Diversity and Pathogenicity of Pythium Species Associated with Reduced Yields of Processing Tomatoes (Solanum lycopersicum) in Victoria, Australia

Tomatoes are the second most profitable vegetable commodity in Australia, worth $674.2 million in 2018 to 2019 (Horticulture Innovation Limited Australia 2019). The Australian Processing Tomato Industry (APTI), based primarily in northern Victoria and southern New South Wales, makes up a small portion of this amount, worth between 15 and 30 million Australian dollars annually (Plant Health Australia 2017). Agronomic improvements and the gradual mechanization of the industry have helped improve productivity over time from 50 tons/ha to 95 tons/ha; however, in recent years productivity has plateaued and in some fields decreased (Gray 2019). This effect has been linked to the introduction of subsurface drip irrigation (SDI), which is kept in place between 5 and 20 years (Ashcroft et al. 2003).

Yield decline in the APTI is characterized by poor crop establishment, stunting, wilting, and reduced root systems of mature plants. The problem has been attributed partially to root and collar disease caused by soilborne pathogens (Ashcroft et al. 2003). Poor crop establishment has led some growers to switch from direct seeding to transplanting as a partial solution; however, the problem of crop stunting and suboptimal yields remains a key issue for the APTI.

Previously, Phytophthora nicotianae was identified as the primary pathogen causing root rot disease of processing tomatoes in Victoria (Flett 1986; Washington et al. 2001). However, preliminary surveys of Victorian tomato fields in the 2015 to 2016 season suggested Phytophthora root rot was no longer as prevalent, as had been reported in previous decades, possibly as the result of the shift from furrow irrigation to SDI and effective management in the form of phosphonate distributed via drip lines (Callaghan et al. 2019). In contrast, Pythium spp. and Fusarium oxysporum were consistently isolated from the root systems and collars of plants exhibiting poor growth in preliminary studies (Callaghan et al. 2019).

Pythium identification can be difficult because of overlapping morphological characteristics between species, as well as the diversity within some species groups. In recent decades, DNA-based methods have improved Pythium taxonomy. Some of the most commonly used molecular markers for the identification of oomycetes, including Pythium, are the internal transcribed spacer (ITS) region of the rDNA (Robideau et al. 2011) and the mitochondrial genes Cytochrome c oxidase subunit I (Cox-1) and Cytochrome c oxidase subunit II (Cox-2) (Martin 2000). Some reports have noted the need for additional molecular markers based on highly variable genes in order to resolve some of the more blurred taxon boundaries within the genus (Hyde et al. 2014; LéVesque and De Cock 2004). Generally, however, morphology coupled with ITS, Cox-1, and Cox-2 sequencing can resolve the identification of most Pythium species.

Of the approximately 20 different Pythium species that have been isolated from tomato plants globally, P. aphanidermatum, P. irregulare, and P. ultimum appear to be the most important pathogens (Blancard 2012; Deniel et al. 2011; Jenkins and Averre 1983; LéVesque and De Cock 2004; Manorantitham et al. 2001; Robertson 1973, 1976). Most of these reports do not come from Australia, but some of these species have been reported in Australia on hosts other than tomato (Harvey et al. 2008; Li et al. 2014; Petkowski et al. 2013; Tesoriero 2011). Most previous reports of Pythium root rot disease of tomato plants globally come from glasshouse rather than field systems and from seedlings rather than mature plants. Additionally, many previous reports on identification and taxonomy were based only on morphological characteristics.

The aim of this study was to use a combination of morphological and molecular methods to determine the incidence and diversity of Pythium species associated with Victorian processing tomatoes and to assess the relative pathogenicity of these species on processing tomato plants at different stages of growth from seed through to maturity. The overall objective was to ascertain whether Pythium species might be contributing to the poor growth and suboptimal yields in processing tomato crops.

Materials and Methods

Sampling and isolation.

During the tomato growing seasons (November to February) of 2016 to 2017 and 2017 to 2018, surveys were conducted at 20 tomato growing farms in north-central Victoria (Loddon-Campaspe and Goulburn regions), Australia. The sites had variable histories of tomato cropping, ranging from land with no prior history of cultivation to land with many decades of tomato cropping. Soil across the region is variable, including both gray self-mulching vertosols and red clay chromosols. Crop rotation varied, with some growers having ≤6 years of consecutive tomato crops on a site, others rotating every second year, and variations in between.

In each crop, observations for diseased plants were made via the entire length of five interrow spaces selected on an ad hoc basis, with scans of the adjacent three rows on each side. Plants showing aboveground symptoms of stunting, abnormal foliage color, or wilting were carefully removed, inspected, and transported in cool boxes to the laboratory for processing. We made isolations from root lesions, necrotic root tips, and the discolored inner collar and lower stem tissue of stunted plants by plating small segments of tissue onto water agar, potato dextrose agar amended with streptomycin (0.1 mg/ml) (PDA-S), and a vegetable juice (V8)–based oomycete selective medium amended with pimaricin (0.01 mg/ml), ampicillin (0.25 mg/ml), and rifampicin (0.01 mg/ml) (V8-PAR) (Jeffers and Martin 1986).

