High genetic diversity and different distributions of glycosyl hydrolase family 10 and 11 xylanases in the goat rumen.

Abstract

The rumen harbors a complex microbial ecosystem for efficient hydrolysis of plant polysaccharides which are the main constituent of the diet. Xylanase is crucial for hemicellulose hydrolysis and plays an important role in the plant cell wall degradation. Xylanases of ruminal strains were widely studied, but few studies have focused on their diversity in rumen microenvironment.
We explored the genetic diversity of xylanases belonging to two major glycosyl hydrolase families (GH 10 and 11) in goat rumen contents by analyzing the amplicons generated with two degenerate primer sets. Fifty-two distinct GH 10 and 35 GH 11 xylanase gene fragments (similarity <95%) were retrieved, and most had low identities with known sequences. Based on phylogenetic analysis, all GH 10 xylanase sequences fell into seven clusters, and 88.5% of them were related to xylanases from Bacteroidetes. Five clusters of GH 11 xylanase sequences were identified. Of these, 85.7% were related to xylanases from Firmicutes, and 14.3% were related to those of rumen fungi. Two full-length xylanase genes (one for each family) were directly cloned and expressed in Escherichia coli. Both the recombinant enzymes showed substantial xylanase activity, and were purified and characterized. Combined with the results of sheep rumen, Bacteroidetes and Firmicutes are the two major phyla of xylan-degrading microorganisms in rumen, which is distinct from the representatives of other environments such as soil and termite hindgut, suggesting that xylan-degrading microorganisms are environment specific.
The numerous new xylanase genes suggested the functional diversity of xylanase in the rumen microenvironment which may have great potential applications in industry and agriculture. The phylogenetic diversity and different distributions of xylanase genes will help us understand their roles in plant cell wall degradation in the rumen microenvironment.

Full text

Page 1

High Genetic Diversity and Different Distributions of
Glycosyl Hydrolase Family 10 and 11 Xylanases in the
Goat Rumen
Guozeng Wang1, Huiying Luo1, Kun Meng1, Yaru Wang1, Huoqing Huang1, Pengjun Shi1, Xia Pan1,
Peilong Yang1, Qiyu Diao1, Hongfu Zhang2, Bin Yao1*
1Key Laboratory for Feed Biotechnology of the Ministry of Agriculture, Feed Research Institute, Chinese Academy of Agricultural Sciences, Beijing, People’s Republic of
China, 2State Key Lab of Animal Nutrition, Institute of Animal Science, Chinese Academy of Agricultural Sciences, Beijing, People’s Republic of China
Abstract
Background: The rumen harbors a complex microbial ecosystem for efficient hydrolysis of plant polysaccharides which are
the main constituent of the diet. Xylanase is crucial for hemicellulose hydrolysis and plays an important role in the plant cell
wall degradation. Xylanases of ruminal strains were widely studied, but few studies have focused on their diversity in rumen
microenvironment.
Methodology/Principal Findings: We explored the genetic diversity of xylanases belonging to two major glycosyl hydrolase
families (GH 10 and 11) in goat rumen contents by analyzing the amplicons generated with two degenerate primer sets.
Fifty-two distinct GH 10 and 35 GH 11 xylanase gene fragments (similarity ,95%) were retrieved, and most had low
identities with known sequences. Based on phylogenetic analysis, all GH 10 xylanase sequences fell into seven clusters, and
88.5% of them were related to xylanases from Bacteroidetes. Five clusters of GH 11 xylanase sequences were identified. Of
these, 85.7% were related to xylanases from Firmicutes, and 14.3% were related to those of rumen fungi. Two full-length
xylanase genes (one for each family) were directly cloned and expressed in Escherichia coli. Both the recombinant enzymes
showed substantial xylanase activity, and were purified and characterized. Combined with the results of sheep rumen,
Bacteroidetes and Firmicutes are the two major phyla of xylan-degrading microorganisms in rumen, which is distinct from
the representatives of other environments such as soil and termite hindgut, suggesting that xylan-degrading
microorganisms are environment specific.
Conclusion/Significance: The numerous new xylanase genes suggested the functional diversity of xylanase in the rumen
microenvironment which may have great potential applications in industry and agriculture. The phylogenetic diversity and
different distributions of xylanase genes will help us understand their roles in plant cell wall degradation in the rumen
microenvironment.
Citation: Wang G, Luo H, Meng K, Wang Y, Huang H, et al. (2011) High Genetic Diversity and Different Distributions of Glycosyl Hydrolase Family 10 and 11
Xylanases in the Goat Rumen. PLoS ONE 6(2): e16731. doi:10.1371/journal.pone.0016731
Editor: Shuang-yong Xu, New England Biolabs, Inc., United States of America
Received October 9, 2010; Accepted December 24, 2010; Published February 3, 2011
Copyright: ? 2011 Wang et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was supported by the Key Program of Transgenic Plant Breeding (2008ZX08011-005), the Earmarked Fund for Modern Agro-industry
Technology Research System (NYCYTX-42-G2-05), and the Agricultural Science and Technology Conversion Funds (grant 2008GB23260388). The funders had no
role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: yaobin@caas-bio.net.cn
Introduction
Plant cell walls mainly consist of cellulose, hemicelluloses, and
lignin, and are a major component of the diet of grazing ruminants
[1]. The rumen ecosystem consists of various anaerobic microor-
ganisms including bacteria, fungi, protozoa, and archaea that are
characterized by high population density, wide diversity, and
complexity of interactions [2]. These rumen microbes work
synergistically to efficiently hydrolyze plant cell wall polysaccha-
rides by producing various enzymes, such as cellulases, hemi-
cellulases, polyphenol oxidases, esterases, etc [2], [3], [4].
Hydrolysis of the plant cell wall is a complex process in which
hemicellulose digestion is the initial step with subsequent
hydrolysis of the cellulose [5], [6]. Xylan is the major component
of hemicellulose, and its complete hydrolysis requires a crucial
enzyme—endo-1,4-b-D-xylanase (EC 3.2.1.8)—to cleave xylan
into short xylooligosaccharides of varying lengths [7], [8].
Xylanases have been found in rumen bacteria, fungi, and protozoa
but not yet archaea [2], and some xylanases display unique
structures and characteristics [9]. Using protein purification and
molecular cloning methods, researchers have detected multiple
xylanases in rumen bacteria and fungi [10], [11], [12]. Moreover,
whole-genome sequencing data of rumen bacteria [13] (http://
www.jcvi.org/rumenomics/) further provide a more comprehen-
sive understanding of the diversity of xylanase genes, their
distribution in various bacteria, and their roles in degrading plant
cell wall polysaccharides [14], [15], [16].
Thegeneticdiversityofrumenmicrobialcommunitiesbasedon16S
rDNA sequences has been widely studied, and results have suggested a
high diversity of rumen microbes with the majority of them not yet
PLoS ONE | www.plosone.org1February 2011 | Volume 6 | Issue 2 | e16731

