequenced and expressed in E. coli, reportedly producing active linoleate isomerase. However, when we and others (G.T. Attwood, personal communication) BLASTed the deposited sequence against protein and nucleotide databases, a high similarity to oxaloacetate decarboxylase was revealed. Indeed, when we investigated the genomes of the previously described human bacteria, greater than 80% identity was obtained with a gene annotated as an oxaloacetate decarboxylase alpha subunit OadA. Clustering with this gene, rather than finding an ORF of unknown function, we found genes encoding transport and other genes relating to oxaloacetate metabolism. Oxaloacetate decarboxylase is very well characterized. Once again, therefore, the published sequence seems to be in error.

1.3.2 Microbiology of CLA Biosynthesis

There is little doubt that bacteria are the main organisms responsible for biohydrogenation in the rumen, but equally there is no debate that ruminal fungi and particularly protozoa have roles in determining the amounts of CLA and trans 18:1 isomers leaving the rumen.106

1.3.2.1 Role of Bacteria

Polan et al.162 identified B. fibrisolvens as a bacterium that rapidly hydrogenates 18:2 n-6, forming cis-9, trans-11 18:2 and trans-11 18:1 as intermediates. Biohydrogenation of B. fibrisolvens did not result in the formation of 18:0. Subsequent studies115,163 identified other bacteria that were capable of biohydrogenation, but the results did not provide much information about relative activities. The method used radio-labelled fatty acids as substrates and was therefore highly sensitive, but the concentrations of labelled acids used as substrates were very low (2 μg ml-1), making comparisons of specific activity and relative contribution of different species difficult. Even minor activity could have led to a positive result. Among the bacteria capable of forming 18:0, two were identified as ‘Fusocillus’ spp.115Fusocillus is not a genus that has endured in bacterial taxonomy, nor did any of the cultures survive into the DNA-sequencing era. Van de Vossenberg and Joblin164 isolated a bacterium from a cow at pasture that could also form 18:0 from 18:2 n-6. It was phenotypically similar to ‘Fusocillus' and 16S rRNA analysis indicated that it was phylogenetically close to Butyrivibrio hungatei. Subsequently, a species named Clostridium proteoclasticum was identified as a 18:0 producer with morphological and metabolic properties that were indistinguishable from those reported for Fusocillus.165 Moon et al.166 reclassified C. proteoclasticum as Butyrivibrio proteoclasticus from its 16S rRNA gene sequence. The bacteria involved in the different steps of the biohydrogenation process were classified for many years as group A and B.48 Group A bacteria hydrogenated 18:2 n-6 and 18:3 n-3 to trans-11 18:1, whereas group B bacteria converted both fatty acids to 18:0. It is now more appropriate to describe the bacteria based on their correct taxonomy (Figure 1.5).

Fig. 1.5 Phylogenetic tree of the Butyrivibrio and some key phenotypic properties. Abbreviations: CLA, conjugated linoleic acid (cis-9, trans-11 18:2); LA, linoleic acid (cis-9, cis-12 18:2); VA, vaccenic acid (trans-11 18:1).

Phenotypically, the B. hungatei and B. proteoclasticus groups are much more sensitive to the toxic effects of unsaturated fatty acids than the rest of the Butyrivibrio/Pseudobutyrivibrio cluster, such that their isolation from media containing unsaturated fatty acids is made much more difficult.82,165 They can also be distinguished phenotypically and genetically based on the mechanism by which they form 4:0 (butyrate), their second most abundant fermentation product after 2:0 (acetate). B. hungatei and B. proteoclasticus had a butyrate kinase activity >600 U per mg protein, while the others had much lower activity82 (Figure 1.5). The butyrate kinase gene was present in B. hungatei and B. proteoclasticus but not in the other group. It has been suggested that the different sensitivities to the toxic effects of CLA isomers and trans-11 18:1 may be linked to the enzyme mechanism by which butyrate is produced. Intracellular acyl-CoA concentrations, principally the precursors of 4:0, in B. fibrisolvens are depleted when the bacterium is exposed to 18:2 n-6.167

Metabolism of 18:2 n-6 by Butyrivibrio results in the formation of cis-9, trans-11 18:2, with smaller amounts of trans-9, trans-11 18:2, and trans-11 18:1, but no trans-10,cis-12 18:2 or trans-10 18:1 is formed.134 The bacteria responsible for trans-10 18:1 must therefore be different to the Butyrivibrio spp. The formation of trans-10, cis-12 18:2 also occurs by a different enzymic mechanism to that of cis-9, trans-11 18:2.142 Enrichment cultures with starch were reported to isomerize 18:2 n-6 to trans-10, cis-12 18:2,168 but these also contained abundant large cocci identified as Megasphaera elsdenii. Authors concluded that M. elsdenii as capable of trans-10, cis-12 CLA production in the rumen. Subsequent studies indicated that P. acnes could be responsible for the formation of trans-10, cis-12 18:2.165 Wallace et al.165 found no lineate isomerase activity in any pure culture of M. elsdenii. Furthermore, when digesta samples from cows producing high amounts of trans-10, cis-12 18:2 were analysed for M. elsdenii by qPCR of 16S rRNA genes, numbers were <103 g-1, while much larger numbers of P. acnes were detectable (R.J. Wallace and S. Muetzel, unpublished observation). P. acnes does not, however, convert trans-10, cis-12 18:2 to trans-10 18:1.134Butyrivibrio spp. are capable of reducing trans-10, cis-12 18:2 and trans-10, trans-12 18:2.134 However, it remains difficult to understand how Butyrivibrio would catalyse the reduction of trans-10 18:1 but not the isomerization of 18:2 n-6 to trans-10, cis-12-18:2. Shifts in biohydrogenation pathways favouring the production of trans-10 18:1 are known to occur in ruminants fed high concentrate diets containing unsaturated fatty acids169–171 or during incubations of 18:2 n-6 with mixed ruminal bacteria containing starch172,173 or maintained at a low pH.127 Recently, switching from high forage to high concentrate diets was shown to be accompanied by major changes in the bacterial community in lactating cows,174 conditions under which a shift towards the accumulation of trans-10 18:1 at the expense of trans-11 18:1 could be expected.175,176 Non-ruminal Lactobacillus plantarum converts 18:2 n-6 to cis-9, trans-11 18:2 and trans-9, trans-11 18:2.177 Of particular relevance to the trans-10 shift, L. plantarum reduces geometric 9,11 18:2 isomers to trans-10 18:1 rather than trans-11 18:1. Lactobacillus spp. have commonly been isolated from the rumen.178 Their numbers increase with dietary starch content.179 However, they have been associated with hydration rather than hydrogenation of unsaturated fatty acids.180 A future challenge will be to associate with certainty the role of individual microbial species with alterations in rumen function and biohydrogenation pathways, such as those indicated from ribosomal intergenic spacer analysis.174

There are often significant differences between bacterial communities attached to solids and those free-swimming, ‘planktonic’ communities that inhabit the liquid phase of the fermentation mixture. The same is true for ruminal lipid metabolism, with the planktonic community converting 18:2 n-6 as far as 18:1 intermediates, whereas the solids-associated community reduced 18:2 n-6 to 18:0181. B. proteoclasticus was present only in the solids-associated community, at 12% of Butyrivibrio-related clones.181

Li et al.182 carried out an interesting enrichment study by sequentially subculturing bovine ruminal digesta in the presence of 50 µg per ml trans-11 18:1 in order to investigate species capable of the final reduction of trans 18:1 to 18:0. Amounts of 18:0 were produced at progressively faster rates as the subculturing progressed. 16S rRNA amplicon analysis indicated that bacteria closely related to B. proteoclasticus were enriched, along with a number of previously uncultured bacteria. It was not possible to establish if any of these bacteria hydrogenated trans-11 18:1. Furthermore, there is no obvious reason why trans-11 18:1 would enrich for biohydrogenating bacteria as there would be no benefit conferred to those bacteria able to do so. More likely was that the soluble sugars present in the medium selected in favour of these bacteria.

Several studies have indicated that bacteria other than the Butyrivibrio spp. might be involved in biohydrogenation, particularly in the final reduction of cis 18:1 and trans 18:1 to 18:0. The long-chain unsaturated fatty acids in marine algae inhibit the biohydrogenation of 18-carbon fatty acids in the rumen and alter the ruminal microbial community.183 These changes have been taken, by association, to indicate bacterial species involved in biohydrogenation. Inhibition of biohydrogenation by marine algae was found to induce changes in uncultivated bacteria clustering with Butyrivibrio and Pseudobutyrivibrio. Boeckaert et al.184 found that this changed when biohydrogenation was inhibited. In contrast, no significant decrease in numbers of the known Butyrivibrio-related 18:0 producers were detected in the rumen of sheep fed marine algae.120 However, only five animals per treatment were investigated and numerical decreases were detected as the amount of algae supplement increased. Restriction fragment length polymorphism (RFLP) of 16S rRNA amplicons indicated that Lactospiracaea spp. or Quinella-like bacteria increased, which could have occurred for other reasons. In experiments investigating the effects of fish oil supplements on the rumen bacterial community, only weak associations have been observed between digesta 18:0 concentration or ruminal 18:0 outflow and B. proteoclasticus group 16S rRNA concentration.122,185 Further investigation based on multivariate analysis of data obtained from terminal RFLP (tRFLP) and denaturing gradient gel electrophoresis (DGGE) revealed many terminal restriction fragments (T-RFs) and DGGE bands that were correlated with ruminal cis-9, trans-11 18:2, trans-11 18:1 and 18:0 concentrations.185 Predictive identification revealed that these linked tRFs were likely to originate from as yet uncultured bacteria, including Prevotella, Lachnospiraceae incertae sedis, and unclassified Bacteroidales, Clostridiales and Ruminococcaceae. DGGE bands led to similar conclusions. Consistent with other recent studies, the bacteria have not been isolated and possible isomerase or reductase activity remains to be confirmed.

