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Department of Plant Biology and Pathology, Cook College, Rutgers University, 59 Dudley Road, New Brunswick, New Jersey 08903
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Many wild and cultivated cool-season grass species are naturally infected with fungal endophytes of the genera Neotyphodium and Epichloë. These associations generally are considered mutualistic with the plants benefiting from reduced herbivory and the fungi benefiting from nutrients supplied by the plants. The fungi secrete proteins that might have a role in the interspecies symbiosis. In the interaction between Poa ampla Merr. and the endophyte Neotyphodium sp., a fungal chitinase was detected in the apoplastic protein fraction. The chitinase was also the major protein secreted in culture. Sequence analysis of the chitinase revealed it has a low level of amino acid sequence identity to other fungal chitinases and one of the conserved active site residues is altered. DNA gel-blot analysis indicated the chitinase was encoded by a single gene. Expression of similar chitinases also was detected in endophyte-infected tall fescue (Festuca arundinacea Schreb.), perennial ryegrass (Lolium perenne L.) and Chewings fescue (Festuca rubra L. subsp. fallax [Thuill] Nyman). This is the first report of an endophyte chitinase expressed in the infected host grass. As a secreted hydrolytic enzyme, the chitinase might have roles in the nutrition, growth or defense of the endophyte.
Key words: Epichloë, mutualistic symbiosis, Neotyphodium
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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) or the related teleomorphic genus Epichloë (Schardl 2001, Scott 2001). In nature these fungi are found exclusively in association with their plant hosts. Infection often results in benefits to the host, such as reduced herbivory by insects and animals (Breen 1994, Bush et al 1997). Other physiological benefits to the plants, such as disease resistance and drought resistance, also have been reported (Clay and Schardl 2002, Latch 1997). The fungi benefit from nutrients supplied by the plants. Because of the benefits provided to both fungi and plants, these associations generally are considered to be mutualistic symbioses (Clay 1988). These associations are important agriculturally, both positively and negatively, because infection can protect turf and forage grasses from insect attack (Funk et al 1983, Barker et al 1984) but also can result in toxic syndromes in grazing animals (Ball et al 1993). Both of these effects result from the production of toxic alkaloids in the infected grasses (Bush et al 1997).
Within the infected plants the fungal hyphae are found in the intercellular spaces of the leaf sheaths and blades running longitudinally between the plant cells. They often form close associations with the plant cell walls (Christensen et al 2002). However, they do not invade the plant cells. Very little is known regarding the factors important in host colonization or nutrient exchange between plant and fungus. Fungal secreted proteins are expected to be synthesized for growth and nutrient acquisition and perhaps for defense. They are likely to be important components of the mutualistic interaction because they are located at the interface of the two species. We previously have reported the expression of a fungal subtilisin-like proteinase, invertase, and ß-1,6-glucanase in endophyte-infected plants (Lam et al 1995, Moy et al 2002, Reddy et al 1996). These enzymes might function in the extraction of nutrients from the plant apoplast. Fungi rely on secreted hydrolytic enzymes to degrade environmental polymers to smaller molecules that can be used for nutrition (Wessels 1993). Secreted enzymes also might be important in fungal growth. Secreted chitinases have been reported from a number of filamentous fungal species, and roles in hyphal branching and autolysis have been proposed (Gooday et al 1992). Secreted chitinases also are considered to be important in the mycoparasitic and entomopathogenic processes of some fungi (Garcia et al 1994, St. Leger et al 1996). Here we report the characterization of a Neotyphodium sp. secreted chitinase that is the major secreted protein in culture and that also is present in the apoplastic fluid of the infected host grass, Poa ampla.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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).
Isolation and peptide sequencing of secreted proteins. –
For isolation of secreted proteins, the endophyte was grown in 30 mL liquid tryptone-sucrose medium (Lam et al 1995) for 6–12 d. Mycelial plugs of approximately 5 mm diam, taken from the growing margins of a colony growing on a PDA plate, were used to inoculate tryptone-sucrose medium. The culture filtrate was collected by filtration through two layers of Miracloth (Calbiochem, Darmstadt, Germany) followed by centrifugation at 12 000 x g. The proteins in the culture filtrate were concentrated to 60–200 µL in Centriprep-30 and Centricon-30 concentrators (Amicon, Beverly, Massachusetts). The final protein concentrations were determined by the Bio-Rad Protein Assay Reagent (Bio-Rad, Hercules, California) with bovine serum albumin as a standard.
The concentrated secreted proteins were separated by tri-cine-SDS-PAGE, without glycerol (Schagger and von Jagow 1987), and stained with Coomassie Brilliant Blue. The 52-kDa protein band was excised and the gel slices washed with 50% (v/v) acetonitrile in water. Sequence analysis by Ed-man degradation of peptides generated by trypsin digestion was performed by the Texas Microchemistry Facility (University of Texas Medical Branch, Galveston, Texas).
