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. 2009 Feb 23;10(3):389–401. doi: 10.1111/j.1364-3703.2009.00539.x

Characterization of a putative endoxylanase in the migratory plant‐parasitic nematode Radopholus similis

ANNELIES HAEGEMAN 1, BARTEL VANHOLME 2, GODELIEVE GHEYSEN 1,✉
PMCID: PMC6640231  PMID: 19400841

SUMMARY

Plant‐parasitic nematodes have developed an arsenal of enzymes to degrade the rigid plant cell wall. In this article, we report the presence of a putative endoxylanase in the migratory endoparasitic nematode Radopholus similis. This enzyme is thought to facilitate the migration of the nematode, as it breaks down xylan, the major component of hemicellulose. The corresponding gene (Rs‐xyl1) was cloned and the sequence revealed three small introns. Interestingly, the position of all three introns was conserved in a putative endoxylanase from Meloidogyne hapla, and the position of one intron was conserved in two endoxylanases from Meloidogyne incognita, which suggests a common ancestral gene. The spatial and temporal expression of the Rs‐xyl1 gene was examined by in situ hybridization and semi‐quantitative reverse transcriptase‐polymerase chain reaction. The putative protein consists of a signal peptide, a catalytic domain and a carbohydrate‐binding module (CBM). The catalytic domain showed similarity to both glycosyl hydrolase family 5 (GHF5) and GHF30 enzymes. Using Hidden Markov Model profiles and phylogenetic analysis, we were able to show that Rs‐XYL1 and its closest homologues are not members of GHF5, as suggested previously, but rather form a subclass within GHF30. Silencing the putative endoxylanase by double‐stranded RNA targeting of the CBM region resulted in an average decrease in infection of 60%, indicating that the gene is important for the nematode to complete its life cycle.

INTRODUCTION

During evolution, nematodes adapted to a wide range of environmental conditions. A subset of nematode species evolved into effective plant parasites with various life styles. They all developed a stylet, a hollow needle‐like structure that is used to withdraw the contents of plant cells on which they feed. In addition to a role in feeding, the stylet is used to mechanically damage the rigid cell wall, as well as to inject enzymes into the plant tissue to weaken the plant cell wall during migration. These cell wall‐degrading enzymes are produced in large pharyngeal glands located in the anterior half of the nematode's body. The most extensively studied nematode cell wall‐degrading enzymes, endo‐1,4‐β‐glucanases, have been identified in both sedentary (Rosso et al., 1999; Smant et al., 1998) and migratory (Haegeman et al., 2008; Kikuchi et al., 2004; Uehara et al., 2001) species. Several other cell wall‐degrading enzymes have been identified, such as pectate lyase (de Boer et al., 2002; Doyle and Lambert, 2002; Kikuchi et al., 2006; Vanholme et al., 2007), polygalacturonase (Jaubert et al., 2002) and recently a xylanase (Mitreva‐Dautova et al., 2006). The latter enzymes have only been found in sedentary nematodes, with the exception of pectate lyase, which was also identified in the migratory nematode Bursaphelenchus xylophilus (Kikuchi et al., 2006).

Xylan is composed of β‐1,4‐linked xylopyranose units and can have various substituents and variable structures according to the plant species (Collins et al., 2005). It is the major component of hemicellulose, a complex of polymeric carbohydrates in plant cell walls, and, next to cellulose, the second most abundant polysaccharide in nature. Xylan is found at the interface between lignin and cellulose, where it plays a role in fibre cohesion and plant cell wall integrity. In monocotyledons, it is located in the primary cell wall, whereas, in dicotyledons, it is the major constituent of the secondary cell wall. Endo‐1,4‐β‐xylanases (EC 3.2.1.8) depolymerize the non‐hydrolysed xylan polymer by the random cleavage of the xylan backbone (Subramaniyan and Prema, 2002). These enzymes are mainly produced by plant pathogenic bacteria and fungi, as well as by endosymbionts located in the digestive tracts of various wood‐boring insects (Brennan et al., 2004). Glycosyl hydrolases are classified into different families according to their sequence similarity (Henrissat and Bairoch, 1996). Endo‐1,4‐β‐xylanases are mostly classified in glycosyl hydrolase families 10 and 11 (GHF10 and GHF11), but some have also been designated to GHF5, GHF7, GHF8, GHF16, GHF26, GHF43, GHF51 and GHF62 (Collins et al., 2005) (Carbohydrate Active Enzymes website http://www.cazy.org) (Coutinho and Henrissat, 1999). To our knowledge, only three endoxylanases have been found in animals: one in Gastropoda (Ampullaria crossean AAY46801, GHF10), one in Insecta (Phaedon cochleariae CAA76932, GHF11) and one in Nematoda (Meloidogyne incognita AAF37276, GHF5). The nematode endoxylanase, Mi‐XYL1, has similarity to a range of bacterial endoxylanases. Most of these have been designated as GHF5, although there is some confusion about this classification as the proteins also show similarity to GHF30 enzymes. Originally, the first enzyme of this group was classified into a new family located between GHF5 and GHF30 (Keen et al., 1996). Other authors also suggested that they should be classified into a new family (Hurlbert and Preston, 2001; Suzuki et al., 1997). However, since the first crystallographic structure of one of these xylanases was unravelled, they have been inconsistently classified as GHF5 (John et al., 2006; Larson et al., 2003; Mitreva‐Dautova et al., 2006).

