Research Article

Plant Carbohydrate Scavenging through TonB-Dependent Receptors: A Feature Shared by Phytopathogenic and Aquatic Bacteria

  • Servane Blanvillain equal contributor,

    equal contributor Contributed equally to this work with: Servane Blanvillain, Damien Meyer, Emmanuelle Lauber, Matthieu Arlat

    Affiliation: Laboratoire des Interactions Plantes-Microorganismes, Centre National de la Recherche Scientifique (CNRS)/Institut National de la Recherche Agronomique (INRA) UMR2594/441, Castanet-Tolosan, France

  • Damien Meyer equal contributor,

    equal contributor Contributed equally to this work with: Servane Blanvillain, Damien Meyer, Emmanuelle Lauber, Matthieu Arlat

    Affiliation: Laboratoire des Interactions Plantes-Microorganismes, Centre National de la Recherche Scientifique (CNRS)/Institut National de la Recherche Agronomique (INRA) UMR2594/441, Castanet-Tolosan, France

    Current address: UMR PaVe, Centre INRA, Beaucouze, France

  • Alice Boulanger,

    Affiliation: Laboratoire des Interactions Plantes-Microorganismes, Centre National de la Recherche Scientifique (CNRS)/Institut National de la Recherche Agronomique (INRA) UMR2594/441, Castanet-Tolosan, France

  • Martine Lautier,

    Affiliations: Laboratoire des Interactions Plantes-Microorganismes, Centre National de la Recherche Scientifique (CNRS)/Institut National de la Recherche Agronomique (INRA) UMR2594/441, Castanet-Tolosan, France, Université Paul Sabatier, Toulouse III, Toulouse, France

  • Catherine Guynet,

    Affiliation: Laboratoire des Interactions Plantes-Microorganismes, Centre National de la Recherche Scientifique (CNRS)/Institut National de la Recherche Agronomique (INRA) UMR2594/441, Castanet-Tolosan, France

    Current address: Laboratoire de Microbiologie et Génétique Moléculaires, Centre National de la Recherche Scientifique (CNRS) UMR5100, Toulouse, France

  • Nicolas Denancé,

    Affiliation: Laboratoire des Interactions Plantes-Microorganismes, Centre National de la Recherche Scientifique (CNRS)/Institut National de la Recherche Agronomique (INRA) UMR2594/441, Castanet-Tolosan, France

    Current address: Signalisation chez les Végétaux, Pole de Biotechnologie Végétale, Centre National de la Recherche Scientifique (CNRS) UMR5546, Castanet-Tolosan, France

  • Jacques Vasse,

    Affiliation: Laboratoire des Interactions Plantes-Microorganismes, Centre National de la Recherche Scientifique (CNRS)/Institut National de la Recherche Agronomique (INRA) UMR2594/441, Castanet-Tolosan, France

  • Emmanuelle Lauber equal contributor mail,

    equal contributor Contributed equally to this work with: Servane Blanvillain, Damien Meyer, Emmanuelle Lauber, Matthieu Arlat

    To whom correspondence should be addressed. E-mail: (EL); (MA)

    Affiliation: Laboratoire des Interactions Plantes-Microorganismes, Centre National de la Recherche Scientifique (CNRS)/Institut National de la Recherche Agronomique (INRA) UMR2594/441, Castanet-Tolosan, France

  • Matthieu Arlat equal contributor mail

    equal contributor Contributed equally to this work with: Servane Blanvillain, Damien Meyer, Emmanuelle Lauber, Matthieu Arlat

    To whom correspondence should be addressed. E-mail: (EL); (MA)

    Affiliations: Laboratoire des Interactions Plantes-Microorganismes, Centre National de la Recherche Scientifique (CNRS)/Institut National de la Recherche Agronomique (INRA) UMR2594/441, Castanet-Tolosan, France, Université Paul Sabatier, Toulouse III, Toulouse, France

  • Published: February 21, 2007
  • DOI: 10.1371/journal.pone.0000224


TonB-dependent receptors (TBDRs) are outer membrane proteins mainly known for the active transport of iron siderophore complexes in Gram-negative bacteria. Analysis of the genome of the phytopathogenic bacterium Xanthomonas campestris pv. campestris (Xcc), predicts 72 TBDRs. Such an overrepresentation is common in Xanthomonas species but is limited to only a small number of bacteria. Here, we show that one Xcc TBDR transports sucrose with a very high affinity, suggesting that it might be a sucrose scavenger. This TBDR acts with an inner membrane transporter, an amylosucrase and a regulator to utilize sucrose, thus defining a new type of carbohydrate utilization locus, named CUT locus, involving a TBDR for the transport of substrate across the outer membrane. This sucrose CUT locus is required for full pathogenicity on Arabidopsis, showing its importance for the adaptation to host plants. A systematic analysis of Xcc TBDR genes and a genome context survey suggested that several Xcc TBDRs belong to other CUT loci involved in the utilization of various plant carbohydrates. Interestingly, several Xcc TBDRs and CUT loci are conserved in aquatic bacteria such as Caulobacter crescentus, Colwellia psychrerythraea, Saccharophagus degradans, Shewanella spp., Sphingomonas spp. or Pseudoalteromonas spp., which share the ability to degrade a wide variety of complex carbohydrates and display TBDR overrepresentation. We therefore propose that TBDR overrepresentation and the presence of CUT loci designate the ability to scavenge carbohydrates. Thus CUT loci, which seem to participate to the adaptation of phytopathogenic bacteria to their host plants, might also play a very important role in the biogeochemical cycling of plant-derived nutrients in marine environments. Moreover, the TBDRs and CUT loci identified in this study are clearly different from those characterized in the human gut symbiont Bacteroides thetaiotaomicron, which allow glycan foraging, suggesting a convergent evolution of TBDRs in Proteobacteria and Bacteroidetes.


Bacteria are able to colonize a wide variety of habitats, including the most extreme environments and even living organisms. This remarkable feature likely reflects a high degree of adaptability and the presence of specific genetic programs devoted to the exploitation of nutrients present in these diverse habitats. The bacterial ability to use defined carbohydrates to support cell survival or growth implies the availability of these substrates in their habitat. Moreover, it requires the recognition of these molecules and the coordinated induction of particular uptake systems and of metabolic enzymes [1], [2]. Characterization of the repertoire of genes involved in signal perception or transduction and in carbohydrate utilization, in conjunction with analysis of the regulation of their expression, is likely to provide key information about the interaction and adaptation of bacteria with their environment [3].

The analysis and comparison of complete genomic sequences of numerous bacteria from diverse phylogenies and habitats have shown a relationship between the ecological niches that a bacterium occupies and the proportion of genes involved in signal perception and transduction. Bacteria that inhabit stable environments (extremophiles, obligate parasites or symbionts), which generally have small genomes, possess fewer sensory and regulatory genes than free-living bacteria found in complex and changing environments such as those living in soil or in association with plants [2]. Therefore, it was proposed that sensors and regulators can be used as “descriptors of bacterial lifestyle” [2].

With the aim to study the molecular mechanisms controlling adaptation of phytopathogenic bacteria to their host-plants, we undertook a global analysis of receptors and regulators of Xanthomonas campestris pv. campestris (Xcc), the causal agent of black rot of crucifers. This pathogen infects a wide range of Brassicaceae plants of economic interest, including cabbage, cauliflower and radish as well as the model plant Arabidopsis thaliana. This epiphytic bacterium naturally infects host plants via wounds in the leaves or hydathodes which are specialized pores on the leaf margins of higher plants that connect to the vascular system. Then bacteria multiply and progress in vascular tissues [4], [5]. During the past two decades, classical molecular and genetic studies led to the characterization of several determinants controlling pathogenicity of Xcc, such as secretion of extracellular plant cell wall degrading enzymes [6][8], cell-cell signaling [9], [10], biofilm formation [11] and hrp genes coding for a type III secretion system [12], [13]. The characterization of new virulence factors should now be greatly facilitated by the availability of complete genome sequences of two Xcc strains (strain ATCC33913 [14] and strain 8004 [15]). Moreover, comparative genomics would improve the analysis of virulence and host adaptation in Xanthomonas, since the genomic sequences of four other Xanthomonas strains displaying different host specificities and representing three other species are also available, i.e. Xanthomonas axonopodis pv. citri (Xac), the causal agent of citrus canker [14], Xanthomonas campestris pv. vesicatoria (Xcv), the causative agent of bacterial spot disease on pepper and tomato plants [16], and Xanthomonas oryzae pv. oryzae (Xoo) (strain Kacc10331 [17] and strain MAFF311018 [18]), the causal agent of bacterial blight of rice. Thus, the Xanthomonas genus, which affects two major model plants (Arabidopsis and rice), constitutes a very attractive model to study plant-pathogen interactions.

Our analysis of the Xcc (ATCC33913) genome revealed an overrepresentation of a particular family of receptors, named TonB-dependent receptors (TBDRs). These proteins are located in the outer membrane of Gram-negative bacteria and are mainly known to transport iron-siderophore complexes and vitamin B12 into the periplasm [19]. In most cases, the expression of the genes encoding these receptors is under the control of the Fur (Ferric uptake regulator) repressor and activated under conditions of iron starvation [20]. In contrast to diffusion through porins, transport via TBDRs requires energy which is provided by the inner membrane energy-coupling TonB-ExbB-ExbD protein complex [21].

The exploration of complete genome sequences of 226 Gram negative bacteria showed that the overrepresentation of TBDRs is restricted to a small proportion of these bacteria, but is a common trait of all sequenced Xanthomonas species. Interestingly, most of the bacteria displaying this particularity have diverse lifestyles and belong to different taxonomical lineages, but they all share the ability to exploit complex carbohydrates. Therefore, we postulated that some Xcc TBDRs might be involved in the transport of plant-derived molecules. This hypothesis was recently reinforced with the characterization of a TBDR, named MalA, in the oligotrophic aquatic bacterium Caulobacter crescentus, which transports maltodextrins [22].

A systematic study of Xcc TBDRs, based on mutagenesis, expression analyses and transport assays identified one Xcc TBDR involved in the transport of sucrose, a major plant sugar. This TBDR gene is required for full virulence on Arabidopsis and is associated with genes required for sucrose metabolism, thus forming a “sucrose utilization locus”. Our study also suggests the existence of other TBDR-containing loci involved in the utilization of plant cell wall compounds which are conserved in a wide range of bacteria displaying TBDR overrepresentation.


Identification of Xcc TBDRs

TBDR proteins consist of two domains, with a C-terminal membrane embedded β-barrel domain that is sealed by the N-terminal plug domain [23][27]. Hidden Markov models (HMMs), PF00593 and PF07715, corresponding to these two domains, are available in the Pfam database [28]. Seventy-six proteins carrying one or both domains were detected in the proteome of the Xcc ATCC33913 strain (Table S1). Among these proteins, 64 possess both domains. The remaining 12 proteins, which possess only one of the two domains, can be divided into three groups: 3 proteins (XCC2208, XCC4131 and XCC4132) which do not seem to have a canonical plug domain, one protein (XCC2497) which has no domain detected by the PF00593 HMM, and finally 8 proteins (XCC0304-0305, XCC1750-1751, XCC3215-3216 and XCC3270-3271) which are truncated and correspond to 4 pseudogenes having a frame-shift in their coding region (Table S1). Each of these 4 pseudogenes was thereafter studied as a unique entity.

Other conserved features of TBDRs were then used to further characterize these identified proteins. In the N-terminal part of the plug domain, TBDRs display a conserved sequence, the TonB-box, that interacts with the TonB protein [26], [27], [29]. A TonB-box, with the Xcc consensus sequence tLDXVXV (lower case indicates less highly conserved amino acid, X indicates any amino acid) was detected in all studied proteins (Table S1). The localization of TBDRs in the outer membrane implies their transport across the inner membrane and thus the presence of a signal peptide at their N-terminal end. The proteins were therefore analyzed for the presence of such a motif. TBDRs which were not predicted to possess a signal peptide in the annotation proposed by da Silva and colleagues [14] were manually re-annotated, thus revealing putative signal peptides (Table S1). Out of 72 proteins, 1 TBDR (XCC2385) does not have a signal peptide. Finally, the 12 C-terminal amino acids of outer membrane proteins (OMPs) form a membrane anchoring β-sheet with the last residue being aromatic (F in the vast majority of OMPs) [30], [31]. The 12 C-terminal amino acid of 55 of the proteins studied here are predicted to form a potential β-sheet, ending with an aromatic residue (Table S1). Among the TBDRs which did not display a typical C-terminal sequence, 8 have a β-sheet ending with an aromatic amino acid followed by a short extension of 1 to 4 amino acids (Table S1).

Taking all this information together, 48 proteins possess all typical features of TBDRs in Xcc ATCC33913 and were thus considered as putative functional TBDRs. The other proteins lacking or having degenerated TBDR domains were also included in our study and then named Ps-TBDRs (for Pseudo-TBDRs) (Table S1).

Finally, some TBDRs contain a N-terminal extension, located between the signal peptide and the plug domain [32]. Two types of N-terminal extension have been identified: the transducer and the Oar-like extensions [33]. In Xcc, among the 9 TBDRs having an N-terminal extension, one is a member of the TBDR-transducer subclass, seven are in the Oar-like subclass and one displays a N-terminal extension that does not correspond to the two types already identified (Table S1). Among these TBDRs, only two have all canonical features of TBDRs.

TBDRs are overrepresented in Xanthomonas spp. and in bacteria scavenging complex carbohydrates

A survey of TBDRs was performed in 226 eubacterial completely sequenced genomes, by identifying proteins detected by both PF07715 and PF00593 HMMs. This analysis showed that most bacteria (71%) hold less than 14 TBDRs per proteome, whereas the remaining bacteria have a very broad variation in TBDR number, ranging from 14 to 120 (Figure 1 and Table S2). In fact, 27% of the bacteria studied here have no TBDR proteins and 43% possess between 1 and 13 TBDRs. Only a very small proportion of bacteria (15.5%) possesses more than 30 TBDRs, thus forming a particular class in which TBDRs seem to be over represented. It is worth noting that most bacteria in this class either belong to the α or γ-Proteobacteria classes or to the Bacteroides genus (Figure 1 and Table S2). The ratios between the number of TBDRs found in each proteome and the genome size or the number of annotated coding sequences (CDS) displayed a very similar distribution pattern (Table S2), thus showing that there was no major bias due to annotation. Therefore, the TBDR/Mbp ratio was used to define several bacterial classes: bacteria displaying a ratio higher than 5 were considered as members of a class showing TBDR overrepresentation, while those having a ratio ranging from 3 to 5 belong to an intermediary class. The 4 Xanthomonas species whose genomes have been sequenced belong to the class displaying TBDR overrepresentation (Table S2). It is worth noting that phytopathogenic bacteria, such as Pseudomonas syringae pathovars or Erwinia carotovora subsp. Atroseptica belong to the intermediary class.