1.3.2.2 Role of Ciliate Protozoa

Since up to half of the rumen microbial biomass may be protozoal in origin186 and ca. 75% of the microbial fatty acids present in the rumen may be in protozoa,125 it follows that protozoa could represent a very important source of several biohydrogenation intermediates, including isomers of CLA and trans-11 18:1. Early studies concluded that both protozoa and bacteria were involved in biohydrogenation.187,188 However, the extensive ingestion of bacteria by protozoa caused Dawson and Kemp189 to doubt this conclusion. Biohydrogenation in ruminal digesta was only slightly decreased following defaunation (i.e. removal of protozoa from the rumen) and the presence of protozoa was not necessary for biohydrogenation to occur.189 Others also suggested that the minor contribution of protozoa to the biohydrogenation process was due to the activity of ingested or associated bacteria.190,191 Yet it has been known for a long time that protozoal lipids contain proportionally more unsaturated fatty acids than the bacterial fraction.48,192 Recently, it was established that these unsaturated fatty acids include cis-9, trans-11 18:2 and trans-11 18:1,193 further highlighting the possible significance of protozoa as a means of facilitating the escape of biohydrogenation intermediates from the rumen. Protozoal species differ in fatty acid composition, with larger species like Ophryoscolex caudatus containing more than ten times higher concentrations of cis-9, trans-11 18:2 and trans-11 18:1 than small species such as Entodinium nannelum.193 However, the lipid of Isotricha prostoma, a large species and the only holotrich examined, had low concentrations of cis-9, trans-11 18:2 and trans-11 18:1. Holotrichs do not ingest the large particles seen with entodiniomorphs. In incubations with fractionated ruminal digesta, 18:2 n-6 metabolism was similar between strained ruminal fluid and the bacterial enriched fraction, while the fraction containing protozoa had a much lower activity. There was no evidence of 14C enrichment in trans-11 18:1 or cis-9, trans-11 18:2 during incubations of protozoa with 14C-18:0. No genes with sequence similarity to fatty acid desaturases from other organisms were found in cDNA libraries from ruminal protozoa (E. Devillard, personal communication). Thus, the protozoa are relatively abundant in trans-11 18:1 or cis-9, trans-11 18:2, but do not synthesize these from 18:2 n-6 or 18:0, confirming earlier considerations.189 It could be argued that the high unsaturated fatty acid content of protozoa arises from the ingestion of plant particles, especially chloroplasts.187,194 A recent study showed that the engulfment of chloroplasts is a major contributor to the high 18:3 n-3 concentration of protozoa.78 However, these observations do not explain the relatively high concentrations of trans-11 18:1 or cis-9, trans-11 18:2 that originate from biohydrogenation. Lourenço et al.195 suggested that these findings reflect protozoa preferentially accumulating trans-11 18:1 or cis-9, trans-11 18:2 formed by bacteria associated with lipids in ingested plant cell organelles (Figure 1.6).

Fig. 1.6 Confocal microscopy images of an Epidinium sp. isolated from the rumen of Hereford x Friesian steers 2 h after being offered fresh perennial ryegrass saturated with intracellular chloroplasts (adapted from Huws et al.78). Figure 1.6a is the light microscopy image and Figure 1.6b is the contrasting fluorescent image taken at a depth of 2.24 μm.

The relatively low conversion of these intermediates to 18:0 may arise because the bacteria responsible for the reduction of 18:1 intermediates are more vulnerable to protozoal digestive activities. Increased accumulation of 9,11 and 10,12 18:2 isomers in washed protozoa to antibiotic treatment196 reflects a similar phenomenon, in as much that the enrichment of biohydrogenation intermediates is not due to endogenous protozoal activity. Until the genes encoding the enzymes involved in both bacteria and protozoa are identified, it will be difficult to resolve the issue unambiguously. Recently, I. prostoma was shown incapable of hydrogenating 18:2 n-6,181 but considering its low trans-11 18:1 or cis-9, trans-11 18:2 CLA concentrations, this observation may not be relevant to other species, particularly entodiniomorphs.

Determinations of protozoal fatty acid composition imply that the availability of unsaturated fatty acids, including trans-11 18:1 or cis-9, trans-11 18:2, for absorption by the host animal could depend more on the flow of protozoa rather than bacteria from the rumen. Some ciliate protozoa are retained selectively within the rumen by a migration/sequestration mechanism that depends on chemotaxis.197,198 As a consequence, protozoal biomass at the duodenum is proportionally less than would be the case if protozoa escaped the rumen attached to digesta particles.199,200 The flow of microbial nitrogen at the duodenum of steers was recently shown to be 12–15% protozoal in origin. However, protozoal lipid accounted for between 30% and 43% of cis-9, trans-11 18:2 and 40% of trans-11 18:1 reaching the duodenum.201 The contribution of protozoa to the flows of 16:0 and 18:0 to the duodenum was less than 20% and 10%, respectively. While there is no doubt that chloroplasts accumulate inside protozoal cells, particularly entodiomorphs, translating that capability to increase unsaturated fatty acids, including trans-11 18:1 or cis-9, trans-11 18:2, available for absorption may prove to be elusive.202

1.3.2.3 Role of Other Ruminal Microorganisms

Anaerobic fungi were first discovered in the rumen before being found in other environments.203 It is estimated that they comprise around 7% of rumen microbial biomass, but this value is difficult to ascertain and undoubtedly highly variable.204 Mixed ruminal fungi are capable of the isomerization of 18:2 n-6 to cis-9, trans-11 18:2,205,206 but activity is very low in comparison with B. fibrisolvens.205,207Orpinomyces is the most active genus in biohydrogenation.206

Methanogenic archaea are also very significant members of the rumen microbial community, comprising up to perhaps 3–4% of the biomass.208 Until recently, the mechanisms of methane formation have formed the focus for research on ruminal archaea.209 This is changing, however, as genomic analysis reveals new, unsuspected activities.210 Lipase genes were not noted in the Methanobrevibacterium ruminantium genome, but many ORFs of unknown function were present.210 Whether these may also include enzymes involved in biohydrogenation is not clear.

1.3.3 Measuring Ruminal Synthesis

Quantitative assessment of ruminal CLA biosynthesis requires the measurement of DM flow at the omasum, abomasum or duodenum and a detailed analysis of lipid in digesta entering the sampling site. Measurement of nutrient flow in the ruminant gastro-intestinal tract is reliant on the use of indigestible marker systems to account for sampling errors that occur due to the tendency for digesta to separate during collection.211 Obtaining representative samples of digesta for the analysis of CLA content is of particular importance owing to the heterogeneous distribution of fatty acids in digesta. It is possible to account for errors due to unrepresentative sampling using a combination of indigestible markers that specifically associate with liquid or particulate fractions of digesta that exhibit different flow characteristics.211

Unless a single marker is uniformly distributed across all digesta phases, unrepresentative sampling will inevitably introduce errors in estimated flows.212 In a number of experiments, Cr2O3 has been used as a single marker to determine CLA at the duodenum in lactating cows,175,213 growing cattle214–217 and sheep.104,218,219 In other studies multiple markers have been used for the measurement of CLA at the omasum54,61,67,122,220 or duodenum.68,221–229

Analysis of CLA in sampled digesta is reliant on reliable methods for lipid extraction and the preparation of fatty acid derivatives. Conjugated fatty acids are susceptible to isomerization and may disappear during prolonged exposure to acid catalysts.230 Under these conditions there is also a risk that conjugated products are formed from endogenous sources during methylation. Base-catalysed transesterification has been shown to be the most accurate method for determining CLA composition in range of biological samples.231 However, NEFA are not methylated under these conditions. Virtually all of the CLA leaving the rumen is non-esterified,54,61 and therefore this catalyst is ineffective for the analysis of digesta fatty acid composition. Isomerization and the production of artifacts with acid-based catalysts can be minimized using lower temperatures during methylation,232 but under these conditions the methylation of PL is incomplete.231 Preparation of FAME using methanolic sulphuric acid,220,233 diazomethane followed by sodium methoxide,232 or methanolic sulphuric acid followed by sodium methoxide121 can be used while avoiding isomerization or the synthesis of allylic methoxy derivatives.

Determination of the relative distribution and abundance of specific CLA isomers also requires the use of long (≥100 m) highly polar capillary columns during gas chromatography (GC) analysis in combination with silver-ion high performance liquid chromatography (HPLC).234 During GC analysis, cis-9, trans-11 18:2 elutes with the same retention time as trans-8, cis-10 18:2 and trans-7, cis-9 18:2. The trans-11, cis-13 18:2 peak may contain minor amounts of cis-9, cis-11 18:2, while trans, trans isomers with double bonds from 7,9 to 10,12 typically elute as a single peak. The occurrence of 21:0 also complicates the determination of CLA by GC. Depending on the GC column used and the temperature programme applied 21:0 elutes anywhere between cis-11, trans-13 18:2 and cis-10, cis-12 18:2, and may therefore, be erroneously identified as an isomer of CLA.234–236 An additional complication is that the retention of 21:0 relative to several minor CLA isomers may also differ between GC columns of the same type and changes over the lifetime of the GC column.234

1.3.4 Formation of CLA Isomers

The amount of CLA formed in the rumen has been determined for lactating cows, non–lactating cows, growing cattle and non-lactating sheep (Table 1.3). There are no estimates for lactating sheep, goats or other ruminant species. Synthesis of CLA in the rumen of sheep, growing cattle, non-lactating cows or lactating cows varies between 0.02 and 7.14, 0.14 and 6.59, 0.68 and 3.00 and 1.02 and 15.3 g per day, respectively. It should be noted that the estimates based on GC analysis alone ignore the contribution of minor isomers and are subject to possible errors due to the interference of other fatty acids. Nevertheless, it is clear that diet composition rather than the level of intake, physiological state or species is the major determinant of CLA synthesis in the rumen.

