Molecular analysis of two glycoside hydrolase family 8 enzymes from the human gut bacterium, Bacteroides intestinalis DSM17393.

Abstract

Xylan is an abundant plant cell wall polysaccharide and is a dominant component of dietary fiber. Bacteria in the distal human
gastrointestinal tract produce xylanase enzymes to initiate the degradation of this complex heteropolymer. These xylanases
typically derive from glycoside hydrolase (GH) families 10 and 11; however, analysis of the genome sequence of the xylan-degrading
human gut bacterium Bacteroides intestinalis DSM 17393 revealed the presence of two putative GH8 xylanases. In the current study, we demonstrate that the two genes encode
enzymes that differ in activity. The xyn8A gene encodes an endoxylanase (Xyn8A), and rex8A encodes a reducing-end xylose-releasing exo-oligoxylanase (Rex8A). Xyn8A hydrolyzed both xylopentaose (X5) and xylohexaose (X6) to a mixture of xylobiose (X2) and xylotriose (X3), while Rex8A hydrolyzed X3 through X6 to a mixture of xylose (X1) and X2. Moreover, rex8A is located downstream of a GH3 gene (xyl3A) that was demonstrated to exhibit β-xylosidase activity and would be able to further hydrolyze X2 to X1. Mutational analyses of putative active site residues of both Xyn8A and Rex8A confirm their importance in catalysis by these
enzymes. Recent genome sequences of gut bacteria reveal an increase in GH8 Rex enzymes, especially among the Bacteroidetes, indicating that these genes contribute to xylan utilization in the human gut.

Full text

Two New Xylanases with Different Substrate Specificities from the
Human Gut Bacterium Bacteroides intestinalis DSM 17393
Pei-Ying Hong,
a
* Michael Iakiviak,
a,b
Dylan Dodd,
b,c
* Meiling Zhang,
a,b
* Roderick I. Mackie,
a,b
Isaac Cann
a,b,c
Department of Animal Sciences,
a
Institute for Genomic Biology,
b
and Department of Microbiology,
c
University of Illinois at Urbana-Champaign, Urbana, Illinois, USA
Xylan is an abundant plant cell wall polysaccharide and is a dominant component of dietary fiber. Bacteria in the distal human
gastrointestinal tract produce xylanase enzymes to initiate the degradation of this complex heteropolymer. These xylanases typi-
cally derive from glycoside hydrolase (GH) families 10 and 11; however, analysis of the genome sequence of the xylan-degrading
human gut bacterium Bacteroides intestinalis DSM 17393 revealed the presence of two putative GH8 xylanases. In the current
study, we demonstrate that the two genes encode enzymes that differ in activity. The xyn8A gene encodes an endoxylanase
(Xyn8A), and rex8A encodes a reducing-end xylose-releasing exo-oligoxylanase (Rex8A). Xyn8A hydrolyzed both xylopentaose
(X
5
) and xylohexaose (X
6
) to a mixture of xylobiose (X
2
) and xylotriose (X
3
), while Rex8A hydrolyzed X
3
through X
6
to a mixture
of xylose (X
1
) and X
2
. Moreover, rex8A is located downstream of a GH3 gene (xyl3A) that was demonstrated to exhibit ␤-xylosi-
dase activity and would be able to further hydrolyze X
2
to X
1
. Mutational analyses of putative active site residues of both
Xyn8A and Rex8A confirm their importance in catalysis by these enzymes. Recent genome sequences of gut bacteria reveal
an increase in GH8 Rex enzymes, especially among the Bacteroidetes, indicating that these genes contribute to xylan utiliza-
tion in the human gut.
Xylan is an abundant plant cell wall polysaccharide that consists
ofa␤-(1,4)-linked xylosyl backbone with various degrees of
polymerization and substitution. Xylans are the major polysac-
charides in cereal-derived food products, fruits, and vegetables
that are consumed daily by humans. The hydrolysis and fermen-
tation of plant cell wall polysaccharides, inclusive of xylan, are
important metabolic processes that contribute approximately
10% of the caloric requirements in human hosts (1). The fermen-
tation process yields volatile fatty acids, including butyrate, which
is a compound essential for maintaining the integrity of colonic
epithelial cells (2–4). Due to the heterogeneous nature of xylan,
its degradation requires a combination of different enzymes, in-
cluding endoxylanases, ␤-xylosidases, ␣-L-arabinofuranosidases,
␣-glucuronidases, ferulic acid esterases, and acetylxylan esterases
(5). Humans do not possess these enzymes; therefore, degradation
of xylan in the intestine is a function exclusively undertaken by
commensal microorganisms (6,7).
Among the human gut microbiota, Bacteroides spp. are the
most numerically dominant xylanolytic bacteria (8,9). Past stud-
ies have identified Bacteroides eggerthii,B. ovatus,B. fragilis,B.
vulgatus,B. intestinalis,B. cellulosilyticus, and B. xylanisolvens as
the major xylanolytic members of this genus (10–17). Many of the
Bacteroides spp. possess xylanases, which are required to depoly-
merize xylans by cleaving the long polymeric chain into shorter
chains of xylose units. The genome sequences of Bacteroides ovatus
ATCC 8483 and Bacteroides intestinalis DSM 17393 revealed that
these organisms possess the most highly expanded repertoire of
glycoside hydrolase and polysaccharide lyase genes among all gut
bacteria sequenced to date (18). These genes are arranged in poly-
saccharide utilization loci (PULs) that are specifically regulated at
the transcriptional level during growth with the cognate polysac-
charides (19). In B. ovatus ATCC 8483, a relatively large number of
genes are regulated at the transcriptional level during growth on
xylan and these genes are highly expressed in vivo (20), indicating
that they are important for xylan degradation by this bacterium.
Despite the large number of genes induced by xylan, biochemical
evidence to define the substrate specificities of these enzymes is
lacking. This information is particularly important in helping to
define the metabolic potential of these abundant gut bacteria.
The majority of xylanases that have been studied derive from
the glycoside hydrolase (GH) families 10 and 11, with a relative
minority belonging to GH families 5, 8, 30, and 43 (21,22). Com-
pared to xylanases in the GH10 and GH11 families, the substrate
preference and hydrolysis product profiles of xylanases in GH
families 5, 8, 30, and 43 have not been extensively studied. As of
January 2014, the CAZy database has 729 entries in the GH8 fam-
ily, with a total of 56 enzymes listed as characterized. Among these
entries, six have been shown to degrade xylan, including endoxy-
lanases (23–26) and reducing-end xylose-releasing exo-oligoxyla-
nases (27,28). The genome map of B. intestinalis DSM 17393
revealed the presence of two GH family 8 genes (BACINT_04210
and BACINT_00927) (Fig. 1). BACINT_04210 is located in a
polysaccharide utilization locus (PUL) consisting of 11 genes
(BACINT_04220 to BACINT_ 04210). BACINT_00927 is located
downstream of a predicted GH3 glycosidase (BACINT_00926).
Received 25 September 2013 Accepted 10 January 2014
Published ahead of print 24 January 2014
Editor: M. J. Pettinari
Address correspondence to Isaac Cann, icann@illinois.edu.
* Present address: Pei-Ying Hong, King Abdullah University of Science and
Technology (KAUST), Environmental Science and Engineering, Water Desalination
and Reuse Center, Thuwal, Saudi Arabia; Meiling Zhang, School of Life Science,
East China Normal University, Shanghai, China; Dylan Dodd, Department of
Pathology, Stanford University School of Medicine, Stanford, California, USA.
P.-Y.H. and M.I. contributed equally to this article.
Supplemental material for this article may be found at http://dx.doi.org/10.1128
/AEM.03176-13.
Copyright © 2014, American Society for Microbiology. All Rights Reserved.
doi:10.1128/AEM.03176-13
2084 aem.asm.org Applied and Environmental Microbiology p. 2084 –2093 April 2014 Volume 80 Number 7

The genomic context of these genes indicates possible roles in
xylan degradation.
