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ANIMAL PRODUCTION |
* Texas Agricultural Experiment Station, P.O. Box 1658, Vernon 76385;
and
United States Department of Agriculture, Agricultural Research Service, Southern Plains Agricultural Research Center, Food and Feed Safety Research Unit, 2881 F & B Rd., College Station, TX 77845
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Abstract |
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 Top Abstract INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Key Words: bacterial diversity • frothy bloat • gas production • methane • rumen
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INTRODUCTION |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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; Min et al., 2005b). Total and soluble plant proteins have been identified as precursors to bloat on wheat pasture (Bartely et al., 1975; Min et al., 2005b). Gutierrez et al. (1963) reported that the rapid release of soluble protein into rumen environment caused the capture of ruminal gases in a low-gas permeable polysaccharide slime (referred to as biofilm) layer that promotes frothy bloat.
Considerable research on the rate of ruminal fermentation and ruminal bacterial populations (Hungate et al., 1955; Howarth et al., 1981, 1991) has documented many unique differences in bloating and nonbloating cattle fed legume forages and showing microbial differences due to frothy bloat. The potential role of ruminal microorganisms in frothy bloat in cattle grazing wheat forage has not been well established. Recently, RNA-and DNA-based approaches for characterizing ruminal bacterial diversity have been evaluated such as competitive- and real-time-PCR, and denaturing gradient gel electrophoresis (DGGE; Quwerkerk et al., 2002; Reilly et al., 2002; Hume et al., 2003). Both real-time PCR and DGGE are the methods of choice research involving large sample sizes. The DGGE used in present study allows for rapid screening of bacterial populations and visualization of PCR products representing predominant ruminal bacterial communities (Hume et al., 2003), while not being subject to selective pressures inherent to traditional medium-based culturing techniques. A series of in vitro and in vivo experiments were designed to elucidate the effect of wheat forage on the bacterial growth, biofilm complexes, ruminal fermentation end products, ruminal bacterial diversity, and bloat potential.
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MATERIALS AND METHODS |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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In the current study, M. smithii was chosen as a coculture strain because ruminal gases, particularly CH4, H2, and acetate, are related to dietary energy loss (Johnson and Johnson, 1995; Miller, 1995). Strain purity was ensured by determining single colony morphology on agar plates and by a single consistent cellular morphology in liquid cultures examined under the microscope (personal communication, J. Yanke). All isolates were inoculated from their respective long-term storage (lyophilization) vials into anaerobic basal medium in Hungate tubes for 24 h at 39 ° C. The isolates were reinoculated into anaerobic plant protein medium and incubated for 24 h at 39 ° C. At the end of the incubation period, concentrations of VFA, H2, CO2, and CH4 gases present in the headspace of the tubes were determined as described later.
Preparation of Soluble Plant Protein
Preparation, distribution, and inoculation of a basal growth medium containing soluble plant protein (referred to as plant protein medium) were carried out under a stream of CO2 or in an anaerobic hood (Coy Laboratory Products Inc., Grass Lake, MI) with a 95% CO2/5% H2 atmosphere. Total soluble plant protein was extracted from fresh wheat forage by blending 100 g of plant material in 300 mL of artificial saliva (pH 6.8; McDougall, 1948), and squeezing the extract through 4 layers of cheesecloth. The filtrate was collected and centrifuged at 16,000 x g for 20 min at 5 ° C, and the supernatant fraction was filter-sterilized by passing through a 0.45-µm syringe filter (Millipore Co., Bedford, MA). All preparation and handling procedures for plant protein medium were conducted anaerobically and filter-sterilized before addition to the autoclaved basal growth medium.
