CSIRO Livestock Industries, Long Pocket Laboratories, Indooroopilly, Brisbane, Qld 4068, Australia1
Department of Animal Sciences, University of Illinios, Urbana, IL, USA2
Author for correspondence: Denis O. Krause. Tel: +61 7 3214 2723. Fax: +61 7 3214 2881. e-mail: denis.krause{at}li.csiro.au
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Keywords: Fibrobacter succinogenes, eukaryotes, ecology, fibre, cellulose digestibility
Abbreviations: BCVFA, branched-chain volatile fatty acids; DDMI, digestible dry matter intake; DMD, dry matter digestibility; DMI, dry matter intake; TCC, total culturable count
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The observations that faeces may contain potentially fermentable fibre were supported by the work of Van Gylswyk (1970) , who demonstrated that nutritional inadequacy could limit ruminal fibre fermentation. Ruminococcus (one of the most fibrolytic rumen bacteria; Hespell et al., 1997
) numbers were increased when poor quality forage was supplemented with urea and branched-chain volatile fatty acids (BCVFA). There was an associated improvement in feed intake and some improvement in forage digestibility, a result which suggested that the abundance of fibrolytic bacteria in the rumen was limiting cell-wall digestibility (Van Gylswyk, 1970
). In contrast, Dehority & Tirabasso (1998)
increased the numbers of fibrolytic bacteria in the rumen by feeding a high-cellulose diet composed of purified wood cellulose. There was a 10-fold increase in the number of cellulolytic bacteria, but no significant increase in the digestion of alfalfa (lucerne) cellulose when it was placed in a nylon bag and suspended in the rumen for 24 h.
If fibre digestibility is to be improved by microbial manipulation, then two questions need to be answered. The first is whether it is the cellulolytic bacteria or the actual nature of plant cell walls that are the limiting factor to improving fibre digestion. A second, and very significant, issue is whether introduced organisms can multiply and persist at levels in the rumen that are sufficient to improve fibre digestion. To address these questions, we increased the relative abundance of highly fibrolytic Ruminococcus by daily dosing over a period of 8 d. Introduced strains of ruminal bacteria often decline rapidly after dosing (Flint et al., 1989 ; Miyagi et al., 1995
; Attwood et al., 1988
), making it difficult to measure the effect of the dosed strains on fibre digestion. We hoped that our protocol would enable us to measure fibre digestion in the presence of high numbers of dosed strains and that repeated dosing would enable inoculants to establish in the rumen.
We designed and characterized unique 16S-rRNA-based oligonucleotide probes to each of the dosed strains in order to track these organisms in the rumen. In addition, we used higher-level probes to Ruminococcus, Fibrobacter and eukaryotes to explore the effects of dosing on these populations. Various physiological parameters of fibre digestion [in situ nylon bag digestibility, whole-tract dry matter digestibility (DMD), dry matter intake (DMI) and digestible dry matter intake (DDMI)] were used to measure fibrolytic activity in the rumen.
![]() |
METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Housing and feeding
Experiment 1.
Adult Merino sheep (3035 kg) were maintained in individual metabolism crates in a containment animal house. Six sheep divided into dosing and control groups were housed in metabolism cages in separate rooms. The two groups were completely isolated from one another (separate feed and water supply and ventilation system) to prevent contamination of controls by dosed sheep. Dosing and sample collection was always carried out on control animals first, followed by the dosed animals.
In situ nylon bag DMD was only measured in experiment 1 and during the last 24 h of each of the pre-dosing, dosing and post-dosing periods. Air-dried rhodes grass was ground through a 2 mm mesh screen and 3 g was weighed into a Dacron bag (19x9·5 cm, pore size 50 µm). A single steel marble was inserted and the bag tied off before being suspended in the rumen. Bags were removed from the rumen and washed together with the control bag (not suspended in the rumen) in water while gently squeezing until no colour was visible in the wash. Bags were then dried at 65 °C and in situ nylon bag digestibility determined by difference.
Experiment 2.
Ten adult cannulated sheep were used in the control and dosed groups. The number of animals was greater than that in experiment 1 to increase the possibility of obtaining statistically significant results for DMI, DMD and DDMI. Rotary feeders were not used and animals received feed ad libitum. Diets were identical to those in experiment 1.
Sheep were fitted with rubber rings glued to wool around the anus to allow attachment of faecal collection bags (Raabe, 1968 ). Total faecal collections were made for 10 d, after a training period of 2 weeks to accustom sheep to the diet and daily faecal collection procedures. There were three intake and collection periods of 10 d each, designated pre-dosing, dosing and post-dosing. During the dosing period, sheep were dosed daily (500 ml of dose) through the rumen fistula, and feed was immediately offered to provide 150200 g in excess of the amount eaten during the previous day. Calculations of DDMI and DMI were made during the three periods and data from days 1 and 2 of each period were excluded to allow for adaptation to treatment. Total faecal output was collected from each sheep for each period, dried at 65 °C for 48 h and weighed. DMI was calculated as the total intake of rhodes grass corrected for feed refusal and wastage.
