Skip to main content
  • Loading metrics

Variability in an effector gene promoter of a necrotrophic fungal pathogen dictates epistasis and effector-triggered susceptibility in wheat

  • Evan John,

    Roles Formal analysis, Investigation, Methodology, Writing – original draft, Writing – review & editing

    Affiliations Centre for Crop and Disease Management, Curtin University, Bentley, Perth, Western Australia, Australia, Curtin University, Bentley, Perth, Western Australia, Australia

  • Silke Jacques,

    Roles Investigation, Methodology, Writing – review & editing

    Affiliations Centre for Crop and Disease Management, Curtin University, Bentley, Perth, Western Australia, Australia, Curtin University, Bentley, Perth, Western Australia, Australia

  • Huyen T. T. Phan,

    Roles Formal analysis, Writing – review & editing

    Affiliations Centre for Crop and Disease Management, Curtin University, Bentley, Perth, Western Australia, Australia, Curtin University, Bentley, Perth, Western Australia, Australia

  • Lifang Liu,

    Roles Investigation, Writing – review & editing

    Affiliations Centre for Crop and Disease Management, Curtin University, Bentley, Perth, Western Australia, Australia, Curtin University, Bentley, Perth, Western Australia, Australia

  • Danilo Pereira,

    Roles Data curation, Resources, Writing – review & editing

    Current address: Environmental Genomics, Max Planck Institute for Evolutionary Biology, Ploen, Germany.; Environmental Genomics, Christian-Albrechts University of Kiel, Kiel, Germany.

    Affiliation Plant Pathology, Institute of Integrative Biology, ETH Zurich, Zurich, Switzerland

  • Daniel Croll,

    Roles Formal analysis, Resources, Writing – review & editing

    Affiliation Laboratory of Evolutionary Genetics, Institute of Biology, University of Neuchâtel, Neuchâtel, Switzerland

  • Karam B. Singh,

    Roles Supervision, Writing – review & editing

    Affiliations Centre for Crop and Disease Management, Curtin University, Bentley, Perth, Western Australia, Australia, Agriculture and Food, Commonwealth Scientific and Industrial Research Organisation, Floreat, Western Australia, Australia

  • Richard P. Oliver,

    Roles Supervision, Writing – review & editing

    Affiliation Curtin University, Bentley, Perth, Western Australia, Australia

  • Kar-Chun Tan

    Roles Conceptualization, Formal analysis, Funding acquisition, Project administration, Supervision, Writing – original draft, Writing – review & editing

    Affiliations Centre for Crop and Disease Management, Curtin University, Bentley, Perth, Western Australia, Australia, Curtin University, Bentley, Perth, Western Australia, Australia


The fungus Parastagonospora nodorum uses proteinaceous necrotrophic effectors (NEs) to induce tissue necrosis on wheat leaves during infection, leading to the symptoms of septoria nodorum blotch (SNB). The NEs Tox1 and Tox3 induce necrosis on wheat possessing the dominant susceptibility genes Snn1 and Snn3B1/Snn3D1, respectively. We previously observed that Tox1 is epistatic to the expression of Tox3 and a quantitative trait locus (QTL) on chromosome 2A that contributes to SNB resistance/susceptibility. The expression of Tox1 is significantly higher in the Australian strain SN15 compared to the American strain SN4. Inspection of the Tox1 promoter region revealed a 401 bp promoter genetic element in SN4 positioned 267 bp upstream of the start codon that is absent in SN15, called PE401. Analysis of the world-wide P. nodorum population revealed that a high proportion of Northern Hemisphere isolates possess PE401 whereas the opposite was observed in representative P. nodorum isolates from Australia and South Africa. The presence of PE401 removed the epistatic effect of Tox1 on the contribution of the SNB 2A QTL but not Tox3. PE401 was introduced into the Tox1 promoter regulatory region in SN15 to test for direct regulatory roles. Tox1 expression was markedly reduced in the presence of PE401. This suggests a repressor molecule(s) binds PE401 and inhibits Tox1 transcription. Infection assays also demonstrated that P. nodorum which lacks PE401 is more pathogenic on Snn1 wheat varieties than P. nodorum carrying PE401. An infection competition assay between P. nodorum isogenic strains with and without PE401 indicated that the higher Tox1-expressing strain rescued the reduced virulence of the lower Tox1-expressing strain on Snn1 wheat. Our study demonstrated that Tox1 exhibits both ‘selfish’ and ‘altruistic’ characteristics. This offers an insight into a complex NE-NE interaction that is occurring within the P. nodorum population. The importance of PE401 in breeding for SNB resistance in wheat is discussed.

Author summary

Breeding for durable resistance to fungal diseases in crops is a continual challenge for crop breeders. Fungal pathogens evolve ways to overcome host resistance by masking themselves through effector evolution and evasion of broad-spectrum defense responses. Association studies on mapping populations infected by isolate mixtures are often used by researchers to seek out novel sources of genetic resistance. Disease resistance quantitative trait loci (QTL) are often minor or inconsistent across environments. This is a particular problem with septoria diseases of cereals such as septoria nodorum blotch (SNB) of wheat caused by Parastagonospora nodorum. The fungus uses a suite of necrotrophic effectors (NEs) to cause SNB. We characterised a genetic element, called PE401, in the promoter of the major NE gene Tox1, which is present in some P. nodorum isolates. PE401 functions as a transcriptional repressor of Tox1 and exerts epistatic control on another major SNB resistance QTL in the host. In the context of crop protection, constant surveillance of the pathogen population for the frequency of PE401 in conjunction with NE diversity will enable agronomists to provide the best advice to growers on which wheat varieties can be tailored to provide optimal SNB resistance to regional pathogen population genotypes.


Breeding for effective resistance to fungal diseases in crops is a continual challenge. Fungal pathogens, namely biotrophs and hemibiotrophs, evolve ways to overcome host resistance by masking themselves to evade pathogen-triggered immunity. This can occur through mutations in Avr-effector genes, leading to the loss of recognition and the subsequent breakdown of host resistance that ultimately results in ‘boom and bust’ cycles [1]. This is manifested through the mass adoption of a resistant cultivar (boom). The pathogen population exposed to the resistant gene will then adapt by selecting favourable mutations in the Avr gene that leads to a loss of host recognition and resistance (bust). Notable examples occur in the rust and blackleg pathosystems [2,3]. In contrast, necrotrophic fungal pathogens belonging to the Pleosporales such as the tan spot of wheat fungus Pyrenophora tritici-repentis (Ptr) and the septoria nodorum blotch (SNB) of wheat fungus Parastanonospora nodorum secrete host-specific necrotrophic effectors (NEs) to cause tissue necrosis on host plants possessing a matching dominant susceptibility gene. This results in effector-triggered susceptibility, enabling the pathogen to proliferate [4,5]. Improvements in host resistance to necrotrophic fungal pathogens rely on the removal of the host dominant-susceptibility genes [6]. However, necrotrophic fungal pathogens often possess multiple NEs that exploit different dominant-susceptibility genes carried by the host [7]. Hence, these functional redundancies often require breeders to remove multiple host genes or stack the required number of desirable disease resistance alleles to be effective [8,9]. These situations make it difficult for breeders to provide long-lasting and durable resistance.

P. nodorum utilises proteinaceous necrotrophic effectors (NE)s to cause necrosis on host plants with a susceptible genotype, resulting in necrosis that facilitates host infection [1012]. Genetic evidence has revealed at least 10 NE-dominant susceptibility gene interactions in the P. nodorum-wheat pathosystem. These are ToxA-Tsn1 [13,14], Tox1-Snn1 [15,16], Tox267-Snn2/Snn6/Snn7 [17], Tox3-Snn3B1/Snn3D1 [18,19], Tox4-Snn4 [20], Tox5-Snn5 [21] and Tox2A-Qsnb.cur–2AS1 [22]. Of the NEs, genes for ToxA [13], Tox1 [16], Tox3 [18], Tox267 [17] and Tox5 [21] have been cloned and characterised for their roles in virulence. These NEs are secreted, generally have a high cysteine content and are relatively small in molecular mass (<30kDa). Of the host dominant-susceptibility receptors, the genes for Tsn1 [14], Snn1 [15] and Snn3D1 [19] have been cloned. These genes encode proteins similar to canonical nucleotide-binding leucine-rich repeat resistance gene receptors that typically possess a protein kinase domain coupled with nucleotide-binding leucine-rich repeats or major sperm protein domains. Under controlled settings, the removal of a NE or the host receptor from the pathosystem results in the loss of ETS and a drastic reduction of P. nodorum virulence on wheat [eg. 13,16,18].

SNB resistance is quantitatively inherited [23] and improvements in resistance have been, in part, mediated through the removal of dominant susceptibility genes in the host [6]. However, the situation is complex. Multiple NEs expressed by P. nodorum afford the pathogen functional redundancies on wheat varieties that possess multiple dominant susceptibility genes [8,24]. For instance, Tan et al. [8] demonstrated that the deletion of ToxA, Tox1 and Tox3 in P. nodorum SN15 did not significantly alter virulence on a diverse collection of modern wheat varieties [8,9]. A further complication in breeding for SNB resistance is that some NE-host gene interactions were not consistently detected in field trials using mapping populations with known susceptibility genes [25,26]. It is intuitive to hypothesise that NE epistasis plays a significant role in shaping the variations observed in host resistance to SNB (Table 1) [27]. NE epistasis can be defined as interactions between NE genes where the effect conferred by one is masked by the presence of another [27]. NE epistasis in the P. nodorum-wheat pathosystem can be divided into two broad regulatory categories. Firstly, the contribution of a NE-host dominant susceptibility gene interaction is suppressed by another competing NE-host gene interaction possibly through host-gene action or cross-talk among NE recognition pathways [eg. 28,29]. Secondly, NE gene repression mediated by the expression of another NE [eg. 22,30]. For the latter, a clear-cut example involves the suppression of the Tox3-Snn3 interaction in SNB by Tox1-Snn1 described by Phan et al. [22]. This was explored as part of a study that mapped SNB on a double-haploid (DH) wheat population derived from two major Australian commercial wheat cultivars. SNB quantitative trait loci (QTL) detected during infection by the Australian P. nodorum reference isolate SN15 were compared with a Tox1 deletion mutant (tox1-6) [22]. This revealed the Tox1-Snn1 NE-host receptor interaction was epistatic to Tox3-Snn3B1 and a major SNB QTL located on chromosome 2A in the DH population that confers sensitivity to a novel NE which we coined as Tox2A. Quantitative RT-PCR revealed that Tox3 expression was significantly higher in tox1-6 than in the wildtype, which presented a basis for NE epistasis mediated by gene repression [22].

