V. Seidl, Research Area Gene Technology and Applied Biochemistry, Institute of Chemical Engineering, TU Vienna, Getreidemarkt 9-166-5, A-1060 Vienna, Austria Fax: +43 1 58801 17299 Tel: +43 1 58801 17263 E-mail: firstname.lastname@example.org Website: http://www.vt.tuwien.ac.at/
Genome-wide analysis of chitinase genes in the Hypocrea jecorina (anamorph: Trichoderma reesei) genome database revealed the presence of 18 ORFs encoding putative chitinases, all of them belonging to glycoside hydrolase family 18. Eleven of these encode yet undescribed chitinases. A systematic nomenclature for the H. jecorina chitinases is proposed, which designates the chitinases corresponding to their glycoside hydrolase family and numbers the isoenzymes according to their pI from Chi18-1 to Chi18-18. Phylogenetic analysis of H. jecorina chitinases, and those from other filamentous fungi, including hypothetical proteins of annotated fungal genome databases, showed that the fungal chitinases can be divided into three groups: groups A and B (corresponding to class V and III chitinases, respectively) also contained the so Trichoderma chitinases identified to date, whereas a novel group C comprises high molecular weight chitinases that have a domain structure similar to Kluyveromyces lactis killer toxins. Five chitinase genes, representing members of groups A–C, were cloned from the mycoparasitic species H. atroviridis (anamorph: T. atroviride). Transcription of chi18-10 (belonging to group C) and chi18-13 (belonging to a novel clade in group B) was triggered upon growth on Rhizoctonia solani cell walls, and during plate confrontation tests with the plant pathogen R. solani. Therefore, group C and the novel clade in group B may contain chitinases of potential relevance for the biocontrol properties of Trichoderma.
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After cellulose, chitin is the second most abundant organic source in nature . The polymer is composed of β-(1,4)-linked units of the amino sugar N-acetylglucosamine. It is a renewable resource, extracted mainly from shellfish waste, and can be processed into many derivatives, which are used for a number of commercial products such as medical applications (e.g. surgical thread), cosmetics, dietary supplements, agriculture and water treatment [1–3].
Various organisms produce chitinolytic enzymes (EC 188.8.131.52), which hydrolyze the β-1,4-glycosidic linkage . The chitinases currently known are divided into two families (family 18 and family 19) on the basis of their amino acid sequences . These two families do not share sequence similarity and display different 3D structures: family 18 chitinases have a catalytic (α/β)8-barrel domain [6–9], while family 19 enzymes have a bilobal structure and are predominantly composed of α-helices [10–12]. They also differ in their enzymatic mechanism: family 18 chitinases have a retaining mechanism, which results in chito-oligosaccharides being in the β-anomeric configuration, whereas family 19 chitinases have an inverting mechanism and consequently the products are α-anomers. Another difference is the sensitivity to allosamidin, which inhibits only family 18 chitinases . N-acetylhexosaminidases (EC 184.108.40.206), which cleave chito-oligomers and also chitin progressively from the nonreducing end and release only N-acetylglucosamine monomers, belong to glycoside hydrolase family 20 .
Some species of the imperfect soil fungus, Trichoderma[e.g. T. harzianum (teleomorph Hypocrea lixii), T. virens (teleomorph H. virens), T. asperellum and T. atroviride (teleomorph H. atroviridis)], are potent mycoparasites of several plant pathogenic fungi that cause severe crop losses each year, and are therefore used in agriculture as biocontrol agents. Biocontrol is considered to be an attractive alternative to the strong dependence of modern agriculture on fungicides, which may cause environmental pollution and selection of resistant strains. Lysis of the host cell wall of the plant pathogenic fungi has been demonstrated to be an important step in the mycoparasitic attack [14–17]. Consequently, with chitin being a major cell wall component of plant pathogens like, for example, Rhizoctonia solani, Botrytis cinerea and Sclerotinia sclerotium, several chitinase genes have been cloned from Trichoderma spp. [18–25] and, for some, the encoded protein has also been characterized [26,27]. Recently, the chitinase, Ech30, from H. atroviridis was overexpressed in Escherichia coli and characterized , but neither its expression pattern nor its biological relevance were studied. The possible roles of the endochitinases, Ech42 and Chit33, and the N-acetylglucosaminidase, Nag1, in mycoparasitism have been investigated [29–34].
In order to obtain a comprehensive insight into the chitinolytic potential of Trichoderma, we screened the recently published genome sequence of H. jecorina (anamorph: T. reesei) for chitinase-encoding genes. In this study, we present a supposedly complete list of chitinases of Trichoderma, and demonstrate their evolutionary relationships to each other and to those from other fungi. The chitinases were characterized in silico and we propose a unifying nomenclature for the large number of chitinase-encoding genes that can be found in the H. jecorina genome. Finally, five selected chitinase genes were cloned from the mycoparasitic species H. atroviridis and their transcription studied under conditions relevant for chitinase formation and mycoparasitism. A member of a new group of high-molecular-weight chitinases (chi18-10), unidentified, to date, in filamentous fungi, thereby shows a transcription profile which suggests that it may be relevant for biocontrol.