Figure 1.

Occurrence of TonB-dependent receptor (TBDR) genes in 226 completely sequenced Gram negative bacterial genomes, in ecological (A) or phylogenetic (B) contexts. TBDRs were detected by screening the Pfam database and the UniprotKB protein knowledge database using the two Pfam HMMs (PF07715, Plug; PF00593, TonB_dep_Rec). Only proteins displaying the two Pfam domains were considered. The habitat of bacteria (A) and their phylogeny (B) are represented by colour codes (see the legend boxes).


Seven Xcc TBDR and two Ps-TBDR genes are Fur-regulated

As TBDRs are mainly known to be involved in iron uptake, it was important to determine how many of these receptors are assigned to this function in Xcc. In most cases, the expression of genes encoding TBDRs involved in iron transport is regulated by the iron status in the medium. These genes are activated under iron depletion conditions and repressed under iron repletion conditions via the Fur repressor (Ferric-uptake regulator), that binds to a specific DNA sequence element called the “Fur-box”, which is found in the target promoters of iron-regulated genes [20], [34], [35]. These features led us to analyze the regulation by the iron status for all identified Xcc TBDR and Ps-TBDR genes.

The 76 Xcc TBDR and Ps-TBDR genes were mutated by insertion of the suicide plasmid pVO155 [36], leading to transcriptional fusion with the promoterless uidA reporter gene. Two insertions (in XCC1990 and XCC3209) were found unstable. Thus, we constructed deletion-mutations in these genes using the cre-lox system [37], [38]. In order to analyze the expression of these 2 deleted TBDR genes, their promoter regions were cloned upstream of a promoterless lacZ gene in a reporter plasmid (see Materials and Methods), and these constructions were introduced into the wild-type strain. Using β-glucuronidase or β-galactosidase expression assays, we monitored the expression of all Xcc TBDRs and Ps-TBDRs in iron-repleted and -depleted media. Seven genes (XCC0158, XCC0768, XCC1391, XCC2772, XCC3050, XCC3518, and XCC4162) and 2 Ps-TBDR genes (XCC3216-3215 and XCC3595) are induced by iron starvation, compared to iron-replete conditions (Figure 2A). With the exception of Ps-TBDR XCC3216-3215, this regulation pattern was confirmed by quantitative real-time reverse transcriptase polymerase chain reaction (qRT-PCR) in a wild-type background (data not shown). We then checked whether these genes are under the control of the Fur regulator. XCC1470, the orthologue of the Fur regulator identified in Xoo [39] and in Xanthomonas campestris pv. phaseoli (Xap) [40], was mutated using the manganese method (see Materials and Methods). A point mutant in this latter gene was obtained (fur1). As expected for a fur mutation, this mutant produced more siderophores than the wild-type strain and this production remained unaffected in response to increased iron levels (from 1 to 50 µM), as determined by using the chrome azurol S assay (data not shown). It is worth noting that the fur mutant is unable to induce any disease symptoms (data not shown), as already reported in Xoo [39].


Figure 2.

Expression levels of 9 Xanthomonas campestris pv. campestris TonB-dependent receptor genes. (A) β-glucuronidase assays, performed in at least two independent experiments with pVO155 insertion mutant strains, cultivated in minimal medium supplemented with iron (100 µM FeSO4) or containing an iron chelator (100 µM 2,2′-dipyridyl; DPD). (B) Real-time quantitative RT-PCR (qRT-PCR), performed in at least three independent experiments, on RNA extracted from the wild-type strain, a fur mutant strain (fur1) or from a complemented strain [fur1 (pL-fur)], cultivated in minimal medium containing 100 µM FeSO4. Calculation of relative expression includes normalization against the endogenous control gene 16S RNA.


We then compared by qRT-PCR analysis the relative expression of the TBDR and Ps-TBDR genes induced by iron starvation, in the wild-type strain versus a fur mutant, in an iron-containing medium. As presented in Figure 2B, the expression of all these genes was repressed by the Fur protein. However, the repression level of the 2 Ps-TBDR genes was low (less than twofold). We confirmed that the deregulation phenotype of the fur mutant was the result of mutation in the fur gene by complementation experiments: the repression by iron of the 7 TBDR and the 2 Ps-TBDR genes was restored by providing the wild-type Fur protein on the pL-XCC1470 plasmid (see Materials and Methods) in the fur mutant (Figure 2B).

The 9 Fur-regulated TBDR/Ps-TBDR genes, but not the other Xcc TBDR genes, possess a putative Fur-box

The DNA regions located upstream of the 9 Fur-repressed TBDR and Ps-TBDR genes were analyzed using the MotifSampler program [41], [42] to identify putative Fur-boxes. For genes arranged in putative operons (XCC3050 and XCC0768), the DNA region located upstream of the first gene of the putative operon was also studied. XCC3595, which belongs to the TBDR transducer subclass, is preceded by two genes displaying significant similarities with the fecI (XCC3593) and fecR (XCC3594) genes, coding for a sigma factor of the ECF subfamily and its anti-sigma factor, respectively. In most cases, the FecI/FecR system is associated with a TBDR of the transducer subclass and is involved in iron signaling and transport by regulating the expression of the TBDR gene and other “iron-associated” genes (for review see [33], [43]). We thus also analyzed the DNA regions upstream of these two genes. This analysis allowed the identification of a single significant 19-bp palindromic motif (9-1-9 inverted repeat) in the upstream region of most genes or operons studied here (Figure 3A, Table 1). For XCC3595, the palindromic motif was detected in the upstream region of XCC3593 (fecI). For XCC0768, two motifs were identified: one in the upstream region of XCC0767 and one in the coding region of this gene (Table 1). This motif (AATGAgAATcATTctCATT; lower case indicates less highly conserved) only showed a weak similarity with the E. coli 19 bp Fur-box consensus sequence (GATAATGATAATCATTATC) [44]: it matched this sequence in 10 bp positions out of 19 (52% identity). However, the conservation was significantly higher with Fur-box consensus sequences identified in Shewanella oneidensis (AATGATAATAATTATCATT; 84% identity) [45] and Bacillus subtilis (aaTGAtAATnATTaTCAtt; 84% identity) [46]. Interestingly, a multiple alignment of these 4 Fur-box consensus sequences showed that although the E. coli consensus sequence seems more divergent, it perfectly matches the 3 other sequences after a 3 bp shift (Figure 3B and C). These results suggest that the palindromic motif identified in this study might correspond to a possible Xcc Fur-box. We then explored DNA sequences located upstream of the other Xcc TBDR or Ps-TBDR genes as well as the entire Xcc genome sequence to detect putative Fur-boxes, using the MotifScanner program [42] or the PatScan pattern matcher software [47]. No putative Fur-box was detected in the promoter region of any other TBDR gene. These analyses identified more than 20 genes having candidate Fur-boxes in the region from −300 to +20 relative to the start of translation (data not shown). Several of these genes are orthologs or homologs of genes regulated by Fur in other bacteria, such as feoA (XCC1834) [48] and bfd (XCC0481) [49], or involved in iron storage such as piuB (XCC3955) [50], thus reinforcing the validity of our Fur-box consensus sequence. Altogether, this analysis suggests that only 7 TBDR and 2 Ps-TBDR are directly associated with iron uptake in Xcc. The role of the other TBDRs remained to be deciphered.


Figure 3.

Palindromic consensus Fur-box sequence identified upstream of 9 Xanthomonas campestris pv. campestris Fur regulated TonB-dependent receptor genes. (A) Sequence logos generated by WebLogo (, [123]) of the Xcc Fur-box motif as predicted by the MotifSampler program [41], [42]. (B and C) Multiple alignement generated with Multalin program [116] with previously proposed Fur-box consensus sequences of Bacillus subtilis [46], Shewanella oneidensis [45], X. campestris pv. campestris and Escherichia coli [44] (B) and after a 3 bp shift of the E. coli sequence (C).


Table 1. Candidate Fur-boxes identified in the promoter regions of Xanthomonas campestris pv. campestris Fur regulated TonB-dependent receptor (TBDR) genes, TBDR containing operons and the XCC3593 FecI sigma factor gene.


Two TBDR genes belong to the hrp regulon of Xcc

During the study of TBDR gene promoters, we identified a pip/hrpII box in the promoter of 2 TBDR genes: XCC1041 and XCC1719. In Xanthomonas, hrp genes coding for a TTSS, as well as other functions related to pathogenicity are positively regulated by hrpG/hrpX regulatory genes [51]. HrpG regulates the expression of hrpX, which controls the expression of genes possessing a pip-box (TTCGCN15TTCGC) or a hrpII-box (TTCGN16TTCG) [52], [53]. By qRT-PCR experiments, we showed that XCC1041 and XCC1719 TBDR genes are activated by HrpG and HrpX (Figure 4). Interestingly, a mutant in the XCC1719 TBDR gene is weakly affected in pathogenicity (data not shown, see below). This gene seems to be specific to Xcc as it is absent in the genomes of Xcv, Xac, Xoo and Xf, although its surrounding genes are conserved in syntenic regions in all these strains (Table S3). On the other hand, XCC1041 is present in all Xanthomonas species (but absent in Xf) and encodes a TBDR with an atypical N-terminal extension.


Figure 4.

Regulation of Xanthomonas campestris pv. campestris TonB-dependent receptor genes XCC1041 and XCC1719 by hrp regulators. The relative expression was analyzed by real-time quantitaive RT-PCR (qRT-PCR), perfomed on RNA extracted from the wild-type strain and hrpG or hrpX insertion mutants (strains XP082 and XP083, respectively). Experiments were repeated at least three times. Calculation of relative expression includes normalization against the endogenous control gene 16S RNA.


Xcc TBDRs and plant interactions: XCC3358 plays a major role in pathogenicity

In order to assess whether Xcc TBDRs control the interaction with plants, we tested the 76 TBDR mutants constructed in this study for pathogenicity on the Arabidopsis thaliana Sf-2 ecotype. Among these mutants, 17 were altered in symptom development (data not shown). However, the alteration was weak and was not reproducible in all the tests we performed, suggesting either that their involvement is not crucial or that functional redundancy mask their individual contribution. However, the XCC3358 insertion mutant was clearly and reproducibly altered in pathogenicity, showing a clear delay in symptom development in comparison to the wild-type strain (Figure 5A). The XCC3358 gene is likely to form an operon with the downstream gene, XCC3359 (Figure 6). The ortholog of the XCC3359 gene, named suh, has been characterized in Xanthomonas axonopodis pv. glycines (Xag), the causal agent of bacterial pustule disease on soybean. This gene codes for a sucrose hydrolase, SUH, which plays a major role in sucrose metabolism in Xag. A mutant in this gene is moderately affected in pathogenicity on soybean [54]. This prompted us to construct a non polar deletion-mutation in XCC3358 (XCC3358Δ1, see Materials and Methods and Figure 6). A deletion-mutant was also generated into XCC3359 (XCC3359Δ1, see Materials and Methods and Figure 6). These two mutants showed an altered phenotype, i.e. delayed symptom development, similar to that observed with the pVO155 insertion mutant in XCC3358 (Figure 5B). The non polar mutation in XCC3358 was complemented by introducing the plasmid pL-XCC3358, which carries a functional XCC3358 gene, into XCC3358Δ1 mutant (Figure 5B). This result confirms that the XCC3358 TBDR plays a role in virulence. Similarly, the deleted mutant in XCC3359 was complemented by a plasmid carrying this gene (Figure 5B).


Figure 5.

Quantitative analysis of the interaction between Xanthomonas campestris pv. campestris (Xcc) and Arabidopsis thaliana Sf-2 plants. (A) Pathogenicity tests with the Xcc wild-type strain and the XCC3358 insertion mutant (XCC3358::pVO). (B) Pathogenicity and complementation tests with the Xcc wild-type strain, the XCC3356 and XCC3357 insertion mutants (XCC3356::pVO and XCC3357::pVO, respectively), the XCC3358 and XCC3359 deleted mutants (XCC3358D1 and XCC3359D1, respectively) and their corresponding complemented strains [XCC3358D1 (pL-XCC3358) and XCC3359D1 (pL-XCC3359), respectively]. Disease symptoms were scored 5 to 8 days after inoculation. Each inoculated leaf was individually scored as: no symptom = 0; weak chlorosis surrounding the wound sites = 1; strong V-shaped chlorosis = 2; developing necrosis = 3; leaf death = 4. The represented average disease scores and the standard deviations were calculated from the values of four plants with four inoculated leaves per plant.


Figure 6.

Genetic organization of the Xanthomonas campestris pv. campestris sux locus. Location of mutations are indicated above the map: arrowheads indicate pVO155 insertions and deleted sequences are represented by horizontal dotted bars. Regions cloned in plasmids are indicated below the map: horizontal thick black bars indicate sequences used for plasmid constructions.


XCC3358 TBDR belongs to a locus required for full pathogenicity

In Xcc genome, XCC3358 and XCC3359 are preceded by two other genes that might be related to sugar metabolism: XCC3356 codes for a putative transcriptional repressor of the LacI/GalR-family and XCC3357 encodes a putative sugar transporter of the major facilitator family, allowing the transport of substrate molecules through the inner membrane. We constructed insertion mutants in these two genes, presuming that they are monocistronic (Figure 6). These insertions lead to transcriptional fusions with the promoterless uidA reporter gene. The insertion mutant in XCC3357 showed an altered phenotype on the A. thaliana Sf-2 ecotype, resembling that using XCC3358 or XCC3359 mutants, whereas the insertion mutant in XCC3356 was not affected in pathogenicity (Figure 5B). These results suggest that XCC3357, XCC3358 and XCC3359 are required in the same process related to sucrose metabolism and/or transport. The phenotype of a mutant in XCC3356 was not surprising since this gene encodes a putative repressor, which might negatively control the expression of the other genes of the locus. A mutation in this gene should induce the constitutive expression of the other genes without affecting their function (see below).