Medium composition.
Basal medium composition was (per litre): 150 ml clarified rumen fluid, 150 ml mineral solution A (contents per 100 ml: 3 g K2HPO4 . 3H2O), 150 ml minerals solution B (contents per 100 ml: 3 g KH2PO4, 6 g (NH4)2SO4, 6 g NaCl, 1·23 g MgSO4 . 7H2O, 1·58 g CaCl2 . 2H2O), 2 ml trace mineral salts [contents per 100 ml: 0·5 mg ZnSO4 . 7H2O, 0·15 mg MnCl2 . 4H2O, 1·5 mg H3BO3, 1·0 mg CoCl2 . 6H2O, 0·05 mg CaCl2 . 2H2O, 0·1 mg NiCl2 . H2O, 0·15 mg Na2MO4 . 2H2O, 7·5 mg FeCl2 . 4H2O), 3·1 ml volatile fatty acid solution (contents per 100 ml: 0·68 ml acetic acid, 0·3 ml propionic acid, 0·18 ml butyric acid, 0·05 ml isobutyric acid, 0·06 ml methylbutyric acid, 0·06 ml valeric acid, 0·06 ml isovaleric acid, 0·1 g phenylacetic acid), 1 g L-cysteine . HCl and 0·01% resazurin. Media were prepared anaerobically according to the methods of Hungate (1950) as modified by Bryant (1972)
. The anaerobic gas was a 95% CO2:5% H2 mix, and 4 g Na2CO3 l-1 was included to buffer the medium at pH 6·7. Aliquots (9 ml) of anaerobically prepared medium were dispensed into 25 ml Balch tubes (18 mmx250 mm) inside the anaerobic cabinet, stoppered and autoclaved for 15 min at 100 kPa. Medium for enumeration of total culturable counts (TCC) contained in addition to the basal medium (per litre): 150 ml clarified rumen fluid (total of 300 ml), 20 ml DL-lactic acid (10%, v/v), 0·4 g Casitone, 0·4 g cellobiose, 0·4 g soluble starch, 0·4 g maltose, 0·4 g birchwood xylan and 2·0 g agar. Anaerobic diluent was made up as described previously (Mackie & Wilkens, 1988
).
Cultures used for dosing.
Strains of Ruminococcus albus (SY3 and AR67) and Ruminococcus flavefaciens (Y1, LP9155 and AR72) were selected for dosing based on their superior ability to degrade dry matter and neutral detergent fibre of two tropical grasses [rhodes grass and spear grass (Heteropogon contortus)] and a temperate legume [lucerne (Medicago sativa)]. These were laboratory strains that had been in culture for at least 3 years. Their ability to degrade purified cellulose was also evaluated (Krause et al., 1999a ).
Preparation of Ruminococcus spp. for dosing.
Ruminococcus strains used for dosing were maintained as axenic cultures on basal medium plus 50 mg rhodes grass (in 10 ml medium) for at least 2 weeks to ensure that they were growing well on the same complex carbohydrate sources as fed to sheep. Medium for dosing was made up anaerobically in 2 l bottles and contained basal medium plus 10 g rhodes grass. In experiment 1, Ruminococcus strains were inoculated into the same 2 l medium bottle in an anaerobic chamber and were allowed to incubate at 39 °C for 24 h, at which time 0·1% cellobiose was added to the medium. Bacteria were allowed to grow for an additional 24 h before dosing to sheep. Growth before the addition of cellobiose ensured that rhodes grass would be utilized, while growth on cellobiose increased cell yield. In experiment 2, the individual Ruminococcus strains were always grown separately, and were mixed together in an anaerobic cabinet immediately prior to dosing.
Sheep (experiments 1 and 2) were dosed with 500 ml medium containing the designated Ruminococcus strains. Control sheep received 500 ml fresh uninoculated medium. Direct microscopic counts indicated that Ruminococcus grew to approximately 1x1010 cells ml-1 (estimated from direct microscopic counts of several doses), so that each animal received approximately 5x1012 cells. Sheep were dosed consecutively for 9 d, 1 h after the morning feeding. Representative rumen digesta samples (approx. 100 g) were taken with a stomach tube (20 mm diameter) immediately prior to dosing, placed on ice and transported to the laboratory for further processing (only experiment 1).
Enumeration of TCC.