Chitinase activity assay. –
Chitinase activity was detected in SDS-PAGE essentially as described by Tronsmo and Harman (1993). Ten µg of culture filtrate protein was mixed with an equal volume of 2x SDS sample buffer, without 2-mercap-toethanol and subjected to SDS-PAGE at 4 C without boiling. After electrophoresis, gels were washed twice with 2.5% (v/ v) Triton-X-100 and then incubated at 4 C in 2.5% (v/v) Triton-X-100 for 1 h to allow enzyme renaturation (Zhang et al 2001). Enzyme activity was detected in agarose gel overlays of 1% (w/v) low melting point agarose (Bio-Rad) in 50 mM sodium acetate, pH 5.0 containing 300 µg mL–1 of the fluorescent substrates 4-methylumbelliferyl-ß-D-N,N'-diacetylchitobioside (4-MU-[GlcNAc]2), or 4-methylumbelliferyl-ß-D-N-N'N''triacetylchitotrioside (4-MU-[GlcNAc]3). The agarosesubstrate suspensions were dissolved in a microwave oven and cooled to 42 C before application to the gels. The agarosegel overlays were incubated at 37 C for 30 min and the fluorescent activity bands visualized on a UV transilluminator.
To determine the effect of heating on the migration in SDS-PAGE of the protein in the active bands, the fluorescent bands were excised from the activity gels. The gel slices were ground in 50 µL 2x SDS sample buffer, heated at 90 C for 5 min and centrifuged at 12 000 x g for 5 min. The supernatants were subjected to SDS-PAGE and the gels stained with Coomassie Brilliant Blue.
Chitinase activity in solution was determined by a colorimetric assay modified from Roberts and Selitrennikoff (1988) in which p-nitrophenyl-N-acetyl-ß-D-glucosaminide (pNp-GlcNAc), p-nitrophenyl- ß-D-N,N'-diacetylchitobio-side (pNp-[GlcNAc]2) and p-nitrophenyl- ß-D-N,N',N''-tri-acetylchitotriose (pNp-[GlcNAc]3) (Sigma-Aldrich) serve as substrates. A typical enzyme reaction contained 100 µL of a 300 µg mL–1 solution of substrate in 50 mM sodium acetate, pH 5.0 and 10 µL of protein sample. After incubation at 37 C for 30 min, the reaction was stopped by adding 10 µL of 1 N NaOH. The release of p-nitrophenol was monitored spectrophotometrically at 405 nm. One unit of enzyme activity was defined as the amount of enzyme that catalyzes the release of 1 nmol p-nitrophenol per min at 37 C.
Purification of the 52-kDa chitinase. –
The Neotyphodium sp. endophyte was grown in 100 mL tryptone-0.1% (w/v) sucrose medium (Lam et al 1995) for 7 d. The culture filtrate was separated from the fungal mycelium by filtering the culture through Miracloth followed by centrifugation at 5000 x g for 10 min. The secreted proteins in the culture filtrate were concentrated 100-fold to approximately 1 mL using Centriprep-30 and Centricon-30 concentrators. The buffer of the concentrated protein was changed to 50 mM potassium phosphate buffer, pH 7.0 using a Centricon-30. The concentrated culture filtrate was mixed with 4 g (wet weight) of colloidal chitin and incubated at 4 C for 3 h to allow adsorption of the chitinase. Colloidal chitin was prepared from crude chitin (crab shell flakes, Sigma-Aldrich) (Lingappa and Lockwood 1962). Before use, the colloidal chitin was neutralized with NaOH to pH 7.0 and washed three times with 50 mM potassium phosphate buffer, pH 7.0. After adsorption, the colloidal chitin was pelleted by centrifugation and washed twice with 100 mM NaCl, 50 mM potassium phosphate buffer, pH 7.0. The bound protein was eluted from the chitin pellet by sequential washes of 10 mL each of 1 M NaCl, 50 mM potassium phosphate, pH 7.0 and 1M NaCl, 50 mM potassium phosphate, pH 3.0. The combined eluted proteins were concentrated and the buffer changed to 10 mM MES, pH 6.8, with Centricon-30 concentrators. The concentrated chitin-bound protein solution was adjusted to 1 M ammonium sulfate and subjected to hydrophobic interaction chromatography. The sample was applied to a 1 mL HiTrap Phenyl HP column (Amersham Pharmacia Biotech Inc., Piscataway, New Jersey) and washed with 3 mL of 1 M ammonium sulfate, 10 mM MES, pH 6.8. The effluent was collected as the unbound fraction. The unbound fraction was concentrated and the buffer changed to 10 mM MES, pH 6.8, with Centricon-30 concentrators. Aliquots were assayed for chitinase activity spectrophotometrically.
The purified protein was subjected to SDS-PAGE (Laemmli 1970) and electroblotted to a PVDF membrane (ProBlott, Applied Biosystems Inc., Foster City, California) for protein sequencing. Transferred proteins were visualized by staining the membrane with 0.2% (w/v) Ponceau S (Sigma-Aldrich) in 1% (v/v) acetic acid. The membrane was destained with H2O and the 52-kDa band excised. N-terminal sequencing was performed by the Texas Micro-chemistry Facility (University of Texas Medical Branch, Gal-veston, Texas).
Nucleic acid manipulations. –
Fungal DNA and plant RNA isolations were as previously described (Moy et al 2002). For DNA gel blot analysis, 12 µg of DNA from the Neotyphodium sp. endophyte was digested with BamHI, ClaI, EcoRI, EcoRV, HindIII, KpnI, SalI, or XbaI in a 50 µl total reaction volume at 37 C for 12 h. The DNA was subjected to electrophoresis through a 0.8% agarose gel. DNA in the gel was depurinated by washing in 0.25 N HCl for 12 min. The gel then was washed in H2O and the DNA was transferred to a nylon membrane (Zeta-Probe, Bio-Rad) overnight with 0.4 M NaOH (Reed and Mann 1985). The membrane was washed in 2x SSC and fixed by drying.
