Skip main navigation
Previous
Next
RESEARCHFree Access icon

Brassica Cover Crops and Natural Spongospora subterranea Infestation of Peat-Based Potting Mix May Increase Powdery Scab Risk on Potato

    Affiliations
    Authors and Affiliations
    • Maryam M. Alaryan1 †
    • Yuan Zeng2 3
    • Ana Cristina Fulladolsa1
    • Amy O. Charkowski1
    1. 1Department of Agricultural Biology, Colorado State University, Fort Collins, CO 80523
    2. 2School of Plant and Environmental Sciences, Virginia Tech, Blacksburg, VA 24061
    3. 3Southern Piedmont Agricultural Research and Extension Center, Virginia Tech, Blackstone, VA 23824
    Published Online:1 Sep 2023https://doi.org/10.1094/PDIS-04-22-0863-RE

    Abstract

    Spongospora subterranea is a soilborne plasmodiophorid that causes powdery scab and root gall formation in potato. In this study, 18 cover crops suitable for use in dry, high-altitude potato production regions were assessed in potting mix trials to determine whether these cover crops altered S. subterranea population levels. Although S. subterranea appeared to invade roots of all plant species tested, the pathogen was unable to complete its life cycle on 11 of 18 cover crops based on postharvest qPCR and microscopy results. Buckwheat, legumes, and scarlet barley do not appear to support pathogen replication, but the pathogen may be able to complete its life cycle in some mustards. High variability occurred in the experiments and part of this may be due to the natural infestations of peat-based potting mix with S. subterranea. A tomato bioassay was used to confirm that commercial sources of peat-based potting mix were infested with S. subterranea. Dry heat and autoclaving were tested as sanitation methods and multiple rounds of autoclaving were required to reduce viable S. subterranea in potting mix. A second cover crop experiment with autoclaved potting mix was conducted and it confirmed that buckwheat, legumes, and barley do not support S. subterranea replication but that some brassica crops may be hosts of this pathogen. The results suggest that buckwheat, legumes, and barley pose the least risk as cover crops in S. subterranea infested fields and show that peat-based potting mix should not be used in seed potato production.

    Powdery scab of potatoes caused by the plasmodiophorid, S. subterranea (Wallr.) Lagerh., impacts potato production in many countries (Tsror et al. 2020a). Powdery scab reduces potato tuber quality and marketability by causing small lesions on the tuber skin that may develop into wart-like masses filled with sporosori (Balendres et al. 2017; Tsror et al. 2020a). These tuber lesions can be severe enough to cause tubers to dehydrate during storage and to restrict both domestic and export sales of fresh, processing, and seed potatoes. The pathogen also forms galls on the roots that reduce potato growth (Balendres et al. 2018a; Falloon et al. 2016). S. subterranea may also cause losses in other crops, such as greenhouse-grown tomatoes, where it affects root function and reduces plant growth (Balendres et al. 2018b; Thangavel et al. 2015). S. subterranea also transmits potato mop-top virus (PMTV), the type member of the Pomovirus genus, which causes necrotic arcs (spraing) in potato tuber flesh (Steven 2002; Thangavel et al. 2015).

    S. subterranea has a complex biphasic life cycle, allowing it to persist in soil with the production of resting spore aggregates (sporosori; Merz 2008). The sporosori have some resistance to fumigation and can survive in soil for over 10 years (Balendres et al. 2017; Tsror et al. 2019). Each resting spore within a sporosorus can release a biflagellate motile primary zoospore. The number of primary zoospores per sporosorus ranges from approximately 150 to 1,500 and not all zoospores in a sporosorus germinate simultaneously (Falloon et al. 2011), which makes it difficult to accurately quantify viable soil inoculum. Primary zoospores swim toward host roots, stolons, and tubers if free water is present, and then the zoospores encyst and penetrate plant cells to initiate infection (Amponsah et al. 2022; Balendres et al. 2018a; Merz 2008; Thangavel et al. 2015).

