Previous Article | Next Article 
Applied and Environmental Microbiology, July 1999, p. 3075-3083, Vol. 65, No. 7
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Isolation and Characterization of Proteolytic
Ruminal Bacteria from Sheep and Goats Fed the Tannin-Containing Shrub
Legume Calliandra calothyrsus
Christopher S.
McSweeney,1,*
Brian
Palmer,2
Rowan
Bunch,1 and
Denis O.
Krause1
CSIRO Tropical Agriculture, Long Pocket
Laboratories, Indooroopilly, 4068 Queensland,1
and CSIRO Tropical Agriculture, Davies Laboratory, Aitkenvale,
Townsville, 4814 Queensland,2 Australia
Received 8 December 1998/Accepted 29 March 1999
 |
ABSTRACT |
Tannins in forages complex with protein and reduce the availability
of nitrogen to ruminants. Ruminal bacteria that ferment protein or
peptides in the presence of tannins may benefit digestion of these
diets. Bacteria from the rumina of sheep and goats fed Calliandra
calothyrsus (3.6% N and 6% condensed tannin) were isolated on
proteinaceous agar medium overlaid with either condensed (calliandra tannin) or hydrolyzable (tannic acid) tannin. Fifteen genotypes were
identified, based on 16S ribosomal DNA-restriction fragment length
polymorphism analysis, and all were proteolytic and fermented peptides
to ammonia. Ten of the isolates grew to high optical density (OD) on
carbohydrates (glucose, cellobiose, xylose, xylan, starch, and
maltose), while the other isolates did not utilize or had low growth on
these substrates. In pure culture, representative isolates were unable
to ferment protein that was present in calliandra or had been complexed
with tannin. One isolate, Lp1284, had high protease activity (80 U), a
high specific growth rate (0.28), and a high rate of ammonia production
(734 nmol/min/ml/OD unit) on Casamino Acids and Trypticase Peptone.
Phylogenetic analysis of the 16S ribosomal DNA sequence showed that
Lp1284 was related (97.6%) to Clostridium botulinum NCTC
7273. Purified plant protein and casein also supported growth of Lp1284
and were fermented to ammonia. This is the first report of a
proteolytic, ammonia-hyperproducing bacterium from the rumen. In
conclusion, a diverse group of proteolytic and peptidolytic bacteria
were present in the rumen, but the isolates could not digest protein
that was complexed with condensed tannin.
| |
INTRODUCTION |
Ruminant production in dry tropical
regions is commonly limited by adequate protein supply to the rumen.
Supplementary protein can be made available via shrub and tree legumes,
such as Calliandra calothyrsus, that are high in protein,
produce large amounts of leaf material, and can be readily introduced
into tropical environments (1). However, these plants often
contain condensed tannins (polyphenolics) which complex with protein,
thus reducing nitrogen availability to rumen microorganisms.
Polyphenolics also inhibit growth of predominant rumen bacteria
(Fibrobacter succinogenes, Butyrivibrio
fibrisolvens, Ruminobacter amylophilus, and
Streptococcus bovis) (4, 22), but tolerance to
tannins has been demonstrated in some rumen microorganisms
(35). Strains of bacteria capable of degrading protein may
therefore proliferate in response to tannin-rich diets such as
calliandra. We examined whether proteolytic bacteria, which are present
as significant populations in the rumina of animals fed calliandra, are
able to ferment amino acids in the presence of tannins or hydrolyze
protein that is complexed with tannin.
Nutrient medium containing precipitated tannin-protein complexes has
been used previously to isolate enteric bacteria capable of degrading
these complexes (37). Brooker et al. (6) used this technique to isolate a ruminal bacterium, Streptococcus
caprinus (gallolyticus) (42), which produced
clearing zones in this medium and was tolerant of tannins.
Streptococcus gallolyticus appears to be widely distributed
to ruminants fed tannin-rich diets, but their ecological role in
digestion of these forages is yet to be defined (32). In
this study we used similar media (37) in an attempt to
identify other bacteria that may be beneficial in the digestion of
protein in tanniniferous diets.
| |
MATERIALS AND METHODS |
The anaerobic techniques of Hungate (19) as modified
by Bryant (7) were used for the growth of organisms and
preparation of media. The media were gassed with CO2, and
10-ml aliquots were dispensed into 25-ml Balch tubes (18 by 250 mm),
which were stoppered and autoclaved for 15 min at 100 kPa. B vitamins
(28) were added to each tube of medium just prior to
inoculation, and incubations were at 39°C.
Isolation procedures.
Rumen digesta (100 g) was taken from
four sheep and four goats (2-year-old male castrates, rumen fistulated)
held in metabolism crates and fed a 100% fresh-calliandra diet (3.6%
N) ad libitum. Calliandra fed to animals included leaf and stem
material from young regrowth material, which was cut from trees and
offered to the animals within an hour of harvest. The animals were
sampled 18 h after feed was offered, and digesta was transferred
immediately to an anaerobic chamber containing a gas phase of
CO2-H2 (95:5). A 5-g subsample was diluted
(1:10) with cold anaerobic diluent (29), homogenized for 1 min (Bamix, Mettlen, Switzerland), and serially diluted in diluent to a
10
9 dilution. Tannin overlay agar plates were prepared in
an anaerobic chamber as follows (i) a 2% calliandra condensed tannin
extract or tannic acid (Ajax Chemicals, Sydney, Australia) which was
filter sterilized (22-µm pore size) was poured onto brain heart
infusion (BHI) medium or rumen habitat-simulating (RHS) medium
(30) agar plates (Table 1),
(ii) the tannin solution was allowed to precipitate with protein on the
plate surface for 20 min, (iii) the tannin solution was then aspirated
off, and (iv) the plate was rinsed with diluent (37).
Droplets (20 µl) were pipetted onto dried plates in an anaerobic
chamber from the 105, 106, and
107 dilutions (29). More colonies grew on BHI
medium, and therefore isolations were made only from these plates.
Individual colonies that produced clearing zones or had distinct
morphologies but did not produce zones were picked and inoculated into
liquid BHI medium without tannin and restreaked onto BHI agar plates.
