| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
1 Citrus Research and Education Center, Institute of Food and Agricultural Sciences (IFAS), University of Florida, 700 Experiment Station Road, Lake Alfred, FL 33850, USA
2 Department of Plant Pathology, National Chung-Hsing University, Taichung 402, Taiwan
3 Department of Plant Pathology, Institute of Food and Agricultural Sciences (IFAS), University of Florida, 700 Experiment Station Road, Lake Alfred, FL 33850, USA
Correspondence
Kuang-Ren Chung
krchung{at}ufl.edu
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
The GenBank/EMBL/DDBJ accession numbers. for the CTB5, CTB6 and CTB7 sequences of Cercospora nicotianae are DQ991507, DQ991508 and DQ991509 respectively.
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
). Host-selective toxins kill plant cells by targeting a specific cellular enzyme or component, and thus are toxic only to a limited range of host cultivars (Walton, 1996). In contrast, non-host-selective toxins, targeting various cellular components, enable the producing pathogens to have wide host ranges. Cercosporin (Fig. 1a) is a non-host-selective perylenequinone toxin produced by many phytopathogenic Cercospora species, which have been reported to affect several hundred plant species, including many major crops such as corn, rice, banana, coffee, sugar beet, soybean, peanut and tobacco (reviewed by Daub & Ehrenshaft, 2000; Daub et al., 2005). Cercosporin is an important virulence determinant in Cercospora species (Callahan et al., 1999; Choquer et al., 2005, 2007; Dekkers et al., 2007; Shim & Dunkle, 2003; Upchurch et al., 1991). Compared to other non-host-selective phytotoxins, cercosporin has several unique features, including light activation for its biosynthesis (Ehrenshaft & Upchurch, 1991), light- and oxygen-dependent cytotoxicity (Yamazaki et al., 1975), and the production of reactive oxygen species such as singlet oxygen and superoxide (Daub & Hangarter, 1983). In the past two decades, intensive research has been focused on elucidation of the genetic mechanisms involved in self-protection from cercosporin and singlet-oxygen-generating compounds (Chung et al., 1999, 2003a; Daub et al., 1992, 2000; Ehrenshaft et al., 1998, 1999; Jenns & Daub, 1995; Sollod et al., 1992).
|
) and its structure was chemically determined by Yamazaki & Ogawa (1972). In addition to plants, cercosporin has been shown to be toxic to various cell types, including bacteria and many fungi and animal cells, due to the production of reactive oxygen species (Daub et al., 2005). Cercosporin is capable of breaking down different cellular components such as lipids, proteins and nucleic acids, depending on its localization in cells. During pathogenesis in host plants, cercosporin has been shown to damage cell membranes by causing lipid peroxidation, electrolyte leakage and eventually cell death (Daub, 1982; Daub & Briggs, 1983).
Based on substrate feeding experiments (Okubo et al., 1975) cercosporin has been proposed to be synthesized by a fungal polyketide pathway. However, the genes or enzymes involved in cercosporin production remain largely unknown. Recent studies with molecular and genetic tools began to uncover the cercosporin biosynthetic pathway and regulation network (Daub et al., 2005). Light has long been known to be required for cercosporin toxicity. Biosynthesis of cercosporin is also primarily triggered by light (Jenns et al., 1989). In order to elucidate cercosporin biosynthesis and regulation at molecular levels, several cercosporin-deficient mutants were identified via a restriction-enzyme-mediated mutagenesis approach (Chung et al., 2003b). As a result, two linked genes, CTB1 (encoding a polyketide synthase) and CTB3 (encoding a dual methyltransferase/monooxygenase) that were required for cercosporin biosynthesis were identified and characterized from Cercospora nicotianae (Choquer et al., 2005; Dekkers et al., 2007). We later obtained nine additional ORFs beyond the boundaries of CTB1 and CTB3 by combining chromosome walking and sequence analysis. Six of them (CTB2, CTB4, CTB5, CTB6, CTB7 and CTB8) encode polypeptides proposed to be involved in cercosporin production (Chen et al., 2007). Expression of eight of the genes was induced under cercosporin-producing conditions and was co-ordinately regulated by the Zn(II)Cys6 transcriptional activator, CTB8 (Chen et al., 2007). We hypothesize that cercosporin, like many fungal secondary metabolites (Keller et al., 2005), is synthesized by a cluster of co-regulated genes, in this case designated the cercosporin toxin biosynthesis (CTB) gene cluster. The functions of CTB1, CTB2, CTB3, CTB4 and CTB8 in cercosporin biosynthesis have been unambiguously demonstrated by analysing the respective null mutants (Chen et al., 2007; Choquer et al., 2005, 2007; Dekkers et al., 2007). The functions of CTB5, CTB6 and CTB7, which encode proteins similar to numerous FAD/FMN- or NADPH-dependent oxidoreductases or dehydrogenases, remain to be elucidated. In the present study, we characterized these genes by creating loss- and gain-of-function strains; we present conclusive evidence to demonstrate their roles in cercosporin biosynthesis, and completely define the core CTB gene cluster.
