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Keywords:

  • Colletotrichum coccodes;
  • diagnostics;
  • internal transcribed spacer regions;
  • potato black dot;
  • quantitative (TaqMan) PCR;
  • soil

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Colletotrichum coccodes is the causal agent of the potato blemish disease black dot. Two PCR primer sets were designed to sequences of the ribosomal internal transcribed spacer (ITS1 and ITS2) regions for use in a nested PCR. The genus-specific outer primers (Cc1F1/Cc2R1) were designed to regions common to Colletotrichum spp., and the species-specific nested primers (Cc1NF1/Cc2NR1) were designed to sequences unique to C. coccodes. The primer sets amplified single products of 447 bp (Cc1F1/Cc2R1) and 349 bp (Cc1NF1/Cc2NR1) with DNA extracted from 33 European and North American isolates of C. coccodes. The specificity of primers Cc1NF1/Cc2NR1 was confirmed by the absence of amplified product with DNA of other species representing the six phylogenetic groups of the genus Colletotrichum and 46 other eukaryotic and prokaryotic plant pathogenic species. A rapid procedure for the direct extraction of DNA from soil and potato tubers was used to verify the PCR assay for detecting C. coccodes in environmental samples. The limit of sensitivity of PCR for the specific detection of C. coccodes when inoculum was added to soils was 3·0 spores per g, or the equivalent of 0·06 microsclerotia per g soil, the lowest level of inoculum tested. Colletotrichum coccodes was also detected by PCR in naturally infested soil and from both potato peel and peel extract from infected and apparently healthy tubers. Specific primers and a TaqMan fluorogenic probe were designed to perform quantitative real-time (TaqMan) PCR to obtain the same levels of sensitivity for detection of C. coccodes in soil and tubers during a first-round PCR as with conventional nested PCR and gel electrophoresis. This rapid and quantitative PCR diagnostic assay allows an accurate estimation of tuber and soil contamination by C. coccodes.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

The genus Colletotrichum comprises a diverse range of important plant pathogenic fungi that cause pre- and postharvest crop losses worldwide in cereals, grasses, fruits, legumes, vegetables and perennial crops (Sutton, 1992; Waller, 1992). One of these species, C. coccodes, causes black dot of potato (Solanum tuberosum) and is also pathogenic on several other plants in the Solanaceae and Cucurbitaceae (Dillard, 1992). Black dot is a component of the tuber-blemish disease complex of seed and ware potatoes, and has a wide geographical distribution (Read et al., 1995; Andrivon et al., 1997; Johnson et al., 1997; Denner et al., 1998; Tsror et al., 1999). At present, most widely grown cultivars of potato are susceptible to the pathogen (Read et al., 1995; Andrivon et al., 1997).

Black dot symptoms appear as a dark brownish-grey blemish of potato tubers, progressing to lesions covered with minute black dots of sclerotia. Colletotrichum coccodes can colonize all underground parts (daughter tubers, stolons and roots), basal stems (Andrivon et al., 1997, 1998) and also foliage (Mohan et al., 1992; Johnson & Miliczky, 1993; Johnson, 1994; Andrivon et al., 1998) of the potato plant, with the roots reported to be the most susceptible organs to infection (Komm & Stevenson, 1978; Andrivon et al., 1997; Andrivon et al., 1998).

Potato blemish diseases are of increasing concern because of the rise in sales of washed potatoes with a high quality skin appearance for the fresh market, and black dot symptoms can be responsible for a decrease in the market value of the crop (Jellis & Taylor, 1974; Read & Hide, 1995a). Colletotrichum coccodes was also reported to reduce potato yields under controlled greenhouse and field conditions by 7–30% (Stevenson et al., 1976; Barkdoll & Davis, 1992; Mohan et al., 1992; Johnson, 1994; Tsror et al., 1999). Infection by C. coccodes not only affects the quality and yield of potatoes for seed and consumption, but also serves as an important source of inoculum for future crops through the sowing of contaminated seed tubers (Jellis & Taylor, 1974; Read & Hide, 1988; Barkdoll & Davis, 1992; Johnson et al., 1997) and through C. coccodes surviving free in soil or on colonized plant debris for at least 2 years (Blakeman & Hornby, 1966; Coley-Smith & Cooke, 1971; Farley, 1976; Dillard, 1990; Dillard & Cobb, 1993) and up to 8 years in field soils (Dillard & Cobb, 1998). It is apparent that lengthy crop rotations are required to significantly decrease the viable inoculum of C. coccodes and therefore alleviate inoculum pressure.

To date, there are no effective control measures for C. coccodes. No specific fungicides have been developed to control black dot, and even chemicals that proved effective against C. coccodes in vitro and when applied to seed tubers under cropping conditions failed to control the disease during field experiments (Marais, 1990; Read & Hide, 1995b) and in national surveys of UK potato crops (Read et al., 1995).

No quantitative data are available on the extent of C. coccodes on infected seed tubers or on the current distribution of the fungus in UK field soils. An improved understanding is needed of the relationship of seed- and soilborne inoculum of C. coccodes to tuber infection, together with data concerning the quantitative contribution of these sources of inoculum to the incidence of black dot. This will permit accurate evaluations of the benefits of ‘clean’ seed and define the maximum levels of the pathogen on seed and in soil for the production of ware crops of the required quality. Diagnostic assays are therefore required to detect and quantify C. coccodes on potatoes and in soil. The polymerase chain reaction (PCR) is a diagnostic method widely used for the detection of plant pathogenic fungi (Miller, 1996). Real-time PCR is a modified PCR technique that uses two primers and an additional dual-labelled fluorogenic probe to allow the continuous monitoring of amplicon synthesis during thermocycling, and requires no post-PCR sample handling for target quantification (Orlando et al., 1998). The objectives of this work were to develop a sensitive and rapid PCR diagnostic assay for C. coccodes; to provide an improved understanding of the epidemiology of black dot; and to supply information for the development of effective disease control strategies. This study presents the development of specific primers for both conventional PCR and quantitative real-time (TaqMan) PCR to allow rapid and sensitive detection and monitoring of C. coccodes on potato tubers and in soil.