For RNA gel-blot analyses, RNA was subjected to electrophoresis in formaldehyde agarose gels and transferred to nylon membranes (Magnagraph, Osmonics, Minnetonka, Minnesota) as described by Selden (1987). RNA was fixed to the membrane with a UV Crosslinker (Fisher Scientific, Pittsburgh, Pennsylvania).
A 1.2 kb EcoRI/XhoI restriction fragment from the cDNA clone was labeled with [
32P]dCTP using a commercial kit (Prime-It II Random Primer Labeling Kit, Stratagene, La-Jolla, California) for use as a probe for all hybridization reactions.
For both DNA and RNA gel blots, filters were prehybridized at 42 C in 50% (v/v) formamide, 5x SSC, 5x Denhardt’s solution (1x Denhardt’s solution is 0.02% [w/v] Ficoll, 0.02% [w/v] PVP, 0.02% [w/v] BSA), 50 mM sodium phosphate, pH 6.8, 1% (w/v) SDS, 100 mg µL–1 calf thymus DNA, and 2.5 % (w/v) dextran sulfate. The hybridization solution was 5 x 105 cpm mL–1 of 32P-labeled fragment, 50% (v/v) formamide, 5X SSC, 1x Denhardt’s solution, 20 mM sodium phosphate, pH 6.8, 1% (w/v) SDS, 100 µg mL–1 calf thymus DNA, and 5% (w/v) dextran sulfate. Hybridized membranes were washed with 2x SSPE, 0.5% (w/ v) SDS for 15 min at room temperature, 2x SSPE, 0.5% (w/v) SDS for 15 min at 65 C, and 0.2x SSPE, 0.2% (w/v) SDS for 15 min at 65 C. The washed membranes were exposed to X-Ray film (XOMAT-AR, Kodak, Rochester, New York) with an intensifying screen.
cDNA library screening was as described previously (Moy et al 2002). Hybridization conditions were the same as for the gel blots.
PCR amplification of a chitinase genomic DNA fragment. –
Degenerate oligonucleotide primers for PCR were designed based on two of the peptide sequences obtained from the 52-kDa extracellular protein band. The sequences of the degenerate primers were: primer A, 5'-GTIGCIGAYYT-IGGNYTNGAYGG-3' and primer B, 5'-TTRAAIGGIGT-NGCRAANGGRTT-3'. The symbols used for the mixed bases are: I = deoxyinosine; Y = C, T; N = A, C, G, T; and R = A, G. Amino acid sequences encoded by primers A and B are VADLGLDG and NPFATPFN, respectively. The degenerate oligonucleotide primers were used in PCR ampli-fication of Neotyphodium sp. genomic DNA. PCR reactions were performed in volumes of 50 µL and contained 10 mM Tris-HCl, pH 8.3, 50 mM KCl, 2.5 mM MgCl2, 0.2 mM each dNTP, 0.5 µg of each primer, and 1 µl (2 U) Taq polymer-ase (Life Technologies, Gaithersburg, Maryland). PCR was carried out in a GeneAmp 9600 thermocycler (Perkin El-mer Corp., Foster City, California). The PCR cycling parameters were 30 s at 94 C, 30 s at 58 C and 1 min at 72 C for 35 cycles. The resulting PCR amplification products were separated on a 1.2% agarose gel, and a DNA band of approximately 330 bp was excised from the gel and extracted using a commercial kit (QIAquick Gel Extraction Kit, Qia-gen). The eluted DNA was ligated into pGEM-T Easy vector (Promega, Madison, Wisconsin), and transformed into DH5 E. coli competent cells (PGC Scientifics, Frederick, Maryland). Plasmids were purified from E. coli transformants using a commercial kit (QIAprep Spin Miniprep Kit, Qiagen). Sequencing was performed by Davis Sequencing (Davis, California).
Isolation and peptide sequencing of apoplastic proteins. – Leaf sheaths were cut to 2 cm lengths, cleaned, and vacuum in-filtrated for 30 min with 100 mM Tris-HCl, pH 8.0, 50 mM DTT, 10 mM ascorbic acid and 5 mM PMSF. The leaf sheaths were washed with H2O three times, blotted dry, and collected in a 3 cc syringe. The syringe was placed in a 50 ml tube and centrifuged at 2000 x g for 10 min at 4 C. The apoplastic fluid was collected and concentrated with a Microcon YM-30 (Millipore Corporation, Bedford, Massachusetts). Thirty µg of protein was mixed with an equal volume of 2x SDS sample buffer, heated at 100 C for 5 min and subjected to SDS-PAGE. A 52-kDa band was excised and the gel slice was washed with 50% (v/v) acetonitrile in water. Sequence analysis of peptides generated by trypsin digestion of the protein in the excised band was performed by the Harvard Microchemistry Facility (Harvard University, Cambridge, Massachusetts) by microcapillary reverse-phase HPLC nanoelectrospray tandem mass spectrometry (µLC/ MS/MS) on a Finnigan LCQ quadrupole ion trap mass spectrometer.