|
METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
). Cercosporin-production mutants were screened daily for the lack of red pigment production on thin potato dextrose (glucose) agar (PDA, Difco, Becton, Dickinson and Company) plates by a method described previously (Chung et al., 2003b). We found that a thin PDA plate (less than 15 ml medium in a 100x15 mm Petri dish) supported the highest production of cercosporin under illumination. Assays for sensitivity to photosensitizing compounds (cercosporin, eosin Y, haematoporphyrin, methylene blue or toluidine blue) were performed by growing fungal isolates on CM containing 10 or 100 µM test compound under continuous light as described previously (Jenns & Daub, 1995). All chemicals were purchased from Sigma-Aldrich unless otherwise stated, and dissolved appropriately in acetone or water to make a 10 or 100 mM stock solution.
Cercosporin analysis.
Cercosporin was extracted with 5 M KOH or with ethyl acetate from agar plugs with mycelia as described previously (Choquer et al., 2005; Chung, 2003). Cercosporin in KOH extracts was quantified by measuring A480 using a Genesys 5 spectrophotometer (Spectronic Instruments). Ethyl acetate extracts were analysed on a TLC plate coated with a 60 F254 fluorescent silica gel with ethyl acetate/hexane/methanol/H2O (6 : 4 : 1.5 : 1, by vol.) as a solvent (Choquer et al., 2005).
CTB5, CTB6 and CTB7 gene disruption.
A strategy employing the split-hygromycin phosphotransferase B gene (HYG) marker fused with truncated CTB5, CTB6 or CTB7 fragments was used for targeted gene disruption in C. nicotianae as described previously (Choquer et al., 2005). All DNA clones were built on the backbone of pGEM-T easy vector (Promega). For CTB5 disruption, a 3.3 kb fragment was amplified with primers CTB5R (5'-GCTACAGTGCGACGGAGTCCTG-3') and CTB7I (5'-CTCGGCCGCGAGAAGGCTT-3') from genomic DNA and cloned to yield pCTB5. A 1.0 kb HindIII–AgeI fragment in pCTB5 was replaced with a 1.6 kb BamHI, end-filled fragment harbouring the HYG cassette from pUCATPH [obtained from the Fungal Genetics Stock Center (FGSC), University of Missouri, Kansas City, MO, USA] to generate the disruption construct, pctb5. Two truncated HYG and CTB5 fusion fragments overlapping within the HYG region (800 bp) were amplified by PCR from pctb5, and directly transformed into wild-type C. nicotianae for gene disruption. A 2.3 kb fragment containing 5' CTB5 fused with 3' HYG was amplified with primers CTB7I and Hygsplit2 (5'-CCGACAGTCCCGGCTCCGGATCGG-3'); a 2.1 kb fragment containing 5' HYG fused with 3' CTB5 was amplified with primers CTB5R and Hygsplit1 (5'-AGGAGGGCGTGGATATGTCCTGCGGG-3').
For CTB6 disruption, a 2 kb fragment was amplified with primers ctb6F (5'-CAAACGCAGATACCTCGCCGCATG-3') and mfs4 (5'-GCAAATTCTGAGGATTTCCCTTG-3'), and cloned to form pCTB6. A 0.5 kb BglII–HindIII fragment in pCTB6 was replaced with the HYG cassette to generate p
ctb6. Two fragments, of 1.7 and 1.9 kb, overlapping within HYG, were obtained from pctb6 with primers ctb6F and hygsplit1 and mfs4 and hygsplit2, respectively.
For CTB7 disruption, a 5 kb fragment was amplified with primers tf4 (5'-CCATGAAGCGAGATGC-3') and ord3 (5'-CGTATACCGCTACCCATGTCGTAC-3'), and cloned to become pCTB7. A 0.9 kb Eco47III fragment was replaced with the HYG cassette to yield the disruption construct, pctb7. Split-HYG marker fragments, of 2.5 and 1.5 kb, were amplified from pctb7 with primers tf4 and hygsplit1 and ord3 and hygsplit2, respectively.