Radopholus similis, or the burrowing nematode, is a (sub)tropical migratory endoparasitic nematode which belongs to the family Pratylenchidae. It has many different host plants, both dicotyledons and monocotyledons, with banana being the most economically important crop. Radopholus similis infection causes bad anchorage of the banana root system as a result of necrosis of the root cortex, which can lead to severe toppling (Fogain, 2000). Juveniles sometimes feed ectoparasitically on plant root hairs, whereas adults generally feed and migrate within the root tissue. As opposed to females, adult males do not feed and have a degenerate stylet and reduced pharyngeal glands, questioning their parasitic status (Duncan and Moens, 2006; Trinh et al., 2004). Only recently, some molecular information has become available about this widespread and economically important nematode species. The first described cell wall‐degrading enzymes of R. similis were four different endo‐1,4‐β‐glucanases (Haegeman et al., 2008). A recent expressed sequence tag (EST) study revealed some other interesting genes, including an EST with homology to an endo‐1,4‐β‐xylanase (Jacob et al., 2008). In this study, we report the further characterization of the corresponding gene. It is the first putative endoxylanase in a migratory plant‐parasitic nematode; moreover, it is the first characterized animal endoxylanase that includes a putative carbohydrate‐binding module (CBM).

RESULTS

Putative endoxylanases in R. similis and Meloidogyne species

The xylan activity assay on nematode homogenate from R. similis showed a clear halo, which was not present in the negative control, indicating that R. similis contains active enzymes that are able to break down xylan (Fig. 1). Screening over 7000 ESTs of R. similis revealed a tag of 521 bp with homology to an endoxylanase of Aeromonas punctata (blastx E‐value = 2e−34) (Jacob et al., 2008). Based on this sequence, primers were constructed to clone both the genomic (1337 bp) and coding (1203 bp) sequences of the corresponding gene. The gene was named Rs‐xyl1 and was submitted to the GenBank database (EU190885). Comparing the genomic DNA with the cDNA revealed three short introns (43, 44 and 47 bp). The first two introns, in phases 2 and 0, respectively, contain the consensus GT/AG splice site (Blumenthal and Steward, 1997), whereas the third intron (phase 0) has the alternative splice site GC/AG. The coding sequence has an overall GC percentage of 61%, with GC1 and GC2 each 49% and GC3 85%. The introns have a GC percentage of 60%.

Figure 1.

Figure 1

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Xylan activity assay. Left: nematode homogenate of Radopholus similis; right: negative control.

Nematode endoxylanases have been described previously only for the sedentary nematode M. incognita (Mitreva‐Dautova et al., 2006) and, in the recently published genome of M. incognita, six endoxylanases were identified (Abad et al., 2008). As, to date, only coding sequences are publicly available for these genes and we were interested in the number and specific position of introns, primers were constructed to isolate the corresponding genomic clones. Two genomic fragments of 1503 and 1357 bp were amplified from M. incognita, and were named Mi‐xyl2 (EU475875) and Mi‐xyl3 (EU475876), respectively. Their deduced coding sequences showed minor differences to the original cDNA sequence of Mi‐xyl1 as presented by Mitreva‐Dautova et al. (2006) (8 bp in the case of Mi‐xyl2 and 15 bp in the case of Mi‐xyl3). These isolated genomic sequences are probably other copies of the gene in the genome, or allelic variants. Both genomic sequences contain a single intron in phase 0, holding the canonical GT/AG donor/acceptor motif. The intron of Mi‐xyl2 is 642 bp, whereas that of Mi‐xyl3 is shorter (496 bp). The introns only show similarity at the extensions: approximately 80 nucleotides at the 5′ end of the introns and 120 nucleotides at the 3′ end. Interestingly, both introns were located at the same position as the second intron of Rs‐xyl1.

A tblastn with Rs‐XYL1 against the genome of Meloidogyne hapla revealed one contig (contig 2188) with significant similarity (E‐value = 4e−24). This putative endoxylanase of M. hapla includes the putative CBM and contains three predicted introns of 148, 214 and 275 bp, all with a GT/AG splice site. The introns are located at exactly the same positions as the three introns from Rs‐xyl1.

Protein properties and structure

The putative protein sequence of 400 amino acids, called Rs‐XYL1, contains a putative 21‐amino‐acid signal peptide for secretion. The predicted molecular mass of the mature endoxylanase is 40 kDa. NetNGlyc predictions revealed one putative N‐glycosylation site at position 270 of the mature protein.

A search for protein domains resulted in hits for a glucosylceramidase of GHF30 (EC 3.2.1.45) (E‐value = 3.3e−07) and, with low support, a CBM of family II (E‐value = 3.7e+02). Alternatively, the CATH database (Pearl et al., 2003) was searched, and high support was found for a glycoside hydrolase domain (E‐value = 1.4e−68). Thus, the protein consists of an N‐terminal signal peptide for secretion of 21 amino acids, and two putative modules: a glycoside hydrolase catalytic domain of 274 amino acids and a C‐terminal CBM of 90 amino acids. This putative endoxylanase is the first animal xylanase found with a putative CBM.

The putative endoxylanase found in the M. hapla genome has a predicted signal peptide of 24 amino acids, a catalytic domain of 272 amino acids and a putative CBM of 91 amino acids. One possible N‐glycosylation site is present. The protein sequence deduced from Mi‐xyl2 of M. incognita is identical to Mi‐XYL1, whereas Mi‐XYL3 differs in three amino acids from Mi‐XYL1. An alignment of Rs‐XYL1 with the endoxylanase sequences of M. incognita and M. hapla is shown in Fig. 2.

Figure 2.

Figure 2

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Alignment of Rs‐XYL1 with Mi‐XYL1, the isolated sequences Mi‐XYL2 and Mi‐XYL3 and the putative Mh‐XYL. The locations of the signal peptide, catalytic domain, the putative glycosyl hydrolase family 5 (GHF5) signature and the putative carbohydrate‐binding module (CBM) are indicated. Stars indicate the two catalytic residues. The three arrows indicate the positions of the introns in Rs‐XYL1 and Mh‐XYL. Possible N‐glycosylation sites are boxed. The threshold for shading is 100%.

