Plasmodiophora brassicae in its environment-effects of temperature and light on resting spore survival in soil

Clubroot caused by Plasmodiophora brassicae is an important disease on cruciferous crops worldwide. Management of clubroot has been challenging, due largely to the millions of resting spores produced within an infected root that can survive dormant in the soil for many years. This study was conducted to investigate some of the environmental conditions that may affect the survival of resting spores in the soil. Soil samples containing clubroot resting spores (1 × 107 spores g-1 soil) were stored at various temperatures for two years. Additionally, other samples were buried in soil, or kept on the soil surface in the field. The content of P. brassicae DNA and the numbers of viable spores in the samples were assessed by quantitative polymerase chain reaction (qPCR) and pathogenicity bioassays, respectively. The results indicated that 4°C, 20°C and being buried in the soil were better conditions for spore survival than were −20°C, 30°C and at the soil surface. Most of the spores kept on the soil surface were killed, suggesting the negative effect of light on spore viability. Additional experiments confirmed that ultraviolet (UV) light contributed a large negative effect on spore viability as lower pathogenicity and less DNA content were observed from the 2-and 3-hour UV light treated spores compared to the untreated control. Finally, this work demonstrated that DNA-based quantification methods such as qPCR can be poor predictors of P. brassicae disease potential due to the presence and persistence of DNA from dead spores.

impacted by soil type, soil pH values, water content, temperature and light [14]. Other factors 64 may include the number of freeze-thaw cycles, rapid temperature shifts, microbial activity, 65 population size, premature signals to exit dormancy, etc [14]. There is almost no information on 66 how these soil environment parameters may affect the long-term viability of P. brassicae resting 67 spores. In one study, the soil pH was found to play an important role in spore viability, and that 68 soil temperature and moisture had much less effect [21]. However, the duration of the 69 environmental exposure was only 30-days leaving us without an understanding of long-term 70 effects of the soil environment on resting spore survival. survival of resting spores using qPCR analysis side-by-side with pathogenicity bioassays. The 83 results from the two approaches were compared and any inconsistency was further investigated.

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The objectives of this study were 1) to assess the viability of resting spores after being stored in 85 different conditions for two years, 2) to identify any differential effects of environmental 86 conditions on the survival of spores, 3) to illustrate the difference between DNA content and the 87 biological potential (number of living spores) by using the same samples assessed by qPCR and 88 pathogenicity bioassays, and 4) improve our understanding of P. brassicae survival, biomass 89 assessments, and clubroot epidemiology.  Two P. brassicae populations, one pathotype 3H and the other composed mainly of pathotype 100 5X, were used exclusively in this study. The pathotype 3H population was the Led09 strain [22] 101 and was determined to be pathotype 3 by its reactions on the Williams The dried sample was ground with a coffee grinder and then aliquoted into 1.5-mL 116 microcentrifuge tubes with 1 g soil per tube. The tubes were stored at -20°C.  The spore suspension was adjusted to 2 × 10 8 mL -1 and from which dilutions were prepared. For 121 soil inoculation, 50 µL of a spore suspension was added into one of the 1.5-mL tubes containing 122 1 g soil, which ensured that all inoculated soil samples had the same water content (50 µL g -1 soil) 123 despite different spore concentrations. After inoculation, the tubes were kept at room temperature 124 for one hour to allow the inoculum to be absorbed evenly in the soil sample. The tubes were then 125 vortexed for 10 seconds and immediately used in the subsequent experiments. For simplicity, 126 these tubes will be referred to as 'original-tubes' throughout the rest of this paper. The original-tubes were prepared at the final concentration of 1 × 10 7 spores g -1 soil. One   After autoclave, the tubes were kept at 4°C for no more than 24 hours before subjected to qPCR 144 analyses. For UV light treatment, original-tubes were prepared at the final concentration of 1 × 10 6 spores 148 g -1 soil. Then the soil from each tube was transferred into a 5-cm petri dish. With the lid closed, 149 the petri dish was shaken by hand for 30 seconds to make sure that the soil and the inoculum 150 were well-mixed. After removing the lid, the petri dish was placed in a PCR workstation (Model  The qPCR program consisted of 40 cycles of denaturation at 95°C for 10 sec (3 min for the 178 initial denaturation) and annealing/extension at 60°C for 30 sec. with tap water at pH 6.4 (adjusted with HCl). After 42 days, the plants were separated into 191 classes using a 0-to-3 scale [28], where 0 = no clubbing, 1 < one-third of the root with symptoms 192 of clubbing, 2 = one-thirds to two-thirds clubbed, and 3 > two-thirds clubbed (Fig. 1C). templates extracted from soil samples containing a 10-fold dilution series (from 1 × 10 7 to 1 × 217 10 3 spores g -1 soil) of P. brassicae resting spores (Fig. 2). The regression equation of the 218 standard curve is Y = -3.0650X + 40.7583 with R 2 = 0.9968. The calculated primer efficiency is 11 219 112%. For any soil sample, if the Cq value was obtained, its spore concentration or the 220 equivalent of spore concentration could be calculated using the regression equation.

