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The xylan utilization system of the plant pathogen Xanthomonas
campestris pv campestris controls epiphytic life and reveals
common features with oligotrophic bacteria and animal gut
symbionts
Guillaume Dejean1,2*, Servane Blanvillain-Baufume1,2*, Alice Boulanger1,2, Armelle Darrasse3, Thomas Duge de
Bernonville1,2, Anne-Laure Girard3, Sebastien Carrere1,2, Stevie Jamet1,2, Claudine Zischek1,2, Martine Lautier1,2,4,
uttner5, Marie-Agnes Jacques3, Emmanuelle Lauber1,2 and Matthieu Arlat1,2,4
Magali Sole5, Daniela B€
1
INRA, Laboratoire des Interactions Plantes–Microorganismes (LIPM), UMR441, F-31326, Castanet-Tolosan, France; 2CNRS, Laboratoire des Interactions Plantes–Microorganismes (LIPM),
UMR2594, F-31326, Castanet-Tolosan, France; 3INRA, UMR 1345, Institut de Recherche en Horticulture et Semences (IRHS), 42 rue Georges Morel, 49071, Beaucouze CEDEX 01, France;
4
Universite de Toulouse, Universite Paul Sabatier, Toulouse, France; 5Institut f€ur Biologie, Bereich Genetik, Martin-Luther-Universit€at Halle-Wittenberg, D–06099, Halle (Saale), Germany
Summary
Author for correspondence:
Matthieu Arlat
Tel: +33 561 285 047
Email: [email protected]
Received: 7 December 2012
Accepted: 9 January 2013
New Phytologist (2013) 198: 899–915
doi: 10.1111/nph.12187
Key words: epiphytic, gut symbiont,
oligotrophy, TonB-dependent transporter,
transport, xylan, xylanase.
Xylan is a major structural component of plant cell wall and the second most abundant plant
polysaccharide in nature.
Here, by combining genomic and functional analyses, we provide a comprehensive picture
of xylan utilization by Xanthomonas campestris pv campestris (Xcc) and highlight its role in
the adaptation of this epiphytic phytopathogen to the phyllosphere.
The xylanolytic activity of Xcc depends on xylan-deconstruction enzymes but also on transporters, including two TonB-dependent outer membrane transporters (TBDTs) which belong
to operons necessary for efficient growth in the presence of xylo-oligosaccharides and for
optimal survival on plant leaves. Genes of this xylan utilization system are specifically induced
by xylo-oligosaccharides and repressed by a LacI-family regulator named XylR.
Part of the xylanolytic machinery of Xcc, including TBDT genes, displays a high degree of
conservation with the xylose-regulon of the oligotrophic aquatic bacterium Caulobacter
crescentus. Moreover, it shares common features, including the presence of TBDTs, with the
xylan utilization systems of Bacteroides ovatus and Prevotella bryantii, two gut symbionts.
These similarities and our results support an important role for TBDTs and xylan utilization systems for bacterial adaptation in the phyllosphere, oligotrophic environments and animal guts.
Introduction
Xylans represent the predominant hemicelluloses in the cell wall
of terrestrial plants. They comprise a conserved backbone composed of 1,4-linked b-D-xylose residues which may be substituted
with glucuronic acid, 4-O-methyl-glucuronic acid, arabinose or a
combination of types of decorations (Burton et al., 2010; Scheller
& Ulvskov, 2010; Fig. 1a). Altogether, xylans account for
approximately one-third of all renewable organic carbon on
earth. They therefore represent a substantial source of nutriment
and many bacteria are able to degrade this hemicellulolytic substrate (Kulkarni et al., 1999; Saha, 2003; Dodd & Cann, 2009).
These xylanolytic microbes can be found in diverse ecological
niches, which typically comprise environments where plant material accumulates and deteriorates, including plant debris, soil,
aquatic environments and the digestive tract of animals
*These authors contributed equally to this work.
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(Collins et al., 2005). Plant pathogenic bacteria also display xylanolytic activities, which may help them to breach the cell wall obstacle and to release nutrients during the colonization of plants.
Bioconversion of xylans has been intensively studied in the past
decade because of its potential applications in agro-industrial processes, such as the pulp and paper industry and biofuel production.
These studies have shown that xylan bioconversion is mediated by
a wide array of enzymes (Collins et al., 2005; Dodd & Cann,
2009). Although the xylanolytic systems of bacteria isolated from
soil or from digestive tracts of animals have been studied in detail,
there is only limited information regarding the xylanolytic systems
of plant pathogenic bacteria. Moreover, little is known about transport into bacterial cells of xylan deconstruction products.
The Xanthomonas genus comprises an important group of
plant pathogenic bacteria that together affect c. 400 plant hosts,
including agronomically important crops (Buttner & Bonas,
2009; Ryan et al., 2011). Most Xanthomonas species are able to
survive on the aerial part of plants (phyllosphere), a feature that
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β-Xylosidase
(a)
4-O-methyl-glucuronic acid
– OOC
H3CO
HO
O
HO
O
GH67
XCC4102
XCC1283
XCC1404
XCC1775
XCC2892
XCC3814
XCC4106
XCC0149
XCC1178
XCC4064
XCC4105
XCC4122
XCC3975
Acetate
O
O
O
OH
O
OH
O
HO
GH39
β-D-Xylose
O
H3CO
GH43
H 3C
OH
O
α-Glucuronidase
GH3
O
GH43
GH51
XCC0149
XCC1178
XCC4064
XCC4105
XCC4122
XCC1191
XCC1759
xylB
xylA1
xylE
Xylose
isomerase
D-xylose
n
GH30
XCC4115
XCC4118
XCC0857
xytA locus
(c)
D-xylulokinase
O
Xylanase
Arabinofuranosidase
XCC1757
XCC4103
O
GH10
α-L-Arabinose
(b) xylE locus
Acetyl xylan esterase
O
HO
O
OH
Ferulic acid
O
XCC2825
xyaC
XCC2828
xyaB xyaA
xytA
Tryptophan Hyp
Hyp
halogenase protein protein
transporter
TonB-dependent
Transporter
(d) xylR locus
XCC4100
xylA2
XCC4107
GH67
agu67A
xylR
axeXA
Xylose
LacI
α-Glucuronidase
isomerase repressor
Acetyl
esterase
uxuA
GH43
gly43E
GH3
xyl3A
Mannonate β-Xylosidase/
dehydratase arabinosidase
uxuB
β-Xylosidase
Fructuronate
reductase
(e) xytB locus
XCC4115
GH10
XCC4122
G H2
xyn10C
gly2A
Xylanase
Glycosyl
hydrolase
GH10
uxaC
xyn10A
GH43
xypA
Glucuronate Xylanase
MFS
isomerase
transporter
xytB
TonB-dependent
Transporter
plays an important role in the early stages of infection (Rigano
et al., 2007; Li & Wang, 2011). Xylanases have been shown to
control the virulence of two members of this genus, Xanthomonas
oryzae pv oryzae (Xoo) and Xanthomonas campestis pv vesicatoria
(Xcv) (Rajeshwari et al., 2005; Szczesny et al., 2010). The aim of
this study was to characterize the xylan utilization system of
Xanthomonas campestris pv campestris (Xcc) the causal bacterium
of black rot disease of Brassica. Xcc harbours CUT systems (Carbohydrate Utilization with TBDT systems) which are involved in
plant carbohydrate scavenging (Blanvillain et al., 2007). These
systems comprise inner membrane transporters, degrading
enzymes, transcriptional regulators and TonB-dependent outer
membrane transporters (TBDTs; Blanvillain et al., 2007). In
contrast to passive transport mediated by porins, TBDTs allow
high-affinity and active transport of bigger substrate molecules
(Cornelis, 2010; Krewulak & Vogel, 2011). TBDTs have been
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xypB
gly43F
Symporter β-Xylosidase/
transporter arabinosidase
Fig. 1 General structure of xylans and
putative xylan-degrading enzymes of
Xanthomonas campestris pv campestris
ATCC33913 (LMG568) (Xcc-568) and their
genetic organization. (a) The major enzymes
degrading xylan found in Xcc-568 and their
sites of action are depicted with arrows. For
each enzymatic activity, the corresponding
families listed in the CAZy database are
shown and the Xcc-568 proteins belonging
to each family are listed beneath: glycosyl
hydrolase (GH). Proteins belonging to Xcc568 xylan CUT system are indicated in red.
(b–e) Genetic organization of Xcc-568 xylE
(b), xytA (c), xylR (d), and xytB (e) loci.
Genes are represented by arrows, their
names and putative functions are indicated
beneath. Perfect xyl-boxes are represented
by white circles. Genes encoding predicted
enzymatic functions are annotated according
to their CAZy family number. Genes coding
for enzymes involved in xylose metabolism
are in yellow. Genes involved in glucuronic
acid metabolism are in blue. Inner membrane
transporter genes are indicated by a pink
colour. TBDT genes are represented in red.
Other enzymes putatively involved in xylan
or xylo-oligosaccharides degradation are
shown in green.
shown to transport iron-siderophore complexes, vitamin B12 and,
more recently, various carbohydrates (Neugebauer et al., 2005;
Blanvillain et al., 2007; Eisenbeis et al., 2008; Schauer et al., 2008).
A global study of Xcc ATCC33913 (LMG568) (Xcc-568) TBDT
genes has shown that the expression of two of them, XCC2828 and
XCC4120, is specifically induced by xylan and xylose (Blanvillain
et al., 2007). In this study we show that they belong to a complex
CUT system involved in the uptake and utilization of xylan. This
system is important for fitness of Xcc-568 in the phyllosphere.
Materials and Methods
Bacterial strains, plasmids and growth conditions
The Xcc-568 strains and plasmids used in this study are listed in
Supporting Information Table S1. Xcc-568 cells were grown at
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28°C in MOKA rich medium (Blanvillain et al., 2007) or in
minimal medium (MME; Arlat et al., 1991). Escherichia coli cells
were grown on Luria–Bertani medium at 37°C.
Antibiotics were used at the following concentrations: for
Xcc-568, 50 lg ml 1 rifampicin, 50 lg ml 1 kanamycin, and
5 lg ml 1 tetracycline; for E. coli, 50 lg ml 1 ampicillin,
25 lg ml 1 kanamycin, and 10 lg ml 1 tetracycline.
Construction of Xanthomonas campestris pv campestris
mutants
Insertion mutants were constructed using the suicide plasmid
pVO155 (Oke & Long, 1999) with a 300- to 500-bp PCR
amplicon internal to each open reading frame (ORF). Deletion
mutants were constructed by using the cre-lox system adapted by
Angot et al. (2006) from the system of Marx & Lidstrom (2002)
or by using the sacB system (Schafer et al., 1994). Deleted regions
and pVO155 plasmid insertions are indicated in Table S1 and
represented on Fig. S1. Oligonucleotide primers used for PCR
amplification will be provided upon request.
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using the RNeasy Mini Kit (Qiagen). A total of 5 lg of RNA was
reverse transcribed with Transcriptor Reverse Transcriptase
enzyme (Roche Diagnostics, Meylan, France) using random
hexamers (Biolabs, Evry, France) for 10 min at 25°C and then
for 40 min at 55°C. The resulting cDNAs were used as a template for PCR amplification with Taq polymerase using specific
primer pairs for each gene (as indicated in Fig. S2) and analysed
by agarose-gel electrophoresis.
Quantitative reverse transcription-PCR (qRT-PCR) experiments were performed essentially as previously described (Blanvillain et al., 2007). For qRT-PCR, experiments were performed
on bacteria grown on solid medium containing 4-O-Methyl-Dglucurono-D-xylan-Remazol brilliant blue R (RBB-Xylan;
Sigma), colonies obtained after 48 h growth were resuspended in
1 ml of water. A 1 lg sample of RNA was treated with RNasefree DNase I (Sigma) for 20 min at room temperature. After
DNase inactivation (10 min at 70°C), RNAs were reverse transcribed as indicated above. Oligonucleotide primers used for
quantitative PCR amplification will be provided upon request.
16S rRNA was used as a control for real-time PCR (Morales
et al., 2005; Blanvillain et al., 2007).