In some cases, soil samples were collected from the drip line of diseased tomatoes at 5 to 10 cm depth and baited for Pythium species. White rose petals were used as baits, and V8-PAR was used for plating infected material as described by Callaghan et al. (2016).

Putative Pythium isolates were hyphal-tipped onto V8-PAR and given University of Melbourne (UM) accession numbers. For storage, McCartney bottles containing tap water (about 5 ml) and 10 hemp (Cannabis sativa) seeds were autoclaved twice at 121°C for 20 min. In a sterile laminar flow cabinet, seven plugs (2 × 2 mm) from fully colonized V8 plates were excised and transferred aseptically into the McCartney bottles. The bottles were stored in darkness at room temperature.

Molecular identification and phylogenetics.

For DNA extraction, mycelia were scraped from 7-day-old hyphal-tipped cultures on PDA-S. DNA was extracted with a DNeasy Plant Mini Kit (Qiagen Pty. Ltd., Australia) according to the manufacturer’s instructions. DNA quality and concentration were assessed, then DNA concentrations were adjusted to 25 ng/μl suspended in sterilized deionized water (Milli-Q, Millipore Corporation).

The ITS gene region was amplified with primers ITS-1 (TCCGTAGGTGAACCTGCGG) and ITS-4 (TCCTCCGCTTATTGATATGC) (White et al. 1990). The PCR mixture consisted of 0.18 μl of Taq DNA polymerase (5 u/μl), 0.36 μl of 50 mM MgCl2, 0.75 μl of each primer (10 μM), 1.05 μl of 10 mM deoxynucleoside triphosphate, 1 μl of DNA, and Milli-Q water up to a final volume of 20 μl. The PCR cycle was a denaturation step of 5 min at 94°C; 35 cycles of 30 s at 94°C, 30 s at 55°C, and 90 s at 72°C; then a final elongation step at 68°C for 10 min. For Cox-2, the primers Cox2-F (GGCAAATGGGTTTTCAAGATCC) and Cox2-R (CCATGATTAATACCACAAATTTCACTAC) (Choi et al. 2015; Hudspeth et al. 2003) were used with the same PCR mixture and cycle as used for ITS. Cox-1 was amplified with OomCoxI-Levup (5′-TCAWCWMGATGGCTTTTTTCAAC-3′) and OomCoxI-Levlo (5′-CYTCHGGRTGWCCRAAAAACCAAA-3′) primers (Robideau et al. 2011). The PCR mixture was the same as that for ITS and Cox-2, except only 0.5 μl of DNA was added.

All PCRs were performed with an Eppendorf thermal cycler (Eppendorf Pty. Ltd. Australia). PCR products were then run alongside a Lambda DNA HindIII ladder (Thermo Scientific) at 85 V in a 1.5% (wt/vol) agarose gel stained with SYBR Safe DNA gel stain (Thermo Scientific) (0.1 μl/ml) for 45 min and then were visualized with a GelDoc (Bio-Rad Laboratories Inc., Australia) to confirm the presence of the amplified product. A NanoDrop One Microvolume UV-Vis Spectrophotometer (Thermo Scientific) was used to ascertain the concentration of product. PCR products were purified with a QIAquick PCR purification kit (Qiagen Pty. Ltd., Australia) and sequenced in both forward and reverse directions at the Australian Genome Research Facility (Melbourne, Australia).

Consensus sequences were created through ClustalW alignment of the forward and reverse sequences. Isolates were tentatively identified to species level via Basic Local Alignment Search Tool searches of ITS, Cox-1, and Cox-2 sequence data in GenBank. If isolates were 99 to 100% similar to type or ex-type species from reputable sources in GenBank, species identity was tentatively confirmed. Sequences of Pythium isolates from various studies of LéVesque and De Cock (2004), Robideau et al. (2011), Moralejo et al. (2008), and Spies et al. (2011) were downloaded from GenBank to be used as reference isolates in subsequent phylogenetic analysis (Supplementary Material S1) (LéVesque and De Cock 2004; Moralejo et al. 2008; Robideau et al. 2011; Spies et al. 2011).

