Science, Irving K. Barber Faculty of (Okanagan)
Biology, Department of (Okanagan)
DSpace
UBCO
Hrycan, Jared
Doctor of Philosophy - PhD
University of British Columbia
Young vine decline (YVD) fungal pathogens cause internal necrosis, leading to stunted growth and eventual plant mortality within the first few years after planting. Canada’s fast-growing grape and wine industry depends on the import of nursery propagated material but the health status of ready-to-plant nursery grapevines in Canada regarding YVD fungi is unclear. A nursery survey was conducted to investigate the presence or absence and quantity of the most prevalent YVD pathogens reported in other countries. 99% of plants contained at least one YVD pathogen, and overall, a mean of 3 YVD fungal species were found in each grapevine. Pathogen abundance for each fungus in single plants was conducted using droplet digital PCR. Results showed fungal abundance to vary widely between nurseries, grapevines, and different parts of the same grapevines. Additionally, several YVD pathogens were isolated that had not previously been reported in Canada, showing that nursery material can be the source or new fungal pathogens in Canadian vineyards. Phaeomoniella chlamydospora (Pc), one of the main agents of the YVD Petri disease, was found in up to 80% of ready-to-plant nursery grapevines. The presence of Pc in asymptomatic nursery grapevines has led to the hypothesis that it may be a latent pathogen, transitioning from an endophyte to a pathogen under plant stress. Two greenhouse and four field experiments were conducted to determine the effect of water stress, heat stress, j-rooting, and ring nematode infestation on Pc abundance and disease development in young grapevines. Arbuscular mycorrhizal (AM) fungi were inoculated in the greenhouse to investigate whether a reduction in plant stress due to AM fungal colonization would reduce Pc abundance. In the greenhouse, a combination of water stress and AM fungal colonization increased Pc abundance. Young vine decline like symptom development was observed in the field and higher mortality occurred in grapevines subjected to the 50% water treatment compared to plants subjected to the 100% water treatment. Ring nematode infestation increased Pc abundance in the greenhouse in AM inoculated plants compared to nematode negative AM inoculated plants. In the field, plants inoculated with 5,000 Pc spores that were inoculated with ring nematodes had higher overwinter mortality than nematode negative 5,000 Pc inoculated plants These findings suggest a relationship between plant stress, Pc abundance, and subsequent disease development in young grapevines within a few years after planting.
THE EFFECT OF ABIOTIC AND BIOTIC PLANT STRESS ON PHAEOMONIELLA CHLAMYDOSPORA ABUNDANCE AND DISEASE DEVELOPMENT IN YOUNG GRAPEVINES by Jared Hrycan B.Sc. University of British Columbia Okanagan, 2018 A THESIS SUBMITTED IN PARTIAL FULLFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE COLLEGE OF GRADUATE STUDIES (Biology) THE UNIVERSITY OF BRITISH COLUMBIA (Okanagan) July 2024 © Jared Hrycan, 2024 ii The following individuals certify that they have read, and recommend to the College of Graduate Studies for acceptance, a thesis/dissertation entitled: THE EFFECT OF ABIOTIC AND BIOTIC PLANT STRESS ON PHAEOMONIELLA CHLAMYDOSPORA ABUNDANCE AND DISEASE DEVELOPMENT IN YOUNG GRAPEVINES submitted by Jared Hrycan in partial fulfillment of the requirements of the degree of Doctor of Philosophy Dr. Miranda Hart, Irving K. Barber School of Arts and Sciences Supervisor Dr. José Ramón Úrbez-Torres, Agriculture and Agri-Food Canada, Summerland Co-Supervisor Dr. Patricia Bowen, Agriculture and Agri-Food Canada, Summerland Supervisory Committee Member Dr. Thomas Forge, Agriculture and Agri-Food Canada, Summerland Supervisory Committee Member Dr. Louise Nelson, Irving K. Barber School of Arts and Sciences Supervisory Committee Member Dr. Gurcharn Singh Brar, University of British Columbia University Examiner Dr. Philippe E. Rolshausen, University of California, Riverside External Examiner iii Abstract Young vine decline (YVD) fungal pathogens cause internal necrosis, leading to stunted growth and eventual plant mortality within the first few years after planting. Canada’s fast-growing grape and wine industry depends on the import of nursery propagated material but the health status of ready-to-plant nursery grapevines in Canada regarding YVD fungi is unclear. A nursery survey was conducted to investigate the presence or absence and quantity of the most prevalent YVD pathogens reported in other countries. 99% of plants contained at least one YVD pathogen, and overall, a mean of 3 YVD fungal species were found in each grapevine. Pathogen abundance for each fungus in single plants was conducted using droplet digital PCR. Results showed fungal abundance to vary widely between nurseries, grapevines, and different parts of the same grapevines. Additionally, several YVD pathogens were isolated that had not previously been reported in Canada, showing that nursery material can be the source or new fungal pathogens in Canadian vineyards. Phaeomoniella chlamydospora (Pc), one of the main agents of the YVD Petri disease, was found in up to 80% of ready-to-plant nursery grapevines. The presence of Pc in asymptomatic nursery grapevines has led to the hypothesis that it may be a latent pathogen, transitioning from an endophyte to a pathogen under plant stress. Two greenhouse and four field experiments were conducted to determine the effect of water stress, heat stress, j-rooting, and ring nematode infestation on Pc abundance and disease development in young grapevines. Arbuscular mycorrhizal (AM) fungi were inoculated in the greenhouse to investigate whether a reduction in plant stress due to AM fungal colonization would reduce Pc abundance. In the greenhouse, a combination of water stress and AM fungal colonization increased Pc abundance. Young vine decline like symptom development was observed in the field and higher mortality iv occurred in grapevines subjected to the 50% water treatment compared to plants subjected to the 100% water treatment. Ring nematode infestation increased Pc abundance in the greenhouse in AM inoculated plants compared to nematode negative AM inoculated plants. In the field, plants inoculated with 5,000 Pc spores that were inoculated with ring nematodes had higher overwinter mortality than nematode negative 5,000 Pc inoculated plants These findings suggest a relationship between plant stress, Pc abundance, and subsequent disease development in young grapevines within a few years after planting. v Lay Summary The health status of imported ready-to-plant grapevines from nurseries regarding fungal pathogens responsible for young vine decline is unclear in Canada. I tested grapevines from four nurseries using molecular tools to determine whether they were infected with these pathogens and to determine the level of infection. I found that 99% of tested plants were infected with a mean of three fungal pathogen species per plant. The level of infection of these pathogens varied widely from very low to very high quantities of these pathogens. One of the most common young vine decline pathogens was chosen to investigate whether a combination of infection and plant stress would lead to pathogen growth, resulting in plant death. We found that a combination of water stress and a fungal bio-fertilizer containing a beneficial root colonizing fungus as well as ring nematode infestation and this fungal bio-fertilizer increased the growth of this pathogen. We observed young vine decline like symptom development in the field and observed higher mortality in grapevines subjected to the 50% water reduction treatment compared to plants subjected to the 100% water treatment and in plants inoculated with a medium quantity of pathogen spores with ring nematode infestation of the roots compared to nematode free plants. This work suggests that a combination of infection level and plant stress could increase the growth of the pathogen and result in symptoms development and plant death. vi Preface A version of chapter 1 has been published. Hrycan, J., Hart, M., Bowen, P., Forge, T., and Úrbez-Torres, J. R. 2020. Grapevine trunk disease fungi: their roles as latent pathogens and stress factors that favour disease development and symptom expression. Phytopathol. Mediterr. 59:395–424. A version of chapter 2 has been published. Hrycan, J., Theilmann, J., Mahovlic, A., Boulé, J., and Úrbez-Torres, J. R. 2023. Health status of ready-to-plant grapevine nursery material in Canada regarding young vine decline fungi. Plant Dis. 107:3708-3717. I wrote the review paper featured in chapter 1 with guidance and edits from Dr. José Ramón Úrbez-Torres, Dr. Miranda Hart, Dr. Patricia Bowen, and Dr. Thomas Forge. The nursery survey described in chapter 2 was carried out by myself with help from Jane Theilmann, April Mahovlic, and Julie Boulé. I wrote the manuscript with guidance from Dr. José Ramón Úrbez-Torres. The work was conducted at Agriculture and Agri-Food Canada’s Summerland Research and Development Centre (AAFC-SuRDC). Chapter 3, 4, and 5 were designed in consultation with Dr. José Ramón Úrbez-Torres, Dr. Miranda Hart, Dr. Patricia Bowen, and Dr. Thomas Forge. Greenhouse, field, and laboratory work were primarily carried out at AAFC-SuRDC with some work conducted in Dr. Miranda Hart’s laboratory at the University of British Columbia Okanagan. The Plant Pathology Laboratory at AAFC-SuRDC helped at various times with these experiments. Dr. Thomas Forge and Paige Munro helped with chapter 4 and the Nematology Laboratory at AAFC-SuRDC was used for all nematode work. Dr. Patricia Bowen and Carl Bogdanoff aided in rooting and maintenance of plants and with conducting plant health measurement for all experiments through guidance or the use of their laboratories. Writing of chapters 3, 4, and 5 were carried out by myself with guidance from Dr. José Ramón Úrbez-Torres, Dr. Miranda Hart, Dr. Patricia Bowen, and Dr. Thomas Forge. All tables and figures are listed at the end of each chapter. Supplementary tables and figures are listed at the end of this thesis. Generative AI was not used in any aspect of this work. vii Table of Contents Abstract ......................................................................................................................................... iii Lay Summary .................................................................................................................................v Preface ........................................................................................................................................... vi Table of Contents ........................................................................................................................ vii List of Tables ............................................................................................................................... xii List of Figures ........................................................................................................................... xviii Supplementary Files ................................................................................................................. xxii Acknowledgements .................................................................................................................. xxiii Dedication ................................................................................................................................. xxiv 1 Introduction .................................................................................................................................1 1.1 Grapevine trunk disease fungi: their roles as latent pathogens and stress factors that favour disease development and symptom expression ...............................................................1 1.1.1 Endophytes and latent pathogens: from endophytic to pathogenic phase ...........................2 1.1.2 Endophytes in grapevines ....................................................................................................5 1.2 Grapevine trunk disease fungi as latent pathogens ..............................................................6 1.2.1 Botryosphaeriaceae .............................................................................................................7 1.2.2 Ilyonectria ...........................................................................................................................9 1.2.3 Phaeomoniella chlamydospora and Phaeoacremonium species ......................................10 1.2.4 Diatrypaceae .....................................................................................................................11 1.2.5 Diaporthe ..........................................................................................................................12 1.3 Young vine decline and grapevines nursery stock ..............................................................13 1.3.1 Infection during the grapevine propagation ......................................................................15 1.3.2 Rootstock and scion resistance to GTD fungi ...................................................................16 1.4 Abiotic stress factors and GTD symptom expression .........................................................19 1.4.1 Botryosphaeriaceae ...........................................................................................................20 1.4.2 Ilyonectria .........................................................................................................................22 1.4.3 Phaeomoniella chlamydospora and Phaeoacremonium species ......................................23 1.4.4 Diatrypaceae .....................................................................................................................24 1.4.5 Diaporthe ..........................................................................................................................25 1.5 Roles of biotic stress factors on GTD symptom expression ...............................................26 1.5.1 Parasitic nematodes affecting Ilyonectria symptom expression .......................................26 viii 1.5.2 Fungal pathogen interactions and infection thresholds .....................................................28 1.6 Climate change and viticulture practices: impacts on abiotic and biotic stress factors and GTD symptom expression ....................................................................................................30 1.6.1 Water deficit irrigation ......................................................................................................31 1.6.2 Vine pruning and training practices ..................................................................................32 1.6.3 Endophytes as potential reduces of host stress ..................................................................33 1.6.4 Arbuscular mycorrhizal fungi as potential stress reducers ................................................34 1.7 Conclusions and future directions ........................................................................................37 1.7.1 GTD as latent pathogens ...................................................................................................37 1.7.2 Latent pathogens in young vineyards ................................................................................38 1.7.3 Methods for host stress reductions ....................................................................................38 1.8 Research objectives ................................................................................................................47 1.8.1 Are young vine decline fungi present in asymptomatic grapevine nursery material planted in Canada and what is their abundance within individual grapevines ..........................47 1.8.2 Does Phaeomoniella chlamydospora abundance within grapevines impact disease development in young grapevines .............................................................................................48 1.8.3 Does abiotic and biotic plant stress influence Phaeomoniella chlamydospora abundance and Petri disease symptom development in young grapevines .................................................48 1.8.4 Do arbuscular mycorrhizal fungi reduce plant stress and subsequent pathogen abundance and disease development in young grapevines .........................................................................49 2 Health status of ready-to-plant grapevine nursery material in Canada regarding young vine decline fungi .........................................................................................................................51 2.1 Background ............................................................................................................................51 2.2 Materials and methods ..........................................................................................................54 2.2.1 Plant material and sample preparation .............................................................................54 2.2.2 Morphological and molecular characterization of isolated fungi .....................................55 2.2.3 Symptom analyses ............................................................................................................56 2.2.4 DNA extraction and absolute fungal quantification by Droplet Digital™ PCR ..............56 2.2.5 ddPCR reactions and conditions ......................................................................................58 2.2.6 Statistical Analysis ...........................................................................................................59 2.3 Results .....................................................................................................................................59 2.3.1 Morphological and molecular characterization of isolated fungi .....................................59 2.3.2 Symptom analyses ............................................................................................................60 ix 2.3.3 Absolute fungal quantification by ddPCR .......................................................................61 2.3.4 Fungal abundance in different rootstocks ........................................................................64 2.4 Discussion................................................................................................................................64 3 Impact of water stress on Phaeomoniella chlamydospora abundance and Petri disease symptom development in young grapevines .............................................................................83 3.1 Background ............................................................................................................................83 3.2 Materials and methods ..........................................................................................................87 3.2.1 Greenhouse experiment ....................................................................................................87 3.2.1.1 Plant material, inoculum preparation and vacuum inoculation ..................................87 3.2.1.2 Experimental design and plant health measurements ................................................88 3.2.1.3 Phaeomoniella chlamydospora absolute quantification ............................................90 3.2.1.4 Arbuscular mycorrhizal fungal colonization .............................................................90 3.2.2 Field experiment ...............................................................................................................91 3.2.2.1 Plant material, inoculum preparation and vacuum inoculation ..................................91 3.2.2.2 Experimental design and plant health measurements ................................................92 3.2.2.3 Phaeomoniella chlamydospora absolute quantification ............................................93 3.2.3 Statistical analysis .............................................................................................................94 3.3 Results .....................................................................................................................................94 3.3.1 Greenhouse ........................................................................................................................94 3.3.1.1 Plant health measurements ........................................................................................94 3.3.1.2 Quantification of Phaeomoniella chlamydospora .....................................................95 3.3.1.3 Trunk measurements .................................................................................................96 3.2.2 Field ...................................................................................................................................96 3.3.1.1 Plant health measurements ........................................................................................96 3.3.1.2 Quantification of Phaeomoniella chlamydospora .....................................................97 3.3.1.3 Trunk measurements .................................................................................................98 3.2.2.3 Plant symptom development and mortality ...............................................................98 3.4 Discussion................................................................................................................................99 4 Investigating the interaction between ring nematode infestation and Petri disease development in young ‘Merlot’ grapevines infected with Phaeomoniella chlamydospora .120 4.1 Background ..........................................................................................................................120 4.2 Materials and methods ........................................................................................................123 x 4.2.1 Greenhouse experiment ..................................................................................................124 4.2.1.1 Plant material, inoculum preparation and vacuum inoculation ................................124 4.2.1.2 Experimental design and plant health measurements ..............................................126 4.2.1.3 Phaeomoniella chlamydospora absolute quantification ..........................................127 4.2.1.4 Arbuscular mycorrhizal fungal colonization ...........................................................128 4.2.2 Field experiment .............................................................................................................128 4.2.2.1 Plant material, inoculum preparation and vacuum inoculation ................................128 4.2.2.2 Experimental design and plant health measurements ..............................................129 4.2.2.3 Phaeomoniella chlamydospora absolute quantification ..........................................130 4.2.3 Statistical analysis ...........................................................................................................131 4.3 Results ...................................................................................................................................132 4.3.1 Greenhouse ......................................................................................................................132 4.3.1.1 Plant health measurements ......................................................................................132 4.3.1.2 Quantification of Phaeomoniella chlamydospora ....................................................133 4.3.1.3 Trunk measurements ...............................................................................................134 4.2.2 Field .................................................................................................................................134 4.3.1.1 Plant health measurements ......................................................................................134 4.3.1.2 Quantification of Phaeomoniella chlamydospora ...................................................135 4.3.1.3 Trunk measurements ...............................................................................................136 4.3.1.3 Plant mortality .........................................................................................................136 4.4 Discussion..............................................................................................................................137 5 Effects of j-rooting and heat stress on Phaeomoniella chlamydospora abundance and Petri disease development in young ‘Merlot’ grapevines ...............................................................158 5.1 Background ..........................................................................................................................158 5.2 Materials and methods ........................................................................................................161 5.2.1 Plant material, inoculum preparation and vacuum inoculation ......................................161 5.2.2 J-Root .............................................................................................................................163 5.2.2.1 Experimental design and measurements .................................................................163 5.2.3 Heat Stress ......................................................................................................................164 5.2.3.1 Experimental design and measurements .................................................................164 5.2.4 Quantification of Phaeomoniella chlamydospora ..........................................................165 5.2.5 Statistical analysis ..........................................................................................................167 xi 5.3 Results ...................................................................................................................................167 5.3.1 J-root experiment .............................................................................................................167 5.3.1.1 Plant health measurements ......................................................................................168 5.3.1.2 Quantification of Phaeomoniella chlamydospora ...................................................168 5.3.1.3 Trunk measurements ...............................................................................................168 5.2.2.4 Plant symptom development and mortality .............................................................168 3.2.2 Heat stress experiment ....................................................................................................169 3.3.1.1 Plant health measurements ......................................................................................169 3.3.1.2 Quantification of Phaeomoniella chlamydospora ...................................................170 3.3.1.3 Trunk measurements ...............................................................................................171 3.2.2.3 Plant symptom development and mortality .............................................................171 5.4 Discussion..............................................................................................................................172 6 Conclusions ..............................................................................................................................189 6.1 Are young vine decline fungi present in asymptomatic grapevine nursery material planted in Canada and what is their abundance within individual grapevines? .................189 6.2 Does Phaeomoniella chlamydospora abundance within grapevines impact disease development in young grapevines?...........................................................................................190 6.3 Do abiotic and biotic plant stress influence Phaeomoniella chlamydospora abundance and Petri disease symptom development in young grapevines? ............................................191 6.4 Do arbuscular mycorrhizal fungi reduce plant stress and subsequent pathogen abundance and disease development in young grapevines? ..................................................192 6.5 Concluding remarks ...........................................................................................................193 Bibliography ...............................................................................................................................194 Appendices ..................................................................................................................................223 Appendix A: Supplementary file for Chapter 2 ......................................................................223 Appendix B: Supplementary material for Chapter 3 .............................................................225 Appendix C: Supplementary material for Chapter 4 .............................................................231 Appendix D: Supplementary material for Chapter 5……………………………………….235 xii List of Tables Table 2.1. Mean sum of infection (copies/ng DNA) of all fungal species in the basal end of each grapevine and percent necrosis at the base of each grapevine for each nursery individually ......73 Table 2.2. Maximum, median, and minimum level of infection (copies/ng DNA) of each individual fungal species tested and all fungal species combined in infected grapevines and percent infection of all grapevines in each nursery individually and all nurseries combineda ......74 Table 2.3. Mean copies/ng DNA and standard deviation of Botryosphaeriaceae spp. in each section of each cultivar and in total in each cultivar for all nurseries ...........................................75 Table 2.4. Mean copies/ng DNA and standard deviation of Cadophora luteo-olivacea in each section of each cultivar and in total in each cultivar for all nurseries ...........................................76 Table 2.5. Mean copies/ng DNA and standard deviation of Dactylonectria macrodidyma in each section of each cultivar and in total in each cultivar for all nurseries ...........................................77 Table 2.6. Mean copies/ng DNA and standard deviation of Dactylonectria torresensis in each section of each cultivar and in total in each cultivar for all nurseries ...........................................78 Table 2.7. Mean copies/ng DNA and standard deviation of Phaeoacremonium minimum in each section of each cultivar and in total in each cultivar for all nurseries ...........................................79 Table 2.8. Mean copies/ng DNA and standard deviation of each Phaeomoniella chlamydospora in each section of each cultivar and in total in each cultivar for all nurseries ..............................80 Table 2.9. Mean sum copies/ng DNA and standard deviation of each fungal species in the basal end and roots of each rootstock for Nursery A2014y ....................................................................81 xiii Table 3.1. Analysis of inoculum, irrigation treatment, AM, time and their interactions on leaf chlorophyll, stem water potential, and soil matric potential in year 2 using an analysis of variance of a linear mixed effects model for greenhouse. ..........................................................................108 Table 3.2. Analysis of inoculum, irrigation treatment, AM fungi, year and their interactions on dry pruning weight, node number, and dry root weight using an analysis of variance for greenhouse. ..................................................................................................................................109 Table 3.3. Analysis of inoculum, irrigation treatment, AM, and their interactions on Pc abundance at harvest, class 1, 2, and total necrosis, and basal surface area, at harvest using an analysis of variance for greenhouse. ............................................................................................110 Table 3.4. Percent class 1, class 2, total necrosis, and mean basal surface area (mm) and standard error recorded at the base of each grapevines at the end of the greenhouse experiment for each level of inoculum and in each level of irrigation treatment in the greenhouse. ...........................111 Table 3.5. Analysis of cultivar, inoculum, stress, and time for each year and their interactions on leaf chlorophyll and stem water potential, using an analysis of variance of a linear mixed effects model for the field ........................................................................................................................112 Table 3.6. Analysis of cultivar, inoculum, irrigation treatment, year and their interactions on dry pruning weight, node number, and dry root weight using an analysis of variance of a linear mixed effects model for the field. ................................................................................................113 Table 3.7. Analysis of cultivar, inoculum, irrigation treatment, and their interactions on Pc abundance at harvest, class 1, 2, and total necrosis, and basal surface area, at the end of the experiment using an analysis of variance of a linear mixed effects model for the field. .............114 xiv Table 3.8. Chi-squared analysis of plant total (reps), plant mortality and symptom expression for each year and cumulatively for each cultivar, level of inoculum x irrigation treatment and level of inoculum overall. .....................................................................................................................115 Table 4.1. Analysis of variance summary of inoculum and AM fungi on ring nematode populations in the greenhouse using an analysis of variance and analysis of inoculum, year, on ring nematode populations, inoculum and stress on root-knot nematode populations in the field using an analysis of variance on a linear mixed effects model. ...................................................146 Table 4.2. Analysis of inoculum, ring nematode infestation (Nem), AM fungi, time, on leaf chlorophyll content in year 2 of the greenhouse experiment using an analysis of variance of a linear mixed effects model. ..........................................................................................................147 Table 4.3. Analysis of inoculum, ring nematode infestation (Nem), AM fungi, year, and their interactions on dry pruning weight, node number, and root weight in the greenhouse using an analysis of variance. .....................................................................................................................148 Table 4.4. Analysis of inoculum, ring nematode infestation (Nem), AM fungi, and their interactions on fungal abundance, class 1, 2, and total necrosis, and basal surface area at harvest in the greenhouse using an analysis of variance. .........................................................................149 Table 4.5. Percent class 1, class 2, total necrosis, mean basal surface area (mm) and standard error at the base of each grapevine at the end of the experiment containing each level of inoculum in the greenhouse and field. .........................................................................................150 Table 4.6. Analysis of inoculum, ring nematode infestation (Nem), and time for each year and their interactions on leaf chlorophyll and stem water potential each year in the field using an analysis of variance of a linear mixed effects model. ..................................................................151 xv Table 4.7. Analysis of inoculum, ring nematode infestation (Nem), and year on dry pruning weight, node number, and root weight in the field using an analysis of variance on a linear mixed effects model. ...............................................................................................................................152 Table 4.8. Analysis of inoculum, stress, and their interactions on pathogen abundance, class 1, 2, and total necrosis, basal surface area, and trunk diameter at harvest in the field using an analysis of variance on a linear mixed effects model ................................................................................153 Table 4.9. Chi-squared analysis of plant total (reps) and mortality for each year and cumulatively for each level of inoculum x nematode treatment and each level of inoculum overall ...............154 Table 5.1. Analysis of inoculum, stress treatment, time, and their interactions on leaf chlorophyll content each year using an analysis of variance on a linear mixed effect model for the j-root experiment....................................................................................................................................179 Table 5.2. Mean leaf chlorophyll content and standard error for each level of inoculum and j-rooting or enclosure treatment each year for the j-root and heat stress experiments. ..................180 Table 5.3. Analysis of inoculum, treatment, year, and their interactive effects on pruning weight, node number, and root weight for the j-root and heat stress experiments using an analysis of variance of a linear mixed effect model. ......................................................................................181 Table 5.4. Analysis of inoculum, stress treatment, and their interactions on pathogen fungal abundance at harvest, class 1, 2, and total percent necrosis, basal surface area, and trunk diameter using an analysis of variance of a linear mixed effect model for the j-root and heat stress experiments. .................................................................................................................................182 xvi Table 5.5. Mean percent class 1, 2, total necrosis, mean basal surface area (mm) and standard errors recorded at the end of the experiment for each level of inoculum and stress treatment in the j-root and heat stress experiments. .........................................................................................183 Table 5.6. Chi-squared analysis of plant mortality and symptom expression each year and cumulatively for each level of inoculum x stress treatment and inoculum level overall for the j-root and heat stress experiments ..................................................................................................184 Table 5.7. Analysis of inoculum, treatment, time, and their interactions on leaf chlorophyll content and stem water potential for each year of the heat stress experiment using an analysis of variance of a linear mixed effect model. ......................................................................................185 Table S2.2. Primers and probe sequences of targeted fungal pathogens used in each ddPCR assay for absolute quantification in ready-to-plant grapevine nursery material .........................224 Table S3.1. Mean SPAD leaf chlorophyll and standard error over time in 2021 for each level of inoculum and each level of stress in the greenhouse ...................................................................225 Table S3.2. Mean dry pruning weight, dry root weight, and node number for each year and standard error for year 1 and year 2 for each level stress and AM fungi in the greenhouse .......226 Table S3.3. Mean SPAD leaf chlorophyll and standard error each year for each cultivar, level of inoculum, and level of stress in the field .....................................................................................227 Table S3.4. Mean dry pruning weight and node number, for 2020 and 2021 and standard error for each cultivar, level of infection, and stress in the field ..........................................................228 Table S3.5. Mean dry pruning weight, node number, trunk diameter, and dry root weight for 2022 and 2023 and standard error for each cultivar, level of infection, and stress in the field. .229 xvii Table S3.6. Percent class 1, class 2, total necrosis, and mean basal surface area and standard error of each cultivar containing each level of inoculum and in each level of stress in the greenhouse ...................................................................................................................................230 Table S4.1. Mean leaf chlorophyll, and standard error each year in the greenhouse for each level of inoculum and ring nematode infestation treatment .................................................................231 Table S4.2. Mean pruning weight, node number, and root weight, and standard error each year in the greenhouse for each level of inoculum, AM treatment, and ring nematode infestation treatment ......................................................................................................................................232 Table S4.3. Mean SPAD leaf chlorophyll, stem water potential (MPa), and standard error each year in the field for each level of inoculum and ring nematode infestation treatment ................233 Table S4.4. Mean pruning weight, node number, trunk diameter, and root weight and standard error each year in the field for each level of inoculum and ring nematode infestation treatment. ......................................................................................................................................................234 Table S5.1. Mean pruning weight, node number, trunk diameter, and root weight and standard error each year in the field for each level of inoculum and j-rooting treatment ..........................235 Table S5.2. Mean pruning weight, node number, trunk diameter, and root weight and standard error each year in the field for each level of inoculum and enclosure treatment .........................236 xviii List of Figures Figure 1.1. Illustration of the Threshold Model in grapevines, showing the relationship between the grapevine age and GTD fungal colonization density. Under normal conditions, fungal growth is slow, only reaching the colonization threshold in old grapevines during natural senescence. Under stress conditions, fungal growth is accelerated, and this threshold is reached rapidly in the grapevine lifespan, leading to disease progression and premature host death. This threshold is hypothesized to be reached sooner when multiple GTD fungi are present than when few fungi occur (adapted with permission from Sieber 2007) .......................................................................40 Figure 1.2. Botryosphaeria dieback, Eutypa, Phomopsis dieback, and Esca symptoms. Botryosphaeria dieback symptoms on wine (A) and table grapevines (B) are characterized by dead spurs and cordons with no spring growth. Characteristic foliar symptoms of Eutypa dieback are shoots with short internodes and chlorotic and cupped leaves (C and D). Phomopsis dieback symptoms resemble those of Botryosphaeria dieback, with dead spurs and lack of spring growth (E). Vines affected by Botryosphaeria, Eutypa and/or Phomopsis dieback have perennial cankers in spurs, cordons and trunks, often with wedge shapes (F). Tiger-striped leaves (G) and characteristic soft yellowish wood rot (H) caused by Basidiomycete fungi are commonly associated with Esca disease ..........................................................................................................41 Figure 1.3. Decline of young vines caused by Petri disease and/or black foot. Overall symptoms in the field include poor vigour, leaf chlorosis, short internodes (A and B) and eventual or sudden collapse I during the growing season. Internal wood necrosis observed at the basal end of the rootstock (D) commonly associated with black foot. Tylosis plugged xylem vessels and necrosis observed at a graft uniI(E) and trunk base, usually associated with Petri disease in young vines (F). Longitudinal section of a ready-to-plant nursery vine showing vascular necrosis originating xix from the basal end of the rootstock (G and H). Phaeomoniella chlamydospora, Phaeoacremonium spp., Ilyonectria spp., and some Botryosphaeriaceae spp. can be isolated from asymptomatic (I) and symptomatic (J) wood tissues from nursery propagated material, which reveals their potential latent phases in grapevines ..............................................................43 Figure 1.4. Abiotic and biotic stress factors hypothezised to contribute to grapevine trunk disease fungal development and symptom expression. A. Leaf scorching as a result of severe vine water stress. B and C. Leaves of young vines showing nutrient deficiency. D. Leaf deformities caused by herbicide damage. E. Severe over-cropping during the first years of vineyard establishment (courtesy of D. Gramaje, ICVV, Logroño, Spain). F. Winter damage and/or spring frost could further abiotic factors favouring disease development. G. J-rooting as a result of poor planting conditions. H. Low vigour vines caused by a nematode infestation. I. Ring nematode (Mesocriconema xenoplax) feeding on a root by introducing the stylet (black arrow). J. Close up of the ring nematode head. K. Leaf galls caused by erineum mite (Colomerus vitis). High levels of insect damage could increase vine stress. L. Grapevine crown gall caused by Agrobacterium vitis. M. Grapevine showing of grapevine leaf roll symptoms caused by Grapevine leafroll-associated virus 3. Grapevines affected by biotic stress factors (fungal, bacterial or virus diseases) may favour GTD fungal infections, wood colonization and symptom expression ......................................................................................................................................45 Figure 2.1. Scan showing nursery A2019 basal necrosis from the plant with the highest total fungal abundance at the basal end (724.05 copies/ng) with 50% necrosis (A) and the plant with the lowest total fungal abundance at the basal end (2.90 copies/ng) with 43% necrosis (B) ........82 xx Figure 3.1. Scan showing basal necrosis of an inoculated plant (1,2,3) and non-inoculated plant (4,5,6) showing necrosis (1,2), threshold analysis of class 1 necrosis set at 5-45 (2,4), and threshold analysis of class 2 necrosis set at 46-80 (3,6) using ImageJ software .........................116 Figure 3.2. Stem water potential recorded at mid-day in 1. Year 2 in the greenhouse, 2. 2022 in the field, 3. Soil matric potential recorded every 15 minutes in the greenhouse for 100% watered (I1) and 50% watered (I2) plants. Significantly different results indicated by * at each time point.......................................................................................................................................................117 Figure 3.3. Pc abundance (copies/ng DNA) at the base of plants at the end of the greenhouse experiment in infected plants under Irrigation (I1, I2) x AM fungal inoculation (AM-, AM+) treatments. Treatments with the same letter are not significantly different according to Tukey’s significance test. ...........................................................................................................................118 Figure 3.4. Photographs of young vine decline-like symptom progression in 2022 on 1. August 9th, 2. August 30th, and 3. September 27th for a grapevine infected with Pc subject to irrigation treatment. .....................................................................................................................................119 Figure 4.1. Pc abundance (copies/ng DNA) at the end of the greenhouse experiment in Pc inoculated plants with, nematode infestation (Nem+, Nem-) x AM fungal inoculation (AM+, AM-) treatments. Treatments with the same letter are not significantly different according to Tukey’s significance test..............................................................................................................155 Figure 4.2. Mean root-knot nematode populations per 100mL of soil for plants in each level of inoculum treatment. Treatments with the same letter are not significantly different from each other according to Tukey’s significance test. ..............................................................................156 xxi Figure 4.3. Pc abundance (copies/ng DNA) at the end of the field experiment for each inoculum level in ‘Merlot’ grapevines. Treatments with the same letter are not significantly different from each other according to Tukey’s significance test. ......................................................................157 Figure 5.1. Photographs of 1. Planting of j-rooted ‘Merlot’ grapevine on July 10, 2020, 2. A j-rooted grapevine harvested in Spring 2023, 3. Set-up of enclosure treatment on July 27, 2022.186 Figure 5.2. Photographs of young vine decline symptom progression of a 2.5k-JR+ plant on 1. June 2. 