Growth of Organisms
Six individual strains, in monoculture or in coculture with M. smithii, were grown in vitro in plant protein medium. The composition of plant protein basal medium was resazurin (Eastman Kodak), 0.2 mL; clarified rumen fluid, 40 mL; mineral solutions numbers 1 and 2, 8 mL each; glucose, 0.2 g; cellobiose, 0.2 g; starch, 0.2 g; xylose, 0.2 g; trypticase peptone (BBL), 0.1 g; Na2CO3, 0.8 g; distilled water, 134 mL, in a final volume of 200 mL. Mineral solution number 1 contained 0.6% K2HPO4, and mineral solution number 2 contained 0.6% KH2PO4, 0.6% (NH4)2SO4, 1.2% NaCl, 0.25% MgSO4 7H2O, and 0.16% CaCl2. The pH was adjusted to pH 6.8 with 30% NaOH before dispensing 9 mL of basal medium into test tubes under constant CO2 gas streams. Cystein-HCl solution (2.5%) was then added (0.2 mL) to each tube after autoclaving. Pure cultures of M. smithii were grown in methanogen medium (Balch et al., 1979); cocultures of M. smithii with respective test bacteria were grown under an oxygen-free H2-CO2 (50:50, vol/vol) gas mixture in plant protein medium. Pure cultures of other strains were grown under an oxygen-free CO2 in plant protein medium (9 mL of basal growth medium together with 3.5 mL of soluble protein extracted from wheat forage, which contained 3.27% soluble-nitrogen (N) as the major N source) in Hungate tubes.
The specific growth rate, based on optical density using a spectrophotometer (Milton Roy Co., Spectronic 20D, Rochester, NY) at 600 nm, in plant protein medium was measured after 0, 2, 4, 6, 8, 12, and 24 h of incubation at 39 ° C in Hungate tubes. The specific growth rate was calculated using the following equation:
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where Z and Z0 correspond to the growth rate of rumen bacteria at times t (maximum) and t0 (time 0, start time), respectively (Stanier et al., 1976).
Experiment 2: Forages and Animals
The Texas A&M University Animal Care Committee approved the experimental protocol. Wheat pastures were fertilized before planting at a rate of 56 and 17.8 kg of N and S/ha, respectively. Wheat seed (Triticum aestivum var. Cutter) was sown on September 24, 2004, at a rate of 67 kg of seed/ha. Wheat forage biomass was collected during the experimental period by hand-plucking and was collected from the same pasture grazed by ruminally cannulated steers. Samples from wheat forage and bermudagrass hay were taken twice monthly and dried in a forced-air oven at 60 ° C for 48 h and ground (Cyclone sample mill, Udy Co., Fort Collins, CO) to pass a 1-mm sieve for CP, NDF, ADF, and IVDMD analyses.
In the in vivo experiment, 6 healthy ruminally cannulated steers (Angus x Hereford x Brangus; 375 ± 30 kg) were used to quantify biofilm complexes, ruminal microbiota profiles, and ruminal fluid protein fractions associated with frothy bloat in steers grazing winter wheat forage. Rumen samples were collected after 1 mo on a bermudagrass (Cynodon dactylon) hay diet (i.e., a nonbloat promoting diet) and served as a background sample (d 0) for each animal. Steers were then transferred with an additional 12 nontest ruminally cannulated cohorts to a wheat pasture (14.1 ha) and allowed to graze wheat forage for up to 50 d.
Rumen contents (about 500 g/steer) were collected from the 6 steers on d 30 (no bloated steers observed), 40, and 50 (bloated steers observed) for analysis of biofilm complexes, ruminal microbiota populations, and ruminal fluid protein fractions associated with frothy bloat. Rumen microbiota population analyses were conducted on d 30 and 50 only. The grazing period was from December 10, 2004, to January 30, 2005. A pure sward of vegetative stage of fresh wheat forage was managed under ad libitum access [18 kg of DM/(100 kg of BW · d)] during the experimental period (Pinchak et al., 1996). All steers were provided ad libitum access to a free choice of mineral supplement (ACCO Wheat Advantage Mineral, Minneapolis, MN) and water.
The degree of bloat was determined daily by visual observation at approximately 0800 and was classified into 4 bloat score (BS) categories: BS 0 indicated no visible distension on the left side of the animal; BS 1 indicated noticeable distension on the left side; BS 2 indicated severe distension on the left side; and BS 3 indicated severe distention on the left side and noticeable distension on the right side. The scoring system followed those of Paisley and Horn (1998) and Min et al. (2005b).
Ruminal Microbial Protein Fractions and Biofilm Measurement
Analyses of rumen content of microbial protein fractions was determined on d 0, 40, and 50. Rumen fluid was assayed using the method described by Min et al. (2005a). The biofilm complexes in all experiments were measured as described by Gutierrez et al. (1963). Rumen samples were transported to the laboratory within 30 min of collection, strained with 4 layers of cheesecloth to remove most of the large debris, and centrifuged in a microcentrifuge (Sorvall Centrifuge, Kendro Lab. Products, Heraeus, Germany) at 16,000 x g for 30 min to remove most of the bacteria, protozoa, and small debris. To the supernatant (1 mL) was added 1 mL of absolute ethanol; addition of the absolute ethanol precipitated the biofilm, a viscid material that floated to the surface of the mixture during 24 h at 5 ° C. Precipitate was harvested by centrifugation at 16,000 x g for 15 min. The biofilm fraction was dried for 24 h at 60 ° C and weighed for DM determination.