Plasmid constructions
DNA manipulations were performed using standard procedures
(Sambrook et al., 1989).
For complementation studies, PCR amplicons presented in
Fig. S1 (oligonucleotide primers used for PCR amplification will
be provided upon request) were cloned into pCZ1016, a derivative of pFAJ1700 containing the Ptac promoter, multiple cloning
sites and the T7 terminator from pSC150 (Dombrecht et al.,
2001; Cunnac, 2004). To perform chromosomal complementations, PCR amplicons were cloned into pCZ1034, a derivative of
pK18mobsacB (Schafer et al., 1994) with the MCS replaced by a
Ptac promoter, a MCS and a T7 terminator flanked by a 700-bp
fragment corresponding to the region upstream from the open
reading frame XCC0127 and a 700-bp fragment corresponding to
the region downstream from the open reading frame XCC0128.
The XCC4120 promoter region (see Table S1) was PCR
amplified with appropriately designed primers. This promoter
region was cloned as HindIII–XbaI fragment, into the pCZ962
plasmid, a pFAJ1700 (Dombrecht et al., 2001) derivative
containing the KpnI–AscI lacZ gene from the pCZ367 plasmid
(Cunnac, 2004), giving pPr-xytB.
Calculation of maximal growth rate
Growth curves of Xcc-568 strains grown at 28°C in MME liquid
culture in the presence of xylose or xylo-oligosaccharides were
generated using a FLUOStar Omega apparatus (BMG Labtech,
Offenburg, Germany) with four replicates. Growth was monitored by measuring OD600 using 96-well flat-bottom microtiter
plates with 200 ll preparations inoculated at OD600 of 0.1 from
four independent washed overnight precultures. The microplates
were shaken continuously at 700 rpm using the linear-shaking
mode. Generation time (G), defined as doubling time, was calculated during the exponential phase of growth using the following
formula: G = tf t0/n where n is equal to (logNf logN0)/log2
(N0, initial number of bacteria at the initial time point considered (t0); Nf, final number of bacteria at the final time point considered (tf)). The maximum specific growth rate (lmax), defined
as the increase in cell mass per time unit, was calculated as follows: lmax = ln 2/G. Statistical analysis was performed using the
RGUI software (GNU General Public License; Free Software
Foundation Inc., Boston, MA, USA).
[14C] xylose transport experiments
Expression studies, RNA isolation and operon mapping
b-galactosidase and b-glucuronidase assays: bacterial cultures in
the appropriate medium were harvested at different time points
and b-galactosidase and b-glucuronidase (GUS) assays were performed as previously described (Blanvillain et al., 2007).
In order to investigate the transcriptional organization, reverse
transcription-PCR (RT-PCR) experiments were performed.
Bacterial cultures from xylR mutant of Xcc-568 grown in minimal
medium (MME) were harvested after 6 h of incubation at an
optical density at 600 nm (OD600) of 0.6. RNAs were extracted
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[14C] xylose transport assays were conducted as previously
described (Blanvillain et al., 2007; Boulanger et al., 2010).
[14C] xylose (Amersham Biosciences, specific activity of
3.15 GBq mmol 1) was added to a final concentration of
0.5 lM. For competition experiments, unlabelled sugars were
added to [14C] xylose at final concentrations of 0.5, 5, 50 and
500 lM, and cells were incubated for 1 h before collection.
The initial concentration-dependent xylose transport was determined using the rapid dilution method as previously described
(Neugebauer et al., 2005; Blanvillain et al., 2007).
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Plate assays for detection of xylanase activity
The plate assay for xylanase activity was performed using
MME-agar plates containing 0.1% RBB-xylan (Sigma). Overnight cultures of Xcc-568 strains grown in MOKA medium were
centrifuged. Pellets were resuspended in MME medium and the
OD600 was adjusted to 0.4. Five microlitres of bacterial suspension were spotted on plates that were incubated at 28°C. The
detection of xylanase activity was examined periodically by checking the halo against the blue background.
Pathogenicity tests
Pathogenicity tests were conducted on Arabidopsis thaliana Sf-2
ecotype as previously described (Meyer et al., 2005).
Dynamics of bacterial population densities in the
phyllosphere of cabbage and bean
Experiments on cabbage (Brassica oleracea cv Bartolo) and dry
bean (Phaseolus vulgaris cv Flavert) as well as statistical analyses
were performed at IRHS as previously described (Darsonval
et al., 2008).
In silico analyses
The presence of signal peptides and protein localization were
determined using the SignalP 3.0 server (http://www.cbs.dtu.dk/
services/SignalP/; Emanuelsson et al., 2007).
Patscan and Predetector software (Dsouza et al., 1997; Hiard
et al., 2007) were used to identify xyl-boxes.
Results
XCC2828 and XCC4120 TBDTs genes are located in loci
putatively involved in xylan/xylose metabolism
The XCC2828 and XCC4120 TBDT genes, whose expression is
specifically induced by xylan and xylose, display significant
homologies with TBDTs genes from the aquatic bacterium
Caulobacter crescentus CB15 (Cc-CB15), CC0999 and CC2832,
respectively, (Fig. 2a, Table S2). Interestingly, a transcriptomic
analysis showed that these two Cc-CB15 genes belong to a complex xylose regulon whose repression is mediated by CC3065, a
LacI-family regulator named XylR. This repressor was shown to
recognize a specific 14 bp-operator motif (Hottes et al., 2004;
Stephens et al., 2007a,b; Fig. S3). This motif, found upstream
from both CC0999 and CC2832 TBDT genes has several close
matches in the genome of Xcc-568, two of which are located
upstream from XCC2828 and XCC4120 TBDT genes (Hottes
et al., 2004). The screening of the Xcc-568 genome sequence predicted two additional motifs perfectly matching the 14 bp palindromic motif, named xyl-box, located upstream from XCC2828
and XCC4120 (Table S3). One motif is located upstream from
the XCC4119 gene, encoding a putative inner membrane transporter of the major facilitator superfamily (MFS) and the other
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upstream from the XCC4100 gene, encoding a putative xylose
isomerase (Table 1). Interestingly, several genes surrounding
XCC2828, XCC4100, XCC4119 and XCC4120 genes code for
proteins displaying high similarities to proteins of the xylose
regulon of Cc-CB15 (Fig. 2a; Table S2). Among these proteins,
XCC4101, a putative LacI family regulator, is very well
conserved to XylR from Cc-CB15 and was therefore named
XylR. Moreover, several genes located in these loci have predicted
functions associated with the utilization of xylan, xylose or glucuronic acid (Table 1; Fig. 1). The major enzymes that attack xylan
backbone are classified in the carbohydrate-active enzyme
(CAZy) database (http://www.cazy.org; Cantarel et al., 2009).
They comprise endo-1,4-b-D-xylanases (EC 3.2.1.8), which generate xylo-oligosaccharides (Table S4). The degradation of xylooligosaccharides is mediated by b-D-xylosidases, whereas elimination of the side groups is catalysed by a-L-arabinofuranosidases,
a-D-glucuronidases, acetylxylanesterases, ferulic acid esterases and
p-coumaric acid esterases (Table S4). The genes of Xcc-568 found
in these different glycosyl hydrolases (GH) or carboxylesterase
(CE) families were named to indicate their activity and CAZy
family, as previously described for Cellvibrio japonicus (DeBoy
et al., 2008) and according to the nomenclature recently proposed by Potnis et al. (2011; Table 1). This analysis allowed us to
define three loci, named xytA, xytB and xylR, containing xyl-boxes
and enzymes putatively associated with xylan deconstruction and
glucuronic acid metabolism, as well as inner membrane transporters and TBDTs. They might therefore form a xylan CUT
system (Fig. 1). Moreover, the xylR locus contains xylaA2 gene
encoding a putative xylose isomerase, an enzyme which carries
out the first step in xylose metabolism (Lawlis et al., 1984;
Fig. 3). The analysis of the Xcc-568 proteome showed that this
pathogen possesses a second xylose isomerase gene (named xylA1)
which displays high similarity to xylA2 (97% identity at DNA
level). xylA1 does not belong to the xylan CUT system defined
above. It is located between XCC1759 (xylE) gene, encoding a
putative MFS inner membrane transporter and xylB, a putative
D-xylulokinase gene, thus suggesting that Xcc-568 possesses a classical two-step xylose utilization pathway (Lawlis et al., 1984;
Fig. 3; Table 1). No perfect or even degenerated xyl-box was
found in this locus, named xylE (Fig. 1).
Genes belonging to xytA, xytB and xylR loci are specifically
induced by xylo-oligosaccharides
The expression of most genes located in the xytA, xytB or xylR loci
was studied in the presence of xylan, xylose or xylo-oligosaccharides (xylobiose, X2; xylotriose, X3; xylotetraose, X4). These experiments were performed by using pVO155 insertion mutants which
carry transcriptional fusions between the targeted genes and the
uidA reporter gene (Oke & Long, 1999) or by qRT-PCR.
Most genes in these three loci display a similar expression
pattern: their expression is specifically and highly induced by
xylo-oligosaccharides. They are also induced to a lesser extent by
xylan, and xylose (Tables 2, 3).
xylR regulatory gene and xypB, that code for a putative inner
membrane transporter, showed distinctive expression induction.
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(a)
Xanthomonas
campestris
pv. campestris
ATTCC33913
C l b t
Caulobacter
crescentus
CB15
xylE locus
xytA locus
xylB xylA1 xylE
xylR locus
xyaC xyaB xyaA xytA
xylA2 xylR agu67A
1759
2825
xytB locus
uxuA gly43E
PF03629
GH67
XC
XCC1757
axeXA
xyl3A
GH43
uxuB
GH3
GH10
4107
410 0
2828
xyn10C gly2A
uxaC xyn10A xypA
GH2
xytB
xypB gly43F
GH10
GH43
4115
4122
GH43
GH10
CC0814 0813
3065
xylR
0999
1002
GH67
2812
G
H
GH10
2811
2804
1487
GH43
1490
1508
3042
2832
2802
(b)
xylE locus
Xanthomonas
campestris
pv. campestris
ATTCC33913
Bacteroides
ovatus
ATCC8483
Prevotella
bryantii
B1 4
xyn30A
GH30
XCC0857
GH43
Bacova_02534
02535
Bac
xylR locus
xylB xylA1 xylE
xylA2 xylR agu67A
1757
4100
axeXA
PF03629
GH67
GH3
1759
GH43
GH31
xytB locus
uxuA gly43E
xyl3A
GH43
uxuB
GH3
xyn10C gly2A
GH2
GH10
4107
GH43
GH97
GH67 GH43
0882
0398
GH10
xusA
xusB
0391
0381
xusC
xypB gly43F
GH43
4122
GH97 GH43 GH43
GH10 GH43
xytB
GH10
4115
03417
Pbr_0883
uxaC xyn10A xypA
xusD
GH30
PF03629
03432
03450
GH67 GH43 GH10
PF03629 GH10
04385
04393
xusE xyn10C
GH10
GH43
0377
Core xylan cluster
Legend
Glycosyl hydrolases
Putative acetylesterase
Glucuronate metabolism
Inner membrane transporters
GH2
GH31
PF03629
Glucuronate isomerase
GH3
GH43
Xylose metabolism
Fructuronate reductase
GH10
GH67
Xylose isomerase
GH30
GH97
Xylulokinase
Major facilitator
superfamily
Outer membrane transporters
Regulators
TBDT Xanthomonas/Caulobacter
LacI
TBDT SusC/RagA family
HTCS
Mannonate dehydratase
Fig. 2 Conservation of the xylan CUT system of Xanthomonas campestris pv campestris ATCC33913 with the xylose regulon of Caulobacter crescentus
CB15 (a) and xylan regulons of Bacteroides ovatus ATCC8483 and Prevotella bryantii B14 (b). The genes are colour-coded based on their predicted roles
as indicated in the legend. Genes encoding predicted enzymatic functions are annotated according to their CAZy family number. Transparent stained zones
show conserved genes or loci. ORF numbers are from genome projects hosted in the GenBankTM database. (a) For C. crescentus CB15, genes induced by
xylose (Hottes et al., 2004) are indicated by a purple halo. Blue circles indicate xylose operator motifs of C. crescentus CB15; white circles show perfect
Xcc-568 xyl-boxes. (b) B. ovatus and P. bryantii genes whose expression is induced by xylan are indicated by a blue halo.
