The complete coat protein nucleotide sequences of 11 Potato virus X isolates from Australia and two from Britain were compared to those of 72 others. On phylogenetic analysis, clade I contained all 11 Australian sequences, and sub-clade II-1 contained the two new British sequences. Clade I isolates were from six different continents, but those in sub-clades II-1 and II-2 were only from Europe and the Americas, respectively. Clade I contained isolates in strain groups 1, 3 and 4, and sub-clades II-1 and II-2, isolates in strain groups 2 and 4. Thus, strain group 4 now occurs within both clades.
The complete coat protein (CP) nucleotide sequences of 13 Potato virus S (PVS) isolates from Australia and three from Europe were compared to those of 37 others. On phylogenetic analysis, the Australian sequences were in PVS(O) sub-clades III and IV, and the European isolates were in sub-clades I and VII. The European isolates invaded Chenopodium spp. systemically, but eight Australian isolates did not. Amino acid sequence differences at the N-terminal ends of the CPs were unrelated to the ability to invade Chenopodium spp. systemically. The acronym PVS(O-CS) is suggested for isolates that invade Chenopodium spp. systemically but are not within clade PVS(A).
The increased occurrence of triazole fungicide resistant strains of Blumeria graminis f. sp. hordei (Bgh) is an economic concern for the barley industry in Australia and elsewhere. High levels of resistance to triazoles in the field are caused by two separate point mutations in the Cyp51 gene, Y136F and S509T. Early detection of these mutations arising in pathogen field populations is important as this allows time for changes in fungicide practices to be adopted, thus mitigating potential yield losses due to fungicide failure and preventing the resistance from becoming dominant. A digital PCR (dPCR) assay has been developed for the detection and quantification of the Y136F and S509T mutations in the Bgh Cyp51 gene. Mutation levels were quantifiable as low as 0.2% in genomic DNA extractions and field samples. This assay was applied to the high throughput screening of Bgh field and bait trial samples from barley growing regions across Australia in the 2015 and 2016 growing seasons and identified the S509T mutation for the first time in the Eastern states of Australia. This is the first report on the use of digital PCR technology for fungicide resistance detection and monitoring in agriculture. Here we describe the potential application of dPCR for the screening of fungicide resistance mutations in a network of specifically designed bait trials. The combination of these two tools constitute an early warning system for the development of fungicide resistance that allows for the timely adjustment of management practices.
As the incidence of fungicide resistance in plant pathogens continues to increase, control of diseases and the management of resistance would be greatly aided by rapid diagnostic methods. Quantitative allele-specific PCR (ASqPCR) is an ideal technique for the in-field analysis of fungicide resistance as it can quantify the frequency of mutations in fungicide targets. We have applied this technique to the fungal pathogen Blumeria graminis f. sp. tritici (Bgt), the causal agent of wheat powdery mildew. In Australia, strobilurin-resistant Bgt was first discovered in 2016. Molecular analysis revealed a nucleotide transversion in the cytochrome b (cytb) gene in the cytochrome bc1 enzyme complex, resulting in a substitution of alanine for glycine at position 143 (G143A). We have developed an in-field ASqPCR assay that can quantify both the resistant (A143) and sensitive (G143) cytb alleles down to 1.67% in host and Bgt DNA mixtures, within 90 min of sample collection. The in situ analysis of samples collected during a survey in Tasmania revealed A143 frequencies ranging between 9–100%. Validation of the analysis with a newly developed laboratory based digital PCR assay found no significant differences between the two methods. We have successfully developed an in-field quantification method, for a strobilurin-resistant allele, by pairing the ASqPCR assay on a lightweight qPCR instrument with a quick DNA extraction method. The deployment of these type of methodologies in the field can contribute to the effective in-season management of fungicide resistance.
