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1. Australia: A Continent Without Native Powdery Mildews? The First Comprehensive Catalog Indicates Recent Introductions and Multiple Host Range Expansion Events, and Leads to the Re-discovery of Salmonomyces as a New Lineage of the Erysiphales

Author

Levente Kiss, Niloofar Vaghefi, Kaylene Bransgrove, John D. W. Dearnaley, Susumu Takamatsu, Yu Pei Tan, Craig Marston, Shu-Yan Liu, Dan-Ni Jin, Dante L. Adorada, Jordan Bailey, Maria Graciela Cabrera de Álvarez, Andrew Daly, Pamela Maia Dirchwolf, Lynne Jones, Thuan Dat Nguyen, Jacqueline Edwards, Wellcome Ho, Lisa Kelly, Sharl J. L. Mintoff, Jennifer Morrison, Márk Z. Németh, Sandy Perkins, Roger G. Shivas, Reannon Smith, Kara Stuart, Ronald Southwell, Unaisi Turaganivalu, Kálmán Zoltán Váczy, Annie Van Blommestein, Dominie Wright, Anthony Young, and Uwe Braun

Subjects
host jumps, host-pathogen interactions, invasive species, obligate biotrophs, plant-microbe interactions, rapid evolution, Microbiology, QR1-502
Abstract

In contrast to Eurasia and North America, powdery mildews (Ascomycota, Erysiphales) are understudied in Australia. There are over 900 species known globally, with fewer than currently 60 recorded from Australia. Some of the Australian records are doubtful as the identifications were presumptive, being based on host plant-pathogen lists from overseas. The goal of this study was to provide the first comprehensive catalog of all powdery mildew species present in Australia. The project resulted in (i) an up-to-date list of all the taxa that have been identified in Australia based on published DNA barcode sequences prior to this study; (ii) the precise identification of 117 specimens freshly collected from across the country; and (iii) the precise identification of 30 herbarium specimens collected between 1975 and 2013. This study confirmed 42 species representing 10 genera, including two genera and 13 species recorded for the first time in Australia. In Eurasia and North America, the number of powdery mildew species is much higher. Phylogenetic analyses of powdery mildews collected from Acalypha spp. resulted in the transfer of Erysiphe acalyphae to Salmonomyces, a resurrected genus. Salmonomyces acalyphae comb. nov. represents a newly discovered lineage of the Erysiphales. Another taxonomic change is the transfer of Oidium ixodiae to Golovinomyces. Powdery mildew infections have been confirmed on 13 native Australian plant species in the genera Acacia, Acalypha, Cephalotus, Convolvulus, Eucalyptus, Hardenbergia, Ixodia, Jagera, Senecio, and Trema. Most of the causal agents were polyphagous species that infect many other host plants both overseas and in Australia. All powdery mildews infecting native plants in Australia were phylogenetically closely related to species known overseas. The data indicate that Australia is a continent without native powdery mildews, and most, if not all, species have been introduced since the European colonization of the continent.

Published
2020
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2. Guidelines for the reliable use of high throughput sequencing technologies to detect plant pathogens and pests

Author

Sebastien Massart, Ian Adams, Maher Al Rwahnih, Steve Baeyen, Guillaume J. Bilodeau, Arnaud G. Blouin, Neil Boonham, Thierry Candresse, Anne Chandellier, Kris De Jonghe, Adrian Fox, Yahya Z.A. Gaafar, Pascal Gentit, Annelies Haegeman, Wellcome Ho, Oscar Hurtado-Gonzales, Wilfried Jonkers, Jan Kreuze, Denis Kutjnak, Blanca B. Landa, Mingxin Liu, François Maclot, Marta Malapi-Wight, Hans J. Maree, Francesco Martoni, Natasa Mehle, Angelantonio Minafra, Dimitre Mollov, Adriana G. Moreira, Mark Nakhla, Françoise Petter, Alexander M. Piper, Julien P. Ponchart, Robbie Rae, Benoit Remenant, Yazmin Rivera, Brendan Rodoni, Marleen Botermans, J.W. Roenhorst, Johan Rollin, Pasquale Saldarelli, Johanna Santala, Rose Souza-Richards, Davide Spadaro, David J. Studholme, Stefanie Sultmanis, René van der Vlugt, Lucie Tamisier, Charlotte Trontin, Ines Vazquez-Iglesias, Claudia S.L. Vicente, Bart T.L.H. van de Vossenberg, Marcel Westenberg, Thierry Wetzel, Heiko Ziebell, and Benedicte S. M. Lebas

Subjects
Biointeractions and Plant Health, fungi, food and beverages, Life Science, PE&RC
Abstract