When monitored in the xylR::pVO insertion mutant, the expression of xylR was not induced by xylan or xylo-oligosaccharides
(Table 2), whereas its expression is induced by xylan and X3 when
monitored by qRT-PCR in a wild-type background (Table 3;
Fig. S4b). Similarly, when monitored in xypB::pVO insertion
mutant, the expression of xypB gene was not induced by xylan,
X3, X4, or xylose (Table 2) whereas its expression followed the
general induction pattern (i.e. high induction by xylo-oligosaccharides and weaker induction by xylose) when monitored by
qRT-PCR in a wild-type background (Table 3). These observations suggested that a functional copy of this gene might be
required for its own induction by xylan and/or X3 or X4. This
hypothesis was confirmed by introducing the (pC-xypB) plasmid,
expressing xypB constitutively, into the xypB::pVO mutant
(Fig. 4a). Moreover, the induction by X2, X3 and X4 of xytB
TBDT promoter fused to the lacZ reporter gene on the pPr-xytB
plasmid was abolished in the DxypB deletion mutant and recovered by introducing a functional copy of xypB into the DxypB
chromosome (Fig. 4b).
The expression pattern of the xylA1, xylB and xylE genes of the
xylE locus is clearly different from that observed for the CUT
xylan utilization system because they are generally equally
induced by xylose and xylo-oligosaccharides (Table 3). Finally,
the expression of genes coding for other enzymes located outside
the xytA, xylR, xytB and xylE loci, including xyn30A putative
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xylanase, is not induced by xylose, xylo-oligosaccharides or xylans
(data not shown).
XylR represses the expression of genes/operons preceded
by a xyl-box
The conservation of XylR and xyl-boxes in Cc-CB15 and Xcc-568
prompted us to compare the expression of genes located in xyl E,
xytA, xytB and xylR loci in the wild-type strain or in a xylR::pVO
insertion mutant by qRT-PCR analysis. XylR represses the
expression of all genes located immediately downstream from
putative xyl-boxes (i.e. xytA, xytB, xypA and xylA2; Table 4). The
expression of the four genes located downstream of xypA (xyn10A
to xyn10C) is also repressed by XylR (Table 4), suggesting that
they form an operon with xypA. Operon mapping by RT-PCR
analysis confirmed this hypothesis (Fig. S2). Similarly, it
appeared that xytA, xyaA and xyaB, on the one hand, and xytB,
xypB, and gly43F, on the other, form two operons negatively regulated by XylR (Table 4, Fig. S2).
The expression status was clearly different in the xylR locus.
xylA2 which is the unique gene of this locus displaying a xyl-box
is the only one whose expression is repressed by XylR in this
locus. The expression of agu67A, axeXA, uxuA, gly43E, xyl3A and
xylR itself is not repressed by XylR in MME (Table 4). As
the expression of all these genes is specifically induced by
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XytB
XypB
XCC4121
UxaC
XCC4117
XCC4120
Gly2A
XCC4116
XypA
Xyn10C
XCC4119
900/Yesd
UxuB
XCC4107
xytB locus
XCC4115
Xyn10A
GH2
385/Yes
Xyl3A
XCC4106
XCC4118
GH10
487/No
Gly43E
XCC4105
495/No
980/Yes
501/No
330/yes
471/No
896/Yes
565/Yes
419/No
GH10
GH3
GH43
UxuA
GH67
XCC4104
654/Yes
446/No
366/No
739/Yes
313/No
1047/Yes
AxeXA
XyaA
XytA
XCC2827
XCC2828
343/No
XCC4103
XyaB
XCC2826
498/Yes
XylA2
XylR
Agu67A
XyaC
xytA locus
XCC2825
CAZy
family
xylR locus
XCC4100
XCC4101
XCC4102
Name
ORF
Protein size
(aa)/Signal
peptidea
PF00593
-PF07715/TIGR01782
PF07690/COG2211/
TIGR00792
PF07690/TIGR00893
PF00331
PF02836-PF02837
-PF00703
PF02614
PF00331
PF00933-PF01915
-PF07691
PF01232-PF08125
PF04616
PF02746-PF01188
PF01261
PF03566-PF0532
PF03648-PF7477
-PF07488
PF03629
PF07277
PF00593-PF07715
No conserved domain
PF04820
Pfam/COG//TIGRb
GusB (E. coli/YP_001458395.1) (Liang et al., 2005)
Caul_1838 (Caulobacter sp. K31/ABZ70967)
UxaC (Geobacillus stearothermophilus T6/ABI49945)
(Shulami et al., 1999)
XynB, Xyn10B, CJA3280 (Cellvibrio japonicus Ueda107/P23030)
(Kellett et al., 1990; DeBoy et al., 2008)
ExuT (Ralstonia solanaceaum/AAL24034) (Gonzalez & Allen, 2003)
Xyn10A (Bacteroides xylanisolvens XB1A/CBH32823)
(Mirande et al., 2010)
XylC (Cellvibrio mixtus/AAD09439) (Fontes et al., 2000)
Xyn10D, CJA2888 (Cellvibrio japonicus Ueda107/YP_001983344)
(DeBoy et al., 2008)
OTER_3378 (Opitutus terrae PB90-1/ACB76655)
UxuB (Escherichia coli K12/BAA02591) (Blanco et al., 1986)
Xyl3C (P. Bryantii B14/ADD92016) (Dodd et al., 2010a)
ManD (Novosphingobium aromaticivorans/2QJJ_A)
(Rakus et al., 2007)
XylB (Butyvibrio fibrisolvens/P45982) (Utt et al., 1991)
SiaE (Mus musculus/CAA67214) (Stoddart et al., 1996)
XylA (Piromyces sp. E2/CAB76571) (Harhangi et al., 2003)
XylR (C. crescentus CB15/NP421859) (Stephens et al., 2007b)
GlcA67A (Cellvibrio japonicus/AAL5772) (Nurizzo et al., 2002)
PHZ_c2924 (Phenylobacterium zucineum HLK1/YP_002131762.1)
Patl_3278 (Pseudoalteromonas atlantica T6c/YP_662838.1)
Pass1 (Rattus Norvegicus/Q5BKC6) (Liu et al., 2000)
PyrH (Streptomyces rugosporus/AAU95674) (Zehner et al., 2005)
Representative homologous protein (species/accession no)
(reference)c
29/444
51/950
34/389
457/no
979/Yes
439/No
599/Yes
473/No
25/453
39/282
919/Yes
379/Yes
378/Yes
45/359
43/377
68/877
378/Yes
486/No
857/Yes
517/No
402/No
541/Yes
437/No
351/No
732/Yes
238/No
1006/Yes
479/No
519/No
Protein size
(aa)/Signal
peptidea
45/359
41/457
42/821
38/509
32/202
32/262
71/401
61/435
50/351
55/719
48/231
42/999
39/126
34/491
Identity
(%)/amino
acid overlap
Table 1 Identification and properties of the relevant ORFs from Xanthomonas campestris pv campestris ATCC33913 (Xcc-568) xylan CUT system
Putative hexuronate
transporter
TonB-dependent
transporter
Xylo-oligosaccharides
inner membrane
transporter
Endo-1,4-beta-xylanase
Putative glycoside
hydrolase
Glucuronate isomerase
Putative endo
-1,4-beta-xylanase
Putative D-mannonate
dehydratase
Putative beta
-xylosidase-alpha/L
-arabinofuranosidase
Putative beta-D
-xylosidase
Fructuronate reductase
Putative acetylesterase
Xylose isomerase
LacI family repressor
Alpha-D-glucuronidase
Putative Tryptophan
halogenase
Hypothetical
Pass1-related protein
SapC-related protein
TonB-dependent
transporter
Proposed annotation
in Xcc-568
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XylB
XylA1
XylE
xylE locus
XCC1757
XCC1758
XCC1759
Gly43B
Abf51A
Xyl39A
Gly43C
XCC1178
XCC1191
XCC3975
XCC4064
GH51
508/Yesd
544/Yes
GH43
GH39
GH43
549/Yesd
521/Yes
GH30
GH43
GH43
CAZy
family
405/Yes
526/Yes
497/No
446/No
481/No
344/No
Protein size
(aa)/Signal
peptidea
PF04616/COG3507
PF01229/COG3664
PF06964/COG3534
PF04616/COG3507
PF02055/COG5520
PF04616/COG3507
PF00370-PF02782
PF01261
PF00083
PF04616
Pfam/COG//TIGRb
Abf51A CJA_2769 (Cellvibrio japonicus/AAK84947)
(Beylot et al., 2001)
XynB1 (Geobacillus stearothermophilus T/ABI49941)
(Shulami et al., 1999)
XynB (Paenibacillus sp. JDR-2/ABV90487) (Chow et al., 2007)
XynC (Xanthomonas campestris pv vesiscatoria/YP362696)
(Szczesny et al., 2010)
XynC (Erwinia chrysanthemi/AAB53151) (Keen et al., 1996)
XynB (Paenibacillus sp. JDR-2/ABV90487) (Chow et al., 2007)
XynB (Paenibacillus sp. JDR-2/ABV90487) (Chow et al., 2007)
XylB (Piromyces sp. E2/CAB76752) (Harhangi et al., 2003)
XylA (Piromyces sp. E2/CAB76571) (Harhangi et al., 2003)
GlcP (Synechocystis PCC6803/P15729.2) (Zhang et al., 1989)
Xsa (Bacteroides ovatus V975/P49943) (Whitehead, 1995)
BACOVA_04386 (Bacteroides ovatus 8483/ZP_02067379)
(Martens et al., 2011)
XynB, PBR0394 (P. bryantii B14/P48791) (Gasparic et al., 1995)
Representative homologous protein (species/accession no)
(reference)c
29/515
36/483
521/No
504/No
517/Yes
413/Yes
521/No
57/397
33/515
53/508
406/Yes
521/No
81/400
31/427
494/No
437/No
468/No
319/no
54/309
45/494
61/435
52/455
325/no
325/no
Protein size
(aa)/Signal
peptidea
57/313
57/313
Identity
(%)/amino
acid overlap
Putative Beta
-xylosidase/alpha-L
-arabinofuranosidase
Putative Beta
-xylosidase/alpha-L
-arabinofuranosidase
Putative alpha-L
-arabinofuranosidase
Putative Beta-xylosidase
Putative Beta
-xylosidase/alpha
-L-arabinofuranosidase
Putative endo
-1,4-beta-xylanase
D-xylose
Xylose isomerase
inner
membrane transporter
D-xylulokinase
Putative exoxylanase
Proposed annotation
in Xcc-568
b
Signal peptide prediction using SignalP (http://www.cbs.dtu.dk/services/SignalP/; Emanuelsson et al., 2007).