    After a successful infection, the pathogen forms plasmodia that develop into zoosporangia, which produce secondary zoospores that lead to further infections when released into the soil (Merz 2008; Thangavel et al. 2015). The secondary zoospores are also biflagellate and motile, and they can infect the host roots, tubers, or stolons, where they form plasmodia that may develop into zoospores or sporosori. In addition, primary and/or secondary zoospores may fuse via plasmogamy to form a binucleate spore that will invade plant cells and form a binucleate plasmodium that can also develop into a sporosorus. Unlike the resting spores, the zoospores cannot tolerate harsh environmental conditions. Once emerged, zoospores have approximately 2 to 5 h in soil to locate a host before they die (Balendres et al. 2018a; Merz 2008).

    Cover cropping is an effective and important practice for managing soilborne diseases and improving soil structure (Larkin 2015). Disease management can occur through loss of spore viability in the absence of a host crop, biofumigation, or trap crops, which support spore germination or infection but not completion of the pathogen’s life cycle (Tsror et al. 2020b). However, cover crops in which the pathogen can complete its life cycle are inappropriate for rotation in pathogen-infested fields. Common cover crops in the Brassicaceae, Asteraceae, Solanaceae, and Poaceae families may serve as S. subterranea hosts (Qu and Christ 2006; Simango et al. 2020; Tsror et al. 2020b). Previous cover crop studies rarely reported which varieties were tested and inconsistent results were observed.

    The initial goal of this study was to use a greenhouse assay to identify cover crops suitable for management of powdery scab in dry, high-altitude potato production regions in order to guide on-farm S. subterranea management. During our initial work in this study, we found that commercial peat-based potting mix is frequently contaminated with a low level of S. subterranea. This was consistent with the findings of Zeng et al. (2020), where a low level of inoculum was present in potting mix to which no inoculum had been added. Since the initial generation of seed potatoes is frequently grown in peat-based potting mix (Frost et al. 2013) and this substrate is often used for S. subterranea experiments, potting mix sanitation is essential. Therefore, an additional goal of this study was to test methods for potting mix sanitation.

    Materials and Methods

    Plants and planting materials

    The cover crop seeds were provided by a crop consultant cooperator and are listed in Table 1. Solanum chacoense line M6 seeds were obtained from the USDA germplasm center. The cold-adapted tomato varieties “Anna Russian” and “Alaskan Fancy” were obtained from commercial sources. The peat-based potting mix for these experiments was collected from four commercial potato growers and purchased from commercial suppliers of potting mix.

    Table 1. The mean ± standard deviation (SD) of postharvest Spongospora subterranea population (sporosori/g of potting mix) in peat-based potting mix, resulting from growing 18 cover crops in experiment one, and the presence of plasmodia and zoosporangia in their roots

    Cover crop experimental design

    In the first greenhouse experiment, 18 cover crops used in rotation with potato in the San Luis Valley were evaluated for their ability to reduce or increase soil inoculum level (Table 1). Then, a subset of 11 cover crops (Table 2) that grew well under greenhouse conditions and that appeared to be either the most resistant (buckwheat and legumes) or the most susceptible (mustards) to S. subterranea were evaluated in a second greenhouse experiment. Both experiments were conducted in a greenhouse at the San Luis Valley Research Center (SLVRC). The first was conducted from February 5 to May 7, 2019, and the second from December 18, 2020, to March 18, 2021. The greenhouse conditions were set as described in Houser and Davidson (2010) with some modifications. The greenhouse temperature was set to 10 to 15.5°C with 70% relative humidity and 12 h of light.

    Table 2. The mean ± SD of postharvest Spongospora subterranea population (sporosori/g of potting mix) in peat-based potting mix, resulting from growing 11 cover crops in experiment two, and the presence of zoosporangium in their roots

    In the first experiment, the positive control, Solanum chacoense line M6, and cover crop seeds were grown in a greenhouse room in the Plant Growth Facilities at Colorado State University. The seedlings were transferred to the SLVRC after 3 to 5 weeks and planted in prepared potting mix (1:1:2 vermiculite/peat moss/sand) with or without addition of 10 sporosori/g of potting mix per pot (Zeng et al. 2020). In the second experiment, seeds of selected cover crops and single-drop tubers of “Cherry Red” potato (Solanum tuberosum L., positive control) were planted directly into the prepared potting mix with or without the addition of 40 sporosori/g of potting mix per pot at the SLVRC. Each 15-cm diameter pot was filled with 2 liters of prepared potting mix with 3 seed/mini tubers planted 5 cm below the potting mix surface level, and later the plants were thinned to one plant per pot. The seed tubers were surface sterilized with a 10% dilute solution of commercial bleach (sodium hypochlorite) for 1 to 3 min, rinsed with distilled water, and dried before planting. The seed tubers did not have visible powdery scab symptoms. During the first experiment, we discovered that S. subterranea is common in commercial peat used in potting mix. To reduce S. subterranea inoculum in the potting mix, the peat moss used to prepare the potting mix in the second experiment was autoclaved three times for 60 min at 121°C with slow exhaust prior to use.