High-throughput sequencing (HTS) technologies have the potential to become one of the most significant advances in molecular diagnostics. Their use by researchers to detect and characterize plant pathogens and pests has been growing steadily for more than a decade and they are now envisioned as a routine diagnostic test to be deployed by plant pest diagnostics laboratories. Nevertheless, HTS technologies and downstream bioinformatics analysis of the generated datasets represent a complex process including many steps whose reliability must be ensured. The aim of the present guidelines is to provide recommendations for researchers and diagnosticians aiming to reliably use HTS technologies to detect plant pathogens and pests. These guidelines are generic and do not depend on the sequencing technology or platform. They cover all the adoption processes of HTS technologies from test selection to test validation as well as their routine implementation. A special emphasis is given to key elements to be considered: undertaking a risk analysis, designing sample panels for validation, using proper controls, evaluating performance criteria, confirming and interpreting results. These guidelines cover any HTS test used for the detection and identification of any plant pest (viroid, virus, bacteria, phytoplasma, mycetes, nematodes, arthropods, plants) from any type of matrix. Overall, their adoption by diagnosticians and researchers should greatly improve the reliability of pathogens and pest diagnostics and foster the use of HTS technologies in plant health., Scientific publication adapted from the deliverable 2.2 of the Valitest project - www.valitest.eu

Published
2022
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3. Facilitating the adoption of high-throughput sequencing technologies as a plant pest diagnostic test in laboratories : A step-by-step description

Author

Benedicte Lebas, Ian Adams, Maher Al Rwahnih, Steve Baeyen, Guillaume J. Bilodeau, Arnaud G. Blouin, Neil Boonham, Thierry Candresse, Anne Chandelier, Kris De Jonghe, Adrian Fox, Yahya Z. A. Gaafar, Pascal Gentit, Annelies Haegeman, Wellcome Ho, Oscar Hurtado‐Gonzales, Wilfried Jonkers, Jan Kreuze, Denis Kutjnak, Blanca Landa, Mingxin Liu, François Maclot, Martha Malapi‐Wight, Hano J. Maree, Francesco Martoni, Natasha Mehle, Angelantonio Minafra, Dimitre Mollov, Adriana Moreira, Mark Nakhla, Françoise Petter, Alexander M. Piper, Julien Ponchart, Robbie Rae, Benoit Remenant, Yazmin Rivera, Brendan Rodoni, Johanna W. Roenhorst, Johan Rollin, Pasquale Saldarelli, Johanna Santala, Rose Souza‐Richards, Davide Spadaro, David J. Studholme, Stefanie Sultmanis, René van der Vlugt, Lucie Tamisier, Charlotte Trontin, Ines Vazquez‐Iglesias, Claudia S. L. Vicente, Bart T. L. H. Vossenberg, Thierry Wetzel, Heiko Ziebell, Sebastien Massart, Gembloux Agro-Bio Tech [Gembloux], Université de Liège, Fera Science Ltd, University of California [Davis] (UC Davis), University of California (UC), Research Institute for Agricultural, Fisheries and Food (ILVO), Canadian Food Inspection Agency (CFIA), Newcastle University [Newcastle], Biologie du fruit et pathologie (BFP), Université de Bordeaux (UB)-Institut National de Recherche pour l’Agriculture, l’Alimentation et l’Environnement (INRAE), 1Walloon Agricultural Research Centre, Gembloux, Belgium 2The Research Institute of Organic Agriculture (FIBL), Frick, Switzerland, Institute for Epidemiology and Pathogen Diagnostics, Julius Kühn-Institut - Federal Research Centre for Cultivated Plants, Laboratoire de santé des végétaux (LSV Angers), Laboratoire de la santé des végétaux (LSV), Agence nationale de sécurité sanitaire de l'alimentation, de l'environnement et du travail (ANSES)-Agence nationale de sécurité sanitaire de l'alimentation, de l'environnement et du travail (ANSES), Ministry for Primary Industries, Animal Plant Health Inspection Service, APHIS-USDA, Humble, Texas, 77338, USA., World Agroforestry Center [CGIAR, Pérou] (ICRAF), World Agroforestry Center [CGIAR, Kenya] (ICRAF), Consultative Group on International Agricultural Research [CGIAR] (CGIAR)-Consultative Group on International Agricultural Research [CGIAR] (CGIAR), National Institute of Biology [Ljubljana] (NIB), Centro de Investigaciones Biológicas (CSIC), Consejo Superior de Investigaciones Científicas [Madrid] (CSIC), University of Tasmania [Hobart, Australia] (UTAS), Stellenbosch University, Citrus Research International, Partenaires INRAE, Agriculture Victoria (AgriBio), CNR Istituto per la Protezione Sostenibile delle Piante [Torino, Italia] (IPSP), National Research Council of Italy | Consiglio Nazionale delle Ricerche (CNR), Horticultural Crops Research Unit, USDA-ARS : Agricultural Research Service, International Plant Protection Convention, European and Mediterranean Plant Protection Organization - Organisation Européenne et Méditerranéenne pour la Protection des Plantes (EPPO), Liverpool John Moores University (LJMU), Netherlands Food and Consumer Product Safety Authority (NVWA), Finnish Food Authority, International Seed Federation (ISF), Università degli studi di Torino = University of Turin (UNITO), University of Exeter, Wageningen University and Research [Wageningen] (WUR), Instituto Nacional de Investigação Agrária e Veterinária = National Institute for Agrarian and Veterinary Research [Oeiras, Portugal] (INIAV), Dienstleistungszentrum Ländlicher Raum DLR Rheinpfalz (DLR Rheinpfalz), Julius Kühn-Institute, Federal Research Centre for Cultivated Plants-Institute for Biological Control, European Project: 773139,VALITEST, and European Commission