Materials and methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Design of C. coccodes-specific PCR primers

The internal transcribed spacer regions (ITS1 and ITS2) of the rDNA gene repeat of two or three isolates of each of the following 19 Colletotrichum species, previously characterized by Sreenivasaprasad et al. (1996), were aligned using the clustal v package (Higgins et al., 1992): C. coccodes (isolates 527·77, LC and NFT), C. acutatum, C. capsici, C. dematium, C. destructivum, C. fragariae, C. fructigenum, C. fuscum, C. gloeosporioides, C. graminicola, C. kahawae, C. linicola, C. lindemuthianum, C. musae, C. orbiculare, C. sublineolum, C. trichellum, C. trifolii and C. truncatum. Only one unique region existed for the design of a C. coccodes-specific primer in the ITS1 sequence, and none was identified in the ITS2 sequence. The 20-base region specific to C. coccodes was used to design the nested forward primer Cc1NF1, and regions conserved within the genus Colletotrichum were selected for the design of the nested reverse (Cc2NR1) and outer primer set (Cc1F1/Cc2R1). Two sets of 20-mer primers were designed (Table 1), and expected amplicon sizes for the outer and nested PCR primers were 447 and 349 bp, respectively. The Primer Express software (Applied Biosystems, Warrington, UK) was used to design primers and a TaqMan probe (Table 1) based on the alignment of ITS1 sequences to develop quantitative real-time PCR. The TaqMan probe (CcTqP1) was based on the same region used in the design of the C. coccodes-specific primer (Cc1NF1). The original primers developed were not used because real-time PCR requires small amplicons of 50–150 bp in length to yield consistent results (Applied Biosystems). The fluorogenic probe (CcTqP1) was labelled at the 5′ end with the fluorescent reporter dye VIC (Applied Biosystems), while the 3′ end was modified with the quencher dye TAMRA (6-carboxy-tetramethylrhodamine) (Applied Biosystems). The starting concentration of target sequence present in each reaction was calculated by comparing threshold cycle (Ct) values of unknown samples to the Ct values of standards with known amounts of C. coccodes DNA, where the Ct value is defined as the cycle number at which a statistically significant increase in the reporter fluorescence (i.e. exceeds the threshold) is first detected, and is dependent on the input of starting copies of target. Ct values were plotted against the log of the initial concentration of C. coccodes DNA to produce a standard curve. The specificity of all designed primer sequences for both conventional and real-time PCR were confirmed before synthesis following blast (www.ncbi.nlm.nih.gov/BLAST/) and fasta (www2.ebi.ac.uk/fasta3/) database searches of DNA sequences.

Table 1.  PCR primers specific for Colletotrichum spp. and C. coccodes
PrimersSequencea (5′ to 3′)Target DNASize of product (bp)
  • a

    Bases in bold and italics denote those used in the design of both conventional PCR primers and TaqMan primers/probes.

  • b

    The reverse strand of DNA was used in the design of the TaqMan probe in order to meet the optimal requirements for fluorescent PCR technology (PE Applied Biosystems).

Conventional PCR
First-round PCR (Colletotrichum-specific)  447
Cc1F1ACCTAACTGTTGCTTCGGCGITS1 
Cc2R1AAATTTGGGGGTTTTACGGCITS2 
Nested PCR (C. coccodes-specific)  349
Cc1NF1TGCCGCCTGCGGACCCCCCTITS1 
Cc2NR1GGCTCCGAGAGGGTCCGCCAITS2 
Real-time quantitative PCR
(C. coccodes-specific)  145
CcTqF1TCTATAACCCTTTGTGAACATACCTAACTGITS1 
CcTqR1CACTCAGAAGAAACGTCGTTAAAATAGAGITS1 
TaqMan probeb
CcTqP1CGCAGGCGGCACCCCCTITS1 

Extraction of microbial DNA from pure cultures, soil and potato tubers

Following growth of C. coccodes on potato dextrose agar (PDA, Oxoid, Basingstoke, UK) at 25°C for 7 days, a 6 mm2 plug was taken from the margin of colony growth and used to inoculate 50 mL potato dextrose broth (PDB, Oxoid). Total DNA was extracted by the method of Nicholson et al. (1996) from mycelium obtained from cultures grown in PDB at ambient temperature on an orbital shaker (100 r.p.m.) in darkness for up to 7 days. For the inoculation of nonsterile soil samples, spores or sclerotia were removed from the surface of colonies grown on PDA, resuspended in sterile distilled water (dH2O), and the concentration determined by direct microscopic counting. Separate suspensions of spores and sclerotia were made in sterile dH2O and used to inoculate 8 kg soil collected from the Scottish Agricultural Science Agency (SASA) Farm, Edinburgh and Pentland Hill, Edinburgh to a concentration of 106 spores and 2 × 104 sclerotia of C. coccodes per kg, respectively. These soils were then diluted serially with virgin soil to produce 8 kg samples that contained 1000, 300, 100, 30, 10 or 3 spores per g, or 40, 12, 4, 1·2, 0·4, 0·12 or 0·06 sclerotia of C. coccodes per g, or the equivalent volume of dH2O as an unseeded control. DNA was quantified using a TKO 100 Mini-Fluorometer (Hoefer Scientific Instruments, San Francisco, CA, USA) following the manufacturer’s instructions, and the quality was checked by agarose gel electrophoresis. Serial dilutions (10 ng µL−1 to 1 fg µL−1) genomic DNA in ultra-pure dH2O (HPLC-grade, Sigma-Aldrich, Poole, UK) were used as template to assess the sensitivity of PCR.

Soil samples

Samples were collected from three sites in Scotland and two in England, representing a total of four different soil types. Sites in Edinburgh, Scotland were located at Gogar Bank and the SASA Farm (both clay-loam soils), and Pentland Hill (a silty clay-loam); potatoes had not been planted at these sites for up to 13, 20 and 50 years, respectively. Soil samples were also obtained from Agricultural Development and Advisory Service (ADAS) survey sites in England at Arthur Rickwood Experimental Farm, Norfolk (fen peat soil) and Gleadthorpe, Nottinghamshire (sandy soil), sites which had not had a potato crop for 5 and 8 years, respectively. Duplicate or triplicate soil samples (10 g each) were resuspended in 20 mL extraction buffer [SPCB: 120 mm sodium phosphate, 2% hexadecyltrimethylammonium bromide (CTAB), 1·5 m NaCl pH 8·0] and fungal DNA was extracted from soil suspensions (1·5 mL) by physical disruption in a Mini-BeadBeater (Bio-Spec Products, Bartlesville, OK, USA) and purified by polyvinylpolypyrrolidone (PVPP) spin-column chromatography by the methods of Cullen et al. (2001).