A three‐dimensional model was created using a xylanase from Erwinia chrysanthemi as template [Protein Data Bank (PDB) identifier 1NOF] (Fig. 3). As the proteins are 45% identical and 75% similar, the simplest homology modelling scenario could be used. The α‐ and β‐chains of the catalytic domain fold into an (α/β)8‐barrel structure (indicated in green in Fig. 3), whereas eight consecutive β‐strands of the CBM form a β‐sandwich (indicated in yellow in Fig. 3). One β‐strand at the N‐terminal end of the mature protein (13 amino acids) contributes to the C‐terminal β‐sandwich (indicated in purple in Fig. 3).

Figure 3.

Figure 3

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Three‐dimensional model of the putative endoxylanase Rs‐XYL1 from Radopholus similis, based on the three‐dimensional model of the Erwinia chrysanthemi endoxylanase 1NOF (top, front view; bottom, side view). The catalytic domain is indicated in green and the carbohydrate‐binding module (CBM) in yellow. The β‐strand at the N‐terminal end of the protein, which contributes to the CBM, is shown in purple. The catalytic residues (Glu153 and Glu240) are shown in red and the possible N‐glycosylation site (Asn291) is indicated in blue and with a star.

Database searches and phylogenetic analysis

A blastp search was performed using the catalytic domain as query. There were two groups of blast results. The first group included the top hits (E‐value between 1e−73 and 1e−22), and consisted of 24 bacterial sequences and the nematode endoxylanases. Both the literature and CAZy website (http://www.cazy.org) classified most of these proteins as GHF5 enzymes, although they all showed higher similarity to GHF30 enzymes according to the Conserved Domain Database (CDD) (E‐value < 1e−3). Similarly, HMMer searches (HMM, Hidden Markov Model), using both the GHF5 and GHF30 profiles, classified all proteins (including Rs‐XYL1) as GHF30, except for Mi‐XYL1. No significant matches were found when using the GHF5 profile. These observations question the classification of the proteins to the GHF5 subclass. The second group of blast hits, which show less homology to Rs‐XYL1 (E‐value between 1e−14 and 1e−4), appeared to be GHF30 enzymes from both bacteria and fungi, mostly named glycosyl hydrolase or glucosylceramidase. It should, however, be noted that only two of these hits are classified as GHF30 according to the CAZy database. The others have not been classified at all, although they all have a significant match to the GHF30 profile.

A phylogenetic tree, including the two groups of blast hits and some GHF5 enzymes, revealed a clear separation of two clusters, highly supported with a posterior probability value of 1.00 (Fig. 4). The first cluster includes the ‘true’ GHF5 enzymes, whereas the second cluster includes all proteins which showed homology to the catalytic domain of Rs‐XYL1. From now on, we will call the first cluster the ‘GHF5 cluster’ and the second cluster the ‘GHF30 cluster’. Within the GHF30 cluster, a first group is separated from the other proteins with high support (posterior probability of 1.00). This group consists of all fungal sequences and one bacterial sequence. The rest of the GHF30 cluster is not well resolved (posterior probability values less than 0.60). However, Rs‐XYL1 and all the blast top hits, with very low E‐values, form a monophyletic group. This group is not well resolved from the other blast hits with higher E‐values, as the posterior probability is only 0.56. This low posterior probability is mainly a result of the protein ZP_01576533. Discarding this protein from the dataset results in a monophyletic cluster of all other top blast hits with high support (posterior probability of 1.00). Despite the fact that more basal branches within the GHF30 clusters are not well resolved, the GHF30 cluster itself forms a monophyletic group with high support.

Figure 4.

Figure 4

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Phylogenetic tree including the closest homologues of the catalytic domain of Rs‐XYL1. Ten glycosyl hydrolase family 5 (GHF5) enzymes were included and three GHF10 enzymes were chosen as outgroup. The tree was deduced by Bayesian analysis with 106 generations with a sample frequency of 100. Posterior probabilities are shown on the branches. The GHF5 cluster includes true GHF5 enzymes selected from the PFAM seed alignment for GHF5. The lower cluster, indicated as Rs‐XYL1 top blast hits, includes all blast hits from a blastp search with the catalytic domain of Rs‐XYL1 as query with E‐values between 1e−73 and 1e−22. The other sequences within the GHF30 cluster include blastp hits with E‐values between 1e−14 and 1e−4. Fungal and nematode sequences are indicated; all other sequences are bacterial. Stars indicate enzymes that are classified as GHF5 according to the CAZy classification.

A blastp search with the putative CBM resulted in fewer hits, all from bacteria (Table 1). An alignment is shown in Fig. 5. Only the first two hits (both xylanases from A. punctata) have a significant match in CDD and are classified as CBM6 (E‐values = 1e−15 and 7e−10). The other blast hits have no matching domains in CDD, although some are classified in different CBM groups by the CAZy classification (Table 1). A HMMer search was performed with profiles for different CBMs against the different proteins retained by the blast search. HMM profiles for which a significant hit was found (positive bit score) are indicated in Table 1. The closest homologues, from A. punctata, have been assigned to CBM35 according to CAZy. Nevertheless, some homologues belong to CBM13 and CBM22, whereas others have not been assigned to a CBM family at all. A HMMer search with different CBM profiles resulted in even more confusion, as some CBMs showed positive bit scores to different CBM families, for example CBM35 members were also recognized as CBM6 (Table 1). None of the HMM profiles matched the putative CBM of Rs‐XYL1.

Table 1.

Closest homologues to the Rs‐XYL1 carbohydrate‐binding module (CBM), as identified by blastp (E < 1e−4). The species name, accession number and description of the protein are given. The CAZy classification (http://www.cazy.org) and Hidden Markov Model (HMM) profiles that resulted in a significant hit (positive bit score) are indicated.

