1 Department of Biology, University of Saskatchewan, 112 Science Place, Saskatoon, SK, Canada S7N 5E2
2 Department of Plant Sciences, University of Saskatchewan, 51 Campus Drive, Saskatoon, SK, Canada S7N 5A8
3 Plant Biotechnology Institute, National Research Council of Canada, 110 Gymnasium Place, Saskatoon, SK, Canada S7N 0W9
Correspondence
Yangdou Wei
yangdou.wei{at}usask.ca
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ABSTRACT |
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INTRODUCTION |
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Plant-pathogenic fungi produce an array of extracellular hydrolytic enzymes that enable them to penetrate and infect the host tissue (Knogge, 1996; Kolattukudy, 1985
; Oliver & Osbourn, 1995
) and are collectively called cell-wall-degrading enzymes (CWDEs). These enzymes may contribute to pathogenesis by degrading wax, cuticle and cell walls, thus aiding tissue invasion and pathogen dissemination. Furthermore, they can act as elicitors of host defence reactions and may also play a nutritional role during certain stages of the fungal life cycle. Early studies suggested that fungal cutinases played an important role in the penetration of plant surfaces. However, contradictory evidence as to the importance of cutinases in disease establishment has emerged from disruption studies of individual cutinase-encoding genes in several pathosystems (Crowhurst et al., 1997
; Stahl & Schäfer, 1992
; Sweigard et al., 1992
; van Kan et al., 1997
). It has been proposed that cutinolytic activities expressed by pathogenic fungi during the infection process could be contributed by other enzymes (Comménil et al., 1995
, 1998
, 1999
; Nasser Eddine et al., 2001
; van Kan et al., 1997
). Upon contact with the host surface, plant-pathogenic fungi often produce an extracellular matrix underneath the fungal germling, a phenomenon of the prepenetration process that determines the success of infection and disease development. Production of secreted lipolytic activities associated with the extracellular matrix has been reported among obligately biotrophic powdery mildew (Roberts & Mims, 1998
) and rust (Deising et al., 1992
), hemibiotrophic Colletrotrichum (Pain et al., 1996
) and necrotrophic Botrytis (Doss, 1999
). Thus, it is likely that secreted fungal lipolytic activities play an important role in the infection processes of these pathogens.
Fusarium head blight (FHB) has emerged in recent years as a major disease causing damage on wheat, barley and other small grains in North America (McMullen et al., 1997). FHB can reduce seed quality and yield significantly due to the production of discoloured, shrivelled tombstone kernels. Fusarium-infected grain is also often contaminated with the mycotoxins trichothecenes and zearalenone (McMullen et al., 1997
), making it unsuitable for food or feed. Fusarium graminearum Schwabe [teleomorph Gibberella zeae (Schw.) Petch] is the major pathogen responsible for this disease, but other Fusarium species, especially Fusarium culmorum, also play a role (Gilbert & Tekauz, 2000
). Pathogens of the genus Fusarium can also cause head blight and root rot on many other plant species. The importance of F. graminearum as the most agriculturally significant pathogen is also highlighted by the recently coordinated efforts of the F. graminearum genome project. Through the USDA/NSF microbial genome sequencing program, a 36 Mb assembly has been released based on
10x genome coverage from shotgun sequencing of the F. graminearum genome (http://www.broad.mit.edu/annotation/fungi/fusarium). The rapid availability of the genome sequences of F. graminearum and other fungal species will immediately promote functional identification of many fungal genes, specifically for those fungi in which gene disruption is feasible through transformation approaches.
Based on the unique catalytic property of fungal lipase and its possible role in pathogenesis, we have identified and characterized a secreted lipase gene (LIP1) from F. graminearum strain PH-1. Expression levels of LIP1 were examined in vitro and in planta during infection. The function of LIP1 was further investigated by gene replacement studies, indicating that LIP1 was essential for lipid utilization rather than for fungal pathogenicity.
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METHODS |
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Chemicals.
All restriction enzymes were purchased from New England Biolabs. Agar, PDA, tryptone, peptone, yeast extract and yeast nitrogen base without amino acids and without ammonium sulfate (YNB) were purchased from Difco. -D-Glucanase and Driselase were purchased from Interspex Products. Other chemicals, unless stated otherwise, were purchased from Sigma-Aldrich.
Cloning of LIP1.
Three genomic sequences were amplified from F. graminearum by PCR using Pfu polymerase (Stratagene) and specific primer pairs (Table 1). F1/R1 were used to amplify LIP1 including its coding region and 5'- and 3'-flanking sequences. F2/R2 and F3/R3 were used to amplify the LIP1 coding region with and without the predicted signal peptide, respectively. The 4532 bp fragment amplified by F1/R1 was cut with SacII/ApaI and cloned into the same restriction sites of pBluescript II KS+ (Stratagene). The other two fragments were cloned into TA vector (pBluescript II KS+). The resultant plasmids were named pLIP1, pLIP1ORF and pLIP1ORF-SP and transformed into Escherichia coli strain DH5
. All PCR products were confirmed by sequencing.