Ten grams of rumen digesta was weighed out into a 300 ml beaker and diluted (1:10) with chilled anaerobic diluent. This mixture was blended for 1 min (Bamix) and serially diluted to the 10-10 dilution (Mackie & Wilkens, 1988 ). Droplets (20 µl) were pipetted onto TCC plates in an anaerobic chamber from the 10-5 to 10-9 dilutions. Plates were incubated for approximately 48 h before colonies were counted.
RNA extraction.
RNA was extracted according to the procedure of Stahl et al. (1988) with some modifications. A 1 ml subsample of crude rumen digesta was taken with a wide-bore pipette (5 mm) so that sufficient plant material was included in the sample. The sample was pipetted into a 2 ml screw-cap tube containing 0·5 g zirconium beads (75200 µm diameter). The tube was centrifuged at 10000 g for 1 min to pellet digesta, the supernatant was discarded and 700 µl phenol/chloroform (4:1, pH 5·1) was added. Mechanical disruption of microbial biomass was done by bead-beating (Biospec) for 5 min. The nucleic acid was precipitated with a one-tenth volume of sodium acetate (3 M), resuspended in RNase-free water and incubated at 39 °C with 1 µg ml-1 (final concentration) RNase-free DNase (Promega) to remove contaminating RNA. The RNA concentration was measured spectrophotometrically at 260 nm and adjusted to a final concentration of 100 ng µl-1. At least two subsamples from each sample of ruminal digesta were extracted and pooled.
Probe hybridization protocol.
Oligonucleotides were labelled with digoxigenin (Roche Diagnostics) and analysis was carried out as previously described (Krause & Russell, 1996 ). RNA was denatured by incubation for 10 min at 25 °C with 3 vols 2% glutaraldehyde. A sample volume equivalent to 1 µg per slot was diluted with 0·0002% bromophenol blue and 1 µg polyadenylic acid ml-1 before application to positively charged nylon membranes (Roche Diagnostics). The membranes were baked at 120 °C for 30 min to covalently cross-link the rRNA to the membrane. Prehybridization solution [25% formamide, 5x SSC (1x SSC: 0·15 M sodium chloride and 0·015 sodium citrate), 50 mM Na2HPO4, 2% blocking reagent (Roche Diagnostics), 2% SDS, 0·1% N-lauroylsarcosine] was incubated with membranes for at least 2 h before the addition of the labelled probe (10 ng ml-1). Probes were allowed to hybridize overnight and membranes were then washed at the appropriate stringency for each probe with 1x SSC. Membranes were subsequently processed according to the manufacturers instructions (Roche Diagnostics). Total 16S rRNA was determined by hybridization with a universal eubacterial probe. To prevent contamination, pre-dosing samples from the dosed sheep and all the samples from the control sheep were blotted onto separate membranes. Blotting was done in duplicate, with each duplicate blotted onto a separate membrane. A dilution series of the reference organism was included on the membrane with the samples or alternatively the dilution series was done on a membrane with only selected samples and then cross-referenced to the master membrane.
Optimization of wash temperature.
Denatured RNA samples (1000 ng) were applied by slot-blotting to positively charged nylon membranes (Roche Diagnostics) and hybridized as described above. After hybridization, the membranes were cut into individual slots and each membrane (consisting of duplicate hybridization slots) was washed in 1x SSC for at least 10 min at 34 °C. This process was repeated 12 times at increasing temperatures (34, 37, 40, 43, 46, 49, 52, 55, 58, 61, 63 and 65 °C). Each membrane was then processed as described above. The hybridization intensity (probe remaining on blot) was plotted against the wash temperature and the dissociation temperature (Td) was defined as the temperature at which 50% of the duplex remained bound.
Cross-hybridization assay.
Probes to the dosed ruminococci were synthesized and tested for specificity against a phylogenetically diverse group of ruminal bacteria. These bacteria included R. flavefaciens AR71, R. flavefaciens AR72, R. flavefaciens R13e2, R. flavefaciens LP-9155, R. flavefaciens RF1Ba, R. flavefaciens R1-addax, R. flavefaciens C14-addax, R. flavefaciens AR69, R. flavefaciens B146, R. flavefaciens AR46, R. flavefaciens AR47, R. flavefaciens Y1, R. flavefaciens FD-1, R. albus B199, R. albus Ra8, R. albus AR67, R. albus SY3, Ruminococcus callidus ATCC 27760, Ruminococcus bromii ATCC 27255, Butyrivibrio fibrisolvens OB156, B. fibrisolvens H17c, Escherichia coli K-12, Eubacterium cellulosolvens 5494, Eubacterium ruminantium GA195, Fibrobacter succinogenes S85, Lactobacillus vitulinus B26, Megasphaera elsdenii B159, Prevotella ruminicola 23, Prevotella ruminicola GA33, Ruminobacter amylophilus 70, Selenomonas ruminantium HD4, Streptococcus bovis JB1, Streptococcus bovis K11-21-09, Succinimonas amylolytica B24, Succinivibrio dextrinosolvens 22B, Treponema bryantii B25.