Species ACC Description blastp E‐value CAZy HMM search
Radopholus similis EU190885 Endo‐1,4‐β‐xylanase — — —
Aeromonas punctata AAB63573 Xylanase D 2.00e−14 CBM35 CBM6, CBM35
Aeromonas punctata BAA13641 Endo‐xylanase 9.00e−14 CBM35 CBM6, CBM35
Clostridium cellulolyticum ZP_01574162 Carbohydrate‐binding family 6 2.00e−12 — —
Bacillus subtilis NP_389697 Hypothetical protein 4.00e−11 — —
Bacillus subtilis Q6YK37 Glucuronoxylanase xynC precursor 5.00e−11 — —
Bacillus amyloliquefaciens YP_001421408 YnfF 7.00e−11 — —
Clostridium acetobutylicum NP_149281 Possible xylan degradation enzyme 3.00e−09 CBM13 CBM13
Clostridium acetobutylicum NP_149282 Possible xylan degradation enzyme 3.00e−08 CBM13 CBM13
Clostridium cellulolyticum ZP_01576533 Cellulosome enzyme, dockerin type I 8.00e−08 — CBM6, CBM35
Ruminococcus albus BAB39494 xynC 8.00e−08 CBM22 CBM4(9)
Ruminococcus albus BAB39495 Xylanase C 3.00e−07 CBM22 CBM4(9)
Clostridium thermocellum YP_001039401 Carbohydrate‐binding family 6 5.00e−07 — CBM6
Cytophaga hutchinsonii YP_678647 CHU large protein, candidate xylanase 7.00e−07 — —
Bacillus subtilis AAN07016 YnfF 2.00e−05 — —
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Figure 5.

Figure 5

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Alignment of the carbohydrate binding module of Rs‐XYL1 with its closest homologues (blastp hits with E‐value < 1e−4), Mh‐XYL and two translated expressed sequence tags (ESTs) from Meloidogyne chitwoodi. Ap, Aeromonas punctata; Ba, Bacillus amyloliquefaciens; Bs, Bacillus subtilis; Ca, Clostridium acetobutylicum; Cc, Clostridium cellulolyticum; Ch, Cytophaga hutchinsonii; Ct, Clostridium thermocellum; Mc, Meloidogyne chitwoodi; Mh, Meloidogyne hapla; Ra, Ruminococcus albus. The threshold for shading is 75%.

All nematode ESTs were searched for homologues to the endoxylanase domains. The top hits (E‐value < 1e−20) of the tblastn search with the catalytic domain were all ESTs derived from the genus Meloidogyne: M. chitwoodi (CB931318, CD418879), M. arenaria (CF357210, CF357155) and M. javanica (CF350477, CF350376). Other hits (E‐value < 1e−4) were from Globodera pallida (BM415401, CV578872) and Caenorhabditis elegans (AU208008, BJ762920, CB388002). The latter show less similarity and are most probably not endoxylanases, but other glycosyl hydrolases and glucosylceramidases, probably involved in glycan degradation and sphingolipid metabolism (EC 3.2.1.45, BRENDA database) (Barthelmes et al., 2007). A tblastn search with the CBM resulted in only two hits (E‐value < 1e−3), both from M. chitwoodi (CB931318, CD418879).

Temporal and spatial expression of Rs‐xyl1

The expression level of Rs‐xyl1 was compared for different life stages by semi‐quantitative reverse transcriptase‐polymerase chain reaction (RT‐PCR) (Fig. 6). In developing embryos and juveniles, no expression of Rs‐xyl1 was detected, whereas males and females revealed approximately similar expression levels. To localize the transcripts, a whole‐mount in situ hybridization was carried out (Fig. 7). The antisense probe showed clear staining in the gland cell area. The sense probe used as negative control showed no staining.

Figure 6.

Figure 6

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Expression of Rs‐xyl1 and actin in developing embryos (27, 30, 33 and 36 cycles), juveniles (30, 33, 36 and 39 cycles), females (21, 24, 27 and 30 cycles) and males (24, 27, 30 and 33 cycles) of Radopholus similis.

Figure 7.

Figure 7

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Whole‐mount in situ hybridization on Radopholus similis: (a) Rs‐xyl1 antisense probe; (b) Rs‐xyl1 sense probe. Scale bar, 20 µm.

Silencing of Rs‐xyl1 by RNA interference (RNAi) and infection tests

Batches of freshly harvested nematodes were soaked in double‐stranded RNA (dsRNA) against green fluorescent protein (dsgfp), water or dsRNA against Rs‐xyl1 (either targeting the catalytic domain or the putative CBM). After soaking, the expression level of Rs‐xyl1 was checked by semi‐quantitative RT‐PCR. Actin was amplified to approximately the same level in all samples.

In one of the three experiments, no differences in the expression level of Rs‐xyl1 could be detected between the different treatments. The corresponding infection experiment showed no significant differences in infection of the soaked nematodes compared with the controls.

In the other two experiments, Rs‐xyl1 showed a lower expression level in the nematodes soaked in dsRNA targeted to the CBM when compared with the controls (dsgfp and water). No silencing of Rs‐xyl1 could be detected in nematodes soaked in dsRNA targeted to the catalytic domain. Both corresponding infection experiments showed a significant (P < 0.01) decrease in the infection (53% and 66%) of nematodes soaked in dsRNA targeted against the CBM of Rs‐xyl1 compared with nematodes soaked in dsgfp or water. Interestingly, nematodes soaked in dsRNA targeting the catalytic domain of Rs‐xyl1 appeared to show a slight decrease in infection, but this was not significantly different from the controls. The data of one of these experiments are graphically represented in Fig. 8.

Figure 8.