Page 2

cultured [17]. Rumen metagenomics have revealed the diversity of
glycosyl hydrolases (GH) of rumen microbes [3], [18], but far fewer
xylanase genes have been obtained using these methods relative to the
large number of uncultured microorganisms in the rumen ecosystem.
Culture-independent study of functional gene diversity has allowed
access to the uncultured rumen microenvironment and provided
insight into metabolic capabilities of uncultivated microbial commu-
nities [19]. Moreover, xylanases from these uncultured microbes may
be of special interest because of their functional diversity and potential
applications [18], [19].
The objective of this study was to explore the genetic diversity of
xylanases in goat rumen using culture-independent molecular
approaches. Sequence analysis showed that most of the xylanase
gene fragments have low identities to known xylanases, suggesting
the existence of a large number of uncharacterized xylanase genes
in rumen. To confirm that these genes encode active enzymes, one
GH 10 and one GH 11 full-length xylanase genes were directly
cloned from the rumen genomic DNA and expressed in Escherichia
coli. One of the xylanases showed high activity even at relatively
low temperatures. Our study provides new insight into the genetic
diversity and distribution of xylanases, which will help us
understand their roles in the rumen microenvironment.
Results
Amplification and sequence analysis of GH 10 xylanase
gene fragments
Using degenerate primers X10-F and X10-R [20], gene fragments
of about 260 bp for GH 10 xylanases were amplified directly from
the total genomic DNA of goat rumen contents. A clone library was
constructed, and positive transformants (white colonies) were
confirmed by PCR with primers M13F and M13R. Of the 310
clones sequenced, 236 sequences showed 35–84% amino acid
identity with known GH 10 xylanases based on BLAST analysis,
and no fragment contained introns. The highly conserved Asn
residue of GH 10 xylanases [21] was identified in all of these protein
sequences. The result suggested that these sequences were partial
xylanases and that some of them might be previously unidentified.
After removing the redundant sequences using CD-hit program
[22], 52 sequences showed divergence (sharing ,95% identity)
(Table S1). The length of these sequences varied a lot, with the
range from 84 to 111 residues. Abundance analysis using distance-
based operational taxonomic unit and richness determination
(DOTUR) software [23] showed that GR117 was the predomi-
nant operational taxonomic unit (OUT) that represented 30
sequences. Twelve OTUs contained only one sequence (Table S1).
Phylogenetic analysis of GH 10 xylanase gene fragments
An unrooted protein-level phylogenetic tree for GH 10
xylanases was constructed with the 52 divergent sequences from
the GH 10 clone library and 13 reference sequences from
GenBank. All the sequences were confined to seven clusters,
denoted A, B, C, D, E, F, and G, indicating substantial diversity
among GH 10 xylanases in goat rumen (Figure 1). Many
sequences had no close relatives.
Cluster A contained 34 sequences from the goat rumen and two
reference sequences from Prevotella ruminicola and Bacteroides cellulosi-
lyticus. Cluster B contained the sequences of six clones and one
xylanase directly cloned from an uncultured human gut bacterium.
This reference sequence was highly similar to xylanases from
anaerobic intestinal bacteria such as Bacteroides spp. and Prevotella
spp. [24]. Four sequences as well as three reference sequences from
Prevotella buccae, Prevotella bergensis, and Prevotella copri formed cluster C.
ThreesequencesinclusterDwerecloselyrelatedtothexylanasefrom
Xanthomonas axonopodis. In cluster E, there were two sequences that
shared highest identity with the xylanase from Cellulosilyticum
ruminicola. Two sequences in cluster F were closely related to the
xylanases from P. ruminicola and Bacteroides sp. 2_2_4. Cluster G
containedonesequencefromourlibraryandtworeferencesequences
from Ruminococcus flavefaciens FD-1 and Epidinium ecaudatum.
Amplification and sequence analysis of GH 11 xylanase
gene fragments
Amplicons with the size of about 210 bp for GH 11 xylanases
were obtained from the metagenomic DNA of goat rumen
contents using degenerate primers X11-F and X11-R [20]. Of the
clone library constructed with PCR product recovered, 200 clones
were randomly sequenced, and 172 sequences showed 56–95%
amino acid identity with known GH 11 xylanases. The highly
conserved catalytic residue, Glu, of GH 11 xylanases [25], [26]
was found in all the protein sequences of 70–81 residues in length
(Table S2). Thus these sequences were identified to be partial GH
11 xylanases, and some of them were yet undiscovered.
After removing the redundant sequences using CD-hit program
[22], 35 sequences showed divergence (sharing ,95% identity)
(Table S2). DOTUR abundance analysis showed that R8 was the
predominant OTU that represented 15 sequences. Four OTUs
contained only one sequence (Table S2).
Phylogenetic analysis of GH 11 xylanase gene fragments
The 35 distinct partial sequences of GH 11 xylanases were used
to construct an unrooted phylogenetic tree with 12 reference
sequences (Figure 2). Five clusters (I, II, III, IV, and V) were formed
based on high bootstrap values. Many clades formed without closely
related references, suggesting that these sequences were probably
different from known xylanases. Twenty-two sequences shared
highest identity with xylanases from Ruminococcus albus, R. flavefaciens,
and Ruminococcus sp., and fell into clusters I and III. Five sequences
and that of Clostridium stercorarium weregrouped into cluster II.Seven
sequences closely related to xylanases from Neocallimastix patriciarum,
Orpinomyces sp. PC-2, and R. flavefaciens fell into cluster IV. Cluster V
only contained two references from N. patriciarum.
Cloning and expression of xylanase genes
Two full-length xylanase genes were directly cloned from the
metagenomic DNA of goat rumen contents. The complete
sequence of xynGR67 contained an open reading frame of
1,239 bp encoding a 412-residue polypeptide with a typical signal
peptide (residues 1–40). Sequence similarity searches showed that
deduced XynGR67 shared highest identity (45%) with the GH 10
xylanase from Flavobacteriaceae bacterium 3519-10, an isolate
recovered from a deep Antarctic ice core [27].
The complete sequence of xynR8 contained an open reading
frame of 807 bp that encoded a 268-residue polypeptide. No
signal peptide was predicted. The deduced protein shared highest
identity with the GH 11 xylanases from R. flavefaciens FD-1 [14]
and R. albus 8 (75% and 64%, respectively).
Both genes encoding the mature proteins were expressed in E.
coli BL21 (DE3). After induction with IPTG at 25uC for 12 h,
substantial xylanase activity was detected in the culture superna-
tant of recombinant cells (11.6 and 27.6 U ml–1for XynGR67 and
XynR8, respectively).
Biochemical characterization of purified recombinant
XynGR67 and XynR8
Using birchwood xylan as the substrate, XynGR67 showed the
highest activity at pH 6.0, and .75% of the maximum activity
Xylanases Diversity in Goat Rumen
PLoS ONE | www.plosone.org2February 2011 | Volume 6 | Issue 2 | e16731