Denaturing Gradient Gel Electrophoresis for Rumen Microbial Population Analysis
Changes in predominant microbial populations were evaluated by banding patterns detected after DGGE. Amplicon melting (separation of the double-stranded DNA) domains and migration in the polyacrylamide gel, urea-formamide denaturing gradient were determined by the unique guanosine + cytosine (G+C) content, primary sequences, and interactions between associated bp (Muyzer et al., 1993). Genomic bacterial DNA was isolated from 0.5 mL of each rumen sample according to the method described in the QIAamp DNA Mini Kit (Qiagen, Valencia, CA). Concentrations of DNA were measured using a GeneQuant pro (GE Health Care Life Sciences, Piscataway, NJ).
The PCR amplifications were conducted using bacteria-specific PCR primers to conserved regions flanking the variable V3 region of 16S rDNA genes. Primers [50 pmol of each per reaction mixture; primer 2 and primer 3 (Integrated DNA Technologies Inc., Coralville, IA)] with a 40-bp G+C clamp (Sheffield et al., 1989; Muyzer et al., 1993) were mixed with Jump Start Red-Taq Ready Mix (Sigma Chemical Company, St. Louis, MO), according to the kit instructions, 250 ng of template DNA from rumen digesta of individual steers, and 5% (wt/vol) acetamide to eliminate preferential annealing (Reysenbach et al., 1992). The PCR amplifications were conducted on a PTC-200 Peltier Thermal Cycler (MJ Research Inc., Waltham, MA) with the following program: 1) denaturation at 94.9 ° C for 2 min; 2) subsequent denaturing at 94.0 ° C for 1 min; 3) annealing at 67.0 ° C for 45 s, –0.5 ° C per cycle (Wawer and Muyzer, 1995); 4) extension at 72.0 ° C for 2 min; 5) repeat steps 2 to 4 for 17 cycles; 6) denaturation at 94 ° C for 1 min; 7) annealing at 58.0 ° C for 45 s; 8) repeat steps 6 to 7 for 12 cycles; 9) extension at 72.0 ° C for 7 min; 10) 4.0 ° C final.
Denaturing gradient gel electrophoresis was run according to the method of Muyzer et al. (1993). Polyacrylamide gels (8% [vol/vol] acrylamidebisacrylamide ratio 37.5:1; BioRad Lab., Richmond, CA) were cast with a 35 to 60% urea, deionized formamide (USA Amersham Life Sciences, Cleveland, OH) gradient; 100% denaturing acrylamide was 7 M urea and 40% deionized formamide. Amplified samples were mixed with an equal volume of 2 x loading buffer [0.05% (wt/vol) bromophenol blue, 0.05% (wt/vol) xylene cyanol, and 70% (vol/vol) glycerol], and 4 µL were placed in each sample well (16-well comb). Gels were placed in a DeCode Universal Mutation Detection System (BioRad Lab.) for electrophoresis in 0.5 x TAE (20 mM Tris (pH 7.4), 10 mM sodium acetate, 0.5 M EDTA) at 59 ° C for 17 h at 60 V. Gels were stained with SYBR Green I (1:10,000 dilution; Sigma), and the fragment-pattern relatedness was determined with Molecular Analysis Fingerprinting Software, Version 1.6 (BioRad Lab., Hercules, CA) based on the Dice similarity coefficient and the un-weighted pair group method using arithmetic averages for clustering.
The Dice coefficient (values between 0 and 1) is an arithmetic determination of the degree to which banding patterns are alike (i.e., contain the same bands). Clusters (groups) are determined by sequentially comparing the patterns and the construction of a relatedness tree (dendrogram) reflecting the relative similarities. The amount of similarity is reflected by the relative closeness or grouping and is indicated by the percentage similarity coefficient bar located above each dendrogram.
