Science, Irving K. Barber Faculty of (Okanagan)
Biology, Department of (Okanagan)
DSpace
UBCO
O'Farrell, Corynne
Master of Science - MSc
University of British Columbia
Brassica plants with biofumigant properties have been used to mitigate soil borne pathogens for years. This is common in till systems where brassicas are incorporated into the soil. The effect of these crops as living mulch on soil microbes is not as well studied. This study compared the response of the soil microbial community to four brassica species, two commonly used in vineyards (Sinapis alba L. (white mustard) and Raphanus sativus (L.) Domin) and two native/naturalized to the Okanagan (Capsella bursa-pastoris (L.) Medik. And Boechera holboelli (Hornem.) Á.Löve & D.Löve (Holbøll’s rockcress)). These cover crops were grown in a vineyard for one growing season. Soil samples were taken at the end of the season for DNA and nematode extraction. The fungal community was analyzed by sequencing of the ITS2 sub-region. Species evenness was higher in brassicas compared to the control. I found no differences in community composition. Changes to the arbuscular mycorrhizal fungi (AMF) community were analyzed by spore count; I found no differences. The nematode community was analyzed by counting total, plant parasitic (PPN), and free-living nematodes (FLN). There were no differences in abundance of nematodes. I found the relative abundance of PPN to be significantly lower in the C. bursa-pastoris and S. alba treatments compared to the control. A greenhouse experiment was conducted to determine if any of the four brassicas act as hosts for fungal pathogen Ilyonectria liriodendri (PARC60 and PARC393), nematode pathogen Mesocriconema xenoplax, or AMF Rhizophagus irregularis. The same four cover crops as well as a grapevine control were grown in soil inoculated with the organism of interest for four months, at the end of which abundance of I. liriodendri and AMF in the roots was measured as well as M. xenoplax abundance in the soil. There were no differences in I. liriodendri colonization between cover crops, but all had some colonization suggesting they may be able to act as hosts. None of the brassicas were colonized by AMF, although B. holboelli was colonized by another fungal endophyte. Abundance of M. xenoplax in the soil was significantly lower in soil of all brassicas compared to the grapevine.
DO BRASSICA COVER CROPS INFLUENCE SOIL FUNGAL AND NEMATODE COMMUNITIES IN A VINEYARD? by Corynne O'Farrell B.Sc. (Hons), The University of Calgary, 2016 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in THE COLLEGE OF GRADUATE STUDIES (Biology) THE UNIVERSITY OF BRITISH COLUMBIA (Okanagan) January 2022 © Corynne O'Farrell, 2022 ii The following individuals certify that they have read, and recommend to the College of Graduate Studies for acceptance, a thesis/dissertation entitled: Do brassica cover crops influence soil fungal and nematode communities in a vineyard? submitted by Corynne O'Farrell in partial fulfillment of the requirements of the degree of Master of Science . Dr. Miranda Hart, Department of Biology, Irving K. Barber Faculty of Science Supervisor Dr. Tomas Forge, Adjunct Professor, Summerland Research and Development Centre Supervisory Committee Member Dr. Melanie Jones, Department of Biology, Irving K. Barber Faculty of Science Supervisory Committee Member Dr. Cara Haney, Department of Microbiology and Immunology University Examiner iii Abstract Brassica plants with biofumigant properties have been used to mitigate soil borne pathogens in agricultural settings for years. This practice is especially common in till systems where the brassicas are incorporated into the soil as green manure or seed meal. The effect of these crops on soil microbes in the field, including both pathogens and beneficials, is not as well studied in no-till systems. This study compared the response of the soil microbial community to four different brassica cover crops, two of which are commonly used in Okanagan vineyards (Sinapis alba L. (white mustard) and Raphanus sativus (L.) Domin (tillage radish)) as well as two brassicas that are native or naturalized to the Okanagan (Capsella bursa-pastoris (L.) Medik. (Shepherd’s purse) and Boechera holboelli (Hornem.) Á.Löve & D.Löve (Holbøll’s rockcress)). These cover crops were grown in a field in a vineyard for one full growing season. Soil samples were taken at the end of the season for DNA and nematode extraction. The fungal community was analyzed by sequencing of the ITS2 sub-region with Illumina. I found no differences in the diversity or composition of the soil fungal community. Changes to the arbuscular mycorrhizal fungi (AMF) community were analyzed by spore count of which I found no differences between treatments. The nematode community was analyzed by counting total community and relative abundance of plant parasitic (PPN) to free-living nematodes (FLN). There were no differences in total abundance of nematodes between treatments or in abundance of PPN or FLN. However, I found the relative abundance of PPN to be significantly lower in the C. bursa-pastoris and S. alba treatments compared to the control. Additionally, a greenhouse experiment was conducted to determine if any of the cover crops used can act as hosts for fungal pathogen Ilyonectria liriodendri (PARC60 and PARC393), nematode pathogen Mesocriconema xenoplax, or beneficial AMF Rhizophagus irregularis. The same four cover crops as well as a control of grapevines were grown in soil inoculated with the organism of interest for four months, at the end of which abundance of I. liriodendri and AMF in iv the roots was measured as well as M. xenoplax abundance in the soil. There were no differences in I. liriodendri colonization between cover crops, but all had some colonization suggesting they may be able to act as hosts. None of the brassica cover crops were colonized by AMF, although B. holboelli was colonized by another fungal endophyte. Abundance of M. xenoplax in the soil was significantly lower in soil of all brassica cover crops compared to the grapevine control. v Lay Summary Disease incidence and soil health are intertwined in viticulture as many grapevine pathogens are soil-borne. Brassica plants have biofumigant properties that are known to suppress pathogens. I looked at how using brassica plants as groundcover can alter soil fungal and nematode communities. I grew four brassicas as cover crops in a vineyard. After a season of growth, fungal and nematode communities were analyzed using molecular and microscopy techniques. There were no significant differences in fungal community or nematode abundance. Additionally, I looked at interactions between the same brassicas and soil microbes of interest (two plant pathogens (fungus and nematode), and a beneficial fungus). These plants were inoculated with the microbe and grown in a greenhouse for three months. Interactions were analyzed using molecular and microscopy techniques. None of the brassicas were hosts for the nematode pathogen or beneficial fungus. All brassicas show potential as a host for the fungal pathogen. vi Table of Contents Abstract .................................................................................................................................... iii Lay Summary ............................................................................................................................ v Preface ....................................................................................................................................... v Table of Contents ..................................................................................................................... vi List of Tables ............................................................................................................................ xi List of Figures ........................................................................................................................ xiv List of Abbreviations ............................................................................................................. xix Acknowledgements ................................................................................................................ xx Chapter 1: Background ............................................................................................................ 1 1.1 Soil borne pathogens in vineyards .............................................................................1 1.2 Biofumigant plants .....................................................................................................1 1.2.1 Fungicidal activities ..............................................................................................3 1.2.1.1 Effects on beneficial fungi.............................................................................4 1.2.2 Nematocidal activities ..........................................................................................5 1.2.2.1 Effects on beneficial nematodes ...................................................................6 1.2.3 Bactericidal activities ............................................................................................6 1.2.4 Effects of biofumigants on the overall soil microbial community ...........................7 1.2.4.1 Indirect effects on soil-borne pathogens via changes to the microbial community ...............................................................................................................8 1.2.4.2 Other factors influencing the soil microbial community ..............................9 1.2.5 Variation in activity among brassica species ......................................................10 1.3 Use of biofumigant green manures and seed meals in viticulture .............................11 1.3.1 Grapevine fungal diseases .................................................................................12 1.3.2 Grapevine plant parasitic nematodes .................................................................12 vii 1.3.3 Limitations to biofumigation in vineyards ............................................................13 1.4 Objectives ................................................................................................................14 Chapter 2: Growing brassica cover crops in a vineyard ..................................................... 16 2.1 Background .............................................................................................................16 2.2 Methods ...................................................................................................................19 2.2.1 Vineyard location and site conditions .................................................................19 2.2.2 Cover crops .......................................................................................................19 2.2.3 Establishment of the experiment ........................................................................20 2.2.4 Plot maintenance ...............................................................................................21 2.2.5 Estimated biomass of brassica cover crops .......................................................21 2.2.6 Soil sampling .....................................................................................................21 2.2.7 Do intact brassica cover crops affect soil fungal diversity and community composition? If so, are there differences in effects among brassica species? ...............22 2.2.7.1 Effect of cover crops on AMF abundance ...................................................22 2.2.7.2 Fungal community analysis ........................................................................22 2.2.7.3 Fungal community sequencing ...................................................................22 2.2.8 Do intact brassica cover crops affect soil nematode abundance and community composition? If so, are there differences in effects among brassica species? ...............23 2.2.8.1 Nematode extraction and counting .............................................................23 2.3 Statistical analysis ...................................................................................................23 2.3.1 Fungal community ..............................................................................................23 2.3.2 Nematode community ........................................................................................25 2.4 Results ....................................................................................................................26 2.4.1 Cover crop establishment ..................................................................................26 viii 2.4.2 Do intact brassica cover crops affect soil fungal diversity and community composition? If so, are there differences in effects among brassica species? ...............26 2.4.3 Do intact brassica cover crops affect soil nematode abundance and community composition? If so, are there differences in effects among brassica species? ...............38 2.5 Discussion ...............................................................................................................48 2.5.1 Brassica cover crop growth ................................................................................48 2.5.2 Do intact brassica cover crops affect soil fungal diversity and community composition? If so, are there differences in effects among brassica species? ...............48 2.5.4 How do brassicas affect beneficial fungi AMF in the field? .................................54 2.5.5 Do intact brassica cover crops affect soil nematode abundance and community composition? If so, are there differences in effects among brassica species? ...............55 Chapter 3: Brassica species host status ............................................................................... 