Complementation and fungal transformation.
The full-length CTB5, CTB6 or CTB7 ORF, including the corresponding endogenous promoter, was independently amplified with gene-specific primers by a high-fidelity DNA polymerase (Roche Applied Science). Genetic complementation was performed by co-transformation of a PCR fragment with the pCB1532 plasmid carrying the Magnaporthe grisea acetolactate synthase gene (SUR) cassette for sulfonylurea resistance (Sweigard et al., 1997, obtained from FGSC) into ctb5-D8, ctb6-D18 or ctb7-D2 null mutants. Transformants were selected against sulfonylurea and screened for cercosporin production. Fungal protoplasts were prepared and transformed using CaCl2 and PEG by methods described previously (Chung et al., 2002). Transformants were selected on medium containing 250 µg hygromycin ml–1 (Roche), or 5 µg chlorimuron ethyl ml–1 (Chem Service) as appropriate and tested for cercosporin production on PDA plates (Chung, 2003).
Sequence analysis.
Fungal DNA was isolated with a DNeasy Plant Mini kit (Qiagen). Full-length CTB5, CTB6 or CTB7 was amplified with the respective primers as described above and cloned into pGEM-T easy vector (Promega) for sequence analysis from both directions at Eton Bioscience. PCR primers were synthesized by Integrated DNA Technologies. Searches for sequence similarity and functional domains were performed with web-based software programs as described previously (Chen et al., 2007). Sequence data from this study have been deposited with the EMBL/GenBank Data Libraries under accession nos. DQ991507 (CTB5), DQ991508 (CTB6) and DQ991509 (CTB7).
Molecular techniques.
Standard procedures were used for endonuclease digestion of DNA, electrophoresis, and Southern and Northern blot hybridization. The hybridization probes were generated by PCR with gene-specific primers to incorporate DIG-11-dUTPs (Roche) into CTB1, CTB5, CTB6, CTB7 or CTB8 DNA fragments as previously described (Choquer et al., 2005; Chen et al., 2007). The conditions and procedures for probe labelling, hybridization, post-hybridization washing and immunological detection of the probe with a disodium 3-[4-methoxyspiro{1,2-dioxetane-3,2'-(5'-chloro)tricyclo[3.3.1.1]decan}-4-yl]phenyl phosphate (CSPD) chemiluminescent substrate for alkaline phosphatase were carried out according to the manufacturer's instructions (Roche).
Pathogenicity assay.
Assay for fungal pathogenicity was carried out on detached tobacco leaves (Nicotiana tabacum ‘Burley 21’) with conidia suspensions (5x104 conidia ml–1) as described previously (Choquer et al., 2005).
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
). In this study we utilized a genetic approach to further define the roles of CTB5, CTB6 and CTB7 in cercosporin biosynthesis. Both CTB5 and CTB7 are located near the left border of the cercosporin biosynthetic gene cluster (Fig. 1b). The coding regions of CTB5 and CTB7 are separated by 867 bp and are transcribed in the same direction. The CTB5 ORF contains 1380 bp with no introns and is predicted to encode a polypeptide of 459 aa. The translation product of CTB5 displays 30–59 % identity and 50–72 % similarity to numerous uncharacterized, conserved, hypothetical proteins from sequenced genomes of fungi (data not shown). The amino acid sequence deduced from the CTB5 ORF also resembles various oxygen- and FAD/FMN-dependent oxidoreductases of various bacteria in the databases. CTB5 has a putative TonB-dependent receptor protein signature, a potential oxygen-interacting site, a FAD-binding site, a tyrosine sulfation site, and a putative NADH-binding site (Fig. 2).
|
), consists of 1074 bp with no introns, and presumably encodes a 357 aa polypeptide. The translation product of CTB6 has 26–40 % identity and 48–61 % similarity to numerous NADPH-dependent reductases, oxidoreductases or dehydrogenases of various micro-organisms. An alcohol dehydrogenase family signature and a motif that probably interacts with NADPH were identified in CTB6 (Fig. 2).
The CTB7 ORF consists of 1401 bp interrupted by a single intron of 48 bp and is predicted to encode a protein of 450 aa. The translation product of CTB7 has 22–33 % identity and 38–51 % similarity to a wide variety of FAD/FMN-dependent oxidoreductases, hydrolases, or monooxygenases of fungi and bacteria. CTB7 has two FMN/FAD or flavin-containing monooxygenase-binding sites and an amidation site (Fig. 2).