Cultures were considered to be pure after three successive isolations
from agar plates showed a single colony and cell morphological types. Cells were treated with Gram stain and examined by light microscopy. The number of proteolytic bacteria in sheep was estimated on RHS medium
(30), and proteolytic colonies were counted as those with
zones of hydrolysis after the plates were flooded with 1 M HCl.
View this table:
[in this window]
[in a new window]
|
TABLE 1.
Compositions of media used to select bacteria and for
growth and metabolism experiments under various conditions
|
|
Growth and nitrogen digestion studies in the absence of
tannin.
Growth, ammonia production, and protease activity were
determined for individual isolates grown on (i) peptide medium (PM), which contained primarily Casamino Acids (Difco) and Trypticase Peptone
(Becton Dickinson) as substrates for growth, and (ii) PM with
carbohydrates (PMC), which contained cellobiose, birchwood xylan,
xylose, starch, and maltose (Table 1). Bacteria that showed little
growth response to carbohydrates were grown on PMC containing 0.5%
glucose to simulate rumen conditions on a concentrate diet. Isolate
Lp1284 was also grown in PM that lacked yeast extract but contained
1.5% Trypticase Peptone and 1.5% Casamino Acids, or the peptides were
replaced with either 1.5% casein (BDH Laboratory Supplies) or 3%
fraction 1 leaf protein so that specific rates of ammonia production
from peptides and growth on protein could be compared with published
data (3). Fraction 1 leaf protein, which is
ribulose-1,5-bisphosphate carboxylase-oxygenase (Rubisco), was isolated
and partially purified from the leguminous plant lucerne
(Medicago sativa) by gel filtration (23, 24). PM
which lacked both yeast extract and rumen fluid was used to
definitively confirm that Lp1284 could grow on peptides without carbohydrate.
Growth was measured as change in optical density (OD) at 600 nm
(Spectronic spectrophotometer; Milton Roy Co.). Aliquots of culture
fluid were taken at regular intervals during growth for measurements of
ammonia production. The indophenol method for the determination of
ammonia as described by Chaney and Marbach (9) was used to
estimate the rate and amount of ammonia production in cultures.
Protease activity was measured during exponential growth in PM and PMC.
Cells for protease assays were separated from culture
fluid by
centrifugation (7,000 ×
g for 20 min at 4°C),
washed,
and suspended in 0.1 M Bis-Tris
[bis(2-hydroxyethyl)imino-tris(hydroxymethyl)methane
(pH 7)] and
recentrifuged as described above. The cell pellet
was resuspended in
0.01 of the original volume and disrupted twice
by ultrasonication with
a Sanophon ultrasonic disintegrator (Ultrasonic
Industries Pty. Ltd.,
Sydney, Australia) at 60 W for 10 min at
a time. Centrifuged culture
fluid was also assayed for protease
activity.
Proteolytic activities in cell-associated and extracellular fractions
of cultures were determined spectrophotometrically with
azocasein as
the substrate (
5). Control experiments were performed
by
incubating enzyme samples and the azocasein substrate separately
and by
combining these solutions at the time of acid addition
as described by
Cotta and Hespell (
13). One proteolytic enzyme
unit equalled
1 µg of azocasein digested per h at 39°C. Extracellular
proteolytic
activity was expressed per milliliter of culture fluid
per the OD of
the culture prior to centrifugation. Cell-associated
proteolytic
activity per milliliter of culture fluid was also
expressed per the OD
of the culture. Assays were performed on
centrifuged cell pellets that
had been resuspended in buffer to
the volume of the original culture.
Enzyme activities were not
expressed per milligram of microbial protein
since the media contained
large concentrations of the substrate
nitrogen, some of which
appeared to precipitate and could interfere
with analysis of pelleted
cell protein. The proteolytic ruminal
bacterium
Prevotella ruminicola B
14 was used for
comparison of protease activities, with azocasein
as the
substrate.
Volatile fatty acids, ethanol, formate, lactate, succinate, and ethanol
in the PMC culture fluid were analyzed by high-performance
liquid
chromatography with a Waters System equipped with an Aminex
HPX-87
cation-exchange column (300 by 7.8 mm) for organic acids
and a
microguard column (Bio-Rad Laboratories, Richmond, Calif.)
with a
column heater (Waters model 1122/WTC-120). Organic acids
and ethanol
were eluted with a mobile phase of 2.5% acetonitrile
in 0.2%
(vol/vol) phosphoric acid at a flow rate of 0.7 ml/min
and a column
temperature of 60°C, with UV detection at 210 nm.
All assays were
performed at least in
duplicate.
Protein digestion in the presence of tannins.
Bacterial
isolates were also screened for the ability to degrade protein that was
complexed with tannin. These isolates were grown with medium in which
substrate protein was included only as a tannin-protein complex, and
evolution of ammonia was used as an indicator of fermentation of
protein as described previously.
Tannin-protein complexes (50 mg) were added to PMC (5 ml), but Casamino
Acids, Trypticase peptone, and yeast extract were
not included so that
the only protein available was in a complexed
form. The complexes were
weighed into sterile Balch tubes, and
autoclaved medium was added
before each tube was closed with a
sterile stopper. Tannin-protein
complexes were mainly insoluble
and could not be sterilized by heat, UV
light, or ethylene oxide
gas, because each of these processes causes
changes to the tannin
molecule, or by filtration. Duplicate cultures
were each inoculated
with 0.1 ml of individual isolates which had been
grown to late
log phase on PMC. Uninoculated controls were routinely
included
in all assays to account for any microbial activity introduced
by the tannin-protein complexes, all of which were not
sterilized.
The ability to ferment protein in calliandra was also examined with
selected isolates alone or in combination with a fiber-degrading
microbe,
Ruminococcus flavefaciens AR67, and results were
compared
with results with a mixed rumen fluid inoculum. Cocultures
with
AR67 were performed because this strain degrades fiber in
calliandra,
and it is expected that the availability of plant protein
is enhanced
by the activities of polysaccharide-hydrolyzing bacteria
(
16).
