A continuous-culture experiment was performed to investigate the effects of tall fescue (TF)-to-legume ratios (TF:legume = 75:25, 50:50, or 25:75 on a DM basis) of 3 different TF-legume mixed diets [TF-alfalfa (TF+AF), TFbirdsfoot trefoil (TF+BT), or TF-cicer milkvetch (TF+CM)] on in vitro fermentation characteristics. Nine dietary treatments were tested in a 3 (TF-legume ratio) x 3 (TF-legume mixed diet) splitplot design. Dietary treatments did not affect the concentrations of total VFA, acetate, and butyrate, whereas increasing legume proportion increased propionate concentration (P = 0.03). Regardless of TF-legume ratio, feeding TF+CM resuited in the greatest propionate concentration, whereas TF+AF and TF+BT maintained a similar concentration of propionate. The TF+AF combination resulted in a greater acetate-to-propionate ratio than TF+BT or TF+CM (P = 0.03). Decrease in ammonia-N concentration (P < 0.01) was noticed when legume proportion decreased. AmmoniaN concentration of TF+CM decreased (P < 0.01) compared with TF+AF, and it further decreased in cultures receiving TF+BT (P < 0.01). Methane production was decreased by increasing legume proportions, and the result was particularly notable in TF+BT due to increased condensed tannin concentration. In addition, TF+BT increased the proportion of C18:1 trans-11 and decreased C18:0 in the culture (P < 0.01), but no effects were detected because of legume ratio. Results of this experiment indicate that increasing the proportion of legume in combination with TF favorably shifted in vitro fermentation pathways by producing more propionate and less ammonia-N and methane.
Key words: continuous culture, in vitro fermentation, legume, tall fescue
Tall fescue [Schedonorus arundinaceus (Schreb.); TF] is the most widely planted grass species in the humid pasture region of the United States. However, the nutritive quality of TF decreases as the grazing season progresses (Noviandi et al., 2012b). Mixed grass-legume pastures may be a good pasture management option to improve the nutritive value of pastures by increasing CP concentration and digestibility and animal growth performance. Birdsfoot trefoil (Lotus comiculatus L.; BT) and cicer milkvetch (Astragalus cicer, CM) are nonbloating forage legumes that are similar to alfalfa (Medicago sativa L.; AF) in feeding value. Alfalfa contains low concentration of condensed tannins (CT), whereas BT has high concentration of CT. Although CM does not contain CT, it possesses a unique plant structure that alters microbial digestion in the rumen; the rate of microbial digestion is decreased by its thick epidermal layers and vein pattern of the leaf (Lees et ah, 1982). Birdsfoot trefoil in mixed grass-legume pastures not only increases CP concentration and total forage yield, but also increases total BW gain by steers (Wen et ah, 2002). In addition, in vitro research by our laboratory showed that TF and BT mixed diets improved nutrient utilization by decreasing ammonia-N (NH3-N) and methane (CH4) production (Noviandi et ah, 2012a).
Despite some advantages of TFlegume mixed pastures, there are still some limitations on their application to grazing cattle. One of the most significant limitations is legume's poor stand persistence. Wen et al. (2002) found that there was a rapid decline in total yield of BT during a spring to fall grazing season, which led to a 57% reduction of BT in a TF and BT mixed pasture. In contrast, observations by producers and plant geneticists indicate an increased proportion of legumes in TF-legume mixed pasture from spring to fall (May to September) in northern Utah. To our knowledge, no research has evaluated the effect of legume species in combination with TF in varying proportions on microbial fermentation in continuous-culture system. Thus, it was hypothesized that TF-legume species treatments with greater proportion of legume would improve in vitro ruminal fermentation characteristics. Additionally, CT in BT and CM resistance to microbial digestion would result in beneficial effects by reducing NH.(-N concentration and CH4 production in continuous cultures. Therefore, the objectives of this experiment were to determine in vitro ruminal fermentation profiles in response to feeding various TF-legume mixed diets and assess how the dietary treatments would affect the ruminal fermentation under varying TF-legume ratios in a continuous-culture system.
MATERIALS AND METHODS
Pasture Forages, Dietary Treatments, and Experimental Design
Forages assessed in this experiment were planted in a randomized complete block design with 4 replications on August 4, 2010, at the Utah State University Experiment Station Intermountain Irrigated Pasture Project Farm in Lewiston, Utah. Irrigation was used for establishment of the pastures and during production. Nitrogen fertilizer was applied to TF monocultures at 68 kg of N/ha in 3 applications during the growing season, and the applications were made the first week of April before plant growth and following the first (June 4) and third harvest (July 31). All pasture forages tested in the current experiment were harvested July 2, 2012. Plots were harvested to a height of 8 cm with a Swift Current sickle bar harvester (Swift Machine k Welding LTD, Swift Current, SK, Canada). This was the second harvest of the season, and the TF was in the vegetative stage, and AF, BT, and CM were at approximately 5% bloom, late bud to 1% bloom, and mid to late bud development, respectively. The nutrient composition of the forages is presented in Table 1.
After fresh forage samples were collected, they were immediately cooled, transported to the laboratory, and freeze-dried (FreeZone 12 L Freeze Dry Systems, Labconco Corp., Kansas City, MO). Forage samples for dietary treatments were ground to pass a 4.0- mm screen (Standard Model 4; Arthur H. Thomas Co., Swedesboro, NJ), and those for proximate and fatty acid analyses were ground to pass a 1.0- mm screen (Standard Model 4).