Table 1. Epistasis of P. nodorum NE-wheat dominant susceptibility gene interactions.

*RI, recombinant inbred; DH, doubled haploid; ITMI, International Triticeae Mapping Initiative.

The expression of Tox1 also varies among isolates. As Tox1 production was much higher in SN2000 than both SN4 and SN6, the former was used to enable its detection and functional characterisation [16,32,33]. This contrasts with other USA isolates such as SN4 and SN6 where Tox1 expression is low relative to SN2000 [34]. A recent study also demonstrated that the Australian P. nodorum SN15 isolate expressed Tox1 significantly higher than SN4, which correlated with a greater SNB disease contribution mediated by the Tox1-Snn1 interaction [31]. The mechanisms behind this differential expression remain unexplored however, it was noted that the Tox1 promoter region was found to be polymorphic between these isolates [31]. These observations prompted us to ask several key questions. Firstly, does the genetic polymorphism in the promoter region drive differential Tox1 expression and affect epistatic outcome of Tox3-Snn3B1 and Tox2A-Qsnb.cur–2AS1 interactions. Secondly, what is the possible mechanism of Tox1 regulation? Thirdly, is there a fitness penalty associated with lower Tox1 expression in P. nodorum isolates? And lastly, what is the international distribution of Tox1-expressing P. nodorum variants? The study presented here was undertaken to characterise the gene regulatory elements controlling Tox1 expression in P. nodorum and explore their role in NE epistasis with end-goals of 1. improving SNB management strategies in wheat growing regions where the disease is prominent and 2. to provide an insight into a mechanism of NE epistasis that may dictate the outcome of fungal-host plant pathosystems.


Genetic variability in the Tox1 promoter region

To determine why both Tox1-mediated epistasis and Tox1 expression differed between P. nodorum isolates, we aligned Tox1 promoters sequences from a geographically diverse set of 20 P. nodorum genomes [3537] (Fig 1A). The alignment revealed a 401 bp genetic element that was absent in four isolates within the collection. This element, henceforth called PE401 was absent in the Australian P. nodorum SN15 and SN2000 isolates that were associated with a higher level of Tox1 expression (Fig 1A) [16,31,34]. The size and proximity to the gene within the Tox1 promoter suggested PE401 was the most plausible variant involved in gene repression, despite the presence of other smaller genetic polymorphisms.

Fig 1. Genetic variation in the Tox1 promoter region.

A. An alignment highlighting promoter variants in a geographically diverse selection of 20 Tox1-containing P. nodorum isolates. Within the 1,500 bp promoter region, the 401 bp PE401 and a 14 bp GTTTTCGGCCGTAT tandem repeat polymorphism are indicated. B. A GC% plot using a 50 bp average sliding window covering the Tox1 promoter from the USA reference isolate SN4. PE401 has a low GC% (blue) relative to directly adjacent promoter regions. AT% (green) is also indicated. C. A dot plot of PE401 using a 5 bp sliding window demonstrating the paucity of tandem/inverted repeat stretches. Genome coverage for of the promoter region of Tox1 in NOR-4 is incomplete.

PE401 was then examined for features of a mobile genetic element (MGE), as they are known to modulate the expression of proximal fungal genes [3841]. The SN4 promoter sequence was queried against the Repbase and Rfam databases. No matches to characterised MGEs or expressed non-coding elements were observed. BLAST analysis of PE401 identified a single region with 63% identity in the SN15 genome, partially overlapping the annotated gene SNOG_30065 [35]. However, the SNOG_30065 predicted peptide sequence did not contain annotated protein domains, nor was the gene expressed under in vitro conditions and during host infection [42]. Therefore, SNOG_30065 is likely a pseudogene. No further similar matches to PE401 were detected when queried against the NCBI nucleotide and whole-genome shotgun collections, suggesting the sequence was not a repetitive element. Terminal repeats characteristic of fungal MGEs were not identified, although the GC% was low (41.9%) relative to the rest of the SN4 promoter region (48.9%) [43] (Fig 1B and 1C). We concluded that PE401 is unlikely to be a transposable element or functional gene in P. nodorum.

Evidence of regional distribution for PE401 in the international P. nodorum population

The distribution of PE401 was determined in a international panel of 484 Tox1-containing P. nodorum isolates. It was observed that isolates carrying PE401 were ubiquitous in Europe (Switzerland, Denmark, Finland, Netherlands and Norway), most sampling sites in North America (USA and Canada) and also in Iran. The latter region was previously proposed as a centre of origin for P. nodorum NEs including Tox1 [44,45]. Isolates that lacked PE401 were dominant in Australia (87% of 180 isolates) and South Africa (100% of 22 isolates), while 27% of all isolates from North Dakota (USA) and a single isolate from Texas (USA) also lacked PE401 (Fig 2).

Fig 2. The international distribution of PE401 in the P. nodorum Tox1 promoter.

Presence/absence of PE401 in a international collection of 484 P. nodorum isolates. Spheres represent individual populations grouped based on the state/country of origin. % values represent the proportion of (-) PE401 isolates where it was identified in that population. Population and group sizes are indicated (n). Isolate details are provided in full in S1 File. The world map image was obtained from generated with ‘maps’ (

PE401 functions as a repressor of Tox1 expression

We hypothesised that PE401 dictates Tox1 expression based on the differential expression profile between SN4 (+PE401) and SN15 (-PE401) carrying the two versions of the promoter [31]. Therefore, Tox1 expression was assessed in SN4, SN15 and three representative Australian P. nodorum wildtype isolates with or without PE401 (Fig 3A). Quantitative RT-PCR analysis revealed the two Australian isolates carrying PE401, WAC13443 and WAC13072, shared a low Tox1 expression profile comparable to SN4. On the other hand, Tox1 expression was significantly higher in SN15 and WAC13690 which lacked PE401 (Fig 3B). Sequence alignment of the promoter region revealed WAC13443 and WAC13072 does not carry the 14 bp tandem repeat found in SN4, suggesting that PE401 was the variant responsible (Fig 3A).

Fig 3. Sequence specificity of the 401 bp PE401 dictates Tox1 repression.

A. Alignment of the promoter regions comparing the USA reference isolate SN4 with four Australian isolates +/- the 401 bp PE401. B. Tox1 expression [2dCt(Tox1-Actin)] of P. nodorum wildtype isolates +/- PE401 in vitro. Error bars indicate standard deviations from biological triplicates. Letters not connected by the same letter are significantly different (P < 0.05) based on ANOVA. C. Schematic overview of the three Tox1 promoter replacement strategies with (+), without (-) or using a spacer (S) 401 bp sequence into SN15 and WAC13443 (WAC) strains (S2 File). D. Tox1 expression (2dCt(Tox1-Actin)) in SN15 and WAC mutant strains carrying promoter replacements grown in vitro. Error bars indicate standard deviations from combined averages (n = 2) of independently verified isogenic promoter replacement mutants. Gene expression levels not connected by the same letter are significantly different (P < 0.05) based on ANOVA.

We then created isogenic strains with and without PE401 to eliminate the possibility of strain-specific genetic factors located outside of the Tox1 locus involved in gene repression. The constructs used for promoter replacement included 873 bp of the Tox1 promoter either with PE401 (+), without PE401 (-) or with a 401 bp spacer (S) (Fig 3C). These were independently transformed into SN15 and WAC13443 at the Tox1 locus through homologous recombination. Quantitative RT-PCR analysis of the mutants revealed a 40-fold (SN15) and 38-fold (WAC13443) reduction in Tox1 gene expression occurred in the presence of PE401 when grown in Fries3 broth in vitro. Tox1 was not repressed in mutants that carry the spacer sequence, suggesting the effect was attributable to sequence specificity conferred by PE401 (Fig 3D). We concluded that PE401 is associated with Tox1 repression, perhaps through an interaction with a trans-acting regulatory factor.

We then determined the regulatory role of PE401 in Tox1 expression during host infection using qRT-PCR. Infection time points were chosen to represent major stages of infection such as penetration (3 dpi), colonisation (5 dpi) and sporulation (7 dpi) [46] (Fig 4). As expected, Tox1 expression was much lower (log power) in SN15 and WAC13443 mutants carrying PE401 than mutants carrying either the spacer sequence or lacking PE401 throughout the entire infection period (Fig 4).

Fig 4. PE401 represses Tox1 expression during host infection.

Quantitative RT-PCR determination of Tox1 and Tox3 expression in promoter replacement mutants of SN15 and WAC13343 sampled at three (early penetration), five (colonisation) and seven (sporulation) days post-infection of wheat cv. Halberd. The average values [2dCt(Target-Actin)] of two biological replicates were used. Error bars indicate standard deviations.

It was previously observed that deletion of Tox1 in SN15 caused a significant increase in Tox3 expression [22]. Therefore, we hypothesised that the inclusion of PE401 would alleviate Tox3 repression by Tox1. However, the decrease in Tox1 expression conferred by PE401 did not affect Tox3 expression (Fig 4). This indicated Tox1 repression caused by PE401 was insufficient to alleviate epistasis of Tox3 and that full suppression of Tox1 expression is needed to achieve this.

PE401 influences Tox1-Snn1-mediated ETS in wheat

The contribution of Tox1-Snn1 to SNB symptoms caused by SN15 (+) and SN15 (-) mutant strains were determined using an association mapping approach. We used a Calingiri (Snn1, snn3b1, tsn1) and Wyalkatchem (snn1, Snn3B1, tsn1) (CxW) DH wheat population infected with mutant spores at the seedling stage to directly compare the effect of PE401 repression on Tox1-Snn1 and other important SNB interactions. SNB seedling resistance QTL can be detected in the CxW population on chromosomes 1BS (Snn1), 5BS (Snn3B1), 2A, 2D, 3A and 4B when infected with SN15 or its derivative NE deletion mutants [22]. The seedling infection assay revealed the average disease score on the 179 wheat lines (including parental lines) for the SN15 (+) mutant was 4.4. This is significantly lower than SN15 (-) at 4.8 (Table 2). We then compared the SNB severity of SN15 (+) and (-) on genotype combinations that possess Snn1 and observed that the former isolates were significantly more pathogenic (Table 2). In contrast, no significant difference was observed between SN15 (-) and SN15 (+) on snn1 wheat lines.