Phylogenetic analyses. –
The ClustalW program was used to align amino acid sequences using the BLOSUM62 scoring matrices (Henikoff and Henikoff 1992). Ambiguously aligned regions were removed from the alignment. Phylogenetic analysis was performed using PAUP (version 4.0b10 for Macintosh) (Swofford 2002). Heuristic searches using random stepwise additions (100 replicates) with maximum-parsimony criterion was performed. Gaps were excluded from the analysis. Branch support was obtained by bootstrapping (1000 replicates) also using random step-wise additions (10 replicates) with maximum parsimony criterion.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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) to ensure that all protein in the culture filtrate originated from fungal secretion and not from the medium itself. The Neotyphodium sp. endophyte was grown in medium containing either 0.1% or 1% sucrose for 8 d and the proteins in the culture filtrate subjected to SDS-PAGE (FIG. 1A). There was no difference in the protein pattern, and in both medium formulations there was a major protein band at approximately 52-kDa. The pattern of protein secretion from 6 to 12 d in culture was compared in the tryptone-0.1% sucrose medium (FIG. 1B). The 52-kDa band was a prominent band at all time points and by 8 d was the single most abundant band. The abundant 52-kDa protein band was excised from a gel and subjected to trypsin digestion and the sequence of some resulting peptides determined by Edman degradation. Ten peptide sequences were obtained, three of which, NPFAXPFNT, LLTFASPAGEQR and LVADLGLDGLDLDYEY-VANSK, were similar to an endochitinase, CHIT42, from the mycoparasitic fungus Trichoderma harzianum (Garcia et al 1994).
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). In the boiled sample the major stained band was at 52-kDa, whereas in the unboiled sample the major stained band migrated higher in the gel, identical to the migration of the activity bands. The migration position of the other proteins in the culture filtrate was the same in both the boiled and unboiled samples. These results suggested that the activity bands corresponded to the major stained protein band in the unboiled sample which, in turn, corresponded to the 52 kD major stained band seen in the boiled sample. This was confirmed by cutting out the active regions from the activity gels, and heating the extracted proteins in 2x SDS buffer followed by SDS-PAGE (FIG. 2C). Bands at 52 kD were seen from both activity bands.
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, FIG. 3). To distinguish between N-acetyl-ß-D-glucosaminidase, exochitinase, and endochitinase activities, three chromogenic substrates, pNp-GlcNAc, pNp-(GlcNAc)2 and pNp-(GlcNAc)3, were used at all steps in the purification. The crude culture filtrate had activity against all three substrates. Affinity binding to chitin removed most of the proteins in the culture filtrate leaving the 52-kDa protein and a few other proteins (FIG. 3, lane 2). The chitin-bound fraction had activity against all three substrates in the same ratios as the crude culture filtrate. All three types of chitinolytic enzymes would be expected to bind to colloidal chitin. Passage through phenyl sepharose further purified the 52-kDa protein band and removed the N-acetyl-ß-D-glucosaminidase activity. The 52-kDa chitinase did not bind to the phenyl sepharose column and was recovered in the effluent. By SDS-PAGE analysis the purified 52-kDa chitinase was a single protein (FIG. 3, lane 3). Because the chitinase protein was the major protein in the crude culture filtrate, the final purification was only 1.6-fold. By gel analysis, however, the protein was highly purified. The reduction in specific activity against the substrates pNp-(GlcNAc)2 and pNp-(GlcNAc)3 from the chitin-bound fraction to the phenyl sepharose unbound fraction was attributable to the removal of the N-acetyl-ß-D-glucosaminidase, which also has activity against those substrates. Activity against both pNp-(GlcNAc)2 and pNp-(GlcNAc)3 by the purified chitinase was indicative of endochitinase activity (Tronsmo and Harman 1993).
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Isolation of a cDNA clone for the Neotyphodium chitinase. – Degenerate oligonucleotides based on the peptide sequences VADLGLDG and NPFATPFN were used in PCR of fungal genomic DNA. A 330-bp am-plified band was cloned that encoded an amino acid sequence that was similar to other fungal chitinases. The PCR clone was used to screen a cDNA library prepared from endophyte-infected (E+) P. ampla leaf sheath tissue and a clone containing the complete coding sequence was obtained. Isolation of the clone from the cDNA library indicated the Neotyphodium sp. chitinase was expressed in the infected grass host as well as in culture.
The nucleotide and deduced amino acid sequences of the cDNA clone are shown in FIG. 4. The clone contained an untranslated 5'-upstream sequence of 115 bp, an open-reading frame of 1377 bp, and an untranslated 3'-sequence of 197 bp. A 458-amino acid protein with a molecular weight of 50 301 daltons is predicted from the cDNA sequence. There was one amino acid difference between the peptide sequence NPFATPFN that was used to generate the PCR clone and the corresponding region of the cDNA clone, NPDATPFN. The seven peptide sequences obtained from the 52-kDa protein band that were not originally recognized as having similarity to other fungal chitinases all are encoded by the Neotyphodium sp. clone (FIG. 4). That all of the peptide sequences obtained from the protein band are encoded by the cDNA suggested that the Neotyphodium sp. chitinase was the major component of the 52-kDa protein band obtained from the culture filtrate.