Nine treatments were randomly applied to an 8-unit, dual-flow, continuous-culture fermentor system using a split-plot design with 3 TF-legume ratios as a whole plot and 3 TF-legume species as a subplot. The continuousculture fermentors were considered as experimental units, and 3 independent runs were used as replicates (n = 3). The ratios of TF:legume were 75:25 (G75L25), 50:50 (G50L50), and 25:75 (G25L75), and TFlegume species treatment consisted of TF and AF (TF+AF), TF and BT (TF+BT), and TF and CM (TF+CM). Because in practical situations it is not possible to maintain the exact TF-legume ratios, the mixed diets were prepared by combining pasture forages from monocultures to mimic the grass-to-Iegume ratios we chose to test. The grass-to-legume ratios were based on each species' DM (Table 1).
Eight 1,000-mL, dual-flow, continuous-culture fermentors (Prism Research Glass Inc., Research Triangle Park, NC) designed according to Teat her and Sauer (1988) were used in 3 replicated periods of 8 d (5 d of adaptation and 3 d of data and sample collection). Each fermentor was inoculated with pooled ruminal fluid obtained from 3 rumen-fistulated beef cows fed a forage diet (AF and TF hay) ad libitum. Care, handling, and sampling of the donor cows were approved by the Utah State University Institutional Animal Care and Use Committee. Ruminal fluid was collected from various locations within the rumen, strained through a polyester material (PeCAP, pore size of 355 pm; B & SH Thompson, Ville MontRoyal, QC, Canada) into preheated insulated containers, and transported to the laboratory. Approximately 700 mL of strained ruminal fluid was added into each continuous-culture fermentor.
Anaerobic condition in the fermentors was maintained by infusion of C02 at a rate of 20 mL/min. Artificial saliva prepared according to Slyter et al. (1966) was continuously infused into fermentors at a rate of 0.78 mL/min using a pump (Model 323, Watson-Marlow Inc., Wilmington, MA) to maintain a fractional dilution rate of 6.3%/h. To mimic rumen motility, cultures were continuously stirred by a central paddle attached to an electric motor. Each fermentor received a total of 15 g of DM/d that was fed in 4 equal portions at 0600, 1200, 1800, and 2400 h.
All data collection, sampling, and analysis of culture content from continuous-culture fermentors were independently performed in each run. At d 6 and 7 of each run, ruminal culture pH data and 2 sets of 5-mL culture fluid samples for VFA and NH3-N analysis were collected. Culture pH was measured hourly through a pH electrode connected to a pH meter (Model 63, Jenco Instruments Inc., San Diego, CA). At 0600, 0900, 1200, 1500, and 1800 h, CH4 samples were collected from the headspace gas of each fermentor using a 10 pL gastight syringe (Hamilton Co., Reno, NV) and analyzed for CH( with a GLC (Model CP-3900, VarÍan, Walnut Creek, CA). Daily CH4 production (mili/d) was calculated as reported by Jenkins et al. (2003) using the following equation: CH( proportion in fermentor headspace (milf/mL) x C02 gas flow through the fermentor headspace (20 mL/min) x 60 min x 24 h.
Immediately after CH4 sampling at 0900 and 1500 h, 5 mL of culture contents was taken, added to 1 mL of 25% meta-phosphoric acid, and stored at -40°C for VFA determination. At the same times as VFA sample collection, another 5 mL of culture content was collected from each fermentor, mixed with 1% sulfuric acid, and stored frozen (-40°C) for NH3-N analysis.
On the final day of each run (d 8), the total volume of fermentor contents was collected and blended using a blender (Master Prep, EURO-PRO Operating LLC, Boston, MA) for 1 min. The remaining ruminal fluid was filtered through polyester material (PeCAP, pore size of 355 pm). For fatty acid analysis, approximately 150 mL of filtrate was collected, freezedried (FreeZone 12 L Freeze Dry Systems), and ground with a mortar and pestle. Another 500 mL of filtrate was centrifuged at 800 x g for 15 min at 4°C to remove solids, and then the supernatant fraction was centrifuged at 27,000 x g for 30 min at 4°C to obtain a bacterial pellet (Yang et ah, 2004). The pellets were freeze-dried and ground using a ball mill (Mixer Mill MM2000; Retsch, Haan, Germany) at 25 MHz for 4 min to a fine powder for determination of N content using an organic elemental analyzer (Flash 2000 N/Protein Analyzer, Thermo Scientific, Cambridge, UK).
Analytical DM concentration of samples was determined by oven drying at 105°C for 3 h (AOAC International, 2000; method 930.15), and OM was determined by ashing at 550°C for 5 h (AOAC International, 2000; method 942.05). Concentration of N was determined using an organic elemental analyzer (Flash 2000 N/ Protein Analyzer, Thermo Scientific). Concentrations of NDF and ADF were sequentially determined using an ANKOM200/220 Fiber Analyzer (ANKOM Technology, MacedÓn, NY) according to the methodology supplied by the company, which is based on the methods described by Van Soest et al. (1991). Sodium sulfite was used in the procedure for NDF determination and pretreated with heat-stable amylase (Type XI-A from Bacillus subtilis; Sigma-Aldrich Corporation, St. Louis, MO). Ether extract was measured (AOAC International, 2000; method 2003.05) using an AnkomXT20 Fat Analyzer (ANKOM Technology). Total extractable CT concentration in forage samples and experimental diets was determined using a butanol-HCl colorimetric procedure (Terrill et ah, 1992). Concentration of nonfiber carbohydrates (NFC) was calculated using the following formula: NFC, % = 100 - (% NDF + % CP + % ether extract + % ash).