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Quantitative PCR analysis of spores stored in different conditions for two years 223 After stored in different environmental conditions for two years, soil samples inoculated with P. 224 brassicae resting spore were analysed by qPCR. Based on the Cq values, the equivalent spore 225 concentration was calculated (Fig. 3). Compared to the fresh prepared 1 × 10 7 spores g -1 soil  series (1 × 10 7 to 1 × 10 3 spores g -1 soil) of resting spores were conducted in parallel with the 252 pathogenicity assays of the stored samples. The results showed that a spore concentration ≥ 5 × 253 10 3 spores g -1 soil could cause clubroot with same severity as the spore concentration at 1 × 10 7 254 spores g -1 soil (Fig. 4B). Thus, we could conclude that the concentration of living spores in the 255 soil samples kept on soil surface, at 30°C or at -20°C were lower than 5 × 10 3 spores g -1 soil 256 while the concentration in the sample on soil surface was lower than 1 × 10 3 spores g -1 soil.

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Pathogenicity assays vs. quantitative PCR 259 Comparing the results from the qPCR analysis to those of the pathogenicity bioassays indicated 260 that qPCR should not be used to assess the numbers of living spores or disease potential. When 261 spores die, their DNA can remain in the sample and produce qPCR amplicons, although more or 262 less of this DNA would degrade depending on the environmental conditions. This is illustrated 263 by the qPCR and pathogenicity bioassay results of the soil sample stored at -20°C. The qPCR 264 data indicated that the amount of DNA in the -20°C sample was equivalent to 1 × 10 6.39 spores 13 265 g -1 soil, which is the highest mean among all treatments except the fresh prepared 1 × 10 7 spores 266 g -1 soil control (Fig. 3). However, pathogenicity bioassays indicated that less than 5 × 10 3 living 267 spores g -1 soil were present in this sample. The difference between the qPCR result and the 268 pathogenicity bioassay result of this sample was greater than those of the samples on the soil 269 surface, or at 20°C or 30°C. This could be explained by the assumption that most spores in soil were treated with UV light and subjected to qPCR analyses and pathogenicity bioassay.

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After treatment with UV light for one or two hours, a decrease in the quantity of DNA was 291 observed (Fig. 6A). When the treatment time was extended to 3 hours, only 50% (10 5.66-5.94 ) of 292 DNA, compared to the non-treated control, could be identified from the soil sample, which 293 indicated that more than half of the resting spores were killed by the 3-h UV light treatment. This 294 result was supported by the pathogenicity bioassays (Fig. 6B), in which 2-and 3-hour UV light 295 treated samples showed significantly reduced pathogenicity compared to the non-treated control. In this study we used primer pair CrqF2/CrqR2 and probe PB1 for qPCR assessment of P. 300 brassicae spores. The probe has been used with the primer pair DC1F/DC1mR [27,30], which 301 was derived from Rennie et al.
[31] but with the reverse primer modified. We redesigned the 302 primers to avoid potential primer dimers as predicted by the Multiple Primer Analyzer 303 (https://www.thermofisher.com). Compared to other published primers, this primer pair showed 304 similar specificity and sensitivity to P. brassicae when used in SYBR green based qPCR (Feng,305 unpublished data). Since the sequence of the probe PB1 doesn't create more specificity, we 306 believed that this primer pair would be as efficient as others when used in probe-based qPCR.

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The efficiency of the primers used for the template DNA in this study was 112%. This could be 309 attributed to the PCR inhibitors present in the DNA template solution. For example, if the 310 standard curve was created from a dilution series of one genomic DNA preparation with inhibitors (generally chemicals used for DNA extraction), primer efficiency larger than 100% 312 would be expected, due to the fact that the inhibitors in the original template solution were 313 diluted along with DNA dilution. However, in this study, the standard curve was created by 314 serial DNA preparations from soil samples inoculated with spore dilutions. Thus, we concluded 315 that at least some of the PCR inhibitors were derived from P. brassicae spores. The more spores 316 from which the DNA derived, the more inhibitors in the DNA solution.