    The inoculum used in this study was scraped from scabby lesions on potato tubers, sieved through three different mesh sizes (850, 150, and 74 μm), and air dried before storing at 4°C. Before inoculation, sporosori were suspended in water and the number of sporosori per microliter of water was determined with a hemocytometer under a light microscope (Qu and Christ 2006). The quantified sporosori inoculum was mixed with the prepared potting mix (Zeng et al. 2020), and each seed tuber or seedling was planted approximately 5 cm below the surface level of potting mix. After planting, each pot was watered to saturation using approximately 500 ml of water, and every two days each pot was watered by hand (200 ml/pot) until the plants emerged. After that, a drip system connected with 1/2 gallon per hour (GPH) pressure pinch-drip emitters (DripWorks, Willits, CA) was installed to automatically apply water for 2 min every 8 h. At 75 days after planting (DAP), the drip system was adjusted to apply water (200 ml/pot) for 5 min every 8 h until harvest (Zeng et al. 2020). In addition, water-soluble fertilizer “Peter’s Blossom 10-30-20” (Peters Professional, Earth City, MO) was applied every 7 days, at a rate of 200 ppm nitrogen/pot, starting from 35 DAP until harvest.

    In both greenhouse experiments, five plants of each species were inoculated and five were reserved as controls (noninoculated). Pots were arranged in a randomized complete block design.

    Plants were harvested approximately 90 days after planting, and the potting mix, tubers, and roots were collected from each pot and transferred to the laboratory for further analysis. The collected potting mix samples were tested for pathogen DNA as described below.

    Tomato plant bioassay

    Four 10-cm diameter pots were filled with 25 g (dry weight) of each potting mix sample, which was previously saturated with distilled water. Three tomato bait plants were grown from seed in each pot. Cold-tolerant varieties Anna Russian and/or Alaskan Fancy were used for these assays. Each pot was placed on top of a 100- × 15-mm Pyrex Petri dish to contain excess water and avoid cross-contamination. Plants were maintained in a growth chamber for 4 to 6 weeks (∼8 cm height), at 18 ± 0.2°C, with a 12-h light/dark cycle. Plants were watered with distilled water as necessary to maintain saturation. Osmocote 14-14-14 fertilizer was added at a rate of 1 g/pot after 1 to 2 weeks. Tomato plants grown in triple-autoclaved potting mix, vermiculite, and sand served as negative controls. In some assays, 9 or 140 sporosori/g of potting mix was added as a positive control.

    Preparation of potting mix samples for DNA extraction

    The potting mix samples were collected after harvest and left for approximately 2 months to air dry. Large pieces were broken up by smashing the potting mix in sealed plastic bags with a plastic mallet. The potting mix was then passed through two kitchen sieves (4 and 2 mm) to remove larger fragments of roots and potting mix. The sieved potting mix was ground for two 8-s pulses in a Blade Coffee Grinder (KitchenAid, St. Joseph, MI).

    Spongospora subterranea detection using qPCR

    DNA was extracted from a 0.25-g potting mix sample with a DNeasy Powersoil Kit (Qiagen, Hilden, Germany), following the manufacturer’s protocol with modifications described by Mallik et al. (2019). A standard curve was generated to quantify S. subterranea as described by Mallik et al. (2019). For the cover crop study, potting mix (1:1:2 vermiculite/peat moss/sand) was prepared and mixed well in a tray for each pot individually, and the desired amount of inoculum was added to the potting mix and mixed thoroughly to make sure that each pot had the same amount of inoculum. For the tomato bioassay, the standard curve was generated as described in Mallik et al. (2019), using a commercial potting mix instead of soil. The potting mix was autoclaved three times to eliminate naturally occurring S. subterranea prior to the addition of S. subterranea sporosori. Quantitative PCR (qPCR) was performed using the SsTQF1 and SsTQR1 primers and SsTQP1 probe designed for S. subterranea detection (van de Graaf et al. 2003). Each 20 μl qPCR reaction consisted of 2 μl of DNA, 10 μl of PrimeTime Gene Expression Master Mix (Integrated DNA Technologies, Coralville, Iowa), 0.5 μl of each primer (10 μM), 0.25 μl of the probe (10 μM), and 6.75 μl of sterile distilled water. The thermocycler protocol was 95°C for 2 min, followed by 40 cycles of 94°C for 15 s and 58°C for 45 s. Reactions were performed in a QuantStudio 3 Real-Time PCR System (Thermo Fisher).