Tuber samples

Tubers of the potato cultivars Maris Piper and Home Guard (which are classified as susceptible and resistant to black dot, respectively; Read, 1991) and of cv. Shula were obtained from field sites at the Scottish Crop Research Institute (SCRI). Tubers were washed under running water to remove soil particles, and peel strips (1–2 mm thick) were removed from the length of single tubers from the stolon heel end to the rose end, and also across a lateral section, with a hand-held potato peeler. Two methods for the extraction of DNA from tubers were compared to determine the most efficient and reliable for PCR analysis. DNA was extracted from tuber extract collected after passage of peel strips through a sap press or from diced frozen peel tissue using the methods described by Cullen et al. (2001). Percentage disease cover of potato tubers was visually estimated by the method of Andrivon et al. (1997).

PCR methods

PCR amplification of samples with the outer primer set (Cc1F1/Cc2R1) was based on an initial denaturation step at 95°C (2 min), followed by 35 cycles of denaturation at 95°C for 45 s, annealing at 61°C for 1 min and extension at 72°C for 90 s, plus a final elongation step at 72°C for 5 min, in a reaction volume of 25 µL using a GeneAmp PCR System 9600 thermal cycler (Applied Biosystems). A two-step program was used for nested PCR because of the high melting temperature (Tm) values of the primers Cc1NF1 (Tm = 72°C) and Cc2NR1 (Tm = 70°C), and involved an initial denaturation step at 95°C (2 min), followed by 35 cycles of denaturation at 95°C for 45 s and annealing and extension at 72°C for 135 s, plus a final elongation at 72°C for 5 min, in a reaction volume of 25 µL. Optimal conditions for PCR (single-round and nested) contained a master mix of the following components: 1× reaction buffer (16 mm[NH4]2SO4, 67 mm Tris-HCl pH 8·8, 0·1% Tween-20; Bioline Ltd, London, UK); deoxynucleoside triphosphates (dNTPS; each 200 µm; Bioline); primers (each 0·3 µm; MWG-Biotech UK Ltd, Milton Keynes, UK); 5·0 mm MgCl2; 250 µg mL−1 BSA (Roche Diagnostics, Lewes, UK); and 1 or 2 U Biolase Diamond (Bioline; the latter amount was used for soil and potato DNA extracts). One µL of undiluted or 1/10-diluted DNA (representing 10–100 ng) was used as template and 1 µL of first-round PCR product was used for nested PCR. Colletotrichum coccodes DNA (10 ng) was used as a positive control in the PCR assay; negative controls were carried out with PCR reagents and 1 µL dH2O or nontarget DNA. PCR products were analysed by electrophoresis on a 2% agarose gel (Bioline) in 1× TBE buffer (89 mm Tris base, 89 mm boric acid, 2 mm EDTA pH 8·0), stained with ethidium bromide (0·5 mg L−1) and photographed under UV illumination (Sambrook et al., 1989).

Reaction components for quantitative real-time PCR were obtained from the TaqMan Universal PCR Master Mix (Applied Biosystems), and the reaction was performed in MicroAmp optical 96-well plates using the automated ABI Prism 7700 sequence detector (Applied Biosystems). Primers CcTqF1/CcTqR1 were included at a final concentration of 0·3 µm per reaction and the Taqman probe (CcTqP1) was used at 0·1 µm. Reaction volumes of 25 and 50 µL were compared using 2 µL undiluted and diluted (1/10, 1/20, 1/40, 1/50 and 1/100 dilutions) template DNA to determine the optimum concentrations for reliable quantification. The manufacturer’s recommended universal thermal cycle protocol (Applied Biosystems) was used for PCR amplification: stage 1 [50°C for 2 min; AmpErase uracil-N-glycosylase (UNG) digestion]; stage 2 (95°C for 10 min; denaturation of UNG and activation of AmpliTaq Gold DNA polymerase); and stage 3 (45 cycles at 95°C for 15 s and 60°C for 1 min). The Ct values for each PCR reaction were automatically calculated and analysed by the ABI prism sequence detection systems software.

The specificity of all primer sets was tested against genomic DNA from 33 European and North American isolates of C. coccodes (Table 2); other isolates of Colletotrichum spp. (C. acutatum, C. capsici, C. dematium, C. fragariae, C. gloeosporioides, C. graminicola, C. linicola and C. trifolii) representing the six phylogenetic groups used to divide the genus (Sreenivasaprasad et al., 1996); and also from a range of other plant pathogens in the SCRI culture collection (Table 3). To exclude false-negative results with C. coccodes-specific primers, all template DNA samples were tested for PCR amplification using universal primers ITS5/4 for fungal 18S and 28S rDNA (White et al., 1990), and V3-region primers 341F/534R for 16S bacterial rDNA following the method of Muyzer et al. (1993).

Table 2.  Isolates of Colletotrichum coccodes obtained from potato, listed according to origin and date of isolation
IsolateOriginaDate of isolation
  • a

    BPC, British Potato Council; CSL, Central Science Laboratory; SASA, Scottish Agricultural Science Agency; SCRI, Scottish Crop Research Institute.

C1SCRI, Dundee, Scotland1996
C2Washington State University, USA1997
C3Washington State University, USA1997
C4Washington State University, USA1997
C5SCRI, Dundee, Scotland1985
C6SCRI, Dundee, Scotland1989
C7SCRI, Dundee, Scotland1988
C8Dept. Agriculture, N. Ireland1997
C9SASA, Montrose, Scotland1997
C10SASA, Newmill, Scotland1998
C11SASA, Brichie, Scotland1998
C12SASA, Scotland1998
C13Dept. Agriculture, N. Ireland1998
C14Dept. Agriculture, N. Ireland1998
C15Dept. Agriculture, N. Ireland1998
C16SCRI, Dundee, Scotland1998
C17TESCO, Scotland1998
C18TESCO, Scotland1998
C19Rothamsted Farm, England1983
C20Rothamsted Farm, England1988
C21Rothamsted Farm, England1988
C22Rosefarm, Scotland1991
C23Dalreoch Farm, Scotland1991
C24BPC, Sutton Bridge, England1997
C25BPC, Sutton Bridge, England1997
C26BPC, Sutton Bridge, England1997
C27BPC, Sutton Bridge, England1997
C28BPC, Sutton Bridge, England1997
C29BPC, Sutton Bridge, England1997
C30SCRI, Dundee, Scotland1998
C31SCRI, Dundee, Scotland1998
C32SCRI, Dundee, Scotland1998
C33CSL, York, England1999
Table 3.  Fungal, oomycete, plasmodiophorid and bacterial species used in this study to test the specificity of Colletotrichum coccodes primers
Fungal and other eukaryotic speciesBacterial species
  • a

    DNA samples were obtained from S. Foster, IACR-Rothamsted, Harpenden, UK.