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It has not escaped our notice that the standard deviations of qPCR data from each treatment in 319 the UV light experiment (Fig. 6A) were smaller than those in the 2-year-storage experiments 320 (Fig. 3). We inferred that in the 2-year-storage experiments, stochastic factors had more time to 321 interact with the spore samples. Therefore, more variability was created within the different 322 tubes, although these tubes were kept in the same bottle. In this study, we provided the evidence that using qPCR data to enumerate or estimate living 325 spores can, in some cases, dramatically overestimate the numbers of viable resting spores and 326 disease potential. Few studies have been conducted to evaluate differences between using qPCR 327 results and pathogenicity bioassay results to assess P. brassicae resting spore viability. PMA-328 PCR has been used for the assessment of P. brassicae living spores [32,33], however while this 329 method has the potential to improve the measurement of viable spores via qPCR, the usefulness suggest that the best way to measure spore viability and disease potential is pathogenicity 335 bioassays, with vital staining of spores being a next best option [33].

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Survival of Plasmodiophora brassicae spores in different temperatures 338 Among the four controlled temperatures, 4°C and 20°C were better than -20°C and 30°C on 339 maintaining spore viability (Fig. 4A). These results indicated that compared to 4°C and 20°C, some experiments have suggested that the decay may not be linear, but rather a rapid decline in 365 viability occurs over the first year or two without a host, followed by a much slower decline over 366 the next 10 to 20 years [12,20]. In this study, we found that most spores can maintain full 367 viability and pathogenicity after being buried in the soil for two years. In contrast, most spores at 368 the soil surface totally lost their viability and pathogenicity after two years. However, since 369 spores were kept in 1.5-mL tubes and the tubes were kept in bottles, results from either treatment 370 cannot be used to predict the fate of spores in real field conditions. Nevertheless, the differences 371 between the two treatments provided evidence that spores buried in the soil had more chances to 372 survive than those on soil surface. According to Kim et al. [36], more than 97% of the total of P. It has been known for many years that UV light has various effects on fungi and other 385 microorganisms [38]. When DNA is exposed to UV radiation adjacent thymine bases will be 386 induced to form cyclobutane pyrimidine dimers by the condensation of two ethylene groups at C-387 5 and C-6. Additionally, adjacent thymines can be linked between the C-4 residue and the C-6 of 388 its neighbor. In either case, a "kink" is introduced into the DNA. Therefore, by exposing 389 microorganisms to UV radiation, their DNA will be photo-damaged and will not be amplified by 390 DNA polymerase [39]. As another consequence, the DNA can't replicate and thus the cells die. Dopachrome biosynthesis pathway, which is involved in melanin synthesis, has been predicted 397 by a bioinformatic study [42]. However, in natural conditions, especially outside agriculture, 398 most resting spores of P. brassicae would be buried in the soil for their entire life. Thus, it is 399 likely that P. brassicae did not evolve a strongly expressed UV light resistance during evolution. This was supported by the current study, in which the P. brassicae resisting spores were found to 401 be sensitive to UV light treatment. To prevent the spread of clubroot, sanitation of field equipment is important. Currently bleach is 404 recommended as one of the most effective chemicals for inactivation of clubroot resting spores 405 (https://www.canolawatch.org/2018/06/27/clubroot-disinfectants-bleach-is-best/). However, 406 bleach is corrosive to metals, causes rubber to harden, stains and damages clothing and footwear.   Inoculating canola with Plasmodiophora brassicae-infected soil 423 As with other biotrophic plant pathogens, plant inoculation with P. brassicae can be challenging.

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In diagnosis service activities and studies on the pathogenicity of multiple strains (e.g. 425 pathotyping, screening for resistant germplasms), inoculum sources are generally limited by the 20 426 amount of P. brassicae-containing soil or root galls. Using small amounts of inoculum allows for 427 more experimental replications to be set up, but often decreases the efficiency of infection. In 428 this study, we developed and optimized a simple procedure for canola seedling inoculation (Fig. 429 1B and C; termed tube-method), which is extremely useful when the inoculum source is limited.

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Compared to the soil inoculation method [45], the tube-method uses much less inoculum;  shown as mean ± standard deviation (n=3).  spores. Means in the plot topped by the same letter do not differ based on Fisher's LSD test at P 599 ≤ 0.05 (n = 5). A, Two experiments on soil samples containing 1 × 10 7 spores g -1 soil spores that 600 had been stored in different conditions for two years. Negative CK, an uninoculated soil sample 601 that has been stored at -20°C for two years; Positive CK, a fresh prepared soil sample containing