Finely milled lyophilized calliandra (50 mg, 3.64% N)
was added
to the basal medium (10 ml) used for fermentation experiments
of tannin-protein complexes, but NH
4Cl was also included at
a
final concentration of 3 mM as a nitrogen source for the AR67.
Polyethylene glycol 4000 (PEG 4000; 1 mg/10 mg of calliandra)
was also
included in some tubes to counteract the effect of tannin
on protein
(see reference
23). PEG 4000 at this concentration
does not affect the rate of ammonia production in cultures where
tannin
is not present (
30a). Ammonia production was monitored
during growth for 72 to 96 h. Procedures for inoculation of
cultures
were essentially the same as those described for experiments
with
tannin-protein complexes, except triplicate assays and
uninoculated
control experiments were performed. The rumen fluid
inoculum (0.1
ml) was obtained from a steer fed a diet comprising
(70%) rhodes
grass (
Chloris gayana) and (30%) lucerne.
Rumen digesta was strained
through muslin cloth and incubated
anaerobically at 39°C for 30
min so that the larger particulate
matter would float to the surface,
and aliquots for inoculation were
taken from the fluid phase beneath
this
layer.
Tannin purification and complexes.
Total condensed tannin in
calliandra was 6% as determined by the butanol-HCl method
(46). Condensed tannin in calliandra used in these studies
was obtained by extraction with 7:3 (vol/vol) acetone-water from fresh
calliandra leaves that had been lyophilized and ground through a
1-mm-pore-size screen (46). The acetone was removed by
rotary evaporation, and the aqueous solution was washed with diethyl
ether to remove nontanniniferous material before chromatographic
purification on Sephadex LH-20 as described by Terrill et al.
(47).
Tannin-protein complexes were made as follows: calliandra tannin which
had been purified as described previously was dissolved
(3%
[wt/vol]) in 0.2 M sodium acetate buffer (pH 5) and then added
slowly
to a solution of BHI medium (2% [wt/vol] BHI, 0.2 M sodium
acetate
buffer [pH 5]), allowing tannin-protein complexes to form.
The
complexes were precipitated by centrifugation (2,000 ×
g,
10 min), washed in the same buffer, and then centrifuged
again
and washed in culture medium to remove any soluble complexes
before
a final centrifugation and lyophilization of the
pellet.
Bacterial genotyping and extraction of genomic DNA.
The
genotypic diversity and phylogeny of the isolates were determined with
restriction fragment length polymorphisms (RFLP) of 16S ribosomal DNA
(rDNA) amplified by PCR which had been digested with restriction
endonucleases (33). In addition, repetitive extragenic
palindromic sequences (REP), enterobacterial repetitive intergenic
consensus sequences (ERIC), and amplified rDNA intergenic spacer
regions between 16 and 23S rDNAs (IR) were also used for genotyping
(15, 21, 49). The REP, ERIC, and IR techniques are able to
differentiate bacteria at the subspecies level and can be used to
rapidly confirm the identity and purity of strains.
Bacterial isolates were grown on PMC, and cells were centrifuged at
high speed (10,000 ×
g, 10 min). The pellet was
resuspended
and washed in 500 µl of TE (10 mM Tris-Cl, 1 mM
Na
2-EDTA), and
DNA was extracted according to the method of
Stahl et al. (
44)
as modified by Krause et al.
(
27). DNA concentration was measured
at 260 nm and adjusted
to a final concentration of 10 ng/µl.
ERIC, REP, and IR PCR protocols.
Each 50-µl PCR mixture
included a 1/100 dilution of bacterial cells, 5 µl of 10× reaction
buffer (Bresatec, Adelaide, Australia), 2.0 mM MgCl2 (ERIC
and REP) or 3 mM MgCl2 (IR), 0.2 mM each deoxynucleotide triphosphate, 10 pmol of each primer, and 2.5 U of Taq
polymerase (Promega, Sydney, Australia). For ERIC- and REP-PCR, cycling
conditions were denaturation for 1 min at 94°C, annealing for 1 min
at 47°C (ERIC) or 40°C (REP), and a final extension at 72°C for 2 min. This cycle was repeated 30 times. PCR cycling conditions for the 16-to-23S spacer were denaturation at 94°C for 5 min for the first cycle only and, 1 min thereafter, annealing at 50°C for 1 min, extension at 72°C for 1.5 min for 30 cycles, and a final extension at
72°C for 7 min. ERIC and REP primers were as previously described (15). The 16-to-23S rDNA spacer was amplified with a
conserved 16S forward primer (5'-AAG TCG TAA CAA GGT AG/AC CGT A-3'),
and a conserved 23S reverse primer (5'-GGG TTT/G/C CCC CAT TCG G-3').
16S rDNA RFLP analysis.
The 16S rDNA was amplified from a
1/100 dilution of overnight culture with universal primers (27f and
1492r). PCR mixtures contained (per 20 µl) 5 µl of 10× PCR buffer,
0.5 µl of MgCl2 (250 mM), 1 µl of deoxynucleoside
triphosphates (10 mM), 10 pmol of each primer, 1 U of Taq
polymerase (Promega), and 0.5 µl of a 1/100 dilution of culture.
Cycling conditions were one cycle of 94°C for 5 min, 60°C for 1 min, and 72°C for 90 s, and then 31 cycles of 94°C for 1 min,
60°C for 1 min, and 72°C for 90 s. The final PCR cycle was
94°C for 1 min, 60°C for 1 min, and 72°C for 8 min. Approximately
100 ng of the 16S rDNA PCR product was digested with tetrameric
restriction enzymes (AluI, DdeI, and Sau3a) for at least 2 h according to the
manufacturer's instructions. Five microliters of the restricted
product was run on a 1% (0.5× Tris-borate-EDTA) agarose gel.
Multivariate cluster analysis.
The isolates were grouped on
similarity of patterns of DNA fragments in agarose gels by the 16S rDNA
RFLP typing method described above. Individual isolates were scored
visually for the presence or absence of DNA fragments (range, 70 to 1.4 kb) generated by this typing method. Cluster analysis of similarity
matrices was performed by the unweighted-pair-group method with
arithmetic averages (43).