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Transcript induction of LIP1.
CDC Teal wheat spikes were inoculated with wild-type strain PH-1 by the spore-droplet method following Jenczmionka et al. (2003). The samples were collected 5 days post-inoculation from healthy and infected spikes. For in vitro induction tests, PH-1 was grown at 24 °C and shaken at 130 r.p.m. in minimal medium supplemented with 1 % (w/v) carbon sources as required. Wheat cell-wall material, used as a potential inducer of CWDE gene expression, was prepared from leaves of CDC Teal by the method described by Lehtinen (1993)
.
Construction of gene replacement vector and fungal transformation.
To construct the gene replacement vector pLIP1, a 3·2 kb cassette containing the E. coli hygromycin B phosphotransferase gene (hygR) that has been rendered suitable for fungal expression was cut from vector pGC1-1 (Rikkerink et al., 1994
) by SalI/HindIII and inserted into the same sites of the pLIP1 backbone. As a result, hygR replaced the LIP1 coding region and was flanked by border sequences from the wild-type genomic locus. The left and right flanking sequences were 1·2 kb (SacIISalI fragment) and 1·0 kb (HindIIIKpnI fragment), respectively (see Fig. 4a
). For fungal transformation, protoplasts were prepared as described previously (Hohn & Desjardins, 1992
) except that the enzymes used to degrade the fungal cell wall were 1 %
-D-glucanase, 0·1 %
-glucuronidase and 2 % Driselase. For reliable transformation, a concentration of at least 108 protoplasts ml1 was used. Transformation of the protoplasts with the p
LIP1 plasmids was conducted according to the protocols described by Wei et al. (2004)
. After transformation, hygromycin B-resistant mutants were preliminarily screened by PCR amplification with primers F3/R3.
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Northern and Southern hybridization.
Total RNA was extracted from 5 g fresh mycelium grown in minimal medium with different treatments. Genomic DNAs extracted from the wild-type or mutant mycelium were digested for 5 h with KpnI before being subjected to electrophoretic separation. Northern and Southern hybridizations were conducted following standard techniques (Sambrook & Russell, 2001).
Confocal microscopy.
In the rhodamine B plate assays, oil droplets produced during hydrolysis of triglyceride lipids by secreted fungal lipases were examined under a confocal laser scanning electron microscope (LSM510; Zeiss) using excitation/emission wavelengths of 405/560615 nm.
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RESULTS |
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A dendrogram was constructed based on comparison of the deduced amino acid sequences of LIP1, the closest BLAST hits from other fungal species and 11 putative C. rugosa lipase-like genes of F. graminearum (Fig. 1a). LIP1 clustered with and showed 61, 61 and 57 % identity to the putative lipase sequences of Neurospora crassa, Magnaporthe grisea and B. cinerea, respectively, suggesting that they are orthologues of LIP1 in other fungal genomes. The other 11 putative lipases from F. graminearum showed little similarity (<37 %) both to each other and to the lipase cluster containing LIP1. At the time of analysis, none of these putative genes had been investigated further.
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LIP1 encodes a triglyceride lipase
P. pastoris strain X-33 does not produce secreted lipolytic activity on culture plates. We introduced the LIP1 gene with or without the sequence encoding the signal peptide into vectors pPICZA and pPICZA, respectively. Extracellular lipolytic activity of the transformed yeast was determined by substrate hydrolysis, which produced a clear zone around the colony on tributyrin emulsion plates. Clear zones formed only around colonies of transformants with the pPICZ
A-Lip1ORF-SP vector (Fig. 2
a), but not in yeast transformed with pPICZA-Lip1ORF or with the empty vectors. It is therefore likely that the putative LIP1 signal peptide of F. graminearum could not function in P. pastoris strain X-33.
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Expression of LIP1 in planta and in vitro
To test whether LIP1 expression occurs during the infection process, we performed Northern blot analysis using total RNA extracted from CDC Teal wheat spikes 5 days post-inoculation with F. graminearum wild-type strain PH-1 and from uninoculated healthy spikes. LIP1 transcripts were detected in the infected wheat spikes but not in healthy spikes (Fig. 3a). Induction of LIP1 expression was also examined in liquid minimal medium supplemented with various carbon sources. Northern blot analysis revealed that LIP1 expression was induced dramatically in medium supplemented with WGO. Lower expression of LIP1 was also detected when the fungus was grown in medium containing olive oil or triolein. In contrast, no expression was detected from fungal cultures grown in the minimal medium supplemented with glucose, sucrose, tributyrin, cell-wall material or apple pectin as the sole carbon source. To test whether prepared cell-wall material is an efficient inducer of the fungal CWDEs, the same blot was further probed with an F. graminearum gene (FgCel; EAA73192) encoding a cellulase. Expression of FgCel was detected only when the fungus was grown in a medium containing cell-wall material prepared from wheat leaves. An expression time course was conducted in minimal medium supplemented with either WGO or glucose as the sole carbon source. No expression of LIP1 was detected in fungal cultures grown in the minimal medium supplemented with glucose during the time course. Induction of LIP1 by WGO started 12 h after culturing and the expression level increased progressively up to 72 h (Fig. 3c
).