Maximum likelihood phylogenetic trees were constructed based on the Tamura and Nei (1993) model in Mega 7 (Kumar et al. 2015) to confirm species identification and examine phylogenetic relationships. Single-gene maximum likelihood trees that proved concordant were combined to produce a concatenated, multigene maximum likelihood tree. All gaps were treated as missing values. Support for the tree topology was assessed via bootstrap analysis with 1,000 replications. Consensus ITS, Cox-1, and Cox-2 sequences were deposited in the GenBank database under the accession numbers MT819392 to MT819428 and MT940649 to MT940652 (ITS), MT981111 to MT981147 (Cox-1), and MW006634 to MW006666 (Cox-2). The multigene tree is deposited in TreeBASE (submission 26710). Representative isolates of all Pythium species were lodged with the National Collection of Fungi, Bundoora Herbarium, under the reference numbers VPRI43923 to 43947.

Morphology.

In cases where the molecular data did not provide certainty of identification (i.e., for Pythium sp. nov., P. dissotocum, and P. catenulatum), cultures were also examined morphologically. Plugs (5 to 10 mm²) of 1- to 2-week-old Pythium cultures were floated in Petri plates with sterilized soil extract water (Jeffers 2006), and boiled pieces of grass (Pennisetum clandestinum) or sterile hemp seeds were added to the water. These floating cultures were maintained at room temperature (22 to 24°C) on a bench and checked after 3 days, and each day thereafter, for taxonomic features. Examinations were made with a DM2900 Leica compound microscope under various magnifications (100× to 400×). Petri plates were observed directly, and then mycelial mats of interest were removed with tweezers and mounted in water or toluidine blue dye for examination in reference to morphological keys (Pennsylvania State University 2021; Van der Plaats-Niterink 1981).

Colony growth rates.

Daily mycelial growth was measured at 2.5°C intervals between 7.5 and 40°C. Radial growth was measured for one representative isolate of each of nine Pythium species. Plugs (0.5 cm²) of 1-week-old cultures were subcultured onto PDA contained in 9-cm Petri plates. The plugs were placed 2 mm from the edge of the Petri plate so that the fast-growing species did not cover the plate too quickly. After culturing, the plates were left overnight (approximately 14 h) at 23°C because earlier trials showed that initial growth after subculturing was sometimes slower than subsequent growth (results not shown). The edge of each colony was then marked with a fine permanent pen. Plates were incubated for a further 24 h at the temperature of interest under UV light. The edge of the colony was then marked again, and the growth between the two lines was measured at two points. We calculated daily growth by averaging the two measurements. There were three replicate plates per isolate, and the experiment was repeated three times.

Pathogenicity tests.

Pythium isolates.

Although 11 species of Pythium were originally isolated and identified, the growth rate tests and pathogenicity tests (in vitro and pot) included only nine species because cultures of the other two species (P. rhizosaccharum and P. catenulatum) died. Tests included one representative isolate of each Pythium species unless otherwise stated.

In vitro pregermination pathogenicity test.

Tomato seeds of five processing cultivars (H1015, H1175, H2401, H3402, and H4401) were surface-sterilized by immersion in 80% ethanol for 30 s, 25% sodium hypochlorite (1% a.i. HOCl) for 2 min, and twice in H₂O for 1 min. Three seeds were then planted into each Petri plate, which contained water agar entirely colonized by 1- or 2-week-old samples of each Pythium culture.

The seeds were assessed for disease severity 10 days after planting. The severity rating score was from 0 to 5, where 0 = seed germinated and symptomless; 1 = seed germinated and seedling shows a few light brown lesions on the radicle, shoot asymptomatic or slightly stunted; 2 = seed germinated and seedling shows brown, enlarging or coalescing lesions on radicle, shoot present but clearly stunted and unthrifty; 3 = seedling died after germination, radicle length >3 mm, radicle brown and necrotic, shoot not emerged or partially emerged but leaves still encased in seed; 4 = seedling died shortly after germination, radicle <3 mm, radicle dark brown and shoot not emerged; 5 = seed failed to germinate and discolored.

Logistic regression of the ordinal response variable of disease severity score, with the factors of isolate, host cultivar, and isolate–host interaction, was performed via the logistic procedure of SAS version 9.4 (SAS Institute, Cary, NC). This model used the nonproportional adjacent-category logit link function, which generates separate log odds ratios (rather than cumulative log odds ratios) for the response categories (Agresti and Tarantola 2018).

Pot trials.

All pot trials took place in a temperature-controlled glasshouse with natural light. The ambient temperature ranged from 22 to 25°C, and day length was approximately 12 h. In each experiment, the pots were arranged in a completely randomized design on the bench, and moisture was maintained at field capacity.

Experiments were conducted in twice-pasteurized potting mix (composted bark, 90%; and sand, 10%) or in twice-pasteurized soil (Victorian red clay soil, 90%; sand, 8%; and perlite, 2%). The inoculum and pot setup procedures were the same for all trials and were as follows. Pythium sp. inoculum was prepared by soaking Trill Budgerigar millet seed mix (Trill, Australia) in tap water for 16 h. The millet was strained and transferred into 500-ml bottles, then autoclaved three times at 121°C for 20 min. Each bottle of medium was inoculated with 10 agar squares (2 cm²) cut from a 1-week-old culture. The bottles were shaken to integrate the plugs and were then incubated at 25°C under continuous light until the millet was fully colonized (2 to 4 weeks).