The XCC3356-3359 locus is required for sucrose utilization

Insertion mutants in XCC3357, XCC3358 and XCC3359 and the deletion mutant XCC3359Δ1 grew like the wild-type strain on minimal medium containing glucose or fructose (data not shown), but were all affected in growth on sucrose (20 mM) (Figure 7A). The growth of the insertion mutant in XCC3356 and the XCC3358Δ1 mutant was not impaired in minimal medium containing sucrose. The fact that the insertion mutant in XCC3358 was impaired in growth on sucrose whereas the deletion mutant in the same gene was not, suggested that the phenotype of the insertion mutant was due to a polar effect of the insertion on XCC3359 gene expression. This hypothesis was confirmed by complementation experiments. The growth of insertion mutants in XCC3358 and XCC3359 on sucrose media was restored when XCC3359 was supplied in trans on the expression plasmid pL-XCC3359, which allows constitutive expression of the XCC3359 gene. No complementation was observed when the XCC3358 gene was supplied in trans in the XCC3358 insertion mutant (Figure 7B). These results confirmed that XCC3358 and XCC3359 form an operon. They also pointed out that this putative operon and the entire locus is important for sucrose utilization in Xcc. Thus, we renamed this locus sux, for sucrose utilization in Xanthomonas. For the purpose of this study, the TBDR-amylosucrase operon was renamed suxAB, the putative sugar transporter gene, suxC, and the putative regulatory gene, suxR (see Figure 6). However, the role of the TBDR SuxA in sucrose utilization remained elusive: this transporter gene was not required for growth on sucrose, but a non polar mutant in this gene was altered in pathogenicity, like suxB and suxC insertion mutants. This observation suggested that this suxA gene might have a particular function.


Figure 7.

Growth of Xanthomonas campestris pv. campestris wild-type strain and mutants in minimal medium containing 20 mM sucrose. (A) Bacterial growth of insertion mutants XCC3356::pVO to XCC3359::pVO or deletion mutants XCC3358Δ1 and XCC3359Δ1, compared to the wild-type strain. (B) Growth of strains carrying plamids pL-XCC3358 or pL-XCC3359 allowing the constitutive expression of XCC3358 and XCC3359 genes, respectively. Bars represent standard deviations from 3 independent experiments.


SuxA and SuxC allow sucrose uptake

To clarify the role of the SuxA TBDR, we investigated its involvement in sucrose transport, by performing sucrose uptake experiments using [14C]sucrose.

First, the uptake rates into the Xcc wild-type strain were compared after an overnight preculture in the presence (induced) or absence (uninduced) of sucrose. As shown in figure 8A, induced cells took-up [14C]sucrose quicker than uninduced cells, but the values obtained after 2 hours of incubation were lower for induced cells. For further experiments, we worked with uninduced cells, in order to study the effect of different mutations on the transport induction.


Figure 8.

[14C]sucrose transport into Xanthomonas campestris pv. campestris (Xcc). (A) Transport of [14C]sucrose over 120 minutes into the Xcc wild-type strain precultured in minimal medium without sugar (uninduced) or supplemented with 20 mM sucrose (induced). (B and C) Transport of [14C]sucrose over 8 minutes into the Xcc wild-type strain, the insertion mutants in suxR and in suxC (suxR::pVO and suxC::pVO, respectively), the deleted mutants in suxA and suxBsuxA and ΔsuxB, respectively) and their corresponding complemented strains [ΔsuxA (pL-suxA) and ΔsuxB (pL-suxB), respectively]. Cells were grown in minimal medium without sucrose. Transport was measured using 0.5 µM [14C]sucrose.


Competition experiments were performed to test transport specificity. A 10-fold excess of unlabelled sucrose reduced the transport rate to 52% of the level obtained without unlabelled sucrose, and with a 100-fold or 200-fold excess, the transport rates were reduced to 10.5 and 7% respectively (Table 2). Moreover, the addition of unlabelled fructose or glucose (sucrose degradation sub-products) slightly affected sucrose transport; a 200-fold excess of these carbon sources reduced the sucrose transport to about 80% of the control level (Table 2).


Table 2. [14C]sucrose transport rates related to uninduced Xanthomonas campestris pv. campestris wild-type strain (Xcc568).


The involvement of the SuxA TBDR in sucrose transport was then checked by comparing the uptake rate of [14C]sucrose into the wild-type strain and into a suxA non polar mutant (ΔsuxA). Sucrose transport was much lower in the ΔsuxA strain, compared to the wild-type strain (Figure 8B and Table 2): after 8 minutes, it reached only 3.5% of the transport rate into the wild-type strain. When suxA was supplied in trans on a constitutive expression plasmid, sucrose transport was more efficient and rapid, with a transport rate value at 8 minutes corresponding to 484% of that of the wild-type strain. This result confirms that SuxA is required for sucrose entry into Xcc.

We then studied sucrose transport in suxR and suxC insertion mutants (suxR::pVO and suxC::pVO, respectively), and in a suxB deletion mutant (ΔsuxB). When the inner membrane transporter encoded by suxC was absent, the sucrose transport rate reached only 2.1% of the value obtained for the wild-type strain. Thus, this protein is necessary for sucrose entry into Xcc. On the contrary, sucrose transport was greatly enhanced in the suxR repressor mutant and in the ΔsuxB amylosucrase mutant (488% and 241% respectively of the value obtained for the wild-type strain after 8 minutes) (Figure 8C and Table 2). These results confirm the repressor function of SuxR (see below) and suggest that SuxB also has a repressor activity on suxA expression. When suxB was supplied in trans in the ΔsuxB mutant, the sucrose transport was reduced to about the same level as the wild-type strain (76.4% of the transport rate level of the wild-type strain).

Concentration-dependent sucrose transport experiments showed a biphasic kinetics (Figure 9), with a fast rate between 0.01 and 0.1 µM sucrose (Kd = 0.033 µM), and a slow rate between 0.25 and 5 µM sucrose (Kd = 0.59 µM). This biphasic pattern is very similar to those observed in energy coupled transports of vitamin B12 into E. coli through the BtuB TBDR [55], [56] and more recently of maltose transport into C. crescentus through the MalA TBDR [22]. In both systems, it was concluded that the first phase reflects binding of the transported molecule to the OM TBDR and that the second slower phase reflects binding to a cytoplasmic membrane transporter. Thus, we presume that the low Kd value (0.033 µM) mainly reflects binding to SuxA and that the higher Kd value (0.59 µM) binding to SuxC.


Figure 9.

Concentration-dependent [14C]sucrose transport into wild-type Xanthomonas campestris pv. campestris. Cells were grown in minimal medium without sucrose and transport was measured for 15 sec at the indicated [14C]sucrose concentrations. The insert shows the Scatchard transformation of the binding data.


Altogether, these data indicate that both SuxA and SuxC are required for sucrose transport. It is worth noting that if the ΔsuxA mutant and the suxC insertion mutant have a different phenotype regarding their growth ability in presence of sucrose (see figure 7A), both mutants show a very similar alteration in sucrose transport rate after 8 minutes. However, after 240 minutes, sucrose transport rate in the suxA-deleted mutant reached almost 23% of the value obtained for the wild-type strain, whereas in the suxC insertion mutant, this rate was less than 7% of that of the wild-type strain. This result suggested the existence of two sucrose uptake pathways, an active and a passive pathway, both requiring transport through SuxC. The active transport pathway depends on SuxA for translocation through the outer membrane, whereas the passive pathway does not. It is assumed that the sucrose uptake rate observed into the suxA-deleted mutant is determined by slow and passive diffusion of sucrose across the outer membrane, which is sufficient to support growth in the presence of 20 mM sucrose in the medium.

Sucrose uptake is energy-dependent

Active iron transport through TBDRs depends on the proton motive force (PMF) [56], [57]. Thus, we performed sucrose transport experiments in the presence of a PMF inhibitor, carbonyl cyanide 3-chlorophenyl-hydrazone (CCCP). Addition of 20 µM CCCP inhibited [14C]sucrose transport rate to 1.13% of the transport rate in the absence of CCCP (Table 2), demonstrating that sucrose transport through SuxA and/or SuxC is dependent on the proton motive force and does not occur by facilitated diffusion. These experiments did not permit us to conclude whether SuxA-mediated transport depended on the PMF, since SuxC belongs to the Na+-melibiose cotransporter and H+ cotransporters family, which require the transmembrane potential for transport across the cytoplasmic membrane. We then tested whether TonB was required for sucrose uptake. Eight genes coding for proteins displaying significant similarities to TonB are predicted in the genome of Xcc (Table 2). These 8 genes were mutated by insertion of the pVO155 plasmid. As the insertion in XCC2927 was found to be unstable, we constructed a deletion mutant in this gene (Materials and Methods). Sucrose transport was not significantly impaired in the 8 putative tonB mutants (Table 2). This result suggests a complete or partial functional redundancy between at least two Xcc TonB proteins. Such a redundancy has already been observed in various other bacteria, e.g. Serratia marcescens [58], Vibrio cholerae [59] and Pseudomonas aeruginosa [60], [61].

The expression of sux genes is induced by sucrose and repressed by SuxR

Using qRT-PCR analyses, we observed that the expression of suxR, suxC and suxAB is specifically induced by the presence of sucrose in the medium (Table 3).


Table 3. Relative expression ratios of suxR, suxC, suxA and suxB in the wild-type strain, in minimal medium containing 20 mM sucrose vs minimal medium without sucrose.


As SuxR has a high degree of similarity with members of the LacI/GalR family of transcriptional repressors [62], [63], we tested whether this protein regulates the expression of sux genes by comparing their expression by qRT-PCR in the suxR insertion mutant and in the wild-type strain, cultivated in minimal medium with or without sucrose. suxAB, suxC and suxR were over-expressed in a suxR mutant as compared to the wild-type strain in the absence of sucrose (Table 4), suggesting a negative and effector-dependent control of sux genes by SuxR.


Table 4. Relative expression ratios of suxR, suxC, suxA and suxB in the suxR insertion mutant (suxR::pVO) vs wild-type strain.


We also studied the expression of the suxAB operon in the presence of different sucrose concentrations. For this purpose, the promoter region of this operon was cloned upstream of the promoterless LacZ gene in a reporter plasmid (see Materials and Methods). This plasmid, named pPr-suxA, was introduced into the Xcc wild-type strain, the suxR and suxC insertion mutants (suxR::pVO and suxC::pVO, respectively), and the suxA and suxB deletion mutants (ΔsuxA and ΔsuxB, respectively).

These strains were used to perform β-galactosidase assays after 6 hours growth in minimal medium containing a range of sucrose concentrations (Figure 10A). Induction of the expression of the reporter gene was detected for sucrose concentrations ranging from 20 µM to 20 mM. Maximal induction (3.5 to 4-fold induction) was observed for concentrations higher than 200 µM in the wild-type background (Figure 10A). As expected, the expression of suxA was highly induced in the suxR mutant, confirming the repressor activity of this gene. However, we observed a significant diminution of the induction level at 2 and 20 mM sucrose in this mutant, suggesting the existence of a second level of control. A high and constitutive induction of suxA expression was also observed in the suxB deletion mutant, suggesting that the amylosucrase gene plays a role in the repression of the suxAB operon. At 2 mM sucrose concentration, a reduction in suxA expression level was also observed in this mutant, but it was smaller than that detected in the suxR mutant. The deregulation of suxA expression in the suxB mutant is in agreement with transport studies which showed a higher sucrose transport rate in this mutant (Figure 8C). No induction of suxA expression was observed in the suxC mutant, whatever sucrose concentration was used. This lack of induction certainly reflects the absence of sucrose entry into the cytoplasm of this mutant, thus preventing the alleviation of SuxR repression. This result clearly shows that the induction of suxA expression by sucrose requires the entry of this molecule into the cell. The pattern and level of expression of suxA observed in the suxA mutant were very similar to those observed in the wild-type strain, except that this expression was significantly higher in the mutant in the presence of 2 mM sucrose. This difference was reproducibly observed and further investigations are needed to understand this phenomenon.


Figure 10.

Expression of the Xanthomonas campestris pv. campestris suxA gene in the presence of sucrose, fructose or glucose. The pPr-suxA plasmid carrying the promoterless LacZ reporter gene under the suxA promoter region was used to monitor suxA expression in different genetic backgrounds. Expression was measured in minimal medium and cells were harvested at the indicated times. (A) Expression after 6 hours induction in the presence of different sucrose concentrations, in the wild-type background, the suxR and suxC insertion mutant backgrounds (suxR::pVO and suxC::pVO, respectively), and the suxA and suxB deletion mutant backgrounds (ΔsuxA and ΔsuxB, respectively). (B) Kinetics of LacZ expression in the wild-type background or in the suxA deletion mutant background (ΔsuxA), in presence of 20 mM (triangles) or 100 µM (squares) sucrose. (C) Kinetics of LacZ expression in the wild-type background in the presence of 100 µM sucrose, fructose or glucose.


Expression assays suggested that sucrose transport through SuxA is not high enough to influence the expression of the suxAB operon. Sucrose entry across the outer membrane by slow diffusion seems to be sufficient to promote induction of sux genes after 6 hours growth in the presence of sucrose.

suxA is required for its rapid induction as revealed by using low sucrose concentrations

We performed time-course experiments comparing suxA expression in the wild-type strain and the suxA deletion mutant grown in presence of different sucrose concentrations (0, 20 µM, 50 µM, 100 µM, 200 µM 500 µM, 1 mM, 2 mM, 5 mM and 20 mM). For clarity, only results obtained with 100 µM and 20 mM are shown in Figure 10B. In the presence of 20 mM sucrose (triangles on Figure 10B), suxA expression was not significantly different after 6 hours growth in the wild-type and the suxA mutant backgrounds. However, a slight difference was observed in the induction level of suxA, from 60 to 120 minutes after sucrose addition. The expression level of suxA was 1.5 to 1.6 fold higher in the wild type strain than in the suxA mutant. Later, this difference tended to decrease and the ratio reached a value of 1.1 after 6 hours. A reproducible and clear difference in curve profiles was noticed in the presence of 100 µM sucrose (squares on Figure 10B). A rapid induction was observed in the wild-type context, from 30 to 120 minutes after sucrose addition, followed by a plateauing, in which expression seems to remain constant. On the other hand, in the suxA deletion mutant background, the induction was linear and less rapid. Between 30 and 120 minutes after sucrose addition, the expression level of suxA was 2 fold higher in the wild-type strain background than in the suxA deletion mutant background, although after 360 minutes, the expression levels were identical in both strains. Similar results were obtained with sucrose concentrations ranging from 20 to 200 µM (data not shown). We propose that these results could reflect the existence of two pathways controlling suxA expression, which can be clearly differentiated with low sucrose concentrations. One of these pathways is controlled by SuxA. These results can be related to the transport analyses showing the existence of two types of sucrose transport across the outer membrane, one being SuxA-dependent and the other one, slower, being dependent on passive diffusion. We thus postulate that the induction of suxA indirectly reflects these two means of transport. Moreover, this induction is specific to sucrose and is not due to its degradation products glucose and fructose, as no induction was observed following addition of 100 µM of these molecules in the wild-type background (Figure 10C). Therefore, these experiments suggest that SuxA plays a role in sucrose transport which might be crucial at low concentrations of this sugar.