    Microscopy assays

    For tomato bioassay experiments, the tomato roots were carefully harvested and gently rinsed in water to dislodge debris. The two longest roots from each pot were then stained with lactophenol trypan blue stain at 95°C for 5 min (Hernandez Maldonado et al. 2013; Koch and Slusarenko 1990). The stain solution was prepared by mixing three parts ethanol with one-part lactophenol trypan blue stock solution (10 g of phenol, 10 ml of glycerol, 10 ml of lactic acid, 10 ml of sterile distilled water, and 0.02 g of trypan blue stain; Hernandez Maldonado et al. 2013; Koch and Slusarenko 1990). The roots were destained overnight with a solution of one part glacial acetic acid to three parts ethanol. Stained roots were mounted onto slides immediately or stored in 50% glycerol. The roots were observed for the presence of S. subterranea under a light microscope at 400 and 1,000× magnification. The presence of plasmodia or zoosporangia was recorded. The remaining root tissue from each pot was combined and DNA was extracted for subsequent qPCR detection as previously described.

    For the first cover crop experiment a portion of the roots was randomly chosen and stained as described above. Stained roots from each replicate were mounted onto three slides to observe the presence of plasmodia, zoosporangia, and sporosori under the light microscope. In the second experiment, a portion of the roots was arbitrarily chosen from each replicate and stained with 1× DAPI staining solution at room temperature and incubated in the dark for 15 min. The stained roots from each replicate were mounted onto one slide with antifade media to observe the sporangial stage under the fluorescence microscope (PureBlu DAPI Nuclear Staining Dye, BIO-RAD). DAPI was used for better visualization of the sporangial stage of S. subterranea since zoosporangia are more distinctive than plasmodia.

    Potting mix sanitation assays

    Potting mix was subjected to dry heat at 105°C for 2, 5, or 8 days, or autoclaving at 121°C and 15 psi once, twice, or three times for 60 min. Samples were tested with qPCR and the tomato bioassay after the sanitation treatment to determine its effects. The dry heat treatments were assessed once, and the autoclave treatment experiment was performed twice.

    Statistical analysis

    Because the datasets on postharvest S. subterranea inoculum in the cover crop experiments were not normally distributed, the data were transformed using a Box-Cox power transformation before subjecting them to a paired t test using R software (package: MASS) to compare the means of the control and the inoculated samples. An ANOVA test was then applied to the transformed datasets to check the significant changes in sporosori amount after harvest among the tested cover crops. In addition, for each plant species, least squares means (LS-means) of the inoculum change for each treatment were compared among each other (control to control, inoculated to inoculated, and control to inoculated) using the Tukey method. For the potting mix sanitation experiments, generalized linear mixed models (GLMM) were used to determine if there was a significance of each fixed effect of a model by a type III test using the statistical software SAS OnDemand for Academics (https://www.sas.com/en_us/software/on-demand-for-academics.html). These fixed effects are tomato variety, types of potting mix, or their interactions in the tomato baiting assay and sanitation method or different levels of a sanitation method in the sanitation experiment. Specifically, PROC GLIMMIX was used to examine each fixed effect in the tomato baiting assay on the log odds of plants with plasmodia with a binomial distribution, whereas PROC GENMOD was used to conduct a Poisson regression analysis on S. subterranea sporosorus population in tomato roots, postharvest sporosorus population in potting mix, or the changes of sporosorus population levels in potting mix with a log link function. The effect of sanitation method or different levels of dry heat or autoclaving on the incidence of tomato plants with plasmodia was examined as described above using the GLIMMIX procedure. A multinomial model with a cumulative logit link function was constructed to determine the effect of different dry heat levels on S. subterranea sporosorus populations in tomato roots, or several linear models with a normal distribution with the identity link function were constructed to investigate other fixed effects on the changes of sporosorus population levels in potting mix using the GENMOD procedure. LS-means for each fixed effect were compared via Tukey’s method for multiple comparisons. All statistical analyses were performed at a significance level of α = 0.05.