Figure 8

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Silencing of Rs‐xyl1. (a) Semi‐quantitative reverse transcriptase‐polymerase chain reaction (RT‐PCR) with primers against actin (30 cycles) and Rs‐xyl1 (36 cycles) on RNA extracted from nematodes soaked in double‐stranded RNA (dsRNA) against Rs‐xyl1 located in the catalytic domain (Rsxyl‐catdom), dsRNA against Rs‐xyl1 located in the putative carbohydrate‐binding module (CBM) (Rsxyl‐cbm), double‐stranded RNA against green fluorescent protein (gfp) and water (– control). (b) Box‐plot of the number of Radopholus similis in the root tissue per plate 10 days after inoculation with soaked nematodes. Eleven plants were used for each treatment. The length of the vertical bar denotes the interquartile range, the middle bar represents the median, and the upper and lower horizontal bars denote the upper and lower quartiles, respectively.

DISCUSSION

During the last decade, different endogenous cell wall‐degrading enzymes have been identified in plant‐parasitic nematodes. These enzymes facilitate the penetration and migration of the nematode through the rigid plant tissue by softening the plant cell walls. Although xylan is an important component of this cell wall, endoxylanases have only been described for the sedentary nematode species Meloidogyne (Mitreva‐Dautova et al., 2006).

In this study, we report the identification and characterization of the first putative endoxylanase (Rs‐xyl1) of a migratory nematode, R. similis. The overall GC content of the coding sequence (61%) is higher than the average value (54%) (Jacob et al., 2008), and is a result of the high GC3 (85% rather than 65%). This high GC3 content was also observed for the secreted endo‐1,4‐β‐glucanases of this species (Haegeman et al., 2008). The gene contains three short introns (less than 50 bp). Interestingly, a putative endoxylanase found in the genome sequence of M. hapla (contig 2188, http://www.hapla.org) revealed three predicted introns at the same position as the introns of Rs‐xyl1, and two isolated genomic endoxylanase sequences of the root‐knot nematode M. incognita showed a single intron at the same position as the second intron of Rs‐xyl1. This indicates that these introns were probably already present in the common ancestor xylanase gene of these species. The phylogenetic analysis confirmed the close phylogenetic relationship of the nematode endoxylanases. The isolated genomic sequences of M. incognita xylanases (Mi‐xyl2, Mi‐xyl3) are very similar to the previously described xylanase Mi‐xyl1 (Mitreva‐Dautova et al., 2006). If these genes are not allelic variants, they probably originated from the same ancestral gene through duplications.

The main difference between the putative endoxylanase of R. similis and the previously identified endoxylanase of M. incognita is the presence of a putative CBM in Rs‐XYL1 (Fig. 2). This putative CBM was also present in the genomic sequence found in the M. hapla genome, and blast searches against the M. incognita genome revealed that it is probably also associated with the six identified endoxylanases in the genome (Abad et al., 2008). Three‐dimensional modelling of the protein revealed that the CBM consists of eight β‐strands, and that the first β‐strand at the N‐terminal end of the mature protein contributes to the β‐sandwich, as is the case for the Erwinia chrysanthemi xylanase 1NOF. The presence of a putative CBM in Rs‐XYL1 is not surprising, because most bacterial endoxylanases are also associated with a CBM. Although difficult to prove, it is a possibility that nematode endoxylanases have originated through the horizontal gene transfer of a bacterial endoxylanase, as also suggested for nematode endoglucanases (Keen and Roberts, 1998). In this view, it is more likely to find an endoxylanase with a CBM in nematodes. The putative endoxylanase found in the genome of M. hapla also includes a putative CBM. A search through the nematode ESTs with the putative CBM of Rs‐XYL1 resulted in two hits from M. chitwoodi. These tags were also retained by a blastx search using the catalytic domain as query. Therefore, it seems that other nematode endoxylanases possibly also have a CBM. Endoxylanases without a CBM, such as Mi‐XYL1, could have lost this domain during evolution.

The amino acid sequence of the catalytic domain showed significant similarity to the M. incognita xylanase Mi‐XYL1 as well as to different bacterial endoxylanases of GHF5 and GHF30. As the closest homologues of the protein have been assigned to GHF5, it is tempting to classify Rs‐XYL1 in the same subclass. However, it should be noted that the GHF5 signature, which consists of a consensus sequence motif of 10 amino acids, shows three mismatches in both Rs‐XYL1 and Mi‐XYL1, and two mismatches in Mh‐XYL and most bacterial endoxylanases that appeared as best hits in our blast search, questioning their classification as GHF5 proteins. Moreover, none of these proteins shows significant similarity to the HMM GHF5 profile, whereas all proteins (except for Mi‐XYL1) have a significant match to the GHF30 profile. The fact that Mi‐XYL1 is not recognized by the HMM profile of GHF30 is probably a result of the limited seed alignment from only three sequences on which the profile has been based. Keen et al. (1996) suggested that this group of endoxylanases should be classified in an intermediate class between GHF5 and GHF30. Unfortunately, this idea never caught on, and the enzymes were later assigned to GHF5, resulting in an inconsistent classification. In the phylogenetic analysis of this study, we were able to show that Rs‐XYL1 and related endoxylanases form a monophyletic cluster with other GHF30 enzymes. We can conclude that the endoxylanase homologues are a subclass of GHF30, rather than GHF5, and that previous classifications of some of these enzymes into GHF5 are incorrect.

In analogy with the difficulties in classifying the catalytic domain, it is not clear to which family the CBM domain belongs. Similarity was found to proteins belonging to different CBM families, and a number of homologues are not properly classified themselves. This implies that the CBM classification needs to be revised for some families.