2. August 9. 3. August 30 and in a 25k-LT plant on 4. August 9. 5. August 30. 6. September 27, 2022 .....................................................................................................................187 Figure 5.3. Mean hourly temperature recorded by iButton for from July 27 to August 1 during enclosure treatment in five PFE+ and five PFE- grapevines. Dashed line represents PFE+ and solid line represents PFE-. * indicates significant difference between temperatures according to Tukey’s significance test set at 0.05. ...........................................................................................188 Figure S5.1. Mean hourly temperature recorded by iButton for the duration of the heat stress treatment five PFE- grapevines and adjacent weather station. Dashed line represents the weather station and solid line represents PFE-. * indicates significant difference between temperatures according to Tukey’s significance test .........................................................................................237 xxii Supplementary Files Table S2.1. Total number of plants tested and infected with young vine decline fungi based on morphological and molecular identification of fungal colonies obtained from re-isolations File: Table S2.1.xlsx.............................................................................................................................221 xxiii Acknowledgements I would first like to sincerely thank my co-supervisors, Dr. José Ramón Úrbez-Torres and Dr. Miranda Hart. Your guidance and support throughout these past few years has meant a great deal. I am extremely grateful to have had the opportunity to learn from you. I also want to thank my committee members, Dr. Pat Bowen, Dr. Tom Forge, and Dr. Louise Nelson for all your support. Your doors were always open when I had questions, and I had many. I sincerely appreciate all the guidance and support you have provided me since I started. Thank you to the Plant Pathology laboratory at the Summerland Research and Development Centre, Jane Theilmann, Melanie Walker, Julie Boulé, Isidora Silva-Valderrama, April Mahovlic, and Dan O’Gorman, as well as the co-op students and casuals including Mia Alexander, Cassandra Bruce, and Brett Pfilger for your help. I would also like to thank the lab technicians and researchers who aided me in my research, in particular, Paige Munro, Steve Marsh, Brad Estergaard, and Carl Bogdanoff for providing me with varying levels of guidance and technical support. Lastly, I would like to sincerely thank the nurseries who provided me with the grapevine material used in chapter 2 of this dissertation as well as Agriculture and Agri-Food Canada’s Summerland Research Centre, the British Columbia Wine Grape Council, and the Canadian Grapevine Certification Network for funding this research. xxiv Dedication To my parents, Anne-Marie and David Hrycan, Thank you for always supporting me. 1 1 Introduction 1.1 Grapevine trunk disease fungi: their roles as latent pathogens and stress factors that favour disease development and symptom expression Grapevine trunk diseases (GTDs) are major threats to the economic sustainability of viticulture; they cause significant economic losses due to reduced yields, increased crop management costs and shortened life spans for vineyards (Bertsch et al. 2013; Gramaje et al. 2018; Kaplan et al. 2016; Siebert 2001; Wicks and Davies 1999). The GTD complex includes Black foot disease, Botryosphaeria dieback, Esca, Eutypa dieback, Petri disease, and Phomopsis dieback (Agustí-Brisach and Armengol 2013; Bertsch et al. 2013; Gramaje and Armengol 2011; Gramaje et al. 2018; Mondello et al. 2018; Úrbez-Torres 2011). These diseases are primarily caused by taxonomically unrelated Ascomycete fungi, and to lesser extent by several Basidiomycetes which are primarily associated with the Esca complex (Gramaje et al. 2018). Black foot disease and Petri disease typically occur in grapevines aged 5 years or younger, while Botryosphaeria dieback, Esca, Eutypa dieback, and Phomopsis dieback are usually seen in older vineyards, although these diseases may also occur in young grapevines (Agustí-Brisach and Armengol 2013; Bertsch et al. 2013; Gramaje and Armengol 2011; Úrbez-Torres et al. 2008). Pruning wounds serve as major routes of infection by these pathogens in established vineyards. The discharge of spores from fruiting bodies and airborne transmission of these spores from plant to plant have primarily been associated with rainfall, with the exception of black foot pathogens, which are known to be soil-borne (Agustí-Brisach and Armengol 2013; Agustí‐Brisach et al. 2014; Eskalen and Gubler 2001; Rooney-Latham et al. 2005; Trese 1980; Úrbez-Torres et al. 2010a; Úrbez-Torres et al. 2010b). 2 Grapevine trunk diseases are caused by xylem-colonizing fungi and are categorized as slow-progression diseases with symptoms sometimes taking several years to appear after infection. In addition, GTD symptoms are known to be expressed inconsistently from year to year on individual grapevines (Bertsch et al. 2013; Calzarano et al. 2018; Songy et al. 2019). General symptoms of GTDs include delayed bud-break, leaf chlorosis, reduced vigour, stunted growth, wood necroses, canker formation, dieback, and eventual death of affected grapevine plants (Agustí-Brisach and Armengol 2013; Bertsch et al. 2013; Billones-Baaijens and Savocchia 2019; Gramaje and Armengol 2011; Gramaje et al. 2018; Mondello et al. 2018; Reis et al. 2019; Songy et al. 2019; Úrbez-Torres 2011). Due to the inconsistent symptom expression from year to year for some GTDs, and the reported isolation of GTD fungi from asymptomatic tissues, it has been proposed that some of these fungi may act as latent pathogens (Gramaje et al. 2018; Úrbez-Torres 2011). The presence of GTD fungi in asymptomatic tissues can be very problematic for growers and propagators, as infections can spread unnoticed in the field or in nurseries. Asymptomatic infections in rootstock and scion mother plants can provide possible sources of infection in nurseries (Fourie and Halleen 2004; Mondello et al. 2018). In addition, abiotic and biotic stresses are likely to play important roles in GTD symptom development (Gramaje et al. 2018; Reis et al. 2019; Songy et al. 2019; Úrbez-Torres 2011). 1.1.1 Endophytes and latent pathogens: from endophytic to pathogenic phases Endophytes are organisms found in the internal tissues of plants, including in roots, xylem, phloem, and/or leaves, and these organisms may act as saprobes, mutualists, or latent pathogens (Hyde and Soytong 2008; Rodriguez et al. 2009; Wilson 1995). The definition of endophyte has changed several times; however, for this review, the definition of Petrini (1991) is followed, as “all organisms inhabiting plant organs that at some time in their life, can colonize internal plant 3 tissues without causing apparent harm to their host.” This definition includes latent pathogens during their asymptomatic phases (Sieber 2007; Stone et al. 2000; Wilson 2000; Wilson 1995). The term “endophyte” is used here for the endophytic phase of an organism’s life cycle, the life cycle of which may include several other phases that may be transitory and/or pathogenic (Stone et al. 2000; Wilson 2000; Wilson 1995). Most fungal endophytes are in Ascomycota R.H. Whittaker, although some are in Basidiomycota Whittaker ex R.T. Moore, Deuteromycota R.T. Moore, or Oomycota Arx (Hyde and Soytong 2008; Petrini 1986). Fungal endophytes can also be classified into two broad categories, as clavicipitalean endophytes, which live in the internal tissues of grasses and have been thoroughly researched, and non-clavicipatalean endophytes, which live in a wide range of hosts including woody perennials and non-vascular plants (Hyde and Soytong 2008; Rodriguez et al. 2009; Sieber 2007). Rodriguez et al. (2009) suggested an endophyte classification system with four classes. This included Class 1 as clavicipitaceous endophytes which are grass endophytes. Classes 2 to 4 are non-clavicipitaceous endophytes. Class 2 endophytes are distinguished from the rest by their colonization of roots, stems, and leaves, and their transmission via seed coats and rhizomes. Class 3 endophytes have diverse plant host ranges, including woody perennials, and these endophytes live in the inner bark, wood, fruit, or flowers of their hosts, and can reproduce via sporulation on dead plant tissues. Class 3 endophytes include xylem-colonizing endophytes. Class 4 endophytes are only found in host roots, where they form melanised structures. The focus of the present review is on xylem-colonizing Class 3 endophytes, which make up a distinct guild of xylem-inhabiting endophytes (Stone et al. 2000). These include species in the order Hypocreales Lindau and in the genus Hypoxylon Bull, and to a lesser extent some Basidiomycetes. Survival strategies of many xylem-inhabiting endophytes consist of initial 4 infection of host plants, followed by indeterminate periods of interrupted growth allowing for further invasion and exploitation of the substrates upon occurrence of favourable conditions, such as host stress (Stone et al. 2000). These xylem-colonizing endophytes typically disperse through horizontal transmission, with drop-splash from rainfall for spore dispersal. Wind and/or other vectors such as animals or insects may also facilitate transmission of these fungi (Petrini 1991; Wilson 2000). Endophytes can either have wide host ranges, or as suggested by Petrini (1991), they may have coevolved with their host plants, allowing the endophytes to infect their hosts without activating host defense mechanisms. There are three hypotheses for the causes of the transition from endophytic to pathogenic phases. The first is that some event occurs which increases host susceptibility, facilitating the transition. These events are most likely to be abiotic and/or biotic stressors, including drought, poor host nutrient supply, and/or host wounding (Petrini 1991; Stone, et al. 2000). The second hypothesis includes endophyte changes, including single point mutations, the transfer of virulence genes, and/or virus infections causing change to a pathogenic state (Sieber 2007). The third hypothesis is the threshold model developed by Sieber (2007) for conifer needle endophytes, and this could be relevant to other types of endophytes. This model suggests that endophytes live in needle tissues, which senesce once a colonization threshold has been reached. This threshold is only reached under normal circumstances during the natural senescence of the needles; however, under stress conditions, the colonization threshold is reached sooner, leading to premature death (Sieber 2007). All these scenarios could be applied to GTD fungi thought to have endophytic phases before becoming pathogenic. Most GTD fungi are slow wood colonizers causing slow-developing diseases (Agusti-Brisach et al. 2019; Bradshaw et al. 2005; Troccoli et al. 2001; Mugnai et al. 1999). In healthy grapevines, fungal growth is characteristically slow. When vines 5 become stressed, however, fungal growth is accelerated and colonization thresholds are reached sooner than under non-stress conditions. Additionally, this threshold could be reached sooner when multiple GTD fungi are present within a grapevine (Figure 1.1). 1.1.2 Endophytes in grapevines Characterization of bacterial and fungal endophytes of grapevines has been a major focus of research, to identify potential biocontrol agents against pathogens (Bruisson et al. 2019). Endophytes colonize above- and below-ground plant organs, including, in grapevines, flowers, berries, xylem, roots, and seeds (Bruisson et al. 2019; Compant et al. 2011). Bacterial endophyte richness is greater than fungal endophyte richness in grapevines, and Proteobacteria Garrity et al. and Ascomycota species, respectively are the most abundant bacteria and fungi in grapevines (Deyett and Rolshausen 2020). In Vitis vinifera L., Ascomycota endophytes are primarily Class 3 xylem-colonizing endophytes in the Hypocreales, Pleosporales Luttrell ex M.E. Barr, and Xylariales Nannf., which is consistent with other woody perennial hosts (González and Tello 2011; Hyde and Soytong 2008; Petrini 1986). Among these Ascomycetes, species in the Hypocreales and Pleosporales were the most abundant endophytes found across multiple grapevine cultivars in a study in Spain (González and Tello 2011). However, the cultivars were colonized by different endophyte populations, showing that grapevines cannot be viewed as a homogeneous group of hosts, but cultivars must be studied individually. For example, González and Tello (2011) found ‘Tempranillo’ and ‘Merlot’ cultivars harboured high levels of Eurotiales G.W. Martin ex Benny & Kimbr., while ‘Cabernet Sauvignon’ harboured the least variable endophyte diversity. This study also showed high prevalence of the GTD pathogen Diaporthe ampelina (Berkeley & M.A. Curtis) R.R. Gomes, C. Glienke & Crous, 6 (syn. Phomopsis viticola Sacc.) in asymptomatic and symptomatic grapevines (González and Tello 2011). Other GTD pathogens found less widely in asymptomatic vines include Phaeomoniella chlamydospora (W. Gams, Crous, M.J. Wingf. & Mugnai) Crous & W. Gams, Phaeoacremonium minimum (Tul. & C. Tul.) D. Gramaje, L. Mostert & Crous, and Phaeoacremonium inflatipes W. Gams, Crous & M.J. Wingf. (involved in Petri disease and Esca), Ilyonectria destructans (Zinssm.) Rossman, L. Lombard & Crous (associated with Black foot disease), and species in the Botryosphaeriaceae Theiss. & P. Syd. (causing Botryosphaeria dieback (Gonzalez and Tello, 2011). These results, along with other reports of GTD fungi isolated from asymptomatic grapevines (Aroca et al. 2010; Carlucci et al. 2017; Halleen and Petrini 2003; Kaliterna et al. 2009; Zanzotto et al. 2007), support the hypothesis that some of these fungi may have roles as latent pathogens in grapevines (González and Tello 2011; Gramaje and Armengol 2011; Gramaje et al. 2018; Úrbez-Torres 2011). The objectives of this review are to provide up-to-date information indicating that GTD fungi may function as latent grapevine pathogens, to develop a synthesis of current knowledge available on the different abiotic and biotic stress factors that may influence the transition from endophytic to pathogenic phases, and to indicate future research directions for advancing understanding and management of these complex diseases. 1.2 Grapevine trunk disease fungi as latent pathogens Most knowledge on the potential of GTD fungi to act as latent pathogens comes from studies in two main groups of Ascomycetes: Botryosphaeriaceae and Ilyonectria P. Chaverri & C. Salgado (“Cylindrocarpon” Wollenw.) spp., on grapevines as well as other woody perennials (Fourie and Halleen 2004; Slippers and Wingfield 2007; Úrbez-Torres 2011). This paper also 7 reviews the endophytic-pathogenic continuum of Phaeomoniella chlamydospora, Diatrypaceae and species of Phaeoacremonium and Diaporthe on grapevines. 1.2.1 Botryosphaeriaceae Botryosphaeriaceae includes morphologically diverse fungi which have been found in gymnosperms and angiosperms in every climatic region except the polar regions (Phillips et al. 2013; Slippers and Wingfield 2007). Not all fungi in the Botryosphaeriaceae produce disease symptoms, and some species may be true endophytes, which never produce disease symptoms (Slippers and Wingfield 2007). Some Botryosphaeriaceae have been shown to be important pathogens and have become problems in economically important agricultural crops, with several different species found to be associated with decline symptoms in apple trees (Brown-Rytlewski and McManus 2000; Úrbez-Torres et al. 2016), Eucalyptus (Burgess et al. 2005), dwarf cashew (Cardoso et al. 2006), hickory (Dai et al. 2017), macadamia (Jeff-Ego and Akinsanmi 2019), bay laurel (Lawrence et al. 2017), almond, dried plum, pistachio, walnut (Luo et al. 2019), peach, pear, blueberry (Sessa et al. 2018), black currant (Singer and Cox, 2010), olive (Úrbez-Torres et al. 2013b), and grapevine (Úrbez-Torres 2011). However, even in the most severe diseases caused by Botryosphaeriaceae in some hosts, virulence is probably favoured by abiotic and/or biotic stresses suffered by host plants (Slippers and Wingfield 2007). Fungi in the Botryosphaeriaceae have been found in several hosts which were asymptomatic and symptomatic (Brown-Rytlewski and McManus 2000; Panahandeh et al. 2019; Scala et al. 2019; Tennakoon et al. 2018). Botryosphaeriaceae isolated from asymptomatic hosts have been found to be pathogenic in pathogenicity trials conducted under controlled greenhouse conditions (Manzanos et al. 2017; Steinrucken et al. 2017). The Botryosphaeriaceae have gone through extensive taxonomic changes and revisions since first introduced in 1918 and have primarily been considered to be saprophytes 8 (Phillips et al. 2013). Only in the last two decades have detailed studies on the Botryosphaeriaceae, supported by the use of molecular techniques, revealed the widespread presence of these fungi in asymptomatic hosts, which has led to further investigations of the hypothesis that some Botryosphaeriaceae species may act as latent pathogens. In grapevines, up to 26 species of Botryosphaeriaceae have been identified as pathogens responsible for symptoms associated with the GTD Botryosphaeria dieback. These symptoms include leaf chlorosis, bud and wood necrosis, weak spring growth, and vascular cankers primarily in the shape of wedges (Billones-Baaijens and Savocchia 2019; Gramaje et al. 2018; Úrbez-Torres 2011) (Figure 1.2). Among all species known to be associated with Botryosphaeria dieback, Botryosphaeria dothidea (Moug. ex Fr.) Ces. & De Not. has been described as a latent pathogen of global importance in woody perennials (Marsberg et al. 2017). Several other Botryosphaeriaceae species have been described as latent pathogens in other woody hosts but have not been confirmed in grapevines (Billones-Baaijens and Savocchia 2019; Carlucci et al. 2015; Marsberg et al. 2017; Slippers and Wingfield 2007). Lecomte and Bailey (2011) found Botryosphaeriaceae species, primarily Diplodia seriata De Not., in wounded host tissues, which was attributed to presence of these fungi in the bark prior to wounding, suggesting that the fungi may act as latent pathogens in grapevines. This is important for grape production, as Botryosphaeria dieback is one of the most prevalent diseases of grapevines, and the causal pathogens may not be detected due to their latent nature (Agusti-Brisach et al. 2019; Billones-Baaijens and Savocchia 2019; Marsberg et al. 2017). Neofusiccocum parvum (Pennycook & Samuels) Crous, Slippers & A.J.L. Phillips remained latent in ‘Cabernet Sauvignon’ grapevines, with limited pathogen spread or lesion growth for 1.5 months after inoculation; thereafter, lesion lengths increased (Czemmel et al. 2015). However, no cause for the transition from endophyte to 9 pathogen was investigated, and more research should be conducted to determine whether this is the normal latency period of N. parvum, and/or if some external factors played roles in the transition, such as abiotic or biotic stress. Some GTD fungi can be detected in asymptomatic host tissues far beyond necrotic or cankered wood. For example, B. dothidea colonizes asymptomatic tissues beyond canker margins in many different hosts, which supports the hypothesis of its endophytic nature (Schoeneweiss 1981; Wene 1979). Similarly, Botryosphaeriaceae known to be pathogenic in grapevines have been found in asymptomatic and symptomatic grapevines (Abreo et al. 2013; Aroca et al. 2010; Halleen and Petrini 2003; Úrbez-Torres et al. 2015). Other Botryosphaeriaceae have also been shown to infect asymptomatic tissues beyond lesions in grapevines (Amponsah et al. 2011; Billones‐Baaijens et al. 2013a). As in other hosts, the relationship between symptomatic tissue and/or hosts in grapevines has also become an area of interest, with particular emphasis on the role that water stress may play on symptom expression and disease development. 1.2.2 Ilyonectria Species of Ilyonectria are cosmopolitan soil-borne fungi, and the fungi are weak and/or opportunistic pathogens associated with root rot diseases in several economically important hosts, including apple (Manici et al. 2013), avocado (Parkinson et al. 2017), banana (Booth and Stover 1974), Mexican blue palm, Guadalupe palm, Kentia palm (Aiello et al. 2014), clover (Barbetti et al. 2007), ginseng (Rahman and Punja 2005), hazelnut (Guerrero et al. 2014), kiwifruit (Erper et al. 2013), loquat (Agustí-Brisach et al. 2016), olive (Úrbez-Torres et al. 2012), strawberry (Ceja-Torres et al. 2008), and other fruit trees and nut producing hosts (Lawrence et al. 2019). In addition, several Ilyonectria species and closely related Campylocarpon Halleen, Schroers & Crous, 10 Cylindrocladiella Boesew., Dactylonectria L. Lombard & Crous, Neonectria Wollenw., and Thelonectria P. Chaverri & C. Salgado, are known to cause black foot of grapevines (Agusti-Brisach et al. 2013). Black foot pathogens have been isolated from asymptomatic and symptomatic rootstock mother-plants, rootstock cuttings, and young grafted grapevines, in nurseries and young vineyards, as well as from other hosts, which has led to the hypothesis that Black foot pathogens act as latent and/or weak pathogens in grapevines (Agustí-Brisach and Armengol 2013; Carlucci et al. 2017; Dubrovsky and Fabritius 2007; Dumroese et al. 2002; Halleen and Petrini 2003; Úrbez-Torres et al. 2015). 1.2.3 Phaeomoniella chlamydospora and Phaeoacremonium species Petri disease is mainly caused by P. chlamydospora and Phaeoacremonium spp. in young grapevines, while Esca is thought to be caused by the same species of fungi along with colonization of various Basidiomycetes in older grapevines (Gramaje et al. 2018). While Petri disease and Esca are different, for the purpose of this review these two diseases are considered together (Figure 1.3). Phaeomoniella chlamydospora and Phaeoacremonium spp. have been isolated from asymptomatic and symptomatic wood from grapevines and other hosts (Abreo et al. 2011; Edwards and Pascoe 2004; Halleen and Petrini 2003; Panahandeh et al. 2019; Zanzotto et al. 2007). Some grapevines infected with Esca pathogens have displayed symptoms one year but not in the next, which has been proposed to correspond to differences in environmental conditions, primarily temperature and rainfall (Péros et al. 2008; Surico et al. 2000; Calzarano et al. 2018; Marchi et al. 2006). In addition, P. chlamydospora and Phaeoacremonium spp. have been found to occur widely in grapevines in several regions such as in Australia and British Columbia (Canada). However, Esca foliar symptoms are rarely observed in these regions in comparison with Europe, where these symptoms are common (M. Sosnowski, Personal Communication; Úrbez-Torres et al. 2014a). 11 Though not yet studied, the lack of Esca symptom expression in Australia and/or Canada may result from the lack of Basidiomycete species associated with Esca symptoms in Europe, primarily those in Fomitiporia Murrill (Fischer 2002; Fischer and Kassemeyer 2003). On the other hand, other studies have tended to exclude a direct role of Fomitiporia in Esca leaf symptom expression (Calzarano and Di Marco 2007). In a study in Australia, P. chlamydospora was widely found in south-eastern Australia, less commonly in western Australia, and absent from the northern region, indicating that host range may be mainly influenced by climatic factors (Edwards and Pascoe 2004). Nevertheless, these observations suggest that Esca internal symptoms are not necessarily indicative of external symptoms, and that foliar symptom expression may rely also on external abiotic or biotic stress factors (Surico et al. 2000; Zanzotto et al. 2007; Calzarano and Di Marco 2018). Lack of correlation between extent of vascular alteration and foliar symptom expression in Esca may indicate a latent phase (Calzarano and Di Marco 2007). In addition, rainfall has been suggested to be involved in foliar symptom expression regardless of the extent of fungal colonization (Calzarano et al. 2018). 1.2.4 Diatrypaceae The causal organism for Eutypa dieback is the diatrypaceous fungus Eutypa lata (Pers.) Tul. & C. Tul. Although other 23 species in the Diatrypaceae Nitschke have been isolated from grapevine cankers, only E. lata has been proven to cause the foliar symptoms associated with Eutypa dieback (Gramaje et al. 2018; Trouillas and Gubler 2010). These symptoms include internal wood wedge-shaped necroses, cordon dieback, leaf chlorosis, and stunted shoots (Rolshausen et al. 2015) (Figure 1.2). These symptoms mainly appear on grapevines that are 7 years or older, and foliar symptoms typically do not appear for 3 to 8 years post-infection (Rolshausen et al. 2015). Symptoms have also been observed to be inconsistent from year to year, with individual grapevines 12 displaying symptoms one year, while appearing asymptomatic the next (Bertsch et al. 2013; Mundy and McLachlan 2016; Sosnowski et al. 2007b). Several hypotheses have been proposed to explain why symptoms do not appear for extended periods, including the influence of environmental factors. Bruez et al. (2016) found E. lata among other GTD pathogens in asymptomatic grapevines aged 40 years or older and suggested that there was a balance between beneficial organisms and pathogenic fungi, which prevented the pathogens from developing further in grapevines. Eutypa lata and other Diatrypaceae have been isolated from asymptomatic or healthy tissues several centimeters ahead of disease margins, indicating pathogen latency (Pitt et al. 2013). Other hypotheses, including the threshold model (above), could also explain these observations. 1.2.5 Diaporthe The main causal agent of Phomopsis dieback is D. ampelina, which has recently been added to the list of pathogens in the GTD complex (Úrbez-Torres et al. 2013a). Symptoms of Phomopsis dieback are similar to those observed in Botryosphaeria dieback, and include perennial cankers, lack of or delayed bud break and leaf chlorosis (Úrbez-Torres et al. 2013a) (Figure 1.2). Fungi in Diaporthe Nitschke are diverse, inhabiting many different hosts including grapevines, as pathogens, true endophytes, or saprobes (Gomes et al. 2013; Úrbez-Torres et al. 2013a). In grapevines, a diversity of Diaporthe species, including D. ampelina, have been found in asymptomatic and symptomatic grapevine tissues from mature vines and especially from nursery propagated plants, leading to the suggestion that some Phomopsis dieback pathogens may have endophytic phases. However, research into this is limited (Guarnaccia et al. 2018; Kaliterna et al. 2009; Król 2012; Mostert et al. 2000). 13 In general, knowledge of the role that species in the Diaporthaceae Höhn. ex Wehm. may have as latent pathogens comes from hosts other than grapevines. For example, Diaporthe toxica P.M. Will., Highet, W. Gams & Sivasith., the cause of Phomopsis stem blight in Lupinus albus L., has been suggested as a latent pathogen, due to a prolonged latent period under which no visible symptoms occur (Cowley et al. 2012). Tongsri et al. (2016) also suggested an endophytic phase for Diaporthe pathogens in durian leaves and flowers, with symptoms typically occurring 32 d after infection. Similarly, lupins infected with Phomopsis leptostromiformis (J.G. Kühn) Bubák, exhibited a 20 d asymptomatic period after infection (Williamson et al. 1991). Furthermore, symptom expression from D. toxica and P. leptostromiformis infections in lupins has typically been associated with host senescence (Cowling et al. 1984; Williamson and Sivasithamparam 1994). Mature durian leaves also contained a higher density of Diaporthe pathogens than immature leaves (Tongsri et al. 2016). This evidence indicates that these pathogens reach their thresholds for transition from endophytic to pathogenic phase during natural senescence in mature leaves and/or hosts, as suggested by Sieber (2007). However, further research is required to confirm these observations, particularly in grapevines, to determine if Phomopsis dieback pathogens act similarly across different host plants. 1.3 Young vine decline and grapevine nursery stock As outlined above, some of the pathogens responsible for Botryosphaeria dieback, Black foot and Petri disease in grapevines have been isolated from ready-to-plant nursery material and from plants in young vineyards, either separately or together and either from symptomatic or asymptomatic tissues. Accordingly, some of these pathogens may often be latent or weak pathogens, remaining in asymptomatic grapevines for indeterminate periods, and being widespread in vineyards (Agustí-Brisach and Armengol 2013; Carlucci et al. 2017; Dubrovsky and 14 Fabritius 2007; Giménez‐Jaime et al. 2006; Gramaje and Armengol 2011; Halleen et al. 2007a; Halleen and Petrini 2003; Rumbos and Rumbou 2001; Úrbez-Torres et al. 2014a, 2014b). Black foot pathogens are soil-borne and can be present and ready to infect young grapevines when new vineyards are established (Agusti-Brisach et al. 2019; Carlucci et al. 2017; Halleen and Petrini 2003). However, serious infections and disease outbreaks by these pathogens have been found shortly after grapevines were planted (Úrbez-Torres et al. 2014b). This rapid disease development has been suggested to result from: i) exposure to abiotic and/or biotic stress factors during planting in material that is already contaminated with Ilyonectria spp. (Halleen et al. 2006); or ii) greater than normal infection thresholds that host plants cannot withstand (Gramaje et al. 2018; Úrbez-Torres et al. 2014b). In addition, Black foot pathogens have been found in soil in rootstock mother fields and young plant nurseries (Agustí‐Brisach et al. 2014; Cardoso et al. 2013; Whitelaw‐Weckert et al. 2013). These pathogens are prevalent in asymptomatic inner tissues of nursery stock, as shown in 15 nurseries in Northern Spain (Berlanas et al. 2020). A total 1,427 isolates of Black foot pathogens were found, including species of Dactylonectria, Ilyonectria, Neonectria, and Thelonectria from 3,426 ready-to-plant grafted grapevines. Of these pathogens, Dactylonectria torresensis (A. Cabral, Rego & Crous) L. Lombard & Crous made up 75% of all isolates. Further research should be conducted in other regions on the incidence of Black foot and Petri disease pathogens in nurseries and young vineyards to better determine the scale of this problem. In particular, species of Cadophora Lagerb. & Melin have come to attention, since they have been found in grapevines in many regions, including Africa, Europe, North America, and South America (Casieri et al. 2009; Halleen et al. 2007b; Travadon et al. 2015; Úrbez-Torres et al. 2014a). Cadophora species have also been shown to be occur in nursery material in Northern Spain (Maldonado-Gonzalez et al. 15 2020), with Cadophora luteo-olivacea (J.F.H. Beyma) T.C. Harr. & McNew found in 27% of ‘Tempranillo’/’110R’ dormant grapevines. Recent studies have also identified species in Fusarium Link to occur in high numbers in nursery material and vineyards in Canada (Úrbez-Torres et al. 2017). Pathogenicity studies have shown some Fusarium species to be capable of causing necrosis in grapevines to the same extent as Ilyonectria spp. under favourable conditions. This indicates that Fusarium species potentially act as weak or latent pathogens in grapevines, transitioning from endophytic to pathogenic phases under stress conditions, thus playing roles in young vine decline symptom development (Úrbez-Torres et al. 2017). The suggestion that Fusarium is involved in grapevine decline is not new, and had been proposed by Highet and Nair (1995), when a clear association between Fusarium oxysporum Schlecht. emend. Snyder & Hansen and vine decline was found in vineyards in New South Wales, Australia. However, the roles of Cadophora and Fusarium in nursery material and young vineyards, and any interactions of these fungi with Petri disease and Black foot pathogens is not well-understood, although the high incidence of these fungi in nursery material warrants further investigation of them as potentially relevant to young vine decline. In addition, the roles of different types of stress, and the effects of intermittent versus prolonged stress on young grapevines remains unknown. Research on the roles of Cadophora and Fusarium spp. in association with Black foot and Petri disease pathogens should be further investigated. 1.3.1 Infection during the grapevine propagation Several Black foot pathogens have also been discovered in nurseries on grapevine scion cuttings, cutting tools, in water from hydration tanks, and in callusing medium, indicating that infections can occur at several steps in the propagation process, and in the field (Cardoso et al. 16 2013; Gramaje and Armengol 2011). The presence of pathogens during several steps of nursery processes has also been observed with several Botryosphaeriaceae species associated with Botryosphaeria dieback in New Zealand. In a study of three commercial nurseries, 33 to 100% of canes contained detectable amounts of Botryosphaeriaceae on the surfaces of the canes, and 15 to 68% of canes were internally infected with these fungi (Billones-Baaijens et al. 2013b). In a separate study in New Zealand, three of ten apparently healthy ‘3309’ mother grapevines and two out of ten ‘101-14’ grapevines were internally infected with Botryosphaeriaceae in nursery mother blocks. External infection rates in plant bark were also high, with the bark in one out of three shoots infected in one grapevine. Therefore, it was suggested that these Botryosphaeriaceae remained latent in the bark of dormant cuttings, which may be a potential route of infection and spread during nursery propagation (Billones-Baaijens et al. 2015). These results also indicate that disease control should begin in mother blocks from which scion and rootstock material is obtained, and precautions should also be taken during the propagation processes, in which there are several possible avenues of spread of these pathogens. Current best practices for the prevention of transmission have been outlined (Gramaje and Armengol 2011; Waite et al. 2018). In addition to adopting these best practices, rootstocks are a promising avenue to reduce transmission. Berlanas et al. (2019) found that ‘161-49C’, ‘140 Ru’, ‘1103P’, and ‘110R’ had lower abundance of “Cylindrocarpon”-like asexual morphs, compared to ‘41B’ rootstocks. 1.3.2 Rootstock and scion resistance to GTD fungi In grapevines infected with Botryosphaeria dieback pathogens, it has been shown that some scion varieties are more susceptible than others to these pathogens (Úrbez-Torres and Gubler 2009). Studies in New Zealand vineyards showed ‘Sauvignon Blanc’, with 83% incidence of infection, was more susceptible to species of Botryosphaeriaceae compared to ‘Pinot noir’, which 17 had 42% incidence (Baskarathevan et al. 2012). Similarly, ‘Shiraz’ and ‘Sauvignon Blanc’ had greater susceptibility to D. seriata (pathogen recovery 12 to 21 mm from inoculation sites, while ‘Green Veltliner’ and ‘Muscadelle’ were less susceptible (pathogen recovery 8 to 17 mm from inoculation sites (Sosnowski et al. 2017c). Pintos et al. (2018) showed that ‘Savagnin’ grafted plants had greater Botryosphaeria dieback pathogen infections (66%) than other cultivars. Rootstock susceptibility to Black foot has also been evaluated, and among the screened rootstocks, a majority were found to be susceptible, in particular ‘110R’. This rootstock was the most susceptible to infection by Ilyonectria liriodendri (Halleen, Rego & Crous) Chaverri & C. Salgado and Ilyonectria macrodidyma (Halleen, Schroers & Crous) P. Chaverri & C. Salgado, which are considered the main pathogens of Black foot. These results were similar to those for susceptibility of ‘110R’ to P. chlamydospora (Alaniz et al. 2010; Nguyen 2013; Zanzotto et al. 2008). ‘SO4’, ‘Freedom’, and ‘Riparia Glorie’ were also susceptible to I. liriodendri and I. macrodidyma, while ‘44-53’ and ‘St. George’ were more tolerant to these fungi (Nguyen 2013). Susceptibility of different rootstocks to co-infection of Black foot pathogens and those causing either Petri disease or Botryosphaeria dieback should be studied further, along with the impacts of nematode damage on rootstocks infected by GTD pathogens. Symptom expression of grapevines infected with Esca and/or Petri disease pathogens may be influenced by scion variety, with ‘Merlot’ found to be very tolerant to Esca disease (1.5 to 3.6% incidence), while ‘Cabernet Sauvignon’ was less resistant (20.2 to 27.8% incidence) (Christen et al. 2007). Andreini et al. (2009) showed that ‘Trebbiano’ and ‘Sangiovese’ grapevines were found to have lower disease incidence than ‘Cabernet Sauvignon’, which is probably one of the most affected cultivars. Several other studies have noted varying degrees of susceptibility between rootstocks and scions; however, the factors influencing scion and rootstock susceptibility remain 18 unknown (Edwards and Pascoe 2004; Edwards et al. 2007a, 2007b). A detached cane assay study conducted by Martinez-Diz et al. (2019) found variations in colonization of xylem tissues among 20 cultivars, varying from 3 to 34 mm when infected with P. chlamydospora and 9 to 48 mm when infected with P. minimum. Gramaje et al. (2010) showed that 1-year-old rootstocks of ‘110R’ and ‘140Ru’ were very susceptible to P. chlamydospora, Phaeoacremonium spp., and Cadophora spp., while ‘161-49C’ was the most tolerant rootstock to these pathogens. ‘Fiano’ and ‘Sauvignon Blanc’ scions grafted onto ‘SO4’ rootstock, which is intolerant to water stress, displayed increased disease incidence compared to when grafted onto ‘1103P’ rootstock, which is more drought tolerant (Murolo and Romanazzi, 2014). These results are similar to those of Cardoso et al. (2006), in that scion and/or rootstock resistance to abiotic or biotic stress factors may influence disease susceptibility. This requires further investigation, however. Cultivar variety may affect symptom expression in grapevines infected with E. lata, with studies showing that different cultivars were more resistant or susceptible than others to Eutypa dieback (Baumgartner et al. 2019; Cardot et al. 2019). ‘Merlot’ was the most resistant to Eutypa dieback (38% plants asymptomatic), while ‘Ugni Blanc’ was very susceptible (87% of plants with symptoms). (Cardot et al. 2019). Péros and Berger (1994) assessed different cultivars for foliar symptoms 5- and 10-weeks post-inoculation. ‘Cabernet Sauvignon’ was the most susceptible, (90% of plants displaying foliar symptoms), followed by ‘Ugni Blanc’, while ‘Sauvignon’ was the least susceptible. Research is being instigated on rootstock tolerance to GTD pathogens under abiotic or biotic stress conditions. However, further studies are required on the tolerance of different rootstock and cultivars to Esca disease while under stress (Sosnowski et al. 2017a; Sosnowski et al. 2017b). In a field grapevine study, trunk disease severity was greater in red than white cultivars 19 when infected with E. lata. ‘Petit Verdot’ had the least trunk symptom severity while ‘Odola’ had the greatest for red cultivars, while ‘Traminer’, ‘Petit Meslier’, ‘Muscadelle’, and ‘Green Veltliner’ displayed no trunk symptoms. Among white cultivars infected with E. lata ‘Sauvignon Blanc’ had the greatest trunk symptom severity (Sosnowski et al. 2017c). In a follow-up detached cane assay (Sosnowski et al. 2017c), E. lata was recovered 20 to 23 mm from inoculation sites in ‘Shiraz’ and ‘Sauvignon Blanc’ compared to 4 to 14 mm in ‘Petit Meslier’, ‘Petit Verdot’, and ‘Muscadelle’ grapevines, confirming the field assessments. A later field inoculation experiment (Sosnowski et al. 2017c) also showed that ‘Petit Verdot’ and ‘Muscadelle’ had increased resistance to pathogen colonization. Clonal susceptibility for ‘Shiraz’ varied between clones aged 21 to 34 years, for ‘Sauvignon Blanc’ clones between 20 to 44 years, and ‘Chardonnay’ clones between 18 to 45 years, with less symptom severity in younger than older grapevines. Plant age and resistance to environmental stresses in relation to pathogen resistance should be further explored, to determine how resistance varies as grapevines age. Rootstock and scion varieties present a promising avenue for tolerance to GTD infection. However, further research is required on this subject, including host tolerance to infection under abiotic and biotic stress conditions. Research has been conducted on rootstock performance in varying growing condition, but the roles that abiotic and biotic stress factors play for grapevine latent infections by GTD pathogens in rootstocks, particularly during the first few years after planting, have not been explored (Cuneo et al. 2020; Fort et al. 2017). In particular, rootstock and scion tolerance to stress and symptom expression should be assessed at the regional levels, to determine the best rootstock selections, and which ones are best suited for unique stress factors encountered in each region. 1.4 Abiotic stress factors and GTD symptom expression 20 1.4.1 Botryosphaeriaceae Many members of the Botryosphaeriaceae are not host specific, but rather the environment probably plays a major role in host affinity (Ibrahim et al. 2017; Slippers and Wingfield 2007). Symptom expression in many different hosts has been found to coincide with periods of drought or other extreme weather events (Acimovic et al. 2018; Bendixsen et al. 2015; Brown-Rytlewski and McManus 2000; Crous et al. 2017; Golzar and Burgess 2011; Jeff-Ego and Akinsanmi 2019; Jurc et al. 2006; Ma et al. 2001; Slippers and Wingfield 2007). Direct evidence for stress induced symptom expression includes the research of Old et al. (1990), which showed that Neofusicoccum ribis (Slippers, Crous & M.J. Wingf.) Crous, Slippers & A.J.L. Phillips, caused larger cankers in Eucalyptus when trees were defoliated or attacked by insects, than where insect attack was absent. Water stress is probably the most important factor in symptom expression, and has been the most extensively researched out of all potential stress factors (Brown-Rytlewski and McManus 2000; Scala et al. 2019; Slippers and Wingfield 2007). Similarly, water stress which reduced leaf water potentials increased the symptoms caused by B. dothidea. Plants remained resistant to B. dothidea infection until a water potential threshold of -12 to -13 MPa was reached (Schoeneweiss 1981; Wene 1979). However, it was necessary to sustain stem water potentials at less than -12 MPa for at least 3 d to allow for predisposition to B. dothidea, and that under field conditions, the effects were reversible if drought was not sustained. Similarly, Ma et al. (2001) in a greenhouse study showed that old pistachio trees could not recover from the combined effects of water stress and B. dothidea infection, while young trees were more resilient. This may be explained by the ability of B. dothidea to grow rapidly when pistachio tree stem water potentials were decreased from 0 MPa to -2 MPa (Ma et al. 2001). Pusey (1989) also found that drought conditions resulting in leaf water potential of -3.0 MPa increased lesion length in peach trees. These results indicate that symptom 21 expression is a combination of reduced host resistance due to stress and increased pathogen colonization due to optimal growth conditions. Similarly, Cardoso et al. (2006) hypothesized that resistance in a dwarf cashew clone to Lasiodiplodia theobromae (Pat.) Griff. & Maubl. was due to indirect mechanisms related to the drought tolerance of the clone. In grapevines, under drought conditions, ‘Cabernet Sauvignon’ grapevines infected with N. parvum had larger lesions than plants provided with adequate water; however, grapevines infected with Neofusiccocum luteum (Pennycook & Samuels) Crous, Slippers & A.J.L. Phillips had larger lesions when grown at high soil moisture levels, indicating physiological conditions induced by low moisture content may affect wood symptom expression, but other factors may be involved that influence lesion length (Galarneau et al. 2019). Qiu et al. (2016) reported similar results, where ‘Chardonnay’ grapevines infected with N. parvum, B. dothidea, L. theobromae, and D. seriata produced larger necrotic wood lesions in water stressed grapevines than in unstressed vines. Van Niekerk et al. (2011a) also showed that ‘Shiraz’ grapevines infected with Neofusiccocum australe (Slippers, Crous & M.J. Wingf.) Crous, Slippers & A.J.L. Phillips, N. parvum, L. theobromae, and D. seriata developed larger lesions and reduced shoot mass when water stressed compared with unstressed plants. Other host stress factors, including freezing damage, root damage from transplanting, pathogenic nematode infestations, and suboptimal environmental conditions for growth (nutrient-poor or high-salt soils, excessive cold or heat) have also been associated with GTD symptom expression in multiple hosts. In addition to these associations, some research has been completed on freezing stress, showing similar effects to water stress in Cornus sericea L. Localized cold exposure of this host to -30℃ produced no visible signs of freeze damage, but colonization by B. dothidea occurred solely in the cold exposed areas and did not extend significantly beyond the margins. Similar results were also found in Sorbus aucuparia L. infected with B. dothidea 22 (Schoeneweiss and Wene 1980; Wene 1979). Manawasinghe et al. (2018) also suggested that B. dothidea outbreaks in China were associated with other stress factors, such as increased temperature and pest pressure. 1.4.2 Ilyonectria Several factors have been shown to affect symptom expression in hosts infected with Ilyonectria spp. In apple and ginseng, root age affects lesion severity, with young roots developing more severe lesions compared to older roots, which displayed shallow lesions (Manici et al. 2018; Rahman and Punja 2005). Abiotic and biotic factors also probably affect symptom expression, including root damage, acidic soils, temperature, and drought. (Barbetti et al. 2007; Rahman and Punja 2005; Ruiz-Gómez et al. 2019). Black foot symptom expression on grapevines has been associated with three categories of stress factor: (1) nursery-related factors such as cold storage, grapevine defects, and low light and other conditions leading to low carbohydrate levels in the plants; (2) vineyard conditions and plant establishment stress factors, including nutrient deficient soils, poor soil drainage leading to low oxygen levels, drought, soil compaction, J-rooting, and other factors contributing to poor root development and temperature extremes; and (3) biotic factors, including fungal pathogens, pathogenic nematodes, and insect pests (Agusti-Brisach et al. 2019; Barbetti et al. 2007; Gramaje and Armengol 2011; Halleen et al. 2006; Nguyen 2013; Rahman and Punja 2005; Ruiz-Gómez et al. 2019; Stamp 2001) (Figure 1.4). Moreno-Sanz et al. (2013) suggested that symptom expression primarily occurs when grapevines are under stress, as defined by hindering of normal growth and development and photosynthesis, and causing physiological responses, e.g. increased stress hormones that lead to numerous responses that enhance plant survival at the expense of crop 23 performance. As Ilyonectria spp. are likely weak or opportunistic pathogens, they probably only invade weakened or dead roots (Halleen et al. 2006; Jankowiak et al. 2016; Scheck et al. 1998). These conclusions were further supported by Probst et al. (2012), who found that increased cold storage time correlated with increased disease incidence and severity for several Black foot pathogens. Partial defoliation also decreased root dry weight and increased disease severity resulting in increased canker development in young grapevines infected with I. destructans, which was probably due to reduced carbohydrate levels (Brown et al. 2012). This is similar to the results of Old et al. (1990) stated above. 