58 3.1 Background .............................................................................................................58 3.2 Methods ...................................................................................................................61 3.2.1 Grapevine propagation.......................................................................................61 3.2.2 Greenhouse setup .............................................................................................61 3.2.3 Microbes ............................................................................................................61 3.2.3.1 Fungal pathogen ........................................................................................61 3.2.3.2 Nematode pathogen ...................................................................................62 3.2.3.3 Plant symbiotic fungi ..................................................................................63 3.2.4 Setup and growing conditions ............................................................................63 3.2.5 Sampling and harvest ........................................................................................64 3.2.5.1 Plant measurements ..................................................................................64 3.2.5.2 Soil measurements.....................................................................................64 ix 3.2.6 Do these brassicas act as hosts for Ilyonectria liriodendri? If so, do some brassicas allow for higher levels of colonization than others? ........................................64 3.2.6.1 DNA extraction ...........................................................................................64 3.2.6.2 Digital droplet polymerase chain reaction (ddPCR) assay ..........................65 3.2.7 Do these brassicas act as hosts for Mesocriconema xenoplax (ring nematode)? 65 3.2.8 Do these brassicas act as hosts for AMF Rhizophagus irregularis? ...................66 3.3 Statistical analysis ...................................................................................................66 3.3.1 Ilyonectria ..........................................................................................................66 3.3.2 Ring nematode ..................................................................................................67 3.3.3 AMF ...................................................................................................................68 3.4 Results ....................................................................................................................68 3.4.1 Do these brassicas act as hosts for Ilyonectria liriodendri? If so, do some brassicas allow for higher levels of colonization than others? ........................................68 3.4.2 Do these brassicas act as hosts for Mesocriconema xenoplax (ring nematode)? 70 3.4.3 Do these brassicas act as hosts for AMF Rhizophagus irregularis? ...................72 3.5 Discussion ...............................................................................................................72 3.5.1 Do these brassicas act as hosts for Ilyonectria liriodendri? ................................72 3.5.2 Do some species of brassica allow for higher levels of colonization than others? 73 3.5.3 Do these brassicas act as hosts for Mesocriconema xenoplax (ring nematode)? 76 3.5.4 Do these brassicas act as hosts for AMF Rhizophagus irregularis? ...................77 Chapter 4: Conclusion ............................................................................................................ 81 x 4.1 Do intact brassica cover crops affect soil fungal diversity and community composition? If so, are there differences in effects among brassica species? ...................81 4.2 How do brassicas affect AMF in a field? ..................................................................83 4.3 Do intact brassica cover crops affect soil nematode abundance and community composition? If so, are there differences in effects among brassica species? ...................84 4.4 Do these brassicas act as hosts for Ilyonectria liriodendri? ......................................85 4.5 Do these brassicas act as hosts for Mesocriconema xenoplax (ring nematode)? ....86 4.6 Do these brassicas act as hosts for AMF Rhizophagus irregularis? .........................87 4.7 Strengths and limitations..........................................................................................87 4.8 Concluding remarks .................................................................................................88 Bibliography ............................................................................................................................ 90 Appendix A – Supporting material for chapter 2 .................................................................. 107 A.1 Figures and tables .................................................................................................... 107 A.2 R code ................................................................................................................... 113 A.2.1 Fungal community analyses ............................................................................. 113 A.2.2 Nematode analyses ......................................................................................... 115 Appendix B – Supporting material for chapter 3 .................................................................. 117 B.1 Plant growth .............................................................................................................. 117 B.2 Figures and tables .................................................................................................... 118 B.3 R code ................................................................................................................... 120 B.3.1 Ilyonectria analyses ......................................................................................... 120 B.3.2 Ring nematode analysis ................................................................................... 121 xi List of Tables Table 1. Seeding rates for each brassica species used in the field experiment. Calculations based on recommendations from seed suppliers. .....................................................................20 Table 2. Mean biomass (grams) for individual plants from each brassica treatment. A random individual was chosen from each plot to get the mean weight of individuals grown in the field. Samples collected September 2020. .........................................................................................28 Table 3. Planned comparision results for indices of diversity of the soil fungal community between all brassica cover crop treatments (tillage radish, white mustard, shepherd’s purse, rockcress) and the undistrubed control (n = 3 for control, n = 5 for all other treatments). Diversity measurements as follows: Pielou’s evenness Index, Shannon Diversity Index (SDI), Species richness, and Faith Phylogenetic Diversity (FPD). .....................................................................29 Table 4. Planned comparision results for indices of diversity of the soil fungal community between native brassica cover crop treatments (shepherd’s purse and rockcress) and exotic brassica cover crop treatments (tillage radish and white mustard) (n = 5 for each brassica species). Diversity measurements as follows: Pielou’s evenness Index, Shannon Diversity Index (SDI), Species richness, and Faith Phylogenetic Diversity (FPD). .............................................29 Table 5. One-way ANOVA results for indices of diversity of the soil fungal community among different cover crop treatments (tillage radish, white mustard, shepherd’s purse, rockcress) (n = 5). Block used as a random factor. Diversity measurements as follows: Pielou’s evenness Index, Shannon Diversity Index (SDI), Species richness, and Faith Phylogenetic Diversity (FPD). ......30 Table 6. Spearman correlation coefficients relating estimated brassica cover crop biomass to various diversity measurements (Pielou’s evenness Index, Shannon Diversity Index (SDI), Species richness, and Faith Phylogenetic Diversity (FPD)) (n = 20). Includes all brassica treatments (tillage radish, white mustard, shepherd’s purse, and rockcress). ............................30 xii Table 7. PERMANOVA results for fungal community composition of different cover crop treatments (tillage radish, white mustard, shepherd’s purse, rockcress, and control) at different taxonomic levels (n = 3 for control, n = 5 for all other treatments). Species level analysis includes all detected phylotypes (not all detected to species level). Permutations = 999. ..........31 Table 8. PERMANOVA results for beta diversity of detected fungal OTUs of different cover crop treatments (tillage radish, white mustard, shepherd’s purse, rockcress, and control) (n = 3 for control, n = 5 for all other treatments). Permutations = 999. ......................................................31 Table 9. P-values for analyses run on nematode abundance data. Planned contrasts for Control vs. Brassicas and Native vs. Exotic brassica species. ANOVA run on all brassica species (tillage radish, white mustard, shepherd’s purse, and rockcress) using block as a random factor. PPN (Plant Parasitic Nematode), FLN (Free-living Nemtode). ...........................................................40 Table 10. Spearman correlation analysis comparing estimated brassica cover crop biomass to each nematode measurement (n = 20). Includes all brassica treatments (tillage radish, white mustard, shepherd’s purse, and rockcress). PPN (Plant Parasitic Nematode), FLN (Free-living Nemtode). .................................................................................................................................47 Table 11. Results of a zero inflation model for prediction of excess zeros (no Ilyonectria inside the roots) based on treatment species (tillage radish, white mustard, shepherd’s purse, rockcress, and grapevine) (n = 8). Estimate is binomial logit coefficient for predicting excess zeros. Log likelihood: -153.6. ....................................................................................................69 Table 12. Mean reproductive factor (RF) of ring nematodes grown with different plant species (n = 8). Treatments: tillage radish, white mustard, shepherd’s purse, rockcress, and grapevine). .72 Table A1. Mean biomass (grams) for individual plants from each brassica treatment. A random individual was chosen from each plot to get the mean weight of individuals grown in the field. Samples collected September 2020…………………………………………………………………111 xiii Table A2. Number of reads retained at each step in the DADA2 pipeline. From left to right are initial read counts (input), reads after filtering (filtered), reads after denoising (denoised), and number of reads after chimera removal (non-chimeric)………………………………………...