Amplification and sequencing of 16S rDNA.
Bacterial biomass
cultured in PMC for 2 days was centrifuged, washed once in sterile
distilled water, and frozen at 
20°C. Methods detailed in references
26 and 27 were employed for isolation and storage of DNA and PCR amplification of the 16S rDNA.
Automated sequencing (27) was employed to obtain the nearly complete sequences (1,450 bp) of the 16S rDNAs from strains Lp1265, Lp1275, Lp1276, and Lp1284. Primers 27f and 1492r were employed for
PCR, while primers 343r, 519r, 787r, 907r, 1100r, 1241r, 1385r, and
1492r were employed for sequencing.
Comparative sequence analysis.
Sequence data was aligned
with CLUSTAL W (48), and the alignments were manually
adjusted to take account of the conserved nature of helices.
Phylogenetic analysis was by the distance methods of Jukes and Cantor
(25), and tree topology was inferred by using the
neighbor-joining algorithm (34) with Treecon. All sequence-based trees were analyzed by bootstrapping of 1,000 trees.
Statistical analysis.
Statistical analysis of the effects of
carbohydrates on growth, ammonia production, and protease activity was
by analysis of variance, with differences being determined by the
method of least significant difference at the 5% level (P < 0.05).
| |
RESULTS |
Isolation, morphology, and genotype.
Proteolytic bacteria were
present in the rumina of calliandra-fed sheep at 1.5 × 108 cells/g of digesta. Fifteen (data for Lp1272 are not
shown) different proteolytic isolates were identified based on
polymorphisms of IR, ERIC, and REP sequences (Fig.
1; Table
2). The bacteria were grouped on the
basis of similarity according to results of cluster analysis of 16S
rDNA RFLP patterns (Fig. 2 and
3). Several strains (Lp1268, Lp1269,
Lp1272, Lp1276, Lp1275, and Lp1289) were closely related
phylogenetically based on 16S rDNA RFLP patterns, and all produced
lactate, formate, and acetate as end products of fermentation (Table
2). Strain Lp1284 produced both isobutyrate and isovalerate, while
Lp1311 was the only isolate that formed succinate. Both strains Lp1265
and Lp1266 produced butyrate as a major end products. Two bacteria
(Lp1265 and Lp1284) produced clearing zones on both tannic acid and
calliandra tannin plates, while the other isolates produced zones with
only one tannin type or no zones at all (Table 2).
View larger version (75K):
[in this window]
[in a new window]
|
FIG. 1.
DNA fragments amplified by primers specific for ERIC,
REP, and 16-to-23S IR sequences for proteolytic isolates.
|
|
View this table:
[in this window]
[in a new window]
|
TABLE 2.
Characteristics of bacterial strains isolated on agar
plates of BHI medium overlaid with tannic acid (hydrolyzable
tannin) and calliandra tannin (condensed tannin)
|
|
View larger version (69K):
[in this window]
[in a new window]
|
FIG. 2.
Restriction fragment length patterns of 16S rDNAs
digested with AluI, DdeI, and Sau3 for
proteolytic isolates.
|
|
View larger version (13K):
[in this window]
[in a new window]
|
FIG. 3.
Cluster analysis of restriction patterns of the
amplified 16S rDNAs shown in Fig. 2 for proteolytic isolates.
|
|
Carbohydrate fermentation and deaminase and protease
activities.
The isolates were placed into two groups (high and low
fermentation of carbohydrate) based on a significantly higher
(P < 0.05) growth response to inclusion of
carbohydrates other than glucose in the medium (Table
3). Inclusion of glucose of PMC increased
the maximum OD by only 0.2 to 0.4 for two of the bacteria with low
levels of carbohydrate fermentation (Lp1280 and Lp1283B), which was
less than the growth response to sugars in the group with high levels
of carbohydrate fermentation. Deletion of yeast extract and rumen fluid
from PM resulted in a decrease in growth of Lp1284 from 0.70 to 0.42 OD
units.
View this table:
[in this window]
[in a new window]
|
TABLE 3.
Growth, ammonia production, and protease activities of
bacteria grown on peptide medium with and
without carbohydratea
|
|
All isolates showed deaminase activity when they were grown on PM, but
strain Lp1284 grew to the highest OD (
P < 0.05) and
produced the greatest amount of ammonia (Table
3) (
P < 0.05).
Specific growth rate, OD, linear rate of ammonia
production, and
total ammonia production for Lp1284 in PM that
contained 1.5%
Trypticase peptone and 1.5% Casamino Acids were 0.28, 1.45, 734
µmol/min/ml/OD unit and 62.3 mM, respectively. Strain
Lp1284 also
produced ammonia from growth on casein (OD, 0.65) and
fraction
1 leaf protein (Fig.
4).
Approximately 18.2 and 31.5% of N from
Trypticase-Casamino Acids and
casein, respectively, were fermented
to ammonia N by Lp1284.
Proteolysis of azocasein was detected
for all isolates and varied from
<10 to >160 U compared with 105
U of activity for
P. ruminicola B
14 (Table
3). Protease activity
in
bacteria with low carbohydrate fermentation was predominantly
extracellular, whereas carbohydrate fermenters produced both
cell-associated
and extracellular activities. The protease activity of
carbohydrate-fermenting
bacteria (except for Lp1269) was not
significantly (
P < 0.05)
affected by the presence or
absence of carbohydrate in PM, although
this activity tended to be
lower in the absence of sugar in the
bacteria that fermented
carbohydrate at low levels (Table
3).
Protein digestion in the presence of tannins.
Initial screens
showed that two isolates (Lp1284 and Lp1265) produce ammonia (>1 mM)
from calliandra, and these isolates were thus compared with mixed rumen
fluid. Both isolates and mixed rumen fluid fermented protein in
calliandra to ammonia (5 to 10 mM) when PEG 4000 was included in the
medium, but the cultures accumulated significantly less ammonia (1.5 mM) when PEG 4000 was absent (Fig. 5).