1.4.3 Phaeomoniella chlamydospora and Phaeoacremonium species Several stress factors have been associated with, or are thought to influence, symptom expression from Esca and Petri disease pathogens, including biological factors such as nematode or other fungal pathogens, and abiotic factors such as cold exposure, and those associated with poor soil quality including low water and nutrient supply, extreme pH, and excess salt (Corino et al. 2004; Fischer and Ashnaei 2019; Oliveira et al. 2009; Oliveira et al. 2013; Stamp 2001). Drought has received the most attention. In young grapevines, wood symptom expression, plant growth, and likely survival of grapevines infected with Esca and/or Petri disease pathogens depend on sufficient water supply to avoid stress (Fischer and Kassemeyer 2012). In two separate studies conducted in Australia, ‘Cabernet Sauvignon’ and ‘Zinfandel’ grapevines were infected with P. chlamydospora and subjected to either an irrigation regime that resulted in no measurable water stress, 50% irrigation, or 25% irrigation. While stomatal conductance and leaf water potentials were both progressively reduced in the 50% and 25% irrigation regimes, for both non-infected and P. chlamydospora infected grapevines, in each irrigation treatment the infected grapevines displayed greater stomatal conductance and less leaf water potential compared to non-infected 24 grapevines. Furthermore, ‘Chardonnay’ grapevines displayed grater leaf water potentials than ‘Cabernet Sauvignon’ and ‘Zinfandel’, which may indicate ‘Chardonnay’ is more tolerant of P. chlamydospora infection (Edwards et al. 2007a, 2007b). On potato dextrose agar (PDA) amended with either potassium chloride or sucrose to generate osmotic water potentials between -0.3 and -8.3 MPa, two out of three isolates of P. chlamydospora grew at down to -8.3 MPa (Whiting et al. 2001), indicating that the the fungus could grow in severely water-stressed plant tissues. Results by Lima et al. (2017) indicated that a combination of water stress and Esca infection increased xylem sap nutrient concentrations, allowing for increased growth of the pathogens. In contrast, heavy rainfall from fruit set until veraison were associated with increasing Esca foliar symptoms, reducing grape quality (Calzarano and Di Marco 2018). These results show that water stress may not be the only water status condition in grapevines that favour Esca development. 1.4.4 Diatrypaceae It is well known that precipitation plays a role in E. lata distribution and frequency of occurrence. However, the effects of environmental conditions on symptom expression are not well understood. In a survey in France, the incidence and frequency of Eutypa dieback varied among different regions, indicating that climatic conditions, particularly temperature regimes which varied among regions, were major causes of differences in Eutypa occurrence (Guérin-Dubrana et al. 2013). Magarey and Carter (1986) also found E. lata to be restricted to geographical locations where annual rainfall exceeded 350 mm, or in vineyards subject to overhead watering systems, indicating that precipitation affects E. lata distribution. Likewise, winter precipitation may be an additional requirement for the full development of E. lata in particular regions (Petzoldt et al. 1983; Úrbez-Torres et al. 2019; van Niekerk et al. 2011b). 25 Guérin-Dubrana et al. (2013) and Sosnowski et al. (2007a) found that symptom expression and severity were influenced by climactic conditions in different regions. In Australia, E. lata isolates obtained from regions with intermediate climate had greater effects on symptom expression in a shadehouse experiment (Sosnowski et al. 2007a). These isolates may have had greater virulence due to more favourable climactic conditions in the shade-house compared to the field. In a separate study in Australia, grapevines grown in extreme temperature and moisture conditions had increased foliar symptoms, but internal wood staining was not related to disease severity (Sosnowski et al. 2007a; Sosnowski et al. 2011). This is consistent with other reports on Botryosphaeriaceae, that indicate wood staining does not relate to colonization rates or correspond to foliar symptoms, but may be a consequence of wounding (Travadon et al. 2013). Grapevines infected with E. lata also displayed decreased leaf water potential compared with non-infected grapevines when exposed to moisture deficits, suggesting that the pathogen increased sensitivity to water stress which may facilitate symptom development (Sosnowski et al. 2011). Decreased leaf water potentials may be due to Eutypa dieback pathogens occluding the xylem vessels, but this needs to be confirmed (Sosnowski et al. 2011). 1.4.5 Diaporthe Host stress may also affect the transition from the endophytic to pathogenic phase. Hulke et al. (2019) found a correlation between increased disease incidence of Phomopsis stem blight of sunflowers and climate data over two decades. Diaporthe citri F.A. Wolf, J. Agric. also has been suggested to cause increased symptom expression in yuzu trees (Citrus ichangensis x Citrus reticulata var. austera) under adverse weather conditions (Kim et al. 2015). However, investigations into the effects of stress on Diaporthe symptom expression are recent and limited, especially with grapevines. 26 1.5 Roles of biotic stress factors on GTD symptom expression Though still limited, the number of studies investigating the roles that abiotic stress factors play in GTD symptom expression and disease progression is far greater than those investigating effects of biotic factors. Among the few studies available, two biotic factors have been identified: nematodes and other fungal pathogen interactions. In addition, few studies have primarily focused on the roles of biotic factors on GTD caused by Ilyonectria fungi. 1.5.1 Parasitic nematodes affecting Ilyonectria symptom expression Plant parasitic nematodes have been shown to affect disease development and symptom expression across a range of pathosystems (Powelson and Rowe 1993; LaMondia 2003; Cao et al. 2006), but data are scarce on their interactions with Ilyonectria spp. (Hastings and Bosher 1938; Booth and Stover 1974; Sutherland 1977; Rahman and Punja 2005). While not involving Ilyonectria, bacterial canker of Prunus species fruit and nut trees provides an example of how nematodes can influence development and severity of a disease of woody perennials. Ring nematode infestation of roots has been shown to increase lesion length of bacterial canker in peach, caused by Pseudomonas syringae van Hall, (Cao et al. 2005; Cao et al. 2006). Ring nematode infestations lead to nitrogen deficiency in peach trees, which was suggested as the mechanism for nematodes increasing susceptibility to infection (Cao et al. 2011). The effects of nitrogen deficiency on disease severity, measured by lesion length are mixed, however, with Cao et al. (2011) finding no association between host nitrogen levels and lesion length in peach, while in a separate study, Cao et al. (2013) showed that nitrogen fertilization led to significantly decreased lesion lengths in almond trees. The mechanisms for nitrogen fertilization effects were 27 not determined, but it was suggested that the nitrogen may decrease nematode populations in the soil or by indirect effects such as increased host vigour. Hastings and Bosher (1938) showed that growth of seedlings of several hosts was inhibited by a combined treatment of root lesion nematodes (Pratylenchus pratensis) (de Man) Filipjev and I. destructans to a greater extent than either pathogen or nematode alone. Rahman and Punja (2005) also found that development of root rot in ginseng was contingent on minor wounds, which could occur from nematode damage. This was also shown for root rot in clovers, where increased stress symptoms occurred when root systems were damaged (Barbetti et al. 2007). Sutherland (1977) inoculated Douglas fir seedlings with dagger nematodes (Xiphinema bakeri Williams) and I. destructans, separately or in combination, and showed that while the nematode caused corky root symptoms on its own, the combined inoculation did not give a synergistic interaction. In vineyards, several genera of plant parasitic nematodes are ubiquitous, including root-knot nematodes (Meloidogyne Goeldi), ring nematodes (Mesocriconema xenoplax Andrassy), root lesion nematodes (Pratylenchus Filipjev), and dagger nematodes (Xiphinema Cobb) (Storey et al. 2017) (Figure 1.4). Recently research has begun to focus on nematode/pathogen relationships. Rahman et al. (2014) assessed population densities of the ring nematode (Mesocriconema xenoplax (Raski) Loof & De Grisse) and the citrus nematode (Tylenchulus semipenetrans Cobb) under symptomatic and non-symptomatic vines infected with Ilyonectria spp. Nematode population densities did not differ between the symptomatic and asymptomatic vines. However, this study was conducted during a drought period, which is not considered conducive to the spread of Ilyonectria spp. or to the buildup of nematode populations, and was carried out with ‘Ramsey’ rootstock, which is resistant to ring nematodes. Similarly, co-infections of Black foot pathogens 28 with other pathogenic fungi have also been associated with an increase in disease incidence and severity (Probst et al. 2012). 1.5.2 Fungal pathogen interactions and infection thresholds The interactions between Ilyonectria spp. and other fungal pathogens has been documented in apples. For apple replant disease (ARD), a combination of I. macrodidyma and Pythium irregulare Buisman reduced seedling growth when compared to single inoculations of each pathogen (Tewoldemedhin et al. 2011). Braun (1991) also measured reductions in plant weight and size when apple trees were co-inoculated with P. irregulare and Thelonectria lucida (Höhn.) P. Chaverri & C. Salgado, which had previously been shown to be pathogenic to apple seedlings (Jaffee 1982). Research on effects of co-infections on expression of GTDs has recently increased. Young ‘Chardonnay’ grapevines infected with pathogens causing Botryosphaeria dieback and Black foot gave more severe decline compared to single pathogen inoculation, while ‘Sauvignon Blanc’ grafted onto ‘101-14’ infected with Petri disease and Black foot pathogens also had increased disease incidence and severity than with single pathogen infections (Probst et al. 2012; Whitelaw‐Weckert et al. 2013). This is important, because co-infections with Black foot and Botryosphaeria dieback pathogens, and especially with those causing Black foot and Petri disease, are common in nurseries and young vineyards (Halleen et al. 2007a; Halleen and Petrini 2003; Úrbez-Torres et al. 2014a; 2014b). Fungal interactions in infected grapevines probably affect disease development. Several Basidiomycetes, particularly Fomitiporia species, have been associated with Esca symptoms in old grapevines, and are commonly associated with P. chlamydospora and Phaeoacremonium spp. 29 (Cloete et al. 2014; Fischer 2002; Fischer and Kassemeyer 2003; White et al. 2011). Fischer and Kassemeyer (2003) isolated F. mediterranea M. Fisch., along with P. chlamydospora and Phaeocremonium spp. from symptomatic tissues. In addition to F. mediterranea, other species of Hymenchaetales Oberw, have been associated with Esca symptoms, including species in Fomitiporella Murrill, Inocutis Fiasson & Niemela, Inonotus P. Karst., and Phellinus Quél. (Cloete et al. 2015a; Cloete et al. 2015b). This has led to the hypothesis that some Basidiomycetes are involved in the development of Esca symptoms, but has yet to be confirmed (Gramaje et al. 2018). Similar to Ilyonectria spp. co-infections with other GTD pathogens discussed above, the presence of Basidiomycetes may act as inciting factors in symptom expression. However, further research is required to demonstrate specific roles that Basidiomycetes play in Esca disease of grapevines. While environmental and biotic triggering factors are likely to be important, threshold fungal concentrations within grapevines may also affect the switch from endophyte to pathogen. Inoculation of up to 104 conidia mL-1 suspensions of N. luteum increased lesion lengths, but greater inoculum concentrations decreased lesion lengths (Amponsah et al. 2014). This contrasts with B. dothidea, where spore concentrations up to 106 conidia mL-1 gave increasing lesion lengths in non-wounded peach bark (Pusey 1989). Reduced lesion length may be due to competition in small inoculation sites, which was 3 mm diam. In this case, further research into effects of spore concentration on disease severity and grapevine mortality has been suggested (Amponsah et al. 2014). Additionally, differences were observed in virulence among different isolates of individual species of Botryosphaeriaceae (Úrbez-Torres and Gubler 2009). Secondary metabolites produced by different GTD fungi, such as Eutypines produced by E. lata, are known to be phytotoxic (Andolfi et al. 2011). Other species producing secondary metabolites include P. chlamydospora and several Botryosphaeriaceae involved in Botryosphaeria dieback (Andolfi et al. 2011). 30 However, the roles these secondary metabolites play in symptom expression and differences in virulence, in particular among the other GTD fungi, are unclear. In addition, how secondary metabolites may respond in grapevines under either abiotic or biotic stress needs to be elucidated. 1.6 Climate change and viticulture practices: impacts on abiotic and biotic stress factors and GTD symptom expression The predicted effects of climate change vary from variations in average temperatures and precipitation, to extreme weather events such as flooding or droughts (Ali 2013; Dixon 2012). Several projections have predicted increasing global temperatures and variations in precipitation, as well as expressing concerns about these changes affecting global food security. Temperature increases associated with climate change are expected to increase occurrence of heat waves, agricultural droughts, and river floods, which will likely impact grape-growing regions (Arnell et al. 2019). Environmental stress factors such as drought, temperature increases, and salinity are of concern for plant health, particularly relating to climate change (Ahuja et al. 2010). Grapevines are sensitive to climate change, particularly temperature, which may put them at increased risk in major grape-growing regions such as the Mediterranean basin (Biasi et al. 2019; Giorgi and Lionello 2008). In addition to likely crop stress due to climate change, it is expected that these climate changes will lead to increased pathogen incidence in crops, due to improved survivability in milder winters, altering the geographic distribution of pathogens, and possible modifications in host resistance to pathogens and plant-pathogen relationships (Dixon 2012). In particular, climate change may affect grapevines grown in optimal grape-growing regions such as California, where increasingly mild winters may prevent latent bud hardening and pests and pathogens reduced by cold winters may increase (Dixon 2012). Similar effects have been observed for Fusarium infections in maize, where mild winters resulted in increased biomass of Fusarium in maize debris 31 while frost-thaw cycles reduced biomass and reduced risks of infection (Lukas et al. 2014). Extreme weather conditions such as drought may impact plant defenses, and increased temperatures may lead to plant stress in summers when temperatures peak. However, predictions on effects of climate change on plants are difficult to confirm, as experiments are typically carried out in controlled environments rather than in the field (Elad and Pertot 2014). 1.6.1 Water deficit irrigation Some viticulture practices employed to improve grape and wine quality may be stress factors with unintended consequences for GTD’s. One example which requires further investigation is regulated deficit irrigation, which involves maintaining water deficits for particular seasonal development periods. For wine grapes, deficit irrigation is often employed from after fruit set until veraison, when normal irrigation recommences (Chaves et al. 2010; Intrigliolo and Castel 2010; Santesteban et al. 2011). Deficit irrigation possibly increases grape and wine quality (Acevedo-Opazo et al. 2010; Santesteban et al. 2011), although there are conflicting reports on the effectiveness of regulated deficit irrigation for improved quality of grapes in all situations (Acevedo-Opazo et al. 2010; Chaves et al. 2010; Hepner et al. 1985; Lauer 2012; Santesteban et al. 2011). Úrbez-Torres (Personal communication) observed a difference in GTD incidence on mature ‘Cabernet Sauvignon’ vineyards grown for different wine quality in Chile. Vineyard blocks grown for premium quality wines at high elevation with poor soils and subjected to a severe water deficit irrigation to improve quality, showed a much greater incidence of Botryosphaeria dieback than other blocks cultivated for the production of bulk table wines grown in valleys on fertile soils and subjected to no water deficit irrigation. Although these observations require explanation, they are consistent with the hypothesis that water stress may enhance GTD development. In contrast, Sosnowski et al. (2016), studying ‘Cabernet Sauvignon’ in South Australia to assess effects of 32 water stress on pruning wound susceptibility to E. lata and D. seriata, showed that water stress did not increase susceptibility to pruning wound infections. Their results indicated that drought and deficit irrigation practices were not likely to contribute to increased prevalence of GTD in vineyards. Regulated deficit irrigation for grapevines involves moderate water stress, but the effects of this stress on GTD symptom expression and disease development are unclear. These should be further investigated as a potential grapevine health concern for plants that may contain asymptomatic infections (Santesteban et al. 2011; Tarara and Perez Peña 2015). 1.6.2 Vine pruning and training practices Travadon et al. (2016), in a study in France assessed incidence of Esca on ‘Syrah’ ‘Mourvèdre’ vines grafted onto ’140Ru’ rootstocks, to assess effects of minimal pruning and spur pruning on a bilateral cordon system. Both cultivars displayed more symptomatic vines when spur pruned than those that had minimal pruning. Mean necrosis was likewise greater (35%) for spur pruned vines than those receiving minimal pruning (21%). Similar results were obtained in California, United States of America, in ‘Cabernet Sauvignon’ responses to Eutypa dieback symptoms, with minimal pruning in the summer producing less dieback than the pruning of dormant vines or using the Sylvoz system (Gu et al. 2015). Lecomte et al. (2012) observed that vineyards with the Lyra training system had foliar symptoms in 56% of grapevines, but also had a low proportion of trunks affected. This indicated that the long cordons of the Lyra training system may have been one of the causes. In a follow-up study in France, it was confirmed that different training practices affected Esca foliar symptom severity. Grapevines with short cordons had more severe symptoms than grapevines with long cordons. In addition, plants which were pruned less had less severe symptoms than those that were heavily pruned (Lecomte et al. 2018). These surveys only recorded foliar symptoms, but the results were similar to those of Travadon et al. (2016), who 33 recorded decreased necrosis in minimally pruned grapevines. While these results are promising, further research is required for other GTD fungi, particularly for those causing Black foot, Botryosphaeria dieback, and Phomopsis. Furthermore, the effects of pruning and training practices require further investigation to determine if and/or how they influence wood necrosis. In addition, grapevines under different pruning regimes and training systems may respond differently to stress conditions, and thus may develop GTD symptoms differently. No studies have been reported on this subject, but these would likely provide greater understanding of these interactions, which would improve management of GTD under different pruning and training systems. 1.6.3 Endophytes as potential reducers of host stress Endophytes may increase plant tolerance to abiotic and biotic stresses associated with climate change (Chakraborty and Newton 2011). Endophytes possibly contribute several beneficial attributes to their hosts, including enhanced water use efficiency leading to improved survival during drought, increased nutrient uptake and recycling, reduced stress associated with temperature increases or decreases, and increased tolerance to soil salinity, alkalinity, and heavy metals (Lata et al. 2018). In two varieties of rice plants not well adapted to high soil salinity or drought, class 2 fungal endophytes enhanced host water use efficiency, and increased growth rates, reproductive yield, and biomass under drought and high salinity in greenhouse experiments. Class 2 endophytes conferred cold tolerance to rice plants subjected to temperatures ranging from 5℃ to 20℃, in a plant growth chamber (Redman et al. 2011). Similarly, fungal endophytes increased grain yields and second-generation seed viability in wheat subject to drought and increased temperature (Hubbard et al. 2014). 34 The use of Pythium oligandrum Drechsler is another example of reduction of disease progression caused by P. chlamydospora. In a 4-month greenhouse assay with ‘Cabernet Sauvignon’ canes infected with P. chlamydospora, colonization of roots by P. oligandrum resulted in up to 50% reduction of necrosis, compared to grapevines infected with P. chlamydospora without P. oligandrum (Yacoub, et al. 2016). In addition, in ‘Cabernet Sauvignon’, expression of several genes associated with P. chlamydospora infection was greater in vines with P. oligandrum colonization of the roots than in vines without P. oligandrum, suggesting that P. oligandrum induced resistance in grapevines. In addition, P. oligandrum may also reduce biotic stress by the way of fungus-produced tryptamine, an auxin compound involved in plant growth (Benhamou et al. 2012; Floch et al. 2003). Further studies, in particular in other grapevine cultivars and during prolonged intervals are required to determine the efficacy of P. oligandrum as a potential biocontrol agent. Promotion of plant growth may aid reduction in the effects of abiotic and biotic plant stress, but this requires further confirmation to determine what effects this endophyte may have on grapevines infected with GTDs. In grapevines, Pseudomonas fluorescens Migula promoted high salinity tolerance via production of endogenous melatonin in host roots, and promoted plant growth (Ma et al. 2016). Bacterial and fungal endophytes have been associated with grapevine tolerance to drought, high salinity, heavy metals, and high and low temperatures (Pacifico et al. 2019). These factors include associated biotic tolerance to some GTD pathogens such as P. chlamydospora, and N. parvum. The use of endophytes to reduce environmental stress should be further investigated in relation to GTD disease development, to determine the roles that bacterial and fungal endophytes may have. 1.6.4 Arbuscular mycorrhizal fungi as potential stress reducers 35 While not endophytes by definition, arbuscular mycorrhizal (AM) fungi (Glomeromycota Walker & A. Schüßler) colonize roots, and these organisms can reduce stress associated with high and low temperatures, as well as water stress, soil compaction, and salinity (Charest et al. 1993; Ellis et al. 1985; Harrier and Watson 2003; Lu et al. 2007; Zhu et al. 2010). Suggested mechanisms for increased tolerance include improved plant nutrition, compensation for damaged roots, competition for infection sites, alterations in root anatomy or morphology, alterations in the mycorrhizosphere, and activation of plant defenses (Azcón-Aguilar and Barea 1997). In addition to increased environmental stress tolerance, some evidence suggests that AM fungi increase tolerance to pathogens, in particular root pathogens, although the mechanisms are as yet unclear (Comby et al. 2017; Whipps 2004). AM fungi may also affect tolerance to airborne pathogens, and plants have been reported to have increased resistance to hemibiotrophic and necrotrophic fungal infection, although results have been varied and some have reported increased susceptibility to these pathogens (Comby et al. 2017). In grapevines, AM fungi were shown to increase root and shoot biomass in ‘110R’ rootstocks infected with Armillaria mellea (Vahl) P. Kumm., compared to plants without AM fungi (Nogales et al. 2009). In greenhouse studies, AM fungi have been shown to influence disease severity in hosts infected by GTD pathogens. Petit and Gubler (2006) inoculated Vitis rupestris Scheele with the AM fungus Rhizophagus irregularis (Błaszk., Wubet, Renker & Buscot) C. Walker & A. Schüßler and I. macrodidyma, a causal agent of Black foot, in a controlled greenhouse environment. While I. macrodidyma was recovered from both AM fungus-inoculated roots and the non-inoculated roots, grapevines inoculated with R. irregularis displayed decreased disease severity. Also, apple trees with roots inoculated with AM fungi had decreased symptom expression and increased survival when infected with N. ribis compared to experimental controls (Krishna et al. 2010). 36 However, research into the effects of AM fungi on disease severity has given mixed responses, and, in some cases, these fungi have been shown to be detrimental to the plant health. In ‘Riparia gloire’ rootstocks infected with I. liriodendri, colonization by R. irregularis increased abundance of the pathogen in the roots, and had no effect on plant growth (Holland et al. 2019). Likewise, in ‘101-14’ rootstocks planted in soil inoculated with I. liriodendri, a positive linear relationship was demonstrated between AM fungus colonization of roots and proportions of necrosis found (Vukicevich et al. 2018). These results indicate that the use of commercial AM fungi inoculants for Vitis vinifera should be further investigated to determine the impacts of these fungi on GTD development. The use of AM fungi to reduce plant stress is a potential avenue of research, and should be considered for its potential to increase environmental stress and disease tolerances, and the mechanisms of these tolerances should be further investigated. This should particularly be in relation to the potential environmental stress and symptom expression relationships for many GTDs. Field studies are required to determine effects of AM fungi on health of GTD-infected grapevines in field conditions. Field inoculations with commercial AM fungi have also given inconsistent response, and inoculum may spread beyond the intended fields. Further research should therefore be conducted on the use of regionally-based AM fungi inoculants (Farmer et al. 2007; Kokkoris et al. 2019). This is of particular importance for Vitis, as AM fungal communities differ between grapevine rows and inter-row plants. This indicates that grapevines may select for particular AM fungal communities (Holland et al. 2014). Increasing the diversity of cover crops may increase the diversity of AM fungal communities, allowing greater chances of grapevine root colonization, and also reduce soil-borne pathogen populations (Holland et al. 2014; Vukicevich et al. 2016). 37 1.7 Conclusions and future directions Limited research has been conducted on endophytes in economically important crops, particularly on their roles as latent pathogens. This is an increasing area of interest particularly in grapevines, where many GTD fungi have been suggested to have endophytic phases. Many of the hypotheses addressing transition from the endophytic to the pathogenic phases remain to be tested, in particular, the threshold model developed by Sieber (2007). 1.7.1 GTD as latent pathogens The roles of Botryosphaeriaceae as potential latent pathogens has been well-documented particularly in forest trees. However, research has just begun in grapevines. Several Botryosphaeriaceae have been discovered in the asymptomatic and symptomatic grapevine wood. Effects of abiotic and biotic stress factors on disease development are still largely unknown, and research has only recently begun on the effect of pathogen quantities on symptom expression. Likewise, the role of Ilyonectria spp. has been examined for a number of hosts, with several factors influencing symptom expression. Research on Ilyonectria spp. in grapevines is limited, however, with the most studies conducted in nurseries and young vineyards where these pathogens are mostly found. Biotic factors affecting nematode infection of the roots and co-infections of GTD fungi are likely to be important and productive areas of research. Petri disease pathogens in grapevine nurseries and young vineyards have been associated with stress factors in young grapevines, while Esca has been associated with biotic factors including Basidiomycete co-infections. Several abiotic factors have been associated with Petri disease and Esca. Eutypa dieback has received more research attention than other GTDs, but knowledge is limited and associations between symptom expression and climatic factors have only recently been assessed. 38 Phomopsis dieback as a GTD is a new area of research, and most investigations of this disease have been in other hosts. Several species of Diaporthe have been suggested as latent pathogens, and their discovery in asymptomatic and symptomatic grapevines indicates that they are also latent pathogens in grapevines. 1.7.2 Latent pathogens in young vineyards The roles of latent pathogens, particularly those causing Black foot and Petri disease in mother blocks, nurseries and young vineyards, are of particular importance. Many of these pathogens are widespread, and conditions during nursery propagation processes are optimal for the latent transmission of these fungi. Young grapevines may also be more susceptible to disease development and less able to recover, compared with mature grapevines which display symptoms erratically. Further study of the stress factors which may induce disease development is required to determine the nature of these pathogens, and indicate potential treatment options, including best planting conditions to avoid stress and the potential transition from endophytic to pathogenic phases by these fungi. Further study is required of abiotic stress, particularly in nurseries. Research on biotic stress, particularly from pathogenic nematodes and fungal interactions will also be worthwhile. Recently, work has begun on fungal interactions in GTDs, with grapevine initially infected with D. seriata followed by infection by one of N. parvum, or P. chlamydospora. Initial results have shown reduced lesion lengths and changes in host physiology (Wallis et al. 2019). These indicate an urgent need for further research, particularly focusing on fungal interactions involving Black foot and Petri disease pathogens in young grapevines. 1.7.3 Methods for host stress reduction 39 Reducing stress on grapevines is an important first step, but not all stress in the field can be controlled. Therefore, bacterial and fungal endophytes and AM fungi known to enhance resistance of plants to some stress factors are likely to be productive research candidates. However, the effects of AM fungi on disease development are variable, and regionally-based approaches to AM fungi are recommended for further study. Rootstock and scion variety is also likely to be a productive area of study, and research is required to show how resistance of rootstocks to abiotic and biotic stress factors affect GTD development in grapevines. While GTD fungi potentially acting as latent pathogens have long been the subject of speculation, recent discoveries aided by molecular biology research tools have led to the resurrection of this hypothesis. Study on latent pathogens has recently commenced, and requires urgent focus given the growing impacts of GTDs in most grape production areas. Endophytic phases may lead to uncontrolled spread of potential pathogens throughout nurseries and vineyards and may cause harm within young vineyards before grape growers can recoup capital investments in new plantings, or may shorten the economic lives of vineyards. Effects of climate change along with specific viticulture practices may further increase the incidence and severity of GTDs, causing increased economic losses. Understanding the effects of these stress factors and development of methods to minimize stress on grapevines in relation to GTD disease development, are crucial first steps to reducing the effects of future climate change and increasing the economic lifespan of vineyards containing asymptomatically infected grapevines. 40 Figure 1.1. Illustration of the Threshold Model in grapevines, showing the relationship between the grapevine age and GTD fungal colonization density. Under normal conditions, fungal growth is slow, only reaching the colonization threshold in old grapevines during natural senescence. Under stress conditions, fungal growth is accelerated, and this threshold is reached rapidly in the grapevine lifespan, leading to disease progression and premature host death. This threshold is hypothesized to be reached sooner when multiple GTD fungi are present than when few fungi occur (adapted with permission from Sieber 2007). 41 Figure 1.2. Botryosphaeria dieback, Eutypa, Phomopsis dieback, and Esca symptoms. Botryosphaeria dieback symptoms on wine (A) and table grapevines (B) are characterized by dead spurs and cordons with no spring growth. Characteristic foliar symptoms of Eutypa dieback are shoots with short internodes and chlorotic and cupped leaves (C and D). Phomopsis dieback symptoms resemble those of Botryosphaeria dieback, with dead spurs and lack of spring growth (E). Vines affected by Botryosphaeria, Eutypa and/or Phomopsis dieback have perennial cankers in spurs, cordons and trunks, often with wedge shapes (F). Tiger-striped leaves (G) and characteristic soft yellowish wood rot (H) caused by Basidiomycete fungi are commonly associated with Esca disease. 42 43 Figure 1.3. Decline of young vines caused by Petri disease and/or black foot. Overall symptoms in the field include poor vigour, leaf chlorosis, short internodes (A and B) and eventual or sudden collapse I during the growing season. Internal wood necrosis observed at the basal end of the rootstock (D) commonly associated with black foot. Tylosis plugged xylem vessels and necrosis observed at a graft uniI(E) and trunk base, usually associated with Petri disease in young vines (F). Longitudinal section of a ready-to-plant nursery vine showing vascular necrosis originating from the basal end of the rootstock (G and H). Phaeomoniella chlamydospora, Phaeoacremonium spp., Ilyonectria spp., and some Botryosphaeriaceae spp. can be isolated from asymptomatic (I) and symptomatic (J) wood tissues from nursery propagated material, which reveals their potential latent phases in grapevines. 44 45 Figure 1.4. Abiotic and biotic stress factors hypothezised to contribute to grapevine trunk disease fungal development and symptom expression. A. Leaf scorching as a result of severe vine water stress. B and C. Leaves of young vines showing nutrient deficiency. D. Leaf deformities caused by herbicide damage. E. Severe over-cropping during the first years of vineyard establishment (courtesy of D. Gramaje, ICVV, Logroño, Spain). F. Winter damage and/or spring frost could further abiotic factors favouring disease development. G. J-rooting as a result of poor planting conditions. H. Low vigour vines caused by a nematode infestation. I. Ring nematode (Mesocriconema xenoplax) feeding on a root by introducing the stylet (black arrow). J. Close up of the ring nematode head. K. Leaf galls caused by erineum mite (Colomerus vitis). High levels of insect damage could increase vine stress. L. Grapevine crown gall caused by Agrobacterium vitis. M. Grapevine showing of grapevine leaf roll symptoms caused by Grapevine leafroll-associated virus 3. Grapevines affected by biotic stress factors (fungal, bacterial or virus diseases) may favour GTD fungal infections, wood colonization and symptom expression. 46 47 1.8 Research objectives Ready-to-plant nursery grapevines were tested for the most prevalent young vine decline fungi to determine the health status of nursery material sold and planted in Canada. A combination of greenhouse and field experiments were conducted to investigate the potential relationship between plant stress and disease development in grapevines infected with Petri disease fungi. This was addressed through four main questions: 1.8.1 Are young vine decline fungi present in asymptomatic grapevine nursery material planted in Canada and what is their abundance within individual grapevines? This question is addressed in chapter 2 of this dissertation. Ready-to-plant grapevines from four commercial nurseries were sent to the Summerland Research and Development Centre for analysis. Pathogen presence and absence and quantification with droplet digital PCR focused on the most prevalent young vine decline pathogens, including Cadophora luteo-olivacea, Phaeoacremonium minimum, and Phaeomoniella chlamydospora for Petri disease, Dactylonectria torresensis and Dactylonectria macrodidyma for black foot and Botryosphaeriacaea spp. for Botryosphaeria dieback. Samples were taken from four different sections of each grapevine to evaluate whether pathogen presence and abundance was consistent throughout the plant. Additionally, wood from each section was plated onto a nutrient medium to evaluate how traditional plating methodology compared to molecular techniques and to screen for other pathogens not selected for molecular identification. I expected young vine decline pathogens to be widely present in ready-to-plant nurseries with wide variability within and between plants for the abundance of each pathogen. 48 1.8.2 Does Phaeomoniella chlamydospora abundance in grapevines impact disease development in young vines? This question was addressed in chapters 3, 4, and 5 of this dissertation, which included two greenhouse experiments and four field experiments conducted at the Summerland Research and Development Centre. I inoculated dormant grapevines with different quantities of P. chlamydospora spores prior to planting the grapevines in the greenhouse (chapters 3 and 4), and in the field (chapters 3, 4, and 5) to understand how initial pathogen abundance impacts plant health each growing season. Each year I measured the aboveground growth of the plants. Upon conclusion of the experiment, I weighed the roots and scanned a cross-section of the trunk near the base to determine how different initial pathogen quantity affects internal trunk necrosis. I also extracted DNA from the wood near the base of each trunk and used droplet digital PCR to quantify the pathogen to understand how initial pathogen quantity affects pathogen abundance in the first few years after planting. I expected a higher initial pathogen inoculation quantity to negatively affect plant health and lead to greater pathogen abundance, greater necrosis, and disease development. 1.8.3 Do abiotic and biotic plant stress influence Phaeomoniella chlamydospora abundance and Petri disease symptom development in young grapevines? This question was addressed in chapters 3, 4, and 5 of this dissertation, which included two greenhouse and four field experiments. In chapter 3, I conducted a greenhouse and field experiment to determine the effect of water stress on P. chlamydospora abundance and disease development. In chapter 4, I conducted a greenhouse and field experiment to determine the effect of plant-parasitic ring nematodes on P. chlamydospora abundance and disease development. 49 Lastly, in chapter 5, I conducted two field experiments involving heat stress and j-rooting (forcing grapevine roots into an upward position at planting), to understand their effects on P. chlamydospora abundance and disease development. I monitored above-ground plant growth measurements to understand how stress and the pathogen separately and together affected the plant. Upon conclusion of the experiments, I quantified the pathogen at the base of each grapevine, measured internal trunk necrosis, and conducted above- and below-ground measurements of the plant to determine the effects of both stress and the pathogen on several plant health parameters. I expected plant stress to result in increased P. chlamydospora abundance resulting in increased internal necrosis and disease development. 1.8.4 Do arbuscular mycorrhizal fungi reduce plant stress and subsequent pathogen abundance and disease development in young grapevines? This question was addressed in chapters 3 and 4 of this dissertation in two separate greenhouse experiments. In chapter 3, half the plants from each treatment were inoculated with a commercial arbuscular mycorrhizal fungi biofertilizer to determine whether AM fungi could reduce the effect of water stress on the plant and subsequently reduce pathogen abundance compared to non-mycorrhizal plants. In chapter 4, the commercial mycorrhizal fungi were inoculated into half the plants of each treatment to determine whether the AM fungi could reduce the effects of ring nematodes on plant health and subsequently reduce P. chlamydospora abundance. AM fungal colonization of the roots was determined via molecular droplet digital PCR and by staining of the roots to visually confirm AM fungal structures within the roots. Phaeomoniella chlamydospora abundance was quantified with droplet digital PCR upon conclusion of the experiment to determine the effect AM fungi by itself and in combination with water stress or nematode infestation had on pathogen abundance. I expected AM fungi to reduce 50 the impact of ring nematodes and water stress on the plant, subsequently reducing P. chlamydospora abundance and disease development. 51 2 Health status of ready-to-plant grapevine nursery material in Canada regarding young vine decline fungi. 2.1 Background Young vine decline (YVD) occurs worldwide, resulting in poor vigor, lower yield, and premature death of grapevines, usually within five years after planting (Gramaje and Armengol 2011). The YVD complex is made up of Petri disease, caused primarily by the fungal pathogens Cadophora luteo-olivacea, Phaeoacremonium spp., and Phaeomoniella chlamydospora; black foot disease, caused by several species from the genera Campylocarpon, ‘Cylindrocarpon’, Cylindrocladiella, Dactylonectria, and Ilyonectria; and Botryosphaeria dieback, caused by several fungi from the Botryosphaeriaceae family (Agustí-Brisach and Armengol 2013; Gramaje and Armengol 2011; Úrbez-Torres 2011). Fungal species responsible for Botryosphaeria dieback and Petri disease can infect grapevines through pruning wounds, while fungi responsible for black foot disease are soil-borne and infect grapes via the roots or basal end of the rootstock or self-rooted cultivar. However, it has been well-demonstrated that many of these fungi can also be present in ready-to-plant nursery material (Agustí-Brisach and Armengol 2013; Gramaje and Armengol 2011; Gramaje et al. 2018). Infection by YVD fungi has been shown to occur at different stages during the nursery propagation process starting at the nursery mother blocks, where pruning wounds can facilitate the direct entry of fungal spores into plants (Gramaje and di Marco 2015; Waite et al. 2018). Nursery mother block surveys have found high incidences of black foot disease, Botryosphaeria dieback and Petri disease fungi present in plants as well as Botryosphaeria dieback and Petri disease fungi as aerial inoculum (Aroca et al. 2010; Halleen et al. 2020). In addition, several 52 practices have been identified as potential inoculum sources during the nursery propagation process, including the use of hydration tanks, cold storage of grapevines, grafting, and callusing, in which routine cuts along with poor or lack of sanitation practices may facilitate the infection and spread of YVD fungi (Gramaje and Armengol 2011; Gramaje and di Marco 2015). Sources of rootstock and scion material, as well as production practices, vary widely among nurseries and thus, a single cause for the entry and spread of YVD fungi in nurseries is very difficult to determine (Gramaje et al. 2018; Waite et al. 2013). Among all fungal pathogens isolated from nursery material in different countries, the most frequently identified include the black foot disease pathogens Dactylonectria macrodidyma and Dactylonectria torresensis, the Botryosphaeria dieback fungi Botryosphaeria dothidea, Diplodia seriata, Diplodia mutila, Neofusicoccum luteum, and Neofusicoccum parvum, and the Petri disease fungi C. luteo-olivacea, Phaeoacremonium minimum, and P. chlamydospora (Aigoun-Mouhous et al. 2019, 2021; Akgül et al. 2022; Berlanas et al. 2020; Billones-Baaijens et al. 2013c; Carbone et al. 2022; Dubrovsky and Fabritius 2007; Gramaje et al. 2009; Langenhoven et al. 2018; Maldonado-González et al. 2020; Pintos et al. 2018; Rego et al. 2000; Tosun and Yildiz 2022; Whitelaw-Weckert et al. 2013). Black foot disease fungi have been more commonly isolated from the roots or base of grapevines, Botryosphaeria dieback fungi have been primarily found in the graft-union, and Petri disease fungi have been commonly isolated from the base and graft-union. Nursery surveys have been conducted in different countries to better understand where in the nursery process infection may occur, and which YVD fungi are primarily found in nursery material. Infection can be widespread across nurseries. For instance, Akgül et al. (2022) reported 93% of nurseries in Türkiye having material infected with black foot disease pathogens. However, incidence of infection varied significantly between nurseries. In a study conducted 53 across 15 nurseries in Spain, asymptomatic infection of black foot fungi in ready-to-plant nursery material ranged from 5 to 90% of plants depending on the nursery (Berlanas et al. 2020). Similarly, incidence of Botryosphaeriaceae spp. in nursery material in New Zealand varied from 5 to 63% of plants infected, depending on the nursery (Billones-Baaijens et al. 2013c). Overall, high incidence of infection has been reported in nursery material with most available studies reporting up to 100% of plants tested yielding one or more YVD fungi (Carbone et al. 2022; Carlucci et al. 2017; Pintos et al. 2018; Whitelaw-Weckert et al. 2013). With such high levels of infection reported, it can be hypothesized that the presence of the fungus in the ready-to-plant nursery material may not directly imply the death of the plant or otherwise most of the vines would eventually die across vineyards worldwide. Young vine decline fungi are commonly isolated from grapevines showing vascular discoloration and/or necrosis in roots, rootstock, graft-union, and scion. However, they have also been reported from asymptomatic nursery material and from asymptomatic grapevines in vineyards, leading to the hypothesis that some of these fungi could act as latent pathogens (Hrycan et al. 2020; Waite et al. 2018). As such, some of these fungi could reside inside the plant as endophytes, causing no symptoms until changes in the host, most likely in the form of abiotic and/or biotic stress factors, may result in their transition from an endophytic to a pathogenic phase (Hrycan et al. 2020; Slippers and Wingfield 2007). Available studies reporting the incidence of YVD infection in nursery material have primarily focused on the presence or absence of the different fungi by conducting isolations from wood tissues. In addition, identification of the different fungi has primarily relied on traditional plating of the wood, and subsequent morphological and molecular identification from pure colonies. In this study, Droplet Digital™ PCR (ddPCR) assays were developed to determine the 54 absolute abundance of YVD fungi. To date, very few studies have focused on determining the absolute fungal abundance in grapevine nursery material in regards to YVD fungi (Billones-Baaijens et al. 2013c; Lade et al. 2022; Maldonado-González et al. 2020). Determining fungal abundance within grapevines is an important step in understanding the health status of nursery material, as it may have implications on plant survivability when planted in vineyards. The Canadian grape and wine industry is a relatively young industry and its growth relies on the import of nursery material from certified and accredited nurseries. The health status of nursery material planted in Canadian vineyards is currently unknown with regard to YVD fungal presence and abundance. The main objective of this study was to quantify the fungal abundance of the most important YVD fungi from different nurseries selling plants in Canada using ddPCR. 