Culture VFA were separated and quantified using a GLC (Model 6890 series II, Hewlett Packard Co., Avandale, PA) with a capillary column (30 m x 0.32 mm i.d., 1-pm phase thickness, Zebron ZB-FAAP, Phenomenex, Torrance, CA) and flame ionization detection. The oven temperature was held at 170°C for 4 min, increased to 185°C at a rate of 5°C/min, then increased by 3°C/min to 220°C and held at this temperature for 1 min. The injector and the detector temperatures were 225 and 250°C, respectively, and the carrier gas was helium (Eun and Beauchemin, 2007). Concentration of NH3-N was determined as described by Rhine et ah (1998) using a plate reader (MRXC, Dynex Technologies, Chantilly, VA).
Fatty acid extraction of diets was performed according to procedures of O'Fallon et ah (2007), whereas extraction of ruminal-fluid fatty acids was done based on a one-step mÉthylation method (Sukhija and Palmquist, 1988; Jenkins, 2010). Analysis of fatty acid methyl esters was performed using a GLC equipped with an autoinjector, autosampler, and flame ionization detector (HP 6890N, Agilent Technologies Inc, Wilmington, DE). Samples containing methyl esters in hexane (1 pL) were injected through the split injection port (25:1) onto the column (HP 88, Agilent Technologies Inc.). Oven temperature was set at 35°C and held for 2 min and then increased to 190°C at 12°C/min for 39 min. The temperature was then increased again to 218°C at 20°C/min and held for 16 min. Injector and detector were set at 250°C. Total run time was 66 min. Individual fatty acid proportions were obtained by taking the specific fatty acids area as a percentage of total fatty acids and were reported as grams per 100 g of total fatty acids. Fatty acid identification and quantification were performed using Agilent Chem Station Software (Agilent Technologies Inc.) by comparison with GLC-603 standards (Nu-Chek Prep Inc, Elysian, MN).
Data were analyzed using the MIXED procedure of SAS (SAS Institute Inc, Cary, NC) using the model described below:
Y., = y + GLR, + Run/GLR,) + MPFt + GLR, x MPFk + eijk,
where Yjjk is the individual response variable measured, y is the overall mean, GLR is the fixed effect of TFdegume ratio, Run (GLR,) is the whole plot error, MPI^. is the fixed effect of TF-legume species, GLR, x MPF, is the fixed effect of interaction k between TF-legume ratio and TFlegume species, and e..k is the subplot error.
The TF-legume ratio was compared with the whole plot error term, whereas TF-legume mixed diets and interaction between TF-legume ratio and TF-legume mixed diets were tested using the subplot error term. Comparison of TF-legume ratio and TF-legume mixed diets means were done by contrast test with Tukey's HSD test, when the effect of TFlegume ratio and TF-legume mixed diets (P < 0.10) was detected by the model. Significant effects were accepted when P < 0.05, and trends were discussed when P < 0.10.
RESULTS AND DISCUSSION
Nutrient Composition of Diets
In comparison with TF, legumes contained greater CP and less fiber (Table 1). Thus, when TF-legume diets contained a greater proportion of legume, CP concentration increased, whereas NDF and ADF concentrations decreased (Table 2). Regardless of TF-legume ratio, TF+AF and TF+CM had greater CP concentrations compared with TF+BT (averaged 14.2 and 14.3 vs. 12.7%, respectively), whereas fiber concentrations of TF+CM were the lowest (43.9% NDF and 26.9% ADF on average). The differences in nutrient composition among TF-legume diets were due to legume type; AF and CM showed similar CP concentration, but BT had the lowest concentration (averaged 19.4, 18.8 vs. 16.4%, respectively). Additionally, CM had the lowest NDF and ADF concentrations (averaged 25.9 and 20.4%, respectively). Similar CP concentrations between AF and CM with less fiber concentration in CM have been reported by others (Chang et al, 2002; Acharya et al, 2006; Williams et al, 2011). Differences in fiber concentrations between CM and the other 2 legumes (AF and BT) can be accounted for by the different leaf-to-stem ratio of the plants; CM has high leaf-to-stem ratio, which is 40% greater than that of AF (Baldridge and Lohmiller, 1990).
A noticeable CT concentration was only detected in BT among pasture forages (3.25% DM; Table 1). Consequently, dietary treatments containing BT had accountable concentrations of CT ranging from 0.91 to 2.43% (Table 2), and the CT concentration increased with increasing proportion of BT in the diet. The CT concentration in BT has been reported at 0.97 to 7.31% DM (Scharenberg et al, 2007; Williams et al, 2010; Lyman et al, 2012).
Primary fatty acids observed in all diets were C18:3n-3 followed by C16:0 and C18:2n-6 at 48.3, 20.3, and 13.4 g/100 g of total fatty acids on average, respectively. The C18:2n-6 proportion slightly increased with increasing proportion of legume, but no noticeable differences were detected in the other fatty acid profiles because of different TF-legume ratios or TF-legume species. The increased C18:2n-6 proportion in diets with greater legume proportion was related to the differences of fatty acid profiles between grass and legume; it was reported that legumes normally comprised greater amounts of C18:2n-6 than grass species (Boufaied et al, 2003; Dierking et al, 2010).
Culture pH and VFA Production
Although culture pH was affected by TF-legume ratio and TF-legume species (Table 3), all cultures maintained a pH of at least 6.10, and the largest difference in culture pH was 0.19 unit between TF+AF in the G50L50 and TF+CM in the G75L25. Therefore, biological importance of the effect of culture pH in response to dietary treatments would be minimal.
