Development of conventional and quantitative real-time PCR assays for the detection and identification of Rhizoctonia solani AG-3 in potato and soil
Article first published online: 29 MAY 2002
Volume 51, Issue 3, pages 293–302, June 2002
How to Cite
Lees, A. K., Cullen, D. W., Sullivan, L. and Nicolson, M. J. (2002), Development of conventional and quantitative real-time PCR assays for the detection and identification of Rhizoctonia solani AG-3 in potato and soil. Plant Pathology, 51: 293–302. doi: 10.1046/j.1365-3059.2002.00712.x
- Issue published online: 29 MAY 2002
- Article first published online: 29 MAY 2002
- Accepted 7 December 2001
- Rhizoctonia solani
A specific and sensitive PCR assay was developed for the detection and identification of Rhizoctonia solani AG-3, the main causal pathogen of stem canker and black scurf of potato. A conventional primer set (Rs1F2 and Rs2R1) was designed from the nuclear ribosomal internal transcribed spacer (ITS1 and ITS2) regions of R. solani. Following PCR amplification, a 0·5-kb product was amplified from DNA of all isolates of AG-3 using primers Rs1F2 and Rs2R1. No product was amplified when DNA from isolates belonging to a range of other R. solani anastomosis groups or from a selection of other potato pathogens was tested, confirming the specificity of the primers for AG-3 only. Rhizoctonia solani AG-3 was also detected in potato tissue with varying black scurf severity, and in soil inoculated with sclerotia of R. solani to a minimum detection level of 5 × 10−4 g sclerotia/g soil. In addition, specific primers RsTqF1 (based on the Rs1F2 sequence) and RsTqR1, and a TaqMan™ fluorogenic probe RQP1, were designed to perform real-time quantitative (TaqMan) PCR. The conventional PCR and real-time PCR assays were compared and combined with direct DNA extraction from soil and a seed-baiting method to determine the most reliable method for the detection and quantification of AG-3 in both artificially inoculated field soil and naturally infested soils. It was shown that direct DNA extractions from soil could be problematic, although AG-3 was detectable using this method combined with the real-time PCR assay. The amplification of Rhizoctonia solani by seed baiting increased the sensitivity of the assay compared with direct extraction of DNA from the soil, and AG-3 was detectable in artificially inoculated and naturally infested soils when seed baiting was combined with either the conventional PCR or the real-time PCR assay. The potential for using these rapid and quantitative AG-3-specific assays to address epidemiological questions and as tools for decision-making in disease management is discussed.
Rhizoctonia solani (teleomorph: Thanatephorus cucumeris) is a soilborne pathogen that comprises several groups which are pathogenic to different host species. In potato, infection of shoots by R. solani soon after planting causes stem canker and can significantly delay plant emergence. Stolon infection can influence the number and size distribution of harvested tubers, and in severe cases of disease total yield can be reduced (Hide & Horrocks, 1994; Hide et al., 1996). Sclerotia of R. solani can develop on progeny tubers, resulting in the tuber blemish disease black scurf, thus affecting tuber quality and acting as an inoculum source for subsequent crops (Banville, 1989). Isolates of R. solani causing potato disease have been ascribed mainly to anastomosis group 3 (AG-3), although several other AG groups, including AG-4, AG-5 and AG-9, can also infect potato (Anguiz & Martin, 1989; Balali et al., 1995).
Anastomosis grouping of isolates is assigned on the basis that only hyphae of isolates belonging to the same group can fuse (as described by Parmeter et al., 1969). This technique has remained the most frequently used method for differentiating anastomosis groups of R. solani, despite being time-consuming and requiring some degree of skill, and has resulted in the recognition of 12 individual anastomosis groups (Carling et al., 1994). Rhizoctonia solani anastomosis groups can also be defined using many other methods, including assessments of cultural and pathogenic variation (Anderson, 1982; Chand & Logan, 1983; Neate & Warcup, 1985). In addition, biochemical and molecular methods, such as electrophoresis of soluble proteins (Reynolds et al., 1983), the examination of pectic zymograms (Sweetingham et al., 1986; Neate et al., 1988) and restriction fragment length polymorphism (RFLP), have been used to reveal variation among R. solani groups (Vilgalys & Gonzalez, 1990). Balali et al. (1996) generated a DNA probe specific to isolates of AG-3, but did not describe its use for diagnostic purposes.