2.2 Materials and Methods 2.2.1 Plant material and sample preparation The health status of dormant ready-to-plant Vitis vinifera L. cultivars sold in Canada was evaluated from four nurseries. In total, 30 plants per nursery were tested, 10 each of ‘Chardonnay’, ‘Merlot’, and ‘Pinot noir’. Vines from nurseries A2014, A2019, C and D were grafted onto ‘3309 Couderc’ (‘3309C’) rootstock, while those from nursery B consisted of self-rooted plants. Grapevines were collected and analyzed from nursery A in 2014 and in 2019; nursery B in 2014, and nurseries C and D in 2021. In nurseries A2014, A2019, C, and D, samples were collected from the roots, basal end of the rootstock, graft-union, and scion. In nursery B samples were taken from the roots, basal end of the scion, and scion. In total, 150 plants were collected from all nurseries and four samples were tested from the roots, base, graft union, and scion, with the exception of nursery B which was self-rooted for a total of 570 samples analyzed. 55 Additionally, the basal end and roots of five of each ‘Riparia Gloire,’ ‘SO4,’ ‘101-14,’ and ‘161-49’ rootstocks and self-rooted ‘Baco noir,’ were analyzed from nursery A2014. The roots were removed from the basal end of each grapevine. A random selection of root segments of approximately 0.5 g was taken from each plant. Roots were placed in 70% ethanol for 30 s, 10% bleach for 1 min, and sterile water for 2 x 1 min washes. Half of the root selection was finely sliced with a razor blade and 0.2 g were placed in Qiagen DNeasy PowerSoil DNA extraction tubes (Qiagen, Hilden, Germany) containing one ¼” ceramic sphere (MP Biomedicals, Irvine, CA) above, and one below plant material. Tubes were stored at -20°C until DNA was extracted. The remaining half of the root sample was plated in duplicate on potato dextrose agar (PDA, HiMedia Laboratories, India) amended with 0.1 mg/mL tetracycline hydrochloride (PDA-tet, Sigma-Aldrich, St Louis, MO) and incubated without light at 20°C until fungal colonies were observed. The basal end of each grapevine was removed at 1 cm from the base and placed in storage at 4°C without light in clear plastic bags for analysis of the necrosis. The bark from 1-2 cm of wood from the basal end, graft-union, and scion was shaved off and discarded. Segments of 1 cm in length were cut from the basal end, graft-union, and scion and split in half longitudinally. One half was surface sterilized via flame sterilization with 95% ethanol and shavings were plated on PDA-tet Petri plates in duplicates and incubated without light at 20°C until fungal colonies were observed. The other half of each sample was finely cut and 0.2 g of each sample was placed in Qiagen DNeasy PowerSoil DNA extraction tubes as previously described. 2.2.2 Morphological and molecular characterization of isolated fungi 56 Morphological identification to the genus level was conducted on the fungal colonies grown from each plant section on PDA-tet as described in Úrbez-Torres et al. (2014a, b). Representative colonies from morphologically similar fungi were selected and transferred to fresh PDA. Pure colonies were obtained from hyphal tips from actively growing colonies. The DNA for sequencing was extracted from pure colonies using the Qiagen DNeasy PowerSoil kit according to the instructions of the manufacturer. Oligonucleotide primers ITS1/ITS4 (White et al. 1990), EF1-728F/EF1-986R (Carbone and Kohn 1999), T1/Bt2b, Bt2a/Bt2b (Glass and Donaldson 1995), and EF1α/EF2α (O’Donnell et al. 1998) were used to confirm the identity of the isolated fungi depending on the preliminary morphological identification to genus level (Gramaje et al. 2019). Sequencing and analyses were completed as described in Úrbez-Torres et al. (2014a, b). Consensus sequences were compared with those available in the GenBank database by using the Basic Local Alignment Search Tool (BLAST) to identify the different fungi to species level. DNA sequences were deposited in GenBank. 2.2.3 Symptom analyses The axial cross sections from wood 1 cm above the basal end of rootstock or self-rooted scion of each grapevine were scanned using an Epson Expression 1689 scanner set to 600 dpi, 48 x 24 color, reflective positive using Adobe Photoshop CS2 Version 9.0.2. Total necrosis was measured using the Fiji ImageJ version 1.53t software program with distance in pixels set to 23.62 mm (Schindelin et al. 2012). Total percent necrosis was measured as 𝑁𝑒𝑐𝑟𝑜𝑠𝑖𝑠−𝑃𝑖𝑡ℎ𝑇𝑜𝑡𝑎𝑙 𝑠𝑢𝑟𝑓𝑎𝑐𝑒 𝑎𝑟𝑒𝑎−𝑃𝑖𝑡ℎ∗ 100. 2.2.4 DNA extraction and absolute fungal quantification by Droplet Digital™ PCR 57 Plant material collected in Qiagen DNeasy PowerSoil DNA extraction tubes was macerated in a FastPrep-24 High-Speed Benchtop Homogenizer (MP Biomedicals, Irvine, CA) at 6 m/s for 40 s twice, followed by DNA extraction according to the Qiagen manufacturer’s protocol. The DNA concentrations were determined using a Qubit 1.0 Flurometer with the dsDNA HS Qubit assay (Thermofisher, Waltham, MA). Fungal quantification for six pathogens, including Botrysophaeriaceae spp. (Bot), C. luteo-olivacea (Clo), D. macrodidyma (Dm), D. torresensis (Dt), P. minimum (Pm), and P. chlamydospora (Pc) was conducted using a Droplet Digital™ PCR (ddPCR) system (Bio-Rad Laboratory, Hercules, CA). Primers and probes for each targeted fungus are listed in Table S2.1. Three of the assays (Bot, Clo, Pm) were adapted directly from previously published studies (Billones-Baaijens et al. 2018; Maldonado-González et al. 2020; Martin et al. 2012). For the Pc assay, primers published by Martín et al. (2012) were used, with the addition of a probe designed in the region between the published primers by using the IDT PrimerQuest™ tool (DNA Technologies, Inc.) (Table S2.1). Assays for Dm and Dt were developed specifically for this study. Initially, the complete histone 3 (H3) gene sequences of 16 Dm species from locations worldwide (Berlanas et al. 2020; Cabral et al. 2012) were retrieved from GenBank and aligned using Clustal W (Larkin et al. 2007). Four different H3 sequences were identified and further aligned with the type species from Dactylonectria alcaracerensis (CBS 129087), Dactylonectria novozelandica (CBS 112608), and D. torresensis (CBS 129086). Primers and probes were identified from a 220 bp region of the H3 gene using IDT PrimerQuest™. A highly variable section within this region was used for selection of the probe, as it was able to differentiate Dm from closely related species. A second probe from this variable region was selected to specifically identify Dt (Table S2.1). For all assays, primers/probes were tested and optimized in ddPCR for determination/confirmation of 58 annealing temperatures and specificity. A temperature gradient from 55°C to 65°C was used to test the ddPCR conditions using DNA obtained from pure fungal spores of the target species. The final annealing/extension temperatures are indicated in Table S2.1. The assays were validated for specificity using a series of closely related and unrelated fungal species from the Fungal Collection at the Summerland Research and Development Centre (SuRDC). Results were consistent with the positive and negative species assignments that had been determined through prior sequencing and plating assessments of these collection samples. 2.2.5 ddPCR reactions and conditions. Droplet digital PCR reactions were prepared according to the protocols for primer or probe assays (Bio-Rad). Briefly, ddPCR “probe” reactions (Bot, Clo, Dm, Dt, Pc) contained 11 ul 2 x ddPCR™ Supermix for probes (Bio-Rad Laboratory, Hercules, CA), 900 nM of each forward and reverse primer, 250 nM of probe and 5.5 ul DNA in a final volume of 22 ul. Droplet generation was conducted using a Bio-Rad QX200™ Automated Droplet Generator (Bio-Rad), sealed with foil using a PX1 PCR Plate Sealer (Bio-Rad), and loaded into a C1000 Touch Thermal Cycler (Bio-Rad). PCR conditions were as follows: initial denaturation at 95oC for 10 min; 40 cycles of 94oC for 30 s, followed by the specific annealing/extension temperature (Table S2.1) for 1 min; an enzyme deactivation step of 98oC for 10 min; and a 4oC hold. Ramp rate was 2°C/s. The ddPCR “primer” reactions (Pm) were similar, but contained 2 x EvaGreen Supermix for primers (Bio-Rad), 150 nM of each primer, with the following PCR conditions: initial denaturation at 95oC for 5 min; 40 cycles of 95oC for 30 s, followed by the specific annealing/extension temperature for 1 min; a signal stabilization step of 4oC for 5 min then 90oC for 5 min; and a 4oC hold. Ramp rate was 2 oC/s. 59 Droplets were read in a Bio-Rad QX200™ Droplet Reader and analyses conducted using QuantaSoft software version 1.7.4 (Bio-Rad). Droplet counts were examined to ensure all samples were above 10,000 total droplets and the amplitude threshold was set in order to separate positive from negative droplets. Samples were identified as positive if >1 positive droplet occurred. Results were normalized to DNA concentration input and reported as gene copies/ng DNA. 2.2.6 Statistical Analysis Data analyses were conducted to determine whether there were differences in fungal abundance for each pathogen between different sections of each grapevine, differences in fungal abundance between cultivars within each nursery, and differences in necrosis between each cultivar in each nursery. Data from fungal quantification and symptom analyses were analyzed by Kruskal-Wallis rank sum test and Dunn’s test of multiple comparisons adjusted with the Benjamini-Hochberg method with a p value set at 0.05 with the dunn.test package (Dinno 2017). Correlation of total fungal abundance at the basal end of each grapevine and percent necrosis was analyzed via Spearman’s rank correlation coefficient in R Studio version 4.0.4 (R Core Team 2021). 2.3 Results 2.3.1 Morphological and molecular characterization of isolated fungi Sequencing results revealed YVD fungi that were not quantified in this study by ddPCR, including the black foot fungi ‘Cylindrocarpon’ sp., Dactylonectria pauciseptata, Ilyonectria crassa, Ilyonectria destructans, and Ilyonectria robusta, and the Petri disease fungi Cadophora malorum, Phaeoacremonium fraxinopennsylvanica, Phaeoacremonium iranianum, and 60 Pleurostoma richardsiae (Table S2.2). While the general Botryosphaeriaceae spp. ddPCR assay was used for quantification; the following Botryosphaeria dieback fungi were identified through sequencing: Botryosphaeria dothidea, Macrophomina phaseolina, Neofusicoccum mediterraneum, and Neofusicoccum parvum (Table S2.2). Other grapevine pathogenic fungi identified included Diaporthe ampelina and Acremonium sp. (Table S2.2). Based on the isolation results, 95 out of a total of 150 vines (63%), and 145 out of a total of 570 samples (25%) were positive for at least one fungal pathogen known to be associated with YVD. Isolation results showed 74 out of 150 vines (49%) and 104 out of 570 samples (18%) positive for at least one of the fungal species quantified using ddPCR. Droplet digital PCR results showed 149 out of 150 vines (99%) and 497 out of 570 samples (87%) were positive for at least one of the fungal species studied. 2.3.2 Symptom analyses Percent necrosis measured at the base of the rootstock or self-rooted cultivar was lowest in nursery B and highest in nurseries A2014, A2019, C, and D, which were not significantly different from each other. Percent necrosis was not significantly different between cultivars within nurseries A2014, A2019, C, and D. In nursery B, ‘Pinot noir’ percent necrosis was higher compared to ‘Chardonnay’ (Table 2.1). Percent necrosis varied widely within nurseries A2014 (26 to 78%), A2019 (27 to 72%), B (10 to 46%), C (26 to 66%), and D (31 to 74%). Correlation between percent necrosis and total fungal abundance at the basal end was not significant within nursery A2014 (p=0.5383), A2019 (p=0.6849), B (p=0.1413), C (p=0.8319), and D (p=0.8005) (Figure 2.1). Within individual cultivars in nursery B, no significant correlations were found between percent necrosis and total fungal abundance at the basal end for self-rooted ‘Chardonnay’ (p=0.5538), ‘Merlot’ (p=0.4483), or ‘Pinot noir’ (p=0.1413). 61 Percent necrosis did not differ between the different rootstocks in nursery A2014 (p=0.2084). Percent necrosis varied widely within ‘Riparia Gloire’ (29 to 55%), self-rooted ‘Baco noir’ (25 to 53%), ‘SO4’ (36 to 65%), ‘101-14’ (35 to 62%), and ‘161-49’ (29 to 39%). However, total fungal abundance at the basal end of each rootstock was not correlated with percent necrosis in ‘Riparia Gloire’ (p=0.35), ‘161-49’ (p=0.3217), ‘SO4’ (p=0.08333), ‘101-14’ (p=0.2333), or self-rooted ‘Baco noir’ (p=0.95). 2.3.3 Absolute fungal quantification by ddPCR Fungal abundance varied significantly between nurseries for each fungal species tested (Tables 2.2). Across nurseries and cultivars, 7 to 87% of plants were positive for Botryosphaeriaceae spp., 93 to 100% for C. luteo-olivacea, 0 to 40% for D. macrodidyma, 0 to 97% for D. torresensis, 33 to 100% for P. minimum, and 3 to 80% for P. chlamydospora (Table 2.2). All plants in nurseries A2014, A2019, B and D, and 97% of plants in nursery C were infected with at least 1 YVD fungi (149 out of 150 plants, Table 2.2). A mean of three YVD fungal species were identified in all grapevines from all nurseries. Overall, fungal abundance varied significantly between infected plants within and between nurseries (Table 2.2). Nursery A2014 had the largest variation in abundance among individual grapevines; ranging from 17.1 to 48,829.7 copies/ng DNA. Nursery A2014 also showed the highest median level of infection with 1,409.6 copies/ng DNA, while nursery B had the lowest median level of infection with 32.5 copies/ng DNA (Table 2.2). Botryosphaeriaceae spp. were present in all nurseries sampled. Incidence of plants infected with Botryosphaeriaceae spp. was highest in nursery A2014 (87%) and lowest in nursery A2019 (7%) (Table 2.2). Plants from nursery A2014 contained the highest 62 Botryosphaeriaceae spp. mean abundance, and plants from nursery A2019 had the lowest mean abundance. Plant material from nurseries B, C, and D contained low Botryosphaeriaceae spp. mean abundances. Total Botryosphaeriaceae abundance varied between cultivars within nurseries A2014, B, C, and D and was significantly higher in ‘Merlot’ compared to ‘Pinot noir’ in nursery B, in ‘Merlot’ compared to all cultivars in nurseries A2014 and C, and in ‘Pinot noir’ compared to all cultivars in nursery D (Table 2.3). Botryosphaeriaceae abundance varied significantly among individual plants from each nursery and they were present in all plant sections in nurseries A2014 and B, but primarily in the graft union and scion in nurseries A2019, C, and D (Table 2.3). Cadophora luteo-olivacea was the most ubiquitous of all fungi tested and was present in 97% of plants tested (Table 2.2). Incidence of plants infected was highest in nurseries A2014 and A2019 (100%) and lowest in nurseries B and C (93%) (Table 2.2). Mean C. luteo-olivacea abundance was highest in nursery C and lowest in nursery A2019 (Table 2.4). Total C. luteo-olivacea abundance varied between cultivars within nurseries and was greater in ‘Pinot noir’ than ‘Chardonnay’ in nursery A2014, was greatest in ‘Chardonnay’ in nursery A2019, and was greatest in ‘Pinot noir’ in nursery C (Table 2.4). Cadophora luteo-olivacea abundance varied significantly among individual plants across all nurseries and the fungus was present in all plant sections from all nurseries in variable abundances (Table 2.4). Dactylonectria macrodidyma was the least abundant species among all tested with a 13% incidence of plants infected (Table 2.2). Incidence of plants infected with D. macrodidyma was highest in nursery B (40%) and lowest in nursery A2019 (0%, Table 2.2). The fungus was present in a low quantity only in ‘Merlot’ in nurseries A2014 and C, and only in ‘Chardonnay’ in nursery D (Table 2.5). There were no significant differences in fungal abundance between cultivars for 63 nurseries A2014, B, or D (Table 2.5). Fungal abundance varied significantly among individual plants in nurseries A2014, B, C, and D and the fungus was only present in the roots or base in nurseries A2014, B, C, and D (Table 2.5). Dactylonectria torresensis was present in all nurseries except in nursery D (Table 2.2). Incidence of plants infected with D. torresensis was highest in nursery A2014 (97%, Table 2.2). Dactylonectria torresensis abundance in nursery A2014 was significantly higher compared to A2019 (Table 2.6). Total fungal abundance was lower in ‘Merlot’ compared to ‘Pinot noir’ in nursery A2014, and ‘Chardonnay’ compared to ‘Merlot’ and ‘Pinot noir’ in nursery A2019, but no significant difference between cultivars was observed in nurseries B or C (Table 2.6). Fungal abundance varied significantly among individual plants across nurseries A-C, and the fungus was primarily present in the roots and base (Table 2.6). Phaeoacremonium minimum was present in all nurseries sampled. Incidence of plants infected with P. minimum was highest in nursery D (100%) and lowest in nursery A2019 (33%, Table 2.2). The fungus was present in low numbers in nurseries A-C (Table 2.7). Total fungal abundance was not significantly different between cultivars for all nurseries. Phaeoacremonium minimum abundance varied significantly among individual plants within each nursery and cultivar and the fungus was present in all sections in variable abundance (Table 2.7). Phaeomoniella chlamydospora was present in all nurseries sampled. Incidence of plants infected with P. chlamydospora was highest in nursery A2019 (80%) and lowest in nursery B (3%) (Table 2.2). Fungal abundance was high in both nurseries A2014 and A2019, but was significantly higher in 2014 (Table 2.8). Phaeomoniella chlamydospora abundance was higher in ‘Pinot noir’ compared to ‘Chardonnay’ in nursery A2014, and highest in ‘Chardonnay’ in nursery 64 A2019 compared to the other cultivars. No significant differences among cultivars were observed in nurseries B, C, and D. There was significant variability in fungal abundance between individual plants for all nurseries. The fungus was present in variable abundance in all plant sections, but was not present in ‘Merlot’ roots in nursery A2014 or in any roots in nursery A2019, B, or C (Table 2.8). 2.3.4 Fungal abundance in different rootstocks Analysis of rootstock material obtained from nursery A2014 revealed significant variation in the incidence of rootstocks infected for all fungal species tested. Botryosphaeriaceae spp. abundance was higher in ‘101-14’ compared to ‘Baco noir’ and ‘Riparia Gloire’, but was not statistically different from ‘SO4’ or ‘161-49’ (Table 2.9). Cadophora luteo-olivacea abundance was highest in ‘101-14’ but was not statistically significant from ‘Riparia Gloire’ (Table 2.9). Phaeomoniella chlamydospora was present in high abundances in self-rooted ‘Baco noir’, ‘SO4’ and ‘101-14’, and was not present in ‘Riparia Gloire’ and ‘161-49’. Dactylonectria macrodidyma was not present in the rootstock material tested and D. torresensis fungal abundance was not statistically different between rootstocks (Table 2.9). Phaeoacremonium minimum had the highest abundance in self-rooted ‘Baco noir’ and the fungus was not present in ‘Riparia Gloire’ (Table 2.9). 2.4 Discussion This study represents the first attempt to determine the health status of ready-to-plant nursery material planted in Canada in regards to YVD fungi. In addition, this work is the first to use ddPCR for identification and absolute quantification of a wide range of taxonomically different fungi associated with YVD in ready-to-plant nursery material. 65 Sequencing of fungi isolated from grapevine wood by the traditional plating method revealed the presence of YVD fungi that had not been previously identified in Canada, including the black foot disease pathogens ‘Cylindrocarpon’ sp., Ilyonectria crassa, and Ilyonectria destructans, the Petri disease fungi Cadophora malorum and Pleurostoma richardsiae and the Botryosphaeriaceae spp. Macrophomina phaseolina and Neofusicoccum mediterraneum, supporting that nursery material is a source of new invasive pathogens for young vineyards (Aroca et al. 2010; Berlanas et al. 2020; Billones-Baaijens et al. 2013c; Carbone et al. 2022; Carlucci et al. 2017; Langenhoven et al. 2018; Úrbez-Torres et al. 2015). Additionally, sequencing confirmed the presence of fungi not quantified by ddPCR, revealing that this study does not present the full extent of fungal presence in nursery material planted in Canada. Prior to this study, identification of YVD associated fungi from plant material has primarily relied on their isolation from the wood, and subsequent morphological and molecular characterization (Akgül et al. 2022; Berlanas et al. 2020; Billones-Baaijens et al. 2013c; Carbone et al. 2022; Carlucci et al. 2017; Giménez-Jaime et al. 2006; Langenhoven et al. 2018; Pintos et al. 2018), and only recently molecular methods for the direct identification of YVD fungi from nursery material have been introduced (Maldonado-González et al. 2020; Úrbez-Torres et al. 2015). This study found that 18% of samples obtained through traditional plating harbored at least one of the fungal species targeted, while 87% of samples were positive using ddPCR. These findings indicate that traditional plating methods are not sufficient to determine the extent of infection in plant material. Traditional plating and sequencing from pure cultures requires several time-consuming steps. Sequencing of YVD fungi also requires more than one loci to discriminate among species, further lengthening the process (Aigoun-Mouhous et al. 2021; Aroca et al 2010; Berlanas et al. 2020; Gramaje et al. 2019; Úrbez-Torres et al. 2014a, b). Due to 66 the skills and time required, the process to screen for YVD fungi is difficult for nurseries to implement. The use of ddPCR provides nurseries with more accurate information to determine the health status of material, both in terms of identification and quantification of the pathogens. This information can be used by nurseries to identify main sources of infection and implement proper sanitation and/or management strategies. Absolute quantification using ddPCR showed that 99% of plants tested in this study were infected with at least one YVD fungi. The implications of this finding are significant for the Canadian grape and wine industry, which is reliant on healthy nursery material for grapevine replacement and industry growth. This study suggests that 100% of ready-to-plant nursery material planted in Canada may be infected with at least one YVD fungi. However, only 7.8% of young grapevines displayed symptoms of YVD in a recent survey conducted in British Columbia, Canada, which is a relatively small percentage compared to a possible 100% infection rate of ready-to-plant nursery material. A high degree of variability in the number of YVD symptomatic plants among blocks in British Columbia was identified, ranging from 5% to 54% incidence (Úrbez-Torres et al. 2014b). As not every grapevine that is infected with YVD fungi is symptomatic, it has been hypothesized that some of these fungi may be latent pathogens, living as endophytes in the grapevine until triggered to transition to a pathogenic state (Hrycan et al. 2020). The trigger for this transition is thought to be plant stress, which may explain the disparity between the pathogen abundance observed in this study, and symptomatic grapevines reported in young vineyards (Brown et al. 2012; Slippers and Wingfield 2007; van Niekerk et al. 2011c). A second potential explanation for the comparatively low incidence of YVD symptomatic plants in vineyards compared to the high percent infection of nursery material is the abundance of fungi within each individual ready-to-plant grapevine. This study revealed significant 67 variability in fungal abundance among individual plants within all nurseries sampled. It is unclear what effect, if any, a high initial abundance of YVD fungi may have on the health of young grapevines when planted; however, fungal abundance has been correlated with symptom expression in other patho-systems such as in Douglas-fir Swiss Needle Cast Disease, caused by Phaeocryptopus gaeumannii (Manter et al. 2003). Similarly, some susceptible soybean cultivars had higher Phialophora gregata densities and expressed greater brown stem rot symptom severity than resistant cultivars which contained lower fungal abundance and lesser symptom severity (Impullitti et al. 2009). It has been hypothesized that a threshold is required for the transition from an endophytic to a pathogenic phase (Sieber 2007). A higher initial YVD fungal abundance may reach this threshold sooner, leading to premature death (Hrycan et al. 2020). An alternative hypothesis is that the plant cannot contain these YVD fungi when fungal abundance is too great initially, leading to YVD symptoms and eventual death of the plant regardless of plant stress. The underlying reason for the significant variability in fungal abundance among individual plants is unclear. Infection can occur at several stages of the propagation process, starting from the mother block to infection during their first year of growth (Aroca et al. 2010; Gramaje and Armengol 2011; Gramaje and di Marco 2015; Halleen et al. 2020). Infection in the mother block or at an earlier stage in the propagation process may result in an increased fungal abundance as conditions in the nursery propagation process are ideal for fungal growth while the grapevine is dormant, rooting or growing. Infection later on in the propagation process may result in a lower fungal abundance as there is less time for the fungus to grow. Alternatively, variations in the microbiome of individual plants may impact YVD fungal abundance as some fungi and/or bacteria are antagonistic to various YVD fungi (Álvarez-Pérez et al. 2017; Berbegal 68 et al. 2020; Pertot et al. 2016; van Jaarsveld et al. 2021). However, these factors do not completely account for variations in YVD fungal abundance between cultivars within nurseries or between nurseries. The phytosanitary practices of individual nurseries may play a significant role in fungal growth in different sample sets of plants within nurseries and between nurseries (Waite et al. 2013). For example, variations or improvement in phytosanitary practices and control strategies in the mother blocks and nursery may explain the fungal abundance differences observed between nursery A2014 and A2019. Plant material may have also originated from different mother blocks in 2014 and 2019. Presence and abundance of individual YVD fungal species was highly variable between nurseries. Our findings revealed a high degree of variability in the number of plants infected with certain YVD fungal species. In support of previous findings, variation in percent infection among individual plants for Botryosphaeriaceae spp. (Billones-Baaijens et al. 2013c) and black foot fungi (Berlanas et al. 2020) was observed. Incidence of Botryosphaeriaceae spp. in young vines in New Zealand ranged from 5% to 63% of all plants in individual nurseries with an average incidence of 23% (Billones-Baaijens et al. 2013c). This study revealed a range of 7% to 87% in incidence for individual nurseries with an average incidence of 36%, which is higher, but generally similar with those reported from New Zealand. The higher incidence observed in this study may be a result of increased detection sensitivity of ddPCR compared to traditional plating methods and differences in source material for nurseries. In Spanish nurseries, D. torresensis accounted for 75% of all fungi identified from grafted plants, followed by 4% for D. macrodidyma, while the overall incidence of black foot fungi ranged from 5-90% among nursery fields sampled (Berlanas et al. 2020). Our findings revealed 0-40% incidence of D. macrodidyma and 0-97% incidence of D. torresensis among 69 nurseries. Planting of material in in-field nurseries prior to selling to commercial vineyards may play a significant role in the increased incidence of black foot fungi within plant material. Dactylonectria macrodidyma was observed solely in the roots and basal ends of ready-to-plant nursery material; however, D. torresensis was observed in all plant sections, indicating infection may occur in the mother blocks or during the nursery propagation process (Gramaje and Armengol 2011). Petri disease fungi were found in high abundance in some nursery material, but incidence in individual nurseries varied. Our study revealed a high incidence and abundance of C. luteo-olivacea, P. minimum, and P. chlamydospora in accordance with previous findings (Aigoun-Mouhous et al. 2021; Dubrovsky and Fabritius 2007; Gramaje et al. 2009; Maldonado-González et al. 2020; Rego et al. 2000; Whitelaw-Weckert et al. 2013). We found C. luteo-olivacea in the majority of nursery material from all nurseries and in high abundances, contrary to the findings of Maldonado-González et al. (2020), who found 80% of plants contained C. luteo-olivacea, but only 25% of samples contained more than three copy numbers of the targeted pathogen in nursery material from Spain. Incidence of nursery material for P. minimum and P. chlamydospora varied significantly between individual nurseries in which the infected planting material ranged from 3-80% and 33-100% between nurseries. The presence or absence of specific YVD fungi may be due to their presence or absence in the mother block, where incidence of YVD fungi may be high or low (Aroca et al. 2010; Halleen et al. 2020). As mother blocks vary in location and age, the health status of mother blocks may play an important role in the subsequent infection of nursery material. Variations in the presence or absence and fungal abundance may also be related to whether the same mother blocks are continuously used or the use varies between blocks. 70 Variability in YVD fungal abundance was significant between individual plants of each sample set, but also within individual plant sections. Fungal presence has previously been indicated to be higher in sections of the plant that are wounded during the propagation process, such as the graft union and base (Lade et al. 2022; Pintos et al. 2018; Rego et al. 2000; Whitelaw-Weckert et al. 2013). Results from this study revealed significant variability among individual plants, but no clear distinction for fungal abundance between sections of grapevines, with the exception of black foot fungi that were more prominent in the roots and base. Black foot disease fungi may occur more prominently in the roots and basal end as high-density planting of nursery plant material in the first year is a potential source of infection (Gramaje and Armengol 2011). Nursery A2014 rootstock material fungal abundance was comparable to cultivar fungal abundance. However, a key difference was observed with regard to P. chlamydospora abundance since it was not present in ‘Riparia Gloire’ and ‘161-49’ rootstocks, despite high fungal abundance in the nursery. Cadophora luteo-olivacea abundance was significantly higher in ‘101-14’ compared to ‘161-49’, ‘SO4’, and self-rooted ‘Baco noir’. This may be due to differences in phytosanitary practices between mother blocks and at what stage in the nursery propagation process infection occurs. The microbiomes of different rootstocks can vary which may also influence the presence or absence of certain YVD fungi, in which microbial communities differed between 18 to 44% among rootstocks (Gramaje et al. 2022; Lade et al. 2022). Fungal microbiota associated with the potential biocontrol of YVD fungi have been identified and variations in the abundance or composition of these communities may impact the presence and fungal abundance of YVD fungi within certain plants and between rootstocks (Gramaje et al. 2022). Rootstock xylem anatomy may also impact YVD fungal abundance. Smaller rootstock 71 xylem diameter has been associated with a lower abundance of P. minimum; however, xylem diameter is unlikely to prevent infection (Ramsing et al. 2021). Necrosis at the basal end of nursery material has commonly been thought an indicator of YVD infection, although it was reported recently that necrosis was correlated to individual nurseries and rootstocks (Carbone et al. 2022; Dubrovsky and Fabritius 2007; Gramaje and Armengol 2011; Lade et al. 2022). Percent necrosis at the basal end of grapevines was consistent across cultivars within nurseries except in nursery B where ‘Pinot noir’ percent necrosis was higher than ‘Chardonnay’. The mean sum of fungal abundance at the basal end of each grapevine for all YVD fungi tested did not correlate with percent necrosis. The consistent level of necrosis within individual nurseries indicates that nursery practices may play a larger role in initiating internal necrosis than YVD fungi in the early stages of infection. Necrosis in self-rooted plant material from nursery B was significantly lower compared to nurseries A2014, A2019, C, and D, where nursery material was grafted. Percent necrosis was not statistically different among grafted material between nurseries, further indicating plant source material may play a significant role in initial necrosis development. The reasoning for this lower percent necrosis in self-rooted plant material from nursery B is unclear; however, rootstock material can be difficult to callus and root (Wurz et al. 2022; Yin et al. 2023). Poor callusing and rooting may leave the basal end open to abiotic and biotic factors which may impact the level of necrosis at the basal end of plant material and could account for variations in percent necrosis among plants within nurseries. This research reveals that ready-to-plant nursery material is a source of new pathogen entry into young vineyards and that YVD fungi are prevalent in nursery material planted in Canada. Young vine decline fungal incidence and abundance varied between sections of individual grapevines, individual grapevines, cultivars, and nurseries. Based on these findings, if 72 presence of the fungi resulted in disease and eventual death in all cases, viticulture in Canada would not exist in its current form. Investigation into the potential reasons for this variability in fungal abundance among plant material may reveal why some plants are symptomatic in the first few years after planting, while others are not. Once planted in a vineyard, a high initial YVD fungal abundance in ready-to-plant nursery material may impact grapevine health, as it may lead to plant mortality in the first few years after planting. Plant stressors may also impact fungal growth, leading to symptomatic infection and plant mortality in young grapevines; thus, future studies should evaluate the effects of different thresholds of fungal abundance on plant health, and the effect that stress may play in favoring disease development. Nursery material is also likely to be infected with several YVD fungi at one time and the effect of co-infection of several taxonomically different fungi on plant health should be further investigated. 73 Table 2.1. Mean sum of infection (copies/ng DNA) of all fungal species in the basal end of each grapevine and percent necrosis at the base of each grapevine for each nursery individually. Chardonnay Merlot Pinot noir Overallz Nurseryy copies/ng DNA Necrosis copies/ng DNA Necrosis copies/ng DNA Necrosis copies/ng DNA Necrosis A2014 88.6 48.7% a 96.9 47.3% a 2,293.2 51.4% a 2478.7 49.1% A A2019 197.7 49.5% a 31.9 48.7% a 40.5 48.0% a 270.2 48.7% A B 50.6 19.5% a 21.9 23.9% ab 12.9 31.9% b 85.5 25.1% B C 0.1 46.1% a 231.3 49.3% a 83.3 41.2% a 314.4 45.5% A D 98.3 43.2% a 75.6 45.1% a 42.6 51.4% a 216.5 46.6% A y For each row within nurseries, means with the same lowercase letter are not significantly different (P = 0.05) according to Dunn's test z For the overall column, means between nurseries with the same uppercase letter are not significantly different (P = 0.05) according to Dunn’s test. 74 Table 2.2. Maximum, median, and minimum level of infection (copies/ng DNA) of each individual fungal species tested and all fungal species combined in infected grapevines and percent infection of all grapevines in each nursery individually and all nurseries combineda. Nursery Max-min % Bot Bot % Clo Clo % Dm Dm % Dt Dt % Pm Pm % Pc Pc % Total Total A2014 Maximum 87% 711.8 100% 4,555.9 3% 5.19 97% 306.7 53% 11.6 70% 48,161.6 100% 48,829.7 Median 64.7 353.8 5.19 23.9 1.3 1401.5 1,409.6 Minimum 1.5 5.5 5.19 2.6 0.5 0.5 17.1 A2019 Maximum 7% 1.6 100% 303.5 0% 0.00 87% 14.5 33% 121.9 80% 2,252.8 100% 2,371.8 Median 1.6 37.3 0.00 3.6 1.7 22.6 89.0 Minimum 1.6 3.6 0.00 0.9 0.3 1.0 7.2 B Maximum 40% 16.9 93% 13,028.6 40% 16.83 63% 52.0 37% 6.2 3% 3.5 100% 13,036.1 Median 4.2 35.5 2.53 4.7 0.6 3.5 32.5 Minimum 0.8 0.7 0.90 0.5 0.1 3.5 1.9 C Maximum 20% 141.3 93% 6,525.8 20% 5.13 43% 7.4 83% 134.0 17% 394.3 97% 6,531.1 Median 78.2 110.6 3.80 0.6 0.6 0.9 193.3 Minimum 18.1 0.5 0.50 0.1 0.1 0.5 0.5 D Maximum 27% 73.1 97% 3,220.6 3% 3.94 0% 0.0 100% 2,822.3 7% 0.6 100% 3,522.3 Median 0.7 102.5 3.94 0.0 73.2 0.5 280.8 Minimum 0.3 17.5 3.94 0.0 0.4 0.4 31.4 Total Maximum 36% 711.8 97% 13,028.6 13% 16.83 58% 306.7 61% 2,822.3 35% 48,161.6 99% 48,829.7 Median 10.2 97.0 3.62 6.1 2.4 64.8 208.4 Minimum 0.3 0.5 0.50 0.1 0.1 0.4 0.5 a Bot = Botryosphaeriaceae spp., Clo = Cadophora luteo-olivacea, Dm = Dactylonectria macrodidyma, Dt = Dactylonectria torresensis, Pm = Phaeoacremonium minimum, Pc = Phaeomoniella chlamydospora. 75 Table 2.3. Mean copies/ng DNA and standard deviation of Botryosphaeriaceae spp. in each section of each cultivar and in total in each cultivar for all nurseries. y For each column within cultivars, means with the same lowercase letter are not significantly different (P = 0.05) according to Dunn's test. z For each row with totals, means with the same uppercase letter are not significantly different (P = 0.05) according to Dunn’s test. Cultivary Section Nursery A2014 Nursery A2019 Nursery B Nursery C Nursery D Chardonnay Scion 5.7 ± 7.1 a 0 ± 0 a 0 ± 0 a 0 ± 0 a 0 ± 0 a Graft Union 13.8 ± 27.2 a 0.2 ± 0.5 a N/A 0 ± 0 a 0 ± 0 a Base 1.7 ± 2.0 a 0 ± 0 a 0.7 ± 2.2 a 0 ± 0 a 0 ± 0 a Roots 5.7 ± 12.8 a 0 ± 0 a 0.3 ± 0.8 a 0 ± 0 a 0 ± 0 a Totalz 26.9 ± 42.0 A 0.2 ± 0.5 A 1.1 ± 2.3 AB 0 ± 0 A 0 ± 0 A Merlot Scion 68.6 ± 36.5 a 0 ± 0 a 2.4 ± 4.3 a 20.5 ± 30.7 a 0 ± 0 a Graft Union 81.7 ± 124.1 a 0 ± 0 a N/A 21.4 ± 29.3 ab 0 ± 0 a Base 80.3 ± 100.7 a 0 ± 0 a 2.1 ± 2.4 a 0.1 ± 0.2 ab 0.1 ± 0.1 a Roots 79.9 ± 95.4 a 0 ± 0 a 0.3 ± 0.6 a 0 ± 0 b 0 ± 0 a Total 310.5 ± 192.8 B 0 ± 0 A 4.9 ± 5.3 A 42.0 ± 50.9 B 0.1 ± 0.1 A Pinot noir Scion 4.9 ± 8.6 ab 0.2 ± 0.5 a 0.2 ± 0.5 a 2.5 ± 7.4 a 1.3 ± 3.3 a Graft Union 16.7 ± 34.9 b 0 ± 0 a N/A 4.3 ± 12.9 a 6.2 ± 18.6 a Base 0.2 ± 0.5 a 0 ± 0 a 0.4 ± 0.9 a 0 ± 0 a 0.1 ± 0.1 a Roots 1.3 ± 1.9 ab 0 ± 0 a 0 ± 0 a 0 ± 0 a 0.2 ± 0.4 a Total 23.1 ± 42.9 A 0.2 ± 0.5 A 0.6 ± 1.3 B 6.7 ± 20.3 A 7.8 ± 21.8 B All TOTAL 120.2 ± 178.0 0.1 ± 0.5 2.2 ± 3.9 16.3 ± 36.6 2.6 ± 13.1 76 Table 2.4. Mean copies/ng DNA and standard deviation of Cadophora luteo-olivacea in each section of each cultivar and in total in each cultivar for all nurseries. y For each column within cultivars, means with the same lowercase letter are not significantly different (P = 0.05) according to Dunn's test. z For each row with totals, means with the same uppercase letter are not significantly different (P = 0.05) according to Dunn’s test. Cultivary Section Nursery A2014 Nursery A2019 Nursery B Nursery C Nursery D Chardonnay Scion 30.5 ± 31.8 ab 0.5 ± 0.8 a 1,466.8 ± 3,863.3 a 0.5 ± 0.8 a 47.1 ± 69.6 ab Graft Union 298.3 ± 456.5 a 96.2 ± 72.5 b N/A 4.6 ± 7.4 a 120.5 ± 141.1 a Base 11.6 ± 18.4 b 27.9 ± 26.4 b 42.6 ± 64.6 a 0.1 ± 0.1 a 0.5 ± 0.7 b Roots 5.2 ± 5.8 b 0.1 ± 0.4 a 67.9 ± 181.6 a 27.1 ± 73.9 a 57.1 ± 145.2 ab Totalz 345.7 ± 465.4 A 124.7 ± 84.3 A 1,584.2 ± 3,838.1 A 32.3 ± 80.0 A 225.1 ± 222.0 A Merlot Scion 185.1 ± 110.1 a 2.1 ± 5.0 a 8.1 ± 18.0 a 4.1 ± 6.3 a 8.5 ± 12.5 a Graft Union 198.4 ± 295.4 a 17.5 ± 19.8 b N/A 6.8 ± 9.2 a 46.2 ± 35.8 b Base 5.5 ± 12.2 b 15.7 ± 9.0 b 16.3 ± 37.8 a 182.1 ± 535.6 a 0.2 ± 0.3 a Roots 1.7 ± 5.1 b 0.2 ± 0.3 a 9.2 ± 26.2 a 55.2 ± 40.0 b 406.7 ± 923.0 b Total 390.7 ± 352.60 AB 35.5 ± 28.9 B 33.6 ± 43.0 A 248.2 ± 528.5 A 461.8 ± 929.2 A Pinot noir Scion 247.9 ± 82.9 a 0.2 ± 0.7 a 450.5 ± 745.7 a 34.4 ± 72.3 a 20.2 ± 19.8 ab Graft Union 548.3 ± 1262.3 ab 12.0 ± 11.5 b N/A 44.0 ± 89.9 a 103.2 ± 75.3 a Base 32.8 ± 55.6 c 17.7 ± 28.1 b 2.8 ± 3.7 a 82.6 ± 164.2 a 2.9 ± 7.6 c Roots 152.9 ± 296.2 bc 3.5 ± 4.7 ab 8.3 ± 23.1 a 3,587.0 ± 1,141.5 b 10.2 ± 18.7 bc Total 981.9 ± 1225.2 B 34.5 ± 26.7 B 461.6 ± 757.9 A 3,748.1 ± 1,212.0 B 136.5 ± 92.0 A All TOTAL 572.8 ± 835.5 64.9 ± 68.4 693.1 ± 2,351.6 1,342.9 ± 1,866.9 274.5 ± 570.9 77 Table 2.5. Mean copies/ng DNA and standard deviation of Dactylonectria macrodidyma in each section of each cultivar and in total in each cultivar for all nurseries. Cultivary Section Nursery A2014 Nursery A2019 Nursery B Nursery C Nursery D Chardonnay Scion 0 ± 0 a 0 ± 0 a 0 ± 0 a 0 ± 0 a 0 ± 0 a Graft Union 0 ± 0 a 0 ± 0 a N/A 0 ± 0 a 0 ± 0 a Base 0 ± 0 a 0 ± 0 a 3.7 ± 6.0 a 0 ± 0 a 0.4 ± 1.2 a Roots 0 ± 0 a 0 ± 0 a 0.6 ± 0.8 a 0 ± 0 a 0 ± 0 a Totalz 0 ± 0 A 0 ± 0 A 4.3 ± 6.3 A 0 ± 0 A 0.4 ± 1.2 A Merlot Scion 0 ± 0 a 0 ± 0 a 0 ± 0 a 0 ± 0 a 0 ± 0 a Graft Union 0 ± 0 a 0 ± 0 a N/A 0 ± 0 a 0 ± 0 a Base 0.5 ± 1.5 a 0 ± 0 a 0.6 ± 1.9 a 0 ± 0 a 0 ± 0 a Roots 0 ± 0 a 0 ± 0 a 0.7 ± 1.0 a 1.9 ± 2.1 b 0 ± 0 a Total 0.5 ± 1.5 A 0 ± 0 A 1.4 ± 1.9 A 1.9 ± 2.1 B 0 ± 0 A Pinot noir Scion 0 ± 0 a 0 ± 0 a 0 ± 0 a 0 ± 0 a 0 ± 0 a Graft Union 0 ± 0 a 0 ± 0 a N/A 0 ± 0 a 0 ± 0 a Base 0 ± 0 a 0 ± 0 a 0.3 ± 1.1 a 0 ± 0 a 0 ± 0 a Roots 0 ± 0 a 0 ± 0 a 0.3 ± 0.8 a 0 ± 0 a 0 ± 0 a Total 0 ± 0 A 0 ± 0 A 0.7 ± 1.5 A 0 ± 0 A 0 ± 0 A All TOTAL 0.2 ± 0.9 0 ± 0 2.1 ± 4.2 0.6 ± 1.5 0.1 ± 0.7 y For each column within cultivars, means with the same lowercase letter are not significantly different (P = 0.05) according to Dunn's test. z For each row with totals, means with the same uppercase letter are not significantly different (P = 0.05) according to Dunn’s test. 78 Table 2.6. Mean copies/ng DNA and standard deviation of Dactylonectria torresensis in each section of each cultivar and in total in each cultivar for all nurseries. Cultivary Section Nursery A2014 Nursery A2019 Nursery B Nursery C Nursery D Chardonnay Scion 0.1 ± 0.3 a 0 ± 0 a 0 ± 0 a 0.1 ± 0.2 a 0 ± 0 a Graft Union 0.1 ± 0.4 a 0 ± 0 a N/A 0 ± 0 a 0 ± 0 a Base 10.7 ± 14.2 b 2.4 ± 4.2 b 3.0 ± 7.4 ab 0 ± 0 a 0 ± 0 a Roots 63.2 ± 86.2 b 0 ± 0 a 0.5 ± 0.6 b 0.8 ± 1.9 a 0 ± 0 a Totalz 74.1 ± 87.1 AB 2.4 ± 4.2 A 3.6 ± 7.6 A 0.8 ± 1.9 A 0 ± 0 A Merlot Scion 0.1 ± 0.3 a 0 ± 0 a 0.1 ± 0.4 a 0 ± 0 a 0 ± 0 a Graft Union 0 ± 0 a 0 ± 0 a N/A 0 ± 0 a 0 ± 0 a Base 10.1 ± 6.0 b 4.6 ± 3.3 b 2.1 ± 3.3 a 0 ± 0 a 0 ± 0 a Roots 3.2 ± 7.1 a 1.2 ± 0.6 b 1.0 ± 1.5 a 1.3 ± 2.2 b 0 ± 0 a Total 13.4 ± 10.9 A 5.8 ± 3.3 B 3.3 ± 3.1 A 1.3 ± 2.2 A 0 ± 0 A Pinot noir Scion 0.7 ± 0.9 ab 0.0 ± 0.0 a 0 ± 0 a 0 ± 0 a 0 ± 0 a Graft Union 0 ± 0 b 0 ± 0 a N/A 0 ± 0 a 0 ± 0 a Base 27.7 ± 67.5 a 4.2 ± 3.7 b 9.2 ± 15.0 b 0 ± 0 a 0 ± 0 a Roots 40.9 ± 26.2 c 1.6 ± 1.4 b 0.7 ± 0.9 ab 0.1 ± 0.1 b 0 ± 0 a Total 69.3 ± 83.4 B 5.9 ± 3.6 B 9.9 ± 15.6 A 0.1 ± 0.1 A 0 ± 0 A All TOTAL 52.3 ± 75.1 4.7 ± 4.1 5.6 ± 10.6 0.8 ± 1.8 0 ± 0 y For each column within cultivars, means with the same lowercase letter are not significantly different (P = 0.05) according to Dunn's test. z For each row with totals, means with the same uppercase letter are not significantly different (P = 0.05) according to Dunn’s test. 79 Table 2.7. Mean copies/ng DNA and standard deviation of Phaeoacremonium minimum in each section of each cultivar and in total in each cultivar for all nurseries. Cultivary Section Nursery A2014 Nursery A2019 Nursery B Nursery C Nursery D Chardonnay Scion 0 ± 0 a 0.1 ± 0.1 a 0 ± 0 a 0.2 ± 0.4 a 0.4 ± 0.6 a Graft Union 0.2 ± 0.5 a 0.1 ± 0.1 a N/A 0 ± 0 a 0.2 ± 0.4 a Base 1.3 ± 2.9 a 0.7 ± 1.1 a 0.4 ± 1.4 a 0 ± 0 a 97.4 ± 124.7 b Roots 0.6 ± 1.5 a 0.1 ± 0.3 a 0.1 ± 0.2 a 0.1 ± 0.3 a 0.4 ± 0.7 a Totalz 2.2 ± 3.9 A 0.9 ± 1.2 A 0.6 ± 1.3 A 0.4 ± 0.5 A 98.4 ± 124.5 A Merlot Scion 0.9 ± 2.1 a 0 ± 0 a 0.3 ± 0.9 a 0.1 ± 0.3 a 98.9 ± 174.8 a Graft Union 0.4 ± 0.6 a 5.8 ± 16.4 a N/A 22.7 ± 32.4 b 441.4 ± 666.9 a Base 0 ± 0 a 0 ± 0 a 0.6 ± 1.8 a 9.4 ± 19.7 a 75.4 ± 172.5 a Roots 0 ± 0 a 0 ± 0 a 0 ± 0 a 0.3 ± 0.65a 9.1 ± 21.9 a Total 1.3 ± 2.5 A 5.8 ± 16.4 A 0.9 ± 1.9 A 32.6 ± 43.8 A 624.8 ± 798.8 A Pinot noir Scion 0 ± 0 a 0 ± 0 a 0 ± 0 a 0.1 ± 0.2 a 0.8 ± 1.5 a Graft Union 0 ± 0 a 12.2 ± 36.6 a N/A 0.1 ± 0.1 a 128.6 ± 384.2 a Base 0.2 ± 0.6 ab 0 ± 0 a 0.1 ± 0.2 a 0.5 ± 1.3 a 39.6 ± 56.9 b Roots 2.1 ± 3.6 b 0 ± 0 a 0.1 ± 0.3 a 0.1 ± 0.1 a 0.1 ± 0.1 a Total 2.4 ± 3.7 A 12.2 ± 36.5 A 0 .2 ± 0.3 A 0.8 ± 1.4 A 169.0 ± 393.2 A All TOTAL 2.0 ± 3.4 6.3 ± 23.6 0.6 ± 1.4 11.3 ± 29.4 297.4 ± 569.1 y For each column within cultivars, means with the same lowercase letter are not significantly different (P = 0.05) according to Dunn's test. z For each row with totals, means with the same uppercase letter are not significantly different (P = 0.05) according to Dunn’s test. 80 Table 2.8. Mean copies/ng DNA and standard deviation of each Phaeomoniella chlamydospora in each section of each cultivar and in total in each cultivar for all nurseries. Cultivary Section Nursery A2014 Nursery A2019 Nursery B Nursery C Nursery D Chardonnay Scion 242.5 ± 516.2 a 20.7 ± 60.4 a 0.35 ± 1.05 a 0 ± 0 a 0 ± 0 a Graft Union 637.6 ± 1533.4 a 347.91 ± 535.6 b N/A 0 ± 0 a 0 ± 0 a Base 63.2 ± 132.8 a 166.61 ± 249.8 b 0 ± 0 a 0 ± 0 a 0 ± 0 a Roots 516.0 ± 1196.3 a 0 ± 0 a 0 ± 0 a 0 ± 0 a 0 ± 0 a Totalz 1,459.4 ± 2,918.8 A 535.2 ± 670.19 A 0.35 ± 1.05 A 0 ± 0 A 0 ± 0 A Merlot Scion 987.6 ± 1719.0 a 0.2 ± 0.5 a 0 ± 0 a 0 ± 0 a 0 ± 0 a Graft Union 830.0 ± 1407.0 ab 225.4 ± 675.3 ab N/A 0 ± 0 a 0 ± 0 a Base 0.5 ± 1.1 bc 11.6 ± 18.6 b 0 ± 0 a 39.4 ± 118.3 a 0 ± 0 a Roots 0 ± 0 c 0 ± 0 a 0 ± 0 a 0 ± 0 a 0.1 ± 0.2 a Total 1,818.2 ± 2,793.5 AB 237.2 ± 672.11 B 0 ± 0 A 39.4 ± 118.3 A 0.1 ± 0.2 A Pinot noir Scion 2,843.9 ± 7,160.1 a 0.1 ± 0.3 a 0 ± 0 a 0 ± 0 a 0 ± 0 a Graft Union 1,537.6 ± 4,035.8 a 0.7 ± 1.6 a N/A 0.1 ± 0.3 a 0 ± 0 a Base 2,232.2 ± 3,400.0 a 18.5 ± 45.9 a 0 ± 0 a 0.1 ± 0.3 a 0 ± 0 a Roots 68.4 ± 150.2 a 0 ± 0 a 0 ± 0 a 0.1 ± 0.1 a 0 ± 0 a Total 6,682.1 ± 14,056.6 B 19.4 ± 45.6 B 0 ± 0 A 0.3 ± 0.4 A 0 ± 0 A All TOTAL 3,319.9 ± 8,773.7 263.9 ± 588.0 0.12 ± 0.63 13.2 ± 70.8 0.1 ± 0.2 y For each column within cultivars, means with the same lowercase letter are not significantly different (P = 0.05) according to Dunn's test. z For each row with totals, means with the same uppercase letter are not significantly different (P = 0.05) according to Dunn’s test. 81 Table 2.9. Mean sum copies/ng DNA and standard deviation of each fungal species in the basal end and roots of each rootstock for Nursery A2014y. Rootstockz Bot Clo Dm Dt Pm Pc Riparia Gloire 1.9 ± 3.7 a 10.7 ± 10.9 ab 0 ± 0 a 46.8 ± 42.7 a 0 ± 0 a 0 ± 0 a Self-rooted Baco noir 0.2 ± 0.3 a 5.4 ± 8.4 a 0 ± 0 a 21.3 ± 17.1 a 220.8 ± 269.3 b 1,598.4 ± 778.2 b SO4 8.7 ± 11.5 ab 11.9 ± 20.2 a 0 ± 0 a 9.4 ± 4.6 a 2.3 ± 4.6 a 1,156.1 ± 795.3 b 101-14 13.6 ± 7.9 b 117.0 ± 80.0 b 0 ± 0 a 14.1 ± 8.8 a 1.4 ± 2.5 a 1,851.4 ± 775.2 b 161-49 2.6 ± 2.1 ab 3.3 ± 2.8 a 0 ± 0 a 15.0 ± 4.5 a 0.4 ± 0.4 a 0 ± 0 a y Bot = Botryosphaeriaceae spp., Clo = Cadophora luteo-olivacea, Dm = Dactylonectria macrodidyma, Dt = Dactylonectria torresensis, Pm = Phaeoacremonium minimum, Pc = Phaeomoniella chlamydospora. z For each row within rootstocks, means with the same letter are not significantly different (P = 0.05) according to Dunn's test. 82 Figure 2.1. Scan showing nursery A2019 basal necrosis from the plant with the highest total fungal abundance at the basal end (724.05 copies/ng) with 50% necrosis (A) and the plant with the lowest total fungal abundance at the basal end (2.90 copies/ng) with 43% necrosis (B). 83 3 Impact of water stress on Phaeomoniella chlamydospora abundance and Petri disease symptom development in young grapevines. 