Table 1.3 Synthesis of CLA in the rumen of growing cattle, lactating cows and sheep.
DMIa (kg/d) Forageb F : Cc Supplementd (g/kg DM) CLAe (g/d) No. of isomers detected Analytical methodf Referenceg
Non-lactating sheep
0.86 GH 18:82-73:27 SBO (31–46) 0.35–.23 3 GC Kucuk et al.218
0.56 GH 18:82 SBO (0–94) 0.08–0.41 3 GC Kucuk et al.219
0.80 GH 18:82 SAF (0–90) 0.02–7.14 4 GC Atkinson et al.104
Non-lactating cows
5.90 GS 55:45 0.68–0.82 2 GC Lock and Garnsworthy240
10.5-9.9 MS+LH 70:30 LO (0–49) 1.0–3.0 4 GC Doreau et al.228
Growing cattle
3.60–4.78 GS/RCS 100:0 0.17–.25 4 GC Lee et al.224
4.15–8.48 GS/RCS/WCS 60:40 0.74–2.66 1 GC Lee et al.221
6.11–6.69 GS 80:20-20:80 LO (25–33) 1.94–1.80 7 GC Lee et al.225
12.0–11.4 GH 82:18 HOS (131) 0.80–0.40 3 GC Scholljegerdes et al.216
12.0–11.6 GH 82:18 HLS (137) 0.80–1.30 3 GC Scholljegerdes et al.216
10.4–10.7 GH 14:86 MO (0–24) 0.63–1.25 7 GC Duckett et al.214
5.54–6.19 GH 12:88-36:64 SFO (20–40) 0.68–0.88 9 GC Sackmann et al.215
8.33–8.77 GH 12:88 RO/MO (30–40)+FO (0-10) 0.97–2.96 9 GC Duckett and Gillis217
5.18–5.20 GS 100:0 FO (0–30) 0.14–0.52 16 GC+HPLC Lee et al.227
6.59–5.09 RCS 100:0 FO (0–30) 0.30–0.48 16 GC+HPLC Lee et al.227
7.55–7.45 GS 60:40 FO (0–40) 2.65–6.59 4 GC Lee et al.222
10.5–9.77 MS 60:40 FO (0–24) 0.39–0.52 10 GC+HPLC Shingfield et al.229
8.88–8.34 MS 60:40 FO (0–30)/LO (0–30) 0.38–3.18 GC+HPLC Shingfield et al.68
Lactating cows
15.7/15.5 HSG/HNG 100:0 1.45/2.50 3 GC Doreau et al.226
16.7–21.8 FG/GH/GS 60:40 3.70–5.38 12 GC+HPLC Halmemies-Beauchet-Filleau et al.61
19.9–18.4 GS/RCS 60:40 3.94–4.12 15 GC+HPLC Halmemies-Beauchet-Filleau et al.54
20.6–24.1 MS+LH 60:40-25:75 1.02–1.84 12 GC+HPLC Piperova et al.175
19.6–20.5 GH 65:35-35:65 LO (0–30) 1.70–.71 8 GC Loor et al.176
15.1–15.9 GS 60:40 SFO (0–50) 3.0–15.3 13 GC+HPLC Shingfield et al.67
17:1/17:2/19.3 GH 35:65 FO(25)/LO(50)/SFO(50) 4.01/6.90/8.30 8 GC Loor et al.223
15.7–17.7 GS 60:40 FO (0–16) 4.44–3.46 11 GC+HPLC Shingfield et al.220
18.7–15.6 GS 60:40 FO (0–19) 3.82–5.13 12 GC+HPLC Shingfield et al.122
15.0–18.5 LS+GH+MS 36:64 FO (20–5)+SFO (0–20) 6.04–.89 NR GC Qiu et al.213
a DMI, dry matter intake. b Forage in the diet: GH, grass hay; GS, grass silage; FG, fresh grass; HNG, high nitrogen grass; HSG, high sugar grass; LH, lucerne haylage; MS, maize silage; RCS, red clover silage; WCS, white clover silage. c Dietary forage:concentrate ratio on a dry matter basis. d Lipid supplements: FO, Fish oil; LO, linseed oil; MO, Maize oil; RO, rapeseed oil; SAF, safflower oil; SBO, soyabean oil; SFO, sunflower oil. e Based on sampling at the duodenum or omasum and measurements of dry matter flow at the sampling site using indigestible markers. f Determination of total CLA based on gas chromatography alone or in combination with silver-ion high performance liquid chromatography. g Numbers refer to citations listed in the reference section.

Thus far, 16 isomers of CLA have been detected in ruminal,119,120 omasal54,61,67,122,220 or duodenal68,175,227,229 digesta with double bonds located at 7,9 through to 13,15. Most reports indicate that ruminal digesta contains only trace amounts of trans-7, cis-9 18:2. Not all 16 isomers have been isolated in a single sample of digesta, indicating that the formation of specific CLA isomers is dependent on the composition of the basal diet and the intake of fatty acid substrates. In lactating cows offered diets based on grass, grass hay, or red clover forages supplemented with cereals and solvent extracted plant protein supplements (Forage: Concentrate (F:C) ratio on a DM basis; 60:40), cis-9, trans-11 18:2 is the major isomer accounting for 48–84% of total CLA at the omasum.54,61,67,122,220Cis-9, trans-11 18:2 is the main CLA isomer formed in the rumen of cattle fed maize silage based diets68,229 or in sheep fed diets based on lucerne hay.119,120 However, the relative abundance of cis-9, trans-11 18:2 was found to be lower (27–31% of total CLA) in lactating cows fed diets (F:C ratio, 60:40) containing lucerne hay and maize silage.175 Under these circumstances the amounts of trans, trans (7,9 through to 12,14) isomers accounted for 57–63% of total CLA at the duodenum. Reports on ruminal synthesis of specific CLA isomers in non-lactating sheep fed high concentrate diets (F:C 18:82) containing no additional lipid are conflicting. In one investigation, cis-9, trans-11 18:2 was reported to be the major CLA isomer at the duodenum,219 whereas in a follow-up experiment trans, trans isomers were quantitatively more important.104

In ruminants fed high proportions of grass, grass silage or red clover silage, where 18:3 n-3 is the predominant dietary fatty acid, trans-11, trans-13 18:2 or trans-11, cis-13 18:2 are the principal CLA isomers formed in the rumen.227,228,237Trans-12, trans-14 18:2 is also relatively abundant, whereas cis-9, trans-11 18:2 accounts for between 6.7% and 32% of total CLA synthesis.227,228

A number of investigations have explored the potential to increase ruminal CLA synthesis through changes in diet composition. Emphasis has been placed on alterations in the relative proportions of forages and concentrates or marine lipid supplements to manipulate rumen environment or supplementing the diet with plant oils to increase the supply of substrates for ruminal CLA synthesis. For diets containing no additional lipid supplements, increases in concentrate supplementation have variable and rather marginal effects on the amount of CLA at the duodenum (Table 1.3). Decreases in F:C ratio from 60:40 to 25:75 were shown to promote ruminal synthesis of geometric isomers of 9,11 and 10,12 18:2.175 In contrast, decreases from 65:35 to 35:65 had no effect other than decreasing trans- 9, cis-11 18:2 and increasing formation of several trans, trans isomers.176

Studies of 18:2 n-6 metabolism in vitro indicate that the amount of cis-9, trans-11 18:2 leaving the rumen would be enhanced in diets that promote high passage rates, maintain rumen pH and contain high amounts of 18:2 n-6.127,129,130,132 Increasing the intake of 18:2 n-6 on ruminal CLA synthesis has been investigated in cattle and sheep. Dietary supplements of sunflower oil, soyabean oil or safflower as a source of 18:2 n-6 has been shown to increase the amount of 8,10, 9,11 and 10,12 18:2 isomers and total CLA at the omasum or duodenum (Table 1.3). However, increases in the the formation of specific isomers to 18:2 n-6 supply is dependent on the composition of the basal diet. In lactating cows fed diets based on grass silage (F:C ratio 60:40), supplements of sunflower oil from 0 to 750 g per day progressively increased cis-9, trans-11 18:2, trans-9, trans-11 18:2, trans-10, cis-12 18:2 and trans-10, trans-12 18:2 at the omasum67 (Table 1.4). In contrast, adding higher amounts of sunflower oil from 20 to 40 g per kg DM in high concentrate diets (average F:C ratio 24:76) stimulated ruminal trans-10, cis-12 18:2 formation (0.21–0.37 g per day) but had no effect on other measured CLA isomers at the duodenum in growing cattle.215 Ruminal synthesis of trans-10,cis-12 CLA has been reported to be as high as 1.83 g per day in lactating cows fed high concentrate diets (F:C ratio 35:65) containing 50 g per kg DM of sunflower oil.223 Consistent with these findings, ruminal infusion of safflower oil in non-lactating sheep fed high concentrate diets (F:C ratio 18:82) at a rate equivalent to 0 to 90 g per kg DM progressively increased the amount (g per day) of cis-9, trans-11 18:2 (<0.01–0.38), trans-9, trans-11 18:2 (0.01–2.08), trans-10, cis-12 18:2 (<0.01–4.49) and trans-10, trans-12 18:2 (0.03–0.42) recovered at the duodenum.104 At the highest level of infusion, trans-10,cis-12 CLA was identified as the principal isomer formed in the rumen. However, an earlier experiment in non-lactating sheep fed a similar diet reported that incremental soyabean oil supplements (0-94 g per kg DM) had no influence on cis-9, trans-11 18:2 and resulted in limited increases in trans-10, cis-12 18:2 from 0 to 0.22 g per day at the duodenum.219

Table 1.4 Synthesis of CLA isomers in the rumen of growing cattle and lactating cows.
DM Intake (kg/d) Foragea F:Cb Supplement (g/kg DM)c Conjugated linoleic acid isomer (mg/d)d Referencee
cis, trans trans, cis trans, trans
9,11 11,13 12,14 7,9 8,10 9,11 10,12 11,13 12,14 7,9 8,10 9,11 10,12 11,13 12,14
Lactating cows
20.6 MS+LH [60:40] 330 21 5 9 86 9 14 38 200 109 156 88 Piperova et al.175
21.9 MS+LH [60:40] Buffer (20)f 276 20 7 13 54 7 13 73 130 140 179 107
23.7 MS+LH [25:75] 529 35 5 22 256 13 21 96 391 234 154 82
24.1 MS+LH [25:75] Buffer (20)f 244 21 6 17 78 12 16 21 291 121 141 67
17.7 GS [60:40] 2858 13 52 95 460 0 9 224 47 403 193 Shingfield et al.220
15.7 GS [60:40] FO (16) 2077 11 2 21 197 46 99 552 57 89 78
15.1 GS [60:40] 1927 36 84 334 5 184 30 221 132 Shingfield et al.67
15.0 GS [60:40] SFO (17) 4768 14 144 237 26 444 131 241 139
14.7 GS [60:40] SFO (34) 9228 17 182 143 37 930 226 210 121
14.9 GS [60:40] SFO (50) 11571 23 396 322 36 1675 424 337 218
18.7 GS [60:40] 4246 69 104 299 95 64 286 65 559 244 Shingfield et al.122
18.8 GS [60:40] FO (4.0) 4091 26 24 256 109 76 232 31 217 140
17.8 GS [60:40] FO (8.4) 3741 21 30 194 269 134 331 33 94 68
15.6 GS [60:40] FO (19) 3457 17 49 55 201 86 264 58 60 4.9
16.7 FG [60:40] 2430 20 90 370 3 180 30 380 120 Halmemies-Beauchet-Filleau et al.61
18.4 GH [60:40] 4350 20 140 110 240 30 200 80
19.8 GH [60:40] 2710 60 280 160 10 250 80 370 140
20.2 GS [60:40] 2710 30 170 360 20 210 80 760 330
21.8 GS [60:40] 2710 40 240 580 20 240 80 880 400
19.9 GS [60:40] 1880 20 20 30 160 720 70 10 30 120 60 580 180 Halmemies-Beauchet- Filleau et al.54
18.4 RCS [60:40] 2320 10 20 30 110 510 90 0 20 90 60 600 190
Growing cattle
5.18 GS [100:0] 18 3 6 <1 5 5 6 1 1 4 1 78 32 Lee et al.227
5.11 GS [100:0] FO (10) 23 <1 4 2 1 6 49 2 1 0.7 6 4 40 24
4.87 GS [100:0] FO (20) 66 <1 <1 16 3 <1 12 74 7 1 2 21 7 58 55
5.20 GS [100:0] FO (30) 84 1 <1 15 10 2 17 126 14 8 7 25 16 74 53
6.59 RCS [100:0] 16 2 3 5 1 1 5 2 12 3 10 7 113 59
6.35 RCS [100:0] FO (10) 37 1 10 8 1 6 55 17 1 7 17 12 180 81
5.88 RCS [100:0] FO (20) 71 2 1 9 1 2 8 65 11 11 11 32 17 151 73
5.09 RCS [100:0] FO (30) 98 <1 26 6 7 7 110 6 8 8 31 15 143 95
10.5 MS [60:40] 230 3 6 10 26 12 48 26 22 15 Shingfield et al.229
10.6 MS [60:40] FO (8) 284 3 12 17 33 13 75 29 29 25
10.3 MS [60:40] FO (16) 278 3 13 7 25 15 72 20 22 18
9.8 MS [60:40] FO (24) 212 4 16 8 13 19 51 16 15 14
8.88 MS [60:40] 194 1 17 30 8 54 27 35 13 Shingfield et al.68
8.83 MS [60:40] FO (30) 107 5 5 29 8 12 56 14 11 9
8.34 MS [60:40] LO (30) 628 37 7 24 1194 22 163 58 690 289
8.83 MS [60:40] FO+LO (30) 264 4 10 12 96 14 80 10 27 30
a Forage in the diet: FG, fresh grass; GH, grass hay; GS, grass silage; LH, lucerne haylage; MS, maize silage; RCS, red clover silage. b Forage:concentrate ratio of the diet on a dry matter basis. c Lipid supplements: FO, Fish oil; LO, linseed oil; SFO, sunflower oil. d Determined based on sampling at the duodenum or omasum and measurements of dry matter flow at the sampling site using indigestible markers. Amounts of individual CLA isomers determined by complimentary gas chromatography and silver-ion high performance liquid chromatography. e Numbers refer to citations listed in the reference section. f Comprised of a mixture (3:1 by weight) of NaHCO3 and MgO.