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; http://www.cbs.dtu.dk/services/SignalP-2.0/). Secreted chitinases from other fungi often have an additional proteolytic processing step after cleavage of the signal peptide. The Neotyphodium sp. chitinase apparently also has such a processing step since the experimentally determined N-terminus of the purified protein was downstream of the signal peptide cleavage site. The calculated molecular weight Mr of the mature protein is 46532 and the pI is 8.78. The calculated Mr is smaller than that estimated from SDS–PAGE of the purified protein. This discrepancy may be due to protein glycosylation or anomalous electrophoretic mobility. There are two potential N-glycosylation sites of NXT/S in the amino acid sequence at residues 94 and 319.
Similarity of the Neotyphodium sp. chitinase to other sequences. –
The Neotyphodium sp. sequence is similar to chitinases from other fungal species, but the level of identity is not high to any other sequences currently in the database. Phylogenetic analysis comparing 15 of the most similar glycosyl hydrolase family 18 chitinases currently in the database illustrates the relationship of the Neotyphodium sp. chitinase sequence to other characterized fungal chitinases (FIG. 5). The data matrix contained 16 taxa and 408 total characters, of which 264 characters were parsimony informative. The 100 searches using random stepwise additions resulted in three equally parsimonious trees with consistency and homoplasy indices of 0.70 and 0.30, respectively, one of which is depicted in the phylogram shown in FIG. 5. The position of the Neotyphodium sp. chitinase relative to the other sequences was the same in all three trees. The branching of the Neotyphodium and Candida chitinases from the same point in the phylogenetic tree reflects the dissimilarity of both sequences to the others in the comparison and not their similarity to each other, which is only 28% identity.
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, Robertus and Monzingo 1999), which were found between amino acids 145–156 and 182–191 (FIG. 6). The active site amino acid residues of the Coccidioides immitis chitinase have been determined from crystallography (Hollis et al 2000) and seven of the eight amino acids were conserved in the Neotyphodium sp. sequence (FIG. 6). The conserved glutamic acid at position 190 and aspartic acid at position 188 are believed to be the residues directly involved in catalysis (Hollis et al 2000, Bortone et al 2002).
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). The most prominent band at approximately 34-kDa in the E+ sample was the fungal subtilisin-like proteinase which previously was characterized (Lindstrom and Belanger 1994, Reddy et al 1996). A component of the protein band at approximately 47-kDa was identified as a fungal ß-1,6-glucanase (Moy et al 2002). Another apoplastic protein band that was present only in the E+ sample had a Mr of 52-kDa, similar to the chitinase identified from the fungal culture filtrate. The 52-kDa protein band therefore was excised from the gel and subjected to trypsin digestion and peptide sequencing by tandem mass spectrometry (MS/MS). The deduced amino acid sequence of the chitinase cDNA clone was included in the database used for analysis of the resulting peptides. Ninety-five peptide sequences from the 52-kDa apoplastic protein band were identified that were identical to the chitinase sequence deduced from the cDNA clone. The peptide sequences obtained from the 52-kDa band spanned the chitinase protein sequence from amino acid 46 to amino acid 453. These results indicated that the fungal chitinase protein accumulated to detectable levels in the apoplast of the infected host plant.
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). An intense hybridizing band was detectable from both total RNA and poly(A+) RNA. No hybridization was detectable in RNA from endophyte-free plants, indicating that the host, P. ampla, did not express a highly similar chitinase.
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, Latch et al 1984, Leuchtmann et al 1994). Hybridization also was detected in RNA from a Chewings fescue artificially inoculated with the Neotyphodium sp. endophyte that infects P. ampla ( Johnson-Cicalese et al 2000). These results indicated that transcripts similar to the Neotyphodium sp. chitinase were being produced by other endophyte species infecting other grass species.
Neotyphodium sp. chitinase is encoded by a single gene. –
Gel-blot analysis of the Neotyphodium sp. DNA hybridized with the 1.2-kb chitinase cDNA fragment indicated that the chitinase is encoded by a single gene. When the DNA was digested with restriction enzymes for which there are no sites in the chitinase cDNA, EcoRI, KpnI and XbaI, only single hybridizing bands were detected (data not shown). When the DNA was digested with enzymes for which there are restriction sites within the chitinase cDNA, BamHI, ClaI, EcoRV, HindIII and SalI, the predicted number of hybridizing bands were detected. These data indicated that there are no additional closely related chitinase genes in the Neotyphodium sp. genome. Multiple chitinase genes that do not cross-hybridize have been reported from Trichoderma virens (Kim et al 2002). Whether the Neotyphodium sp. has other chitinase genes is not known.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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). Exochitinases progressively release only the dimer diace-tylchitobiose from the nonreducing end of the chitin polymer (Sahai and Manocha 1993). This latter type of chitinase activity also has been termed chitobiosidase (Tronsmo and Harman 1993). N-acetylglucosaminidases release GlcNAc monomers from the oligomeric products generated by endochitinases and exochitinases (Cohen-Kupiec and Chet 1998). 4-MU-(GlcNAc)3 is considered a substrate for endochitinases, whereas 4-MU-(GlcNAc)2 is a substrate for both endochitinases and exochitinases (De la Cruz et al 1992, McCreath and Gooday 1992, Tronsmo and Harman 1993). Activity of the Neotyphodium sp. chitinase against both 4-MU-(GlcNAc)2 and 4-MU-(GlcNAc)3 suggested that it had endochitinase activity. The sequence of the chitinase was similar to other fungal endochitinases. The active site residues identified from crystallography of the C. immitis chitinase all were conserved in the Neotyphodium sp. sequence except the valine at position 258. The corresponding position is a tyrosine in C. immitis and in more than 200 other chitinase sequences in the NCBI database. The assignment of a valine at that position in the Neotyphodium sp. chitinase is supported by multiple lines of evidence. The corresponding DNA sequence was obtained both by PCR of genomic DNA and from the cDNA clone. At the protein level, MS/MS peptide sequencing of the 52-kDa apoplastic protein identified two peptides YVDLYNFMAVDYAGPSFSK and YVDLYNFMAVDYAGPSFSKK, containing the valine. Together, these three sequence approaches confirm the assignment of a valine at position 258 in the Neotyphodium chitinase.