Statistical analysis.
Experiments followed a repeated-measures factorial arrangement of treatments (dosed animal group and one control group) and time (levels) (Littell et al., 1998 ). Total sums of squares were partitioned between the sums of squares for treatments and time (level). Only F-values for model effects with an alpha-value greater than 0·1 were considered significant. The standard error of the mean (SEM) was computed for each analysis. All statistical analyses were carried out with Statistica 5.0 (StatSoft, Tulsa, USA).
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
|
|
One-quarter dilutions were made of 1000 ng rRNA extracted from strains SY3, AR67, Y1, LP9155 and AR72, and blotted onto nylon membranes. Hybridization with probes to SY3, AR67, Y1, LP9155 and AR72 indicated that as little as 0·9 ng 16S rRNA could be detected (Fig. 2). This value constituted approximately 106 cells ml-1 and represented the probe detection limit.
|
|
During the dosing period, TCC increased significantly (P<0·05) from approximately 7x109 to 1·6x1010 ml-1 (Fig. 4). This was an increase of 128% over that during the pre-dosing period. The increase in TCC was in general agreement with that of Fibrobacter and eukaryotic populations (Fig. 3c
), which increased by approximately 100%. In contrast, the Ruminococcus increased by almost 200% (Fig. 3c
).
|
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Probes to SY3, AR67 and Y1 performed well and there was very little cross-hybridization with non-target strains. In comparison, probes to LP9155 and AR72 cross-reacted and probe design relied, in some cases, on only 1 bp mismatches with non-target strains. A possible means of increasing probe specificity is the use of peptide nucleic acids (PNA), in which the sugar-phosphate backbone is replaced by peptide moieties (Nielsen, 1999 ; Von Wintzingerode et al., 2000
). The stability of the PNADNA duplex is considerably greater than that of the RNADNA duplex (Nielsen, 1999
).
Results from individual strain probes (Fig. 3a, b
) showed that dosed organisms declined post-dosing, which is typical of dosing studies (Flint et al., 1989
; Miyagi et al., 1995
; Attwood et al., 1988
), illustrating that ecological principles governing the persistence of bacterial inoculants in complex microbial communities are not well understood. It is, however, apparent that persistence is probably a consequence of community-level reproductive strategies in which community processes limit the relative abundance of individual organisms (Caldwell et al., 1997
). These strategies are based on the evolution of cooperative networks of micro-organisms in which some members cleave specific bonds, others utilize particular substrates and still others produce inhibitors (Caldwell et al., 1997
). A ruminal example is the production of cellodextrins by cellulolytic bacteria (Russell, 1985
), which are utilized by non-structural carbohydrate-fermenting bacteria (NSC). The NSC in turn produce ammonia and BCVFA that are consumed by cellulolytic bacteria (Miller & Wolin, 1979
; Wolin & Miller, 1988
). It is unlikely that the inability to produce cellodextins or the lack of a requirement for BCVFA are the reason that the dosed strains did not persist. However, the long time that these strains have been in culture, inhibition by bacteriocin-producing organisms (Odenyo et al., 1994a
, b
) and protozoal predation (Sharp et al., 1994
) may be significant factors in thwarting their successful reproduction in the rumen.
Establishment of an exogenous organism in the rumen is complicated by the fact that the rumen is not a closed ecosystem and micro-organisms are continually entering, making it a dynamic system that significantly impacts on the persistence of the inoculant. This was demonstrated by Varel et al. (1995) who removed the rumen contents from three cows and replaced it with 20 l medium buffer and 6 l Clostridium longisporum. At the initiation of the experiment, C. longisporum was the predominant cellulolytic bacterium, but it decreased to below the detection limit after 48 h. In similar experiments in our laboratory (D. O. Krause & C. S. McSweeney, unpublished) we grew R. albus AR67 in 20 l fermenters containing a nutrient medium and rhodes grass. The rumen contents of cannulated cattle were completely removed and replaced by the fermenter contents. Animals were immediately allowed to consume a rhodes grass diet, but we found that gastric stasis was often the consequence and the dosed strain declined rapidly. The reasons for the decline are not known, but the predatory behaviour of protozoa may be significant (Coleman & Sandford, 1979
; Sharp et al., 1994
; Newbold & Hillman, 1990
).
