A complex gene locus enables xyloglucan utilization in the model saprophyte Cellvibrio japonicus

Abstract

The degradation of plant biomass by saprophytes is an ecologically important part of the global carbon cycle, which has also inspired a vast diversity of industrial enzyme applications. The xyloglucans (XyGs) constitute a family of ubiquitous and abundant plant cell wall polysaccharides, yet the enzymology of XyG saccharification is poorly studied. Here, we present the identification and molecular characterization of a complex genetic locus that is required for xyloglucan utilization by the model saprophyte Cellvibrio japonicus. In harness, transcriptomics, reverse genetics, enzyme kinetics, and structural biology indicate that the encoded cohort of an α-xylosidase, a β-galactosidase, and an α-l-fucosidase is specifically adapted for efficient, concerted saccharification of dicot (fucogalacto)xyloglucan oligosaccharides following import into the periplasm via an associated TonB dependent receptor. The data support a biological model of xyloglucan degradation by C. japonicus with striking similarities to - and notable differences with - the complex polysaccharide utilization loci of the Bacteroidetes.

Full text

A complex gene locus enables xyloglucan utilization in the
model saprophyte
Cellvibrio japonicus
Johan Larsbrink,1Andrew J. Thompson,2
Magnus Lundqvist,1Jeffrey G. Gardner,3
Gideon J. Davies2and Harry Brumer1,4*
1Division of Glycoscience, School of Biotechnology,
Royal Institute of Technology (KTH), AlbaNova
University Centre,106 91 Stockholm, Sweden.
2Department of Chemistry,University of York,
Heslington, York YO10 5DD, UK.
3Department of Biological Sciences,University of
Maryland – Baltimore County,1000 Hilltop Circle,
Baltimore, MD 21250, USA.
4Michael Smith Laboratories and Department of
Chemistry,University of British Columbia,2185 East
Mall, Vancouver, BC, V6T 1Z4, Canada.
Summary
The degradation of plant biomass by saprophytes is
an ecologically important part of the global carbon
cycle, which has also inspired a vast diversity of
industrial enzyme applications. The xyloglucans
(XyGs) constitute a family of ubiquitous and abundant
plant cell wall polysaccharides, yet the enzymology of
XyG saccharification is poorly studied. Here, we
present the identification and molecular characteriza-
tion of a complex genetic locus that is required for
xyloglucan utilization by the model saprophyte Cell-
vibrio japonicus. In harness, transcriptomics, reverse
genetics, enzyme kinetics, and structural biology
indicate that the encoded cohort of an α-xylosidase, a
β-galactosidase, and an α-L-fucosidase is specifically
adapted for efficient, concerted saccharification of
dicot (fucogalacto)xyloglucan oligosaccharides fol-
lowing import into the periplasm via an associated
TonB-dependent receptor. The data support a biologi-
cal model of xyloglucan degradation by C. japonicus
with striking similarities – and notable differences – to
the complex polysaccharide utilization loci of the
Bacteroidetes.
Introduction
The saccharification of diverse types of plant biomass is
both an ecologically important part of the global carbon
cycle and a biotechnologically relevant aspect of the food,
feed, biofuel, biomaterials, and cleaning-product indus-
tries. Hydrolysis of the complex plant cell wall to acquire
sugars for growth presents a significant challenge for
microorganisms, and requires a wide range of enzyme
activities to address the tremendous diversity of glycosidic,
peptide, polyphenolic, and ester linkages present (Carpita
and McCann, 2000; Jovanovic et al., 2009). Whereas the
complete hydrolysis of cellulose microfibres is difficult due
to their semi-crystalline nature, the monosaccharide and
linkage complexity of the hemicelluloses and pectins addi-
tionally confounds enzymatic attack. Moreover, the inter-
twining of plant cell wall biomolecules into a composite
material provides an additional level of structural complex-
ity mitigating biomass saccharification, and there is
growing evidence that addition of hemicellulases to indus-
trial enzyme cocktails can improve efficacy (Hu et al.,
2013; Jabbour et al., 2013). As such, carbohydrate-active
enzyme (CAZyme) discovery remains a vibrant area of
research (Jovanovic et al., 2009; Mewis et al., 2013;
Hemsworth et al., 2014; del Pulgar and Saadeddin, 2014).
Cellvibrio japonicus (previously Pseudomonas fluores-
cens subsp. cellulosa) is a Gram-negative bacterium that
was first isolated from Japanese soil in the 1950s and has
since become intensely studied due to its ability to degrade
all common plant cell wall polysaccharides (Hazlewood
and Gilbert, 1998; Deboy et al., 2008). As a model sapro-
phytic organism, the complete genome sequence of C.
japonicus has been determined and genetic tools have
been established (Deboy et al., 2008; Gardner and
Keating, 2012). C. japonicus is thus important both for
CAZyme discovery and for fundamental molecular studies
of polysaccharide degradation by Gram-negative environ-
mental bacteria. Furthermore, recent metabolic engineer-
ing of a strain with the ability to produce ethanol has
demonstrated the potential of C. japonicus for industrial
chemical production in consolidated bioprocessing
(Gardner and Keating, 2010). Whereas the mannan-,
xylan-, and arabinan-degrading systems have been inten-
sively studied (Cartmell et al., 2008; 2011; Deboy et al.,
2008; Emami et al., 2009 and references therein], the
Accepted 25 August, 2014. *For correspondence. E-mail brumer@
msl.ubc.ca; Tel. (+1) 6048273738; Fax (+1) 6048222114.
Molecular Microbiology (2014) ■doi:10.1111/mmi.12776
© 2014 The Authors. Molecular Microbiology published by John Wiley & Sons Ltd.
This is an open access article under the terms of the Creative Commons Attribution License, which permits use, distribution and
reproduction in any medium, provided the original work is properly cited.

capacity of C. japonicus to degrade the ubiquitous hemi-
cellulose family of xyloglucans has received little attention.
The xyloglucans (XyG) are a structurally complex family
of polysaccharides found in all terrestrial plants (Carpita
and McCann, 2000; Popper et al., 2011). In dicots alone,
XyGs may account for 20–25% of primary cell wall dry
weight (Scheller and Ulvskov, 2010) and, as such, consti-
tute an important terrestrial carbon sink. XyGs are com-
posed of a cellulose-like linear β(1→4)-glucan backbone
appended with branching α(1→6)-xylosyl moieties,
which can in turn be extended with additional glycosides
(e.g. galacto-, fuco- and arabinosyl moieties) in a tissue-
and species-dependent manner (Peña et al., 2008;
Tuomivaara et al., 2014). Thus, the complete saccharifica-
tion of XyGs necessitates a cohort of enzymes to address
the linkage diversity present in individual XyG variants.
Aligned with our continuing interest in xyloglucan-active
enzymes (Gilbert et al., 2008; Ariza et al., 2011; Mark
et al., 2011; Eklöf et al., 2012; 2013; Larsbrink et al., 2014),
we previously reported the biochemical and structural
characterization of the main α-xylosidase of C. japonicus,
CjXyl31A, which specifically hydrolyses terminal non-
reducing-end xylose moieties of xyloglucan oligosaccha-
rides (XyGOs) (Larsbrink et al., 2011). In the same study,
we also identified that C. japonicus produces at least one
extracellular endo-xyloglucanase capable of generating
XyGOs by xyloglucan backbone cleavage. However, a key
remaining question is, which are the other players in C.
japonicus that might work in concert with these enzymes to
fully deconstruct complex xyloglucans?
