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Keywords:

  • bioassay;
  • black dot;
  • microplants;
  • minitubers;
  • PCR;
  • potato;
  • silver scurf;
  • soil inoculum

Abstract

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

The sensitivity of a bioassay in detecting soil inoculum of Colletotrichum coccodes and Helminthosporium solani was examined using potato minitubers and microplants. Tests were conducted on soils which were collected from fields in which the interval after a previous potato crop differed, and which were also artificially infested with conidia or microsclerotia. For C. coccodes, determining plant infection based on the occurrence of infected roots after 9–12 weeks was a sensitive method for detecting and quantifying the amount of inoculum in soil. Infestations of less than 0·4 microsclerotia per g soil were detected in artificially infested soils. A semiselective medium, developed for isolating C. gloeosporioides from pepper, detected soil infestations by C. coccodes as low as nine conidia or one microsclerotium per g soil in artificially infested soil. For H. solani, infection on minitubers was a sensitive measure, with soil inoculum of fewer than 10 conidia per g soil being detected. Soil infestation could be quantified by assessing the percentage surface area of minitubers covered by sporulating lesions, which was strongly related to the amount of soil infestation. The results of these bioassay tests were compared with published results for real-time quantitative PCR assays on the same soils. The two methods were in good agreement in artificially infested soils, but the bioassay appeared to be more sensitive with naturally infested soils.


Introduction

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

The increase in the sale of potatoes as a washed and prepacked product in the UK (Read et al., 1995) and elsewhere (Andrivon et al., 1997) has meant that skin blemish diseases have become economically important, as they affect appearance and therefore quality. Two of the most common diseases are black dot caused by Colletotrichum coccodes and silver scurf caused by Helminthosporium solani (Read et al., 1995). Both pathogens cause a superficial blackish or silvery discoloration of the tuber skin, producing an unattractive appearance and an increase in skin permeability which can result in water loss and shrivelling of the tubers (Read & Hide, 1984). Errampalli et al. (2001) also reported that processing tubers affected by silver scurf had thicker skins which peeled less satisfactorily and produced chips with unacceptable burnt edges. On seed tubers, Rudkiewicz (1986) concluded that severe silver scurf infection delayed plant emergence, increased blanking and decreased yield, but Read & Hide (1984) found that silver scurf had only a transient effect on plant growth.

Jellis & Taylor (1974) considered that the primary source of inoculum in the spread of H. solani was the infected mother tuber, whereas the spread of C. coccodes occurred from both infected seed tuber and soil. In South Africa, Denner et al. (1998) concluded that the contribution of soil inoculum of C. coccodes to the development of black dot was twice as great as that of seedborne inoculum. Read et al. (1995) indicated that there was no relationship between the amount of black dot on seed tubers and that developing on the daughter tubers. With silver scurf, the relationship between disease on seed and daughter tuber has commonly been found to be inverse, particularly when silver scurf was very severe on the seed (Adams & Hide, 1980; Lennard, 1980; Read & Hide, 1984). Sporulation from large, old lesions is potentially very limited compared with that from small lesions (Jellis & Taylor, 1977; Read & Hide, 1984). However, Firman & Allen (1995) suggested that the transmission of H. solani from seed tubers could be minimized if the severity of silver scurf on the seed was very low. They considered that very small lesions might not produce large amounts of conidia before tuber decay halted the growth, and hence the sporulation of H. solani. Although Jellis & Taylor (1977) did not detect H. solani in soils collected 1 year after an infected potato crop, Kamara & Huguelet (1972) and Mérida & Loria (1994) reported that the fungus could survive over winter in North America. Daughter tubers derived from healthy tubers became infected by H. solani when grown in fields at a range of intervals after a potato crop (Mérida & Loria, 1994; Bains et al., 1996). Furthermore, Carnegie et al. (1996) and Hall (1996) showed that H. solani could survive in dry soils in potato stores and this inoculum could contaminate healthy tubers.