  • b

    DNA samples were obtained from K. Hughes, Central Science Laboratories, York, UK.

  • c

    Isolates were obtained from R. Loria, Cornell University, Ithaca, NY, USA.

Alternaria brassicaeaEnterobacter aerogenes
Botrytis cinereaErwinia carotovora ssp. atroseptica
Colletotrichum acutatumbErwinia carotovora ssp. betavasculorum
Colletotrichum capsicibErwinia carotovora ssp. carotovora
Colletotrichum coccodesErwinia carotovora ssp. odorifera
Colletotrichum dematiumbErwinia carotovora ssp. wasabiae
Colletotrichum fragariaebErwinia chrysanthemi
Colletotrichum gloeosporioidesbErwinia rhapontici
Colletotrichum graminicolabErwinia stewartii
Colletotrichum linicolabPseudomonas syringae pv. morsprunorum
Colletotrichum trifoliibPseudomonas syringae pv. syringae
Erysiphe graminis f.sp. avenaeRalstonia solanacearum
Erysiphe graminis f.sp. hordeiStreptomyces acidiscabiesc
Fusarium coeruleumStreptomyces scabiesc
Fusarium oxysporumaXanthomonas albilineans
Fusarium sulphureumXanthomonas campestris pv. campestris
Helminthosporium solaniXanthomonas campestris pv. phaseoli
Leptosphaeria maculansaXanthomonas campestris pv. phaseoli var. fuscans
Nectria haematococcaaXanthomonas campestris pv. vesicatoria
Neurospora crassaa 
Phoma exigua var. foveata 
Phytophthora cactorum 
Phytophthora cryptogea 
Phytophthora erythroseptica 
Phytophthora fragariae 
Phytophthora infestans 
Polyscytalum pustulans 
Puccinia recondita f.sp. tritici 
Pyrenopeziza brassicaea 
Pythium ultimum 
Rhizoctonia solani 
Sclerotinia sclerotioruma 
Spongospora subterranea 
Tapesia yallundaea 
Verticillium dahliaea 
Verticillium lecanii 

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Design, specificity and sensitivity of C. coccodes-specific primers for conventional PCR

Two sets of PCR primers were designed to sequences within the ITS1/ITS2 regions of C. coccodes isolates 527·77, LC and NFT (Sreenivasaprasad et al., 1996) after alignment of the ITS sequences with other previously characterized isolates of Colletotrichum spp., representing 18 species (see Materials and methods). The outer primers (Cc1F1/Cc2R1) were designed in conserved regions to be genus-specific for Colletotrichum spp., while the nested primers (Cc1NF1/Cc2NR1) were designed to be specific for C. coccodes based on a unique sequence in primer Cc1NF1. On testing the PCR for the detection of C. coccodes, a specific product (349 bp) was reliably detected following a nested PCR (primers Cc1NF1/Cc2NR1) when genomic DNA was included at concentrations down to and including 1 fg per reaction (data not shown). Outer and nested sets of primers amplified product of the correct size during a first-round PCR from the corresponding genomic DNA of all 33 isolates of C. coccodes tested (Table 2; Fig. 1). Comparisons between primer sequences Cc1F1, Cc2R1 and Cc2NR1 with DNA database sequences (blast and fasta programs) of other plant pathogenic fungi and bacteria revealed no significant levels of similarity, apart from 90–100% homology to ITS sequences in EMBL and GenBank for other isolates of Colletotrichum spp. (see Materials and methods). The C. coccodes-specific primer (Cc1NF1) showed homology (100%) only to the unique 20-base sequence in the ITS1 region of C. coccodes following a database search. Primer specificity was confirmed for both outer and nested primer sets by the absence of PCR product when testing genomic DNA from a wide range of other fungal and bacterial plant pathogens (Table 3; Fig. 1). The specificity of the nested primers (Cc1NF1/Cc2NR1) to C. coccodes was also demonstrated by the absence of PCR product when testing DNA from eight other Colletotrichum taxa representing the six phylogenetic groups of the genus (Table 3; Fig. 1). The quality of all DNA preparations tested for amplification by PCR throughout this study was confirmed by the detection of product using either primers Cc1F1/Cc2R1 for all Colletotrichum spp., universal eukaryotic primers (ITS4/5), or universal prokaryotic (341F/534R) primers (data not shown).

image

Figure 1. Specificity testing of Colletotrichum coccodes-specific primers against genomic DNA from different isolates of C. coccodes and a wide range of other plant pathogens. Upper row, lanes (4 µL product): 1, DNA marker; 2, negative control (dH2O); 3–10 (amplification with Cc1F1/Cc2R1), C. coccodes isolates C13, C15, C18, C20, C22, C23, C24 and C27; 11–20 (amplification with Cc1NF1/Cc2NR1), C. coccodes isolates C1, C2, C3, C4, C5, C6, C7, C8, C9 and C10. Lower row, lanes (5 µL product): 1, DNA marker; 2, negative control (dH2O); 3–20 (amplification with Cc1NF1/Cc2NR1), C. coccodes isolate C1, C. acutatum, C. dematium, C. gloeosporioides, C. graminicola, C. linicola, C. trifolii, Fusarium coeruleum, Fusarium sulphureum, Helminthosporium solani, Phoma exigua var. foveata, Phytophthora erythroseptica, Phytophthora infestans, Polyscytalum pustulans, Pythium ultimum, Rhizoctonia solani, Spongospora subterranea and C. coccodes isolate C2.