Here, we report the identification and molecular charac-
terization of a unique xyloglucan utilization locus (XyGUL)
in the genome of C. japonicus (Fig. 1). In addition to the
α-xylosidase CjXyl31A, this locus contains a predicted
β-galactosidase, a predicted α-L-fucosidase, and a
predicted TonB-dependent receptor (Ferguson and
Deisenhofer, 2002; Koebnik, 2005), whose collective pres-
ence indicates a role in dicot (fucogalacto)xyloglucan deg-
radation and uptake. Using a combination of transcriptional
analysis, reverse genetics, subcellular protein localization,
recombinant enzyme kinetics, and structural biology, we
were able to verify these predictions of glycoside function
and extend a biological model of XyG utilization by this
important saprophyte.
Results
Genome walking reveals a putative xyloglucan
utilization locus (XyGUL) in C. japonicus
Genome walking in silico revealed the presence of open
reading frames encoding a predicted β-galactosidase from
glycoside hydrolase (GH) family 35 (CjBgl35A, encoded
by locus tag CJA_2707, bgl35A), a predicted TonB-
dependent receptor (TBDR, encoded by CJA_2709), and a
predicted α-L-fucosidase from glycoside hydrolase family
95 (CjAfc95A, encoded by CJA_2710, afc95A) upstream
from the gene encoding the known α-xylosidase CjXyl31A
(CJA_2706, xyl31A) (Fig. 1). Protein and gene names
used here correspond to those proposed in Table S2 of
Deboy et al. (2008). The predicted protein sequences of
both CjBgl35A and CjAfc95A were compared to previously
characterized enzymes in GH35 and GH95, respectively,
using BLAST. Interestingly, CjBgl35Ashowed a high homol-
ogy only to the Xanthomonas campestris pv. campestris
GalD β-galactosidase, with a protein identity of 55% and
similarity of 70% (over 536 residues), while similarity to
other characterized GH35 enzymes catalogued in the
CAZy database (Lombard et al., 2014; http://www.cazy
.org/GH35_characterized.html) was poor. Moreover, the
β-galactosidase activity of GalD has been indicated primar-
ily through genetic studies and minimal specificity data
exists for this enzyme (Yang et al., 2007), such that no
reliable prediction of substrate specificity for CjBgl35A
can be made. CjAfc95A, in contrast, had high similarity
to all five characterized α(1→2)-L-fucosidases of
GH95 (Lombard et al., 2014; http://www.cazy.org/GH95
_characterized.html), with identities of 31–35% and simi-
larities of 47–52%. In particular, the two plant enzymes
(from Arabidopsis thaliana and Lilium longiflorum) and the
fungal enzyme (from Aspergillus nidulans) have been
shown to cleave the α(1→2)-fucosyl residues from XyGOs
xyl31A tbdrbgl35A afc95A
bp 3175359 bp 3185628
CJA_2706 CJA_2707
CJA_2708
CJA_2709 CJA_2710
XGLF
(Galpβ1)0-1
2
66
2
6
2
(Fucpα1)0-1
[ ]
4Glcpβ1 4Glcpβ1 4Glcpβ1 4Glcpβ1
Xylpα1 Xylpα1 Xylpα1
(Galpβ1)0-1
n
A
B
Fig. 1. Xyloglucan structure and the arrangement of xyloglucan-
degrading genes within the C. japonicus genome.
A. Typical (fucogalacto)xyloglucan structure, with abbreviated motif
nomenclature according to Tuomivaara et al. (2014) indicated.
B. The xyloglucan utilization locus (XyGUL) in the genome of C.
japonicus; hypothetical proteins are shown in grey.
2
J. Larsbrink
et al
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© 2014 The Authors. Molecular Microbiology published by John Wiley & Sons Ltd., Molecular Microbiology

(Bauer et al., 2006; Ishimizu et al., 2007; Nagae et al.,
2007; Leonard et al., 2008). On the basis of these bioinfor-
matics analyses, we hypothesized that the GHs were
involved in the cleavage of β(1→2)-galactosyl and α(1→2)-
L-fucosyl residues of dicot (fucogalacto)xyloglucan,
respectively, while theTBDR may function in carbohydrate
import (Dejean et al., 2013).
CjBgl35A and CjAfc95A are XyG-specific glycosidases
To test predictions of the catalytic specificity of the GHs,
we performed initial substrate screens on aryl glycosides
(Glcp/Galp/Xylp-β-PNP, L-Araf-α-PNP and L-Fuc-α-CNP)
with both enzymes produced recombinantly in Escheri-
chia coli. To further investigate the potential roles of the
enzymes in vivo, xyloglucan polysaccharides (XyG) and
oligosaccharides (XyGOs) from tamarind and iceberg
lettuce were examined as natural substrates, including
product analysis by HPAEC-PAD and MALDI-TOF. Full
kinetic data for both enzymes are listed in Table 1.
CjBgl35A. CjBgl35A was able to hydrolyse Gal-β-PNP
but did not act upon any of the other substrates in the
initial screen. The pH optimum of the enzyme was deter-
mined to be pH 5.5 in acetate buffer (Supplemental
Fig. S1), but the pH-rate profile deviated from the bell-
shaped profile expected for an enzyme mechanism
involving two ionizing residues. The kcat/Kmvalue for the
reaction on Gal-β-PNP was 12.5 s−1mM−1, which is in the
same range as another bacterial GH35 β-galactosidase,
BgaC from Streptococcus pneumonia (49.3 s−1mM−1) and
the A. thaliana BGAL4 (19.1 s−1mM−1) (Ahn et al., 2007;
Cheng et al., 2012). Hydrolysis of the substrate lactose,
which is the preferred substrate for many enzymes in
GH35, could not be observed; CjBgl35A is thus likely not
able to generally address β(1→4)-linkages.
In contrast, HPAEC-PAD indicated that the enzyme
released galactose from both tamarind seed (galacto)xy-
loglucan and the doubly galactosylated XLLG nonasac-
charide (see Fig. 1 for XyGO nomenclature). Enzyme
saturation could not be reached for either substrate (Sup-
plemental Fig. S2). Notably, a nasturtium (Tropaeolum
majus)β-galactosidase also exhibited a strictly linear
dependence of rate on substrate concentration using
either nasturtium or tamarind xyloglucan as substrates
(Edwards et al., 1988) (it is likely, but not conclusive, that
this is a GH35 enzyme: GenBank CAW88932). Apparent
kcat/Kmvalues were nonetheless calculated by linear
regression of the initial velocities versus substrate concen-
tration plot. Notably, the apparent kcat/Kmvalue for the
degalactosylation of tamarind xyloglucan is c. 1000-fold
lower than that for XLLG (Table 1). This difference, esti-
mated from the total galactose content of the polysaccha-
ride, is likely to represent a lower limit, given the possibility
of contaminating galactosylated oligosaccharides (acting
as alternate substrates) below our limits of detection and
the clear preference of CjBgl35A for one of the two galac-
tosyl residues (vide infra). Regardless, the data indicate
that the galactosidase is most likely to act subsequent to
the hydrolysis of xyloglucan polysaccharide chains into
component oligosaccharides.