Table 2. Seedling SNB rating of the CxW DH population grouped by Tox1 and Tox3 sensitivity conferred by Snn1 and Snn3B1, respectively.

SN15 isogenic mutants with (+) and without (-) PE401 were used to assess seedling wheat infection. Mean SNB scores and SD were determined for both promoter replacement mutants. A paired Student’s T-test (n = 3) was used to compare the mean between strains for each wheat genotype groups. *Indicates significant differences between the mutants at the P<0.05 significance threshold. Disease scores for all lines are provided in S3 File.

There was no difference in disease severity on Snn3 versus snn3 wheat for either mutant, which had previously been observed for the tox1 mutant compared to SN15 [22]. This indicated that Tox1 remained epistatic to Tox3 when expressed at the lower level conferred by the PE401. Reliable markers for the 2A, 2D, 3A and 4B resistance loci are not yet available which meant the disease severity could not be directly compared between the SN15 (-) and (+) mutants. Nevertheless, the results suggest that the Tox1-Snn1 interaction contributed significantly to SNB severity, and that higher expression of Tox1 conferred by the absence of PE401 produced a significantly higher level of disease on Snn1 wheat.

The absence of PE401 results in higher biomass on Snn1 wheat lines

We hypothesised that P. nodorum isolates lacking PE401 resulting in higher Tox1 expression are fitter on Snn1 wheats. However, low Tox1-expressing P. nodorum isolates that possess PE401 are maintained in the population pool at a low frequency, S1 Fig and [47]. To determine evidence of fitness penalty on Snn1 wheat, we used a digital PCR approach to compare the DNA biomass of SN15 (+) and (-) during infection of Snn1 (H086, H336 and H091) and snn1 (H213 and H324) wheat varieties deriving from the CxW population that lacked Tox3 and Tox2A sensitivity (Fig 5). Fungal biomass was determined at two and four dpi where Tox1 is maximally expressed. For snn1 wheat varieties infected with either SN15 (+) or (-), there was no significant difference (P > 0.05) between the biomass of SN15 (+) and (-) at two or four dpi, with an expected increase of biomass over time. For wheat varieties with Snn1, the biomass of SN15 (-) was significantly higher than SN15 (+) at two and four dpi (Fig 5). We then co-infected all five wheat varieties with an equal ratio of SN15 (+) and (-) to determine if SN15 (-) would outcompete SN15 (+) during infection on Snn1 wheats using a basic De Wit replacement series of 50:50 input replacement combining SN15 (-) and (+) [48]. We observed the proportion of SN15 (+) and (-) biomass was near-identical at both time points in contrast to the initial hypothesis. This indicates that SN15 (-) assisted SN15 (+) infection by driving Tox1-Snn1 ETS of the host through elevated Tox1 expression. Moreover, in two Snn1 wheat lines (H086 and H336) the total biomass at four dpi of co-infection was slightly but significantly elevated (P < 0.05) compared to SN15 (-) (Fig 5). This suggests that SN15 (-) assists SN15 (+) in host colonisation possibly through a mutualistic interaction.

Fig 5. Digital droplet PCR analysis of SN15 (+) and (-) the 401 bp PE401 biomass during infection of wheat varieties with and without Snn1 (bar graph) at 2 and 4 dpi where Tox1 expression is maximal.

Relative biomass was also determined on a 50:50 co-infection of SN15 (+) and (-) (stacking bar graph). All lines selected were insensitive to Tox3 (snn3b1) and P. nodorum toxa13 culture filtrate containing Tox2A [22]. A one-way ANOVA with posthoc Tukey-HSD testing was used to identify significant (P < 0.05) differences across treatments of day two (lowercase letter) and four (uppercase letter). Letters not connected by the same letter are significantly different.

Tox1 repression by PE401 alleviates epistasis of a major SNB QTL on chromosome 2A

Tox1 is epistatic to the SNB QTL Qsnb.cur–2AS1 detected on chromosome 2A in addition to Tox3-Snn3B1 [22]. Since neither the NE or host receptor on chromosome 2A have been cloned, epistasis can only be studied through genetic mapping using wheat mapping populations that segregate for Qsnb.cur–2AS1. Therefore, interval mapping and QTL analysis were undertaken using the markers previously developed for the CxW population [22] to explore any differences in the disease interactions produced by the SN15 (-) and (+) PE401 mutant strains. The QTL detected on chromosome 1B, where Snn1 is located, explained the highest phenotypic contribution to SNB in the SN15 (-) mutant (47.9%) compared with SN15 (+) (30.1%) (Table 3). However, the Qsnb.cur–2AS1 SNB QTL was detected in the SN15 (+) mutant only, where it contributed 12.1% to the disease. Qsnb.cur–2AS1 was the most prominent QTL previously detected during infection with the Tox1 knockout mutant tox1-6 (S2 Fig) [22]. Despite Wyalkatchem being a donor of Snn3B1 in the CxW population [22], association mapping did not detect the Snn3B1 QTL on chromosome 5B. Therefore, we concluded that Tox1 repression by PE401 alleviated the epistatic effect on Tox2A-Qsnb.cur–2AS1 but not Tox3-Snn3B1.

Table 3. A summary of SNB QTL detected with the CxW wheat population.

Details of the flanking markers, LOD scores and phenotype contribution (R2) are indicated. Composite interval mapping of SNB QTL is provided in S2 Fig.

Promoter substitution identifies the positive regulatory region of Tox1

To characterise the core regulatory regions involved in gene expression, we generated SN15 promoter replacement mutants that carry sequentially truncated Tox1 promoter sequences (Fig 6). The shortest truncation included only the Tox1 5’ UTR (untranslated region) from the annotated genome which encompassed 86 bp upstream of the Tox1 start codon [35]. Further truncations were made at 107 bp and 157 bp upstream of the ATG, either side of a TATATAA sequence typical of a core eukaryotic TATA box element [49]. Additional truncations were made at 263 bp and 310 bp upstream to exclude/include the site of the PE401 variant. In addition, 598 bp and 873 bp upstream mutants were selected to exclude possible undiscovered regulatory elements; the 873 bp region corresponding to the entire sequence of the SN15 (-) mutant used in the preceding analyses. Furthermore, SN15 (+) mutants carrying PE401 were generated in each of the 310, 598 and 873 bp mutant backgrounds to assess any effect of the upstream elements on the repressor activity (Fig 6A).

Fig 6. Analysis of Tox1 expression regulation through promoter sequential deletions.

A. Schematic overview of the Tox1 sequential promoter deletions in SN15 either with (+) or without (-) the 401 bp PE401. Indicated are the predicted TATA box and 5’ UTR derived from the reference SN15 genome. B. Tox1 gene expression [2dCt(Tox1-Actin)] in the corresponding SN15 mutants after 72 hrs growth in Fries3 liquid culture. Error bars indicate standard deviations from the combined averages two biological replicate of two independently verified isogenic promoter replacement mutants. Letters not connected by the same letter are significantly different (P < 0.05) based on ANOVA.

Gene expression analysis revealed that rather than any single element being essential, an additive effect on Tox1 gene expression was observed by the inclusion of regions up to 310 bp, where the expression was maximal (Fig 6B). In particular, significant increases were observed by the inclusion of the putative TATA box region (107–157 bp) and the upstream region (157–263 bp). PE401 acted as a strong repressor in each of the promoter deletion mutants tested. The inclusion of longer promoter regions did not significantly increase Tox1 gene expression. This suggested that positive regulatory elements are located directly adjacent to the 3’ end of the PE401 which functions as a repressor sequence independent of upstream sequences.


In this study, we sought to characterise the gene regulatory elements controlling Tox1 expression and explore their role in NE epistasis and the virulence of P. nodorum. An important finding was the clear role in Tox1 repression conferred by PE401 located 267 bp upstream of the start codon of Tox1 (Fig 6). Analysis of the truncated promoter mutants indicated PE401 could repress the transcriptional activators driving Tox1 expression (Fig 6). The exact mechanism remains obscure, as there were no distinctive features of a non-coding element or MGE. Nonetheless, we can conclude that a novel polymorphic NE regulatory element has been identified in P. nodorum, likely targeted by sequence-specific repressor proteins.

Genetic studies indicate that field populations of P. nodorum possess a high level of genetic diversity contributed from frequent gene flow that occurs between populations in most cases. In addition, random mating, large effective population sizes and diversifying selection of virulence loci were frequently observed in P. nodorum [1,23]. Hence, it is generally considered that P. nodorum possess a high evolutionary potential that allows the fungus to adapt to environmental changes [1]. Therefore, there have been historical interests in understanding the population structure and evolutionary history of P. nodorum summarised in [23]. In a recent study, Pereira et al. [50] used a pan-genomics approach to perform a population structure analysis from an international collection of P. nodorum isolates. Four distinct genotype groups were observed consisting of Australia/South Africa, Europe/Southern US, Northern US/Iran and Oklahoma. Similarly, Phan et al. [47] analysed the population structure of Australian P. nodorum using microsatellite markers and observed significant changes in pathogen genotype and aggressiveness which coincided with shifts in the mass adoption of commercially planted wheat cultivars over four decades.