None of the dietary treatments influenced concentrations of total VFA, acetate, and butyrate. Similarly, Tavendale et al. (2005) and Williams et al. (2010) reported that there was no effect on total VFA concentration in vitro between AF and BT diets. In contrast, propionate concentration increased when fermentors were fed with a greater proportion of legumes in our experiment. The increased concentration of propionate in cultures associated with greater proportions of legumes may have resulted from NFC concentration of legumes. As shown in Table 2, the NFC concentration increased with increasing legume proportion. In this experiment, the greatest concentration of NFC was detected in CM followed by BT and AF (averaged 40.5, 39.8, and 34.6%, respectively; Table 1). Regardless of TF-legume ratio, feeding TF+CM consistently elicited increased propionate concentration compared with TF+AF and TF+BT (11.9 vs. 9.94 and 10.6 mM, respectively; P < 0.01), resulting in decreased acetateto-propionate ratio. The differences in propionate concentration due to TF-legume mixed diets was related to their NFC concentration, which was greater in TF+CM than TF+AF and TF+BT (averaged 25.1, 18.2, and 20.2%, respectively; Table 2). In an in vitro experiment using 5 different forage-based diets, Williams et al. (2011) reported that fermentors offered CM-based diet had greater propionate concentration and lower acetate-to-propionate ratio.
Within TF-legume ratios tested, G25L75 resulted in the greatest NH.,N N followed by G50L50 and G75L25 (18.9, 16.9, and 13.7 mg/100 mL on average, respectively; P < 0.01; Table 3). The increased NH3-N concentration due to greater legume proportion was associated with the greater CP concentration in the corresponding diets. As shown in Table 2, CP concentrations of G75L25, G50L50, and G25L75 increased with increasing legume proportion in the diets (averaged at 11.4, 13.1, and 16.7%, respectively). Ruminal NH3-N concentration is a result of a balance between production (proteolysis) and assimilation (De Visser et al., 1997). Thus, increased ruminal NH3-N concentration due to high legume proportion in the diets may reflect an imbalance between dietary protein degradation and ruminal capture of NH.-N for microbial protein synthesis. Although increasing legume proportion in the diets increased NFC concentration, it is likely that the increased NFC concentration would not be sufficient to incorporate the NH3-N into microbial protein. Increased NH.(-N concentration-to-VFA concentration ratio due to increased legume proportion in the diets supports the inefficient utilization of NH3-N for microbial protein synthesis. It is believed that energy is the most limiting factor in microbial growth (Bach et ah, 2005), and thus, increasing NFC as a proportion of carbohydrates typically has positive effects on carbohydrate utilization for microbial protein synthesis. However, Lykos et al. (1997) found that inclusion of NFC in the range of 35 to 42% DM was needed to increase energy density in the diets. In our experiment, the NFC concentrations were between 15 and 26% DM, and therefore, it is unlikely that the NFC concentrations would affect NH3-N utilization.
Feeding TF+BT resulted in lower NH3-N concentration compared with TF+AF and TF+CM (14.7 vs. 18.0 and 16.8 mg/100 mL on average, respectively; P < 0.01; Table 3). However, TF+BT did not consistently increase microbial N concentration. Therefore, the reduced NH3-N concentration in cultures offered TF+BT may have resulted from inhibitory effects of CT on the proteolysis of dietary soluble protein rather than an increase in microbial protein synthesis. Condensed tannins affect ruminal NH3-N concentration in 2 ways: 1) reducing dietary protein degradation via formation of insoluble tanninprotein complexes or decreasing the solubility of protein (Tanner et ah, 1994; Min et ah, 2000) and 2) inhibiting proteolytic bacteria or proteolytic enzymatic activity or both (Patra et al., 2012). Under typical cattle-feeding conditions, manipulation of ruminal protein degradation or the efficiency of N utilization in the rumen is the most effective strategy to reduce N losses (Tamminga, 1996). Using data obtained from continuous-culture studies, Bach et al. (2005) reported that as efficiency of N utilization increases, NH3-N accumulation in the fermentors decreases (R2 = 0.78). Thus, the reduction in the NH3-N concentration through CT in TF+BT can contribute to improving utilization of dietary N in ruminal fermentation and reducing N excretion.
Although TF+AF and TF+CM had similar CP concentrations (averaged 14.2 and 14.3%, respectively; Table 2), TF+CM produced less NH3-N concentration than TF+AF (averaged 16.8 vs. 18.0 mg/100 mL, respectively; Table 3). In an experiment using 6 different legumes, Lees et al. (1982) observed that CM had a thicker epidermal layer containing smaller epidermal cells, which may increase CM resistance to mechanical damage occurring during microbial digestion. Furthermore, the authors also reported that the venation pattern and vein structure in CM were very effective in restricting digestion by rumen microbes (Lees et al., 1982). In addition, Broderick and Albrecht (1997) reported that rate of ruminal CP degradation was slower in CM than AF. Thus, the slower ruminal degradation of CM may limit the availability of dietary N in the rumen, resulting in less NH,t-N concentration in the rumen.
Cultures fed TF+BT with the greatest portion of legume (G25L75) showed a lower increase of NH3-N concentration than those with G50L50 (7.29 vs. 21.8%), which resulted in an interaction between TF-legume ratio and TF-legume species treatments (Table 3). As shown in Table 2, TF+BT under G25L75 contained 2.43% CT, whereas TF+BT under G50L50 contained 1.65% CT. The results in our experiment suggest that inhibitory effects of CT on dietary protein degradation may occur when CT concentration was at 1.65% and could further decrease protein degradation at 2.43% CT. It has been reported that the minimum concentration of CT needed to decrease protein degradation was about 0.4% in in vitro studies (Aerts et al., 1999; Molan et al., 2000; Williams et al., 2010), whereas in in vivo studies the required CT concentration was between 2.0 to 3.2% (Min et al., 2002; Al-Dobaib, 2009).