Biomining the H. jecorina genome for chitinase genes
Chitinase genes, present in the H. jecorina genome sequence, were identified by using an iterative strategy of Blast searches with fungal chitinases, as described in the Experimental procedures. We were able to identify 18 ORFs encoding putative chitinases (Table 1), including orthologues of all chitinases described, to date, from Trichoderma (ech42, Tv-ech2, Tv-ech3, chit33, Tv-cht2, ech36 and ech30). In addition to these seven known chitinases there are 11 novel, as yet undescribed/unknown, chitinase-encoding genes present in the H. jecorina genome. interproscan predicted all of them to encode a family 18 chitinase.
Table 1. Properties of Hypocrea jecorina chitinases. The theoretical pI, molecular mass, subcellular localization of the H. jecorina chitinases and the number of expressed sequence tags (ESTs) found in the H. jecorina genome database for the respective genes are given. Novel chitinases are shown in bold. Orthologues already cloned from other Trichoderma spp. and the orthologues from the mycoparasitic strain H. atroviridis, cloned in this study, are listed. The affiliation to the phylogenetic group, as determined in this study, is also given. EC, extracellular; ER, endoplasmic reticulum.
H. jecorina chitinase
Molecular mass (kDa)
Previously cloned orthologues in otherTrichoderma spp.
Cloned from H. atroviridis in this study
Ech42, Chit42, Tv-ech1 var. Trichoderma spp. (Fig. 2)
Chit36 (H. lixii, AY028421) Chit36y (T. asperellum, AAL01372)
Tv-Cht2 (H. virens, AAL78811)
EC/cell wall bound (?)
To identify potential chitinases of glycoside hydrolase family 19, a chitinase from Hordeum vulgare[GenBank accession number (acc. no.): P11955] and a chitinase from Encephalitozoon cuniculi (GenBank acc. no.: Q8STP5) were used for a tBlastn search. This strategy was unable to produce any hits, however.
tBlastn search of the H. jecorina genome database with N-acetylglucosaminidase Nag1 of H. atroviridis, which is a member of glycoside hydrolase family 20 , produced two hits that corresponded to the two N-acetylglucosaminidase-encoding genes previously cloned from H. lixii and T. asperellum. Using the same iterative Blast strategy as for the family 18 chitinases, we were unable to identify further members of the glycoside hydrolase family 20 in H. jecorina.
Having presumably identified the whole chitinase spectrum of H. jecorina, we used the following nomenclature, which is based on the proposal of Henrissat , to name chitinases according to their glycoside hydrolase family, and on the International Union of Biochemistry (IUB) nomenclature for numbering isoenzymes, which starts with the protein having the lowest pI . Therefore, the H. jecorina family 18 chitinases are named chi18-1 to chi18-18. Numbers were used instead of letters to follow the nomenclature for genes from pyrenomycetes. Table 1 shows a list of all chitinase-encoding genes of H. jecorina, including the pI and Mr of the hypothetical proteins. Also given are the hitherto existing names of chitinases that are already known in other Trichoderma spp. and the number of H. jecorina expressed sequence tags (ESTs) [38–40] that have been sequenced for the respective genes (giving an estimate of their level of expression).
Properties of the H. jecorina chitinase proteins
We used interproscan to predict the domain structure of the identified chitinase sequences and the presence of potential target sequences for cellular traffic and location (Fig. 1). The high molecular mass (>136 kDa) chitinases – Chi18-1, Chi18-8, Chi18-9 and Chi18-10 (Table 1) – are predicted to contain two LysM domains (InterPro acc. no.: IPR002482) that are suggested to bind to peptidoglycan-like structures  and a chitin-binding domain 1 (InterPro acc. no.: IPR001002) [42,43]. This type of chitin-binding domain corresponds to carbohydrate-binding module (CBM) 18 in the carbohydrate-active enzymes (CAZy) classification (CAZy database: http://afmb.cnrs-mrs.fr/CAZY/) . In addition, Chi18-10 also displays an epidermal growth factor-1-like domain known to be involved in protein–protein interactions (InterPro acc. no.: IPR001336) . For the four chitinases Chi18-1, Chi18-8, Chi18-9 and Chi18-10, considerable similarity (e−100, about 55% functionally identical amino acids on ≈ 50% of the length of the Hypocrea proteins) was obtained with the α- and β-subunits of the Kluyveromyces lactis-type killer toxins of yeasts (K. lactis, Pichia etchellsii, P. acaciae, P. inositovora, Debaromyces robertsiae and D. hansenii). These toxins consist of three subunits (α, β, γ) with α and β encoded by one ORF and the γ subunit by a separate ORF. The α-subunit has chitinase activity that is required for the toxin to act on susceptible yeast cells. The β subunit may – together with α– play a role in binding and translocation of the toxin, allowing the γ subunit to enter the cell, which leads to cell cycle arrest .