That the Neotyphodium sp. chitinase was the major secreted protein produced in culture suggested it was not regulated by either catabolite repression or induced by chitin because the medium contained sucrose and was not supplemented with chitin. This is in contrast to the regulation of expression of some other secreted fungal chitinases that are subject to catabolite repression and chitin induction (Blaiseu et al 1992, Garcia et al 1994).
The chitinase also was highly expressed within the infected plant as evidenced by the high message level seen in RNA blots and the high representation in the cDNA library. In screening 150 000 plaques, 81 positives were obtained, suggesting that 1 out of every 2000 cDNA molecules in the library was from the fungal chitinase even though most of the clones in the library must have originated from the host. In leaf sheaths of perennial ryegrass infected by N. lolii, the fungal biomass constituted 0.2% of the infected tissue biomass (Tan et al 2001). Since the fungal mycelial mass is a small component of the mass of the infected grass tissue used for RNA isolation, the fungal chitinase message must constitute a major component of the fungal messages in the Neotyphodium sp./P. ampla interaction. Expression of similar chitinases was detected in other grass species infected with other fungal endophyte species, suggesting it might be a general feature of endophyte infection.
The Neotyphodium sp. chitinase was secreted actively from the fungal cells as evidenced by its recovery from the culture filtrate and the plant apoplast and by the presence of a signal sequence. Some fungal chitinases have been reported that have amino acid similarity in the regions of the mature protein but which have no signal sequence and are not secreted from the cell (Kim et al 2002, Takaya et al 1998). Takaya et al (1998) suggested that such intracellular chitinases play a role in fungal morphogenesis.
Fungal secreted proteins are of interest in endophyte-host interactions because they act at the interface of the two species. In natural environments, fungi rely on a range of secreted hydrolytic enzymes to supply the small molecules required for their nutritional needs. Secreted chitinases have been studied from numerous fungal species in the context of degradation of environmental chitin. Chitin is an abundant biopolymer in nature (Robertus and Monzingo 1999), being a major component of insect exoskeletons, crustacean shells and fungal cell walls. Ecological recycling of chitin occurs through chitinase-mediated degradation carried out by saprophytic organisms, including fungi, which use chitin as a nutritional source (Gooday 1990). Fungal chitinases also are associated with the process of mycoparasitism (Goldman et al 1994, Herrera-Estrella and Chet 1999) and can be considered a special case of secreted enzymes involved in fungal nutrition. Similarly, entomopathogenic fungi secrete chitinases that are believed to be involved in digestion of the insect cuticle (St. Leger et al 1996).
Aging fungal cultures experiencing nutrient limitation undergo a process of autolysis, which releases nutrients from older cells to support new growth at the hyphal apex (White et al 2002). Secreted chitinases and other hydrolytic enzymes including proteases and glucanases are associated with this process in many fungal species (White et al 2002). In nature, this also is likely to occur to allow fungal mycelia to continue growth through the environmental substrate. The role of chitinases in autolysis can be considered another special case of a nutritional function.
Secreted chitinases also have been found to have roles in fungal morphogenesis. In Saccharomyces cerevisiae, chitin is localized to the junction between mother and daughter cells and a secreted chitinase is required to release the newly formed daughter cells (Kuranda and Robbins 1991). Secreted chitinases also are involved in hyphal branching and in spore germination (Gooday et al 1992). Distal to the plastic growing hyphal tip region, the mature cell wall of filamentous fungi generally is composed of chitin and ß-1,3-glucans that are cross-linked by ß-1,6-glucans resulting in a rigid structure (Wessels 1993, 1994). The structures of the cross-links among cell wall components have been identified in mature yeast cell walls (Kollar et al 1997). In filamentous fungi, for hyphal branching to occur wall loosening of the rigid mature wall by the concerted action of several hydrolytic enzymes, including chitinases, is required (Gooday et al 1992, Rast et al 1991, Wessels 1999).
