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Australasian Plant Pathology, 2004, 33, 173–181

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Genetic relationships among isolates of Phoma ligulicola from pyrethrum and chrysanthemum based on ITS sequences and its detection by PCR S. J. PethybridgeA,C, J. B. ScottB and F. S. HayA A

Tasmanian Institute of Agricultural Research, University of Tasmania — North West Centre, PO Box 3523, Burnie, Tas. 7320, Australia. B CSIRO Plant Industry, 306 Carmody Road, St Lucia, Qld 4072, Australia. C Corresponding author; email: [email protected]

Abstract. Variation within the internal transcribed spacer (ITS1, 5.8S gene and ITS2) region of the rDNA (ITS) was used to characterise the phylogenetic relationships among Phoma ligulicola isolates infecting pyrethrum crops in Tasmania, P. ligulicola isolates from the USA, Germany and mainland Australia, and other closely related fungal species. This study reports the first characterisation of the ITS region of P. ligulicola. Sequence homology within P. ligulicola isolates varied between 99.3 and 100%. For 9 of the 11 isolates from Tasmania, Australia, the nucleotide sequences in this region were identical, whereas the sequences for the remaining two isolates differed only by two nucleotides in the ITS1 region. Isolates from Australia and the USA failed to metabolise NaOH on malt-extract agar and were characterised as P. ligulicola var. inoxydablis. The two isolates from ray blight disease of chrysanthemum in Germany (DSMZ 63133 and DSMZ 62547) were classified as P. ligulicola var. ligulicola. Phylogenetic analyses suggested that the ITS sequences of P. ligulicola isolates were more similar to other Phoma species than selected representatives of the Mycosphaerella genus. Didymella bryoniae had the greatest interspecific homology with P. ligulicola of the fungi used in this study. This information was used to design specific primers within the ITS regions for the detection of P. ligulicola. AP03097 eSITt.alJ.DPrNtehAysbreidqugencedviesrtiyin Phoma ligulicoal

Introduction Pyrethrum (Tanacetum cineariifolium) is a perennial, herbaceous flowering plant, grown for insecticidal pyrethrins produced in glands on the achenes. Pyrethrins are composed of a mixture of six different esters that are active against a broad range of insects. They are incorporated into a wide range of products, such as fly sprays, fumigants, mosquito coils and pet flea collars (Casida and Quistad 1995). Tasmania is the only state in Australia in which pyrethrum is produced, and the second biggest producer of pyrethrins in the world after Kenya. In the 2001/02 season, 2150 ha of pyrethrum were harvested in Tasmania, of which 95% was exported, mainly to the USA. The most serious fungal foliar disease of pyrethrum in Tasmania is ray blight, caused by Phoma ligulicola, the anamorph of Didymella ligulicola. The fungus was first reported in commercial crops in Tasmania in 1995 (Pethybridge and Wilson 1998). P. ligulicola has also been reported to cause disease in commercial pyrethrum crops in Kenya (Robinson 1963), Tanzania (Peregrine and Watson 1964) and Papua New Guinea (Shaw 1984). Severe losses have been attributed to this disease, up to 100% on some © Australasian Plant Pathology Society 2004

occasions (Pethybridge and Hay 2001). Although the most striking symptom associated with this disease is necrosis of the flowers (or ray florets), symptoms can also to be found on the leaves and stems. On the flower, necrotic tissues extend into the peduncle of the flower or developing bud and up to 30 mm along the flowering stem, causing the flower or bud to droop (Pethybridge and Wilson 1998). On the stem, the fungus causes necrotic lesions which under conducive conditions can cover a large proportion of the tissue. On the leaves, symptoms begin as necrotic spots that coalesce and irregularly encompass the entire leaf area. Similar symptoms have also been described from glasshouse- and field-grown chrysanthemum crops in the USA (Baker et al. 1949; Stevens 1907), Japan (Walker and Baker 1983), the United Kingdom (Walker and Baker 1983), Italy (Garibaldi and Gullino 1971) and New South Wales, Australia (Anonymous 1956). Although conidia and ascospores are the primary means of pathogen dispersal (Baker et al. 1949; Blakeman and Fraser 1969; McCoy et al. 1972), the fungus can also survive as epiphytic mycelium (Chesters and Blakeman 1966) and pseudosclerotia within the soil (Blakeman and Hornby 1966). To date, only the lifecycle of the anamorph 10.1071/AP03097