Sterile pots (1.5 liter capacity) were filled with 5% (vol/vol) colonized millet, mixed thoroughly with twice-pasteurized soil or potting mix. The control treatments contained the equivalent amount of sterile millet mixed through soil or potting mix. Two 3-week-old tomato seedlings (cultivar H3402) were carefully transplanted into each pot.

The postgermination damping-off assessment was conducted four times (twice in potting mix and twice in soil), and the postgermination pathogenicity assessment was conducted three times in soil only. Potting medium was included as a factor in the statistical analyses of the postgermination damping-off test.

Postgermination damping-off.

Seven days after planting, postgermination damping-off was assessed, where seedlings were rated as either alive or dead. Binary logistic regression was performed initially with the factors of trial, pot, species, isolate, and various interactions between them via the logistic procedure in SAS version 9.4 (SAS Institute, Cary, NC). Factors were tested for significance (maximum likelihood) and were kept in the model if significant (P < 0.05), and if they were not significant (P > 0.05) they were omitted from subsequent iterations of the model. The binary response variable was seedling outcome (dead, 0; or alive, 1).

Postgermination pathogenicity.

One week after planting, pots were harvested so that only one seedling remained in each pot. One month after inoculation, plants were harvested and washed gently in tap water, with care taken not to damage the roots. Root rot severity was rated for each plant on a scale of 0 to 4 where 0 = roots white colored and healthy in appearance (<5% necrotic), 1 = roots mildly necrotic (6 to 20% necrotic), 2 = moderately necrotic (21 to 55% necrotic), 3 = severely necrotic (56 to 89% necrotic), and 4 = very severely necrotic (90 to 100% necrotic).

Roots were then cut from shoots. Three 5-mm pieces of lateral root tissue were excised from each root system along with a 3-mm piece of collar tissue for culturing onto PDA-S to confirm Pythium infection. Roots and shoots were then placed in the oven at 70°C for 48 h for dry root and shoot weight measurement.

Height and dry-weight data were analyzed on Minitab 19 via Welch’s analysis of variance and the Games–Howell post hoc test, which does not assume equal variances and sample size. The root rot severity data scores were converted into root disease indices (DI) via the method described by Foster et al. (2017) with the following formula:

where a, b, c, d, and e are the number of plants with each disease score (0 to 4). The results of the two trials were combined after statistical analysis showed results were not significantly different.

Results

Isolations and identification.

A total of 143 Pythium isolates representing 11 species were recovered from plants and soil baiting across the 20 crops surveyed over two seasons (Table 1). P. aphanidermatum, P. catenulatum, P. dissotocum, P. heterothallicum, P. irregulare, Pythium sp. nov., and P. ultimum were isolated from both plant collars and roots; P. inflatum and P. rhizosaccharum were isolated only from plant roots; P. carolinianum and P. recalcitrans were isolated only by soil baiting.

Table 1. Pythium species isolated from processing tomato plants and soil during surveys between 2016 and 2018

Speciesw	Total number of isolatesx	Number of sites Pythium sp. was isolated	Number of soil samples or plants from which Pythium sp. was isolated	Isolation mediay	Originz
P. aphanidermatum	19	3	9	PDA-S, V8-PAR	SR, LR, LS
P. carolinianum	2	1	1	Rose petals	Soil
P. catenulatum	2	2	2	PDA-S	SR, LR, LS
P. dissotocum	70	14	36	PDA-S, V8-PAR, WA	SR, LR, LS
P. heterothallicum	7	4	6	PDA-S, V8-PAR	SR, LR, LS
P. inflatum	9	6	7	V8-PAR, WA	SR, LR
P. irregulare	16	5	10	PDA-S, V8-PAR, WA	SR, LR, LS
P. recalcitrans	4	1	1	Rose petals	Soil
P. rhizosaccharum	1	1	1	PDA-S	LR
Pythium sp. nov.	11	2	5	PDA-S, V8-PAR, WA	LS, SR
P. ultimum var. ultimum	2	1	1	V8-PAR, WA	LS

^wSpecies as determined by morphology or sequence-based similarity (ITS, Cox-1, and Cox-2).

^xIsolated from different pieces of plant tissue.

^yPDA-S, potato dextrose agar amended with streptomycin; WA, water agar. V8-PAR is defined in Materials and Methods.

^zLR, lateral roots/feeder roots; LS, low stem/collar region; soil, baited from soil sample as described in methods; SR, small roots/rootlets.