Chi18-14, Chi18-16 and Chi18-17 contain a cellulose-binding domain (CBD) (InterPro acc. no.: IPR000254; CBM 1 in the CAZy classification) [47,48], and Chi18-14 has additionally a subtilisin-like serine protease domain (InterPro acc. no. IPR000209) .
All except three chitinases (Chi18-2, Chi18-3 and Chi18-7) show the presence of a typical signal peptide, and often also a dibasic or basic-acid Kex2-like cleavage site [50,51], and are therefore likely to be secreted proteins. Chi18-3 is predicted to be located in the mitochondrion, whereas the highest subcellular localization probability for Chi18-2 and Chi18-7 is the cytoplasm. Interestingly, the putative mitochondrial location of Chi18-3 is also predicted for its orthologues from other fungi (Fig. 2). This protein also has two S-globulin domains (InterPro acc. no.: IPR000677) , which are frequently reported in association with glycoside hydrolase family 18 domains. Chi18-4 contains an endoplasmic reticulum (ER) retention signal (KDEL) which causes a relocalization of the post-translationally modified protein in the ER .
Chi18-18 consists of two domains (one being the glycoside family 18 domain, the other of unknown function), which are linked through a large unstructured region of ≈ 40 kDa that may be a cell wall anchor . This region consists of only the four amino acid residues K, A, S and T. The large number of K residues is also responsible for the unusually high theoretical pI of 9.69 of Chi18-18.
Phylogenetic relationship of the H. jecorina chitinases
The 18 chitinases were aligned with putative ortho- and paralogues present in the databases from Neurospora crassa, Gibberella zeae, Magnaporthe grisea and Aspergillus nidulans, and from other filamentous fungi found in GenBank. Also, the deduced protein sequences of five chitinases from H. atroviridis, which were cloned in this study, are included. A reliable alignment of all these protein sequences together was not possible owing to insufficient similarity between some members, and consequently three separate alignments were made. Group A contains proteins showing similarity to Ech42, group B consists of chitinases similar to Chit33 and group C comprises several, so far unknown, chitinase proteins. These groups were subjected to neighbour-joining analysis using mega2.1. Corresponding phylogenetic trees are shown in Figs 2–4. The phylogenetic relationship of the fungal chitinases (Figs 2–4) is also represented by characteristic amino acid exchanges in the consensus motifs of these family 18 chitinases [9,55]. However, the E residue in motif 2 that has been shown to be essential for catalytic activity is conserved in all chitinases . Chi18-15 is not included in any of the trees because it did not show any similarity to fungal chitinases, except to its orthologues from different Trichoderma spp. and to one chitinase from Cordyceps bassiana (GenBank acc. no.: AAN41259; e−157 and 88% functionally identical amino acids; 100% of the amino acid sequence of H. jecorina Chi18-15 was used for the significant alignment). It should be noted that the only other proteins with high similarity to Chi18-15 were chitinases from the Gram-positive bacterium Streptomyces (GenBank acc. no. CAB61702 and BAC67710; e−151 and 87% functionally identical amino acids; 100% of the amino acid sequence of H. jecorina Chi18-15 was used for the significant alignment).
The group A tree (Fig. 2) contained eight of the H. jecorina chitinases, of which three are already known in other Trichoderma spp. [Chi18-5 (= Ech42), Chi18-6 and Chi18-7)] and five are new, including the intracellular Chi18-2, mitochondrial Chi18-3, ER-targeted Chi18-4 and extracellular Chi18-11 and Chi18-18. The latter occurred in a basal position (clade A-I) and had an orthologue only in G. zeae (EAA72615). The remainder of the tree displayed five strongly supported clades: A-III, consisting of Chi18-4 and Chi18-11 as sister clades; A-IV, containing the two intracellular chitinases Chi18-2 and Chi18-3; and A-V, which also bifurcated into two sister clades, one containing Chi18-6 and the other containing Chi18-5 (Ech42) as well as the intracellular Chi18-7 in a terminal branch. The topology of the group A tree suggests that none of the H. jecorina chitinases are the products of gene duplication events, although such cases are seen for M. grisea and G. zeae (e.g. in the Chi18-6 branch of clade A-V).
The group B tree (Fig. 3) contained five chitinases, of which three (Chi18-13, Chi18-14 and Chi18-16) were new. All of the cellulose-binding domain-containing chitinases occur in this tree, which splits into two major clades: B-I branching into two subclades, each containing also chitinases from Metarhizium anisopliae, which have orthologues in H. jecorina. Chit18-13 is the orthologue of Ech30, for which enzymatic properties were recently described . The other branch contains Chi18-16 and Chi18-14, the latter apparently having arisen by gene duplication. Clade B-II bifurcates into two subclades containing the orthologues of the previously cloned H. virens Tv-cht1 and Tv-cht2 , Chi18-12 and Chi18-17.