As determined by using the Conserved Domain Database (Marchler-Bauer et al., 2011) and the Pfam database (Finn et al., 2010).
c
The reported homologous proteins are those showing the highest score among proteins with an experimentally defined function. In the absence of relevant biochemical data, the most similar
protein from bacteria outside the Xanthomonadaceae family was reported.
d
Start codon prediction revised in this work. All other start codons are from GenBank (da Silva et al., 2002) or from Blanvillain et al. (2007).
a
Xyn30A
XCC0857
Other genes
XCC0149 Gly43A
Gly43F
Name
XCC4122
ORF
Table 1 (Continued)
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Table 2 Relative expression ratios measured by using pVO155 reporter plasmid insertions in genes of the xylan utilization system grown in the presence of
xylan, xylose or xylo-oligosaccharides
Expression ratiosb (SDc)
2 mM
Locus
Gene IDa
Name
Orienta
0, 125%
MME Xnd/MME
xytA
xylR
XCC2828
XCC4101
XCC4102
XCC4103
XCC4104
XCC4105
XCC4106
XCC4107
XCC4115
XCC4116
XCC4117
XCC4118
XCC4120
XCC4121
XCC4122
xytA
xylR
agu67A
axeXA
uxuA
gly43E
xyl3A
uxuB
xyn10C
gly2A
uxaC
xyn10A
xytB
xypB
gly43F
R
R
F
F
F
F
F
F
R
R
R
R
F
F
F
20.85 (2.02)
0.81 (0.02)
0.95 (0.09)
1.71 (0.04)
1.56 (0.21)
4.80 (0.04)
3.76 (0.002)
4.08 (0.02)
11.68 (1.31)
12.90 (0.26)
11.58 (0.85)
5.78 (1.23)
39.56 (5.31)
1.42 (0.06)
48.51 (2.02)
xytB
20 mM
MME X1d/MME
MME X1d/MME
MME X2d/MME
MME X3d/MME
MME X4d/MME
15.66 (2.76)
0.37 (0.01)
6.87 (0.95)
4.31 (0.15)
3.28 (0.32)
3.43 (0.38)
3.29 (0.06)
3.34 (0.17)
1.95 (0.14)
2.00 (0.19)
2.42 (0.03)
1.17 (0.18)
29.20 (1.49)
1.08 (0.25)
30.02 (9.18)
6.68 (0.27)
0.55 (0.03)
1.61 (0.14)
1.87 (0.23)
1.33 (0.05)
1.12 (0.04)
1.20 (0.14)
1.40 (0.06)
0.92 (0.12)
1.21 (0.04)
1.47 (0.21)
0.97 (0.15)
1.65 (0.12)
0.44 (0.01)
1.06 (0.01)
59.51 (4.39)
0.73 (0.01)
27.15 (6.29)
19.69 (3.41)
11.01 (0.29)
21.89 (6.53)
13.48 (1.86)
15.49 (1.51)
18.23 (1.73)
29.73 (1.11)
31.39 (1.89)
4.00 (0.28)
129.75 (2.81)
4.48 (0.13)
10.64 (2.42)
67.42 (1.10)
0.63 (0.02)
24.79 (6.36)
19.07 (1.77)
9.04 (0.12)
23.27 (3.57)
13.03 (1.19)
14.84 (0.39)
15.07 (1.03)
23.42 (2.74)
27.16 (2.85)
4.59 (0.35)
152.15 (2.03)
0.80 (0.02)
11.26 (1.65)
63.14 (4.60)
0.64 (0.02)
23.97 (5.74)
17.79 (1.56)
9.97 (0.59)
21.28 (6.04)
13.03 (2.40)
14.67 (0.25)
15.57 (0.55)
26.07 (2.60)
28.05 (2.31)
5.01 (0.70)
137.63 (3.53)
0.67 (0.03)
5.19 (0.91)
a
Gene ID and transcriptional orientation are from Xanthomonas campestris pv campestris strain ATCC33913 (da Silva et al., 2002). F, forward; R, reverse.
All ratios are from expression monitored by measuring b-glucuronidase activity of mutants carrying pVO155 insertion in the tested genes.
c
SD, standard deviation obtained from values of three independent experiments.
d
Minimal medium (MME) was supplemented with xylan (Xn), xylose (X1), xylobiose (X2), xylotriose (X3) or xylotetraose (X4).
b
Table 3 Relative expression ratios measured by qRT-PCR for genes in the xylan utilization system in the presence of xylan, xylose or xylo-oligosaccharides
Expression ratiosb (SDc)
2 mM
Locus
Gene IDa
Name
Orient.a
0, 125%
MME Xnd/MME
xytA
XCC2825
XCC2826
XCC2828
XCC4100
XCC4101
XCC4102
XCC4103
XCC4107
XCC4119
XCC4120
XCC4121
XCC4122
XCC1757
XCC1758
XCC1759
xyaC
xyaB
xytA
xylA2
xylR
agu67A
axeXA
uxuB
xypA
xytB
xypB
gly43F
xylB
xylA1
xylE
R
R
R
F
R
F
F
F
R
F
F
F
F
F
F
2.16 (0.06)
2.54 (0.83)
20.32 (1.42)
1.36 (0.07)
2.54 (0.11)
6.25 (1.06)
4.25 (1.15)
4.37 (0.68)
23.47 (0.4)
43.49 (6.32)
3.19 (0.69)
17.17 (3.65)
1.47 (0.55)
1.37 (0.03)
0.99 (0.21)
xylR
xytB
xylE
20 mM
MME X1d/MME
MME X1d/MME
MME X2d/MME
MME X3d/MME
MME X4d/MME
nde
nd
nd
3.03 (0.39)
nd
nd
nd
nd
3.96 (0.62)
nd
nd
2.31 (0.66)
4.55 (0.87)
20.52 (0.64)
6.04 (0.54)
0.92 (0.04)
nd
nd
3.21 (0.23)
1.08 (0.29)
5.15 (2.22)
2.60 (0.62)
3.39 (1.40)
1.02 (0.06)
3.07 (1.01)
1.48 (0.23)
0.87 (0.18)
6.00 (0.64)
3.23 (1.70)
4.54 (0.25)
3.61 (0.76)
nd
nd
20.25 (2.70)
nd
nd
nd
nd
14.24 (1.25)
nd
7.13 (3.26)
25.77 (9.58)
4.91 (2.23)
3.24 (1.94)
2.82 (0.80)
4.78 (0.72)
7,48 (1.03)
34.65 (1.54)
19.37 (2.99)
10.57 (1.92)
69.37 (11.35)
54.07 (13.73)
30.62 (2.06)
15.95 (1.24)
266.69 (28.91)
5.41 (0.99)
20.13 (6.32)
4.03 (1.33)
2.42 (0.99)
2.46 (0.43)
2.53 (0.63)
nd
nd
18.08 (2.61)
nd
nd
nd
nd
16.52 (1.48)
nd
3.43 (1.00)
15.95 (5.20)
2.67 (0.23)
1.15 (0.49)
1.73 (0.15)
a
Gene ID and transcriptional orientation are from Xanthomonas campestris pv campestris strain ATCC33913 (da Silva et al., 2002). F, forward; R, reverse.
Expression was determined by qRT-PCR in the wild-type strain; calculation of relative expression includes normalisation against the 16S rRNA endogenous
control.
c
SD, standard deviation obtained from values of three independent experiments.
d
Minimal medium (MME) was supplemented with xylan (Xn), xylose (X1), xylobiose (X2), xylotriose (X3) or xylotetraose (X4).
e
nd, not determined.
b
xylo-oligosaccharides and to a lesser extent by xylose, we compared their expression by qRT-PCR in wild-type or xylR::pVO
genetic backgrounds. Our data clearly show that the induction of
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xylR expression by X3 depends on a functional copy of xylR (Fig.
S4b). Similarly, the induction of agu67A, axeXA and uxuB by X1
and X3 appears to be positively influenced by XylR (Fig. S4).
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Xylan
Xyn10A
(XCC4118)
AX2
GAX3
GA
X3
?
Xyn10C
(XCC4115)
Agu67A
AxeXA
Gly2A
Gly43E
XyaC
Xyl3A
Xyn30A
XytA
?
?
xylose
X2
?
?
?
XytB
Periplasm
?
XypB
XypA
?
XylE
Cytoplasm
D-glucuronate
D-xylose
Glucuronate
isomerase
Xylose isomerase
UxaC
Gly43F
G
XylR
(XCC4117)
(XCC4122)
(XCC4101)
D-fructuronate
XylA1
XylA2
(XCC1758)
(XCC4100)
D-xylulose
Fructuronate
reductase
Xylulokinase
(XCC4107)
F-6-P
D-mannonate
Mannonate
dehydratase
GA3P
UxuA
Glycolysis
G-6-P
UxuB
XylB
(XCC1757)
D-xylulose-5-P
Pentose
cycle
(XCC4104)
2-keto-3-deoxygluconate (KDG)
Pyruvate
KDG kinase
KdgK
(XCC0118)
KDGP aldolase
2-keto-3-deoxy6phosphogluconate (KDGP)
KdgA
(XCC2140)
-D-Xylopyranose
-D-Glucopyranuronic acid
also with 4-O-methyl groups (
)
-L-Arabinofuranose
O-acetyl groups
Fig. 3 Model of xylan degradation pathway in Xanthomonas campestris pv campestris ATCC33913. Xyn10A is a key extracellular enzyme in the
degradation of xylan. This endo-1,4-b-xylanase of family GH10 releases short to medium-sized xylo-oligosaccharides that can be substituted with various
side chains such as L-arabinose, D-glucoronic acid or its 4-O-methyl ether, thus generating decorated or nondecorated xylo-oligosaccharides such as
glucuronoxylotriose (GAX3), arabinoxylobiose (AX2), xylotriose (X3) or xylobiose (X2), for example. These compounds are either directly taken up into the
periplasm or further degraded in the extracellular medium to generate transportable molecules. The transport of some hydrolysis products might be
mediated by XytA, XytB or as yet unidentified TBDTs or unknown porins. The transported degradation products are further degraded in the periplasm to
generate short xylo-oligosaccharides (X2, X3 …). The exact location of the different degradation steps is not yet known. Enzymes displaying a signal
peptide are active either in the periplasm or in the extracellular medium or even bound to membranes. They are shown in the yellow box that crosses the
outer membrane. The xylo-oligosaccharides are then transported into the cytoplasm by XypB inner membrane transporter. Xylose monomers present in the
periplasm are taken up through XylE, whereas glucuronic acid might be transported by XypA putative hexuronate transporter. Inside the cell, xylooligomers are hydrolysed to xylose by Gly43F putative exoxylanase. Xylose is converted into xylulose-5-phosphate, which can enter the pentose cycle.
D-glucoronic acid is converted to glyceraldehyde 3-P and pyruvate by a five-step pathway catalysed by three enzymes of the xylan/xylose CUT system,
UxaC, UxuB, and UxuA and two other enzymes KdgK, and KdgA. Glyceraldehyde 3-P and pyruvate can enter the Embden–Meyerhof–Parnas pathway.
This might also be the case for the uxuA, gly43E and xyl3A genes
which are locaded between axeXA and uxuB and that seem to
form a large operon with agu67A. Indeed, expression experiments
carried out by qRT-PCR analysis with the wild-type strain and
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the agu67A::pVO mutant suggest that the pVO insertion into
agu67A, the first gene of this putative operon, has a polar effect
on the transcription of axeXA and uxuB (Fig. S5) as well as uxuA,
gly43E and xyl3A (data not shown). This insertion into agu67A
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xypB expression
No supplement
250
2 mM xylose
2 mM xylobiose
200
2 mM xylotriose
2 mM xylotetraose
150
100
50
0
xypB::pVO
xypB::pVO
/pC-xypB
(b)
β-galactosidase activity (Miller units)
β-glucuronidase activity (Miller units)
(a)
xytB promoter activity (pPr-xytB)
1000
No supplement
2 mM xylose
2 mM xylobiose
800
2 mM xylotriose
2 mM xylotetraose
600
400
200
0
Wild-type
ΔxypA
ΔxypB
ΔxypB::xypB
Genetic background
Genetic background
Fig. 4 Expression of the Xanthomonas campestris pv campestris ATCC33913 (LMG568) xypB and xytB genes in presence of xylose or xylooligosaccharides. (a) The expression of xypB was monitored in xypB::pVO insertion mutant or in xypB::pVO strain carrying the complementation plasmid
pC-xypB (xypB::pVO/pC-xypB) by measuring the b-glucuronidase activity after 6 h of growth in MME supplemented with xylose or xylo-oligosaccharides
at a final concentration of 2 mM. (b) The pPr-xytB plasmid carrying the promoterless lacZ reporter gene under the xytB promoter region was used to
monitor xytB expression in presence of xylose or xylo-ol igosaccharides in different genetic backgrounds. b-galactosidase activity was measured after 6 h
induction in MME supplemented with xylose or xylo-oligosaccharides at a final concentration of 2 mM. Bars, SD calculated from at least three different
biological repetitions.