…112 Table A3. Pairwise PERMANOVA results of fungal community beta diversity using Bray-Curtis distance. Permutations = 999………………………………………………………………………...112 Table A4. Pairwise PERMANOVA results of fungal community beta diversity using Unweighted UniFrac. Permutations = 999.....................................................................................................113 Table A5. Pairwise PERMANOVA results of fungal community beta diversity using Weighted UniFrac. Permutations = 999.....................................................................................................113 xiv List of Figures Figure 1. Examples of the diversity of glucosinolate breakdown products. MYR, myrosinase. Adapted from "Glucosinolate structural diversity, identification, chemical synthesis and metabolism in plants," by Blaževića et al., 2020, Phytochemistry, 169, p. 7. ...............................3 Figure 2. Species evenness (Pielou’s evenness index) of fungal communities in soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Values were obtained in QIIME2. Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. Group means sharing the same letter are not significantly different according to the Tukey-Kramer post-hoc test. ............................................................................................................................32 Figure 3. Shannon diversity index of fungal communities in soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; ; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Values were obtained in QIIME2. Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. ANOVA; F = 1.737; df = 4, 18; P = 0.1857; R2 = 0.279. ...................................................................................................................33 Figure 4. Species richness as measured by number of observed OTUs in fungal communities in soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Values were obtained in QIIME2. Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. ANOVA; F = 0.50; df = 4, 18; P = 0.735; R2 = 0.100. ...............................................................................34 Figure 5. Faith’s Phylogenetic Diversity index of fungal communities in soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, xv Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Values were obtained in QIIME2. Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. ANOVA; F = 0.426; df = 4, 18; P = 0.7881; R2 = 0.086. .............................................................................................................35 Figure 6. Mean abundance of OTU’s of different phyla in fungal communities in soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Values were obtained in QIIME2. Black bars represent standard error. PERMANOVA; F = 1.47; P = 0.207; R2 = 0.2463. Pairwise comparisons of treatments that differed significantly; TR vs SP, P = 0.048; RC vs SP, P = 0.025. Dispersion; P = 0.971. .........36 Figure 7. Relative abundance of OTU’s of the ten most abundant classes in fungal communities in soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard. Values were obtained in QIIME2. PERMANOVA; F = 1.320; P = 0.237; R2 = 0.227. Dispersion; P = 0.918. ........................................................................................................................................37 Figure 8. Count of AMF spores in soil samples from plots with different cover crop treatments. Spores extracted from 20 g soil samples. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. .................................................................................39 Figure 9. Abundance of total nematodes in 100 cm3 soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 5). Values were obtained from extracting nematodes from the soil samples and counting under a light microscope. Hollow red circles xvi represent individual replicates, solid red dots and black lines represent mean and standard error. ANOVA; F = 1.08; df = 4, 20; P = 0.392. ..........................................................................41 Figure 10. Abundance of free-living nematodes in 100 cm3 soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 5). Values were obtained from extracting nematodes from the soil samples and counting under a light microscope. Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. ANOVA; F = 1.48; df = 4; P = 0.246; R2 = 0.228. .............................................................42 Figure 11. Abundance of plant parasitic nematodes in 100 cm3 soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 5). Values were obtained from extracting nematodes from the soil samples and counting under a light microscope. Values were log transformed before analysis. Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. ANOVA; F = 1.70; df = 4; P = 0.189; R2 = 0.253. ........................................................................................................................................43 Figure 12. Effect of cover crop treatment on the PPN to FLN ratio. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 5). Data obtained using a mixed effect logistic regression model with plot number as a random factor. Blue points represent mean PPN to FLN ratio and pink lines represent standard error. ANOVA; χ2 = 13.87; df = 4; P = 0.008. Group means sharing the same letter are not significantly different. ....................................................................................................................................44 Figure 13. Odds ratio comparing the occurrence of PPN over FLN given exposure to a brassica cover crop treatment compared to the undisturbed control. Treatments: RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 5). Data obtained using a mixed xvii effect logistic regression model with block as a random factor. Asterisks represent significance levels (* P ≤ 0.05, ** P ≤ 0.01, *** P ≤ 0.001). Lines represent 95% confidence limits. ..............45 Figure 14. Scatterplot of the total nematodes counted and the estimated biomass of brassicas in each plot (n = 20). Black dots represent one plot, blue line represents the least squares line and gray shading represents confidence interval. Spearman rs = 0.49, P = 0.030. ...........................46 Figure 15. Scatterplot of the FLNs counted and the estimated biomass of brassicas in each plot (n = 20). Black dots represent one plot, blue line represents the least squares line and gray shading represents confidence interval. Spearman rs = 0.46, P = 0.043....................................47 Figure 16. Abundance of Ilyonectria beta-tubulin copies per g root in roots of cover crop plants. Only samples with non-zero value are shown. Data was transformed (Box Cox) to achieve normality. Values were obtained by ddPCR. Treatments: RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard. ANOVA; F = 0.68; df = 4; P = 0.621; R2 = 0.214. ........70 Figure 17. Abundance of ring nematodes (nematodes per 100 ml soil) in soil samples taken from pots grown with different plants. Treatments: RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 8). Values were obtained from extracting nematodes from soil samples and counting under a light microscope. Data were log transformed before statistical analysis. Individual replicates are represented by black dots while means are represented by red dots with red lines as standard error. ANOVA; F = 17.90; df = 4; P < 0.001; R2 = 0.672. Group means sharing the same letter are not significantly different according to the Tukey-Kramer post-hoc test. .....................................................................................................71 Figure A1. Principal coordinates analysis of fungal communities from soil of plots treated with growth of different brassica cover crop species. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Bray curtis dissimilarity…………………………………………………108 xviii Figure A2. Principal coordinates analysis of fungal communities from soil of plots treated with growth of different brassica cover crop species. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Weighted unifrac………………………………………………………..109 Figure A3. Principal coordinate analysis of fungal communities from soil of plots treated with growth of different brassica cover crop species. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Unweighted unifrac……………………………………………………..110 Figure B1. Microscopy images of stained grapevine roots containing AMF structures.............119 Figure B2. Microscopy images of stained rockcress roots containing spores of an unknown fungus........................................................................................................................................120 xix List of Abbreviations AMF Arbuscular mycorrhizal fungi ANOVA Analysis of variance cpgr Copies per gram root ddPCR Digital droplet polymerase chain reaction DNA Deoxyribose nucleic acid FLN Free-living nematodes GSL Glucosinolate ITC Isothiocyanate ITS Internal transcribed spacer OTU Operational taxonomic unit PCR Polymerase chain reaction PERMANOVA Permutational analysis of variance PPN Plant parasitic nematodes RF Reproductive factor RO Reverse osmosis SuRDC Summerland Research and Development Centre xx Acknowledgements I would like to thank my fellow grad students and colleagues in the Hart lab as well as those outside the lab who were always there to provide both moral and academic support. I also thank my supervisor Miranda Hart and my committee members Melanie Jones and Tom Forge for always providing valuable insight and feedback and helping me when I had questions. I am grateful for the assistance of Felix from Tantalus Vineyards for allowing me to use his fields for my experiment and always being helpful and easy to work with, as well as Tanis from SeedsCo for providing some of the harder to find seeds essential for my project. A special thank you to my husband who has been nothing but supportive throughout my degree and even helped me with some of the field work. And of course, my mom who has always supported me throughout my education both morally and financially. 1 Chapter 1: Background 1.1 Soil borne pathogens in vineyards The Canadian wine industry is a significant contributor to the Canadian economy, with an estimated economic impact of $9.04 billion in 2015 (www.canadianvintners.com). Soil-borne root diseases result in significant economic loss across vineyards in Canada every year (Úrbez-Torres et al., 2014). This includes grapevine trunk diseases found commonly in the Okanagan such as Black-foot disease and Petri disease caused by soil-borne fungal pathogens (mainly Ilyonectria spp.). In addition to fungal pathogens, plant parasitic nematodes (PPN) cause damage to grapevine roots. Infection with these pathogens can result in reduced yield, stunted growth, and eventual death of the grapevine. 1.2 Biofumigant plants If these pathogens exist in nature, how do plants deal with them? There are some species of plants, found mainly in the Brassicaceae family, that can produce biocidal chemicals known as glucosinolates as a method of defense against herbivores (Matthiessen & Shackleton, 2005). The glucosinolates produced by such plants are broad spectrum biocides that have been shown to also successfully suppress pathogenic bacteria (Park et al., 2012), fungi (Tiznado-Hernandez & Troncoso-Rojas, 2008), nematodes (Dutta et al., 2019), weeds (Norsworthy et al., 2005), and insect pests (Hopkins et al., 2008) when plant tissue is incorporated into soil. Glucosinolates (GSLs) are the main biofumigant compound in brassica plants. GSLs are a class of natural compounds that contain sulphur and nitrogen and are formed by combination of glucose and an amino acid (Sønderby et al., 2010). GSLs alone have no biocidal effects; they must be activated by the enzyme myrosinase (Lazzeri et al., 1993). When a GSL comes into contact with myrosinase in the presence of water, the glucose group is cleaved off. The resulting hydrolysis product is most commonly isothiocyanate (ITC) but can also result in conversion to a nitrile, 2 epithionitrile, or thiocyanate, depending on soil conditions or the presence of specialized plant proteins (Fahey et al., 2001). These GSL hydrolysis products, mainly ITCs, are the compounds responsible for biocidal activity (Figure 1). To prevent constant activation of GSLs, myrosinase is stored separately from GSLs in specialized idioblasts called myrosin cells (Hoglund et al., 1991). Because of this compartmentalization within the plant, some sort of damage to the cells is required to release myrosinase into the environment to allow it to interact with GSLs. 3 Figure 1. Examples of the diversity of glucosinolate breakdown products. MYR, myrosinase. Adapted from "Glucosinolate structural diversity, identification, chemical synthesis and metabolism in plants," by Blaževića et al., 2020, Phytochemistry, 169, p. 7. 