Page 3

Xylanases Diversity in Goat Rumen
PLoS ONE | www.plosone.org3 February 2011 | Volume 6 | Issue 2 | e16731

Page 4

was retained at pH 5.5–7.5 (Figure 3A). The enzyme was stable at
pH 5.0–8.0, retaining more than 80% of the initial activity after
incubation at 25uC for 1 h. The optimal temperature for enzyme
activity of XynGR67 was 40uC (Figure 3B). At 0uC and 10uC, the
enzyme still exhibited 8.2% and 22.5% of the maximal activity,
respectively. XynGR67 was very thermolabile, retaining 65% of
the activity after 1 h incubation at 30uC, and losing activity rapidly
when incubated at 40uC (a half-life of 15 min).
XynR8 showed the highest activity at pH 5.5, and .80% of the
maximum activity was retained at pH 5.0–8.0 (Figure 3C). The
enzyme was stable at pH 5.0–9.0, retaining more than 80% of the
initial activity after incubation at 37uC for 1 h. The optimal
temperature for enzyme activity of XynR8 was 55uC (Figure 3D).
XynR8 was thermostable, retaining 85% of the activity after 1 h
incubation at 50uC, and remaining 31% activity when incubated
at 60uC for 1 h.
Kinetic parameters of purified recombinant XynGR67 and
XynR8 on birchwood xylan were shown in Table 1. Both enzymes
showed substantial xylanase activity to birchwood xylan. The
substrate specificity of XynGR67 and XynR8 were shown in
Table 2. Of four types of xylan tested, XynGR67 had the highest
activity toward beechwood xylan while XynR8 displayed the
highest activity toward soluble wheat arabinoxylan.
Discussion
In this study, xylanase genes were targeted for diversity analysis
because of their key roles in the initial steps of plant cell wall
breakdown and their great potential for industrial and agricultural
applications [7], [8], [28]. The microorganisms in the rumen are
immensely diverse, and the microbial-mediated hydrolysis of plant
cell wall polysaccharides is highly efficient [2]. Thus, the rumen is
an ideal microenvironment to study the genetic diversity of
functional xylanases.
Application of degenerate primers designed based on CODE-
HOP principles [20] resulted in the discovery of 52 distinct GH 10
and 35 GH 11 xylanase gene fragments from goat rumen contents.
Most sequences had low identities with known xylanases in
GenBank, implying that these xylanases may be as yet undiscov-
ered. Moreover, these amplified partial sequences had low
similarities and were distantly related based on phylogenetic
analysis (Figures 1 and 2). These results suggest that there are a
large number of unidentified xylanase genes in rumen, corre-
sponding to the large number of uncultured microorganisms in the
rumen. Compared with previous culture-independent methods
such as metagenomic library construction and screening [18],
pyrosequenced rumen metagenome [3], and termite hindgut
metagenomics [29], far more xylanase genes were documented in
our present study, and they were more diverse (Table 3),
suggesting that the PCR-based culture-independent method is
an efficient way to identify xylanase genes in the rumen.
Rumen bacteria and fungi are the key agents to degrade plant
fiber [2], [30]. It has been proposed that rumen fungi play an
important role in the initiation of plant fiber degradation during
fermentative digestion in ruminants [2], [12]. Pyrosequenced
rumen metagenome analysis by Brulc et al. [3] showed that all the
xylanases they examined were bacterial, and no sequence was
closely related to those of fungi. In our present study, however,
seven GH 11 xylanase gene fragments were retrieved directly from
the rumen and showed a close relationship with rumen fungi based
on phylogenetic analysis (Figure 2). Among these seven xylanase
sequences, five had the highest identity to xylanases from N.
patriciarum and Orpinomyces sp. PC-2, and two were closely related to
those from R. flavefaciens FD-1 (Table S2). This result suggests that
horizontal gene transfer from rumen bacteria to fungi may have
occurred [31], [32]. In addition, the distribution of GH 11
xylanases in rumen differed from that in both tundra soil (in which
more fungal than bacterial xylanase genes were detected) [20] and
termite hindgut (in which GH 11 xylanases were mainly from the
phyla Fibrobacteres and Spirochaetes) [29]. This result suggests
that the rumen microenvironment differs from other ecosystems
and has specific xylan-degrading microbial community.
Firmicutes is one of the two most abundant bacteria in rumen
based on 16S rDNA library-based analysis [2], [17]. The
representative genus of xylan-degrading Firmicutes in rumen is
Ruminococcus [14]. Our current study showed that 33 distinct
sequences (30 of GH 11 and 3 of GH 10) are closely related to the
xylanases from Firmicutes, which comprises 38% of the total
distinct sequences. Among them, 25 sequences (24 of GH 11 and 1
of GH 10) are related to Ruminococcus. Abundance analysis also
showed that xylanase genes related to Ruminococcus are dominant in
goat rumen (Table S2). The OTU representative of GH 11
xylanase gene fragments, R8, was subjected to full-length gene
cloning and heterogeneous expression. The recombinant enzyme
showed high activity to xylan. All these results suggest that the
genus Ruminococcus plays an important role in xylan degradation in
the goat rumen.
The Cytophaga-Flexibacter-Bacteroides group represents an-
other abundant group of bacteria in rumen [2], [17]. Based on
phylogenetic analysis, 46 GH 10 xylanase sequences were closely
related to the xylanases from Bacteroides, comprising 53% of the
total distinct sequences. Abundance analysis also showed that
xylanase genes related to Bacteroides were dominant in goat
rumen (Table S1). More than half of them (26/46) were closely
related to the xylanases from Prevotella, which is one of the most
important xylan-degrading bacterial genera in rumen [33], [34].
Many new clades were formed despite a lack of relatives,
suggesting that there are other unidentified xylanase-producing
Bacteroides in goat rumen. In addition to xylanases from
Bacteroides, xylanase fragments from other phyla were also
detected in the goat rumen. Three sequences (GR112, GR126,
and GR146) in the GH 10 clone library had the highest similarity
with xylanases from X. axonopodis and Verrucomicrobiae bacteri-
um. This is the first report of xylanase genes obtained from rumen
that is homologous to Verrucomicrobia and Proteobacteria
xylanase.
In general, rumen microorganisms can be classified into two
major groups: cellulolytic and noncellulolytic [2]. The cellulolytic
microorganisms include cellulolytic rumen bacteria, fungi, and
some protozoa that can utilize both cellulose and xylan as a carbon
source. These microorganisms, such as Fibrobacter succinogenes, R.
flavefaciens, and Neocallimastix frontalis, express both GH 10 and GH
11 xylanases [12], [15, [16]. The noncellulolytic microorganisms
Figure 1. Phylogenetic analysis based on the partial amino acid sequences of GH 10 xylanase genes detected in the goat rumen
contents and their relationship with the reference sequences retrieved from GenBank. This tree was constructed using the neighbor-
joining method (MEGA 4.0). The lengths of the branches indicate the relative divergence among the amino acid sequences. The reference sequences
are marked with a closed diamond (¤) with source strains and GenBank accession numbers in parentheses. The gene fragments (GR67) used for full-
length cloning were marked with a solid square (&). The numbers at the nodes indicate bootstrap values based on 1,000 bootstrap replications and
bootstrap values (.50) are displayed. The scale bar represents 0.1 amino acid substitutions per position.
doi:10.1371/journal.pone.0016731.g001
Xylanases Diversity in Goat Rumen
PLoS ONE | www.plosone.org4 February 2011 | Volume 6 | Issue 2 | e16731