3.1 Background Young vine decline (YVD) is caused by several taxonomically unrelated xylem-colonizing fungi, which cause vascular necrosis resulting in poor grapevine vigor, lower fruit yield, and eventual plant mortality within the first few years after planting (Gramaje and Armengol 2011). Young vine decline can be a consequence of up to three diseases, including black foot disease, Botryosphaeria dieback, and Petri disease (Agustí-Brisach and Armengol 2013; Gramaje and Armengol 2011; Úrbez-Torres 2011). Fungal pathogens causing the above-mentioned diseases can infect grapevine nursery material through various stages of the propagation process and have been reported from ready-to-plant nursery material around the world (Akgül et al. 2022; Berlanas et al. 2020; Billones-Baaijens et al. 2013c; Carbone et al. 2022; Pintos et al. 2018). Among all fungi known to cause YVD, the Petri disease pathogens Cadophora luteo-olivacea, Phaeoacremonium spp., and Phaeomoniella chlamydospora (Pc) have been largely found in asymptomatic ready-to-plant nursery material (Aigoun-Mouhous et al. 2021; Gramaje et al. 2009; Hrycan et al. 2023; Maldonado-González et al. 2020). Up to 100% of ready-to-plant nursery vines can be infected with Petri disease fungi, with significant variation in pathogen abundance between plants, different parts of the plant, and between nurseries (Hrycan et al. 2023). If infection always led to symptom development and plant mortality, viticulture would not exist in its current form today. For instance, studies conducted in British Columbia (BC) showed that approximately 8% of young grapevines were symptomatic for YVD across the province (Úrbez-Torres et al. 2014a), raising the question why mortality is not higher 84 when up to 100% of ready-to-plant nursery plants sold in Canada are infected with YVD fungi (Hrycan et al. 2023). Furthermore, several studies have identified many of these fungi in asymptomatic grapevine tissue in mature grapevines in vineyards (Abreo et al. 2011; Edwards and Pascoe 2004; Gainza-Cortes et al. 2020; González and Tello 2011; Halleen and Petrini 2003; Nerva et al. 2019; Zanzotto et al. 2007). Thus, it has been hypothesized that some of these YVD fungi may act as latent pathogens, living as endophytes within the grapevine microbiome until a trigger occurs, resulting in a transition to a pathogenic lifestyle (Hrycan et al. 2020). It is thought that the transition to a pathogenic phase occurs as a result of plant stress (Brown et al. 2012; Hrycan et al. 2020; Sieber 2007, Slippers and Wingfield 2007; van Niekerk et al. 2011b). Several observations have led to the hypothesis of an interaction between plant stress and disease development in grapevines infected with Petri disease fungi, which are also involved with Esca disease in older grapevines (Gramaje et al. 2018). For instance, Esca symptoms can vary yearly and have been correlated with climatic conditions (Marchi et al. 2006; Péros et al. 2008; Surico et al. 2000). Esca foliar symptom development has also been correlated with higher internal necrosis (Maher et al. 2012). A combination of pathogen infection level and plant stress is hypothesized to increase pathogen growth in the plant, which can result in the pathogen reaching a colonization threshold the plant cannot contain, leading to mortality, primarily due to increased internal necrosis development (Hrycan et al. 2020). However, the impact of plant stress on internal necrosis development in Pc infected plants is unclear. Recently, further investigation into Esca symptom development and water stress found that Esca foliar symptoms were inhibited in water stressed plants (Bortolami et al. 2021). However, Esca is a complex disease with two forms, characterized as chronic foliar symptom development, which can vary year to year, and 85 apoplectic, resulting in plant wilting (Gramaje et al. 2018). Esca-related apoplexy has been associated with high temperature and low rainfall conditions (Surico et al. 2006). The differing observations between the chronic and apoplectic form of Esca suggest the mechanisms for chronic symptom development may be different than apoplexy, which could be the result of plant stress. Like the apoplectic form in Esca, Petri disease in younger grapevines can result in sudden grapevine collapse (Gramaje and Armengol, 2011). While Esca and Petri disease are different, Pc is involved in both diseases and could respond similarly to plant stress in both young and older grapevines. Associations between plant stress and factors influencing disease development in grapevines infected with Esca and Petri disease associated fungi have been reported, primarily focused on water stress (Fischer and Ashnaei 2019). Whiting et al. (2001) observed that Pc can grow at osmotic water potentials of up to -8.3 MPa on potato dextrose agar, indicating the fungus can grow under severe water deficit. In addition, Lima et al. (2017) reported an increase in xylem sap and nutrient concentration in infected grapevines subjected to water stress, which may provide the fungus with increased carbohydrates under water stress conditions. Phaeomoniella chlamydospora can also impact plant-water relations. ‘Cabernet sauvignon’ and ‘Zinfandel’ vines infected with Pc and subjected to water stress displayed lower leaf water potential and greater stomatal conductance compared to non-infected water stressed plants, indicating that these fungi may increase susceptibility to water stress (Edwards et al. 2007a, b). Effects of Pc on plant-water relations may be due to the formation of tyloses in the xylem, which can impact stem hydraulic conductivity and may impact grapevine survival during periods of stress (Bortolami et al. 2021). In combination, these studies indicate Pc can grow during periods of severe water stress, increase plant susceptibility to water stress, and may gain access to increased resources for pathogen 86 growth. While conditions for pathogen growth may improve under water stress, pathogen abundance in plants under water stress has not been investigated. Arbuscular mycorrhizal (AM) fungi have been shown to mitigate abiotic and biotic stress factors in plants, improve plant growth and nutrition and can be applied to the soil using commercial fertilizers (Trouvelot et al. 2015). In water stressed grapevines, AM fungi have been reported to increase water content in the leaves, chlorophyll concentration, and dry root and shoot weight compared to non-AM grapevines (Ye et al. 2022). It has also been shown that AM can reduce black foot disease severity in grapevines, suggesting a potential use in disease management (Petit and Gubler 2006). However, recent work has reported increased internal necrosis and pathogen abundance in mycorrhizal grapevines infected with black foot disease fungi (Holland et al. 2019; Vukicevich et al. 2018). There is a lack of knowledge overall on whether water stress influences disease development in grapevines, the effects of which may vary for different fungal pathogens. Studies investigating water stress on Eutypa dieback reported an increase of Eutypa lata colonization in well-watered plants and drought did not increase pruning wound susceptibility to E. lata infection (Sosnowski et al. 2021). Similarly, high temperatures and high soil water content led to increased symptoms in E. lata infected grapevines (Sosnowski et al. 2011). Understanding how plant stress factors affect the growth of YVD fungi is essential in limiting the impact of disease in young vineyards. Previous studies on Pc infection of grapevines have primarily focused on foliar symptom development, lesion length and phenotypical measurements to record the effect of water stress on disease development. Currently, no studies have investigated whether water stress increases Pc abundance in grapevines. Accordingly, the main objectives of this study were to investigate if water stress increases Pc abundance and disease development in both the 87 greenhouse and field and to determine if AM fungi can mitigate stress under greenhouse conditions, thus reducing disease development. 3.2 Materials and methods 3.2.1 Greenhouse Experiment 3.2.1.1 Plant material, inoculum preparation and vacuum inoculation Dormant ‘Merlot’ canes were collected from one of the experimental vineyard blocks located at the Summerland Research and Development Centre (SuRDC), BC, Canada in February 2020 and stored in the dark at 2.5°C until used. Phaeomoniella chlamydospora isolate SuRDC-1070, obtained from grapevines showing Petri disease symptoms in BC and kept in the SuRDC Plant Pathology fungal collection was used for this study (Úrbez-Torres et al. 2014a). Mycelial plugs from the isolate were retrieved from the collection and plated onto potato dextrose agar (PDA; Sigma-Aldrich, St Louis, MO) petri plates incubated at 22°C in the dark for three weeks until a mycelial colony developed. Two mL of sterile distilled water amended with a drop of Tween 80 were pipetted onto the fresh mycelial colony and gently scraped with a sterile spoonula. The resulting spore suspension was vortexed and adjusted to 25,000, 5,000, and 1,000 spores per 10 uL with a haemocytometer. ‘Merlot’ canes were pruned to three nodes (approximately 20 cm) and were vacuum inoculated from the base with either 10 uL of sterile distilled water amended with a drop of Tween 80 or 10 uL of a spore suspension of either 25,000, 5,000, or 1,000 spores of Pc (Rooney et al. 2001). Inoculations were made by pipetting onto the freshly cut base of each grapevine cutting and introduced into the vascular system by applying vacuum at the other end of the cane for 1 s. Vacuum pressure was 71.5 mmHg measured with a model 06-664-19 Manometer (Fisher 88 Scientific, Waltham, MA). Viability of Pc was determined by plating a 10 uL suspension from each inoculum concentration in triplicate immediately following the procedure. Plates were incubated at 22°C in the dark for seven days until fungal colonies were observed. Dormant canes were transferred to the greenhouse and rooted in 2 L buckets, separated by treatment, containing 750 mL of water, changed twice weekly. Buckets were placed on rooting mats set to 20℃. Upon budbreak, greenhouse lights were turned on and the greenhouse temperature was set to 20℃. Rooted cuttings were planted in 1-gallon pots containing 3,000 g of pasteurized soil on March 31, 2020. Soil was pasteurized twice in a LANSA model LA soil pasteurizer (Johnson Machine Company LTD, Burlington, Ontario) set at a temperature exceeding 72℃ (Trevors 1996), following manufacturer’s instructions. Half of the pots were amended with 0.25g of AGTIV Specialty Crops Powder (Premier Tech Growers and Consumers, Quebec, Canada) containing 3,000 Rhizophagus irregularis spores following manufacturer’s instructions. 3.2.1.2 Experimental design and plant health measurements The factorial set of treatments was carried out in a complete randomized design and consisted of five levels of inoculum, 25,000 (25k), 5,000 (5k), 1,000 (1k) Pc spores, water-inoculated (WIC), and non-inoculated (NIC), two levels of AM fungi, AM inoculated (AM+), and non-inoculated (AM-), and two levels of irrigation, 100% water (I1) and 50% water (I2) by volume and included seven reps per treatment with 140 plants total. As a result of pandemic restrictions during the first year of the experiment, grapevines were grown to one shoot during 2020 and maintained in the greenhouse equally watered with 500 mL twice weekly until July 10, and three times a week onward until mid-September when water was gradually reduced to 50 mL three times a week in preparation for dormancy. Grapevines were placed in cold storage in the dark at 2.5°C on December 18, 2020, and returned to the same greenhouse compartment on 89 March 5, 2021. In year two (2021), plants were grown to two shoots and watered every second day. Water stress in year two was imposed via a 50% reduction in water volume. NS plants received 500mL water and S plants 250mL every second day starting June 18, 2021. Water volume was adjusted according to plant measurements and was reduced to NS: 400mL, S: 200mL on August 14th, NS: 300mL, S:150mL on August 18th, NS: 200mL, S:100mL, on August 28th, and raised to NS: 300mL, S:150mL on September 1st until experiment concluded on September 13th, 2021. Grapevines were fertilized with equal amounts of granular 20-20-20 (Master Plant-Prod Inc., ON, Canada) bi-weekly according to manufacturer’s instructions. Stem water potential, leaf chlorophyll content, and soil matric potential were measured in year two. Mid-day stem water potential was measured bi-weekly from June 29 to September 9, 2021, with a model 610 pressure chamber (PMS Instrument Co. Corvallis, OR). Leaf chlorophyll concentration was measured bi-weekly from May 25 to September 9, 2021with a SPAD 502 Plus Chlorophyll Meter (Spectrum Technologies, Aurora, IL). Soil matric potential was measured in the soil of six NS and six S pots randomly distributed in the greenhouse with T5 Tensiometers (Meter Group, Pullman, WA) every 15 m and averaged daily from June 8 to September 9, 2021. Pruning weights and node number were collected in both 2020 and 2021 and root weight was collected upon conclusion of the experiments. Shoots and roots were dried in an Environmental Chamber model 217502 (Hotpack, Anaheim, CA) at 65°C until dry weights were consistent over a 24 h period. Necrosis measurements were conducted upon conclusion of the experiment at one cm from the base of each grapevine as described in Hrycan et al. (2023) with the following alterations: Images were set at 8-bit colour and a threshold analysis was conducted in Fiji ImageJ version 1.53t. Threshold numbers are set values for the degree of shading from black to white (0-255). Black wood discoloration (Class 1 necrosis) was set at 5-45, brown wood discoloration 90 (Class 2 necrosis) was set at 46-80, and black and brown wood discoloration (total necrosis) was 5-80 (Figure 3.1). 3.2.1.3 Phaeomoniella chlamydospora absolute quantification In order to verify presence of the pathogen in the plant and possible Pc growth between inoculation and planting, three grapevines from each treatment were randomly selected 24 h after vacuum inoculation and at time of planting (six weeks post-inoculation). Plants were cut into two cm sections from 0-18 cm. At harvest, a two cm section of wood was cut two cm above the base. Individual sections were surface sterilized via submersion in 70% ethanol for 30 s, 10% bleach for 1 min, and sterile water 2 x 1 min washes. Sections were cut in half longitudinally, finely chopped with a razor blade, and 0.2g was placed in Qiagen DNeasy PowerSoil DNA extraction tubes (Qiagen, Hilden, Germany). The other half of each section was finely cut and plated on PDA amended with 0.1mg/mL tetracycline hydrochloride (PDA-tet) to re-isolate Pc as described in Hrycan et al. (2023). A Qubit 1.0 Fluorometer dsDNA HS Qubit assay (Thermofisher, Waltham, MA) was used to calculate DNA concentration. Quantification of Pc was conducted with a Droplet Digital™ PCR (ddPCR) system (Bio-Rad Laboratory, Hercules, CA) using ITS primers (Martín et al. 2012) with the addition of a probe (Hrycan et al. 2023). Reactions and analysis were conducted as described in Hrycan et al. (2023). Pathogen abundance quantification results were standardized via calculation of gene copies/ng of DNA (Hrycan et al. 2023). 3.2.1.4 Arbuscular mycorrhizal fungal colonization Fresh roots were collected from the first rep of each treatment at harvest. Roots were frozen in liquid nitrogen, crushed with a pestle and mortar, and 0.2g of crushed root material was placed in Lysing Matrix A (MP Biomedicals Irvine, CA), tubes and extracted using the Qiagen 91 DNeasy Powerlyzer Powersoil kit according to manufacturer’s specifications. The Qiagen DNeasy Powerlyzer Powersoil kit was used due to manufacturer discontinuation of the DNeasy Powersoil kit. DNA concentration was calculated using a Qubit 1.0 fluorometer as previously described and ddPCR was conducted on each sample to determine presence or absence of the AM fungi using primers and probe described in Kokkoris et al. (2019). Black Sheaffer ink root staining was conducted to visually confirm presence of AM fungi via microscope (Vierheilig et a. 1998). 3.2.2 Field experiment 3.2.2.1 Plant material, inoculum preparation and vacuum inoculation Dormant ‘Merlot’ and dormant ‘SO4’ canes were collected from an experimental block at the SuRDC in February 2020 and stored in the dark at 2.5°C. Mycelial plugs containing Pc (SuRDC-1070) obtained from the SuRDC plant pathology fungal collection were plated and incubated followed by the creation of spore suspensions containing 25,000 (25k) and 2,500 (2.5k) spores using a haemocytometer as described in section 3.2.1.1. Canes were pruned to three nodes (22 cm) and were vacuum inoculated from the base as described in 3.2.1.1 with sterile distilled water, 2.5k or 25k Pc spores. Phaeomonella chlamydospora viability was determined by plating 10 uL of each suspension in triplicate onto PDA, followed by incubation at 22°C for seven days until fungal colonies were observed. Inoculated canes were immediately placed in cold storage at 2.5°C for 24 h. Dormant ‘SO4’ canes, self-rooted ‘Merlot’ canes and ‘Merlot’ scion material were dipped in 10% bleach for 1 s, disbudded, and omega grafted together. The grafted ‘SO4’ and self-rooted ‘Merlot’ were dipped in Stim-Root rooting hormone (Premier Tech Home and Garden, Brantford, 92 ON) and layered in bins containing moist perlite, separated by treatment, and stored at room temperature in the dark for four weeks. On April 6, 2020, callused grapevines were removed from perlite, the top bud was dipped in wax, and planted in individual cartons containing 50% perlite and 50% Sunshine Mix #4 (Sun Gro Horticulture, Agawam, MA). Grapevines were placed in the greenhouse at 20℃ and watered evenly 3x per week. Rooted grapevines were planted on July 9, 2020, in an experimental field plot at the SuRDC. 3.2.2.2 Experimental Design and plant health measurements Grapevines were planted in a split-plot randomized block design, split by stress treatment. The block contained loamy sand soil and row x plant spacing was 2.5m x 0.93m. Factorial treatments included Cultivar: self-rooted ‘Merlot’ and ‘Merlot’ grafted onto ‘SO4’ (‘Merlot’ / ‘SO4’), inoculum: water-inoculated control (WIC), 2.5k, and 25k, and irrigation treatment: 100% water (I1) and 50% water (I2) with eight replicates per treatment. Plants were grown to one shoot in year one (2020), and three shoots in years two (2021) and three (2022). The plants were removed in the spring of year four (2023). Non-stressed plants received irrigation from two Toro 0.5 GPH pressure compensating emitters (Toro, Bloomington, MI) while water stressed plants were irrigated with one emitter. Irrigation timing to reduce water volume was adjusted according to stem water potential measurements. Plants were irrigated Mon-Wed-Fri. Plants were fertilized each spring with equal amounts of 14-14-14 fertilizer (ICL Group LTD, Tel-Aviv, Israel) according to manufacturer’s instructions. No measurements were taken in 2020 due to pandemic restrictions. Mid-day stem water potential was measured with a model 610 pressure chamber (PMS Instrument Co. Corvallis, OR) in the second and third year. In the second year, mid-day stem water potential was measured on one leaf per plant on September 9, 2021. In the third-year, mid-day stem water potential was measured bi-weekly on 93 Tuesday or Thursday from July 19 to August 16, 2022, and then weekly until September 15, 2022. Leaf chlorophyll concentration was measured on 10 leaves per plant and averaged and was conducted on August 17 and September 8 in the second year (2021), weekly from July 11 to September 26 in the third year (2022), and on May 29 in the fourth year (2023) with a SPAD 502 Plus Chlorophyll Meter (Spectrum Technologies, Aurora, IL). Pruning weights and node numbers were collected throughout each growing season and dried at 65°C as described in chapter 3.2.1.2. Symptom development and plant mortality were recorded each year. Plants were uprooted and placed in cold storage in the dark at 2.5℃ on May 30, 2023, for analysis. Roots were cleaned in fresh water, removed from the plant, and dried at 65℃ until weights were consistent over two days. Necrosis measurements were taken one cm from the base as described in chapter 3.2.1.2. Trunk diameter was measured below the primary bud of the initial cane with CD-6” C digital caliper (Mitutoyo Corp. Japan). 3.2.2.3 Phaeomoniella chlamydospora absolute quantification To verify presence of the pathogen, three grapevines from each treatment were randomly selected at time of planting, 13 weeks post-vacuum inoculation. The Qiagen DNeasy Powersoil kit was used for DNA extractions and DNA concentration and pathogen abundance was analyzed as described in chapter 3.2.1.3. At harvest, plants were drilled at 0-1cm, 2-4cm, and 4-6cm locations from the base using a sterile 1/8” drill-bit for re-isolation of the pathogen as described in chapter 3.2.1.3. 0.1g of wood obtained from each drilling was used, wood was placed in Lysing Matrix A tubes, frozen in liquid nitrogen, and macerated in a FastPrep-24™ at 6.0 m/s x 40 s x3. DNA was extracted using the Qiagen DNeasy Plant Mini kit according to manufacturer’s instructions. The DNA extraction kit was changed due to manufacturer discontinuation of the Qiagen DNeasy Powersoil kit. The DNA concentration and ddPCR were 94 conducted as described in chapter 3.2.1.3. Each plant section was quantified separately and the sum of copies/ng DNA for 0-1, 2-4, and 4-6cm was used for analysis to compare Pc abundance between grapevines. 3.2.3 Statistical analysis Data analyses were conducted to determine whether inoculum level, stress, and/or AM fungi affected fungal abundance, necrosis, stem water potential, leaf chlorophyll content, dry shoot and root weight, and node number for each experiment. Greenhouse fungal abundance, necrosis, pruning and root weights, and node number were analyzed with a two-way ANOVA followed by Tukey’s Honest Significant Difference test set at 5% significance. Residuals were tested with the Shapiro-Wilk test. Greenhouse leaf chlorophyll, stem water potential, soil matric potential and all field data were analyzed with a linear mixed effect model considering the split-plot randomized block design and fit with REML with the package nlme. Significance was tested with a Type 3 ANOVA. Residuals were inspected visually. Tukey’s Honest Significant Difference test set at 5% was conducted on significant interactions using the emmeans package in R Version 4.3.0 (Lenth 2023; Pinheiro and Bates 2000; R Core Team, 2023). A chi-squared test was conducted to determine effects of cultivar, inoculum, and treatment on symptom expression and plant mortality. 3.3 Results 3.3.1 Greenhouse 3.3.1.1 Plant health measurements In year two (2021), leaf chlorophyll content was 4.72%, 4.11%, and 2.45% lower in Il2 than I1 plants on June 30, July 29 and September 9 (Table S3.1) respectively, while no other 95 treatment effects were observed (Table 3.1). Mid-day stem water potential was significantly lower in I2 than I1 plants at each time point (Figure 3.2) and overall (I2: -1.9MPa; I1: -1.32MPa), showing higher stress in I2 plants, but no effects of inoculum or AM fungi were observed (Table 3.1). Soil matric potential was lower in soil in I2 treated pots (-19.75 kPa) compared to I1 treated pots (-7.58 kPa) overall (Table 3.1). Soil matric potential daily average was lower in I2 soil compared to non-stressed soil at several time points, reaching a minimum of -72.35 kPa in stressed pots (Figure 3.2). In year two (2021), I2 treatment reduced dry pruning weight by 10.4% and node number by 22.4% in I2 compared to I1 plants. Mean node number was higher in AM+ I2 plants than AM- I2 and both AM+ and AM- I1 plants in year two (Table S3.2). At harvest, I1 plants showed 17.7% lower dry root weight than I2 plants. Inoculum had no effect on pruning weight, node number, or root weight in either year (Table 3.2). Root molecular analysis determined that 27% of AM inoculated plants were colonized, and 0% of non-mycorrhizal plants were colonized by the end of the experiment. Visual confirmation via compound microscope confirmed the presence of AM fungi in the roots. 3.3.1.2 Quantification of Phaeomoniella chlamydospora Phaeomoniella chlamydospora was not detected at 2-4cm from the base at time of inoculation using ddPCR, however, re-isolations confirmed the presence of Pc in 100% of inoculated plants. Molecular detection at time of planting detected higher Pc abundance at 2-4cm from the base in 25k compared to 5k and 1k inoculated plants, while NIC and WIC plants were negative for Pc. Re-isolations confirmed the presence of Pc at time of planting in 100% of inoculated plants while the fungus was not isolated from NIC or WIC plants. At harvest, Pc was 96 not detected in NIC or WIC plants via ddPCR or traditional plating, but Pc was isolated from 96% of 25k, 93% of 5k, and 90% of 1k plants. Initial inoculum quantity did not affect pathogen abundance at harvest. Irrigation treatment and AM fungi had a positive effect on pathogen growth in Pc inoculated plants (Table 3.3). Inoculated plants subjected to I2 treatment and inoculated with AM fungi had significantly higher Pc abundance than non-stressed and stressed non-mycorrhizal plants (Figure 3.4). 3.3.1.3 Trunk measurements Basal necrosis measurements indicated effects of inoculum and stress on class 1 necrosis, total necrosis, and basal surface area. Class 2 necrosis was impacted by inoculum alone (Table 3.3). Class 1 and total necrosis were significantly higher in plants containing Pc and in I2 plants, while plants containing Pc had lower Class 2 necrosis compared to WIC plants (Table 3.4). Plants inoculated with 1k Pc spores had higher Class 2 necrosis compared to 25k spores (Table 3.4). Basal surface area was higher in plants inoculated with 25k spores compared to WIC plants and in I1 compared to I2 plants (Table 3.4). 3.3.2 Field 3.3.2.1 Plant health measurements In year two (2021), 25k ‘SO4’ plants showed 6.25% lower leaf chlorophyll content than ‘SO4’ WIC plants. I2 treatment plants showed overall a 4.76% higher leaf chlorophyll content than I1 plants. In year three (2022), plants inoculated with 25k spores showed 3.15% and 5.00% lower leaf chlorophyll content than WIC and 2.5k spore plants respectively, while ‘Merlot’ plants had 2.39% higher chlorophyll content than ‘SO4’ plants. Leaf chlorophyll content was not significantly higher between treatments in 2023 (Table 3.5; S3.3). Mid-day stem water potential 97 was significantly lower in I2 plants (-0.85 MPa) than I1 plants (-0.79 MPa) in 2021, and in 2022 (I2 = -1.11 MPa, I1 = -0.93 MPa). In 2022, I2 plants had significantly lower MPa on August 25 and September 6th (Figure 3.2). ‘Merlot’ mean stem water potential (-0.99 MPa) was significantly higher than ‘SO4’ (-1.11 MPa) in year three. Dry pruning weight was significantly lower in ‘Merlot’ than ‘SO4’ in 2021, 2022, and 2023 and node number was significantly lower in ‘Merlot’ than ‘SO4’ in 2022 and 2023. ‘Merlot’ dry pruning weight was 41.4% lower than ‘SO4’ in 2021, 44.5% in 2022, and 64.7% in (Table S3.4; S3.5). ‘Merlot’ node number was 30.15% lower in 2022 (‘Merlot’: 26.6, ‘SO4’ 38.1), and 38.79% lower in 2023 (‘Merlot’: 42.9. ‘SO4’: 70). Dry root weight was 45% lower in ‘Merlot’ than ‘SO4’ upon conclusion of the experiment, but water stress and inoculum had no effect (Table 3.6; S3.4) .3.3.2.2 Quantification of Phaeomoniella chlamydospora Phaeomoniella chlamydospora abundance was higher in plants inoculated with 25k spores compared to 2.5k and WIC plants at time of planting. However, WIC plants were infected with Pc at time of planting. Re-isolations confirmed the presence and viability from the first 0-6cm of Pc in 66% of 25k and 2.5k plants, and 0% of WIC plants. Pc abundance was impacted by initial inoculum quantity only (Table 3.7). Plants initially inoculated with 25k spores contained 65,605.95 ± 14,669.94 copies/ng DNA at harvest, which was not significantly different than plants inoculated with 2.5k spores containing 44,227.35 ± 9,889.54 copies/ng DNA. At harvest, Pc inoculated plants contained significantly higher Pc abundance than WIC plants, which contained 21,167.32 ± 4,233.45 copies/ng DNA (Table 3.7). 98 ‘Merlot’ plants had a Pc abundance of 45,828.4 ± 9290.8 copies/ng DNA which was not significantly different than ‘SO4’ which contained 38,800.7 ± 7110 copies/ng DNA. 3.3.2.3 Trunk measurements Class 1 necrosis was significantly lower in WIC plants (8.96%) compared to plants inoculated with 25k (17.7%) or 2.5k (10.87%) Pc spores and total necrosis was significantly higher in plants inoculated with 25k Pc spores (32.81%) compared to 2.5k (21.14%) and WIC plants (19.36%). Basal surface area was significantly lower overall in plants inoculated with 25k spores (‘Merlot’: 131.57mm, ‘SO4’: 120.37mm, overall: 125.17mm) compared to 2.5k spores (‘Merlot: 195.44mm, ‘SO4’: 131.45mm; overall: 157.05mm), but 2.5k inoculated plant basal surface area was not significantly different from WIC plants (‘Merlot’: 178.11mm, ‘SO4’: 131.39mm, overall: 153.74mm). Total necrosis was higher in ‘Merlot’ (31.78%) compared to ‘SO4’ (18.44%) and basal surface area was higher in ‘Merlot’ (168.1mm) than ‘SO4’ (127.94mm). Class 2 necrosis and trunk diameter were not significantly different between treatments (Table 3.7). 3.3.2.4 Plant symptom development and mortality In 2021, cumulatively plant mortality was higher in ‘Merlot’ plants inoculated with 25k and 2.5k spores compared to 100% survival of WIC plants (Table 3.8). In 2021, 100% of ‘SO4’ plants survived. In 2022, WIC plants in I1 treatment had significantly higher mortality compared to I2 WIC plants (Table 3.8). In 2022 and cumulatively, ‘Merlot’ plants subjected to I1 treatment had significantly higher mortality (47% and 62% respectively) compared to I2 plants (13% and 28% respectively), while no significant effects were observed for ‘SO4’ (Table S3.3). In 2021 and 2022, ‘Merlot’ and ‘SO4’ plant mortality occurred but was not significantly different 99 between treatments (Table 3.8). Across all years, ‘Merlot’ overall had 46.15% mortality which was significantly higher than ‘SO4’ plant mortality at 17.95%. Six plants were symptomatic for young vine decline in 2022 (Figure 3.4). Symptomatic plant mean stem water potential was -0.71 MPa compared to -1.04 MPa in non-symptomatic plants and leaf chlorophyll readings in 2022 were 18.81 in symptomatic grapevines and 27.25 in asymptomatic. Three of the six symptomatic plants collapsed prior to the conclusion of the experiment in 2023, when most grapevine mortality occurred, thus these three plants were excluded from overall plant health and pathogen abundance analysis. The three plants that collapsed were 1) ‘Merlot’ WIC I2, 2) ‘SO4’ 2.5k I2, and 3) ‘SO4’ WIC I2. Pathogen abundance was 1) 33,700, 2) 11,384, and 3) 17,466 copies/ng DNA. For the three plants mean dry shoot weight was 2.82g, mean node number was 32, and mean dry root weight was 11.05g. Mean class 1 necrosis was 25.8%, class 2 necrosis was 42.2%, and total necrosis was 68%. Mean basal surface area was 129.04 mm. 3.4 Discussion This is the first study to investigate the effects of water stress and different inoculum levels on the Petri disease pathogen Phaeomoniella chlamydospora development in grapevines. Greenhouse mid-day stem water potential and tensiometer readings indicated soil moisture was low and plants were subject to high to severe water stress. In the field, mid-day stem water potential indicated mild to moderate plant water stress (Deloire et al. 2020). In the field in 2022, leaf chlorophyll levels were lower in 25k plants than WIC, but different inoculum levels had no effect on stem water potential. Dry pruning weight, node number, and dry root weight in the greenhouse were reduced due to I2 treatment while in the field, node number was 100 reduced in I2 plants in 2022 and 2023 only and no other effects of irrigation treatment on plant growth measurements were observed. In contrast, I2 treatments in the greenhouse had no effect on plant growth measurements. Field growth measurements indicated no effect of Pc infection. Petri disease symptoms include stunted growth and plants infected with Pc have been reported to show lower stem water potential (Edwards et al. 2007a, b; Gramaje et al. 2018), but no effects of Pc were observed in our study in the greenhouse, nor in the field until 2022. Foliar symptoms developed in the field in August and September of 2022, primarily in 25k and 2.5k inoculated ‘SO4’ plants, leading to sudden vine collapse, a characteristic symptom of severe Petri disease (Gramaje et al. 2018). Symptoms included reddening of the leaves, lower leaf chlorophyll content, and greater stem water potential in 2022, indicating leaf dysfunction. In older grapevines with Esca, in which Pc is involved, water stress has been correlated with the inhibition of symptoms, in contrast to the findings of this study (Bortolami et al. 2012). However, Esca is a separate and complex disease, and has two forms, chronic and apoplectic (Gramaje et al. 2018), and symptom development in this study was followed by sudden vine collapse, like the Esca apoplectic form. Chronic Esca symptom development may be unrelated to internal necrosis, as observed in other grapevine trunk diseases (Travadon et al. 2023; Sosnowski et al. 2011) and instead could be due to leaf vascular occlusions or toxin production (Bortolami et al. 2019, 2022). On the other hand, Esca apoplexy has been associated with high temperatures and lack of rainfall (Surico et al. 2006). In this study ‘Merlot’ mortality was significantly higher in I2 plants than I1. In contrast, symptom development and plant mortality did not occur in the greenhouse, but the greenhouse experiment was significantly shorter than the field experiment and was conducted under greenhouse conditions in pasteurized soil, which may have resulted in more favourable conditions for plant survival due to exposure to fewer external factors than field 101 plants. I2 treatment was applied for a longer period in the field and external factors in the field such as plant-parasitic nematode infestation (Forge et. 2021), cold events, and heatwaves may have weakened the plants further in the field compared to the greenhouse. The findings of this study suggest pathogen abundance is correlated with symptom development, which has been observed in other diseases in other hosts (Impullitti et al. 2009; Manter et al. 2003). Sudden drops in temperature during the winters of 2021 and 2022 may have contributed to vine mortality or had an impact on the pathogen abundance. An association between localized cold exposure and B. dothidea colonization has been reported in other hosts (Schoeneweiss and Wene 1980; Wene 1979). Phaeomoniella chlamydospora has often been isolated from asymptomatic grapevines and nursery material, indicating it can have a limited impact on phenotypical plant measurements for a period (Abreo et al. 2011; Edwards and Pascoe 2004; Gainza-Cortes et al. 2020; González and Tello 2011; Halleen and Petrini 2003; Nerva et al. 2019; Zanzotto et al. 2007). The results of this study indicate this could be related to pathogen abundance, as seen in other hosts, but further work is required to understand the levels of Pc abundance required for symptom development in young grapevines. Variations in rootstock susceptibility to Pc infection have been previously reported (Eskalen et al. 2001), but rootstock tolerance to infection under stress has not been investigated. In this study ‘Merlot’ plants had lower dry pruning weight, node numbers, dry root weight, higher pathogen abundance, total necrosis, and overall plant mortality than ‘SO4’. These findings suggest self-rooted ‘Merlot’ has higher susceptibility to Pc infection compared to ‘SO4’ rootstock under natural field conditions. However, this study found ‘SO4’ had lower stem water potential compared to ‘Merlot’ indicating a higher level of stress in ‘SO4’. Murolo and Romanazzi (2014) observed increased disease incidence in ‘SO4’ compared to ‘1103P’ rootstock, 102 which is thought to be more drought tolerant, but the results of this study indicate plant stress may not be the only factor. Xylem diameter can also play a role in pathogen abundance and plant susceptibility to Pc (Pouzoulet et al. 2017). In grapevines infected with P. minimum, another Petri disease associated fungi, smaller xylem diameter has been associated with lower pathogen abundance (Ramsing et al. 2021). Xylem diameter in combination with plant stress should be investigated to determine how pathogen growth and symptom expression are impacted. Previous work has reported no correlation between internal necrosis and pathogen abundance in ready-to-plant nursery grapevines (Hrycan et al. 2023). One objective of this study was to determine whether initial pathogen abundance had an impact on internal necrosis within the plant after planting. In both greenhouse and field experiments, pathogen abundance affected internal necrosis. In the greenhouse, class 1 and total necrosis were significantly higher in inoculated plants compared to non-inoculated, while class 2 necrosis was higher in non-inoculated plants. Recent nursery surveys have reported a correlation between necrosis and individual nurseries and rootstocks rather than pathogen abundance or infection (Carbone et al. 2022; Dubrovsky and Fabritius 2007; Gramaje and Armengol 2011; Hrycan et al. 2023; Lade et al. 2022). However, previous studies have primarily investigated total necrosis without separating black necrosis from browning of the wood (Hrycan et al. 2023). Necrosis from both greenhouse and field plants suggests browning of the wood should be excluded from pathogen necrosis analysis for Pc. In the field, class 1 necrosis was higher in 25k and 2.5k inoculated plants than WIC plants, and total necrosis was higher in 25k inoculated plants compared to 2.5k and WIC plants, while no differences between inoculum levels were observed in the greenhouse. In combination with higher total necrosis in 25k plants, basal surface area was lower. Water stress alone was 103 mild to moderate and unlikely to induce plant death, instead, increased necrosis and lower basal surface area could have impacted plant symptom development and mortality in combination with water stress (Maher et al. 2012; Oliva et al. 2014). In addition, plant stress was higher in the greenhouse than the field, and stress induced higher necrosis in the greenhouse, potentially due to xylem cavitation (Gambetta et al. 2020). Interestingly, in ‘Merlot’ plants inoculated with 25k Pc spores opposing effects on basal surface area were observed, increasing in the greenhouse and decreasing in the field, although in field plants, the effect of Pc on basal surface area was not statistically significant in ’Merlot’ alone. Pc infection can result in xylem blockage due to necrosis or tylose formation (Bortolami et al. 2021), which the plant could respond to by the growth of new xylem vessels as observed in other host responses to xylem dysfunction after water stress (Gauthey et al. 2022). A decrease of basal surface area in 25k field plants could be due to increased pathogen growth in the field over the longer-term. Phaeomoniella chlamydospora can grow under severe water deficit and may have better access to carbohydrates under drought conditions (Lima et al. 2017; Whiting et al. 2001), indicating water stress may favor pathogen growth and colonization (Hrycan et al. 2020). The main objective of this study was to understand whether water stress increases pathogen growth within the plant. In the greenhouse, Pc pathogen abundance was greater in Pc inoculated water stressed plants, but only when AM fungi were present. Arbuscular mycorrhizal fungi can increase plant growth and improve plant resistance to abiotic and biotic stress factors (Trouvelot et al. 2015; Ye et al. 2022). Thus, it was hypothesized that AM fungi would reduce plant stress and decrease disease development in grapevines infected with Pc. However, the results of this study indicate AM fungi to be beneficial to pathogen growth under plant stress. The findings of the greenhouse experiment on the interaction between pathogen abundance and presence of AM 104 fungi are in accordance with Holland et al (2019) and Vukicevich et al. (2018) who observed an increase in fungal abundance and necrosis in plants infected with black foot fungi and colonized by AM fungi. In contrast, a link between internal necrosis in Pc infected grapevines was not observed with AM fungi. The mechanism behind the influence of AM colonization and drought on Pc fungal abundance is unclear but, an interactive effect of AM fungi and drought on plant defenses may explain Pc growth. Water stress alone and in combination with AM fungi has been reported to increase salicylic acid production in other hosts (Choudhary and Senthil-Kumar 2022; Sanchez-Romera et al. 2018). An increase in salicylic acid, which is antagonistic to jasmonic acid involved in induced systemic resistance (Thakur and Sohal 2013), could result in reduced plant resistance against necrotrophic pathogens (Rahman et al. 2012). Additionally, plants subjected to the 50% irrigation treatment in the greenhouse had lower leaf chlorophyll, which could indicate a reduction in nutrient uptake due to water stress. Reduced nutrient uptake and changes to the grapevine microbiome due to water stress in infected plants (Leal et al. 2024) could further promote pathogen growth in plants due to reduced capacity for plant defense and reduced microbial diversity. If Pc is a latent pathogen, increased pathogen growth under water stress conditions will reach the colonization threshold in which plant mortality occurs sooner (Hrycan et al. 2020). Overall, greenhouse pathogen abundance results support the hypothesis of increased growth under plant stress, but other factors may be involved in Pc growth, as stress alone did not influence pathogen abundance. It is unclear whether greater Pc abundance is detrimental to plant health, but pathogen abundance has been correlated with symptom development in other hosts such as Swiss Needle Cast disease in Douglas-Fir and brown stem rot of soy (Impullitti et al. 2009; Manter et al. 2003). Further investigation is required to understand how high Pc abundance affects plant health. 105 In contrast to the greenhouse, pathogen abundance was not impacted by irrigation treatment under natural field conditions. Observed differences between the field and greenhouse may be due to the greater severity of water stress observed in the greenhouse compared to field plants. While Pc abundance was not affected by irrigation treatment in the field, initial inoculum quantity influenced pathogen abundance at harvest. In the field, 25k inoculated plants had greater Pc abundance than WIC plants which were positive for P. chlamydospora. This contrasts with pathogen abundance in the greenhouse where inoculum levels were not significantly different at harvest. However, pathogen abundance varied greatly between plants, which may have impacted significance. Variation in pathogen abundance is in accordance with the findings of Hrycan et al. (2023) in ready-to-plant nursery material, however, while variation in nursery material could be attributed to natural infection, it is unclear why we obtained such high Pc variation in both greenhouse and field plants inoculated with known spore quantities. Further work is required to understand the underlying cause of this variation between plants. The microbiome of individual plants plays a role in protection of wood against pathogens (Xiong and Yang 2023) and is a source of biocontrol agents used in the protection of grapevines against grapevine trunk disease, of which Petri disease is a part (Cobos et al. 2022). In grapevines infected with Pc, water stress altered the microbiome by reducing microbial diversity and composition and increasing Pc abundance (Leal et al. 2024), suggesting the microbiome plays a role in Pc establishment and growth within the plant. Variation in the microbiomes of individual plants could similarly influence Pc establishment and growth within the plant, but further investigation is required to determine if the individual plant microbiome is the cause of this variation in pathogen abundance. 106 Several Petri disease fungi have been hypothesized to be latent pathogens, in part due to isolating them from both asymptomatic plant wood tissue and their ability to grow under low osmotic water potentials, which affect nutrient levels in the xylem (Abreo et al. 2011; Edwards and Pascoe 2004; Halleen and Petrini 2003; Lima et al. 2017; Hrycan et al. 2020; Whiting et al. 2001; Zanzotto et al. 2007). This study revealed a correlation between water stress and AM fungal inoculation with increased pathogen abundance in the greenhouse. In the field, all plants contained Pc and overall water stressed plants had greater mortality than non-stressed plants. Symptom development was similarly observed in field plants however was not statistically significant between treatments. These results support the hypothesis that Pc is a latent pathogen and increased pathogen abundance leads to plant mortality in young grapevines under stress (Hrycan et al. 2020; Sieber 2007). To our knowledge, this is the first study to induce young vine decline foliar symptom expression, and sudden vine collapse, with a link between pathogen abundance and grapevine mortality within the first few years after planting. However, the underlying mechanisms for symptom development and pathogen growth remain unclear. Increased initial pathogen inoculation was correlated with an increase in internal necrosis which could lead to heightened risk of severe plant stress. Plants under severe stress could upregulate anthocyanin biosynthesis genes resulting in anthocyanin production in the leaves, and sudden vine collapse during periods of high stress (Dabravolski and Isayenkov 2023; Gramaje et al. 2018; Haider et al. 2017). A combination of plant stress and AM colonization increased Pc abundance, but water stress alone did not. Further work is required to understand the underlying reasons behind Pc growth as stress alone may not be sufficient to induce pathogen growth. Investigation of pathogen and host gene expression under plant stress requires further research for wood 107 pathogens affecting grapevines (Galarneau et al. 2019). Additionally, a link between pathogen growth and AM fungi has been observed in this study and in plants infected with black foot fungi (Holland et al 2019; Vukicevich et al. 2018). Investigation into gene expression in mycorrhizal plants infected with young vine decline fungi is another avenue of research to investigate the underlying mechanisms behind this pathogen growth. The impact of rootstock or cultivar under plant stress on disease development in Pc infection plants is likewise unknown. This study suggests ‘SO4’ may be less susceptible to disease development and mortality compared to self-rooted ‘Merlot’ in natural environments, but further investigation into the factors’ influencing susceptibility is required, particularly under natural field conditions where plants are exposed to various abiotic and biotic stressors. 