In this study, the protein coding sequences for BACINT_
00926, BACINT_00927, and BACINT_04210 were cloned, and
the proteins were expressed in Escherichia coli and purified to near
homogeneity. It is hypothesized that these three genes are in-
volved in xylan degradation, and therefore, the activities of the
three proteins against xylo-oligosaccharides and natural xylans
were evaluated. The important catalytic residues in the two GH8
enzymes were also evaluated by site-directed mutagenesis. Results
from this study reveal that BACINT_04210 encodes an endoxyla-
nase (Xyn8A), BACINT_00926 encodes a ␤-xylosidase (Xyl3A),
and BACINT_00927 encodes a reducing-end xylose-releasing
exo-oligoxylanase (Rex8A). Xyl3A cleaves xylobiose released by
Rex8A, thus representing an alternative xylan-degrading pathway
in gut bacteria involving GH8 and GH3 enzymes.
MATERIALS AND METHODS
Materials and strains. Bacteroides intestinalis DSM 17393 (29) was ob-
tained from the DSMZ (Braunschweig, Germany). Escherichia coli XL-10
Gold competent cells and E. coli BL-21 CodonPlus(DE3) RIL competent
cells were obtained from Agilent (Santa Clara, CA). Medium viscosity
wheat arabinoxylan (WAX) and xylo-oligosaccharides were obtained
from Megazyme (Bray, Ireland). All other reagents were obtained from
Sigma-Aldrich or Fisher Scientific.
Gene cloning, expression, and protein purification. B. intestinalis
DSM 17393 genomic DNA was extracted using the UltraClean Soil DNA
isolation kit from Mo-Bio (Carlsbad, CA) according to the manufactur-
er’s protocol. The concentrations of total DNA were quantified using the
Qubit dsDNA BR assay kit (Invitrogen, Grand Island, NY). Oligonucleo-
tide primers used for amplifying xyl3A,rex8A, and xyn8A (Table 1) were
engineered to include 5=and 3=extensions for subsequent ligation-inde-
pendent cloning (LIC). Signal peptide cleavage sites were predicted at the
N terminus of each protein using SignalP v4.1 (http://www.cbs.dtu.dk
/services/SignalP/)(30). Thus, to ensure that the protein accumulates
within the E. coli cells, the forward primers were designed to amplify the
genes beginning with the codon immediately downstream of the pre-
dicted peptidase cleavage site. The coding sequences for these three genes
were amplified by PCR using the PicoMaxx high-fidelity PCR mix from
Agilent, and the resulting amplicons were purified using the Wizard DNA
purification kit (Promega, Madison, WI). The purified amplicons were
digested using the exonuclease activity of T4 DNA polymerase, annealed
with a similarly digested pET-46b vector (EMD Chemicals, Darmstadt,
Germany), and introduced into E. coli XL10 Gold competent cells by
chemical transformation. Individual colonies were picked and cultured
overnight in lysogeny broth (LB) supplemented with ampicillin (100 ␮g/
ml), and plasmid DNA was purified using a Plasmid Minikit from Qiagen
4220 4219 4218 4217 4216 4215 4213 4212 4211 4210 4209 4208
50799005084000508810050922005096300
5100400 5075800
0926 0927 0928
0924 0925
0923
10000 120008000 14000 16000 18000 20000 22000
0929 0930 0931 0932
0921
xusC xusD xusC xusD hyp. xyn10A xyn5A CE1 CE6/GH95 xyn8A integrase xynR
xyl3A rex8A arylsulfatase
ATPase methyl
transferase
hyp. GH43 GH43 hyp. hyp.
D-ala-D-ala
dipeptidase
Contig 8
Contig 7
A
.
B.
FIG 1 Genomic context for the two Bacteroides intestinalis GH8 genes. (A) The xyn8A gene (BACINT_04210) is located within a large xylan-specific polysac-
charide utilization locus (BACINT_04220 to BACINT_04210). An integrase gene (BACINT_04209) and an xynR transcriptional regulator homolog
(BACINT_04208) are also located in close proximity to the xyn8A gene. (B) The rex8A gene (BACINT_00927) is not located within a polysaccharide utilization
locus but is preceded immediately by xyl3A (BACINT_00926), which encodes a GH family 3 ␤-xylosidase.
TABLE 1 Primers used in this study
Gene use and name Orientation Sequence(5=–3=)
a
Desired mutation
Cloning
a
xyl3A (BACINT_00926) Forward GACGACGACAAGATGCAACCTCCCTACAAAAACCC
Reverse GAGGAGAAGCCCGGTTATTTAACAATGACTGGTATCGCC
rex8A (BACINT_00927) Forward GACGACGACAAGATGGACCCGACAAAGCCCTGGGATAAAG
Reverse GAGGAGAAGCCCGGTTACTTTTCAGGGAAGATGATGCGGTAG
xyn8A (BACINT_04210) Forward GACGACGACAAGATGCATCCTGTTCAGGAAGACAGTAGTGGGG
Reverse GAGGAGAAGCCCGGTTACTTGATAATCCGGAAATTGCCACTGACATGC
Mutagenesis
b
rex8A E90A Forward CATGATGTCCGCACCGCAGGTATGTCTTACGGA Glu90Ala
rex8A D148A Forward GGCCCCGCCTCCGCCGGAGAACTTTACT Asp148Ala
rex8A D286A Forward GATGCTTTCCGCTTCGCTTCTTGGCGTGTACCG Asp286Ala
xyn8A E104A Forward AATCAGGATGTACGTACAGCAGGAATGTCCTATGGAATG Glu104Ala
xyn8A D164A Forward AGGAGCCAAGTTGCGCTTCTGCTGGTGAAATTTATTTTATAACT Asp164Ala
xyn8A D303A Forward CAAGAAGATATCAGTTTGCTGCTCTTCGCTGTGCCAT Asp303Ala
a
Underlined sequences indicate the incorporated T4 exonuclease digestion sites from the pET-46b Ek-LIC cloning kit.
b
Underlined sequences indicate the substituted codon. Primers were designed using the Agilent QuikChange Primer Design tool
(http://www.genomics.agilent.com/primerDesignProgram.jsp).
GH8 Enzymes from B. intestinalis DSM 17393
April 2014 Volume 80 Number 7 aem.asm.org 2085

(Valencia, CA). The cloned inserts were then sequenced to confirm the
integrity of the genes (W. M. Keck Center for Comparative and Func-
tional Genomics at the University of Illinois).
The recombinant pET-46 EK/LIC plasmids containing the cloned
genes (xyl3A,rex8A,orxyn8A) were introduced into E. coli BL-21 Codon-
Plus(DE3) RIL chemically competent cells (Agilent, Santa Clara, CA) by
the heat shock transformation method and cultured overnight on LB agar
plates supplemented with ampicillin (100 ␮g/ml) and chloramphenicol
(50 ␮g/ml) at 37°C. After 12 h, a single colony was used to inoculate fresh
LB (10 ml) supplemented with the same antibiotics and cultured with
vigorous aeration at 37°C for 8 h. The culture was then diluted into 1 liter
of fresh LB supplemented with ampicillin (100 ␮g/ml) and chloramphen-
icol (50 ␮g/ml) in 2.8-liter Fernbach flasks, and the cultures were incu-
bated at 37°C with vigorous aeration by shaking at 250 rpm. When the
culture reached an optical density at 600 nm (OD
600
) of 0.3, isopropyl
␤-D-thiogalactopyranoside (IPTG) was added to a final concentration of
0.1 mM, and the culture was incubated at 16°C for an additional 16 h. The
cells were then harvested by centrifugation (4,000 ⫻g, 15 min, 4°C). The
cell pellets were resuspended in 30 ml of ice-cold lysis buffer (50 mM
Tris-HCl, 300 mM NaCl, pH 7.5) and ruptured by two passages through
an EmulsiFlex C-3 cell homogenizer from Avestin (Ottawa, Canada). The
cell lysates were clarified by centrifugation at 12,000 ⫻gfor 30 min at 4°C.
The recombinant proteins were then purified from the clarified lysates
using Talon metal affinity resin (ClonTech, Mountain View, CA) accord-
ing to the supplier’s protocol with slight modifications to the binding (50
mM Tris-HCl, 300 mM NaCl, pH 7.5) and elution (50 mM Tris-HCl, 300
mM NaCl, 250 mM imidazole, pH 7.5) buffers. Aliquots of eluted frac-
tions were analyzed by sodium dodecyl sulfate-polyacrylamide gel elec-
trophoresis (SDS-PAGE) according to Laemmli’s method (31), and pro-
tein bands were visualized by staining with Coomassie brilliant blue
G-250. The protein concentrations were calculated by absorbance spec-
troscopy at 280 nm using a NanoDrop 1000 instrument (Thermo Scien-
tific, Waltham, MA) with extinction coefficients of 120,670 M
⫺1
cm
⫺1
,
117,940 M
⫺1
cm
⫺1
, and 103,625 M
⫺1
cm
⫺1
for Xyl3A, Rex8A, and
Xyn8A, respectively.