Product analysis by MALDI-TOF MS showed that
CjBgl35A was able to hydrolyse both galactosyl moieties
from XLLG to form XXXG (Fig. 2A). A time-course study
using HPAEC-PAD revealed an order of preference for the
galactosyl residues on the two L (Gal–Xyl–Glc) moieties:
XXLG was produced essentially exclusively during the
initial stages of the reaction, with XXXG appearing only
when a majority of the XLLG had been converted into
XXLG. Given this preference, the initial-rate the kinetic
data (Table 1, Supplemental Fig. S2) likely reflect the
hydrolysis of the galactosyl moiety most distal from the
reducing-end (underlined: XLLG). To assess whether the
enzyme could cleave galactosyl moieties underlying a
terminal fucosyl moiety, a 1:2 mixture of the oligosaccha-
rides XXFG and XLFG (Fig. 1A) was prepared from fuco-
sylated XyG extracted from iceberg lettuce by digestion
with a Bacteroides ovatus GH5 endo-xyloglucanase
(Larsbrink et al., 2014) and size-exclusion chromatogra-
phy. Extended incubation of a high concentration of
CjBgl35A with this mixture (10 μM enzyme, c. 0.5 mM
substrate, 16 h, 25°C) followed by MALDI-TOF MS
demonstrated that only XLFG was hydrolysed, yielding
XXFG, which was not further transformed (Fig. 2C). The
Table 1. Activity of CjBgl35A and CjAfc95A on various substrates.
Enzyme Substrate kcat (s−1)Km(mM) kcat/Km(s−1mM−1)
CjBgl35A Gal-β-PNP 10.1 ±0.2 0.81 ±0.05 12.5
XLLG 3.8 ±0.08
Tamarind xyloglucana0.0031 ±(6.5 ×10−5)
CjAfc95A L-Fuc-α-CNP 26.5 ±1.1 2.1 ±0.2 12.9
XLFG 19.2 ±0.5 0.21 ±0.017 89.8
Lettuce xyloglucanb1.85 ±0.084 0.26 ±0.03 7.1
a. Kinetic parameters calculated from the available galactosyl moieties of the substrate (16%) as reported by the manufacturer.
b. Kinetic parameters calculated from the available fucosyl moieties of the substrate, as determined by end-point hydrolysis by CjAfc95A.
Cellvibrio japonicus
xyloglucan utilization locus
3
© 2014 The Authors. Molecular Microbiology published by John Wiley & Sons Ltd., Molecular Microbiology

Intensity (a.u.)
m/z
XLLG
1408
XLXG/XXLG
1246 1701
1114 1570
A
Intensity (a.u.)
m/z
XXXG
1084
B
0
100
200
300
10.0 10.5
PAD (nC)
t (min)
XXXG
XLLGXXLG
XLXG
C
Intensity (a.u.)
m/z
XLFG
1555
XXFG
1393
XXFG
1393
Intensity (a.u.)
m/z
*
*
Fig. 2. Analysis of products from CjBgl35A activity on galactosylated xyloglucan oligosaccharides.
A. MALDI-TOF spectra of a preparation of XLLG containing a minor amount of XXLG/XLXG before and after incubation with CjBgl35A,
indicating complete hydrolysis of both galactosyl residues. Observed m/z values are consistent with [M +Na]+adducts. Minor peaks at m/z
1570 and 1701 correspond to unidentified Hex7Pen3and Hex7Pen4oligosaccharides (Tuomivaara et al., 2014).
B. HPAEC-PAD chromatograms of a time-course study of the conversion of XLLG into XXXG via XXLG as a primary product. Arrows indicate
trends in peak intensity changes for the starting material and major products. XLLG (bold line) was incubated with CjBgl35A and samples
were analysed after 5 min (thin line), 20 min (dotted line) and 2 h (dashed line). Formation of XLXG is not observed, while XXXG appears only
after the majority of the XLLG has been converted into XXLG, revealing that the galactosyl moiety closest to the non-reducing end (cf. Fig. 1)
is the preferred substrate for the enzyme. Asterisks indicate minor components, which may correspond to unidentified oligosaccharides
observed by MALDI-TOF MS.
C. MALDI-TOF spectra of the hydrolysis of a preparation containing XLFG and XXFG by CjBgl35A. XLFG is completely converted into XXFG
while the ‘F’ unit is unchanged, demonstrating that fucosylation blocks the strictly exo-acting galactosidase. Observed m/z values are
consistent with [M+Na]+adducts.
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J. Larsbrink
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data indicate that CjBgl35A acts strictly as an exo-
galactosidase that requires terminal galactosyl moieties on
XyGOs.
At present, no other bacterial enzymes from GH35 have
been assayed for XyG specificity, to our knowledge, so
direct comparisons with CjBgl35A are difficult to make.
However, many of these are specific for lactose, a sub-
strate not hydrolysed by CjBgl35A, which suggests dis-
tinct activity profiles. Rather, the specificity of CjBgl35A is
similar to plant β-galactosidases with a general ability to
remove side-chain galactosyl units from xyloglucan poly-
saccharides; the GH35 member from A. thaliana, Bgal10
(At5g63810) produces XXLG and XXXG sequentially due
to a preference for the underlined side-chain in XLLG
(Sampedro et al., 2012 and references therein).
CjAfc95A. CjAfc95A did not cleave any of the four PNP
glycosides tested (vide supra), but readily hydrolysed
L-Fuc-α-CNP. Using this substrate, the pH-rate profile was
classically bell-shaped with a pH optimum of pH 6.5 in
citrate buffer (Supplemental Fig. S3). The observed kcat/Km
value for CjAfc95A (12.9 mM−1s−1; Table 1 and Fig. S4)
compares favourably with that of a Bifidobacterium longum
subsp. infantis GH95 member (0.12 mM−1s−1) (Sela et al.,
2012). On natural substrates, product analysis by HPAEC-
PAD and MALDI-TOF (Fig. 3) indicated that XLFG and
XXFG were converted to XLLG and XXLG respectively.
This specificity is similar to characterized GH95 α-L-
fucosidases from the plants A. thaliana and L. longiflorum,
both of which are active on fucosylated XyGOs (Ishimizu
et al., 2007; Leonard et al., 2008).
The release of fucose from both the polysaccharide and
the mixture of fucosylated XyGOs was also quantified by
HPAEC-PAD (Table 1). Although the kinetics are con-
founded by the two-component XyGO mixture, the relative
apparent kcat/Kmvalue for the oligosaccharides was seven-
fold higher than that for L-Fuc-α-CNP, due to a 10-fold
lower apparent Kmvalue. This suggests that carbohydrate-
binding interactions in the positive enzyme subsites signifi-
cantly enhance catalysis (for subsite nomenclature see
Davies et al., 1997). Indeed, the intrinsic leaving-group
ability of 2-chloro-4-nitrophenol [pKa5.45 (Ibatullin et al.,
2008)] is potentially 10 orders of magnitude greater than a
sugar hydroxyl group [pKac. 16 (Damude et al., 1996)].
Similar strong effects of extended XyGO binding in positive
enzyme subsites are exhibited by the α-xylosidase in the
locus (Larsbrink et al., 2011; Silipo et al., 2012).
The tertiary structure of CjBgl35A reveals the molecular
basis for substrate recognition
To complement our structure–function analysis of
CjXyl31A (Larsbrink et al., 2011; Silipo et al., 2012), we
wished to complete the tertiary structural characterization
A
B
Intensity (a.u.)
m/z
XLFG
1552
XXFG
1390
1406
1568
Intensity (a.u.)
m/z
XLLG
1407
1423
XXLG
1245
PAD (nC)
t (min)
150
100
50
0
9.5 10.0 10.5
XLFG
XXLG
XLLG
XXFG
Fig. 3. Analysis of products from CjAfc95A activity on fucosylated
xyloglucan oligosaccharides.
A. MALDI-TOF spectra of the hydrolysis of a preparation containing
XLFG and XXFG into XLLG and XXLG, respectively, after
incubation with CjAfc95A.
B. HPAEC-PAD chromatograms of the same conversion. Observed
m/z values are consistent with [M+Na]+adducts; minor amounts of
[M+K]+adducts are also observed.