The expression of Rs‐xyl1 could only be demonstrated in the adult stages of R. similis and not in eggs or juveniles. This suggests that the putative endoxylanase only plays a role in later life stages, which is not what is expected, as both juvenile and adult stages migrate and feed within the root tissue. Although adult males of R. similis do not feed, the expression level of Rs‐xyl1 in males is the same as that in females. This expression pattern differs from the expression of endoglucanases, which show a lower level in males (Haegeman et al., 2008). The expression of Rs‐xyl1 is located in the gland cell area of R. similis, as shown by in situ hybridization (Fig. 7). The fact that the putative protein sequence contains a signal peptide for secretion, and that the gene is expressed in the gland cell area, suggests that the protein is secreted through the stylet of the nematode. Once released into the plant tissue, the putative endoxylanase can break down xylan present in the plant cell wall. The functional importance of the putative endoxylanase was proven by RNAi and infection experiments. When dsRNA targeted to the putative CBM was applied to nematodes, partial silencing of Rs‐xyl1 occurred, whereas no silencing could be observed when dsRNA was based on the catalytic domain. This difference in silencing effect caused by different target regions in the same gene has also been observed for a pectate lyase of Heterodera glycines (Sukno et al., 2007). Moreover, in one of the experiments, no silencing could be detected at all. Therefore, it seems that the silencing effect can vary according to the position of the target sequence of the gene, but also according to the replication of the experiment itself. When Rs‐xyl1 had a lower expression level after treatment with dsRNA targeted to CBM, a significant decrease in infection of, on average, 60% was detected. In the one experiment in which there was no silencing effect, no significant decrease in infection was observed. This suggests that Rs‐xyl1 indeed plays an important role in the infection process. We observed a slight, but statistically insignificant, decrease in infection with the nematodes were soaked in dsRNA targeted to the catalytic domain but, as we did not see a clear reduction in expression, we cannot draw any conclusions for this construct. Nevertheless, this experiment proves that the RNAi technique, so far almost exclusively studied in sedentary nematodes (Lilley et al., 2007), can be successful in reducing the infection of migratory nematodes as well. In one other migratory nematode, B. xylophilus, it has been shown recently that RNAi can effectively reduce gene expression and cause lethality, but effects on plant parasitism were not studied (Park et al., 2008).

A search for other xylanase homologues in nematode ESTs did not reveal any new candidate xylanase genes. Homologues were found in M. chitwoodi, M. arenaria and M. javanica, as also described by Mitreva‐Dautova et al. (2006). Despite the available EST sequence information, endo‐1,4‐β‐xylanases have not been found extensively, in contrast with, for example, endoglucanases. Nevertheless, in this study, we have shown that a putative endoxylanase of a migratory nematode, R. similis, is important in the infection process.

EXPERIMENTAL PROCEDURES

Nematode cultures, DNA extraction, RNA extraction and cDNA synthesis

Radopholus similis was cultured on carrot discs in small Petri dishes, as described by Jacob et al. (2007). Mixed stages were collected by rinsing the Petri dishes with distilled water 6–8 weeks after inoculation. Meloidogyne incognita was cultured on the roots of pea (Pisum sativa). The seeds were sterilized by soaking them for 30 min in sterile distilled water, for 5 min in 100% ethanol, for 15 min in 5% NaOCl and 1% Tween‐20, and afterwards washing them five times in sterile water. The sterile seeds were germinated on KNOP (2.4 mm KNO3, 160 µm MgSO4.7H2O, 1.0 mm Ca(NO3)2.4H2O, 400 µm KH2PO4, 7.0 µm EDTA ferric sodium salt, 0.25 µm ZnSO4.7H2O, 0.058 µm CuSO4.5H2O, 0.043 µm Na2MoO4.2H2O, 0.025 µm CoCl2.6H2O, 8.6 µm H3BO3, 2.9 µm MnCl2.4H2O, 6.8 µm NaCl, 0.56 mm myo‐inositol, 8.1 µm nicotinic acid, 4.9 µm pyridoxine HCl, 30 µm thiamine HCl, 1% sucrose, 0.8% agar; adjusted to pH 6.4) medium and grown at 21 °C. After 2–3 weeks, the root tips were infected with nematodes of the J2 stage and the plants were incubated at 28 °C. About 7 weeks after inoculation, galls with egg masses were dissected and allowed to hatch in sieves in sterile water at 28 °C.

Genomic DNA was isolated from approximately 5000 individuals, as described by Bolla et al. (1988). RNA was extracted with TRIzol (Invitrogen, Carlsbad, CA, USA), according to the manufacturer's instructions. RNA was treated with DNAse (Fermentas, St. Leon‐Rot, Germany) to avoid DNA contamination. First‐strand cDNA was synthesized from 1 µg of RNA in the presence of 4 mm deoxynucleoside triphosphates (dNTPs), 0.5 µm oligodT primer, 10 mm dithiothreitol (DTT), 50 mm Tris‐HCl (pH 8.3), 75 mm KCl, 3 mm MgCl2 and 200 U Superscript II Reverse Transcriptase (Invitrogen). The reaction mixture was incubated for 2 h at 42 °C. For RT‐PCR analysis, RNA was extracted from approximately 100 individuals, and cDNA was obtained and amplified with the SMART PCR cDNA synthesis kit (Clontech, Palo Alto, CA, USA).

Activity assay and cloning of Rs‐xyl1

Approximately 10 000 freshly harvested nematodes were crushed in 0.5 mL TE buffer [10 mm Tris‐HCl, 1.0 mm ethylenediaminetetraacetic acid (EDTA), pH 8.0] to which 0.5 mm DTT was added. The sample was centrifuged for 5 min at 2000 g in a table‐top centrifuge, and 2 µL of the supernatant was loaded onto a 1.5% agar plate containing 0.5% xylan from beechwood (Sigma, St. Louis, MO, USA). After overnight incubation at 37 °C, the plate was stained for 15 min with 0.1% Congo Red and subsequently washed for 15 min with 1.0 m NaCl. Water rinsed from uninfected carrot discs was used as a negative control.