Disruption of the CTB5, CTB6 and CTB7 genes
Targeted gene disruption was performed to evaluate the functions of CTB5, CTB6 and CTB7 associated with cercosporin production in C. nicotianae. Transformants were screened for cercosporin production on a thin PDA plate. In total, 53 of 202 (26 %) transformants recovered from CTB5 disruption, 10 of 141 (7.1 %) from CTB6 disruption, and 3 of 16 (19 %) from CTB7 disruption were completely defective in cercosporin accumulation.
Successful disruption of each CTB locus was validated by Southern blot analysis. Hybridization of EcoRI/HindIII-digested genomic DNA from the wild-type and six putative ctb5 disruptants to a CTB5 gene probe identified a 3.0 kb band in wild-type (Fig. 3a, b). In contrast, all transformants had a 2.4 kb hybridizing band instead, due to the insertion of HYG within CTB5 and the presence of an additional EcoRI site in the HYG gene cassette. Three hybridizing bands, >4 kb (indicated by arrowheads in Fig. 3b), in DNA prepared from the ctb5-D1 and D9 disruptants, were due to the ectopic integrations of split marker fragments in the genome. Northern blot analysis of total RNA from two ctb5 disruptants also confirmed the null mutation at CTB5 (Fig. 3c).
|
, hybridization of NcoI/XhoI-digested genomic DNA to a CTB6 gene probe yielded a 2.0 kb hybridizing band in wild-type and a 1.8 kb band in three putative ctb6 disruptants due to the presence of an extra NcoI site in the inserted HYG fragment. The hybridization patterns confirmed a successful disruption at the CTB6 locus. Hybridizing bands (>2.0 kb and <1.5 kb) due to the ectopic integration of PCR fragments were detected in DNA prepared from the ctb6-D4 disruptant. Northern blot hybridization of total RNA from two ctb6 disruptants further validated the null mutation at CTB6 (Fig. 4c).
|
also confirmed targeted disruption of CTB7. The CTB7 probe hybridized to 2.7 and 3.4 kb BclI fragments in DNA purified from the wild-type and five putative ctb7 disruptants, respectively. Ectopic insertions were also detected in DNA from the ctb7-D1, D4 and D7 disruptants. Hybridization of total RNA from wild-type and two ctb7 disruptants to a CTB7 probe also confirmed the null mutation at CTB7 (Fig. 5c).
|
Cercosporin production
The amounts of cercosporin in KOH extracts were determined, revealing that the absorbance values obtained from the ctb5, ctb6 or ctb7 disruptants were indistinguishable from the controls (agar plugs only) and were considered to be zero (Table 1). To determine if a trace amount of cercosporin was produced by the disrupted mutants, cercosporin was extracted by ethyl acetate from agar plugs with fungal mycelia and the extracts were analysed by TLC. As shown in Fig. 6, the wild-type and the CTB5-, CTB6- and CTB7-complemented strains produced a red pigment (cercosporin), whereas the ctb5, ctb6 or ctb7 disruptants produced no detectable cercosporin. Thus, disruption of the CTB5, CTB6 or CTB7 gene completely blocked cercosporin biosynthesis, but disruptants accumulated a yellowish or purplish pigment that was barely visible on fluorescent-TLC plates (Fig. 6).
|
|
). The results indicated that expression of CTB5 was almost undetectable in the ctb6 and ctb7 null mutants (Fig. 7a). Accumulation of the CTB6 transcript was slightly reduced in two ctb5 null mutants, but completely abolished in two ctb7 null mutants (Fig. 7b). Expression of CTB7 was barely detectable in the ctb5 and ctb6 null mutants (Fig. 7c). Similarly, expression of CTB1 and CTB8 was drastically down-regulated in the ctb5 and ctb6 null mutants, and accumulation of the CTB1 but not the CTB8 gene transcript was also reduced in two ctb7 null mutants (Fig. 8).
|
|
). For comparison purpose, strain 205C3, which is deficient in a zinc finger transcriptional factor, CRG1, required for normal cercosporin production and resistance (Chung et al., 2003a), was tested. In agreement with the previous findings (Chung et al., 2003a), the 205C3 mutant exhibited partial sensitivity to cercosporin but not to other photosensitizers, and produced less than 50 % of the cercosporin produced by the wild-type (Table 1).