The availability of simple and specific diagnostic assays is important for investigations of the ecology and epidemiology of plant pathogens. The ability to detect the presence of a particular pathogen in the crop or the soil, to determine threshold levels of inoculum and to investigate the relative importance of seed- and soilborne inoculum provides information on which disease management decisions can be based. The objective of this work was to develop an R. solani AG-3-specific assay to allow the unequivocal typing of AG-3 isolates, and to use the assay as a quantitative tool for the detection of the pathogen in plant tissue and soil, in order that investigations of the epidemiology of AG-3 and predictions of disease incidence could be made. The development of such an assay is described.
Materials and methods
Origin and maintenance of fungal isolates and anastomosis testing
Isolates of R. solani and other fungi used in this study, along with their sources, are given in Table 1. The anastomosis group of each isolate of R. solani isolated from tubers or obtained from another source was tested using the method of Parmeter et al. (1969), except that hyphal fusion was observed under the microscope without lactophenol/cotton blue staining.
|Code||Original code||Sourcea||Origin||Host||Anastomosis group/species|
|R58||H5-537||1||Kyushu, Japan||Paddy rice||AG 1–1 A|
|R61||RH 367||1||Hokkaido, Japan||Sugar beet||AG 1–1 B|
|R62||RH 28||1||Hokkaido, Japan||Sugar beet||AG 1–1 C|
|R7||–||–||Portsoy, Scotland||Potato||AG 2–1|
|R22||Unknown||2||Colorado, USA||Unknown||AG 2–1|
|R38||22RG2||3||the Netherlands||Sugar beet||AG 2–2|
|R50||22R02||3||Japan||Sugar beet||AG 2–2 IV|
|R1||–||–||Dundee, Scotland||Potato||AG 3|
|R2||–||–||Dundee, Scotland||Potato||AG 3|
|R3||–||–||Dundee, Scotland||Potato||AG 3|
|R4||–||–||Dundee, Scotland||Potato||AG 3|
|R5||–||–||Dundee, Scotland||Potato||AG 3|
|R8||–||–||Dundee, Scotland||Potato||AG 3|
|R9||–||–||Portsoy, Scotland||Potato||AG 3|
|R10||–||–||Ormskirk, England||Potato||AG 3|
|R12||–||–||Tayside, Scotland||Potato||AG 3|
|R13||–||–||Tayside, Scotland||Potato||AG 3|
|R14||1/97 T1||4||Northern Ireland||Potato||AG 3|
|R15||2/97 T1||4||Northern Ireland||Potato||AG 3|
|R16||4/97 T1||4||Northern Ireland||Potato||AG 3|
|R17||4/97 T3||4||Northern Ireland||Potato||AG 3|
|R18||Rz 1058||5||Christchurch, New Zealand||Potato||AG 3|
|R19||Rz Linc 1||5||Lincoln, New Zealand||Potato||AG 3|
|R20||Rz Linc 2||5||Lincoln, New Zealand||Potato||AG 3|
|R31||Rs 1||6||Rothamsted, England||Potato||AG 3|
|R32||Rs 2||6||Rothamsted, England||Potato||AG 3|
|R33||USDA 3||2||Unknown||Unknown||AG 3|
|R36||–||–||Dundee, Scotland||Potato||AG 3|
|R37||–||–||Aberdeen, Scotland||Potato||AG 3|
|R43||9729-1||7||Finistere, France||Potato stem||AG 3|
|R45||9776-8||7||Seine Maritime, France||Potato tuber||AG 3|
|R47||9845-4||7||Loiret, France||Potato tuber||AG 3|
|R81||PLB-20||1||Hokkaido, Japan||Potato||AG 3|
|R85||PF 59||1||Hokkaido, Japan||Potato||AG 3|
|R51||R3||2||Colorado State University, USA||Unknown||AG 4|
|R91||RH 164||1||Hokkaido, Japan||Sugar beet||AG 5|
|R96||AI 1-4||1||Hokkaido, Japan||Soil||AG 6|
|R97||305551||1||Kagawa, Japan||Soil||AG 7|
|R28||SH 41||2||University of Hull, England||Unknown||AG 8|
|R56||08 R01||2||Unknown||Unknown||AG 8|
|R98||TE2-4||1||Hokkaido, Japan||Soil||AG BI|
Cultures were maintained at 15°C on potato dextrose agar (PDA) (Difco, West Molesey, UK). For DNA preparations, mycelium from 4-day-old-colonies of R. solani and other fungi grown on PDA was harvested and DNA was extracted using a Puregene® Genomic DNA isolation kit (Gentra Systems, Minneapolis, MN, USA), according to the manufacturer’s instructions.