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In both seasons, P. aphanidermatum, P. inflatum, P. heterothallicum, P. irregulare, and a group of isolates that formed a clade with P. coloratum, P. diclinum, P. dissotocum, P. dictyosporum, P. lutarium, and P. marinum were isolated, whereas P. recalcitrans, P. carolinianum, and P. ultimum var. ultimum were isolated only in the 2016 to 2017 survey, and P. catenulatum, P. rhizosaccharum, and Pythium sp. nov. were isolated only during the 2017 to 2018 survey.

In both years, P. dissotocum was the most abundant and widespread Pythium species, with a total of 70 isolates recovered from 70% of the surveyed sites. P. aphanidermatum was the next most abundant species, with 19 isolates found, but only across 15% of sites. After P. dissotocum, P. inflatum was the next most widespread species isolated from 30% of sites, followed by P. irregulare, isolated from 25% of the sites.

Species identification was tentatively confirmed based on 99 to 100% similarity to reference isolates via Basic Local Alignment Search Tool search, with some exceptions. The identity of isolate UM1333 could not be confirmed based on ITS alone, because the sequence was equally similar to P. catenulatum, P. intermedium, P. torulosum, and P. pyrilobum. However, Cox-1 and Cox-2 sequences coupled with morphological observations confirmed that the isolate was P. catenulatum. Similarly, the identity of isolates UM519 and UM524 could not be resolved via ITS sequences, which were equally similar to P. heterothallicum and P. glomeratum. However, Cox-1 and Cox-2 sequences resolved that these isolates were P. heterothallicum.

Eleven isolates (including UM1308 and UM1007) were only 96% similar (based on ITS, Cox-1, and Cox-2) to a named Pythium species on GenBank, P. catenulatum; however, they were 99 to 100% similar to isolates of a putative new Pythium species labeled Pythium sp. nov. P8209 and Pythium sp. nov. P8207 in the phylogenetic study by Robideau et al. (2011). Attempts at morphological description of these isolates was unsuccessful because although hyphal swellings and sporangial vesicles were observed, oogonia could not be produced. Isolates UM1308 and UM1007 are hence referred to as Pythium sp. nov.

For P. dissotocum, ITS sequences were >99% similar to two to five species in most cases, including P. dissotocum, P. coloratum, P. diclinum, P. dictyosporum, P. lutarium, and P. marinum. The Cox-1 and Cox-2 sequences of these isolates matched to fewer species; however, most isolates were >98 to 100% similar to at least two of the aforementioned species. Morphological examination of these isolates led to their identification as P. dissotocum. Distinguishing features included the slightly inflated, branched filamentous sporangia, the stalked and both monoclinous and diclinous antheridia, and aplerotic oospores (Van der Plaats-Niterink 1981). P. marinum is rarely encountered and has been recovered only from water. It also has plerotic oospores. P. dictyosporum has reticulate oospores, and P. coloratum has oospores with a lilac hue. P. diclinum has diclinous antheridia and aplerotic oospores (Van der Plaats-Niterink 1981).

Phylogenetic analysis.

ITS consensus sequences were 855 bp long, the Cox-1 sequences were 706 bp long, and the Cox-2 sequences were 598 bp long. The three individual maximum likelihood trees (ITS, Cox-1, and Cox-2) containing representative Pythium isolates from these surveys and reference isolates were constructed, and species identification was confirmed (Supplementary Materials S2, S3, and S4).

The three trees were concordant, and thus sequences were concatenated to construct a multigene tree (Fig. 1). The Pythium species represented five of the 11 described clades (A, B, E, F, and I) sensu LéVesque and De Cock (2004). The five clades were each well supported, with bootstrap values of 99 to 100 for all except clade B, which was 95. Similarly, the bootstrap values for most individual species clades (containing UM isolates and reference isolates) were mostly high, at 99 to 100, excluding the P. dissotocum clade and the P. catenulatum clade. The bootstrap support for the P. dissotocum species complex as a whole was high, at 100; however, there were some subclades, and the species identity of the UM isolates was not apparent based on either the individual or concatenated gene trees.

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Colony growth rates.

Pythium species had optimal growth rates on PDA between 25 and 32°C with the exception of P. aphanidermatum, which had an optimal growth temperature of 37.5°C (Fig. 2). Specifically, the optimum growth temperature for P. heterothallicum, P. irregulare, and P. recalcitrans was 25°C; the optimum growth temperature for P. carolinianum and P. ultimum was 27.5°C; the optimum growth temperature for P. inflatum and P. dissotocum was 30°C; and the optimum growth temperature for Pythium sp. nov. was 32.5°C.

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Excluding P. aphanidermatum, most species grew slowly at 40°C for the first 24 h of incubation but died within 5 days of incubation at this temperature (data not shown). All species could grow slowly and survive at 7.5°C for ≥5 days (data not shown).