The tree of group C (Fig. 4) contains one major supported clade (C-II), which separates from a poorly resolved clade (C-I) containing several putative chitinases from A. nidulans, G. zeae and M. grisea. All group C H. jecorina chitinases (Chi18-1, Chi18-8, Chi18-9, and Chi18-10) – which contain class I chitin-binding domains – are located in C-II, but the branches are mostly poorly supported, and it is thus unclear whether Chi18-8 and Chi18-10 are also a consequence of gene duplication.
Cloning and characterization of five novel chitinases from H. atroviridis
H. atroviridis P1 is a powerful biocontrol agent. To investigate whether some of the new genes would eventually be relevant for biocontrol, we cloned five representatives of those phylogenetic clusters which contained yet-uncharacterized chitinase-encoding genes: chi18-2, chi18-3, chi18-4, chi18-10 and chi18-13. The coding regions and 5′- and 3′-UTRs of the five chitinases were determined by RT-PCR and RACE (for details see Table 2).
Table 2. Transcription products of the new Hypocrea atroviridis chitinase-encoding genes. The 5′- and 3′-UTRs and coding regions were determined using RACE and RT-PCR.
H. atroviridis chitinase gene
Coding region (bp)
The domain structure of the novel H. atroviridis chitinases is similar to their H. jecorina orthologues, which are shown in Fig. 1. H. atroviridis Chi18-10 has an additional gamma-crystallin like element (amino acids 77–117), which can also be found in yeast killer toxins, and in antifungal and antimicrobial proteins (InterPro acc. no.: IPR011024) . In all three phylogenetic trees (Figs 2–4), the five cloned chitinases from H. atroviridis clustered immediately beneath the corresponding H. jecorina protein, proving that they are true orthologues of them.
Sequence analysis of the 5′ noncoding regions of the novel H. atroviridis chitinases identified numerous consensus binding sites for fungal transcription factors that have previously been associated with the regulation of chitinases or other polysaccharide degrading enzymes (Fig. 5). Consensus sites for the transcription factors AbaA (5′-CATTAY-3′) , BrlA (5′-MRGAGGGR-3′) , AceI (5′-AGGCA-3′) , AreA (5′-WGATAR-3′) , Cre1 (5′-SYRGGRG-3′) [62,63], PacC (5′-GCCARG-3′)  and STRE elements (5′-AGGGG-3′) [65–67], are present in the 5′ noncoding regions of the novel H. atroviridis chitinase genes. The putative Trichoderma mycoparasitism-related consensus sites, MYC1–3  were also detected in some of the 5′ noncoding regions. We used the meme motif discovery tool  to identify additional motifs in the upstream regions of the cloned H. atroviridis chitinases. However, the only highly conserved regions that were detected were chitinase consensus region 1 (CCR1) (5′-GAGACGTGCTAC-3′), which is present upstream of chi18-3 and chi18-13, and chitinase consensus region 2 (CCR2) (5′-CACTCTCAGATC-3′), which was found in the 5′ noncoding regions of chi18-3 and chi18-10 (Fig. 5).
The length of the 5′- and 3′-UTRs of the new chitinases was very variable, ranging from 52 bp to 196 bp for the 5′-UTRs and 66 bp to 466 bp for 3′-UTRs (Table 2). Interestingly, the 3′-UTR of chi18-13 contains the motif 5′-UGUANAUA-3′, which has been shown to be involved in post-transcriptional regulation. In Saccharomyces cerevisiae, binding of the RNA-binding protein, Puf3p, results in rapid deadenylation and decay of the respective mRNA [69,70].
Transcription profiles of five new chitinases from H. atroviridis
We examined the transcription of the new H. atroviridis chitinases under several conditions relevant for chitinase induction and biocontrol/mycoparasitism: various stages of plate confrontation assays with the fungal host R. solani; growth on chitin and R. solani cell walls; presence of the putative inducer, N-acetylglucosamine; and starvation for carbon and/or nitrogen. Chi18-5 (= ech42), whose transcription profile had previously been studied in this regard [18,71–73], and the constitutively expressed translation elongation factor 1-alpha (tef1)  were used as controls. A preliminary analysis showed that most of the transcripts were of too low abundance to be detected by northern analysis, therefore we used RT-PCR instead (Fig. 6). The results show that H. atroviridis chi18-10 and chi18-13 strongly respond to mycoparasitic conditions: both are up-regulated during growth on fungal cell walls and before contact with the host, respectively, chi18-10 also after contact. The transcription of these two genes was not triggered by chitin, N-acetylglucosamine or starvation for carbon or nitrogen. This is in contrast to chi18-5, which showed a constitutive basal transcription level and induction by chitin, R. solani cell walls and carbon starvation, but was only moderately transcribed in confrontation assays. Transcription of chi18-5 was even stronger when H. atroviridis grew on plates in the absence of its host than during confrontations. Similarly, chi18-4, whose translation product is ER-targeted, was transcribed constitutively and – although its transcription varied under the different conditions to some degree – no clear triggering by any of the conditions tested was found. The two putatively intracellular chitinases, chi18-2 and chi18-3, were also constitutively transcribed.