Potato tubers of various cultivars harvested from field trials at the Scottish Crop Research Institute (SCRI) in 1999, with varying levels of visual black scurf, were selected. Disease levels were as follows: (i) no visual disease symptoms (two samples having no symptoms were taken from different sites for comparison); (ii) low infection level (∼ 1 sclerotium cm−2); (iii) high infection level (∼10 sclerotia cm−2). Tubers were peeled very thinly using a scalpel and the peel ground to a fine powder in liquid nitrogen using a pestle and mortar. Duplicate 0·5 g samples of the resulting powder were transferred to 2 mL screw-cap tubes containing 1 mL SPCB buffer (120 mm sodium phosphate, 2% CTAB, 1·5 m NaCl, pH 8·0) and two 4 mm steel balls, and the samples blended at 4600 rpm for 60 s in a mini-beadbeater. DNA was then extracted according to the method described below for soil.
For the direct extraction of DNA from soil, two replicate subsamples (10 g) were taken from each soil sample. Soil was resuspended in 20 mL SPCB extraction buffer (as above), sonicated in a water bath for 15 min, then shaken (5 min) in the presence of 1·5 g sterile glass beads (1·0 mm diameter) on a flask shaker. Soil suspensions were allowed to settle for 1 min before 1·0 mL aliquots were transferred to 2 mL screw-cap tubes containing 0·2 g each of zirconia/silica beads and 1·0-mm-diameter sterile glass beads, and were then blended in a mini-beadbeater (Biospec Products, Bartlesville, OK, USA) at 5000 rpm for 60 s. Samples were centrifuged (2460 g for 5 min) and the supernatant was extracted with an equal volume of chloroform, mixed and centrifuged (11 500 g for 5 min). DNA was precipitated with 0·3 m sodium acetate (pH 5·2) and an equal volume of isopropanol for 1 h at room temperature. The DNA was pelleted by centrifugation (11 500 g for 5 min), washed in 70% ethanol, repelleted and resuspended in 75 µL TE (pH 8·0).
DNA extracts from soil were purified through a Micro Bio-Spin column (0·8 mL capacity; Bio-Rad Laboratories, Hemel Hempsted, UK) that contained water-insoluble polyvinylpolypyrrolidone (PVPP; Sigma-Aldrich, Poole, UK) using the method described by Cullen & Hirsch (1998), except that a Biofuge 13 (Heraeus, Hanau, Germany) microfuge was used.
Soil baiting procedure
Soil-bait tests were carried out following the method of Thornton et al. (1999). However, following the baiting period, the nine seeds of Chenopodium quinoa were removed from the surface of the soil samples and DNA was extracted from them using a Puregene® Genomic DNA isolation kit as described previously. In order to compare sensitivity, the effect of isolating DNA from individual seeds and bulks of all nine bait seeds was examined. In addition, the effect of the baiting period on detection of R. solani AG-3 was examined by taking samples of seeds following 48 and 96 h of incubation.
The internal transcribed spacer regions (ITS1 and ITS2) of R. solani were accessed on the GenBank and EMBL databases, and sequences of different AG groups were compared using the CLUSTAL V package (Higgins et al., 1992). Putative AG-3-specific regions were selected and AG-3-specific forward and reverse primers Rs1F2 (5′-TTGGTTGTAGCTGGTCTATTT-3′) and Rs2R1 (5′-TATCACGCTGAGTGGAACCA-3′) were designed.
The Primer Express® software (PE Applied Biosystems, Foster City, CA, USA) was used to design a forward primer RsTqF1 (5′- AAGAGTTTGGTTGTAGCTGGTCTATTT-3′) based on the original specific Rs1F2 primer and a reverse primer RsTqR1 (5′-AATTCCCCAACTGTCTCACAAGTT-3′) for use in a real-time quantitative PCR assay. The original primers were not used, as real-time PCR requires small amplicons of 50–150 bp in length to yield consistent results (PE Applied Biosystems). A TaqMan™ fluorescent probe RQP1 (5′- TTTAGGCATGTGCACACCTCCCTCTTTC-3′) was also designed and was labelled at the 5′ end with the reporter dye FAM (6-carboxy-fluorescein), while the 3′ end was modified with the quencher dye TAMRA (6-carboxy-tetramethylrhodamine) (PE Biosystems, Warrington, UK).