Several investigations have examined the effects of dietary linseed oil supplements as a means to increase 18:2 n-6 and 18:3 n-3 intake on ruminal CLA synthesis. In non-lactating cows fed high forage diets (F:C ratio 70:30) supplements of linseed oil (49 g per kg DM) increased duodenal flows of unresolved trans-8, cis-10 18:2 and cis-9, trans-11 18:2, trans-11, cis-13 18:2 and trans-11, trans-13 18:2 from 0.22, 0.12 and 0.13 to 0.64, 0.46 and 0.64 g per day, respectively.228 In growing cattle fed maize silage based diets, linseed oil (30 g per kg DM) stimulated ruminal cis-9, trans-11 18:2, trans-8, cis-10 18:2, trans-11, cis-13 18:2, trans-13, cis-15 18:2 and trans, trans (9,11 to 13,15) 18:2 synthesis68 (Table 1.4). Consistent with reports on the influence of 18:2 n-6 supply, the composition of the basal diet is also a major determinant of the changes in ruminal CLA synthesis to linseed oil. In lactating cows fed high or low forage diets (F:C 65:35 vs. 35:65), linseed oil resulted in similar increases in ruminal cis-9, trans-11 18:2, trans-11, cis-13 18:2 and trans-11, trans-13 18:2 synthesis, irrespective of diet composition.176 However, trans-8, cis-10 18:2, trans-9, cis-11 18:2 and cis-11, trans-13 18:2 formation was increased on the low forage diet, but remained unchanged on the high forage diet.

It is well established that the concentrations of cis-9, trans-11 18:2 and total CLA are higher in milk in grazing ruminants compared with that on diets based on conserved forages.45,47,49 Relatively few studies have compared the synthesis of CLA isomers in the rumen of grazing ruminants compared with diets containing conserved forages.61,237 Even though the number of observations that have been made is limited, there is little evidence that the effects on milk fat CLA concentrations are related to differences in the synthesis of CLA isomers in the rumen. Similarly, replacing grass silage with grass hay61 or red clover silage54,227 has limited influence on total CLA synthesis, but may result in differences in the amounts of minor isomers formed in the rumen (Table 1.4).

The effect of dietary marine lipid supplements on ruminal CLA synthesis has also been investigated. Even though the highly unsaturated fatty acids in fish oil119,122,220 or marine algae120,184,238 inhibit the complete biohydrogenation of unsaturated 18-, 20-, 21- and 22-carbon unsaturated fatty acids in the rumen, the influence on the accumulation of total CLA is rather small. In some experiments, supplementing grass silage, red clover silage or grass silage based diets with fish oil has been demonstrated to increase trans-7, trans-9 18:2, trans-8, trans-10 18:2 and trans-9, trans-11 18:2 formation and decrease cis-12, trans-14 18:2 synthesis,220,227 but not in all cases122 (Table 1.4). Studies in growing cattle fed maize silage-based diets indicated that fish oil supplementation up to 16 g per kg diet DM resulted in marginal increases in cis-9, trans-11 18:2 and trans-9, trans-11 18:2 at the duodenum, changes that were not evident when the amount of fish oil in the diet was increased above 24 g per kg diet DM.68,229

1.3.5 Formation of CLA Precursors

Secretion of trans-7, cis-11 18:2 and cis-9, trans-11 18:2 in milk is known to be several-fold higher compared with the amounts of these isomers at the duodenum in lactating cows.45,239,240 These differences are explained by endogenous synthesis in ruminant tissues. It now known that the majority of cis-9, trans-11 18:2 in milk is formed from the desaturation of trans-11 18:1 in the mammary glands. Most, if not all, of the trans-7, cis-11 18:2 found in ruminant milk also originates from the desaturation using trans-7 18:1 as a substrate. The potential to increase the synthesis of CLA in the rumen is relatively limited compared with projections of the enrichment in ruminant foods required to confer benefits to human health. For this reason, considerable emphasis has been placed on understanding the role of diet on the synthesis of CLA precursors, trans-11 18:1 in particular, in the rumen. The reduction of trans 18:1 intermediates to 18:0 is considered rate limiting for the complete biohydrogenation of 18-carbon unsaturated fatty acids in the rumen.125 Studies in vitro have demonstrated that increases in the amount of 18:2 n-6 or 18:3 n-3 incubated with mixed ruminal bacteria results in the accumulation of trans 18:1 (Table 1.2). In vivo changes in diet composition tend to have a much greater influence on the amount and relative proportions of trans 18:1 leaving the rumen (Table 1.5) compared with ruminal CLA synthesis (Table 1.3). Therefore, formulating diets that increase ruminal formation of trans-11 18:1 rather than specifically targeting cis-9, trans-11 18:2 synthesis is more effective for increasing the CLA content of ruminant foods.

Table 1.5 Synthesis of trans 18:1 isomers in the rumen of growing cattle, lactating cows and sheep.
Foragea F:Cb Supplementc Trans 18:1 isomer (g/d)d Referencef
(g/kg DM) Δ4 Δ5 Δ6–8e Δ9 Δ10 Δ11 Δ12 Δ13/14e Δ15 Δ16
Non-lactating sheep
GH 18:82 Control (0) 0.23 2.62 0.22 Atkinson et al.104
SAF (90) 3.07 40.8 1.92
Growing cattle
HSG 100:0 0.45 0.23 0.33 7.85 0.48 0.69 0.88 Lee et al.224
GS 100:0 0.26 0.14 0.25 4.60 0.29 0.51 0.55
RCS 100:0 0.26 0.15 0.26 3.83 0.49 0.74 0.92
GS 80:20 0.08 0.05 2.24 1.18 1.57 24.3 2.66 1.84 4.14 0.23 Lee et al.225
20:80 0.15 0.10 2.92 1.28 4.68 29.2 3.57 2.98 5.56 1.22
GH 12:88 30 0.70 41.4 3.48 1.29 Sackmann et al.215
36:64 30 2.14 15.5 11.8 2.53
GS 100:0 Control <0.01 0.15 0.1 0.17 2.28 0.29 0.26 0.5 0.92 <0.01 Lee et al.227
100:0 FO (30) 0.06 0.03 0.77 0.67 1.08 7.87 1.49 0.69 1.28 1.22
RCS 100:0 Control 0.02 0.01 0.34 0.23 0.47 3.02 0.94 0.62 1.7 2.29
100:0 FO (30) 0.09 0.04 1.26 1.11 1.8 11.2 2.47 0.7 1.94 1.75
MS 60:40 Control 0.23 0.16 1.58 1.09 2.05 13.8 1.47 1.01 1.4 1.63 Shingfield et al.68
LO (30) 0.69 0.42 4.67 2.84 3.91 51.2 4.79 4.47 5.76 5.63
FO (30) 0.15 0.11 5.08 4.45 19.2 66.7 5.85 2.68 3.25 0.98
Non-lactating cows
HNG 100:0 0.36 1.63 1.97 1.03 2.85 62.0 2.75 13.9 0.30 5.65 Doreau et al.226
HSG 100:0 0.27 1.26 1.51 0.78 2.05 41.6 2.25 9.89 0.20 4.07
Lactating cows
FG 60:40 0.24 0.19 1.37 0.96 2.26 20.1 1.95 4.47 2.02 2.59 Halmemies-Beauchet-Filleau et al.61
GH 60:40 0.12 0.15 1.14 0.75 1.89 12.5 1.83 2.93 1.66 2.06
GH 60:40 0.20 0.19 1.44 0.85 2.72 15.0 2.15 4.16 1.84 2.27
GS 60:40 0.24 0.25 1.97 1.3 2.84 18.9 3.53 8.74 4.54 5.45
GS 60:40 0.29 0.29 2.16 1.43 3.53 22.9 4.12 10.3 4.81 5.81
GS 60:40 0.27 0.20 2.08 1.57 2.93 25.8 3.56 9.04 4.95 5.09 Halmemies-Beauchet- Filleau et al.54
RCS 60:40 0.32 0.23 2.36 1.71 2.92 25.8 4.10 10.8 5.80 5.69
MS+LH 60:40 1.02 2.41 5.73 20.76 5.68 14.11 5.74 5.45 Piperova et al.175
25:75 3.72 4.55 29.13 33.61 9.52 22.86 8.53 7.98
GS 60:40 Control 0.3 0.3 1.3 0.8 1.3 14.9 1.7 4.5 2.3 2.9 Shingfield et al.67
SFO (50) 1.9 1.3 10.5 7.0 20.6 126.2 12.9 22.6 10.7 12.3
GH 65:35 Control 0.37 1.29 1.83 1.38 1.46 21.4 1.93 4.17 1.95 2.34 Loor et al.176
LO (30) 1.06 3.12 6.75 3.89 6.61 61.7 8.32 29.6 12.1 11.5
GH 35:65 Control 0.88 1.81 5.98 2.96 20.2 26.0 3.78 10.3 4.84 3.98
LO (30) 1.87 3.36 16.2 13.1 50.6 139 9.57 42.9 16.8 10.7
GS 60:40 Control 0.62 0.44 3.46 2.02 4.25 22 4.31 11.3 5.24 5.99 Shingfield et al.122
60:40 FO (8.4) 0.74 0.62 7.94 6.37 9.95 80.8 11.8 21 9.2 7.59
60:40 FO (19) 0.4 0.4 6.83 5.72 56.4 66.5 9.41 15.9 6.68 2.84
a Forage in the diet: FG, fresh grass; GH, grass hay; GS, grass silage; HNG, high nitrogen grass; HSG, high sugar grass; LH, lucerne haylage; MS, maize silage; RCS, red clover silage. b Forage:concentrate ratio of the diet (on a dry matter basis). c Lipid supplements; FO, Fish oil; LO, linseed oil; SAF, safflower oil; SFO, sunflower oil. d Determined based on sampling at the duodenum or omasum and measurements of dry matter flow at the sampling site using indigestible markers. e Isomers not resolved during analysis. f Numbers refer to citations listed in the reference section.