Primers Rs‐xyl‐F and Rs‐xyl‐R (Table 2) constructed on EST EY194441 were used in PCR. The reaction mixture contained 150 ng of DNA as template, 0.5 µm of each primer, 4 mm dNTPs, 10 mm DTT, 50 mm Tris‐HCl (pH 8.3), 75 mm KCl and 1 U of Taq DNA polymerase (Invitrogen). The PCR conditions were as follows: 35 cycles of 1 min at 94 °C, 1 min at 54 °C and 1 min at 72 °C. PCR products were loaded onto a 0.5 × TAE 1.5% agarose gel, and the fragments of interest were gel excised and purified with the QIAquick gel extraction kit (Qiagen, Hilden, Germany). The purified fragments were ligated into pGEM‐T (Promega, Madison, WI, USA), and the ligation mixture was used for transformation into Escherichia coli DH5α cells. Transformed cells were selected on Luria–Bertani (LB) agar plates supplemented with 100 µg/mL carbenicillin. Positive colonies were identified by a colony PCR under standard conditions using SP6 and T7 primers (Table 2). Plasmids were extracted using the Nucleobond AX kit (Machery‐Nagel, Düren, Germany), and the inserts were sequenced at the VIB Genetic Service Facility (VIB‐GSF, Antwerp, Belgium). Based on the sequence obtained, primers were developed for upstream and downstream walking using the Genome Walker Universal Kit (Clontech), according to the manufacturer's instructions, in two successive PCRs. The primers used for upstream walking were Rs‐xyl‐up1 and Rs‐xyl‐up2; those for downstream walking were Rs‐xyl‐down1 and Rs‐xyl‐down2 (Table 2). The fragments of interest were cloned and sequenced as described above. An additional primer set (Rs‐xyl‐start and Rs‐xyl‐stop, Table 2) was developed to amplify the full‐length coding sequence of the putative endoxylanase from a cDNA pool. The resulting cDNA fragment was cloned in pGEM‐T and sequenced as described above.

Table 2.

Primers used for cloning, in situ hybridization, reverse transcriptase‐polymerase chain reaction (RT‐PCR) and RNA interference (RNAi) of Radopholus similis and Meloidogyne incognita endoxylanases (Invitrogen).

Primer Sequence
Rs‐xyl‐F AAGGGCAAGCAGGTGTGGATG
Rs‐xyl‐R TGATGCTCAGTTGGAGACGAAG
Rs‐xyl‐up1 CATCATACTGGTCCAGTCGAAGC
Rs‐xyl‐up2 TAGTGCTCCGTCATCCACACCTGC
Rs‐xyl‐down1 AAGAGCGGCGCATTCACCTACTC
Rs‐xyl‐down2 CTTCGTCTCCAACTGAGCATCA
Rs‐xyl‐start ATGTTCGCCCTTCTCGTTCCTG
Rs‐xyl‐stop TCAGTTGGAGACGAAGGTGGTGACG
ACT‐F GAAAGAGGGCCGGAAGAG
ACT‐R AGATCGTCCGCGACATAAAG
Mi‐xyl‐F TTTGGTGGTTCTAGTGCTTGG
Mi‐xyl‐R TGTTGTTGGATTTTCAGTAGCA
SP6 ATTTAGGTGACACTATAGAATACTCAAGC
T7 TAATACGACTCACTATAGGGCGAATTGG
OligodT TTTTTTTTTTTTTTTTTTTTTTTTTVN
Rs‐xyl‐catdomF CCGCCTCGATGAAGTCCAACAAC
Rs‐xyl‐catdomR GGGTCAGTGAGTGCCTTGTTG
Rs‐xyl‐CBMF CCAACGTGTTTCTGAGCGCCTAC
Rs‐xyl‐CBMR TCAGTTGGAGACGAAGGTGGTGAC
GFP‐F ATCCGCCACAACATCGAGG
GFP‐R TTGTACAGCTCGTCCATGC
Rs‐xyl‐F2 GGTATGCAGTTGGCTCTGGT
Rs‐xyl‐R2 GACACAACGCTGACAGTTGG
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Primers Mi‐xyl‐F and Mi‐xyl‐R (Table 2) were developed based on the endoxylanase coding sequence of M. incognita (AF224342). These primers were used in a standard PCR reaction on DNA of M. incognita, and the resulting fragments were cloned and sequenced as described above.