A collaborative project to examine the role of soil inoculum in the development of silver scurf and black dot was undertaken by ADAS Consulting Ltd, Scottish Crop Research Institute (SCRI) and the Scottish Agricultural Science Agency (SASA), particularly in relation to the impact of crop rotation and husbandry practices on inoculum surviving in field soil. A key part of this work was to evaluate a range of techniques for detecting and quantifying propagules of C. coccodes and H. solani present in field soils. Selective media for use in detecting C. coccodes (Farley, 1972) and H. solani (Singh, 1972) have been reported, but preliminary tests revealed that these were relatively insensitive and variable in their performance. However, a semiselective medium (CGPI) developed by Manandhar et al. (1995) for isolating C. gloeosporioides from pepper plants was found to have potential for detecting C. coccodes, and further tests were conducted in conjunction with the bioassays. In order to detect a range of tuber pathogens (including C. coccodes and H. solani) in dry soil from potato stores, Carnegie et al. (1996) and Hall (1996) used minitubers planted in test soil layered with vermiculite in plastic pots in a glasshouse. A series of experiments was therefore conducted to assess the sensitivity of a modification of this bioassay method and the CGPI medium in detecting C. coccodes or H. solani in naturally or artificially infested soils.

Materials and methods

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

Inoculum and test soils

Three isolates of C. coccodes and H. solani recently recovered from potatoes were grown on potato dextrose agar (PDA) (Oxoid, Basingstoke, UK) at room temperature (18–22°C) for 2 and 4 weeks, respectively. Suspensions of conidia or microsclerotia of C. coccodes and conidia of H. solani were prepared by pouring water-suspended scrapings from the surface of colonies on to coarse muslin. Microsclerotia trapped in the muslin were resuspended in sterile distilled water. The concentration was then estimated by dispensing 10 100 µL drops of the suspension onto glass slides and counting, under a microscope, the number of microsclerotia in each drop. The number of conidia of H. solani or C. coccodes in the filtrate was counted using a haemocytometer. The suspensions of propagules were then adjusted to the desired concentration.

Three soils were used in the tests. Soils A and B (both clay loams) were sampled from fields at SASA's Gogarbank Farm, Midlothian, in which a potato crop had not been grown for 6 months and 10 years, respectively. Soil C (silty clay loam) was sampled from a field at Causewood Farm, West Lothian, in which potatoes had not been grown for at least 45 years. Each soil was air-dried in a glasshouse for 24 h and passed through a sterilized coarse sieve to remove stones and plant debris. Samples of soils B and C were infested by adding the suspension of propagules to give the highest soil concentration in any series (Table 1). After thoroughly mixing the infested soil for 30 min, a subsample was mixed with unamended soil to give the next soil infestation. This process of serial dilution was repeated to create the series of soil infestations for each experiment (Table 1). In addition, to assess the efficacy of the test method in detecting natural soil inoculum, soils A (6 months after potatoes) and B (10 years after potatoes) were mixed in a range of proportions (Table 1).

Table 1.  Details of bioassay test soils infested with fungal propagules of Helminthosporium solani or Colletotrichum coccodes, or prepared by mixing soils from fields sampled at various intervals after a previous potato crop
Number of conidia of H. solani per g soilNumber of microsclerotia of C. coccodes per g soilProportion, by weight, of soil A : soil B in test soil
Soil BSoil CSoil BSoil C
Exp 1Exp 2
  1. Soil A, 6 months after potatoes; soil B, 10 years after potatoes; soil C, c. 45 years after potatoes. potatoes.