In ruminants fed diets containing forages and concentrate ingredients, trans-11 18:1 is typically the major biohydrogenation intermediate leaving the rumen. Increases in concentrate supplementation have been shown to cause the accumulation of trans 18:1 intermediates in the rumen.176,218,241 Switching from grass hay based diets containing 35% to 65% of concentrates increased the flow of trans 18:1 (Δ4 to 16) at the duodenum in lactating cows.176 Changes in trans-11 18:1 in response to concentrate supplementation were marginal, with most of the increase being associated with the trans-10 isomer (Table 1.5). More extreme increases in the amount of concentrates offered to lactating cows, from 60% to 75%, caused all trans 18:1 isomers to accumulate, other than trans-11 and trans-16.176 Under these conditions, the most marked influence was on trans-10 18:1 at the duodenum, being increased four-fold compared with the high forage diets (Table 1.5). Switching from a diet based on dehydrated grass to concentrate ingredients was found to result in the almost complete disappearance of trans-11 18:1 and a substantial increase in trans-10 18:1 concentration of abomasal digesta in growing lambs.242

Examination of the effects of pH (6.4 vs. 5.6) or dietary F:C ratio (70:30 vs. 30:70) on the production of biohydrogenation intermediates in dual-flow continuous culture suggested that decreases in pH rather that increases in concentrate per se was the principal factor responsible for the shift from trans-11 18:1 to trans-10 18:1 accumulation.127 However, this interpretation has been challenged based on recent reports of 18:2 n-6 metabolism by ruminal bacteria243 and measurements of ruminal fatty acid composition in cows fed diets containing low or high amounts of starch.173 The findings from both investigations led to the conclusion that the influence of starch on trans-10 18:1 formation was independent of the decreases in pH that occur during starch fermentation.

Under most situations, metabolism of 18:2 n-6 and 18:3 n-3 in the rumen leads to the formation of trans-11 18:1 during the complete hydrogenation to 18:0 (Figures 1.3 and 1.4). It is perhaps unsurprising that dietary supplements of oils enriched in 18:2 n-6 or 18:3 n-3 have used to increase the amounts of trans-11 18:1 formed in the rumen. Including plant oils in the diet have been shown to result in dose dependent increases in the amount of trans 18:1 intermediates leaving the rumen in sheep104,218,219, growing cattle,68,214–217 lactating67,176 and non-lactating cows.228 However, the profile of trans 18:1 intermediates that accumulate is dependent on a complex interaction between the relative proportions of forages and concentrates in the diet and the amount and source of oil supplement (Table 1.5). On high forage diets, supplements of sunflower oil67 or linseed oil176 increase the amount of trans-11 18:1 escaping the rumen. In marked contrast, inclusion of the same oils in high concentrate diets has minimal influence on trans-11 18:1, but results in trans-10 18:1 accumulating several-fold.176,215 Increases in the concentration of trans-10 18:1 are also accompanied by decreases in the synthesis of fatty acids in the mammary glands of cows and sheep, but not in goats.107 Changes in milk fatty acid composition also serve as a proxy of the relative amounts of trans 18:1 escaping the rumen, indicating that the major pathways of biohydrogenation are altered under these conditions causing trans-10 18:1 to displace trans-11 18:1 as the principal intermediate formed in the rumen. The most recent evidence has indicated that a low rumen pH and high dietary concentrations of starch and oil are prerequisite in promoting the ‘trans-10 shift’.172,173

Palmquist et al.45 proposed that ruminal trans-11 18:1 formation is dependent on three inter-dependent processes; 1) substrate supply, 2) inhibition of the reduction of trans 18:1 to 18:0 and 3) prevention of the trans-10 shift. It was argued that substrate supply has a typically permissive role in determining the extent of trans 18:1 fatty acid accumulation in the rumen in response to the other two processes, such that the balance between the inhibition of trans 18:1 reduction and induction of the shift towards trans-10 18:1 at the expense of trans-11 18:1 regulates the magnitude of the overall response to changes in diet composition.

Fish oil and marine algae contain relatively high amounts of long chain (≥ 20-carbon) polyunsaturated fatty acids. When added to the diet of ruminants, both lipid sources lead to an inhibition of the complete biohydrogenation of 18-carbon unsaturated fatty acids causing trans 18:1 and trans 18:2 intermediates to accumulate.68,120,122 Fish oil is an effective means to increase the amount of trans-11 18:1 leaving the rumen (Table 1.5), but in high amounts can induce a shift towards trans-10 18:1 formation.122 Incubations with mixed ruminal bacteria have shown that both 20:5 n-3 and 22:6 n-3 cause trans 18:1 to accumulate.244–246 It is possible that other 20-, 22- or 24-carbon unsaturated fatty acids contained in fish oil and marine algae may also inhibit the reduction of trans 18:1 to 18:0 in the rumen.

Relatively few studies have examined the effects of forage conservation or forage species on ruminal biohydrogenation. The most recent investigations indicate that the relative abundance of trans-11 18:1 in the rumen of lactating cows is higher on pasture compared with zero-grazing or grass silage.237 Flows of trans-11 18:1 at the omasum have also been shown to be higher in lactating cows during zero-grazing compared with diets based on hay or silage prepared at the same time from the same grass swards.61 Conservation of grass by drying or ensiling had no effect on the amount of trans-11 18:1 leaving the rumen.61 Replacing grass silage with red clover silage54,227 also has little influence on trans-11 18:1 or total trans-18:1 formation in the rumen (Table 1.5).

Studies involving the incubation of fatty acids with mixed ruminal bacteria have demonstrated that the biohydrogenation of cis-9 18:1, 18:2 n-6 and 18:3 n-3 results in the formation of trans-7 18:1 as an intermediate (Table 1.2). Because trans-6, -7 and -8 elute as a single peak during GC analysis there are no definitive reports on the effect of diet on trans-7 18:1 formation in the rumen. Direct comparisons of milk fat trans-7, cis-9 18:2 concentrations in ruminants offered plant oil or oilseed supplements have shown a consistently higher enrichment when rapeseed lipids are fed.46,47 By implication, biohydrogenation of cis-9 18:1 appears to be the main source of trans-7 18:1 in the rumen.

Recent studies have also provided evidence that trans-9 16:1 (palmitelaidic acid) may also serve as a substrate for endogenous cis-9, trans-11 18:2 synthesis in ruminant tissues.247 The metabolic origins of trans 16:1 formed in the rumen have not been extensively investigated. A range of trans 16:1 with ethylenic bonds between positions Δ3 to 14 are known to be formed in the rumen and incorporated into milk fat.54,248 Fish oil is known to cause trans-9 16:1 to accumulate in the rumen.68,122,227,229 Under these situations, it seems plausible that trans-9 16:1 originates from the incomplete biohydrogenation of cis-9, cis-12 16:2 or cis-6, cis-9, cis-12, cis-15 16:4. On conventional diets, trans 16:1 isomers are thought to originate from the incomplete biohydrogenation of cis-6, cis-9, cis-12 16:3 present in trace amounts in the ruminant diet.248

1.4 Endogenous CLA Synthesis in Ruminants

Early reports indicated that the concentrations of cis-9, trans-11 18:2 in milk were increased on pasture249,250 or on diets supplemented with linseed oil251 or fish oil.252 Such findings were perplexing. The convention at the time was that ruminal biohydrogenation of 18:2 n-6 was responsible for cis-9, trans-11 18:2 formation in ruminants, and yet pasture and linseed oil contain high proportions of 18:3 n-3 and fish oil contains few 18-carbon unsaturated fatty acids. Much earlier studies had demonstrated that several trans 18:1 isomers could be converted to cis-9 containing 18:2 products during incubations with rat liver microsomal preparations.253,254

These seminal investigations confirmed that trans 18:1 fatty acids with double bonds at Δ4–7 and Δ11–13 were substrates for the enzyme stearoyl-CoA desaturase (SCD; E.C. 1.14.99.5). Based on the observed changes in milk fat composition to diet and the substrate specificity of the SCD enzyme, Griinari and Bauman255 proposed that the rumen was not the sole source of cis-9, trans-11 18:2 and that endogenous synthesis due to the desaturation of trans-11 18:1 also occurred in ruminant tissues. An overview of the metabolic pathways of endogenous CLA synthesis in ruminants is described in Figure 1.7.

Fig. 1.7 Metabolic pathways of endogenous conjugated linoleic acid synthesis in ruminants. Most of the cis-9, trans-11 18:2 incorporated into tissue lipids and milk fat is synthesized by the action of stearoyl-CoA desaturase (SCD) on trans-11 18:1 in adipose and the mammary glands. Trans-11 18:1 is formed as the penultimate intermediate of 18:2 n-6 and 18:3 n-3 metabolism in the rumen. Cis-9, trans-11 18:2 in adipose may also be synthesized from the elongation and desaturation of trans-9 16:1 formed in the rumen during the biohydrogenation of 16:3 n-4. Ruminal synthesis of trans-7, cis-9 18:2 is negligible. The activity of SCD on trans-7 18:1 formed during the biohydrogenation of cis-9 18:1 in the rumen is the main source of trans-7, cis-9 18:2 in ruminant tissues and milk fat.