Size exclusion chromatography. The quaternary structures of Xyn8A
and Rex8A were analyzed by size exclusion chromatography using a Su-
perdex 200 10/300 GL size exclusion column affixed to an AKTAxpress
FPLC unit. One hundred microliters of Xyn8A (1 mg/ml), Rex8A (1 mg/
ml), or a gel filtration standard mixture (Bio-Rad, Hercules, CA) was
loaded onto the column preequilibrated with a buffer composed of 50
mM sodium phosphate–150 mM NaCl. The pH of the buffer was adjusted
to 6.0 for Rex8A and 6.5 for Xyn8A. The proteins were eluted in the same
buffer at a flow rate of 0.5 ml/min. A calibration curve of molecular mass
versus retention time was constructed with the gel filtration standards,
and the apparent molecular masses of the two proteins were calculated by
comparison of experimental retention times with calibration standards.
CD spectroscopy. Circular dichroism (CD) spectra were obtained us-
ing a J-815 spectropolarimeter from Jasco (Easton, MD). The enzymes
were exchanged into CD buffer (50 mM sodium phosphate, pH 6.0 for
Rex8A or pH 6.5 for Xyn8A) using a HiPrep 26/10 desalting column
affixed to an AKTAxpress FPLC unit. The proteins were then diluted in
CD buffer to a final concentration of 1 ␮M, and the far-UV CD spectra
were recorded from 260 to 190 nm with a wavelength step of 0.1 nm. The
secondary-structure contents of Xyn8A and Rex8A were calculated using
the DichroWeb online circular dichroism analysis server with the refer-
ence set 4 optimized for 190 to 240 nm (32).
Evaluation of hydrolysis of xylo-oligosaccharides. The capacities of
Xyn8A, Rex8A, and Xyl3A to hydrolyze xylo-oligosaccharides were tested
with xylose (X
1
), xylobiose (X
2
), xylotriose (X
3
), xylotetraose (X
4
), xylo-
Rex8A
Xyn8A
A
.B. C.
D. E. F.
X1
A1
X2
X3
X4
X5
Rex8A
Xyn8A
----
-- --
++
++
WAX OSX
MW
(kDa)
66
45
31
X1
Xyn8A -+
X2
-+
X3
-+
X4
-+
X5
-+
X6
-+
X1
X2
X3
X4
X5
X6
X1
Rex8A -+
X2
-+
X3
-+
X4
-+
X5
-+
X6
-+
X1
X2
X3
X4
X5
X6
Xyn8A
Rex8A
wavelength (nm)
0
30.9 ± 0.1 kDa
elution volume (mL)
A280nm (mAU)
Xyn8A
020
0
38.6 ± 0.2 kDa
Rex8A
10 30
100
200
100
200
X6
200
220 240 260
-20
0
20
40
CD (mdeg)
FIG 2 Thetwo Bacteroides intestinalis GH8 genes encode xylan-degrading enzymes with distinct properties. (A) Purification of the two GH8 proteins, Xyn8A and
Rex8A. The proteins were produced in their recombinant forms in E. coli, purified by immobilized metal affinity chromatography, resolved by 12% SDS-PAGE,
and stained with Coomassie brilliant blue G250. (B) Predicted secondary structure of the two GH8 proteins. Circular dichroism (CD) spectra were obtained for
Rex8A and Xyn8A in the far-UV region (190 to 260 nm). (C) Gel filtration chromatography. The sizes of purified Rex8A and Xyn8A were determined by size
exclusion fast-protein liquid chromatography (FPLC). The molecular masses of Rex8A and Xyn8A were calculated from the retention time of the peak
absorbance by comparison with calibration standards having known molecular masses. (D) Hydrolysis of model xylans. Rex8A and Xyn8A were incubated with
wheat arabinoxylan (WAX) or oat spelt xylan (OSX) for 16 h, and the products of hydrolysis were analyzed by thin-layer chromatography (TLC). (E and F)
Hydrolysis of xylo-oligosaccharides (XOS). Rex8A (E) or Xyn8A (F) was incubated with XOS of different lengths, and the products were analyzed by TLC.
Hong et al.
2086 aem.asm.org Applied and Environmental Microbiology

pentaose (X
5
), and xylohexaose (X
6
). Substrates (5 mM) were diluted in
phosphate reaction buffer (50 mM sodium phosphate, 150 mM NaCl, pH
6.0 for Rex8A or pH 6.5 for Xyn8A and Xyl3A); reactions were initiated by
the addition of enzyme (0.5 ␮M final concentration), and the mixtures
were incubated at 37°C for 16 h. Preliminary experiments revealed pH
optima of 6.0 for Rex8A and 6.5 for Xyn8A and Xyl3A (see Fig. S1 in the
supplemental material). Reaction mixtures were terminated by heating at
99°C for 10 min, and 1-␮l aliquots of samples were spotted onto silica gel
thin-layer chromatography (TLC) plates (60 Å, 250-␮m thickness)
(Whatman, Piscataway, NJ). The products of hydrolysis were then re-
solved by one ascent with a mobile phase composed of n-butanol, acetic
acid, and water (10:5:1). The plates were then dried, and products were
visualized by spraying with a mixture of sulfuric acid (10%, vol/vol), or-
cinol (0.1%, wt/vol), and methanol (50%, vol/vol) and developed by heat-
ing at 80°C for 15 min.
To evaluate the hydrolysis patterns over time, enzyme concentrations
that gave near-complete hydrolysis over the 3-h reaction period for each
enzyme-substrate combination were first determined. For Rex8A, the en-
zyme concentration used was 50 nM (X
3
through X
6
), whereas for Xyn8A
the concentration was 500 nM (X
4
), 20 nM (X
5
),or2nM(X
6
). Reaction
mixtures were prepared with substrate (1 mM) and enzyme, incubated at
37°C, and were then taken at 10-, 30-, 60-, 120-, 180-, and 360-min inter-
vals for heat inactivation. The products of hydrolysis were first diluted
50-fold and then analyzed using high-performance anion exchange chro-
matography (HPAEC) with a System Gold high-performance liquid chro-
matography (HPLC) instrument (Beckman Coulter, Fullerton, CA). The
instrument was fitted with a CarboPac PA1 guard column (4 by 50 mm)
and a CarboPac PA1 analytical column (4 by 250 mm) from Dionex Cor-
poration (Sunnyvale, CA) and with a Coulochem III electrochemical de-
tector (ESA Biosciences, Chelmsford, MA). Peak retention times and peak
areas from sample chromatograms were compared to those obtained us-
ing commercially available xylo-oligosaccharides analyzed as standards.
Evaluation of hydrolysis of natural xylans. The capacity of the en-
zymes to hydrolyze natural xylans (i.e., wheat arabinoxylan [WAX] and
oat spelt xylan [OSX]) was assessed by dissolving WAX or OSX (2.5 mg/
ml) in phosphate buffer (50 mM, 150 mM NaCl,; pH 6.0 for Rex8A or pH
6.5 for Xyn8A; final volume, 100 ␮l). Reactions were initiated by the
addition of enzyme (final concentration, 0.5 ␮M), and reaction mixtures
were incubated at 37°C for 16 h. The reaction mixtures were then heat
inactivated at 99°C for 10 min and centrifuged for 10 min at 15,000 ⫻g,
and hydrolysis products in the supernatants were evaluated by TLC as
described above. The concentrations of reducing sugars were quantified
by the PAHBAH (para-hydroxybenzoic acid hydrazide) method as previ-
ously described (33).
Site-directed mutagenesis. Mutagenesis was performed using the
QuikChange Lightning Multi Site-directed mutagenesis kit (Agilent
Technologies, Santa Clara, CA). Mutagenic primers were designed with
the QuikChange Primer Design tool (Agilent Technologies) (Table 1).