Page 5

Xylanases Diversity in Goat Rumen
PLoS ONE | www.plosone.org5 February 2011 | Volume 6 | Issue 2 | e16731

Page 6

include members of the dominant Gram-negative Cytophaga-
Flexibacter-Bacteroides phylum and Gram-positive Firmicutes,
such as Butyrivibrio fibrisolvens, Roseburia sp., and Eubacterium rectale,
which can use soluble products resulting from plant cell wall
breakdown [2]. These rumen microorganisms produce only GH
10 xylanases. In our study, about 88.5% (46/52) of the GH 10
xylanase sequences were closely related to Bacteroides xylanases,
whereas 85% (30/35) of the GH 11 xylanase sequences were
related to xylanases from cellulolytic Firmicutes, confirming the
primary distribution of GH 10 xylanase genes in noncellulolytic
microorganisms and GH 11 xylanase genes in cellulolytic-
xylanolytic microorganisms. GH 10 and GH 11 xylanases are
distinct from each other in both three-dimensional structure [35]
and mechanism of action [25]. The products of GH 11 xylanases
can be further hydrolyzed by GH 10 enzymes [7]. Therefore, the
different distributions of these two families imply their different
roles in xylan degradation in the rumen.
To date, only a few xylanase genes have been obtained from
uncultured microorganisms in rumen by direct cloning [36] or
screening from the metagenomic library of rumen [18]. In our
current study, two full-length xylanase genes were directly
obtained from the metagenomic DNA of goat rumen contents.
Both enzymes showed high activity towards different natural
xylans and displayed different characteristics (Tables 1 and 2).
XynGR67 had low identity (45% at maximum) with known
xylanases in GenBank and similarity to a characterized cold-active
xylanase [37]. Although the optimum temperature of XynGR67
was 40uC, substantial activity remained at low temperatures
(22.5% at 10uC and 8.2% at 0uC). The enzyme was also
thermolabile, showing a half-life of 15 min at 40uC. Compared
with the cold-active xylanase XynA from Glaciecola mesophila [37],
XynGR67 displayed a much higher kcat/Kmvalue (178.5 vs. 15.4)
at 4uC. These data suggest that XynGR67 has some cold-active
characteristics and is distinct from other known ruminal xylanases.
Using molecular methods, bacterial composition in the
gastrointestinal tract are found to be dependent on host, diet,
age and so on [38], [39], [40]. Of these factors, host species is the
most important one [41], [42]. To expand our assessment of the
xylanase gene diversity in rumen microevironment, the rumen
contents of a Small Tail Han sheep that grazed under the same
conditions as the Boer goat under study were subjected to xylanase
gene analysis (Tables S3 and S4). Both rumens harbor similar
xylan-degrading communities based on the sequence comparison
analysis (Table S5, Figures S1 and S2). Similar or identical
sequences were found in both rumens, and the vast majority of the
amplified xylanase fragments belonged to the two major phyla of
Firmicutes and Bacteroides. Based on UniFrac analysis [43], the
GH 10 xylanase gene fragments in both rumens were marginally
different (P=0.02) while there was no significant difference in that
of GH 11 (P=0.35). Although the abundant sequences in each
rumen are different (Table S5), the predominant OTUs of
different rumens shared high identities (68% for GH 10 and
89% for GH 11, respectively). A small part of sequences are
unique in each rumen (Figure S2), implying that some xylan-
degrading microorganisms might be host specific [41]. The
strategy reported here can be applied to explore other gastroin-
testinal tract ecosystems known to be highly specialized for raw
biomass degradation (i.e., human and insect gut microbiomes) for
novel xylanases.
In conclusion, an efficient culture-independent molecular
method was used to explore the diversity of xylanases in the
rumen ecosystem. Sequence analysis revealed a large number of
unidentified and potential new xylanases in the goat rumen. Full-
length cloning and heterologous expression of some genes further
confirmed their function as active xylanases. The majority of GH
10 xylanase sequences were from noncellulolytic microorganisms,
whereas most of the GH 11 sequences were from cellulolytic
microorganisms, implying their different roles in xylan degrada-
tion in the rumen ecosystem. Moreover, xylanase distribution in
different environments is variable, suggesting that xylan-degrading
microorganisms are environment specific.
Materials and Methods
Ethics statement
All animal studies were followed the regulation for the review
committee of laboratory animal welfare and ethics and protocol
for the review on laboratory animal welfare and ethics, Beijing
Administration Office of Laboratory Animal. The animal
experimentation was approved by the Committee of Laboratory
Animal Welfare and Ethics, Beijing Administration Office of
Laboratory Animal with the approval No. SYXK2008-0007.
Sample collection and DNA extraction
A three-year-old male Boer goat that had grazed on late-fall
pasture in Inner Mongolia, China was selected. Total rumen
contents were collected immediately after slaughtering and stored
at 270uC. The biomass was collected by centrifuging at 17,0006g,
4uC for 30 min. The sample used for DNA extraction were frozen
in liquid nitrogen and ground to a fine powder with a mortar.
Total genomic DNA was extracted following a protocol specific for
high molecular weight DNA from environmental samples [44] and
purified with an Agarose Gel DNA Purification kit (TaKaRa,
Japan). The final DNA concentration was ,80 ng ml–1.
PCR amplification of xylanase gene fragments
Two degenerate primer sets (X10-F: 59-CTACGACTGG-
GAYGTNIBSAAYGA-39 and X10-R: 59-GTGACTCTGGAWR-
CCIABNCCRT-39; and X11-F: 59-AACTGCTACCTGKC-
NITNTAYGGNTGG-39 and X11-R: 59-CCGCACGGACCAG-
TAYTGNKIRAANGT-39) specific for GH 10 and GH 11
xylanases, respectively, were used to amplify xylanase gene
fragments with the purified DNA as a template following the
reaction system and PCR conditions as reported [20]. PCR
products were visualized on an agarose gel, and bands of the
expected size (,260 bp for GH 10 and 210 bp for GH 11
xylanases) were excised and purified with the TaKaRa Agarose
Gel DNA Purification kit.
Cloning and sequencing of PCR products
To construct the clone library for each xylanase family, the
purified PCR products were ligated into vector pGEM-T Easy
Figure 2. Phylogenetic analysis based on the partial amino acid sequences of GH 11 xylanase genes detected in the goat rumen
contents and their relationship with the reference sequences retrieved from GenBank. This tree was constructed using the neighbor-
joining method (MEGA 4.0). The lengths of the branches indicate the relative divergence among the amino acid sequences. The reference sequences
are marked with a closed diamond (¤) with source strains and GenBank accession numbers in parentheses. The gene fragment (R8) used for full-
length retrieving is marked with a solid square (&). The numbers at the nodes indicate bootstrap values based on 1,000 bootstrap replications and
bootstrap values (.50) are displayed. The scale bar represents 0.05 amino acid substitutions per position.
doi:10.1371/journal.pone.0016731.g002
Xylanases Diversity in Goat Rumen
PLoS ONE | www.plosone.org6 February 2011 | Volume 6 | Issue 2 | e16731