During this study, we observed that chi18-3 and chi18-13 produced two cDNA bands of different size. Sequencing showed that the larger products still contained introns. Tests for contamination with genomic DNA were negative, therefore implying the presence of two mRNA species. Interestingly, for chi18-13, only the unspliced mRNA was detected when the mycelium was grown on glucose, whereas under other conditions (e.g. when the H. atroviridis was grown on plates) the spliced transcript was predominantly present (Fig. 6). This suggests post-transcriptional regulation mechanisms for chi18-13. The presence of different levels of spliced and unspliced mRNAs has already been reported in other organisms [75–77]. Similarly, for chi18-3 the ratio of spliced to unspliced transcript and their abundance seemed to depend on growth conditions. RT-PCR products of the other chitinase genes did not contain introns and the possibility of differential mRNA splicing could therefore not be investigated. Some contained introns at the 5′ ends of the coding regions, but primers for transcript analysis were placed close to the 3′ end of the coding region to rule out differences in RT-PCR owing to inefficient reverse transcription.
In this study we identified 18 genes encoding proteins belonging to glycoside hydrolase family 18 and two members of family 20 in the H. jecorina genome, whereas no members of family 19, primarily found in plants, were detected. Previously, most authors named Trichoderma chitinases according to the putative Mr, thereby frequently also attaching an abbreviation of the species from which it was cloned [23,25,35]. However, the large number of chitinases in H. jecorina presented in this study, and the clear presence of orthologues in other filamentous fungi, makes a more systematic nomenclature for these proteins necessary. In this article we have therefore applied the rules of the IUPAC-IUB Commission on Biochemical Nomenclature (CBN) to the Trichoderma chitinases, and numbered the isoenzymes starting with the protein having the lowest theoretical pI . As we assume that we have assessed the complete chitinase spectrum of H. jecorina, we propose that the names of Trichoderma chitinases should be based on their H. jecorina orthologue and then be numbered accordingly. In addition, we follow the proposal of Henrissat , to include the glycoside hydrolase family identification number after the three letter code of the gene (chi). Chi was chosen because it is already the most commonly used name for chitinases from other organisms.
Seventeen of the H. jecorina family 18 chitinases members could be classified into three phylogenetic groups also containing several chitinases from other filamentous fungi, whereas Chi18-15 could not be aligned with any of them. Chi18-15 was previously cloned from T. asperellum and characterized, by Viterbo et al., as Chit36 [24,25]. The only orthologues that could be found in other organisms are a chitinase from the entomopathogen C. bassiana, which has been demonstrated to be involved in the attack of the fungus on insects  and two chitinases from Streptomyces spp. These data suggest that the occurrence of chi18-15 in the genome of H. jecorina, H. atroviridis and C. bassiana is caused by horizontal transfer, which – because C. bassiana and Trichoderma are both members of the Hypocreaceae– has apparently taken place rather recently (110–150 million years ago) .
All other family 18 chitinases have orthologues in filamentous fungi, including the phylogenetically diverse ascomycetes A. nidulans, N. crassa and G. zeae. This indicates that the ancestors of these genes/proteins were formed very early during the evolution of ascomycetes and their gene products therefore very likely fulfil vital functions in the fungal life cycle and/or ecology.
Particularly for chitinases of group A, orthologues were found in almost all other filamentous fungi. The closest neighbours to Trichoderma chitinases were mostly the G. zeae orthologues, indicating that evolution of these genes parallels the evolution of these species. In fact, one of these genes, chi18-5 (ech42), is used as a locus for phylogenetic analysis of the genus Trichoderma[80,81]. Chi18-5 is a chitinase that is well conserved throughout the ascomycetes, and is therefore likely to have a vital function in them. This is supported by the finding that for H. jecorina chi18-5, and the closely related chi18-7, encoding a putatively intracellular chitinase, a large number of ESTs can be found in the H. jecorina genome database, whereas none, or only two to four ESTs, were sequenced from other chitinases. It is intriguing that this gene has also been frequently investigated with respect to its involvement in mycoparasitism and biocontrol by H. atroviridis, H. lixii and H. virens[29,33,34,73,82]. Knockouts of this gene resulted in some, albeit small, reduction in biocontrol of the corresponding strains [29,34], consistent with the interpretation that chi18-5 has a rather different function in Trichoderma. As transcription of chi18-5 is triggered by carbon starvation, Brunner et al.  speculated that its main function may be associated with mycelial autolysis.