Table 4 Regulation of genes in the xylan CUT system by XylR
Locus
Gene IDa
Name
Orientationa
Expression ratiosb (SDc)
xylR::pVO mutant in
MME/Wild type in MME
xytA locus
XCC2825
XCC2826
XCC2827
XCC2828d
XCC4100d
XCC4101
XCC4102
XCC4103
XCC4104
XCC4105
XCC4106
XCC4107
XCC4115
XCC4116
XCC4117
XCC4118
XCC4119d
XCC4120d
XCC4121
XCC4122
xyaA
xyaB
xyaC
xytA
xylA2
xylR
agu67A
axeXA
uxuA
gly43E
xyl3A
uxuB
xyn10C
Gly2A
uxaC
xyn10A
xypA
xytB
xypB
gly43F
R
R
R
R
F
R
F
F
F
F
F
F
R
R
R
R
R
F
F
F
5.99 (1.21)
3.98 (1.79)
110.33 (31.91)
685.25 (142.12)
7.16 (2.60)
1.57 (0.60)
1.31 (0.24)
0.35 (0.10)
0.78 (0.19)
0.62 (0.06)
1.00 (0.57)
0.97 (0.01)
10.28 (0.75)
61.8 (12.12)
129.34 (21.81)
54.98 (12.15)
20.09 (2,27)
358.6 (74.96)
85.25 (5.95)
47.94 (3.79)
xylR locus
xytB locus
a
Gene ID and transcriptional orientation are from Xanthomonas campestris pv campestris strain ATCC33913 (da Silva et al., 2002). F, forward; R, reverse.
Expression was obtained by qRT-PCR with bacteria grown in MME; calculation of relative expression includes normalisation against the 16S rRNA
endogenous control.
c
SD, standard deviation calculated from values of at least three independent experiments.
d
Contains a xyl-box motif upstream.
b
has no effect on the transcription of xylR, xyn10A or xytB
(Fig. S5).
Finally, the expression of genes of the xylE locus (Fig. S4a and
data not shown) or coding for other enzymes putatively involved
in xylan deconstruction but located outside the xytA, xylR or xytB
loci, including xyn30A putative xylanase, is not controlled by
XylR (data not shown).
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Xyn10A, Agu67A, Gly43F, XylR and XypB control the
production of extracellular xylanase activity
In order to see whether genes belonging to the xytA, xylR, and
xytB loci are involved in the production of the extracellular xylanolytic activity produced by Xcc-568, mutants in these loci were
tested for the production of extracellular xylanase activity. Most
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of the studied mutants displayed xylanase activities similar to that
of the wild-type strain (data not shown). However, some mutants
were significantly affected (Table 5). The level of xylanase activity
was increased in the xylR repressor mutant, thus confirming that
this gene represses the expression of genes required for xylan degradation. More surprisingly, the activity was also significantly
higher in the Dgly43F deletion mutant than in the wild-type
strain (Table 5). This mutant was the only mutant of family
GH43 to show a modification in xylanase activity. The introduction of the pC-gly43F complementation plasmid into the
Dgly43F mutant significantly reduced the level of xylanase activity, confirming the role of this enzyme in the production of
xylanase activity.
No activity was detected for Dxyn10A mutant, which carries a
deletion of xyn10A xylanase gene. Complementation experiments
conducted with pC-xyn10A plasmid, confirmed that the extracellular activity detected in these conditions is coded by xyn10A
gene (Table 5). Accordingly, we did not observe any significant
reduction in extracellular xylanase activity in Dxyn10C or
xyn30A::pVO mutants affected in the two other putative xylanase
genes of Xcc (Table 5). The level of extracellular xylanase activity
was also significantly lower in agu67A::pVO mutant. This
reduced phenotype was complemented by the introduction of
pC-agu67A plasmid (Table 5). This suggests that the putative
a-glucuronidase encoded by this gene is mandatory to get full
extracellular xylanase activity. Finally, the activity was severely
decreased in DxypB inner membrane transporter mutant but not
in any other transporter mutants (Table 5). Complementation
experiments carried out with pC-xypB plasmid confirmed that
the reduction in xylanase activity is due to the mutation in this
gene. This result correlates well with expression results suggesting
that this putative transporter plays a crucial role in the induction
of the system.
Xylose is transported across Xcc-568 inner membrane by
XylE
Table 5 Production of extracellular Xylanase by Xanthomonas campestris
pv campestris strains
Strain
Xcc-568 (wild-type)
Putative xylanase mutants
xyn30A::pVO
Dxyn10C
Dxyn10A
DXyn10A/pC-xyn10A
WT/pC-xyn10A
Other xylan degradation associated mutants
agu67A::pVO
agu67A::pVO/pCZ1016b
agu67A::pVO/pC-agu67A
Dgly43F
Dgly43F/pC-gly43F
Inner membrane transporter mutants
DxypA
DxypB
DxypB/pC-xypB
xypB ::pVO
xypB ::pVO/pC-xypB
xylE::pVO
TonB-dependent transporter mutants
DxytA
DxytB
DxytADxytB
Regulatory mutants
xylR::pVO
xylR::pVO/pC-xylR
Xcc-568/pC-xylR
Xylanase relative
level (plate assay)a
+
+
+
++
++
+/
+/
+
+++
+/
+
+/
+
+/
+++
+
+
+
+
+++
+/
+/
Xylanase relative activity was estimated by calculating the (H2–C2)/C2
ratio, where H is the diameter of the halo and C the diameter of the
bacterial colony, measured 4 d after spotting. The symbols +++, ++, +, +/
or refer to the production of very high, high, medium, low or
nonproduction of xylanase relative activity by the different strains.
b
pCZ1016 is the empty expression vector for complementation
experiments. This empty vector was introduced into all tested mutants
without affecting xylanase activity (data not shown) as shown for
agu67A:: pVO mutant.
a
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The phenotype of xypB mutants, the presence of TBDT and
other inner membrane transporter genes in the XylR regulon
prompted us to study the transport of xylose and xylo-oligosaccharides by Xcc.
The initial concentration-dependent [14C]xylose transport,
reflecting the dissociation constant (Kd) for xylose uptake was
determined using the previously described rapid dilution method
(Neugebauer et al., 2005; Blanvillain et al., 2007). The deduced
Kd (122 lM) is in a range similar to that of Kd values obtained
for passive diffusion through porins (Boulanger et al., 2010).
Moreover, the kinetic values showed that the uptake rate was low
and monophasic (Fig. 5), suggesting passive diffusion. In agreement with these data the transport of xylose is not depending on
XytA and XytB TBDTs (Table 6). Experiments performed with
mutants in xypA, xypB or xylE, the inner membrane transporter
genes identified in the xylan/xylose CUT system, showed that
XylE only is required for xylose transport across the inner membrane. The uptake rate of labelled xylose obtained for xylE
mutants represented only c. 20% of the rate obtained for the
wild-type strain (Table 6). These results were confirmed by comparing maximum specific growth rates (lmax) of the wild-type
strain and mutants in transporter genes in MME supplemented
with xylose. Growth of the xylE::pVO mutant was impaired on
MME containing xylose, contrary to the xypA, xypB, DxytA or
DxytB1 mutants (Fig. 6a). In the xylE::pVO-complemented
strain, xylose transport capacity and growth on MME-xylose
were both restored (Table 6; Fig. 6a).
xytB locus is required for normal growth in presence of
xylo-oligosaccharides
Because growth of the wild-type strain and xylE mutants in the
presence of xylose corroborated the transport status observed for
[14C] xylose uptake by these strains, we speculated that growth
rate studies might indirectly allow us to study the transport of
xylo-oligosaccharides. We focused this analysis on the xytA and
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Table 6 Rates of 14C-labelled xylose transport of mutants compared to the
rate in Xanthomonas campestris pv campestris ATCC33913 wild-type
straina
Strain
Mean%
transport (SD)b
Protein family
Wild-type
xylR::pVO
DxytA-DxytB1
DxypA
DxypB
DxypA-DxypB
xylE::pVO
xylE::pVO/pC-xylE
DxypA-DxypB-xylE::pVO
LacI family regulator
TBDT
MFS transporter
Sugar-cation symporter
Inner membrane
transporters
MFS transporter
Inner membrane
transporters
100 (6.4)
110.3 (19.6)
92.5 (16.3)
106.3 (9.4)
97 (4.2)
109.1 (3.8)
19.8 (3.3)
101.4 (7.6)
20 (2.7)
a
Transport rates were measured 60 min after addition of 14C-labelled
xylose.
b
Standard deviations were calculated from three independent
experiments.
[14C] xylose molecules per cell (×1000)
450
400
350
300
250
type growth rate in the presence of X3 was restored to the
gly43F::pVO insertion mutant by introducing the pC-gly43F
complementation plasmid. Similarly, the DxypB deletion mutant
could be complemented by introducing the pC-xypB plasmid.
However, the xypB::pVO insertion mutant was only partially
complemented by the introduction of pC-xypB whereas it was
fully complemented by the pC-xypB-gly43F plasmid (Fig. 6b).
These results confirm that xypB and gly43F are co-transcribed. In
the presence of X3, the lmax of DxytB1 and xytB::pVO mutants
was also reduced but remained higher than that of the xypB or
gly43F insertion mutants (Fig. 6b). Surprisingly, although considered as nonpolar, the deletion introduced into xytB mutants
could not be complemented by introducing the complementation plasmid pC-xytB (Fig. 6b). Smaller deletions were constructed (Fig. S1) and similar results were obtained even with the
smaller deletion mutant (DxytB3) (data not shown). The DxytB1
and xytB::pVO mutants were fully complemented by the pCxypB-gly43F plasmid (Fig. 6b). Previous data obtained with the
XylR::pVO mutant showed that xytB, xypB and gly43F form an
operon. However, the fact that the lmax of the xytB insertion or
deletion mutants is similar and higher than that of the xypB or
gly43F insertion mutants in the presence of X3 suggests that xytB
is not fully co-transcribed with xypB and gly43F in these conditions. Moreover, complementation experiments suggest a cis-regulatory effect. Altogether, these data show that that xypB and
gly43F may play an important role in xylo-oligosaccharide transport and metabolism. The phenotype of xytB deletion mutants
renders the study of the role of this TBTD in xylo-oligosaccharide transport difficult to assess.
200
xytA and xytB loci are important for growth on plant leaves
150
100
50
0
0
20
40
60
[14C] xylose (μM)
Fig. 5 Concentration-dependent transport of 14C-labelled xylose into
Xanthomonas campestris pv campestris. Cells were grown in minimal
medium without xylose, and transport was measured for 15 s at the [14C]
xylose concentrations indicated. The error bars indicate SD obtained
from three independent experiments.
xytB loci because they both contain transporter genes. The wildtype strain and mutants in these loci were grown in MME supplemented with X2, X3 or X4, at a final concentration of 2 mM, a
concentration that induced the expression of most genes of the
xylan utilization system. Data obtained with X3 are presented
(Fig. 6b). Similar results were obtained with X2 and X4 (data not
shown).
