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© 2008 American Society for Nutrition J. Nutr. 138:889-896, May 2008

Nutrient Physiology, Metabolism, and Nutrient-Nutrient Interactions

Fish Oil Increases the Duodenal Flow of Long Chain Polyunsaturated Fatty Acids and trans-11 18:1 and Decreases 18:0 in Steers via Changes in the Rumen Bacterial Community1,2

Eun J. Kim3,6, Sharon A. Huws3,6, Michael R. F. Lee3, Jeff D. Wood4, Stefan M. Muetzel5, R. John Wallace5 and Nigel D. Scollan3,*

3 Institute of Grassland and Environmental Research, Plas Gogerddan, Aberystwyth, SY23 3EB, UK; 4 Division of Food Animal Science, University of Bristol, Langford, Bristol, BS40 5DU, UK; and 5 Rowett Research Institute, Bucksburn, Aberdeen, AB21 9SB, UK

* To whom correspondence should be addressed. E-mail: nigel.scollan{at}bbsrc.ac.uk.


   ABSTRACT
TOP
 ABSTRACT
Introduction
 Materials and Methods
 Results
 Discussion
 LITERATURE CITED
 
Ruminant fat is rich in SFA, partly due to the biohydrogenation of dietary PUFA to SFA in the rumen. This process can be inhibited by the dietary inclusion of fish oil. The only bacteria isolated from the rumen capable of converting PUFA to SFA are closely related to Clostridium proteoclasticum. The aim of this study was to investigate if a correlation could be found in vivo between dietary fish oil inclusions and the composition of the ruminal bacterial community and specifically of C. proteoclasticum. Six Hereford x Friesian steers, prepared with ruminal and duodenal cannulae, received grass silage plus 1 of 3 concentrates resulting in total dietary fish oil contents of 0, 1, or 3% of dry matter. A dual flow marker technique was employed to estimate the relative flow of fatty acids. Steers fed the 3% fish oil diet had 100% increases in trans 18:1 flow, whereas 18:0 flow declined to 39% of steers fed the control diet. 16S ribosomal RNA-based denaturing gradient gel electrophoresis profiles obtained from ruminal digesta showed major changes in the bacterial community within steers fed the 3% fish oil diet. Quantitative PCR indicated only a weak relation between numbers of C. proteoclasticum and 18:0 flow between treatments and in individual steers (P < 0.05, but the percentage variance accounted for only 22.8) and did not provide unambiguous evidence that numbers of C. proteoclasticum in the rumen dictate the ratios of SFA:PUFA available for absorption by the animal. Understanding which microbes biohydrogenate PUFA in the rumen is key to developing novel strategies to improve the quality of ruminant products.


   Introduction
TOP
 ABSTRACT
 Introduction
Materials and Methods
 Results
 Discussion
 LITERATURE CITED
 
Recent nutritional advice has emphasized the need to decrease intake of SFA and increase intakes of beneficial PUFA, in particular {alpha}-linolenic acid [18:3(n-3)] and the longer chain (n-3) PUFA [20:5(n-3) and 22:6(n-3)], in the diet (1). Ruminant products make an important contribution to the human diet but have caused concern due to their enriched SFA content (2). This is largely due to microbial biohydrogenation of dietary unsaturated fatty acids in the rumen, although some intermediates of biohydrogenation such as conjugated linolenic acid (cis-9, trans-11 CLA)7 and trans-11 18:1 could be important in human health (3). Previous studies have shown that including fish oil in the diet of beef cattle resulted in increased long chain (longer than 20 C) PUFA (LCPUFA) in muscle resulting in a lower (n-6):(n-3) fatty acid ratio (4). Fish oil has also been shown to interrupt the complete biohydrogenation of C18 PUFA, resulting in increased production of trans-11 18:1 (5–7), the precursor for CLA (cis-9, trans-11) in the mammary gland (8).

The bacteria involved in the different steps of the biohydrogenation pathway have been categorized as Group A and B (9): group A bacteria hydrogenate 18:2(n-6) and 18:3(n-3) to trans-11 18:1; in contrast, group B bacteria convert the same fatty acids to 18:0. The only Group B bacteria identified for many years was ‘Fusocillus’ spp. (10,11). Modern phylogenetic analysis of recent isolates has now shown that 18:0-forming bacteria, like the most active Group A bacteria, are part of the Butyrivibrio fibrisolvens group, an ill-defined taxon that includes the genera Butyrivibrio and Pseudobutyrivibrio and the species Clostridium proteoclasticum (12,13). Group B bacteria (18:0 producers) form a tight grouping in which strains cluster together close to C. proteoclasticum (14,15). For this reason, in this article, the 18:0 producers are described as C. proteoclasticum.