108 Table 3.1. Analysis of inoculum, irrigation treatment, AM, time and their interactions on leaf chlorophyll, stem water potential, and soil matric potential in year 2 using an analysis of variance of a linear mixed effects model for greenhouse. Leaf chlorophyll Stem water potential Soil matric potential Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Inoculum 4 0.55 0.6964 4 0.386 0.8172 N/A N/A N/A Irrigation 1 3.12 0.0803 1 179.964 <.0001 1 75.308 <.0001 AM 1 0.02 0.8832 1 0.141 0.7093 N/A N/A N/A Time 7 363.13 <.0001 5 137.332 <.0001 96 20.1579 <.0001 Inoculum:Irrigation 4 0.44 0.7806 4 0.244 0.9118 N/A N/A N/A Inoculum:AM 4 0.78 0.5434 4 0.654 0.6274 N/A N/A N/A Irrigation:AM 1 0.6 0.4393 1 0.369 0.5469 N/A N/A N/A Inoculum:Time 28 0.64 0.9263 20 0.646 0.8742 N/A N/A N/A Irrigation:Time 7 5.7 <.0001 5 17.377 <.0001 96 10.7317 <.0001 AM:Time 7 1.38 0.2109 5 1.868 0.1015 N/A N/A N/A Inoculum:Irrigation:AM 4 0.13 0.9694 4 1.05 0.3936 N/A N/A N/A Inoculum:Irrigation:Time 28 1.44 0.0649 20 0.649 0.8717 N/A N/A N/A Inoculum:AM:Time 28 0.8 0.7623 20 1.321 0.1688 N/A N/A N/A Irrigation:AM:Time 7 1.88 0.0696 5 1.397 0.2268 N/A N/A N/A Inoculum:Irrigation:AM:Time 28 0.99 0.4774 20 1.82 0.0206 N/A N/A N/A 109 Table 3.2. Analysis of inoculum, irrigation treatment, AM fungi, year and their interactions on dry pruning weight, node number, and dry root weight using an analysis of variance for greenhouse. Pruning weight Node number Root weight Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Inoculum 4 0.861 0.4884 4 0.365 0.83323 4 1.346 0.2595 Irrigation 1 6.124 0.0141 1 33.797 2.10E-08 1 31.937 2.08E-07 AM 1 1.283 0.2585 1 0.262 0.609 1 0.014 0.9075 Year 1 1094.171 < 2e-16 1 161.244 < 2e-16 4 0.266 0.8991 Inoculum:Irrigation 4 0.917 0.4549 4 0.464 0.76238 4 1.528 0.2013 Inoculum:AM 4 0.429 0.7874 4 0.637 0.63691 1 2.914 0.0915 Irrigation:AM 1 0.489 0.4853 1 0.022 0.88095 4 2.202 0.0755 Inoculum:Year 4 0.676 0.6095 4 1.74 0.14209 N/A N/A N/A Irrigation:Year 1 17.702 3.74E-05 1 45.246 1.44E-10 N/A N/A N/A AM:Year 1 5.643 0.0184 1 1.626 0.20363 N/A N/A N/A Inoculum:Irrigation:AM 4 2.801 0.0268 4 3.803 0.00518 N/A N/A N/A Inoculum:Irrigation:Year 4 0.304 0.8749 4 1.234 0.29725 N/A N/A N/A Inoculum:AM:Year 4 0.892 0.4695 4 1.057 0.37851 N/A N/A N/A Irrigation:AM:Year 1 2.778 0.097 1 3.976 0.04736 N/A N/A N/A Inoculum:Irrigation:AM:Year 4 0.639 0.6351 4 0.998 0.40973 N/A N/A N/A 110 Table 3.3. Analysis of inoculum, irrigation treatment, AM, and their interactions on Pc abundance at harvest, class 1, 2, and total necrosis, and basal surface area, at harvest using an analysis of variance for greenhouse. Inoculum Irrigation AM Inoculum: Irrigation Inoculum: AM Irrigation: AM Inoculum: Irrigation: AM Harvest Pc abundance Df 2 1 1 2 2 1 2 F value 0.989 23.373 4.416 2.463 0.12 5.857 0.401 Pr(>F) 0.3771 7.48E-06 0.0392 0.0924 0.8869 0.0181 0.6711 Class 1 necrosis Df 4 1 1 4 4 1 4 F value 240.757 10.094 3.452 1.221 1.802 1.127 0.315 Pr(>F) < 2e-16 0.00192 0.0658 0.30598 0.13339 0.29066 0.86768 Class 2 necrosis Df 4 1 1 4 4 1 4 F value 66.728 1.576 1.673 2.182 2.144 0.378 1.443 Pr(>F) <2e-16 0.2119 0.1985 0.0755 0.0801 0.5398 0.2246 Total necrosis Df 4 1 1 4 4 1 4 F value 10.473 4.758 1.172 0.434 2.12 0.682 0.94 Pr(>F) 3.04E-07 0.0313 0.2813 0.7842 0.0829 0.4107 0.4438 Basal surface area Df 4 1 1 4 4 1 4 F value 3.98 10.74 1.402 1.028 2.36 0.691 1.476 Pr(>F) 0.00467 0.0014 0.23883 0.39588 0.05761 0.40759 0.21423 111 Table 3.4. Percent class 1, class 2, total necrosis, and mean basal surface area (mm) and standard error recorded at the base of each grapevines at the end of the greenhouse experiment for each level of inoculum and in each level of irrigation treatment in the greenhouse. Treatment Class 1 necrosis Class 2 necrosis Total necrosis Basal surface area NIC 1.28% ± 0.23 b 17.82% ± 1.28 c 19.1% ± 1.32 b 419.09 ± 22.18 ab WIC 0.96% ± 0.11 b 16.13% ± 0.87 c 17.1% ± 0.89 b 352.9 ± 15.54 b 1k 15.91% ± 1.01 a 8.08% ± 0.41 b 23.98% ± 1.1 a 385.1 ± 16.71 ab 5k 17.94% ± 1.02 a 7.66% ± 0.47 ab 25.6% ± 1.28 a 362.49 ± 16.95 ab 25k 15.31% ± 0.95 a 6.39% ± 0.37 a 21.7% ± 1.06 a 431.2 ± 18.75 a I1 9.5% ± 0.93 B 10.95% ± 0.62 A 20.44% ± 0.7 B 416.08 ± 12.48 B I2 11.95% ± 1.14 A 10.91% ± 0.85 A 22.86% ± 0.89 A 364.24 ± 10.7 A For each column, percent necrosis or surface area with the same lowercase letter are not significantly different (p = 0.05) according to Tukey’s significance test. For each column, percent necrosis or surface area with the same uppercase letter are not significantly different (p = 0.05) according to Tukey’s significance test. 112 Table 3.5. Analysis of cultivar, inoculum, irrigation treatment, and time for each year and their interactions on leaf chlorophyll and stem water potential each year, using an analysis of variance of a linear mixed effects model for the field. 2021 2022 2023 Leaf chlorophyll Stem water potential Leaf chlorophyll Stem water potential Leaf chlorophyll Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Cultivar 1 0.682 0.4103 1 6.1463 0.0164 1 6.2507 0.0128 1 8.8888 0.0033 1 26.6147 <.0001 Inoculum 2 1.885 0.1557 2 0.6006 0.5522 2 5.808 0.0032 2 3.3865 0.0364 2 1.6701 0.2053 Irrigation 1 11.104 0.0126 1 9.0363 0.0198 1 0.0442 0.8418 1 7.6669 0.0394 1 0.2106 0.6655 Time 1 25.718 0.0002 N/A N/A N/A 9 4.1585 0.0001 6 43.9486 <.0001 N/A N/A N/A Cultivar:Inoculum 2 3.934 0.0218 2 0.2789 0.7577 2 1.6192 0.1992 2 2.861 0.0603 2 4.1823 0.025 Cultivar:Irrigation 1 0.117 0.7324 1 0.0022 0.9631 1 0.0282 0.8667 1 10.7414 0.0013 1 0.2127 0.648 Inoculum:Irrigation 2 1.545 0.217 2 0.109 0.897 2 0.9113 0.4028 2 3.8964 0.0224 2 0.7898 0.4632 Cultivar:Time 1 6.853 0.0098 N/A N/A N/A 9 0.9983 0.4406 6 3.9422 0.0011 N/A N/A N/A Inoculum:Time 2 0.18 0.8353 N/A N/A N/A 18 0.4735 0.9684 12 0.9748 0.4754 N/A N/A N/A Irrigation:Time 1 1.658 0.2187 N/A N/A N/A 9 0.4684 0.8926 6 3.7167 0.003 N/A N/A N/A Cultivar:Inoculum:Irrigation 2 1.538 0.2186 2 0.5296 0.592 2 2.7338 0.0661 2 1.4448 0.239 2 0.1854 0.8317 Cultivar:Inoculum:Time 2 0.542 0.5828 N/A N/A N/A 18 0.4201 0.9836 12 0.6833 0.7657 N/A N/A N/A Cultivar:Irrigation:Time 1 0.09 0.7647 N/A N/A N/A 9 0.2258 0.9908 6 1.9834 0.0713 N/A N/A N/A Inoculum:Irrigation:Time 2 0.014 0.9864 N/A N/A N/A 18 0.312 0.9973 12 1.1284 0.3412 N/A N/A N/A Cultivar:Inoculum:Irrigation:Time 2 0.047 0.9542 N/A N/A N/A 18 0.3404 0.9953 12 0.2875 0.9906 N/A N/A N/A 113 Table 3.6. Analysis of cultivar, inoculum, irrigation treatment, year and their interactions on dry pruning weight, node number, and dry root weight using an analysis of variance of a linear mixed effects model for the field. Pruning weight Node number Root weight Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Cultivar 1 43.80605 <.0001 1 54.1974 <.0001 1 7.95834 0.0073 Inoculum 2 3.04469 0.0494 2 2.4006 0.0938 2 2.0946 0.1358 Irrigation 1 1.16566 0.3161 1 1.0838 0.3325 1 0.09298 0.7727 Year 3 145.06338 <.0001 2 51.6458 <.0001 2 0.01633 0.9838 Cultivar:Inoculum 2 0.11886 0.888 2 0.7449 0.4764 1 2.43886 0.1259 Cultivar:Irrigation 1 2.25635 0.1343 1 0.0748 0.7849 2 0.75768 0.4751 Inoculum:Irrigation 2 0.75799 0.4697 2 2.205 0.1135 2 0.85775 0.4314 Cultivar:Year 3 16.06971 <.0001 2 6.6503 0.0017 N/A N/A N/A Inoculum:Year 6 0.14534 0.9898 4 0.097 0.9833 N/A N/A N/A Irrigation:Year 3 1.24546 0.2938 2 1.4549 0.2364 N/A N/A N/A Cultivar:Inoculum:Irrigation 2 0.73585 0.4801 2 0.9663 0.3826 N/A N/A N/A Cultivar:Inoculum:Year 6 0.92214 0.4795 4 0.6675 0.6154 N/A N/A N/A Cultivar:Irrigation:Year 3 0.03283 0.992 2 0.0321 0.9684 N/A N/A N/A Inoculum:Irrigation:Year 6 0.79943 0.5712 4 0.5423 0.7049 N/A N/A N/A Cultivar:Inoculum:Irrigation:Year 6 0.93975 0.4671 4 0.3551 0.8402 N/A N/A N/A 114 Table 3.7. Analysis of cultivar, inoculum, irrigation treatment, and their interactions on Pc abundance at harvest, class 1, 2, and total necrosis, and basal surface area, at the end of the experiment using an analysis of variance of a linear mixed effects model for the field. Cultivar Inoculum Irrigation Cultivar: Inoculum Cultivar: Irrigation Inoculum: Irrigation Cultivar: Inoculum: Irrigation Harvest Pc abundance Df 1 2 1 2 1 2 2 F value 3.105 8.737 0.055 1.951 0.425 0.165 0.069 Pr(>F) 0.0853 0.0007 0.8242 0.1547 0.518 0.8487 0.933 Class 1 necrosis Df 1 2 1 2 1 2 2 F value 15.8283 7.28659 0.89884 2.29077 0.8757 0.14904 0.44416 Pr(>F) 0.0003 0.0019 0.3866 0.1137 0.3547 0.862 0.6443 Class 2 necrosis Df 1 2 1 2 1 2 2 F value 2.3194 1.5396 0.0109 0.353 0.7924 3.9304 0.241 Pr(>F) 0.1353 0.2263 0.9209 0.7046 0.3784 0.0272 0.7869 Total necrosis Df 1 2 1 2 1 2 2 F value 14.2663 6.6057 2.309 0.1978 2.3818 1.4639 1.5015 Pr(>F) 0.0005 0.0032 0.1891 0.8213 0.1303 0.2429 0.2345 Basal surface area Df 1 2 1 2 1 2 2 F value 9.642 7.289 1.355 2.876 1.929 0.557 3.067 Pr(>F) 0.0034 0.0019 0.297 0.0675 0.1722 0.5768 0.0571 Trunk diameter Df 1 2 1 2 1 2 2 F value 0.9262 1.9179 0.8388 0.5474 1.2205 0.2345 2.6761 Pr(>F) 0.3414 0.1596 0.4017 0.5825 0.2756 0.792 0.0806 115 Table 3.8. Chi-squared analysis of plant total (reps), plant mortality and symptom expression for each year and cumulatively for each cultivar, level of inoculum x irrigation treatment and level of inoculum overall. 2021 2022 Cumulative Cultivar Inoculum Treatment Reps % Mortality % Symptoms Reps % Mortality % Symptoms Reps % Mortality M WIC I1 6 0 a N/A 6 0 a 0 a 6 0 a I2 7 0 a N/A 7 43 a 14 a 7 43 a Overall 13 0 B N/A 13 23 A 8 A 13 23 B 2.5k I1 6 33 a N/A 4 25 a 0 a 6 50 a I2 7 43 a N/A 4 75 a 0 a 7 86 a Overall 13 38 A N/A 8 50 A 0 A 13 69 A 25k I1 6 17 a N/A 5 20 a 0 a 6 33 a I2 7 43 a N/A 4 25 a 0 a 7 57 a Overall 13 31 A N/A 9 22 A 0 A 13 46 A RS WIC I1 6 0 a N/A 6 0 a 0 a 6 0 a I2 7 0 a N/A 7 14 a 14 a 7 14 a Overall 13 0 A N/A 13 8 A 8 A 13 8 A 2.5k I1 6 0 a N/A 6 0 a 0 a 6 0 a I2 7 0 a N/A 7 29 a 29 a 7 29 a Overall 13 0 A N/A 13 15 A 15 A 13 15 A 25k I1 6 0 a N/A 6 50 a 40 a 6 50 a I2 7 0 a N/A 7 14 a 14 a 7 14 a Overall 13 0 A N/A 13 31 A 25 A 13 31 A Overall WIC I1 12 0 a N/A 12 0 b 0 a 12 0 b I2 14 0 a N/A 14 29 a 14 a 14 29 a Overall 26 0 A N/A 26 15 A 8 A 26 15 A 2.5k I1 12 17 a N/A 10 10 a 0 a 12 25 a I2 14 21 a N/A 11 45 a 18 a 14 57 a Overall 26 19 A N/A 21 29 A 10 A 26 42 A 25k I1 12 8 a N/A 11 36 a 18 a 12 42 a I2 14 21 a N/A 11 18 a 9 a 14 36 a Overall 26 15 A N/A 22 27 A 14 A 26 38 A For each column percent mortality and symptoms between Irrigation within each inoculum and cultivar with the same lowercase letter are not significantly different (p = 0.05) according to Chi-squared test. For each column, percent mortality and symptoms between levels of inoculum within each cultivar with the same upper-case letter are not significantly different (p= 0.05) according to Chi-squared test. 116 Figure 3.1. Scan showing basal necrosis of an inoculated plant (1,2,3) and non-inoculated plant (4,5,6) showing necrosis (1,2), threshold analysis of class 1 necrosis set at 5-45 (2,4), and threshold analysis of class 2 necrosis set at 46-80 (3,6) using ImageJ software. 117 Figure 3.2. Stem water potential recorded at mid-day in 1. Year 2 in the greenhouse, 2. 2022 in the field, 3. Soil matric potential recorded every 15 minutes in the greenhouse for 100% watered (I1) and 50% watered (I2) plants. Significantly different results indicated by * at each time point. 118 Figure 3.3. Pc abundance (copies/ng DNA) at the base of plants at the end of the greenhouse experiment in infected plants under irrigation treatment (I1, I2) x AM fungal inoculation (AM-, AM+) treatments. Treatments with the same letter are not significantly different according to Tukey’s significance test. 119 Figure 3.4. Photographs of young vine decline-like symptom progression in 2022 on 1. August 9th, 2. August 30th, and 3. September 27th for a grapevine infected with Pc subject to irrigation treatment. 120 4 Investigating the interaction between ring nematode infestation and Petri disease development in young ‘Merlot’ grapevines infected with Phaeomoniella chlamydospora. 4.1 Background Petri disease is part of the young vine decline complex, which impacts young grapevines within the first few years after planting. The primary causal agents of Petri disease are Cadophora luteo-olivacea, Phaeoacremonium spp., and Phaeomoniella chlamydospora (Pc), which cause internal necrosis, leading to stunted growth and vine collapse (Gramaje and Armengol 2011; Gramaje et al. 2018). Petri disease fungal infection can occur at various stages throughout the nursery propagation process and through infection of open pruning wounds in the vineyard (Carbone et al. 2022; Gramaje and Armengol 2011; Pintos et al. 2018; Waite et al. 2018). Among the different Petri disease fungi detected in established young grapevines in the vineyard, Phaeomoniella chlamydospora has been found to be the most prevalent and isolated on most occasions from asymptomatic vascular tissue in both nursery material and the vineyard (Abreo et al. 2011; Edwards and Pascoe 2004; Halleen and Petrini 2003; Gramaje et al. 2009; Hrycan et al. 2023). These observations have led me to hypothesize that some Petri disease fungi, including Pc may act as latent pathogens with stress thought to be the trigger transitioning them from endophytic to pathogenic (Hrycan et al. 2020). The underlying cause of any transition from an endophytic to pathogenic lifestyle is unknown; however, it is thought that there is a colonization density threshold for endophytes where, once reached, plant senescence results (Sieber 2007). Similarly, in grapevines it is thought that a colonization density threshold is required for plant mortality, and that plant stress results in increased pathogen growth, leading to plant mortality (Hrycan et al. 2020). 121 In older grapevines, Esca symptoms, also caused by P. chlamydospora and Phaeoacremonium spp., have been associated with climatic conditions. Several studies have shown Esca symptoms to vary year to year, this variation correlated primarily with temperature and rainfall (Marchi et al. 2006; Péros et al. 2008; Surico et al. 2000). However, several abiotic and biotic stress factors, including nematodes, are thought to influence Esca and Petri disease symptom expression (Hrycan et al. 2020). While the effect of plant stress on Petri disease development is unknown in young grapevines, a potential interaction between climatic conditions in older Esca infected grapevines suggests that a relationship between plant stress and disease development may be present for these fungi which may translate to infection in young grapevines as well. Plant-parasitic nematodes have been hypothesized to be a potential biotic stress influencing the transition of latent pathogens in grapevines (Hrycan et al. 2020). Plant-parasitic nematodes, including dagger, ring, root-knot, and root lesion, vary between geographical areas worldwide but are widespread in vineyards (Forge et al. 2021; Nicol et al. 1999; Zasada et al. 2012). These plant-parasitic nematodes feed on the roots, reducing water and nutrient uptake by the plant, resulting in a reduced plant vigour (Nicol et al. 1999). The ring nematode, Mesocriconema xenoplax, is prevalent in Canadian vineyards, reported in 82% of vineyards in British Columbia (BC), Canada (Forge et al. 2021). The ring nematode has been shown to reduce plant vigour in young grapevines (Forge et al. 2020, 2021; Pinkerton et al. 2005; Schreiner et al. 2012a, b; McKenry et al. 2001). In BC, a four-year microplot study reported that, M. xenoplax significantly reduced above-ground biomass, trunk diameter, and root biomass in self-rooted ‘Merlot’ indicating that the ring nematode increases plant stress in Canadian vineyards (Forge et al. 2020). 122 Several factors can influence the impact of ring nematode on plant health, such as nematode population densities and resistance to nematodes (Ferris et al. 2012; McKenry and Anwar 2006; Nicol et al. 1999). However, in a microplot study conducted in BC, Canada, mean ring nematode population densities were between two to five per cm3 of soil depending on rootstock variety (Forge et al. 2020). In addition to the impact of ring nematode on plant health measurements, ring nematodes have been associated with disease development in other fruit crops (Cao et al. 2005; 2006). In a study investigating the interaction between black foot fungi and ring nematodes in grapevines, nematode populations did not differ between plants symptomatic for black foot disease compared to non-symptomatic grapevines, which may indicate no direct relationship between ring nematodes and fungal pathogens (Rahman et al. 2014). However, this study did not investigate the effect of M. xenoplax on fungal pathogen abundance and it is yet unclear what effect ring nematodes may have on disease development in plants infected with Petri disease fungal pathogens. The effect of ring nematode on plant health and its association with disease development in other crops makes M. xenoplax a biotic stress of interest in the development of Petri disease in young grapevines. Arbuscular mycorrhizal fungi are commonly used as a commercial bio-fertilizer to improve plant nutrition and growth, and improve tolerances to stresses in various crops, including grapevines (Trouvelot et al. 2015). In one study, ring nematode infestation decreased the number of fine roots containing AM arbuscules, while in another study ring nematodes reduced arbuscule concentration, indicating AM fungi and ring nematodes compete for resources (Pinkerton et al. 2004; Schreiner and Pinkerton 2008; Schreiner et al. 2012b). AM fungi have been considered as an inoculant of interest for black foot disease in grapevines, but contrary results have been reported on its impact on disease development. In two recent studies, AM fungi 123 colonization of the roots resulted in an increase in pathogen abundance and lesion length in grapevines infected with Ilyonectria spp., the causal agent of black foot disease (Holland et al. 2019; Vukicevich et al. 2018). In contrast, AM fungi were found to decrease disease severity in AM colonized plants infected with black foot fungi indicating they may be beneficial (Petit and Gubler 2006). While opposing results have been reported on the impact of AM fungi in black foot disease, no work has been conducted on the impact of AM fungal colonization in Petri disease development in grapevines. Currently, there is a lack of knowledge on the effect that biotic stress factors may play in Petri disease development in young grapevines. Nematodes are commonly found in vineyards worldwide, but understanding of the interaction between fungal disease and nematode infestation is limited. Similarly, the interaction between AM fungi, nematodes, and fungal wood pathogens requires further investigation to clarify the role AM fungi may play in disease development or suppression. Accordingly, the main objective of this study was to investigate whether the ring nematode M. xenoplax plays a role in Petri disease development in grapevines infected with P. chlamydospora and to determine whether AM fungi can mitigate plant stress and subsequently reduce disease development. To accomplish these objectives, a two-year greenhouse experiment studying the effect of ring nematode infestation and AM fungal colonization on P. chlamydospora pathogen abundance and plant health measurements was conducted. In addition, a three-year field study evaluating the effects of ring nematode infestation on P. chlamydospora abundance and plant health measurements was completed. My hypothesis for both experiments was that the presence of Mx would increase colonization of plant xylem tissues by Pc and/or result in greater levels of necrosis than in vines unexposed to Mx. 4.2 Materials and methods 124 4.2.1 Greenhouse experiment 4.2.1.1 Plant material, inoculum preparation and vacuum inoculation Dormant ‘Merlot’ canes were collected from an experimental vineyard block located at the Summerland Research and Development Centre (SuRDC), BC, Canada in November 2019 and stored in a cold room in the dark at 2.5℃ until used. Phaeomoniella chlamydospora isolate SuRDC-1070, obtained from the SuRDC Plant Pathology fungal collection and initially obtained from Petri disease symptomatic grapevines was used in this study (Úrbez-Torres et al. 2014a). Mycelial plugs were obtained from the collection and grown on potato dextrose agar (PDA, Sigma-Aldrich, St Louis, MO) petri plates at 22℃ in the dark for two weeks, until mycelial colony development. Two mL of sterile distilled water amended with a drop of Tween 80 were pipetted onto the colony and gently scraped with a sterile spoonula to suspend the spores. The spore suspension was then vortexed and adjusted to 25,000, 5,000, and 1,000 spores per 10 µL with a haemocytometer. Dormant ‘Merlot’ canes were pruned to three nodes (approximately 20 cm) and vacuum inoculated by pipetting 10 µL of sterile distilled water amended with a drop of Tween 80, or 10 µL of spore suspensions containing either 25,000, 5,000, or 1,000 Pc conidia onto the base of each freshly cut cane. A vacuum with a pressure of 71.5 mmHg measured with a 06-664-19 Manometer (Fisher Scientific, Waltham, MA) was applied to each cane for 1 s to inoculate the cane with the spore suspension (Rooney et al. 2001). Immediately following inoculation, Pc viability and the sterile water solution were tested by pipetting 10 µL of each suspension onto fresh PDA in triplicate and incubated for one week at 22℃ until mycelial colonies were observed. Immediately following inoculation, the dormant canes were transferred to a turned off greenhouse and placed in 2 L buckets, separated by treatment, containing 750 mL water, changed 125 twice weekly. The canes were rooted on heating mats set to 20℃. At budbreak greenhouse lights were turned on to 16 h a day and temperature was set to 20℃. Once cuttings developed sufficient and strong roots, they were transferred to one-gallon pots containing 3,000 g of pasteurized soil on February 10, 2020. Loamy sand soil was obtained from a fallow field at SuRDC and was pasteurized 2x to temperatures exceeding 72℃ according to manufacturer’s instructions in a LANSA model LA soil pasteurizer (Johnson Machine Company LTD, Burlington, Ontario; Trevors 1996). At planting, half of the pots were amended with 0.25g (3,000 spores) of AGTIV Specialty Crops Powder (Premier Tech Growers and Consumer, Quebec, Canada), containing the AM fungus Rhizophagus irregularis following manufacturer’s instructions to inoculate each pot with one spore per gram of soil. A population of the ring nematode M. xenoplax previously obtained from SuRDC grapevines was initiated and maintained in the greenhouse. Nematodes were transferred to sterile soil containing 50% loamy sand and 50% Sunshine Mix #4 (Sun Gro Horticulture, Agawam, MA) in greenhouse pots containing self-rooted ‘Merlot’ at SuRDC to build up the population prior to this experiment. A ring nematode suspension was made by extracting the nematodes from the soil by first passing a soil-water suspension through a 25-mesh sieve to remove large debris. The suspension was subsequently passed through a 400-mesh sieve, and remaining residue was rinsed into a 50 mL centrifuge tube. The sample was centrifuged for 5 min at 3000 rpm and the supernatant was poured off. A sucrose solution (454g sugar/1L water) was added to each tube and the mixture was re-suspended and subsequently centrifuged for 60 s at 3000 rpm. The resulting supernatant was poured into a 500-mesh sieve rinsed with tap water, and then rinsed into a scintillation vial. An inverted microscope was used to count the number of ring nematodes per sample, and then samples were combined to create a nematode suspension 126 containing 3,000 nematodes / 5mL. Half the pots were inoculated with 5 ml of this suspension by pipetting 2.5 ml of the suspension into each of 2 holes in the soil at the same time AM fungi inoculation occurred. Ring nematodes and AM fungi were lightly watered post-inoculation. 4.2.1.2 Experimental design and plant health measurements. The greenhouse experiment was set up March, 2020 with a factorial treatment design consisting of five levels of inoculum: 25,000 (25k), 5,000 (5k), 1,000 (1k) Pc spores, water inoculated control (WIC), and non-inoculated control (NIC); two levels of AM fungi: inoculated (AM+), and non-inoculated (AM-); and two levels of ring nematode infestation: ring nematode inoculated (Nem+), and non-inoculated control (Nem-). There were seven replicate pots of each of the twenty treatment combinations, arranged in a completely randomized experimental design on the greenhouse bench. Grapevines were watered Mon-Wed-Fri with 500 mL water using one Toro 0.5 GPH pressure compensating emitter (Toro, Bloomington, MI) per pot and grown to one shoot in the first year during 2020 until mid-September when water was gradually reduced in preparation of dormancy. On December 18, 2020, grapevines were transferred to a dark cold storage room set to 2.5℃ and then returned to the same greenhouse compartment on March 5, 2021. In year two (2021), grapevines were grown to three shoots and watered equally on Mon-Wed-Fri with 500mL. Grapevines were fertilized with granular 20-20-20 (Master Plant-Prod Inc., ON, Canada) dissolved in water bi-weekly according to manufacturer’s instructions. Due to COVID19 restrictions leaf chlorophyll content was measured only in year two (2021) on 10 leaves per plant bi-weekly with a SPAD 502 Plus Chlorophyll Meter (Spectrum Technologies, Aurora, IL). Pruning weights and node numbers were collected in both years and root weight was measured upon conclusion of the experiment on October 18, 2021. Roots were washed in water to remove soil immediately after uprooting. Prior to weighing, pruning and root 127 weights were dried at 65℃ until weights were consistent over a 24 h period using an Environmental Chamber model 217502 (Hotpack, Anaheim, CA). Necrosis measurements were taken one cm from the base of the trunk using an Epson Expression 1689 scanner and analyzed with ImageJ version 1.53t as described in Hrycan et al. (2023) with the following modification: Images were set to 8-bit colour and analysis was conducted with a threshold analysis to identify different types of wood shading. Threshold numbers indicate the degree of shading from zero (darkest) to 255 (lightest). For black wood discoloration, classified as class 1 necrosis, the threshold was set from 5-45, while brown wood discoloration, labeled class 2 necrosis was set from 46-80. Total necrosis was calculated by the sum of class 1 and class 2 necrosis (chapter 3.2.1.2) 4.2.1.3 Phaeomoniella chlamydospora absolute quantification Pathogen viability and presence were verified at time of inoculation and time of planting by the random selection of three grapevines 24 h post-inoculation and at time of planting (eight weeks post-inoculation). Two cm sections from 0-18cm were cut for each grapevine and split in half longitudinally. At harvest, a two cm section from 2-4cm from the base of each grapevine was selected. Each section was surface sterilized in 70% ethanol for 30 s, 10% bleach for 1 min, and sterile water 2 x 1 min. Following sterilization, the first section was finely cut with a sterile scalpel and 0.2g of wood was placed in a Qiagen DNeasy PowerSoil DNA extraction tube (Qiagen, Hilden, Germany). The other half of each section was finely cut and placed on PDA amended with 0.1mg/mL tetracycline hydrochloride (PDA-tet) to isolate fungi from the wood (chapter 3.2.1.3; Hrycan et al. 2023). DNA extraction tubes were placed in a FastPrep-24™ (MP Biomedicals, Santa Ana, CA) and macerated at 6.0 m/s x 40 s x2. DNA extractions were conducted according to manufacturer’s instructions. DNA was quantified using a Qubit 1.0 128 Fluorometer dsDNA HS Assay (Thermofisher, Waltham, MA) and Pc quantification was carried out using a Droplet Digital™ PCR (ddPCR) system (Bio-Rad Laboratory, Hercules, CA) with ITS primers with the addition of a probe (Hrycan et al. 2023; Martín et al. 2012). Analysis was conducted as described in Hrycan et al. (2023). 4.2.1.4 Arbuscular mycorrhizal fungal colonization At harvest, new growth roots were collected from the first rep of each treatment, frozen in liquid nitrogen, and crushed with a pestle and mortar. 0.2g of crushed root material was placed in lysing matrix A tubes (MP Biomedicals, Santa Ana, CA) and extracted using a Qiagen DNeasy Powerlyzer Powersoil extraction kit according to manufacturer’s instructions. The Qiagen DNeasy Powerlyzer Powersoil extraction kit was used due to manufacturer discontinuation of the DNeasy Powersoil kit. DNA concentration was calculated with a Qubit 1.0 fluorometer. AM fungal quantification was conducted using ddPCR with AM fungal primers and a probe to determine presence or absence of the AM fungi Rhizophagus irregularis (Kokkoris et al. 2019). A selection of roots from the first rep of each treatment was stained with black Sheaffer ink and visually confirmed via microscope (Vierheilig et al. 1998). 4.2.2 Field experiment 4.2.2.1 Plant material, inoculum preparation and vacuum inoculation Collection of dormant ‘Merlot’ canes from an experimental block at the SuRDC occurred on March 3, 2021. Canes were stored in a cold room at 2.5℃ in the dark until March 18, 2021. Pc (SuRDC-1070) mycelial plugs obtained from the SuRDC plant pathology fungal collection were incubated for four weeks prior to creation of the spore suspensions. Suspensions containing 25k, 5k, 1k, and water amended with a drop of Tween-80 (WIC) were created using a 129 haemocytometer as described in chapter 4.2.1.1. Dormant ‘Merlot’ canes were cut to three nodes (22 cm) and vacuum inoculated from the base on March 18, 2021, as described in chapter 4.2.1.1 with 10 µL of each spore suspension and water control. Suspensions and water were plated on fresh PDA plates in triplicate to verify viability of spores. Plates were incubated at 22℃ for one week until fungal colonies were observed. Post-inoculation, grapevines were immediately transferred to 2 L buckets containing 750 mL of water in the greenhouse. Buckets were set on heating mats at 20℃ and water was changed twice weekly. At budbreak, greenhouse lights were turned on to 16 h per day, and greenhouse temperature was set to 20℃. HAV-4500 80x50 cm, 178L nursery pots were dug into the ground and filled with loamy sand soil at a field plot at the SuRDC. There were sixteen pots positioned in each of four adjacent rows. The spacing (pot centre to pot centre) between-rows and between-plants spacings were 2.5 m x 1.12 m, respectively. The pots were dug to a depth of 45 cm, leaving an approximately 5 cm lip projecting above the soil surface. Basamid® Granular Soil Fumigant was mixed into the soil in each pot at 24.5 g Basamid/pot, watered, and covered with clear plastic. One-month post-application, composite soil core samples were taken from each rep to determine success of fumigation on ring-nematode populations. Nematodes were extracted from 100ml aliquots as described in chapter 4.2.1.1. A cress test was performed to confirm the fumigant had dissipated and the soil was suitable for planting (Kehres et al. 2006). 4.2.2.2 Experimental design and plant health measurements The field experiment was set up May, 2021 in a randomized complete block design. Treatments consisted of a factorial combination of 4 levels of inoculum: 25,000 (25k), 5,000 (5k), 1,000 (1k) Pc spores, and water inoculated (WIC), with two levels of ring nematode: ring nematode inoculated (Nem+) and non-inoculated control (Nem-). The eight treatment 130 combinations were randomized into each of the eight rows (blocks). The experiment was conducted over 3 growing seasons from Spring 2021 to Fall 2023. Grapevines were watered with approximately 4L of water on Mon-Wed-Fri using two Toro 0.5 GPH pressure compensating emitters. Grapevines were equally fertilized with granular 14-14-14 fertilizer (ICL Group LTD, Tel-Aviv, Israel) each spring. Grapevines were grown to one shoot in the first year (2021), three shoots in the second year (2022), and five shoots in the third year (2023). Vine survival, dry pruning weights, node numbers and leaf chlorophyll were measured each year as previously described in chapter 4.2.1.2. Mid-day stem water potential was measured bi-weekly from 1 leaf per plant in year two from July 19 to September 15, 2022, and year three from July 13 to September 7, 2023, with a model 610 pressure chamber (PMS Instrument Co. Corvallis, OR). Leaf chlorophyll was measured on ten leaves per plant in year one on August 17 and September 8, 2021, year two from July 11 to September 26, 2022, and year three from July 13 to September 7, 2023, using a SPAD 502 Plus Chlorophyll Meter. Pruning weights and node numbers were collected each year and dried at 65℃ until weights were consistent over 24 h. In the second week of October each year, three core samples were taken from each pot and nematode extractions were conducted on 100mL of soil as described in chapter 4.2.1.1. Following the final core sample taken on October 12, 2023, plants were uprooted and placed in a cold storage room in the dark set to 2.5℃ on November 14, 2023. Roots were removed and cleaned in fresh water and shoots and node numbers were collected and dried at 65℃. Class 1, 2 and total necrosis measurements were taken at one cm from the base, and a CD-6C” C digital caliper (Mitutoyo Corp. Japan) was used to measure trunk diameter below the first bud (chapter 3.2.2.2). 4.2.2.3 Phaeomoniella chlamydospora absolute quantification 131 Pathogen presence and viability were verified on three randomly selected grapevines from each treatment at 24 h post-inoculation and at time of planting, eight weeks post-inoculation. Isolation of the pathogen onto PDA-tet, DNA extractions, and absolute quantification using ddPCR were conducted as described in chapter 4.2.1.3 using the Qiagen DNeasy Powersoil kit. At harvest necrosis was measured one cm from the base as described in chapter 4.2.1.3. The 0-1 cm, 2-4 cm, and 4-6 cm sections were then surface sterilized and drilled with a 1/8” drill-bit for re-isolation of the pathogen and collection of wood for DNA extractions as described in chapter 3.2.2.3. Wood was placed in lysing matrix A tubes (MP Biomedicals, Irvine, CA), frozen in liquid nitrogen and macerated at 6.0m/s x 40 s in a FastPrep-24™ x3. A Qubit 1.0 fluorometer was used to determine DNA concentration. Due to manufacturer discontinuation of the Qiagen DNeasy Powersoil kit, DNA extractions were conducted with the Qiagen DNeasy Plant Mini kit according to manufacturer’s instructions. Each plant section was quantified and the sum of copies/ng DNA for 0-1, 2-4, and 4-6 cm was used for analysis (chapter 3.2.2.3). Absolute quantification of the pathogen was conducted using ddPCR and samples were standardized and analyzed as described in Hrycan et al. (2023). 4.2.3 Statistical analysis For the greenhouse experiment, a 3-way ANOVA model was used to test the main-factor and interaction effects of Pc inoculum level, AMF inoculation and Mx inoculation on all parameters: class 1, 2 and total necrosis, stem cross-sectional surface area, dry shoot and root weight, node number, and leaf chlorophyll content. A 2-way ANOVA model was used to test the main-factor and interaction effects of Pc inoculum level and AMF inoculation on Mx population at harvest. Analyses were conducted in R version 4.3.0 (R Core Team, 2023). A Shapiro-Wilk test was performed on each parameter to test residuals for normalcy. Nematode population and 132 pathogen abundance data were log transformed and node number data were square-root transformed. Tukey’s Honest Significant Difference test set to 5% significance was used to assess differences between Pc inoculum levels and interaction effects. Field experiment data were analyzed using a 2-way randomized complete block ANOVA model. The analyses were conducted using a linear mixed effect model from the nlme package in R (Pinheiro and Bates 2000), and type 3 sums of squares were used to assess effects. Blocks were entered into the model as random factors, and sample date was analyzed to determine time variation and pooled if no time variation occurred. Residuals were inspected visually, and log transformations were performed on nematode population data and pathogen abundance data while square root transformations were performed on node number data prior to final analyses. Significant effects were examined with a type 3 sums of squares. Tukey’s Honest Significant Difference test set to 5% was used to analyze significant interactions using the emmeans package in R version 4.3.0 (Lenth 2023; R Core Team, 2023). A chi-square test to determine the effect of initial level of Pc inoculum and ring nematode infestation on plant mortality in 2021. 4.3 Results 4.3.1 Greenhouse 4.3.1.1 Plant health measurements At harvest mean ring nematode population increased to 696 nematodes per 100mL of soil of Nem+ pots, while ring nematodes were not present in the soil of Nem- pots. Neither AMF nor Pc inoculum level affected ring nematode population size (Table 4.1). 133 Leaf chlorophyll content in year two varied over time with leaf chlorophyll values between 36.08 to 37.98 (Table 4.2), and ring nematodes decreased leaf chlorophyll content in May 26 and June 15 (Table S4.1). There was a significant main-factor effect of Nem on pruning weights (Table 4.3) which were reduced by 7.06% in Nem+ plants compared to Nem- plants (Table S4.2). The average number of nodes per vine for Nem+ plants was 10.23% lower than in Nem- plants in the first year but not in the second year (Table S4.2). A post-hoc test revealed no significant interactions between AM treatments. Dry root weight at harvest was not significantly different between treatments (Table 4.3; S4.2). Droplet digital™ PCR analysis of fresh roots revealed AM colonization in 27% of AM+ plants, and 0% colonization in AM- plants. Visual confirmation by electron microscope verified the presence of AM fungi in AM+ plant roots and absence of the fungus in AM- plant roots. 4.3.1.2 Quantification of Phaeomoniella chlamydospora Pathogen viability plates at time of inoculation confirmed growth of Pc on PDA from 25k, 5k, and 1k spore suspensions, and no growth in the WIC suspension. At time of inoculation, ddPCR detected mean pathogen abundance of 11.31 copies/ng DNA in 25k inoculated plants and did not detect Pc in 5k, 1k, WIC, or NIC. Pc abundance was not significantly different between inoculated plants at time of inoculation (p value = 0.0906). Re-isolation of the pathogen at time of inoculation revealed 100% of 25k and 5k, 66% of 1k, and 0% of WIC and NIC positive for pathogen growth. At time of planting, ddPCR detected mean pathogen abundance of 4,997.16 copies/ng DNA in 25k, 4,848.6 copies/ng DNA in 5k, 6,398.51 copies/ng in 1k, 140.66 copies/ng DNA in WIC, while Pc was not detected in NIC plants. Pc abundance in inoculated plants was not 134 significantly different at time of planting (p value = 0.81) but was significantly higher in Pc inoculated than WIC and NIC plants. Re-isolation of the pathogen at time of planting found 100% of 25k, 5k, and 1k, and 0% of WIC and NIC positive for pathogen growth. At harvest Pc was detected in all inoculated plants and was not detected in WIC or NIC plants via ddPCR or pathogen re-isolation. An ANOVA indicated an interaction between nematode infestation and AM fungal inoculation (Table 4.4). Pc inoculated Nem+/AM+ plants had higher pathogen abundance than Pc inoculated Nem-/AM+ plants (Figure 4.1). Re-isolation of the pathogen onto PDA-tet at harvest found 100% of 25k, 5k, 1k and 0% of WIC and NIC plants. 4.3.1.3 Trunk measurements Pathogen inoculation influenced class 1 and 2 necrosis, and basal surface area (Table 4.4). Class 1 necrosis was significantly greater in inoculated plants compared to WIC and NIC plants, but did not vary between levels of Pc inoculum, while class 2 necrosis was significantly greater in WIC and NIC plants compared to Pc inoculated plants. Total necrosis did not differ between levels of inoculum. Basal surface area was lower in WIC plants than 25k, 5k, and 1k Pc inoculated plants, while NIC plants had decreased mean basal surface area compared to 25k and 5k inoculated plants (Table 4.5). 4.3.2 Field 4.3.2.1 Plant health measurements Ring nematode population varied by year (Table 4.1), increasing in population from 29 nematodes/100mL of soil in 2021 to 66 in 2022 and 258 in 2023. Ring nematode population did not differ due to level of Pc inoculation. In 2023, second-stage juveniles of the northern root- 135 knot nematode, Meloidogyne hapla, were detected in the soil of all pots and population level was found to vary by Pc inoculum level (Table 4.1). Northern root-knot nematode population in the soil was significantly higher in pots containing WIC inoculated plants compared to 25k or 1k Pc inoculated plants (Figure 4.2). Leaf chlorophyll content varied by time in 2021, by time and by ring nematode infestation in 2022, and by ring nematode infestation and Pc inoculum level in 2023 (Table 4.6). Leaf chlorophyll content was 3.45% lower in Nem- plants compared to Nem+ plants in 2022, but leaf chlorophyll values were 4.48% higher compared to ring nematode inoculated in 2023 (Table S4.3). A post-hoc test found no effect of inoculum on leaf chlorophyll content in 2023. Stem water potential varied by time and ring nematode infestation in 2022 and by time in 2023 (Table 4.6). In 2022, stem water potential was 5.02% lower in Nem+ plants compared to Nem- plants (Table S4.1). Dry shoot weight and node number varied between years but was not impacted by treatment (Table 4.7; S4.4). Root weight and trunk diameter at harvest were not significantly different between plants in any treatment group (Table 4.7; S4.4). 4.3.2.2 Quantification of Phaeomoniella chlamydospora Viability plates of Pc spore inoculum at time of inoculation confirmed growth of the pathogen on PDA from 25k, 5k, and 1k spore suspensions, while no growth was observed from WIC suspension plates. At time of inoculation, ddPCR absolute quantification revealed significantly higher pathogen abundance in 25k plants (63.64 copies/ng DNA) than 5k (3.6 copies/ng DNA) or 1k and WIC plants (0 copies/ng DNA). At time of inoculation, pathogen re- 136 isolation onto PDA-tet was successful in 100% of 25k, 5k, and 1k inoculated plants and 0% of WIC plants. At time of planting, ddPCR pathogen quantification found mean copies/ng DNA in 25k plants was 15,120, 5k plants was 5,934, 1k plants was 6,439, and WIC was 0. Pathogen abundance between 25k, 5k, and 1k plants was not statistically different but pathogen abundance in inoculated plants was significantly higher than WIC plants. Pathogen re-isolation from the wood was successful in 100% of 25k and 5k plants, 66% of 1k plants, and 0% of WIC plants. At harvest inoculated plants had significantly higher pathogen abundance than WIC plants, which contained detectable levels of Pc in 13/15 plants. There was no significant difference in pathogen abundance between inoculum levels at the end of the experiment (Figure 4.3). Pc was re-isolated from 92% of 25k, 5k and 1k plants, and 27% of WIC plants. 4.3.2.3 Trunk measurements Level of inoculum interacted with class 1, 2 and total necrosis at harvest, but basal surface area and trunk diameter had no significant interactions (Table 4.8). Percent class 1 and total necrosis were significantly higher in Pc inoculated plants than WIC plants, while 25k inoculated plants had greater class 2 necrosis than WIC plants (Table 4.5). 4.3.2.4 Plant mortality Plant mortality occurred in the winters of 2022 and 2023. Inoculum level nor nematode inoculation was correlated with mortality in 2022 or 2023 but mortality was higher in 1k Pc inoculated plants than WIC controls, and in 5k/Nem+ plants compared to 5k/Nem- cumulatively (Table 4.9). 137 4.4 Discussion In this study ring nematode population at harvest in both the greenhouse and field were not altered by initial pathogen abundance, suggesting ring nematodes may not benefit from Pc infection in young ‘Merlot’ grapevines. These findings are comparable to Rahman et al. (2014) who did not detect differences in ring nematode population levels between symptomatic and non-symptomatic grapevines infected with Ilyonectria spp. Similarly, ring nematode populations in the greenhouse did not differ due to AM fungal inoculation which contrasts with previous studies reporting a detrimental impact on AM fungi due to ring nematode infestation (Pinkerton et al. 2004; Schreiner and Pinkerton 2008; Schreiner et al. 2012b). However, the low level of AM fungal colonization in this study may have reduced any significant interactions between ring nematode population and AM fungal colonization. Root colonization by AM fungi may have been reduced due to optimal watering and fertilization of the plants in combination with the low level of plant stress achieved. My hypothesis was that M. xenoplax would impact plant growth measurements, increase the level of colonization by Pc and/or increase the amount of necrosis in vine stems. In my greenhouse experiment M. xenoplax reduced leaf chlorophyll on May 26 and June 15 measurement dates and reduced pruning weights and node numbers in the first year. In my field experiment M. xenoplax reduced stem water potentials and leaf chlorophyll in one year but I was not able to measure effects on plant biomass measurements. In the greenhouse, Pc inoculated Nem+/AM+ plants had higher pathogen abundance than Nem-/AM+, but M. xenoplax did not affect pathogen abundance in the field, nor did M. xenoplax impact internal trunk necrosis in either experiment. The impact of M. xenoplax on pathogen abundance in the greenhouse is comparable to the impact of irrigation treatment reported in chapter 3 which showed 50% 138 irrigation AM+ plants had higher pathogen abundance than all other irrigation and AM treatments. The effect of ring nematode infestation on pathogen abundance was less pronounced than irrigation treatment, however, irrigation treatment had greater effects on plant growth measurements, indicating the plants were under greater stress due to irrigation treatment than ring nematode infestation. The lack of effects of M. xenoplax on vine growth parameters in my field microplot experiment contrasts with previous field microplot studies in Oregon (Pinkerton et al. 2004) and BC (Forge et al. 2020), in which ring nematodes did reduce vine growth. However, the previous studies involved larger M. xenoplax population densities and were conducted over longer time periods than my study. For example, in the BC study (Forge et al. 2020) effects on pruning weights and trunk diameters were observed after four years of growth in soil with average M. xenoplax population densities of 367 nematodes/ 100 ml soil whereas my microplot experiment measured growth over three years in soil with an average M. xenoplax population density of 118 nematodes/ 100 ml soil population densities in the three years of my study were 29, 67, and 258 nematodes/100mL soil. Variation between the effects of ring nematode infestation on plant health may be due to ring nematode population density differences which can impact symptom severity (Pinkerton et al. 1999, 2004). Greenhouse ring nematode population density at the end of the experiment was greater at 696 nematodes per 100mL of soil, however, the experiment was conducted over a shorter period and greenhouse plants were not subject to the same external factors as field plants. Importantly, the vine root systems were likely pot-bound in the second year of the experiment which could have masked effects of M. xenoplax on vine growth in the second year. Additionally, greenhouse plants were pruned to fewer shoots each year, resulting in less above-ground biomass compared 139 to field plants, which may have influenced the impact of ring nematode infestation on plant health in the greenhouse. Although M. xenoplax infestation did not reduce vegetative vine growth in the field microplots, I did measure effects on stem water potentials. A decrease in stem water potential may indicate ring nematode infestation results in increased plant susceptibility to other stress factors in young grapevines. Mean mid-day stem water potentials of control vines ranged from -0.77 MPa to -0.80 MPa in 2022 to 2023 indicating that the vines were overall sufficiently well-watered (Deloire et al. 2020). While the reduction in stem water potentials caused by ring nematodes was small in magnitude, it indicates that the presence of nematodes could exacerbate water stress caused by inadequate irrigation or other factors. Furthermore, incremental water stress caused by ring nematodes could indicate incremental reduction in photosynthesis. Such reductions, when integrated over multiple years, could have important cumulative impacts on vine carbon allocation and suggest the likelihood of longer-term impacts. I am unaware of any previous studies of the impact of M. xenoplax on grapevine water relation is however, similar studies have been conducted in tree fruits. In peach trees, no variation in stem water potential due to ring nematode infestation was reported (Cao et al. 2006), in contrast with this study. Phenotypical measurements in both the greenhouse and field show ring nematode infestation had an impact on plant health in the greenhouse and in the field. The effect of ring nematode infestation on mid-day stem water potential indicates a relationship with plant water relations and may increase plant susceptibility to other stress factors. Similarly, Pc infection is well-known to cause internal necrosis in plants which could increase susceptibility to hydraulic failure under high stress conditions (Bortolami et al. 2019, 2023; Gramaje and Armengol 2011; Gramaje et al. 2018). 