1.2.1 Fungicidal activities ITC’s ability to inhibit growth of fungi has been known for decades (Walker et al., 1937). Because of the broad-spectrum suppression mechanism, no one species of fungi is targeted. 4 Studies have found significant variance in fungal toxicity dependent on the structure of the ITC and the species of fungi (Drobnica et al., 1967; Manici et al., 1997; Sarwar et al., 1998). It has been consistently observed that an increase in side chain length of aliphatic ITCs corresponds with a decrease in toxicity (Manici et al., 1997; Sarwar et al., 1998; Stotz et al., 2011). However, correlations between other structure variations and fungal toxicity have been inconsistent. The variation in fungal toxicity among ITCs, combined with the variation in effects on different fungal species, suggests it is likely that the biocidal mechanism may vary from case to case as well. Kojima and Oawa reported that ITCs inhibit the coupling between electron transport and phosphorylation reactions, hindering ATP synthesis (Kojima & Oawa, 1971). Manici et al. (1997) found some ITCs able to inhibit fungal growth but have little to no effect on sporulation. These results combined with their findings on correlations between side chain structure and toxicity lead them to suggest that ITCs work by degrading the lipid bilayer of cell membranes. ITCs have been found to have a growth inhibiting effect rather than a biocidal one unless the concentration is high enough (Smolinska et al., 2003). 1.2.1.1 Effects on beneficial fungi Effects of brassica green manure, seed meal, and ITCs on pathogenic fungi has been the focus in the literature. However, effects on beneficial fungi is less clear. If ITCs have such broad-spectrum toxic effects on soil pathogens, it may similarly inhibit beneficial fungal populations such as arbuscular mycorrhizal fungi (AMF). AMF are symbiotic fungi that penetrate the roots of grapevines and help the plant capture nutrients (Cheng & Baumgartner, 2006). Brassicas are not mycorrhizal due to GSL hydrolysis products inhibiting germination of AMF spores (Vierheilig & Ocampo, 1990). Even without forced activation of GSLs through mulching or otherwise wounding the plant, intact brassicas are able to produce and activate enough GSLs to inhibit AMF spore germination (Schreiner & Koide, 1993a). 5 AMF are considered highly important beneficials for a vineyard system (Trouvelot et al., 2015). Grapevines inoculated with AMF tend to have increased biomass, while grapevines grown in the absence of AMF can become stunted (Menge et al., 1983). Mycorrhizal cover crops are implicated in increased nutrient transfer to grapevines via AMF by creating fungal links between the plants (Cheng & Baumgartner, 2004). Additionally, AMF root colonization has been found to be related to root-knot nematode infection rates. Roots colonized with AMF tend to be more resistant to infection by root-knot nematode (Li et al., 2006). Choosing non-mycorrhizal brassicas as cover crops potentially exchanges the benefits of AMF for increased disease suppression. Although given the inherent variation in GSLs produced by different species of brassica, it may be possible to select for a species that is able to suppress soil-borne pathogens while having minimum inhibitory effects on AMF. 1.2.2 Nematocidal activities Although some PPN are able to reproduce on brassica species (Kruger et al., 2015a), the nematocidal effects of brassica green manures were noted as early as 1925 (Morgan, 1925) and have been extensively studied since (Dutta et al., 2019). ITCs can inhibit nematode growth and motility, preventing them from reproducing or attacking the plant. The effectiveness of ITCs on suppression of nematodes can vary greatly depending on the structure of the amino acid side chain and the species of nematode in question (Lazzeri et al., 1993; Buskov et al., 2002; Zasada & Ferris, 2003). The concentration of ITCs present plays a significant role in nematocidal efficacy (Donkin et al., 1995; Buskov et al., 2002; Wu et al., 2011). Early studies measured nematocidal effect through methods that counted immobile nematodes as dead. In 2004 a study by Lazzeri et al., (2004) found that if the concentration of ITCs is not high enough, the immobilized nematodes could regain mobility. Multiple studies since have confirmed ITCs immobilizing effect on nematodes (Lord et al., 2011; Wu et al., 2011). ITCs can also inhibit nematode eggs from hatching; this was found to occur even at low concentrations (Morgan, 1925; Ellenby, 1945, 1951). 6 More recent studies have confirmed the ovicidal effects of ITCs (Lord et al., 2011; Oliveira et al., 2011; Avato et al., 2013). The biocidal mechanism proposed involves the central carbon of the thiocyanate group (–N=C=S) of ITCs reacting with the target nematode’s essential enzymes causing them to become irreversibly alkylated, inhibiting their functionality (Avato et al., 2013). 1.2.2.1 Effects on beneficial nematodes Although the focus in the literature tends to be on PPNs, there are many free-living species of nematodes that are beneficial for soil health. Free-living nematodes (FLN) influence soil processes by grazing on bacteria, fungi, or other nematodes, thereby playing a role in decomposition and nutrient cycling (Ingham et al., 1985; Freckman, 1988). Studies looking at the effects of biofumigation on FLN populations have had mixed results, some reporting an increase in FLN (Stirling et al., 2003; Valdes et al., 2012), while others reporting no change (Gruver et al., 2010; Vervoort et al., 2014). Daneel et al., (2018) applied brassica cover crops as green manure treatments in two field experiments, one with potatoes and one with tomatoes. They found an increase in beneficial nematodes over time in the potato experiment, but a decrease in beneficial nematodes over time in the tomato experiment. A paper by Gruver et al. (2010), studying the effects of brassica incorporations on nematodes looked in more detail at changes in the FLN community by categorizing them by trophic groups or functional guilds. They found changes within the FLN community varied depending on the species of brassica. Given the various reactions of different nematodes to different brassicas/ITCs, part of this discrepancy may be related to the choice of brassica in each study. Many studies have shown nematode communities to respond differently depending on the species of brassica or type of ITC added (Potter et al., 1999; Zasada & Ferris, 2003, 2004; Reardon et al., 2013; Mocali et al., 2015). 1.2.3 Bactericidal activities ITCs are known inhibitors of bacterial activity and have found uses across industries (Pal Vig et al., 2009). They have mainly been studied for use in the medical industry as antibacterial 7 treatments in humans (Mennicke et al., 1988). Like with other microbes, the type and concentration of ITC as well as the target species plays a significant role in biocidal activities (Wilson et al., 2013). Correlations found between gram negative vs. gram positive susceptibility to ITCs have been inconsistent throughout the literature (Jang et al., 2010; Park et al., 2012; Wilson et al., 2013). Side chain variations such as presence of sulphonyl groups, molecule size, and hydrocarbon chain length seem to have the most influence on the toxicity of a given ITC (Wilson et al., 2013). Evidence has been found for multiple mechanisms through which ITCs inhibit bacterial growth. Most noteworthy include disruption of the cell membrane, inhibition of bacterial metabolic enzymes, and interference with ATP production (Ahn et al., 2001; Luciano & Holley, 2009). The mechanism is likely dependent on the ITC and bacterium involved. Bacterial species that are resistant to ITCs biocidal effects tend to be species that are beneficial for the brassica (Siebers et al., 2018). 1.2.4 Effects of biofumigants on the overall soil microbial community The effects of brassica green manures and seed meals on pathogens and beneficial soil organisms have been considered separately, but it is important to consider the overall influence on diversity and composition of microorganisms in the soil as well. In general, microbial species diversity and composition in soil directly affects soil health (Garbeva et al., 2004). There are many examples in the literature that have found treatment of soils with brassica green manure or seed meal results in changes to the fungal community (Cohen et al., 2005; Larkin et al., 2010; Omirou et al., 2011; Wang et al., 2012, 2014; Mocali et al., 2015). When considering why this pattern occurs, the actual compositional changes should be considered. However, there have been few comprehensive studies on the effects of GSLs and their hydrolysis products on soil microbial diversity that look at species composition changes using high resolution techniques. Hollister et al. (2012) were one of the first to study the effects of ITCs from seed meal extracts on microbial communities using high-throughput sequencing. They found that the addition of ITCs significantly 8 influenced both the fungal and bacterial community compositions, with >60% reduction in fungal diversity and an increase in bacterial taxa related to fungal disease suppression. More recent studies that used high resolution techniques similarly observed a decrease in diversity and a shift in species composition to favour microbes with plant pathogen (including fungal and nematode) suppression properties (Mazzola et al., 2015; Hu et al., 2015; Siebers et al., 2018; Ren et al., 2018b). Regarding nematode populations, a study by Gruver et al., (2010) found R. sativus to increase bacterivore populations and Brassica napus to increase fungivore populations. As bacterivore and fungivore nematodes can prey on plant pathogens, stimulating an increase in their populations may contribute to pathogen suppression abilities of the soil. The results from these studies indicate selection for organisms with plant pathogen suppression capabilities is an important contributor for disease suppression and diversity is not. Additionally, these results show bacteria and fungi with biofumigant resistance tend to be species that are antagonistic to plant pathogens. 1.2.4.1 Indirect effects on soil-borne pathogens via changes to the microbial community When it comes to long term disease suppression, changes to diversity and composition of soil microbial communities may contribute more than direct effects of GSL hydrolysis products alone. There are inconsistencies in the literature on whether community changes resulting from the addition of brassicas can persist or not. Studies by Potgieter et al. (2013) and Fouché et al. (2016) found soil communities influenced by brassicas to be transient, while others found the community changes to persist (Hollister et al., 2012; Mazzola et al., 2015). These studies were done using either brassica green manure or brassica oilseed meal, meaning one-time additions of a large amount of biomass. The community would react and change due to the sudden addition of biofumigants and biomass. Once the GSLs/ITCs are depleted, the soil community may revert to its initial composition, which thrived before addition of brassica biomass. If this is the case, consistent reapplication would be required to maintain disease resistance in the soil. Using 9 brassica cover crops would mean a continuous low level of GSLs and ITCs being present in the soil which may be more efficient at maintaining a disease suppressive community. 