Laboratory Measurements
The VFA concentrations were determined by gas chromatography (580-Gow, Mac Instr. Co., Bethlehem, PN) using the method of Hinton et al. (1990), and H2, CO2, and CH4 concentrations were measured by gas chromatography as described by Allison et al. (1992 and 2005). The concentration of CP in the plant extract and rumen fluid was determined using the Kjeldahl digestion procedure (AOAC, 1990). The NDF, ADF, and IVDMD of dried forage samples were determined using the Filter Bag Technique (Ankom Technology Co., Fairport, NY).
Statistical Analysis
Data were analyzed as a repeated-measures analysis using the MIXED procedure of SAS (SAS Inst. Inc., Cary, NC). Data are presented as least squares means and the associated SEM. The variables in Exp. 1 included in vitro rumen gas products, VFA, and specific growth rate. The model included bacterial strains, with replicates as the random effect. Variables in Exp. 2 included biofilm complexes, ruminal microbial biodiversity, and protein fractions associated with bloat in steers grazing winter wheat. The model included sampling day (d 0, 30, 40, and 50), BS (0 vs. 1), and associated interactions. Animals were the experimental units and were treated as a random effect.
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RESULTS |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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and 2. In monoculture, total gas production was greatest for R. albus (P < 0.01), intermediate for S. bovis (P < 0.05), and least for P. ruminicola, F. succinogenes, E. ruminantium, and R. flavefaciens among strains during 24-h rumen incubation. In mixed culture with M. smithii, total gas production was also greater (P < 0.01) for R. albus and S. bovis than for other bacterial strains. Growth in the presence of the methanogenic bacteria resulted in a considerable shift in fermentation gas products. As expected, R. albus (P < 0.01) and R. flavefaciens (P < 0.05) produced the most H2 among strains and supported greater (P < 0.01) CH4 production when cocultured with M. smithii than other coculture combinations.
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). In the absence of an exogenous supply of H2, the M. smithii was incapable of growth as a monoculture in the plant protein medium (data not shown).
When wheat soluble protein was added as a major fermentation substrate, E. ruminantium, R. albus, F. succinogenes, and R. flavefaciens + M. smithii produced more (P < 0.05) acetate than other strains and cocultures (Table 2). Cultures of P. ruminicola, E. ruminantium, and F. succinogenes produced more (P < 0.05) butyrate than any of the respective monocultures. Propionate production among strains was similar. When cocultured with M. smithii, E. ruminantium and F. succinogenes exhibited reduced (P < 0.05) acetate production compared with monocultures, indicating that methane was produced by methanogenesis from acetate. However, the mixed culture of R. flavefaciens and M. smithii produced more (P < 0.05) acetate than did the monoculture of R. flavefaciens (Tables 1 and 2). The acetate to propionate ratio was greater for R. albus and R. albus + M smithii than for other respective mono or simple cocultures with M. smithii (Table 2).
Bio-Film Complexes and Genomic DNA Concentrations
In vitro production of biofilm complexes varied among bacterial strains (Table 3) and animal diets in vivo (Table 4). Biofilm production was greatest (P < 0.01) for S. bovis, intermediate (P < 0.05) for R. albus and F. succinogenes, and lowest for P. ruminicola, E. ruminantium, R. flavefaciens, and M. smithii (Table 3). In mixed cultures with M. smithii, biofilm production varied among strains. Culture of E. ruminantium with M. smithii was the only mixed culture to produce more (P < 0.05) biofilm than the respective monoculture did.
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). There was no difference in biofilm production between bloated and nonbloated steers grazed at d 40 of wheat grazing. Biofilm production, however, was greater (P < 0.05) on d 50 in steers that subsequently bloated than in those that did not, with BS by grazing date interactions (P < 0.03). There was no difference in rumen DNA concentrations between bloated and nonbloated steers on d 50. Average DNA concentrations were lower on d 30 than 50 (Table 4).
Experiment 2: Chemical Compositions of Forages and Rumen Fluid
The chemical concentrations of CP, NDF, ADF, and IVDMD in bermudagrass hay were 12, 61, 32, and 67% DM, respectively (data not shown). In the winter wheat forage, forage chemical compositions of CP, NDF, ADF, and IVDMD were 28, 44, 29, and 89% DM, respectively (data not shown).