Cellvibrio japonicus
xyloglucan utilization locus
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© 2014 The Authors. Molecular Microbiology published by John Wiley & Sons Ltd., Molecular Microbiology

of the remaining GHs encoded by the putative C. japonicus
XyGUL to reveal the determinants of xyloglucan recogni-
tion. Whereas CjBgl35A has proven amenable to crystal-
lography, CjAfc95A has thus far resisted our attempts,
despite extensive effort.
The three-dimensional structure of CjBgl35A reveals a
relatively unusual two-domain architecture comprising an
N-terminal catalytic (β/α)8(TIM) barrel domain (residues
37–419), appended to a smaller, C-terminal, mixed α/β
structure (residues 420–575) consisting of two short α
helices and nine βstrands (Fig. 4 and Supplemental
Table S2; PDB ID 4D1I). Given its poor sequence similarity
to other GH35 enzymes (vide supra), this two-domain fold
exhibits low homology to other currently known protein
structures; only one significant structural match to another
bacterial GH35 β-galactosidase, from Caulobacter cres-
centus strain CB15, was found. Analysis using the Dali
server (Holm and Rosenström, 2010) shows Bgl35A and
the C. crescentus enzyme (PDB accession code 3U7V) to
have a Z-score of 61.5, an r.m.s.d. value of 1.2 Å mapped
across 508 Cαpositions, and a sequence identity of 56%.
The next most-similar protein (GH5 endo-β-mannanase,
PDB code 1QNO) has a Z-score of 25.1, an r.m.s.d. value
of 3.0 Å across 289 Cαpositions, and a sequence identity
of just 12%.
The (β/α)8structure of the CjBgl35A N-terminal catalytic
domain reveals a centrally positioned cleft running later-
ally across the open end of the barrel. This topology
appears to be consistent with the requirement for binding
of extended, branched oligo/polysaccharide substrates
such as XyGOs and/or XyG and thus the cleft was antici-
pated to contain the catalytic active site. Subsequent
soaking of native Bgl35A crystals with the iminosugar
1-deoxygalactonojirimycin [DGJ; 1,5-Dideoxy-1,5-imino-
D-galactitol (Paulsen et al., 1980)] confirmed both the
location of the active centre and, importantly, unveiled
crucial ligand-protein interactions including the identity of
the likely catalytic amino acids (Fig. 4B and C, and Sup-
plemental Table S2; PDB ID 4D1J). The Kdfor this inter-
action in solution was determined to be 485 ±42 nM by
ITC (Fig. S5).
In the complex structure, DGJ is located within a deep
cavity, approximately at the centre of the proposed active-
site cleft. DGJ is co-ordinated in this −1 subsite by a
complex hydrogen-bonding network mediated by the side-
chain groups of residues N67 (interacting with O3), K134
(O3 and O4), N135 (O4), N204 (O2), and N383 (O6)
(Fig. 4C). The side-chains of E205 and E349 occupy clas-
sical positions expected for the catalytic pair, comprising a
proton donor-acceptor and nucleophile consistent with the
retaining mechanism expected for all GH35 members
(Fig. 4B). Furthermore, the position of the catalytic
proton donor-acceptor residue is consistent with an anti-
protonation trajectory (Heightman and Vasella, 1999).
A pocket positioned directly adjacent to the pseudoano-
meric position of DGJ is proposed to be the +1 catalytic
subsite that is responsible for binding side-chain xylosyl
moieties present on XyG/XyGOs. Curiously, the apparent
solvent-exposed nature of the non-reducing end of the
ligand suggests the possibility to accommodate extended
substrates, since the axial 4-OH position of the six-
membered ring points directly out into the large cleft
(Fig. 4D). However, in fucosylated XyGOs such as XXFG
and XLFG, the pendant fucosyl residue is α(1→2)-linked to
the galactosyl residue (Fig. 1). Steric considerations
(Fig. 4D) therefore explain the requirement for CjAfc95A to
first remove the terminal Fuc from such XyGOs before the
underlying Gal can be addressed by CjBgl35A (Fig. 2).
Despite the observation of separate protein domains
(Fig. 4A), it seems likely that these together solely consti-
tute a catalytic module, i.e. a distinct biochemical/biological
function is not indicated. Structure/sequence homology
searching of the C-terminal region in isolation produced no
significant results that would indicate similarity to known
regulatory or carbohydrate-binding modules (CBMs),
despite the observation of a trough-like surface somewhat
reminiscent of a substrate recognition cleft (Gilbert et al.,
2013). In addition, the observation that the early part of the
C-terminal domain is formed from a distortion of the final
helix of the barrel domain, and that the initial β-strand of this
domain stacks directly against the penultimate barrel helix,
strongly suggests a rigid, monolithic structure.
Genetic analysis defines the biological importance of
the XyGUL
With a firm understanding of the determinants of substrate
recognition by the glycoside hydrolases CjXyl31A
(Larsbrink et al., 2011), CjBgl35A, and CjAfc95A, we
sought to confirm that the putative XyGUL indeed consti-
tuted a co-ordinately regulated locus. Further, we were
interested to determine whether this multi-gene locus was
exclusively responsible for XyG saccharification by C.
japonicus.
Transcriptional analysis
Quantitative PCR (qPCR) was performed on all four pre-
dicted XyGUL genes (Fig. 1; xyl31A, bgl35A, tbdr, afc95A)
to monitor possible upregulation during growth on minimal
medium containing tamarind XyGOs versus a glucose
control. Additional predicted β-galactosidases and α-L-
fucosidases found in the C. japonicus genome (Deboy
et al., 2008) (bgl2A, bgl2B, bgl2C and afc95B), but not in
the putative XyGUL, were also included to determine their
potential involvement in XyG deconstruction. Of these,
the predicted β-galactosidase gene, bgl2B, and the other
predicted α-L-fucosidase gene, afc95B, showed the same
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level of expression in both growth conditions, and were
therefore used as reference genes. The four XyGUL
genes were all clearly upregulated 12- to 45-fold when C.
japonicus was grown in M9-XyGO medium (Fig. 5A).
The predicted β-galactosidase genes bgl2A and bgl2C,
which are not in the putative XyGUL were also slightly
(<5-fold) upregulated in the M9-XyGO cultures (Fig. 5A).
However, genome analysis suggests that these genes are
Fig. 4. Crystallography of CjBgl35A.
A. The tertiary structure viewed along the barrel axis of the N-terminal catalytic domain. The mixed α/βmotif of the C-terminal domain is
indicated, while a transparent surface overlay illustrates the rigid nature of the domain interface.
B. 1-Deoxygalactonojirimycin (DGJ) bound in the active site (carbon atoms in grey); the proposed catalytic residues, E349 and E205, are also
illustrated (carbon atoms in green). The pseudoanomeric position of the ligand can be observed approximately 3.14 Å from the carboxylate
group of E349, the likely catalytic nucleophile. Electron density is depicted as a maximum-likelihood weighted 2Fo−Fcsynthesis contoured at
0.29 e Å−3(1.0 σ).
C. Additional residues in the −1 subsite interacting with GDJ.
D. Surface representation, coloured according to electrostatic potential, showing the position of DGJ within the −1 subsite. A further cavity
immediately adjacent to the pseudoanomeric position of DGJ is suggested to be the +1 subsite, accommodating the branched oligosaccharide
substrates within the active-site cleft.
Panel A was assembled using PyMOL version 1.3r1 (Schrödinger, LLC), while panels B–D were created using CCP4Mg (McNicholas et al.,
2011).