Page 7

(Promega, USA) and electroporated into E. coli DH5a competent
cells (TaKaRa). Cells were grown on Luria-Bertani agar plates
containing 100 mg ml21ampicillin, 80 mg ml21X-Gal, and
0.5 mM isopropyl-b-D-1-thiogalactopyranoside (IPTG) at 37uC for
15 h. The positive transformants (white clones) in each library
were randomly picked, amplified with primers M13F (59-
GTAAAACGACGGCCAGT-39)
CAATTTCACACAGGA-39), and sequenced by Sunbiotech
(China) for confirmation.
andM13R(59-GGATAA-
Phylogenetic analysis
Nucleotide sequences of the xylanase gene fragments were
translated into amino acid sequences with EMBOSS Transeq
(http://www.ebi.ac.uk/emboss/transeq) and aligned at the
protein level with known sequences in the GenBank database
with ClustalW. Redundant amino acid sequences were removed
using CD-hit [22] with a 95% sequence identity cut-off. The
protein sequence similarities were assessed by using the
BLASTpprograms(http://www.ncbi.nlm.nih.gov/BLAST/;
until August 15, 2010). Phylogenetic trees were constructed
with MEGA 4.0 [45] using the neighbor-joining method [46].
Confidence for tree topologies was estimated by bootstrap
values based on 1,000 replicates. Twenty-five representative
sequences originating from the rumens or human guts identified
by BLASTp analysis were selected and used as references for
tree construction.
Abundance analysis
Gene abundance of each GH family was estimated using
distance-based operational taxonomic unit and richness determi-
nation (DOTUR) software [23]. Distance matrices of the fragment
sequences were calculated at the protein level with the default
parameters of protdist in PHYLIP (http://evolution.genetics.
washington.edu/phylip.html). Sequences were then assigned to
OTUs based on UPGMA (average linkage clustering) implement-
ed in DOTUR with default parameters of precision (0.01) and
1,000 bootstrap replicates.
Cloning of full-length xylanase genes
Fragments GR67, which showed the phylogenetic novelty
(Figure 1) and only appeared once in the GH 10 library (see Table
S1), and R8, which represented the most abundant OTU in the
GH 11 library (Table S2), were subjected to full-length gene
cloning. The flanking regions of these xylanase gene fragments
were cloned with eight specific primers for each fragment (Table
S6) following the protocol for modified TAIL-PCR [47]. PCR
products were electrophoresed on 1.3% agarose gels, cloned into
pGEM-T Easy vectors, sequenced, and assembled with the known
fragment sequences.
Assembly of the xylanase gene sequences and identification of the
open reading frames were performed with programs in Vector NTI
10.3 (InforMax, USA). The signal peptide sequence was predicted
with SignalP(http://www.cbs.dtu.dk/services/SignalP/). The
Figure 3. pH and temperature activity profiles of purified recombinant XynGR67 and XynR8. A Effect of pH on XynGR67 activity.
Activities at various pHs were assayed at 30uC. B Effect of temperature on XynGR67 activity in McIlvaine buffer (pH 6.0). C Effect of pH on XynR8
activity. Activities at various pHs were assayed at 30uC. D Effect of temperature on XynR8 activity in McIlvaine buffer (pH 5.5). The error bars represent
the mean 6 SD (n=3).
doi:10.1371/journal.pone.0016731.g003
Xylanases Diversity in Goat Rumen
PLoS ONE | www.plosone.org7February 2011 | Volume 6 | Issue 2 | e16731