Sequence analysis

To identify introns, cDNA and genomic sequences were compared with ClustalW (Thompson et al., 1994) using default parameters. The overall GC content and the GC content of the first, second and third positions of the codons (GC1, GC2 and GC3) were calculated by an in‐house Perl program. The cDNA sequence was translated to protein using the emboss program ‘Transeq’ (Rice et al., 2000). The molecular weight of the putative protein was calculated with the ‘pI/Mw’ tool on the ExPASy Proteomics server (Gasteiger et al., 2003). The presence of signal peptides was predicted with SignalP 3.0 configured for eukaryotic sequences (Bendtsen et al., 2004), and N‐glycosylation sites were predicted by NetNGlyc (http://www.cbs.dtu.dk/services/NetNGlyc/). Tertiary structure predictions were performed in swiss‐model (Guex and Peitsch, 1997) using the first approach mode and the crystal structure of a xylanase of Erwinia chrysanthemi (PDB entry 1NOF) (Larson et al., 2003) as template (E‐value = 2.0e−81). The PDB coordinates obtained were visualized using Deepview spdbv 3.7 (Swiss‐PdbViewer) and rendered with POV‐Ray version 3.6. Because of a lack of similarity to 1NOF, the first 12 amino acids of the mature protein were not modelled by the first approach mode, and these were manually fitted into the model. To identify possible domains in the putative protein sequence of the endoxylanase, a domain search was performed with InterProScan (Zdobnov and Apweiler, 2001). A blastp search with default settings was performed with the catalytic domain and the putative CBM (September 2008). Sequences of the closest homologues were retrieved from GenBank and a protein alignment was constructed with the ClustalW algorithm (Thompson et al., 1994) in BioEdit 7.0.5.3 (Hall, 1999). A local HMM search was performed with HMMER 2.3.2 (http://hmmer.janelia.org) against the closest homologues of the Rs‐XYL1 catalytic domain with profiles from GHF5 and GHF30 downloaded from the PFAM website (Finn et al., 2006). The CAZy website (http://www.cazy.org) was searched to find to which CBMs the closest homologues of the Rs‐XYL1 CBM could be assigned. A local HMMer search was performed against the closest homologues of the CBM with the following CBM profiles downloaded from the PFAM website: CBM4_9 (PF02018), CBM6 (PF03422), CBM13 (PF00652). The HMM profile of CBM35 was constructed locally from raw sequences. To identify homologous sequences in other nematode species, tblastn searches with the catalytic domain and the CBM were performed against all nematode ESTs. The nematode genome of M. hapla was searched for putative endoxylanases by tblastn with Rs‐XYL1 as query on the website of the genome project (http://www.hapla.org) (Opperman et al., 2008).

A phylogenetic tree was constructed including the 100 best hits from a blastp search with the catalytic domain. Ten protein sequences which belong to the PFAM seed alignment from GHF5 were selected, as diverse as possible (both mannanases and endoglucanases from bacteria and fungi). Three protein sequences belonging to GHF10 were chosen as outgroup. The phylogenetic analysis was performed by Bayesian statistics using the WAG‐model in MrBayes 3.1.2 (Ronquist and Huelsenbeck, 2003), with 106 generations and a sample frequency of 100. The first 2500 generations were considered as burn‐in. The resulting tree was visualized in TreeView 1.6.6 (Page, 1996).

Spatial and temporal expression of Rs‐xyl1

An in situ hybridization was performed according to Jacob et al. (2007). Probes were generated by a linear PCR with a single primer (Rs‐xyl‐F, sense probe; Rs‐xyl‐R, antisense probe) in the presence of digoxigenin (DIG)‐labelled oligonucleotides (Roche, Mannheim, Germany). As template for the probe generating PCR, the product of a first PCR was used. This PCR was performed on a plasmid containing the corresponding xylanase cDNA fragment with primers Rs‐xyl‐F and Rs‐xyl‐R. All PCRs were carried out as described above.

A semi‐quantitative RT‐PCR on different life stages was performed according to Haegeman et al. (2008). The primers used were Rs‐xyl‐F and Rs‐xyl‐R and, as a positive control, actin (EU000540) was amplified using the primers ACT‐F and ACT‐R (Table 2).

Generation of dsRNA, soaking, infection tests and semi‐quantitative RT‐PCR

Two different regions of Rs‐xyl1 were selected as target for silencing by RNAi, one in the catalytic domain (286 bp) and one in the putative CBM (242 bp) of Rs‐xyl1. These regions were amplified by PCR under standard conditions with the T7 promoter sequence incorporated at the 5′ end of either the sense or antisense strand. Primers used for the first region were Rs‐xyl‐catdomF and Rs‐xyl‐catdomR, and primers for the second region were Rs‐xyl‐CBMF and Rs‐xyl‐CBMR. dsRNA against gfp was prepared from a gfp‐containing construct using primers GFP‐F and GFP‐R. PCR products were employed as templates for in vitro transcription reactions using a Megascript RNAi kit (Ambion, Huntingdon, Cambridgeshire, UK). The dsRNA was quantified spectrophotometrically. A few thousand nematodes of mixed stages were soaked in 50 mm octopamine, 3 mm spermidine, 0.05% gelatin and 0.5 mg/mL dsRNA for 24 h at room temperature on a rotator in the dark. As an extra negative control, nematodes were incubated in the same solution, but without dsRNA. After soaking, the nematodes were sterilized for 1 h in 0.33% hospital antiseptic concentrate (HAC) on a rotator at room temperature and subsequently washed three times with sterile water. Seedlings from Medicago truncatula L. var. Jemalong were grown in six‐well plates on modified Strullu Romand medium (Elsen et al., 2000). After 3 weeks, the seedlings were infected with 50 sterilized soaked nematodes per plant, 11 plants for each condition. The plants were grown at 22 °C and light/dark cycles of 16 h/8 h. Ten days later, the roots were stained with acid fuchsin (Byrd et al., 1983), destained using acidified glycerol and the nematodes inside the roots were counted using a dissection microscope. The results were statistically analysed in S‐PLUS 7.0. Normality was checked with the Kolmogorov–Smirnov test, homoscedasticity with a modified Levene test and analysis of variance (anova) was performed using the Tukey method. The soaking and infection tests were repeated three times. For semi‐quantitative RT‐PCR, approximately 5000 soaked nematodes were used. RNA extraction and cDNA synthesis were performed as described above. The primers ACT‐F and ACT‐R were used to amplify actin, whereas Rs‐xyl‐F2 and Rs‐xyl‐R2 were used to amplify Rs‐xyl1. The amount of cDNA added as template and the number of PCR cycles were optimized for each gene product to detect the exponential phase of the reaction. The resulting products were separated on a 0.5 × TAE gel.

ACKNOWLEDGEMENTS

This research was supported by a grant from Ghent University (GOA 01G00805) and a grant to Annelies Haegeman from the Institute for the Promotion of Innovation through Science and Technology in Flanders (IWT‐Vlaanderen). Bartel Vanholme is a postdoctoral researcher at Ghent University (BOF).

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