1.4.1 Stearoyl CoA Desaturase

1.4.1.1 Tissue Specific Expression

The SCD gene encodes a protein of 359 amino acid residues located in the endoplasmic reticulum (ER) that catalyzes the desaturation of 10- to 19-carbon fatty acyl-CoA substrates, resulting in the formation of a cis double bond located between carbons 9 and 10 of the fatty acid moiety.256,257 Homologues of the SCD gene have been characterized in several mammalian species, including the mouse, rat, hamster, pig and human. Some mammalian genomes have been shown to contain multiple SCD isoforms. In mice, four isoforms (SCD1, SCD2, SCD3 and SCD4) have been documented.258–261 Until recently only one isoform of the SCD gene, now referred to as SCD1, had been characterized in sheep,262 cows263 and goats.264 A second SCD isoform (SCD5) was recently identified in cattle265 and subsequently found to be expressed in the goat.265

Expression of the SCD1 gene is several-fold higher in adipose compared with brain, heart, liver, lung, muscle, spleen or intestinal mucosa,265,267 with adipose tissue being the major site of desaturase activity in growing ruminants.268,269 Abundance of SCD1 mRNA in subcutaneous adipose tissue increases after weaning270 with activity of the desaturase enzyme increasing during growth even after SCD gene expression starts to decline.271 Catalytic activity of SCD also varies between tissues. Activity of SCD was found to be higher in intestinal mucosa, interfasicular and subcutaneous adipose compared with liver (18.1, 10.6, 5.24 and 1.81 nmol per mg protein per min, respectively).272 Immediately after parturition, there is a substantial upregulation of SCD1 expression in the mammary glands of sheep262 and cattle.273 In lactating goats, relative expression of SCD1 is higher in mammary tissue compared with the liver, subcutaneous, omental and perirenal adipose274–277 consistent with the mammary glands being the major site of fatty acid desaturation in lactating ruminants.278,279

Transcript abundance of SCD5 is several-fold higher in the brain compared with heart, liver, lung, muscle and spleen in growing cattle.265 SCD5 is also expressed in the bovine mammary gland.280 In lactating goats, abundance of SCD5 mRNA is higher in mammary tissue compared with the liver, omental and perirenal adipose.277,281 Even though SCD5 is expressed in several tissues, the contribution of this SCD isoform to the desaturation of fatty acids in ruminants remains unclear.

1.4.1.2 Catalytic Activity

The SCD enzyme is the rate-limiting step in the synthesis of monoenoic fatty acids that are utilized as substrates for TAG, PL and cholesterol ester (CE) formation.282 Stearoyl and palmitoyl- CoA are the preferred substrates but a range of 10- to 19-carbon fatty acyl-CoA are desaturated in various ruminant tissues.46,256Cis-9 18:1 is the major product of the SCD enzyme in mammals, the incorporation of which into PL contributes to the maintenance of membrane fluidity.282 Activity of SCD in the ruminant mammary gland is thought to occur as a mechanism to maintain and regulate the fluidity of milk.283

Fatty acid desaturases are nonheme iron-containing enzymes that introduce a double bond between defined carbons of fatty acyl chains. In common with other desaturases, SCD catalyses a highly regio- and stereo-selective reaction on long-chain fatty acids that consist of essentially equivalent methylene chains in the absence of a distinguishing feature close to the site of desaturation. The Δ9 desaturase homologous to SCD isolated from Ricinus communis (castor) is a homodimeric protein with each monomer folded into a compact single domain comprised of nine helices.284 A diiron active site is located within a core four-helix bundle that is positioned alongside a deep, bent, narrow hydrophobic channel in which the substrate is bound during catalysis.285 It is thought that SCD has 4 transmembrane domains in the ER membrane and that both the -NH2 and -COOH terminal groups are oriented toward the cytosol.286

Oxidation of fatty acyl-CoA substrates catalysed by SCD in mammals is an aerobic process with an absolute requirement for molecular oxygen, cytochrome b5 reductase and the electron acceptor cytochrome b5.287 Desaturation is initiated by the energy demanding abstraction of a hydrogen atom from the methylene group at the Δ9 position followed by the abstraction of a second hydrogen atom from the Δ10 position.288 This is made possible by the recruitment and activation of molecular oxygen with the use of an active-site diiron cluster.289 Electrons are transferred sequentially from NAD(P)H, via NADH-cytochrome b5 reductase and cytochrome b5, to the terminal desaturase SCD, and finally to active oxygen, which is reduced to H2O287 (Figure 1.8).

Fig. 1.8 Scheme for the transfer of electrons during the oxidation of acyl trans-11 18:1-CoA yielding acyl cis-9, trans-11 18:2 CoA by stearoyl-CoA desaturase (SCD) adapted from Paton and Ntambi.288 Adipose is the major site of desaturation of fatty acids in growing ruminants, whereas the mammary glands are the main site in lactating ruminants.

For membrane bound desaturases the initial breaking of the C–H bond is rate-limiting.290 Electron transfer is facilitated by a catalytic di iron protein complex of the terminal desaturase.291 Insertion of Fe to the di-iron centre is thought to occur spontaneously and anchored to the active site by binding with carboxyl and nitrogen groups of glutamine and histidine residues, with evidence that the affinity of the two iron atoms comprising the di-iron center differ.289

The promoter region of the bovine SCD1 gene was shown to contain a 36-bp sequence of critical importance, designated the stearoyl-CoA desaturase transcriptional enhancer element (STE) that contains three putative transcription factor binding complexes.292 In the bovine MAC-T cell, the SCD gene promoter was shown to be up-regulated by insulin and down-regulated by cis-9 18:1, with the STE region shown to play a key role in the inhibitory effect of trans-10, cis-12 18:2 on SCD gene transcription. There is no known allosteric or feedback inhibition by substrates or products but the SCD enzyme is regulated by several dietary factors including glucose and PUFA, hormones such as insulin, glucagon and thyroid hormone and the sterol responsive element binding protein-1c transcription factor.257,282,288 Transcriptional regulation is made possible due to the relatively short half-life (3–4 h) of the mammalian SCD protein.293 In ruminants, SCD mRNA abundance and activity in the mammary glands has in some, but not all studies, been reported to be decreased on diets containing plant oils, oilseeds or marine lipid supplements, whereas corresponding effects on the transcription and activity of the SCD enzyme in adipose are marginal.274–277 In growing cattle, plant oils and fish oil have been shown to lower SCD gene and protein expression in adipose but not the amount of substrate desaturated per unit SCD protein.47 Abomasal infusion of trans-10, cis-12 18:2280,294 or oral dosing or ruminal administration of cobalt EDTA or cobalt acetate295,296 have been shown to decrease fatty acid desaturation in the bovine mammary glands.

1.4.2 Endogenous Synthesis in the Mammary Glands

It is well established that fatty acids are extensively desaturated in the mammary glands of ruminants. The activity of the SCD enzyme contributes to ca. 56%, 60% and 90% of cis-9 18:1, cis-9 16:1 and cis-9 14:1, respectively, in bovine milk.239,297–299 Several approaches have been used to estimate the contribution of endogenous cis-9, trans-11 18:2 synthesis in lactating ruminants. These have involved: 1) postruminal infusion of sterculic oil to inhibit SCD activity in the mammary glands and measuring subsequent changes in milk fatty acid composition, 2) postruminal infusions of trans-11 18:1 and measuring the output of cis-9, trans-11 18:2 in milk, 3) comparison of ruminal synthesis and output of secretion of CLA isomers in milk, 4) measurements of enrichment of cis-9, trans-11 18:2 in milk following the administration of 13C-labelled trans-11 18:1 and 5) measuring arterio-venous differences of circulating NEFA and TAG across the mammary glands and the output of fatty acids in milk fat (Table 1.6).

Table 1.6 Endogenous cis-9, trans-11 18:2 synthesis in the mammary glands of lactating cows, goats and sheep. Values determined based on 1) post-ruminal infusion of sterculic oil to inhibit stearoyl-CoA desaturase activity and measuring the subsequent changes in milk fatty acid composition, 2) post-ruminal infusion of trans-11 18:1 and measuring changes in cis-9, trans-11 18:2 in milk, 3) administration of 13C-labelled trans-11 18:1 and determination of 13C enrichment in cis-9, trans-11 18:2, 4) measuring arterio-venous differences across the mammary glands and secretion of fatty acids in milk and 5) comparison of ruminal synthesis and output of trans-11 18:1 and cis-9, trans-11 18:2 in milk.
Species Foragea F:Cb Supplementc (g/kg DM) Methodology Desaturation of trans-11 18:1, % Endogenous cis-9, trans-11 18:2 synthesis, % Referenced
Bovine LH 45:55 Abomasal infusion of sterculic oil ≥64 Griinari et al.300
LH 45:55 Abomasal infusion of sterculic oil 78 Corl et al.301
Pasture 100:0 Abomasal infusion of sterculic oil 24 ≥91 Kay et al.306
Pasture 100:0 SFO (28) Abomasal infusion of sterculic oil 20 ≥91 Kay et al.306
LH 47:53 Abomasal infusion of 12.5 g trans-11 18:1/d Griinari et al.300
LH 45:55 Abomasal infusion of 23.5 g trans-11 18:1/d 23.4 Corl et al.301
GS 75:25 Abomasal infusion of 7.5-30 g trans-11 18:1/d 28.9 84.4 Shingfield et al.302
LS/BS/LH 53:47 Abomasal administration of 1.5 g 1-13C trans-11 18:1 25.7 83.1 Mosley et al.279
FG/GH/GS 60:40 Arterio-venous differences across the mammary glands 18.6–34.1 78.0 Halmemies-Beauchet- Filleau et al.299
Multiple 35:65 100:0 Meta-analysis of fatty acid flow at the duodenum and secretion in milk 21.0 94.7 Glasser et al.239
Caprine GH 66:34 SFO (46) i.v. administration of 1.5 g of 1-13C trans-11 18:1 31.7 73.1 Bernard et al.266
GH 68:3 2 SFO (30)+FO (15) i.v. administration of 1.5 g of 1-13C trans-11 18:1 31.6 62.9 Bernard et al.266
Ovine Pasture 100:0 i.v. administration of sterculic oil 74 Bichi et al.307
a Forage in the diet: BS, barley silage; FG, fresh grass; GH, grass hay; GS, grass silage; LH, lucerne haylage; LS, lucerne silage. b Forage:concentrate ratio of the diet (on a dry matter basis). c Lipid supplements: FO, Fish oil; SFO, sunflower oil. d Numbers refer to citations listed in the reference section.