Subjects
plant pest diagnostic test, Biointeractions and Plant Health, [SDV]Life Sciences [q-bio], Life Science, high-throughput sequencing, Plant Science, Horticulture, PE&RC, Agronomy and Crop Science, [SDV.BV.PEP]Life Sciences [q-bio]/Vegetal Biology/Phytopathology and phytopharmacy
Abstract

High-throughput sequencing (HTS) is a powerful tool that enables the simultaneous detection and potential identification of any organisms present in a sample. The growing interest in the application of HTS technologies for routine diagnostics in plant health laboratories is triggering the development of guidelines on how to prepare laboratories for performing HTS testing. This paper describes general and technical recommendations to guide laboratories through the complex process of preparing a laboratory for HTS tests within existing quality assurance systems. From nucleic acid extractions to data analysis and interpretation, all of the steps are covered to ensure reliable and reproducible results. These guidelines are relevant for the detection and identification of any plant pest (e.g. arthropods, bacteria, fungi, nematodes, invasive plants or weeds, protozoa, viroids, viruses), and from any type of matrix (e.g. pure microbial culture, plant tissue, soil, water), regardless of the HTS technology (e.g. amplicon sequencing, shotgun sequencing) and of the application (e.g. surveillance programme, phytosanitary certification, quarantine, import control). These guidelines are written in general terms to facilitate the adoption of HTS technologies in plant pest routine diagnostics and enable broader application in all plant health fields, including research. A glossary of relevant terms is provided among the Supplementary Material., This article is based upon work from the work package 2 of the project VALITEST (https://www.valitest.eu/), supported by the European Union’s Horizon 2020 research and innovation programme under grant agreement no. 773139.

Published
2022
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4. First record of Bursaphelenchus hildegardae Braasch et al., 2006 (Nematoda) in New Zealand with updated information on morphology, sequencing and a key to species of the eggersi-group

Author

Michael Surrey, Wellcome Ho, Zeng Qi Zhao, Brent Rogan, B. J. R. Alexander, Carolyn Bleach, and Milen Marinov

Subjects
Bark beetle, Nematoda, Zoology, Bursaphelenchus, Biology, DNA, Ribosomal, Rhabditida, Animals, Animalia, Hylastes ater, Internal transcribed spacer, Ecology, Evolution, Behavior and Systematics, Phylogeny, Taxonomy, Aphelenchidae, Morphometrics, Pinus radiata, Biodiversity, biology.organism_classification, Pinus, Secernentea, Key (lock), Animal Science and Zoology, Taxonomy (biology), New Zealand, Aphelenchida
Abstract

Bursaphelenchus hildegardae Braasch et al., 2006 was collected from pine wood (Pinus radiata) growing in Kaingaroa Timberlands, and a bark beetle, Hylastes ater Paykull, 1800 in New Zealand. This is a new record for B. hildegardae, occuring in New Zealand, and the second report from the southern hemisphere in addition to Australia. In general, the New Zealand isolate of B. hildegardae corresponds well with the description of B. hildegardae given by Braasch et al. (2006) from Germany. The New Zealand isolate is characterized by having an adult body length of 807–1190 μm, medium a ratios (47.5–58.5 for female and 44.6–60.1 for male), b ratios of 9.8–14.5 (female) and 10.2–12.7 (male), c ratios of 18.8–25.2 (female) and 21.6–32.4 (male), c’ ratios of 4.0–4.4 (female) and 2.1–2.7 (male), and is characterised by having three incisures in the lateral fields, thorn-shaped spicules with a distinctly dorsally-bent thin hook-like condylus, and a dorso-ventally visible terminal bursa. In addition, molecular phylogeny using near full length small subunit (SSU), D2/D3 expansion segments of the large subunit (LSU) and the internal transcribed spacer region (ITS1 and 2) of the ribosomal rDNA supports the identification. A key to Bursaphelenchus species in the eggersi-group is given.

Published
2021

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