10001000100010·0010·00   100 : 0
200 300 300 2·00 3·0020·0 : 80·0
40 100 100 0·40 1·00 4·0 : 96·0
8  30  30 0·08 0·30 0·8 : 99·2
2  10  10 0·02 0·10 0·2 : 99·8
0   0   0 0 0·03 0·01 0   0 : 100

Bioassay tests

Minitubers of cultivars Maris Piper and Desiree were obtained from a commercial producer of Pre-basic TC seed potatoes (GenTech Propagation Ltd, Dundee, UK). Those of cv. Desirée were used only in the experiments to detect C. coccodes in artificially infested soil B and in the mixtures of soils A and B. Samples of 50 tubers from each purchased lot were tested for the presence of H. solani and C. coccodes as described later.

A 3 cm layer of coarse-grade vermiculite was placed in the bottom of a plastic pot 15 cm in diameter. This was covered with a layer of test soil c. 5 cm deep (450–500 g) into which a minituber was planted. Finally, another 3 cm layer of vermiculite was placed on top of the soil. Each experiment was laid out in a glasshouse in a randomized complete block design with 12 replicate pots. The pots were watered twice weekly. Initially, the soil moisture was maintained at c. 80% field capacity, as measured by weighing the pots, until the plants had emerged; thereafter, soils were maintained at 95 or 90% field capacity to detect C. coccodes and H. solani, respectively. In those experiments in which there were two assessment times, six replicate pots selected at random were harvested on each occasion.

Using artificially infested soil C, the effectiveness of microplants in detecting C. coccodes was compared with that of plants from minitubers. Microplants of cv. Maris Piper from SASA's seed potato nuclear stock collection were grown in vitro for 3 weeks to allow roots to develop. The rooted microplants were transplanted in 15 cm diameter pots containing a 1 : 1 mixture of test soil and perlite. Each pot contained c. 500 g test soil and six microplants. The experiment was laid out in a randomized complete block design with three replicate pots. One microplant was harvested from each pot at intervals of 2, 4, 6, 8 and 9 weeks after planting.

Disease and pathogen assessment

Minitubers and daughter tubers were placed in sealed containers and incubated at 18–22°C and c. 95% relative humidity for 7 days before examining the tubers for setose microsclerotia of C. coccodes or the characteristic conidiophores of H. solani. The proportion of infected daughter tubers was recorded for each replicate, except in the experiment using soil C infested with C. coccodes, in which plants were recorded as infected if at least one daughter tuber was infected. In addition, for H. solani the number of lesions was counted and/or the percentage of the surface area covered by sporulating lesions was estimated for each tuber.

Underground stems and roots were examined for disease symptoms. One diseased root, or the longest healthy root and one stem, were selected from each plant. The plant pieces were dipped in 96% ethanol for 0·5 min, rinsed in sterile distilled water, cut into 10 pieces and placed on PDA containing 200 mg streptomycin L−1. After incubation for 7 days at 18–22°C, the number of plants with infected stems or roots was determined by recording the presence of colonies of C. coccodes and H. solani developing from the individual plant pieces. The assessment was similar for the microplants, except that stems and roots were treated as one entity.

Statistical analysis

As the data were derived from relatively small samples (percentages based on six plants), the analyses were conducted using a number of nonparametric and exact tests (Conover, 1980). Spearman's rank correlation coefficient (rs) was used to evaluate the relationships between the various disease assessments and the number of propagules in the test soils. The incidence of disease within the range of soil infestations was compared with that for the unamended soil using the one-sided Fisher's exact test, and McNemar's test was used to compare the incidence of disease on the plant parts.

Detection of C. coccodes using a semiselective medium

For the first test, another series of infested soils was prepared as described previously by adding conidia to soil B. The concentrations of conidia per g soil were 870, 260, 87, 26, 9 and 0 (unamended soil B). A second test was conducted in conjunction with a bioassay test, using soil C infested with a range of concentrations of microsclerotia (Table 1). In the first test, a suspension of 10 g of each test soil in 500 mL sterile distilled water was shaken vigorously by hand for 1 min. In the second experiment, 10 g soil was suspended and shaken in 100 mL water. Aliquots of 0·5 mL were then spread on the surface of Petri dishes containing C. gloeosporioides pepper isolate (CGPI) medium (Manandhar et al., 1995). There were 20 replicate dishes for each soil. After incubation for 14 days at 18–22°C, the number of colonies of C. coccodes was recorded on each dish.