The most parsimonious explanation regarding the origins of PE401 is that the sequence was present in the ancestral Tox1 gene promoter and was subsequently lost in a subset of isolates. This stems from the fact that it was only detected in isolates outside the proposed centres of origin for SNB [44,45]. Regions with the highest proportion of isolates without PE401 were from South Africa and Australia. There is little data regarding South African wheat but it is known that Snn1 wheat in Australia has been widely sown throughout the period corresponding to the isolate collections assessed [24,47]. No detectable shifts in the frequency of the element were observed during this time 2001 and 2015 (S1 Fig). This indicates either that selection was neutral throughout this period, or that the maintenance of some variation in the population is beneficial for the pathogen. This situation is supported by the results of our co-infection assay, which demonstrated PE401 did not cause a fitness penalty in the presence of the strain lacking PE401. The other major P. nodorum population where PE401 was absent is the North Dakota (Northern US) population, albeit at a reduced frequency (27%) compared to Australia (87%) and the South African field sample (100%). In this region, it has been reported that Snn1 is prevalent in widely grown durum wheat varieties [16,51]. Interestingly, P. nodorum population structure analyses have suggested the North Dakota population is unrelated to isolates collected in Australia and South Africa, which are closely related to each other [44,50]. It is therefore possible that PE401 was lost on separate occasions in these populations. On the other hand, it is also possible that genetic exchange has occurred between the populations followed by selection. Both possibilities are speculative and require further investigation. Accordingly, if Snn1 is removed from cultivars sown in these and other regions, it will be important to monitor the frequency of the element in addition to the Tox1 gene in fungal isolates.

The infection assay using the CxW DH population demonstrated that higher expression of Tox1 conferred a higher average level of virulence on Snn1 wheat. While it is known that a fungal NE repertoire is associated with phytopathogenicity [13,16,18], the role of gene expression had not been extensively examined [52]. In the case of the tan spot of wheat fungus Pyrenophora tritici-repentis, a much higher level of ToxA expression than in P. nodorum is maintained during infection [53]. It was proposed this can compensate for the reduced potency of the PtrToxA isoform relative to P. nodorum variants [53] and demonstrates an example in another pathosystem where NE expression change can have significant disease outcomes.

The genetic mapping results comparing SN15 (-) and SN15 (+) on the CxW DH wheat population suggested removal of PE401 did not affect Tox1-Snn1 epistasis over Tox3-Snn3B1. This indicated that Tox1-Snn1 must entirely be removed before Tox3-Snn3B1 compensates for virulence. However, the Qsnb.cur–2AS1 SNB QTL was only detected by the inclusion of PE401. The physical interval for Qsnb.cur–2AS1 is notably large [22]. Nevertheless, several recent studies suggest the locus contains a gene conferring sensitivity to an undiscovered NE, that functions as an important determinant of both seedling, adult leaf and glume SNB resistance since these QTL overlap with Qsnb.cur–2AS1 [20,22,26,54]. In addition, a minor seedling resistance QTL on chromosome 2A was detected in a recombinant inbred population derived from Swiss winter wheat varieties Arina and Forno [20]. Therefore, the Tox2A-Qsnb.cur–2AS1 interaction provides a layer of NE redundancy that maintains virulence on wheat, albeit at a lower level, in the absence of other interactions such as Tox1-Snn1. In addition to SNB, a co-localised Qsnb.cur-2AS1 QTL is a major contributor to tan spot resistance in wheat [55]. Interestingly, tan spot is the most damaging necrotrophic fungal disease of wheat in Australia [56] and the fungus frequently co-infects wheat with P. nodorum [5759].

Regional P. nodorum populations differ greatly in NE gene frequency [60,61], possess high haplotype variability [45], differential isoform activity [52], variable NE expression [62; this study] and high evolutionary potential [1]. This makes SNB a difficult disease to eliminate using a common genetic resistance background in wheat across different cereal growing regions where SNB is endemic.

PE401 provides a novel example of a population variant regulating NE epistasis through gene repression. Epistasis between NE-host gene interactions was frequently observed through the use of association mapping with wheat genetic mapping populations. A complex picture has now emerged as some interactions are additive in SNB [23] while others are epistatic (Table 1). Several instances of epistasis, where one fungal NE-host receptor interaction suppresses the disease contribution of another, are now documented (Table 1). In some cases, differential NE gene expression underpins epistasis, but the regulatory elements were unclear. In one example, the expression of ToxA was reported to be two-fold higher during infection in the P. nodorum isolate SN5 compared with SN4 [62]. Higher ToxA expression correlated with a greater contribution of ToxA-Tsn1 to the disease phenotype of SN5 on the BR34 x Grandin RI wheat population that segregates for Tsn1 and Snn2. Conversely, the phenotypic contribution of Tox2-Snn2 was reduced [62]. Recently, a gene encoding the NE Tox267, which interacts with Snn2, has been cloned and functionally characterised [17]. The deletion of Tox267 in SN4 resulted in up-regulation of the NEs ToxA, Tox1, and Tox3, demonstrating further connectivity between the expression of these genes. Another study that used an RI wheat population (ITMI) segregating for Tsn1, Snn1 and Snn3 reported that the presence of ToxA in SN2000 suppressed Tox1-Snn1 ETS [31]. Gene deletion of ToxA produced only a mild reduction in disease severity, which was compensated by increased Tox1 expression. Collectively, these studies suggest gene repression as the basis for NE epistasis is a common mechanism, for which the results presented here have provided a novel mechanistic insight.

The phenomenon of NE epistasis is also observed in other fungal-plant pathosystems, dictating the contribution of effector-susceptibility/resistance gene interactions during disease [27]. P. tritici-repentis possesses a near-identical copy of ToxA to P. nodorum and infects the host through Tsn1-mediated ETS [13]. The deletion of ToxA increased the virulence of P. tritici-repentis on some Tsn1 wheat lines and unmasked the effect of a novel chlorosis-inducing factor [63,64]. In the Brassicae black leg Leptosphaeria maculans-Brassica napus pathosystem, Rlm3-mediated resistance through recognition of the avirulence effector protein AvrLm3, was suppressed by another avirulence effector AvrLm4-7 [65].

Further evidence was provided here using a competitive De Wit replacement assay that demonstrated a direct selective advantage in fungal proliferation in the absence of the PE401 on Snn1 wheat lines since the high-expressing Tox1 variant confers a virulence advantage over low Tox1 expression. In Australia, Snn1 is continually being maintained in wheat varieties grown in Western Australia (WA), a region where SNB is endemic [47]. Association mapping analyses of wheat infected with P. nodorum from Australian field trials frequently detect a QTL on chromosome 1B where Snn1 is located [22,6669] summarised in [23]. This suggests that the Tox1-Snn1 interaction plays a role in SNB in Australia. Therefore, it is paramount to remove Snn1 from commercial wheat to improve their genetic potential for SNB resistance. Maintenance of high Tox1 expression can be bioenergetically taxing if it does not confer a selective advantage for the pathogen. Therefore, it remains to be seen if the complete removal of Snn1 in commercial wheat through germplasm improvements will affect the frequency of PE401 within the Australian P. nodorum population in the future. Tox1 and the other two well-characterised NEs ToxA and Tox3 are present in almost all P. nodorum isolates in Australia but in other parts of the world, ToxA, 1 and 3 frequencies are much lower [45,60]. Outside of Australia, recent association studies using both GWAS and biparental mapping detected the 1B SNB QTL from field trials conducted in Norway [25,70], but the detection of Tox1-Snn1 interactions in field studies can be variable [summarised in 23]. For instance, it was previously observed that an eight-way Multiparent Advanced Generation Inter-Cross (MAGIC) population segregated for Tox1 sensitivity at the 1B locus where Snn1 is situated [71,72]. However, Lin et al. [54] carried out a field study using a sub-set of MAGIC lines naturally infected with P. nodorum stubble across Norway and the UK during a four year trial period. Surprisingly, the Tox1-Snn1 interaction was not detected in SNB of wheat across all the trial periods. Interestingly, QTLs located on chromosome 2A were detected across all trial seasons [54]. There are several possible explanations. Firstly, natural isolates present at both trial sites may lack Tox1 [60]. A recent survey of the Norwegian P. nodorum population revealed that Tox1 prevalence in isolates collected from different sampling sites varied from 7% to 45% [61]. Second, other NE-Snn interactions may be active and epistatic to Tox1-Snn1 (Table 1). Lastly, isolates at both trial sites may possess PE401 and thus represses the role of Tox1-Snn1 in SNB. Hence, we can only speculate that there is less dependency for Tox1-Snn1 ETS outside of Australia. It was also hypothesised by Haugrud et al. [30] that P. nodorum down-regulates Tox1 as a means to prevent antagonism with the activity of other NEs that subvert alternative host-immune responses. This hypothesis requires further testing.

From the co-infection assay, we anticipated that SN15 (-) will outcompete SN15 (+) when infecting Snn1 wheats when all other known NE-host dominant susceptibility gene interactions are removed in CxW DH lines. In contrast, SN15 (-) appears to ‘rescue’ SN15(+) when co-infecting. In two of the three Snn1 wheat lines, the pathogen complex accumulated more biomass than a single infection. McDonald et al. [73] observed different genotypes of P. nodorum on a single infected wheat plant which suggests that P. nodorum exists as a fungal disease complex to ensure maximal infection capabilities on susceptible wheat. This observation accounts for a mixed Tox1 expressing genotype in the Australian P. nodorum population where the high expressing variant remains dominant [47 and S1 Fig].

In this study, we observed that Tox1 exhibits both ‘selfish’ and ‘selfless’ (altruistic) characteristics. We hypothesise that the loss of PE401 allow the gene to exert dominance or epistasis over both Tox3-Snn3 and Tox2A-Qsnb.cur-2AS1 interactions which is driven by Snn1 and results in Tox1 exhibiting a ‘selfish’ character [74]. However, our coinfection competition assay demonstrates evidence of altruism at the isolate level where one isolate must have a higher Tox1 expressing variant to help maintain the virulence and prevalence of isolates with lower Tox1 expression, which is mediated by the presence of PE401 in the P. nodorum population. Hence, this study has provided a unique mechanistic insight into a complex NE-NE interaction that is occurring within the P. nodorum population.

The identity of trans-acting regulatory molecules that are associated with Tox1 regulation remains unknown at this stage. However, key results from this study allow us to propose a hypothetical regulatory model for Tox1 and NE-Snn epistasis detected using the CxW population (Fig 7). In the absence of the PE401, transcriptional activator(s) can drive high Tox1 expression (Fig 7A). As a result, Tox1-Snn1 contributes strongly to SNB while the Qsnb.cur-2AS1 QTL is suppressed. An unidentified repressor binds to the PE401 and inhibits the transcriptional activator(s) bound to the TATA-region [75,76], thereby repressing Tox1 expression and the consequent epistatic effect on the Tox2A-Qsnb.cur-2AS1 interaction during SNB (Fig 7B). Although the proposed hypothetical regulatory mechanism remains speculative at this stage, it provides a framework that will allow future studies of regulatory components. As such, affinity purification assays using these sequences as bait represent a promising avenue to identify the regulators involved.