Pathogenicity
As assayed on detached tobacco leaves, the ctb5, ctb6 or ctb7 disruptants caused fewer lesions compared to the wild-type. However, the complementation strains incited necrotic lesions on tobacco, indistinguishable from those induced by the wild-type (data not shown).
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
). Chromosome walking coupled with sequence analysis led to identification of the cercosporin toxin biosynthesis (CTB) cluster. The core CTB gene cluster in C. nicotianae consists of a transcriptional regulator gene, a potential transporter gene, and six biosynthetic genes (Chen et al., 2007). To fully determine the function of the CTB1–8 genes associated with cercosporin production, we performed genetic and molecular analysis of C. nicotianae strains with loss- and gain-of-function mutations in each CTB gene (Chen et al., 2007; Choquer et al., 2005, 2007; Dekkers et al., 2007). In this present study we analysed three putative oxidoreductase-encoding genes (CTB5, CTB6 and CTB7) that were localized in the CTB cluster, and obtained experimental evidence to support their crucial roles in cercosporin biosynthesis.
Although the biochemical function of CTB5, CTB6 and CTB7 as oxidoreductases remains to be proven, molecular and genetic analyses clearly indicated that they were required for cercosporin biosynthesis. A prior study revealed that accumulation of CTB5, CTB6 and CTB7 transcripts was co-ordinately controlled by the CTB8 transcriptional activator (Chen et al., 2007). In the present study, disruption of CTB5, CTB6 or CTB7 yielded mutants that were completely impaired in cercosporin production, yet retained the wild-type level of resistance to cercosporin and other singlet-oxygen-generating photosensitizers. We have also demonstrated that disruption of each of the CTB5, CTB6 or CTB7 genes markedly reduces transcriptional accumulation of the other CTB gene transcripts, consistent with the previous finding of the presence of a feedback inhibition mechanism (Chen et al., 2007). However, it appears that such inhibition was not completely stringent since expression of CTB6 was slightly reduced in two ctb5 null mutants, but completely undetectable in the ctb7 null mutants (Fig. 7). Furthermore, expression of CTB8 was apparently down-regulated in the ctb5 or ctb6 null mutants, but was normal in the ctb7 mutants (Fig. 8).
CTB5 has amino acid similarity to many oxygen- and FAD/FMN-dependent oxidoreductases of bacteria and fungi, including vanillyl-alcohol oxidases, D-lactate dehydrogenases, 6-hydroxy-D-nicotine oxidases (accession nos. P56216, P06149, ZP_00522304, and CAA29416), and mitomycin radical oxidases of Streptomyces (accession no. P43485). These enzymes catalyse the oxidation of a wide variety of substrates for energy production and conversion (Rule et al., 1985; van den Heuvel et al., 2000). The mitomycin radical oxidase, however, oxidizes the reduced form of mitomycins and is involved in cellular self-defence against mitomycin in Streptomyces lavendulae (August et al., 1994). The enzyme encoded by CTB5 is proposed to utilize FAD and/or FMN as a cofactor and catalyse the oxidation steps in the cercosporin biosynthetic pathway. CTB5 has a putative TonB-dependent receptor signature. The TonB protein is involved in the passive uptake of large and low-affinity substrates by interacting with outer-membrane receptors in Escherichia coli (Bell et al., 1990). Without TonB, the receptors bind their substrates but fail to transport them into the periplasmic space. In addition, CTB5 also has a tyrosine sulfation site that is physiologically associated only with proteins or domains that are transported or reside in the Golgi lumen (Huttner, 1988). It will be interesting to determine how those domains contribute to CTB5 function.
The protein encoded by CTB6 displays considerable similarity to numerous NADPH-dependent reductases, oxidoreductases or dehydrogenases in bacteria, yeasts and plants. Some members include aldehyde reductases, nucleoside-diphosphate sugar epimerases, carbonyl reductases, dihydroflavonol 4-reductases and cinnamyl-alcohol dehydrogenases (accession nos. 1Y1PA, XP_71038, ZP_00592614, and ZP_00528210). A computer search identified an alcohol dehydrogenase (adh) family signature and a NADPH-binding motif in CTB6. A CTB6 homologue in Saccharomyces cerevisiae (accession no. NP_011476) was thought to catalyse NADPH-dependent reduction of the bicyclic diketone bicyclo[2,2,2]octane-2,6-dione to the chiral ketoalcohol-6-hydroxybicyclo[2,2,2]octane-2-one (Goffeau et al., 1996). It is tempting to speculate that the CTB6 enzyme catalyses an NADPH-dependent reduction or hydration step during ring closure of pentaketide in the cercosporin biosynthetic pathway.