Comparison of detection by PCR and bioassay

Samples of unamended and artificially infested soils were also provided to SCRI, where detection tests were conducted using real-time quantitative (TaqMan) PCR (Cullen et al., 2001; Cullen et al., 2002). Before testing, soils infested with either of the pathogens were mixed to give soils containing both. Therefore the series of soil inoculum concentrations in these tests did not match those in the bioassay tests; however the results for the detection of the two pathogens by bioassay were compared with those obtained by PCR using the concentrations of soil infestation, which were identical or very similar in each test soil series.

Results

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

Neither C. coccodes nor H. solani was detected in the test samples of minitubers.

Bioassay and C. coccodes

Symptoms of infection by C. coccodes were observed on minitubers, and on stems, roots and daughter tubers of plants derived from minitubers. Six weeks after planting minitubers in soil B infested with microsclerotia of C. coccodes, the incidence of infection from infested soils differed significantly (P < 0·05) from that in unamended soil only for stems and roots grown in soil containing 10 microsclerotia per g soil (Fig. 1). At two microsclerotia per g soil, infection was detected on minitubers, stems and roots but none was found on daughter tubers. At 0·4 microsclerotia per g soil, 33% of plants had infected roots but no infection was found on any of the other plant parts. Although a low incidence of infection was recorded on roots of plants grown in unamended soil B, C. coccodes was not detected on plants grown in soils with the two lowest concentrations of inoculum. Twelve weeks after planting the minitubers had decayed so that it was not possible to assess them for black dot. The incidence of plant infection at all concentrations of soil infestation was greater (P < 0·05) for roots than for stems (Fig. 2). The incidence of plant infection on stems and roots and of infection on daughter tubers increased significantly (P < 0·05) with the degree of soil infestation, although the correlation was highest with roots (rs = 0·941, n = 6). The proportion of plants with infected stems, roots and daughter tubers differed significantly (P < 0·05) from that recorded for unamended soil B only at concentrations of ≥ 0·4 microsclerotia per g soil. Infection of stems, roots and daughter tubers occurred on plants grown in unamended soil B.

image

Figure 1. Frequency of plant infection by Colletotrichum coccodes, 6 weeks after planting, on plants grown from minitubers in soil B (10 years after potatoes) infested with differing concentrations of microsclerotia.

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image

Figure 2. Frequency of infection by Colletotrichum coccodes, 12 weeks after planting, on stems, roots and daughter tubers of plants grown from minitubers in soil B (10 years after potatoes) infested with differing concentrations of microsclerotia. Minitubers had decayed during this time period.

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With infested soil C, infection by C. coccodes was recorded after 9 weeks on plants grown from minitubers at the five highest concentrations; however, too many minitubers had decayed to give a meaningful assessment (Fig. 3). At the two highest concentrations of soil infestation, infection occurred on all plant parts. Infection on the roots of plants derived from minitubers also occurred at 0·1, 0·3 and one microsclerotia per g soil, but infection was not recorded on daughter tubers, and was recorded only once on stems. Colletotrichum coccodes was not found on plants in soils with ≤ 0·03 microsclerotia per g soil. The mean incidence of infection on stems, roots and daughter tubers increased significantly (P < 0·05) with the degree of soil infestation, and the correlation was highest for roots (rs = 0·98, n = 8). With the microplants, infection was detected 2 weeks after planting only on two plants at the greatest inoculum concentration (10 microsclerotia per g soil), and after 4 weeks 29% of plants were infected. The incidence of infection on microplants examined 6, 8 and 9 weeks after planting was similar, with an overall mean of 36%. The detection of C. coccodes in soil using microplants was similar to that for plants grown from minitubers, except that one microplant from soil containing 0·03 microsclerotia per g soil was found to be infected, and the roots of one plant grown from a minituber in soil containing 0·1 microsclerotia per g soil were infected (Fig. 3). The incidence of microplant infection increased (rs = 0·87, n = 8) with the amount of soil infestation. Colletotrichum coccodes was not detected on any plants grown in unamended soil C.