The average maximum daily growth rate for P. inflatum, P. heterothallicum, P. recalcitrans, and Pythium sp. nov. was 10 to 15 mm/24 h. The average maximum daily growth for P. dissotocum, P. ultimum, and P. irregulare was 25 to 30 mm/24 h. The average maximum daily growth for P. aphanidermatum was the highest of all species, at 42 mm per 24 h. The average maximum daily growth for P. carolinianum was lowest of all species at 7 mm/24 h. Confidence intervals for growth rate measurements were small for most species except for P. aphanidermatum, which exhibited slightly more variation between replicates.

Pathogenicity tests.

In vitro pregermination pathogenicity test.

The null hypothesis that the regression coefficients in the model are equal to zero was rejected $({\mathrm{\chi }}^2=287.06,\text{df}=44,P<0.001).$ The levels of disease severity induced by infection with different Pythium spp. were significantly different (P < 0.0001), the susceptibility of the different tomato cultivars to Pythium disease were significantly different (P = 0.0024), and there was a significant interaction between different Pythium spp. and tomato cultivars (P < 0.0001). The concordance value was 0.83, showing that the adjacent categories logistic model had high predictive power (Agresti and Tarantola 2018).

P. aphanidermatum, P. ultimum var. ultimum, P. dissotocum, and P. irregulare were the most aggressive pathogens, with infection by these species most likely to result in a disease severity score of 4 (seed death) on all tomato cultivars (Fig. 3). Pythium sp. nov. was moderately aggressive, with infection by this species most likely to result in a disease score of 3 or 4 (severe seedling stunting or seed death, respectively) on all tomato cultivars. P. carolinianum, P. heterothallicum, P. inflatum, and P. recalcitrans were weaker pathogens. Although infection often resulted in disease symptoms and could result in seedling death in some cases, disease scores of ≤3 were collectively much more likely than disease scores of 4 or 5.

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The more aggressive pathogens (P. aphanidermatum, P. dissotocum, P. irregulare, P. ultimum, and Pythium sp. nov.) tended to have largely consistent effects across the different tomato cultivars. One exception to this was the P. ultimum interaction with H4401, with this cultivar appearing significantly more tolerant of infection than all other cultivars. On the other hand, the less aggressive pathogens (P. carolinianum, P. heterothallicum, P. inflatum, and P. recalcitrans) had more variable effects on the different tomato cultivars. For example, P. carolinianum had >75% predicted probability of disease severity score 0 (no symptoms) on cultivar H1015 but <25% predicted probability of disease severity score 0 on cultivar H4401, which was apparently much more susceptible to disease caused by this organism. Similarly, P. heterothallicum was more likely to cause a disease score of 0 on H1015 and H3402 than on the other tomato cultivars (H4401, H1175, and H2401).

None of the tomato cultivars performed consistently better than others. H1015 and H3402 were more resistant when infected by P. carolinianum and P. heterothallicum, and H1015 was also most resistant of all the cultivars when infected by P. inflatum. However, H1015 and H2401 were most susceptible when infected by the Pythium sp. nov., with H4401, H1175, and H3402 more resistant. H1175 and H2401 were slightly more resistant than the other cultivars when infected by P. recalcitrans.

Pot trials.

i. Damping-off.

Initial testing identified that there was no significant effect of isolates (different isolates of the same species) and no significant interaction between species and trial (P > 0.05; data not shown), so these terms were excluded from the final model. On the other hand, the individual factors of species and trial were found to have significant effects (both P < 0.001) and so were kept as terms in the model. The four trials were analyzed collectively because although trials differed significantly, the lack of interaction between trial and species indicated that the relative pathogenicity of species was consistent across trials. The differences between trials could be explained by seasonal variation in the greenhouse and by the fact that two tests were performed in potting mix and two in soil. Throughout this series of experiments, pathogenicity in soil was generally higher than pathogenicity in potting mix.

P. ultimum and P. aphanidermatum had the highest least squares means probabilities of causing damping-off, at 88 and 73%, respectively (Fig. 4). The next most aggressive damping-off pathogens had much lower probabilities of causing damping-off and were P. irregulare (37%) and P. dissotocum (25%). Pythium sp. nov. had a 6% probability of causing damping-off. P. carolinianum, P. heterothallicum, and P. inflatum had a 1.5% probability of causing damping-off.

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ii. Postgermination pathogenicity.

One month after inoculation, when plants were 2 months old, P. aphanidermatum, P. dissotocum, P. irregulare, P. ultimum, and P. recalcitrans caused significant stunting (height reduction) compared with controls (Table 2). Significant shoot dry weight reduction was observed in plants infected by four of these species: P. ultimum, P. irregulare, P. recalcitrans, and P. aphanidermatum. The most severe effects were observed in the roots, where P. ultimum, P. irregulare, P. recalcitrans, P. aphanidermatum, P. dissotocum, and P. inflatum caused significant dry weight reduction (Table 2).