Mutations were prepared according to the QuikChange protocol with
either the rex8A or the xyn8A pET-46b plasmid as the DNA template. The
residual parent plasmid was then digested by incubation with DpnI over-
night at 37°C, and the resulting DNA was transformed into E. coli XL10-
Gold ultracompetent cells by the heat shock method. Colonies were
picked and cultured, and plasmids were purified using a Plasmid Minikit
(Qiagen, Valencia, CA). The purified plasmid DNA was then sequenced to
ensure that the appropriate mutations were introduced and that the rest of
the gene sequences remained unchanged. Expression and purification of
the mutant recombinant proteins were performed as described above for
the wild-type (WT) proteins.
TABLE 2 Analysis of CD spectra using DichroWeb
a
Protein (ORF no.)
%
␣-helix
%
␤-sheet
%
␤-turn
%
Unordered
Xyn8A (BACINT_04210) 71 ⫾69⫾39⫾410⫾3
Rex8A (BACINT_00927) 81 ⫾26⫾16⫾26⫾2
a
CD spectra were recorded in the far-UV range utilizing a J-815 CD
spectropolarimeter. The buffer composition was 50 mM sodium phosphate, pH 6.0 for
Rex8A or pH 6.5 for Xyn3A. The spectra were recorded from 190 to 260 nm at a scan
rate of 50 nm/s and a 0.1-nm wavelength step with five accumulations. Data are from
three independent experiments and are presented as means ⫾standard deviations of
the means. The spectra were uploaded onto the DichroWeb online server and analyzed
as described in Materials and Methods. ORF, open reading frame.
0
0.2
0.4
0.6
0.8
1.0
0
0.5
1.0
1.5
0
0.5
1.0
1.5
2.0
X1
X2
X3
X4
X5
X6
1
2
3
[Sugar] (mM)[Sugar] (mM)
A
.B.
C. D.
Time (min)
0 60 120 180 0 60 120 180
0
0 60 120 180 0 60 120 180
Time (min)
FIG 3 Hydrolysis of XOS by Rex8A (BACINT_00927). Rex8A was incubated with xylotriose (A), xylotetraose (B), xylopentaose (C), and xylohexaose (D), and
products were analyzed at designated intervals using high-performance anion exchange chromatography (HPAEC). Abbreviations are as follows: X6, xylo-
hexaose; X5, xylopentaose; X4, xylotetraose; X3, xylotriose; X2, xylobiose; and X1, xylose). Concentrations were determined by comparison to calibration curves
constructed with known concentrations of sugars. Results are presented as means and standard deviations from three individual experiments.
GH8 Enzymes from B. intestinalis DSM 17393
April 2014 Volume 80 Number 7 aem.asm.org 2087

Hydrolysis of pNP-linked sugars. The enzyme-catalyzed hydrolysis
of para-nitrophenyl (pNP)-linked monosaccharide substrates was as-
sayed by using a thermostated Synergy II multimode microplate reader
(BioTek Instruments Inc., Winooski, VT). A library of pNP substrates was
screened for activity as described previously (34). The substrates (1 mM)
in 100 ␮l phosphate buffer (50 mM sodium phosphate, 150 mM NaCl, pH
6.5) were incubated at 37°C in the presence or absence of Xyl3A (0.1 ␮M
for pNPXyl; 0.5 ␮M for pNPAra; 1 ␮M for pNPGal and pNPGlu) for 30
min, and the amount of pNP release was determined by continuously
monitoring the absorbance at 400 nm. The path length correction feature
of the instrument was employed to convert the absorbance values re-
corded to correspond to those for a 1-cm path length. The extinction
coefficient for pNP at pH 6.5 and at a wavelength of 400 nm was measured
to be 3.179 mM
⫺1
cm
⫺1
.
RESULTS
Identification of two xylan-specific GH8 genes in Bacteroides
intestinalis.Genomic analysis of Bacteroides intestinalis DSM
17393 revealed the presence of two GH8 genes, BACINT_04210
and BACINT_00927, which encode the proteins Xyn8A and
Rex8A, respectively. The genomic context for these genes supports
their predicted roles in xylan degradation. To illustrate,
BACINT_04210 is located in a polysaccharide utilization locus
(PUL) consisting of 11 genes (BACINT_04220 to BACINT_
04210). Included in this PUL is a large gene cluster consisting of
two tandem repeats of susC and susD orthologs (xusC and xusD)
followed by a hypothetical gene (BACINT_04216) and a GH10
endoxylanase (BACINT_04215) (Fig. 1A). Downstream are a hy-
pothetical protein (BACINT_04214), a GH5 endoxylanase
(xyn5A, BACINT_04213) (17), a family 1 carbohydrate esterase
(BACINT_04212), a two-domain CE6/GH95 gene (BACINT_
04211), and the GH8 gene BACINT_04210. Following this cluster
is a predicted integrase, an observation that suggests that this locus
may be part of an integrative and conjugative element. Diver-
gently transcribed relative to the gene cluster described above is a
homolog of xynR, a hybrid two-component system regulator that
regulates xylanase gene expression in Prevotella bryantii B
1
4(35).
The second GH8 gene (BACINT_00927) is located directly down-
stream of a predicted GH3 glycosidase (Fig. 1B).
Both of these proteins, Xyn8A and Rex8A, possess putative
signal peptides with predicted signal peptidase II cleavage sites,
followed by GH8 domains. Amino acid sequence alignments of
the GH8 domains demonstrated 39% amino acid sequence iden-
tity (see Fig. S1 in the supplemental material).
Cloning, expression, and purification of Xyn8A and Rex8A.
To determine whether or not the two GH8 genes encode proteins
with redundant biochemical properties, they were individually
cloned into an E. coli expression vector. The two proteins were
then expressed as soluble recombinant hexahistidine fusion pro-
teins for subsequent purification via cobalt immobilized metal
affinity chromatography (IMAC). In a single IMAC step, the two
recombinant proteins were purified to near homogeneity as as-
sessed by SDS-PAGE (Fig. 2A).
Recombinant Xyn8A and Rex8A have similar secondary
structures. To compare the secondary structural compositions of
the two recombinant proteins, CD spectra in the far-UV region
(190 to 260 nm) were collected (Fig. 2B). The raw data were then
uploaded onto the DichroWeb server, and secondary structural
elements were predicted by comparison with a standardized data
set. The recombinant proteins both yielded CD spectra indicating
the presence of ␣-helix, ␤-sheet, and ␤-turn secondary structural
elements. Comparison of the DichroWeb results using the Stu-
dent ttest revealed no significant differences in ␣-helices,
␤-sheets, ␤-turns, or unordered regions between the two proteins
[sugar] (mM)
A
.B.
C.
0
0.5
1.0
1.5
2.0
0
0.3
0.6
0.9
1.2
0
0.5
1.0
1.5
Time (min)
Time (min)Time (min)
0 60 120 180 0 60 120 180
060120180
[sugar] (mM)
X1
X2
X3
X4
X5
X6
FIG 4 Hydrolysis of XOS by Xyn8A (BACINT_04210). Xyn8A was incubated with xylotetraose (A), xylopentaose (B), and xylohexaose (C), and products were
analyzed at designated intervals using high-performance anion exchange chromatography (HPAEC). Abbreviations are as follows: X6, xylohexaose; X5, xylo-
pentaose; X4, xylotetraose; X3, xylotriose; X2, xylobiose; and X1, xylose. Concentrations were determined by comparison to calibration curves constructed with
known concentrations of sugars. Results are presented as means and standard deviations from three individual experiments.
Hong et al.
2088 aem.asm.org Applied and Environmental Microbiology

(Table 2). These data suggest that when expressed as recombinant
proteins, Xyn8A and Rex8A have similar overall secondary struc-
tural properties.
Xyn8A and Rex8A are both monomers in solution. To evalu-
ate whether the two proteins exhibited similar quaternary struc-
tures and to rule out whether the two proteins exist as aggregates,
their apparent molecular weights were determined by gel filtration
analysis. Both proteins eluted from the column well after the size
exclusion limit, indicating that neither of the two proteins existed
as large aggregates in solution. The elution volumes of the two
proteins revealed apparent molecular weights slightly lower than
those demonstrated by SDS-PAGE. This discrepancy suggests that
the proteins may have different shapes from those of the proteins
in the calibration standards. Nevertheless, these results indicate
that both proteins exist as monomers in solution (Fig. 2C).
Xyn8A and Rex8A exhibit different biochemical properties.