image

Figure 3. Frequency of plant infection by Colletotrichum coccodes, 9 weeks after planting, on stems, roots and tubers of plants from minitubers and on microplants (mean of assessments made 6, 8 and 9 weeks after planting) grown in soil C (c. 45 years after potatoes) infested with differing concentrations of microsclerotia. Minitubers had decayed during this period.

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Infection by C. coccodes was detected on stems, roots and daughter tubers of plants grown in a mixture of unamended soil B (10 years after a potato crop) and soil A (6 months after a potato crop) (Fig. 4). The incidence of infection was significantly (P < 0·05) greater on plants grown in unamended soil A than in soil B or any of the mixtures, but was not correlated with the proportion of soil A in the growing medium.

image

Figure 4. The development of Colletotrichum coccodes, 12 weeks after planting, on plants grown from minitubers in soils A (6 months after potatoes) and B (10 years after potatoes) mixed in differing proportions. Minitubers had decayed during this period.

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Detection of C. coccodes using CGPI medium

Colletotrichum coccodes on CGPI medium produced sparse mycelial growth, with acervuli bearing setae developing most profusely at the centre and edge of a colony. When the rate of soil dilution was low, 1 : 10 as in the test with infested soil C, the amount of other fungal growth was relatively high and the development of colonies of C. coccodes was restricted. However, the fungus could still be distinguished on the reverse of the plate by the presence of black mycelium with microsclerotial initials. In the first test, C. coccodes was not detected in unamended soil B, but was recovered from all soils that had been artificially infested with conidia (Table 2). There was a close relationship between number of colonies per g soil detected using CGPI medium, and the estimated number of conidia in the soil. In test 2 with soil C, the soil populations were much lower and C. coccodes was not detected in soils containing less than one microsclerotium per g soil.

Table 2.  Populations of Collectotrichum coccodes in soils artificially infested with conidia or microsclerotia as estimated by plating diluted suspensions on a semiselective medium
Soil BSoil C
Estimated number of conidia per g soilMean number of colonies per g soilEstimated number of microsclerotia per g soilMean number of colonies per g soil
870789 ± 631010
260210 ± 32 3 5
87 42 ± 12 1 1
26 53 ± 18 0·3 0
9  5 ± 5 0·1 0
0  0 0·03 0
   0·01 0
   0 0

Bioassay and H. solani

Helminthosporium solani was not recovered from the stem or roots of plants grown from minitubers planted in any of the soils to which conidia had been added. However, infection on incubated minitubers and daughter tubers developed as black circles of conidiophores of H. solani. Six weeks after planting, all minitubers grown in soil B infested with 40, 200 or 1000 conidia per g soil (experiment 1) were infected by H. solani (Table 3). The number of colonies could not be counted on minitubers from the soil containing the most conidia, as the colonies had coalesced. Incidence of tuber infection and mean number of colonies on daughter tubers 12 weeks after planting were lower than on minitubers (Table 3). Overall on minitubers and daughter tubers, the incidence of infection and the number of silver scurf colonies increased significantly (P < 0·05) with the level of soil infestation. With amended soil, the incidence of infection on minitubers differed significantly (P < 0·05) from that with unamended soil only at infestations of 40 conidia per g soil or more, but on daughter tubers this difference occurred only at 1000 conidia per g soil. In the second experiment with soil B, all minitubers from the amended soils were infected and 50% from the unamended soil were infected (Table 4). The surface area affected by silver scurf on minitubers increased significantly (rs = 1·0, n = 6) with the concentration of H. solani propagules in the soil.