The translation product of CTB7 has an amidation site and two FMN/FAD or flavin-containing monooxygenase-binding sites. Proteins having similarity to the CTB7 include flavoprotein monooxygenases and pyridine nucleotide-disulfide oxidoreductases of Pseudomonas (accession nos. YP_2333205 and YP_272409), and FAD-dependent oxidoreductases of Xanthomonas (accession nos. YP_363434 and AAM36534). CTB7 also has similarity to a Xanthomonas campestris pv. zinniae oxidoreductase (accession no. AAY86766) which has recently been shown to be involved in cercosporin degradation (Taylor et al., 2006). We propose that the function of CTB7 in the cercosporin biosynthetic pathway is probably to catalyse a hydration or reduction step during ring closure to form the polyketomethylene skeleton of cercosporin prior to the methylation steps.
Biosynthetic gene clusters often include one or more genes for cellular self-protection in some fungi. For example, TRI12 is an efflux pump that is involved in self-protection against trichothecene in Fusarium sporotrichioides (Alexander et al., 1999), and TOXA is a putative HC-toxin transporter in Cochliobolus carbonum (Pitkin et al., 1996). Cercosporin is toxic to many cells due to the production of singlet oxygen; however, Cercospora species are very resistant to cercosporin (Daub et al., 2005). The mechanisms involved in cercosporin resistance have been attributed to the ability of Cercospora species to transiently reduce cercosporin, perhaps via membrane reductases (Daub et al., 1992, 2000; Leisman & Daub, 1992; Sollod et al., 1992). Recently, expression of a yeast FAD-dependent reductase in tobacco conferred resistance to cercosporin (Panagiotis et al., 2007). Despite the fact that the CTB5, CTB6 and CTB7 genes encode putative oxidoreductases, targeted gene disruption indicated that none of them is responsible for cercosporin self-resistance. Loss of the ability to synthesize cercosporin was the only phenotypic change caused by mutation of the CTB5, CTB6 or CTB7 genes. Conidiation of ctb5, ctb6 and ctb7 disruptants was not affected (data not shown). Finally, the ctb5, ctb6 and ctb7 disruptants produced fewer lesions compared to the wild-type on tobacco leaves, consistent with previous findings that cercosporin is an important virulence factor (Chen et al., 2007; Choquer et al., 2005, 2007; Dekkers et al., 2007).
|
ACKNOWLEDGEMENTS |
|---|
Edited by: M. Tien
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
|---|
August, P. R., Flickinger, M. C. & Sherman, D. H. (1994). Cloning and analysis of a locus (mcr) involved in mitomycin C resistance in Streptomyces lavendulae. J Bacteriol 176, 4448–4454.
Bell, P. E., Nau, C. D., Brown, J. T., Konisky, J. & Kadner, R. J. (1990). Genetic suppression demonstrates interaction of TonB protein with outer membrane transport proteins in Escherichia coli. J Bacteriol 172, 3826–3829.
Callahan, T. M., Rose, M. S., Meade, M. J., Ehrenshaft, M. & Upchurch, R. G. (1999). CFP, the putative cercosporin transporter of Cercospora kikuchii, is required for wild type cercosporin production, resistance, and virulence on soybean. Mol Plant Microbe Interact 12, 901–910.[Medline]
Chen, H., Lee, M.-H., Daub, M. E. & Chung, K.-R. (2007). Molecular analysis of the cercosporin biosynthetic gene cluster in Cercospora nicotianae. Mol Microbiol 64, 755–770
Choquer, M., Dekkers, K. A., Chen, H.-Q., Ueng, P. P., Daub, M. E. & Chung, K.-R. (2005). The CTB1 gene encoding a fungal polyketide synthase is required for cercosporin biosynthesis and fungal virulence of Cercospora nicotianae. Mol Plant Microbe Interact 18, 468–476.[Medline]
Choquer, M., Lee, M. H., Bau, H. J. & Chung, K. R. (2007). Deletion of a MFS transporter-like gene in Cercospora nicotianae reduces cercosporin toxin accumulation and fungal virulence. FEBS Lett 581, 489–494.[CrossRef][Medline]
Chung, K.-R. (2003). Involvement of calcium/calmodulin signaling in cercosporin toxin biosynthesis by Cercospora nicotianae. Appl Environ Microbiol 69, 1187–1196.