The percentages of calliandra protein N fermented to ammonia N by
Lp1284, Lp1265, and rumen fluid in the presence of PEG 4000 were 52.2, 71.4, and 64.8% respectively. Coculturing of either Lp1284 or Lp1265
with the fibrolytic strain R. flavefaciens AR67 did not
significantly affect the rate or extent of protein fermentation (data
not shown).
View larger version (11K):
[in this window]
[in a new window]
|
FIG. 5.
Ammonia production by Lp1284 and Lp1265 and mixed rumen
microorganisms grown on calliandra with and without PEG 4000 (1 mg/10
mg of calliandra).
|
|
Ammonia production from growth on tannin-protein complexes (containing
22% protein) was not greater than that of the control
cultures, which
lacked the complexes (data not
shown).
Nucleic acids and 16S rDNA comparisons.
The GenBank accession
numbers for the nucleotide sequences used are AF105402 (Lp1284) and
AF105403 (Lp1265). Lp1284 was closely related to
Clostridium botulinum NCTC 7273 (97.6%). The sequence of
the closest relative to strain Lp1265 was Pseudobutyrivibrio ruminis DSM 9798 (96.3%). Strains Lp1275 (accession no.
AF135452) and Lp1276 (accession no. AF135453) grouped with
Eubacterium spp. and Streptococcus spp.,
respectively, and the closest relatives were Eubacterium
gallinarium (accession no. AF03990) (98.3%) and Streptococcus bovis JB1 (99.1%). Phylogenetic trees based
on 16S rDNA sequences of these bacteria and closely related organisms are shown in Fig. 6.
View larger version (38K):
[in this window]
[in a new window]
|
FIG. 6.
Phylogenetic tree of Lp1265, Lp1275, Lp1276, and Lp1284
and related organisms based on 16S rDNA sequences. Bootstrap values are
given as percentages of 1,000 random trees. The scale bar represents
0.1 mutation per site and is expressed as a percentage.
|
|
| |
DISCUSSION |
This study demonstrates that proteolytic bacteria which ferment
peptides to ammonia occur in relatively high numbers (108/g
of digesta) in ruminants fed a highly tanniniferous diet and that the
isolated organisms fall into two classes: those with high and low
levels of carbohydrate fermentation. However, it cannot be definitively
stated that the presence of these bacteria was due to the
tannin-containing calliandra since animals not fed a tannin-enriched
diet may also harbor these microorganisms. Therefore, the
characterization of these proteolytic bacteria is discussed (i) within
the broader context of rumen microbial ecology and (ii) in relation to
their potential role in digesting tannin-rich forage.
Hungate (18) stated that he had "encountered" rumen
bacteria "able to digest casein and requiring no carbohydrate," but he did not isolate these bacteria. This is the first report of a
proteolytic ruminal bacterium that grows rapidly on peptides and amino
acids and does not require carbohydrates for growth. Previously, only
obligate amino-acid-fermenting bacteria that were not proteolytic had
been isolated (14). Strain Lp1284 is an ammonia
hyperproducer, grows rapidly on peptides and amino acids, and is highly
proteolytic. This organism also ferments peptides to branched-chain
fatty acids and propionate, which has also been observed with the
obligate amino-acid-fermenting rumen bacterium Clostridium
sticklandii SR (11), which is not proteolytic. The rate
and amount of ammonia production of Lp1284 grown on peptide medium were
similar to those of the obligate amino-acid-fermenting rumen bacteria
Peptostreptococcus sp. strain D1 and strain D4 (3). The high specific rates of ammonia production by Lp1284 are also comparable with those of Peptostreptococcus
anaerobius (type C), C. sticklandii (type SR) and
Clostridium aminophilum (type F) (10, 11, 26).
Strain Lp1284 apparently converted approximately 18.3% of peptide N
from Trypticase and Casamino Acids to ammonia and grew to an OD similar
to that of Peptostreptococcus anaerobius (type C), which
utilized 23 to 31% of the N from these nitrogen sources
(10). A highly proteolytic clostridium (C. proteoclasticum) has also been isolated from the bovine rumen, but
this organism grows on carbohydrate and does not produce ammonia (2). Most of these ammonia-hyperproducing bacteria fall
within the genus Clostridium, as described by Collins and
coworkers (12) (Fig. 6).
Phylogenetic analysis based on 16S rDNA sequence indicates that Lp1284
is related to C. botulinum group 1. This group of bacteria contain proteolytic C. botulinum types A, B, and F
(20). They are also closely related to Clostridium
sporogenes and fall within cluster 1 of the polysaccharolytic
clostridia (40). Phylogenetic analysis and phenotypic
characteristics demonstrate that strain Lp1265 belongs to cluster XIVa
of the Clostridium subphylum (12, 50) and that it
is closely related to several strains of B. fibrisolvens
(17) but that Lp1268, Lp1269, Lp1272, and Lp1276 are
Streptococcus spp.
This is the first report of an attempt to isolate rumen bacteria with
an ability to digest protein in the presence of condensed tannins.
Although the bacteria isolated were proteolytic, we were unable to
demonstrate significant degradation of calliandra-protein complexes or
fermentation of in situ complexed calliandra protein. However, when the
interaction between tannin and protein in calliandra was counteracted
with PEG 4000, then proteolytic bacteria were able to readily degrade
protein. This result indicates that a substantial amount of calliandra
protein is readily available for fermentation, provided that it is not
totally complexed with tannin, which occurs in fresh plant material.
Anaerobic bacteria that degrade hydrolyzable tannins or hydrolyzable
tannin-protein complexes have been isolated from the digesta of many
nonruminant species, including koala (Phascolarctos cinereus), by using tannic acid as the model tannin in the
selection medium (36-39). A reason why bacteria that
degrade calliandra tannin-protein complex were not isolated is probably
due to the difference in chemical structures of these two classes of
tannins. Hydrolyzable tannins (e.g., tannic acid) are polymerized units
of glucose esterified to gallic and hexahydroxydiphenic acid, whereas
condensed tannins (e.g., calliandra tannin) are polymers of
flavan-3-ols or flavon-3,4-diols (Fig.