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Detection of C. coccodes from potato tubers and soil by conventional PCR

The assay detected a single product of the expected size following both a first-round PCR (Cc1F1/Cc2R1; 447 bp) and nested PCR (Cc1NF1/Cc2NR1; 349 bp) when tested against DNA extracted from tubers with no visible disease symptoms, or with either 5% (Maris Piper) or 50% (Home Guard) cover of black dot (Fig. 2). No amplification products were detected using DNA extracts obtained from tubers showing symptoms of silver scurf (cv. Shula) or common scab (cv. Maris Piper), confirming both the specificity of the primers to detect C. coccodes and the absence of C. coccodes on the tubers tested (Fig. 2). There was no difference in the sensitivity of detection of C. coccodes after nested PCR when DNA was extracted from either peel extract or diced and frozen tuber peel from cultivars Maris Piper or Home Guard, or when undiluted and diluted (1/10) DNA extracts were compared (Fig. 2).

image

Figure 2. Detection of Colletotrichum coccodes by nested PCR from naturally infected potato tubers. Upper row, lanes 4–15 show results after first-round amplification (primers Cc1F1/Cc2R1) of DNA extracted from frozen peel (lanes 4, 5, 6, 8, 10, 12 and 14) and peel extract (lanes 7, 9, 11, 13 and 15). Lanes (4 µL undiluted product): 1 and 16, DNA marker; 2, negative control (dH2O); 3, positive control (C. coccodes isolate C1); 4–15, undiluted DNA extracts from cvs Shula (lane 4), Maris Piper (lanes 5–9) and Home Guard (lanes 10–15): 4, symptoms of silver scurf; 5, symptoms of common scab; 6–7, zero control; 8–9, 5% disease cover; 10–11, zero control; 12–13, no visible symptoms; 14–15, 50% disease cover. Lower row, lanes (4 µL 1/10-diluted product) 2–15 show amplification results using nested PCR (primers Cc1NF1/Cc2NR1) following the amplification of first-round products (1 µL) shown in lanes 2–15 of the upper row. PCR products in the lower row were diluted for visualizing on gel because of the high concentration amplified during nested PCR.

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Colletotrichum coccodes was detected consistently by nested PCR (primers Cc1NF1/Cc2NR1) in seeded soil samples down to and including a level of three spores per g (data not shown) or the equivalent of 0·12 sclerotia per g (Fig. 3) using two different soil types (see Materials and methods), and no signals were detected from unseeded controls (Fig. 3). In addition, product of the correct size after nested PCR was also detected from the same soil samples (usually two out of three replicates) seeded with the equivalent of 0·06 sclerotia per g, the detection limit of the PCR system. Colletotrichum coccodes was also detected by PCR in naturally infested field soils collected from ADAS Arthur Rickwood, ADAS Gleadthorpe and Gogar Bank Farm, representing three different soil types that had not been planted with potatoes for 5, 8 and 13 years, respectively (Fig. 3). Stronger signals were obtained during first-round PCR (genus-specific primers) than after nested PCR from soils collected from Gogar Bank Farm and Gleadthorpe, indicating the presence of other Colletotrichum spp. in addition to C. coccodes (Fig. 3). Viable propagules of C. coccodes were also detected in the same soil samples from Gogar Bank Farm by a minituber baiting test, and from conventional seed potatoes (including stems and roots) grown at Arthur Rickwood and Gleadthorpe (unpublished results).

image

Figure 3. Detection of Colletotrichum coccodes in seeded soil and naturally infested soils by nested PCR. Upper row, lanes 2–20 show results after first-round amplification (primers Cc1F1/Cc2R1) of undiluted soil DNA extracts. Lanes (4 µL product): 1, DNA marker; 2, negative control (dH2O); 3, positive control (C. coccodes isolate C1); 4, unseeded Arthur Rickwood soil; 5–6, unseeded Gogar Bank soil; 7–8, unseeded Pentland Hill soil; 9–18, seeded (C. coccodes sclerotia per g soil) Pentland Hill soil samples as follows: 9–10, 12 sclerotia; 11–12, four sclerotia, 13–14, 1·2 sclerotia; 15–16, 0·12 sclerotia; 17–18, 0·06 sclerotia; 19–20, unseeded Gleadthorpe soil. Lower row, lanes (4 µL product) 2–20 show amplification results of nested PCR (primers Cc1NF1/Cc2NR1) following the amplification of first-round products (1 µL) shown in lanes 2–20 of the upper row.

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Real-time quantification of C. coccodes DNA extracted from potato tubers and soil samples

An automated ABI Prism 7700 sequence detector, together with the primers (CcTqF1/CcTqR1) and the fluorogenic probe (CcTqP1) developed in this study, were used to perform quantitative real-time PCR. The standard curve used to calculate the starting concentration of C. coccodes template DNA had a linear correlation coefficient of r = 0·978 (Fig. 4), demonstrating a high reproducibility among different replicates and the accuracy of this PCR-based quantification assay. During the optimization of real-time PCR, it was determined that 2 µL of 1/20-diluted soil DNA extracts or potato tuber DNA extracts in a reaction volume of 50 µL provided the optimal conditions for accurate quantification of C. coccodes DNA. No signals were obtained for the same undiluted DNA extracts (2 µL) in a reaction volume of 50 µL or from undiluted and diluted DNA extracts in 25 µL volumes.

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Figure 4. Standard graph used for the absolute quantification of Colletotrichum coccodes target DNA during real-time PCR.

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The specificity of the primers/probe developed for real-time quantitative detection of C. coccodes was confirmed when no signals were generated for DNA extracts of eight different Colletotrichum spp. and 46 other plant pathogenic species of fungi, oomycetes a plasmodiophorid and bacteria (Table 3). All potato tuber extracts and soil samples tested by conventional PCR were also analysed and quantified by real-time PCR (Tables 4 and 5). The same high level of sensitivity was achieved when testing tuber extracts (symptomless) and seeded soil samples (3·0 spores per g or 0·06 sclerotia per g soil) by real-time PCR after a first round of 45 cycles, as during conventional nested PCR (a total of 70 cycles), confirming that both methods validate each other. Colletotrichum coccodes target DNA was detected in apparently healthy potato tubers during real-time PCR, and the lowest concentration of target DNA was detected in peel extract from symptomless tubers at concentrations of 3·26 and 6·20 pg for cvs Maris Piper and Home Guard, respectively (Table 4). A higher concentration of target DNA was recovered when DNA was extracted from frozen peel tissue rather than peel extract (Table 4) and, as expected, higher levels of target DNA were detected as percentage disease cover increased from 5 to 50%. No signals specific to C. coccodes were detected in control tubers or those showing symptoms of silver scurf or common scab (Table 4).

Table 4.  Quantification of the concentration of Colletotrichum coccodes template DNA in naturally infected potato tubers
Cultivar/sampleaDisease cover (%)bConcentration ofC. coccodes DNAc (pg)
  • a

    DNA was extracted from frozen peel or peel extract as described in Materials and methods.

  • b

    Disease cover as estimated by Andrivon et al. (1997).

  • c

    Mean concentration ± SD from three replications.