1.2.4.2 Other factors influencing the soil microbial community Factors other than GSL hydrolysis products will also have an influence on the diversity and composition of the soil microbial community after the addition of brassica cover crops. One such factor is the GSLs before hydrolysis. Although they may not be active from a biocidal perspective, organisms in the soil may interact with them in different ways. Hanschen et al. (2015) observed that the presence of certain intact GSLs can have an even more significant influence on the bacterial community than GSL degradation products, specifically selecting for species that are able to use the GSL as a carbon source. The method used for treatment with brassicas should also be taken into consideration. Typically, the addition of biofumigants is done through addition of plant biomass such as green manure or brassica seed meal. Omirou et al., (2011) suggested the microbial community changes may be a result of the addition of organic matter (i.e. a sudden shift in nutrient sources) rather than the toxic effects of the ITCs alone. Gruver et al. (2010) also found this to be the case for shifts in the FLN population after brassica incorporation. Brassica plant biomass not only contains components of biofumigation but a variety of nutrients as well. The soil microbial community would respond to not only the addition of GSLs/ITCs but also the addition of a new source of nutrients. Mocali et al., (2015) designed a study to determine if changes in the microbial community were due mainly to ITCs themselves, or the addition of organic matter. They found that although addition of non-biofumigant organic matter did result in significant microbial community changes, the ITCs were necessary for suppression of the pathogen. A study by Mattner et al. (2008) incorporated brassica crops into the soil but did not detect ITCs in the soil, yet they still found effects including weed suppression and differences in pathogen growth. They suggest that allelochemicals other than ITCs may be responsible for some effects of biofumigation. In relation to nematode populations, a study by Vervoort et al. (2014) 10 concluded any effects on the nematode community were in response to the addition of green manure and tillage rather than the ITCs themselves. It is likely both the addition of organic matter as well as the addition of ITCs play a role in shaping the soil microbial community. 1.2.5 Variation in activity among brassica species Although biocidal activities of brassica green manures have been well documented, less is known about variation in efficacy among species. There are many reasons why biocidal activities might differ among species. For example, it is known that different brassica species produce different GSLs (Fahey et al., 2001; Rangkadilok et al., 2002; Bellostas et al., 2004). The biological activity of ITCs is dependent on the structure of the amino acid side chain of its GSL precursor. The variation in structure comes from binding of different amino acids to the glucose molecule as well as secondary modifications to the amino acid side chain. Amino acids used to form GSLs include alanine, leucine, isoleucine, valine, methionine, phenylalanine, tyrosine, and tryptophan (Sønderby et al., 2010). Methionine and phenylalanine may undergo chain elongation before being used for GSL biosynthesis, the amount of elongation is another factor that changes the function of the resulting GSL. Different GSLs may result in ITCs with varying biocidal abilities for specific microorganisms (Manici et al., 1997; Koenraad et al., 2001; Lazzeri et al., 2004). Different brassica species have been found to produce varying amounts of a range of GSLs (Larkin & Griffin, 2007). There is additional variation in both concentration and types of GSLs when looking at different parts of the plant (roots vs. shoots) and in different stages of life cycle (Bellostas et al., 2004; Van Dam et al., 2009; Malik et al., 2010; Bhandari et al., 2015). It must also be considered that different species of plants will produce different levels of biomass that would contribute to the different levels of GSL being produced and expelled into the surrounding soils. This suggests phenology and morphology of brassicas would have an impact on the GSL exudation pattern. These variations in GSL exudation patterns are reflected in the different compositional changes in soil microbial community depending on the types of brassicas added 11 (Hollister et al., 2012; Hu et al., 2015, 2019; Fouché et al., 2016). This has also been reflected in nematode communities that have been reported to vary depending on the brassica/ITC added as well as the family of nematode responding to the treatment (Zasada et al., 2009; Gruver et al., 2010; Reardon et al., 2013; Mocali et al., 2015; Dahlin & Hallmann, 2020). This consistent discrepancy in microbial soil community response suggests it may be possible to select brassica species, based on their GSL exudation profiles, for the needs of an individual field – whether it be in relation to pathogen suppression efficacy, beneficial organism response, or some other factor. 1.3 Use of biofumigant green manures and seed meals in viticulture The potential benefits of brassica green manures and seed meals and their biofumigant properties are well known and have been studied for use in agricultural settings for decades (Ellenby, 1945). Proficiency of brassica green manures and seed meals on suppression of various grapevine pathogens of interest has been explored in the literature. Although studies have found success in suppression of a wide range of grapevine diseases using brassica green manure and seed meal treatments, these treatment methods are not ideal for a vineyard setting. Perennials crops, like grapevines, remain in the soil for years after initial planting. The plants establish a deep root system in the soil that should avoid being disturbed, and makes the soil around the roots prone to build up of soil-borne pathgens. The simplest way to treat soil for soil-borne pathogens is to till fumigants (or biofumigants such as brassica green manure or seed meal) into the soil before planting or after complete harvest which is not possible in an established vineyard. That said, the literature shows successful suppression of grapevine diseases using these brassica treatments. It would be ideal if these suppressive effects could be seen without the need for tilling the brassica plants into the soil; however, there are few studies that look at the exudation of GSLs/ITCs from intact brassica plants. First, I will focus on what we know about suppression of grapevine diseases caused by fungal or nematode pathogens using brassica green manures or seed meals. 12 1.3.1 Grapevine fungal diseases Young vine decline is a grapevine trunk disease that stunts growth, reduces yield, and causes root necrosis in young grapevines (Úrbez-Torres et al., 2014). Young vine decline encompasses two diseases: Petri and black-foot disease. Both of these are fungal disease complexes in which multiple pathogens can be the cause. There are few studies that look at the effects of brassica treatments in suppression of pathogens related to these diseases; however, those that do so show promising results. A study by Bleach (2013) used brassica green manures and seed meals on various black-foot disease isolates and found successful inhibition of germination, colony development, and mycelium growth in all isolates. Berlanas et al. (2018) performed a field study in Spain that applied brassica green manure to infected grapevine soils and found reduced incidence and severity of black-foot disease in the biofumigant treated soils. Richards et al., (2020) found growth of grapevines infected with the black-foot pathogen Ilyonectria liriodendri with white mustard resulted in a lower abundance of the pathogen in the grapevine roots. 1.3.2 Grapevine plant parasitic nematodes There are many PPN that have been reported to cause damage to grapevines, reducing their yield, and leading to economic loss. Some of the most common reported in the Okanagan region include root-knot nematode (Meloidogyne spp.), ring nematode (Mesocriconema xenoplax (Raski) Loof & Degrisse), and lesion nematode (Pratylenchus spp.) (Forge et al., 2021). These PPN are known to infect a variety of hosts, including grapevines. Highly controlled lab studies have been done using GSLs isolated from brassicas and their hydrolysis products and applying them directly to root-knot nematode infested soil. The isolated compounds were consistently found to be effective in suppressing root-knot nematode growth in a lab environment (Lazzeri et al., 2004; Wu et al., 2011; Ren et al., 2018a). Greenhouse studies using brassica green manures or seed meals on soils infested with root-knot, ring, or lesion nematode have shown suppression 13 abilities that appear dependent on the species of brassica used (McLeod & Steel, 1999; Zasada et al., 2009; Kruger et al., 2015a). There are few vineyard field studies that look at the suppression abilities of brassica green manures on PPN and the results are inconsistent. Rahman & Somers (2005) found significant decrease in root-knot nematode populations after application of a brassica green manure that included shallow tillage and incorporation of the green manure. Another field study by Kruger et al. (2015b) grew different brassica cover crops in a vineyard then incorporated them into the soil as green manure. They found no effects on the root-knot nematode population and some brassica species to decrease the ring nematode population, but only in the alleys, not in the vine row. There are many variables that might cause discrepancies in field studies including different application methods, temperatures, soil conditions, and native soil microbial communities. The nematodes having different vulnerabilities to different GSL profiles (from different brassica species) would also play a significant role. Depending on the vulnerability, higher levels of the GSLs may be required before seeing a significant suppression effect. Therefore, the choice of brassica species as well as the amount of green tissue produced at the site before incorporation would be an important factor in determining efficacy of field application of brassicas for PPN suppression. 1.3.3 Limitations to biofumigation in vineyards In perennial systems such as vineyards, chemical fumigant or biofumigant incorporation is done as a pre-plant treatment. Once the grapevines are established, tillage can only occur in the inter-rows as to avoid disturbing the grapevine roots. Incorporation of any treatment into the soil requires some level of tillage. This means the soil immediately surrounding the grapevine roots – where soil-borne pathogens are most likely to gather – would be left untreated. Although treatment of the surrounding soils may still be beneficial, no-till systems have become increasingly common over the years due to the findings of potential detrimental impact of tillage on the health of soil (Sharley et al., 2008; Peregrina et al., 2012). These limitations allow for the accumulation 14 of grapevine trunk diseases and other pathogenic soil organisms. The challenge of suppressing soil-borne pathogen accumulation in vineyards is a constant topic of interest for farmers. Despite this increase in popularity of no-till systems, there are few studies which consider the effects of non-green manure brassica treatments. 1.4 Objectives The aim of this project was to investigate how different species of brassica cover crops influence the soil fungal and nematode communities, including both pathogenic and beneficial organisms, in a vineyard setting. A field study was designed to address the following questions and hypotheses: 1. Do intact brassica cover crops affect soil fungal diversity and community composition? If so, are there differences in effects among brassica species? H0: Soil fungal diversity and community composition will not differ between the control and the brassica treatments or among brassica treatments. H1: All brassica treatments will decrease soil fungal diversity and alter community composition when compared to the control. Differences will be greater in soil under white mustard and tillage radish (exotic species) than in soil under shepherd’s purse and rockcress (native/nativized species). 2. Do intact brassica cover crops affect soil nematode abundance and community composition? If so, are there differences in effects among brassica species? H0: Soil nematode abundance and community composition will not differ between the control and brassica treatments or among brassica treatments. H1: All brassica treatments will decrease soil nematode abundance and reduce PPN in the community. 