PCR amplification of all samples using primers Rs1F2 and Rs2R1 was based on a standard set of conditions [initial denaturation at 95°C for 2 min, followed by 35 cycles of 95°C for 45 s, 65°C for 60 s and 72°C for 90 s, with a final extension step of 72°C for 5 min, in a reaction volume of 25 µL using an MWG-Biotech Primus 96 Thermal cycler (MWG-Biotech, Ebersberg, Germany)]. The master mix contained the following components: 1 × reaction buffer (16 mm[NH4]2SO4, 67 mm Tris-HCl, pH 8·8, 0·1% Tween-20; Bioline UK Ltd, London, UK), dNTPs (Bioline) at 200 µm each, primers (MWG-Biotech) at 0·3 µm each, 5·0 mm MgCl2, 250 µg mL−1 BSA (Roche Diagnostics Ltd, Lewes, UK), 1 U Biolase Diamond (Bioline). One microlitre of DNA (representing 10–100 ng) was used as template. Aliquots (10 µL) of amplification products were electrophoresed through agarose gels (1·5% w/v) in 1 × TBE buffer (89 mm Tris-HCl, 89 mm boric acid, 2 mm EDTA, pH 8·3) and photographed under UV illumination after staining in ethidium bromide (0·5 mg L−1).
The specificity of the primer pair (Rs1F2 and Rs2R1) was tested against 30 other R. solani AG-3 and 18 non-AG-3 isolates and several other fungi associated with potato diseases (see Table 1 for sources). The ability of the primers to amplify AG-3 from DNA extracted from potato tissue and from soil inoculated with different concentrations of sclerotia was also examined using the PCR conditions described. For assessing the sensitivity of the assay, autoclaved soil was inoculated with sclerotia of R. solani AG-3 at 5 × 10−2 g sclerotia/g soil. A dilution series was prepared by the addition of 9 g sterile soil to 1 g inoculated soil to give samples with final concentrations of 5 × 10−3, 5 × 10−4 and 5 × 10−5 g sclerotia/g soil from which DNA was then extracted. All experiments were repeated for confirmation of results.
Real-time quantitative (TaqMan) PCR was performed in MicroAmp optical 96-well plates using the automated ABI Prism 7700 sequence detector (PE Applied Biosystems). The 25 µL reaction mix included 1 µL template DNA, TaqMan Universal PCR Master Mix (PE Applied Biosystems), primers RsTqF1/RsTqR1 at a final concentration of 0·2 µm per reaction, and the Taqman probe (RQP1) at 0·2 µm. The manufacturer’s recommended universal thermal cycle protocol (PE Applied Biosystems) was used for PCR amplification: stage 1 [50°C for 2 min; 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 critical threshold (Ct) values for each PCR reaction were automatically calculated and analysed by the ABI prism sequence detection systems software (version 1·6). The starting concentration of target sequence present in each reaction was calculated by comparing Ct values of unknown samples to those of standards with known amounts of R. solani AG-3 DNA; Ct values were plotted against the log of the initial concentration of R. solani DNA to produce a standard curve. The Ct value is dependent upon the input of starting copies of target and is defined as the cycle number at which a statistically significant increase in the reporter fluorescence can first be detected (i.e. exceeds the threshold).
The specificity of the real-time primer set for the detection of R. solani AG-3 was tested against 18 non-AG-3 isolates and 30 isolates of AG-3 (see Table 1 for sources). The ability of the real-time assay to amplify AG-3 DNA extracted from a dilution series of inoculated soil (5 × 10−3, 5 × 10−4 and 5 × 10−5 g sclerotia/g soil as described above) was also measured.
Comparison of methods
Direct extraction of DNA from soil followed by conventional PCR and real-time PCR amplification was compared to amplification of DNA extracted from seed following a soil-baiting period followed by conventional and real-time PCR assays.
Five samples (SCRI 1–5, see Table 2) of artificially inoculated soil were obtained from a disease nursery at SCRI that had been inoculated with several isolates of R. solani AG-3 in April 2000 and on which a potato crop had subsequently been grown. Samples of soil were taken at random from the field in October 2000 and two replicate subsamples (reps 1 and 2) were taken from each of these five samples.