Maris Piper
Frozen peelControl (0)   0
Peel extractControl (0)   0
Frozen peel 0   4·86 ± 0·42
Peel extract 0   3·26 ± 0·30
Frozen peel 5 135·3 ± 14·2
Peel extract 5  14·6 ± 1·25
Common scab symptoms70   0
Home Guard
Frozen peelControl (0)   0
Peel extractControl (0)   0
Frozen peel 0 188·0 ± 10
Peel extract 0   6·20 ± 1·11
Frozen peel501680·0 ± 220
Peel extract50 400·0 ± 60
Shula
Silver scurf symptoms50   0
Table 5.  Quantification of the starting concentration of Colletotrichum coccodes template DNA in soil samples
Number of spores or sclerotia added to soil (g−1)Concentration ofC. coccodes DNAa
  • a

    Mean concentration ± SD from six (*) or nine (†) replications as indicated.

Gogar Bank Farm*
0 spores/sclerotia  432 ± 52·0 fg
ADAS Gleadthorpe*
0 spores/sclerotia  25·7 ± 2·05 pg
ADAS Arthur Rickwood*
0 spores/sclerotia 11·52 ± 0·44 pg
SASA Farm*
0 spores0
3 spores 118·0 ± 25·7 fg
30 spores 883·3 ± 129·2 fg
300 spores 5133·3 ± 281 fg
Pentland Hill
0 sclerotia0
0·06 sclerotia 23·0 ± 1·41 fg
0·12 sclerotia 43·0 ± 26·6 fg
1·2 sclerotia 154·0 ± 54·9 fg
12 sclerotia 1573·3 ± 178 fg

Quantitative real-time PCR was sensitive enough to detect the lowest numbers of C. coccodes spores (3·0 spores per g) and sclerotia (0·06 sclerotia per g) added to soil samples, and detected target DNA down to fg levels (Table 5). It was also demonstrated that there was no difference in DNA extraction efficiency from these two types of fungal propagule when added to soil at low concentrations. It was possible to detect the equivalent of 3·0 C. coccodes spores per g in six out of six replicate seeded samples of SASA farm soil by real-time PCR (Table 5). The mean quantity of target DNA detected from this seeded soil, based on a comparison with the standard curve, was 118 fg (equivalent to 5·9 fg per PCR reaction at a 1/20 DNA dilution). In addition, C. coccodes target DNA was also detected from soil samples (usually three out of nine replicates) seeded at rates down to and including the equivalent of 0·06 sclerotia per g, which was equal to an extracted quantity of 23·0 fg (equivalent to 1·15 fg per PCR reaction at a 1/20 DNA dilution). The concentration of C. coccodes target DNA in the unseeded soil sample of the Gogar Bank farm was estimated at 432 fg (equivalent to 21·6 fg per PCR reaction at a 1/20 dilution) and therefore equivalent to three to 30 spores per g or 1·2 to 12 sclerotia per g when compared to the amounts estimated for seeded soil samples (Table 5). The highest levels of C. coccodes DNA were extracted and detected in the naturally infested soils at the ADAS field sites of Arthur Rickwood (11·52 pg DNA per g soil) and Gleadthorpe (25·7 pg DNA per g soil), and by extrapolating from the standard curve, these levels represent the equivalent of approximately 700–1500 spores or 90–200 sclerotia per g.

Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

This paper describes the development of simple, robust and quantitative PCR techniques for the specific detection of C. coccodes. These assays were based on the design of two sets of primers (outer and nested) for conventional PCR and one primer/probe set for quantitative real-time (TaqMan) PCR. Together with the DNA-extraction protocols, these diagnostic methods will allow rapid and accurate assessment of tuber and soil contamination by C. coccodes. The specificity of both PCR systems was confirmed by testing against 33 isolates of C. coccodes, eight different Colletotrichum spp. and 46 other plant pathogenic eukaryotes and prokaryotes (Tables 2 and 3). Both PCR assays were of a high sensitivity and specificity to allow the detection of C. coccodes on potato tubers in the absence of visible symptoms of black dot, and in soil seeded with inoculum at rates down to and including three spores per g, or the equivalent of 0·06 sclerotia per g soil. Real-time PCR has the advantage of calculating the absolute starting quantity of C. coccodes DNA in tuber or soil samples within 3 h, in contrast to the 6 h taken by conventional nested PCR, which can only indicate the presence or absence of the target organism. The data obtained from real-time PCR also showed that the reproducibility among replicates was high (Tables 4 and 5), demonstrating that the assay was accurate in terms of DNA extraction and recovery rates from all potato tuber and seeded soil samples. The choice between the two PCR systems will be dictated by cost as the ABI Prism 7700 sequence detector system (Applied Biosystems) is more expensive than the equipment used for conventional PCR, even following the further development of this latter system to allow quantification, for example, based on the use of competitor template fragments (Bell et al., 1999; Hyman et al., 2000).

The sensitivity of the PCR assay described here is revealed when the number of spores or sclerotia included per PCR reaction is considered. Assuming 100% recovery of DNA after extraction from soil seeded with three spores per g (2·25 spores per 0·75 g sampled; DNA resuspended in 75 µL) or 0·06 sclerotia per g (0·045 sclerotia per 0·75 g sampled; DNA resuspended in 75 µL), 1 µL of undiluted soil DNA extract would contain the equivalent of only 0·03 spores or 0·0006 sclerotia per PCR reaction. It was still possible to detect a signal from the 1/20 dilutions (equivalent to 0·0015 spores or 0·00003 sclerotia per reaction) of these seeded-soil DNA extracts by conventional and real-time PCR.

Although the blemish diseases black dot and silver scurf produce similar symptoms on potato, the PCR assay was able to distinguish C. coccodes from Helminthosporium solani (silver scurf; Errampalli et al., 2001), and can thus be used to differentiate symptoms of black dot from those of silver scurf. The detection of C. coccodes by PCR, minituber baiting, or on harvested potato tubers in naturally infested field soils at ADAS Arthur Rickwood, ADAS Gleadthorpe and Gogar Bank, which had not been planted with potatoes for 5, 8 and 13 years, respectively, further demonstrated the organism’s ability to survive in soil for long periods (Blakeman & Hornby, 1966; Coley-Smith & Cooke, 1971; Farley, 1976; Dillard, 1990; Dillard & Cobb, 1993; Dillard & Cobb, 1998). The infection of minitubers and conventional seed potatoes by C. coccodes confirmed that viable propagules existed, and highlighted the potential threat of disease development in potato crops if planted in these three field soils. Such a PCR test may thus be crucial when making crop management decisions regarding suitable field sites for planting potatoes. The PCR detection limits set for C. coccodes sclerotia in soil are more meaningful, as sclerotia serve as overwintering and survival structures for the fungus (Dillard & Cobb, 1998).