In eastern Australia, there have been several as yet unconfirmed reports of Wheat mosaic virus (WMoV) infecting wheat (3). WMoV, previously known as High plains virus (HPV), is transmitted by the wheat curl mite (WCM, Aceria tosichella). It is often found in mixed infections with Wheat streak mosaic virus (WSMV), also transmitted by WCM (2,3). WSMV was first identified in Australia in 2003 (3). In October 2012, stunted wheat plants with severe yellow leaf streaking were common in a field experiment near Corrigin in Western Australia consisting of nine wheat cultivars. These symptoms were also common in two commercial crops of wheat cv. Mace near Kulin. Leaf samples (one per plant) from each location were tested by ELISA using specific antiserum to WMoV (syn. HPV 17200, Agdia, Elkhart, IN). At the field experiment, 20 leaf samples were collected at random from each wheat plot (4 replicates) and tested individually by ELISA. WMoV incidence was 5% for cv. Yipti, 16% for cvs Emu Rock, Wyalkatchem and Mace, 22% for cvs. Corack, Fortune, Calingiri, and Magenta, and 55% for cv. Cobra. From the two commercial wheat crops, 100 leaf samples were collected at random from each and tested by ELISA. WMoV incidence was 2 and 4%. In addition, 50 leaf samples of Hordeum leporinum (barley grass) and 20 of Lolium rigidum (annual ryegrass) were collected and tested by ELISA. WMoV incidence was 2% in H. leporinum, but 0% in L. rigidum. Infected H. leporinum plants were symptomless. Symptomatic wheat leaf samples from both sites were tested by RT-PCR using WMoV specific primers designed from its RNA3 sequence (1). The PCR products (339 bp) were sequenced and lodged in GenBank (Accession Nos KC337341 and KC337342). WMoV isolates from Corrigin (WA-CG12) and Kulin (WA-KU12) had identical sequences. When the nucleic acid sequences of WA-CG12 and WA-KU12 were compared with those of the three other WMoV isolates on GenBank, they had 100% nucleotide sequence identity with a Nebraska isolate (U60141), and 99.7% identity to two United States sweet corn isolates (AY836524 and AY836525). Ten symptomatic wheat plants were collected from each location, transplanted into pots and leaf samples tested individually for WMoV and WSMV (07048, Loewe, Germany) by ELISA. All were infected with both viruses and infested with WCM. WCM-infested glumes (>10 WCM/glume) were placed on the leaf sheaths of 60 wheat plants cv. Calingiri (35 with WA-CG12 and 25 with WA-KU12) and 13 sweet corn plants cv. Snow Gold (WA-CG12 only). In addition, 20 wheat and 10 sweet corn plants were left without infested glumes to be uninoculated controls. All 60 WCM-inoculated wheat plants became stunted with severe leaf streaking. When leaf samples from each plant were tested by ELISA 18 to 30 days later, both viruses were detected. WMoV was detected in all 13 WCM-inoculated sweet corn plants and WSMV in two of them. Plants with WMoV alone initially had short chlorotic leaf streaks that subsequently combined, causing broad streaks. These are typical WMoV symptoms for sweet corn (1). No symptoms developed and no virus was detected in any of the uninoculated wheat or sweet corn control plants. The WMoV nucleotide sequence obtained from an infected sweet corn plant was identical to those of WA-CG12 and WA-KU12. To our knowledge, this is the first confirmed report of WMoV presence in Australia. References: (1) B. S. M. Lebas et al. Plant Dis. 89:1103, 2005. (2) D. Navia et al. Exp. Appl. Acarol. 59:95, 2013. (3) J. M. Skare et al. Virology 347:343, 2006.
scite is a Brooklyn-based organization that helps researchers better discover and understand research articles through Smart Citations–citations that display the context of the citation and describe whether the article provides supporting or contrasting evidence. scite is used by students and researchers from around the world and is funded in part by the National Science Foundation and the National Institute on Drug Abuse of the National Institutes of Health.
hi@scite.ai
10624 S. Eastern Ave., Ste. A-614
Henderson, NV 89052, USA
Copyright © 2024 scite LLC. All rights reserved.
Made with 💙 for researchers
Part of the Research Solutions Family.