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has been described in Tasmanian pyrethrum crops (Pethybridge and Wilson 1998; Pethybridge and Hay 2001; Pethybridge et al. 2003). The taxonomy of the phytopathogenic Phoma species has been well studied by Boerema and co-workers based upon morphology, cultural characteristics on at least three culture media, in vitro macrochemical reactions, and references to specific ecological and phytopathological features (Dorenbosch 1970; Boerema and Bollen 1975; Boerema and Loerakker 1985; Van der Aa et al. 1990). The identification of P. ligulicola by traditional morphological examination methods is time consuming. Also, identification is often complicated by the failure of some isolates to reliably produce fruiting structures in vitro, and frequently observed variation within both conidial size and septation between individual pycnidia in the same culture. Two variants of P. ligulicola have been described based upon their reaction to the addition of NaOH to cultures on malt-extract agar (Van der Aa et al. 1990). Isolates which are able to utilise NaOH indicate the presence of the antibiotic metabolite ‘E’, similar to that produced by Phoma exigua var. exigua, and are termed P. ligulicola var. ligulicola. Isolates that are not able to utilise NaOH but are indistinguishable morphologically from the former, are termed P. ligulicola var. inoxydablis. Isolates from the USA and Japan have been characterised as P. ligulicola var. ligulicola, whereas P. ligulicola var. inoxydablis has been reported from Europe and Australia (Van der Aa et al. 1990). In this study we report the phylogenetic relationships between P. ligulicola isolates from Tasmania, Europe and the USA using sequence information from the internal transcribed region (ITS1, 5.8S gene and ITS2) (ITS) of the rDNA. The development of PCR-based species-specific primers for use in the polymerase chain reaction (PCR) for the specific detection of P. ligulicola is described. Methods Fungal isolates Isolates of P. ligulicola were collected for DNA sequencing and added to other fungal sequences for phylogenetic analysis (Table 1). Isolates of other fungal species not used in phylogenetic analysis were also collected for use in the development of a PCR detection technique (Table 2). Fungal isolates were maintained on potato-dextrose agar (PDA) (Difco Laboratories, Detroit, MI). Herbarium specimens of P. ligulicola were sourced from Dr Michael Priest, New South Wales Agriculture, Australia (DAR 70020), the American Type Culture Collection, Manassas, VA, USA (ATCC 10748), and the Deutsche Sammlung von Mikroorganismen und Zellkulturen (DSMZ), Braunschweig, Germany (DSMZ 63133 and DSMZ 62547). Herbarium specimens from the USA and Germany were freeze-dried and re-hydrated in 5 mL sterile distilled water prior to transfer to PDA. For long-term preservation, small blocks (10 mm2) of PDA were placed in sterile distilled water and stored at 4°C. Tasmanian cultures were isolated from diseased plant tissue. Plant tissue was surface sterilised in 1% sodium hypochlorite solution for 2 min, followed by three rinses in sterile distilled water. Small (< 2 mm) pieces of tissue at the junction between the healthy and necrotic tissue were excised under aseptic