The success and reliability of any PCR assay will be dependent on obtaining high yields and representative samples of target DNA from environmental samples. A simple and inexpensive extraction method was used in this study with a small number of efficient lysis and purification steps to allow rapid processing of many samples. The extraction procedure was based on the physical disruption of microbes within the resuspended soil or tuber extract sample using a beadbeater. Lysis based on physical disruption is the method previously reported to lyse the majority of cell types within a sample and to produce the highest yields of DNA (Borneman et al., 1996; Yeates et al., 1997; Cullen & Hirsch, 1998; Miller et al., 1999). High molecular-weight (23 kb) DNA of a suitable purity for PCR was recovered from soil (four different soil types) and potato tubers within 3 h by the modified extraction method (Cullen et al., 2001). Potato peel extracts (obtained by passing peel strips through a sap press) and frozen and diced peel tissue samples were both used successfully for extraction and PCR detection of C. coccodes, although the former method was less laborious and time-consuming. It was possible to detect C. coccodes in soil spiked with as few as three spores per g or the equivalent of 0·06 sclerotia per g in a single day with the DNA-extraction and PCR protocols developed. The same DNA-extraction methods have been used successfully to detect other potato blemish pathogens such as H. solani (Cullen et al., 2001), Streptomyces scabies (Cullen et al., 1999) and Spongospora subterranea (Bell et al., 1999) in soil and plant material.

The development of management strategies requires a precise estimate of the role of seed inoculum relative to soil inoculum in order to evaluate the benefits of ‘clean’ seed, as well as to study the effect of closer rotations on the tuber health of ware crops. Denner et al. (1998) reported that the contribution of soilborne inoculum (P < 0·001) to the incidence of black dot on progeny tubers was at least twice that of seedborne inoculum, even when seed tubers were severely infected with C. coccodes. In addition, the maximum levels of disease on seed allowing the production of ware crops of the required quality need to be defined, together with an estimation of soil inoculum levels, in order to relate these to disease incidence in the crop.

The sensitive and accurate quantitative PCR assay developed for C. coccodes in this study will permit more discriminating investigations on factors such as soil type and soilborne inoculum, seed inoculum, previous cropping history, irrigation and the efficacy of fungicides, which may all have an impact on the incidence and severity of black dot. This will also assist in epidemiological studies, offering the advantage of achieving results in a single day compared to up to 6–12 weeks with the present glasshouse-based bait tests. A rapid detection assay would also provide the industry with a tool to enable growers to avoid high-risk situations and apply a rational selection of potential fungicides to control black dot, and would permit more detailed monitoring of fungal populations.

PCR diagnostics thus offer the potential to allow rapid and accurate identification of diseases and to quantify the presence of a pathogen in presymptomatic potato stocks and in soil. The molecular tools developed are currently being used to measure the inoculum levels of C. coccodes, H. solani and S. scabies in several field soils and potato crops as part of an epidemiological survey involving crop rotation. The aim is to determine the threshold levels necessary for the development of potato blemish diseases, and to study the interactions between cultural and husbandry factors. If successful, this work will result in a predictive diagnostic test that will enable the industry to identify high-risk fields and/or seed stocks where populations of blemish pathogens are above threshold values.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

We gratefully acknowledge the following for the generous provision of isolates: Stuart Carnegie, James Choiseul, Louise Cooke, Jane Etheridge, Simon Foster, Kelvin Hughes, Graham Jellis and Rosemary Loria. We thank Mairi Nicolson for assistance with DNA extractions. We are also grateful to Vanessa Young and Lillian Yengi for guidance and assistance with real-time PCR analysis, and the Biomedical Research Centre, Ninewells Hospital, Dundee for the use of the ABI Prism 7700 sequence detector. This work was funded by the former Ministry of Agriculture, Fisheries and Food (MAFF), UK (grant number HP0125T).

