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Scald on gramineous hosts in Iran and their potential threat to cultivated barley

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Abstract

There are five described Rhynchosporium species, Rhynchosporium commune, R. secalis, R. agropyri, R. orthosporum and R. lolii, that cause scald diseases on Poaceae. This study used morphological (conidial shape and size) and phylogenetic analyses of two loci (the internal transcribed spacer region (ITS) and β-tubulin (TUBB)) to identify Rhynchosporium species and their host ranges in Iran. Despite the large variation observed for Rhynchosporium conidial dimensions, the phylogenetic analyses of the ITS region and concatenated ITS and TUBB loci revealed that all isolates from wild grasses in Iran belong to R. commune. R. commune was isolated from Hordeum murinum ssp. glaucum, Hordeum vulgare ssp. spontaneum, Lolium multiflorum and Avena sativa in Iran. A. sativa has only been reported from Iran as a host for R. commune. After cross inoculation, A. sativa was considered as the most resistant host showing the lowest susceptibility to R. commune isolates. Of the grass hosts tested, H. vulgare ssp. spontaneum was the most susceptible. The most aggressive isolate across all tested hosts was isolated from Hordeum murinum ssp. glaucum. Cross-infection of the R. commune isolates from all hosts onto uncultivated grasses and cultivated barley suggests the potential of the uncultivated grasses as inoculum sources for cultivated barley epidemics and pathogen evolution. Thus, management of uncultivated grasses in the vicinity of barley fields should assist in managing the disease on cultivated barley.

Introduction

Different species of Rhynchosporium cause scald disease on barley and other members of the Poaceae. Lesions have a pale green or pale brown centre with dark brown margins on leaves, leaf sheaths, and ears (Caldwell 1937; Shipton et al. 1974). The damage of scald on barley results in high yield losses (Brown 1985). In addition to quantity, the quality of grain is also reduced due to a reduced malting capacity (Avrova and Knogge 2012).

Rhynchosporium was first reported from rye in the Netherlands (Oudemans 1897), in the same year it was isolated from barley in Germany (Frank 1897). It also was observed on triticale and other grasses such as Agropyron spp., Hordeum spp. and Bromus diandrus (Caldwell 1937; Shipton et al. 1974; Welty and Metzger 1996). Initially, only two species of Rhynchosporium were described, Rhynchosporium secalis and R. orthosporum. R. secalis was split into Rhynchosporium commune, R. agropyri and R. secalis (Zaffarano et al. 2011). These species all have beak-shaped conidia (Caldwell 1937; Shipton et al. 1974), whereas R. orthosporum has cylindrical conidia (Caldwell 1937). R. orthosporum is known as the scald agent on Dactylis glomerata (Caldwell 1937) and R. secalis was known as a causing agent of leaf blotch on species of Hordeum, Agropyron, Bromus, Lolium and other plants of Poaceae (Caldwell 1937; Shipton et al. 1974). Phylogenetic analysis of multilocus DNA sequence data of Rhynchosporium from barley, rye and wild grasses displayed three monophylectic groups. Based on their respective hosts, each group was considered as a new species, i.e. R. agropyri from Agropyron, R. secalis from rye and R. commune from Hordeum and Bromus. Furthermore, host specificity of these species was confirmed using pathogenicity tests; R. secalis on rye and triticale, R. commune on barley and other Hordeum spp. and B. diandrus, and R. agropyri could not infect Hordeum or rye (Zaffarano et al. 2011). King et al. (2013) identified Lolium multiflorum as a new host of R. commune. They also described a new species from Lolium, Rhynchosporium lolii, with cylindrical conidia which was the most closely related species to R. orthosporum but with clear molecular, morphological and host differences. Sequencing of the internal transcribed spacer region (ITS) and mating-type loci revealed that R. commune is an anamorphic ascomycete and closely related to genera Oculimacula and Pyrenopeziza (Goodwin 2002; Foster and Fitt 2003).

A morphological study on Rhynchosporium species by King et al. (2013) showed the agents of scald on gramineous hosts are divided into two groups based on conidial shape, including group 1 with beak-shaped conidia and group 2 with cylindrical conidia. R. agropyri, R. secalis and R. commune all have beak-shaped conidia which are indistinguishable based on conidial dimension. Both R. orthosporum and R. lolii have cylindrical conidia which are distinguishable based on conidial dimension.