Advances in molecular microbial technology based on 16S ribosomal RNA (rRNA) genes mean that we are now able to quantify these bacterial groups using quantitative PCR (QPCR) and to investigate total eubacterial and Butyrivibrio-specific population changes using denaturing gradient gel electrophoresis (DGGE) (16). One of the major advantages of these molecular methodologies is the avoidance of an often laborious cultivation step that is frequently error-prone due to media selectivity and the suspected existence of noncultivable bacteria.

The aims of this study were to assess the involvement of C. proteoclasticum, other Butyrivibrio-related spp. and eubacteria in general in the biohydrogenation pathways operating in the rumen.


   Materials and Methods
TOP
 ABSTRACT
 Introduction
 Materials and Methods
Results
 Discussion
 LITERATURE CITED
 
    Animals, diets, and experimental design. The experiment was conducted under the authorities of the UK Animal (Scientific Procedures) Act (1986). Six Hereford x Friesian steers (609 ± 6.9 kg) prepared with ruminal cannulae and simple T-piece cannulae in the proximal duodenum [immediately post-pylorus and pre-common bile duct; (17)] were offered grass silage plus 1 of 3 concentrates: F0, F1, and F3 that provided increasing amounts of fish oil: 0, 23, and 69 g · kg–1 dry matter (DM), respectively (0, 1, and 3% of DM intake, respectively). The grass silage was prepared from first-cut perennial ryegrass (Lolium perenne) in May 2002. Salmon (Salmo salar) oil was purchased from the International Fish Meal and Oil Manufacturers Association. To maintain an iso-lipid nature of the diets, the addition of fish oil was balanced by the removal of Megalac (Volac; Table 1). The total daily allowance was set at 14 g DM · kg–1 live weight to ensure complete daily consumption with a forage:concentrate ratio of 60:40 (DM basis). Steers were housed in individual pens and transferred to stalls for each measurement period. The building was well ventilated and steers had free access to fresh water and mineral blocks (Baby Red Rockies, Tithebarn; composed of 380 g · kg–1 Na, 5000 mg · kg–1 Mg, 1500 mg · kg–1 Fe, 300 mg · kg–1 Cu, 300 mg · kg–1 Zn, 200 mg · kg–1 Mn, 150 mg · kg–1 I, 50 mg · kg–1 Co, and 10 mg · g–1 Se).


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  		TABLE 1 Formulation of the concentrate component of experimental diets

 
The experimental design consisted of a replicated 3 x 3 Latin Square with 2 steers per treatment. Each 21-d period consisted of 17 d of adaptation to the experimental diets and 4 d for sample collection. Steers received their daily allocations in 2 equal meals at 0900 and 1600. Digesta flow at the duodenum was estimated using a dual-phase marker technique with ytterbium acetate and chromium EDTA as the particulate and liquid phase markers, respectively (18). Ytterbium acetate (375 mg · d–1 Yb) and chromium EDTA (2401 mg · d–1 Cr) were infused via separate lines intraruminally at a rate of 28 mL · h–1 commencing on d 8. On d 18 and 19, 400 mL of duodenal digesta were collected manually every 3 h over a 24-h period, bulked, and stored at 4°C for further fractionation (see below). Samples of strained ruminal fluid (1 L) were taken 2 h post-feeding, for microbial analysis on d 21 and 10 mL strained ruminal fluid were taken every 2 h over a 12-h period (0830 to 2030) to measure pH, ammonia-nitrogen (ammonia-N), and volatile fatty acids (VFA) concentration in the rumen.