15 Additionally, I wanted to look at interactions between brassica species and particular organisms of interest. For this, a greenhouse study was designed to address the following objectives: 1. Do white mustard, tillage radish, shepherd’s purse, or rockcress act as hosts for Ilyonectria liriodendri? If so, do some brassicas allow for higher levels of colonization than others? H0: None of the brassica treatments will act as hosts for Ilyonectria. H1: Tillage radish will be the only brassica that will show colonization of Ilyonectria as it has been found as a host in previous literature (Langehoven et al. 2018). Tillae radish will have lower levels of colonization than grapevines. Since the strain of Ilyonectria used was isolated from grapevines, I expect it would be adapted to grow better in grapevines. 2. Do white mustard, tillage radish, shepherd’s purse, or rockcress act as hosts for Mesocriconema xenoplax (ring nematode)? H0: None of the brassica treatments will act as hosts for ring nematode. H1: None of the brassica treatments will act as hosts for ring nematode. I expect this as previous studies have not found brassica species to be suitable hosts. 3. Do white mustard, tillage radish, shepherd’s purse, or rockcress act as hosts for AMF? H0: None of the brassica treatments will act as hosts for AMF. H1: Some of the brassica treatments will act as hosts for AMF. Results from this study could potentially be adapted for use in commercial vineyards to help mitigate disease incidence and maximize soil health in an organic and cost-efficient manner. 16 Chapter 2: Growing brassica cover crops in a vineyard 2.1 Background The biofumigant effects of brassica green manure or seed meal have been studied extensively. In vineyards, tillage, particularly around grape root systems, is not desirable, which limits the potential utility of brassica green manures as biofumigants. Consequently, it is of interest to determine if brassica cover crops that are not incorporated into the soil can affect soil-borne pathogens and the overall soil community in desirable ways. The focus tends to be on interactions with target organisms rather than the soil community. Even fewer studies analyzed the fungal community using high-throughput sequencing techniques. Studies that did use high-throughput sequencing techniques, used brassicas in the form of green manure or seed meal (Hollister et al., 2012; Hu et al., 2015; Siebers et al., 2018). Currently only Cloutier et al. (2020) has done a study that looks at changes to the soil microbial community from intact brassicas using high-throughput sequencing techniques. Previous studies looking at fungal community changes from brassica cover crops alone have not reported diversity changes (Larkin et al., 2010; Wen et al., 2017; Cloutier et al., 2020). However, other studies have consistently demonstrated a decrease in fungal diversity in response to addition of brassicas in the form of seed meal or green manure (Hollister et al., 2012; Hu et al., 2015; Siebers et al., 2018). The literature focus is similar when considering the nematode soil community; biofumigation studies tend to focus on the effects on PPN (Dutta et al., 2019). There are some studies that report on changes to the FLN population as a result of brassica cover crop incorporation, but only one study by Gruver et al. (2010) that considered changes to the FLN using intact brassicas. It is important to understand how using brassica cover crops might alter the soil community. If brassica cover crops can provide the desired disease suppression effects with minimal negative influence on the rest of the soil community, this may provide farmers with a cost effective and organic alternative to chemical fumigation. 17 Brassica cover crops are typically incorporated via tillage into the soil as green manure to maximize the benefits of biofumigation. For a perennial system such as vineyards, this tillage could not happen close to the grapevine roots where the soil-borne pathogens tend to accumulate. In addition, many vineyards avoid tillage outright, even in the interrows. That said, little is known about any impact brassica cover crops may be having on the soil microbial community without incorporation. In this study I wanted to determine if non-green manure brassica cover crops can alter the soil fungal community in a vineyard, and if different species of brassica might have different effects. Four different brassica species were selected for this study: two that are commonly used in Okanagan vineyards but are considered exotic to the region (Sinapis alba L. (white mustard) and Raphanus sativus (L.) Domin (tillage radish)) as well as two that are native or naturalized to the Okanagan (Capsella bursa-pastoris (L.) Medik. (shepherd’s purse) and Boechera holboelli (Hornem.) Á.Löve & D.Löve (rockcress)). The control plots for this study consisted of whatever growth was present to begin with (containing many different plant species), while the treatment plots were managed to ensure the intentionally planted brassica was the dominant species. Therefore, we also needed to consider the effects of plant diversity on the soil fungal community. There is evidence to suggest increased plant diversity results in increased soil microbial diversity (Turrini et al., 2016; Steinauer et al., 2016). Lower plant diversity than the control combined with GSL/ITC production in the brassica treatments lead to the prediction that the fungal community diversity would decrease in response to all brassica treatments compared to the control. Similarly, it was predicted that fungal community composition would be different in all brassica treatments when compared to the control. This prediction was based off results from Cloutier et al. (2020) who studied fungal community changes because of brassica cover crops and found that even if there was no change in overall fungal diversity, there were changes to the community composition. Among brassica species, it was predicted that differences would be due to the exotic vs. native/nativized nature of 18 the cultivars used. A study by Checinska Sielaff et al., (2018) comparing the fungal community in soils grown with either native or exotic plant species dominating the plant community found a significant difference in community composition and function, but not in diversity. As the fungal community present in the current study is likely more adapted to the native or nativized brassica species, I expected the exotic cultivars (tillage radish and white mustard) would result in greater changes to the fungal community composition than the native/nativized (rockcress and shepherd’s purse) brassicas when compared to the control. Shepherd’s purse and rockcress are commonly found throughout the Okanagan. In particular, Shepherd’s purse appeared in abundance in the vineyard used for this study. Regarding beneficial fungi, AMF are highly important in agriculture. AMF are obligate symbiotic fungi that penetrate the roots of a plants and are associated with 80% of terrestrial plant species, including grapevines (Wang & Qiu, 2006). They are known to enhance nutrient uptake and increase biomass of grapevines (Menge et al., 1983). Additionally, grapevine roots colonized with AMF have been found to have increased resistance to some soil-borne plant pathogens (Azcón-Aguilar & Barea, 1996; Li et al., 2006; Petit & Gubler, 2006). Therefore, it is important to understand how our brassica cover crops might interact with AMF in the soil. I predicted that brassica treatments would result in a lower abundance of AMF spores in the soil when compared to the control. This prediction is based on evidence that intact brassica plants, which do not host AMF, have also been able to successfully suppress AMF germination in the soil (Schreiner & Koide, 1993b; Cantor et al., 2011). Reported effects of brassica treatments on the soil nematode community under field conditions are fewer. We know brassica species tend to be uncommon hosts for PPN (Kruger et al., 2015a) and that the ITCs produced by brassicas have been found to be toxic to some nematodes (Dutta et al., 2019). That said, there is a minimum GSL/ITC requirement needed to reduce nematode abundance (Ramirez et al., 2009; Wu et al., 2011; Reardon et al., 2013) and a 19 study by Zasada & Ferris (2003) found that the amount of GSLs added to the soil in field studies had to be 70% higher than that applied in in vitro experiments in order to achieve the same result. All the studies that report changes to the nematode community because of brassica addition under field conditions were using seed meal or green manure. Cover crops that are not tilled into the soil produce less GSLs/ITCs than a seed meal or green manure application (Wen et al., 2017). Therefore, I predicted that the brassica cover crops would not significantly alter the overall nematode abundance or the nematode community composition. 2.2 Methods 2.2.1 Vineyard location and site conditions The field study took place in Tantalus Vineyards, Kelowna, BC, Canada (latitude 49o 50’ 5” N, longitude -119o 27’ 33” W) from May 2020 to September 2020. This region lies within the Okanagan Valley geographical indication for viticulture (http://www.bcvqa.ca/wine-regions-of-bc/). Annual rainfall in this region averages 311.3 mm (Environment Canada). The field in question had been used as a horse pasture until 2012 and was fallow from 2012 to 2020. Grapevines planted in this field were Vitis vinifera cv. Pinot noir (ungrafted) and were planted with one shovel scoop of compost per vine (Glengrow: https://www.kelowna.ca/city-services/okanagan-compost/compost-we-produce/glengrow) at the beginning of the experiment in early June 2020. 2.2.2 Cover crops I used four brassica species to determine the effects of brassica cover crop species on soil fungal community. Two species were cultivars commonly used in agriculture: Tillage radish (Raphanus sativus L.) and white mustard (Sinapis alba L.). The other two species were native or naturalized to the Okanagan Valley: Shepherd’s purse (Capsella bursa-pastoris (L.) Medik.) and rockcress (Boechera holboelli (Hornem.) Á. Löve & D. Löve). S. alba seeds were obtained from Richter’s, Ontario, Canada. R. sativus were seeds obtained from William Dam Seeds, Dundas, 20 Ontario, Canada. C. bursa-pastoris seeds were obtained from Xeriscape Endemic Nursery, West Kelowna, BC, Canada. B. holboelli seeds were obtained from SeedsCo Community Conservation, Kelowna, BC, Canada. 2.2.3 Establishment of the experiment Field plots (approximately 2 metres wide each, centred on the vines) were prepared in May 2020 by removing all growth in the plots by finger weeder (machine with a rotating disk made of strong rubber fingers that rip into the soil approximately 2-4 cm deep) and manual removal of all vegetation apart from control plots where vegetation was not removed. Vegetation in the interrow remained undisturbed for all treatments. Cover crops in the interrow included proso and golden millet, radish, crimson clover, phacelia, forage peas, buckwheat, turnip, mustard, and berseem clover. Seeds for each treatment were weighed ahead of time to achieve their respective seeding rates (Table 1). B. holboelli and C. bursa-pastoris were planted by sprinkling the seeds onto the soil while S. alba and R. sativus seeds were planted 2-3 cm deep individually and covered. Treatments were planted in a 5x5 randomized block design, with each row representing a block and plots within blocks representing experimental units for a total of 25 experimental units. Each plot contained five grapevines evenly spaced which were planted 2 weeks after cover crop seeding. Table 1. Seeding rates for each brassica species used in the field experiment. Calculations based on recommendations from seed suppliers. Brassica species Seeding rate Rockcress 1 g m-2 Shepherd’s purse 1 g m-2 Tillage radish 0.7 g m-2 White mustard 1.1 g m-2 21 2.2.4 Plot maintenance Plants were watered and fertilized as per vineyard management protocols (4.5 metric tonnes/acre of compost (Glengrow) in spring - no additional fertilizer). Vines were irrigated by drip irrigation every other day for two hours at 1.89 L per hour. Interrow cover crops were irrigated by microsprinklers for four hours every two weeks. Water came on in May and continued through the season. Plots were hand-weeded as needed to ensure the treatment species was the dominant undervine cover crop. 2.2.5 Estimated biomass of brassica cover crops To estimate the amount of biomass produced by each of our brassica cover crops, I determined the dry biomass of a randomly chosen single plant from each of the plots (roots included) at the end of the growing season (September 2020). I then estimated the number of individuals in each of the plots using a 50 x 50 cm quadrat, then multiplied the mean dry biomass of the representative individuals by the total number of individuals to obtain a measure of dry biomass/ m2. Vegetation biomass was not measured for control plots. 