The two proteins were incubated with either wheat arabinoxylan
(WAX), oat spelts xylan (OSX), or xylo-oligosaccharides (XOS),
and their capacity to degrade the substrates was assessed by ana-
lyzing the hydrolysates by TLC. In the absence of enzyme, no
oligosaccharides were present in either the WAX or the OSX mix-
ture (Fig. 2D). However, upon the addition of Xyn8A, several
spots appeared, corresponding to shorter xylo-oligosaccharides
being released, an activity characteristic of endoxylanase enzymes
(Fig. 2D). The spots did not clearly comigrate with the standard
XOS, which is likely explained by the fact that these products are
xylo-oligosaccharides substituted with arabinosyl side chains. In
contrast, no detectable degradation of the two polysaccharides
was noted for Rex8A.
Both enzymes degraded XOS (Fig. 2E and Fig. 2F); however,
Xyn8A exhibited no detectable activity toward X
2
and X
3
. Upon
incubation with xylotetraose, a mixture of X
3
,X
2
, and X
1
was
observed, and a mixture of X
3
and X
2
was seen following incuba-
tion with xylopentaose and xylohexaose. In contrast, Rex8A com-
pletely converted X
3
through X
6
to a mixture of X
2
and X
1
. Most
notably, the two enzymes displayed differences in the product
distribution profiles for X
5
, with Xyn8A cleaving X
5
into X
2
and
X
3
, whereas Rex8A converted X
5
to X
1
and X
2
. This result suggests
that the two enzymes have different cleavage site preferences, with
Xyn8A cleaving the middle glycosidic linkage within X
5
, and
Rex8A cleaving one of the terminal glycosidic linkages in X
5
, pro-
ducing X
4
and X
1
and then converting X
4
to X
3
and X
1
and so on.
To further evaluate the hydrolytic activity of Rex8A with XOS,
the enzyme was incubated with X
3
through X
6
, and the concen-
trations of X
1
through X
6
were followed over time by HPLC. These
experiments confirmed results from the TLC plates, which
showed that the predominant products at the final time point of
the reaction were X
1
and X
2
for all substrates tested (Fig. 3). Rex8A
converted all of the X
3
to a mixture of X
1
and X
2
by 180 min of
incubation (Fig. 3A). Rex8A hydrolyzed all of the X
4
substrate
after 180 min of incubation by first accumulating X
1
and X
3
and
further hydrolyzing the accumulated X
3
to X
2
and X
1
(Fig. 3B).
With X
5
as a substrate, there was an initial accumulation of X
4
and
X
1
and a subsequent hydrolysis of X
4
to accumulate X
3
(Fig. 3C).
FIG 5 Mutational analysis reveals residues important for catalysis in Xyn8A (BACINT_04210) and Rex8A (BACINT_00927). The residues are Glu104, Asp164,
and Asp303 for Xyn8A. The residues are Glu90, Asp148, and Asp286 for Rex8A. (A) Three-dimensional homology modeling. Homology models were built for
Xyn8A and Rex8A using the ModWeb server (http://modbase.compbio.ucsf.edu/ModWeb20-html/modweb.html) with Pseudoalteromonas haloplanktis Xyn8A
(PDB accession no. 1H12) and Bacillus halodurans Rex (PDB accession no. 1WU4) as the templates, respectively. (B) Purification of mutants. Three mutants for
each Xyn8A and Rex8A were expressed as recombinant proteins, purified by cobalt affinity chromatography, and analyzed by SDS-PAGE. (C) Hydrolysis of
xylohexaose. Wild-type and mutant proteins were incubated with xylohexaose (X
6
) for 16 h, and the reaction products were analyzed by thin-layer
chromatography.
GH8 Enzymes from B. intestinalis DSM 17393
April 2014 Volume 80 Number 7 aem.asm.org 2089

Similarly, in the hydrolysis of X
6
,X
5
first accumulated during 20
min of incubation and was subsequently hydrolyzed to X
4
and X
3
and finally to X
1
and X
2
by the end of the 180-min incubation (Fig.
3D). At a relatively high concentration of Xyn8A (500 nM), X
4
was
converted first to X
3
and X
1
, and then the X
3
was eventually hy-
drolyzed to X
2
and X
1
(Fig. 4A). The enzyme exhibited much
higher activity with longer oligosaccharides; therefore, the
amount of enzyme added was decreased to observe the initial hy-
drolysis products (X
5
, 20 nM; X
6
, 2 nM). Under these conditions,
X
5
was converted to stoichiometric amounts of X
3
and X
2
, whereas
X
6
was cleaved, accumulating mainly X
3
(Fig. 4B and C).
Taken together, the activities seen for Xyn8A with polysaccha-
rides and XOS indicate that this enzyme is an endoxylanase,
whereas the activity for Rex8A was clearly very different from that
of Xyn8A and is consistent with the activity seen for reducing-end
xylose-releasing exo-oligoxylanases, which have been demon-
strated for GH8 proteins (27,28).
Site-specific mutagenesis to elucidate catalytic residues. To
predict amino acid residues important for catalysis, three-dimen-
sional homology models were constructed for Rex8A and Xyn8A
with the reducing-end xylose-releasing exo-oligoxylanase (Rex)
from Bacillus halodurans (PDB accession number 1WU4) and the
cold-adapted endoxylanase from Pseudoalteromonas haloplanktis
Xyn8A (PDB accession number 1H12), respectively. These struc-
tures revealed that the three catalytic residues proposed previously
for REX (27,36) (Glu90, Asp148, Asp286) were conserved in both
Rex8A and Xyn8A (Fig. 5A). Therefore, to evaluate whether these
residues (Glu104, Asp164, and Asp303 for Xyn8A and Glu90,
Asp148, and Asp286 for Rex8A) were also important for Xyn8A
and Rex8A, these three residues were changed to alanine by site-
directed mutagenesis.
The three mutants for each protein were expressed in E. coli
and purified by cobalt IMAC as described for the wild-type pro-
tein. All six proteins were expressed in the soluble fraction, indi-
cating that the mutations did not cause the proteins to form in-
soluble aggregates. Similar to the wild-type protein, the mutants
were purified after a single chromatography step (Fig. 5B). The
proportions of secondary structural elements between the WT
and the mutants were determined and compared by circular-di-
chroism (CD) spectroscopy to ensure that the change in enzyme
activity was not caused by gross structural differences. Compari-
son of the ␣-helix, ␤-sheet, ␤-turn, and unordered regions using
the Student ttest revealed no significant differences between the
wild-type and mutant forms of Rex8A and Xyn8A (see Table S1 in
the supplemental material). These data demonstrate that muta-
tion of these active-site residues did not appreciably alter the sec-
ondary structures of the proteins.
Following 16 h of incubation at 37°C with X
6
, only the D164A
mutant of Xyn8A exhibited minor residual activity compared to
the wild type, with a trace amount of X
3
visualized on the TLC
plate (Fig. 5C). For the three Rex8A mutants, no activity was de-
tected (Fig. 5C). These results therefore show that these three car-
boxylate-containing residues are important for catalysis in both
Rex8A and Xyn8A.
Identification of Xyl3A as a GH3 ␤-xylosidase. Rex8A was
observed to hydrolyze X
3
through X
6
to a mixture of xylose and
xylobiose (Fig. 2F). Given that rex8A is located immediately
downstream of a predicted GH3 glycosidase gene, it is possible
that the xylobiose produced by Rex8A would be converted to xy-
lose by this GH3 enzyme. Because GH3 glycosidases have a very
broad substrate range, it was decided to clone this GH3 glycosi-
dase gene and study its biochemical properties. The protein was
produced recombinantly in E. coli as described for Xyn8A and
Rex8A and purified to near homogeneity by cobalt IMAC (Fig.
6A). Biochemical activity assays with a library of para-nitrophenyl
(pNP)-linked substrates revealed that the enzyme had highest ac-
tivity with pNP-␤-D-xylopyranoside as a substrate (Fig. 6B). These
results suggest that the enzyme is a ␤-xylosidase. To further con-
firm this with natural substrates, the enzyme was incubated with
XOS and the capacity to hydrolyze these substrates was assessed by
TLC. Following incubation of the purified protein with XOS rang-
ing in length from X
2
through X
6
, the sole product visualized was
xylose (Fig. 6C). These results indicate that BACINT_00926 en-
codes a GH3 ␤-xylosidase, and the associated enzyme was there-
fore named Xyl3A.