Weedy host species act as reservoirs of inoculum and may harbour genetically diverse pathogen populations. They also, by using these hosts as a green bridge, can survive in the off-seasons. In other words, the weeds associated with the main crop will allow the pathogen to survive in different conditions because of different phenological cycles (Burdon and Thrall 2008). Uncultivated hosts of pathogens can act as alternative hosts and inoculum sources between or within growth seasons as for Fusarium graminearum (Mourelos et al. 2014) and Puccinia tritici (Burdon and Roelfs 1985). Recently, it was shown that weedy Hordeum hosts of R. commune act as alternative hosts which harbour more pathotypes, more variation in the nip1 virulence gene and has high transmission rates of the pathogen from the weed to cultivated barley (Linde et al. 2016). Pathogens may also have higher rates of sexual reproduction on alternate hosts such as for Puccinia striiformis f. sp. tritici (Ali et al. 2014) and Pyrenophora teres (Linde and Smith 2019), although this could not be shown for R. commune (Linde et al., 2016).

Previous studies in Iran have only reported R. commune from barley (Ershad 2009), with a recent report on oats and L. multiflorum (Seifollahi et al., 2018). This recent report identified R. commune on ryegrass and oats using microsattelite markers. Here we aim to (1) confirm the identity of scald agents on Poaceae in Iran using molecular and morphological methods and (2) investigate whether R. commune from uncultivated grasses can infect cultivated barley and thus whether uncultivated grasses act as possible inoculum sources on cultivated barley. This is especially important given the large acreage of barley and high potential yield losses caused by the disease in Iran (Beigi et al. 2013).

Material and methods

Collection of infected leaves from wild grasses and barley

Infected leaves from uncultivated grasses (Fig. 1) and barley from Eyvan, Baghmalek and Gorgan regions of Iran were collected in spring of 2014 and 2015. These regions are located in the provinces of Ilam, Khuzestan and Golestan, respectively. The number of uncultivated grass leaves collected varied depending on disease incidence (2–12). Leaves from uncultivated grasses were collected from inside or adjacent to barley fields. Infected leaves of L. multiflorum were obtained from Gorgan, whereas infected leaves of Hordeum vulgare ssp. spontaneum and Avena sativa were obtained from Eyvan, and leaves of H. murinum ssp. glaucum were from Eyvan and Baghmalek.

Fig. 1
figure1

Symptoms of scald caused by Rhynchosporium commune on uncultivated grasses in barley fields (a) Lolium multiflorum (b) Hordeum vulgare ssp. spontaneum (c) Hordeum murinum ssp. glaucum, (d) complex scald lesions with other diseases on Avena sativa leaves which scald lesions highlighted with the red arrows

Fungal isolation

Infected leaves were surface disinfected in 1:1 solution of 70% ethanol and 1% sodium hypochlorite, then washed twice with sterile ionized water for 1 min, placed on 1% water agar medium (Brown 1985) and incubated under continuous dark and 17 °C for 10 days for sporulation. After sporulation, all samples were purified by single-spore method. Purified fungal isolates were kept on 1% water agar medium and maintained in the Fungal Culture Collection of Iran at the Iranian Research Institute of Plant Protection (Accession numbers of isolates in Supplementary data, Table 1S).

Table 1 Rhynchosporium isolate groups based on conidial dimensions. Groups were constructed based on significant differences in spore dimensions using a Tukey test

Identification of uncultivated plants

Host plants from which Rhynchosporium isolates were obtained from, were identified to species level with DNA sequencing. DNA of all hosts was extracted using the method of Murray and Thompson (1980). The matK and rbcL loci were amplified using the primer pairs matK390F (5′-cgatctattcattcaatatttc-3′), matK1326R (5′-tctagcacacgaaagtcgaagt-3′) (Cuénoud et al. 2002) and rbcL-F (5′-ttgcaaaggtttcatttacgc-3′), rbcL-R (5′-tacctgcagtcgcattcaag-3′), respectively, to identify Poaceae plants using DNA barcoding (Drumwright et al. 2011). Each 20 μL of PCR reaction contained 4 μL Amplicon taq DNA polymerase red (Amplicon), 0.5 μL of each of two primers matK390F/matK1326R or rbcL-F/rbcL-R (20 pM concentration of each primer), 12 μL sterile deionized water and 3 μL of genomic DNA (5–20 ng). PCR was done in MJ Mini Bio-Rad thermocycler with PCR condition described by Drumwright et al. (2011). PCR products were sent to Macrogen Company in South Korea and sequenced bidirectionally using the Sanger method. Sequences were compared with sequences available in GenBank in BLAST searches to infer host species identity.