2.2.6 Soil sampling Three soil cores (3 cm x 20 cm deep) were taken from each plot at the end of the growing season (September 2020), combined in one plastic zipper storage bag and homogenized by shaking the bag before subsequent analyses. A subsample was stored in a paper bag and dried at 40 oC for 48 hours for DNA (Deoxyribose nucleic acid) extraction, soil chemistry, and spore extraction. The dried soil was stored at -20 oC. The remaining wet soil was stored at 4 oC for nematode extraction. Soil pH was determined by mixing a 1:1 ratio of dried soil (10 g) with RO water and measuring using an Orion Star A111 benchtop pH meter (Thermo Fisher Scientific, Waltham MA). 22 2.2.7 Do intact brassica cover crops affect soil fungal diversity and community composition? If so, are there differences in effects among brassica species? 2.2.7.1 Effect of cover crops on AMF abundance To determine the influence of brassica species on AMF abundance in the soil, spore extractions were performed using 20 mg of dried soil from each plot using the protocol described in Gerdemann & Nicolson (1963). 2.2.7.2 Fungal community analysis DNA was extracted from each dried soil sample (500 mg) using FastDNA Spin Kit for Soil (MP Biomedicals, Irvine CA) and eluted with 100 µl DNase-free water. DNA quantification was done using a Qubit 4 Fluorometer (Thermo Fisher Scientific, Waltham MA). Extracted DNA was then sent to the Centre for Comparative Genomics and Evolutionary Bioinformatics (Dalhousie University, Halifax Nova Scotia) for NextGen sequencing with the MiSeq (Illumina, Inc., San Diego CA). Amplicon sequencing of the ITS2 (Internal Transcribed Spacer) sub-region was performed for each plot using primers ITS86F 5′-GTGAATCATCGAATCTTTGAA-3′ and ITS4R 5′-TCCTCCGCTTATTGATATGC-3′ to target all fungi (Vancov & Keen, 2009). Raw reads uploaded on NCBI (http://www.ncbi.nlm.nih.gov/bioproject/753154). 2.2.7.3 Fungal community sequencing Raw sequence data was processed in QIIME2 (version 2020.2, https://qiime2.org). Due to the poor quality of the reverse reads, only forward reads were used for analyses. Of the five control plots, only three plots were successfully sequenced. Sequences were denoised using Divisive Amplicon Denoising Algorithm (DADA2 pipeline) (Callahan et al., 2016) with the specifications: fragment length 6 to 279. Truncation length was determined by where the quality score starts to drop below 25. Features were identified for taxa comparison by using the QIIME2 q2-feature-classifier plugin and the naïve Bayes classifier trained on the UNITE database for 23 QIIME reference sequences for fungi (version 8.2, dynamic) (Abarenkov et al., 2020). Alpha and beta diversity analyses were performed in QIIME2 using the core-metrics-phylogenetics plugin with a sampling depth of 2355 that retained 42.60% of all features in 100% of samples. Alpha diversity metrics used included Shannon diversity index (SDI), Faith’s phylogenetic diversity index (FPD), Pielou’s evenness, and Observed Operational Taxonomic Units (OTU) (species richness). Beta diversity compared differences in OTUs in each plot based on cover crop treatment and metrics included Weighted UniFrac (Lozupone et al., 2007), Unweighted UniFrac (Lozupone & Knight, 2005), and Bray Curtis (Sorenson, 1948) dissimilarities. 2.2.8 Do intact brassica cover crops affect soil nematode abundance and community composition? If so, are there differences in effects among brassica species? 2.2.8.1 Nematode extraction and counting All soil nematodes were extracted from 100 cm3 of wet soil within one month of sampling using a modified wet sieving and sucrose flotation technique (Jenkins, 1964). Nematodes extracted from each sample were counted using a gridded counting dish under an Olympus CK 2 inverted light microscope at 40x magnification (Olympus, Tokyo Japan). To determine the individual abundances of PPN and FLN as well as the ratio of PPN to FLN, PPN and FLN were separately counted in one transect of the counting plate representing 1/12 of the total sample. PPN were distinguished by the presence of a distinct stylet while all other nematodes were considered FLN. 2.3 Statistical analysis 2.3.1 Fungal community Correlation testing to relate mean diversity per cover crop (SDI, FPD, evenness, and richness) with estimated mean brassica biomass per crover crop was done in QIIME2 using the 24 alpha diversity metrics generated with core-metrics-phylogenetics and the qiime diversity alpha-correlation command (Spearman, 1904). Differences in alpha diversity among cover crop treatments were tested using one-way analysis of variance (ANOVA) with cover crop treatment as a fixed factor and block as a random factor (stats package version 4.0.5) (R Core Team, 2021) (R statistical software (version 1.4.1106)). Post hoc comparisions of means were performed using Tukey-Kramer post-hoc test with p set at 0.05 (stats package version 2.0.5) (R Core Team, 2021). Planned contrasts were conducted using the contrasts function (stats package version 4.0.5) (R Core Team, 2021) and the aov function from the car package (version 3.0-10) (Fox & Weisberg, 2019) to compare the control to all brassica treatments and to compare native brassica treatments to exotic brassica treatments. Planned contrast were conducted for fungal species evenness, SDI, FPD, and observed OTUs. Differences in beta diversity of OTUs among cover crop treatments was analyzed in QIIME2 using beta-group-significance (permutational analysis of variance (PERMANOVA), permutations = 999) (Anderson, 2001). Differential abundance of features among cover crop treatments was done using analysis of composition of microbiomes (ANCOM) in QIIME2. Before analysis, features were filtered to those that had a minimum frequency of 10 and appeared in a minimum of 2 samples. ANCOM was run on all features based on differences between treatments. Taxonomic data for the Phylum, Class, and Species levels (levels 2, 3, and 7) were exported from QIIME2 for further analysis in R. PERMANOVA using adonis2 (vegan package version 2.5-7) (Oksanen et al., 2020) was run at each of these taxonomic levels to determine differences in composition among cover crop treatments. A square root transformation was performed on the data before running the analysis to minimize the influence of the most abundant groups. A PERMANOVA was also run on all the phylotypes within each of the classes. If any significance was found, it was followed by a pairwise analysis using the vegan package add-on 25 pairwise.adonis (Martinez Arbizu). All PERMANOVA tests used 999 permutations and Bray for calculating pairwise distances. 2.3.2 Nematode community A one-way ANOVA was run in R studio (stats package version 4.0.5) (R Core Team, 2021) with cover crop treatment as a fixed factor and block as a random factor to determine if brassica treatment had an effect on the nematode population. Brassica species was compared with total nematode abundance, estimated abundance of PPN, and estimated abundance of FLN in the soil. To meet the assumption of normality, FLN values were log transformed before final analysis. To determine if brassicas had an effect on the nematode population when compared to the control, or if native brassica treatments differed compared to exotic brassica treatments, a planned contrast was conducted using the contrasts function (stats package version 4.0.5) (R Core Team, 2021) and the aov function from the car package (version 3.0-10) (Fox & Weisberg, 2019). These planned contrasts were conducted on total nematode abundance, PPN abundance, and FLN abundance. To determine how the treatment species influenced the nematode community composition in terms of PPN to FLN ratio (a binary outcome), a mixed effect logistic regression model was done in R (package lme4 version 1.1-26) (Bates et al., 2015) using block as a random factor and cover crop species as a fixed effect. This model was then run through Anova from the car package (version 3.0-10) (Fox & Weisberg, 2019). This model was selected to show which treatments were more or less likely to alter the relative abundance of PPN to FLN compared to the control. Correlation testing to relate each nematode measurement (total, PPN, FLN, and PPN/FLN ratio) with estimated brassica biomass was done in R studio using the cor_test fuction from the rstatix package (version 0.7.0) (Kassambara, 2021). Estimated biomass did not meet the normality assumption so a non-parametric Spearman correlation was conducted with a confidence level of 0.95 (Spearman, 1904). 26 2.4 Results 2.4.1 Cover crop establishment All brassica cover crops within treatment groups grew at similar rates and with a similar abundance. Seeds started sprouting within the first two weeks of planting, but a lot of weeding was required to maintain the brassica treatment as the dominant species. Tillage radish and white mustard both grew to similar heights and were the tallest of the brassica crops planted. Rockcress was the shortest in height but grew clustered together forming a mat on the soil. At the time of harvest in September 2020, tillage radish had the highest estimated biomass per individual (54.9 g ± 10.4 g), followed by white mustard (19.5 g ± 6.9 g), then rockcress (1.1 g ± 0.2 g), and shepherd’s purse (0.8 g ± 0.2 g) (Table 2). At the time of havest, white mustard and shepherd’s purse were at senescence but shepherd’s purse was further into degradation as it had started senescing in August 2020. Tillage radish was green and seeding, and rockcress was green and at maturity. 2.4.2 Do intact brassica cover crops affect soil fungal diversity and community composition? If so, are there differences in effects among brassica species? In order to determine if brassica cover crops selectively killed certain fungal taxa, I measured four measures of species diversity including SDI, FPD, Pielou’s evenness index, and species richness (total OTUs). I used planned comparisons to compare these diversity metrics between the control and all brassica treatments as well as between native and exotic brassica treatments. I detected a trend for significant differences in fungal community evenness and SDI between brassica cover crops and the control (Table 3). Rockcress treatments had significantly higher evenness than control plots evenness (Tukey-Kramer; P = 0.054) (Figure 2). To compare soil fungal diversity among different brassica cover crop treatments, I used a one-way ANOVA with brassica species as the fixed factors (no control) and block as a random 27 factor. I found no statistical difference among cover crop treatments in terms of species evenness, SDI, species richness, or FPD (Table 5). Across all treatments, average evenness was 0.79±0.03 (Figure 2), average SDI was 5.84±0.29 (Figure 3), average species richness was 173±32 (Figure 4), and average FPD was 46.95±8.01 (Figure 5). In order to test if there was a correlation between the biomass of brassicas in the plot and fungal diversity metrics, I collected the mean weight of each brassica species and multiplied it by the mean number of plants in each plot. I then compared this estimated brassica biomass with species evenness, SDI, species richness, and FPD using Spearman correlation. There was no correlation between estimated brassica biomass and any of the fungal diversity metrics (Table 6). I used the soil fungal community sequencing data to determine fungal species composition. In order to determine if the fungal species composition changed in the brassica treated plots compared to the control plots, I used a planned contrast to compare all brassica treatments to the control plots. No differences were detected in fungal community species composition between control and brassica treatments (P = 0.166). In order to test if there were any differences in fungal community composition between brassica species, I compared all four brassica treatments at the level of Phylum, Class, and