Identification of other Xcc TBDRs in loci probably involved in plant carbohydrates utilization

The genome of Xcc was then explored to see whether other TBDRs could belong to loci involved in the utilization of other plant compounds. Xcc has an extensive repertoire of plant cell-wall degrading enzymes, with cellulolytic, pectinolytic and hemicellulolytic activities [14]. The analysis of carbohydrate active enzymes identified in the predicted proteomes of 209 Gram negative bacteria and referenced in the CAZy database ( showed that, after Bacteroides sp. and S. degradans, which are well known specialists for polysaccharide degradation, Xcc has one of the highest number of genes involved in polysaccharide metabolism per megabase (29.9, total 152) (Table S2). These genes encode 82 predicted glycosyl hydrolases, 45 glycosyl transferases, 5 polysaccharide lyases, 18 carbohydrate esterases and 2 carbohydrate binding proteins. Interestingly, 46 of these proteins are encoded in the vicinity of 24 TBDR/Ps-TBDR genes, thus suggesting the existence of 19 new loci putatively involved in carbohydrate utilization (Table S3). Among those loci, 7 also contain an inner membrane transporter coding gene and a regulatory gene. For 3 loci identified in this analysis, the substrate probably utilized and transported could be easily deduced from the nature of the degradative enzymes: the XCC0120 TBDR gene is localized upstream of genes coding for a pectin methyl esterase and a pectate lyase and might belong to a locus involved in pectin utilization (Figure 11A); the XCC4120 TBDR gene is found in a cluster of genes probably involved in xylan metabolism (Figure 11B); and the XCC2469 TBDR gene might be related to maltose/maltodextrin utilization, since it is associated with genes coding for a cyclomaltodextrin glucanotransferase, two α-glucosidases, a sugar transporter and a maltose transport gene repressor (Figure 11C).

To verify whether these TBDRs are really associated with carbohydrate utilization, we studied their regulation by plant compounds. For this purpose, we performed β-glucuronidase expression assays using all Xcc TBDR pVO155 insertion mutants. Cells were grown in rich medium or in minimal medium supplemented or not with polygalacturonic acid (PGA), arabinose, glucose, maltose, sucrose, xylose or xylan. Most TBDR (or Ps-TBDR) genes (44 out of 74) were repressed in rich medium, compared to minimal medium (Table S4). We also observed that in minimal medium, 48 Xcc TBDR genes are repressed in the presence of sucrose, xylose, arabinose and/or glucose, suggesting that these genes are submitted to catabolic repression (Table S4). However, the repression patterns were variable, suggesting the existence of several repression pathways. It is worth noting that most Fur-regulated TBDR genes were submitted to catabolic repression.


Figure 11.

Genetic organization of putative Xanthomonas campestris pv. campestris CUT loci and specific induction of the TonB-dependent receptor gene by plant carbohydrates. Putative pectin (A), xylan (B) and maltodextrin (C) utilization CUT loci and specific induction of the corresponding TBDR gene in the presence of PGA, xylose and maltose, respectively. β-glucuronidase assays were performed in at least two independent experiments with TBDR insertion mutants cultivated in minimal medium supplemented with glutamate 20 mM (GT) or specific carbohydrates (PGA: polygalacturonic acid 0.125%; XYL: xylose 20 mM; MAL: maltose 20 mM). TBDR: TonB-dependent receptor; PME: pectin methyl esterase; PL: pectate lyase; CGTase: cyclomaltodextrin glucanotransferase.


Three TBDR/Ps-TBDR genes were induced by PGA (XCC0120, XCC1749 and XCC1750-1751), one by maltose (XCC2469), seven by arabinose (XCC0050, XCC1749, XCC1750, XCC1892, XCC2828, XCC4120 and XCC4222), three by xylan (XCC2828, XCC4120 and XCC4237) and finally two by xylose (XCC2828 and XCC4120) (Table S4). Interestingly, the 2 TBDRs induced by xylose were also induced by xylan and arabinose, and XCC1749 and XCC1750 were both induced by PGA and arabinose. suxA is the only TBDR gene for which expression is specifically induced by sucrose, and XCC2469, the orthologue of C. crescentus malA TBDR gene [22], is the only one induced by maltose. It is worth noting that the expression of the XCC0120, XCC4120 and XCC2469 TBDR genes was specifically induced by the postulated substrate of the associated CAZy referenced enzymes (see Figure 11). Altogether, these data suggest that several TBDRs are part of loci which seem to be involved in carbohydrate utilization. We thus propose the existence of 6 putative loci named carbohydrate utilization containing TBDRs (CUT) loci, which were defined by the presence of genes coding for carbohydrate degradative enzymes, inner membrane transporters and sugar related regulators beside TBDR genes. We also identified 15 putative partial CUT loci in the Xcc genome (see Table S3).

Conservation and distribution of TBDRs and CUT loci in Xanthomonads

When compared with proteins in the databases using BlastP and the “distance tree of results option” displayed on the BlastP result page, the best homologous genes of Xcc TBDRs/Ps-TBDRs were putative TBDRs of other Xanthomonas species and in some cases of Xyllela strains. These conserved genes might be considered as orthologs. We compared the distribution of Xcc TBDRs among Xanthomonads strains for which genome sequences are available, i.e. Xcc strain 8004, Xcv (strain 85–10), Xac (strain 306), Xoo (strains KACC10331 and MAFF311018) and Xf (strains 9a5c and PD). A search similar to that used for Xcc allowed us to identify 72 TBDR/Ps-TBDR in the proteome of Xcc strain 8004, 74 in that of Xac, 61 in Xcv, 41 and 42 in Xoo strains KACC10331 and MAFF311018 respectively, and finally 10 in each Xf strain. To further study the relationship between these TBDRs and Xcc TBDRs, we performed a comparative study using alignments generated by ClustalW [64]. To carry out this comparative analysis, we only used TBDRs which seemed to be complete and Ps-TBDRs over 600 amino acids, in order to avoid bias in alignments. Examination of the phylogenetic tree, coupled with Blast results analysis, clearly confirmed that most Xcc TBDRs are well conserved in other Xanthomonas strains (Figure S1 and Table S3). All TBDR or Ps-TBDR genes detected in Xcc strain ATCC33913 are present in strain 8004; 55 seem to have orthologs in Xac; 49 are conserved in Xcv and 34 only are conserved in Xoo strains which possess significantly less TBDRs. In most instances, the grouping and distribution pattern of orthologous TBDRs matched the phylogenetic classification of Xanthomonas species [65][67]. This observation suggests that this protein family is ancient in the Xanthomonas genus. It is worth noting that branch lengths were variable, suggesting differential evolution rates. Moreover, we noticed that some truncated Xcc Ps-TBDRs could be functional in other strains: the Ps-TBDR XCC1750-1751 doublet corresponds to unique and complete TBDR proteins in Xcv, Xac and Xcc strain 8004. Similarly, the 2 Ps-TBDRs XCC3215-3216 and XCC3270-3271, which are also truncated in Xcc strain 8004, correspond to unique proteins in both Xac and Xcv. Interestingly, Xac, Xcv or Xoo TBDR orthologs of Xcc Ps-TBDR having non-canonical C-terminal regions (β-sheet ended by an aromatic amino acid followed by a short extension of 1 to 4 amino acids), also displayed this feature (data not shown). In most cases, these non canonical C-terminal domains were identical in all strains. This conservation might reflect a specific functional feature of Xanthomonas TBDRs.

The analysis of the phylogenetic tree also showed that Xanthomonads TBDRs can be divided into three main groups, with most TBDRs belonging to group 1. This group can be divided into 8 subgroups (1A to 1H). We noticed that (i) Xcc Oar-TBDRs are clustered in subgroup 1D, (ii) most plant carbohydrate induced TBDRs (8 out of 11) are grouped in subgroup 1C, and (iii) most Xcc Fur-regulated TBDRs (6 out of 8) are grouped in subgroup 1F. Among these Fur-regulated TBDR genes, XCC3518, XCC3595 and XCC4162, which belong to subgroup 1F, seem specific to Xcc strains.

The comparison of genes located adjacent to orthologous TBDR genes in the different Xanthomonas strains showed that in most cases, these regions are syntenic (Table S3). Thus, all putative CUT loci identified in Xcc genome are well conserved in Xac and Xcv but only 2 out of 6 were found in Xoo. Similarly, almost all putative partial CUT loci are present in both Xcv and Xac, but only 9 out of 15 are conserved in Xoo strains. The putative polygalacturonate utilization locus (XCC0120-XCC0122) seems unique to Xcc. The genes bordering this locus are conserved and contiguous in Xcv and Xac, thus suggesting insertion or deletion events (Table S3).

Finally, among the 10 TBDRs identified in Xf strains, 6 are conserved in Xcc, Xac, Xcv and Xoo, 1 is conserved in Xcc, Xac and Xcv, whereas only 1 is present in Xcc. The 2 remaining Xf TBDRs (XF0339/PD1711; XF0599/PD1552) seem more divergent and thus specific to Xf strains. However, XF0599 displays weak similarities with XCC2772, a Fur-regulated TBDR of Xcc. None of the other Xf TBDRs are related to Xcc Fur-regulated TBDRs nor to Xcc TBDRs belonging to putative CUT loci identified in this study. Only 2 Xf TBDR genes (XF2713 and XF1036) correspond to Xcc TBDRs present in partial CUT loci.

Xcc TBDRs are conserved in aquatic bacteria and/or phytopathogenic bacteria

For each Xcc TBDR included in the phylogenetic study, we analyzed the BlastP results obtained on the nr databank, to characterize the next best homologous genes after Xanthomonads orthologs. This analysis allowed us to cluster Xcc TBDRs on the basis of the bacterial origin of the homologous TBDR. Thus, Xcc TBDRs could be divided into 4 main groups (Table S5). The first group corresponds to TBDRs showing their next best homologies (after Xanthomonads orthologs) with other Xanthomonas TBDRs rather than with TBDRs from other genera. This observation mainly concerned TBDRs found in subgroups 1B, 1C and 1D of the phylogenetic tree. Their positions in the tree suggested that they could be generated by successive duplication events that occurred before Xanthomonas speciation (Figure S1). The second group corresponds to TBDRs which display high similarities with TBDRs of β-Proteobacteria and/or Pseudomonas species. Interestingly most of these TBDRs are clustered in subgroup 3 of the phylogenetic tree. Moreover, some of these TBDRs showed high similarities with TBDRs of phytopathogenic bacteria such as Acidovorax avenae subsp. citrulli, Pseudomonas syringae pathovars or Ralstonia solanacearum. The third group contains two Xcc TBDRs proposed to be involved in iron uptake and showing significant similarities with putative TBDRs from Cyanobacteria. The fourth group, which is the largest one (with 35 TBDRs out of 70), corresponds to TBDRs which displayed very significant homologies with putative TBDRs of a wide range of aquatic bacteria belonging to the α or γ classes of Proteobacteria. It is worth noting that in most cases, Xcc TBDRs were not affiliated to TBDRs of a specific bacterial class, but there was rather a variety of origins. Thus, for 16 Xcc TBDRs the best similarities (after Xanthomonads similarities) were obtained with putative TBDRs of C. crescentus CB15 and Caulobacter sp. K31 strains, which are aquatic oligotrophs belonging to the Caulobacterales order of the α-Proteobacteria. Homologies were also observed with TBDRs of other α-Proteobacteria living in aquatic habitats, such as Oceanicaulis alexandrii, Maricaulis maris or Parvilarcula bermudensis HTCC2503, or found in multiple environments like Sphingomonas sp. SKA58, Sphingopyxis alaskensis or Novosphingobium aromacitovorans. Significant similarities were also obtained with putative TBDRs of aquatic γ-Proteobacteria classified in the Alteromonadales, including Alteromonas macleodii, Saccharophagus degradans 2–40, Pseudoalteromonas and Shewanella species. One common trait of most of these bacteria is that they show TBDRs overrepresentation with values of TBDR number per megabase ranging from 7.4 to 15.7 (Table S2). This raised the question of whether homologies between these TBDRs were fortuitous and a consequence of their large number or whether they reflect common biological functions.

Conservation of TBDR regions and CUT loci in Gram-negative bacteria

Some regions surrounding TBDR genes were found to be conserved in closely related bacteria such as Vibrio parahaemolyticus or P. aeruginosa. Thus, the XCC3050 Fur-regulated TBDR belongs to a cluster of 6 genes showing similarities with the pvuA-pvsABCDE gene cluster of V. parahaemolyticus, involved in the uptake and biosynthesis of the siderophore vibrioferrin [68] (Table S3). Similarly, a group of proteins encompassing XCC3067 TBDR and putatively involved in cobalamin uptake and biosynthesis in Xcc was conserved in Pseudomonas species (Table S3). Preliminary experiments showed that the expression of this TBDR gene is repressed by the presence of vitamin B12 in Xcc, thus suggesting that this cluster is functional (data not shown).

Several CUT loci identified in Xcc were not conserved in taxonomically related bacteria but rather in the group of aquatic bacteria showing TBDR conservation and in particular in C. crescentus. The sux locus, 3 of the putative CUT loci and 4 of the putative partial CUT loci identified in this study were entirely or partially conserved in this group of bacteria (Table S3). Thus, the putative maltose CUT locus was partially conserved with the mal locus recently identified in C. crescentus, which contains the MalA TBDR proposed to be involved in the uptake of maltodextrins [22] (Figure S2B). Interestingly, MalA shows significant similarities with the XCC2469 TBDR whose expression is induced by maltose and maps in the maltose CUT locus (Figure 11C).

Similarly, the putative xylose locus, containing the XCC2828 TBDR gene, is also very well conserved in C. crescentus (Figure S2C). The corresponding CC0999 TBDR gene was shown to be induced by xylose in this bacterium [69]. Moreover, CC2832, which was also shown to be xylose-induced in C. crescentus, displays significant homologies with the xylose-induced TBDR gene XCC4120 belonging to a xylose CUT locus. Furthermore, the 20-bp palindromic motif conserved upstream of genes of the C. crescentus xylose regulon, was also found upstream of genes putatively involved in xylose metabolism in Xcc, as well as in the promoter region of XCC2828 and XCC4120 TBDR genes [69].