S. J. Pethybridge et al.

conditions and placed onto water agar in Petri plates and incubated at 20°C in the dark. After 5 days, mycelium growing from the tissue was transferred to PDA, malt-extract agar (MEA) (Acumedia Manufacturers Inc., Baltimore, Maryland) and oatmeal agar (OA) (Difco Laboratories) for identification (Punithalingam 1980). Typing of P. ligulicola to variety Reaction to 5 M NaOH on the outer boundary of the colony was used to differentiate the P. ligulicola varieties, inoxydablis and ligulicola (Van der Aa et al. 1990) using three replicate plates of 14-day-old cultures growing on MEA. NaOH (2 mL) was added drop-wise at the edge of the cultures and colour development was assessed within 30 min. Colour was assessed by the presence or absence of a red pigment in the agar. Cultures in which no colour development was observed by the naked eye were identified as P. ligulicola var. inoxydablis, whereas cultures that developed a red pigment were designated var. ligulicola. Deoxyribonucleic acid isolation from fungal mycelium Mycelial plugs were added to 20 mL of 2% malt-extract broth in Petri plates, agitated using an orbital shaker set at 42 rpm and incubated at 25°C under constant illumination by fluorescent lights. Mycelium (0.05–1 g) was removed after 10 days, transferred to an eppendorf tube for DNA extraction using a modification of the CTAB method described by Moller et al. (1992). Mycelial samples were suspended in hot 2% CTAB buffer (Moller et al. 1992) and 800 PL phenol:chloroform:isoamyl alcohol (25:24:1) and homogenised using 500 mg of glass beads (0.5 mm diameter) in a Mini-Beadbeater (Biospec Products) for 3 min at 30 s intervals. Following homogenisation, samples were centrifuged at 13 000 g for 5 min and the aqueous layer transferred to a fresh tube. Nucleic acids were extracted from the aqueous layer with an equal volume of chloroform:isoamyl alcohol (24:1). Deoxyribonucleic acid was precipitated using cold 95% ethanol and washed with 70% ethanol. Deoxyribonucleic acid was dissolved in 50 μL of TE (10 mM Tris-HCl, 1 mM EDTA, pH 8.0) buffer and stored at –20°C until use. Amplification and sequencing of ITS regions The ITS region was amplified using the previously described universal primers, ITS1 and ITS4 (Table 3), targeted to conserved regions in the 18S and 28S rRNA genes (White et al. 1990). Amplifications were performed in a total reaction volume of 50 μL. The PCR reaction mixture contained 1 mmol dNTPs, 0.5 μmol primers, 1 unit Taq DNA polymerase (Applied Biosystems Inc., Foster City, CA), 1 u PCR Buffer (Applied Biosystems Inc.) and 1.5 mmol MgCl2. One PL containing 10–50 ng of genomic DNA was used as the template for each reaction. PCR reactions were conducted with a GeneAmp PCR System 2400 (Perkin Elmer, Norwalk, CT) thermo-cycler using the following conditions: 5 min at 94°C for initial denaturation followed by 35 cycles consisting of 30 s of denaturation at 94°C, 1 min of annealing at 50°C, and 2 min extension at 72°C. Reactions concluded with a final extension at 72°C for 8 min. The PCR product was separated by electrophoresis using a 2% agarose (ICN Biochemicals, Ohio) gel and 1 u TAE (40 mM Tris-acetate, 1 mM EDTA, pH 8.0) buffer stained with ethidium bromide, and visualised under UV light. Excess primers and unincorporated nucleotides were removed upon completion of the PCR using the QIAQuick PCR Purification Kit (QIAGEN, Hilden, Germany) according to the manufacturer’s instructions. Direct sequencing was conducted in both directions using an ABI Prism BigDye Terminator Cycle Sequencing Ready Reaction Kit and an ABI 377 automatic sequencer (PE Applied Biosystems). Deoxyribonucleic acid sequence information has been deposited in GenBank.

Australasian Plant Pathology

ITS rDNA sequence diversity in Phoma ligulicola

Table 1.