Morphological characterization

Sixty eight purified isolates were grown on Lima bean agar (62.5 g Lima beans and 16 g agar per L) medium (LBA62.5) for 10 days in continuous dark at 15 °C (King et al. 2013). After 10 days, microscopic slides were prepared using fungal tissue and a drop of lactophenol cotton blue. To identify species based on morphological characteristics, conidial shape (beak- or cylindrical shape of conidia) and dimension were investigated. Conidial dimensions were measured using an Olympus CX21FS1 light microscope. Both length and width of 30 mature conidia for each isolate were measured by using an ocular micrometre. At first, total means of conidial lengths and widths (30 conidial sizes for each 68 isolate, totally mean of 2040 conidial lengths and widths) with their standard error were calculated using the Minitab v18.1 software. The means of conidial dimension of all isolates were compared using ANOVA and then grouped using pairwise comparison of means in a Tukey test to select isolates with significant differences in length or width means of conidia for molecular identification. ANOVA and Tukey tests were performed using the Minitab v18.1 software. To cluster isolates based on conidium length and width and choose isolates with different conidial dimension, at first the relevant number of clusters using Davies–Bouldin index (DBI) (Davies and Bouldin 1979) was determined for K-means cluster analysis of width and length using the NbClust package (Charrad et al. 2014) in R (Team RCD 2011). K-means cluster analysis of conidial width and length was done using Minitab v18.1 software to consider conidial dimensions simultaneously and cluster isolates based on conidial dimension. Then, isolates which were clustered close to the centroid of each cluster, were selected for further molecular identification.

DNA extraction, amplification and sequencing

Purified isolates were grown on Lima bean agar (80 g Lima bean, 4 g sucrose, 4 g malt extract, 4 g yeast extract, 16 g agar per L) medium (LBA80) at 17 °C. After 14 days, 1 mL sterile water was added to each plate and mycelia were scraped with a scalpel. A suspension of spores and mycelium was added to a 100 mL flask containing 50 mL PDB medium and three drops of Chlobiotic® 0.5% solution (0.5 g chloramphenicol per 100 mL). Inoculated flasks were incubated for 2 weeks in a shaker incubator at 17 °C and continuous dark. Mycelial tufts were collected using filtration through filter paper in a Buchner funnel, frozen in liquid nitrogen and crushed using a mortar and pestle. DNA was extracted using the method of Murray and Thompson (1980).

The ITS-rDNA region was amplified using ITS4 and ITS5 primers (White et al. 1990). Each 20 μL PCR reaction contained 4 μL Amplicon Taq DNA polymerase red (Amplicon), 0.5 μL of each of two primers (20 pM concentration of each primer), 12 μL sterile deionized water and 3 μL of genomic DNA (5–20 ng). Amplification was conducted using a MJ Mini Bio-Rad thermocycler with the following temperature cycles: initial denaturing 94 °C for 5 min, 35 cycles, 94 °C for 30 s, 58 °C for 1 min, 72 °C for 30 s and final extension 72 °C for 5 min.

A partial region of the β-tubulin (TUBB) gene was amplified using primers BTUB-21F (5’-ATGCGTGAAATCGTACGTCAC-3′) and BTUB-615R (5’-TGACCGAAAGGACCAGCACG-3′) (Zaffarano et al. 2008). Each 20 μL PCR reaction contained 4 μL Amplicon Taq DNA polymerase red (Amplicon), 0.5 μL of each of two primers (20 pM concentration of each primer), 12 μL sterile deionized water and 3 μL of genomic DNA (5–20 ng). Amplification was conducted using a MJ Mini Bio-Rad thermocycler with following temperature cycles: initial denaturing in 96 °C for 2 min, 35 cycles, 96 °C for 1 min, 65 °C for 1 min, 72 °C for 1 min and final extension in 72 °C for 5 min. Amplicon sequencing of both loci were performed by Macrogen Company in South Korea.

Phylogenetic analyses

Phylogenetic analyses using sequences of the ITS region and concatenate sequences of ITS and TUBB loci were conducted. ITS sequences obtained for 27 isolates (Supplementary data, Table 1S) and reference ITS sequences from different species of Rhynchosporium by Zaffarano et al. (2008) and King et al. (2013) were imported into MEGA v. 7.0.26 (Kumar et al. 2016) for phylogenetic analyses. All sequences were aligned using CLUSTAL W (Thompson et al. 1994). To determine the best model for inferring phylogenetic trees, a model analysis was conducted based on a maximum likelihood method in MEGA v. 7.0.26 (Kumar et al. 2016). The Kimura 2-parameter model with the lowest Bayesian information criterion (BIC) was considered as the best model. Phylogenetic constructions were performed using maximum likelihood (ML), Kimura 2-parameter model and performing 1000 bootstrap replications in MEGA v. 7.0.26 on single locus and concatenated ITS and TUBB loci (Kumar et al. 2016). To infer phylogenetic trees using both sequences of ITS and TUBB, 17 isolates from this study (Supplementary data, Table 1S) along with reference sequences of 18 haplotypes or isolates by Zaffarano et al. (2008) and King et al. (2013) were included in the analyses.