The first demonstration that endogenous synthesis of CLA occurred in ruminants was reported in lactating cows. Abomasal infusion of a mixture of fatty acids in lactating cows providing 12.5 g per day of trans-11 18:1 were shown to increase milk cis-9, trans-11 18:2 concentrations by 31%.300 Subsequent studies demonstrated that infusions of higher amounts of trans-11 18:1 of up to 30 g per day enriched cis-9, trans-11 18:2 in milk with no evidence that the extent of desaturation varied according to substrate supply.301,302 In goats, intravenous injections of trans-11 18:1 were found to elevate cis-9, trans-11 18:2 concentrations in milk and upregulate SCD gene expression in mammary tissue.303 Administration of trans-11 18:1 was also associated with an upregulation of proteasome (prosome, macropain) subunit α type 5 (PSMA5) and downregulation of peroxiredoxin-1 and translationally controlled tumor protein 1 in mammary tissue. The PSMA5 complex is involved in the proteolytic degradation via the ubiquitin-proteasome pathway. Given that SCD is a short-lived protein and constitutively degraded within the ER by the ubiquitin-proteasome system,304 it has been suggested that PSMA5 is involved in SCD degradation.303

As an alternative to increasing substrate supply, several experiments have used sterculic oil containing 7-2-octyl-1-cyclopropenyl heptanoic acid and 8-2-octyl-1-cyclopropenyl octanoic acid to inhibit SCD activity and measured the changes in trans-11 18:1 and cis-9, trans-11 18:2 concentrations. Using cis-9 14:1/14:0 concentration ratios as a proxy of the extent of SCD inhibition, the contribution of endogenous synthesis to cis-9, trans-11 18:2 in milk was estimated to vary between 64% and 91% in lactating cows.300,301,305,306 Using the same methodology, between 85% and 100% of trans-7, cis-9 18:2 in bovine milk was estimated to originate from the desaturation of trans-7 18:1.305 Similarly intravenous infusion of sterculic oil and measurements of milk fat composition in sheep estimated that endogenous synthesis contributed to 74% of cis-9, trans-11 18:2 milk.307 Quantitative estimates of endogenous CLA synthesis in lactating ruminants have been made using 1-13C labelled trans-11 18:1. Modelling of the exponential decay of 13C enrichment of fatty acids in milk indicated that 26% of trans-11 18:1 taken up by the mammary glands was converted to cis-9, trans-11 18:2.279 Desaturation of trans-11 18:1 was determined as 32% in the caprine mammary glands.266 Endogenous cis-9, trans-11 18:2 synthesis in lactating cows has also been determined from measurements of arterio-venous differences across the mammary glands and the output of fatty acids in milk.299 Such experiments indicated that between 18.6% and 34.1% of trans-11 18:1 extracted from circulating NEFA and TAG and taken up by the mammary glands was converted to cis-9, trans-11 18:2. Overall, these experiments have confirmed that endogenous synthesis is the major source of cis-9, trans-11 18:2 in ruminant milk. Numerous investigations have demonstrated that the concentrations of trans-11 18:1 and cis-9, trans-11 18:2 in bovine,255,308 caprine309,310 and ovine311,312 milk fat are highly correlated over a wide range of diets. Regression analysis indicate that the relationship between cis-9, trans-11 18:2 and trans-11 18:1 is linear, and that the slopes are similar (ca. 0.40), irrespective of the source of milk analysed. Such evidence indicate that increases in trans-11 18:1 supply does not inhibit desaturation in the mammary glands and that the conversion to cis-9, trans-11 18:2 is very similar in the cow, goat and sheep.

1.4.3 Endogenous Synthesis in Other Tissues

Estimating the extent of endogenous CLA synthesis in growing ruminants is particularly challenging given that CLA isomers accumulate in adipose over the life time of the animal. Several studies have attempted to estimate endogenous synthesis of the major CLA isomer in growing cattle based on comparisons of trans-11 18:1 and cis-9, trans-11 18:2 abundance in abomasal digesta and muscle. Such comparisons suggest that ca. 86% of cis-9, trans-11 18:2 in Longissimus dorsi originates from trans-11 18:1 and more than 80% in adipose tissue.313,314 Modelling of the relative proportions of trans-11 18:1 and cis-9, trans-11 18:2 in mesenteric adipose, subcutaneous adipose and longissimus muscle in lambs have also been used to predict endogenous synthesis. Between 45% and 95% of cis-9, trans-11 18:2 deposited in muscle and adipose was estimated to originate from trans-11 18:1 with the extent of conversion ranging between 11% and 22%.315

Recent studies in lactating goats have attempted to quantify endogenous CLA synthesis in several tissues of lactating goats using 1-13C trans-11 18:1 as a chemical tracer.277 Goats were slaughtered 4 d after jugular injection and 13C enrichment of trans-11 18:1 and cis-9, trans-11 18:2 was determined in mammary secretory tissue, liver, omental and perirenal adipose. From these measurements, it was calculated that 27% of trans-11 18:1 was desaturated to cis-9, trans-11 18:2 in perirenal adipose but conversion was only 2% in omental fat. The same experiment also demonstrated a higher enrichment of 13C trans-11 18:1 in the liver compared with mammary or adipose tissues. Due to hepatic uptake and incorporation of 13C cis-9, trans-11 18:2 synthesized in other tissues, no conclusions can be drawn on the role of the liver in endogenous CLA synthesis in ruminants. Even though mRNA encoding for SCD has been identified in the liver of ruminants, reports on the relative importance of hepatic fatty acid desaturation are inconsistent.45 Studies in vitro indicate that trans-11 18:1 is not converted to cis-9, trans-11 18:2 in the bovine liver.316

A recent experiment also provided the first evidence that trans-9 16:1 may also serve as a substrate for endogenous cis-9, trans-11 18:2 synthesis in ruminant tissues. Incubations of bovine primary stromal vascular cells with incremental amounts of trans-9 16:1 resulted in a progressive increase in adipocyte trans-11 18:1 and cis-9, trans-11 18:2 concentrations.247 Measurements indicated that ca. 50% of trans-9 16:1 incorporated into adipocytes was elongated to trans-11 18:1, with about 8% being desaturated to cis-9, trans-11 18:2. It was proposed that trans-9 16:1 served as a substrate for the fatty acid elongases 5 (ELOVL5) and 6 (ELOVL6), which catalysed the addition of two carbon atoms to yield trans-11 18:1 that could subsequently be converted to cis-9, trans-11 18:2.

1.5 Synthesis of CLA in Humans

1.5.1 Endogenous Synthesis

Synthesis of cis-9, trans-11 18:2 via the action of SCD on trans-11 18:1 has been documented in mice,317–319 rats,320–322 hamsters323 and pigs.324 Two isoforms of the SCD gene, SCD1325 and SCD5326 have been characterized in humans. The SCD1 gene is expressed in most tissues, whereas that of SCD5 is essentially confined to the brain and pancreas.288 Several key observations indicated that endogenous CLA synthesis via the activity of the SCD enzyme also occurs in humans. Firstly, the concentration of cis-9, trans-11 18:2 in plasma was found to be elevated in volunteers consuming diets high in trans fatty acids.327,328 Secondly, 13C enrichment of cis-9, trans-11 18:2 was detected in the blood of a single male subject offered 1-13C trans-11 18:1.329 Thirdly, trans-11 18:1 was shown to be converted to cis-9, trans-11 18:2 during incubations with human mammary (MCF-7) and colon (SW480) cancer cell lines.330

The first quantitative estimates of endogenous cis-9, trans-11 18:2 synthesis were derived from an intervention involving 30 healthy subjects.331 Volunteers were offered a diet rich in oleic acid for two weeks, followed by diets providing 1.5, 3.0 or 4.5 g of trans-11 18:1 per day for nine days. Plasma concentrations of trans-11 18:1 were elevated 194%, 407% and 620% above baseline, respectively. These changes were also accompanied by 50%, 169% and 198% increases in cis-9, trans-11 18:2 concentrations. Regression of the change in cis-9, trans-11 18:2 against the change in trans-11 18:1 plus the change in cis-9, trans-11 18:2 in circulating TAG of very low density lipoproteins indicated that 19% of trans-11 18:1 was desaturated. These estimates were supported by a longer intervention in which 12 volunteers were offered 3 g per day of trans-11 18:1 over a 42 d interval.332 Concentrations of trans-11 18:1 and cis-9, trans-11 18:2 were increased eight- and two-fold relative to baseline. Enrichment of trans-11 18:1 and cis-9, trans-11 18:2 estimated that 24% and 19% of trans-11 18:1 was desaturated in plasma and red blood cell membranes, respectively. These observations led the authors to propose that ca. 25% of trans-11 18:1 supplied from the diet was converted to cis-9, trans-11 18:2 in humans.332 Such estimates do not, however, account for the disappearance of trans-11 18:1-derived cis-9, trans-11 18:2 into organs or tissues from the circulation, and are not an indication of the extent of desaturation, but simply a reflection of the net sum of end-products surviving metabolism. Conversion of trans-11 18:1 in humans was confirmed in an intervention involving four lactating women offered 2.5 mg of 13C-labelled trans-11 18:1 per kg body weight.333 Enrichment of 13C was detected for cis-9, trans-11 18:incorporated into TAG, CE and PL fractions in plasma and in milk lipids. Up to 10% of cis-9, trans-11 18:2 in milk was found to originate from trans-11 18:1, indicating that endogenous synthesis of CLA in the mammary glands of humans is considerably lower than for ruminants (Table 1.6).

In all human studies to date, the extent of endogenous cis-9, trans-11 18:2 synthesis has been shown to vary considerably between individuals. These differences have been attributed to variations in diet composition, physiological state or genetics. In rats, the conversion of trans-11 18:1 to cis-9, trans-11 18:2 has been shown to differ between tissues, ranging from 36.2% to 4.2% (testes and kidneys>adipose>ovaries>muscle>liver>heart).334 In addition to tissue specific expression, genetic variations may also influence the SCD enzyme activity and endogenous cis-9, trans-11 18:2 synthesis. A recent study provided the first indications of an association between single nucleotide polymorphisms of the SCD gene and concentration ratios of cis-9, trans-11 18:2/trans-11 18:1 in the plasma of Caucasian and Asian adults.335

Recent studies in vitro have also raised the possibility that trans-9 16:1 is elongated and desaturated to cis-9, trans-11 18:2 in humans.247 There are no reports on the metabolic fate of trans-9 16:1 in humans. Evidence emerging from clinical prospective studies have reported that higher trans-9 16:1 concentrations in blood are associated with a more favourable metabolic profile and lower incidence of diabetes.336,337 Thus far, no cause and effect has been established.