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References
  • Andrivon D, Ramage K, Guérin C, Lucas JM, Jouan B, 1997. Distribution and fungicide sensitivity of Colletotrichum coccodes in French potato-producing areas. Plant Pathology 46, 7228.
  • Andrivon D, Lucas J-M, Guérin C, Jouan B, 1998. Colonization of roots, stolons, tubers and stems of various potato (Solanum tuberosum) cultivars by the black-dot fungus Colletotrichum coccodes. Plant Pathology 47, 4405.
  • Barkdoll AW, Davis JR, 1992. Distribution of Colletotrichum coccodes in Idaho and variation in pathogenicity on potato. Plant Disease 76, 1315.
  • Bell KS, Claxton JR, Roberts J, Cullen DW, Williams NA, Harrison JG, Toth IK, Cooke. DEL, Duncan JD, 1999. Detection and quantification of Spongospora subterranea f. sp. subterranea in soils and on tubers using specific PCR primers. European Journal of Plant Pathology 105, 90515.
  • Blakeman JP, Hornby D, 1966. The persistence of Colletotrichum coccodes and Mycosphaerella ligulicola in soil, with special reference to sclerotia and conidia. Transactions of the British Mycological Society 49, 22740.
  • Borneman J, Skroch PW, O’Sullivan KM, Palus JA, Rumjanek NG, Jansen JL, Nienhuis J, Triplett EW, 1996. Molecular microbial diversity of an agricultural soil in Wisconsin. Applied and Environmental Microbiology 62, 193543.
  • Coley-Smith JR, Cooke RC, 1971. Survival and germination of fungal sclerotia. Annual Review of Phytopathology 9, 6592.
  • Cullen DW, Hirsch PR, 1998. Simple and rapid method for direct extraction of microbial in DNA from soil for PCR. Soil Biology and Biochemistry 30, 98393.
  • Cullen DW, Lees AK, Toth IK, Duncan JM, 1999. Development of a PCR assay for specific detection of the three main pathogens of potato blemish diseases. In: Proceedings: Crop Protection in Northern Britain, Dundee. Farnham, UK: British Crop Protection Council, 2615.
  • Cullen DW, Lees AK, Toth IK, Duncan JM, 2001. Conventional PCR and real-time quantitative PCR detection of Helminthosporium solani in soil and on potato tubers. European Journal of Plant Pathology 107, 38798.
  • Denner FDN, Millard CP, Wehner FC, 1998. The effect of seed- and soilborne inoculum of Colletotrichum coccodes on the incidence of black dot on potatoes. Potato Research 41, 516.
  • Dillard HR, 1990. Survival of Colletotrichum coccodes in New York. Phytopathology 80, 1026 (abstract).
  • Dillard HR, 1992. The pathogen and its hosts. In: BaileyJA, JegerMJ, eds. Colletotrichum: Biology, Pathology and Control. Wallingford, UK: CAB International, 22536.
  • Dillard HR, Cobb AC, 1993. Persistence of Colletotrichum coccodes on tomato roots and in soil. Phytopathology 83, 1345 (abstract).
  • Dillard HR, Cobb AC, 1998. Survival of Colletotrichum coccodes in infected tomato tissue and in soil. Plant Disease 82, 2358.
  • Errampalli D, Saunders JM, Holley JD, 2001. Emergence of silver scurf (Helminthosporium solani) as an economically important disease of potato. Plant Pathology 50, 14153.
  • Farley JD, 1976. Survival of Colletotrichum coccodes in soil. Phytopathology 66, 6401.
  • Higgins DG, Bleasby AJ, Fuchs R, 1992. clustal v– improved software for multiple sequence alignment. Computer Applications in the Biosciences 8, 18991.
  • Hyman LJ, Birch PRJ, Dellagi A, Avrova A, Toth IK, 2000. A competitive PCR-based method for the detection and quantification of Erwinia carotovora subsp. atroseptica on potato tubers. Letters in Applied Microbiology 30, 3305.
  • Jellis GJ, Taylor GS, 1974. The relative importance of silver scurf and black dot: two disfiguring diseases of potato tubers. ADAS Quarterly Review 14, 97112.
  • Johnson DA, 1994. Effect of foliar infection caused by Colletotrichum coccodes on yield of Russet Burbank potato. Plant Disease 78, 10758.
  • Johnson DA, Miliczky ER, 1993. Effects of wounding and wetting duration on infection of potato foliage by Colletotrichum coccodes. Plant Disease 77, 137.
  • Johnson DA, Rowe RC, Cummings TF, 1997. Incidence of Colletotrichum coccodes in certified potato seed tubers planted in Washington state. Plant Disease 81, 1199202.
  • Komm DA, Stevenson WR, 1978. Tuber-borne infection of Solanum tuberosum‘Superior’ by Colletotrichum coccodes. Plant Disease Reporter 62, 6827.
  • Marais L, 1990. Efficacy of fungicides against Colletotrichum coccodes on potato tubers. Potato Research 33, 27581.
  • Miller SA, 1996. Detecting propagules of plant pathogenic fungi. Advances in Botanical Research 23, 73102.
  • Miller DN, Bryant JE, Madsen EL, Ghiorse WC, 1999. Evaluation and optimisation of DNA extraction and purification procedures for soil and sediment samples. Applied and Environmental Microbiology 65, 471524.
  • Mohan SK, Davis JR, Sorensen LH, Schneider AT, 1992. Infection of aerial parts of potato plants by Colletotrichum coccodes and its effects on premature vine death and yield. American Potato Journal 69, 54759.
  • Muyzer G, De Waal EC, Uitterlinden AG, 1993. Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Applied and Environmental Microbiology 59, 695700.
  • Nicholson P, Lees AK, Maurin N, Parry DW, Rezanoor HN, 1996. Development of a PCR assay to identify and quantify Microdochium nivale var. nivale and Microdochium nivale var. majus in wheat. Physiological and Molecular Plant Pathology 48, 25771.
  • Orlando C, Pinzani PP, Pazzagli M, 1998. Developments in quantitative PCR. Clinical Chemistry Laboratory Medicine 36, 25569.
  • Read PJ, 1991. The susceptibility of tubers of potato cultivars to black dot (Colletotrichum coccodes (Wallr.) Hughes). Annals of Applied Biology 119, 47582.
  • Read PJ, Hide GA, 1988. Effects of inoculum source and irrigation on black dot disease of potatoes (Colletotrichum coccodes (Wallr.) Hughes) and its development during storage. Potato Research 31, 493500.
  • Read PJ, Hide GA, 1995a. Development of black dot disease (Colletotrichum coccodes (Wallr.) Hughes) and its effects on the growth and yield of potato plants. Annals of Applied Biology 127, 5772.
  • Read PJ, Hide GA, 1995b. Effects of fungicides on the growth and conidial germination of Colletotrichum coccodes and on the development of black dot disease of potatoes. Annals of Applied Biology 126, 43747.
  • Read PJ, Storey RM, Hudson DR, 1995. A survey of black dot and other fungal tuber blemishing diseases in British potato crops at harvest. Annals of Applied Biology 126, 24958.
  • Sambrook J, Fritsch EF, Maniatis T, 1989. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor, NY, USA: Cold Spring Harbor Laboratory Press.
  • Sreenivasaprasad S, Mills PR, Meehan BM, Brown AE, 1996. Phylogeny and systematics of 18 Colletotrichum species based on ribosomal DNA spacer sequences. Genome 39, 499512.
  • Stevenson WR, Green RJ, Bergesen GB, 1976. Occurrence and control of black dot root in Indiana. Plant Disease Reporter 60, 24851.
  • Sutton BC, 1992. The genus Glomerella and its anamorph Colletotrichum. In: BaileyJA, JegerMJ, eds. Colletotrichum: Biology, Pathology and Control. Wallingford, UK: CAB International, 126.
  • Tsror (Lahkim) L, Erlich O, Hazanovsky M, 1999. Effect of Colletotrichum coccodes on potato yield, tuber quality, and stem colonization during spring and autumn. Plant Disease 83, 5615.
  • Waller JM, 1992. Colletotrichum diseases of perennial and other cash crops. In: BaileyJA, JegerMJ, eds. Colletotrichum: Biology, Pathology and Control. Wallingford, UK: CAB International, 16785.
  • White TJ, Bruns T, Lee S, Taylor J, 1990. Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. In: InnisMA, GelfandDH, SninskyJJ, WhiteTJ, eds. PCR Protocols: A Guide to Methods and Applications. San Diego, CA, USA: Academic Press, 31522.
  • Yeates C, Gillings MR, Davison AD, Altavilla N, Veal DA, 1997. PCR amplification of crude microbial DNA extracted from soil. Letters in Applied Microbiology 25, 3037.