|Code||Location||Sampling date||Soil type||Rhizoctonia history||Last potato crop||2000 crop|
|SCRI 1||SCRI||October 2000||Medium loam||Black scurf||2000||Potato|
|SCRI 2||SCRI||October 2000||Medium loam||Black scurf||2000||Potato|
|SCRI 3||SCRI||October 2000||Medium loam||Black scurf||2000||Potato|
|SCRI 4||SCRI||October 2000||Medium loam||Black scurf||2000||Potato|
|SCRI 5||SCRI||October 2000||Medium loam||Black scurf||2000||Potato|
|M1||E. Suffolk||February 2001||Loamy sand||None||Pre-1990||Linseed|
|M2||E. Suffolk||February 2001||Medium loam||None||1996||Wheat|
|M3||E. Suffolk||February 2001||Loamy sand||None||Pre-1980||Wheat|
|M4||E. Suffolk||February 2001||Sandy loam||None||Pre-1990||Wheat|
|M5||Essex||February 2001||Medium loam||None||None||Wheat|
|M6||E. Suffolk||February 2001||Loamy sand||None||Pre-1980||Wheat|
|M7||W. Suffolk||February 2001||Loamy sand||None||None||Wheat|
|M8||W. Suffolk||February 2001||Medium loam||None||None||Wheat|
|M9||E. Suffolk||February 2001||Loamy sand||Black scurf 1996||1996||Cauliflower|
|M10||E. Suffolk||February 2001||Sandy loam||Black scurf 1996||1996||Cauliflower|
Naturally infested soils (M 1–10) were obtained from 10 field sites in East Anglia, UK, that had a varied cropping history and previous incidence of black scurf (see Table 2). Samples (a maximum of 500 g of soil) were taken in February 2001 and consisted of 20–30 individual subsamples taken from across a field. Two replicate subsamples (reps 1 and 2) were taken from each of these 10 samples.
Anastomosis-group-3-specific PCR and real-time PCR primers
Primers Rs1F2 and Rs2R1 were tested against DNA extracted from 30 isolates of R. solani AG-3 from diverse locations, and from 18 isolates belonging to eight other anastomosis groups (Table 1, Fig. 1). Following PCR, a single fragment of 0·5 kb was amplified from the 30 isolates of AG-3 only. The specificity of the primer set was also confirmed by the absence of product when total genomic DNA extracted from a range of fungal pathogens commonly found infecting potato was tested (i.e. Polyscytalum pustulans, Phytophthora infestans, Fusarium sulphureum, F. coeruleum, Helminthosporium solani, Colletotrichum coccodes, Phoma foveata and Spongospora subterranea).
The PCR primers RsTqF1 and RsTqR1 were also specific to AG-3 in a real-time assay. The Ct values of the 18 non-AG-3 samples showed that DNA from these samples had not amplified, whereas the Ct values of the 30 AG-3 samples were in the range 23–29, equivalent to 1·2 × 104−4·7 × 107 fg DNA µL−1, indicating that the DNA in these samples had been amplified under the conditions used. The correlation coefficient of the standard curve used for the calculations was 0·97 and the minimum amount of DNA detected in the controls was 20 fg µL−1.
Detection of R. solani AG-3 in soil and plant tissue
The DNA extracted from a dilution series of soil seeded with 10−2−10−4 g sclerotia/g soil was amplified both in a conventional PCR assay and a real-time PCR assay using primer pairs Rs1F2/Rs2R1 and RsTqF1/RsTqR1, respectively. In the conventional PCR assay, a single fragment of 0·5 kb was amplified from soil with inoculum concentrations of 10−2, 10−3 and 10−4 g sclerotia/g soil (Fig. 2). No signal was detected when DNA extracted from autoclaved soil was amplified. In the real-time assay, the amount of R. solani AG-3 DNA detected was shown to decrease in relation to soil dilution and none was detected in the sterile soil control samples (Fig. 2). There was variation in the amount of AG-3 DNA detected between duplicate samples in the 5 × 10−3 treatment (Fig. 2).
In addition, amplification of a product of the expected size was also detected from DNA extracted from potato tissue with (i) a low level (1 sclerotium cm−2) and (ii) a high level (10 sclerotia cm−2) of black scurf symptoms using the conventional PCR assay. When DNA from tubers having no symptoms was tested, a signal was obtained in both samples from a first site, but in neither of the samples taken from a second site (Fig. 3).