175

Origin of Phoma ligulicola isolates or fungal sequences used for phylogenetic analyses

IsolateA

LocationB

Host

GenBank accession number

Phoma ligulicola Tas 2 Tas 3 Tas 4 Tas 6 Tas 7 Tas 14 Tas 15 Tas 17 Tas 19 Tas 20 Tas 22 ATCC 10748 DAR 70020 DSMZ 62547 DSMZ 63133

Tanacetum cineariifolium T. cineariifolium T. cineariifolium T. cineariifolium T. cineariifolium T. cineariifolium T. cineariifolium T. cineariifolium T. cineariifolium T. cineariifolium T. cineariifolium Chrysanthemum morifolium T. cineariifolium C. morifolium C. indicum

Scottsdale, Tas. Barrington, Tas. Burnie, Tas. North Motton, Tas. Sisters Creek, Tas. Ulverstone, Tas. Moriarty, Tas. Howth, Tas. Wesley Vale, Tas. Don, Tas. Gawler, Tas. North Carolina, USA Forth, Tas. Germany Germany

Didymella bryoniae Phoma destructiva P. glomerata P. wasabiae P. herbarum P. pinodella P. epicoccina Epicoccum nigrum P. exigua P. medicaginis P. tracheiphilia Mycosphaerella fragariae M. macrospora M. citri M.graminicola M.arachidis M.musicola M. brassicola Stemphylium botryosum

Sequences sourced from GenBank Cucumis melo USA Unknown Unknown Unknown Unknown Eutrema wasabiae Unknown Unknown Unknown Unknown Unknown Unknown Unknown Unknown Unknown Medicago truncatula Unknown Amaranthus powelli Unknown Unknown Unknown Fragariae × ananassa Unknown Unknown Unknown Citrus sinensis Unknown Hordeum vulgare Unknown Arachis hypogaea Unknown Musa sp. Unknown Brassica oleracea Unknown Unknown Unknown

AY157875 AY157877 AY157878 AY157879 AY157882 AY157883 AY157876 AY157884 AY157880 AY157881 AY157885 AY157886 AY157887 AY157889 AY157888 AF297228 AF268191 AF126819 L38711 AF218792 AY131199 AF149933 AF149928 AY131200 AF079775 AF272553 AF297235 AF297231 AF181703 AF181694 AF297224 AF181706 AF297236 Y17068

A

ATCC = American Type Culture Collection; DAR = NSW Agriculture Herbarium, Orange, Australia; DSMZ = Deutsche Sammlung von Mikroorganismen und Zellkulturen, Braunschweig, Germany. B Tas. = Tasmania, Australia.

Phylogenetic analysis

Design and specificity of primers and optimisation of PCR conditions

Additional DNA sequences for related species were obtained from GenBank (Table 1) and included in the phylogenetic analyses. Sequences were aligned using Clustal W (Thompson et al. 1994) and edited visually. Phylogenetic analyses were conducted using programs from the PHYLIP software package (Felsenstein 1995). Distance analysis was conducted using the neighbour-joining method in the program NEIGHBOUR, based on Kimura 2-parameter distances from the DNADIST program. Maximum parsimony analysis was conducted using the program DNAPARS. The topology of the constructed trees was supported by constructing 1000 bootstrap replicates using the program SEQBOOT, analysing as above, and finding the majority and strict concensus trees with the program, CONSENSE. The outgroup, Stemphylium botryosum (GenBank Acc. No. Y17068) was used as a root for all trees.

Unique sequences within the ITS regions of P. ligulicola were identified visually. P. ligulicola specific primers (PL1/PL2) (Table 3) were developed that amplified a 230 bp PCR product. Optimisation of the PCR conditions for reliable and specific detection was done by varying the concentration of MgCl2 between 1 and 5 mM, using the GeneAmp PCR System 2400 (Perkin Elmer) thermo-cycler with the following program: 5 min at 94°C for initial denaturation followed by 30 cycles consisting of 30 s of denaturation at 94°C, 1 min of annealing at 56°C and 1 min of extension at 72°C. A final extension time of 10 min at 72°C followed the end of the last cycle. Products were separated in 2% agarose (ICN Biochemicals) gels in 1 u TAE, stained with ethidium bromide and visualised under UV light. Primers PL1/PL2 were tested against all P. ligulicola isolates used for their development (Table 1) and 30 additional isolates collected from

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Table 2.