Chung, K.-R., Jenns, A. E., Ehrenshaft, M. & Daub, M. E. (1999). A novel gene required for cercosporin toxin resistance in the fungus, Cercospora nicotianae. Mol Gen Genet 262, 382–389.[Medline]
Chung, K.-R., Shilts, T., Li, W. & Timmer, L. W. (2002). Engineering a genetic transformation system for Colletotrichum acutatum, the causal fungus of lime anthracnose and postbloom fruit drop. FEMS Microbiol Lett 213, 33–39.[CrossRef][Medline]
Chung, K.-R., Daub, M. E., Kuchler, K. & Schüller, C. (2003a). The CRG1 gene required for resistance to the singlet oxygen-generating cercosporin toxin in Cercospora nicotianae encodes a putative fungal transcription factor. Biochem Biophys Res Commun 302, 302–310.[CrossRef][Medline]
Chung, K.-R., Ehrenshaft, M., Wetzel, D. K. & Daub, M. E. (2003b). Cercosporin- deficient mutants by plasmid tagging in the asexual fungus Cercospora nicotianae. Mol Genet Genomics 270, 103–113.[CrossRef][Medline]
Daub, M. E. (1982). Peroxidation of tobacco membrane lipids by the photosensitizing toxin, cercosporin. Plant Physiol 69, 1361–1364.
Daub, M. E. & Briggs, S. P. (1983). Changes in tobacco cell membrane composition and structure caused by the fungal toxin, cercosporin. Plant Physiol 71, 763–766.
Daub, M. E. & Ehrenshaft, M. (2000). The photoactivated Cercospora toxin cercosporin: contributions to plant disease and fundamental biology. Annu Rev Phytopathol 38, 461–490.[CrossRef][Medline]
Daub, M. E. & Hangarter, R. P. (1983). Production of singlet oxygen and superoxide by the fungal toxin, cercosporin. Plant Physiol 73, 855–857.
Daub, M. E., Leisman, G. B., Clark, R. A. & Bowden, E. F. (1992). Reduced detoxification as a mechanism of fungal resistance to singlet-oxygen-generating photosensitizers. Proc Natl Acad Sci U S A 89, 9588–9592.
Daub, M. E., Li, M., Bilski, P. & Chignell, C. F. (2000). Dihydrocercosporin singlet oxygen production and subcellular localization: a possible defense against cercosporin phototoxicity in Cercospora. Photochem Photobiol 71, 135–140.[CrossRef][Medline]
Daub, M. E., Herrero, S. & Chung, K.-R. (2005). Photoactivated perylenequinone toxins in fungal pathogenesis of plants. FEMS Microbiol Lett 252, 197–206.[CrossRef][Medline]
Dekkers, K. L., You, B.-J., Gowda, V. S., Liao, H.-L., Lee, M.-H., Bau, H.-J., Ueng, P. P. & Chung, K.-R. (2007). The Cercospora nicotianae gene encoding dual O-methyltransferase and FAD-dependent monooxygenase domains mediates cercosporin toxin biosynthesis. Fungal Genet Biol 44, 444–454.[CrossRef][Medline]
Ehrenshaft, M. & Upchurch, R. G. (1991). Isolation of light-enhanced cDNAs of Cercospora kikuchii. Appl Environ Microbiol 57, 2671–2676.
Ehrenshaft, M., Jenns, A. E., Chung, K.-R. & Daub, M. E. (1998). SOR1, a gene required for photosensitizer and singlet oxygen resistance in Cercospora fungi, is highly conserved in divergent organisms. Mol Cell 1, 603–609.[CrossRef][Medline]
Ehrenshaft, M., Bilski, P., Li, M. Y., Chignell, C. F. & Daub, M. E. (1999). A highly conserved sequence is a novel gene involved in de novo vitamin B6 biosynthesis. Proc Natl Acad Sci U S A 96, 9374–9378.
Goffeau, A., Barrell, B. G., Bussey, H., Davis, R. W., Dujon, B., Feldmann, H., Galibert, F., Hoheisel, J. D., Jacq, C. & other authors (1996). Life with 6000 genes. Science 274, 546–547.