The 5 ruminal protein fractions assayed differed between sampling day and presence of bloat (Table 5). When diets were changed from bermudagrass hay to wheat forage d 0 through 50, ruminal microbial protein fractions (whole ruminal content, particulate matter, cheese-cloth filtrate, and protozoa and plant particle protein fractions) increased (P < 0.01). When steers were grazing on wheat forage, particulate matter, and bacterial protein fractions were similar between the bloated and nonbloated animals (Table 5). However, whole ruminal protein content (P < 0.01) and cheesecloth filtrate protein (P < 0.05) fractions in the rumen were greater for bloated than for nonbloated animals on d 50, suggesting that frothy bloat was related to changes in the total and soluble protein rumen pools. Bloat x sampling day interactions (P < 0.01) existed for protozoa and plant particle protein, suggesting that protozoa protein and plant protein fractions varied more with bloat severity than the other protein fractions in the rumen of steers grazing wheat.
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). Amplicon profiles for ruminal bacterial samples from steers 1, 2, 3, 4, 5, and 6 on d 30 varied from 67 to 87% similarity among animals (Figure 1). Cluster analysis of DGGE bands between d 30 and 50 showed that there were low ( < 83%) similarities between sample periods (data not shown).
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). The non-bloated group clustered from 90 to 97% similarity, with greater proportions of high-G+C-containing bacterial strains and a few low-G+C-containing strains than found in bloated animals. Mid-G+C-containing strains were similar between bloated and nonbloated animals. Overall, the bloated group exhibited less similarity among animals than nonbloated group and clustered from 86 to 89% similarity. The data collectively suggest that ruminal bacterial populations changed when steers experienced frothy bloat while grazing wheat forage. |
DISCUSSION |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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). In the current study, soluble plant protein extract from wheat forage was chosen as a suitable protein substrate for detailed studies on the potential role of microbial activities associated with frothy bloat. When cultured in vitro with soluble protein extract from wheat forage, S. bovis exhibited the greatest specific growth rate and produced more biofilm than other bacteria monocultures or simple cocultures with M. smithii, suggesting S. bovis is a high producer of biofilm in cattle grazing wheat forage. In vivo whole rumen protein content, cheesecloth filtrate protein fraction, and biofilm complexes were greater in rumen fluid from bloated steers than nonbloated steers. Two distinct ruminal bacterial populations developed between bloated and nonbloated steers. Therefore, frothy bloat appears to be associated with specific changes in microbial populations in the rumen when steers graze wheat forage.
Experiment 1: Interaction and Growth Rate of 6 Rumen Bacterial Strains
The main sources of ruminal gasses are from microbial fermentation and acidification of bicarbonate; the major components of ruminal gases are CO2 (45 to 70%) and CH4 (20 to 30%), with N2, O2, H2, and H2S as minor components (Clarke and Reid, 1974). Results from the present experiment show that R. albus and R. flavefaciens produced the most H2 among strains and supported production of CH4 when cocultured with M. smithii utilizing the H2 to reduce CO2 to CH4 (Latham and Wolin, 1977) which is also consistent with reports by Miller and Wolin (1973) and Wolin et al. (1997). However, our understanding of physical and chemical associations between these gases, ruminal microorganisms, and frothy bloat interactions is not clear, and further studies are required on ruminal gases-microorganisms-frothy bloat interactions.
Streptococcus bovis is one of the dominant microbes in the rumen, especially in animals receiving either high soluble carbohydrate diets (Dehority, 2003) or high quality fresh forage diet (Attwood and Reilly, 1996). Attwood and Reilly (1995) reported that 61% of strains isolated from cattle grazing perennial ryegrass/white clover pasture were S. bovis–like strains, whereas E. budayi, B. fibrisolvens, and P. ruminicola-like strains made up to 23, 7, and 2% of the isolated strains, respectively. In the current study, the specific growth rate was greatest for S. bovis among tested strains when cultures were grown with wheat soluble protein, indicating that S. bovis was capable of proliferating on the wheat soluble protein substrate. The role of competitive interactions among ruminal bacterial species in the frothy bloat complex is unknown at this time.