In contrast, group B, which contains chitinases with similarity to Chi18-12 (Chit33), seems to contain proteins with more species-specific functions. One striking feature of this cluster is that we could not detect any orthologue of these proteins in G. zeae, indicating that this group of chitinases is dispensable for a plant pathogenic fungus and therefore probably not essential. With the exception of Chi18-12, all members of this cluster have a fungal cellulose-binding domain (CBD) (InterPro acc. no.: IPR000254), consisting of four strictly conserved aromatic amino acid residues that are implicated in the interaction with cellulose, and four strictly conserved cysteine residues that are predicted to form two disulfide bonds . CBDs occur not only as domains of cellulose-degrading enzymes, but have also been identified in other polysaccharide-degrading enzymes (listed as CBM 1 entries in the CAZy database; http://afmb.cnrs-mrs.fr/CAZY/) . Limon et al.  demonstrated that the addition of a CBD to H. lixii Chit42 (Chi18-5) increased its activity towards high molecular mass insoluble chitin substrates, such as those found in fungal cell walls. It is therefore likely that the presence of CBDs in this cluster of family 18 chitinases may support them in chitin degradation during the mycoparasitic attack.
Interestingly, Kim et al.  reported that the CBD with highest similarity to Chi18-17 (Tv-cht1) was found in an endochitinase from the entomopathogenic fungus M. anisopliae var. acridum (CHI2; GenBank acc. no.: CAC07216). While this was true for the limited sample of chitinases available for the study, we found three chitinases from H. jecorina that are phylogenetically more close to CHI2, and indeed – together with a second chitinase from M. anisopliae (CHIT30; GenBank acc no.: AAS55554) – form a separate clade within group B. The absence of orthologous members of this clade from all other ascomycetous genomes makes it highly likely that these proteins have a special function in chitin degradation by mycoparasitic fungi (like Trichoderma) and entomopathogens (like Metarhizium). Consistent with this assumption, we showed that one member of this cluster (chi18-13) is strongly up-regulated in H. atroviridis in the presence of R. solani cell walls and in plate confrontations before contact. Thus, chi18-13, and probably also chi18-14 and chi18-16, are genes that are potentially involved in mycoparasitism and biocontrol.
It should be noted that groups A and B in the phylogenetic analysis correspond to the family 18 chitinase subgroup classes V and III, respectively. Together with the chitinase classes I, II and IV, which contain members of glycoside hydrolase family 19, this classification was used for plant chitinases prior to the glycoside hydrolase family classification [10,85]. This prompted authors to use names like fungal/plant (class III) and fungal/bacterial (class V) chitinases for these subclasses owing to similarities to either plant chitinases or bacterial chitinases [54,86]. As we detected a third subgroup of glycoside hydrolase family 18 chitinases, but our phylogenetic analysis was restricted to filamentous fungi, we simply called the subgroups (according to the clusters in Figs 2–4) group A (which is consistent with class V, also called fungal/bacterial chitinases), group B (consistent with class III and fungal/plant chitinases) and group C (a novel group of family 18 chitinases).
This third cluster (group C) of chitinases probably contains the most intriguing members of family 18. First, none of these proteins has as yet been characterized from any filamentous fungus, the cluster comprising – with the exception of A. fumigatus Chi100, for which, however, only a GenBank entry is available – only putative proteins from other fungal genome databases. Second, all of its members have a domain structure consisting of a class I chitin-binding domain (InterPro acc. no.: IPR001002; CBM 18 according to the CAZy classification) , comprising eight disulfide-linked cysteines  accompanied by two LysM domains and then followed by the glycoside family 18 domain. Although the occurrence of orthologues of these proteins in other nonmycoparasitic ascomycetes indicates that these proteins have not specifically evolved for antagonism of other fungi by Trichoderma, it is intriguing to note that these high molecular weight chitinases have high similarity to the killer toxins of certain yeasts , and chi18-10 of H. atroviridis is only expressed during growth on fungal cell walls and during plate confrontation assays, and not upon carbon starvation or growth on chitin. No protein with similarity to the γ-subunit of the yeast killer toxins – which is the actual toxicity factor – has been found in the H. jecorina genome. However, as the γ-subunit causes cell cycle arrest in yeast, it is probably dispensable for the antagonization of multicellular fungi. Rather, we speculate that Trichoderma uses a killer-toxin like mechanism to enable the penetration of antifungal molecules into its host. For this reason, we also consider this group of chitinases potentially interesting candidates for proteins that are connected with the biocontrol properties of Trichoderma.