    Sample preparation and chemical analysis. Accumulated samples of daily duodenal digesta were thoroughly mixed and a whole and centrifuged fraction produced as described by Lee et al. (19). Separate samples of silage and concentrate were taken daily (~500 g) during the sampling periods and pooled subsamples were freeze-dried, ground, and retained at –20°C for chemical analysis (19). The microbial fraction of ruminal fluid was obtained as described by Lee et al. (20) and freeze-dried and ground before molecular microbial analysis ensued. The fatty acids in the silage and concentrate were measured using a 1-step extraction-transesterification procedure (21). Digesta fatty acids were obtained by direct hydrolysis at 60°C, with added internal standard (100 µL 21:0 methyl ester, 15 g · L–1 CHCl3), in 5 mol · L–1 KOH in aqueous methanol. Potassium carboxylates were converted into fatty acids by the addition of 5 mol · L–1 H2SO4 and methylated using 5% HCl in methanol at 50°C (22). Samples were analyzed by GC on a CP-Select chemically bonded for FAME column (100-m x 0.25-mm i.d.; Varian), split injection 30:1, helium carrier gas, and a temperature gradient program according to Lee et al. (6). Peaks were identified from external standards (ME61, Larodan Fine Chemicals; S37, Supelco; CLA, Matreya) and quantified using the internal standard (21:0) using the Varian star 6.4.1 software.

    DNA extraction from rumen microbial samples. Genomic DNA was extracted from rumen microbial samples (10 mg DM) using the BIO101 FastDNA SPIN kit for soil (Qbiogene) in conjunction with a FastPrep cell disrupter instrument (Bio101, ThermoSavant, Qbiogene) according to the manufacturer's instructions, except that the samples were processed for 3 x 30 s at speed 6.0 in the FastPrep instrument. The integrity of the DNA was verified by agarose gel electrophoresis.

    PCR-DGGE analysis of the total eubacterial population and the Butyrivibrio group. Amplification of the V6-V8 region of the 16S rRNA gene was carried out with the primer pair F968GC (5'-CGC CCG CCG CGC GCG GCG GGC GGG GCG GGG GCA CGG GGG GAA CGC GAA GAA CCT TAC-3') and R1401 (5'-CGG TGT GTA CAA GAC CC-3') (23,24), and F968GC and B fib (5'-TTC GGG CAT TYC CRA CT-3') for total eubacterial and Butyrivibrio group-specific PCR, respectively. The 16S rRNA-targeted Butyrivibrio spp. specific reverse primer was based on a B. fibrisolvens probe published by Klieve et al. (25), but it was modified slightly so that it would amplify all members of the Butyrivibrio group while still fitting the criteria required for PCR-DGGE. In brief, we checked specificity in silico using an alignment of rumen bacterial sequences previously generated (26) and using the Probe Match tool in the Ribosomal Database Project-II release 9.42 (27). Once specificity for the Butyrivibrio group was determined, coverage of this group was investigated by aligning all Butyrivibrio group 16S rRNA gene sequences deposited in GenBank and EMBL databases using the program ClustalW (28). Based on this analysis, 2 bases were changed to include degeneracy to amplify all members of the Butyrivibrio group while maintaining specificity. Further specificity of the newly developed primer was confirmed by PCR of many pure cultures. Various Mg concentrations and annealing temperatures were investigated so that maximum specificity could be obtained while ensuring sensitivity at the same time. A Mg concentration of 3 mmol · L–1 was required to ensure sensitivity and an annealing temperature of 58°C was required to obtain specificity (data not shown). Specificity was then checked using DGGE analysis of ruminal digesta samples obtained from 2 ruminally cannulated Holstein-Friesian nonlactating dairy cows fed grass silage as described below. Dominant DGGE bands were excised and sequenced revealing that this PCR-DGGE amplified both cultivable and as yet uncultivated members of the Butyrivibrio group. Of 15 clones obtained from 5 dominant bands, 6 had 91% identity to Eubacterium cylindroides L346616, so it should also be noted that this PCR-DGGE may amplify Eubacterium spp. The reverse B fib primer does have one and sometimes 2 mismatches in the middle of the primer for sequences deposited for cultivable Eubacterium spp. and in the presence of eubacterial 16S rRNA having a 100% match with this primer, amplification of Eubacterium spp. should be minimal.

All PCR amplifications were performed using a 2720 thermal cycler (Applied Biosystems) in 50-µL volumes containing 1x PCR buffer (20 mmol · L–1 Tris HCl, pH 8.4, 50 mmol · L–1 KCl), 3 mmol · L–1 MgCl2, 200 µmol/L deoxyribonucleotide triphosphate mix, 500 mmol · L–1 each primer, and 1.25 U of iTaq DNA polymerase (Bio-Rad Laboratories) with ~100 ng of DNA template. Amplification conditions were: an initial denaturation of 95°C for 3 min followed by 35 cycles of 95°C for 30 s, 56°C (total eubacteria) or 58°C (Butyrivibrio group) for 30 s and 72°C for 1 min, and then a final extension of 72°C for 5 min. Amplification of products was verified by agarose gel electrophoresis.