DISCUSSION
A recent analysis of human gut microbiome reference genomes
revealed that of all gut bacterial strains, Bacteroides intestinalis
DSM 17393 exhibited the highest representation of individual gly-
specific activity (mIU/mg)
pNPAra
pNPGal
pNPXyl
pNPGlu
A. B.
C.
MW
(kDa)
66
45
31
200
116
97
X2
Xyl3A -+
X3
-+
X4
-+
X5
-+
X6
-+
X1
X2
X3
X4
X5
X6
X1
-+
100
200
300
400
500
0
FIG 6 BACINT_00926 encodes a GH3 ␤-xylosidase, Xyl3A. (A) Expression
and purification of Xyl3A. Xyl3A was expressed as a recombinant protein,
purified by cobalt affinity chromatography, and analyzed by SDS-PAGE. (B)
Activity with artificial substrates. Purified Xyl3A was screened with a library of
pNP-linked glycans using a continuous spectrophotometric assay, and the
specific activities for a subset of substrates are shown. Abbreviations are as
follows: pNPAra, pNP-␣-L-arabinofuranoside; pNPGal, pNP-␤-D-galactopy-
ranoside; pNPXyl, pNP-␤-D-xylopyranoside; pNPGlu, pNP-␤-D-glucopyran-
oside. (C) Hydrolysis of XOS. Xyl3A was incubated with XOS ranging in length
from xylobiose (X
2
) to xylohexaose (X
6
) for 16 h, and products of hydrolysis
were analyzed by thin-layer chromatography. Specific activity (mIU/mg) is
displayed in units of nmol substrate consumed per minute per mg of protein.
Hong et al.
2090 aem.asm.org Applied and Environmental Microbiology

coside hydrolase (GH) and polysaccharide lyase (PL) families and
the second highest total number of GH and PL genes (18). The
number of genes in the B. intestinalis genome targeted toward
degradation of the plant cell structural polysaccharide xylan is
particularly high. Coupled with previous studies of this organism
(17,37), the current data demonstrate that B. intestinalis DSM
17393 encodes at least nine endoxylanase enzymes deriving from
three different GH families (i.e., GH10, GH5, and GH8).
Expansion of GH families is a characteristic of many gut bac-
teria, most notably those from the Bacteroidetes phylum (18,38,
39). The current study highlights this process by revealing that B.
intestinalis DSM 17393 harbors two copies of genes encoding xy-
lan-specific GH8 enzymes with completely distinct biochemical
properties. Xyn8A is a GH8 endoxylanase enzyme that targets lon-
ger xylan fragments, hydrolyzing them to shorter xylo-oligosac-
charides that can subsequently be degraded by side-chain-cleav-
ing enzymes and ␤-xylosidases. Rex8A, on the other hand, is likely
a reducing-end xylose-releasing exo-oligoxylanase that releases
xylose from the reducing end of xylo-oligosaccharides, liberating
fermentable monosaccharides. The discovery of these two en-
zymes broadens our understanding of xylan degradation by gut
bacteria and provides insight into the highly dynamic genomic
assemblages that gut bacteria possess to capture energy from di-
etary polysaccharides.
Rex8A forms the core of a group of GH8 enzymes that derive
largely from bacteria isolated from human gastrointestinal tract
and rumen sources (Fig. 7A). Despite sharing high amino acid
sequence similarity, the polypeptide domain architectures vary
remarkably among these different organisms. Another human gut
bacterium, Bacteroides eggerthii, possesses a carbohydrate esterase
(CE) family 15 domain at the N terminus, while Prevotella copri,
Prevotella bergensis, and Bacteroides uniformis possess N-terminal
GH43/CBM6 modules (Fig. 7B). These proteins are likely to be
bifunctional enzymes with two distinct active sites. CE family 15
proteins have been demonstrated to cleave the methyl ester link-
age in 4-O-methyl-glucuronyl methyl esters (40,41), an activity
that is important for fermenting the glucuronic acid component
of heteroxylans. GH43/CBM6 proteins are commonly found to
have either ␤-xylosidase or arabinofuranosidase activities, both of
which are associated with the degradation of xylan fragments.
Therefore, it is likely that these alternative forms of GH8 Rex en-
zymes have evolved to possess additional enzymatic activities that
are important for xylan degradation.
The rex8A gene is located immediately downstream of the
xyl3A gene, which encodes a GH3 ␤-xylosidase. This unique pair-
ing of two genes encoding enzymes with activity against xylo-
oligosaccharides is deserving of further discussion. Rex enzymes
possess an active site with a minimum binding requirement for
three xylose sugars that is closed off at the terminal reducing end.
Catalysis then occurs between the sugars occupying the ⫺1 and
⫹1 subsites, liberating xylose and the remaining oligosaccharide
chain (36). Importantly, the minimum requirement of binding 3
sugars for catalysis means that the shortest XOS that this enzyme
can cleave is xylotriose, and therefore xylobiose is not cleaved by
these enzymes. On the other hand, GH3 glycosidases have a short
coin-shaped active site that contains two subsites (42); therefore,
enzymes from this family efficiently cleave xylobiose (34). These
two enzymes have complementary activities that may be impor-
0
length (amino acids)
200 400 600 800
GH 8
BiRex8A
GH 8
Bcel|WP_007211634.1
GH 8
Bole|WP_009130232.1
GH 8
Buni|WP_005834406.1
GH 43 CBM6
GH 8
Begg|WP_004289607.1
CE 15
GH 8
Pcop|WP_006849124.1
GH 43 CBM6
GH 8
Pber|WP_007175202.1
GH 43 CBM6
Bcel|WP_007211634.1
90
93 BiRex8A
Bole|WP_009130232.1
Begg|WP_004289607.1
Buni|WP_005834406.1
Pber|WP_007175202.1
Pcop|WP_006849124.1
Dmos|WP_006843639.1
Prum|YP_003574583.1
Dgad|WP_006799191.1
Mpal|WP_008505011.1
Slin|YP_003386258.1
Fjoh|YP_001196198.1
Bhal|Rex
Bsp|WP_010677526.1
Bcel|WP_007212772.1
BiXyn8A
Pgol|WP_007656390.1
Fsuc|YP_005820971.1
Chut|YP_677852.1
Ccel|YP_002505633.1
Acel|WP_010245038.1
Athe|YP_004175483.1
Phal|Xyn8
Cthe|Cel8A
Ccel|Cel8C
0.1
69
100
100
90
100
87 64
48
100
100
92
100
100
92
94
55
98
74
76
100
GIT REX
enzymes
GIT Xyn
enzymes
A. B.
signal peptide lipoprotein signal
FIG 7 Expansion and diversity of Rex8A homologs in the gut microbiome. (A) Phylogenetic analysis of Xyn8A (BACINT_04210) and Rex8A (BACINT_00927).
The amino acid sequences for GH8 proteins annotated by the CAZy database were aligned with Xyn8A and Rex8A using ClustalW, and a neighbor-joining tree
was constructed using CLC Genomics Workbench v5.0 software. Each alignment was resampled 100 times, and the bootstrap values are indicated on the internal
branches. The branch length is reported as the expected number of substitutions per amino acid position. GIT, gastrointestinal tract. (B) Protein domain
structure of Rex8A homologs. Domain architectures were predicted using dbCAN (48). Signal peptides and lipoprotein signal sequences were predicted using
SignalP v4.1 (30) and LipoP v1.0 (43), respectively. Organism abbreviations: Bcel, Bacteroides cellulosilyticus DSM 14838; Bi, Bacteroides intestinalis; Bole,
Bacteroides oleiciplenus YIT 12058; Begg, Bacteroides eggerthii DSM 20697; Buni, Bacteroides uniformis CL03T12C37; Pber, Prevotella bergensis DSM 17361; Pcop,
Prevotella copri DSM 18205; Dmos, Dysgonomonas mossii DSM 22836; Prum, Prevotella ruminicola 23; Dgad, Dysgonomonas gadei ATCC BAA-286; Mpal,
Mucilaginibacter paludis DSM 18603; Slin, Spirosoma linguale DSM 74; Fjoh, Flavobacterium johnsoniae UW101; Bhal, Bacillus halodurans C-125; Bsp, Bacillus sp.