Fig 7. A hypothetical model for Tox1 regulation and its influence on SNB.

SNB contribution A. in the absence and B. presence of PE401. The effect on gene expression and direct (solid arrows) or epistatic (dashed arrows) contribution to SNB are depicted by the strength of the arrows.

Conclusions and implications

The findings presented here have advanced our knowledge regarding the control of Tox1 expression in P. nodorum and its profound epistatic effect on SNB resistance in wheat. Although the proposed regulatory mechanism remains speculative at this stage, we have defined individual regulatory elements within the promoter region of Tox1. This provides a framework for future studies into the components regulating Tox1. We are currently using a DNA-protein bioassay approach to identify transcriptional activator and repressor proteins that are associated with the promoter region of Tox1 and regulate its expression. However, underlying questions on the mechanism of the epistatic suppression of NE gene expression and phenotypic contributions of SNB remained unanswered. Small interfering RNA may be present within the fungus that regulates gene expression including silencing NE genes [77]. Alternatively, NE-host gene interactions within the same system compete for common plant-signaling pathways, such as the mitogen-activated protein kinase pathway, or are antagonistic in the mechanisms by which ETS is triggered [15,78].

The evidence presented highlights the general importance of understanding NE regulation, the outcomes of which extend beyond the individual NE-host receptor interactions. The evolution of changes in NE gene expression provides a mechanism for P. nodorum to adapt where the complement of host receptors shifts through resistance breeding efforts. More importantly, the identification of PE401 will help to formulate effective crop protection strategies against SNB of wheat. Downie et al. [23] provided a framework of management strategies to minimise the impact of SNB. This includes establishing an extensive isolate collection programme, epidemic monitoring, population genotyping and wheat variety adoption strategies. Based on the outcome of this study, we propose additional guidelines for SNB management to complement the Downie et al. [23] framework. These are;

  1. Monitor for changes in the frequency of PE401 in regional P. nodorum populations.
  2. The maintenance of Snn1 in wheat is a likely driver for P. nodorum to retain PE401 and Tox1 evolution. Therefore, Snn1 removal should be prioritised in cereal breeding programmes through conventional breeding, genome editing or genetic modifications.
  3. Introduce the resistance allele for Qsnb.cur-2AS1 with other SNB resistance alleles (eg. snn1 conferring Tox1 insensitivity) into wheat breeding stocks. Phan et al. [9] and Lin et al. [54] demonstrated that stacking of four regionally relevant SNB resistance alleles provide optimal disease resistance in wheat.
  4. Qsnb.cur-2AS1 also confers resistance to tan spot [55]. Inclusion of the Qsnb.cur-2AS1 insensitivity/SNB resistance allele in breeding programmes should reduce the impact of the SNB-tan spot disease complex.

We anticipate that the outcome of this study will drive a greater level of research into the field of effector regulation and epistasis in other fungal-plant pathosystems to generate similar outcomes to improve existing crop protection strategies.

Materials and methods

Fungal culture and isolate sources

All P. nodorum strains were maintained on V8-PDA agar at 21°C under a 12-h photoperiod or grown in Fries3 broth as previously described [53,79]. All isolates analysed in this study were obtained from random, transects or hierarchical sampling of natural field infections as previously described [47,50,80,81]. Additional information on the sampling locations, dates collected and other relevant information available for the respective isolates is summarised in S1 File.

Compilation of Tox1 promoter variants

The P. nodorum Tox1 sequence was derived from the genome annotation SNOG_20078 of the Australian reference isolate SN15 [35]. A BLAST database was built (Geneious Prime 20.2.5) from the other available published and assembled genomes for 22 P. nodorum isolates [36,37]. The corresponding Tox1 loci, including the 1,500 bp upstream regions, were retrieved using the SN15 Tox1 nucleotide sequence. The 20 sequences retrieved were aligned using the Geneious aligner on the default settings to detect polymorphisms in the promoter region. A collection of 159 Australian isolates from a previous study [47] were then PCR screened (Tox1_screen_F/R) to detect product size shifts corresponding to a 401 bp element presence/absence variant. All primer sequences are shown in S1 Table. Sanger sequencing was undertaken on the region for two Australian isolates with the 401 bp element (WAC13443 and WAC13072) and an additional isolate without (WAC13690). The Tox1 locus and 1,500 bp upstream regions were also retrieved from genomes that were previously assembled for a worldwide collection of 146 P. nodorum isolates (which included 20 additional Australian isolates) [81,82]. The NCBI BLAST service [83] was then used to query the sequence read archives for 184 Illumina sequenced genomes for USA isolates deposited as PRJNA398070 [80], which were non-redundant to the isolates already compiled. The Tox1 nucleotide sequence and 1,000 bp promoter from SN4 was used to query the respective archives for each isolate. Where continuous coverage of reads >95% identity was obtained across the corresponding region, the presence/absence of PE401 was able to be determined. Isolate details and the data sources used for promoter sequencing and alignments are summarised in S1 File.

In silico analyses of the Tox1 promoter variants

A world distribution map was produced from the geographical information assembled for a panel of 484 P. nodorum isolates with or without the Tox1 PE401 (S1 File) using the maps and ggplot2 packages in R [84]. The SN4 Tox1 promoter sequence was then queried for matches in Repbase by submitting the 1500 bp region through the CENSOR portal using the default setting [85,86]. The 401 bp SN4 Tox1 PE401 was also submitted to the Rfam portal [87]. Furthermore, the NCBI BLAST service [83] was used to scan the PE401 sequence against the SN15 genome, the NCBI nucleotide and the whole-genome shotgun collections. Dot plots and GC% graphs were produced using Geneious. The SNOG_30065 expression and gene annotation data were obtained from a previous study [42].

Generating Tox1 promoter replacement mutants

The cloning strategy and an overview of the modified Tox1 promoter locus in fungal transformants are depicted in S3 Fig and the primer details are listed in S1 Table. In summary, promoter replacement constructs were assembled in two stages by attaching flanking regions to a pTef1-HygR-tTef1 resistance marker using a Golden Gate’ (GG) style cloning system [88] established in-house using a pUC19 vector backbone (New England Biolabs, Ipswich, Massachusetts, USA) and kindly provided by Dr. Jordi Muria-Gonzales. In stage one the pTef1-HygR-tTef1 marker was assembled by BbsI-mediated cloning using Tef1 promoter and terminator regions derived from Aspergillus nidulans using pTef1_P_BbsI_F/R and tTef1_T_BbsI_F/R, as well as the hygromycin resistance gene amplified from Pan7 [89] with HygR_B_BbsI_F/R. Stage two involved the assembly and attachment of distinct flanks to the marker cassette using BsaI cloning. The left flank was the same for all constructs, amplified from SN15 using pTox1_HR_FL_Bsa1_F/R. The right flanks were amplified from SN15 gDNA for the (-) PE401 construct variant using pTox1_873_HR_FR_Bsa1_F/pTox1_HR_FR_Bsa1_R. The same primers were used in combination with pTox1_Dom_BsaI_F/R to amplify the right flank from WAC13443 gDNA for PE401 constructs (which allowed fragment domestication for GG cloning by the introduction of a point mutation). PE401 was also substituted in the previous construct by amplifying the GG plasmid with pTox1_indel_3_BbsI_F/pTox1_indel_5_BbsI_R and ligating in a 401 bp spacer (S) element with a similar GC content but an unrelated sequence (S2 File). This was amplified from the pGEM-T-Easy vector (Promega, Madison, WI) using the primer pair spacer_BbsI_F/R. The three resulting homologous recombination (HR) constructs, were amplified and purified from these templates using pTox1_HR_FL_F/R, before polyethylene glycol (PEG) mediated transformation into SN15 and WAC13443. Truncated versions of the Tox1 promoter replacement construct were produced by modifying the forward primer used to amplify the right flank to incorporate either 86, 107, 157, 263, 310 (+/- PE401) or 598 bp (+/- PE401) regions upstream of the translational start site. The resulting constructs were PEG-transformed into SN15 as previously described [90]. All cloned constructs were verified through PCR screening and Sanger sequencing, while the fungal transformants were PCR screened using pTox1_HR_screen_F/tTox1_HR_screen_R and by qPCR (Tox1_qPCR_F/R vs Actin_qPCR_F/R) to confirm single-copy integration using a robust quantitative PCR method [91]. Two single copy mutants for all transformants were retained for gene expression analyses. All genetically modified strains and their background strains are described in Table 4.

Table 4. Fungal mutant strains used in this study.

Wildtype strains used for mutagenesis are indicated.

Gene expression analysis

P. nodorum cDNA was synthesised from the respective strains/mutants grown for three days in sterile Fries3 liquid medium for three days shaken at 120 rpm in the dark as previously described [53]. Quantitative RT-PCR was used to determine the expression level of Tox1 normalised to Act1 (2dCt). For in planta time series analysis of gene expression, lesions were harvested at 3, 5 and 7 days post-infection by the seedling spray method, from which cDNA was synthesised for qPCR as previously described [92]. Tox1_qPCR_F/R and Tox3_qPCR_F/R were used to assess the respective gene expression normalised to Act1 (2dCt). A one-way ANOVA with Tukey-HSD post-hoc test was used to test for differences (p<0.05) between isolates and/or mutants (SPSS version 27.0).