Amplicons were loaded onto 6% polyacrylamide gels with a 35–60% (total eubacteria) or a 35–65% (Butyrivibrio group) denaturing gradient [100% denaturant consisting of 40% (v:v) deionized formamide and 7 mol/L urea] and electrophoresis performed in a D-Code system (Bio-Rad Laboratories) as described previously (29). Gels were then stained with AgNO3 (30). Gels were scanned using a GS-710 calibrated imaging densitometer (Bio-Rad Laboratories) and the saved image imported into the software package Fingerprinting (Bio-Rad Laboratories) for analysis. DGGE banding patterns were analyzed based on the presence and absence of the bands and resultant binary matrices were translated into distance matrices using the Dice similarity coefficient, with a position tolerance of 0.5% and optimization parameter of 1%. Finally, clusters were constructed using the method of unweighted pair group method with arithmetic mean analysis. The binary data generated were used to calculate band number and the Shannon-Weiner diversity index (24,31) using the Fingerprint Analysis with Missing Data software (32).

    QPCR analysis. Total eubacterial amplification was carried out on ruminal digesta samples in a final volume of 25 µL containing 12.5 µL SYBR Green JumpStart Taq ReadyMix (Sigma-Aldrich), 250 nmol · L–1 each of EubF1 5'-GTG STG CAY GGY TGT CGT CA-3' and Eub R1 5'-GAG GAA GGT GKG GAY GAC GT-3' (33), and 2 µL of a 1:100 dilution of extracted genomic DNA. The thermal cycling program was 30 cycles of 94°C for 30 s and 61°C for 30 s with an initial cycle of 94°C for 5 min. After PCR, a dissociation curve (melting curve) was constructed in the range of 55°C to 95°C. All samples were run in triplicate. A bacterial standard was prepared with equal amounts of genomic DNA from 8 different pure cultures of bacteria: Clostridium aminophilum (ATCC 49906), Peptostreptococcus anaerobius (ATCC 27337), Prevotella ruminicola (ATCC 19189), Fibrobacter succinogenes (ATCC 19169), B. fibrisolvens [JW11; Rowett Research Institute (RRI)], Ruminococcus albus (SY3; RRI), Selenomonas ruminantium (Z108; RRI), and Streptococcus bovis (ES1; RRI).

QPCR analysis of 18:0-producing bacteria was conducted according to the method by Paillard et al. (34) on rumen-derived samples. Dilutions of purified genomic DNA from the control strain Clostridium proteoclasticum P-18 (RRI) were used to construct specific calibration curves. All samples were run in triplicate. Amplification was carried out in a final volume of 25 µL containing 12.5 µL of JumpStart Taq ReadyMix (Bio-Rad Laboratories), 400 and 800 nmol · L–1 of forward (SA-FW; 5'-TCC GGT GGT ATG AGA TGG GC) and reverse primers (SA-RV; 5'-GTC GCT GCA TCA GAG TTT CCT-3'), respectively, 250 nmol · L–1 of molecular beacon (5'-6 FAM-CCG CTT GGC CGT CCG ACC TCT CAG TCC GAG CGG-DABCYL-3'), and 2 µL of a 1:10 dilution of extracted genomic DNA. The thermal cycling program was 40 cycles of 30 s at 95°C, 1 min at 55°C, and 30 s at 72°C with an initial cycle of 95°C for 10 min. Fluorescence data were collected at the end of the hybridization step at excitation and emission wavelengths of 490 and 530 nm, respectively.

All QPCR were performed using an iCycler iQ thermal cycler (Bio-Rad Laboratories) and results were analyzed using the iCycler iQ detection system software (Bio-Rad Laboratories).