Milk and dairy products are the major source of CLA in the human diet.31,35–39 However, these foods provide more CLA than would be indicated based on proximal analysis of CLA composition. Irrespective of species, diet and production system, the relative concentrations of trans-11 18:1 and cis-9, trans-11 18:2 in ruminant milk are relatively constant in a ratio of 2.5:1.255,308–312 The relationship between trans-11 18:1 and cis-9, trans-11 18:2 in ruminant muscle, is however, much more variable, which reflects both differences in the relative proportions of TAG and PL in intramuscular lipid and the differential incorporation of trans-11 18:1 and cis-9, trans-11 18:2 in these lipid fractions. In growing cattle, the relative abundance of trans-11 18:1 and cis-9, trans-11 18:2 varies between fat deposits in a ratio ranging between 0.15 and 0.23 compared with 0.17, 0.26 and 0.85 in heart, liver and erythrocyte PL, respectively.338–340 Relative concentrations of cis-9, trans-11 18:2 to trans-11 18:1 also differ between subcutaneous (0.19), omental (0.15), perirenal adipose (0.12) and liver (0.30) in growing lambs.243 However, ruminant meat is not the principal source of CLA in the human diet.34–36,38,44 Assuming a mean conversion of trans-11 18:1 to cis-9, trans-11 18:2 in humans of 20%, it has been proposed that the effective physiological dose supplied by ruminant-derived foods is 1.4–1.5 times the measured CLA content.39,42

1.5.2 Synthesis in the Hindgut

Evidence of CLA formation by human intestinal bacteria was first obtained indirectly in the experiments of Chin et al.,341 who noted that germ-free rats had a lower incorporation of CLA in liver, lung, kidney, skeletal muscle and abdominal adipose tissue than conventional animals. Subsequently, mixed human intestinal flora were shown to convert 18:2 n-6 to isomers of CLA,342 thus offering the possibility that the human gut microbiota could contribute to the CLA status in humans. This prospect was diminished when it was discovered that little or none of the CLA formed in the rat intestine was absorbed and incorporated into host tissues. Absorption of CLA by the colonic epithelium has not been demonstrated, and it is therefore difficult to understand the mechanisms by which CLA synthesis could elicit a systemic effect. Nevertheless, more recent studies suggest a possible important role in situ for CLA produced in the intestine. Anti-inflammatory activity of cis-9, trans-11 18:2 and trans-10, cis-12 18:2 has been reported in the mouse model of inflammatory bowel disease (IBD).343 Moreover, in animal studies, cis-9, trans-11 18:2 has proved to be a potent anti-carcinogen by lowering the incidence of chemically induced mouse aberrant crypt foci in the rat colon.11 Isomers of CLA have also been found to exhibit anti-proliferative properties in vitro, inhibiting the growth of human colon cancer cells.344 The possibility therefore arises that, if 18:2 n-6 can be delivered to the intestine, and the correct bacterial population is present, possibly through the simultaneous delivery of a CLA-producing ‘probiotic’, the susceptibility to IBD and colorectal cancer in humans might be decreased.

A survey of 30 representative strains of human Gram-positive intestinal bacteria indicated that several indigenous human intestinal bacteria could form isomers of CLA from 18:2 n-6.345Roseburia species were among the most active. Different Roseburia spp. formed either trans-11 18:1 or a 10-OH 18:1 intermediate, both of which served as precursors for cis-9, trans-11 18:2 synthesis. Bacteria from other ecosystems and from food products, but that are also found in the human gut, including strains of Lactobacillus, Propionibacterium and Bifidobacterium, have been known for some time to be capable of 18:2 n-6 isomerisation to cis-9, trans-11 18:2.156,346–348

However, the more abundant bacterial species belonging to clostridial clusters IV and XIVa also metabolized 18:2 n-6 at among the fastest rates of all bacteria investigated, forming products that can serve as precursors of CLA synthesis (Figure 1.9). Given the greater abundance of Clostridium-like bacteria present in the human intestinal microbiota,349,350 it may be deduced that 18:2 n-6 metabolism by this major group will be quantitatively more important than that of the Lactobacillus, Propionibacterium and Bifidobacterium groups. Nonetheless, given that the Lactobacillus and Bifidobacterium genera provide many probiotics in common use, and because, unlike the Roseburia, they are not strict anaerobes, these two genera may be of more practical significance.347Bifidobacterium breve has been shown to have multiple effects on tissue concentrations of polyunsaturated fatty acids in rats used as a model for IBS,351 although the ability to hydrogenate fatty acids, if any, in causing these benefits was unclear.

Fig. 1.9 Proposed pathways of cis-9, cis-12 18:2 metabolism by bacterial species isolated from the human gut adapted from Devillard et al.352 The open arrows represent the bacterial activity of Lactobacillus, Propionibacterium and Bifidobacterium species leading to the formation of 9,11 and 10,12 isomers of conjugated linoleic acid (CLA). The shaded arrows represent the bacterial activity of some Lactobacillus, Propionibacterium and Bifidobacterium species and some Clostridium-like bacteria belonging to clusters IV (e.g. Eubacterium siraeum) and XIVa (e.g. R. intestinalis and Roseburia faecis) leading to the formation of hydroxyl fatty acids (HFA). The solid arrows represent the bacterial activity of Clostridium-like bacteria belonging to cluster XIVa leading to the formation of trans-11 18:1 (VA) (e.g. Roseburia hominis and R. inulinivorans). The dotted arrows represent activities observed in faecal microbiota for which the responsible bacterial species are still unknown.

Thus, there appear to be two potential routes of cis-9, trans-11 18:2 formation in the human intestine (Figure 1.9). When faecal bacteria from four human donors and six species of human intestinal bacteria capable of 18:2 n-6 metabolism were incubated with 18:2 n-6 in deuterium oxide-enriched medium, the main products in faecal suspensions were cis-9, trans-11 18:2 and trans-9, trans-11 18:2, which were labelled at Δ13, as were the other 9,11 geometric isomers formed.352 Traces of trans-10, cis-12 18:2 were formed, but this product was labelled to a much lower extent than cis-9, trans-11 18:2. In pure culture, Bifidobacterium breve formed labelled cis-9, trans-11 18:2 and trans-9, trans-11 18:2, while a human faeces-derived B. fibrisolvens, Roseburia hominis, Roseburia inulinivorans and Ruminococcus obeum-like strain A2-162 converted 18:2 n-6 to trans-11 18:1, labelled in a manner indicating that trans-11 18:1 was formed via C-13-labelled cis-9, trans-11 18:2.352Propionibacterium freudenreichii subsp. shermanii, a possible probiotic strain, formed mainly cis-9, trans-11 18:2 with smaller amounts of trans-10, cis-12 18:2 and trans-9, trans-11 18:2, labelled in the same manner as in the mixed microbiota. Ricinoleic acid (12-OH-cis-9 18:1) was not converted to one or more CLA isomers in the mixed microbiota, in contrast to that described for Lactobacillus plantarum.352 These results were similar to those reported for the mixed microbiota of the rumen. Thus, although the bacterial genera and species responsible for biohydrogenation in the human intestine differs from the rumen, and a second route of cis-9, trans-11 18:2 formation via a 10-OH 18:1 is present in the intestine, the overall labelling patterns of the different CLA isomers formed are common to both gut ecosystems. A hydrogen-abstraction enzyme mechanism was proposed to explain the role of a 10-OH 18:1 intermediate in the formation of cis-9, trans-11 18:2 during incubations of 18:2 n-6 with pure and mixed cultures.352

1.6 Conclusions

Following the discovery that isomers of CLA exhibit anti-mutagenic properties in the rodent model of cancer, subsequent investigations have demonstrated that cis-9, trans-11 18:2 and trans-10, cis-12 18:2 elicit a diverse range of biological activities in cell culture and animal models. These findings have served as a catalyst to explore the biochemical, microbial and physiological mechanisms regulating the synthesis of CLA in ruminants, and more specifically, the transformations of dietary lipid that occur in the rumen. Even though cis-9, trans-11 18:2 is the principal isomer in ruminant tissues and milk, numerous isomers are formed in the rumen. Thus far, sixteen isomers have been detected in ruminal digesta that originate from the metabolism of 18:2 n-6 and 18:3 n-3. Bacteria rather than protozoa or fungi are the primary microorganisms responsible for catalysing the cis-trans isomerization yielding conjugated intermediates. Several ruminal bacteria capable of converting 18:2 n-6 to 9,11 or 10,12 geometric 18:2 isomers have been identified. Much less is known about the metabolic origins of other CLA isomers formed in the rumen. Diets rich in 18:3 n-3 promote the synthesis of 11,13 18:2 and 12,14 18:2 isomers in the rumen, but the metabolic pathways explaining their formation are not well characterized. Synthesis of CLA isomers can be increased several fold by supplementing the diet of ruminants with oils enriched in 18:2 n-6 and 18:3 n-3. Secretion of trans-7, cis-9 18:2 and cis-9, trans-11 18:2 in milk or the appearance of these isomers in host tissues is many fold higher compared with their synthesis in the rumen, due to endogenous synthesis catalysed by the stearoyl CoA desaturase (SCD) enzyme. Virtually all of the trans-7, cis-9 18:2 in ruminants is synthesized via the action of SCD on trans-7 18:1 formed as an intermediate of cis-9 18:1 biohydrogenation in the rumen. Endogenous synthesis using trans-11 18:1 as a substrate accounts for at least 60% of cis-9, trans-11 18:2 in ruminant milk. Between 45% and 95% of cis-9, trans-11 18:2 in ruminant tissues originates from trans-11 18:1. Trans-11 18:1 is the penultimate intermediate formed during the biohydrogenation of 18:2 n-6 and 18:3 n-3 in the rumen. Reduction of trans 18:1 intermediates is thought to be rate limiting for the complete biohydrogenation of 18-carbon unsaturated fatty acids to 18:0. Changes in diet composition and increases in 18:2 n-6 and 18:3 n-3 supply have a profound influence on the formation of the trans 18:1 precursors for endogenous synthesis, more so than on the formation of CLA isomers in the rumen. For this reason, nutritional strategies for enhancing the CLA content of ruminant foods have focused on increasing the amount of trans-11 18:1 escaping the rumen and preventing shifts in biohydrogenation pathways leading to the formation of other trans 18:1 isomers. Recent investigations suggest that trans-9 16:1 escaping the rumen may also serve as a substrate for endogenous cis-9, trans-11 18:2 synthesis via a mechanism that involves elongation to trans-11 18:1 followed by SCD catalysed desaturation. Formation of trans-9 16:1 has not been extensively investigated, but is thought to be an intermediate of 16:2 n-4, 16:3 n-4 and 16:4 n-1 metabolism in the rumen. Appearance of other CLA isomers is directly related to their synthesis in the rumen. Endogenous synthesis of cis-9, trans-11 18:2 via the action of SCD on trans-11 18:1 has also been demonstrated in humans. Desaturation of trans-11 18:1 in the mammary glands is much lower in women compared with cows, goats or sheep. Conversion of trans-11 18:1 to cis-9, trans-11 18:2 implies that the effective physiological dose of CLA supplied from ruminant-derived foods is 1.4–1.5 times the measured content.

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