Cellvibrio japonicus
xyloglucan utilization locus
7
© 2014 The Authors. Molecular Microbiology published by John Wiley & Sons Ltd., Molecular Microbiology

unlikely to be purposely involved in XyG catabolism and
may instead be responding to a general sensing signal
due to galactose released by the XyGUL enzymes. For
example, bgl2A (CJA_0496) is located together with a
predicted endo-1,4-galactanase, gal53A-2 (CJA_0497)
and a TonB-dependent receptor (CJA_0498), and might
instead be primarily involved in (arabino)galactan
metabolism. bgl2C (CJA_2610) is co-located with a pre-
dicted chitinase, chi18D (CJA_2611), which may point
towards a principal function related to fungal cell wall or
insect carapace degradation.
To assay whether any of the locus genes were tran-
scribed as a polycistronic mRNA, PCR was performed in
an attempt to amplify the regions between each of the
genes in the cDNA, using the primers from the qPCR study.
The xyl31A and bgl35A genes were found to be
co-transcribed on one bicistronic mRNA strand (data not
shown). This observation is supported by the similar
expression levels of these two genes indicated by qPCR
(Fig. 5A) and the distance between the genes (41 bases),
which is unlikely to accommodate a promoter region.
Similar analyses of the bgl35A/tbdr and tbdr/afc95A pairs
suggested that the TonB-dependent receptor gene and
afc95A are transcribed individually. The non-coding
regions directly upstream of these genes are 88 and 199
bases in length respectively, and are therefore large
enough to contain individual promoters.
Reverse genetics of xyl31A and bgl35A
To assess physiological function, xyl31A and bgl35A gene-
disruption mutants were generated, confirmed by PCR
(Fig. S6), and assayed for growth on tamarind xyloglucan.
Inactivation of xyl31A mutation results in a severe growth
defect, with a growth rate of 0.06 h−1(peak OD600 =0.13)
on xyloglucan, compared to 0.41 h−1(peak OD600 =1.2) of
the wild-type strain (Fig. 5B and C). As expected from its
bicistronic relationship with xyl31A, disruption of bgl35A
likewise results in a severe growth defect (growth rate
0.07 h−1, peak OD600 =0.21). Both mutants grow in a wild-
type manner on the polysaccharides xylan (from beech-
wood), carboxymethyl cellulose, and pectin (from apple).
Subcellular localization of XyGUL function
We previously demonstrated that CjXyl31A is cell
membrane-bound, with attachment likely mediated
through N-terminal lipidation (Larsbrink et al., 2011). In
contrast, LipoP (Juncker et al., 2003) predicts that neither
CjBgl35A nor CjAfc95A are N-terminally lipidated. Further-
more, SignalP (Petersen et al., 2011) fails to identify a
signal peptide in CjAfc95A. To further refine our spatial
model of XyG utilization in C. japonicus, the subcellular
locations of CjBgl35A, and CjAfc95A were investigated by
Glucose
Xyloglucan
OD600 OD600
Time (hr)
1
0.5
0.1
510152025
WT
bgl35A
xyl31A
1
0.5
0.1
510
15 20 25
xyl31A
bgl35A
tbdr
afc95A
bgl2A
bgl2C
Normalized expression
Fold increase in M9-XGO vs. M9-Glc
50
40
30
20
10
0
A
B
C
Fig. 5. Transcriptomic and reverse genetic analysis of the C.
japonicus xyloglucan utilization locus (XyGUL).
A. Normalized increase in expression of the locus genes (cf. Fig. 1)
in cells grown on M9-XyGO medium versus to M9-Glc medium,
compared with other predicted galactosidase-encoding (bgl2A,
bgl2B, bgl2C) and fucosidase-encoding (afc95B) genes (Deboy
et al., 2008). bgl2B and afc95B had identical expression in both
growth conditions and therefore served as reference genes. Error
bars indicate the standard error of the mean.
B and C. Growth analysis of C. japonicus wild-type (WT) and
bgl35A and xyl31A knockout mutant strains in MOPS minimal
medium with either 0.25% glucose (B) or 0.25% xyloglucan (C) as
sole carbon sources. All growth experiments were performed in
biological triplicate; error bars represent the standard deviation of
the mean (in many cases the error bars are smaller than the data
point marker).
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specific chromogenic assays of cell fractions. Notably, no
significant α-L-fucosidase activity could be detected in any
of the protein fractions from the M9-Glc cultures (Table 2).
Likewise, β-galactosidase activity was absent in the
secreted protein fraction from M9-Glc cultures, while only
very weak activity was observed in the periplasmic and
cytoplasmic protein fractions. In contrast, growth on M9
medium containing XyGOs to induce XyGUL expression
resulted in a significant increase in both activities in all
three fractions. The specific activity levels were however
clearly highest in the periplasmic fractions versus the
secreted and cytoplasmic fractions (activity in these frac-
tions, which is at least eightfold lower in all cases, may
represent some degree of cross-contamination).
Discussion
Transcript and reverse genetics analyses allow us to con-
clude that xyl31A, bgl35A, tbdr, and afc95A (Fig. 1) con-
stitute a xyloglucan utilization locus (XyGUL), which is the
primary genetic determinant conferring C. japonicus with
the ability to saccharify this ubiquitous plant cell wall poly-
saccharide. Together with substrate specificity and struc-
tural analysis of the encoded glycoside hydrolases,
subcellular localization data allow us to propose an
updated model of XyG degradation in C. japonicus (Fig. 6).
In this model, XyG polysaccharide is hydrolysed into
component oligosaccharides by one or more endo-
xyloglucanase(s). The liberated XyGOs are then imported
into the periplasm via the TBDR (Ferguson and
Deisenhofer, 2002; Koebnik, 2005; Dejean et al., 2013),
where the exo-glycosidases CjXyl31A, CjBgl35A, and
CjAfc95A work in concert, together with a currently uniden-
tified β-glucosidase(s), to yield monosaccharides for
further catabolism.
Notably, structural enzymology provides some of the
strongest support for this model. The α-xylosidase,
CjXyl31A, has a strict non-reducing-end specificity due
to a deep, pocket-shaped active site and thus cannot
access xylosyl residues positioned along the intact poly-
saccharide chain (Larsbrink et al., 2011). Likewise, the
apparent kcat/Kmvalue of CjBgl35A for the oligosaccha-
ride XLLG is three orders of magnitude higher than for
tamarind XyG polysaccharide (Table 1), which, together
with the observed pocket topology of the active-site
(Fig. 4), suggests that cleavage of the polysaccharide to
XyGOs also occurs prior to the action of this enzyme. As
highlighted above, steric limitations of the CjBgl35A
active-site also indicate that CjAfc95A must remove ter-
minal fucosyl residues prior to fully enable degalacto-
sylation by CjBgl35A.