    Calculations and statistical analysis. Digesta flows were estimated after mathematical reconstitution of true digesta as described by Faichney (18). Biohydrogenation of PUFA and LCPUFA was assessed as the difference between daily intake and duodenal flow (g · d–1) as a proportion of daily intake. Data for intakes and flows of nutrients [organic matter (OM), total N, neutral detergent fiber (NDF), and fatty acids], for rumen fermentation characteristics (pH, ammonia-N, VFA), and for eubacterial QPCR quantification were subjected to Restricted maximum likelihood analysis using GenStat [release 9.1, (35)]. The model included carryover and diet (treatment) as the fixed effect, and steer, period, and their interactions as the random effect. Treatment effects were further partitioned using a polynomial contrast to evaluate the significance of linear and quadratic components of the response to treatments. The level treated as not significant was P > 0.05, but trends were also expressed (P < 0.1). For band number and Shannon-Weiner diversity index data, difference was also inferred by using a Student's t test (32,35). For the correlation between the flow of 18:0 and the DNA concentration of 18:0-producing bacteria, bootstrap analysis (100 bootstrap measures) (35) was conducted to characterize the relationship more accurately, because linear regression demonstrated a few data points with high leverages.


   Results
TOP
 ABSTRACT
 Introduction
 Materials and Methods
 Results
Discussion
 LITERATURE CITED
 
    Chemical composition of the experimental diets. Concentrates were of typical composition with respect to barley and molasses sugar beet pulp with differences in the oil composition only (Table 1). The grass silage was of good quality with a low pH, and had comparatively low ammonia-N · g–1 total-N with 30.1 g total-N · kg–1 DM (Table 2). More than 50% of the fatty acid existed in the form of 18:3(n-3) in the grass silage. LCPUFA such as 20:5(n-3), 22:5(n-3), and 22:6(n-3) were undetectable in F0. Supplementing the concentrate with fish oil increased the LCPUFA concentration roughly proportionally to the dietary concentration of fish oil (Table 2).


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  		TABLE 2 Chemical composition and fatty acid profile of the experimental diets1

 
    Nutrient intake, duodenal flow, and rumen fermentation characteristics. As indicated in the experimental design, DM intake was controlled, with the mean intake of 7.42 kg · d–1 with the targeted forage:concentrate ratio (60:40, DM basis) throughout the experiment. Duodenal flow of total-N indicated a net gain in steers fed diet F0 and F1 compared with the intakes and was not different between treatments (Table 3). Apparently digested NDF in the rumen ranged from 65 to 80% of intakes in all treatments. There were very few changes in rumen pH and the concentrations of ammonia-N and total VFA in the rumen (Table 3).


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  		TABLE 3 Intake and duodenal flow of OM, total-N, NDF, and rumen fermentation parameters in steers receiving F0, F1, or F3 diets1

 
    Fatty acids intake, duodenal flow, and biohydrogenation. Total fatty acid intake was different (P < 0.001) across treatments with steers fed F3 consuming 25 g · d–1 less than steers offered the F0 diets, although the difference was marginal (just 7% of F0) (Table 4). Duodenal flow of total fatty acids indicated a net gain of 52 and 73 g · d–1 in steers fed diets F0 and F1, respectively, but a net loss of 14 g · d–1 in steers fed diet F3 compared with the intakes. Increasing the concentration of fish oil culminated in greater intakes and flows of major LCPUFA, including 20:5(n-3), 22:5(n-3), and 22:6(n-3). Duodenal flow of 18:0 was much higher than intake in steers receiving all diets and decreased extensively with increasing concentration of fish oil from F0 to F3 (P < 0.001). Flow of total trans 18:1 increased 2.0-fold in steers fed diet F3 relative to steers fed diet F0 (P < 0.001) and this was largely due to increased trans-10 and trans-11, with these isomers contributing >50% of total trans 18:1 in digesta (Table 5). Further fractionation of total cis 18:1 showed that the cis-9 was largely responsible for the decrease in the duodenal flow of this fraction. The most commonly reported CLA isomers, namely cis-9, trans-11 and trans-10, cis-12 CLA, combined flow to the duodenum accounted for 52 and 50% of total CLA flow in steers fed the F1 and F3 diets, respectively (Table 5). Fish oil did not affect the duodenal flow of the cis-9, trans-11 isomer, while increasing fish oil had an elevatory effect on trans-10, cis-12 flow (P < 0.001). However, total CLA flow to the duodenum did not change, with trans-11 trans-13 most abundant in steers fed all diets. Biohydrogenation of 18:1(n-9), 18:2(n-6), and 18:3(n-3) ranged between 0.58 and 0.97 among treatments. Likewise, the majority of 20:5(n-3) and 22:6(n-3) was hydrogenated in the rumen and biohydrogenation of 22:5(n-3) was lower than the other LCPUFA (Table 6).