Page 8

DNAandprotein sequenceidentities/similaritieswere assessed with
the BLASTn and BLASTp programs (http://www.ncbi.nlm.nih.
gov/BLAST/), respectively.
Xylanase expression and activity assay
The coding sequences of two representative xylanase genes
without the signal peptide were amplified using two primer sets
(Table S6), cloned into plasmid pET-22b(+), and transformed into
E. coli BL21 (DE3) competent cells for recombinant expression.
Positive transformants were grown in Luria-Bertani medium
containing 100 mg ml21ampicillin at 37uC to an A600of ,0.6.
Expression was induced with 0.8 mM IPTG at 25uC for 12 h.
Xylanase activity was determined by measuring the release of
reducing sugar from substrate with the 3, 5-dinitrosalicylic acid
method [48]. Reactions containing 0.1 ml of enzyme preparation
and 0.9 ml of 1% (w/v) substrate in McIlvaine buffer (pH 6.0)
were incubated at 30uC for 10 min, followed by addition of 1.5 ml
of 3, 5-dinitrosalicylic acid reagent. The mixture was boiled for
5 min, cooled to room temperature, and the absorbance at
540 nm (A540) was measured. Using a standard curve generated
with
D-xylose, the absorbance was converted into moles of
reducing sugars produced.
Purification and characterization of recombinant
XynGR67 and XynR8
To purify the His-tagged recombinant proteins (XynGR67 and
XynR8), culture supernatant was collected after centrifugation
(12,0006g, 4uC for 15 min) and further concentrated with an
ultrafiltration membrane (PES5000; Sartorius Stedim Biotech,
Germany). The concentrated supernatant was loaded onto a Ni2+-
NTA agarose gel column (Qiagen, Germany) with a linear
imidazole gradient of 20–200 mM in Tris-HCl buffer (20 mM
Tris-HCl, 500 mM NaCl, 10% glycerol, pH 7.6). SDS-PAGE was
used to determine the purity and apparent molecular mass of
recombinant XynGR67. The protein concentration was deter-
mined with the Bradford method [49] with bovine serum albumin
as a standard.
The optimal pH for xylanase activity of the purified
recombinant XynGR67 and XynR8 were determined at 30uC
in buffers of pH 4.0 to 9.0. The enzyme stability at different pHs
were estimated by measuring the residual activity after incubating
the enzyme solution in buffers at pH 3.0–10.0 at 25uC for 1 h.
The buffers used were McIlvaine buffer (0.2 M Na2HPO4/0.1 M
citric acid) for pH 3.0–7.5, and 0.1 M Tris-HCl for pH 7.5–9.0.
The optimal temperature for purified recombinant XynGR67
and XynR8 were determined over the range of 0–80uC in
McIlvaine buffer (pH 5.5 for XynR8 and pH 6.0 for XynGR67).
Thermostability of XynGR67 was determined after pre-incubating
the enzyme in McIlvaine buffer (pH 6.0) at 30uC or 40uC without
substrate for various periods while XynR8 was assayed at 55uC
and 60uC in McIlvaine buffer (pH 5.5).
The Km, Vmax, and kcatvalues for both recombinant xylanases
were determined in McIlvaine buffer (pH 5.5 for XynR8 and
pH 6.0 for XynGR67) containing 1–10 mg ml21birchwood xylan
at 40uC and 55uC, respectively. Kmand Vmaxwere determined
from a Lineweaver-Burk plot with the non-linear regression
computer program GraFit (Erithacus Software, UK). Three
independent experiments were averaged, and each experiment
included three replicates.
The substrate specificity of purified recombinant enzymes were
assayed by incubating the enzyme solution with 1% (w/v)
substrates including birchwood xylan, beechwood xylan, wheat
arabinoxylan or wheat arabinoxylan (insoluble) under standard
conditions. Enzymatic activities against p-nitrophenyl cellobioside
Table 3. Amount of GH 10 and 11 xylanases in the genomes
of Ruminococcus flavefaciens FD-1, Prevotella ruminicola 23,
pyrosequenced rumen, termite hindgut and goat rumen in
this study.
Source GH 10GH 11 References
Ruminococcus flavefaciens FD-1 (Rf)6112
Ruminococcus albus 756 GenBank database
Fibrobacter succinogenes S8523GenBank database
Prevotella ruminicola 2320GenBank database
Prevotella bryantii B1420 GenBank database
Pooled liquid 1026
Fiber-85ND6
Fiber-64716
Fiber-714 ND6
Termite hindgut46 14 46
Goat rumen 52 35 This study
doi:10.1371/journal.pone.0016731.t003
Table 1. Kinetic parameters of XynGR67 and XynR8 towards birchwood xylan.
Enzyme Temperature (6C)
Vmax
(m mmol min–1mg–1)
Km
(mg ml–1)
kcat
(s–1)
kcat/Km
(ml s–1mg–1)
XynGR67 40 2,150.0672.41.4360.16 1,505.0650.4 1,052.5
4500.4614.31.9660.17350.0610.1178.6
XynR8 552,647.9660.93.2760.22 1,306.3630.1399.5
doi:10.1371/journal.pone.0016731.t001
Table 2. Substrate specificity of the purified recombinant
XynGR67 and XynR8.
Substrate
Relative activity (%)
XynGR67XynR8
Birchwood xylan100 100
Beechwood xylan150.763.1 196.862.7
Wheat arabinoxylan53.462.5384.463.4
Wheat arabinoxylan (insoluble) 63.161.954.662.2
pNP-cellobiosideNA NA
pNP-xylosideNA NA
doi:10.1371/journal.pone.0016731.t002
Xylanases Diversity in Goat Rumen
PLoS ONE | www.plosone.org8 February 2011 | Volume 6 | Issue 2 | e16731