PCR and real-time PCR amplification of AG-3 from soil using direct DNA extraction and soil-baiting techniques
Direct soil DNA extraction
When DNA extracted directly from artificially inoculated soil (SCRI 1–5, reps 1 and 2) was tested using primers Rs1F2 and Rs2R1, faint amplification products were visible only in the SCRI 2 sample (both reps 1 and 2). Following real-time PCR amplification of six of these samples (SCRI 1–3, reps 1 and 2), only the SCRI 2 (rep 1) sample gave a positive result (Table 3).
|Soil||Rep||Direct soil DNA extraction||Seed DNA extractions|
|PCR||Real-time PCR (fg DNA µL−1)||PCR||Real-time PCR (fg DNA µL−1)|
|48 h||72 h||48 h||72 h|
|SCRI 1||1||−||−||+||+||2·9 × 105||1·7 × 105|
|2||−||−||+||+||3·0 × 105||1·5 × 105|
|3||nt||nt||+||+||7·9 × 104||1·6 × 104|
|4||nt||nt||−||+||1·4 × 104||5·6 × 104|
|5||nt||nt||+||+||7·1 × 103||1·2 × 105|
|SCRI 2||1||+||3·4 × 104||+||+||1·0 × 104||8·9 × 104|
|2||+||−||+||+||4·7 × 104||1·1 × 105|
|3||nt||nt||−||−||1·3 × 104||4·0 × 104|
|4||nt||nt||+||+||2·3 × 104||2·4 × 105|
|5||nt||nt||+||+||3·0 × 103||2·0 × 106|
|SCRI 3||1||−||−||−||−||2·4 × 104||6·2 × 103|
|2||−||−||−||−||3·9 × 104||1·2 × 103|
|3||nt||nt||−||−||nt||1·4 × 104|
|4||nt||nt||−||−||6·6 × 103||3·2 × 103|
|5||nt||nt||+||−||5·0 × 104||4·9 × 104|
In naturally infested soils (M1–10), PCR amplification of DNA extracted directly form soil failed to detect R. solani AG-3 in any of the 10 samples with two replicates tested (Table 4). Real-time PCR amplification of the same samples was able to detect AG-3 in 14 of the 20 samples. However, levels of detection were very low and occurred at the limits of detection of the assay (0–168 fg DNA µL−1) (Table 4).
|Soil||Rep||Direct soil DNA extraction||Seed DNA extraction|
|PCR||Real-time PCR (fg DNA µL−1)||PCR||Real-time PCR (fg DNA µL−1)|
|M1||1||−||5·7||+||1·2 × 103|
|2||−||12·7||+||2·2 × 103|
|M2||1||−||0||+||2·7 × 102|
|2||−||23·5||+||1·2 × 101|
|M3||1||−||58·1||+||2·4 × 103|
|2||−||11·2||+||1·3 × 103|
|M4||1||−||168·6||+||1·9 × 103|
|2||−||0||+||2·9 × 103|
|M5||1||−||30·4||+||1·4 × 103|
|2||−||11·8||+||2·3 × 103|
|M6||1||−||0||+||1·3 × 103|
|2||−||0||+||8·5 × 102|
|M7||1||−||7·9||+||8·8 × 102|
|2||−||36·3||+||1·5 × 103|
|M8||1||−||20·4||+||1·0 × 103|
|2||−||127·6||+||3·0 × 103|
|M9||1||−||0||+||2·2 × 103|
|2||−||59·9||+||2·3 × 103|
|M10||1||−||0||+||2·0 × 103|
|2||−||11·9||+||1·3 × 103|
Following conventional PCR amplification of DNA extracted from seed used to bait the artificially inoculated sample SCRI 1, PCR products were visible in four out of five replicates (reps 1, 2, 3 and 5) after 48 h incubation, and in all samples after 72 h incubation (see Table 3). Results for sample SCRI 2 were consistent over both tests, with four out of five replicates showing amplification product following PCR, while in sample SCRI 3, a signal was only detectable in rep 5 (bulk of nine seeds) on one occasion, indicating that levels of AG-3 were lower in that sample.
In contrast to conventional PCR, it was shown that R. solani AG-3 was detectable in all replicates [sample SCRI 3 (rep 3), 48 h is missing] by real-time PCR. There was no significant difference (P < 0·05) in either PCR in the amount of AG-3 DNA present in the samples, as calculated from the Ct value (correlation coefficient of standard curve = 0·99) following 48 or 72 h incubation, or between reps (where reps 1–4 were individual seed extractions and rep 5 was a bulk DNA extraction from nine seeds). However, DNA concentrations were generally higher in all samples following 72 h incubation, and samples SCRI 1 and 2 had on average 10–100 times as much AG-3 DNA as sample SCRI 3.