Recent data suggest that dosing of ruminococci while the rumen is still immature does not allow the establishment of the introduced strains of bacteria (Krause et al., 1999c ). When rRNA from the rumen of lambs was hybridized with a probe to the small-subunit rDNA of eukaryotes, there was a significant increase in the relative abundance of eukaryotic rRNA in the dosed groups, implying that protozoal predation might have a significant effect on persistence. In the current study, we observed an increase in the eukaryotic population during dosing to approximately 16% of the 16S rRNA population (Fig. 3c
). In the non-dosed animals, eukaryotes ranged between approximately 3 and 9% (Fig. 3c
). Direct observations of protozoal predation of the dosed strains in this study were not made, but in vitro experiments have shown that when a recombinant Lactobacillus plantarum was mixed with rumen fluid containing protozoa, the rate of decline of the recombinant bacterium was far greater than when the protozoa were absent (Sharp et al., 1994
).
The increase in the eukaryote population may not have been specifically a predation response, but may simply have been a response to the nutrients present in the dose. The bacterial dose contained nitrogen (from bacteria) as well as fermentation acids, which have previously been shown to benefit ruminal fibre fermentation (Van Gylswyk, 1970 ). Lin et al. (1996)
, using a eukaryotic signature sequence (S-D-Euca-0502-a-A-16), probed rRNA extracted from rumen contents of animals consuming diets which differed in the proportion of forage and concentrate. When animals were on a 100% forage diet, the eukaryotic population varied between 16·9 and 18·8% of the total population. If concentrate was included in the diet (at least 40%), the eukaryotic population tended to decline.
The probe used to measure the increase in the eukaryote population will hybridize to rRNA from fungi as well as protozoa. Fungi are important inhabitants of the rumen and may make a significant contribution to fibre digestion (Hespell et al., 1997 ). In a previous study (Krause et al., 1999c
), lambs were dosed with these same strains and we measured the fungal population separately from the eukaryotic populations. These data indicated that the fungal response to the dosing regimen was small in comparison to the total eukaryotic response. Future studies should design and validate 16S-like rRNA probes to ruminal protozoa so that this possibility can be examined specifically.
In dosed animals, the Ruminococcus population was as high as 6·5% and in the undosed group varied between 1·2 and 4·0% of the total bacterial population (Fig. 3c). F. succinogenes increased to 4·7% in dosed animals and was low as 0·9% of the bacterial population in control animals (Fig. 3c
). Previous investigations with S-G-RumIV-0132-a-A-17 and S-S-Fsucc-0650-a-A-20 have shown that Ruminococcus and F. succinogenes vary as a proportion of the bacterial population depending on the diet. When animals were fed a diet of 100% rhodes grass, Ruminococcus was 1·31·9% of the population and Fibrobacter 0·82·7% (Krause et al., 1999b
). In contrast, if sheep were fed a diet of 30% Calliandra calothyrsus (tannin-rich) and 70% rhodes grass, Ruminococcus and Fibrobacter were less than 2% of the bacterial population (McSweeney et al. 2001
). When these same animals were placed on a diet of 70% rhodes grass and 30% lucerne (Medicago sativa) the populations rose to approximately 6% for both Ruminococcus and Fibrobacter. Lin et al. (1994)
, using S-S-Fsucc-0650-a-A-20, observed that F. succinogenes varied between approximately 0·5 and 6% of the population depending on the animal and diet consumed.
When cellulolytic bacteria are grown together in diculture, cellulose degradation is often below that of the pure culture alone (Dehority & Scott, 1967 ; Dehority, 1973
), which is probably the result of competitive and non-competitive interactions between cellulolytic bacteria. Shi et al. (1997)
demonstrated that cell numbers of individual species were approximately equal in cellulose-excess dicultures of R. albus plus R. flavefaciens, R. albus plus F. succinogenes, and R. flavefaciens plus F. succinogenes. However, when cellulose was limiting, R. flavefaciens>R. albus, R. flavefaciens>F. succinogenes, and F. succinogenes>R. albus. These competitive outcomes were likely the result of the superior ability of R. flavefaciens to adhere to cellulose (Shi & Weimer, 1996
). It is interesting to note that R. albus survived under cellulose-limited conditions. This was probably a combination of its ability to utilize glucose (R. flavefaciens does not; Helaszek & White, 1991
), to grow at low concentrations of cellobiose (Shi & Weimer, 1997
) and to produce bacteriocins (Odenyo et al., 1994a
).
A fuller understanding of how bacteria survive as members of consortia or cooperative networks is critical if we wish to advance the field of ecosystem biomodification. In relation to fibre degradation, the issue of bacteriocin production by certain cellulolytic bacteria has only recently being explored and is likely to be an essential component in the formation of cooperative microbial networks. R. albus strains can produce bacteriocin-like substances that inhibit the growth of R. flavefaciens but not of F. succinogenes (Odenyo et al., 1994a , b
). There also appears to be an unusually high incidence of bacteriocin-like activity among Butyrivibrio isolates and butyrivibriocin has been isolated from B. fibrisolvens AR10 (Kalmokoff & Teather, 1997
). How the ability to produce bacteriocins or resistance to bacteriocins are involved in the establishment and persistence of dosed ruminal bacteria is not known, but these compounds are likely to have important ecological consequences.