S. J. Pethybridge et al.

Fungal isolates (other than Phoma ligulicola) used in this study for testing of the specificity of primers PL1/PL2 for detection of P. ligulicola by the polymerase chain reaction

Isolate

Host

Phoma destructiva Phoma glomerata Didymella bryoniae Sclerotinia minor Sclerotinia sclerotiorum Alternaria tenuissima Alternaria alternata Stemphylium botryosum Pithomyces sp. Cladosporium cladosporioides Phoma lingam Ulocladium atrum

Location

Lycopersicon esculentum Juglans regia Cucumis sativus T. cineariifolium T. cineariifolium T. cineariifolium T. cineariifolium T. cineariifolium T. cineariifolium T. cineariifolium Solanum tuberosum T. cineariifolium

NSW Agriculture Herbarium DAR accession number

Taree, NSW Northfield, SA Gosford, NSW Forth, Tas. Forth, Tas. Penguin, Tas. Burnie, Tas. Table Cape, Tas. Moriarty, Tas. Don, Tas. Richmond, Tas. Table Cape, Tas.

25413 37805 33707

75573 75572

Table 3. Primers used for amplification of the complete ITS1 and ITS2 regions of Phoma ligulicola, and those designed based on unique sequences in this region for the specific detection of P. ligulicola Primer name ITS1 ITS4 PL1 PL2

Sequence 5c-TCCGTAGGTGAACCTGCGG-3c 5c-TCCTCCGCTTATTGATATGC-3c 5c-CAACACTTAAACCCTTTGTAATTGA-3c 5c-GGACGTCGTCGTTTTGTTGT-3c

Table 4. Pairwise comparisons of ITS similarity between Phoma ligulicola isolates from Tasmania (DAR 70020; Tas 3 and 22), Germany (DSMZ 63133 and 62547) and the USA (ATCC 10748) Isolate

DAR 70020A Tas 3 Tas 22 ATCC 10748 DSMZ 63133 DSMZ 62547

Amplification specificity

DAR 70020

Tas 3

Isolate Tas 22 ATCC DSMZ DSMZ 10748 63133 62547

1B — — — — —

0.997 1 — — — —

0.993 0.995 1 — — —

0.995 0.997 0.993 1 — —

0.995 0.997 0.993 1 1 —

0.995 0.997 0.993 1 1 1

A DAR 70020 is the P. ligulicola culture deposited in the New South Wales Agriculture Herbarium, Orange, Australia. B A perfect match is indicated by 1. Decreasing values indicate less similarity. Tas 3 represents the majority of isolates in Tasmania. Tas 22 represents the second group of isolates.

diseased tissue from 30 pyrethrum crops across northern Tasmania. In addition, the primers were also tested against other Phoma spp., other fungi commonly isolated from diseased pyrethrum tissue (Table 2), and plant DNA extracted from leaves and seeds using the CTAB method (Moller et al. 1992).

Results Typing of P. ligulicola isolates to variety P. ligulicola cultures were identified based on morphological and cultural characteristics according to