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  		TABLE 4 Fatty acid intake and duodenal flows in steers receiving F0, F1, or F3 diets1

 

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  		TABLE 5 Duodenal flow of 18:1 and CLA isomers in steers receiving F0, F1, or F3 diets1

 

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  		TABLE 6 Biohydrogenation of unsaturated fatty acids in the rumen of steers receiving F0, F1, or F3 diets1

 
    Microbial population profiling. PCR-DGGE analysis of 16S rRNA genes using primers universal for eubacteria indicated complex communities in all samples, with many similarities between steers receiving the F0 and F1 diets (Fig. 1A). Diet F3 substantially altered the bacterial composition in the rumen of steers fed this diet. Cluster analysis separated steers fed diet F3 from those fed diets F0 and F1, with 77% of the band positions being the same (Fig. 1A). Band number also decreased (P < 0.05) in rumen samples taken from steers fed diet F3 compared to those fed diets F0 and F1 (Table 7). Similar analysis for members of the Butyrivibrio group showed a less complex pattern than eubacteria (Fig. 1B). Two main clusters were evident, separating steers fed diet F3 from those fed the other 2 diets, with 66% of the band positions being similar (Fig. 1B), and again band numbers were lower (P < 0.05) in F3 (Table 7). Subclusters between rumen samples derived from steers fed diets F0 and F1 were not clear in either analysis, indicating that supplementing fish oil at 1% of DM did not substantially alter microbial population in the rumen. The Shannon-Weiner diversity index showed only minor differences in bacterial diversity for steers fed all diets with respect to both total eubacteria and the Butyrivibrio population (Table 7).


Figure 1
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  		FIGURE 1  PCR-DGGE-derived unweighted pair group method with arithmetic mean dendrogram showing the effect of fish oil on the total eubacterial (A) and Butyrivibrio (B) populations in the rumen. F0, F1, and F3 relate to diets consisting of 0, 1, and 3% fish oil intake expressed in terms of DM intake. Scale relates to percent similarity.

 

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  		TABLE 7 Band number and Shannon-Weiner diversity index calculated from the total eubacterial and the Butyrivibrio group-specific DGGE profiles obtained from rumen samples of steers receiving F0, F1, or F3 diets1

 
QPCR of 16S rRNA genes indicated that total eubacterial DNA concentration was not significantly different as fish oil increased (data not shown). The same was true of C. proteoclasticum 16S rRNA gene concentration (Fig. 2A). When C. proteoclasticum numbers were expressed as a proportion of total eubacteria, there was no significant differences as fish oil increased (data not shown). A regression approach to the relationship between fish oil supplementation in the diets of the steers with the number of C. proteoclasticum in the rumen showed a significant relationship (P < 0.05), but the percentage variance accounted for (equivalent to adjusted r2) was only 22.8, which was not sufficient to provide a reliable interpretation of the relationship (Fig. 2B). Therefore, bootstrap estimates from 100 bootstrap samples were performed to examine the correlation more accurately and the results were not fully supportive of a clear association between the 2 variables (Fig. 2B).


Figure 2
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  		FIGURE 2  18:0-producing bacterial 16S rRNA gene concentration obtained from strained rumen fluid of steers given the experimental diets (A), and correlation between 18:0-producing bacterial 16S rRNA gene concentration and 18:0 flow to the duodenum (B). F0, F1, and F3 relate to diets consisting of 0, 1, and 3% fish oil intake expressed in terms of DM intake, n = 6. L, Linear response; Q, quadratic response.