Association mapping and QTL analysis

The CxW DH wheat population that consisted of 177 lines (Intergrain Pty Ltd, Perth, Australia) was used to carry out association mapping and QTL studies of seedling SNB as previously described [22]. A total of 482 DarT, wheat genomic SSR and EST-SSR markers were used to construct a genetic map for the population [22]. Two-week old seedlings were infected with P. nodorum using a whole plant spray assay was visually assessed for SNB severity on a standardised scale of one to nine as previously described [22]. A score of one indicates no disease symptoms whereas a score of nine indicates a fully necrotised plant [9,22]. QTL mapping was undertaken using MultiQTL v. 2.6-Complete software (MultiQTL Ltd, Institute of Evolution, Haifa University, Israel). The QTL mapping was based on a genetic linkage map previously built for the CxW population using the Kosambi mapping function using MultiPoint v. 3.2 (MultiQTL Ltd, Haifa University, Israel) from maximum recombination frequencies of 0.35 [22]. This included 385 markers polymorphic between the CxW parent lines. Seedling disease scores taken from the average of three randomised biological replicates were used for interval mapping to determine QTL linked to SNB. Markers with the logarithm of the odds (LOD) score set at a > 2.5 cut-off was used to construct an interval model for the corresponding QTL as previously described [22].

Attached leaf infection assay

An attached leaf assay with modifications was used to carry out multi-strain co-infection [93]. Briefly, the first leaves of two-week old wheat seedlings were inoculated with a P. nodorum spore suspension [1 x 106 spores/ml in 0.02% (v/v) Tween 20] using an airbrush kit (6ml/min, Ozito, Perth, Australia). Per treatment (+PE401; -PE401; 50:50 mix) a total of 6 ml spore inoculum was evenly sprayed across five wheat lines with 6 seedlings per line. Tween was used as a negative control. The wheat lines were H213, H324 (snn1, snn3b1, insensitive to Tox2A) and H086, H336, H091 (Snn1, snn3b1, insensitive to Tox2A). Leaves were allowed to dry and incubated in a controlled growth chamber as described previously [93]. Two and four days post-inoculation, infected leaves were harvested with a scalpel and snap-frozen in liquid nitrogen (three biological repeats per time point per wheat line per treatment). DNA was extracted using phenol/chloroform method as described previously [94] and concentrations were measured using a Qubit Fluorometer (ThermoFisher Scientific, MA USA). All samples were diluted to a concentration of 10 ng/ul DNA.

Droplet digital PCR (ddPCR) quantification for fungal biomass

DdPCR was used for absolute quantification of biomass base on fungal DNA copy abundance. This approach allows for the precise and absolute quantification of DNA biomass as it measures and analyses about 20,000 reactions (droplets) in a single well [95]. We used two primer sets, a specific SN15 (+) primer targeting PE401 (pTox1_401_qPCR_F and pTox1_401_qPCR_R) and a general SN15 primer (ddPCRf and ddPCRr) that will detect both SN15 (-) and SN15 (+) DNA (S1 Table). In a single well, all the positive droplets carrying a copy of the target sequence are counted and expressed as the number of copies per input DNA concentration (in our case, 50 ng of total DNA). As such, we determined and compared the number of copies between the treatments and wheat lines as an accurate measure of pathogen biomass. We used the Evagreen (BioRad, IL USA) approach as described previously [96]. Briefly, a 96-well plate was loaded into the QX200 Auto DG to generate droplets (Bio-Rad, IL USA) in each well. Next, plates were sealed using the PX1 PCR Plate Sealer before continuing to PCR (C1000 Touch Thermal Cycler, Bio-Rad, IL USA) which was performed with the primer sets described above using the following program: 95°C for 5 min, 44 cycles of 95°C for 30 s, 63°C for 30 s, 72°C for 30 s and finish with 4°C for 5 min followed by 90°C for 5 min. The plate was subsequently loaded into the QX200 Plate Reader (Bio-Rad, IL USA) and data was analysed using the QuantaSoft Software (Bio-Rad, IL USA). Genomic DNA of SN15 (-) and SN15 (+) strains were included as positive controls and additionally, primer efficiency was tested, thereby ensuring comparability between wells. A one-way ANOVA with posthoc Tukey-HSD testing was used to identify significant (P < 0.05) differences between the three infection treatments (+401 bp; -401 bp; 50:50 mix) per wheat line (SPSS version 27.0).

Supporting information

S1 File. Tox1 isolate metadata describing the isolate ID, location sourced, any available host-plant information and the collection date.

The presence of the 401 bp PE401 in the Tox1 promoter is indicated along with the data source for this assessment and reference from which this was determined.


S2 File. PE401 and the 401 bp spacer sequence used to substitute the element in SN15 (S) control mutants.

A dotplot is provided to demonstrate the sequence dissimilarity.


S3 File. SNB disease score and QTL mapping data.

A table indicating the CxW lines tested, the presence of Snn1 and Snn3-linked marker, the average disease scores for SN15 (-) and SN15 (+) mutants as well as a summary of the disease score statistical analysis and the chromosomal linkage groups analysed.


S1 Fig. Assessment of PE401 frequencies in the Australian P. nodorum isolate collected between 1972 and 2016.

“TRUE” indicates the presence of PE401 whereas “FALSE” indicates the absence of PE401.


S2 Fig. Composite interval mapping of QTL associated with SNB caused by SN15 (-), SN15 (+), SN15 and tox1-6 [22].


S3 Fig. Tox1 promoter replacement strategy.

A. In step 1, Golden Gate cloning was used to assemble constructs for promoter replacement at the Tox1 locus. Linear replacement constructs were amplified for fungal transformation in step 2. B. Promoter replacement mutants generated in the study representing the Tox1 locus in the respective background strains.



We thank Intergrain Pty. Ltd. for the wheat mapping population. We thank Mr. Lincoln Harper and Dr. Noel Knight for the technical advice on ddPCR.