ITS1, universal ITS2, universal P. ligulicola P. ligulicola

Punithalingam (1980). All P. ligulicola isolates from Tasmania and the USA failed to metabolise NaOH and were hence classified as P. ligulicola var. inoxydablis. Both cultures from Germany produced a red pigment in the agar on application of NaOH and were classified as P. ligulicola var. ligulicola. Phylogenetic analysis Primers ITS1 and ITS4 were used to amplify the ITS as a single fragment from all isolates (Table 3). The intraspecific variability within P. ligulicola isolates was low. Two of the isolates from pyrethrum in Tasmania (17 and 22) differed from the majority by only two nucleotides within the ITS1 region and were identical to each other. The two groups of Tasmanian P. ligulicola isolates showed 99.7 and 99.3% homology with the DNA sequence of a representative culture of P. ligulicola from Tasmania, DAR 70020, respectively. The homology between DAR 70020 and USA and German isolates was 99.5% in all cases (Table 4). Of all the other sequences used in phylogenetic analyses, P. ligulicola displayed the greatest homology with D. bryoniae (97.1–97.5%) (Table 5). Results from both parsimony and distance methods for phylogenetic analyses of interspecific ITS variability were similar. Maximum-parsimony analysis generated 33 equally most parsimonious phylogenetic trees. The most

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Table 5. Pairwise comparison of ITS sequences of Phoma ligulicola isolates from Tasmania (DAR 70020; Tas 3 and 22), Germany (DSMZ 63133 and 62547) and the USA (ATCC 10748) and other fungal sequences Isolate

Didymella bryoniae Phoma wasabiae P. glomerata P. destructiva P. herbarum P. pinodella P. epicoccina E. nigrum P. exigua P. medicaginis P. tracheiphilia Mycosphaerella fragariae M. macrospora M. citri M. graminicola M. arachidis M. musicola M. brassicola

Tas 3A

Tas 22

0.975C 0.710 0.945 0.954 0.964 0.938 0.93 0.932 0.954 0.934 0.724 0.548 0.539 0.537 0.551 0.55 0.539 0.538

0.971 0.712 0.94 0.949 0.96 0.934 0.925 0.928 0.949 0.93 0.724 0.548 0.539 0.537 0.551 0.552 0.541 0.538

Phoma ligulicola isolates DAR 70020B ATCC 10748 0.973 0.708 0.943 0.951 0.962 0.936 0.928 0.930 0.951 0.932 0.722 0.548 0.539 0.537 0.549 0.55 0.539 0.538

0.973 0.71 0.943 0.951 0.967 0.941 0.928 0.93 0.951 0.936 0.722 0.548 0.539 0.537 0.551 0.55 0.539 0.538

DSMZ 63133

DSMZ 62547

0.973 0.71 0.943 0.951 0.967 0.941 0.928 0.93 0.951 0.936 0.722 0.548 0.539 0.537 0.551 0.55 0.539 0.538

0.973 0.710 0.943 0.951 0.967 0.941 0.928 0.930 0.951 0.936 0.722 0.548 0.539 0.537 0.551 0.550 0.539 0.538

A Tas 3 represents the majority of isolates in Tasmania. Tas 22 represents the second group of isolates (containing itself and Tas 17). B DAR 70020 is the P. ligulicola culture deposited in the New South Wales Agriculture Herbarium, Orange, Australia C A perfect match is indicated by 1. Decreasing values indicate less similarity.

parsimonious tree (Fig. 1) was determined by applying the 50% majority rule (strict consensus). When selecting S. botryosum as the outgroup, parsimony analysis split isolates into two clades, with the exception of P. wasabiae and P. tracheiphilia. One of these clades consisted of all Mycosphaerella spp., whereas the other included Didymella bryoniae and Phoma spp. (including P. ligulicola). All P. ligulicola isolates were grouped together, with one branch consisting of isolates from Tasmania. Representatives of the two different varieties of P. ligulicola (var. inoxydablis and var. ligulicola) were not strongly differentiated upon the basis of ITS phylogenetics (Fig. 1). Distance analysis, also using S. botryosum as the outgroup, split isolates in a manner similar to parsimony analysis. Similarly, Mycosphaerella spp. formed one clade, whereas P. ligulicola isolates fell into another clade together with the other Phoma spp. and Didymella bryoniae, with the exception of P. wasabiae and P. tracheiphilia (Fig. 1). Detection of P. ligulicola by PCR The specific primers PL1 and PL2 detected all isolates of P. ligulicola included in this study (Fig. 2). The primers failed to amplify the DNA of the related species Didymella bryoniae, P. lingam, P. glomerata and P. destructiva (Fig. 3). The primers also failed to amplify the DNA from other fungi