The Neotyphodium sp. endochitinase might be involved in any of the above processes. In culture, hyphal branching and conidiospore development and germination are expected as well as possibly wall re-assimilation. Within the apoplast of the infected plant, occasional branching of the fungal hyphae does occur but conidia formation has not been observed. Since the apoplast is a nutrient poor environment, autolysis of older mycelia and re-assimilation of nutrients for continued growth may be occurring. The presence of the endophyte chitinase within the apoplast of the host tissues also might be a factor in the reported resistance of some endophyte-infected grasses to fungal pathogens (Funk et al 1994). The endophytic subtilisin-like proteinase (Reddy et al 1996) and ß-1,6-glucanase (Moy et al 2002) that also are secreted into the apoplast might be acting synergistically with the chitinase in any or all of the above possible physiological functions.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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1 Corresponding author. E-mail: belanger{at}aesop.rutgers.edu
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LITERATURE CITED |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
|---|
Barker GM, Pottinger RP, Addition PJ, Prestidge RA. 1984. Effect of Lolium endophyte fungus infections on behavior of adult Argentine stem weevil. NZ J Agric Res 27:271–277.
Blaiseau P-L, Kunz C, Grison R, Bertheau Y, Brygoo Y. 1992. Cloning and expression of a chitinase gene from the hyperparasitic fungus Aphanocladium album. Curr Genet 21:61–66.[Medline]
Bortone K, Monzingo AF, Ernst S, Robertus JD. 2002. The structure of an allosamidin complex with the Coccidioides immitis chitinase defines a role for a second acid residue in substrate-assisted mechanism. J Mol Biol 320: 293–302.[Medline]
Breen JP. 1994. Acremonium endophyte interactions with enhanced plant resistance to insects. Annu Rev Entomol 39:401–423.
Bush LP, Wilkinson HH, Schardl CL. 1997. Bioprotective alkaloids of grass-fungal endophyte symbioses. Plant Physiol 114:1–7.[Medline]
Christensen MJ, Bennett RJ, Schmid J. 2002. Growth of Epichloë/Neotyphodium and p-endophytes in leaves of Lolium and Festuca grasses. Mycol Res 106:93–106.
Clay K. 1988. Fungal endophytes of grasses: a defensive mutualism between plants and fungi. Ecology 69:10–16.
———, Schardl C. 2002. Evolutionary origins and ecological consequences of endophyte symbiosis with grasses. Am Nat 160:S99–S127.[Medline]
Cohen-Kupiec R, Chet I. 1998. The molecular biology of chitin digestion. Curr Opin Biotechnol 9:270–277.[Medline]
De La Cruz J, Hidalgo-Gallego A, Lora JM, Benitez T, Pintor-Toro JA, Llobell A. 1992. Isolation and characterization of three chitinases from Trichodema harzianum. Eur J Biochem 206:859–867.[Medline]
Funk CR, Belanger FC, Murphy JA. 1994. Role of endophytes in grasses used for turf and soil conservation. In: Bacon CW, White JF Jr, eds. Biotechnology of endophytic fungi of grasses. Boca Raton, Florida: CRC Press. p 201–209.
———, Halisky PM, Johnson MC, Siegel MR, Stewart AV, Ahmad S, Hurley RH, Harvey IC. 1983. An endophytic fungus and resistance to sod webworms: association in Lolium perenne. Bio/Technology 1:189–191.
Garcia I, Lora JM, de la Cruz J, Benitez T, Llobell A, Pintor-Toro J. 1994. Cloning and characterization of a chitinase (CHIT42) cDNA from the mycoparasitic fungus Trichoderma harzianum. Curr Genet 27:83–89.[Medline]
Glenn AE, Bacon CW, Price R, Hanlin RT. 1996. Molecular phylogeny of Acremonium and its taxonomic implications. Mycologia 88:369–383.
Goldman GH, Hayes C, Harman GE. 1994. Molecular and cellular biology of biocontrol by Trichoderma spp. Trends Biotechnol 12:478–482.[Medline]
Gooday GW. 1990. Physiology of microbial degradation of chitin and chitosan. Biodegradation 1:177–190.
———, Zhu W-Y, O’Donnell RW. 1992. What are the roles of chitinases in the growing fungus? FEMS Microbiol Lett 100:387–392.
Henikoff S, Henikoff JG. 1992. Amino acid substitution matrices from protein blocks. Proc Natl Acad Sci USA 89: 10915–10919.
Henrissat B, Bairoch A. 1993. New families in the classification of glycosyl hydrolases based on amino acid sequence similarities. Biochem J 293:781–788.
Herera-Estrella A, Chet I. 1999. Chitinases in biological control. In: Jolles P, Muzzarelli RAA, eds. Chitin and Chitinases. Basel, Switzerland: Birkhauser Verlag. p 171–183.
Hollis T, Monzingo AF, Bortone K, Ernst S, Cox R, Robertus JD. 2000. The X-ray structure of a chitinase from the pathogenic fungus Coccidioides immitis. Protein Sci 9: 544–551.[Medline]
Johnson-Cicalese J, Secks ME, Lam CK, Meyer WA, Murphy JA, Belanger FC. 2000. Cross species inoculation of Chewings and strong creeping red fescues with fungal endophytes. Crop Sci 40:1485–1489.