    Results

    Brassica cover crops were potential Spongospora subterranea hosts (first experiment)

    Root galls were not found on any of the cover crops but were observed on Solanum chacoense M6. The lack of galls on cover crop species is consistent with most previous host range studies with other nonsolanaceous crops and weeds (Qu and Christ 2006; Simango et al. 2020; Tsror et al. 2020b). The light microscopy results showed that all the cover crop roots had plasmodia (Fig. 1C and D), but zoosporangia were only found in inoculated alfalfa and yellow blossom sweet clover (YBSC) in the legume family, and in some mustards (Brassicaceae family) in both inoculated and noninoculated treatments (Fig. 1E and F; Table 1).

    Fig. 1.

    Fig. 1. Representative images of lactophenol trypan blue-stained roots from experiment one. A, Solanum chacoense line M6 root cells containing S. subterranea plasmodia (P) and B, zoosporangia (Z). C, “Dixie” clover root cells and root hair containing S. subterranea plasmodia (P). D, “Pacific Gold” mustard root cells containing S. subterranea plasmodia (P). E, “Caliente” mustard root hairs and F, root cells containing S. subterranea zoosporangia (Z). All images were captured at 1,000× magnification under the light microscope. Similar results were observed in the other cover crops, as reported in Table 1.

    Download as PowerPoint

    There was no direct relationship between qPCR results measuring postharvest sporosori/g present in each sample and the presence of zoosporangia. For example, “Winfred” turnip had 35 sporosori/g potting mix but lacked zoosporangia in the roots, and the potato plants had a similar level of zoosporangia in the roots as some cover crops but had significantly higher postharvest S. subterranea DNA in the potting mix compared with the cover crops. Other plants that had low numbers of sporosori/g of potting mix had zoosporangia in their roots, such as “Pacific Gold” mustard, with 8 sporosori/g of potting mix.

    The mean postharvest inoculum in potting mix did not increase following planting of scarlet barley, “Pacific Gold” mustard, buckwheat, and the majority of legumes, with approximately 10 sporosori/g of potting mix or less detected by qPCR (Table 1). The variability was high and as a result, even cover crops that had more than 10 sporosori/g of potting mix postharvest did not show significant differences between the inoculated and noninoculated treatments. However, significant differences were observed in the inoculated treatments of different plant groups, with higher sporosori levels observed in the mustards compared with the other cover crops (P(daikon radish - cahaba vetch)inoculated = 0.05; P(caliente - sordan 79)inoculated = 0.05). Cover crops in the Brassicaceae family may have potential to increase soil inoculum levels since the postharvest sporosori level was above 10 sporosori/g of potting mix in almost all samples (Table 1).

    The mean postharvest S. subterranea sporosori level was higher in Solanum chacoense line M6 than in any of the cover crops that we examined in the inoculated treatments (Table 1). The microscopy results showed that potato roots were infected with S. subterranea, evidenced by the presence of plasmodia and zoosporangia in both inoculated and noninoculated (control) treatments (Fig. 1A and B; Table 1).

    Autoclaving peat moss prior to inoculation reduced experimental variability (second experiment)

    For the second cover crop experiment, we autoclaved the peat moss used in the potting mix three times to eliminate viable S. subterranea naturally present in the peat. Unlike the first experiment, the qPCR results showed a higher level of S. subterranea DNA in the inoculated samples compared with the noninoculated samples for almost all plant species tested. Among the 11 cover crops tested in this experiment, buckwheat had the lowest postharvest inoculum mean in the inoculated samples (Table 2). All the legumes, as well as buckwheat and barley, had a higher postharvest S. subterranea mean in the potting mix than the amount inoculated into the mix at the start of the experiment (40 sporosori/g potting mix). All the control samples had a low postharvest inoculum mean ranging from 0.6 sporosori/g for buckwheat to 29 sporosori/g for common vetch. “Caliente” mustard had the highest postharvest inoculum mean of the cover crops in both control and inoculated samples (114 sporosori/g and 347 sporosori/g respectively), suggesting that S. subterranea multiplied in this cover crop more than in the others tested. As in the first experiment, the sporosori level in the potato control was much higher than in any of the cover crops, demonstrating the high potential for a susceptible potato cultivar to increase soilborne inoculum compared with the cover crops tested.