Pathogenicity tests

Seven isolates representing all hosts from Iran were grown on LBA80 medium. Plates were incubated at 17 °C in the dark for 21 days. Then the spore suspension was prepared by adding 2 mL sterilized distilled water and scraping off the mycelia with a microscope slide. The suspension of spores and mycelia was filtered through double cheesecloth and propagules counted using a haemocytometer. Spore suspensions were diluted to a final concentration 5 × 105 spores per mL.

For pathogenicity trials, four barley cultivars, Bahman, Ansar, Local and Digger which were previously identified as respectively semi-resistant, resistant, susceptible and susceptible to barley scald, were used. Furthermore, three uncultivated grass species, namely, A. sativa, H. vulgare ssp. spontaneum and H. murinum were grown at 25 °C for 27 days for inoculation. All uncultivated hosts used in the pathogenicity trials were obtained from the Iran-National Plant Gene Bank. Uncultivated grasses were planted 6 days prior to barley cultivars to reach similar developmental stages for inoculation. Twenty microliters of Tween-20 was added to 50 mL spore suspension. Then the plants were sprayed with mentioned suspension until run-off. After inoculation, plants were incubated at 15 °C saturated humidity for 90 h. Then humidity was decreased to 75%, and the conditions were changed to 14 h light at 17 °C and 10 h darkness at 13 °C. To perform Koch’s postulates, fungi were re-isolated from infected leaves following the methods described above. Re-isolations were performed for all observed lesions. Isolates were identified using molecular phylogenetic methods as described above.

Pathogenicity data analyses

The percentage of leaf area infected was measured 28 days after inoculation. Images were captured from the infected leaves using a SAMSUNG Galaxy J5 (2016) smartphone with CCD sensor camera was used. A script was implemented in the image processing software MATLAB R2013a (Matlab 2013) to process the captured images. The leaf (target region of image) was well separated from the background (nontarget region) via the Otsu algorithm (Otsu, 1979). The percentage of leaf area infected was calculated as the ratio of the number of pixels in the infected area to the total leaf area.

Percentage leaf areas infected was Arc Sin transformed to secure normal distribution, and the new data were considered as the response variable. The effect of host type on resistance against different isolates was studied using a factorial experiment in a completely randomized design with three replications. The hosts that were inoculated with isolates, the isolates and interaction between hosts, and isolates were considered as the fixed factors. Analysis of variance was performed using the SAS statistical programme v9.4 to test if there is a significant difference among the percentage leaf areas infected of hosts and caused by isolates, as well as for the interaction between hosts and isolates. Post hoc tests using a t-test (LSD) were conducted to test for significant differences between hosts or isolates. Also, ANOVA and mean comparisons in each host for different isolates and for each isolate for different hosts were done. Then, post hoc tests using a t-test (LSD) were conducted to test for significant differences between hosts or isolates separately.

Results

Identification of hosts

Plants from which scald-like symptoms were observed, showed closest GenBank matches to H. murinum ssp. glaucum, H. vulgare ssp. spontaneum, L. multiflorum and A. sativa. The closest matches for the partial sequences of rbcL locus between one host and L. multiflorum shared 99% identity (GenBank accession number MG021809). Other hosts showed 99–100% identity for partial rbcL sequences to H. murinum ssp. glucum (GenBank accession numbers MG021810–12). Also, 100% identity was shared in partial sequences of the matK locus between one host and A. sativa (GenBank accession number MG021815). The same identity was shared between one host and H. vulgare ssp. spontaneum for partial sequences of rbcL and matkK loci (GenBank accession number MG021813–14).