Within the TF-legume ratio treatments, G75L25 showed the lowest NH.t-N:VFA ratio, whereas no difference was detected between G50L50 and G25L75. The lowest NH"-N:VFA ratio in G75L25 resulted from lower dietary concentration of CP and resultant lower NH3-N concentration in addition to lower VFA concentration, and therefore, it does not imply an improvement of dietary N utilization by G75L25.
Regardless of TF-legume ratio, fermentors fed TF+BT or TF+CM resulted in lower NH3-N:VFA ratios compared with TF+AF (averaged 0.18 vs. 0.21, respectively; P < 0.01). The low NH3-N:VFA ratios due to TF+BT and TF+CM were related to their lower NH3-N concentrations. The lower NH.,-N concentration in TF+BT may be a direct effect of CT, whereas in TF+CM it may have resulted from thick epidermal layer of CM that can improve its resistance to ruminal microbial digestion. This finding also suggests that both TF+BT and TF+CM may have a similar efficiency in N utilization in the rumen.
As the legume proportion in the diets increased, CH4 production decreased (Table 3). Methane generated in the rumen is formed primarily from hydrogen produced during the fermentation of feed, particularly high-fiber diets. The amount of CH4 produced is therefore dependent upon the amount of hexose fermented and the amount of individual VFA produced. Changing the fermentation stoichiometry to produce more propionate at the expense of acetate and butyrate typically results in less CH4 from fermentation. In this experiment, AF, BT, and CM contained greater concentrations of NFC than TF (34.6, 39.8, and 40.5 vs. 19.0%, respectively; Table 1), and consequently, a greater proportion of legume in diets led to greater propionate concentration and less CH4 production. It has been reported that soluble sugars yield less CH4 than plant fiber (Moss et al., 1995; Hindrichsen et al., 2004), and therefore, increasing concentrations of soluble sugars in forage plants can be a way of reducing ruminal CH4 production.
Cultures fed CT-containing diets (TF+BT) produced less CH4, whereas there was no difference in CH4 production between TF+AF and TF+CM. There is a body of evidence to demonstrate that feeding CTcontaining forages or supplementing CT extracts decreases CH4 production in vitro (Huang et al., 2011; Tan et al., 2011; Williams et al., 2011) and in vivo (Woodward et al., 2004; Animut et al., 2008; Sun et al., 2012). The inhibitory effects of CT on rumen methanogenesis have been attributed to direct effects of reducing methanogenic archaea (Finlay et al., 1994; Patra and Saxena, 2009, 2011) and indirect effects through a depression of fiber digestion in the rumen (Patra et al., 2012). In the current experiment, VFA concentration was similar across dietary treatments, and we did not observe any detrimental effect on ruminal fermentation. Therefore, the decreased CH4 production may have resulted from direct effects of CT on methanogenesis rather than indirect effects of depression of ruminal fiber digestion in cultures.
When cultures were fed with an increased dietary portion of BT, reduction of CH4 was more apparent, resulting in an interaction between TF-legume ratio and TF-legume species (Table 3). In this experiment, TF+BT under G25L75 contained a greater concentration of CT (2.43%) than the other diets (Table 2), but TF+BT under G50L50 had CT at 1.65%, leading to no effect on CH4 production. Eun and Min (2012) stated that reliable and distinguishable effects of CT on CH. reduction can be expected only from CT concentrations greater than 2.0% DM.
Fatty Acid Profiles of Ruminal Culture Contents
Overall, C16:0 (ranged from 9.27 to 10.1 g/100 g of total fatty acids) and C18:0 (ranged from 25.4 to 30.9 g/100 g of total fatty acids) composed major proportions of SFA (Table 4). Although C14:0 proportion decreased with decreasing proportion of legume in the diets, there was no effect on the other fatty acid profiles. It has been reported that CT inhibit ruminal biohydrogenation, thus leading to the accumulation of C18:l turns-11 (frans vaccenic acids; TVA) at the cost of C18:0 production (Khiaosa-Ard et ah, 2009; Vasta et al., 2009; Lee et al., 2010); however, it may require a minimum CT concentration at 9.35% to elicit noticeable effects of CT on the ruminal fatty acid profiles (Vasta et al., 2009). In our experiment, the greatest CT concentration in the diets was 2.43% in TF+BT under G25L75.
A greater TVA proportion was achieved when TF+BT was fed to the cultures, whereas TF+AF and TF+CM maintained a similar proportion of TVA. In contrast, feeding TF+BT decreased C18:0 proportion, but there was no effect on C18:0 proportion when TF+AF and TF+CM were fed. These results indicate that incomplete ruminal biohydrogenation may occur due to the presence of CT, particularly in the last step of the hydrogenation process. It was not expected to observe the shift in the ruminal biohydrogenation process due to a relatively low CT concentration in the current experiment. Those in the Butyrivibrio group are the most active species among the group A bacteria, which form TVA from conjugated linoleic acid or Cl8:2 frans-11 as-15 (Harfoot and Hazlewood, 1997), whereas few species of bacteria such as Fusocillus spp. and Clostridium, proteoclasticum (group B) convert TVA to C18:0 (Maia et ah, 2007; Paillard et al., 2007; Durmic et al., 2008). Min et al. (2002) reported that CT from BT reduced the proliferation of C. proteoclasticum. Durmic et al. (2008) tested the inhibitory power upon biohydrogenating bacteria in response to CT and found that the minimum dose of plants needed to inhibit the proliferation of B. fibrisolvens was much greater than the dose needed to inhibit C. proteoclasticum, suggesting that C. proteoclasticum would be more sensitive to CT than B. fibrisolvens (Durmic et ah, 2008). In our case, therefore, a relatively low CT concentration in the diets would inhibit C. proteoclasticum, resulting in increased TVA proportion without affecting overall biohydrogenation.