There were no significant improvements in DMI, DMD or DDMI (Tables 3 and 4
). These results demonstrate that increasing the numbers of cellulolytic bacteria in the rumen to the extent that fibre digestion is improved is very difficult. In the dosing protocol used, we hoped that the microbial population would be perturbed to a sufficient extent to allow the introduced bacteria to establish and multiply. This proved not to be the case as shown by the molecular ecology measurements (Fig. 3
). These data are confirmed by those of Dehority & Tirabasso (1998)
who could not demonstrate any improvement in the proportion of cellulose digested with a 10-fold increase in the number of cellulolytic bacteria in the rumen.
It is clear from these studies that an improvement in fibre digestion in vivo is not a foregone conclusion simply because the dosed strains have been maintained at elevated levels in the rumen. The strains used for inoculation were selected using in vitro criteria and it is not known if these strains could be classified as superior in vivo. For this to be done in situ, techniques for strain evaluation would have to be developed and a functional genomic approach could be taken in which the levels of expression of key enzymes are monitored. It is also known that fibrolytic strains can undergo subtle changes in phenotype because of repeated transfer under laboratory conditions. It is likely that key elements are lost from the strains and many of these could be critical for the ability of strains to colonize and persist in vivo. Future studies should identify and evaluate these elements.
![]() |
ACKNOWLEDGEMENTS |
---|
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Amann, R. I., Krumholz, L. & Stahl, D. A. (1990). Fluorescent-oligonucleotide probing of whole cells for determinative, phylogenetic, and environmental studies in microbiology. J Bacteriol 172, 762-770.[Medline]
Attwood, G. T., Lockington, R. A., Xue, G. P. & Brooker, G. P. (1988). Use of a unique gene sequence as a probe to enumerate a strain of Bacteroides ruminicola introduced into the rumen. Appl Environ Microbiol 54, 534-539.[Medline]
Beever, D. E., Coelho da Silva, J. F., Prescott, J. H. D. & Armstrong, D. G. (1972). The effect in sheep of physical form and stage of growth on the sites of digestion of a dried grass. 1. Sites of digestion of organic matter, energy and carbohydrate. Br J Nutr 28, 347-356.[Medline]
Bryant, M. P. (1972). Commentary on the Hungate technique for culture of anaerobic bacteria. Am J Clin Nutr 25, 1324-1328.[Medline]
Caldwell, D. E., Wolfaardt, G. M., Korber, D. R. & Lawrence, J. R. (1997). Do bacterial communities transcend Darwinism? In Advances in Microbial Ecology , pp. 105-191. Edited by J. W. Jones. New York:Plenum.
Coleman, G. S. & Sandford, D. C. (1979). The engulfment and digestion of mixed rumen bacterial species by single and mixed species of rumen ciliate protozoa grown in vivo. J Agric Sci 92, 729-742.
Dehority, B. A. (1973). Hemicellulose degradation by rumen bacteria. Fed Proc 32, 1819-1824.[Medline]
Dehority, B. A. & Scott, H. W. (1967). Extent of cellulose and hemicellulose digestion in various forages by pure cultures of cellulolytic rumen bacteria. J Dairy Sci 50, 1136-1141.
Dehority, B. A. & Tirabasso, P. A. (1998). Effect of ruminal cellulolytic bacterial concentrations on in situ digestion of forage cellulose. J Anim Sci 76, 2905-2911.
Flint, H. J., Bisset, J. & Webb, J. (1989). Use of antibiotic resistance mutations to track strains of obligately anaerobic bacteria introduced into the rumen of sheep. J Appl Bacteriol 67, 177-183.[Medline]
Forsberg, C. W., Cheng, K. J. & White, B. A. (1997). Polysaccharide degradation in the rumen and large intestine. In Gastrointestinal Microbiology , pp. 319-379. Edited by R. I. Mackie & B. A. White. New York:Chapman & Hall.
Fox, G. F., Wisotzkey, J. D. & Jurtshuk, J. P. (1992). How close is close: 16S rRNA sequence identity may not be sufficient to guarantee species identity. Int J Syst Bacteriol 42, 166-170.[Abstract]
Helaszek, C. T. & White, B. A. (1991). Cellobiose uptake and metabolism by Ruminococcus flavefaciens. Appl Environ Microbiol 57, 64-68.[Medline]
Hespell, R. B., Akin, D. E. & Dehority, B. A. (1997). Bacteria, fungi, and protozoa of the rumen. In Gastrointestinal Microbiology , pp. 59-141. Edited by R. I. Mackie, B. A. White & R. E. Isaacson. New York:Chapman & Hall.