Table 3.  Development of silver scurf on minitubers and daughter tubers 6 or 12 weeks after planting in soil B infested with differing concentrations of conidia of Helminthosporium solani (experiment 1)
Number of conidia per g soilPercentage of tubers infectedMean number of lesions of H. solani per tuber
Minitubers after 6 weeks Daughter tubers after 12 weeksMinitubers after 6 weeks Daughter tubers after 12 weeks
  1. ND, colonies were too numerous to permit an accurate count.

100010050ND1·57
200100238·00·32
40100152·70·23
8 50 00·50
2 33 70·30·07
0 33 00·50
Table 4.  Development of silver scurf on minitubers 6 weeks after planting in soil B infested with differing concentrations of conidia of Helminthosporium solani (experiment 2)
Number of conidia per g soilPercentage of tubers infectedPercentage surface area affected
100010084·2
30010073·3
10010026·0
3010010·0
10100 0·6
0 50 0·3

The incidence of infection on minitubers grown in artificially infested soil C was broadly similar after 3 and 6 weeks, but the surface area covered by lesions was much greater after 6 weeks, when it was impossible to distinguish individual lesions (Table 5). Only one colony developed on minitubers planted in unamended soil C. The surface area of minitubers covered by silver scurf after 6 weeks increased significantly (rs = 0·99, n = 6) with the degree of soil infestation, and on minitubers after 3 weeks increased with the number of lesions (rs = 1·0) and surface area covered (rs = 1·0).

Table 5.  Development of silver scurf on minitubers 3 and 6 weeks after planting in soil C infested with differing concentrations of spores of Helminthosporium solani
 3 weeks after planting 6 weeks after planting
Number of conidia per g soilPercentage of minitubers affectedMean number of lesions per tuberPercentage surface area affectedPercentage of minitubers affectedPercentage surface area affected on minitubers
  1. ND, colonies were too numerous to count.

1000100ND57·510087·5
30010025·441·710087·5
10010013·514·710066·7
30100 2·0 1·8 8313·6
10 67 1·5 1·0100 7·1
0 17 0·2 0·1  0 0

When soils A and B were mixed in various proportions, the incidence of infection by H. solani on minitubers 6 weeks after planting was similar (50%) for unamended soils A and B. The incidence of tuber infection with the various soil mixtures ranged from 16 to 100% in an irregular pattern.

Comparison of PCR and bioassay

Both bioassay tests and real-time quantitative PCR detected C. coccodes and H. solani in artificially infested soil, and for both pathogens each measure was closely related to the concentration of soil inoculum (Table 6). However, the bioassay test detected inoculum of C. coccodes in unamended soil B, 10 years after potatoes, while the PCR test did not. Both methods detected inoculum in unamended soil A and none in unamended soil C. With H. solani the PCR test detected inoculum in unamended soil A, collected 6 months after a potato crop, but not in the other two soils, whereas the bioassay test detected inoculum in all three soils, with the lowest infestation in soil C.

Table 6.  Comparison of the results of detection assays for Collectotrichium coccodes and Helminthosporium solani in naturally and artificially infested soil using real-time quantitative PCR and bait tests with minitubers and microplants
PathogenSoilNumber of propagules per g soilConcentration of DNA (fg)aPercentage infection in bioassay
  • a

    PCR results for C. coccodes and H. solani are taken from Table 5 of Cullen et al. (2002) and Cullen et al. (2001), respectively.

  • b

    Mean of test results using minituber derived plants and microplants assessed 6, 8 and 9 weeks after planting.

  • c

    Not assessed.

  • d

    Mean of assessments made 3 and 6 weeks after planting.