We noticed that lmax of the wild-type strain was slightly lower
in presence of X3 than in the presence of xylose (Fig. 6). The lmax
of the xytB, xypB and gly43F mutants was significantly affected in
the presence of X3 as compared to that of the wild-type strain,
whereas it was not impaired in presence of xylose (Fig. 6). WildNew Phytologist (2013) 198: 899–915
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We studied pathogenicity of pVO155 insertion mutants constructed in this study on cabbage or Arabidopsis thaliana host
plants. These experiments were performed using two distinct
methods: the wound inoculation method, that allows direct
delivery of bacterial cells into the xylem vessels of leaves, or the
infiltration method, which delivers bacteria into the plant leaf
mesophyll (Meyer et al., 2005). None of the mutants tested,
including the xytA, xytB, xypB single mutants and DxytA-DxytB1
double mutants, as well as mutants altered in the three xylanase
genes, were significantly affected in pathogenicity (data not
shown). The growth of xytA::pVO and xytB::pVO mutants in
Arabidopsis plant tissues was also not significantly different from
that of the wild-type (data not shown). We also compared the
survival and the multiplication of the wild-type strain and xytA::
pVO or xytB::pVO mutants in the phyllosphere of cabbage (host
plant) or bean (nonhost plant). The dynamics of bacterial population densities was followed after spray inoculation of the leaves
in conditions that do not favour disease expression (Darsonval
et al., 2008). The multiplication of the xytB::pVO mutant on
cabbage was significantly lower than that of the wild-type strain
only during the first 8 d following the inoculation (Fig. 7a). Cell
densities measured for the xytA::pVO mutant on host plants were
clearly lower than that measured for the wild-type strain and xytB
mutant (Fig. 7a). Interestingly, the survival of both xytA and xytB
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0.20
0.20
–1
Maximum specific growth rate (μmax, h )
(b)
–1
0.15
*
0.10
0.05
0.15
0.10
* *
* *
*
*
0.05
*
*
c-5
68
/pC
Z1
xyl
01
6
E::
pV
O/
pC
xyl
Z1
E::
01
pV
6
O/
pC
-xy
l
E
Δx
ytA
/pC
Z1
01
6
Δx
yp
A/p
CZ
10
16
Δx
y
xyt tB1/
pC
B::
Z1
pV
0
O/
pC 16
Z1
01
Δx
6
y
p
xyp
B/p
B::
C
Z
pV
10
O
/pC 16
gly
43
Z1
F::
01
pV
6
O/
pC
Z1
01
6
Xc
c-5
68
/pC
Z1
01
xyl
6
E::
pV
O/
pC
Z1
01
6
Δx
ytA
/pC
Z1
01
Δx
6
ytA
/pC
-xy
Δx
tA
yp
A/p
CZ
10
16
Δx
ytB
1/p
CZ
Δx
10
Δx
yt
16
ytB
1/p B1/p
CCxyt
xyp
xyt
B
B
B::
pV -gly4
3F
O/
xyt
xyt
p
CZ
B::
B::
10
pV
pV
1
O/
pC O/pC 6
-xy
pB xytB
-gl
y4
3F
Δx
yp
B/p
Δx
CZ
yp
xyp
B/p 1016
B::
Cxyp
pV
O/
B
x
pC
yp
xyp
B::
Z1
B::
p
01
VO
pV
6
O/
/pC
pC
-xy
-xy
pB
pB
gly
-gl
43
y4
F::
3F
pV
gly
O/
43
p
F::
CZ
pV
10
O/
16
pC
-gl
y4
3F
0
0
Xc
Maximum specific growth rate (μmax, h )
(a)
Research 911
Xylose
Xylotriose
Fig. 6 Maximal specific growth rates of Xanthomonas campestris pv campestris wild-type (WT) and mutant strains in the presence of xylose (a) or
xylotriose (b). After overnight growth in rich medium, cells were harvested, washed and resuspended in minimal medium. Xylose and xylotriose were
added at a final concentration of 2 mM. Maximal specific growth rates (lmax) were calculated during the log phase of growth. Hatched bars correspond to
complementation experiments. Colour codes correspond to functional categories as described in Fig. 2. Bars, SD obtained from at least three
independent experiments. The asterisks indicate a significant difference with P < 0.05 as compared to the WT strain in the same culture condition based on
the results of an unpaired Kruskal–Wallis’s test.
mutants was significantly altered on nonhost plant and the defect
of the xytA mutant was again more pronounced than that of the
xytB mutant (Fig. 7b).
Discussion
The phyllosphere represents the aerial parts of terrestrial plants
including leaves, stems, buds, flowers and fruits. This habitat has
been estimated to cover a global surface of c. 1 billion square
kilometres supporting > 1026 bacteria (Morris & Kinkel, 2002;
Lindow & Brandl, 2003; Whipps et al., 2008). Although the
phyllosphere has been less intensively studied than the rhizosphere, metagenomic approaches have recently given interesting
information on bacterial communities colonizing this vast niche
(Vorholt, 2012). Recently, a metaproteogenomics analysis performed on leaves of soybean, clover and Arabidopsis identified
TBDTs as the most prominent group of transport proteins.
These transporters were over-represented among the proteins
assigned to Sphingomonas which was one of the predominant genera identified in a study by Delmotte et al. (2009). It was postulated that this over-representation of TBDTs might play a role in
the successful adaptation of these bacteria on plant leaves. In this
study, by performing functional and genomic analyses of xylan
utilization in Xcc, we identified a xylan CUT system which is
required for optimal colonization of plant leaves. Therefore, our
work seems to confirm the importance of TBDTs for the adaptation of bacteria to the phyllosphere.
The xylan CUT system of Xcc-568 comprises the xytA, xytB
and xylR loci which contain enzymes for the degradation of xylan,
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the metabolism of xylose and glucuronic acid, as well as inner
membrane transporters beside TBDTs. We also identified a
fourth locus, xylE, involved in xylose utilization (see model
Fig. 3).
The expression of most of genes of the xylan CUT system is
specifically and highly induced by short xylo-oligosaccharides
and to lesser extent by xylose. The expression of a large proportion of these genes is repressed by XylR LacI-type repressor. The
regulation mediated by XylR is strictly correlated with the presence of a 14-bp palindromic xyl-box motif in the promoter
region of repressed genes or operons. Interestingly, six contiguous
genes, agu67A, axeXA, uxuA, gly43E, xyl3A and uxuB, which
seem to form an operon, although being induced by xylo-oligosaccharides or xylose, are not under the repression of xylR and no
xyl-box was identified in their promoter regions. On the contrary,
their induction by X3 as well as that of xylR is positively affected
by XylR. This observation shows that the induction by xylooligosaccharides or xylose is not solely under XylR control and
suggests the existence of other regulators controlling the utilization of xylan and xylose in Xcc-568. Further work is needed to
characterize the inducer of XylR in Xcc-568 and to identify other
putative regulators of this system.
Among the three xylanase genes identified in the Xcc-568
genome, xyn10A located in the xytB locus was shown to be
responsible for the detected extracellular activity produced by this
bacterium in our test conditions. No extracellular activity associated with Xyn10C (XCC4118) or Xyn30A (XCC0857) was
detected, although both proteins harbour a signal peptide and
seem to be secreted like Xyn10A (XCC4115) (see Fig. S6 and
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Population sizes
log (CFU g–1 FW)
(a)
7
6
A
A
5
4
xytB::pVO
A
B
B
A
3
B
B
2
WT
A
A
xytA::pVO
B
B
A
A
B
Cabbage
1
0
0
3h
2
4
6
8
10
12
Time post-inoculation (d)
Population sizes
log (CFU g–1 FW)
(b)
7
A
A
A
6
A
A
A
A
B
B
A
A
5
4
A
B
B
3
C
WT
xytB::pVO
xytA::pVO
2
Bean
1
0
0
3h
2
4
6
8
10
12
Time post-inoculation (d)
Fig. 7 Colonization of cabbage and bean leaves by the wild-type strain
Xanthomonas campestris pv campestris ATCC33913 (LMG568) and
strains mutated in xytA or xytB. (a) Bacterial population densities on
cabbage host plants (CFU per gram of fresh weight) were determined on
leaves sampled at 3 h and 1, 4, 8 and 11 d after spray inoculation
(1 9 106 CFU ml 1). (b) Similar experiments were performed on bean
(nonhost plant) leaves. Means and SEMs were calculated for five leaves
per plant species and per sampling date. Mean population densities
followed by different letters are significantly (P < 0.05) different on the
Mann–Whitney test. These experiments were conducted two times
independently and similar results were obtained.
Methods S1). Xyn30A is the orthologue of XynC which is
responsible for the extracellular xylanase activity detected in Xcv
(Szczesny et al., 2010). Despite the high similarity between
Xyn30A and XynC (81% amino acid identity), the presence in
both cases of a signal peptide and the conservation of GH30 family specific catalytic residues (Hurlbert & Preston, 2001; Larson
et al., 2003; St John et al., 2011), we did not detect any extracellular activity associated with the xyn30A gene in Xcc-568. This
gene is neither regulated by XylR nor induced by xylan and xylooligosaccharides in Xcc-568. We observed that beside Xyn10A,
Agu67A, a putative a-glucuronidase involved in the degradation
of glucuronic acid decorations, is also required to get full extracellular xylanase activity. This suggests that removal of these side
chains from the xylan backbone may potentiate the degradation
of xylan. The importance of glucuronic acid liberation during
xylan degradation is underscored by the presence of enzymes
involved in the metabolism of this carboxylic acid in the xylR and
xytB loci (Figs 1, 3).
There might also be a coupling between xylan degradation and
xylose metabolism because the xylan CUT system comprises the
xylA2 gene which codes for a putative xylose isomerase gene. This
gene is duplicated in the Xcc-568 genome and the second copy,
xylA1, maps in the xylE locus with xylE inner membrane transporter gene, required for xylose uptake. The three genes forming
the xylE locus are not under the regulation of XylR and have a
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different induction pattern than that of xylan CUT system genes,
because they are equally induced by xylose and xyl-oligosaccahrides. These observations suggest that there are different regulators for xylose and xylan utilization pathways, but they also
suggest that both pathways are interconnected through the
metabolism of xylo-oligosaccharides. Our results on the xytBxypB-gly43E operon suggest that XypB and Gly43F play essential
functions in this metabolism. Indeed, the XypB putative inner
membrane transporter is required for the production of extracellular xylanase activity, for the induction of the xylan CUT system
by X3, X4 and xylan, and for growth on xylo-oligosaccharides.
These convergent data strongly suggest that this transporter plays
a major role in the transport of xylo-oligosaccharides across the
inner membrane and that this transport is crucial for the induction of the system and for the physiology of Xcc-568. Although
gly43F is also required for normal growth in presence of xylooligosaccharides, a mutation in this gene led to a large increase in
extracellular xylanase activity, unlike what was observed for xypB
mutants. Gly43F is closely related to XynB from Prevotella
bryantii B14 (Table 1). XynB is an intracellular exoxylanase
which was proposed to release xylose progressively from xylo-oligosaccharides, including xylobiose, transported inside the cells
(Gasparic et al., 1995). In Xcc-568, Gly43F is the only protein of
the GH43 family that has no signal peptide, suggesting that it
functions in the cytoplasm. We can speculate that Gly43F
degrades XypB-transported xylo-oligosaccharides to xylose,
thereby promoting bacterial growth. Gly43F may therefore play
a central role in the physiology of Xcc-568 by maintaining a balance between the production of xylose and the maintenance of
xylo-oligosaccharides which induce the CUT system (Fig. 3).
The role of the xytB TBDT gene located upstream from xypB
and gly43F is still elusive. Mutations in this gene have an effect
on growth with xylo-oligosaccharides, but we were unable to
complement these mutations. Our data showed that the xytB,
xypB and gly43F genes form an operon repressed by XylR. However, our growth rate results suggest that the situation is more
complex in the presence of xylo-oligosaccharides. They suggest
the presence of cis-regulatory sequences into xytB driving the
expression of xypB and gly43F. Further work is necessary to fully
characterize this locus.
The xylan CUT system encompasses another TBDT gene,
xytA. This gene is the first gene of a XylR-regulated operon which
comprises two other genes, xyaA and xyaB whose function in
xylan degradation remains unknown. Interestingly, this operon
and the downstream gene are very well conserved with a quartet
of contiguous genes in Cc-CB15 (Fig. 2a, Table S2). The first
two genes of this quartet, including CC0999 TBDT gene, belong
to the xylose regulon. This conservation suggests that this set of
four genes may play an important role in xylan/xylose metabolism in Xcc-568 and Cc-CB15.
The analogy between the Xcc-568 xylan CUT system and the
Cc-CB15 xylose-regulon is not restricted to this locus and 10
other genes of the Xcc-568 xylan/xylose CUT system display significant similarities to proteins of Cc-CB15 (Fig. 2a; Table S2).
Interestingly, six of these genes belong to the xylose regulon
identified in C. cresecentus CB15 (Hottes et al., 2004). This
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conservation includes proteins involved in the removal of substitutions, xylo-oligosaccharides degradation, glucuronate metabolism and TBDTs. With the exception of Xyn10C, which
displays similarities to CC3042, the two other xylanases of Xcc568 are not conserved in C. cresenctus CB15. We identified
another putative xylanase, CC2803, of the GH10 family in the
Cc-CB15 genome which is not conserved in Xcc-568. Like
CC3042, CC2803 was not detected as induced by xylose, but it
is located between two genes induced by this monosaccharide
(Hottes et al., 2004). These observations suggest that Cc-CB15
is able to degrade xylan. Interestingly, the xylose regulon of
Cc-CB15 comprises nine TBDTs, two of which are highly
conserved with XytA and XytB. This high number and the conservation with Xcc-568 TBDTs suggest a crucial role for these
outer membrane transporters in the uptake of molecules during
xylan/xylose catabolism. Recently, sets of genes specifically
induced by xylan have been revealed by transcriptomic studies
on P. bryantii B14 (Pbr) (Dodd et al., 2010b), and Bacteroides
ovatus ATCC8483 (Bov) (Martens et al., 2011), two Bacteroidetes present in the bovine rumen and human gut, respectively.