    In the second experiment, we only looked for zoosporangia since these structures are more distinctive than plasmodia. Zoosporangia were found in 3 of the 11 cover crops tested (alfalfa, chickling vetch, and “Caliente” mustard) and the results were consistent with the first experiment, except that zoosporangium was found in chickling vetch only in the second experiment. Zoosporangia were not as abundant in the inoculated treatments for either alfalfa or chickling vetch compared with “Caliente” mustard and “Cherry Red” potato, and zoosporangia were not found in the noninoculated controls for alfalfa or chickling vetch (Fig. 2A and B). However, zoosporangia were found in both control and inoculated samples in the “Caliente” mustard roots (Fig. 2C and D). As expected, zoosporangia were also found in the roots of “Cherry Red” potato, and zoosporangia were abundant in both control and inoculated samples (Fig. 2E and F). As in the first experiment, there was poor correlation between the microscopy and qPCR results in the second experiment.

    Fig. 2.

    Fig. 2. Representative images of DAPI (4′,6-diamidino-2-phenylindole)-stained roots from experiment two. A, Alfalfa and B, Chickling vetch (legume family) root cells containing S. subterranea zoosporangia in the inoculated samples, captured at 1,000× magnification under the fluorescence microscope. C, “Caliente” (mustard family) root cells and D, root hairs containing S. subterranea zoosporangia in both control and inoculated samples, captured at 1,000× magnification. E, Potato cv. “Cherry Red” (positive control) root cells containing S. subterranea zoosporangia in both control and inoculated samples, captured at 1,000×, and F, potato cv. “Cherry Red” root hairs captured at 360× under the fluorescence microscope.

    Download as PowerPoint

    The ANOVA test for both experiments showed that all variables were significant, including inoculation treatments, cover crops, and their interaction (Table 3). A Tukey test was also performed for a multiple comparison of the transformed dataset to determine whether the cover crops were statistically different from one another. As expected, the postharvest inoculated mean of the positive control Solanum chacoense line M6 was significantly greater than all the cover crops group means (P < 0.05). In addition, there was a significant difference between the control and inoculated group means among all the cover crops tested (all P < 0.05).

    Table 3. ANOVA tests of fixed effects for postharvest Spongospora subterranea population in potting mix

    Potting mixes from multiple sources were infested with Spongospora subterranea

    During the course of the first cover crop experiment, we found that potting mix from multiple sources was infested with S. subterranea. Furthermore, during routine testing of potting mix samples submitted by potato farmers, we found that despite sample homogenization, qPCR assays were variable among potting mix subsamples. Multiple subsamples were tested from four potting mixes and there was a 100-fold variation in the quantitative results from the naturally infested potting mixes (Table 4). Less variation was seen in the inoculated control samples.

    Table 4. Spongospora subterranea detection by qPCR and a tomato bioassay in potting mix samples provided by four commercial potato growers

    A tomato bioassay can detect Spongospora subterranea in infested potting mix

    To determine whether the S. subterranea detected in the commercial potting mix by qPCR was viable, the potting mix samples were also tested with a bioassay using cold-adapted tomato plants as bait (Table 4). Plasmodia were observed in roots of tomatoes grown in two of the four naturally infested potting mix samples and in potting mix samples inoculated with either 9 or 140 sporosori/g of potting mix (Table 4). No plasmodia were observed in roots of plants grown in the negative control sample, which was a potting mix sample that consistently had undetectable levels of S. subterranea by qPCR in assays prior to the start of the experiment.