7). Both tannins complex with protein by
forming hydrogen bonds between the phenolic subunits of the polymer and
aliphatic and aromatic side chains (carbonyl groups of peptides) of the protein. The most likely explanation for degradation of hydrolyzable tannin-protein complexes is that enzymes (tannin acylhydrolases and
esterases) depolymerize the tannin polymer by cleaving the ester
linkages between glucose and the phenolic subunits (41). A
mechanism for the anaerobic degradation of the condensed tannin molecule has not yet been described. However, nonenzymatic cleavage of
both types of tannin-protein may occur under acidic conditions in the
rumen.
View larger version (15K):
[in this window]
[in a new window]
|
FIG. 7.
Diagram of the generic chemical structures of
hydrolyzable (left panel) and condensed (right panel) tannins. n,
repeating subunit of the polymer.
|
|
Further studies should be undertaken to determine whether some ruminal
populations are better adapted to (tolerant of) tannin-containing forages and thus more efficient at digesting protein under those circumstances. Tolerance of the isolates to tannins was not measured, but the isolation procedures have previously yielded tolerant bacteria
(6, 35). However, the predominate use of hydrolyzable tannin
in these experiments rather than condensed tannin should be
reevaluated, considering their relative levels of importance in the
nutrition of herbivores. Condensed tannins are more widely distributed
in plants than the hydrolyzable type and are thus regarded as a more
significant nutritional problem (45). Hydrolyzable tannins
are absent from nonvascular plants, and even in vascular plants, they
are restricted to only dicotyledons and some classes of flowering
plants in monocotyledons.
| |
ACKNOWLEDGMENTS |
This work was partly supported by the Australian Centre for
International Agricultural Research.
We thank Christina Fraser for technical assistance.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: CSIRO Tropical
Agriculture, Private Bag No. 3, P.O., Indooroopilly, 4068 QLD,
Australia. Phone: 61 7 3214 2820. Fax: 61 7 3214 2880. E-mail:
Chris.McSweeney{at}tag.csiro.au.
| |
REFERENCES |
| 1.
|
Ahn, J. H.,
B. M. Robertson,
R. Elliott,
R. C. Gutteridge, and C. W. Ford.
1989.
Quality assessment of tropical browse legumes: tannin content and protein degradation.
Anim. Feed Sci. Technol.
27:147-156.
|
| 2.
|
Attwood, G. T.,
K. Reilly, and B. K. C. Patel.
1996.
Clostridium proteoclasticum sp. nov., a novel proteolytic bacterium from the bovine rumen.
Int. J. Syst. Bacteriol.
46:753-758[Abstract/Free Full Text].
|
| 3.
|
Attwood, G. T.,
A. V. Klieve,
D. Ouwerkerk, and B. K. C. Patel.
1998.
Ammonia-hyperproducing bacteria from New Zealand ruminants.
Appl. Environ. Microbiol.
64:1796-1804[Abstract/Free Full Text].
|
| 4.
|
Bae, H. D.,
T. K. McAllister,
L. J. Yanke,
K.-J. Cheng, and A. Muir.
1993.
Effect of condensed tannins on endoglucanase activity and filter paper digestion by Fibrobacter succinogenes S85.
Appl. Environ. Microbiol.
59:2132-2138[Abstract/Free Full Text].
|
| 5.
|
Brock, F. M.,
C. W. Forsberg, and J. G. Buchanan-Smith.
1982.
Proteolytic activity of rumen microorganisms and effects of proteinase inhibitors.
Appl. Environ. Microbiol.
44:561-569[Abstract/Free Full Text].
|
| 6.
|
Brooker, J. D.,
L. A. O'Donovan,
I. Skene,
K. Clarke,
L. Blackall, and P. Muslera.
1994.
Streptococcus caprinus sp. nov., a tannin-resistant ruminal bacterium from feral goats.
Lett. Appl. Microbiol.
18:313-318.
|
| 7.
|
Bryant, M. P.
1972.
Commentary on the Hungate technique for culture of anaerobic bacteria.
Am. J. Clin. Nutr.
25:1324-1328[Free Full Text].
|
| 8.
|
Caldwell, D. R., and M. P. Bryant.
1966.
Medium without rumen fluid for nonselective enumeration and isolation of rumen bacteria.
Appl. Microbiol.
14:794-801[Medline].
|
| 9.
|
Chaney, A. L., and E. P. Marbach.
1962.
Modified reagents for determination of urea and ammonia.
Clin. Chem.
8:130-132[Abstract].
|
| 10.
|
Chen, G., and J. B. Russell.
1988.
Fermentation of peptides and amino acids by a monensin-sensitive ruminal peptostreptococcus.
Appl. Environ. Microbiol.
54:2742-2749[Abstract/Free Full Text].
|
| 11.
|
Chen, G., and J. B. Russell.
1989.
More monensin-sensitive, ammonia-producing bacteria from the rumen.
Appl. Environ. Microbiol.
55:1052-1057[Abstract/Free Full Text].
|
| 12.
|
Collins, M. D.,
P. A. Lawson,
A. Willems,
J. J. Cordoba,
J. Fernandez-Garayzabal,
P. Garcia,
J. Cai,
H. Hippe, and J. A. E. Farrow.
1994.
The phylogeny of the genus Clostridium: proposal of five new genera and eleven new species combinations.
Int. J. Syst. Bacteriol.
44:812-826[Abstract/Free Full Text].
|
| 13.
|
Cotta, M. A., and R. B. Hespell.
1986.
Proteolytic activity of the ruminal bacterium Butyrivibrio fibrisolvens.
Appl. Environ. Microbiol.
52:51-58[Abstract/Free Full Text].
|
| 14.
|
Cotta, M. A., and J. B. Russell.
1996.
Digestion of nitrogen in the rumen: a model for metabolism of nitrogen compounds in gastrointestinal environments, p. 380-423.
In
R. I. Mackie, and B. S. A. White (ed.), Gastrointestinal microbiology., vol. 1. Chapman and Hall, New York, N.Y.
|
| 15.
|
de Bruijn, F. J.