In the case of the naturally infested soil samples M1–10, conventional PCR amplification of the DNA extracted from bulk seed samples produced signals in all replicates of all soils (Table 3). Using real-time PCR, it was shown that, overall, there was no statistically significant difference (P < 0·05) in the amount of AG-3 DNA detectable in each soil sample. However, both replicates of soil sample M2 showed a 10-fold lower concentration of AG-3 target DNA than the other nine samples. Levels of AG-3 contamination were, as may be expected, lower in the naturally infested soils (M1–10) than in the artificially inoculated soil samples (SCRI 1–3).
Rhizoctonia solani is an important soilborne pathogen and AG-3 is the group predominantly responsible for causing stem canker and black scurf of potato. Previously described methods (Anderson, 1982; Sweetingham et al., 1986; Neate et al., 1988; Vilgalys & Gonzalez, 1990) are useful for the discrimination of anastomosis groups, but are unable to specifically detect particular groups in the environment.
Rhizoctonia solani AG-3 presents several problems for the development of a diagnostic assay. The existence of many anastomosis groups of R. solani, some closely related, potentially makes the identification of anastomosis-group-specific regions of the R. solani genome difficult, and the heterogeneity in R. solani AG-3 noted by several authors (Jabaji-Hare et al., 1990; Duncan et al., 1993; Balali et al., 1996; Bounou et al., 1999) has implications for the location of a universally specific region.
The use of the ITS regions 1 and 2 for the production of specific primers has proved to be a successful strategy for developing diagnostic assays for many pathogenic fungi, including the soilborne potato pathogens Spongospora subterranea (Bell et al., 1999) and Helminthosporium solani (Cullen et al., 2001), and Colletotrichum coccodes (Cullen et al., 2002) and also for the differentiation of Rhizoctonia oryzae from R. solani AG-8 (Mazzola et al., 1996). In the present work, ITS sequence information was used to identify regions specific to AG-3, and conventional PCR primers (Rs1F2 and Rs2R1) and real-time PCR primers (RsTqF1 and RsTqR1) and probe (RQP1) specific to this anastomosis group were designed.
No significant levels of similarity were revealed when comparisons between each primer and probe sequence to DNA database sequences of other fungi and bacteria were made. In addition, the primer pair Rs1F2/Rs2R1 designed in this study was used in PCR to test DNA from isolates of R. solani AG-3 from the UK, mainland Europe and other countries, with amplification of the expected product being achieved in all cases, thus demonstrating the specific nature of the assay.
Two other assays for the detection of R. solani were published recently. Thornton et al. (1999) described a method for the detection of R. solani in glasshouse soils, using a combined baiting and double monoclonal antibody ELISA. This method has the advantage of detecting only live propagules of the fungus, and the inclusion of a baiting step allowed accumulation of the fungus to a level detectable by ELISA. However, unlike the PCR assays developed in the current work, the method is not AG-specific and therefore identification of the AG group by conventional means would still be required for epidemiological studies.
Similarly, Bounou et al. (1999) developed a PCR method for the specific detection of AG-3 isolates, based on the differentiation of AG groups by an RAPD assay together with detection using a PCR-based restriction-mapping technique. This method, although specific for AG-3, requires enzyme restriction of PCR products for the differentiation of the AG-3 group. The authors were able to detect R. solani AG-3 in soil and plant tissue using this method, but did not measure the sensitivity of the detection.
In the present work, ITS-based PCR primers (Rs1F2 and Rs2R1) specific to AG-3 were generated and it was demonstrated that they may be used in a simple assay to detect the pathogen in plant material in the absence of visual black scurf disease symptoms and in soil at a level of 5 × 10−4 g sclerotia/g soil. This primer pair was designed to have amplification requirements similar to those of primer sets developed for the detection of other seed- and soilborne potato pathogens (Cullen et al., 2000) in order that the assays may be integrated in the future to provide a multiplex detection tool.
In addition, a quantitative real-time PCR assay for the sensitive and specific detection of R. solani AG-3 has been described here. Variation between the amount of AG-3 DNA (fg µL−1) detected in replicate samples of artificially inoculated soil using the real-time PCR assay is likely to be due to variation in the amount of R. solani contained in those samples at the DNA extraction stage. Such variation could be reduced by taking more, or larger, samples for DNA extraction. However, it should be noted that some variation at the fg µL−1 level is expected and may not necessarily relate to epidemiologically significant differences of the pathogen in field soils.
In the past, epidemiological studies have been hindered by the lack of such a quantitative assay. As noted by Anderson (1982), one of the factors preventing disease predictions being made has been the absence of AG determinations in R. solani populations. In addition, assessment of yield losses caused by R. solani infection of potato plants in the field would be facilitated by the use of disease-tested potato stocks. Similarly, Neate et al. (1988) discussed the importance of AG identification for the determination of soil inoculum potential.