Interestingly, several genes belonging to Xcc-568 xylan CUT
system display significant similarities with xylan-induced genes
of Pbr and Bov (Fig. 2b, Table S2). Moreover, the xylan regulon
of these latter bacteria share a cluster of conserved genes which
contains two TBDTs belonging to the SusC family. This cluster
is widely conserved among human- and animal-associated
Bacteroides spp. and Prevotella spp. and was proposed to constitute a core set of genes required for xylan fragments uptake by
gut-associated Bacteroidetes (Dodd et al., 2011). Therefore, it
seems that the association between TBDT and xylan utilization
is a common feature shared by bacteria belonging to very different phyla and having apparently different lifestyles. Does this
mean that TBDTs play a very important role in natural conditions? The exact role of these outer membrane transporters has
yet to be determined. However, the involvement of TBDTs
may represent two advantages. First, TBDTs allow the binding
and uptake of larger molecules than porins. Therefore, they
could transport large xylan hydrolysis products thus preventing
release of saccharides in the medium that could be used by
other microorganisms. Second, they allow active transport of
substrate molecules with a very high affinity (Blanvillain et al.,
2007). Therefore, it is possible that TBDTs in these systems
play a crucial role in the transport of xylan breakdown products
when these molecules are present in scarce amounts. This property may be pivotal for oligotrophs such as Caulobacter species
(Hottes et al., 2004). In this study, we showed that XytA and
XytB TBDTs belong to operons that are required for optimal
growth of Xcc-568 on plant leaves. The phyllosphere corresponds to an oligotrophic environment (Lindow & Brandl,
2003) and we can speculate that XytA and XytB may play a
crucial role in the adaptation of Xcc-568 to this niche, which is
an important step for Xcc life cycle. Therefore, it seems that
Xcc-568 and Caulobacter, which belong to different Proteobacteria families, share similar strategies to survive in niches where
nutrients are limited. However, both species most probably possess specific features reflecting their lifestyles. The survival and
Ó 2013 CNRS
New Phytologist Ó 2013 New Phytologist Trust
Research 913
development of Xcc in the phyllosphere may be important to
maintain population sizes sufficient for disease induction. Previous studies on Xoo and Xcv also showed that xylan degradation
play a role in virulence of these strains (Rajeshwari et al., 2005;
Szczesny et al., 2010). Therefore, together with another study
showing the involvement of HrpG and HrpX, the key regulators
of type III secretion system, in the phyllosphere colonization of
Xanthomonas fuscans ssp. fuscans (Darsonval et al., 2008), this
work sheds new light on mechanisms connecting epiphytic colonization to disease induction in plant pathogenic bacteria. It could
also have a significant impact on agro-industrial processes.
Acknowledgements
We thank Annabelle Four-Burgand and Lennart Lessmeier for
technical assistance, Laurent No€el for critical comments on the
manuscript. G.D., S.B-B. and A.B. were funded by the French
Ministry of Research and Technology. We gratefully acknowledge financial support from the Departement Sante des Plantes et
Environnement-Institut National de la Recherche Agronomique
(grant 2007_0441_02) and from the French Agence Nationale
de la Recherche (grant ANR-08-BAN-0193-01). This work is
part of the ‘Laboratoire d’Excellence’ (LABEX) entitled TULIP
(ANR-10-LABX-41).
References
Angot A, Peeters N, Lechner E, Vailleau F, Baud C, Gentzbittel L, Sartorel E,
Genschik P, Boucher C, Genin S. 2006. Ralstonia solanacearum requires Fbox-like domain-containing type III effectors to promote disease on several host
plants. Proceedings of the National Academy of Sciences, USA 103: 14 620–
14 625.
Arlat M, Gough CL, Barber CE, Boucher C, Daniels MJ. 1991. Xanthomonas
campestris contains a cluster of hrp genes related to the larger hrp cluster of
Pseudomonas solanacearum. Molecular Plant–Microbe Interactions 4: 593–601.
Beylot MH, McKie VA, Voragen AG, Doeswijk-Voragen CH, Gilbert HJ.
2001. The Pseudomonas cellulosa glycoside hydrolase family 51
arabinofuranosidase exhibits wide substrate specificity. Biochemical Journal 358:
607–614.
Blanco C, Ritzenthaler P, Kolb A. 1986. The regulatory region of the uxuAB
operon in Escherichia coli K12. Molecular and General Genetics 202: 112–119.
Blanvillain S, Meyer D, Boulanger A, Lautier M, Guynet C, Denance N, Vasse
J, Lauber E, Arlat M. 2007. Plant carbohydrate scavenging through tonBdependent receptors: a feature shared by phytopathogenic and aquatic bacteria.
PLoS ONE 2: e224.
Boulanger A, Dejean G, Lautier M, Glories M, Zischek C, Arlat M, Lauber E.
2010. Identification and regulation of the N-acetylglucosamine utilisation
pathway of the plant pathogenic bacterium Xanthomonas campestris pv.
campestris. Journal of Bacteriology 192: 1487–1497.
Burton RA, Gidley MJ, Fincher GB. 2010. Heterogeneity in the chemistry,
structure and function of plant cell walls. Nature Chemical Biology 6: 724–732.
Buttner D, Bonas U. 2009. Regulation and secretion of Xanthomonas virulence
factors. FEMS Microbiology Reviews 34: 107–133.
Cantarel BL, Coutinho PM, Rancurel C, Bernard T, Lombard V, Henrissat B.
2009. The Carbohydrate-Active EnZymes database (CAZy): an expert resource
for glycogenomics. Nucleic Acids Research 37: D233–D238.
Chow V, Nong G, Preston JF. 2007. Structure, function, and regulation of the
aldouronate utilisation gene cluster from Paenibacillus sp. strain JDR-2. Journal
of Bacteriology 189: 8863–8870.
Collins T, Gerday C, Feller G. 2005. Xylanases, xylanase families and
extremophilic xylanases. FEMS Microbiology Reviews 29: 3–23.
New Phytologist (2013) 198: 899–915
www.newphytologist.com
914 Research
Cornelis P. 2010. Iron uptake and metabolism in pseudomonads. Applied
Microbiology and Biotechnology 86: 1637–1645.
Cunnac S. 2004. Identification a l’e chelle ge nomique des effecteurs de pendant
du syste me de se cre tion de type III de la bacte rie phytopathoge ne Ralstonia
solanacearum. PhD thesis, Universite Toulouse III Paul Sabatier, Toulouse,
France.
Darsonval A, Darrasse A, Meyer D, Demarty M, Durand K, Bureau C,
Manceau C, Jacques MA. 2008. The Type III secretion system of Xanthomonas
fuscans subsp. fuscans is involved in the phyllosphere colonization process and in
transmission to seeds of susceptible beans. Applied and Environment
Microbiology 74: 2669–2678.
DeBoy RT, Mongodin EF, Fouts DE, Tailford LE, Khouri H, Emerson JB,
Mohamoud Y, Watkins K, Henrissat B, Gilbert HJ et al. 2008. Insights into
plant cell wall degradation from the genome sequence of the soil bacterium
Cellvibrio japonicus. Journal of Bacteriology 190: 5455–5463.
Delmotte N, Knief C, Chaffron S, Innerebner G, Roschitzki B, Schlapbach R,
von Mering C, Vorholt JA. 2009. Community proteogenomics reveals insights
into the physiology of phyllosphere bacteria. Proceedings of the National
Academy of Sciences, USA 106: 16 428–16 433.
Dodd D, Cann IK. 2009. Enzymatic deconstruction of xylan for biofuel
production. Global Change Biology Bioenergy 1: 2–17.
Dodd D, Kiyonari S, Mackie RI, Cann IK. 2010a. Functional diversity of four
glycoside hydrolase family 3 enzymes from the rumen bacterium Prevotella
bryantii B14. Journal of Bacteriology 192: 2335–2345.
Dodd D, Mackie RI, Cann IK. 2011. Xylan degradation, a metabolic property
shared by rumen and human colonic Bacteroidetes. Molecular Microbiology 79:
292–304.
Dodd D, Moon YH, Swaminathan K, Mackie RI, Cann IK. 2010b.
Transcriptomic analyses of xylan degradation by Prevotella bryantii and insights
into energy acquisition by xylanolytic bacteroidetes. Journal of Biological
Chemistry 285: 30 261–30 273.
Dombrecht B, Vanderleyden J, Michiels J. 2001. Stable RK2-derived cloning
vectors for the analysis of gene expression and gene function in gram-negative
bacteria. Molecular Plant–Microbe Interactions 14: 426–430.
Dsouza M, Larsen N, Overbeek R. 1997. Searching for patterns in genomic data.
Trends in Genetics 13: 497–498.
Eisenbeis S, Lohmiller S, Valdebenito M, Leicht S, Braun V. 2008. NagAdependent uptake of N-acetyl-glucosamine and N-acetyl-chitin
oligosaccharides across the outer membrane of Caulobacter crescentus. Journal of
Bacteriology 190: 5230–5238.
Emanuelsson O, Brunak S, von Heijne G, Nielsen H. 2007. Locating proteins in
the cell using TargetP, SignalP and related tools. Nature Protocols 2: 953–971.
Finn RD, Mistry J, Tate J, Coggill P, Heger A, Pollington JE, Gavin OL,
Gunasekaran P, Ceric G, Forslund K et al. 2010. The Pfam protein families
database. Nucleic Acids Research 38: D211–D222.
Fontes CM, Gilbert HJ, Hazlewood GP, Clarke JH, Prates JA, McKie VA,
Nagy T, Fernandes TH, Ferreira LM. 2000. A novel Cellvibrio mixtus family
10 xylanase that is both intracellular and expressed under non-inducing
conditions. Microbiology 146: 1959–1967.
Gasparic A, Martin J, Daniel AS, Flint HJ. 1995. A xylan hydrolase gene cluster
in Prevotella ruminicola B(1)4: sequence relationships, synergistic interactions,
and oxygen sensitivity of a novel enzyme with exoxylanase and beta-(1,4)xylosidase activities. Applied and Environment Microbiology 61: 2958–2964.
Gonzalez ET, Allen C. 2003. Characterization of a Ralstonia solanacearum operon
required for polygalacturonate degradation and uptake of galacturonic acid.
Molecular Plant–Microbe Interactions 16: 536–544.
Harhangi HR, Akhmanova AS, Emmens R, van der Drift C, de Laat WT, van
Dijken JP, Jetten MS, Pronk JT, Op den Camp HJ. 2003. Xylose metabolism
in the anaerobic fungus Piromyces sp. strain E2 follows the bacterial pathway.
Archives of Microbiology 180: 134–141.
Hiard S, Maree R, Colson S, Hoskisson PA, Titgemeyer F, van Wezel GP, Joris
B, Wehenkel L, Rigali S. 2007. PREDetector: a new tool to identify regulatory
elements in bacterial genomes. Biochemical and Biophysical Research
Communications 357: 861–864.
Hottes AK, Meewan M, Yang D, Arana N, Romero P, McAdams HH, Stephens
C. 2004. Transcriptional profiling of Caulobacter crescentus during growth on
complex and minimal media. Journal of Bacteriology 186: 1448–1461.
New Phytologist (2013) 198: 899–915
www.newphytologist.com
New
Phytologist
Hurlbert JC, Preston JF III. 2001. Functional characterization of a novel
xylanase from a corn strain of Erwinia chrysanthemi. Journal of Bacteriology 183:
2093–2100.
Keen NT, Boyd C, Henrissat B. 1996. Cloning and characterization of a
xylanase gene from corn strains of Erwinia chrysanthemi. Molecular
Plant–Microbe Interactions 9: 651–657.
Kellett LE, Poole DM, Ferreira LM, Durrant AJ, Hazlewood GP, Gilbert HJ.