Mixed grass-legume pastures have been shown to improve the nutritive quality of pastures as well as animal performance. However, over competition by the grass and reduced persistence of legumes when grown together warrants the need of information on proper grass-to-legume ratios. In addition, there is a need to identify which grasses and legumes are most compatible for growing together and proper grazing management. The current in vitro experiment showed that high proportions of legume in combination with TF had beneficial effects on ruminal fermentation by producing more propionate and less CH4. The TF+BT sizably reduced NH3-N concentration and CH4 production; whereas, TF+CM increased microbial N concentration and showed similar N utilization efficiency compared with TF+BT. Therefore, TF+BT and TF+CM may have a potential to improve ruminal fermentation and nutrient utilization efficiency of grazing cattle, which can contribute to sustainable ruminant production on pasture.
This project was funded by the Western Sustainable Agriculture Research and Education Grants Program (project number: SW10-088) and the Utah State University Irrigated Pasture Grants Program. We thank C. Saunders and W. Burningham (Utah State University, Logan) for their excellent help with the continuousculture operation.
1 Approved as Journal Paper No. 8558 of the Utah Agricultural Experiment Station, Utah State University, Logan.
Acharya, S. N., J. P. Kastelic, K. A. Beauchernin, and D. F. Messenger. 2006. A review of research progress on cicer milkvetch (Astragalus cicer L.). Can. J. Plant Sei. 86:49-62.
Aerts, R. J., W. C. McNabb, A. Molan, A. Brand, T. N. Barry, and J. S. Peters. 1999. Condensed tannins from Lotus comiculatus and Lotus pedunculatus exert different effects on the in vitro rumen degradation of ribulose-l,5-bisphosphate carboxylase/oxygenase (Rubisco) protein. J. Sei. Food Agrie. 79:79-85.
Al-Dobaib, S. N. 2009. Effect of different levels of Quebracho tannin on nitrogen utilization and growth performance of Najdi sheep fed alfalfa (Medicago sativa) hay as a sole diet. Anim. Sei. J. 80:532-541.
Animut, G., R. Puchala, A. L. Goetsch, A. K. Patra, T. Sahlu, V. H. Varel, and J. Wells. 2008. Methane emission by goats consuming diets with different levels of condensed tannins from lespedeza. Anim. Feed Sei. Technol. 144:212-227.
AOAC International. 2000. Official Methods of Analysis. Vol. 1 and 2. 17th ed. AOAC Int., Gaithersburg, MD.
Bach, A., S. Calsamiglia, and M. D. Stern. 2005. Nitrogen metabolism in the rumen. J. Dairy Sei. 88:E9-E21.
Baldridge, D. E., and R. G. Lohmiller. 1990. Montana Interagency Plant Materials Handbook for Forage Production, Conservation, Reclamation, and Wildlife. Montana State Univ., Bozeman.
Boufaied, H., P. Y. Chouinard, G. F. Tremblay, H. V. Petit, R. Michaud, and G. Belanger. 2003. Fatty acids in forages. I. Factors affecting concentrations. Can. J. Anim. Sei. 83:501-511.
Broderick, G. A., and K. A. Albrecht. 1997. Ruminal in vitro degradation of protein in tannin-free and tannin-containing forage legume species. Crop Sei. 37:1884-1891.
Chang, J. S., D. E. Akin, and R. E. Calza. 2002. Effects of cicer milkvetch (Astragalus cicer) on physiology of rumen microorganisms: Protozoa, bacteria and fungi. Asianaustralas. J. Anim. Sei. 15:1147-1155.
De Visser, H., H. Valk, A. Klop, J. Van der Meulen, J. G. M. Bakker, and G. B. Huntington. 1997. Nutrient fluxes in splanchnic tissue of dairy cows: Influence of grass quality. J. Dairy Sei. 80:1666-1673.
Dierking, R. M., R. L. Kallenbach, and C. A. Roberts. 2010. Fatty acid profiles of orchardgrass, tall fescue, perennial ryegrass, and alfalfa. Crop Sei. 50:391-402.
Durmic, Z., C. S. McSweeney, G. W. Kemp, P. Hutton, R. J. Wallace, and P. E. Vercoe. 2008. Australian plants with potential to inhibit bacteria and processes involved in ruminal biohydrogenation of fatty acids. Anim. Feed Sei. Technol. 145:271-284.
Eun, J.-S., and K. A. Beauchemin. 2007. Enhancing in vitro degradation of alfalfa hay and corn silage using feed enzymes. J. Dairy Sei. 90:2839-2851.
Eun, J.-S., and B. R. Min. 2012. Emerging opportunities and challenges on exploitation of bioactive plant secondary compounds to mitigate environmental impacts by ruminants. J. Anim. Sei. 90(Suppl. 3):491. (Abstr.)
Finlay, B. J., G. Esteban, K. J. Clarke, A. G. Williams, T. M. Embley, and R. P. Hirt. 1994. Some rumen ciliates have endosymbiotic methanogens. FEMS Microbiol. Lett. 117:157-161.
Harfoot, G. C., and G. P. Hazlewood. 1997. Lipid metabolism in the rumen. Pages 382-426 in The Rumen Microbial Ecosystem. 2nd ed. P. N. Hobson and C. S. Stewart, ed. Blackie Acad. Prof., New York, NY.