Experiment 2: Bloat Potential, BioFilm Complexes, and Rumen Microbial Biodiversity
Previous research reported that the ethanol-precipitable biofilm fractions from ruminal digesta have been shown to increase during the onset of bloat in steers fed either a high grain ration (Gutierrez et al., 1959) or in cattle grazing Ladino clover forage (Gutierrez et al., 1963). In the present experiment, polysaccharide biofilm production was greatest for S. bovis in monoculture in plant protein medium extracted from wheat forage. This is consistent with findings of Cheng et al. (1973, 1976). A probable source of biofilm precursors are cytoplasmic granules of reserve polysaccharides that frequently occur in rumen bacteria such as S. bovis (Cheng et al., 1976), R. albus (Cheng et al., 1977), Selenomonas ruminantium (Wallace, 1980), M. elsdenii (Brown et al., 1975), and mixed rumen bacterial cells from the rumen of cattle fed a high-energy diet (Cheng et al., 1973, 1976).
In vivo, biofilm production increased when steers were switched from a bermudagrass hay diet to grazing wheat forage. There was 31% increase in biofilm production during the onset of bloat at d 50 in steers grazing wheat forage. Further studies are necessary before one can accurately assess that susceptibility to bloat is linked to microbial interactions that lead to unique production of biofilm complexes.
Differences in steer rumen microbial populations were observed among animals and between bloated and nonbloated animals. The 16S rDNA DGGE technique allowed visualization of microbial diversity patterns in the rumen of steers related to frothy bloat, while not being subject to selective pressures inherent to conventional medium-based culturing techniques (Torsvik et al., 1990; Ward et al., 1990). However, one limitation of 16S gene amplification is that several products of varied G+C content and primary sequences may comigrate in the denaturing gradient, making classification of some changes in individual microbial populations difficult and potentially resulting in an erroneous indication of assortment and abundance (Wintzingerode et al., 1997). Another restriction is that the PCR products apparent on the stained gels are representative of the most abundant bacteria in the population (Muyzer et al., 1993; Murray et al., 1996).
The in vivo study reported is the first to document differences in rumen microbial diversity patterns between bloated and nonbloated steers grazing wheat forage and suggests that bloat might be associated with specific changes in predominant microflora in the reticulorumen. The changes observed in rumen bacterial species are associated with differences in G+C-containing strains between bloated and nonbloated steers and implicates a significant microbial populations correlation with frothy bloat (Fletcher and Hafez, 1960). Recently, molecular diversity of rumen bacteria has been shown to be dominated by bacteria belonging to essentially 4 phyla of low-G+C Gram positive bacteria (52 to 54%), Bacteroides types (Cytophaga Flexibacter-Bacteroides; 38 to 40%), Proteobacteria (4.7%), and Spirochaetes (2.4%; Tajima et al., 1999; Edwards et al., 2004). Further isolation and phenotypic characterization are necessary for identification of individual bacterial strains associated with bloat. In companion research quantifying ingestive grazing behavior patterns associated with bloat, Pinchak and Min (2005), found that bloated steers exhibited altered grazing activities compared with nonbloated cohorts (idling time +28%, ruminating bouts –15%, and jaw movements –20%). These alterations in grazing behavior might modify meal patterns and thereby alter frequency and quantity of fermentable substrate addition to the reticulorumen and as a result affect bacterial population growth patterns and diversity and rumen pH.
The rumen microbiota is highly responsive to changes in diet, age, feed additives, health of the host animals, and season (Ogimoto and Imai, 1981; Stewart and Bryant, 1988; Min et al., 2002). In relation to diet, corn-fed animals displayed more diverse bacterial populations than grass-legume hay diets, which were mostly contributed by Bacteroides-related phylotypes (Kocherginskaya et al., 2001). Findings from our current study show that ruminal bacterial populations from within nonbloated steers grazing wheat forage (d 30) varied from 67 to 87% similarity among animals, indicating that ruminal microbial populations were heterogeneous among study animals. This is similar to reports by Kocherginskaya et al. (2001).
This study has shown that wheat pasture bloat could be related to increased production of biofilm resulting from a switch in the rumen bacterial population resulting from changes in diet composition. Consumption of wheat forage increased biofilm production over time. Experiment 1 clearly shows that some rumen microbial strains produced more biofilm than others. Further research is required to define what kind of microorganisms (bacteria, protozoa, and fungi) promote and produce biofilm in the rumen of cattle grazing wheat pasture and the relationships of these microorganisms to seasonal patterns, forage chemical composition, DMI, and grazing behavior.
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Footnotes |
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2 Corresponding author: bpinchak{at}ag.tamu.edu
Received for publication July 25, 2005. Accepted for publication February 12, 2006.
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LITERATURE CITED |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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