The sux CUT locus seems less well conserved and showed some degree of variation (Figure S2D). The amylosucrase gene (XCC3359) is only well conserved in the corresponding C. crescentus locus. In the loci conserved in other bacteria, the degradation of sucrose seems to involve more classical pathways (for review see [70]). Moreover, the MFS transporter of the Xcc locus is different from that of the C. crescentus sucrose locus encoded by CC1133, although it is well conserved in both Sphingomonas SK58 and Erythrobacter litoralis. These differences show the existence of some degree of plasticity in the evolution of the putative CUT loci. Further investigations are needed to see whether all these partially conserved loci are involved in the utilization of the same molecule. However, this wide conservation of CUT loci is in favor of their existence. These results also confirm that the similarities observed between TBDRs are probably not fortuitous.



The outer membrane (OM) of Gram negative bacteria serves as a selective permeation barrier, excluding hydrophilic solutes, including most nutrients. However, OMs contain embedded integral proteins, named outer membrane proteins (OMPs), which allow sensing and entry of nutrients into the cell. A major class of OMP with a certain substrate specificity, called porins, allows the translocation of hydrophilic solutes through the OM. Another class of OMPs, the TonB-dependent receptors (TBDRs), is mainly known to be involved in iron or vitamin B12 uptake [71]. Recently, the MalA TBDR of C. crescentus was shown to be involved in the ExbBD-dependent uptake of maltodextrins [22]. Here, we show that several Gram negative bacteria, belonging to different lineages and having diverse habitats, display an overrepresentation of TBDRs, which might be related to the uptake of plant-derived carbohydrates.

Xcc TBDRs belonging to the Fur regulon

Our global study of Xcc TBDRs showed that only a small fraction of them seems to be involved in iron uptake. Among the 72 TBDR/Ps-TBDRs identified in the Xcc genome, only 9 are up-regulated under iron-limiting conditions. We established that these 9 genes are repressed by the Fur repressor and that they are the only TBDR genes having a Fur-box in their promoter region. Recently, a proteomic approach carried out in P. aeruginausa PAO1, which contains 34 putative TBDR genes, identified a very similar number of TBDR genes regulated by iron-stavation. In fact, in this bacterium, 7 TBDRs are produced under iron-starvation conditions and 4 others are specifically induced by the presence of heterologous siderophores, under iron-restricted conditions [72]. Therefore, although it is possible that some Xcc TBDR genes need specific heterologous siderophores for their expression, as observed in P. aeruginosa, the number of TBDR genes involved in iron uptake seems very comparable in both bacteria. This suggests that the other putative TBDRs/Ps-TBDRs might be required for different biological functions.

The identification of CUT loci/systems in Xcc suggests a relationships between TBDRs and carbohydrate utilization

Our data reveal that several Xcc TBDRs are related to plant carbohydrate utilization. A large proportion of Xcc TBDR genes are coupled with carbohydrate active enzymes. For 6 of them, an inner membrane transporter gene and a regulatory gene are also present in the same region, thus defining the existence of putative CUT loci involved in the uptake and utilization of plant carbohydrates. Moreover, 15 Xcc TBDR genes belong to partial CUT loci, thus suggesting the existence of CUT systems composed of different parts scattered in the genome. The existence of such multipartite CUT systems was supported by the observation that 11 TBDR/Ps-TBDR genes are specifically induced by plant carbohydrates such as sucrose or plant cell wall derived compounds including arabinose, xylan/xylose, pectin/polygalacturonate or maltose. Moreover, in 5 cases, there is a correlation between the inducing carbohydrate and the degradative enzyme(s) present in the CUT locus. This allows us to propose the existence of a sucrose CUT locus as well as loci involved in the utilization of complex carbohydrates such as pectin, xylan or starch.

The sux CUT locus is functional

The existence of functional CUT loci was confirmed by the detailed study of the sucrose CUT locus, which showed its involvement in the entry and utilization of sucrose in Xcc. This locus comprises four genes, suxA, suxB, suxC and suxR, coding for a TBDR, an amylosucrase, a sugar inner membrane transporter and a regulatory protein, respectively. [14C]sucrose uptake experiments showed that SuxA and SuxC are both required for sucrose entry into the cell. Concentration-dependent sucrose transport experiments showed a biphasic kinetic, with a similar pattern observed for vitamin B12 uptake into E. coli through the BtuB TBDR [55], and more recently for maltose transport into C. crescentus through the MalA TBDR [22]. In both of these systems, it has been concluded that the first phase reflects the binding of the transported molecule to the OM TBDR, and that the second slower phase reflects the binding to a cytoplasmic membrane transporter. Thus, we presume in our experiments that the low Kd value (0.033 µM) mainly reflects binding to SuxA and that the higher Kd value (0.59 µM) binding to SuxC. Sucrose transport was significantly lower in the suxC mutant than in the suxA non polar mutant. This difference might explain the differential phenotype of these two mutants observed for growth on sucrose: the suxA non polar mutant was not impaired in growth on media containing sucrose, whereas the suxC mutant was unable to grow under these conditions. These observations suggest the existence of an alternative pathway that supports facilitated diffusion of sucrose across the OM, whereas it seems that there is a unique route for crossing the inner membrane, depending on SuxC. The existence of this alternative pathway was also revealed by expression analyses of the sux locus. The regulation of sux genes seems to follow a classical inducer/repressor control, mediated by the SuxR repressor, sucrose being the inducer. The expression of sux genes is induced by sucrose but not by fructose or glucose. This induction was detected with sucrose concentrations ranging from 20 µM to 20 mM. These experiments showed that SuxA transport influenced sucrose induction of suxA at low sucrose concentrations (20 to 200 µM), whereas this effect is masked at higher concentrations, probably by interference by passive diffusion.

Our data also indicated that sucrose transport through the sux system is active and depends on the proton motive force. However, we could not conclude whether it depends on the TonB-ExbBD energy coupling system. In addition to TonB, the Xcc genome harbours 7 TonB-like proteins, which might substitute one for another. Another Xcc feature is the presence of a second exbD gene, named exbD2, mapping downstream of the tonB-exbBD1 locus. Interestingly, exbD2 is not required for iron uptake but is essential for HR induction, whereas tonB, exbB and exbD1 genes are necessary for both processes [73], [74]. This differential behavior might be related to the existence of at least two classes of TBDRs in Xcc, one involved in iron uptake and the other one in transport of plant compounds. Further work is needed to address this hypothesis.

The sux locus represents a new sucrose utilization system

The Xcc sux locus is clearly different from other sucrose utilization loci already found in bacteria (for review see [70]). In particular, it differs from the scr sucrose-utilization system from enteric bacteria, which is also present in the plant pathogenic bacterium Erwinia amylovora, where it plays a major role in plant colonization [75]. The differences concern regulation, sucrose utilization and transport. The scr genes are regulated by a LacI/GalR family repressor, but fructose is the inducer [76]. In this system, the degradation of sucrose uses a sucrose-6-phosphate hydrolase/fructokinase degradation pathway [70]. On the contrary, the Xcc sux locus contains an amylosucrase orthologous to SUH, a unique sucrose hydrolase previously characterized in Xag [54]. This enzyme is responsible for intracellular sucrose hydrolysis. It is active on sucrose but not on sucrose-6-phosphate. In the scr system, sucrose metabolism and uptake are coupled by the phosphoeneolpyruvate-dependent carbohydrate:phosphotransferase system (PTS), which controls crossing of the inner membrane and sucrose phosphorylation. The transport across the outer membrane is mediated via a porin, named ScrY [77], [78]. Thus, the transport of sucrose is very different in sux and scr systems. Interestingly, the Kd value of sucrose binding to SuxA is 1500- to 3000-fold lower than that of the E. coli ScrY sucrose porin, which varies from 13 mM [79] to 50 mM [80]. Similarly, C. crescentus MalA TBDR transports maltodextrins with Kd values 1000-fold lower than those of the LamB porin, which facilitates the passive diffusion of maltodextrin [22]. It is worth noting that the Kd values of sucrose binding to SuxA and maltodextrins binding to MalA are comparable. Thus, SuxA and MalA represents a new class of outer membrane carbohydrate transporters showing a much higher affinity for their substrate than porins.

The importance of the sux locus in Xcc is highlighted by the fact that it is required for full virulence on Arabidopsis thaliana. The phenotype of suxB and suxC mutants on plants can be related to their inability to grow on medium containing sucrose (20 mM). However, the non polar suxA mutant, which grows as well as the wild type strain on medium containing sucrose, was also affected in pathogenicity. Thus, it appears that the ability to scavenge sucrose with a very high affinity plays a key role during the interaction with host plants.

In conclusion, data obtained on this sucrose CUT locus strongly support the existence of the other CUT loci identified in Xcc, which might have a similar mode of action (see model presented in Figure 12). This suggests the presence of several systems which seem to partially overlap and which are involved in the scavenging of plant molecules. They might form a complex network required for the exploitation of plant resources but which might also participate in signaling.


Figure 12.

Model of CUT loci functioning based on the Xanthomonas campestris pv. campestris sux locus. This scheme shows sucrose outer membrane transport via the SuxA TBDR or by passive diffusion through a putative porin. After crossing the inner membrane through the SuxC transporter, sucrose is proposed to interact with the SuxR repressor (thus allowing sux gene induction) and also to serve as a substrate for the SuxB amylosucrase. The large double headed arrow below the sux locus represents the balance between metabolic adaptation and virulence control putatively mediated by CUT loci.


TBDRs and CUT loci are ancient in the Xanthomonas genus

The importance of TBDRs and CUT loci is not restricted to Xcc since they are well conserved in Xac and Xcv. A phylogenetic study of Xanthomonads TBDRs suggests that these proteins are an ancient class of proteins in the genus. Moreover, the grouping pattern observed in our phylogenetic study seems to correlate with functional features of Xcc TBDRs, thus suggesting the existence of structure/function relationships. However, it seems that there is some degree of variation in the repertoire of TBDR genes and CUT loci among Xanthomonads, as the number of genes and loci was lower in Xoo strains and also in Xylella. This latter species is considered as a minimal pathogen in the Xanthomonads with a restricted habitat and a reduced genome [81]. Xylella possesses a significantly lower number of TBDRs and only one partially conserved CUT locus.

Conservation of Xcc TBDRs and CUT loci reveals carbohydrate scavenging abilities in other bacteria

One of the main surprises of this work was the observation that the large majority of Xcc TBDRs (35 out of 72) display significant similarities with TBDRs of bacteria which are mostly found in aquatic habitats and which are not closely related to Xanthomonads (see Table S5). Phylogenetic studies place Xanthomonads as a deep branch in the γ-Proteobacteria class, close to that of the β-Proteobacteria [66], [82]. The bacteria possessing similar TBDRs belong either to the α-Proteobacteria class or to the Alteromonadales order of the γ-Proteobacteria. Most bacteria in this latter order have been collected from diverse aquatic environments and belong mainly to 6 different genera: Alteromonas, Colwellia, Idiomarina, Pseudoalteromonas, Saccharophagus and Shewanella. The α-Proteobacteria mentioned in this study are also found mostly in aquatic habitats and are members of several orders, including Caulobacterales, Sphingomonadales, Rhodospirillales and Rhodobacterales. Similarities were not restricted to TBDRs genes, and genome context analyses showed that at least 8 putative (partial) CUT loci identified in Xanthomonas species were conserved among members of these aquatic bacteria. Several of them are found in association with abiotic or biotic surfaces, such as the surface of alga or shellfish. Some are adapted to live in oligotrophic environments, while others are abundant in nutrient-rich habitats or in association with particulate of organic matter. Some others are associated with decaying tissues of plant or algae. Although Xanthomonas seems to be very different from these bacteria in terms of habitat, lifestyle and taxonomy, we identified several common traits. All these bacteria belong to the class showing a TBDR overrepresentation (with TBDRs/Mbp ratios>7.4). Most of them are able to degrade a wide variety of plant molecules or other complex carbohydrates such as chitin, alginate, as well as various aromatic compounds. This combination of characters and the similarities with Xcc suggest the existence of a shared biology which might be related to the ability to scavenge carbohydrates. If the situation in these bacteria is similar to that observed in Xcc, we can speculate that there is a significant proportion of TBDRs involved in the uptake of molecules other than iron-siderophores complexes. Moreover, as previously noticed [33], we observed that bacteria having a restricted habitat, such as obligate parasites or symbionts, have no or only a very small number of TBDRs. Thus, these proteins might be considered as good indicators of bacterial lifestyle.

Association between TBDRs and carbohydrate utilization in other Proteobacteria

A specific role for TBDRs has already been attributed to bacteria displaying an overrepresentation of this protein family. Sphingomonads are widely distributed in nature and are mostly found in soils and aquatic environments. Some strains are associated with plants [83]. Strains like Sphingomonas sp. A1 are able to take up and metabolize macromolecules such as alginate produced by brown seaweed and certain bacteria. This uptake is mediated by large structures, so called superchannels, which form a pit in the outer membrane, and which function as a funnel or concentrator (for review, see [84]). Four TBDRs seem to be part of this superstructure and are thus proposed to participate in alginate transport [85]. Recently, the manipulation of these superchannels revealed the importance of these structures in bioremediation [86].

In C. crescentus, beside the prediction of 67 TBDRs, a genome analysis revealed the presence of several genes for the breakdown of plant polysaccharides as well as transport systems, suggesting that plant polymers are a significant source of nutriment for this organism [87]. Interestingly, several C. crescentus TBDR genes have been shown or proposed to be associated with the uptake of plant compounds. As described above, Neugebauer and colleagues showed that C. crescentus can grow on maltodextrins and that the transport of these molecules is mediated by the MalA TBDR [22]. Proteomic and transcriptomic studies have shown that TBDR genes are expressed at higher levels in minimal medium than in rich medium [69], [88][90], as we observed in Xcc. Moreover, several TBDR genes are specifically induced by the presence of xylose [69]. Interestingly, 2 TBDRs induced by xylose are conserved between C. crescentus and Xcc. Strikingly, putative regulatory cis-element motifs are conserved in the promoters of these homologous genes in both species. Moreover, one of these 2 Xcc TBDR genes (XCC2828) belongs to a putative partial CUT locus which is conserved in C. crescentus (Figure S2C). This suggests a possible lineage between these loci. Moreover, 3 other Xcc CUT loci are entirely or partially conserved in the C. crescentus oligotrophic bacterium, including the maltose and sucrose CUT loci. TBDRs in oligotrophs like C. crescentus might play a very important role by allowing the foraging of carbohydrates in nutrient poor environments.