    DNA was extracted from 25 mg of dry root tissue and from the homogenized potting mix after completion of the bioassay and the qPCR was performed (Table 4). High levels of S. subterranea DNA were detected in all of the tomato roots where plasmodia were observed (except for plants in one pot) and in all of the positive control samples. The qPCR results showed low levels of S. subterranea DNA in roots from samples where no plasmodia were observed. High levels of S. subterranea DNA were only observed in the potting mix after the completion of the bioassay in two of the four control samples infested with 140 sporosori/g of potting mix prior to the start of the assay. Based on these results, we concluded that samples 1 and 2 (Table 4) did not contain viable S. subterranea and that the qPCR assay results of less than 30 sporosori/g of potting mix did not reliably indicate whether viable S. subterranea was present in potting mix samples. The analysis for the varieties was done across all samples, so we had a total of 14 replicates per variety. We also found higher levels of S. subterranea in potting mix after completion of the bioassay when Anna Russian plants were used (P < 0.001), across all samples.

    An additional three commercial potting mixes (PM, PG, and SE) were tested for S. subterranea to further evaluate both the qPCR assay and the tomato bioassay (Table 5). Two subsamples (A and B) per potting mix sample were tested. Of these, two potting mixes may have been free of S. subterranea (PM-A and B, and SE-B), one was infested with viable pathogen (PG), and the results from one subsample of potting mix was inconclusive (SE-A).

    Table 5. Spongospora subterranea detection by qPCR and a tomato bioassay in three commercial potting mix samples

    Potting mix sanitation

    Potting mix containing S. subterranea sporosori was treated with dry heat or autoclaved in an attempt to kill the pathogen. Treatment with dry heat (105°C) for up to 8 days was not effective and the trend was toward a greater number of S. subterranea sporosori in the potting mix after heat treatment (Table 6). The dry heat assay was not repeated because there was no indication that this treatment was effective. Autoclaving the potting mix at least once resulted in little to no detection of S. subterranea in potting mix or in the qPCR assay after completion of the bioassay, but plasmodia were still evident in the tomato roots. The statistical analysis of the autoclaving experiment results indicated no significant difference between autoclaved and nonautoclaved potting mixes.

    Table 6. Spongospora subterranea detection following sanitation by dry heat and autoclaving

    Discussion

    Cover crop strategies are essential for sustainable management of S. subterranea. However, previous work on cover crop susceptibility is contradictory, lacks details on plant varieties used, and only includes a few species suitable for high-altitude agriculture (Arcila Aristizabal et al. 2013; Qu and Christ 2006; Tsror et al. 2020b). We tested cover crop varieties suitable for high-altitude crop production and found plasmodia and zoosporangia in the roots of multiple cover crop species and slight increases in postharvest pathogen levels in the potting mix used to grow the plants for some cover crops. Promisingly, the low postharvest inoculum level observed after planting buckwheat, “Anika” peas, “Cahaba” vetch, and hairy vetch suggest that the soil inoculum level could be reduced over time with use of these cover crops. The cover crop that showed the greatest possibility of pathogen increase was “Caliente” mustard, although the increase was significantly lower than that caused by potato. However, the benefits of this brassica in a cover crop mix such as weed, nematodes, and fungal diseases suppression, great biomass production and nutrient recycling ability, and their ability to produce toxic compounds (isothiocyanates) that act as biofumigants to many soilborne diseases (Haramoto and Gallandt 2007; Wang et al. 2008), may still outweigh the risk of increasing S. subterranea.

    Inconsistencies between qPCR and microscopy assays for S. subterranea occurred in our study and are also common in other S. subterranea studies. This could be because sporosori do not form in or are not released from plant roots back into potting mix with some plant hosts or because the structures observed in the roots are caused by other microbes. For example, although Tsror et al. (2020b) saw that half of their alfalfa samples were positive for microbial structures resembling those of S. subterranea, none of the artificially inoculated alfalfa plants were positive when tested with a PCR assay. There are also inconsistencies among which plants species are reported to be S. subterranea hosts. Neither our study nor Qu and Christ (2006) report alfalfa as a S. subterranea host, but Tsror et al. (2020b) did. Similarly, we found that neither buckwheat nor barley were hosts of this pathogen, but Qu and Christ (2006) saw a low incidence of zoosporangia formation in buckwheat and Tsror et al. (2020b) reported that barley is a potential host. Arcila Aristizabal et al. (2013) reported that S. subterranea could complete its life cycle on Datura stramonium, while White (1954) showed that D. stramonium is an effective trap crop. This variation in results across studies could be due to plant variety differences, a low incidence of infection for some hosts, the presence of other microbes that mimic S. subterranea in microscopy assays, or S. subterranea population differences. This comparison across studies clearly shows that methods must be improved so that there is better correlation among studies and between PCR and microscopy assays.