140 Unusual environmental stresses also could have masked effects of M. xenoplax on vine growth and Petri disease development in the field microplot experiment. Abnormally sudden drops in temperature in winters of 2021-22 and 2022-23 resulted in overwinter vine mortality, Chi-square analyses of the proportional vine mortality indicated that winter kill occurred at a higher frequency in 1k Pc inoculated plants compared to WIC plants and in 5k/Nem+ plants compared to 5k/Nem- suggesting a relationship between Pc inoculum level and ring nematode inoculation with plant mortality. A combination of Pc infection and ring nematode infestation may have made the plants more susceptible to winter kill. In the third year of the field experiment, second-stage juveniles of the northern root-knot nematode, Meloidogyne hapla were found in the soil of all field pots. Because the M. hapla contamination was not detected until the end of the third growing season, we speculate that it had overall minor impacts on the outcome of the experiment, but it could also have masked effects of the intentional M. xenoplax infestation, minimizing the magnitude of M. xenoplax effects. Previous work has reported no correlation between Pc abundance and internal necrosis in nursery material, but the effect initial Pc abundance has on internal necrosis development after planting is unclear (Hrycan et al. 2023). In both the greenhouse and field, Pc inoculated plants had greater class 1 necrosis compared to non-inoculated plants. A correlation between necrosis and Pc infection contrasts with Hrycan et al. (2023), where no correlation was reported, however, the methodology applied in this experiment differs by the distinction between black necrosis and browning of the wood. Total necrosis in the greenhouse did not correlate with infection, which supports previous findings on internal necrosis in nursery material, however, in field plants total necrosis was correlated with infection (Carbone et al. 2022; Hrycan et al. 2023). Correlation of total necrosis in the field may be due to the longer time in which the plants were 141 grown compared to the greenhouse, allowing for increased pathogen growth and internal necrosis development. In addition, pathogen abundance varied widely within nursery material and grapevines often contained several pathogens, while this study used known inoculated amounts of one pathogen into each grapevine, which may have impacted necrosis development compared to necrosis observed in ready-to-plant nursery grapevines (Hrycan et al. 2023). In contrast to class 1 necrosis, in the greenhouse, class 2 necrosis was greater in WIC and NIC plants compared to Pc inoculated, which differs from the field where class 2 necrosis was greater in 25k inoculated compared to WIC plants. While class 2 necrosis was greater in 25k inoculated plants in the field, this may have been due to the initial high pathogen load inoculated into the plant, as no effect was observed at lower inoculum levels or in the greenhouse. The results on class 2 necrosis supports previous work indicating browning of the wood is not related to young vine decline fungal infection and may be associated with rootstock or nursery processes (Carbone et al. 2022; Hrycan et al. 2023; Lade et al. 2022). The findings on class 1, 2, and total necrosis support the classification of necrosis into black necrosis and browning of the wood for the evaluation of Pc infected grapevines. While a Pc inoculum level and ring nematode infestation had an impact on plant mortality, there was no relationship between nematode infestation of pathogen abundance in the field, indicating that mortality may be due to factors unrelated to pathogen growth. Internal necrosis was greater in Pc inoculated plants compared to WIC and non-inoculated plants, suggesting a combination of effects caused by Pc and ring nematodes individually increases plant susceptibility to mortality within the first few years after planting. Arbuscular mycorrhizal fungi are commonly applied as a bio-fertilizer to improve plant health, increase tolerance to stress, reduce nematode infestation, and increase resistance to fungal 142 pathogens (Dey and Ghosh 2022; Trouvelot et al. 2015). However, ring nematodes have been shown to be antagonistic to AM fungi in grapevines (Pinkerton et al. 2004; Schreiner and Pinkerton 2008; Schreiner et al. 2012b). Similarly, conflicting results have been reported for the effect of AM fungal inoculation on black foot fungal pathogens in grapevines (Holland et al. 2019; Petit and Gubler 2006; Vukicevich et al. 2018). Petit and Gubler (2006) reported AM fungi to be beneficial in grapevines infected with black foot fungi, but limited evidence is available for the interaction between AM fungi and Petri disease development. Additionally, recent reports of the negative effects of AM fungal inoculation on black foot disease development suggest further work is required to clarify the potential role AM fungi has in the suppression or promotion of disease development in grapevines (Holland et al. 2019; Vukicevich et al. 2018). Ring nematodes are known to reduce AM fungi in grapevines, which may explain the effects observed in this study compared to chapter 3 where an increase in pathogen abundance was observed in infected plants in the 50% irrigation AM+ treatment compared to all treatments (Pinkerton et al. 2004; Schreiner and Pinkerton 2008; Schreiner et al. 2012b). Recent work has revealed high variability in Pc abundance between and within individual grapevines which may have implications on plant health in both the short and long-term (Hrycan et al. 2023). Plant tolerance to pathogen infection is generally thought to decrease as pathogen load increases and it has been suggested that endophyte density may lead to tissue senescence once a colonization density threshold is reached (Pagán and García-Arenal 2018; Sieber 2007). The impact initial pathogen abundance has on plant health in young grapevines after the first few years of planting is unknown. In the greenhouse, at time of inoculation the pathogen was only detected with ddPCR in 25k Pc plants, while in the field 25k inoculated plants had higher pathogen abundance at 5k or 1k. At time of planting in the field pathogen abundance did not vary 143 between Pc inoculum levels. At harvest in the field 25k inoculated was higher than 5k inoculated. No differences in pathogen abundance were observed in the greenhouse at harvest. The findings of this study are inconclusive and further work is required to determine whether pathogen abundance impacts plant health in the short and long term. Previous work has correlated higher Pc abundance at harvest to initial Pc inoculum level (chapter 3). Fungal pathogen abundance has been shown to impact disease severity in Swiss Needle Cast disease in Douglas-fir and brown stem rot in soy, but further work is required to evaluate the impact of pathogen abundance in Petri disease in grapevines over the long-term (Impullitti et al. 2009; Manter et al. 2003). Pathogen abundance varies significantly between ready-to-plant nursery grapevines and has long lasting impacts on Pc abundance within the first few years after planting (chapter 3.3; Hrycan et al. 2023). In this study we found no impact of Pc on above-ground or below-ground plant measurements in the greenhouse or field, which supports the findings in chapter 3.3 that Pc infection does not always lead to immediate symptom development. However, field plants may have been impacted by Pc infection of the control plants. Pc was not detected in field control plants at time of inoculation or time of planting, indicating infection may have occurred after planting. Phaeomoniella chlamydospora has been reported in the Okanagan Valley, British Columbia, Canada, where this study took place and previous studies have reported Pc from the soil and standing water in vineyards, which is one potential avenue of infection (Agustí-Brisach et al. 2013; Rooney et al. 2001; Saccà et al. 2018; Úrbez-Torres et al. 2014a). Additionally, Pc has been found in a high number of ready-to-plant nursery grapevines sold in Canada and infection of the controls could have resulted due to natural infection (Hrycan et al. 2023). Poor 144 detection of Pc at time of inoculation with ddPCR suggests low levels of infection may not be identified until pathogen abundance has increased. Another nematode found widely in BC vineyards is the northern root-knot which can impact plant growth (East et al. 2021; Forge et al. 2021). In the third year of the field experiment, second-stage juveniles of the northern root-knot nematode, Meloidogyne hapla were found in the soil of all field pots. Because the M. hapla contamination was not detected until the end of the third growing season, we speculate that it had overall minor impacts on the outcome of the experiment, but it could also have masked effects of the intentional M. xenoplax infestation, minimizing the magnitude of M. xenoplax effects. Root-knot populations were higher in pots containing WIC plants compared to pots containing 1k and 25k inoculated plants suggesting a relationship between Pc infection and root-knot nematode populations. Root-knot nematodes are a gall-forming endoparasite, extracting resources from the plant (Kantor et al. 2022). It is unclear why root-knot nematode populations were higher in WIC plants, but plant carbohydrate reserves can be reduced in grapevines affected by Esca, which is a different disease in grapevines caused by various fungi including Pc (Fontaine et al. 2014; Petit et al. 2006). If Pc infection results in a reduction in plant carbohydrate reserves in young grapevines as well, root-knot nematode populations could be reduced due to a reduction in available resources. Further work is required to understand the impact of Pc on carbohydrate reserves in young grapevines and whether the northern root-knot nematode influences Petri disease development in young grapevines. Ring nematode infestation increased Pc abundance in the greenhouse but had no effect on plants in the field, however, winter kill was higher in 5k Pc spore inoculated Nem + plants. Further work is required to understand whether ring nematodes increase plant susceptibility to disease and whether high initial ring nematode populations at planting influence Pc abundance 145 and disease development. Lower mid-day stem water potential in nematode infested plants and an increase in internal necrosis due to Pc infection could indicate increased susceptibility to combined stress, such as drought (Fradin and Thomma 2006; Gauthey et al. 2020; López et al. 2022; Yadeta et al. 2013). Investigation into combined stress involving ring nematodes and another abiotic or biotic stress should be conducted to determine whether ring nematode predispose plants to stress, resulting in pathogen growth and disease development under combined stress. While no significant correlation between pathogen abundance and ring nematode infestation was observed, northern root-knot nematode populations were influenced by initial Pc inoculum and examination into the potential interaction between Pc abundance and the northern root-knot nematode is required to understand this relationship. Additionally, significant variability in pathogen abundance was observed in this study, supporting previous findings (chapter 3; Hrycan et al. 2023). Additional investigation into the variability in pathogen abundance between plants is required to understand how this variability in ready-to-plant nursery material may affect Pc infected plants within the first few years after planting. 146 Table 4.1. Analysis of variance summary of inoculum and AM fungi on ring nematode populations in the greenhouse using an analysis of variance and analysis of inoculum, year, on ring nematode populations, inoculum, and stress on root-knot nematode populations in the field using an analysis of variance on a linear mixed effects model. Experiment Nematode Treatment Df F value Pr(>F) Greenhouse Ring nematode Inoculum 4 1.384 0.254 AM 1 0.242 0.625 Inoculum:AM 4 0.379 0.379 Field Ring nematode Inoculum 3 0.502 0.6824 Year 2 27.6481 <.0001 Inoculum:Year 6 0.3167 0.9257 Root-knot nematode Inoculum 3 4.9264 0.0056 Nem 1 0.0168 0.8976 Inoculum:Nem 3 0.3169 0.813 147 Table 4.2. Analysis of inoculum, ring nematode infestation (Nem), AM fungi, time, on leaf chlorophyll content in year 2 of the greenhouse experiment using an analysis of variance of a linear mixed effects model. Df F value Pr(>F) Inoculum 4 0.262 0.9018 Nem 1 2.092 0.1517 AM 1 1.368 0.2453 Time 4 23.314 <.0001 Inoculum:Nem 4 0.346 0.8463 Inoculum:AM 4 2.035 0.0963 Nem:AM 1 2.042 0.1566 Inoculum:Time 16 1.342 0.1692 Nem:Time 4 2.783 0.0267 AM:Time 4 2.148 0.0745 Inoculum:Nem:AM 4 0.021 0.9991 Inoculum:Nem:Time 16 0.647 0.8446 Inoculum:AM:Time 16 1.023 0.4313 Nem:AM:Time 4 1.004 0.4055 Inoculum:Nem:AM:Time 16 0.45 0.9676 148 Table 4.3. Analysis of inoculum, ring nematode infestation (Nem), AM fungi, year, and their interactions on dry pruning weight, node number, and root weight in the greenhouse using an analysis of variance. Pruning weight Node number Root Weight Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Inoculum 4 0.753 0.55701 4 0.879 0.4778 4 0.802 0.527 Nem 1 8.711 0.00358 1 3.543 0.0614 1 0.328 0.568 AMF 1 0.23 0.63233 1 4.34 0.0389 1 0.322 0.572 Year 1 280.935 < 2e-16 1 240.713 <2e-16 4 0.197 0.939 Inoculum:Nem 4 0.435 0.79045 4 0.539 0.7075 4 0.684 0.605 Inoculum:AMF 4 0.635 0.63836 4 0.799 0.5271 1 0.541 0.464 Nem:AMF 1 0.489 0.48541 1 0 0.9883 4 0.712 0.586 Inoculum:Year 4 0.833 0.50607 4 0.392 0.8138 N/A N/A N/A Nem:Year 1 0.008 0.92922 1 5.342 0.0219 N/A N/A N/A AMF:Year 1 0.017 0.89549 1 0.206 0.6505 N/A N/A N/A Inoculum:Nem:AMF 4 1.513 0.20017 4 1.181 0.3207 N/A N/A N/A Inoculum:Nem:Year 4 0.793 0.53083 4 0.537 0.7088 N/A N/A N/A Inoculum:AMF:Year 4 1.175 0.32343 4 1.126 0.3459 N/A N/A N/A Nem:AMF:Year 1 0.585 0.44529 1 0.059 0.8079 N/A N/A N/A Inoculum:Nem:AMF:Year 4 0.477 0.75275 4 0.592 0.6689 N/A N/A N/A 149 Table 4.4. Analysis of inoculum, ring nematode infestation (Nem), AM fungi, and their interactions on fungal abundance, class 1, 2, and total necrosis, and basal surface area at harvest in the greenhouse using an analysis of variance. Harvest fungal abundance Class 1 necrosis Class 2 necrosis Total necrosis Basal surface area Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Inoculum 2 1.797 0.17506 4 85.907 <2e-16 4 68.198 <2e-16 4 1.724 0.151 4 10.568 4.21E-07 Stress 1 0.922 0.3409 1 0.057 0.811 1 1.62 0.2062 1 0.276 0.601 1 0.191 0.663 AMF 1 0.493 0.4854 1 0.092 0.763 1 0.347 0.5575 1 0.009 0.925 1 0.196 0.659 Inoculum:Stress 2 0.534 0.58918 4 0.331 0.856 4 0.218 0.9277 4 0.592 0.669 4 0.088 0.986 Inoculum:AMF 2 1.819 0.17143 4 0.356 0.839 4 2.793 0.0306 4 0.803 0.526 4 0.202 0.937 Stress:AMF 1 7.577 0.00792 1 0.094 0.76 1 2.989 0.0872 1 1.406 0.239 1 0.029 0.864 Inoculum:Stress:AMF 2 0.205 0.81524 4 0.287 0.886 4 0.84 0.5032 4 0.695 0.597 4 1.777 0.14 150 Table 4.5. Percent class 1, class 2, total necrosis, mean basal surface area (mm) and standard error at the base of each grapevine at the end of the experiment containing each level of inoculum in the greenhouse and field. Experiment Inoculum Class 1 necrosis Class 2 necrosis Total necrosis Basal surface area Greenhouse NIC 0.82 ± 0.21 a 23.1 ± 1.46 a 23.91 ± 1.47 a 246.17 ± 17.4 ab WIC 1.74 ± 0.46 a 23.66 ± 1.52 a 25.4 ± 1.54 a 245.45 ± 12.18 a 1k 21.09 ± 1.45 b 8.06 ± 0.68 b 29.15 ± 1.57 a 343.23 ± 26.33 bc 5k 19.81 ± 2.48 b 9 ± 0.76 b 28.81 ± 2.41 a 369.58 ± 26.67 c 25k 17.6 ± 1.51 b 7.23 ± 0.49 b 24.83 ± 1.75 a 424.32 ± 23.38 c Field WIC 1.84 ± 0.63 a 7.78 ± 1.67 a 9.62 ± 2.17 a 443.52 ± 33.42 a 1k 10.97 ± 4.7 b 18.39 ± 5.58 ab 29.36 ± 9.53 b 358.24 ± 43.9 a 5k 10.62 ± 3.05 b 20.99 ± 5.79 ab 31.61 ± 8.54 b 316.99 ± 35.17 a 25k 17.13 ± 6.71 b 22.09 ± 6.54 b 39.22 ± 10.39 b 318.53 ± 40.96 a For each column, percent necrosis or surface area with the same lowercase letter are not significantly different (p = 0.05) according to Tukey’s significance test. 151 Table 4.6. Analysis of inoculum, ring nematode infestation (Nem), and time for each year and their interactions on leaf chlorophyll and stem water potential each year in the field using an analysis of variance of a linear mixed effects model. 2021 2022 2023 Leaf chlorophyll Leaf chlorophyll Stem water potential Leaf chlorophyll Stem water potential Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Inoculum 3 1.284 0.2841 3 0.531 0.6613 3 1.2978 0.2765 3 3.3815 0.0191 3 1.1463 0.3313 Nem 1 0.852 0.3581 1 7.9 0.0053 1 6.4264 0.012 1 8.1394 0.0048 1 0.2347 0.6285 Time 1 14.746 0.0064 7 8.191 <.0001 4 28.0776 <.0001 5 1.082 0.3873 5 8.0227 <.0001 Inoculum:Nem 3 0.164 0.9201 3 2.232 0.0845 3 0.6363 0.5925 3 0.2118 0.8882 3 1.6573 0.1772 Inoculum:Time 3 0.12 0.9484 21 0.459 0.9815 12 0.784 0.6665 15 0.2852 0.9963 15 0.5756 0.8918 Nem:Time 1 0.343 0.5597 7 0.389 0.9088 4 0.588 0.6717 5 0.7274 0.6035 5 0.4758 0.7941 Inoculum:Nem:Time 3 0.442 0.7234 21 0.641 0.8865 12 0.4239 0.9527 15 0.4201 0.9723 15 0.8458 0.6259 152 Table 4.7. Analysis of inoculum, ring nematode infestation (Nem), and year on dry pruning weight, node number, and root weight in the field using an analysis of variance on a linear mixed effects model. Pruning weight Node number Root weight Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Inoculum 3 2.56159 0.0575 3 1.4998 0.2174 3 2.7799 0.0546 Nem 1 2.46077 0.119 1 1.3511 0.2471 1 0.322 0.5738 Year 2 42.37322 <.0001 2 86.4544 <.0001 3 0.7702 0.5181 Inoculum:Nem 3 1.94059 0.1259 3 1.5535 0.2036 N/A N/A N/A Inoculum:Year 6 0.58285 0.7435 6 0.7406 0.6178 N/A N/A N/A Treatment:Year 2 0.40193 0.6698 2 1.4996 0.2269 N/A N/A N/A Inoculum:Nem:Year 6 1.31535 0.2543 6 0.6946 0.6544 N/A N/A N/A 153 Table 4.8. Analysis of inoculum, stress, and their interactions on pathogen abundance, class 1, 2, and total necrosis, basal surface area, and trunk diameter at harvest in the field using an analysis of variance on a linear mixed effects model. Harvest fungal abundance Class 1 necrosis Class 2 necrosis Total necrosis Basal surface area Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Df F value Pr(>F) Inoculum 2 1.797 0.17506 4 85.907 <2e-16 4 68.198 <2e-16 4 1.724 0.151 4 10.568 4.21E-07 Stress 1 0.922 0.3409 1 0.057 0.811 1 1.62 0.2062 1 0.276 0.601 1 0.191 0.663 AMF 1 0.493 0.4854 1 0.092 0.763 1 0.347 0.5575 1 0.009 0.925 1 0.196 0.659 Inoculum:Stress 2 0.534 0.58918 4 0.331 0.856 4 0.218 0.9277 4 0.592 0.669 4 0.088 0.986 Inoculum:AMF 2 1.819 0.17143 4 0.356 0.839 4 2.793 0.0306 4 0.803 0.526 4 0.202 0.937 Stress:AMF 1 7.577 0.00792 1 0.094 0.76 1 2.989 0.0872 1 1.406 0.239 1 0.029 0.864 Inoculum:Stress:AMF 2 0.205 0.81524 4 0.287 0.886 4 0.84 0.5032 4 0.695 0.597 4 1.777 0.14 154 Table 4.9. Chi-squared analysis of plant total (reps) and mortality for each year and cumulatively for each level of inoculum x nematode treatment and each level of inoculum overall. For each column percent mortality between nematode treatment within each inoculum with the same lowercase letter are not significantly different (p = 0.05) according to Chi-squared test. For each column, percent mortality between levels of inoculum with the same upper-case letter are not significantly different (p= 0.05) according to Chi-squared test. 2022 2023 Cumulative Inoculum Treatment Reps % Mortality Reps % Mortality Reps % Mortality WIC Nem - 8 0 a 8 0 a 8 0 a Nem + 8 13 a 7 0 a 8 13 a Overall 16 6 A 15 0 A 16 6 A 1k Nem - 8 25 a 6 33 a 8 38 a Nem + 8 38 a 5 0 a 8 38 a Overall 16 31 A 11 18 A 16 35 B 5k Nem - 8 0 a 8 0 a 8 0 a Nem + 7 38 a 4 0 a 7 43 b Overall 16 19 A 13 0 A 16 20 AB 25k Nem - 8 13 a 7 14 a 8 25 a Nem + 8 13 a 7 0 a 8 13 a Overall 16 13 A 14 7 A 16 19 AB 155 Figure 4.1 Pc abundance (copies/ng DNA) at the end of the greenhouse experiment in Pc inoculated plants with, nematode infestation (Nem+, Nem-) x AM fungal inoculation (AM+, AM-) treatments. Treatments with the same letter are not significantly different according to Tukey’s significance test. 156 Figure 4.2 Mean root-knot nematode populations per 100mL of soil for plants in each level of inoculum treatment. Treatments with the same letter are not significantly different from each other according to Tukey’s significance test. 157 Figure 4.3. Pc abundance (copies/ng DNA) at the end of the field experiment for each inoculum level in ‘Merlot’ grapevines. Treatments with the same letter are not significantly different from each other according to Tukey’s significance test. 01000020000300004000050000600007000080000 1 5 25 i 158 5 Effects of j-rooting and heat stress on Phaeomoniella chlamydospora abundance and Petri disease development in young ‘Merlot’ grapevines. 5.1 Background Young vine decline (YVD) is caused by three diseases, black foot disease, Botryosphaeria dieback, and Petri disease (Agustí-Brisach and Armengol 2013; Gramaje and Armengol 2011; Úrbez-Torres 2011). Symptoms in all three diseases include internal vascular necrosis, which leads to reduced plant growth, lower fruit yield, and premature plant mortality within the first few years after planting (Gramaje and Armengol 2011; Gramaje et al. 2018). Infection of YVD fungi is primarily thought to occur during the nursery propagation process but can also occur in vineyards through aerial inoculum spread via pruning wounds and through the soil via root infection (Carbone et al. 2022; Gramaje and Armengol 2011; Pintos et al. 2018; Waite et al. 2018). Among the many YVD fungi identified, the Petri disease pathogen P. chlamydospora (Pc), has been reported to be highly prevalent in asymptomatic nursery material worldwide (Aroca et al. 2010; Halleen and Petrini 2003; Hrycan et al. 2023), and in asymptomatic young and mature grapevines in vineyards (Abreo et al. 2011; Edwards and Pascoe 2004; González and Tello 2011; Zanzotto et al. 2007). Pathogen abundance in ready-to-plant nursery grapevines has also been reported to vary widely within and between individual plants, the impact of which on plant health is uncertain (Hrycan et al. 2023). With up to 100% infection of Petri disease fungi in nurseries (Hrycan et al. 2023), if infection always resulted in plant mortality, incidence of YVD symptoms and plant and mortality in vineyards would be expected to be much higher. However, in a survey of young vineyards in British Columbia (BC), Canada, 8% of young grapevines surveyed were symptomatic for YVD, a comparatively low number to the reported incidence of infection in nursery material (Hrycan et al. 2023; Úrbez-Torres et al. 2014a). The high incidence 159 of these fungi within nursery material has led to the hypothesis that some of these fungi, including P. chlamydospora, are latent pathogens, increasing in abundance, resulting in internal necrosis and subsequent plant mortality due to plant stress (Hrycan et al. 2020). Several observations support the hypothesis that pathogen growth and symptoms increase under plant stress in the grapevine trunk disease Esca, in which Pc is known to be one of their several causal agents (Gramaje et al. 2018). Phaeomoniella chlamydospora can grow under severe water deficit (Whiting et al. 2001), which may result in an increased access to carbohydrates and nutrients in plants going under water stress (Lima et al. 2017). Additionally, in mature grapevines with Esca, symptom development has been associated with climatic conditions such as low rainfall and high temperature and thus they have been shown to vary year to year (Marchi et al. 2006; Péros et al. 2008; Surico et al. 2000). J-rooting is the result of roots growing sideways or upward due to poor planting conditions such as improper planting or barriers preventing downward root growth and has been suggested as a potential stress factor influencing the transition of latent pathogens into a pathogenic lifestyle (Gramaje et al. 2018; Hrycan et al. 2020). J-rooting has been associated with black foot disease in young grapevines, but understanding of the role j-rooting has on disease development is unclear (Úrbez-Torres et al. 2014b). Similarly, both j-rooting and water deficit have been hypothesized to impact Petri disease development in California (Gubler et al. 2004). The mechanism of how j-rooting may impact Petri disease development is unknown; however, a limitation of the rootzone due to j-rooting of the plant may increase grapevine susceptibility to other types of stress such as water stress, which has been shown to increase P. chlamydospora abundance in young grapevines (Chapter 3; Leal et al. 2024, Preprint). 160 Climate change is expected to increase the occurrence of extreme weather events such as heatwaves in grape-growing regions (Arnell et al. 2019). Grapevines are temperature sensitive (Biasi et al. 2019), and heat and cold stress have been reported as abiotic stress factors that can favour the transition of latent pathogens to a pathogenic phase in grapevines (Hrycan et al. 2020). Several factors have been reported that indicate disease severity could be impacted by temperature stress in grapevines. In mature grapevines, temperature has been associated with Esca symptom development (Surico et al. 2000). Similar reports of increased symptom development under high temperatures have been reported for Eutypa lata, another xylem-colonizing fungal pathogen in grapevines (Sosnowski et al. 2011). Under extreme temperatures, acceleration of plant cell death can occur, creating more optimal conditions for necrotrophic fungi, while simultaneously inhibiting plant defenses, which may result in increased pathogen growth (Desaint et al. 2021; Elad and Pertot 2014). High growth of Pc has been reported up to 30°C, while temperatures reaching 51°C did not induce mortality, indicating this fungus can tolerate extreme temperatures and continue to grow when plant defenses are inhibited due to heat stress. Heat stress can also result, and is often associated with water stress in plants, including grapevines (Songy et al. 2019; Zhao et al. 2021). Additionally, heat stress may result in increased plant water loss, which could increase the likelihood of xylem embolism, which in combination with xylem dysfunction due to internal necrosis caused by Pc infection (Gauthey et al. 2020; López et al. 2022; Marchin et al. 2022) could lead to hydraulic failure and sudden grapevines collapse, a characteristic symptom of YVD (Gramaje et al. 2018). Similarly, Esca has two forms, chronic and apoplectic (Gramaje and Armengol 2011), of which the apoplectic form has been associated with periods of increased temperature and minimal rainfall (Surico et al. 2006). While 161 Esca is a separate disease to Petri, Pc is involved in both maladies, and both show the sudden collapse of the grapevine. However, the effect of temperature stress on Pc in young grapevines requires further investigation. A transition to a pathogenic lifestyle under heat stress may be optimal for growth if a necrotrophic lifestyle is favoured or plant defenses are weakened (Hrycan et al. 2020). In other hosts, increased pathogen growth has been correlated with symptom development (Impullitti et al. 2009; Manter et al. 2003), and in grapevines it has been hypothesized that increased pathogen growth leads to plant mortality once a colonization threshold has been reached, which is the amount the plant can contain before death occurs (Hrycan et al. 2020). In grapevines displaying symptoms of Esca, symptom development has been correlated with internal necrosis (Maher et al. 2012), which could increase due to higher pathogen abundance in grapevines under stress. The role of these different abiotic plant stress factors on pathogen growth and Petri disease development is however unknown. Hence, the main objective of this study was to determine the effect of long-term (j-root) and short-term (heat) stress on Petri disease development. Additionally, a secondary objective of this study was to investigate the effect of initial inoculum level on Petri disease symptom development in grapevines infected with a high (25,000) or low (2,500) initial level of P. chlamydospora conidia in the first few years after planting under natural field conditions. 5.2 Materials and methods 5.2.1 Plant material, inoculum preparation and vacuum inoculation Dormant ‘Merlot’ canes were collected from an experimental block at the Summerland Research and Development Centre (SuRDC) in February 2020 and stored in a cold room in the 162 dark at 2.5°C until pathogen inoculum was prepared. Mycelial plugs containing Pc (SuRDC-1070), obtained from Petri disease symptomatic grapevines in BC, were incubated on potato dextrose agar (PDA; Sigma-Aldrich, St Louis, MO) petri plates in the dark at 22°C for three weeks until mycelial colonies had developed (Úrbez-Torres et al. 2014a). Two mL of sterile distilled water amended with one drop of Tween 80 was pipetted onto the PDA plate and the colony was scraped with a spoonula (chapter 3.2.1.1). The spore suspension was vortexed and adjusted with a haemocytometer to 25,000 (25k) and 2,500 (2.5k) pores per 10 uL (chapter 3.2.1.1). Dormant canes were pruned to three nodes (22 cm) and immediately vacuum inoculated from the freshly cut base of each grapevine by pipetting either 10 uL of sterile distilled water amended with a drop of Tween 80, 2,5k, or 25k Pc spores (chapter 3.2.1.1; Rooney et al. 2001). Vacuum pressure was applied for 1 s per grapevines (chapter 3.2.1.1) and was measured at 71.5 mmHg with a model 06-664-19 Manometer (Fisher Scientific, Waltham, MA). Immediately following the vacuum inoculation procedure, Pc spore viability was tested by plating 10 uL of each inoculum concentration onto fresh PDA in triplicate and incubated for seven days at 22°C until mycelial colonies were observed. Inoculated canes were placed back in the cold room at 2.5°C for 24 h, after which they were dipped in 10% bleach for 1 s and disbudded below the top bud. The base of each dormant cane was dipped in Stim-Root rooting hormone (Premier Tech Home & Garden, Brantford, ON) and layered in separate bins for each treatment containing moist perlite in the dark at room temperature for four weeks. The apical end of the callused grapevines past the top bud was dipped in wax and each grapevine was planted in individual cartons containing a mixture of 50% perlite and 50% sunshine mix #4 (Sun Gro Horticulture, Agawam, MA). Grapevines were placed in the greenhouse set to 20℃ and watered evenly 3x per 163 week while rooting. On July 9, 2020, rooted grapevines were planted in an experimental field plot at SuRDC (chapter 3.2.2.1). 5.2.2 J-Root 5.2.2.1 Experimental Design and Measurements Rooted grapevines were planted at the SuRDC in a vineyard block over four adjacent rows containing loamy sand soil. Row spacing was 2.5m and plant spacing was 0.93m. At planting, j-rooted grapevines were placed in holes and the roots were manually forced in an upward direction (Figure 5.1). The experiment was carried out in a randomized complete block design. Grapevine treatments consisted of a factorial combination of inoculum: water-inoculated control (WIC), 2,500 (2.5k) Pc spores, and 25,000 (25k) Pc spores, and j-root treatment: j-rooted (JR+), and normal rooted (JR-). Treatments contained eight replicates each. Plants were grown to one shoot in year one (2020), and three shoots in years two (2021) and three (2022). In the spring of year four (2023) the plants were not shoot thinned prior to harvest. Plants were equally fertilized each spring with granular 14-14-14 (ICL Group LTD, Tel-Aviv, Israel) according to manufacturer’s instructions. Plants were watered evenly with approximately 12L of water each Monday, Wednesday, and Friday with two Toro 0.5 GPH pressure compensating emitters (Toro, Bloomington, MI) per plant. Mean leaf chlorophyll content was measured on 10 leaves per plant with a SPAD 502 Plus Chlorophyll Meter (Spectrum Technologies, Aurora, IL) and measured in year two on August 17 and September 8, 2021. In year three, SPAD measurements were taken bi-weekly between July 11 to September 20 with a final measurement on September 26, 2022. In year four, a final SPAD measurement was recorded on May 29, 2023. Pruning weights and node 164 numbers were collected each season and dried at 65°C in a Hotpack Environmental Chamber model 217502 (Hotpack, Anaheim, CA) until dry weights were consistent over a 24 h period (chapter 3.2.1.2). Plant mortality was recorded each year and symptom development was recorded in year three. Plants were removed from the field in year four on May 30, 2023, and placed in the dark at 2.5°C for analysis. At harvest, mean trunk diameter below the primary bud of the initial cane was measured 3x with a CD-6” digital caliper (Mitutoyo Corp, Japan). Roots were washed and removed from each grapevine and dried at 65°C until weights were consistent over 24 h as described in chapter 5.2.2.1. After root removal, necrosis measurements were taken one cm from the base, and necrosis measurements were separated into class 1, 2, and total necrosis (chapter 3.2.1.2). 5.2.3 Heat Stress 5.2.3.1 Experimental Design and Measurements Rooted grapevines were planted in the same experimental plot as the j-rooting experiment with alternating complete blocks for each experiment across the four adjacent rows with row x plant spacing (2.5 x 0.93m). The experiment was conducted in a complete randomized block design. Grapevine treatments consisted of a factorial combination of inoculum: water-inoculated control (WIC), 2,500 (2.5k) Pc spores and 25,000 (25k) Pc spores, and plastic film enclosure treatment to increase ambient temperatures: plastic film enclosure (PFE+) and without enclosure (PFE-) containing eight replicates per treatment. Plants were grown to one shoot in year one (2020), and three shoots in years two (2021) and three (2022). In year four (2023) grapevines were not shoot thinned. Plants were fertilized and watered the same as the j-rooting experiment. Plastic film enclosure treatments were set up in the third year on July 27, 2022. PFE+ grapevines were covered with Ultra+ 6mm thick clear plastic (Duchesne, Yamachiche, QC) measuring h: 165 3.6m, w: 0.7m over each grapevine (Figure 5.1). Two rows of two cm holes were cut near the top of each plastic covering measuring 10 cm x 15 cm between holes to allow airflow. Temperatures were monitored by placing five DS1923-F5# - Hygrochron Temperature & Humidity iButtons (Embedded Data Systems, Lawrenceburg, KY) beside PFE+ and five beside PFE- plants. Temperature iButton probes were protected with solar radiation shields. Temperature was recorded every 15 minutes and the hourly mean was recorded from 12:00 AM on July 27, 2022, until 12:00 AM August 1, 2022. The heat treatment started on July 27, 2022, and the plastic coverings were removed in the morning of August 2, 2022. An adjacent weather station recorded temperature data every 15 minutes using a Campbell Scientific CR1000 data logger for comparison with PFE- iButton temperature readings. Mid-day stem water potential was measured immediately prior to the heat stress treatment on July 26, 2022, during the heat stress treatment on July 28, 2022, and immediately following the experiment on August 2, and 9, 2022, with a model 610 pressure chamber (PMS Instruments Co. Corvallis, OR). Mean plant leaf chlorophyll content was measured with a SPAD 502 Plus Chlorophyll Meter on 10 leaves per plant in the second year on August 17 and September 8, 2021, in the third year on July 11, 25, August 4, 9, 16, 25, 30, and September 15 and 26, 2022, and in the fourth year on May 29, 2023. Pruning weights and node numbers were collected each growing season and dried at 65°C as described in chapter 3.2.2.2. Plant mortality was recorded each growing season, and symptom development was recorded in year three. Plants were removed in year four on May 30, 2023, and stored in the dark at 2.5°C. Mean trunk diameter was measured below the primary bud of the initial cane with a CD-6”C digital caliper 3x. Roots were washed in clean water, removed, and dried as described in chapter 3.2.2.2. Necrosis was separated into class 1, 2, and total necrosis and was measured one cm from the base as described in chapter 3.2.2.2. 166 5.2.4 Quantification of Phaeomoniella chlamydospora For both experiments, pathogen presence, abundance and viability were determined in three grapevines from each treatment selected at random at time of planting (13 weeks post-inoculation) as described in chapter 3.2.2.3. Plants were cut into two cm sections between 0-18 cm at time of planting, and surface sterilized via submersion in 70% ethanol for 30 s, 10% bleach for 1 min, sterile water for 1 min x2 (chapter 3.2.2.3). Sections were split in half with one half finely cut with a sterile razor blade and 0.2g were placed in Qiagen DNeasy Powersoil kit (Qiagen Hilden, Germany) tubes. The other half was finely cut and plated on PDA amended with 0.1mg/mL tetracycline hydrochloride (PDA-tet) to isolate the pathogen from the wood (chapter 3.2.1.3; Hrycan et al. 2023). DNA concentration was determined with a Qubit 1.0 Fluorometer dsDNA HS Qubit assay (Thermofisher, Waltham, MA). Pc quantification was conducted with a Droplet Digital™ PCR (ddPCR) system (Bio-Rad Laboratory, Hercules, CA) using ITS Primers (Martín et al. 2012) and a probe (Hrycan et al. 2023) targeting Pc. Pathogen abundance was standardized to gene copies/ng DNA. At harvest, isolation of the pathogen from the wood onto PDA-tet was conducted as described in chapter 3.2.2.3 from 0-1 cm, 2-4 cm, and 4-6 cm from the base. DNA extractions were conducted using 0.1g of wood drilled with a 1/8” sterile drill-bit from the 0-1 cm, 2-4 cm, and 4-6 cm section from the base and placed in Lysing Matrix A tubes (MP Biomedicals, Irvine, CA). The wood was then frozen in liquid nitrogen and macerated 3x in a FastPrep-24™ (MP Biomedicals, Irvine, CA) set to 6.0 m/s x 40 s (chapter 3.2.2.3). DNA was extracted from macerated wood using a Qiagen DNeasy Plant Mini kit according to manufacturer’s specifications (chapter 3.2.2.3). Extraction kits were changed at harvest due to discontinuation of original DNeasy Powersoil kit by the manufacturer. DNA concentration and pathogen abundance 167 were measured as previously described and standardized to copies/ng DNA. The sum of copies/ng DNA from 0-1, 2-4, and 4-6cm was used for analysis. 5.2.5 Statistical Analysis Data analyses were conducted for both experiments to determine the effect of inoculum level and stress treatment on leaf chlorophyll content, dry shoot weight, and node number throughout each growing season and to determine the effect at harvest on pathogen abundance, internal necrosis, trunk diameter, and root weight. The effect of heat stress on plant health during the temperature treatments was determined by stem water potential and air temperature. Air temperature was analyzed with a generalized linear model fit with gamma family to determine the effects of treatment, time, and date on air temperature. A post-hoc Tukey’s Honest Significant Difference test set to 5% was conducted using the emmeans package in R version 4.3.0 (Lenth 2023; R Core Team, 2023). Pruning weight, node number, leaf chlorophyll content, and mid-day stem water potential measured over time were analyzed with a linear mixed effect model fit with REML considering block design and time/year with the nlme package (Pinheiro and Bates 2000). Residuals were inspected visually, and significance was tested with a Type 3 ANOVA. A post-hoc Tukey’s Honest Significant Difference test set at 5% was conducted using the emmeans package in R. Pathogen abundance, internal necrosis, and root weight were analyzed with a linear mixed effect model considering block design. Symptom development and plant mortality for both experiments were analyzed with a chi-squared test to determine the effect of initial level of Pc inoculum and stress treatment on plant health. 5.3 Results 5.3.1 J-root experiment 168 5.3.1.1 Plant health measurements Leaf chlorophyll content varied over time in 2021 and 2022 (Table 5.1). In year three (2022), leaf chlorophyll content was 4.74% lower in 25k inoculated plants than 2.5k (Table 5.2). In 2022, j-rooted plants had 3.52% lower leaf chlorophyll content than normal rooted plants (Table 5.2). Dry pruning weight, node number, and dry root weight were not affected by inoculum or j-rooting (Table 5.3; S5.1). 5.3.1.2 Quantification of Phaeomoniella chlamydospora At time of planting, Pc was not significantly different between WIC, 2.5k, or 25k inoculated plants, and was detected in 100% of plants. Re-isolation of the pathogen from 0-2, 2-4, and 4-6 cm into PDA-tet confirmed the presence and viability of Pc in 100% of 25k, 66% of 2.5k, and 33% of WIC plants. At harvest (2023), initial Pc inoculation and j-rooting had no effect on pathogen abundance (Table 5.4). Mean pathogen abundance ± standard error was 48,137.00 ± 14,535.51 copies/ng DNA in WIC, 39,330.42 ± 6,813.70 copies/ng DNA in 2.5k, and 53,234.28 ± 11,834.19 copies/ng DNA in 25k inoculated plants. Re-isolation of the pathogen from 0-2, 2-4, and 4-6 cm confirmed the presence and viability of the pathogen in 91% of 25k, 100%% of 2.5k, and 50% of WIC plants at harvest. 5.3.1.3 Trunk measurements Basal surface area, trunk diameter, class 1 necrosis, class 2 necrosis, and total necrosis were not significantly different between treatments (Table 5.4, 5.5). 5.3.1.4 Plant symptom development and mortality 169 In 2022, 29% of 2.5k-JR+ plants, 14% of 25k-JR-, and 17% of 25k-JR+ plants developed YVD symptoms, but treatments were not significantly different from each other (Table 5.6). No symptoms were observed in 2.5k-JR- or WIC. Mean leaf chlorophyll content in symptomatic grapevines was 23.81 compared to 28.21 in non-symptomatic grapevines in 2022. Mortality ranged from 13-25% across all treatments in the first year. In the second year and cumulatively, plant mortality occurred in all treatment groups but was not significantly different between treatments (Table 5.6). Four plants that were symptomatic for young vine decline collapsed prior to the conclusion of the experiment and were analyzed separately. Symptoms included reddening of the leaves in August 2022, followed by sudden vine collapse between August and September 2022 (Figure 5.2). Sudden vine collapse occurred in a 25k-JR-, 25k-JR+, and two 2.5k-JR+ plants and were excluded from overall analysis. In these four plants, pathogen abundance was 35,217 copies/ng DNA in the 25k-JR-, 20,376 copies/ng DNA in the 25k-JR+, 35,373 copies/ng DNA in one 2.5k-JR+, and 11,532 copies/ng DNA in the other 2.5k-JR+. Mean dry shoot weight was 16.34g, mean node number was 34.75, and mean dry root weight was 13.38g. Mean percent class 1 necrosis was 33.31%, class 2 necrosis was 35%, and total percent necrosis was 68.3%. Mean basal surface area was 110.36mm. 5.3.2 Heat stress experiment 5.3.2.1 Phenotypical measurements In 2022, 25k plant leaf chlorophyll content was 2.05% and 4% lower than 2.5k and WIC plant leaf chlorophyll content, respectively (Table 5.2). In 2022, PFE+ plant leaf chlorophyll content was 0.17% lower than plant leaf chlorophyll content in PFE- plants (Table 5.2). Leaf 170 chlorophyll content was not impacted by inoculum in 2021 or 2023 (Table 5.7). Stem water potential before the heat stress treatment was LT: -0.83 MPa and PFE+: -0.81 MPa, during stress treatment was PFE-: -1.02 MPa and PFE+: -0.83 MPa, and after PFE-: -0.82 MPa and PFE+: -0.82 MPa. Stem water potential was significantly lower in PFE- plants during stress treatment and was not significantly different from PFE+ plants before and after stress treatment. Maximum temperature in PFE- grapevines was 41.75°C during the stress treatment compared to 49.05°C in PFE+ grapevines. Minimum temperatures recorded during the stress treatment were 19.69°C in PFE- grapevines and 15.68°C in PFE+ grapevines. PFE+ grapevines reached significantly higher hourly temperatures during the day and significantly lower hourly temperatures during the night for the duration of the heat stress treatment (Figure 5.3). Hourly temperature recordings of PFE- iButton probes was higher than the adjacent weather station at several time points during the heatwave (Figure S5.1). Mean temperature difference between PFE- iButton probes and the weather station was 3.37°C for significantly different time points. Dry pruning weights, node numbers and dry root weight at harvest were unaffected by inoculum or heat stress (Table 5.3; S5.2). 5.3.2.2 Quantification of Phaeomoniella chlamydospora Pathogen abundance was not significantly different between treatments at time of planting and was detected in 100% of 25k, 2.5k, and WIC plants. Re-isolation of the pathogen onto growing media confirmed presence and viability of the pathogen in 100% of 25k and 2.5k plants, and in 33% of WIC inoculated plants. Phaeomoniella chlamydospora abundance at harvest was influenced by initial inoculum level (Table 5.4). Mean pathogen abundance ± standard error was 580,848.9 ± 409,950.2 171 copies/ng DNA in 25k inoculated plants, which was not significantly different from 102,255.8 ± 71,255.5 copies/ng DNA in 2.5k inoculated plants but was significantly greater than 14,565.3 ± 4,851.1 copies/ng DNA in WIC plants. 2.5k and WIC plant pathogen abundances were not significantly different from each other. Re-isolations confirmed the presence of Pc in 79% of 25k plants, 57% of 2.5k plants, and 54% of WIC plants. 5.3.2.3 Trunk measurements Class 1 necrosis was greater in 25k plants than WIC plants (Table 5.5). Basal surface area was significantly greater in WIC plants than 25k plants (Table 5.5). Class 1 and total necrosis were greater in PFE- plants than PFE+ while basal surface area was lower in PFE- plants than PFE+ (Table 5.5). Class 2 necrosis was not significantly different between inoculum or heat stress treatments (Table 5.4). 5.3.2.4 Plant symptom development and mortality In 2022, 38% of 2.5k-PFE+ and 43% of 25k-PFE- plants developed YVD symptoms, while none of the WIC, 2.5k-PFE- or 25k-PFE+ plants were symptomatic. Symptom development was not statistically significant between treatments (Table 5.6). In symptomatic grapevines, mean leaf chlorophyll content was 22.51 compared to 29.45 in asymptomatic plants. Plant symptoms included reddening of the leaves in August of 2022 after the heat stress treatment ended. In 2021 and 2022 plant mortality occurred but was not significantly different between treatments (Table 5.6). Plant mortality was significantly greater in 25k plants than WIC plants in 2022 and cumulatively (Table 5.6). In 2022, 23% of PFE- plants died compared to 32% of PFE+ plants, while symptom development occurred in 14% of PFE- and PFE+ plants. 172 Two 2.5k-PFE+ plants that displayed symptoms of YVD collapsed before conclusion of the experiment in 2022 and were excluded from the overall analysis. Pathogen abundance in the two plants was 21,454.5 and 30,223.9 copies/ng DNA. Mean dry shoot weight was 2.24g, mean node number was 19.5, and mean dry root weight was 13.91g. Mean percent class 1 necrosis was 44.58%, mean percent class 2 necrosis was 29.92%, and mean percent total necrosis was 74.5%. Mean basal surface area was 101.22mm. Symptom development occurred after heat stress treatment was applied in summer of 2022, and grapevine collapse occurred between August and September 2022 (Figure 5.2). 5.4 Discussion This study compares how long-term j-rooting stress and short-term heat stress influence the growth of Pc and Petri disease development in young grapevines within the first four years after planting. J-rooting is thought to impact Petri disease development (Gubler et al. 2004) but its role is unclear (Úrbez-Torres et al. 2014b). It has been suggested that j-rooting can lead to plant stress, resulting in a change to a pathogenic lifestyle by Pc (Hrycan et al. 2020). In this study, j-rooting had a minimal impact on plant health measurements, reducing leaf chlorophyll content in 2022, but plant growth measurements were unaffected. Leaf chlorophyll may be reduced due to reduced nutrient uptake due to j-rooting, but nutrient uptake was not analyzed in this experiment. Similar findings have been reported in loblolly and eastern white pine, showing j-rooting had negligible effects on plant growth and stem water potential within the first three years (Seiler et al. 1990). In contrast to the long-term stress caused by j-rooting, heat stress can be caused by short-term heatwaves, disrupting photosynthesis, and increasing transpiration rates, and leading to increased water requirement by the plant (Venios et al. 2020). In this study, daily temperatures in 173 PFE+ plants exceeded 45℃, which can cause a long-lasting reduction in photosynthetic capability (Kadir and Von Weihe 2007), which indicates the level of heat exposure was extreme which could result in high stress for the plants. Additionally, temperatures varied more widely in enclosed plants due to lower overnight temperatures. The effect of this wider variation in temperature, along with alterations in CO2, light exposure, lack of wind, and humidity due to enclosure treatment could have altered plant responses due to the sudden change in environmental conditions. Plastic sleeves placed over grapevines had similar results as reported by Bowen et al. (2004). Mid-day stem water potential was lower in PFE- plants than PFE+ during the heat stress treatment and returned to similar levels as PFE- plants within one week after the heatwave treatment was removed, while leaf chlorophyll content was negligibly impacted. Higher mid-day stem water potential suggests stomatal conductance was reduced in PFE+ plants compared to PFE-, as has been observed in grapevines when temperatures exceed 40℃ to 45℃ (Luo et al. 2011). However, the heat stress treatment had no long-term recorded impact on plant growth measurements or on mid-day stem water potential. Plant health and growth measurements were unaffected by Pc infection in both the j-rooting and heat stress experiments across all years, supporting the findings of chapter 3 and 4 on the limited impact of Pc infection alone on plant health. Petri disease fungi are often found in asymptomatic nursery material (Carbone et al. 2022; Halleen and Petrini 2003; Hrycan et al. 2023) and have been isolated from asymptomatic young and mature grapevines (Abreo et al. 2011; Gainza-Cortes et al. 2020; González and Tello 2011; Zanzotto et al. 2007). The limited impact on plant health observed suggests that Pc can remain asymptomatic or cause limited phenotypical symptoms within the first few years. 