It is particularly interesting to note the obvious differ-
ences between the XyGUL system of C. japonicus
vis-à-vis the polysaccharide utilization loci (PULs) of Bac-
teroidetes (Koropatkin et al., 2012). A recently character-
ized XyGUL from B. ovatus comprises a complete cohort
of GHs sufficient to address each linkage in solanaceous
(arabinogalacto)xyloglucan, including both a keystone
endo-xyloglucanase and six diverse exo-glycosidases, in
addition to a TBDR, a hybrid two-component sensor/
regulator, and two cell-surface carbohydrate-binding pro-
teins (Larsbrink et al., 2014; Terrapon and Henrissat,
2014). In contrast, the C. japonicus XyGUL – in addition to
being alternatively directed towards dicot (fucogalacto)xy-
loglucan – is significantly less complete: It does not
encode the requisite endo-xyloglucanase(s) necessary to
initiate the saccharification pathway by polysaccharide
backbone cleavage, yet C. japonicus does secrete signifi-
cant endo-xyloglucanase activity into the medium under
XyGO-induction (Larsbrink et al., 2011). Likewise, the C.
japonicus XyGUL does not encode a β-glucosidase,
which would be required to digest the β(1→4)-glucan
backbone of the XyGOs. Finally, the XyGUL appears to
lack a substrate sensor/transcriptional regulator analo-
gous to that which regulates xylan-specific genes in C.
japonicus (Emami et al., 2009). Thus, although the C.
japonicus XyGUL shares a theme of genetic colocaliza-
tion and co-regulation with the Bacteroidetes PULs, it is
notably less evolved with respect to complexity and com-
pleteness. It is also notable that the TBDRs of the C.
japonicus and B. ovatus XyGULs do not share any sig-
nificant sequence similarity, despite predicted functional
homology based on their association with a similar com-
plement of GHs.
In general, the colocalization and co-ordinated regula-
tion of CAZyme-encoding genes in C. japonicus has not
Table 2. Specific galactosidase and fucosidase activities for proteins produced under different growth conditions.
Growth medium Substrate
Specific activity in cell fractions (μM released product·s−1·μg protein−1)
Secreted Periplasmic Intracellular
M9-XyGO Gal-β-PNP 1.25 ±0.14 18.3 ±3 1.0 ±0.3
L-Fuc-α-CNP 160 ±47 1400 ±160 28.8 ±6.6
M9-Glc Gal-β-PNP n/d 0.86 ±0.05 0.058 ±0.003
L-Fuc-α-CNP n/d n/d n/d
Cellvibrio japonicus
xyloglucan utilization locus
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© 2014 The Authors. Molecular Microbiology published by John Wiley & Sons Ltd., Molecular Microbiology

been widely studied. Indeed, annotation of the C. japonicus
genome revealed that many of these genes are not clus-
tered on the chromosome, although interesting exceptions
can be observed (Deboy et al., 2008). For example, early
work by Gilbert, Hazlewood and co-workers revealed
that endo-xylanase B of GH10 (CJA_3280) and α-L-
arabinofuranosidase C of GH62 (CJA_3281) are encoded
by adjacent genes in C. japonicus. Post-genomic re-
examination by McClendon et al. has revealed that these
are likely to constitute part of a Xylan Utilization Locus
that additionally encodes a predicted endo-xylanase of
GH30 (CJA_3279), feruloyl esterase D (Fee1B) of
CE1 (CJA_3282) (Ferreira et al., 1993), a predicted
β-galactosidase of GH98 (CJA_3286), and a second pre-
dicted feruloyl esterase (Fee1A) of CE1 (CJA_3287)
(McClendon et al., 2011). Likewise, the endo-xylanase
Xyn10D (CJA_2888) (Emami et al., 2009) is located
between the α-glucuronidase GlcA67A (CJA_2887) (Nagy
et al., 2003) and the acetylxylan esterase Axe2B
(CJA_2889) (Zhang et al., 2014). On the other hand, the
reason for colocalization of other genes is sometimes
less obvious: the broad-specificity xylan-active α-L-
arabinofuranosidase Abf51A (CJA_2769) (Beylot et al.,
2001) is found in tandem with the endo-mannanase
Man26A (CJA_2770) (Hogg et al., 2001), which perhaps
suggests a greater diversity of mannan substructures
than previously appreciated. Regardless, we posit that
increased consideration of CAZyme colocalization within
genomes can significantly facilitate understanding the
basic biology of C. japonicus and other bacteria, as well as
advance the discovery of novel enzyme cohorts for bio-
technological applications (Martens et al., 2014). For
example, a recent study of a mannan-degrading locus of
Bacteroides fragilis has identified a syntenic locus in
C. japonicus (CJA_0241–0246) putatively comprising a
sugar/cation symporter, a mannosyl-glucose phosphory-
TBDR
H+
H+
TonB-ExbBD-
complex
TBDR
Afc95A
Bgl35A
Xyl31A
β-glucosidase(s)
endo-xyloglucanase(s)
Fig. 6. Proposed pathway of (fucogalacto)xyloglucan degradation by C. japonicus. Sugar symbols are as follows: Glc – blue circles,
Xyl – orange stars, Gal – yellow circles, Fuc – red pentagons. Secreted enzymes with endo-xyloglucanase activity depolymerize the
polysaccharides into xyloglucan oligosaccharides which are imported into the periplasm by the TonB-dependent receptor of the locus. In the
periplasm, CjBgl35A and CjAfc95A strip off galactose and fucose from the oligosaccharides respectively. In concert, CjXyl31A and an
unknown β-glucosidase cleave off terminal xylose and glucose residues from the non-reducing end in an iterative manner.
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lase, a mannobiose 2-epimerase, a GH5 endo-
mannanase and a GH27 α-galactosidase (Deboy et al.,
2008; Senoura et al., 2011).
The data presented here shed new light on the genetic,
biochemical, and structural basis of hemicellulose utiliza-
tion by C. japonicus and, moreover, reveal a novel,
matched set of glycosidases for biotechnological applica-
tions. Indeed, the importance of xyloglucan saccharifica-
tion has been arguably under-appreciated in the context
of biofuel production (Gilbert et al., 2008), although recent
studies are beginning to address this issue (Hu et al.,
2013; Jabbour et al., 2013), buoyed by an increased
understanding of specific, xyloglucan-active enzymes
(Gilbert et al., 2008; Ariza et al., 2011; Larsbrink et al.,
2011; Eklöf et al., 2012). Interestingly, the xyloglucan uti-
lization locus described here encodes neither the endo-
xyloglucanase(s) nor the β-glucosidase(s) required for full
degradation of XyG. The proteins necessary for sensing
XyG in the environment are also presently unknown.
Future studies from our collaboration will focus on the
identification of the corresponding genes elsewhere in the
genome, in addition to functional studies on the TonB-
dependent receptor of the locus.
Experimental procedures
Ultrapure water, purified on a Milli-Q system (Millipore) to a
resistivity of ρ>18.2 MΩcm, was used in all experiments.
Galactose, lactose, Gal-β-PNP, Glc-β-PNP, Xyl-β-PNP and
L-Araf-α-PNP were purchased from Sigma. L-Fuc-α-CNP and
fucose were purchased from Carbosynth. Tamarind xyloglu-
can (XyG) was purchased from Megazyme. XLLG was pre-
pared from tamarind XyG as described previously (Greffe
et al., 2005). Lettuce XyG and XLFG were isolated as
described below. C. japonicus Ueda107 was obtained from
the National Collections of Industrial, Marine, and Food Bac-
teria (Aberdeen, Scotland).
Cloning of CjBgl35A and CjAfc95A
The open reading frames encoding CjBgl35A and CjAfc95A
(GenBank Accession No. ACE85180.1 and ACE83895.1
respectively) were amplified by PCR from genomic DNA of C.
japonicus Ueda107 using Phusion polymerase (Finnzymes),
using forward primers incorporating a CACC overhang to
enable TOPO cloning, and reverse primers excluding stop
codons (Eurofins MWG Operon; Supplemental Table S1). The
forward primer for CjBgl35A was designed to exclude the
predicted native signal peptide (residues 1–36), while no
signal peptide was predicted for CjAfc95A (Petersen et al.,
2011). The PCR products were cloned into the pENTR/SD/D-
TOPO entry vectors (Invitrogen) and recombined into pET-
DEST42 destination vectors (Invitrogen) following the
manufacturer’s instructions.