  1. 1. McDonald BA, Linde C. Pathogen population genetics, evolutionary potential, and durable resistance. Annu Rev Phytopathol. 2002;40:349–79. pmid:12147764
  2. 2. Rouxel T, Balesdent MH. Life, death and rebirth of avirulence effectors in a fungal pathogen of Brassica crops, Leptosphaeria maculans. New Phytol. 2017;214(2):526–32. pmid:28084619
  3. 3. McIntosh RA, Brown GN. Anticipatory breeding for resistance to rust diseases in wheat. Annu Rev Phytopathol. 1997;35:311–26. pmid:15012526
  4. 4. Tan KC, Oliver RP, Solomon PS, Moffat CS. Proteinaceous necrotrophic effectors in fungal virulence. Functional Plant Biology. 2010;37:907–12.
  5. 5. Ciuffetti LM, Tuori RP, Gaventa JM. A single gene encodes a selective toxin causal to the development of tan spot of wheat. Plant Cell. 1997;9(2):135–44. pmid:9061946
  6. 6. Vleeshouwers VG, Oliver RP. Effectors as tools in disease resistance breeding against biotrophic, hemibiotrophic, and necrotrophic plant pathogens. Mol Plant-Microbe Interact. 2014;27(3):196–206. pmid:24405032
  7. 7. Friesen TL, Faris JD, Solomon PS, Oliver RP. Host-specific toxins: effectors of necrotrophic pathogenicity. Cellular Microbiology. 2008;10(7):1421–8. pmid:18384660
  8. 8. Tan KC, Phan HT, Rybak K, John E, Chooi YH, Solomon PS, et al. Functional redundancy of necrotrophic effectors—consequences for exploitation for breeding. Frontiers in Plant Science. 2015;6:501. pmid:26217355
  9. 9. Phan HTT, Rybak K, Bertazzoni S, Furuki E, Dinglasan E, Hickey LT, et al. Novel sources of resistance to Septoria nodorum blotch in the Vavilov wheat collection identified by genome-wide association studies. Theor Appl Genet. 2018. pmid:29470621
  10. 10. Friesen TL, Meinhardt SW, Faris JD. The Stagonospora nodorum-wheat pathosystem involves multiple proteinaceous host-selective toxins and corresponding host sensitivity genes that interact in an inverse gene-for-gene manner. Plant Journal. 2007;51(4):681–92.
  11. 11. Friesen TL, Faris JD. Characterization of the wheat–Stagonospora nodorum disease system: what is the molecular basis of this quantitative necrotrophic disease interaction. Canadian Journal of Plant Pathology. 2010;32(1):20–8.
  12. 12. McDonald MC, Solomon PS. Just the surface: advances in the discovery and characterization of necrotrophic wheat effectors. Curr Opin Microbiol. 2018;46:14–8. pmid:29452845
  13. 13. Friesen TL, Stukenbrock EH, Liu ZH, Meinhardt S, Ling H, Faris JD, et al. Emergence of a new disease as a result of interspecific virulence gene transfer. Nature Genetics. 2006;38(8):953–6. pmid:16832356
  14. 14. Faris JD, Zhang Z, Lu H, Lu S, Reddy L, Cloutier S, et al. A unique wheat disease resistance-like gene governs effector-triggered susceptibility to necrotrophic pathogens. Proceedings of the National Academy of Sciences U S A. 2010;107:13544–9. pmid:20624958
  15. 15. Shi G, Zhang Z, Friesen TL, Raats D, Fahima T, Brueggeman RS, et al. The hijacking of a receptor kinase-driven pathway by a wheat fungal pathogen leads to disease. Sci Adv. 2016;2(10):e1600822. pmid:27819043
  16. 16. Liu Z, Zhang Z, Faris JD, Oliver RP, Syme R, McDonald MC, et al. The cysteine rich necrotrophic effector SnTox1 produced by Stagonospora nodorum triggers susceptibility of wheat lines harboring Snn1. PLoS Pathogens. 2012;8(1):e1002467. pmid:22241993
  17. 17. Richards JK, Kariyawasam G, Seneviratne S, Wyatt NA, Xu SS, Liu Z, et al. A triple threat: the Parastagonospora nodorum SnTox267 effector exploits three distinct host genetic factors to cause disease in wheat. bioRxiv. 2021:2021.02.25.432871. pmid:34227112
  18. 18. Liu Z, Faris JD, Oliver RP, Tan KC, Solomon PS, McDonald MC, et al. SnTox3 acts in effector triggered susceptibility to induce disease on wheat carrying the Snn3 gene. PLoS Pathogens. 2009;5(9):e1000581. pmid:19806176
  19. 19. Zhang Z, Running KLD, Seneviratne S, Peters Haugrud AR, Szabo-Hever A, Shi G, et al. A protein kinase-major sperm protein gene hijacked by a necrotrophic fungal pathogen triggers disease susceptibility in wheat. Plant J. 2021. pmid:33576059
  20. 20. Abeysekara NS, Friesen TL, Keller B, Faris JD. Identification and characterization of a novel host-toxin interaction in the wheat-Stagonospora nodorum pathosystem. Theoretical and Applied Genetics. 2009;120(1):117–26. pmid:19816671
  21. 21. Kariyawasam GK, Richards JK, Wyatt NA, Running K, Xu SS, Liu Z, et al. The Parastagonospora nodorum necrotrophic effector SnTox5 targets the wheat gene Snn5 and facilitates entry into the leaf mesophyll. bioRxiv. 2021:2021.02.26.433117. pmid:34231227
  22. 22. Phan HT, Rybak K, Furuki E, Breen S, Solomon PS, Oliver RP, et al. Differential effector gene expression underpins epistasis in a plant fungal disease. The Plant Journal. 2016;87:343–54. pmid:27133896
  23. 23. Downie RC, Lin M, Corsi B, Ficke A, Lillemo M, Oliver RP, et al. Septoria Nodorum Blotch of Wheat: Disease Management and Resistance Breeding in the Face of Shifting Disease Dynamics and a Changing Environment. Phytopathology. 2021:PHYTO07200280RVW. pmid:33245254
  24. 24. Tan KC, Waters ODC, Rybak K, Antoni E, Furuki E, Oliver RP. Sensitivity to three Parastagonospora nodorum necrotrophic effectors in current Australian wheat cultivars and the presence of further fungal effectors. Crop & Pasture Science. 2014;65:150–8.
  25. 25. Ruud AK, Dieseth JA, Ficke A, Furuki E, Phan HTT, Oliver RP, et al. Genome-Wide Association Mapping of Resistance to Septoria Nodorum Leaf Blotch in a Nordic Spring Wheat Collection. Plant Genome. 2019;12(3).
  26. 26. Lin M, Stadlmeier M, Mohler V, Tan KC, Ficke A, Cockram J, et al. Identification and cross-validation of genetic loci conferring resistance to Septoria nodorum blotch using a German multi-founder winter wheat population. Theor Appl Genet. 2021;134(1):125–42. pmid:33047219
  27. 27. Tan KC, Oliver RP. Regulation of proteinaceous effector expression in phytopathogenic fungi. PLoS Pathog. 2017;13(4):e1006241. pmid:28426760
  28. 28. Friesen TL, Zhang Z, Solomon PS, Oliver RP, Faris JD. Characterization of the interaction of a novel Stagonospora nodorum host-selective toxin with a wheat susceptibility gene. Plant Physiology. 2008;146(2):682–93. pmid:18065563
  29. 29. Friesen TL, Chu C, Xu SS, Faris JD. SnTox5-Snn5: a novel Stagonospora nodorum effector-wheat gene interaction and its relationship with the SnToxA-Tsn1 and SnTox3-Snn3-B1 interactions. Mol Plant Pathol. 2012;13(9):1101–9. pmid:22830423
  30. 30. Haugrud ARP, Zhang ZC, Richards JK, Friesen TL, Faris JD. Genetics of Variable Disease Expression Conferred by Inverse Gene-For-Gene Interactions in the Wheat-Parastagonospora nodorurm Pathosystem. Plant Physiology. 2019;180(1):420–34. pmid:30858234
  31. 31. Peters-Haugrud AR, Zhang Z, Richards JK, Friesen TL, Faris JD. Genetics of variable disease expression conferred by inverse gene-for-gene interactions in the wheat-Parastagonospora nodorum pathosystem. Plant Physiol. 2019. pmid:30858234
  32. 32. Liu ZH, Friesen TL, Rasmussen JB, Ali S, Meinhardt SW, Faris JD. Quantitative trait loci analysis and mapping of seedling resistance to Stagonospora nodorum leaf blotch in wheat. Phytopathology. 2004;94(10):1061–7. pmid:18943794
  33. 33. Chu CG, Faris JD, Xu SS, Friesen TL. Genetic analysis of disease susceptibility contributed by the compatible Tsn1-SnToxA and Snn1-SnTox1 interactions in the wheat-Stagonospora nodorum pathosystem. Theoretical and Applied Genetics. 2010;120(7):1451–9. pmid:20084492
  34. 34. Gao Y, Faris JD, Liu Z, Kim YM, Syme RA, Oliver RP, et al. Identification and characterization of the SnTox6-Snn6 Interaction in the Parastagonospora nodorum-wheat pathosystem. Mol Plant-Microbe Interact. 2015;28(5):615–25. pmid:25608181
  35. 35. Syme RA, Tan KC, Hane JK, Dodhia K, Stoll T, Hastie M, et al. Comprehensive annotation of the Parastagonospora nodorum reference genome using next-generation genomics, transcriptomics and proteogenomics. PLoS One. 2016;11(2):e0147221. pmid:26840125
  36. 36. Syme RA, Tan KC, Rybak K, Friesen TL, McDonald BA, Oliver RP, et al. Pan-Parastagonospora Comparative Genome Analysis-Effector Prediction and Genome Evolution. Genome Biol Evol. 2018;10(9):2443–57. pmid:30184068
  37. 37. Richards JK, Wyatt NA, Liu Z, Faris JD, Friesen TL. Reference Quality Genome Assemblies of Three Parastagonospora nodorum Isolates Differing in Virulence on Wheat. G3 (Bethesda). 2018;8(2):393–9.
  38. 38. Soyer JL, El Ghalid M, Glaser N, Ollivier B, Linglin J, Grandaubert J, et al. Epigenetic control of effector gene expression in the plant pathogenic fungus Leptosphaeria maculans. PLoS Genetics. 2014;10(3):e1004227. pmid:24603691
  39. 39. van Dam P, Rep M. The Distribution of Miniature Impala Elements and SIX Genes in the Fusarium Genus is Suggestive of Horizontal Gene Transfer. J Mol Evol. 2017;85(1–2):14–25. pmid:28744785
  40. 40. Krishnan P, Meile L, Plissonneau C, Ma X, Hartmann FE, Croll D, et al. Transposable element insertions shape gene regulation and melanin production in a fungal pathogen of wheat. BMC Biol. 2018;16(1):78. pmid:30012138
  41. 41. Razali NM, Cheah BH, Nadarajah K. Transposable Elements Adaptive Role in Genome Plasticity, Pathogenicity and Evolution in Fungal Phytopathogens. Int J Mol Sci. 2019;20(14).
  42. 42. Jones DAB, John E, Rybak K, Phan HTT, Singh KB, Lin SY, et al. A specific fungal transcription factor controls effector gene expression and orchestrates the establishment of the necrotrophic pathogen lifestyle on wheat. Sci Rep. 2019;9(1):15884. pmid:31685928
  43. 43. Muszewska A, Steczkiewicz K, Stepniewska-Dziubinska M, Ginalski K. Cut-and-Paste Transposons in Fungi with Diverse Lifestyles. Genome Biol Evol. 2017;9(12):3463–77. pmid:29228286
  44. 44. McDonald MC, Razavi M, Friesen TL, Brunner PC, McDonald BA. Phylogenetic and population genetic analyses of Phaeosphaeria nodorum and its close relatives indicate cryptic species and an origin in the Fertile Crescent. Fungal Genet Biol. 2012;49(11):882–95. pmid:22922546
  45. 45. Ghaderi F, Sharifnabi B, Javan-Nikkhah M, Brunner PC, McDonald BA. SnToxA, SnTox1, and SnTox3 originated in Parastagonospora nodorum in the Fertile Crescent. Plant Pathology. 2020;69(8):1482–91.
  46. 46. Solomon PS, Wilson TJG, Rybak K, Parker K, Lowe RGT, Oliver RP. Structural characterisation of the interaction between Triticum aestivum and the dothideomycete pathogen Stagonospora nodorum. European Journal of Plant Pathology. 2006;114(3):275–82.