Huttner, W. B. (1988). Tyrosine sulfation and the secretary pathway. Annu Rev Physiol 50, 363–376.[CrossRef][Medline]
Jenns, A. E. & Daub, M. E. (1995). Characterization of mutants of Cercospora nicotianae sensitive to the toxin cercosporin. Phytopathology 85, 906–912.[CrossRef]
Jenns, A. E., Daub, M. E. & Upchurch, R. G. (1989). Regulation of cercosporin accumulation in culture by medium and temperature manipulation. Phytopathology 79, 213–219.[CrossRef]
Keller, N. P., Turner, G. & Bennett, J. W. (2005). Fungal secondary metabolism – from biochemistry to genomics. Nat Rev Microbiol 3, 937–947.[CrossRef][Medline]
Kuyama, S. & Tamura, T. (1957). Cercosporin. A pigment of Cercospora kikuchii Matsumoto et Tomoyasu. II. Physical and chemical properties of cercosporin and its derivatives. J Am Chem Soc 79, 5725–5729.[CrossRef]
Leisman, G. B. & Daub, M. E. (1992). Singlet oxygen yields, optical properties, and phototoxicity of reduced derivatives of the photosensitizer cercosporin. Photochem Photobiol 55, 373–379.[CrossRef]
Okubo, A., Yamazaki, S. & Fuwa, K. (1975). Biosynthesis of cercosporin. Agric Biol Chem 39, 1173–1175.
Panagiotis, M., Kritonas, K., Irini, N. O., Kiriaki, C., Nicolaos, P. & Athanasios, T. (2007). Expression of the yeast cpd1 gene in tobacco confers resistance to the fungal toxin cercosporin. Biomol Eng 24, 245–251.[CrossRef][Medline]
Pitkin, J. W., Panaccione, D. G. & Walton, J. D. (1996). A putative cyclic peptide efflux pump encoded by the TOXA gene of the plant-pathogenic fungus Cochliobolus carbonum. Microbiology 142, 1557–1565.
Rule, G. S., Pratt, E. A., Chin, C. C., Wold, F. & Ho, C. (1985). Overproduction and nucleotide sequence of the respiratory D-lactate dehydrogenase of Escherichia coli. J Bacteriol 161, 1059–1068.
Schafer, W. (1994). Molecular mechanisms of fungal pathogenicity to plants. Annu Rev Phytopathol 32, 461–477.
Shim, W.-B. & Dunkle, L. D. (2003). CZK3, a MAP kinase kinase kinase homolog in Cercospora zeae-maydis, regulates cercosporin biosynthesis, fungal development, and pathogenesis. Mol Plant Microbe Interact 16, 760–768.[Medline]
Sollod, C. C., Jenns, A. J. & Daub, M. E. (1992). Cell surface redox potential as a mechanism of defense against photosensitizers in fungi. Appl Environ Microbiol 58, 444–449.
Sweigard, J. A., Chumley, F. C., Carroll, A. M., Farrall, L. & Valent, B. (1997). A series of vectors for fungal transformation. Fungal Genet Newsl 44, 52–53.
Taylor, T. V., Mitchell, T. K. & Daub, M. E. (2006). An oxidoreductase is involved in cercosporin degradation by the bacterium Xanthomonas campestris pv. zinniae. Appl Environ Microbiol 72, 6070–6078.
Upchurch, R. G., Walker, D. C., Rollins, J. A., Ehrenshaft, M. & Daub, M. E. (1991). Mutants of Cercospora kikuchii altered in cercosporin synthesis and pathogenicity. Appl Environ Microbiol 57, 2940–2945.
van den Heuvel, R. H. H., Fraaije, M. W., Mattevi, A. & van Berkel, W. J. H. (2000). Asp-170 is crucial for the redox properties of vanillyl-alcohol oxidase. J Biol Chem 275, 14799–14808.
Walton, J. D. (1996). Host-selective toxins: agents of compatibility. Plant Cell 8, 1723–1733.[CrossRef][Medline]
Yamazaki, S. & Ogawa, T. (1972). The chemistry and stereochemistry of cercosporin. Agric Biol Chem 36, 1707–1718.
Yamazaki, S., Okube, A., Akiyama, Y. & Fuwa, K. (1975). Cercosporin, a novel photodynamic pigment isolated from Cercospora kikuchii. Agric Biol Chem 39, 287–288.
Received 22 February 2007;
revised 3 April 2007;
accepted 17 April 2007.
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| INT J SYST EVOL MICROBIOL | MICROBIOLOGY | J GEN VIROL |
| J MED MICROBIOL | ALL SGM JOURNALS | |