Initial protein crystals produced from the pET-DEST42 con-
struct, featuring a larger C-terminal tag, yielded only relatively
weak diffraction to approximately 4.0 Å (data not shown) and
proved difficult to optimize further. As such, primers were
designed to allow recloning of the gene fragment encoding
CjBgl35A into a pET28a vector modified for Ligation Inde-
pendent Cloning (LIC), and featuring a shorter, N-terminal
hexahistidine tag (Eurofins MWG Operon; Supplemental
Table S1). The PCR product was cloned into a pre-prepared
linear vector stock using the InFusion-HD cloning kit (Clon-
tech) according to the manufacturer’s instructions.
Recombinant gene expression and protein purification
pET-DEST42 constructs were transformed into E. coli
BL21(DE3) by electroporation, and proteins were produced
and purified by immobilized metal affinity chromatography
(IMAC) following an established protocol (Larsbrink et al.,
2011). The modified pET28a-Bgl35A construct was trans-
formed into E. coli TUNER(DE3) cells via the heat-shock
method, with recombinant protein purified by IMAC and size-
exclusion chromatography. Protein purity was verified by
SDS-PAGE. Protein concentration was determined from A280
values; the extinction coefficients used for CjBgl35A and
CjAfc95A were 118260 and 151845 M−1cm−1respectively
[ProtParam server (Gasteiger et al., 2005)].
Crystallization, data collection and structure solution
of CjBgl35A
Crystals of CjBgl35A suitable for full X-ray data collection
were grown using hanging drop vapour diffusion at 19°C,
with equal volumes of pure protein and reservoir solution
(2.6 M sodium acetate pH 7.2). Ligand complex formation
with GDJ was achieved by soaking native crystals in 10 mM
GDJ (final) for a period of approximately 1 h. Since concen-
trated sodium acetate present within the mother-liquor solu-
tion proved an adequate cryo-protectant, no additional
solvents were added to crystals prior to flash cooling in
liquid N2. Full diffraction data for both native and ligand
complex CjBgl35A crystals were collected at beamline I03
of the Diamond Light Source (Didcot, Oxfordshire, UK).
Measured reflection intensities were indexed, integrated
and scaled using XDS (Kabsch, 2010a,b) and the CCP4
suite (Winn et al., 2011) implementation of Aimless. The
structure of CjBgl35A was solved by molecular replacement
with PHASER (McCoy et al., 2007), employing the
co-ordinates of the C. crescentus GH35 (PDB code 3U7V)
as a phasing model. An initial atomic model was con-
structed using the CCP4 (Winn et al., 2011) implementation
of Buccaneer and refined via the maximum-likelihood
method using numerous cycles of REFMAC (Murshudov
et al., 2011; Winn et al., 2011), with additional manual cor-
rection using COOT (Emsley et al., 2010). The structure of
CjBgl35A in complex with GDJ was produced by refining the
scaled data (processed as above) against the finalized
model of the native protein and visually inspecting calcu-
lated electron density maps for evidence of ligand binding.
The final atomic model of the complex was subsequently
refined and corrected as above. Models and structure
factors for both native CjBgl35A and the GDJ complex have
been deposited into the PDB with respective accession
codes 4D1I and 4D1J.
Cellvibrio japonicus
xyloglucan utilization locus
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© 2014 The Authors. Molecular Microbiology published by John Wiley & Sons Ltd., Molecular Microbiology

High-performance anion-exchange chromatography with
pulsed amperometric detection (HPAEC-PAD)
Oligo- and monosaccharides were analysed on a Dionex
ICS-3000 HPLC system operated by Chromelion software
version 7 (Dionex) using a Dionex CarboPac PA200 column
for Gradient Aand a Dionex CarboPac PA1 column for Gradi-
ent B. Solvent Awas water, solvent B was 1 M sodium hydrox-
ide and solvent C was 1 M sodium acetate. The gradients used
were as follows: Gradient A: 0 to 5 min, 10% B, 2% C; 5 to
12 min, 10% B and a linear gradient from 2% to 30% C; 12 to
12.1 min, 50% B, 50% C; 12.1 to 13 min, an exponential
gradient of B and C back to initial conditions; 13 to 17 min,
initial conditions. Gradient B: column pre-conditioned prior to
injection by −13 to −3 min, 12% B, 6.8% C; −3 to 0 min, 100%
A; 0 to 25 min, 100% A.
Matrix-assisted laser desorption/ionization-time-of-flight
(MALDI-TOF) analysis of oligosaccharides
MALDI-TOF was performed on oligosaccharide samples
using a Voyager-DE STR instrument (Applied Biosystems) in
positive linear mode with an acceleration of 20 kV and an
extraction delay time of 150 ns. The samples were prepared
by mixing equal amount of sample (1 μl) with the matrix
(10 mg ml−12,5 dihydroxybenzoic acid in 1:1 acetonitrile :
H2O, containing 0.05% trifluoroacetic acid).
Xyloglucan extraction from iceberg lettuce
Using a modified and scaled-up version of a published proto-
col (Hsieh and Harris, 2009), XyG polysaccharide was
extracted from approximately 500 g of fresh iceberg lettuce
leaves, obtained from a local grocery store. The plant material
was homogenized in 70% aqueous ethanol using a high-
speed blender, collected by filtration using Miracloth (Millipore)
and ground to a fine powder in liquid nitrogen using a ceramic
mortar and pestle. Alcohol-soluble polysaccharides and
smaller sugars were removed by repeated 70% ethanol wash
and filtration steps. Non-cellulosic polysaccharides were
extracted in 6 M sodium hydroxide, containing 1% sodium
borohydride to prevent alkaline peeling, followed by neutrali-
zation with acetic acid.
Ethanol was added to the neutralized solution to a final
concentration of 70% to precipitate polysaccharides, followed
by centrifugation at 24 000 gfor 15 min. The supernatant was
discarded and the pellet washed three times with 70% ethanol.
The washed pellet was dissolved in water and loaded onto a Q
Sepharose column (GE Lifesciences), pre-equilibrated with
10 mM imidazole (pH 7.0) to bind charged polysaccharides
(e.g. pectins) to the matrix. Neutral hemicellulosic polysaccha-
rides were eluted in three column volumes of the same buffer.
The resulting hemicellulose-containing fraction was incubated
with 150 units of the xylanase CjCBM22-GH10 (Xyn10A) and
150 units of mannanase 26A (both purchased from NZYtech)
for 16 h at 37°C, to hydrolyse contaminating xylan and
mannan respectively. The resulting oligosaccharides were
subsequently removed in the supernatant following ethanol
precipitation of the lettuce XyG (final yield 180 mg after lyo-
philization from water). Prior to analysis by HPAEC-PAD
(gradient B), 1 mg of lettuce XyG was hydrolysed by incuba-
tion with 2 M trifluoroacetic acid (TFA) for 3 h at 120°C (1 ml
total volume). Hydrolysis products were vacuum-dried and
re-suspended in deionized water, followed by filtration
(0.2 μm). The lettuce xyloglucan comprised Glc, Xyl, Gal and
Fuc, with pectin derived contaminations of approximately 5%.
In a second extraction, fucosylated xylogluco-
oligosaccharides (XyGOs) were obtained via a similar proce-
dure lacking the ion exchange step. Seven hundred and fifty
milligrams of lyophilized crude lettuce XyG was dissolved in
100 ml 50 mM ammonium acetate, pH 5.5, to which was
added 1.28 mg B. ovatus BoGH5A endo-xyloglucanase
(Larsbrink et al., 2014). The reaction was incubated overnight
at 35°C followed by lyophilization (dry weight 0.3 g). XyGOs
were dissolved in 1 ml water and loaded onto a 100 cm XK16
column packed with 200 ml Bio-Gel P-2 Gel (Bio-Rad), which
had been equilibrated with water. Eluted fractions were ana-
lysed by MALDI-TOF and fractions containing predominantly
XXXG, XXFG and XLFG, respectively, were pooled and
lyophilized.