1992.
Use of repetitive (repetitive extragenic palindromic and enterobacterial repetitive intergeneric consensus) sequences and the polymerase chain reaction to fingerprint the genomes of Rhizobium meliloti isolates and other soil bacteria.
Appl. Environ. Microbiol.
58:2180-2187[Abstract/Free Full Text].
|
| 16.
|
Dolfing, J., and J. C. Gottschal.
1997.
Microbe-microbe interactions, p. 373-433.
In
R. I. Mackie, and B. S. A. White (ed.), Gastrointestinal microbiology, vol. 2. Chapman and Hall, New York, N.Y.
|
| 17.
|
Forster, R. J.,
R. M. Teather,
J. Gong, and S.-J. Deng.
1996.
16S rDNA analysis of Butyrivibrio fibrisolvens: phylogenetic position and relation to butyrate-producing anaerobic bacteria from the rumen of white-tailed deer.
Lett. Appl. Microbiol.
23:218-222[Medline].
|
| 18.
|
Hungate, R. E.
1966.
The rumen and its microbes, p. 297.
Academic Press, New York, N.Y.
|
| 19.
|
Hungate, R. E.
1969.
A roll tube method for cultivation of strict anaerobes.
Methods Microbiol.
3:117-132.
|
| 20.
|
Hutson, R. A.,
D. E. Thompson,
P. A. Lawson,
R. P. Schocken-Itturino,
E. C. Bottger, and M. D. Collins.
1993.
Genetic interrelationships of proteolytic Clostridium botulinum types A, B, and F and other members of the Clostridium botulinum complex as revealed by small-subunit rRNA gene sequences.
Antonie Leeuwenhoek
64:273-283.
|
| 21.
|
Jensen, M. A., and R. J. Hubner.
1996.
Use of homoduplex ribosomal DNA spacer amplification products and heteroduplex cross-hybridization products in identification of Salmonella serovars.
Appl. Environ. Microbiol.
62:2741-2746[Abstract].
|
| 22.
|
Jones, G. A.,
T. A. McAllister,
K.-J. Cheng, and A. D. Muir.
1994.
Effect of sainfoin (Onobrychis viciifolia Scop) on growth and proteolysis by four strains of rumen bacteria: resistance of Prevotella (Bacteroides) ruminicola B14.
Appl. Environ. Microbiol.
60:1374-1378[Abstract/Free Full Text].
|
| 23.
|
Jones, W. T., and J. L. Mangan.
1977.
Complexes of the condensed tannins of sainfoin (Onobrychis viciifolia Scop.) with fraction 1 leaf protein and with submaxillary-mucoprotein, and their reversal by polyethylene glycol and pH.
J. Sci. Food Agric.
28:126-136.
|
| 24.
|
Jones, W. T., and J. W. Lyttleton.
1969.
Bloat in cattle 29. The foaming properties of clover proteins.
N. Z. J. Agric. Res.
12:31-46.
|
| 25.
|
Jukes, T. H., and C. R. Cantor.
1969.
Evolution of protein molecules, p. 21-132.
In
H. N. Munro (ed.), Mammalian protein metabolism. Academic Press, New York, N.Y.
|
| 26.
|
Krause, D. O., and J. B. Russell.
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].
|
| 27.
|
Krause, D. O.,
B. A. White, and R. I. Mackie.
1997.
Ribotyping of adherent Lactobacillus from weaning pigs: a basis for probiotic selection based on diet and gut compartment.
Anaerobe
3:317-325.
|
| 28.
|
Lowe, S. E.,
M. K. Theodorou,
A. P. J. Trinci, and R. B. Hespell.
1985.
Growth of anaerobic rumen fungi on defined and semi-defined media lacking rumen fluid.
J. Gen. Microbiol.
131:2225-2229.
|
| 29.
|
Mackie, R. I.,
F. M. C. Gilchrist,
A. M. Robberts,
P. E. Hannah, and H. M. Schwartz.
1978.
Microbiological and chemical changes in the rumen during the stepwise adaption of sheep to high concentrate diets.
J. Agric. Sci.
90:241-254.
|
| 30.
|
Mackie, R. I., and C. A. Wilkins.
1988.
Enumeration of anaerobic bacterial microflora of the equine gastrointestinal tract.
Appl. Environ. Microbiol.
54:2155-2160[Abstract/Free Full Text].
|
| 30a.
| McSweeney, C. S. Unpublished data.
|
| 31.
|
Menke, K. H.,
L. Raab,
A. Salewski,
H. Steingass,
D. Fritz, and W. Schneider.
1979.
The estimation of the digestibility and metabolisable energy content of ruminant feeding stuffs from the gas production when they are incubated with rumen liquor in vitro.
J. Agric. Sci.
93:217-222.
|
| 32.
|
Miller, S. M.,
J. D. Brooker,
A. Phillips, and L. L. Blackall.
1996.
Streptococcus caprinus is ineffective as a rumen inoculum to improve digestion of mulga (Acacia aneura) by sheep.
Aust. J. Agric. Res.
47:1323-1331.
|
| 33.
|
Moyer, C. L.,
F. C. Dobbs, and D. M. Karl.
1994.
Estimation of diversity and community structure through restriction fragment length polymorphism distribution analysis of bacterial 16S rRNA genes from a microbial mat at an active, hydrothermal vent system, Loihi Seamount, Hawaii.
Appl. Environ. Microbiol.
60:871-879[Abstract/Free Full Text].
|
| 34.
|
Nei, M., and W.-H. Li.
1979.
Mathematical model for studying genetic variation in terms of restriction endonucleases.
Proc. Natl. Acad. Sci. USA
76:5269-5273[Abstract/Free Full Text].
|
| 35.
|
Nelson, K. E.,
M. L. Thonney,
T. K. Woolston,
S. H. Zinder, and A. N. Pell.
1998.
Phenotypic and phylogenetic characterization of ruminal tannin-tolerant bacteria.
Appl. Environ. Microbiol.