Hindrichsen, I. K., H. R. Wettstein, A. Machmuller, C. R. Soliva, K. E. B. Knudsen, J. Madsen, and M. Kreuzer. 2004. Effects of feed carbohydrates with contrasting properties on rumen fermentation and methane release in vitro. Can. J. Anim. Sei. 84:265276.
Huang, X. D.. J. B. Liang, H. Y. Tan, R. Yahya, and Y. W. Ho. 2011. Effects of Leucaena condensed tannins of differing molecular weights on in vitro CH( production. Anim. Feed Sei. Technol. 166-167:373-376.
Jenkins, T. C. 2010. Technical note: Common analytical errors yielding inaccurate results during analysis of fatty acids in feed and digesta samples. J. Dairy Sei. 93:1170-1174.
Jenkins, T. C., V. Fellner, and R. K. McGuffey. 2003. Monensin by fat interactions on trails fatty acids in cultures of mixed ruminal microorganisms grown in continuous fermentors fed corn or barley. J. Dairy Sei. 86:324-330.
Khiaosa-Ard, R., S. F. Bryner, M. R. L. Scheeder, H. R. Wettstein, F. Leiber, M. Kreuzer, and C. R. Soliva. 2009. Evidence for the inhibition of the terminal step of ruminal a-linolenic acid biohydrogenation by condensed tannins. J. Dairy Sei. 92:177-188.
Lee, M. R. F., J. K. S. Tweed, A. Cookson, and M. L. Sullivan. 2010. Immunogold labelling to localize polyphenol oxidase (PPO) during wilting of red clover leaf tissue and the effect of removing cellular matrices on PPO protection of glycerol-based lipid in the rumen. J. Sei. Food Agrie. 90:503-510.
Lees, G. L., R. E. Howarth, and B. P. Goplen. 1982. Morphological characteristics of leaves from some legume forages: Relation to digestion and mechanical strength. Can. J. Bot. 60:2126-2132.
Lykos, T., G. A. Varga, and D. Casper. 1997. Varying degradation rates of total nonstructural carbohydrates: Effects on ruminal fermentation, blood metabolites, and milk production and composition in high producing Holstein cows. J. Dairy Sei. 80:3341-3355.
Lyman, T. D., F. D. Provenza, J. J. Villalba, and R. D. Wiedmeier. 2012. Phytochemical complementarities among endophyte-infected tall fescue, reed canarygrass, birdsfoot trefoil and alfalfa affect cattle foraging. Animal 6:676-682.
Maia, M. R. G., L. C. Chaudhary, L. Figueres, and R. J. Wallace. 2007. Metabolism of polyunsaturated fatty adds and their toxicity to the microflora of the rumen. Antonie van Leeuwenhoek 91:303-314.
Min, B. R., G. T. Attwood, K. Reilly, W. Sun, J. S. Peters, T. N. Barry, and W. C. McNabb. 2002. Lotus comiculatus condensed tannins decrease in vivo populations of proteolytic bacteria and affect nitrogen metabolism in the rumen of sheep. Can. J. Microbiol. 48:911-921.
Min, B. R., W. C. McNabb, T. N. Barry, and J. S. Peters. 2000. Solubilization and degradation of ribulose-l,5-bisphosphate carboxylase/ oxygenase (EC 18.104.22.168; Rubisco) protein from white clover ( Trifolium repens) and Lotus comiculatus by rumen microorganisms and the effect of condensed tannins on these processes. J. Agrie. Sei. 134:305-317.
Molan, A. L., G. C. Waghorn, B. R. Min, and W. C. McNabb. 2000. The effect of condensed tannins from seven herbages on Trichostrongylus colubriformis larval migration in vitro. Folia Parasitol. (Praha) 47:39-44.
Moss, A. R., D. I. Givens, and P. C. Garnsworthy. 1995. The effect of supplementing grass silage with barley on digestibility, in sacco degradability, rumen fermentation and methane production in sheep at two levels of intake. Anim. Feed Sei. Technol. 55:9-33.
Noviandi, C. T., M. N. McDonald, D. R. ZoBell, J.-S. Eun, M. D. Peel, and B. L. Waldron. 2012a. Effects of energy supplementation for pasture forages on in vitro ruminal fermentation in continuous cultures. J. Dairy Sei. 95(E-Suppl. 2):45. (Abstr.)
Noviandi, C. T., B. L. Waldron, J.-S. Eun, D. R. ZoBell, R. D. Stott, and M. D. Peel. 2012b. Growth performance, ruminal fermentation profiles, and carcass characteristics of beef steers grazing tall fescue without or with nitrogen fertilization. Prof. Anim. Sei. 28:519-527.
O'Fallon, J. V., J. R. Busboom, M. L. Nelson, and C. T. Gaskins. 2007. A direct method for fatty acid methyl ester synthesis: Application to wet meat tissues, oils, and feedstuffs. J. Anim. Sei. 85:1511-1521.
Paillard, D., N. McKain, M. T. Rincon, K. J. Shingfield, D. I. Givens, and R. J. Wallace. 2007. Quantification of ruminai Clostridium proteoclasticum by real-time PCR using a molecular beacon approach. J. Appl. Microbiol. 103:1251-1261.
Patra, A. K., B.-R. Min, and J. Saxena. 2012. Dietary tannins on microbial ecology of the gastrointestinal tract in ruminants. Pages 237-262 in Dietary Phytochemicals and Microbes. A. K. Patra, ed. Springer, New York, NY.
Patra, A. K., and J. Saxena. 2009. Dietary phytochemicals as rumen modifiers: A review of the effects on microbial populations. Antonie van Leeuwenhoek 96:363-375.