 

   Discussion
TOP
 ABSTRACT
 Introduction
 Materials and Methods
 Results
 Discussion
LITERATURE CITED
 
Numerous reports have appeared in the literature demonstrating the effects of fish oil on ruminal biohydrogenation. The addition of fish oil to ruminant rations has been used to manipulate LCPUFA concentrations of meat (2,4) and milk (36,37). Upon ingestion by ruminants, dietary lipid undergoes substantial lipolysis followed by microbial biohydrogenation to produce a high concentration of SFA in the rumen. One of the major pathways of biohydrogenation in the rumen is simplified as the conversion of free C18 PUFA to conjugated dienes or trienes, and then to 18:1 and finally by reduction to form 18:0 (11). The results of the present experiment were consistent with earlier findings in that the flow of 18:0 to the duodenum was significantly lower when fish oil was incorporated into the diet of steers. The effect was also dependent on the concentration of the fish oil, as observed in other studies too (5,6,36). Several researchers reported a similar phenomenon with fish oil-supplemented diets (5–7,36) and suggested that the fish oil effect might involve an inhibition of the enzyme that catalyzes the final biohydrogenation step in the rumen. Alternatively, the proliferation of bacteria capable of conversion of 18:3 and 18:2 through to 18:0 might be inhibited by the LCPUFA in fish oil, resulting in increased trans-11 18:1. A recent study by Wasowska et al. (38) showed that the addition of 20:5(n-3) or 22:6(n-3) (50 mg · L–1) to pure cultures inhibited the growth and isomerase activity of B. fibrisolvens, whereas fish oil, in which the fatty acids are present as triacylglycerol, had no effect. Other reports (39,40) suggested that the accumulation of 18:2(n-6) in the rumen [by feeding more 18:2(n-6) to the ruminant] causes incomplete biohydrogenation. However, in the present study, the intake of 18:2(n-6) was similar with increasing concentrations of fish oil, excluding any possibility that 18:2(n-6) in the diets was responsible for the observed effect on biohydrogenation. Our data showed that fish oil at 3% of DM intake decreased the 18:0 flow and also increased the flow of trans-11 18:1 to the duodenum.

Increased flows of 20:5(n-3), 22:5(n-3), and 22:6(n-3) to the duodenum occurred in steers fed fish oil diets. The increased duodenal flow of these fatty acids in steers was lower than the increased dietary intake between diets F1 and F3. For example, the intake of 22:6(n-3) in steers fed diet F3 was 2.9-fold higher than that of steers fed diet F1 while only 1.6-fold higher in the duodenal flow. Biohydrogenation was presumably a major reason for this difference (5–7), although the degree to which these LCPUFA are biohydrogenated and the factors affecting biohydrogenation of LCPUFA in the rumen are still not well understood. Several studies have indicated that 20:5(n-3) and 22:6(n-3) are extensively metabolized in the rumen in vivo (5,6,41). In vitro studies have been less clear with some studies showing limited biohydrogenation (42), while others showed a decrease (43,44) or an increase in the extent of 20:5(n-3) and 22:6(n-3) biohydrogenation in relation to fish oil addition (45). It is also notable that although there was no intake of LCPUFA in the present study for the F0 diet, there were still measurable flows of these LCPUFA at the duodenum, which may reflect endogenous lipid from cell desquamation during digestive processes.

In terms of CLA, cis-9, trans-11 CLA was not the major isomer in duodenal digesta; trans-11, trans-13 CLA isomer accounted for ~50% of total CLA. Increased flow of trans-trans CLA leaving the rumen is consistent with previous reports of diets supplemented with fish oil (5,6,46). In addition, Lee et al. (7) reported that diets with high levels of 18:3(n-3), such as those based on forage or supplemented with oil rich in 18:3(n-3), produced trans-11, trans-13 as the predominant CLA isomer, possibly as a consequence of its involvement in the 18:3(n-3) biohydrogenation pathway. Duodenal flow of trans-10, cis-12 CLA, which is associated with modulating fat deposition (47) and milk fat depression (48), was significantly higher in steers fed the F3 diet. Several bacteria have been reported to convert 18:2(n-6) to trans-10, cis-12 CLA, including Lactobacillus spp. (49), Propionibacterium acnes (50), and Megasphaera elsdenii (51). However, to what extent these particular microorganisms play a role in ruminal biohydrogenation with the diet supplemented with fish oil remains unclear.