commonly associated with disease symptoms in pyrethrum, Sclerotinia minor, S. sclerotiorum (Fig. 3), Ulocladium atrum, Stemphylium botryosum, Alternaria tenuissima, A. alternata, Cladosporium cladosporioides (Fig. 4), and DNA extracted from leaf and seed pyrethrum tissue (results not shown). Varying the concentration of MgCl2 between 1 and 5 mM had little effect on the strength of the reaction (results not shown). To economise on the use of reagent, a MgCl2 concentration of 1.5 mM is recommended. Discussion To our knowledge, this is the first phylogenetic study of P. ligulicola using ITS sequence data. ITS sequences of 15 P. ligulicola isolates were used to assess intraspecific and interspecific phylogenetic relationships and to design primers for the reliable detection of P. ligulicola by PCR. Phylogenetic analyses demonstrated low intraspecific variability within the P. ligulicola isolates used in this study. The majority of isolates from pyrethrum crops in Tasmania were identical, whereas two differed by only two nucleotides within ITS1. The low variability within this region among isolates from Australia, Germany and the USA is notable considering the multiple selection pressures that each would have been exposed to, their different hosts, and geographic separation. It is unknown whether isolates found infecting pyrethrum are able to infect other Asteraceae species

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S. botryosum 100

P. wasabiae P. tracheiphilia M. macrospora 100

M. citri 61

94

M. fragariae M. graminicola

65

M. arachidis 79

100

79

M. musicola M. brassicola

P. herbarum 71 100 86

P. ligulicola DSMZ 62547 P. ligulicola ATCC 10748 P. ligulicola DSMZ 63133 P. ligulicola DAR70020 P. ligulicola Tas 3 P. ligulicola Tas 22

99

Didymella bryoniae 52

99

P. pinodella P. medicaginis

52

P. exigua

58

P. destructiva

66

P. glomerata P. epicoccina

100

E. nigrum

0.1 Fig. 1. Phylogenetic tree generated by Kimura-2 parameter and neighbour joining distance analysis from the ITS sequence data for Phoma ligulicola and other closely related fungi (Mycosphaerella spp. and Phoma/Didymella spp.). Tas 3 represents the majority of isolates in Tasmania. Tas 22 represents the second group of isolates, containing itself and Tas 17. Tree rooted with Stemphylium botryosum as the outgroup. Scale bar represents 0.1 expected nucleotide substitutions per site. Numbers at branches represent bootstrap percentages (only values > 50% shown).

recorded as hosts of P. ligulicola, including Cichorium endiva, Rudbeckia hirta, Helianthus annuus, Zinnia elegans, Dahlia variabilis and Lactuca sativa (Chesters and Blakeman 1967). Other studies of similar species, such as P. exigua, have also not been able to distinguish different varieties on sequence analysis of the ITS, despite clear differences in cultural characteristics (Abeln et al. 2002). In this study, phylogenetic analysis was unable to clearly

differentiate P. ligulicola varieties, determined by their ability to metabolise sodium hydroxide (Van der Aa et al. 1990). This observation agrees with other studies which have defined the ITS region as useful to examine the relationship between species, but not necessarily within species (Crawford et al. 1996; Cooke et al. 2000; Abeln et al. 2002; Nielsen et al. 2002). Differentiating varieties and biotypes of P. ligulicola is the subject of further study.

Australasian Plant Pathology

ITS rDNA sequence diversity in Phoma ligulicola

M

1

2

3

4

5

6

7

8

9

10

11

Fig. 2. Detection of Phoma ligulicola isolates from USA, Germany and Australia by PCR using primer pair PL1/PL2. Lanes: M, 100-bp ladder, where uppermost band is 900 bp; 1, ATCC 10748; 2, DSMZ 62547; 3, DSMZ 63133; 4, DAR70020; 5, Tas 2; 6, Tas 15; 7, Tas 3; 8, Tas 4; 9, Tas 6; 10, Tas 19; 11, Tas 20.