However, in addition to pathogen detection and identification of anastomosis groups, the impact of interactions between many other factors on disease incidence and severity must be considered before predictions of disease can be made. Such factors include variation in pathogenicity between isolates, the spatial distribution of inoculum and the physical and biological environment in which the pathogen exists. The assays described in this work allow disease-causing isolates of R. solani AG-3 to be detected and identified and will facilitate the further research that is needed to understand the epidemiology of the diseases and subsequently to provide a predictive tool for disease incidence or severity.
In this work, the conventional and quantitative real-time PCR assays were combined with the baiting method of Thornton et al. (1999) in order to ascertain the most sensitive and reliable methods for the detection of R. solani AG-3 in soil. The DNA extraction methods and levels of sensitivity of the conventional and real-time assays proved suitable for the detection of R. solani AG-3 in plant and tuber tissue. However, the role of soil inoculum in disease incidence represents an important, but little understood, factor in the epidemiology of the disease, and it was therefore necessary to validate the quantitative assay in naturally infested soils for potential use in epidemiological studies.
It was found that the PCR amplification of DNA extracted directly from soil samples could be problematic, as has been documented in the past (Cullen & Hirsch, 1998). Although signal could be detected using template DNA extracted directly from soil in both the conventional and real-time assays, results were variable in these studies. This variability may be due to the patchy distribution of the pathogen in soil samples and/or the presence of PCR-inhibitory compounds in the soil. It is possible that the use of commercial soil-extraction kits could improve the levels of amplifiable DNA extracted from the soils, as could further attempts to purify the DNA extracts.
Signal was detected in DNA extracted directly from the soil samples M1–10 using real-time PCR, but not using conventional PCR. This is unsurprising, as real-time PCR is more sensitive than conventional PCR. The sensitivity of the conventional PCR assay could perhaps be improved by a second-round nested PCR approach.
Levels of R. solani AG-3 found in naturally infested soils may be quite low and have a patchy distribution, and it is therefore possible that the level of AG-3 DNA in these soils (M1–10) is at the limit of detection by direct DNA extraction and amplification, or that sampling strategies must be addressed in greater detail to ensure representative samples are taken.
In order to increase the sensitivity of the assay for the detection of AG-3 in naturally infested soils and to avoid any possible problems with soil-DNA extraction, the bait test of Thornton et al. (1999) was used in combination with the AG-3-specific and quantitative real-time PCR assay. The bait test allows biological amplification of the fungal populations present in the soil to detectable levels, whereas the real-time assay allows unequivocal and quantitative detection of the pathogen of interest.
It was shown that biological amplification of the fungus in bait seed provided a good way of increasing the target DNA and avoided the need for direct extraction from soil. Soil samples that had given negative results for the presence of AG-3 with both conventional PCR and real-time PCR when tested using direct DNA extraction produced signals using the combined bait/real-time assay. There appeared to be little difference in detection levels for different-length baiting periods and it was found that extraction from a bulk of nine seeds was as efficient as extraction of DNA from individual seeds. However, DNA extraction and amplification from individual seeds proved successful and could have implications for studies into the distribution of the fungus in soil samples.
It is more difficult to ascertain a direct relationship between the amount of DNA present in the soil and the amount detectable by the assay given the inclusion of a biological amplification step. In addition, it is not known whether the levels of AG-3 detected in the naturally infested soils were representative of levels likely to cause disease. It is intended that the disease incidence of black scurf and stem canker will be measured in potato crops grown on the naturally infested sites M1–10 in 2001 in an attempt to relate PCR amplification to disease incidence.
Given the development of these specific, sensitive and quantitative assays for the detection of R. solani AG-3, further work can now be carried out to study the epidemiology of the diseases it causes and to answer questions concerning: (i) the distribution of the fungus in the soil; (ii) suitable sampling strategies for diagnostic tests; and (iii) the relative roles of seed- and soilborne inocula in causing disease.
AKL, MJN and LS thank the Scottish Executive Environment and Rural Affairs Department (SEERAD) for financial support. DWC was supported by the Department for Environment, Food and Rural Affairs (DEFRA). The authors also thank Sandie Linton for technical assistance, Alex Hilton for anastomosis testing, Arthur McCorquodale of MBM for supply of soil samples and colleagues listed in Table 1 for generous provision of isolates.
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