Morphology of conidia

Conidial morphology of 68 isolates of Rhynchosporium spp. from five hosts was examined. All isolates had beak-shaped conidia. The mean conidial length for all isolates was 16.79 with standard error of the mean = 0.0460. The mean of conidial widths was 3.0623 with standard error of the mean 0.00668. The means of conidial length for each isolate varied between 15.00 and 22.53 μm, with conidial width ranging between 2.73 and 3.50 μm. This range of conidial dimension can be observed in sample photos of conidia (Fig. 2). ANOVA analysis showed not only that isolates differed significantly in mean conidial width but also in their mean conidial length (Fwidth = 6.37, P = 0.000; Flength = 11.61, P = 0.000). The isolates grouped into three to four groups based on significant differences (P ≤ 0.05) of a Tukey’s test on conidial width and conidial length, respectively (Table 1). Using the Davies–Bouldin index (DBI), five clusters were determined based on combined conidial dimensions (Table 2). A K-means cluster analysis of length and width determined length and width centroids for each cluster (Table 2).

Fig. 2
figure2

Differences in spore length and width of Rhynchosporium commune isolates can be observed in a–f. Scale bars are 10 μm. All Rhynchosporium commune spores analysed had beak-shaped conidia

Table 2 K-Means cluster analysis of length and width of Rhynchosporium commune conidia

Phylogenetic analyses based on ITS sequences

For phylogenetic analyses, isolates were selected to represent the different groups identified with a Tukey test based on conidial dimensions. Isolates which clustered close to the K-means cluster centroid were included in the analyses. In addition, some isolates representing all hosts were included for molecular identification. In total, 27 isolates were sequenced for the ITS locus (GenBank accession number MG015670–96) (Supplementary data, Table 1S). The phylogenetic trees showed that previously identified species of Rhynchosporium clustered separately with high bootstrap support. An exception was two closely related species of R. lolii and R. orthosporum which clustered together. All isolates from this study formed a monophyletic, well-supported (bootstrap 88%) group with R. commune (Fig. 3).

Fig. 3
figure3

Maximum likelihood phylogeny of the internal transcribed spacer region sequences from Rhynchosporium. The numbers above branches are frequencies of 1000 replicates. Except for Rhynchosporium orthosporum and Rhynchosporium lolii, each species form a monophyletic group. Reference sequences (all H, 7LM11, 4LM11, 13LP11, RS04ITD62, 59DG09) were sourced from (Zaffarano et al. 2008) and (King et al. 2013)

Phylogeny based on ITS and TUBB sequences

The addition of the locus TUBB in an ITS-TUBB concatenated dataset of 17 isolates from this study (GenBank accession number in Table 1S) resulted in the same phylogenetic topology for Rhynchosporium species as with ITS alone. Again all Rhynchosporium species except R. lolii and R. orthosporum formed well-supported clades. Also, all 17 isolates from this study formed a well-supported clade together with R. commune reference isolates based on ML statistical methods (88%) (Fig. 4).

Fig. 4
figure4

Maximum likelihood phylogeny of concatenated the internal transcribed spacer region and beta-tubulin sequences from Rhynchosporium. The numbers above branches are frequencies of 1000 replicates. Except for Rhynchosporium orthosporum and Rhynchosporium lolii, each species form a monophyletic group. Reference sequences (all H, 7LM11, 4LM11, 13LP11, RS04ITD62, 59DG09) were sourced from (Zaffarano et al. 2008) and (King et al. 2013)

Pathogenicity test

The first symptoms appeared 14 days after inoculation on H. vulgare ssp. spontaneum and after 17 days on barley and H. murinum. Twenty-two days after inoculation, small lesions were found on A. sativa leaves (Fig. 5). Re-isolation from all lesions yielded R. commune, confirming Koch’s postulates. R. commune was also re-isolated from both kinds of lesions observed on A. sativa leaves as seen in Fig. 5. Isolate G46 from barley could cause symptoms on four barley cultivars tested, as well as on H. vulgare ssp. spontaneum, H. murinum and A. sativa (Table 3). All Hordeum spp. tested was susceptible to isolates from A. sativa, H. murinum ssp. glaucum and L. multiflorum (Table 3). The one isolate tested from H. vulgare ssp. spontaneum caused infection in all species of Hordeum except H. murinum (Table 3). Only three of the seven isolates tested, one each from barley, H. murinum ssp. glaucum and A. sativa, could infect A. sativa (Table 3).