Transcription analysis of the novel H. atroviridis chitinases chit18-2, chi18-3, chi18-4, chi18-10 and chi18-13 showed that, although transcript levels were generally rather low as they could not be detected by northern analysis and one has to be careful with interpreting the RT-PCR data quantitatively, a clear influence of different growth conditions and carbon sources could be detected. This indicates the functional diversity of the Trichoderma chitinases and that they are not just substitutes for each other, but that they indeed have specific roles in the organism. In particular, the transcript patterns of chi18-10 and chi18-13 were explicitly linked to the presence of components apparently present in the cell wall of R. solani. No striking similarities in the upstream regions of chi18-10 and chi18-13 were detected. The extensive in silico analysis of the novel H. atroviridis chitinase genes (Fig. 5) gives some hints as to which regulatory mechanisms might be important for the respective chitinase genes, but detailed promotor studies are certainly necessary to elucidate any common consensus sites and transcription factors responsible for the regulation of Trichoderma chitinases.
In this study, we showed, for the first time, that post-transcriptional regulation is involved in chitinase expression. We demonstrated that, at least for chi18-3 and chi18-13, different mRNA species were present and that their occurrence was influenced by the growth conditions. Additionally we found a Puf-binding site in the 3′-UTR of chi18-13. It should be noted that proteins with Puf RNA-binding domains (InterPro acc. no.: IPR001313) are indeed present in the H. jecorina genome. The aspect of post-transcriptional regulation has not yet been studied great detail in filamentous fungi. It comprises interesting insights into the actual protein levels that can be observed in vivo and could contribute to a more accurate understanding of enzyme-mediated events, such as mycoparasitism.
H. atroviridis P1 (ATCC 74058) was used in this study and maintained on potato dextrose agar (PDA) (Difco, Franklin Lakes, NJ, USA). E. coli strains ER1647 and BM25.8 (Novagen, Madison, WI, USA) were used for genomic library screening, and JM109 (Promega, Madison, WI, USA) was used for plasmid propagation.
Culture conditions and preparation of special carbon sources
Shake flask cultures were prepared with the medium described by Seidl et al.  and incubated on a rotary shaker (250 r.p.m.) at 28 °C. Cultures were pregrown for 28 h on 1% (w/v) glucose and then harvested by filtering through Miracloth (Calbiochem, Darmstadt, Germany), washed with medium without a nitrogen or carbon source and transferred to a new flask containing 1% (w/v) glucose for 2 h or 1 mmN-acetylglucosaminidase for 30 min, respectively. Starvation was induced by replacing on (a) 0.1% (w/v) glucose (carbon limitation), (b) 1% (w/v) glucose and 0.14 g·L−1 (NH4)2SO4 (nitrogen limitation) or (c) 0.1% (w/v) glucose and 0.14 g·L−1 (NH4)2SO4 for 15 h (carbon and nitrogen starvation). Cultures were grown for 48 h directly on 1% (dry weight) colloidal chitin or R. solani cell walls. Mycelia were harvested by filtration through Miracloth (Calbiochem), washed with cold tap water, squeezed between two sheets of Whatman filter paper, immersed in liquid N2 and stored at −80 °C.
Colloidal chitin was prepared essentially as described by Roberts et al. . Briefly 20 g of crab shell chitin (Sigma, Vienna, Austria) was suspended in 400 mL of concentrated HCl, stirred overnight at 4 °C and filtered through glass wool. The filtrate was precipitated with 2 L of ethanol and washed with distilled water at 4 °C until a pH of 5.0 was reached. R. solani cell walls were prepared by growing R. solani on PDA plates covered with cellophane, grinding the mycelium under liquid nitrogen and suspending it in distilled water containing 0.1% (w/v) SDS (30 mL·g−1 cell wall). The suspension was further homogenized in a Potter-Elvehjem pistill homogenizer, centrifuged (15 min, 18 000 g, 4 °C) and the pellet washed with distilled water to remove attached proteins (the flow through was checked by measuring the absorbance at 280 nm).
For plate confrontation assays, strips of 30 × 3 mm were cut out from the growing front of H. atroviridis and R. solani, and placed on fresh PDA plates (9 cm diameter) covered with cellophane at a distance of 4 cm from each other. The mycelia were harvested at three different time-points (a) before contact, when the mycelia were at a distance of ≈ 10 mm, (b) contact, when the mycelia were just touching, and (c) after contact, when H. atroviridis had overgrown R. solani by ≈ 5–10 mm. Mycelium from the growing front (≈ 7 mm) was harvested with a spatula, frozen in liquid nitrogen and stored at −80 °C. Equivalent zones were collected from control plates, inoculated with H. atroviridis or R. solani only.
Biomining of the H. jecorina genome
The H. jecorina genome (http://gsphere.lanl.gov/trire1/trire1.home.html) was screened for chitinases by using the tBlastn (protein vs. translated nucleotide) program. First, we used the protein sequences of the published chitinase sequences of other Trichoderma spp. (listed in Table 1) as query to search the H. jecorina genome. Then, all chitinases, including that newly identified from H. jecorina, were used to identify further proteins with similar domains, and, finally, all hypothetical proteins encoding chitinases from the annotated genomes of the Broad Institute (http://www.broad.mit.edu/), including Emericella nidulans (A. nidulans), N. crassa, G. zeae (Fusarium graminearum) and M. griseae were used. The loci of the H. jecorina chitinases in the H. jecorina genome database are listed in Table 3.