Table 2. Mean height, dry shoot weight, dry root weight, and root rot disease index of tomato plants (cultivar H3402) 4 weeks after inoculation with Pythium spp.

Treatment	Height at 4 weeks after inoculation (cm)y	Dry shoot weight (g)y	Dry root weight (g)y	Root rot disease indexz
Control	63.6 a	0.73 a	0.17 a	0.05
P. aphanidermatum	38.2 cd	0.20 b	0.02 b	75
P. carolinianum	54.8 abc	0.61 ab	0.13 ab	25
P. dissotocum	40.8 bcd	0.32 ab	0.05 b	40
P. heterothallicum	60 ab	0.66 ab	0.13 ab	20
P. inflatum	46.8 abcd	0.36 ab	0.06 b	40
P. irregulare	30.4 d	0.17 b	0.02 b	80
P. recalcitrans	33.8 d	0.20 b	0.03 b	60
Pythium sp. nov.	55 abc	0.48 ab	0.08 ab	40
P. ultimum	30.4 d	0.21 b	0.02 b	65

^yGames–Howell pairwise comparisons indicated by letters (a to d). Results that do not share the same letter are significantly different (P < 0.05).

^zRoot disease index calculated for 15 plants per treatment. Root disease index is from 0 to 100, where 0 indicates that all 15 plants were healthy with the lowest disease severity score of 0, and 100 indicates that all 15 plants had the maximum disease severity score of 4.

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Root rot disease severity largely reflected the patterns of aggressiveness apparent in quantitative data (heights and dry weights). Plants infected with P. irregulare scored the highest DI (80), followed by P. aphanidermatum (75), P. ultimum (65), and P. recalcitrans (60). Inoculation with each of these species commonly caused >56% necrosis of the roots. The DI for plants inoculated with Pythium sp. nov., P. inflatum, and P. dissotocum was 40, with infection by these species causing root disease severity scores between 0 and 3. The DIs for plants infected with P. carolinianum and P. heterothallicum were lowest, at 25 and 20, respectively. The DI for control plants was 0.5, with these plants largely exhibiting <5% root necrosis. Each of the Pythium species was successfully recovered when the inoculated plants were cultured out at the completion of the trials.

Discussion

A total of 11 Pythium species were associated with processing tomato plants exhibiting symptoms of poor growth at 20 sites in Victoria. Among the Pythium species were P. aphanidermatum, P. irregulare, and P. ultimum, well-known pathogens of tomatoes and other vegetables (Blancard 2012; Deniel et al. 2011; Jenkins and Averre 1983; LéVesque and De Cock 2004; Manorantitham et al. 2001; Robertson 1973, 1976). Some of the other species have been reported on tomatoes infrequently, including P. catenulatum (Frezzi 1956), P. dissotocum (Blancard 2012; LéVesque and De Cock 2004), and P. inflatum (Verma 1987). This is the first report of P. heterothallicum being recovered from tomatoes globally. None of the 11 Pythium spp. have previously been reported in association with field tomatoes in Australia; however, this is probably because of a paucity of relevant studies.

The limited reports of Pythium root rot disease on tomatoes globally could indicate that Pythium species are not commonly associated with tomatoes or that they are considered of minor economic importance. Studies of Pythium disease on other horticultural crops, including some solanaceous species, showed that they could become damaging under particular conditions (Chellemi et al. 2000; Pivonia et al. 2012; Stirling et al. 2004). In the case of Victorian processing tomatoes, there are several biotic, environmental, and agronomic factors potentially conducive to Pythium disease.

Pythium is commonly identified in soilborne disease complexes, where the coinfection of multiple pathogens increases disease potential. In this study, the aggressive Pythium species such as P. irregulare, P. aphanidermatum, and P. ultimum were found in low numbers compared to the moderately aggressive P. dissotocum and mild pathogen P. inflatum, for instance. This indicates that although Pythium may be contributing to the poor condition of the tomatoes, it is unlikely to be the sole cause of disease. It is probable that Pythium species are coinfecting with other soilborne pathogens such as F. oxysporum, which was recovered from sampled roots and collars at a high frequency (Callaghan et al. 2019). A synergistic relationship between Pythium and Fusarium has been noted on other crops (Foster et al. 2017; Harvey et al. 2008; Tesoriero 2011; You et al. 2000).

In terms of environmental factors, soil type, which in much of the region is a gray self-mulching clay, has a tendency for waterlogging (Ashcroft et al. 2001; Flett 1986). Waterlogging is the best known abiotic precursor to Pythium root rot (Akram et al. 2019; Chérif et al. 1997; Hendrix and Campbell 1973; Lookabaugh et al. 2017). In addition, under SDI, plant roots may be in saturated, hypoxic soils for prolonged periods because of the high irrigation frequency and flow rates (Brown 2016; Feld et al. 1990; Yong et al. 2017). Damaged roots can release exudates that are attractive to oomycete zoospores.