Common themes between Xanthomonas and aquatic bacteria

It is clear that the life cycle of Xanthomonas spp. presents common features with the different lifestyles of bacteria described above. The leaf surfaces encountered by Xanthomonas during their epiphytic development might correspond to an oligotrophic environment. Plant leaves release secondary metabolites which can be used by epiphytic microorganisms [83]. However, carbon resources have been shown to be the most limiting resource on plants [91]. Simple sugars, like glucose, fructose and sucrose, are the most dominant carbon sources on plants that have been examined [92]. Studies with bacterial biosensors in situ on plants revealed a high heterogeneity of sucrose availability, with an average accessibility of only about 20 µM on moist bean leaves [93]. Interestingly, this sucrose concentration allowed the induction of suxA gene in our expression assays, thus suggesting that the sux uptake system might function on plant surfaces. The involvement of sux locus in epiphytic life is now being investigated. Furthermore, TBDRs might facilitate the exploitation of plant debris generated during disease development, thus resembling bacteria living on dead tissues. This phenomenon might have an impact on Xanthomonas life cycle by increasing Xanthomonas population size and thus facilitating new infections.

TBDRs and virulence on plants

It appears that in Xanthomonas, some TBDRs such as SuxA play an additional and specific role by controlling virulence. Therefore, it is tempting to propose that a strategy shared with non-pathogenic bacteria and based on the active uptake of plant-derived nutrients, could have been diverted by Xanthomonas for the control of pathogenicity. It is worth noting that several phytopathogenic bacteria such as P. syringae or E. carotovora subsp. Atroseptica, as well as Pseudomonas species associated with plants, have an intermediate overrepresentation of TBDRs, suggesting that this feature could be shared by other bacteria interacting with plants. In R. solanacearum, the expression of hrp genes coding for a type III secretion system (TTSS) which controls disease and HR development, is specifically induced upon contact with plant cells [94]. This signaling is mediated by PrhA, a TBDR belonging to the transducer subclass. This receptor senses contact with plant cells and transduces this signal into the cytoplasm via PrhI and PrhR, which are FecI/FecR homologs [94][96]. Apparently, this regulatory system does not require the transport of plant molecules [94]. Although Xac, Xcc, Xoo or Xcv do not carry any TBDR showing a high homology with PrhA, recently a TBDR controlling both HR and pathogenicity was described in X. oryzae pv. oryzicola, the causal agent of bacterial leaf streak of rice. This TBDR does not belong to the transducer subclass and only shows a weak similarity with PrhA [97]. This TBDR gene is highly conserved in Xoo strain MAFF 311018 (XOO0785), Xcv (XCV3654) and Xcc (XCC0674). Interestingly, in Xoo it is located close to trh (XOO0783), a regulatory gene controlling the expression of hrp genes [53]. The orthologue of this regulatory gene is also conserved in Xcc (XCC0672), in the vicinity of the XCC0674 TBDR gene. Thus, it is possible to speculate that both genes might be involved in a common circuit controlling the expression of hrp genes in Xanthomonas spp. In our study, a mutation in XCC0674 did not affect the pathogenicity of Xcc. However, as there are more TBDRs in Xcc than in Xoo strains, we can speculate that functional redundancy might have masked the effect of the mutation. Further work is needed to see whether these genes regulate hrp genes in Xcc. Nevertheless, we established a link between hrp genes and TBDRs in Xcc. Indeed, we observed that 2 Xcc TBDR genes are regulated by the hrpG and hrpX regulatory genes. Therefore, it seems that there is an overlap between hrp genes and at least 2 TBDR genes. What is the function of these two TBDRs? Are they specifically required during the infection of plants, through the action of the hrp regulon, to exploit specific released plant molecules?

What other functions for Xcc TBDRs?

We have been able to define a putative role for 21 TBDRs out of 72 identified in Xcc. The nature of the molecules putatively transported by the other TBDRs remains to be discovered. It is probable that TBDRs are not restricted to the transport of carbohydrates and that they can take up various other molecules produced by plants. Azospirillum irakensis, a plant associated bacterium, is able to metabolize salicin, a phenolic glycoside produced by plants. Interestingly, the sal operon, which controls the degradation of salicin, contains a TBDR gene which was proposed to be involved in salicin uptake [98]. Thus, secondary metabolites including phenolic compounds might be assimilated through TBDRs. We are now trying to characterize which new molecules may be transported by Xcc TBDRs, to better understand the adaptation of this pathogen to host plants.

TBDRs, CUT loci and evolution in Gram-negative bacteria

This work has identified the existence of an ensemble of bacteria that have an overrepresentation of TBDR genes, and that share specific loci for the scavenging and utilization of carbohydrates. They belong to very different lineages in Proteobacteria and this raises the question of the origin of these TBDRs and CUT loci. Did they arise by convergent evolution or were they transferred from species to species by lateral gene transfer? Our data suggest that this latter hypothesis is most likely. Recently, a genomic comparative study established that Xcc and Xac have close to 40% of their genes showing highest similarities to genes from non γ-Proteobacteria, especially from α-Proteobacteria (20%). These genes seem to belong to genomic islands, denominated “unusual best-match islands” (UBIs) [82]. Interestingly, among 35 UBIs thus identified in Xcc genome, 14 contain TBDRs and/or CUT loci. However, there was no significant difference between the GC content of each of these loci and the overall content of the genome. It is therefore possible that these loci were acquired very early in the evolution of Xanthomonads and thus played a key role in their adaptation to plants. Most bacteria showing a TBDR overrepresentation possess TBDRs or CUT loci conserved with Xcc. However, there is a main exception with members of the Bacteroidetes phylum, as none of the TBDRs characterized in this phylum showed very significant similarities with those identified in Xcc. Bacteroidetes can be encountered in two very different niches, the marine environment and the human intestine [99], [100]. Marine Bacteroidetes such as Gramella forsetii are associated with particulate of organic matter [99], whereas those found in the intestine are assembled on partially digested food particles [101]. In both cases, these bacteria are able to consume biopolymers. Bacteroides thetaiotaomicron, which is a prominent mutualist in the distal intestine of adult humans, has the largest ensemble of TBDR genes and glycobiome yet reported (Table S2). Studies have shown that this bacterium has a carbohydrate foraging behavior [100][102]. It is well known to bind starch through a protein complex of the outer membrane, which comprises SusD and the SusC TBDR protein [103]. One hundred and six paralogs of SusC and fifty three paralogs of SusD were predicted in the B. thetaiotaomicron genome [104]. In our study, we did not detect in Xanthomonas genomes any protein displaying similarities with SusD. Moreover, none of the TBDRs identified in Xcc showed strong similarities with SusC. A Blast analysis suggested that the SusC TBDR and the SusD OMP are specifically conserved in Bacteroidetes (data not shown). These results suggest that TBDRs involved in carbohydrate uptake evolved independently in Proteobacteria and Bacteroidetes. The analysis of the function and evolution of TBDRs in these phyla will certainly help us to better understand the adaptation of bacteria to their environment. This knowledge, which concerns the utilization of plant molecules that are widespread in the environment, will have a major impact not only in plant pathology, but also in human health as well as in the cycling of carbon and geobiology in marine environments.

Materials and Methods

Bacterial strains, plasmids and growth conditions

The Xanthomonas campestris pv. campestris (Xcc) strains and plasmids used in this study are listed in Table S6. Xcc cells were grown at 30°C in MOKA rich medium (Yeast Extract 4 g/l, Casamino acids 8 g/l, K2HPO4 2 g/l, MgSO4.7H2O 0.3 g/l) or in MME minimal medium [12]. E. coli cells were grown on LB medium [105]. Antibiotics were used at the following concentrations for Xcc: rifampicin, 50 µg/ml; kanamycin: 50 µg/ml; tetracycline: 5 µg/ml. Antibiotics were used at the following concentrations for E. coli: ampicillin, 50 µg/ml; kanamycin: 50 µg/ml; tetracycline: 10 µg/ml.

Growth curves were generated using the Bioscreen C instrument (Labsystems, Helsinki, Finland) in three independent experiments. Growth measurements were realized in 200-well microtiter plates on 350 µl volumes of a minimal medium containing 20 mM sucrose, inoculated at an OD600 = 0.15 from a washed starter culture. Non-inoculated wells were used as asepsis controls. Optical densities at 600 nm values were measured every 30 min over a period of 2 to 3 days at 28°C. The microplates were shaken for 5 sec before each measurement.

Construction of Xanthomonas campestris pv. campestris mutants

Insertion mutants were constructed using the suicide plasmid pVO155 [36]. Oligonucleotide primers used for PCR amplification will be provided upon request. Amplicons were 300 bp in average. Location of insertions are indicated in Table S6. Deletion mutants in XCC3358, XCC3359, XCC1990, XCC3209 and XCC2927 were constructed using the cre-lox system adapted from Marx and colleagues [37], [38]. Deleted regions are indicated in Table S6.

A fur mutant strain was obtained using the manganese mutagenesis method [106] in LB medium containing 5 mM MnCl2. After incubation for 48 h at 30°C, surviving colonies were harvested for siderophores over-expression on CAS agar plates [107] adapted for Xanthomonas (K2HPO4 1 g/l; MgSO4 1 mM; Casamino acids 0.15 g/l; (NH4)2SO4 1 g/l; sucrose 20 mM; CAS 60.5 mg/l; HDTMA 72.9 mg/l; FeCl3 10 µM). A resistant strain over-expressing siderophores was selected; the fur gene was sequenced and contains a point mutation (T212A leading to L71Q).

Plasmid constructions

The XCC3358, XCC3359 and XCC1470 genes (suxA, suxB and fur respectively, see Figure 6 and Table S6) were amplified by PCR using appropriately designed primers (Oligonucleotide primers used for PCR amplification will be provided upon request). PCR products corresponding to suxA and suxB genes were cloned into pCZ525, a derivative of pSC154 [108], without cyaA' coding sequence. The obtained plasmids were cloned into pLAFR6 [109] to give pL-XCC3358 and pL-XCC3359 respectively. The PCR product corresponding to the fur gene with its promoter and terminator sequences was cloned into pFAJ1700 [110] to give pL-XCC1470.

The XCC3358, XCC1990 and XCC3209 promoter regions (see Table S6) were PCR amplified with appropriately designed primers. These promoter regions were cloned as HindIII-XbaI fragments, into the pCZ750 plasmid, a pFAJ1700 [110] derivative containing the KpnI-AscI lacZ gene from the pCZ367 plasmid [108].

Expression studies

β-galactosidase and β-glucuronidase assays: bacterial cultures in the appropriate medium were harvested at different time points and β-galactosidase and β-glucuronidase assays were performed as previously described [111], [112].

Quantitative RT-PCR (qRT-PCR): a 6 hour bacterial culture in the appropriate medium was harvested at an OD600 = 0.4 to 0.6. RNA were extracted using the Rneasy Mini Kit (QIAGEN). One µg of RNA was treated with RNase-free DNase I (GE-Healthcare) for 15 min at 37°C. After DNase inactivation (10 min at 75°C), RNA were reverse-transcribed by Superscript II (Invitrogen) using random hexamers (Biolabs), for 2 min at 25°C followed by 1 h at 42°C. Quantitative-PCR amplification was performed on Light Cycler (Roche): 10 min 95°C, 1 cycle; 10 sec 95°C, 10 sec 65°C, 20 sec 72°C, 40 to 50 cycles). Experiments were carried out in three independent biological experiments. Oligonucleotide primers used for quantitative-PCR amplification will be provided upon request. As a control for real-time PCR, we used the 16S rRNA as described [113]. The 16S rRNA forward primer (5′-TGACGGTACCCAAAGAATAAGCA-3′) and 16S rRNA reverse primer (5′-ACGCTTGCACCCTTCGTATTA-3′) amplicon was 72 bp in length.

Pathogenicity tests

Pathogenicity tests were conducted on Arabidopsis thaliana Sf-2 ecotype as previously described [114]. Each strain was tested on sets of 4 plants with 4 leaves per plant. Disease development was scored at days 5, 7 and 9 post-inoculation using a disease index ranging from 0 (no symptom), to 4 (leaf death).

[14C]sucrose transport experiments

Overnight cultures in minimal medium (MME) without carbon source (uninduced) or with 20 mM sucrose (induced) were centrifuged. Pellets were resuspended in MME and the OD600 was adjusted to 1. [14C]sucrose (PerkinElmer, specific activity of 21.8 GBq/mmol) was added to a final concentration of 0.5 µM. For competition experiments, sucrose, fructose or glucose was added to [14C]sucrose at a final concentration ranging from 0.5 to 100 µM. After different times, from 20 sec to 2 hours, samples of 0.2 ml were collected on cellulose nitrate filters, washed with 10 ml water, dried, and finally, the radioactivity was determined in a liquid scintillation counter.

The concentration-dependent initial sucrose transport was determined using the rapid dilution method as described [22]. Cells were precultured in minimal medium without sugar. After centrifugation and adjustment to an OD600 of 1, cells were incubated for 15 sec in presence of 0.01, 0.025, 0.05, 0.1, 0.25, 0.5, 1, 2.5 and 5 µM [14C]sucrose and 0.2 ml samples were diluted into 5 ml MME supplemented with 0.1 mM sucrose. Cells were collected by filtration, washed with 10 ml MME supplemented with 10 mM sucrose, dried and the radioactivity was determined.

For inhibition of the proton motive force (PMF) with carbonyl cyanide 3-chlorophenyl-hydrazone (CCCP), cells were incubated for 10 minutes at 30°C with 20 µM CCCP prior to the addition of [14C]sucrose.

In silico analysis

Location of the signal sequence responsible for the outer membrane localization was determined using the SignalP 3.0 server [115] ( with default parameters for Gram-negative bacteria.

Comparison alignments used to identify the TonB-box were realized using ClustalW [64] ( or Multalin [116] (​multalin.html) softwares.

β-sheets in the last 50 amino-acids of Xcc TBDRs were located using the secondary structure prediction method [117] on the PSIPRED Protein Structure Prediction Server [118], [119] (

The MotifSampler program [41], [42] was used to identify a motif corresponding to the Xcc Fux-box upstream of the 9 Fur-repressed TBDR and Ps-TBDR genes. MotifScanner program [42] or the PatScan pattern matcher software [47] were used with the identified motif to locate all the Fur-boxes in the Xcc genome.

Pip boxes (TTCGCN15TTCGC) [120] and hrpII boxes (TTCGN16TTCG) [121] were identified in the Xcc genome using the PatScan software [47]​can/HTML/patscan.html).

For phylogenetic analysis, amino acid sequences were aligned and phylogenetic trees were reconstructed by the neighbor-joining method as implemented in ClustalX [122].

Supporting Information

Figure S1.