Fig. 5
figure5

Two kinds of symptoms of scald caused by Rhynchosporium commune on the leaves of Avena sativa, 28 days after inoculation with isolate G46. (a) typical scald lesions (b) the reddish brown band on the leaf margin

Table 3 The average percentage leaf infected area of four barley cultivars, Hordeum murinum, Hordeum vulgare ssp. spontaneum and Avena sativa, infected with seven Rhynchosporium commune isolates from cultivated barley and uncultivated grasses in Iran

Analysis of variance showed that hosts, isolates and the interaction between hosts and isolates differed significantly (P < 0.0001) for percentage leaf area infected (Table 4). Comparing the means of percentage leaf area infected in hosts across all isolates showed that H. vulgare ssp. spontaneum had the highest percentage leaf area infection (mean = 82.23) (P < 0.05). The lowest (P < 0.05) percentage leaf area infection was observed on A. sativa (mean = 0.12) (Table 3). Means comparison of the percentage leaf areas infected across all hosts showed that the isolate JM2 (mean = 71.31) produced significantly (P < 0.05) the highest infection, whereas isolate U5 showed the least aggressiveness (46.23) (Table 3). Isolate AG42 obtained from L. multiflorum showed the least aggressiveness in some of the hosts (57.14%). The isolates from H. murinum and H. vulgare ssp. spontaneum showed the most aggressiveness on these hosts, but in general, we found no relation between the aggressiveness of isolates and the host species from which the isolates were obtained from. A. sativa was significantly (P < 0.05) the most resistant host against all isolates (Table 3).

Table 4 Analysis of variance for percentage leaf areas infected for seven Rhynchosporium commune isolates inoculated onto cultivated and uncultivated hosts

Discussion

This study used morphological characterization and phylogenetic analyses of the ITS and TUBB loci to identify Rhynchoporium species from barley and wild grasses. Morphologically, all examined isolates were typical of Rhynchosporium spp. with beak-shaped conidia (Heinsen 1901). Given the reported host specificity of Rhynchosporium species (Zaffarano et al. 2011) and our observation of beak-shaped conidia, as well as preliminary identification of these isolates (Seifollahi et al. 2018) using microsatellites, we hypothesized that the isolates from Hordeum spp., L. multiflorum and A. sativa from Iran belong to R. commune.

In this study, a large and significant variation in conidial width and length was observed for R. commune isolates. Comparison of the mean conidial lengths and widths of our study with mean conidial lengths and widths of King’s et al. study (2013) using a t-test showed that the differences are not significant (P values for widths and lengths were 0.196 and 0.984, respectively). Considering the large sample size measured in this study, compared to small sample sizes in the previous study (King et al. 2013), we suggest that conidial size is not affected by isolate origin (e.g. Iran, France, Switzerland and some parts of UK) or climate.

Rhynchosporium spp. are considered to have a degree of host specificity to Hordeum and Bromus (Zaffarano et al. 2011). However, in this and previous studies, R. commune was also isolated from L. multiflorum (King et al. 2013) and now also confirmed on A. sativa. Furthermore, R. orthosporum also infects L. multiflorum (Wilkins 1973) and is not restricted to Dactylis species, suggesting there is limited host specificity in this genus. However, previously only R. secalis, a pathogen of rye has been reported from A. sativa (Mułenko et al. 2008).

In this study, R. commune isolates from H. vulgare, H. vulgare ssp. spontaneum, H. murinum ssp. glaucum, L. multiflorum and A. sativa could all infect cultivated barley. In Australia, it was found that weedy Hordeum significantly contributed to virulence evolution of R. commune on barley, and pathogen transmission from the weed to cultivated barley was inferred (Linde et al. 2016), posing a risk to cultivated barley production. Confirming previous studies (Zaffarano et al., 2011; King et al., 2013; Seifollahi et al., 2018), we obtained R. commune from Hordeum spp. and L. multiflorum. It has long been hypothesized that alternative hosts, in this case uncultivated grasses, contribute to pathogen evolution (Burdon and Thrall 2008). The weedy species contain many resistant genes (Ali, 1981; Jarosz & Burdon, 1996; Genger. et al., 2005) and thus may harbour more aggressive isolates which have high migration rates into barley (Linde et al. 2016). Here, the most aggressive isolate (JM2) were isolated from H. murinum ssp. glaucum, which may be transmitted to cultivated barley. Furthermore, considering the ability of asexual genetic exchanges and the horizontal gene transmission through the hyphal anastomosis among isolates of R. commune (Forgan et al. 2007), it is possible to transfer virulence genes from isolates of weedy host species to barley isolates.

Weedy hosts may be where fungal sexual reproduction occurs in some pathogens such as Diaporthe phaseolorum var. caulivora (Black et al. 1996). Sexual recombination results in combining virulences to create new pathotypes. Sexual reproduction is feasible on the weeds as both mating types of R. commune were found on them (Seifollahi et al. 2018). If sexual reproduction indeed occurs on these weedy hosts, control of the pathogen will be difficult, so that by combining virulences and increase evolutionary potential via sexual recombination on alternative hosts, the pathogen may be better equipped to deal with, e.g. fungicide applications, resistant cultivars and off-season bottlenecks.