Enzyme assays
All assays were carried out at 25°C, at the pH optimum of the
respective enzyme. Curve fitting and processing of kinetic
data were performed using Origin 8 software (OriginLab).
pH dependence
Measurements of the pH-dependence of CjBgl35A and
CjAfc95A were performed with Gal-β-PNP and L-Fuc-α-CNP
as substrates, respectively, using the assays described
below. Buffers (50 mM) are indicated in Figs S1 and S3.
Chromogenic assays
Activities on PNP glycosides were analysed by a stopped
assay as previously described (Larsbrink et al., 2011), using
enzyme concentrations in the μM range for initial screens and
several hours incubation. For initial rate kinetics on Gal-β-PNP
for CjBgl35A, 42 nM enzyme was used and reactions were
stopped after 10 min. Assays utilizing L-Fuc-α-CNP were
monitored continuously for the release of 2-chloro-4-
nitrophenolate using a Cary 300 spectrophotometer (Agilent
Technologies). For the determination of the kinetic parameters
of CjAfc95A, 22 nM enzyme was used. An extinction coeffi-
cient of 12 936 M−1cm−1, determined from a standard curve,
was used to calculate product concentration from A405 values.
HPAEC-PAD-based assays
Enzymatic reactions on XyG polysaccharides and XyGOs
were performed in 50 μl reactions, containing 50 mM buffer at
the pH optimum of the assayed enzyme, and were stopped
by addition of 2 μl of 5 M sodium hydroxide prior to HPAEC-
PAD analysis. For the reaction of CjBgl35A on XLLG, 2.7 nM
enzyme was used, the reaction was terminated after 10 min.
For the reaction of CjBgl35A on tamarind XyG, 0.16 μM
enzyme was used, the reaction was terminated after 130 min.
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For the reaction of CjAfc95A on XLFG, 0.64 nM enzyme was
used, and the reaction was terminated after 15 min. For the
reaction of CjAfc95A on lettuce XyG, 270 nM enzyme was
used, and the reaction was terminated after 20 min. To quan-
tify the release of galactose or fucose from the reactions, the
respective commercial monosaccharides were used as
standards.
Isothermal titration calorimetry (ITC)
Isothermal titration calorimetry (ITC) was performed with a
MicroCal Auto-iTC200 system (GE Healthcare). Assays were
conducted in triplicate at 25°C, with GDJ (390 μM) titrated
into the ITC cell containing pure CjBgl35A (38.5 μM). Disso-
ciation constants (Kd) for each titration were subsequently
calculated and averaged using the Origin 7 software package
(MicroCal).
Transcription analysis
Total RNA was extracted from cultures of C. japonicus grown
in liquid M9 minimal media supplemented with either a
mixture of tamarind XyGOs (4 g l−1) or glucose (10 g l−1). Five
millilitres overnight cultures were used to inoculate 200 ml
cultures of the same media, and the OD600 of the cultures
were monitored. When the OD600 had reached 1, correspond-
ing to a late exponential growth phase (Gardner and Keating,
2010), 2 ml culture was added to 4 ml RNAprotect Bacteria
Reagent (Qiagen) and the cells were harvested by centrifu-
gation at 4300 gfor 5 min. Total RNAwas extracted using the
RNeasy mini kit (Qiagen), following the manufacturer’s
instructions, including in-solution DNase treatment and sub-
sequent clean-up. The final yield of total RNA was on average
42 μg for XyGO cultures and 21 μg for Glc cultures. One
microgram of each biological sample was reverse transcribed
into cDNA by the iScript cDNASynthesis Kit (Bio-Rad). qPCR
was performed on a CFX96 Touch instrument using Hard-
Shell white well plates (Bio-Rad). Primers for the locus genes
(Eurofins MWG Operon; Table S1), as well as the predicted
β-galactosidase encoding genes CjBgl2A, CjBgl2B, CjBgl2C
and the predicted α-L-fucosidase encoding gene CjAfc95B
were used to analyse the respective genes for each growth
condition in biological duplicates and technical triplicates. The
protocol used for amplification consisted of: 95°C 3 min, 40
cycles of 95°C, 15 s; 58°C, 15 s; 72°C, 15 s, followed by a
melt curve (65°C to 95°C read in 0.5°C steps). The qPCR
results were analysed using the Bio-Rad CFX Manager 2.0
software (Bio-Rad). To analyse whether the genes were tran-
scribed as an operon in a single polycistronic mRNA strand,
the primers from the qPCR study were used to amplify the
regions between adjacent genes using Phusion polymerase
(Finnzymes) and the constructed cDNA as template.
Enzyme localization studies
The remaining cells from the minimal media cultures, above,
were harvested by centrifugation at 4300 gfor 10 min, and
subjected to osmotic shock as described previously (Larsbrink
et al., 2011). The secreted proteins in the supernatant media
were collected, concentrated and washed with 50 mM sodium
phosphate, pH 7.0, using Amicon Ultra filter units (10 kDa
cut-off; Millipore). The remaining cells were re-suspended in
5 ml sodium phosphate, pH 7.0, and lysed by sonication. The
lysate was centrifuged at 25 000 gfor 45 min and the resulting
supernatant liquid was collected and concentrated with the
same filter units and buffer as above to obtain soluble cyto-
plasmic proteins. The three fractions containing secreted,
periplasmic, and cytoplasmic proteins were each assayed for
β-galactosidase and α-L-fucosidase activities using the chro-
mogenic assays described above; protein concentrations
were measured by the Bradford method.
Construction of targeted gene disruptions and strain
growth analysis
Directed gene disruptions from integration of suicide plasmid
pK18mobsacB (Schafer et al., 1994) into the genome were
facilitated by tri-parental mating as done previously (Gardner
and Keating, 2010; 2012). Confirmation of correct gene dis-
ruptions was performed by PCR using primers listed in
Table S1. C. japonicus strains were grown on MOPS minimal
medium (Neidhardt et al., 1974) with glucose [0.25% (wt/vol)]
or tamarind xyloglucan [0.25% (wt/vol)] as sole carbon
sources. Growth assays were performed in a TECAN
M200Pro at 30°C with vigorous shaking. Growth measure-
ments were taken via optical density (OD) at 600 nm. Growth
experiments were performed in biological triplicates.
Acknowledgements
We are grateful to Dr Lauren S. McKee for assistance in
manuscript preparation. Work in the Brumer lab was sup-
ported by the Mizutani Foundation for Glycoscience, and The
Swedish Research Council Formas (via CarboMat – the KTH
Advanced Carbohydrate Materials Centre), faculty funding
from the Michael Smith Laboratories/UBC, the Natural Sci-
ences and Engineering Research Council of Canada (Discov-
ery Grant), the Canada Foundation for Innovation and the
British Columbia Knowledge Development Fund. Work in the
Davies lab was supported by the Biotechnology and Biological
Science Research Council (BBSRC; Grant BB/I014802/1).
The staff of the Diamond Light Source is thanked for provision
of data collection facilities. Work in the Gardner lab was
supported by start-up funds provided by the Provost’s Office at
UMBC. We thank Prof. Harry J. Gilbert for inspiration.
Conflicts of interest
The authors declare no competing interests.
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Supporting information
Additional supporting information may be found in the online
version of this article at the publisher’s web-site.
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