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Targeted gene disruption of LIP1
To investigate the potential function of LIP1 in fungal growth or pathogenicity, targeted gene disruption was conducted by PEG-mediated protoplast transformation. Three independent transformation experiments produced 54 hygromycin-B-resistant transformants. These transformants were preliminarily screened for absence of the native LIP1 sequence by PCR with primers F3/R3 (Table 1). Seven putative gene replacement strains were selected and the remaining transformants were considered ectopic integration strains. Genomic DNAs isolated from these seven strains as well as three ectopic strains and the wild-type PH-1 strain were further analysed by Southern hybridization. Genomic DNA was digested with KpnI, which has one cutting site in the 3' homologous region and an additional cutting site in either the hygR or LIP1 coding region, resulting in a 3·2 or 2·4 kb fragment from replacement and native sequences, respectively. Southern hybridization (Fig. 4b
) showed that only the 3·2 kb fragment was present in the seven gene replacement strains and both the 2·4 and 3·2 kb fragments were present in the three ectopic integration strains. In the wild-type strain, only the 2·4 kb fragment was present. Southern blot analysis also demonstrated the replacement of native LIP1 by vector DNA in all seven transformants. Null mutation of LIP1 in the seven replacement strains was further confirmed by Northern analysis (Fig. 4c
). No LIP1 transcript accumulation was observed in the seven replacement strains grown in minimal medium containing 1 % WGO for 48 h. In contrast, a single transcript was detected from the three ectopic integration strains and the wild-type PH-1 strain (Fig. 4c
).
LIP1 encodes a secreted lipase
Although secretion was predicted for 11 of the 12 lipase gene products in our search, the deduced native signal peptide of LIP1 failed to function in recombinant P. pastoris. We therefore screened the wild-type F. graminearum and its knock-out mutants for utilization of triolein or tristearin in agar plates containing rhodamine B. As an additional control, the ectopic integration strains in which the native gene is intact were also used. After 2 days of incubation, strong fluorescence was observed around the colonies from the wild-type strain PH-1 and ectopic integration strains on both triolein- and tristearin-containing plates, but not from the seven null-mutant strains (Fig. 5a, b). Rhodamine B was not taken up by fungal mycelium, since no fluorescence was detected in the mycelia of either
LIP1 mutants or PH-1 grown on rhodamine B plates (Fig. 5c, d
). Fluorescence was observed from oil droplets surrounding the wild-type mycelium, but absent from that of the
LIP1 mutants, indicating that secreted lipolytic activities were completely abolished in the
LIP1 knock-out mutants. This implied that LIP1 is the major detectable lipase secreted under the growth conditions employed. We continued the incubation to see whether additional lipolytic activity could be detected in the knock-out mutants. Following an extended incubation period, fluorescence was also detected surrounding colonies of the
LIP1 mutants (Fig. 5e
). These results establish that, while LIP1 is the major secreted lipase isoform that is induced by saturated and unsaturated lipid substrates, additional lipolytic activities appear later during growth.
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DISCUSSION |
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Recently, involvement of fungal secreted lipases in infection has been shown using molecular techniques. Comménil et al. (1995, 1998
, 1999)
purified and partially sequenced a 60 kDa extracellular lipase from B. cinerea and showed that a specific antibody raised against this lipase suppressed lesion formation on tomato leaves. A similar lipase gene discovered through expressed sequence tag analyses of Blumeria graminis germinating conidia showed a dramatic increase in transcript abundance during conidium germination and appressorium formation (Thomas et al., 2001
, 2002
; Y. Wei, unpublished results). In the F. graminearum genome database, we identified the gene LIP1, which shares high sequence similarity with these Botrytis and Blumeria lipases. However, our gene disruption study showed that
LIP1 mutants and the wild-type strain caused similar patterns of FHB symptom development on susceptible hosts. Similar results were recently reported for the corresponding Botrytis lipase gene on its host plant bean (Reis et al., 2005
). In contrast, the F. graminearum gene FGL1, encoding a 35 kDa putative secreted lipase, belonging to a Rhizomucor miehei lipase family (Schmidt-Dannert, 1999
), is required for virulence of F. graminearum on wheat and maize (Nasser Eddine et al., 2001
; Voigt et al., 2005
).