Phylogenetic tree of the family of TonB-dependent receptor proteins from Xanthomonas campestris pv. campestris strains ATCC33913 and 8004, Xanthomonas axonopodis pv. citri strain 306, Xanthomonas campestris pv. vesicatoria strain 85–10, Xanthomonas oryzae pv. oryzae strains KACC10331 and MAFF311018 and Xylella fastidiosa strains 9a5c and PD.


(0.04 MB PDF)

Figure S2.

Genome context of genes associated with TonB-dependent receptors present in putative (partial) CUT loci of Xanthomonas campestris pv. campestris (Xcc) and conservation in non related bacteria. Homologous genes are marked by matching colors. White color indicates non conserved genes. For conserved genes, percentages of identity and similarity to the corresponding Xcc gene are indicated beneath.


(0.24 MB PDF)

Table S1.

Regions and domains of Xanthomonas campestris pv. campestris putative TonB-dependent receptors (TBDRs).


(0.04 MB XLS)

Table S2.

Distribution of TonB-dependent receptors in 226 sequenced Gram negative bacterial genomes. The EMBL-EBI, Integr8 web portal (​ and the NCBI ENTREZ Genome Project database (​.fcgi?db = genomeprj) were used to select sequenced bacterial genomes. TonB-dependent receptors (TBDRs) were detected by screening the Pfam database and the UniprotKB protein knowledge database using the two Pfam domains (PF07715, Plug; PF00593, TonB_dep_Rec). Only proteins displaying the two Pfam domains were considered.


(0.08 MB XLS)

Table S3.

Genome context of Xanthomonas campestris pv. campestris (Xcc) TonB-dependent receptor (TBDR) genes and definition of putative CUT loci, conservation in Xanthomonads or in other genera, and TBDR conservation in Pseudomonas aeruginosa PAO1.


(0.18 MB XLS)

Table S4.

Differential expression ratios of Xanthomonas campestris pv. campestris TonB-dependent receptor (TBDR) pVO155 insertion mutants.


(0.03 MB XLS)

Table S5.

Conservation of Xanthomonas campestris pv. campestris (Xcc) TonB-dependent receptors (TBDRs) beside Xanthomonads orthologies.


(0.04 MB XLS)

Table S6.

List of plasmids and Xanthomonas campestris pv. campestris strains used or generated in this study.


(0.03 MB XLS)


We would like to thank J. Cullimore for critical comments on the manuscript, J. Gouzy and S. Carrere for help in sequence analysis.

Author Contributions

Conceived and designed the experiments: EL SB DM MA. Performed the experiments: EL SB DM AB ML CG ND JV. Analyzed the data: EL SB DM MA. Contributed reagents/materials/analysis tools: EL SB DM MA. Wrote the paper: EL MA.