Hicks, R. E., Amann, R. I. & Stahl, D. A. (1992). Dual staining of natural bacterioplankton with 4',6-diamidino-2-phenylindole and fluorescent oligonucleotide probes targeting kingdom-level 16S-rRNA sequences. Appl Environ Microbiol 58, 2158-2163.[Abstract]
Hungate, R. E. (1950). The anaerobic mesophilic cellulolytic bacteria. Bacteriol Rev 14, 1-49.
Hungate, R. E. (1984). Microbes of nutritional importance in the alimentary tract. Proc Nutr Soc 43, 1-11.[Medline]
Kalmokoff, M. L. & Teather, R. M. (1997). Isolation and characterization of a bacteriocin (Butyrivibriocin AR10) from the ruminal anaerobe Butyrivibrio fibrisolvens AR10: evidence in support of the widespread occurrence of bacteriocin-like activity among ruminal isolates of B. fibrisolvens. Appl Environ Microbiol 63, 394-402.[Abstract]
Krause, D. O. & Russell, J. B. (1996). An rRNA approach for assessing the role of obligate amino acid-fermenting bacteria in ruminal amino acid deamination. Appl Environ Microbiol 62, 815-821.[Abstract]
Krause, D. O., Bunch, R. J., Smith, J. M. & McSweeney, C. S. (1999a). Diversity of Ruminococcus strains: a survey of genetic polymorphisms and plant digesting ability. J Appl Bacteriol 86, 487-495.
Krause, D. O., Dalrymple, B. P., Smith, W. J., Mackie, R. I. & McSweeney, C. S. (1999b). 16S rDNA sequencing of Ruminococcus albus and Ruminococcus flavefaciens: design of a signature probe and its application in adult sheep. Microbiology 145, 1797-1807.[Abstract]
Krause, D. O., Smith, W. J. M., Ryan, F. M. E., Mackie, R. I. & McSweeney, C. S. (1999c). Use of 16S-rRNA based techniques to investigate the ecological succession of microbial populations in the immature lamb rumen: tracking of a specific strain of inoculated Ruminococcus and interactions with other microbial populations in vivo. Microb Ecol 38, 365-376.[Medline]
Lin, C., Flesher, B., Capman, W. C., Amann, R. I. & Stahl, D. A. (1994). Taxon specific hybridization probes for fibre-digesting bacteria suggest novel gut-associated Fibrobacter. Syst Appl Microbiol 17, 418-424.
Lin, C., Raskin, L. & Stahl, D. A. (1996). Microbial community structure in gastrointestinal tracts of domestic animals: comparative analysis using rRNA-targeted oligonucleotide probes. FEMS Microbiol Ecol 22, 281-294.
Littell, R. C., Henry, P. R. & Ammerman, C. B. (1998). Statistical analysis of repeated measures data using SAS procedures. J Anim Sci 76, 1216-1231.
Mackie, R. I. & Wilkens, C. A. (1988). Enumeration of anaerobic bacterial microflora of the equine gastrointestinal tract. Appl Environ Microbiol 54, 2155-2160.[Medline]
McSweeney, C. S. (1989). Cannulation of the rumen in cattle and buffaloes. Aust Vet J 66, 266-268.[Medline]
McSweeney, C. S., Palmer, B., Bunch, R. & Krause, D. O. (2001). Effect of the tropical forage calliandra on microbial protein synthesis and ecology in the rumen. J Appl Microbiol 90, 78-88.[Medline]
Miller, T. L. & Wolin, M. J. (1979). Fermentations by saccharolytic intestinal bacteria. Am J Clin Nutr 32, 164-172.[Abstract]
Miyagi, T., Kaneichi, K., Aminov, R. I., Kobayashi, Y., Sakka, K., Hoshino, S. & Ohmiya, K. (1995). Enumeration of transconjugated Ruminococcus albus and its survival in the goat rumen ecosystem. Appl Environ Microbiol 61, 2030-2032.[Abstract]
Newbold, C. J. & Hillman, K. (1990). The effect of ciliate protozoa on the turnover of bacterial and fungal protein in the rumen of sheep. Lett Appl Microbiol 11, 100-102.