    Despite the inconsistencies within and across studies, brassica cover crops are frequently reported to be potential hosts of S. subterranea. For example, multiple studies, including ours, report that mustard and rapeseed are hosts of S. subterranea (Qu and Christ 2006; Simango et al. 2020; Tsror et al. 2020b). Our results and those of others also show that R. sativus is a potential host (Arcila Aristizabal et al. 2013; Tsror et al. 2020b). These results suggest that multiple brassica species commonly used as cover crops with potato have potential to serve as a green bridge for S. subterranea.

    The low, but consistent, levels of S. subterranea associated with some of the cover crops in this study and others suggest that S. subterranea infection of cover crop species is, at best, inefficient. An alternative explanation for this low infection rate, which also takes into account the inconsistency between the qPCR and microscopy results seen in multiple studies, is that multiple Spongospora taxa with varied host ranges could be present in potting mix and soil and a type present at low abundance is infecting mustard and some of the other cover crops. Genome sequencing and further genotyping of S. subterranea populations in fields and potting mix, as well as on roots of cover crops, is needed to clarify the conflicting observations from multiple cover crop studies.

    A challenge that we encountered during this study was the frequency with which we found S. subterranea in commercial peat moss and commercial potting mix from multiple suppliers. Commercial potting mix is essentially unregulated compared with plant propagative materials in the sense that it is not routinely tested for pathogens or weed seeds, even when it crosses national borders. Commercial potting mixes are typically a mix of organic materials, such as peat moss, coco coir, pine bark, or compost, and inorganic materials, such as perlite and vermiculite. A similar problem with potting mix was described in 2012 in South Africa, where powdery scab was found in a potato production greenhouse facility and tentatively traced to the plant growth medium (Wright et al. 2012), and in 2010, when powdery scab was found in a greenhouse in Iran (Norouzian et al. 2010).

    When S. subterranea sporosori were present at low levels in potting mix, our qPCR results were variable among subsamples from the same potting mix sample, even after sample homogenization. This may be caused by sporosori size variability (Falloon et al. 2011), or by clumping of sporosori. We concluded that a tomato bioassay is useful for determining if potting mix samples contained viable S. subterranea and for confirming qPCR results. Based on these data and experience with additional potting mix samples provided by growers, we believe that the reliable detection limit of qPCR assays is approximately 30 sporosori/g of substrate. Unfortunately, in greenhouse assays, as little as 1 sporosorus/g of potting mix can cause severe disease in potato under conducive conditions, so tomato or potato bioassays are likely to be more sensitive in detecting infested potting mix (Zeng et al. 2020).

    S. subterranea is common in potato production fields and it may have spread due to its presence in peat-based potting mix, which is often used to grow the first generation of seed potatoes from micropropagated plantlets (Frost et al. 2013). There are rigorous testing, inspection, and sanitation protocols for seed potato production (Frost et al. 2013), but routine testing for pathogens in potting mix is not required for seed potato certification. We tried to sanitize the potting mix and found that autoclaving reduced inoculum levels, but did not completely eliminate the pathogen from potting mix samples. However, this is not feasible on a commercial scale. Based on our results, peat-based potting mix is a source of S. subterranea inoculum and should not be used in seed potato production unless an effective sanitation method can be developed. Several types of hydroponic or aeroponic systems that do not use potting mix have been developed for seed potato production and use of these systems eliminates the potential for frequent reintroduction of S. subterranea to seed potato production fields caused by the use of potting mix.

    Acknowledgments

    We thank the San Luis Valley Research Center for their collaboration, and we are especially thankful to Dr. Chakradhar Mattupalli and Jeremy Daniel for their great help in the San Luis Valley with our experiments. We also thank Andrew Cordova and Patrick O’Neill for their contributions in this study.

    The author(s) declare no conflict of interest.

    Literature Cited

    Funding: The Agricultural Research Service of the U.S. Department of Agriculture (USDA-ARS; 59-8062-7-001), USDA-NIFA SCRI grant 2020-51181-32136, and the Animal and Plant Health Inspection Service of the U.S. Department of Agriculture (USDA-APHIS; AP19PPQS&T00C182).

    The author(s) declare no conflict of interest.