Apart from their preference for triglycerides, lipases catalyse the hydrolysis and synthesis of a broad range of natural water-insoluble esters, as well as alcoholysis, acidolysis, esterification and aminolysis (Pandey et al., 1999). Two Aspergillus niger enzymes, FAEA and FAEB, originally isolated from a commercial pectinase preparation, showed ferulic acid esterase activity in degradation of complex cell-wall polysaccharides (de Vries et al., 1997
, 2002
). Remarkably, these two proteins have significant sequence similarity to F. graminearum FGL1 and were grouped into the fungal lipase family in the database (http://bioweb.ensam.inra.fr/ESTHER/general). Although C. rugosa and R. miehei lipases belong to the same class of the
/
hydrolase fold family and share similar activation and catalytic mechanisms (Schmidt-Dannert, 1999
; Schmidt-Dannert et al., 1998
), considerably different substrate-binding sites between the two types of lipases explain their varying substrate specificity. For instance, C. rugosa lipases have a tunnel-like binding site and are likely to accept substrates with long-chain fatty acids, whereas R. miehei lipases have a crevice-like binding site and can accept bulkier substrates (Schmidt-Dannert, 1999
; Schmidt-Dannert et al., 1998
). Thus, it will be necessary to have information on the substrate specificity of F. graminearum FGL1 to determine its role in pathogenicity in contrast to the lack of such a role for LIP1. Both F. graminearum lipase genes FGL1 and LIP1 are expressed during infection. The expression of both genes is induced in vitro by WGO and suppressed by sucrose or glucose. In contrast, no FGL1 transcripts were detected in fungal culture incubated in water (Voigt et al., 2005
), whereas expression of LIP1 was significantly induced when the fungus was cultured in sugar-deficient minimal medium (data not shown). These results suggested that overlapping, but distinct regulatory mechanisms are involved in induction of LIP1 and FGL1 gene expression.
Expression of the LIP1 gene was induced strongly by WGO, weakly by triolein and olive oil, but not by tributyrin. Surprisingly, we found that the hydrolytic products had strong and different effects on regulation of LIP1 expression. Long-chain saturated fatty acids such as palmitic acid (C16 : 0; data not shown) and stearic acid (C18 : 0) appeared to be strong inducers, whereas long-chain unsaturated fatty acids such as linoleic acid (C18 : 2) acted as repressors. The association of long-chain saturated fatty acids with induction of LIP1 expression and long-chain unsaturated fatty acids with its repression suggests that the lipolytic products are major regulatory components controlling the transcription of LIP1. In S. cerevisiae, a similar pattern of regulation of the OLE1 gene, which encodes the -9 desaturase, has been reported; the level of OLE1 gene transcription was increased in response to exogenous saturated fatty acids, whereas exposure to unsaturated fatty acids sharply reduced transcription (McDonough et al., 1992
). Although the corresponding positive and negative response elements have been characterized in the OLE1 upstream promoter region (Choi et al., 1996
; McDonough et al., 1992
), preliminary sequence comparison did not reveal the same elements present in the upstream region of the LIP1 gene. The transcription activation and/or repression elements required for fatty-acid-mediated LIP1 expression remain to be determined.
The LIP1 strains were deficient in secreted lipolytic activity on tristearin and showed delayed activity for hydrolysis of triolein. After a long incubation period, lipolytic activities, particularly for triolein, also appeared in the
LIP1 strains, indicating that additional secreted lipases participate in exogenous lipid hydrolysis. A genome-wide survey revealed 11 additional sequences encoding C. rugosa-family lipases present in the F. graminearum genome. Some of these putative lipases might be responsible for the enzyme activity that appeared at late stages of fungal growth under these conditions. In the
LIP1 strains, besides their lack of activity on saturated tristearin, hydrolytic activities on triolein were also delayed significantly. Thus, it is reasonable to expect that the product fatty acids produced by LIP1-catalysed lipid hydrolysis regulate expression of other lipase genes that control lipid hydrolysis in the later stages of growth.
Apart from reduced or abolished pigmentation, colony morphology of LIP1 mutants was identical to the wild-type on minimal and complete media. However, the
LIP1 strains were unable to grow in liquid minimal medium containing tristearin as the sole carbon source. We propose that LIP1 functions primarily when exogenous lipids with long-chain saturated fatty acids are present, acts as a regulator to coordinate other lipase gene expression and plays a role in fungal nutrient acquisition but not in pathogenesis.
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ACKNOWLEDGEMENTS |
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Received 13 June 2005;
revised 23 September 2005;
accepted 27 September 2005.
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