Novel chitinase-encoding genes from H. atroviridis were cloned by using PCR fragments from H. jecorina chitinases as probes. The primers listed in Table 4 were used to amplify the respective fragments from H. jecorina by PCR, which were then isolated and used to screen a genomic λ BlueSTAR library (Novagen) of H. atroviridis P1. Isolated phages were converted to plasmids and sequenced at MWG Biotech AG (Ebersberg, Germany).
Table 4. Primers for amplification of Hypocrea jecorina genomic DNA fragments for phage library screening.
Primer for phage library screening
Fragment from the H. jecorina chitinase gene
Annealing temperature (°C)
Fragment length (bp)
The assembled DNA sequences were deposited in GenBank (acc. nos: DQ068748–DQ68752).
Protein sequences were aligned first with ClustalX 1.8  and then visually adjusted using genedoc 2.6. Phylogenetic analyses were performed in mega 2.1, using Neighbour Joining, a distance algorithmic method. Stability of clades was evaluated by 1000 bootstrap rearrangements. Bootstrap values lower than 50% are not displayed in the cladogram.
PCR reactions were carried out in a total volume of 50 µL containing 2.5 mm MgCl2, 10 mm Tris/HCl, pH 9.0, 50 mm KCl, 0.1% (v/v) Triton X-100, 0.4 µm each primer, 0.2 mm each dNTP and 0.5 U Taq Polymerase (Promega). The amplification program consisted of: 1 min initial denaturation (94 °C), 30 cycles of amplification [1 min at 94 °C, 1 min at the primer-specific annealing temperature (see Table 4), 1 min at 72 °C], and a final extension period of 7 min at 72 °C. For RACE-PCR, amplification cycles were increased to 35 and RT-PCR was carried out over 25 or 35 cycles.
Total RNA was extracted as described previously .
cDNA was synthesized using the Creator SMART cDNA library construction kit (BD Biosciences, Palo Alto, CA, USA) from RNA from H. atroviridis cultures grown on glucose. The primers used for RACE-PCR are listed in Table 5. Amplification of 5′- and 3′ cDNA ends was carried out using the 5′-PCR and CDSIII primers from the cDNA kit and gene-specific primers followed by a second PCR using the 5′-PCR and CDSIII primers and nested gene specific primers.
Table 5. RACE-PCR primers for amplification of Hypocrea jecorina genomic DNA fragments.
Primer for RACE-PCR
Fragment from the H. atroviridis chitinase gene
Annealing temperature (°C)
The resulting fragments were cloned into pGEMT-Easy (Promega, Mannheim, Germany) and sequenced at MWG Biotech (Ebersberg, Germany).
RNA obtained from various cultures was treated with DNAse I (Fermentas, St Leon-Rot, Germany) and purified using the RNeasy MinElute Cleanup Kit (Qiagen, Hilden, Germany). A total of 5 µg of RNA per reaction was reverse transcribed using the RevertAid H Minus First Strand cDNA Synthesis Kit (Fermentas) and the oligo(dT)18 primer.
The cDNA was used for PCR with sequence-specific primers, listed in Table 6, to assess the exon/intron boundaries. For transcript analysis (RTQ-Primers, Table 7), the annealing temperature, RNA concentration and the number of amplification cycles were optimized and, finally, 5 µg of RNA per reaction, 25 cycles (unless otherwise stated) and the temperatures listed in Table 7 were used. A 40 µL sample of each PCR reaction was separated on a 1.5% agarose gel containing 0.5 µg·mL−1 ethidium bromide.
Table 6. RT-PCR primers for identification of coding regions and introns. H. atroviridis, Hypocrea atroviridis.
Primer for RT-PCR
Fragment from the H. atroviridis chitinase gene
Annealing temperature (°C)
Fragment length (bp)
Table 7. RT-PCR primers for transcript analysis under different growth conditions. H. atroviridis, Hypocrea atroviridis.
Primer for RT-PCR (transcript analysis)
Fragment from the H. atroviridis gene
Annealing temperature (°C)
Fragment length (bp)
The following controls were carried out in parallel with each RT-PCR experiment. To ensure the absence of genomic DNA, RNA was treated with DNAse I, purified and subjected to the reverse transcription procedure as described above, but no reverse transcriptase was added during this step. This RNA was subsequently used for PCR under the same conditions that were used for RT-PCR over 35 cycles. Additionally, PCR reactions without template were set up to exclude contamination with other PCR components. In none of the controls was a PCR product detected when they were visualzed by agarose gel electrophoresis.
This study was supported by a grant from the Austrian Science Foundation (P 16601) to CPK. Sequence data were obtained from the Department of Energy Joint Genome Institute (http://www.jgi.doe.gov). The H. jecorina/T. reesei genome sequencing project was funded by the United States Department of Energy.