64:3824-3830[Abstract/Free Full Text].
|
| 36.
|
Nemoto, K.,
R. Osawa,
K. Hirota,
T. Ono, and Y. Miyake.
1995.
An investigation of gram-negative tannin-protein degrading bacteria in fecal flora of various mammals.
J. Vet. Med. Sci.
57:921-926[Medline].
|
| 37.
|
Osawa, R. O.
1990.
Formation of a clear zone on tannin-treated brain heart infusion agar by a Streptococcus sp. isolated from faeces of koalas.
Appl. Environ. Microbiol.
56:829-831[Abstract/Free Full Text].
|
| 38.
|
Osawa, R. O.
1992.
Tannin-protein complex-degrading enterobacteria isolated from the alimentary tracts of koalas and a selective medium for their enumeration.
Appl. Environ. Microbiol.
58:1754-1759[Abstract/Free Full Text].
|
| 39.
|
Osawa, R. O.,
T. P. Walsh, and S. J. Cork.
1993.
Metabolism of tannin-protein complex by facultatively anaerobic bacteria isolated from koala feces.
Biodegradation
4:91-99.
|
| 40.
|
Rainey, F. A., and E. Stackebrandt.
1993.
16S rDNA analysis reveals phylogenetic diversity among the polysaccharolytic clostridia.
FEMS Microbiol. Lett.
113:125-128[Medline].
|
| 41.
|
Skene, I., and J. D. Brooker.
1995.
Characterisation of the tannin acylhydrolase in the ruminal bacterium Selenomonas ruminantium.
Anaerobe
1:321-327.
|
| 42.
|
Sly, I.,
M. M. Cahill,
R. O. Osawa, and T. Fujisawa.
1997.
The tannin-degrading species Streptococcus gallolyticus and Streptococcus caprinus are subjective synonyms.
Int. J. Syst. Bacteriol.
47:893-894[Abstract/Free Full Text].
|
| 43.
|
Sokal, R. R., and C. D. Michener.
1958.
A statistical method for evaluating systematic relationships.
Univ. Kans. Sci. Bull.
38:1409-1438.
|
| 44.
|
Stahl, M.,
G. Molin,
A. Persson,
S. Ahrne, and S. Stahl.
1990.
Restriction endonuclease patterns and multivariate analysis as a classification tool for Lactobacillus spp.
Int. J. Syst. Bacteriol.
40:189-193[Abstract/Free Full Text].
|
| 45.
|
Swain, T.
1979.
Tannin and lignins, p. 657-682.
In
A. Rosenthal, and J. H. Janzen (ed.), Herbivores: their interaction with secondary plant metabolites. Academic Press, New York, N.Y.
|
| 46.
|
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. Sci. Food Agric.
58:321-329.
|
| 47.
|
Terrill, T. H.,
W. R. Windham,
J. J. Evans, and C. S. Hoveland.
1990.
Condensed tannin concentration in Sericea lespedeza as influenced by preservation method.
Crop Sci.
30:219-224.
|
| 48.
|
Thompson, J. D.,
D. G. Higgins, and T. J. Gibson.
1994.
CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choices.
Nucleic Acids Res.
22:4673-4680[Abstract/Free Full Text].
|
| 49.
|
Versalovic, J.,
T. Koeuth, and J. R. Lupski.
1991.
Distribution of repetitive DNA sequences in eubacteria and application to fingerprinting of bacterial genomes.
Nucleic Acids Res.
19:6823-6831[Abstract/Free Full Text].
|
| 50.
|
Willems, A.,
M. Amatmarco, and M. D. Collins.
1996.
Phylogenetic analysis of Butyrivibrio strains reveals three distinct groups of species within the Clostridium subphylum of the gram-positive bacteria.
Int. J. Syst. Bacteriol.
46:195-199[Abstract/Free Full Text].
|
Applied and Environmental Microbiology, July 1999, p. 3075-3083, Vol. 65, No. 7
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
This article has been cited by other articles:
-
Krause, D. O., Smith, W. J. M., McSweeney, C. S.
(2004). Use of community genome arrays (CGAs) to assess the effects of Acacia angustissima on rumen ecology. Microbiology
150: 2899-2909
[Abstract]
[Full Text]
-
Wallace, R. J., Chaudhary, L. C., Miyagawa, E., McKain, N., Walker, N. D.
(2004). Metabolic properties of Eubacterium pyruvativorans, a ruminal 'hyper-ammonia-producing' anaerobe with metabolic properties analogous to those of Clostridium kluyveri. Microbiology
150: 2921-2930
[Abstract]
[Full Text]
-
McIntosh, F. M., Williams, P., Losa, R., Wallace, R. J., Beever, D. A., Newbold, C. J.
(2003). Effects of Essential Oils on Ruminal Microorganisms and Their Protein Metabolism. Appl. Environ. Microbiol.
69: 5011-5014
[Abstract]
[Full Text]
-
Wallace, R. J., McKain, N., McEwan, N. R., Miyagawa, E., Chaudhary, L. C., King, T. P., Walker, N. D., Apajalahti, J. H. A., Newbold, C. J.
(2003). Eubacterium pyruvativorans sp. nov., a novel non-saccharolytic anaerobe from the rumen that ferments pyruvate and amino acids, forms caproate and utilizes acetate and propionate. Int. J. Syst. Evol. Microbiol.
53: 965-970
[Abstract]
[Full Text]
-
Eschenlauer, S. C. P., McKain, N., Walker, N. D., McEwan, N. R., Newbold, C. J., Wallace, R. J.
(2002). Ammonia Production by Ruminal Microorganisms and Enumeration, Isolation, and Characterization of Bacteria Capable of Growth on Peptides and Amino Acids from the Sheep Rumen. Appl. Environ. Microbiol.
68: 4925-4931
[Abstract]
[Full Text]
-
Osawa, R., Kuroiso, K., Goto, S., Shimizu, A.
(2000). Isolation of Tannin-Degrading Lactobacilli from Humans and Fermented Foods. Appl. Environ. Microbiol.
66: 3093-3097
[Abstract]
[Full Text]
Top
Abstract
Introduction