This study suggests that not all R. commune isolates can infect A. sativa, suggesting a differential effector gene interaction. Also, the isolates able to infect A. sativa had different levels of aggressiveness. Thus management of oats as a host of more aggressive isolates in barley fields, as well as uncultivated Hordeum species as a host for R. commune in general, should be considered to protect cultivated barley from R. commune infection and evolution.

This research showed that the host range of R. commune is not limited to Hordeum spp. and Bromus spp., but it can also infect other grasses in areas where the disease level is high. Cross-inoculation results highlight the role of uncultivated grasses as a inoculum source and that they may harbour aggressive isolates. Therefore, the identification and control of secondary hosts will reduce inoculum, disrupt the life cycle of the pathogen and reduce its evolutionary ability. Furthermore, breeding of resistant cultivars will likely be unsuccessful without considering R. commune isolates from uncultivated hosts in breeding programmes. Pathogenicity studies of Iranian R. commune isolates on other cereals is suggested for future research to determine if these isolates have an even broader host range.

References

  1. Ali S, Gladieux P, Rahman H et al (2014) Inferring the contribution of sexual reproduction, migration and off-season survival to the temporal maintenance of microbial populations: a case study on the wheat fungal pathogen Puccinia striiformis f. sp. tritici. Mol Ecol 23:603–617

  2. Ali SM (1981) Barley grass as a source of pathogenic variation in Rhynchosporium secalis. Aust J Agric Res 32:21–25

  3. Avrova A, Knogge W (2012) Rhynchosporium commune: a persistent threat to barley cultivation. Mol Plant Pathol 13:986–997

  4. Beigi S, Zamanizadeh H, Razavi M, Zare R (2013) Genetic diversity of Iranian isolates of barley scald pathogen (Rhynchosporium secalis) making use of molecular markers. J Agric Sci Technol 15:843–854

  5. Black BD, Padgett GB, Russin JS et al (1996) Potential weed hosts for Diaporthe phaseolorum var. caulivora, causal agent for soybean stem canker. Plant Dis 80:763–765

  6. Brown JS (1985) Pathogenic variation among isolates of Rhynchosporium secalis from cultivated barley growing in Victoria, Australia. Euphytica 34:129–133

  7. Burdon JJ, Roelfs AP (1985) The effect of sexual and asexual reproduction on the isozyme structure of populations of Puccinia graminis. Phytopathology 75:1068–1073

  8. Burdon JJ, Thrall PH (2008) Pathogen evolution across the agro-ecological interface: implications for disease management. Evol Appl 1:57–65

  9. Caldwell RM (1937) Rhynchosporium scald of barley, rye, and other grasses. J Agric Res 55:175–198

  10. Charrad M, Ghazzali N, Boiteau V, et al (2014) Package “NbClust.” J Stat Softw 61:1–36

  11. Cuénoud P, Savolainen V, Chatrou LW et al (2002) Molecular phylogenetics of Caryophyllales based on nuclear 18S rDNA and plastid rbcL, atpB, and matK DNA sequences. Am J Bot 89:132–144

  12. Davies DL, Bouldin DW (1979) A cluster separation measure. IEEE Trans Pattern Anal Mach Intell 1:224–227

  13. Drumwright AM, Allen BW, Huff KA et al (2011) Survey and DNA barcoding of Poaceae in flat rock cedar glades and barrens state natural area, Murfreesboro, Tennessee. Castanea 76:300–310

  14. Ershad J (2009) Fungi of Iran, 3rd edn. Agricultural Research, Education and Extention Organization, Tehran

  15. Forgan AH, Knogge W, Anderson PA (2007) Asexual genetic exchange in the barley pathogen Rhynchosporium secalis. Phytopathology 97:650–654

  16. Foster SJ, Fitt BDL (2003) Isolation and characterization of the mating-type (MAT) locus from Rhynchosporium secalis. Curr Genet 44:277–286

  17. Frank AB (1897) Über die Zerstörung der Gerste durch einen neuen Getreidepilz. Wochenschr Brau 42:518–520

  18. Genger RK, Nesbitt K, Brown DA et al (2005) A novel barley scald resistance gene: genetic mapping of the Rrs15 scald resistance gene derived from wild barley, Hordeum vulgare ssp. spontaneum. Plant Breed 124:137–141

  19. Goodwin SB (2002) The barley scald pathogen Rhynchosporium secalis is closely related to the discomycetes Tapesia and Pyrenopeziza. Mycol Res 106:645–654