Kim D-J, Baek J-M, Uribe P, Kenerley CM, Cook DR. 2002. Cloning and characterization of multiple glycosyl hydrolase genes from Trichoderma virens. Curr Genet. 40: 374–384.[Medline]
Kollar R, Reinhold BB, Petrakova E, Yeh HJC, Ashwell G, Drgonova J, Kapteyn JC, Klis FM, Cabib E. 1997. Architecture of the yeast cell wall ß(1–6)-glucan interconnects mannoprotein, ß(1–3)-glucan, and chitin. J Biol Chem 272:17762–17775.
Kuranda MJ, Robbins PW. 1991. Chitinase is required for cell separation during growth of Saccharomyces cerevisiae. J Biol Chem 266:19758–19767.
Laemmli UK. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680–685.[Medline]
Lam CK, Belanger FC, White JF Jr, Daie J. 1995. Invertase activity in Epichloë/Acremonium fungal endophytes and its possible role in choke disease. Mycol Res 99:867–873.
Latch GCM, Christensen MJ, Samuels GJ. 1984. Five endophytes of Lolium and Festuca in New Zealand. Mycotaxon 20:535–550.
———. 1997. An overview of Neotyphodium-grass interactions. In: Bacon CW, Hill NS, eds. Neotyphodium/grass interactions. New York: Plenum Press. p 1–11.
Leuchtmann A, Schardl CL, Siegel MR. 1994. Sexual compatibility and taxonomy of a new species of Epichloë symbiotic with fine fescue grasses. Mycologia 86:802–812.
Lindstrom JT, Belanger FC. 1994. Purification and characterization of an endophytic fungal proteinase that is abundantly expressed in the infected host grass. Plant Physiol 106:7–16.[Abstract]
Lingappa Y, Lockwood JL. 1961. Chitin media for selective isolation and culture of actinomycetes. Phytopathology 52:317–323.
McCreath KJ, Gooday GW. 1992. A rapid and sensitive microassay for determination of chitinolytic activity. J Microbiol Meth 14:229–237.
Moy M, Li HM, Sullivan R, White JF Jr, Belanger FC. 2002. Endophytic fungal ß-1,6-glucanase expression in the infected host grass. Plant Physiol 130:1298–1308.
Nielsen H, Engelbrecht J, Brunak S, von Heijne G. 1997. Identification of prokaryotic and eukaryotic signal peptides and prediction of their cleavage sites. Protein Eng 10:1–6.
Reddy PV, Lam CK, Belanger FC. 1996. Mutualistic fungal endophytes express a proteinase that is homologous to proteases suspected to be important in fungal pathogenicity. Plant Physiol 111:1209–1218.[Abstract]
Roberts WK, Selitrennikoff CP. 1988. Plant and bacterial chitinases differ in antifungal activity. J Gen Microbiol 134:169–176.
Robertus JD, Monzingo AF. 1999. The structure and action of chitinases. In: Jolles P, Muzzarelli RAA, eds. Chitin and chitinases. Basel, Switzerland: Birkhauser Verlag. p 125–135.
Sahai AS, Manocha MS. 1993. Chitinases of fungi and plants: their involvement in morphogenesis and host-parasite interaction. FEMS Microbiol Rev 11:317–338.
Schagger H, von Jagow G. 1987. Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal Biochem 166:368–379.[Medline]
Schardl CL. 2001. Epichloë festucae and related mutualistic symbionts of grasses. Fung Genet Biol 33:69–82.
Scott B. 2001. Epichloë endophytes: fungal symbionts of grasses. Curr Opin Microbiol 4:393–398.[Medline]
St Leger RJ, Joshi L, Bidochka MJ, Rizzo NW, Roberts DW. 1996. Characterization and ultrastructural localization of chitinases from Metarhizium anisopliae, M. flavoviride, and Beauveria bassiana during fungal invasion of host (Manduca sexta) cuticle. Appl Environ Microbiol 62:907–912.
Swofford L. 2002. PAUP*: Phylogenetic analysis using parsimony (* and other methods). Version 4. Sunderland, Massachusetts: Sinauer Associates.
Takaya N, Yamazaki D, Horiuchi H, Ohta A, Takagi M. 1998. Intracellular chitinase gene from Rhizopus oligosporus: molecular cloning and characterization. Microbiology 144:2647–2654.
Tan YT, Spiering MJ, Scott V, Lane GA, Christensen MJ, Schmid J. 2001. In planta regulation of extension of an endophytic fungus and maintenance of high metabolic rates in its mycelium in the absence of apical extension. Appl Environ Microbiol 67:5377–5383.
Tronsmo A, Harman G. 1993. Detection and quantification of N-acetyl-ß-D-glucosaminidase, chitobiosidase, and endochitinase in solutions and on gels. Anal Biochem 208:74–79.[Medline]
Wessels JGH. 1993. Wall growth, protein excretion and morphogenesis in fungi. New Phytol 123:397–413.
———. 1994. Developmental regulation of fungal cell wall formation. Annu Rev Phytopathol 32:413–437.
Wessels JGH. 1999. Fungi in their own right. Fungal Genet Biol 27:134–145.[Medline]
White S, McIntyre M, Berry DR, McNeil B. 2002. The autolysis of industrial filamentous fungi. Crit Rev Biotechnol 22:1–14.[Medline]
Zhang Z, Yuen GY, Sarah G, Penheiter AR. 2001. Chitinases from the plant disease biocontrol agent, Stenotrophomonas maltophilia C3. Phytopathology 91:204–211.[Medline]
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