The microbial ecology of the rumen changed substantially with the addition of fish oil to the diet of steers, particularly at the higher inclusion rate. Many fewer bands were present in DGGE of 16S rRNA genes amplified by universal eubacterial primers. This reflects the toxicity of unsaturated fatty acids to ruminal bacteria, which is particularly severe for cellulolytic species and also butyrate producers (52), although no difference in the concentration of ruminal butyrate was observed in the present study. The altered banding pattern presumably reflects the loss of the most sensitive species. Biohydrogenation occurs to detoxify the fatty acids. DGGE was also used to analyze the impact of fish oil on the Butyrivibrio population. These bacteria are believed to be the most active species involved in fatty acid biohydrogenation (39,53). Their sensitivity to PUFA is highly variable, with some members of the group sensitive to growth inhibition at PUFA concentrations of 5 mg/L (13) while others tolerate concentrations many times higher. The persistence of some bands but not others in the Butyrivibrio DGGE is consistent with this range of sensitivity.

Bacteria forming a small branch of the Butyrivibrio phylogenetic tree, clustering around C. proteoclasticum, are among the most sensitive ruminal species to the toxic effects of PUFA (13,15). They are also the only known ruminal species to convert trans-11 18:1 to 18:0 (10,15). C. proteoclasticum was originally isolated as a proteolytic species (54). Its name does not reflect its taxonomic position accurately, because it is not a spore former and is distantly related to true Clostridium species (15) and its name is in need of revision. Whether C. proteoclasticum truly represents the predominant 18:0 producers in the rumen is by no means certain and it is possible that other 18:0 producers have not yet been cultivated; indeed, some may not be cultivable at all using present culture techniques. If C. proteoclasticum is indeed the main 18:0 producer, a strong correlation might be expected between their numbers and the extent of biohydrogenation to 18:0 in digesta leaving the rumen. Primers and a probe designed to detect the C. proteoclasticum group and to exclude related Butyrivibrio that do not form 18:0 (34) were used in QPCR to assess the influence of fish oil on C. proteoclasticum numbers. The results were equivocal in the sense that a correlation was found, but it was rather weak, particularly when comparing different steers. It may be that other microbial species are involved, although no other ruminal bacteria, protozoa, or fungi are known to carry out the reaction to date. Alternatively, metabolic factors may be involved; for example, 18:0 formation occurs during the growth phase of C. proteoclasticum, not when it reaches stationary phase (15). Thus, the metabolic activity of C. proteoclasticum may not be proportional to 16S rRNA gene concentration and RNA may be a better marker.

In conclusion, these results are consistent with the hypothesis that fish oil has an inhibitory effect on the biohydrogenation of fatty acids in the rumen via its influence on microbial ecology. Total bacteria and Butyrivibrio populations were changed, consistent with the elimination of a number of species by fish oil. Numbers of C. proteoclasticum and duodenal fatty acid composition gave a weak correlation that neither offers strong support for nor against its predominant role in the process vis-à-vis other unknown bacteria that may convert trans-11 18:0 to 18:0. We are now attempting to link microbial changes to differences in fatty acid flow to the duodenum, using multivariate statistical approaches and sequencing of key bands, to identify other key bacteria that may be involved at various points of the biohydrogenation sequence. This information could potentially allow the development of novel strategies for manipulating this process, leading to the beneficial enhancement in nutritional value of ruminant products.


   ACKNOWLEDGMENTS
 
We thank Martin Leyland at Institute of Grassland and Environmental Research, Trawsgoed research farm for the care of the steers, Delma Jones and colleagues in the Analytical Chemistry Department, John Tweed for the skilled GC analysis of lipid samples, and Mark Scott for help with the microbial analysis. We are also thankful for the statistical support of Graham Horgan.


   FOOTNOTES
 
1 Supported by the Department for Environment, Food and Rural Affairs, the Biotechnology and Biological Sciences Research Council, and the Scottish Executive Environment and Rural Affairs Department, UK. Back

2 Author disclosures: E. J. Kim, S. A. Huws, M. R. F. Lee, J. D. Wood, S. M. Muetzel, R. J. Wallace, and N. D. Scollan, no conflicts of interest. Back

6 These authors contributed equally to this study. Back

7 Abbreviations used: CLA, conjugated linoleic acid; DGGE, denaturing gradient gel electrophoresis; DM, dry matter; LCPUFA, long chain PUFA; NDF, neutral detergent fiber; OM, organic matter, QPCR, quantitative PCR; rRNA, ribosomal RNA; VFA, volatile fatty acid. Back

Manuscript received 12 November 2007. Initial review completed 18 December 2007. Revision accepted 6 March 2008.


   LITERATURE CITED
TOP
 ABSTRACT
 Introduction
 Materials and Methods
 Results
 Discussion
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