Nielsen, P. E. (1999). Applications of peptide nucleic acids. Curr Opin Biotechnol 10, 71-75.[Medline]
Odenyo, A. A., Mackie, R. I., Stahl, D. A. & White, B. A. (1994a). The use of 16S rRNA-targeted oligonucleotide probes to study competition between ruminal fibrolytic bacteria: development of probes for Ruminococcus species and evidence for bacteriocin production. Appl Environ Microbiol 60, 3688-3696.[Abstract]
Odenyo, A. A., Mackie, R. I., Stahl, D. A. & White, B. A. (1994b). The use of 16S rRNA-targeted oligonucleotide probes to study competition between ruminal fibrolytic bacteria: pure-culture studies with cellulose and alkaline peroxide-treated wheat straw. Appl Environ Microbiol 60, 3697-3703.[Abstract]
Raabe, R. (1968). An efficient method of excreta collection from caged sheep. Lab Pract 17, 217-218.[Medline]
Rijpens, N. P., Jannes, G., Van Asbroeck, M., Rossau, R. & Herman, L. M. F. (1996). Direct detection of Brucella spp. in raw milk by PCR and reverse hybridization with 16S-23S rRNA spacer probes. Appl Environ Microbiol 62, 1683-1688.[Abstract]
Russell, J. B. (1985). Fermentation of cellodextrins by cellulolytic and non-cellulolytic rumen bacteria. Appl Environ Microbiol 49, 572-576.[Medline]
Sawada, H., Takeuchi, T. & Matsuda, I. (1997). Comparative analysis of Pseudomomonas syringae pv. actinidiae and pv. phaseolicola based on phaseolotoxin-resistant ornithine carbamolytransferase gene (argK) and 16S-23S rRNA intergenic spacer sequences. Appl Environ Microbiol 63, 282-288.[Abstract]
Sharp, R., Hazlewood, G. P., Gilbert, H. J. & ODonnell, A. G. (1994). Unmodified and recombinant strains of Lactobacillus plantarum are rapidly lost from the rumen by protozoal predation. J Appl Bacteriol 76, 110-117.[Medline]
Shi, Y. & Weimer, P. J. (1996). Utilization of individual cellodextrins by three predominant ruminal cellulolytic bacteria. Appl Environ Microbiol 62, 1084-1088.[Abstract]
Shi, Y. & Weimer, P. J. (1997). Competition for cellobiose among three predominant ruminal cellulolytic bacteria under substrate-excess and substrate-limited conditions. Appl Environ Microbiol 63, 743-748.[Abstract]
Shi, Y., Odt, C. L. & Weimer, P. J. (1997). Competition for cellulose among three predominant ruminal cellulolytic bacteria. Appl Environ Microbiol 63, 734-742.[Abstract]
Stahl, D. A., Flesher, B., Mansfield, H. R. & Montgomery, L. (1988). Use of phylogenetically based hybridization probes for studies of ruminal microbial ecology. Appl Environ Microbiol 54, 1079-1084.[Medline]
Ulyatt, M. J. & MacRae, J. C. (1974). Quantitative digestion of fresh herbage by sheep. 1. The sites of digestion of organic matter, energy, readily fermentable carbohydrate, structural carbohydrate, and lipid. J Agric Sci 82, 295-307.
Van Gylswyk, N. O. (1970). The effect of supplementing a low-protein hay on the cellulolytic bacteria in the rumen of sheep and on the digestibility of cellulose and hemicellulose. J Agric Sci 74, 169-180.
Varel, V. H., Yen, J. T. & Kreikemeier, K. K. (1995). Addition of cellulolytic clostridia to the bovine rumen and pig intestinal tract. Appl Environ Microbiol 61, 1116-1119.[Abstract]
Vinuesa, P., Rademaker, J. L. W., De Bruijn, F. J. & Werner, D. (1998). Genotypic characterization of Bradyrhizobium strains nodulating endemic woody legumes of the Canary Islands by PCR-restriction fragment length polymorphism analysis of genes encoding 16S rRNA (16S rDNA) and 16S-23S rDNA intergenic spacers, repetitive extragenic palindromic PCR genomic fingerprinting, and partial 16S rDNA sequencing. Appl Environ Microbiol 64, 2096-2104.
Von Wintzingerode, F., Landt, O., Ehrlich, A. & Göbel, U. B. (2000). Peptide nucleic acid-mediated PCR clamping as a useful supplement in the determination of microbial diversity. Appl Environ Microbiol 66, 549-557.
Weimer, P. J. (1996). Why dont ruminal bacteria digest cellulose faster? J Dairy Sci 79, 1496-1502.
Wolin, M. J. & Miller, T. L. (1988). Microbe-microbe interactions. In The Rumen Microbial Ecosystem , pp. 343-359. Edited by P. N. Hobson. New York:Elsevier.
Zheng, D., Alm, E. W., Stahl, D. A. & Raskin, L. (1996). Characterization of universal small-subunit rRNA hybridization probes for quantitative molecular microbial ecology studies. Appl Environ Microbiol 62, 4504-4513.[Abstract]
Received 11 January 2001;
revised 8 March 2001;
accepted 15 March 2001.