    Plants with infected roots
C. coccodesAUnamended 432100
BUnamended   07
 CUnamended   00b
0·06  236b
0·12  438b
1·2 15452b
  121573100b
    Minitubers/surface area affected
H. solaniAUnamended 13250/–c
BUnamended   050/0·3
301060100/10·0
3005500100/73·3
CUnamended   08/0·05d
30 16892/7·3d
300 872100/64·6d

Discussion

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

These experiments confirm that C. coccodes present in soil can lead to infection of a range of organs of the potato plant (Read & Hide, 1988). Recording root infection was the most sensitive and reliable method for detecting soil infestations by C. coccodes, but did require the plants from minitubers to be grown for at least 9 weeks in the test soil. This time could be reduced using microplants. The sensitivity of the bioassay is not surprising, as roots are susceptible to infection by the fungus. In addition, as they spread through the soil they are likely to come into greater contact with inoculum surviving in soil than either mother tubers or stems. This suggests that if a bioassay test was used to determine the degree of soil infestation by C. coccodes, then tests using microplants may be as effective as those using minitubers. The results also demonstrate that a bioassay test could pick up very low amounts of inoculum of C. coccodes in soil. Infestations of less than 0·4 microsclerotia per g soil were detected in the two artificially infested soils. Plating diluted soil suspensions onto CGPI medium was also effective in detecting infestations of C. coccodes in artificially infested soil as low as nine conidia per g soil or one microsclerotium per g soil.

The bioassay tests for H. solani confirm that minitubers are a sensitive biological tool with which to detect this pathogen, and that exposure of minitubers to test soil for 3 weeks is sufficient to allow infection to develop from soil inoculum. Infestations of ≤ 10 conidia per g soil were apparently detected, but this is likely to overestimate the sensitivity of the method, as all unamended soils gave infection of minitubers and must therefore have contained some inoculum. The degree of infection on minitubers can be quantified by counting the number of colonies or by estimating the surface area covered by sporulating silver scurf lesions. Both assessments were closely related to the degree of soil infestation with H. solani, but the latter measure is more flexible because merging of colonies does not limit recording.

A number of major difficulties exist in employing such bioassays on a routine basis in a test programme. The first is the need to include standard test soils in order to transform the results to a numerical basis. A second difficulty is the space and time required to conduct such bait tests, and these constraints may carry significant costs. A final difficulty is maintaining uniform conditions in a glasshouse, and it may be that variations in environmental conditions affect infection both directly, and indirectly by affecting plant growth. The latter effect is likely to be less critical when infection is measured directly on minitubers after 3 weeks when testing for H. solani. However, the method does have the advantages of allowing relatively large amounts of soil to be tested and detecting infective units in the soil irrespective of how they exist, e.g. single microsclerotium, groups of microsclerotia, or possibly mycelia from groundkeepers. A further constraint is the time required to produce minitubers for bioassay testing. The minitubers also need to be stored for a period after harvesting to allow them to break dormancy naturally. Microplants can be produced much more quickly than minitubers, and were used successfully to detect C. coccodes in soil. However they were not suitable for detecting H. solani, which does not infect roots or stems.