Patra, A. K., and J. Saxena. 2011. Exploitation of dietary tannins to improve rumen metabolism and ruminant nutrition. J. Sei. Food Agrie. 91:24-37.
Rhine, E. D., G. K. Sims, R. L. Mulvaney, and E. J. Pratt. 1998. Improving the Berthelot reaction for determining ammonium in soil extracts and water. Soil Sei. Soc. Am. J. 62:473-480.
Scharenberg, A., Y. Arrigo, A. Gutzwiller, C. R. Soliva, U. Wyss, M. Kreuzer, and F. Dohme. 2007. Palatability in sheep and in vitro nutritional value of dried and ensiled sainfoin (Onobrychis viciifolia), birdsfoot trefoil (Lotus comiculatus), and chicory (Cichorium intybus). Arch. Anim. Nutr. 61:481-496.
Slyter, L. L., M. P. Bryant, and M. J. Wolin. 1966. Effect of pH on population and fermentation in a continuously cultured rumen ecosystem. Appl. Microbiol. 14:573-578.
Sukhija, P. S., and D. L. Palmquist. 1988. Rapid method for determination of total fatty-acid content and composition of feedstuffs and feces. J. Agrie. Food Chem. 36:1202-1206.
Sun, X. Z., G. C. Waghorn, S. O. Hoskin, S. J. Harrison, S. Muetzel, and D. Pacheco. 2012. Methane emissions from sheep fed fresh brassicas (Brassica spp.) compared to perennial ryegrass (Lolium perenne). Anim. Feed Sei. Technol. 176:107-116.
Tamminga, S. 1996. A review on environmental impacts of nutritional strategies in ruminants. J. Anim. Sei. 74:3112-3124.
Tan, H. Y., C. C. Sieo, N. Abdullah, J. B. Liang, X. D. Huang, and Y. W. Ho. 2011. Effects of condensed tannins from Leucaena on methane production, rumen fermentation and populations of methanogens and protozoa in vitro. Anim. Feed Sei. Technol. 169:185-193.
Tanner, G. J., A. E. Moore, and P. J. Larkin. 1994. Proanthocyanidins inhibit hydrolysis of leaf proteins by rumen microflora in vitro. Br. J. Nutr. 71:947-958.
Tavendale, M. H., L. P. Meagher, D. Pacheco, N. Walker, G. T. Attwood, and S. Sivakumaran. 2005. Methane production from in vitro rumen incubations with Lotus pedunculatus and Medicago sativa, and effects of extractable condensed tannin fractions on methanogenesis. Anim. Feed Sei. Technol. 123:403-419.
Teather, R. M., and F. D. Sauer. 1988. A naturally compartmented rumen simulation system for the continuous culture of rumen bacteria and protozoa. J. Dairy Sei. 71:666-673.
Terrill, T. H., A. M. Rowan, G. B. Douglas, and T. N. Barry. 1992. Determination of extractable and bound condensed tannin concentrations in forage plants, protein-concentrate meals and cereal-grains. J. Sei. Food Agrie. 58:321-329.
Van Soest, P. J., J. B. Robertson, and B. A. Lewis. 1991. Methods for dietary fiber, neutral detergent fiber, and nonstarch polysaccharides in relation to animal nutrition. J. Dairy Sei. 74:3583-3597.
Vasta, V., H. P. S. Makkar, M. Mele, and A. Priolo. 2009. Ruminal biohydrogenation as affected by tannins in vitro. Br. J. Nutr. 102:82-92.
Wen, L., R. L. Kallenbach, J. E. Williams, C. A. Roberts, P. R. Beuselinck, R. L. McGraw, and H. R. Benedict. 2002. Performance of steers grazing rhizomatous and nonrhizomatous birdsfoot trefoil in pure stands and in tall fescue mixtures. J. Anim. Sei. 80:19701976.
Williams, C. M., J.-S. Eun, C. M. Dschaak, J. W. MacAdam, B. R. Min, and A. J. Young. 2010. Case study: In vitro ruminal fermentation characteristics of birdsfoot trefoil (Lotus comiculatus L.) hay in continuous cultures. Prof. Anim. Sei. 26:570-576.
Williams, C. M., J.-S. Eun, J. W. MacAdam, A. J. Young, V. Fellner, and B. R. Min. 2011. Effects of forage legumes containing condensed tannins on methane and ammonia production in continuous cultures of mixed ruminal microorganisms. Anim. Feed Sei. Technol. 166-167:364-372.
Woodward, S. L., G. C. Waghorn, and P. G. Laboyrie. 2004. Condensed tannins in birdsfoot trefoil (Lotus comiculatus) reduce methane emissions from dairy cows. N. Z. Soc. Anim. Prod. 64:160-164.
Yang, W. Z., K. A. Beauchemin, D. D. Vedres, G. R. Ghorbani, D. Colombatto, and D. P. Morgavi. 2004. Effects of direct-fed microbial supplementation on ruminal acidosis, digestibility, and bacterial protein synthesis in continuous culture. Anim. Feed Sei. Technol. 114:179-193.
C. T. Noviandi,*2 K. Neal,* J.-S. Eun,*3 M. D. Peel.f B. L. Waldron,f D. R. ZoBell,* and B. R. Min[dagger]
Department of Animal, Dairy, and Veterinary Sciences, Utah State University, Logan 84322; tForage and Range Research Laboratory, USDA-ARS, Logan, UT 84322; and [double dagger]Department of Agricultural and Environmental Sciences, Tuskegee University, Tuskegee, AL 36088
2 Current address: Department of Animal Nutrition and Feed Science, Universitas Gadjah Mada, Yogyakarta, Indonesia.
3 Corresponding author: firstname.lastname@example.org