Other potential factors contributing to an environment conducive to Pythium disease include the increased intensity of cropping, leading to a buildup of inoculum over successive years (Bennett et al. 2012); annual application of metam sodium as a soil fumigant, which can be detrimental for soil microbial communities and lead to the rise of quick-colonizing pathogen species such as Pythium (Hendrix and Campbell 1973); and planting rotation crops such as wheat and maize, which are also hosts for many of the Pythium species identified in this study (Van der Plaats-Niterink 1981).

The Pythium species were most damaging when plants were inoculated before germination (in vitro) compared with when they were inoculated postgermination (pot trials). This may have been because the in vitro inoculation technique was more severe, with seeds being planted directly into a culture compared with a pot where contact with the pathogen would not necessarily have been immediate. In addition, younger plant tissue is generally more susceptible to Pythium infection (Chun and Schneider 1998). The increased susceptibility of younger seedlings was also observed in the pot trials, where plant death occurred only in the first 10 days after inoculation. This observation has implications for management because it suggests that a delay in transplanting seedlings could significantly increase the seedlings’ resistance to some pathogens. Additionally, it confirms that Pythium management strategies would be best focused early in the season, when the impact of the disease is highest.

The Pythium species varied significantly in their aggressiveness. Although the in vitro test results may have magnified the severity of the potential Pythium disease when compared with the pot test, the patterns of relative aggressiveness across the tests and plant growth phases were largely consistent. P. aphanidermatum, P. ultimum, and P. irregulare were the most aggressive; P. dissotocum, Pythium sp. nov., P. recalcitrans, and P. inflatum were in the middle of the spectrum; and P. heterothallicum and P. carolinianum were in the weakly aggressive to avirulent group. This pattern was the same across all pathogenicity tests, validating the in vitro test method, which was still able to convey the range in aggressiveness despite the direct and harsh manner of the seed inoculation. Previous studies have used a similar method to test Pythium pathogenicity on other hosts in vitro (Le et al. 2016). In vitro seed testing might be a helpful tool for tomato breeders when they screen for susceptibility to pathogens.

The five tomato cultivars differed significantly in their susceptibility to infection by the Pythium species. Although they were almost equally susceptible to the aggressive Pythium spp., they varied in their response to infection by the mild and moderate pathogens. This variation in responses between cultivars and within cultivars in reaction to different Pythium species demonstrates that the trait of tolerance to Pythium infection is probably dictated by a complex of genetic factors rather than a single gene. Unfortunately, none of the cultivars were consistently better performing. Hence, decisions about Pythium tolerant cultivars would need to be based on knowledge of which Pythium species predominated in the field before planting. However, further studies are recommended to verify the potential tolerance of processing tomato cultivars to Pythium disease, including pot and field trials.

Although Pythium disease was more severe at early stages of plant growth, surveys and the pot trials suggested some species can also cause disease of mature tomato plants. In glasshouse trials, when the plants were 2 months old, P. aphanidermatum, P. ultimum, P. irregulare, P. recalcitrans, P. inflatum, and P. dissotocum caused significant root weight reduction, and the first four species also caused significant shoot weight reduction. These patterns of aggressiveness were consistent with the in vitro trial and in the damping-off pot trial except for the severity of root disease caused by P. recalcitrans, which was only a mild to moderate pathogen of seedlings. P. recalcitrans was discovered recently on grapevine roots in Spain (Moralejo et al. 2008). It has since been reported as a pathogen of other crops, including cucumber and capsicum in Australia (Stirling 2013; Tesoriero 2011), and carrot (Lu et al. 2013), tobacco (Bian et al. 2016), alfalfa (Berg et al. 2017), and soybean (Radmer et al. 2017) in other countries, but the true extent of its host range is yet to be determined. The fact that it appears to cause more serious disease on mature tomato plants than on seedlings is unusual for a species in this genus and is worthy of further study. The severity of P. dissotocum infection was also noteworthy, given its abundance in surveys. P. dissotocum was first identified in 1930 on sugarcane (Drechsler 1930) and has since been reported as a minor or major pathogen on a large number of hosts including wheat and maize, two of the major rotation crops for tomatoes, and many other plant species (Van der Plaats-Niterink 1981). A study examining the variable effects of the different rotation crops on Pythium inoculum levels in the soil might also be worthwhile. However, the Pythium species identified here have differing host ranges, which would probably influence the efficacy of crop rotation as a management approach.

In conclusion, our findings indicate that Pythium species may be contributing to the poor establishment, stunting, root and collar rot, and yield loss observed in processing tomatoes in Victoria.

Acknowledgments

The authors gratefully acknowledge the Australian Processing Tomato Research Council, the assistance of Liz Mann and Ann Morrison with surveys, and the support of growers for allowing us access to their crops.