  20. Heinsen E (1901) Beobachtungen uiber den neuen Getreidepilz Rhynchosporium graminicola. Jahrb Hamburg Wiss Anstalt 18:43–55

  21. Jarosz AM, Burdon JJ (1996) Resistance to barley scald (Rhynchosporium secalis) in wild barley grass (Hordeum glaucum and Hordeum leporinum) populations in South-Eastern Australia. Aust J Agric Res 47:413–425

  22. King KM, West JS, Brunner PC et al (2013) Evolutionary relationships between Rhynchosporium lolii sp. nov. and other Rhynchosporium species on grasses. PLoS One 8:e72536. https://doi.org/10.1371/journal.pone.0072536

  23. Kumar S, Stecher G, Tamura K (2016) MEGA7: molecular evolutionary genetics analysis version 7.0 for bigger datasets. Mol Biol Evol 33:1870–1874

  24. Linde CC, Smith LM, Peakall R (2016) Weeds, as ancillary hosts, pose disproportionate risk for virulent pathogen transfer to crops. BMC Evol Biol 16:101. https://doi.org/10.1186/s12862-016-0680-6

  25. Linde CC, Smith LM (2019) Host specialisation and disparate evolution of Pyrenophora teres f. teres on barley and barley grass. BMC Evol Biol 19:139. https://doi.org/10.1186/s12862-019-1446-8

  26. Matlab R (2013) Version 8.1. 0.604 (R2013a). Natrick, Massachusetts MathWorks Inc.

  27. Mourelos CA, Malbrán I, Balatti PA et al (2014) Gramineous and non-gramineous weed species as alternative hosts of Fusarium graminearum, causal agent of Fusarium head blight of wheat, in Argentina. Crop Prot 65:100–104

  28. Mułenko W, Majewski T, Ruszkiewicz-Michalska M (2008) A preliminary checklist of micromycetes in Poland. Polish Academy of Sciences, Poland

  29. Murray MG, Thompson WF (1980) Rapid isolation of high molecular weight plant DNA. Nucleic Acids Res 8:4321–4326

  30. Otsu N (1979) A threshold selection method from gray-level histograms. IEEE Trans Syst Man Cybern 9:62–66

  31. Oudemans C (1897) Observations mycologiques. Konink Akad Wetensch Amsterdam 6:86–92

  32. Seifollahi E, Sharifnabi B, Javan-Nikkhah M, Linde CC (2018) Low genetic diversity of Rhynchosporium commune in Iran, a secondary Centre of barley origin. Plant Pathol 67:1725–1734

  33. Shipton WA, Boyd WJR, Ali SM (1974) Scald of barley. Rev Plant Pathol 53:839–861

  34. Team RDC (2011) R development Core team: R: a language and environment for statistical computing. R Foundation for Statistical Computing, Vienna

  35. Thompson JD, Higgins DG, Gibson TJ (1994) CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res 22:4673–4680

  36. Welty RE, Metzger R (1996) First report of scald of triticale caused by Rhynchosporium secalis in North America. Plant Dis 80:1220–1223

  37. White TJ, Bruns T, Lee S, Taylor J (1990) Amplification and direct sequencing of fungal ribosomal RNA genes for phylogenetics. PCR Protoc 18:315–322

  38. Wilkins P (1973) Infection of Lolium multifiorum with Rhynchosporium species. Plant Pathol 22:107–111

  39. Zaffarano PL, McDonald BA, Linde CC (2011) Two new species of Rhynchosporium. Mycologia 103:195–202

  40. Zaffarano PL, McDonald BA, Linde CC (2008) Rapid speciation following recent host shifts in the plant pathogenic fungus Rhynchosporium. Evolution 62:1418–1436

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Acknowledgements

E.S. thanks Isfahan University of Technology for financial support. The authors thank S. Beigi, J. Shokri and A. Ahmadpour who contribute during sampling. Also, we thank J. Gholami for technical assistance, L. Smith for guidance in pathogenicity test, S. Taghadomi-Saberi for image processing, M.R. Sabzalian and M. Isapareh for guidance with statistical analysis.

Funding

This study was funded by Isfahan University of Technology (grant number NA).

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Correspondence to E. Seifollahi.

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Seifollahi, E., Sharifnabi, B., Javan-Nikkhah, M. et al. Scald on gramineous hosts in Iran and their potential threat to cultivated barley. Mycol Progress 19, 223–233 (2020). https://doi.org/10.1007/s11557-019-01553-8

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Keywords

  • Avenae
  • β-tubulin
  • Host specificity
  • ITS
  • Morphology
  • Rhynchosporium