M

1

2

M M

1 1

3

4

2 2

5

3 3

6

11

4 4

5 5

6 6

Fig. 3. Amplification of Phoma ligulicola with the PCR primer pair PL1/PL2. Lanes: M, 100-bp ladder, where uppermost band is 600 bp; 1, Phoma glomerata; 2, P. destructiva; 3, Sclerotinia minor; 4, Sclerotinia sclerotiorum; 5, Didymella bryoniae; 6, P. ligulicola Tas 17.

M

1

2

3

4

5

6

7

8

9

Fig. 4. Reaction of other fungal species found in Tasmanian pyrethrum crops to the primer pair PL1/PL2 using the polymerase chain reaction. Lanes: M, 100-bp ladder, where uppermost band is 400 bp; 1, Phoma ligulicola Tas 14; 2, P. ligulicola Tas 22; 3, P. ligulicola Tas 7; 4, Ulocladium atrum; 5, Alternaria alternata; 6, Alternaria tenuissima; 7, Stemphylium botryosum; 8, Pithomyces sp.; 9, Cladosporium cladosporioides.

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P. ligulicola isolates were consistently grouped closer to other Phoma species and D. bryoniae than the Mycosphaerella species included in this study. This fungus has undergone several changes in nomenclature which were reviewed by Walker and Baker (1983), including Ascochyta chrysanthemi (Stevens 1907), Mycosphaerella ligulicola (Baker et al. 1949; McCoy and Dimock 1972; McCoy et al. 1972) and P. ligulicola (teleomorph Didymella ligulicola). Walker and Baker (1983) determined that the teleomorph is a Didymella species, based on morphological examination of the asci and ascocarp centrum. In Mycosphaerella perithecia, the asci are fasciculate with no pseudoparaphyses, and ascospores are usually not constricted at the central septum. In Didymella, the non-fasciculate asci arise between pseudoparaphyses and contain ascospores that are constricted at a septum slightly above the middle of the spore (Walker and Baker 1983). Our phylogenetic studies support this classification, with P. ligulicola isolates always falling within the Didymella/Phoma clade. The teleomorph stage of this fungus, Didymella ligulicola, has not been found in Tasmanian pyrethrum crops. The separation of P. wasabiae and P. tracheiphilea into a separate clade from the main Didymella/Phoma clade has previously been shown by Reddy et al. (1998). The rapid evolution rate of interspecific variation within the ITS regions make them ideal as a target for primer design for detection of fungi by PCR (White et al. 1990). The primers, PL1 and PL2, were successfully used to detect P. ligulicola whilst failing to amplify DNA from pyrethrum tissue, related species (D. bryoniae and Phoma spp.), or other fungi commonly found in association with foliar diseases of pyrethrum. These primers will be useful for the correct identification of P. ligulicola and discrimination from other fungi when traditional techniques are unable to be implemented or are impractical. Further work will assess their efficacy in the detection of P. ligulicola from extracts of infected plants, on spore traps for epidemiological studies, and to develop a reliable test for P. ligulicola in pyrethrum seed. Identification of P. ligulicola in this way will improve our understanding of the epidemiology of ray blight in Tasmanian pyrethrum crops and the development of efficient management strategies. Acknowledgements We thank Ms Kate Harrison for excellent technical assistance and Mr Tim Groom of Botanical Resources Australia Pty Ltd for constructive discussion, and Dr Kathy Evans and Dr Calum Wilson, Tasmanian Institute of Agricultural Research for comments on the manuscript. This work was funded through Horticulture Australia Limited, the Australian Research Council-Linkage program and Botanical Resources Australia Pty Ltd.

S. J. Pethybridge et al.

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Received 16 June 2003, accepted 5 September 2003

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