  47. 47. Phan HTT, Jones DAB, Rybak K, Dodhia KN, Lopez-Ruiz FJ, Valade R, et al. Low Amplitude Boom-and-Bust Cycles Define the Septoria Nodorum Blotch Interaction. Front Plant Sci. 2020;10:1785. pmid:32082346
  48. 48. de Wit CT. On competition. Verslagen Landbouwkundige Onderzoekigen. 1960;66:1–82.
  49. 49. Basehoar AD, Zanton SJ, Pugh BF. Identification and distinct regulation of yeast TATA box-containing genes. Cell. 2004;116(5):699–709. pmid:15006352
  50. 50. Pereira D, Oggenfuss U, McDonald BA, Croll D. Population genomics of transposable element activation in the highly repressive genome of an agricultural pathogen. Microb Genom. 2021;7(8). pmid:34424154
  51. 51. Cowger C, Ward B, Brown-Guedira G, Brown JKM. Role of Effector-Sensitivity Gene Interactions and Durability of Quantitative Resistance to Septoria Nodorum Blotch in Eastern U.S. Wheat. Front Plant Sci. 2020;11:155. pmid:32210986
  52. 52. Tan KC, Ferguson-Hunt M, Rybak K, Waters OD, Stanley WA, Bond CS, et al. Quantitative variation in effector activity of ToxA isoforms from Stagonospora nodorum and Pyrenophora tritici-repentis. Mol Plant-Microbe Interact. 2012;25(4):515–22. pmid:22250581
  53. 53. Rybak K, See PT, Phan HT, Syme RA, Moffat CS, Oliver RP, et al. A functionally conserved Zn2 Cys6 binuclear cluster transcription factor class regulates necrotrophic effector gene expression and host-specific virulence of two major Pleosporales fungal pathogens of wheat. Mol Plant Pathol. 2017;18(3):420–34. pmid:27860150
  54. 54. Lin M, Corsi B, Ficke A, Tan KC, Cockram J, Lillemo M. Genetic mapping using a wheat multi-founder population reveals a locus on chromosome 2A controlling resistance to both leaf and glume blotch caused by the necrotrophic fungal pathogen Parastagonospora nodorum. Theor Appl Genet. 2020;133(3):785–808. pmid:31996971
  55. 55. Shankar M, Jorgensen D, Taylor J, Chalmers KJ, Fox R, Hollaway GJ, et al. Loci on chromosomes 1A and 2A affect resistance to tan (yellow) spot in wheat populations not segregating for tsn1. Theor Appl Genet. 2017;130(12):2637–54. pmid:28913578
  56. 56. Murray GM, Brennan JP. Estimating disease losses to the Australian wheat industry. Australasian Plant Pathology. 2009;38:558–70.
  57. 57. Abdullah AS, Gibberd MR, Hamblin J. Co-infection of wheat by Pyrenophora tritici-repentis and Parastagonospora nodorum in the wheatbelt of Western Australia. Crop & Pasture Science. 2020;71(2):119–27.
  58. 58. Loughman R, Wilson RE, Thomas GJ. The influence of disease complexes involving Leptosphaeria (Septoria) nodorum on detection of resistance to 3 leaf-spot diseases in wheat. Euphytica. 1993;72(1–2):31–42.
  59. 59. Justesen AF, Corsi B, Ficke A, Hartl L, Holdgate S, Jørgensen LN, et al. Hidden in plain sight: a molecular field survey of three wheat leaf blotch fungal diseases in North-Western Europe shows co-infection is widespread. European Journal of Plant Pathology. 2021. pmid:34465944
  60. 60. McDonald MC, Oliver RP, Friesen TL, Brunner PC, McDonald BA. Global diversity and distribution of three necrotrophic effectors in Phaeosphaeria nodorum and related species. New Phytologist. 2013;199:241–51. pmid:23550706
  61. 61. Lin M, Ficke A, Cockram J, Lillemo M. Genetic structure of the Norwegian Parastagonospora nodorum population. Front Microbiol. 2020. pmid:32612592
  62. 62. Faris JD, Zhang Z, Rasmussen JB, Friesen TL. Variable expression of the Stagonospora nodorum effector SnToxA among isolates is correlated with levels of disease in wheat. Molecular plant-microbe interactions: MPMI. 2011;24(12):1419–26. pmid:21770771
  63. 63. Manning VA, Ciuffetti LM. Necrotrophic effector epistasis in the Pyrenophora tritici-repentis-wheat interaction. PLoS One. 2015;10(4):e0123548. pmid:25845019
  64. 64. Moffat CS, See PT, Oliver RP. Generation of a ToxA knockout strain of the wheat tan spot pathogen Pyrenophora tritici-repentis. Mol Plant Pathol. 2014;15(9):918–26. pmid:24831982
  65. 65. Plissonneau C, Daverdin G, Ollivier B, Blaise F, Degrave A, Fudal I, et al. A game of hide and seek between avirulence genes AvrLm4-7 and AvrLm3 in Leptosphaeria maculans. New Phytologist. 2015. pmid:26592855
  66. 66. Li D, Walker E, Francki M. Genes Associated with Foliar Resistance to Septoria Nodorum Blotch of Hexaploid Wheat (Triticum aestivum L.). Int J Mol Sci. 2021;22(11). pmid:34070394
  67. 67. Francki MG, Walker E, McMullan CJ, Morris WG. Multi-Location Evaluation of Global Wheat Lines Reveal Multiple QTL for Adult Plant Resistance to Septoria Nodorum Blotch (SNB) Detected in Specific Environments and in Response to Different Isolates. Front Plant Sci. 2020;11:771. pmid:32655592
  68. 68. Francki MG, Shankar M, Walker E, Loughman R, Golzar H, Ohm H. New quantitative trait loci in wheat for flag leaf resistance to Stagonospora nodorum blotch. Phytopathology. 2011;101(11):1278–84. pmid:21770777
  69. 69. Francki MG, Walker E, Li DA, Forrest K. High-density SNP mapping reveals closely linked QTL for resistance to Stagonospora nodorum blotch (SNB) in flag leaf and glume of hexaploid wheat. Genome. 2018;61(2):145–9. pmid:29237140
  70. 70. Ruud AK, Windju S, Belova T, Friesen TL, Lillemo M. Mapping of SnTox3-Snn3 as a major determinant of field susceptibility to Septoria nodorum leaf blotch in the SHA3/CBRD x Naxos population. Theor Appl Genet. 2017;130(7):1361–74. pmid:28365817
  71. 71. Cockram J, Scuderi A, Barber T, Furuki E, Gardner KA, Gosman N, et al. Fine-Mapping the Wheat Snn1 Locus Conferring Sensitivity to the Parastagonospora nodorum Necrotrophic Effector SnTox1 Using an Eight Founder Multiparent Advanced Generation Inter-Cross Population. G3 (Bethesda). 2015;5(11):2257–66. pmid:26416667
  72. 72. Mackay IJ, Bansept-Basler P, Barber T, Bentley AR, Cockram J, Gosman N, et al. An eight-parent multiparent advanced generation inter-cross population for winter-sown wheat: creation, properties, and validation. G3 (Bethesda). 2014;4(9):1603–10. pmid:25237112
  73. 73. McDonald BA, Miles T, Nelson LR, Pettway RE. Genetic variability in nuclear DNA in field populations of Stagonospora nodorum. Phytopathology. 1994;84:250–5.
  74. 74. Dawkins R. The selfish gene. New York: Oxford University Press; 1976.
  75. 75. Cooper GM. The Cell: A Molecular Approach. 2nd edition: Sinauer Associates 2000; 2000.
  76. 76. Gaston K, Jayaraman PS. Transcriptional repression in eukaryotes: repressors and repression mechanisms. Cell Mol Life Sci. 2003;60(4):721–41. pmid:12785719
  77. 77. Raman V, Simon SA, Demirci F, Nakano M, Meyers BC, Donofrio NM. Small RNA Functions Are Required for Growth and Development of Magnaporthe oryzae. Mol Plant Microbe Interact. 2017;30(7):517–30. pmid:28504560
  78. 78. Faris JD, Friesen TL. Plant genes hijacked by necrotrophic fungal pathogens. Curr Opin Plant Biol. 2020;56:74–80. pmid:32492572
  79. 79. Liu ZH, Faris JD, Meinhardt SW, Ali S, Rasmussen JB, Friesen TL. Genetic and physical mapping of a gene conditioning sensitivity in wheat to a partially purified host-selective toxin produced by Stagonospora nodorum. Phytopathology. 2004;94(10):1056–60. pmid:18943793
  80. 80. Richards JK, Stukenbrock EH, Carpenter J, Liu Z, Cowger C, Faris JD, et al. Local adaptation drives the diversification of effectors in the fungal wheat pathogen Parastagonospora nodorum in the United States. PLoS Genetics. 2019;15(10):e1008223. pmid:31626626
  81. 81. Stukenbrock EH, Banke S, McDonald BA. Global migration patterns in the fungal wheat pathogen Phaeosphaeria nodorum. Mol Ecol. 2006;15(10):2895–904. pmid:16911209
  82. 82. Pereira D, Oggenfuss U, McDonald BA, Croll D. The population genomics of transposable element activation in the highly repressive genome of an agricultural pathogen. bioRxiv. 2020:2020.11.12.379651.
  83. 83. Johnson M, Zaretskaya I, Raytselis Y, Merezhuk Y, McGinnis S, Madden TL. NCBI BLAST: a better web interface. Nucleic Acids Res. 2008;36(Web Server issue):W5-9. pmid:18440982
  84. 84. Kahle D, Wickham H. ggmap: Spatial Visualization with ggplot2. R J. 2013;5(1):144–61.
  85. 85. Kohany O, Gentles AJ, Hankus L, Jurka J. Annotation, submission and screening of repetitive elements in Repbase: RepbaseSubmitter and Censor. Bmc Bioinformatics. 2006;7. pmid:16401345
  86. 86. Bao WD, Kojima KK, Kohany O. Repbase Update, a database of repetitive elements in eukaryotic genomes. Mobile DNA-Uk. 2015;6. pmid:26045719
  87. 87. Kalvari I, Nawrocki EP, Argasinska J, Quinones-Olvera N, Finn RD, Bateman A, et al. Non-Coding RNA Analysis Using the Rfam Database. Curr Protoc Bioinformatics. 2018;62(1):e51. pmid:29927072
  88. 88. Engler C, Kandzia R, Marillonnet S. A one pot, one step, precision cloning method with high throughput capability. PLoS One. 2008;3(11):e3647. pmid:18985154
  89. 89. Farman ML, Oliver RP. Transformation frequencies are enhanced and vector DNA is targeted during retransformation of Leptosphaeria maculans, a fungal plant pathogen. Mol Gen Genet. 1992;231(2):243–7. pmid:1736094
  90. 90. Solomon PS, Tan KC, Oliver RP. Mannitol 1-phosphate metabolism is required for sporulation in planta of the wheat pathogen Stagonospora nodorum. Mol Plant-Microbe Interact. 2005;18(2):110–5. pmid:15720079
  91. 91. Solomon PS, IpCho SVS, Hane JK, Tan KC, Oliver RP. A quantitative PCR approach to determine gene copy number. Fungal Genetics Report. 2008;55:5–8.
  92. 92. Tan KC, Heazlewood JL, Millar AH, Thomson G, Oliver RP, Solomon PS. A signaling-regulated, short-chain dehydrogenase of Stagonospora nodorum regulates asexual development. Eukaryot Cell. 2008;7(11):1916–29. pmid:18776038
  93. 93. Jacques S, Lenzo L, Stevens K, Lawrence J, Tan KC. An optimized sporulation method for the wheat fungal pathogen Pyrenophora tritici-repentis. Plant Methods. 2021;17(1):52. pmid:34011363
  94. 94. Phan HTT, Rybak K, Bertazzoni S, Furuki E, Dinglasan E, Hickey LT, et al. Novel sources of resistance to Septoria nodorum blotch in the Vavilov wheat collection identified by genome-wide association studies. Theor Appl Genet. 2018;131(6):1223–38. pmid:29470621
  95. 95. Taylor SC, Laperriere G, Germain H. Droplet Digital PCR versus qPCR for gene expression analysis with low abundant targets: from variable nonsense to publication quality data. Sci Rep. 2017;7(1):2409. pmid:28546538
  96. 96. Jiang C, Lian X, Gao C, Sun X, Einkauf KB, Chevalier JM, et al. Distinct viral reservoirs in individuals with spontaneous control of HIV-1. Nature. 2020;585(7824):261–7. pmid:32848246