174 Within one week following the heatwave, which occurred from July 27 to August 1, 2022, plants begun displaying symptoms of YVD. In the heat stress experiment, WIC plants were asymptomatic, while 2.5k-PFE+ and 25k-PFE- plants were symptomatic. Symptom development in the j-rooting experiment was comparable to the heat stress experiment. The treatment groups 2.5k-JR+ and 25k-JR+/JR- plants were symptomatic for YVD while WIC and 2.5k-JR- plants remained asymptomatic. Observed symptom development suggests an association between high initial pathogen abundance and plant stress. Pathogen abundance has been associated with symptom development in other hosts (Impullitti et al. 2009; Manter et al. 2003). Similarly, high temperatures and drought have been associated with Esca symptom development and apoplexy (Surico et al. 2006). While Esca is a separate disease in grapevines, Pc involvement in Esca disease has been widely reported (Gramaje et al. 2018) and the pathogen could respond similarly to stress in both young and old grapevines. Additionally, high temperatures can result in increased symptoms in grapevines infected with Eutypa lata (Sosnowski et al. 2011), another xylem-colonizing fungal pathogen of grapevines (Gramaje et al. 2018). Symptom development can precede the collapse of the grapevine (Gramaje and Armengol 2011; Gramaje et al. 2018), which was observed in this study. Of the symptomatic plants, four heat stress and two j-rooting plants collapsed in August and September 2022, while several symptomatic and non-symptomatic grapevines survived until dormancy but did not survive the winter. In 2022, one 2.5k-PFE- plant died while no mortality was recorded in any other WIC PFE-/JR- or 2.5k JR- plants, compared to plant mortality in three of each 2.5k PFE+/JR+ and one of each WIC PFE+/JR+ which suggests plant stress may have played a role in mortality. Water stress has been correlated with higher mortality in Scots pine infected with 175 Heterobasidion annosum, a wood fungal pathogen (Gomez-Gallego et al. 2022). It was suggested that H. annosum reduces carbon reserves resulting in plant susceptibility to stress (Gomez-Gallego et al. 2022). In grapevines, Pc may have greater access to carbohydrates under water stress (Lima et al. 2017), which could indicate similar effects on carbohydrate reserves. Alternatively, tyloses have been associated with symptom development and apoplexy in older grapevines with Esca disease, which is similarly caused by Pc (Bortolami et al. 2021). An increase in necrosis or tylose formation could result in hydraulic failure as hypothesized by McElrone et al. (2010) in walnut. Restriction of the roots due to j-rooting could result in decreased access to water, while heat stress due to elevated temperatures could increase plant water requirements (Venios et al. 2020). A sudden increase in water requirements may result in hydraulic failure when combined with Pc induced internal necrosis or tylose development due to Pc infection. In contrast to the higher mortality in j-rooted or heat stressed 2.5k inoculated plants, mortality was 50% or greater in 25k plants for both experiments, which may suggest a high initial inoculum level could result in increased internal necrosis and tylose development, resulting in grapevine mortality regardless of stress. In the heat stress experiment, 25k plants had significantly higher mortality than WIC plants, suggesting that initial pathogen level has a long-term impact on plant health and survival. However, sudden drops in temperature in the winters of 2021 and 2022 may have impacted plant mortality or pathogen abundance. Cold damage has been linked to colonization of other hosts by B. dothidea (Schoeneweiss and Wene 1980; Wene 1979) but it is unclear how extreme cold may impact pathogen abundance or plant mortality in grapevines infected with P. chlamydospora. Class 1 necrosis was significantly greater in 25k plants than WIC plants in the heat stress experiment. In both heat stress and j-rooting experiments, 25k plants had greater class 1 and total 176 necrosis than 2.5k plants which were greater than WIC plants. Basal surface area was significantly higher in WIC plants than 25k plants in the heat stress experiment. In the j-rooting experiment, JR+ plants had a lower basal surface area while in the heat stress experiment, PFE+ plants had lower class 1 and total necrosis and greater basal surface area. Phaeomoniella chlamydospora causes internal necrosis (Gramaje et al. 2018) and was higher in 25k inoculated plants compared to WIC plants in the heat stress experiment. Increased internal necrosis has been correlated with Esca symptoms in older grapevines (Maher et al. 2012) and could contribute to increased susceptibility to hydraulic failure (McElrone et al. 2010), particularly in combination with plant stress. Class 2 necrosis was not correlated with plant stress or pathogen inoculation. The results of this study support the findings of chapter 3 and 4 and suggest internal necrosis should be separated into black necrosis (Class 1) and browning of the wood (Class 2) to better understand the effect of Pc infection on the wood (chapter 3.4; 4.4). It is hypothesized that symptom development is due to a transition of Pc from an endophytic phase to a pathogenic, resulting in increased pathogen growth under plant stress (Hrycan et al. 2020). Neither J-rooting nor heat stress impacted Pc abundance at harvest. Pc abundance was similarly unaffected by ring nematode infestation (Chapter 4) and water stress in field plants (chapter 3.3.2.2). In contrast chapter 3.3.1.2 reported increased pathogen growth due to water stress in the greenhouse. It was hypothesized that water stress imposed in the greenhouse was more severe, resulting in increased pathogen growth (chapter 3.4). Similarly, plant growth and health measurements indicate j-rooting and heat stress had minimal long-term impacts on plant health. However, initial Pc quantity did impact Pc abundance at harvest. In the heat stress experiment, plants inoculated with 25k spores had higher pathogen abundance than 177 WIC plants. In contrast, in the j-rooting experiment, at harvest, WIC, 2.5k, and 25k plants all had similar pathogen abundance. Pathogen abundance at harvest in the heat stress experiment suggests initial abundance may have a long-term impact on plant health. Ready-to-plant nursery grapevines have wide variability in pathogen abundance between and within individual plants (Hrycan et al. 2023), and this study suggests plants containing a high level of pathogen abundance may be more susceptible to symptom development and mortality within the first few years after planting. However, other factors may influence pathogen growth within individual plants as Pc abundance varied widely between plants in both the j-rooting and heat stress experiments and pathogen abundance in plants in the j-rooting experiment was not significantly different between treatments. This variability may be due to differences within the microbiome of individual plants (chapter 3.4); however, further investigation into the underlying cause of this variability is required. WIC plants were infected with Pc at time of planting, but the cause of contamination is unclear. Viability plates indicated no contamination in the WIC suspension. However, Hrycan et al (2023) reported high levels of Pc infection in ready-to-plant nursery material sold in Canada and the Pc isolate used in this study was previously obtained from the same geographical region where the ‘Merlot’ canes were collected (Úrbez-Torres et al. 2014a). Infection of the WIC plants may have been due to natural infection, which spread during the propagation process. In a similar study planted in the same geographical location, WIC plants were found to contain Pc at harvest while no infection was present at time of inoculation or time of planting, suggesting Pc is present in the area and may be present in the soil (chapter 4.4). This study suggests a link between initial pathogen abundance, internal necrosis, symptom development, and mortality in young grapevines infected with Pc within the first few 178 years after planting. Further work is required to understand the underlying cause of this mortality in relation to xylem dysfunction and plant stress. Both long-term j-rooting and short-term heat stress appears to affect plant symptom development and mortality, but symptoms appeared only after a heatwave in 2022. Long-term stress and Pc infection resulting in increased internal necrosis may predispose the plant to hydraulic failure when sudden severe stress occurs. Further investigation is required to understand the combined impact of plant stress on grapevines infected with young vine decline fungi and the response of these fungal pathogens to plant stress to understand the relationship between plant stress and symptom development. Additionally, tylose formation has been associated with symptom development and apoplexy in grapevines with Esca (Bortolami et al. 2021) but is less well understood for Petri disease in young grapevines. Petri disease fungi are associated with Esca and understanding whether the lifestyle of these fungi and plant responses to infection are similar in both young grapevines with Petri disease and older grapevines with Esca may help in understanding the underlying cause of sudden grapevine collapse in young grapevines affected by Petri disease. 179 Table 5.1. Analysis of inoculum, stress treatment, time, and their interactions on leaf chlorophyll content each year using an analysis of variance on a linear mixed effect model for the j-root experiment. 2021 2022 2023 f F v lu Pr(>F) f F v lu Pr(>F) f F v lu Pr(>F) Inoculum 2 0.473 0.6252 2 4.042 0.0193 2 2.4527 0.1159 Treatment 1 1.384 0.2435 1 5.627 0.0188 1 0.195 0.6644 Time 1 7.472 0.0292 6 9.557 <.0001 Inoculum:Treatment 2 0.066 0.9366 2 2.381 0.0955 2 1.6324 0.2246 Inoculum:Time 2 0.806 0.4506 12 0.345 0.9793 Treatment:Time 1 6.317 0.0143 6 0.339 0.9154 Inoculum:Treatment:Time 2 0.098 0.9069 12 0.796 0.6546 180 Table 5.2. Mean leaf chlorophyll content and standard error for each level of inoculum and j-rooting or enclosure treatment each year for the j-root and heat stress experiments. Treatment Leaf chlorophyll 2021 Leaf chlorophyll 2022 Leaf chlorophyll 2023 JR 32.97 ± 0.49 28.2 ± 0.66 25.34 ± 1 2.5 32.64 ± 0.47 28.04 ± 0.57 23.07 ± 0.79 25 33.22 ± 0.56 26.71 ± 0.96 22.2 ± 1.43 JR- 32.65 ± 0.52 28.13 ± 0.6 23.24 ± 0.86 JR+ 33.23 ± 0.26 27.14 ± 0.64 B 24 ± 0.93 HS 33.37 ± 0.43 29.24 ± 0.71 25.43 ± 0.76 2.5 34.48 ± 0.72 28.66 ± 0.87 24.76 ± 0.54 25 34.33 ± 0.54 28.07 ± 0.85 26.57 ± 0.82 PFE- 28.67 ± 0.67 25.28 ± 0.57 PFE+ 28.62 ± 0.7 B 25.66 ± 0.64 For each column, percent necrosis or surface area with the same lowercase letter are not significantly different (p = 0.05) according to Tukey’s significance test. For each column, percent necrosis or surface area with the same uppercase letter are not significantly different (p = 0.05) according to Tukey’s significance test. 181 Table 5.3. Analysis of inoculum, treatment, year, and their interactive effects on pruning weight, node number, and root weight for the j-root and heat stress experiments using an analysis of variance of a linear mixed effect model. JR HS Pru i w i ht d um r R t w i ht Pru i w i ht d um r R t w i ht f F v lu Pr(>F) f F v lu Pr(>F) f F v lu Pr(>F) f F v lu Pr(>F) f F v lu Pr(>F) f F v lu Pr(>F) Inoculum 2 0.08484 0.9187 2 0.70653 0.4962 2 0.1241 0.8839 2 0.27028 0.7636 2 0.43703 0.6472 2 1.96004 0.1597 Treatment 1 1.74243 0.1892 1 3.09223 0.0822 1 3.5925 0.0713 1 0.66758 0.4153 1 1.20012 0.2761 1 0.57053 0.4564 Year 3 94.0503 <.0001 2 57.9566 <.0001 2 0.1445 0.8663 3 127.049 <.0001 2 62.2153 <.0001 2 0.03296 0.9676 Inoculum: Treatment 2 0.41278 0.6627 2 0.68027 0.5092 2 0.19306 0.8247 2 0.20319 0.8165 Inoculum: Year 6 2.14949 0.0521 4 2.01061 0.1 6 0.50406 0.8045 4 0.16475 0.9557 Treatment :Year 3 1.0432 0.3758 2 1.16956 0.3153 3 1.09044 0.3554 2 1.03692 0.3585 Inoculum: Treatment: Year 6 0.57073 0.753 4 0.73115 0.5731 6 0.31086 0.9304 4 0.13821 0.9677 182 Table 5.4 Analysis of inoculum, stress treatment, and their interactions on pathogen fungal abundance at harvest, class 1, 2, and total percent necrosis, basal surface area, and trunk diameter using an analysis of variance of a linear mixed effect model for the j-root and heat stress experiments. JR HS culum Tr tm t culum:Tr tm t culum Tr tm t culum:Tr tm t Harvest Pc abundance f 2 1 2 2 1 2 F v lu 0.372 0.498 1.447 4.8679 2.7985 1.6831 Pr(>F) 0.6935 0.4877 0.2569 0.0153 0.1055 0.2041 Class 1 necrosis f 2 1 2 2 1 2 F v lu 1.39673 3.61484 0.13834 4.01604 5.37292 0.69137 Pr(>F) 0.2685 0.0704 0.8716 0.0293 0.028 0.5092 Class 2 necrosis f 2 1 2 2 1 2 F v lu 0.3865 0.6391 0.6048 0.7422 1.2313 0.3887 Pr(>F) 0.6839 0.4326 0.555 0.4852 0.2766 0.6815 Total necrosis f 2 1 2 2 1 2 F v lu 1.4245 2.4716 0.1013 3.4101 4.6904 0.6114 Pr(>F) 0.262 0.1302 0.9041 0.0473 0.039 0.5497 Basal surface area f 2 1 2 2 1 2 F v lu 2.04511 3.57807 0.46663 4.38066 3.08025 0.07002 Pr(>F) 0.1532 0.0718 0.6332 0.0221 0.0902 0.9325 Trunk Diameter f 2 1 2 2 1 2 F v lu 0.093 1.37 0.679 0.0788 0.9887 0.1764 Pr(>F) 0.9117 0.2543 0.5177 0.9244 0.3286 0.8392 183 Table 5.5. Mean percent class 1, 2, total necrosis, mean basal surface area (mm) and standard errors recorded at the end of the experiment for each level of inoculum and stress treatment in the j-root and heat stress experiments. Treatment Class 1 necrosis Class 2 necrosis Total necrosis Basal surface area JR 14.71 ± 4.86 11.79 ± 2.36 26.49 ± 7.09 206.62 ± 23.03 2.5 15.97 ± 5.63 13.28 ± 2.93 29.25 ± 7.26 182.86 ± 19.95 25 20.13 ± 5.28 13.47 ± 1.54 33.6 ± 5.39 152.03 ± 13.33 JR- 14.13 ± 3.99 12.54 ± 2.05 26.67 ± 5.19 199.8 ± 18.78 JR+ 20.06 ± 4.63 13.18 ± 1.8 33.24 ± 5.77 159.37 ± 10.97 HS 9.86 ± 3.44 11.73 ± 2.48 21.59 ± 5.22 251.59 ± 25.89 2.5 17.42 ± 5.67 13.19 ± 2.77 30.61 ± 7.73 193.25 ± 22.21 25 35.64 ± 8.21 14.94 ± 2.67 50.58 ± 9.22 155.06 ± 19.88 PFE- 29.39 ± 6.35 14.04 ± 1.83 43.43 ± 7.27 178.04 ± 20.76 PFE+ 11.82 ± 3.18 B 12.5 ± 2.56 24.31 ± 5.17 B 222.64 ± 18.38 For each column, percent necrosis or surface area with the same lowercase letter are not significantly different (p = 0.05) according to Tukey’s significance test. For each column, percent necrosis or surface area with the same uppercase letter are not significantly different (p = 0.05) according to Tukey’s significance test. 184 Table 5.6. Chi-squared analysis of plant mortality and symptom expression each year and cumulatively for each level of inoculum x stress treatment and inoculum level overall for the j-root and heat stress experiments 2021 2022 Cumulative culum Tr tm t R % M rt lity % Sym t m R % M rt lity % Sym t m R % M rt lity HS PFE- 8 13 7 0 0 8 13 PFE+ 8 13 7 14 0 8 25 Overall 16 13 A N/A 14 7 A 0 A 16 19 A 2.5 PFE- 8 0 8 13 0 8 13 PFE+ 8 0 8 38 38 8 38 Overall 16 0 A N/A 14 25 AB 19 A 16 25 AB 25 PFE- 8 13 7 57 43 8 63 PFE+ 8 13 7 43 0 8 50 Overall 16 13 A N/A 14 50 B 21 A 16 56 B JR JR- 8 25 6 0 0 8 25 JR+ 8 25 6 17 0 8 38 Overall 16 25 A N/A 12 8 A 0 A 16 31 A 2.5 JR- 8 13 7 0 0 8 13 JR+ 8 13 7 43 29 8 50 Overall 16 13 A N/A 14 21 A 14 A 16 31 A 25 JR- 8 13 7 43 14 8 50 JR+ 8 25 6 33 17 8 50 Overall 16 19 A N/A 13 38 A 15 A 16 50 A For each column percent mortality and symptoms between stress within each inoculum and cultivar with the same lowercase letter are not significantly different (p = 0.05) according to Chi-squared test. For each column, percent mortality and symptoms between levels of inoculum within each cultivar with the same upper-case letter are not significantly different (p= 0.05) according to Chi-squared test. 185 Table 5.7. Analysis of inoculum, treatment, time, and their interactions on leaf chlorophyll content and stem water potential for each year of the heat stress experiment using an analysis of variance of a linear mixed effect model. 2021 2022 2023 L f chl r hyll L f chl r hyll St m w t r t ti l L f chl r hyll f F v lu Pr(>F) f F v lu Pr(>F) f F v lu Pr(>F) f F v lu Pr(>F) Inoculum 2 2.443 0.0943 2 7.2688 0.0009 2 2.6005 0.078 2 1.2329 0.3162 Treatment 1 8.0858 0.0048 1 6.6667 0.0109 1 0.0115 0.9159 Time 1 0.784 0.4054 8 14.6284 <.0001 2 23.9551 <.0001 Inoculum:Treatment 2 18.6522 <.0001 2 0.6762 0.5103 2 0.3667 0.6983 Inoculum:Time 2 0.784 0.4604 16 0.4044 0.9808 4 0.551 0.6986 Treatment:Time 8 0.709 0.6835 2 6.949 0.0014 Inoculum:Treatment:Time 16 0.6311 0.8575 4 1.2724 0.2841 186 Figure 5.1. Photographs of 1. Planting of j-rooted ‘Merlot’ grapevine on July 10, 2020, 2. A j-rooted grapevine harvested in Spring 2023, 3. Set-up of enclosure treatment on July 27, 2022. 187 Figure 5.2. Photographs of young vine decline symptom progression of a 2.5k-JR+ plant on 1. June 2. 2. August 9. 3. August 30 and in a 25k-PFE- plant on 4. August 9. 5. August 30. 6. September 27, 2022. 188 Figure 5.3. Mean hourly temperature recorded by iButton for from July 27 to August 1 during enclosure treatment in five PFE+ and five PFE- grapevines. Dashed line represents PFE+ and solid line represents PFE-. * Indicates significant difference between temperatures according to Tukey’s significance test set at 0.05. 189 6 Conclusions In this thesis I assessed the health status of ready-to-plant nursery grapevines imported into Canada regarding several young vine decline pathogens. Overall, pathogen abundance varied widely between nurseries, grapevines within nurseries, and different sections of individual grapevines. Additionally, I aimed to determine the impact plant stress has on disease development in grapevines infected with P. chlamydospora. Pathogen abundance increased in the greenhouse due to a combination of water stress and AM fungal colonization and due to ring nematode infestation and AM fungal treatments. However, neither water stress nor ring nematode infestation in the field, nor j-rooting and heat stress significantly affected P. chlamydospora abundance. In the nematode experiment in the field, plant mortality over winter was higher in plants inoculated with a medium quantity of Pc spores and ring nematodes and mortality was higher in 50% irrigated plants compared to 100% irrigated in the field. This work suggests water stress and ring nematode infestation is detrimental to plant health and that symptom development and plant mortality may be related to plant stress in young grapevines infected with P. chlamydospora. This thesis shows that most grapevines planted in vineyards are likely to be infected with young vine decline fungi and provides new insights into Petri disease symptom development. This work may help in initiating new directions in the management of young vine decline in young vineyards regarding management of plant stress and rootstock selection to reduce the impact young vine decline has on young grapevines. 6.1 Are young vine decline fungi present in asymptomatic grapevines nursery material planted in Canada and what is their abundance within individual grapevines? My research found 99% of grapevines that were tested were infected with at least one young vine decline pathogen, and a mean of three young vine decline fungal species per plant. 190 Pathogen abundance varied widely between nurseries, grapevines, and sections of grapevines with no correlation between cultivars. The grapevines tested in this survey were asymptomatic and internal necrosis did not correlate with pathogen abundance in grapevines from any nursery, however, self-rooted plants from nursery B had lower internal necrosis than plants grafted onto rootstock, indicating internal necrosis is not a good measure of infection at this stage of plant development. Instead, rootstock and nursery may play a larger role in internal necrosis in ready-to-plant grapevines. Additionally, several young vine decline fungal species not previously reported in Canada were isolated from the nursery material and shows that nursery material is a source for the introduction of new pathogens in Canadian vineyards. 6.2 Does Phaeomoniella chlamydospora abundance within grapevines impact disease development in young grapevines? I found internal necrosis did not significantly differ between grapevines initially inoculated with 25k, 5k, 2.5k, or 1k P. chlamydospora conidia. Nor did I find differences in above- or below-ground plant growth measurements due to P. chlamydospora infection. Additionally, pathogen abundance at harvest for all experiments did not significantly differ due to initial P. chlamydospora inoculum level. The findings on pathogen abundance indicate that initial pathogen abundance has no long-term impact on pathogen abundance or plant health when initial spore quantity ranges from between 1-25k under greenhouse-controlled conditions. However, in the field experiments, water-inoculated controls were contaminated with P. chlamydospora and 25k inoculated plants had significantly higher pathogen abundance at harvest compared to WIC in the water stress and ring nematode field experiments. These findings suggest our initial inoculum levels may have been too high and could have resulted in P. chlamydospora quickly saturating the xylem vessels near the base of the grapevines where we 191 tested. The impact initial pathogen quantity has on plant health a few years after planting remains unclear and pathogen quantity varied widely between plants. The underlying reason for this variation was unclear. It was suggested in this thesis that the microbiome of individual grapevines could play a role in P. chlamydospora abundance. Similar levels of pathogen abundance at harvest between inoculated plants may be due to pathogen growth catching up or saturation of the pathogen in the xylem vessels where the DNA extractions from the wood were conducted. 6.3 Do abiotic and biotic plant stress influence Phaeomoniella chlamydospora abundance and Petri disease symptom development in young grapevines? My research shows that water stress can result in increased pathogen abundance, but this only occurred in conjunction with AM fungal colonization of the roots (Chapter 3, 4). However, pathogen abundance was not impacted by water stress in the field, nor due to ring nematodes in the field (Chapter 4) or heat or j-rooting stress (Chapter 5). The more controlled conditions in the greenhouse may explain the observed differences compared to field plants, as field plants were subjected to extreme conditions such as extremely cold winters in 2021 and 2022 as well as heatwaves which could have resulted in higher plant stress or damage. A combination of different stress factors in the field may have masked any treatment effects compared to the greenhouse. While a difference in pathogen abundance was not seen in the field, plant symptom development and mortality were impacted. In the field water stress experiment water stressed ‘Merlot’ plants had significantly higher mortality than non-stressed. The level of stress was not severe in the field which suggests stress alone did not result in mortality; however, due to contamination of the controls with P. chlamydospora I was unable to ascribe mortality to a combination of the pathogen and stress over stress alone The findings in chapters 3 suggest plant 192 stress does impact symptom development and mortality, however, I was unable to attribute symptom development and mortality to an increase in pathogen abundance, suggesting other mechanisms may be responsible, such as hydraulic failure. Ring nematodes had an impact on plant growth measurements and pathogen abundance increased due to ring nematode infestation and AM treatment in the greenhouse. Higher mortality was observed in plants inoculated with 5k Pc spores with ring nematode infestation of the roots compared to ring nematode free plants indicating that they are a potential concern to grapevines. This study revealed lower stem water potential due to ring nematode infestation, which could indicate increased susceptibility of the plant to stress. 6.4 Do arbuscular mycorrhizal fungi reduce plant stress and subsequent pathogen abundance and disease development in young grapevines? In the greenhouse, AM fungal colonization of the roots was associated with an increase in P. chlamydospora under water stress and ring nematode infestation. These findings suggest AM fungi may be detrimental to plant health in plants infected with P. chlamydospora which is comparable to reports of increased black foot pathogen growth in grapevines colonized with AM fungi (Holland et al. 2019; Vukicevich et al. 2018). AM fungi did not significantly reduce plant stress in either greenhouse experiment, however, AM fungal colonization was low in both experiments. It should be noted that plants in both greenhouse experiments received ample fertilization and plants in the nematode greenhouse experiment received sufficient water, which may have impacted AM colonization. Additionally, AM fungi were inoculated into the soil at planting, and in the water stress greenhouse experiment, plants were watered sufficiently in the first year due to pandemic restrictions, which 193 could have impacted AM fungal colonization. Any beneficial effect may have been reduced due to the low level of colonization. 6.5 Concluding remarks I initially expected pathogen abundance to increase in plants under stress, leading to increased internal necrosis and subsequent symptom development and plant mortality. The results of this thesis suggest pathogen abundance can increase due to plant stress but may not be the underlying cause of plant mortality. In the field experiments, pathogen abundance was not altered due to plant stress, however, Petri disease symptoms developed, and plant mortality was observed in the third year. Symptom development occurred immediately following a heatwave in 2022, suggesting sudden onset stress may play a role in disease development. This thesis shows that plant stress and AM fungi can interact with P. chlamydospora and provides evidence to support a potential area of research focused on the management of young vine decline in grapevines through stress reduction. The underlying causes of symptom development remain unclear and this research focused on pathogen abundance in relation to plant stress, but was limited in the other aspects of plant stress on plant responses when infected, alterations to the microbiome, or pathogen responses. 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Primers and probe sequences of targeted fungal pathogens used in each ddPCR assay for absolute quantification in ready-to-plant grapevine nursery material. Assay Name Gene Primer and Probe Name Pathogen Sequence Temperature Reference Bot Bt Probe B-tubulin Bot-Btub-F Bot-Btub-R BOT probe Botryosphaeriaceae spp. For: 5’- GTATGGCAATCTTCTGAACG -3’ Rev: 5’- CAGTTGTTACCGGCRCCAGA -3’ Probe: 5’-/56-FAM/TCG AGC CCG /ZEN/GCA CCA TGG AT/3IABkFQ/-3’ 60°C Billones-Baaijens et al. 2018 Clo Bt Probe B-tubulin Clo-BT-F Clo-BT-R Clo-BT probe Cadophora luteo-olivacea For: 5’-GAATATATGCAGACTCATTCAC-3’ Rev: 5’-AATATTAGGATTTTGTTCATGT-3’ Probe: 5’-/56-FAM/CCAACTCGA/ZEN/GCGCCTGAACGT/31ABkFQ/-3’ 56°C Maldonado-González et al. 2020 Dm H3 Probe Histone 3 Ilyo-H3-F Ilyo-H3-R IlyoMac probe Dactylonectria macrodidyma For: 5’- CTCCTCATCCGCAAGCTC -3’ Rev: 5’- AAGATCGCTCTTGAAGTCCTG -3’ Probe: 5’- ACAACACCTTTAACATTTAACCACACAT –3’ 57°C This study Dt H3 Probe Histone 3 Ilyo-H3-F Ilyo-H3-R IlyoTorr probe Dactylonectria torresensis For: 5’- CTCCTCATCCGCAAGCTC -3’ Rev: 5’- AAGATCGCTCTTGAAGTCCTG -3’ Probe: 5’- CCAAACACAACACCTTTTTACTTTCAACCACAC –3’ 58°C This study Pm Bt Primers B-tubulin Pal1-FOR Pal1-REV Phaeoacremonium minimum For: 5’- CCTCGACGAGCCCAGATTC -3’ Rev: 5’- GTAAAAACCCCACCGGTTAGC -3’ 60°C Martín et al. 2012 Pc ITS Probe ITS Pch-ITS-417F Pch-ITS-495R PCh Probe Phaeomoniella chlamydospora For: 5’- TGGCGGCGTCAAGAATGA -3’ Rev: 5’- CAATAGGGCCAGGCCAAG -3’ Probe: 5’- /56-FAM/AGG TGC AGC /ZEN/GAG CAA TCA AGC ATA CA/3IABkFQ/ -3’ 58°C Martín et al. 2012/ This study 225 Appendix B: Supplementary material for Chapter 3 Table S3.1. Mean SPAD leaf chlorophyll and standard error over time in 2021 for each level of inoculum and each level of stress in the greenhouse. Treatment 25-May 15-Jun 30-Jun 19-Jul 29-Jul 12-Aug 24-Aug 09-Sep NIC 26.96 ± 0.41 a 33.40 ± 0.42 a 32.80 ± 0.50 a 34.95 ± 0.47 a 36.02 ± 0.42 a 35.59 ± 35.59 a 34.42 ± 0.46 a 34.52 ± 0.64 a WIC 27.85 ± 0.35 a 33.46 ± 0.40 a 32.48 ± 0.39 a 35.02 ± 0.35 a 36.16 ± 0.40 a 35.71 ± 35.71 a 34.86 ± 0.43 a 34.20 ± 0.38 a 1k 27.77 ± 0.37 a 33.87 ± 0.66 a 33.07 ± 0.33 a 35.53 ± 0.37 a 35.96 ± 0.42 a 36.28 ± 36.28 a 34.86 ± 0.56 a 33.87 ± 0.44 a 5k 27.17 ± 0.37 a 32.83 ± 0.40 a 32.43 ± 0.47 a 34.51 ± 0.39 a 35.75 ± 0.55 a 35.50 ± 35.50 a 34.39 ± 0.45 a 34.24 ± 0.50 a 25k 27.19 ± 0.43 a 33.71 ± 0.45 a 32.55 ± 0.44 a 34.96 ± 0.45 a 36.06 ± 0.45 a 35.74 ± 35.74 a 34.81 ± 0.49 a 34.04 ± 0.68 a I1 27.36 ± 0.27 A 33.54 ± 0.29 A 33.46 ± 0.27 B 35.26 ± 0.26 A 36.71 ± 0.28 B 35.76 ± 35.76 A 34.52 ± 0.33 A 34.58 ± 0.36 B I2 27.41 ± 0.23 A 33.37 ± 0.33 A 31.88 ± 0.24 A 34.72 ± 0.26 A 35.26 ± 0.27 A 35.77 ± 35.77 A 34.82 ± 0.28 A 33.75 ± 0.33 A Same lowercase and uppercase letter in each column represent no significant differences (p = 0.05) according to Tukey’s significance test. 226 Table S3.2. Mean dry pruning weight, dry root weight, and node number for each year and standard error for year 1 and year 2 for each level stress and AM fungi in the greenhouse. Treatment 2020 2021 Pruning weight (g) Node number Pruning weight (g) Node number Root weight (g) I1:AM- 14.87 ± 0.31 a 69.91 ± 2.76 a 27.41 ± 0.72 a 99.03 ± 2.48 b 37.65 ± 1.34 a I1:AM+ 14.29 ± 0.36 a 63.52 ± 2.89 a 30.11 ± 0.79 a 104.9 ± 2.72 b 39.62 ± 1.52 a I2:AM- 15.11 ± 0.40 a 68.18 ± 3.09 a 25.47 ± 0.55 a 80.27 ± 1.68 a 32.67 ± 1.02 a I2:AM+ 15.09 ± 0.41 a 67.94 ± 2.99 a 26.07 ± 0.62 a 77.88 ± 1.53 a 30.72 ± 1.08 a I1 14.58 ± 0.24 A 66.71 ± 2.04 A 28.76 ± 0.56 B 101.9 ± 1.87 B 38.60 ± 1.02 B I2 15.10 ± 0.29 A 68.06 ± 2.15 A 25.77 ± 0.41 A 79.08 ± 1.15 A 31.76 ± 0.75 A AM- 14.99 ± 0.25 A 69.05 ± 2.07 A 26.44 ± 0.47 A 89.65 ± 1.89 A 34.94 ± 0.89 A AM+ 14.69 ± 0.28 A 65.73 ± 2.10 A 28.09 ± 0.56 A 91.18 ± 2.28 A 34.91 ± 1.11 A Same lowercase and uppercase letter in each column (stress x AM interaction and stress and AM separately) represent no significant differences (p = 0.05) according to Tukey’s significance test. 227 Table S3.3. Mean SPAD leaf chlorophyll and standard error each year for each cultivar, level of inoculum, and level of stress in the field. Cultivar Treatment 2021 2022 2023 M WIC 33.66 ± 0.56 a 27.12 ± 0.87 a 22.8 ± 1.24 a 2.5k 34.78 ± 0.31 a 27.43 ± 0.68 a 25.88 ± 1.25 a 25k 33.97 ± 0.52 a 26.44 ± 0.97 a 25.23 ± 1.25 a I1 33.35 ± 0.39 A 27.45 ± 0.59 A 23.75 ± 0.86 A I2 34.96 ± 0.32 A 26.42 ± 0.84 A 22.23 ± 1.23 A RS WIC 34.82 ± 0.38 b 26.32 ± 1.04 a 28.61 ± 0.6 a 2.5k 33.78 ± 0.91 ab 26.99 ± 0.73 a 27.02 ± 0.96 a 25k 32.64 ± 0.84 a 25.53 ± 1.32 a 27.84 ± 0.68 a I1 32.97 ± 0.69 A 26.58 ± 0.84 A 28.59 ± 0.74 A I2 34.52 ± 0.54 A 26.07 ± 0.87 A 27.19 ± 0.51 A Combined WIC 34.24 ± 0.35 a 26.64 ± 0.66 b 25.97 ± 0.9 a 2.5k 34.28 ± 0.51 a 27.16 ± 0.5 b 26.71 ± 0.79 a 25k 33.28 ± 0.5 a 25.8 ± 0.88 a 25.36 ± 0.84 a I1 33.16 ± 0.39 B 26.99 ± 0.53 A 26.35 ± 0.72 A I2 34.74 ± 0.32 A 26.22 ± 0.62 A 25.6 ± 0.7 A Same lowercase and uppercase letter within each column for each cultivar represent no significant differences (p = 0.05) according to Tukey’s significance test. 228 Table S3.4. Mean dry pruning weight and node number for 2020 and 2021 and standard error for each cultivar, level of infection, and stress in the field. Same lowercase and uppercase letter within each cultivar in each column represent no significant differences (p = 0.05) according to Tukey’s significance test. 2020 2021 Cultivar Inoculum Treatment Pruning weight (g) Node # Pruning weight (g) Node # M WIC I1 1.27 ± 0.24 a N/A 16.92 ± 5.37 a 33.83 ± 6.83 a I2 1.43 ± 0.2 a N/A 11.05 ± 3.89 a 27.71 ± 6.18 a Combined 1.36 ± 0.16 A N/A 13.76 ± 3.34 A 30.54 ± 4.66 A 2.5k I1 1.17 ± 0.33 a N/A 12.39 ± 3.82 a 23.67 ± 3.87 a I2 1.22 ± 0.2 a N/A 12.75 ± 3.53 a 30.86 ± 5.69 a Combined 1.2 ± 0.19 A N/A 12.58 ± 2.59 A 27.54 ± 3.68 A 25k I1 1.98 ± 0.34 a N/A 10.58 ± 3.59 a 23.67 ± 4.75 a I2 1.13 ± 0.24 a N/A 7.73 ± 2.57 a 17.86 ± 4.86 a Combined 1.52 ± 0.23 A N/A 9.05 ± 2.19 A 20.54 ± 3.51 A RS WIC I1 0.78 ± 0.16 a N/A 28.33 ± 3.63 a 36.5 ± 4.75 a I2 0.97 ± 0.17 a N/A 18.04 ± 4.09 a 31.57 ± 4.33 a Combined 0.89 ± 0.12 A N/A 22.79 ± 3.11 A 33.85 ± 3.27 A 2.5k I1 0.85 ± 0.17 a N/A 22.87 ± 8.27 a 25.83 ± 7.86 a I2 1.05 ± 0.14 a N/A 17.38 ± 4.66 a 30.43 ± 5.74 a Combined 0.96 ± 0.11 A N/A 19.92 ± 4.63 A 28.31 ± 4.81 A 25k I1 0.67 ± 0.18 a N/A 16.31 ± 6.46 a 23.5 ± 6.47 a I2 0.76 ± 0.15 a N/A 18.75 ± 6.79 a 31.83 ± 8.98 a Combined 0.72 ± 0.12 A N/A 17.53 ± 4.7 A 27.67 ± 5.66 A Combined WIC I1 1.03 ± 0.16 a N/A 22.63 ± 3.63 a 35.17 ± 4.18 a I2 1.2 ± 0.14 a N/A 14.54 ± 2.97 a 29.64 ± 3.81 a Combined 1.12 ± 0.11 A N/A 18.27 ± 2.45 A 32.19 ± 2.87 A 2.5k I1 1.01 ± 0.19 a N/A 17.63 ± 4.8 a 24.75 ± 4.39 a I2 1.14 ± 0.13 a N/A 15.07 ± 2.99 a 30.64 ± 4.04 a Combined 1.08 ± 0.11 A N/A 16.25 ± 2.75 A 27.92 ± 3.03 A 25k I1 1.32 ± 0.27 a N/A 13.45 ± 3.79 a 23.58 ± 4.01 a I2 1.62 ± 0.6 a N/A 12.82 ± 3.75 a 24.31 ± 5.27 a Combined 1.48 ± 0.34 A N/A 13.12 ± 2.67 A 23.96 ± 3.35 A 229 Table S3.5. Mean dry pruning weight, node number, trunk diameter, and dry root weight for 2022 and 2023 and standard error for each cultivar, level of infection, and stress in the field. Same lowercase and uppercase letter within each cultivar in each column represent no significant differences (p = 0.05) according to Tukey’s significance test. 2022 2023 Cultivar Inoculum Treatment Pruning weight (g) Node # Pruning weight (g) Node # Trunk diameter (mm) Root weight (g) M WIC I1 16.15 ± 6.93 a 32.5 ± 4.81 a 6.53 ± 2.13 a 52.17 ± 5.31 a 12.32 ± 0.76 a 34.77 ± 7.59 a I2 17.63 ± 6.17 a 23.67 ± 4.11 a 5.79 ± 3.5 a 41.25 ± 12.3 a 12.52 ± 0.78 a 25.75 ± 4.78 a Combined 16.89 ± 4.64 A 28.08 ± 3.41 A 6.23 ± 1.9 A 47.8 ± 6.1 A 12.42 ± 0.54 A 30.26 ± 4.67 A 2.5k I1 37.33 ± 16.92 a 29.25 ± 5.1 a 5.54 ± 3.04 a 36.75 ± 9.78 a 13.94 ± 1.11 a 36.13 ± 13.85 a I2 18.53 ± 13.63 a 21 ± 5.69 a 8.66 ± 0 a 65 ± 0 a 12.57 ± 0.93 a 30.31 ± 12.48 a Combined 27.93 ± 11.36 A 25.13 ± 4.09 A 6.17 ± 2.5 A 42.4 ± 9.31 A 13.25 ± 0.76 A 33.22 ± 9.38 A 25k I1 16.72 ± 3.63 a 28.2 ± 4.98 a 3.81 ± 1 a 40.75 ± 3.38 a 12.96 ± 0.6 a 29.18 ± 4.21 a I2 10.63 ± 4.03 a 23.25 ± 3.43 a 0.85 ± 0.1 a 30 ± 0.82 a 11.04 ± 0.86 a 17.56 ± 3.74 a Combined 14.01 ± 2.88 A 26 ± 3.27 A 2.54 ± 0.8 A 36.14 ± 2.81 A 12.11 ± 0.6 A 24.01 ± 3.46 A RS WIC I1 57.23 ± 7.54 a 46.17 ± 3.59 a 16.52 ± 2.92 a 73.17 ± 5.36 a 13.12 ± 0.54 a 47.36 ± 3.85 a I2 33.44 ± 9.83 a 33.86 ± 3.91 a 12.52 ± 1.82 a 67.17 ± 5.22 a 11.88 ± 0.48 a 38.94 ± 7.23 a Combined 44.42 ± 7.14 A 39.54 ± 3.18 A 14.52 ± 1.81 A 70.17 ± 3.84 A 12.45 ± 0.4 A 42.83 ± 4.43 A 2.5k I1 63.88 ± 22.03 a 43.17 ± 2.35 a 14.44 ± 3.3 a 65.83 ± 6.78 a 12.4 ± 0.87 a 39.06 ± 4.98 a I2 40.67 ± 3.24 a 37 ± 1.97 a 13.72 ± 2.79 a 70.6 ± 2.75 a 11.91 ± 0.49 a 48.41 ± 4.46 a Combined 52.28 ± 11.63 A 40.08 ± 1.77 A 14.11 ± 2.21 A 68 ± 3.97 A 12.16 ± 0.5 A 43.74 ± 3.6 A 25k I1 32.62 ± 8.38 a 37 ± 5.85 a 12.47 ± 1.02 a 69.67 ± 5.93 a 10.88 ± 0.92 a 27.63 ± 5.03 a I2 38.98 ± 12.66 a 32 ± 2.87 a 15 ± 2.5 a 73.67 ± 5.92 a 12.09 ± 0.69 a 40.91 ± 5.14 a Combined 36.09 ± 7.6 A 34.27 ± 3.18 A 14.16 ± 1.75 A 72.33 ± 4.46 A 11.54 ± 0.59 A 34.87 ± 4.13 A Combined WIC I1 36.69 ± 7.83 a 39.33 ± 3.59 a 11.53 ± 2.31 a 62.67 ± 4.84 a 11.77 ± 0.49 a 41.06 ± 4.63 a I2 26.15 ± 6.4 a 29.15 ± 3.17 a 9.83 ± 2.06 a 56.8 ± 7.08 a 12.05 ± 0.4 a 32.85 ± 4.83 a Combined 31.21 ± 5.13 A 34.17 ± 2.64 A 10.75 ± 1.58 A 60 ± 4.21 A 12.44 ± 0.33 A 36.79 ± 3.45 A 2.5k I1 53.26 ± 15.41 a 37.6 ± 3.29 a 10.88 ± 2.7 a 54.2 ± 7.22 a 13.39 ± 0.58 a 37.89 ± 6.31 a I2 31.81 ± 6.73 a 30.6 ± 3.57 a 12.88 ± 2.45 a 69.67 ± 2.45 a 11.87 ± 0.75 a 41.17 ± 6.32 a Combined 42.54 ± 8.74 A 34.1 ± 2.55 A 11.63 ± 1.94 A 60 ± 4.97 A 12.6 ± 0.45 A 39.53 ± 4.48 A 25k I1 24.67 ± 5.21 a 32.6 ± 4.09 a 7.52 ± 1.77 a 53.14 ± 6.28 a 12.28 ± 0.61 a 28.4 ± 3.29 a I2 27.64 ± 8.92 a 28.5 ± 2.59 a 10.28 ± 2.78 a 59.11 ± 7.92 a 12.53 ± 0.44 a 31.57 ± 4.99 a Combined 26.16 ± 5.18 A 30.55 ± 2.46 A 9.08 ± 1.78 A 56.5 ± 5.29 A 11.79 ± 0.43 A 29.99 ± 3.01 A 230 Table S3.6. Percent class 1, class 2, total necrosis, and mean basal surface area and standard error of each cultivar containing each level of inoculum and in each level of stress in the greenhouse. Same lowercase and uppercase letter within each cultivar in each column represent no significant differences (p = 0.05) according to Tukey’s significance test. Cultivar Inoculum Treatment Class 1 necrosis Class 2 necrosis Total necrosis Basal surface area M WIC I1 9.14 ± 2.52 a 9.57 ± 1.46 a 18.71 ± 3.7 a 189.52 ± 25.5 a I2 13.1 ± 1.93 a 12.41 ± 2.62 a 24.63 ± 3.71 a 164.41 ± 9.55 a Combined 11.12 ± 1.69 A 10.86 ± 1.52 A 21.4 ± 2.75 A 178.11 ± 14.53 A 2.5k I1 6.65 ± 1.86 a 8.94 ± 2.22 a 15.6 ± 4.08 a 219.22 ± 54.95 a I2 28.98 ± 11.21 a 18.94 ± 6.86 a 47.91 ± 17.82 a 171.66 ± 28.97 a Combined 17.82 ± 6.92 A 13.94 ± 4.01 A 31.75 ± 10.78 A 195.44 ± 32.18 A 25k I1 21.81 ± 5.53 a 17.59 ± 3.19 a 39.41 ± 8.54 a 151.16 ± 18.62 a I2 37.19 ± 11.8 a 13.67 ± 3.49 a 50.87 ± 12.79 a 107.09 ± 20.61 a Combined 28.65 ± 6.59 A 15.85 ± 2.44 A 44.5 ± 7.64 A 131.57 ± 15.63 A RS WIC I1 0.91 ± 0.23 a 8.42 ± 1.06 a 9.32 ± 1.06 a 155.58 ± 14.41 a I2 12.71 ± 8.84 a 12.95 ± 3.14 a 25.66 ± 11.91 a 107.2 ± 7.17 a Combined 6.81 ± 4.87 A 10.68 ± 1.82 A 17.49 ± 6.6 A 131.39 ± 10.33 A 2.5k I1 7.23 ± 1.64 a 7.39 ± 0.72 a 14.62 ± 1.97 a 133.83 ± 16.73 a I2 5.24 ± 1.88 a 8.27 ± 0.63 a 13.51 ± 1.74 a 129.09 ± 3.47 a Combined 6.24 ± 1.28 A 7.83 ± 0.5 A 14.07 ± 1.32 A 131.46 ± 8.57 A 25k I1 7.72 ± 1.17 a 21.34 ± 8.56 a 30.69 ± 9.59 a 111.14 ± 10.76 a I2 9.58 ± 3.23 a 7.81 ± 0.87 a 17.39 ± 2.91 a 129.6 ± 15.77 a Combined 8.74 ± 1.86 A 14.57 ± 4.59 A 24.04 ± 5.22 A 120.37 ± 10.1 A Combined WIC I1 5.02 ± 1.74 a 8.99 ± 0.92 a 14.01 ± 2.35 a 172.55 ± 15.44 a I2 12.58 ± 4.51 a 12.17 ± 2.06 a 24.34 ± 6.44 a 133.36 ± 9.41 a Combined 8.96 ± 2.65 B 10.77 ± 1.2 A 19.36 ± 3.7 B 153.74 ± 9.96 B 2.5k I1 7 ± 1.24 a 8.01 ± 1.02 a 15.01 ± 2.02 a 167.98 ± 27.55 a I2 14.74 ± 5.91 a 12.54 ± 3.22 a 27.27 ± 8.96 a 146.12 ± 13.49 a Combined 10.87 ± 3.14 A 10.28 ± 1.76 A 21.14 ± 4.79 B 157.05 ± 15.53 B 25k I1 14.77 ± 3.6 a 19.64 ± 4.76 a 34.65 ± 6.6 a 129.33 ± 12.25 a I2 20.63 ± 6.66 a 10.15 ± 1.74 a 30.78 ± 7.49 a 120.6 ± 13.03 a Combined 17.7 ± 3.84 A 15.12 ± 2.79 A 32.81 ± 5 A 125.17 ± 8.98 A 231 Appendix C: Supplementary material for Chapter 4 Table S4.1. Mean leaf chlorophyll, and standard error each year in the greenhouse for each level of inoculum and ring nematode infestation treatment. Treatment May 26 June 15 July 22 August 24 September 9 NIC 34.94 ± 0.54 a 35.62 ± 0.64 a 37.18 ± 0.63 a 38.09 ± 0.74 a 37.16 ± 0.88 a WIC 35.43 ± 0.58 a 36.76 ± 0.65 a 37.58 ± 0.7 a 37.37 ± 0.78 a 36.04 ± 0.99 a 1k 35.26 ± 0.44 a 35.89 ± 0.46 a 37.91 ± 0.68 a 39.14 ± 0.63 a 38.23 ± 0.63 a 5k 35.81 ± 0.51 a 36.81 ± 0.66 a 37.73 ± 0.85 a 37.96 ± 0.58 a 37.47 ± 0.78 a 25k 35.71 ± 0.41 a 36.54 ± 0.58 a 37.58 ± 0.72 a 37.41 ± 0.95 a 35.93 ± 0.87 a Nem - 36.08 ± 0.28 B 36.96 ± 0.36 B 37.98 ± 0.41 A 37.91 ± 0.49 A 37.21 ± 0.59 A Nem + 34.69 ± 0.32 A 35.55 ± 0.37 A 37.19 ± 0.5 A 38.21 ± 0.48 A 36.82 ± 0.45 A AMF - 35.49 ± 0.27 A 36.5 ± 0.32 A 37.85 ± 0.37 A 38.7 ± 0.48 A 37.87 ± 0.54 A AMF + 35.36 ± 0.35 A 36.08 ± 0.43 A 37.37 ± 0.52 A 37.4 ± 0.47 A 36.18 ± 0.5 A For each column leaf chlorophyll for each level of inoculum with the same lowercase letter are not significantly different (p = 0.05) according to Tukey’s significance test. For each column, leaf chlorophyll for each level of stress with the same uppercase letter are not significantly different (p = 0.05) according to Tukey’s significance test 232 Table S4.2. Mean pruning weight, node number, and root weight, and standard error each year in the greenhouse for each level of inoculum, AM treatment, and ring nematode infestation treatment. Treatment 2020 2021 Combined Pruning weight (g) Node number Pruning weight (g) Node number Pruning weight (g) Node number Root weight (g) NIC 19.79 ± 0.54 a 81.1 ± 2.46 a 26.96 ± 1.08 a 107.71 ± 3.04 a 23.38 ± 0.59 a 94.4 ± 1.41 a 45.05 ± 3.74 a WIC 18.03 ± 0.81 a 84.74 ± 3.06 a 27.45 ± 0.75 a 112.37 ± 3.54 a 22.74 ± 0.64 a 98.55 ± 2.71 a 46.23 ± 3.32 a 1k 19.16 ± 0.63 a 81.58 ± 2.9 a 27.21 ± 0.77 a 113.69 ± 2.65 a 23.19 ± 0.42 a 97.63 ± 1.73 a 44.24 ± 2.29 a 5k 19.74 ± 0.72 a 82.5 ± 2.57 a 29.04 ± 0.9 a 113.45 ± 3.49 a 24.39 ± 0.58 a 97.98 ± 2.25 a 40.69 ± 1.93 a 25k 19.62 ± 0.7 a 82.43 ± 2.81 a 28.43 ± 1.02 a 115.43 ± 3.73 a 24.02 ± 0.45 a 98.93 ± 2.42 a 40.12 ± 2.48 a Nem - 20 ± 0.42 A 86.53 ± 1.73 A 28.7 ± 0.53 A 112.72 ± 2.01 A 24.35 ± 0.31 B 99.63 ± 1.29 A 43.62 ± 1.45 A Nem + 18.48 ± 0.43 A 77.67 ± 1.53 A 26.77 ± 0.61 A 112.53 ± 2.14 A 22.63 ± 0.33 A 95.1 ± 1.31 43 ± 2.14 A AMF - 19.18 ± 0.39 A 80.94 ± 1.56 A 27.8 ± 0.56 A 110.46 ± 1.95 A 23.49 ± 0.33 A 95.7 ± 1.25 B 43.87 ± 1.44 A AMF + 19.39 ± 0.47 A 83.8 ± 1.91 A 27.79 ± 0.6 A 114.76 ± 2.14 A 23.59 ± 0.35 A 99.28 ± 1.37 A 42.82 ± 2.07 A For each column for each level of inoculum with the same lowercase letter are not significantly different (p = 0.05) according to Tukey’s significance test. For each column, for each level of stress with the same uppercase letter are not significantly different (p = 0.05) according to Tukey’s significance test 233 Table S4.3. Mean SPAD leaf chlorophyll, stem water potential (MPa), and standard error each year in the field for each level of inoculum and ring nematode infestation treatment. 2021 2022 2023 Treatment Leaf chlorophyll Leaf chlorophyll Stem water potential Leaf chlorophyll Stem water potential IC 37.88 ± 0.63 a 27.12 ± 0.63 a N/A 27.29 ± 0.49 a N/A 1k 37.3 ± 0.48 a 27.79 ± 0.59 a N/A 26.45 ± 0.79 b N/A 5k 36.63 ± 0.91 a 27.2 ± 0.53 a N/A 26.19 ± 0.73 ab N/A 25k 36.52 ± 0.67 a 26.96 ± 0.78 a N/A 26.87 ± 0.87 ab N/A Nem- 36.87 ± 0.54 A 26.83 ± 0.44 A -0.77 ± 0.01 A 27.26 ± 0.43 A -.80 ± 0.02 A Nem+ 37.32 ± 0.42 A 27.79 ± 0.48 B -0.82 ± 0.02 B 26.09 ± 0.58 B -0.79 ± 0.02 A For each column leaf chlorophyll and stem water potential readings for each level of inoculum with the same lowercase letter are not significantly different (p = 0.05) according to Tukey’s significance test. For each column, leaf chlorophyll and stem water potential readings for each level of stress with the same uppercase letter are not significantly different (p = 0.05) according to Tukey’s significance test 234 Table S4.4. Mean pruning weight, node number, trunk diameter, and root weight and standard error each year in the field for each level of inoculum and ring nematode infestation treatment. Treatment 2021 2022 2023 Pruning weight (g) Node number Pruning weight (g) Node number Pruning weight (g) Node number Trunk diameter (mm) Root weight (g) WIC 30.46 ± 4.99 a 166.5 ± 13.8 a 74.94 ± 11.38 a 57.27 ± 4.16 a 96.4 ± 13.22 a 95.47 ± 6.7 a 190.09 ± 0.95 a 125.65 ± 9.58 a 1k 18.05 ± 2.34 a 146.35 ± 13.76 a 59.64 ± 11.03 a 56.5 ± 5.67 a 89.34 ± 17.39 a 94.75 ± 9.65 a 189.41 ± 0.89 a 91.08 ± 10.17 a 5k 18.74 ± 4.42 a 133.33 ± 17.6 a 62.74 ± 14.5 a 59.25 ± 4.34 a 88.12 ± 16.23 a 101.17 ± 10.54 a 189.02 ± 0.93 a 105.36 ± 12.36 a 25k 22.32 ± 3.63 a 142 ± 13.06 a 61.65 ± 12.12 a 51.92 ± 4.79 a 73.49 ± 18.84 a 82.77 ± 10.28 a 188.3 ± 1.18 a 92.21 ± 11.07 a Nem - 18.73 ± 2.49 A 136.39 ± 9.83 A 65.15 ± 8.93 A 56.34 ± 3.36 A 84.68 ± 11.77 A 96.72 ± 7.26 A 189.28 ± 0.77 A 103.75 ± 8.67 A Nem + 26.27 ± 3.45 A 158.81 ± 11.07 A 65.43 ± 8.54 A 56.04 ± 3.44 A 90.22 ± 11.52 A 89.3 ± 5.38 A 189.19 ± 0.63 A 105.74 ± 7.17 A For each column for each level of inoculum with the same lowercase letter are not significantly different (p = 0.05) according to Tukey’s significance test. For each column, for each level of stress with the same uppercase letter are not significantly different (p = 0.05) according to Tukey’s significance test 235 Appendix D: Supplementary material for Chapter 5 Table S5.1. Mean pruning weight, node number, trunk diameter, and root weight and standard error each year in the field for each level of inoculum and j-rooting treatment. For each column for each level of inoculum with the same lowercase letter are not significantly different (p = 0.05) according to Tukey’s significance test. For each column, for each level of stress with the same uppercase letter are not significantly different (p = 0.05) according to Tukey’s significance test 2020 2021 2022 2023 Treatment Pruning weight (g) Node number Pruning weight (g) Node number Pruning weight (g) Node number Pruning weight (g) Node number Trunk diameter (mm) Root weight (g) WIC 0.99 ± 0.16 a N/A 13.46 ± 4.31 a 22.13 ± 3.96 a 40.13 ± 10.28 a 42.91 ± 7.72 a 9.2 ± 2.06 a 56.55 ± 5.97 a 13.35 ± 0.66 a 42.86 ± 7.92 a 2.5k 0.97 ± 0.17 a N/A 13.36 ± 2.77 a 26.31 ± 3.3 a 19.35 ± 6.87 a 28.92 ± 3.36 a 4.67 ± 1.25 a 49.82 ± 6.85 a 13.77 ± 0.65 a 38.96 ± 4.41 a 25k 1.46 ± 0.35 a N/A 21.04 ± 6.92 a 31.5 ± 7.08 a 16.65 ± 6.71 a 29.55 ± 4.74 a 4.58 ± 1.38 a 48.88 ± 7.52 a 13.63 ± 0.95 a 36.4 ± 6.34 a JR- 1.16 ± 0.2 A N/A 17.37 ± 3.5 A 27.17 ± 3.43 A 29.72 ± 7.48 A 38.39 ± 4.98 A 7.19 ± 1.64 A 51.65 ± 6.23 A 14.05 ± 0.58 A 46.53 ± 6.21 A JR+ 1.12 ± 0.2 A N/A 14.54 ± 4.53 A 26.13 ± 4.79 A 20.11 ± 6.69 A 28.31 ± 4.25 A 5.16 ± 1.02 A 52.54 ± 4.41 A 13.02 ± 0.61 A 31.13 ± 2.84 A 236 Table S5.2. Mean pruning weight, node number, trunk diameter, and root weight and standard error each year in the field for each level of inoculum and enclosure treatment. For each column for each level of inoculum with the same lowercase letter are not significantly different (p = 0.05) according to Tukey’s significance test. For each column, for each level of stress with the same uppercase letter are not significantly different (p = 0.05) according to Tukey’s significance test 2020 2021 2022 2023 Treatment Pruning weight (g) Node number Pruning weight (g) Node number Pruning weight (g) Node number Pruning weight (g) Node number Trunk diameter (mm) Root weight (g) WIC 1.02 ± 0.18 a N/A 25.37 ± 6.63 a 39.13 ± 6.55 a 31.78 ± 7.74 a 42.15 ± 5.42 a 7.57 ± 1.44 a 60.25 ± 7.09 a 14.25 ± 0.77 a 50.66 ± 6.61 a 2.5k 0.97 ± 0.21 a N/A 26.82 ± 5.67 a 42.75 ± 6.56 a 34.39 ± 9 a 43.86 ± 4.2 a 7.85 ± 1.56 a 63.83 ± 5.96 a 14.67 ± 0.59 a 48.11 ± 7.29 a 25k 0.99 ± 0.22 a N/A 24.99 ± 8.34 a 42.06 ± 10.85 a 26.07 ± 11.31 a 39.79 ± 9.16 a 9.27 ± 2.11 a 66.29 ± 7.5 a 14.46 ± 1.06 a 35.03 ± 7.38 a LT 1.06 ± 0.16 A N/A 26.47 ± 5.76 A 43.04 ± 7.24 A 28.55 ± 7.54 A 38.14 ± 4.95 A 7.68 ± 1.21 A 62.76 ± 6.04 A 14.33 ± 0.66 A 42.62 ± 5.82 A HT 0.93 ± 0.17 A N/A 24.98 ± 5.14 A 39.58 ± 4.96 A 33.23 ± 7.38 A 46.32 ± 4.77 A 8.53 ± 1.52 A 63.29 ± 4.92 A 14.62 ± 0.61 A 46.58 ± 6.19 A 237 Figure S5.1. Mean hourly temperature recorded by iButton for the duration of the heat stress treatment five PFE- grapevines and adjacent weather station. Dashed line represents the weather station and solid line represents PFE-. * Indicates significant difference between temperatures according to Tukey’s significance test.
Thesis/Dissertation
eng
Biology
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Hart, Miranda
Urbez-Torres, Jose Ramon
The effect of abiotic and biotic plant stress on Phaeomoniella chlamydospora abundance and disease development in young grapevines
Text
Dataset