10403023; Pgol, Parabacteroides goldsteinii CL02T12C30; Fsuc, Fibrobacter succinogenes subsp. succinogenes S85; Chut, Cytophaga hutchinsonii ATCC 33406; Ccel,
Clostridium cellulolyticum H10; Acel, Acetivibrio cellulolyticus; Athe, Anaerolinea thermophila UNI-1; Phal, Pseudoalteromonas haloplanktis; Cthe, Clostridium
thermocellum ATCC 27405.
GH8 Enzymes from B. intestinalis DSM 17393
April 2014 Volume 80 Number 7 aem.asm.org 2091

tant for the mechanism of xylan degradation employed by this
organism.
The three proteins described in this study each possess an N-
terminal signal peptide with a signal peptidase cleavage II site,
suggesting that they are transferred across the inner membrane
and anchored to a lipid moiety (43). The so-called “⫹2” rule im-
plies that prolipoproteins containing an aspartic acid at the ⫹2
position relative to the signal peptidase II cleavage site results in
retention of the polypeptide on the inner membrane within the
cytoplasm, whereas a serine residue directs the protein to the outer
membrane (44,45). Xyn8A, Rex8A, and Xyl3A all possess serine
residues at the ⫹2 position, which suggests that these proteins are
all localized to the outer membrane.
Since GH8 Rex enzymes represent a recently discovered group
of enzymes, relatively few studies have evaluated the catalytic res-
idues for these enzymes. In the current study, we identified amino
acid residues (E90, D148, and D286) that were absolutely con-
served among the GH8 Rex and Xyn enzymes and made the cor-
responding mutations in Xyn8A and Rex8A. These mutations had
very large effects on the activities of the two enzymes, despite CD
spectroscopy demonstrating no significant change in secondary
structure (see Table S1 in the supplemental material). These data
confirm results from previous studies that show residues corre-
sponding to E90, D148, and D286 as having roles as catalytic acid,
pKa modulator, and catalytic base, respectively (46).
In summary, this study reports distinct activities for two GH8
enzymes that are present in B. intestinalis DSM 17393, a bacterium
that is endemic to the human gut. Our findings reiterate the pre-
vious observation that this enzyme family contains members with
considerable differences in their substrate specificity (47). Fur-
thermore, our results suggest a xylan degradation pathway active
in gut Bacteroidetes that involves endoxylanases, a reducing-end
oligo-xylanase, and a ␤-xylosidase. These results could be of im-
portance in understanding the pathways of xylan degradation
present in other gut microorganisms harboring GH8 enzymes.
ACKNOWLEDGMENT
This project was partially supported by Agriculture and Food Research
Initiative Competitive Grant no. 2012-67015-19451 from the USDA Na-
tional Institute of Food and Agriculture.
REFERENCES
1. Bergman E. 1990. Energy contributions of volatile fatty acids from the
gastrointestinal tract in various species. Physiol. Rev. 70:567–590.
2. Hague A, Singh B, Paraskeva C. 1997. Butyrate acts as a survival factor for
colonic epithelial cells: further fuel for the in vivo versus in vitro debate.
Gastroenterology 112:1036–1040. http://dx.doi.org/10.1053/gast.1997
.v112.agast971036.
3. Roediger WE. 1980. The colonic epithelium in ulcerative colitis: an ener-
gy-deficiency disease? Lancet ii:712–715.
4. Scheppach W, Bartram HP, Richter F. 1995. Role of short-chain fatty
acids in the prevention of colorectal cancer. Eur. J. Cancer 31A:1077–
1080.
5. Dodd D, Cann IK. 2009. Enzymatic deconstruction of xylan for biofuel
production. Glob. Change Biol. Bioenergy 1:2–17. http://dx.doi.org/10
.1111/j.1757-1707.2009.01004.x.
6. Davison A, Blaxter M. 2005. Ancient origin of glycosyl hydrolase family
9 cellulase genes. Mol. Biol. Evol. 22:1273–1284. http://dx.doi.org/10.1093
/molbev/msi107.
7. Morris SC. 2003. Life’s solution: inevitable humans in a lonely universe.
Cambridge University Press, Cambridge, United Kingdom.
8. Hespell RB, Whitehead TR. 1990. Physiology and genetics of xylan deg-
radation by gastrointestinal tract bacteria. J. Dairy Sci. 73:3013–3022.
http://dx.doi.org/10.3168/jds.S0022-0302(90)78988-6.
9. Chassard C, Goumy V, Leclerc M, Del’homme C, Bernalier-Donadille
A. 2007. Characterization of the xylan-degrading microbial community
from human faeces. FEMS Microbiol. Ecol. 61:121–131. http://dx.doi.org
/10.1111/j.1574-6941.2007.00314.x.
10. Chassard C, Delmas E, Lawson PA, Bernalier-Donadille A. 2008. Bac-
teroides xylanisolvens sp. nov., a xylan-degrading bacterium isolated from
human faeces. Int. J. Syst. Evol. Microbiol. 58:1008–1013. http://dx.doi
.org/10.1099/ijs.0.65504-0.
11. Salyers AA, Vercellotti JR, West SE, Wilkins TD. 1977. Fermentation of
mucin and plant polysaccharides by strains of Bacteroides from the human
colon. Appl. Environ. Microbiol. 33:319–322.
12. Robert C, Chassard C, Lawson PA, Bernalier-Donadille A. 2007. Bac-
teroides cellulosilyticus sp. nov., a cellulolytic bacterium from the human
gut microbial community. Int. J. Syst. Evol. Microbiol. 57:1516–1520.
http://dx.doi.org/10.1099/ijs.0.64998-0.
13. Crittenden R, Karppinen S, Ojanen S, Tenkanen M, Fagerstrom R,
Matto J, Saarela M, Mattila-Sandholm T, Poutanen K. 2002. In vitro
fermentation of cereal dietary fibre carbohydrates by probiotic and intes-
tinal bacteria. J. Sci. Food Agric. 82:781–789. http://dx.doi.org/10.1002
/jsfa.1095.
14. Weaver J, Whitehead TR, Cotta MA, Valentine PC, Salyers AA. 1992.
Genetic analysis of a locus on the Bacteroides ovatus chromosome which
contains xylan utilization genes. Appl. Environ. Microbiol. 58:2764–2770.
15. Whitehead TR. 1995. Nucleotide sequences of xylan-inducible xylanase
and xylosidase/arabinosidase genes from Bacteroides ovatus V975.
Biochim. Biophys. Acta 1244:239–241. http://dx.doi.org/10.1016/0304
-4165(95)00051-C.
16. Mirande C, Mosoni P, Bera-Maillet C, Bernalier-Donadille A, Forano E.
2010. Characterization of Xyn10A, a highly active xylanase from the hu-
man gut bacterium Bacteroides xylanisolvens XB1A. Appl. Microbiol. Bio-
technol. 87:2097–2105. http://dx.doi.org/10.1007/s00253-010-2694-0.
17. Dodd D, Moon YH, Swaminathan K, Mackie RI, Cann IK. 2010.
Transcriptomic analyses of xylan degradation by Prevotella bryantii and
insights into energy acquisition by xylanolytic Bacteroidetes. J. Biol.
Chem. 285:30261–30273. http://dx.doi.org/10.1074/jbc.M110.141788.
18. Kaoutari AE, Armougom F, Gordon JI, Raoult D, Henrissat B. 2013.
The abundance and variety of carbohydrate-active enzymes in the human
gut microbiota. Nat. Rev. Microbiol. 11:497–504. http://dx.doi.org/10
.1038/nrmicro3050.
19. Martens EC, Koropatkin NM, Smith TJ, Gordon JI. 2009. Complex
glycan catabolism by the human gut microbiota: the Bacteroidetes Sus-
like paradigm. J. Biol. Chem. 284:24673–24677. http://dx.doi.org/10.1074
/jbc.R109.022848.
20. Martens EC, Lowe EC, Chiang H, Pudlo NA, Wu M, McNulty NP,
Abbott DW, Henrissat B, Gilbert HJ, Bolam DN, Gordon JI. 2011.
Recognition and degradation of plant cell wall polysaccharides by two
human gut symbionts. PLoS Biol. 9:e1001221. http://dx.doi.org/10.1371
/journal.pbio.1001221.
21. Collins T, Gerday C, Feller G. 2005. Xylanases, xylanase families and
extremophilic xylanases. FEMS Microbiol. Rev. 29:3–23. http://dx.doi.org
/10.1016/j.femsre.2004.06.005.