Bioassay and real-time quantitative PCR were equally effective in detecting C. coccodes and H. solani in artificially infested soil and unamended soil collected 6 months after a potato crop. However, in soils collected at longer intervals after a potato crop, bioassay appeared to be more effective than PCR for detecting soil inoculum. It is likely that surviving propagules in these soils are unevenly distributed, so a relatively large sample may need to be tested in order to detect small pockets of surviving propagules. In the experiments described here, the PCR assay tested the equivalent of 4·5 g soil out of the 20 or 30 g in the original suspensions compared with c. 2·7 kg (six replicates) in the bait test. The same limitation also applies to the use of CGPI medium because the actual amount of soil being plated is less than 1 g out of the suspended sample of 10 g. The sensitivity of the PCR and the semiselective medium assays may therefore be negated to some extent by the volume of soil tested. This is particularly relevant when the methods are used to test naturally infested soil in which the number of surviving propagules is low, and in which the propagules are irregularly distributed. The detection results for C. coccodes in unamended soil B illustrate this point. The proportion of plants developing infected roots in this soil was one in six, both in the artificially infested soil series and in the soil mixture experiment (Figs 2 and 4). This means that only 450 g out of the 2·7 kg tested contained inoculum of C. coccodes, and it is likely that only a portion of this sample contained infective propagules. The probability of detecting inoculum by PCR in a random sample of soil taken from fields a few years after a potato crop may be relatively low because of the amount of soil actually tested. This limitation in detecting inoculum unevenly distributed within a sample collected from one point in a field is likely to be compounded when testing samples collected randomly within a field, particularly if they are bulked. Moreover, the result of a test on a bulked sample may be a poor predictor of the risk of commercially important levels of disease developing in a potato crop from irregularly distributed, low levels of inoculum in a field. These limitations need to be addressed before the method can be employed on a routine basis for advisory purposes.

These experiments demonstrate that C. coccodes can survive in field soils for several years. The results from the field sampled 10 years after potatoes (soil B) suggest that the average amount of inoculum surviving in this soil might be around 0·08 microsclerotia per soil (Figs 1 and 2). However, the amount of inoculum in the soil sampled 6 months after harvesting a potato crop was four to five times greater than that in the soil taken 10 years after potatoes. This very limited testing might suggest that soil inoculum will decline with time, but the results will have been influenced by other factors such as the amount of inoculum originally left in the field, husbandry practices after a potato crop, and soil type. The accompanying study being conducted by ADAS Consulting Ltd may help to elucidate these tentative findings. Farley (1972) found that the number of conidia of C. coccodes in soil declined fairly rapidly in a few weeks but small numbers could still be detected after 52 weeks, whereas with microsclerotia soil populations remained constant. However, Blakeman & Hornby (1966) reported that only 53% of sclerotia remained viable after 83 days in soil in a glasshouse. In the present study this pathogen, which is considered to be a soil organism, was not detected in the soil taken from a field in which potatoes had not been grown for c. 45 years. This may be because the pathogen was not introduced into the soil when the potatoes were last cropped, or because the amount of inoculum was too low to be detected or had not survived.

Helminthosporium solani was recovered from all unamended soils used in these tests, although the recovery from a field (soil C) in which potatoes had not been grown for 45 years was limited to one colony on one minituber. This supports, to some extent, the findings of Bains et al. (1996) who recovered H. solani from a field in which potatoes had not been grown in living memory. However, it is possible that in our experiments some contamination of the soil may have occurred during air-drying in the glasshouse. The incidence of infection on minitubers was similar for soils taken from fields at widely differing intervals after a previous potato crop (6 months and 10 years). The number of conidia per g soil in these fields would appear to be around 10, based on data in Tables 2 and 3. Firman & Allen (1995) noted that the number of viable conidia in an artificially infested soil declined to < 1% of the original number within 10 weeks, and this decline was unaffected by soil moisture but enhanced by increasing temperature. By contrast, Frasier et al. (1998) found that conidia of H. solani survived for at least 9 months in soil from stores, confirming the findings of Carnegie et al. (1996) and Hall (1996) that H. solani could survive in dry store soil. The limited results presented here suggest that H. solani can be present in field soil at considerable intervals after a previous potato crop. Such inoculum may be important in providing the initial infections on healthy stocks (Mérida & Loria, 1994; Bains et al., 1996).

Acknowledgements

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

We thank Mrs J. Craigie and Ms C. Scott for technical assistance, Mr D. W. Cullen, SCRI for help and advice, Mr N. J. Bradshaw, ADAS Consulting Ltd for overall supervision of the project and helpful comments on the paper, and the Ministry of Agriculture, Fisheries and Food (MAFF), now the Department for Environment, Food and Rural Affairs (DEFRA), who funded the work (Grant number HP0125T).

References

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