  1. 1. Galperin MY (2004) Bacterial signal transduction network in a genomic perspective. Environ Microbiol 6: 552–567.
  2. 2. Cases I, de Lorenzo V (2005) Genetically modified organisms for the environment: stories of success and failure and what we have learned from them. Int Microbiol 8: 213–222.
  3. 3. Papa R, Glagla S, Danchin A, Schweder T, Marino G, et al. (2006) Proteomic identification of a two-component regulatory system in Pseudoalteromonas haloplanktis TAC125. Extremophiles 10: 483–91.
  4. 4. Williams PH (1980) Black rot: a continuing threat to world crucifers. Plant Disease 64: 736–742.
  5. 5. Alvarez AM (2000) Black rot of crucifers. In: AJSe al, editor. Mechanisms of resistance to plant diseases. Dordrecht, Netherlands: Kluwer Academic Publications. pp. 21–52.
  6. 6. Gough CL, Dow JM, Keen J, Henrissat B, Daniels MJ (1990) Nucleotide sequence of the engXCA gene encoding the major endoglucanase of Xanthomonas campestris pv. campestris. Gene 89: 53–59.
  7. 7. Tang JL, Gough CL, Daniels MJ (1990) Cloning of genes involved in negative regulation of production of extracellular enzymes and polysaccharide of Xanthomonas campestris pathovar campestris. Mol Gen Genet 222: 157–160.
  8. 8. Dow JM, Clarke BR, Milligan DE, Tang JL, Daniels MJ (1990) Extracellular proteases from Xanthomonas campestris pv. campestris, the black rot pathogen. Appl Environ Microbiol 56: 2994–2998.
  9. 9. Slater H, Alvarez-Morales A, Barber CE, Daniels MJ, Dow JM (2000) A two-component system involving an HD-GYP domain protein links cell-cell signalling to pathogenicity gene expression in Xanthomonas campestris. Mol Microbiol 38: 986–1003.
  10. 10. Ryan RP, Fouhy Y, Lucey JF, Crossman LC, Spiro S, et al. (2006) Cell-cell signaling in Xanthomonas campestris involves an HD-GYP domain protein that functions in cyclic di-GMP turnover. Proc Natl Acad Sci U S A 103: 6712–6717.
  11. 11. Dow JM, Crossman L, Findlay K, He YQ, Feng JX, et al. (2003) Biofilm dispersal in Xanthomonas campestris is controlled by cell-cell signaling and is required for full virulence to plants. Proc Natl Acad Sci U S A 100: 10995–11000.
  12. 12. 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. Mol Plant Microbe Interact 4: 593–601.
  13. 13. Meyer D, Cunnac S, Gueneron M, Declercq C, Van Gijsegem F, et al. (2006) PopF1 and PopF2, two proteins secreted by the type III protein secretion system of Ralstonia solanacearum, are translocators belonging to the HrpF/NopX family. J Bacteriol 188: 4903–4917.
  14. 14. da Silva ACR, Ferro JA, Reinach FC, Farah CS, Furlan LR, et al. (2002) Comparison of the genomes of two Xanthomonas pathogens with differing host specificities. Nature 417: 459–463.
  15. 15. Qian W, Jia Y, Ren SX, He YQ, Feng JX, et al. (2005) Comparative and functional genomic analyses of the pathogenicity of phytopathogen Xanthomonas campestris pv. campestris. Genome Res 15: 757–767.
  16. 16. Thieme F, Koebnik R, Bekel T, Berger C, Boch J, et al. (2005) Insights into genome plasticity and pathogenicity of the plant pathogenic bacterium Xanthomonas campestris pv. vesicatoria revealed by the complete genome sequence. J Bacteriol 187: 7254–7266.
  17. 17. Lee BM, Park YJ, Park DS, Kang HW, Kim JG, et al. (2005) The genome sequence of Xanthomonas oryzae pathovar oryzae KACC10331, the bacterial blight pathogen of rice. Nucleic Acids Res 33: 577–586.
  18. 18. Ochiai H, Inoue Y, Takeya M, Sasaki A, Kaku H (2005) Genome sequence of Xanthomonas oryzae pv. oryzae suggests contribution of large numbers of effector genes and insertion sequences to its race diversity. Japan Agricultural Research Quarterly 39: 275–287.
  19. 19. Postle K, Kadner RJ (2003) Touch and go: tying TonB to transport. Mol Microbiol 49: 869–882.
  20. 20. Bagg A, Neilands JB (1987) Ferric uptake regulation protein acts as a repressor, employing iron (II) as a cofactor to bind the operator of an iron transport operon in Escherichia coli. Biochemistry 26: 5471–5477.
  21. 21. Ahmer BM, Thomas MG, Larsen RA, Postle K (1995) Characterization of the exbBD operon of Escherichia coli and the role of ExbB and ExbD in TonB function and stability. J Bacteriol 177: 4742–4747.
  22. 22. Neugebauer H, Herrmann C, Kammer W, Schwarz G, Nordheim A, et al. (2005) ExbBD-dependent transport of maltodextrins through the novel MalA protein across the outer membrane of Caulobacter crescentus. J Bacteriol 187: 8300–8311.
  23. 23. Chimento DP, Mohanty AK, Kadner RJ, Wiener MC (2003) Substrate-induced transmembrane signaling in the cobalamin transporter BtuB. Nat Struct Biol 10: 394–401.
  24. 24. Ferguson AD, Chakraborty R, Smith BS, Esser L, van der Helm D, et al. (2002) Structural basis of gating by the outer membrane transporter FecA. Science 295: 1715–1719.
  25. 25. Yue WW, Grizot S, Buchanan SK (2003) Structural evidence for iron-free citrate and ferric citrate binding to the TonB-dependent outer membrane transporter FecA. J Mol Biol 332: 353–368.
  26. 26. Pawelek PD, Croteau N, Ng-Thow-Hing C, Khursigara CM, Moiseeva N, et al. (2006) Structure of TonB in complex with FhuA, E. coli outer membrane receptor. Science 312: 1399–1402.
  27. 27. Shultis DD, Purdy MD, Banchs CN, Wiener MC (2006) Outer membrane active transport: structure of the BtuB:TonB complex. Science 312: 1396–1399.
  28. 28. Bateman A, Coin L, Durbin R, Finn RD, Hollich V, et al. (2004) The Pfam protein families database. Nucleic Acids Res 32: D138–141.
  29. 29. Lundrigan MD, Kadner RJ (1986) Nucleotide sequence of the gene for the ferrienterochelin receptor FepA in Escherichia coli. Homology among outer membrane receptors that interact with TonB. J Biol Chem 261: 10797–10801.
  30. 30. Struyve M, Moons M, Tommassen J (1991) Carboxy-terminal phenylalanine is essential for the correct assembly of a bacterial outer membrane protein. J Mol Biol 218: 141–148.
  31. 31. Koebnik R (1993) Structural organization of TonB-dependent receptors. Trends Microbiol 1: 201.
  32. 32. Braun V (1997) Surface signaling: novel transcription initiation mechanism starting from the cell surface. Arch Microbiol 167: 325–331.
  33. 33. Koebnik R (2005) TonB-dependent trans-envelope signalling: the exception or the rule? Trends Microbiol 13: 343–347.
  34. 34. de Lorenzo V, Wee S, Herrero M, Neilands JB (1987) Operator sequences of the aerobactin operon of plasmid ColV-K30 binding the ferric uptake regulation (fur) repressor. J Bacteriol 169: 2624–2630.
  35. 35. de Lorenzo V, Giovannini F, Herrero M, Neilands JB (1988) Metal ion regulation of gene expression. Fur repressor-operator interaction at the promoter region of the aerobactin system of pColV-K30. J Mol Biol 203: 875–884.
  36. 36. Oke V, Long SR (1999) Bacterial genes induced within the nodule during the Rhizobium-legume symbiosis. Mol Microbiol 32: 837–849.
  37. 37. Marx CJ, Lidstrom ME (2002) Broad-host-range cre-lox system for antibiotic marker recycling in gram-negative bacteria. Biotechniques 33: 1062–1067.
  38. 38. Angot A, Peeters N, Lechner E, Vailleau F, Baud C, et al. (2006) Ralstonia solanacearum requires F-box-like domain-containing type III effectors to promote disease on several host plants. Proc Natl Acad Sci U S A 103: 14620–14625.
  39. 39. Subramoni S, Sonti RV (2005) Growth deficiency of a Xanthomonas oryzae pv. oryzae fur mutant in rice leaves is rescued by ascorbic acid supplementation. Mol Plant Microbe Interact 18: 644–651.
  40. 40. Loprasert S, Sallabhan R, Atichartpongkul S, Mongkolsuk S (1999) Characterization of a ferric uptake regulator (fur) gene from Xanthomonas campestris pv. phaseoli with unusual primary structure, genome organization, and expression patterns. Gene 239: 251–258.
  41. 41. Thijs G, Lescot M, Marchal K, Rombauts S, De Moor B, et al. (2001) A higher-order background model improves the detection of promoter regulatory elements by Gibbs sampling. Bioinformatics 17: 1113–1122.
  42. 42. Thijs G, Marchal K, Lescot M, Rombauts S, De Moor B, et al. (2002) A Gibbs sampling method to detect overrepresented motifs in the upstream regions of coexpressed genes. J Comput Biol 9: 447–464.
  43. 43. Braun V, Mahren S (2005) Transmembrane transcriptional control (surface signalling) of the Escherichia coli Fec type. FEMS Microbiol Rev 29: 673–684.
  44. 44. Escolar L, Perez-Martin J, de Lorenzo V (1998) Binding of the fur (ferric uptake regulator) repressor of Escherichia coli to arrays of the GATAAT sequence. J Mol Biol 283: 537–547.
  45. 45. Wan XF, Verberkmoes NC, McCue LA, Stanek D, Connelly H, et al. (2004) Transcriptomic and proteomic characterization of the Fur modulon in the metal-reducing bacterium Shewanella oneidensis. J Bacteriol 186: 8385–8400.
  46. 46. Baichoo N, Helmann JD (2002) Recognition of DNA by Fur: a reinterpretation of the Fur box consensus sequence. J Bacteriol 184: 5826–5832.
  47. 47. Dsouza M, Larsen N, Overbeek R (1997) Searching for patterns in genomic data. Trends Genet 13: 497–498.
  48. 48. Cartron ML, Maddocks S, Gillingham P, Craven CJ, Andrews SC (2006) Feo–transport of ferrous iron into bacteria. Biometals 19: 143–157.
  49. 49. Mey AR, Wyckoff EE, Kanukurthy V, Fisher CR, Payne SM (2005) Iron and fur regulation in Vibrio cholerae and the role of fur in virulence. Infect Immun 73: 8167–8178.
  50. 50. Brown JS, Gilliland SM, Holden DW (2001) A Streptococcus pneumoniae pathogenicity island encoding an ABC transporter involved in iron uptake and virulence. Mol Microbiol 40: 572–585.
  51. 51. Noel L, Thieme F, Nennstiel D, Bonas U (2001) cDNA-AFLP analysis unravels a genome-wide hrpG-regulon in the plant pathogen Xanthomonas campestris pv. vesicatoria. Mol Microbiol 41: 1271–1281.
  52. 52. Koebnik R, Kruger A, Thieme F, Urban A, Bonas U (2006) Specific binding of the Xanthomonas campestris pv. vesicatoria AraC-type transcriptional activator HrpX to plant-inducible promoter boxes. J Bacteriol 188: 7652–7660.
  53. 53. Tsuge S, Nakayama T, Terashima S, Ochiai H, Furutani A, et al. (2006) Gene involved in transcriptional activation of the hrp regulatory gene hrpG in Xanthomonas oryzae pv. oryzae. J Bacteriol 188: 4158–4162.
  54. 54. Kim HS, Park HJ, Heu S, Jung J (2004) Molecular and functional characterization of a unique sucrose hydrolase from Xanthomonas axonopodis pv. glycines. J Bacteriol 186: 411–418.
  55. 55. Di Girolamo PM, Bradbeer C (1971) Transport of vitamin B 12 in Escherichia coli. J Bacteriol 106: 745–750.
  56. 56. Kadner RJ (1990) Vitamin B12 transport in Escherichia coli: energy coupling between membranes. Mol Microbiol 4: 2027–2033.
  57. 57. Bradbeer C, Woodrow ML (1976) Transport of vitamin B12 in Escherichia coli: energy dependence. J Bacteriol 128: 99–104.
  58. 58. Paquelin A, Ghigo JM, Bertin S, Wandersman C (2001) Characterization of HasB, a Serratia marcescens TonB-like protein specifically involved in the haemophore-dependent haem acquisition system. Mol Microbiol 42: 995–1005.
  59. 59. Seliger SS, Mey AR, Valle AM, Payne SM (2001) The two TonB systems of Vibrio cholerae: redundant and specific functions. Mol Microbiol 39: 801–812.
  60. 60. Huang B, Ru K, Yuan Z, Whitchurch CB, Mattick JS (2004) tonB3 is required for normal twitching motility and extracellular assembly of type IV pili. J Bacteriol 186: 4387–4389.
  61. 61. Zhao Q, Poole K (2002) Mutational analysis of the TonB1 energy coupler of Pseudomonas aeruginosa. J Bacteriol 184: 1503–1513.
  62. 62. Weickert MJ, Adhya S (1992) A family of bacterial regulators homologous to Gal and Lac repressors. J Biol Chem 267: 15869–15874.
  63. 63. Somers E, Keijers V, Ptacek D, Halvorsen Ottoy M, Srinivasan M, et al. (2000) The salCAB operon of Azospirillum irakense, required for growth on salicin, is repressed by SalR, a transcriptional regulator that belongs to the Lacl/GalR family. Mol Gen Genet 263: 1038–1046.
  64. 64. Thompson JD, Higgins DG, Gibson TJ (1994) CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res 22: 4673–4680.
  65. 65. Vauterin L, Hoste B, Kersters K, Swings J (1995) Reclassification of Xanthomonas. International Journal of Systematic Bacteriology 45: 472–489.
  66. 66. Anzai Y, Kim H, Park JY, Wakabayashi H, Oyaizu H (2000) Phylogenetic affiliation of the pseudomonads based on 16S rRNA sequence. Int J Syst Evol Microbiol 50 Pt 4: 1563–1589.
  67. 67. Cubero J, Graham JH (2004) The leucine-responsive regulatory protein (lrp) gene for characterization of the relationship among Xanthomonas species. Int J Syst Evol Microbiol 54: 429–437.
  68. 68. Tanabe T, Funahashi T, Nakao H, Miyoshi S, Shinoda S, et al. (2003) Identification and characterization of genes required for biosynthesis and transport of the siderophore vibrioferrin in Vibrio parahaemolyticus. J Bacteriol 185: 6938–6949.
  69. 69. Hottes AK, Meewan M, Yang D, Arana N, Romero P, et al. (2004) Transcriptional profiling of Caulobacter crescentus during growth on complex and minimal media. J Bacteriol 186: 1448–1461.
  70. 70. Reid SJ, Abratt VR (2005) Sucrose utilisation in bacteria: genetic organisation and regulation. Appl Microbiol Biotechnol 67: 312–321.
  71. 71. Nikaido H (2003) Molecular basis of bacterial outer membrane permeability revisited. Microbiol Mol Biol Rev 67: 593–656.
  72. 72. Llamas MA, Sparrius M, Kloet R, Jimenez CR, Vandenbroucke-Grauls C, et al. (2006) The heterologous siderophores ferrioxamine B and ferrichrome activate signaling pathways in Pseudomonas aeruginosa. J Bacteriol 188: 1882–1891.
  73. 73. Wiggerich HG, Klauke B, Koplin R, Priefer UB, Puhler A (1997) Unusual structure of the tonB-exb DNA region of Xanthomonas campestris pv. campestris: tonB, exbB, and exbD1 are essential for ferric iron uptake, but exbD2 is not. J Bacteriol 179: 7103–7110.
  74. 74. Wiggerich HG, Puhler A (2000) The exbD2 gene as well as the iron-uptake genes tonB, exbB and exbD1 of Xanthomonas campestris pv. campestris are essential for the induction of a hypersensitive response on pepper (Capsicum annuum). Microbiology 146 ( Pt 5): 1053–1060.
  75. 75. Bogs J, Geider K (2000) Molecular analysis of sucrose metabolism of Erwinia amylovora and influence on bacterial virulence. J Bacteriol 182: 5351–5358.
  76. 76. Jahreis K, Lengeler JW (1993) Molecular analysis of two ScrR repressors and of a ScrR-FruR hybrid repressor for sucrose and D-fructose specific regulons from enteric bacteria. Mol Microbiol 9: 195–209.
  77. 77. Schmid K, Schupfner M, Schmitt R (1982) Plasmid-mediated uptake and metabolism of sucrose by Escherichia coli K-12. J Bacteriol 151: 68–76.
  78. 78. Sprenger GA, Lengeler JW (1988) Analysis of sucrose catabolism in Klebsiella pneumoniae and in Scr+ derivatives of Escherichia coli K12. J Gen Microbiol 134: 1635–1644.
  79. 79. Van Gelder P, Dutzler R, Dumas F, Koebnik R, Schirmer T (2001) Sucrose transport through maltoporin mutants of Escherichia coli. Protein Eng 14: 943–948.
  80. 80. Schulein K, Schmid K, Benzl R (1991) The sugar-specific outer membrane channel ScrY contains functional characteristics of general diffusion pores and substrate-specific porins. Mol Microbiol 5: 2233–2241.
  81. 81. Puhler A, Arlat M, Becker A, Gottfert M, Morrissey JP, et al. (2004) What can bacterial genome research teach us about bacteria-plant interactions? Curr Opin Plant Biol 7: 137–147.
  82. 82. Lima WC, Van Sluys MA, Menck CF (2005) Non-gamma-proteobacteria gene islands contribute to the Xanthomonas genome. Omics 9: 160–172.
  83. 83. Hashidoko Y (2005) Ecochemical studies of interrelationships between epiphytic bacteria and host plants via secondary metabolites. Biosci Biotechnol Biochem 69: 1427–1441.
  84. 84. Hashimoto W, Momma K, Maruyama Y, Yamasaki M, Mikami B, et al. (2005) Structure and function of bacterial super-biosystem responsible for import and depolymerization of macromolecules. Biosci Biotechnol Biochem 69: 673–692.
  85. 85. Hashimoto W, He J, Wada Y, Nankai H, Mikami B, et al. (2005) Proteomics-based identification of outer-membrane proteins responsible for import of macromolecules in Sphingomonas sp. A1: alginate-binding flagellin on the cell surface. Biochemistry 44: 13783–13794.
  86. 86. Aso Y, Miyamoto Y, Harada KM, Momma K, Kawai S, et al. (2006) Engineered membrane superchannel improves bioremediation potential of dioxin-degrading bacteria. Nat Biotechnol 24: 188–189.
  87. 87. Nierman WC, Feldblyum TV, Laub MT, Paulsen IT, Nelson KE, et al. (2001) Complete genome sequence of Caulobacter crescentus. Proc Natl Acad Sci U S A 98: 4136–4141.
  88. 88. Phadke ND, Molloy MP, Steinhoff SA, Ulintz PJ, Andrews PC, et al. (2001) Analysis of the outer membrane proteome of Caulobacter crescentus by two-dimensional electrophoresis and mass spectrometry. Proteomics 1: 705–720.
  89. 89. Molloy MP, Phadke ND, Chen H, Tyldesley R, Garfin DE, et al. (2002) Profiling the alkaline membrane proteome of Caulobacter crescentus with two-dimensional electrophoresis and mass spectrometry. Proteomics 2: 899–910.
  90. 90. Ireland MM, Karty JA, Quardokus EM, Reilly JP, Brun YV (2002) Proteomic analysis of the Caulobacter crescentus stalk indicates competence for nutrient uptake. Mol Microbiol 45: 1029–1041.
  91. 91. Wilson M, Lindow SE (1994) Coexistence among Epiphytic Bacterial Populations Mediated through Nutritional Resource Partitioning. Appl Environ Microbiol 60: 4468–4477.
  92. 92. Mercier J, Lindow SE (2000) Role of leaf surface sugars in colonization of plants by bacterial epiphytes. Appl Environ Microbiol 66: 369–374.
  93. 93. Leveau JH, Lindow SE (2001) Appetite of an epiphyte: quantitative monitoring of bacterial sugar consumption in the phyllosphere. Proc Natl Acad Sci U S A 98: 3446–3453.
  94. 94. Aldon D, Brito B, Boucher C, Genin S (2000) A bacterial sensor of plant cell contact controls the transcriptional induction of Ralstonia solanacearum pathogenicity genes. Embo J 19: 2304–2314.
  95. 95. Marenda M, Brito B, Callard D, Genin S, Barberis P, et al. (1998) PrhA controls a novel regulatory pathway required for the specific induction of Ralstonia solanacearum hrp genes in the presence of plant cells. Mol Microbiol 27: 437–453.
  96. 96. Brito B, Aldon D, Barberis P, Boucher C, Genin S (2002) A signal transfer system through three compartments transduces the plant cell contact-dependent signal controlling Ralstonia solanacearum hrp genes. Mol Plant Microbe Interact 15: 109–119.
  97. 97. Zou LF, Wang XP, Xiang Y, Zhang B, Li YR, et al. (2006) Elucidation of the hrp clusters of Xanthomonas oryzae pv. oryzicola that control the hypersensitive response in nonhost tobacco and pathogenicity in susceptible host rice. Appl Environ Microbiol 72: 6212–6224.
  98. 98. Faure D, Saier MH Jr, Vanderleyden J (2001) An evolutionary alternative system for aryl beta-glucosides assimilation in bacteria. J Mol Microbiol Biotechnol 3: 467–470.
  99. 99. Bauer M, Kube M, Teeling H, Richter M, Lombardot T, et al. (2006) Whole genome analysis of the marine Bacteroidetes ‘Gramella forsetii’ reveals adaptations to degradation of polymeric organic matter. Environ Microbiol 8: 2201–2213.
  100. 100. Backhed F, Ley RE, Sonnenburg JL, Peterson DA, Gordon JI (2005) Host-bacterial mutualism in the human intestine. Science 307: 1915–1920.
  101. 101. Sonnenburg JL, Xu J, Leip DD, Chen CH, Westover BP, et al. (2005) Glycan foraging in vivo by an intestine-adapted bacterial symbiont. Science 307: 1955–1959.
  102. 102. Xu J, Gordon JI (2003) Inaugural Article: Honor thy symbionts. Proc Natl Acad Sci U S A 100: 10452–10459.
  103. 103. Shipman JA, Berleman JE, Salyers AA (2000) Characterization of four outer membrane proteins involved in binding starch to the cell surface of Bacteroides thetaiotaomicron. J Bacteriol 182: 5365–5372.
  104. 104. Xu J, Bjursell MK, Himrod J, Deng S, Carmichael LK, et al. (2003) A genomic view of the human-Bacteroides thetaiotaomicron symbiosis. Science 299: 2074–2076.
  105. 105. Miller J (1972) Experiments in Molecular Genetics. Cold Spring Harbor, N.Y.: Cold Spring Harbor Laboratory Press.
  106. 106. Hantke K (1987) Selection procedure for deregulated iron transport mutants (fur) in Escherichia coli K 12: fur not only affects iron metabolism. Mol Gen Genet 210: 135–139.
  107. 107. Schwyn B, Neilands JB (1987) Universal chemical assay for the detection and determination of siderophores. Anal Biochem 160: 47–56.
  108. 108. Cunnac S, Occhialini A, Barberis P, Boucher C, Genin S (2004) Inventory and functional analysis of the large Hrp regulon in Ralstonia solanacearum: identification of novel effector proteins translocated to plant host cells through the type III secretion system. Mol Microbiol 53: 115–128.
  109. 109. Huynh TV, Dahlbeck D, Staskawicz BJ (1989) Bacterial blight of soybean: regulation of a pathogen gene determining host cultivar specificity. Science 245: 1374–1377.
  110. 110. 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. Mol Plant Microbe Interact 14: 426–430.
  111. 111. Miller J (1992) A Short Course in Bacterial Genetics. A Laboratory Manual and Handbook for Escherichia coliand Related Bacteria. Cold Spring Harbor, N.Y.: Cold Spring Harbor Laboratory Press.
  112. 112. Jefferson RA, Kavanagh TA, Bevan MW (1987) GUS fusions: beta-glucuronidase as a sensitive and versatile gene fusion marker in higher plants. Embo J 6: 3901–3907.
  113. 113. Morales CQ, Posada J, Macneale E, Franklin D, Rivas I, et al. (2005) Functional analysis of the early chlorosis factor gene. Mol Plant Microbe Interact 18: 477–486.
  114. 114. 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.
  115. 115. Bendtsen JD, Nielsen H, von Heijne G, Brunak S (2004) Improved prediction of signal peptides: SignalP 3.0. J Mol Biol 340: 783–795.
  116. 116. Corpet F (1988) Multiple sequence alignment with hierarchical clustering. Nucleic Acids Res 16: 10881–10890.
  117. 117. Jones DT (1999) Protein secondary structure prediction based on position-specific scoring matrices. J Mol Biol 292: 195–202.
  118. 118. Bryson K, McGuffin LJ, Marsden RL, Ward JJ, Sodhi JS, et al. (2005) Protein structure prediction servers at University College London. Nucleic Acids Res 33: W36–38.
  119. 119. McGuffin LJ, Bryson K, Jones DT (2000) The PSIPRED protein structure prediction server. Bioinformatics 16: 404–405.
  120. 120. Fenselau S, Bonas U (1995) Sequence and expression analysis of the hrpB pathogenicity operon of Xanthomonas campestris pv. vesicatoria which encodes eight proteins with similarity to components of the Hrp, Ysc, Spa, and Fli secretion systems. Mol Plant Microbe Interact 8: 845–854.
  121. 121. Cunnac S, Boucher C, Genin S (2004) Characterization of the cis-acting regulatory element controlling HrpB-mediated activation of the type III secretion system and effector genes in Ralstonia solanacearum. J Bacteriol 186: 2309–2318.
  122. 122. Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F, Higgins DG (1997) The CLUSTAL_X windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res 25: 4876–4882.
  123. 123. Crooks GE, Hon G, Chandonia JM, Brenner SE (2004) WebLogo: a sequence logo generator. Genome Res 14: 1188–1190.