22. Cantarel BL, Coutinho PM, Rancurel C, Bernard T, Lombard V, Hen-
rissat B. 2009. The Carbohydrate-Active EnZymes database (CAZy): an
expert resource for glycogenomics. Nucleic Acids Res. 37:D233–D238.
http://dx.doi.org/10.1093/nar/gkn663.
23. Yoon K-H, Yun HN, Jung KH. 1998. Molecular cloning of a Bacillus sp.
KK-1 xylanase gene and characterization of the gene product. Biochem.
Mol. Biol. Int. 45:337–347.
24. Elleuche S, Piascheck H, Antranikian G. 2011. Fusion of the OsmC
domain from esterase EstO confers thermolability to the cold-active xyla-
nase Xyn8 from Pseudoalteromonas arctica. Extremophiles 15:311–317.
http://dx.doi.org/10.1007/s00792-011-0361-8.
25. Brennan Y, Callen WN, Christoffersen L, Dupree P, Goubet F, Healey
S, Hernandez M, Keller M, Li K, Palackal N, Sittenfeld A, Tamayo G,
Wells S, Hazlewood GP, Mathur EJ, Short JM, Robertson DE, Steer BA.
2004. Unusual microbial xylanases from insect guts. Appl. Environ. Mi-
crobiol. 70:3609–3617. http://dx.doi.org/10.1128/AEM.70.6.3609-3617
.2004.
26. Lee CC, Kibblewhite-Accinelli RE, Wagschal K, Robertson GH, Wong
DW. 2006. Cloning and characterization of a cold-active xylanase enzyme
from an environmental DNA library. Extremophiles 10:295–300. http:
//dx.doi.org/10.1007/s00792-005-0499-3.
27. Honda Y, Kitaoka M. 2004. A family 8 glycoside hydrolase from Bacillus
Hong et al.
2092 aem.asm.org Applied and Environmental Microbiology

halodurans C-125 (BH2105) is a reducing end xylose-releasing exo-
oligoxylanase. J. Biol. Chem. 279:55097–55103. http://dx.doi.org/10.1074
/jbc.M409832200.
28. Lagaert S, Van Campenhout S, Pollet A, Bourgois TM, Delcour JA,
Courtin CM, Volckaert G. 2007. Recombinant expression and character-
ization of a reducing-end xylose-releasing exo-oligoxylanase from Bifido-
bacterium adolescentis. Appl. Environ. Microbiol. 73:5374 –5377. http://dx
.doi.org/10.1128/AEM.00722-07.
29. Bakir MA, Kitahara M, Sakamoto M, Matsumoto M, Benno Y. 2006.
Bacteroides intestinalis sp. nov., isolated from human faeces. Int. J. Syst.
Evol. Microbiol. 56:151–154. http://dx.doi.org/10.1099/ijs.0.63914-0.
30. Petersen TN, Brunak S, von Heijne G, Nielsen H. 2011. SignalP 4.0:
discriminating signal peptides from transmembrane regions. Nat. Meth-
ods 8:785–786. http://dx.doi.org/10.1038/nmeth.1701.
31. Laemmli UK. 1970. Cleavage of structural proteins during the assembly of
the head of bacteriophage T4. Nature 227:680– 685. http://dx.doi.org/10
.1038/227680a0.
32. Whitmore L, Wallace BA. 2004. DICHROWEB, an online server for
protein secondary structure analyses from circular dichroism spectro-
scopic data. Nucleic Acids Res. 32(Suppl 2):W668–W673. http://dx.doi
.org/10.1093/nar/gkh371.
33. Powell JC, Lever M. 1972. A new automated procedure for the colori-
metric determination of glucose. Biochem. Med. 6:543–547. http://dx.doi
.org/10.1016/0006-2944(72)90008-7.
34. Dodd D, Kiyonari S, Mackie RI, Cann IK. 2010. Functional diversity of
four glycoside hydrolase family 3 enzymes from the rumen bacterium
Prevotella bryantii B14. J. Bacteriol. 192:2335–2345. http://dx.doi.org/10
.1128/JB.01654-09.
35. Miyazaki K, Miyamoto H, Mercer DK, Hirase T, Martin JC, Kojima Y,
Flint HJ. 2003. Involvement of the multidomain regulatory protein XynR
in positive control of xylanase gene expression in the ruminal anaerobe
Prevotella bryantii B
1
4. J. Bacteriol. 185:2219–2226. http://dx.doi.org/10
.1128/JB.185.7.2219-2226.2003.
36. Fushinobu S, Hidaka M, Honda Y, Wakagi T, Shoun H, Kitaoka M.
2005. Structural basis for the specificity of the reducing end xylose-
releasing exo-oligoxylanase from Bacillus halodurans C-125. J. Biol.
Chem. 280:17180–17186. http://dx.doi.org/10.1074/jbc.M413693200.
37. Dodd D, Mackie RI, Cann IK. 2011. Xylan degradation, a metabolic
property shared by rumen and human colonic Bacteroidetes. Mol. Micro-
biol. 79:292–304. http://dx.doi.org/10.1111/j.1365-2958.2010.07473.x.
38. Flint HJ, Bayer EA, Rincon MT, Lamed R, White BA. 2008. Polysac-
charide utilization by gut bacteria: potential for new insights from
genomic analysis. Nat. Rev. Microbiol. 6:121–131. http://dx.doi.org/10
.1038/nrmicro1817.
39. Koropatkin NM, Cameron EA, Martens EC. 2012. How glycan metab-
olism shapes the human gut microbiota. Nat. Rev. Microbiol. 10:323–335.
http://dx.doi.org/10.1038/nrmicro2746.
40. Topakas E, Moukouli M, Dimarogona M, Vafiadi C, Christakopoulos
P. 2010. Functional expression of a thermophilic glucuronyl esterase from
Sporotrichum thermophile: identification of the nucleophilic serine. Appl.
Microbiol. Biotechnol. 87:1765–1772. http://dx.doi.org/10.1007/s00253
-010-2655-7.
41. Spanikova S, Biely P. 2006. Glucuronoyl esterase—novel carbohydrate
esterase produced by Schizophyllum commune. FEBS Lett. 580:4597–4601.
http://dx.doi.org/10.1016/j.febslet.2006.07.033.
42. Varghese JN, Hrmova M, Fincher GB. 1999. Three-dimensional struc-
ture of a barley beta-D-glucan exohydrolase, a family 3 glycosyl hydrolase.
Structure 7:179–190. http://dx.doi.org/10.1016/S0969-2126(99)80024-0.
43. Juncker AS, Willenbrock H, Von Heijne G, Brunak S, Nielsen H, Krogh
A. 2003. Prediction of lipoprotein signal peptides in Gram-negative bac-
teria. Protein Sci. 12:1652–1662. http://dx.doi.org/10.1110/ps.0303703.
44. Seydel A, Gounon P, Pugsley AP. 1999. Testing the ‘⫹2 rule’ for lipo-
protein sorting in the Escherichia coli cell envelope with a new genetic
selection. Mol. Microbiol. 34:810– 821. http://dx.doi.org/10.1046/j.1365
-2958.1999.01647.x.
45. Yamaguchi K, Yu F, Inouye M. 1988. A single amino acid determinant of
the membrane localization of lipoproteins in E. coli. Cell 53:423– 432. http:
//dx.doi.org/10.1016/0092-8674(88)90162-6.
46. Collins T, De Vos D, Hoyoux A, Savvides SN, Gerday C, Van Beeumen
J, Feller G. 2005. Study of the active site residues of a glycoside hydrolase
family 8 xylanase. J. Mol. Biol. 354:425–435. http://dx.doi.org/10.1016/j
.jmb.2005.09.064.
47. Pollet A, Schoepe J, Dornez E, Strelkov SV, Delcour JA, Courtin CM.
2010. Functional analysis of glycoside hydrolase family 8 xylanases shows
narrow but distinct substrate specificities and biotechnological potential.
Appl. Microbiol. Biotechnol. 87:2125–2135. http://dx.doi.org/10.1007
/s00253-010-2659-3.
48. Yin Y, Mao X, Yang J, Chen X, Mao F, Xu Y. 2012. dbCAN: a web
resource for automated carbohydrate-active enzyme annotation. Nucleic
Acids Res. 40:W445–W451. http://dx.doi.org/10.1093/nar/gks479.
GH8 Enzymes from B. intestinalis DSM 17393
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