Page 9

and p-nitrophenyl xyloside were examined under standard
conditions at a final concentration of 2 mM.
Nucleotide sequence accession numbers
The nucleotide sequences of GH 10 and 11 xylanase gene
fragments were deposited into the GenBank database under
accession numbers FJ919156–FJ919224, HM773534–HM773543,
and HM773544–HM773578. Accession numbers HQ219689 and
HQ219690 were assigned to the full-length xylanase genes xynR8
and xynGR67 of the goat rumen contents, respectively.
Supporting Information
Figure S1
amino acid sequences of GH 10 xylanase genes detected
in the goat and sheep rumen contents and their
relationship with the reference sequences retrieved
from GenBank. This tree was constructed using the neighbor-
joining method (MEGA 4.0). Sequences from goat rumen were
colored in red and those from sheep were in green. Sequence
clusters that unique in each rumen were marked with Sheep
rumen I–IV or Goat rumen A–C. The lengths of the branches
indicate the relative divergence among the amino acid sequences.
The reference sequences are marked with a closed diamond (¤)
with source strains and GenBank accession numbers in parenthe-
ses. The numbers at the nodes indicate bootstrap values based on
1,000 bootstrap replications and bootstrap values (.50) are
displayed. The scale bar represents 0.1 amino acid substitutions
per position.
(PDF)
Phylogenetic analysis based on the partial
Figure S2
amino acid sequences of GH 11 xylanase genes detected
in the goat and sheep rumen contents and their
relationship with the reference sequences retrieved
from GenBank. This tree was constructed using the neighbor-
joining method (MEGA 4.0). Sequences from goat rumen were
colored in red and those from sheep were colored in green. The
lengths of the branches indicate the relative divergence among the
amino acid sequences. The reference sequences are marked with a
closed diamond (¤) with source strains and GenBank accession
numbers in parentheses. The numbers at the nodes indicate
bootstrap values based on 1,000 bootstrap replications and
Phylogenetic analysis based on the partial
bootstrap values (.50) are displayed. The scale bar represents
0.1 amino acid substitutions per position.
(PDF)
Table S1
in the goat rumen contents and their closest relative
based on amino acid sequence identity and similarity.
(DOC)
The GH 10 xylanase gene fragments detected
Table S2
in the goat rumen contents and their closest relative
based on amino acid sequence identity and similarity.
(DOC)
The GH 11 xylanase gene fragments detected
Table S3
in the sheep rumen contents and their closest relatives
based on amino acid sequence identity and similarity.
(DOC)
The GH 10 xylanase gene fragments detected
Table S4
in the sheep rumen contents and their closest relatives
based on amino acid sequence identity and similarity.
(DOC)
The GH 11 xylanase gene fragments detected
Table S5
fragment sequences obtained from the goat and sheep
rumen contents.
(DOC)
Summary of the GH 10 and GH 11 xylanase
Table S6
expression.
(DOC)
Primers used for xylanase genes cloning and
Acknowledgments
We thank Dr. Chenggang Jiang for his assistance with sample collection.
Author Contributions
Conceived and designed the experiments: GW BY. Performed the
experiments: GW XP HL YW. Analyzed the data: GW HH PS.
Contributed reagents/materials/analysis tools: QD HZ. Wrote the paper:
GW KM PY.
References
1. Bunzel M, Steinhart H (2003) Dietary fibre from plant cell walls: chemical and
structural characteristics. Ernahrungs Umschau 50: 469–475.
2. FlintHJ,BayerEA (2008) Plantcellwallbreakdown by anaerobicmicroorganisms
from the Mammalian digestive tract. Ann NY Acad Sci 1125: 280–288.
3. Brulc JM, Antonopoulos DA, Miller ME, Wilson MK, Yannarell AC, et al. (2009)
Gene-centric metagenomics of the fiber-adherent bovine rumen microbiome reveals
forage specific glycoside hydrolases. Proc Natl Acad Sci USA 106: 1948–1953.
4. Kopecny J, Williams AG (1988) Synergism of rumen microbial hydrolases
during degradation of plant polymers. Folia Microbiol (Praha) 33: 208–212.
5. Ohgren K, Bura R, Saddler J, Zacchi G (2007) Effect of hemicellulose and lignin
removal on enzymatic hydrolysis of steam pretreated corn stover. Bioresour
Technol 98: 2503–2510.
6. Yang B, Wyman CE (2004) Effect of xylan and lignin removal by batch and
flowthrough pretreatment on the enzymatic digestibility of corn stover cellulose.
Bioresour Technol 86: 88–98.
7. Collins T, Gerday C, Feller G (2005) Xylanases, xylanase families and
extremophilic xylanases. FEMS Microbiol Rev 29: 3–23.
8. Kulkarni N, Shendye A, Rao M (1999) Molecular and biotechnological aspects
of xylanases. FEMS Microbiol Rev 23: 411–456.
9. Flint, HJ, Whitehead TR, Martin JC, Gasparic A (1997) Interrupted catalytic
domain structures in xylanases from two distantly related strains of Prevotella
ruminicola. Biochim Biophys Acta 1337: 161–165.
10. Dalrymple BP, Swadling Y, Layton I, Gobius KS, Xue GP (1999) Distribution
and evolution of the xylanase genes xynA and xynB and their homologues in
strains of Butyrivibrio fibrisolvens. Appl Environ Microbiol 65: 3660–3667.
11. Flint HJ,Zhang JX, Martin J (1994) Multiplicity andexpression ofxylanasesin the
rumen cellulolytic bacterium Ruminococcus flavefaciens. Curr Microbiol 29: 139–143.
12. Gomez de Segura B, Durand R, Fevre M (1998) Multiplicity and expression of
xylanases in the rumen fungus Neocallimastix frontalis. FEMS Microbiol Lett 164:
47–53.
13. Morrison M, Daugherty SC, Nelson WC, Davidsen T, Nelson KE, et al. (2010)
The FibRumBa database: a resource for biologists with interests in gastrointes-
tinal microbial ecology, plant biomass degradation, and anaerobic microbiology.
Microb Ecol 59: 212–213.
14. Berg Miller ME, Antonopoulos DA, Rincon MT, Band M, Bari A, et al. (2009)
Diversity and strain specificity of plant cell wall degrading enzymes revealed by
the draft genome of Ruminococcus flavefaciens FD-1. PLoS ONE 4: e6650.
15. Flint HJ, Bayer EA, Rincon MT, Lamed R, White BA (2008) Polysaccharide
utilization by gut bacteria: potential for new insights from genomic analysis. Nat
Rev Micro 6: 121–131.
16. Qi M, Nelson KE, Daugherty SC, Nelson WC, Hance IR, et al. (2008) Genomic
differences between Fibrobacter succinogenes S85 and Fibrobacter intestinalis DR7,
identified by suppression subtractive hybridization. Appl Environ Microbiol 74:
987–993.
17. Edwards JE, McEwan NR, Travis AJ, John Wallace R (2004) 16S rDNA library-
based analysis of ruminal bacterial diversity. Antonie Van Leeuwenhoek 86:
263–281.
18. Ferrer M, Golyshina OV, Chernikova TN, Khachane AN, Reyes-Duarte D,
et al. (2005) Novel hydrolase diversity retrieved from a metagenome library of
bovine rumen microflora. Environ Microbiol 7: 1996–2010.
Xylanases Diversity in Goat Rumen
PLoS ONE | www.plosone.org9 February 2011 | Volume 6 | Issue 2 | e16731