1990. Xylanase B and an arabinofuranosidase from Pseudomonas fluorescens
subsp. cellulosa contain identical cellulose-binding domains and are encoded by
adjacent genes. Biochemical Journal 272: 369–376.
Krewulak KD, Vogel HJ. 2011. TonB or not TonB: is that the question?
Biochemistry and Cell Biology 89: 87–97.
Kulkarni N, Shendye A, Rao M. 1999. Molecular and biotechnological aspects of
xylanases. FEMS Microbiology Reviews 23: 411–456.
Larson SB, Day J, Barba de la Rosa AP, Keen NT, McPherson A. 2003. First
crystallographic structure of a xylanase from glycoside hydrolase family 5:
implications for catalysis. Biochemistry 42: 8411–8422.
Lawlis VB, Dennis MS, Chen EY, Smith DH, Henner DJ. 1984. Cloning and
sequencing of the xylose isomerase and xylulose kinase genes of Escherichia coli.
Applied and Environment Microbiology 47: 15–21.
Li J, Wang N. 2011. The wxacO gene of Xanthomonas citri ssp. citri encodes a
protein with a role in lipopolysaccharide biosynthesis, biofilm formation, stress
tolerance and virulence. Molecular Plant Pathology 12: 381–396.
Liang WJ, Wilson KJ, Xie H, Knol J, Suzuki S, Rutherford NG, Henderson PJ,
Jefferson RA. 2005. The gusBC genes of Escherichia coli encode a glucuronide
transport system. Journal of Bacteriology 187: 2377–2385.
Lindow SE, Brandl MT. 2003. Microbiology of the phyllosphere. Applied and
Environment Microbiology 69: 1875–1883.
Liu C, Gilmont RR, Benndorf R, Welsh MJ. 2000. Identification and
characterization of a novel protein from Sertoli cells, PASS1, that associates
with mammalian small stress protein hsp27. Journal of Biological Chemistry
275: 18 724–18 731.
Marchler-Bauer A, Lu S, Anderson JB, Chitsaz F, Derbyshire MK, DeWeeseScott C, Fong JH, Geer LY, Geer RC, Gonzales NR et al. 2011. CDD: a
Conserved Domain Database for the functional annotation of proteins. Nucleic
Acids Research 39: D225–D229.
Martens EC, Lowe EC, Chiang H, Pudlo NA, Wu M, McNulty NP, Abbott
DW, Henrissat B, Gilbert HJ, Bolam DN et al. 2011. Recognition and
degradation of plant cell wall polysaccharides by two human gut symbionts.
PLoS Biology 9: e1001221.
Marx CJ, Lidstrom ME. 2002. Broad-host-range cre-lox system for
antibiotic marker recycling in gram-negative bacteria. BioTechniques 33:
1062–1067.
Meyer D, Lauber E, Roby D, Arlat M, Kroj T. 2005. Optimization of
pathogenicity assays to study the Arabidopsis thaliana–Xanthomonas campestris
pv. campestris pathosystem. Molecular Plant Pathology 6: 327–333.
Mirande C, Mosoni P, Bera-Maillet C, Bernalier-Donadille A, Forano E. 2010.
Characterization of Xyn10A, a highly active xylanase from the human gut
bacterium Bacteroides xylanisolvens XB1A. Applied Microbiology and
Biotechnology 87: 2097–2105.
Morales CQ, Posada J, Macneale E, Franklin D, Rivas I, Bravo M, Minsavage J,
Stall RE, Whalen MC. 2005. Functional analysis of the early chlorosis factor
gene. Molecular Plant–Microbe Interactions 18: 477–486.
Morris CE, Kinkel LL. 2002. Fifty years of phyllosphere microbiology:
significant contributions to research in related fields. In: Lindow SE, HechtPoinar EI, Elliott VJ, eds. Phyllosphere microbiology. St Paul, MN, USA: APS
Press, 365–375.
Neugebauer H, Herrmann C, Kammer W, Schwarz G, Nordheim A, Braun V.
2005. ExbBD-dependent transport of maltodextrins through the novel MalA
protein across the outer membrane of Caulobacter crescentus. Journal of
Bacteriology 187: 8300–8311.
Nurizzo D, Nagy T, Gilbert HJ, Davies GJ. 2002. The structural basis for
catalysis and specificity of the Pseudomonas cellulosa alpha-glucuronidase,
GlcA67A. Structure 10: 547–556.
Oke V, Long SR. 1999. Bacterial genes induced within the nodule during the
Rhizobium–legume symbiosis. Molecular Microbiology 32: 837–849.
Ó 2013 CNRS
New Phytologist Ó 2013 New Phytologist Trust
New
Phytologist
Potnis N, Krasileva K, Chow V, Almeida NF, Patil PB, Ryan RP, Sharlach M,
Behlau F, Dow JM, Momol M et al. 2011. Comparative genomics reveals diversity
among xanthomonads infecting tomato and pepper. BMC Genomics 12: 146.
Rajeshwari R, Jha G, Sonti RV. 2005. Role of an in planta-expressed xylanase of
Xanthomonas oryzae pv. oryzae in promoting virulence on rice. Molecular Plant–
Microbe Interactions 18: 830–837.
Rakus JF, Fedorov AA, Fedorov EV, Glasner ME, Vick JE, Babbitt PC, Almo
SC, Gerlt JA. 2007. Evolution of enzymatic activities in the enolase
superfamily: D-mannonate dehydratase from Novosphingobium
aromaticivorans. Biochemistry 46: 12 896–12 908.
Rigano LA, Siciliano F, Enrique R, Sendin L, Filippone P, Torres PS, Questa J,
Dow JM, Castagnaro AP, Vojnov AA et al. 2007. Biofilm formation, epiphytic
fitness, and canker development in Xanthomonas axonopodis pv. citri. Molecular
Plant–Microbe Interactions 20: 1222–1230.
Ryan RP, Vorholter FJ, Potnis N, Jones JB, Van Sluys MA, Bogdanove AJ, Dow
JM. 2011. Pathogenomics of Xanthomonas: understanding bacterium–plant
interactions. Nature Reviews Microbiology 9: 344–355.
Saha BC. 2003. Hemicellulose bioconversion. Journal of Industrial Microbiology
and Biotechnology 30: 279–291.
Sambrook J, Fritsch EF, Maniatis T. 1989 Molecular cloning: a laboratory
manual, 2nd edn. New York, NY, USA: Cold Spring Harbor Laboratory Press.
Schafer A, Tauch A, Jager W, Kalinowski J, Thierbach G, Puhler A. 1994. Small
mobilizable multi-purpose cloning vectors derived from the Escherichia coli
plasmids pK18 and pK19: selection of defined deletions in the chromosome of
Corynebacterium glutamicum. Gene 145: 69–73.
Schauer K, Rodionov DA, de Reuse H. 2008. New substrates for TonBdependent transport: do we only see the ‘tip of the iceberg’? Trends in
Biochemical Sciences 33: 330–338.
Scheller HV, Ulvskov P. 2010. Hemicelluloses. Annual Review of Plant Biology
61: 263–289.
Shulami S, Gat O, Sonenshein AL, Shoham Y. 1999. The glucuronic acid
utilisation gene cluster from Bacillus stearothermophilus T-6. Journal of
Bacteriology 181: 3695–3704.
da Silva ACR, Ferro JA, Reinach FC, Farah CS, Furlan LR, Quaggio RB,
Monteiro-Vitorello CB, Sluys MAV, Almeida NF, Alves LMC et al. 2002.
Comparison of the genomes of two Xanthomonas pathogens with differing host
specificities. Nature 417: 459–463.
St John FJ, Hurlbert JC, Rice JD, Preston JF, Pozharski E. 2011. Ligand bound
structures of a glycosyl hydrolase family 30 glucuronoxylan xylanohydrolase.
Journal of Molecular Biology 407: 92–109.
Stephens C, Christen B, Fuchs T, Sundaram V, Watanabe K, Jenal U. 2007a.
Genetic analysis of a novel pathway for D-xylose metabolism in Caulobacter
crescentus. Journal of Bacteriology 189: 2181–2185.
Stephens C, Christen B, Watanabe K, Fuchs T, Jenal U. 2007b. Regulation of
D-xylose metabolism in Caulobacter crescentus by a LacI-type repressor. Journal
of Bacteriology 189: 8828–8834.
Stoddart A, Zhang Y, Paige CJ. 1996. Molecular cloning of the cDNA encoding a
murine sialic acid-specific 9-O-acetylesterase and RNA expression in cells of
hematopoietic and non-hematopoietic origin. Nucleic Acids Research 24: 4003–4008.
Szczesny R, Jordan M, Schramm C, Schulz S, Cogez V, Bonas U, Buttner D.
2010. Functional characterization of the Xcs and Xps type II secretion systems
from the plant pathogenic bacterium Xanthomonas campestris pv vesicatoria.
New Phytologist 187: 983–1002.
Utt EA, Eddy CK, Keshav KF, Ingram LO. 1991. Sequencing and expression of
the Butyrivibrio fibrisolvens xylB gene encoding a novel bifunctional protein
with beta-D-xylosidase and alpha-L-arabinofuranosidase activities. Applied and
Environment Microbiology 57: 1227–1234.
Vorholt JA. 2012. Microbial life in the phyllosphere. Nature Reviews Microbiology
10: 828–840.
Whipps JM, Hand P, Pink D, Bending GD. 2008. Phyllosphere microbiology
with special reference to diversity and plant genotype. Journal of Applied
Microbiology 105: 1744–1755.
Whitehead TR. 1995. Nucleotide sequences of xylan-inducible xylanase and
xylosidase/arabinosidase genes from Bacteroides ovatus V975. Biochimica et
Biophysica Acta 1244: 239–241.
Ó 2013 CNRS
New Phytologist Ó 2013 New Phytologist Trust
Research 915
Zehner S, Kotzsch A, Bister B, Sussmuth RD, Mendez C, Salas JA, van Pee KH.
2005. A regioselective tryptophan 5-halogenase is involved in pyrroindomycin
biosynthesis in Streptomyces rugosporus LL-42D005. Chemistry & Biology 12:
445–452.
Zhang CC, Durand MC, Jeanjean R, Joset F. 1989. Molecular and genetical
analysis of the fructose-glucose transport system in the cyanobacterium
Synechocystis PCC6803. Molecular Microbiology 3: 1221–1229.
Supporting Information
Additional supporting information may be found in the online
version of this article.
Fig. S1 Mutations and plasmids constructed in the xylE (a), xytA
(b), xylR (c) and xytB (d) loci.
Fig. S2 Operons mapping in the xytA and xytB loci in the xylR
mutant.
Fig. S3 Xylose induction motif of Caulobacter crescentus genes
and the xyl-box motif of Xanthomonas campestris pv campestris
ATCC33913 (Xcc-568). Sequence logos were generated by
WebLogo (http://weblogo.berkeley.edu/; Crooks GE, Hon G,
Chandonia JM, Brenner SE. 2004. WebLogo: a sequence logo
generator. Genome Research 14: 1188–1190).
Fig. S4 Effect of pVO155 insertion into the xylR regulatory gene
on expression of xylE, xylR, agu67A, axeXA, uxuB and xytB.
Fig. S5 Effect of pVO155 insertion into the agu67A gene on
expression of xylR, agu67A, axeXA, uxuB, xyn10 and xytB.
Fig. S6 Analysis of in vitro secretion of Xyn10A, Xyn10C and
Xyn30A putative xylanases.
Table S1 List of plasmids and Xanthomonas campestris pv
campestris strains used or generated in this study
Table S2 Conservation of Xanthomonas campestris pv campestris
ATCC33913 (LMG568) proteins encoded by genes induced by
xylan, xylooligosaccharides or xylose, in Caulobacter crescentus
CB15, Prevotella bryantii B14 or Bacteroides ovatus ATCC8483
proteomes
Table S3 Occurrence of perfect xyl-box motif upstream from
Xanthomonas campestris pv campestris ATCC33913 genes
Table S4 Enzymes active on xylan
Method S1 Secretion assays of Xyn10A, Xyn10C and Xyn30A
putative xylanases.
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