Page 10

19. Singh B, Gautam SK, Verma V, Kumar M, Singh B (2008) Metagenomics in
animal gastrointestinal ecosystem: Potential biotechnological prospects. Anaer-
obe 14: 138–144.
20. Wang G, Wang Y, Yang P, Luo H, Huang H, et al. (2010) Molecular detection
and diversity of xylanase genes in alpine tundra soil. Appl Microbiol Biotechnol
87: 1383–1393.
21. Solomon V, Teplitsky A, Shulami S, Zolotnitsky G, Shoham Y, et al. (2007)
Structure-specificity relationships of an intracellular xylanase from Geobacillus
stearothermophilus. Acta Crystallogr D Biol Crystallogr 63: 845–859.
22. Li W, Godzik A (2006) Cd-hit: a fast program for clustering and comparing large
sets of protein or nucleotide sequences. Bioinformatics 22: 1658–1659.
23. Schloss PD, Handelsman J (2005) Introducing DOTUR, a computer program
for defining operational taxonomic units and estimating species richness. Appl
Environ Microbiol 71: 1501–1506.
24. Hayashi H, Abe T, Sakamoto M, Ohara H, Ikemura T, et al. (2005) Direct
cloning of genes encoding novel xylanases from the human gut. Can J Microbiol
51: 251–259.
25. Jeffries TW (1996) Biochemistry and genetics of microbial xylanases. Curr Opin
Biotechnol 7: 337–342.
26. Wouters J, Georis J, Engher D, Vandenhaute J, Dusart J, et al. (2001)
Crystallographic analysis of family 11 endo-b-1,4-xylanase Xyl1 from Streptomyces
sp. S38. Acta Crystallogr D Biol Crystallogr 57: 1813–1819.
27. Raymond JA, Christner BC, Schuster SC (2008) A bacterial ice-binding protein
from the Vostok ice core. Extremophiles 12: 713–717.
28. Beg QK, Kapoor M, Mahajan L, Hoondal GS (2001) Microbial xylanases and
their industrial applications: a review. Appl Microbiol Biotechnol 56: 326–338.
29. Warnecke F, Luginbuhl P, Ivanova N, Ghassemian M, Richardson TH, et al.
(2007) Metagenomic and functional analysis of hindgut microbiota of a wood-
feeding higher termite. Nature 450: 560–565.
30. Dehority BA, Tirabasso PA (2000) Antibiosis between ruminal bacteria and
ruminal fungi. Appl Environ Microbiol 66: 2921–2927.
31. Garcia-Vallve S, Romeu A, Palau J (2000) Horizontal gene transfer of glycosyl
hydrolases of the rumen fungi. Mol Biol Evol 17: 352–361.
32. Gilbert HJ, Hazlewood GP, Laurie JI, Orpin CG, Xue GP (1992) Homologous
catalytic domains in a rumen fungal xylanase: evidence for gene duplication and
prokaryotic origin. Mol Microbiol 6: 2065–2072.
33. Dodd D, Kocherginskaya SA, Spies MA, Beery KE, Abbas CA, et al. (2009)
Biochemical analysis of a b-D-xylosidase and a bifunctional xylanase-ferulic acid
esterase from a xylanolytic gene cluster in Prevotella ruminicola 23. J Bacteriol 191:
3328–3338.
34. Gasparic A, Martin J, Daniel A, Flint HJ (1995) A xylan hydrolase gene cluster
in Prevotella ruminicola B(1)4: sequence relationships, synergistic interactions, and
oxygen sensitivity of a novel enzyme with exoxylanase and b-(1,4)-xylosidase
activities. Appl Environ Microbiol 61: 2958–2964.
35. Biely P, Vrsanska M, Tenkanen M, Kluepfel D (1997) Endo-b-1,4-xylanase
families: differences in catalytic properties. J Biotechnol 57: 151–166.
36. Liu JR, Yu B, Lin SH, Cheng KJ, Chen YC (2005) Direct cloning of a xylanase
gene from the mixed genomic DNA of rumen fungi and its expression in
intestinal Lactobacillus reuteri. FEMS Microbiol Lett 251: 233–241.
37. Guo B, Chen XL, Sun CY, Zhou BC, Zhang YZ (2009) Gene cloning,
expression and characterization of a new cold-active and salt-tolerant endo-b-
1,4-xylanase from marine Glaciecola mesophila KMM 241. Appl Microbiol
Biotechnol 84: 1107–1115.
38. Hopkins MJ, Harp RS, Macfarlane GT (2001) Age and disease related changes
in intestinal bacterial populations assessed by cell culture, 16S rRNA abundance,
and community cellular fatty acid profiles. Gut 48: 198–205.
39. Leser TD, Lindecrona RH, Jensen TK, Jensen BB, Moller K (2000) Changes in
bacterial community structure in the colon of pigs fed different experimental
diets and after infection with Brachyspira hyodysenteriae. Appl Environ Microbiol
66: 3290–3296.
40. Shi PJ, Meng K, Zhou ZG, Wang YR, Diao QY, Yao B (2008) The host species
affects the microbial community in the goat rumen. Lett Appl Microbiol 46:
132–135.
41. Dick LK, Bernhard AE, Brodeur TJ, Santo Domingo JW, Simpson JM, et al.
(2005) Host distributions of uncultivated fecal Bacteroidales bacteria reveal
genetic markers for fecal source identification. Appl Environ Microbiol 71:
3184–3191.
42. Weimer PJ, Stevenson DM, Mantovani HC, Man SL (2010) Host specificity of
the ruminal bacterial community in the dairy cow following near-total exchange
of ruminal contents. J Dairy Sci 93: 5902–5912.
43. Lozupone C, Knight R (2005) UniFrac: a new phylogenetic method for
comparing microbial communities. Appl Environ Microbiol 71: 8228–8235.
44. Brady SF (2007) Construction of soil environmental DNA cosmid libraries and
screening for clones that produce biologically active small molecules. Nat Protoc
2: 1297–1305.
45. Tamura K, Dudley J, Nei M, Kumar S (2007) MEGA4: Molecular Evolutionary
Genetics Analysis (MEGA) Software Version 4.0. Mol Biol Evol 24: 1596–1599.
46. Saitou N, Nei M (1987) The neighbor-joining method: a new method for
reconstructing phylogenetic trees. Mol Biol Evol 4: 406–425.
47. Huang H, Wang G, Zhao Y, Shi P, Luo H, et al. (2010) Direct and efficient
cloning of full-length genes from environmental DNA by RT-qPCR and
modified TAIL-PCR. Appl Microbiol Biotechnol 87: 1141–1149.
48. Miller GL, Blum R, Glennon WE, Burton AL (1960) Measurement of
carboxymethylcellulase activity. Anal Biochem 1: 127–132.
49. Bradford MM (1976) A rapid and sensitive method for the quantitation of
microgram quantities of protein utilizing the principle of protein-dye binding.
Anal Biochem 72: 248–254.
Xylanases Diversity in Goat Rumen
PLoS ONE | www.plosone.org 10 February 2011 | Volume 6 | Issue 2 | e16731