species using PERMANOVAs. I did not detect differences in fungal community composition among cover crop treatments in terms of composition when examined at the level of Phylum (Figure 6) (Table 7), Class (Figure 7) (Table 7), or species (Table 7). This was also found to be the case when running PERMANOVA on beta diversity OTUs, regardless of distance metric (Weighted UniFrac, Unweighted UniFrac, Bray-Curtis) (Table 8) (Figure A1, A2, A3). Of the 127,143 sequences examined, 21.03% could not be identified past the phylum/class level. The most abundant class in our communities was Sordariomycetes (44.50% of total reads), which was primarily represented by the family Nectriaceae (43.54% of Sordariomycetes), family Lasiosphaeriaceae (29.97% of Sordariomycetes), and genus Coniochaeta (13.37% of Sordariomycetes). The class 28 Dothideomycetes (16.85% of total reads) was primarily represented by the genus Curvularia (89.58% of Dothideomycetes) (94% of which was species Curvularia ryleyi). The class Agaricomycetes (12.26% of total reads) was primarily represented by the order Agaricales (30.65% of Agaricomycetes) and the family Cantharellaceae (8.47% of Agaricomycetes). Other classes detected compromised less than 2% of total reads. There were no taxa whose abundance differed significantly among treatments (ANCOM). I extracted AMF spores from the soil of each plot and counted them under a microscope. In order to test if brassicas could alter the number of AMF spores in the soil compared to the undisturbed control, I used a planned contrast. I found no differences between brassica treatments and the control (P = 0.955). I also did a planned contrast to determine if there was any differences in native vs. brassica species and found no significant differences (P = 0.958). To determine if there were any differences among brassica treatments in their effect on AMF spore abundance, I did an ANOVA comparing all four brassica treatments. I found no differences in AMF spore count in the soil among treatments (Figure 8). Table 2. Mean biomass (grams) for individual plants from each brassica treatment. A random individual was chosen from each plot to get the mean weight of individuals grown in the field. Samples collected September 2020. Brassica species Mean biomass (g) Standard Error Mean total biomass per plot Standard error Tillage radish 54.89 10.40 585.45 85.03 Shepherd’s purse 0.80 0.24 18.82 3.73 Rockcress 1.07 0.24 116.49 12.43 White mustard 19.48 6.90 355.83 39.94 29 Table 3. Planned comparision results for indices of diversity of the soil fungal community between all brassica cover crop treatments (tillage radish, white mustard, shepherd’s purse, rockcress) and the undistrubed control (n = 3 for control, n = 5 for all other treatments). Diversity measurements as follows: Pielou’s evenness Index, Shannon Diversity Index (SDI), Species richness, and Faith Phylogenetic Diversity (FPD). Table 4. Planned comparision results for indices of diversity of the soil fungal community between native brassica cover crop treatments (shepherd’s purse and rockcress) and exotic brassica cover crop treatments (tillage radish and white mustard) (n = 5 for each brassica species). Diversity measurements as follows: Pielou’s evenness Index, Shannon Diversity Index (SDI), Species richness, and Faith Phylogenetic Diversity (FPD). Diversity measurement P value F Evenness 0.007 9.169 SDI 0.032 5.431 Richness 0.980 0.001 FPD 0.857 0.033 Diversity measurement P value F Evenness 0.943 0.005 SDI 0.265 1.325 Richness 0.324 1.030 FPD 0.215 1.651 30 Table 5. One-way ANOVA results for indices of diversity of the soil fungal community among different cover crop treatments (tillage radish, white mustard, shepherd’s purse, rockcress) (n = 5). Block used as a random factor. Diversity measurements as follows: Pielou’s evenness Index, Shannon Diversity Index (SDI), Species richness, and Faith Phylogenetic Diversity (FPD). Table 6. Spearman correlation coefficients relating estimated brassica cover crop biomass to various diversity measurements (Pielou’s evenness Index, Shannon Diversity Index (SDI), Species richness, and Faith Phylogenetic Diversity (FPD)) (n = 20). Includes all brassica treatments (tillage radish, white mustard, shepherd’s purse, and rockcress). Diversity measurement P value F df Evenness 0.732 0.434 3, 12 SDI 0.682 0.511 3, 12 Richness 0.594 0.657 3, 12 FPD 0.654 0.556 3, 12 Diversity measurement P value Correlation coefficient Evenness 0.833 -0.050 SDI 0.554 -0.141 Richness 0.332 -0.229 FPD 0.282 -0.253 31 Table 7. PERMANOVA results for fungal community composition of different cover crop treatments (tillage radish, white mustard, shepherd’s purse, rockcress, and control) at different taxonomic levels (n = 3 for control, n = 5 for all other treatments). Species level analysis includes all detected phylotypes (not all detected to species level). Permutations = 999. Table 8. PERMANOVA results for beta diversity of detected fungal OTUs of different cover crop treatments (tillage radish, white mustard, shepherd’s purse, rockcress, and control) (n = 3 for control, n = 5 for all other treatments). Permutations = 999. Taxonomic level P value F df R2 Phylum 0.207 1.471 4 0.246 Class 0.237 1.320 4 0.227 Species 0.217 1.167 4 0.206 Method P value F df Weighted UniFrac 0.295 1.127 4, 18 Unweighted UniFrac 0.327 1.028 4, 18 Bray-Curtis 0.363 1.029 4, 18 32 Figure 2. Species evenness (Pielou’s evenness index) of fungal communities in soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Values were obtained in QIIME2. Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. Group means sharing the same letter are not significantly different. a a a a 33 Figure 3. Shannon diversity index of fungal communities in soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; ; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Values were obtained in QIIME2. Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. Group means sharing the same letter are not significantly different. a a a a 34 Figure 4. Species richness as measured by number of observed OTUs in fungal communities in soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Values were obtained in QIIME2. Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. ANOVA; F = 0.50; df = 4, 18; P = 0.735; R2 = 0.100. Group means sharing the same letter are not significantly different. a a a a 35 Figure 5. Faith’s Phylogenetic Diversity index of fungal communities in soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Values were obtained in QIIME2. Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. Group means sharing the same letter are not significantly different. a a a a 36 Figure 6. Mean abundance of OTU’s of different phyla in fungal communities in soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Values were obtained in QIIME2. Black bars represent standard error. PERMANOVA; F = 1.47; P = 0.207; R2 = 0.2463. Pairwise comparisons of treatments that differed significantly; TR vs SP, P = 0.048; RC vs SP, P = 0.025. Dispersion; P = 0.971. 37 Figure 7. Relative abundance of OTU’s of the ten most abundant classes in fungal communities in soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard. Values were obtained in QIIME2. PERMANOVA; F = 1.320; P = 0.237; R2 = 0.227. Dispersion; P = 0.918. Dothideomycetes Eurotiomycetes Leotiomycetes Sordariomycetes Agariomycetes Microbotryomycetes Rhizophyctidomycetes Spizellomycetes Glomeromycetes Mortierellomycetes 38 Figure 8. Count of AMF spores in soil samples from plots with different cover crop treatments. Spores extracted from 20 g soil samples. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. Group means sharing the same letter are not significantly different. 2.4.3 Do intact brassica cover crops affect soil nematode abundance and community composition? If so, are there differences in effects among brassica species? In order to determine if brassica cover crops influenced the abundance of total nematodes in the soil compared to the undisturbed control, I extracted nematodes from the soil of each plot and counted them under a microscope. I then ran a planned contrast to compare all brassica treatments to the undisturbed control. I found no significant difference between the abundance of nematodes in the soil in the control plots compared to the brassica treatment plots (Figure 9). I then ran a planned contrast to determine whether the native brassicas had a different effect on the total nematode abundance than the exotic brassica species. There was no significant a a a a 39 difference in total nematode abundance between native and exotic brassica species. To test whether there were any differences in total nematode abundance among different brassica species, I ran an ANOVA comparing means from all four brassica treatments. Abundance of all nematodes did not differ significantly among brassica cover crop treatments (P = 0.392) (Figure 9). Abundance ranged from 780 nematodes per 100 ml soil to 1932 nematodes per 100 ml soil, with a mean of 1542 ± 625 nematodes per 100 ml soil. Next, I wanted to consider if the brassica treatments had an effect on the nematode community composition, specifically, the PPN and FLN. To do this, while I was counting total nematodes, I counted the number of nematodes in the population which were PPN and the number which were FLN. I ran a planned contrast to compare all brassica treatments to the undisturbed control for both FLN and PPN total abundance. I found no significant difference between the abundance of FLN or PPN in the soil in the control plots compared to the brassica treatment plots (Figure 10, Figure 11). I then ran a planned contrast to determine whether the native brassicas had a different effect on the total PPN or FLN abundance than the exotic brassica species. There was no significant difference in PPN or FLN abundance between native and exotic brassica species. Then, to test whether there were any differences in total PPN or FLN abundance among different brassica species, I ran an ANOVA comparing means from all four brassica treatments. There were no significant differences in PPN or FLN abundance among cover crop treatments (Figure 10, Figure 11). Beyond the total number of PPN and FLN, I wanted to consider the compositional changes in terms of the ratio of PPN to FLN. To do this I compared the ratios of PPN to FLN among all treatments. When looking at ratio of PPN to FLN there was a significant difference (P = 0.008) (Figure 12). Shepherd’s purse (P = 0.036) and white mustard (P = 0.009) had significantly lower ratio of PPN vs FLN when compared to the control (Figure 12). In comparison to the control, shepherd’s purse lowered the odds of finding a PPN by 46% and the white mustard treatment lowered the odds by 52% (Figure 13). 40 To test whether the brassica biomass was related to any changes in the nematode community, I ran a correlation analysis comparing estimated brassica biomass with total nematode, FLN, and PPN abundance as well as with PPN to FLN ratio. A positive correlation was detected when comparing total nematode abundance with estimated biomass (Figure 14) as well as when comparing FLN abundance with estimated biomass (Figure 15) (Table 10). No significant correlations were detected when comparing PPN abundance with estimated biomass or when comparing PPN:FLN ration with estimated biomass (Table 10). Table 9. P-values for analyses run on nematode abundance data. Planned contrasts for Control vs. Brassicas and Native vs. Exotic brassica species. ANOVA run on all brassica species (tillage radish, white mustard, shepherd’s purse, and rockcress) using block as a random factor. PPN (Plant Parasitic Nematode), FLN (Free-living Nemtode). Nematode measurement Control vs. Brassicas Native vs. Exotic All brassica species Total 0.090 0.136 0.202 PPN 0.197 0.30 0.104 FLN 0.043 0.46 0.254 41 Figure 9. Abundance of total nematodes in 100 cm3 soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 5). Values were obtained from extracting nematodes from the soil samples and counting under a light microscope. Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. ANOVA; F = 1.08; df = 4, 20; P = 0.392. Group means sharing the same letter are not significantly different. a a a a 42 Figure 10. Abundance of free-living nematodes in 100 cm3 soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 5). Values were obtained from extracting nematodes from the soil samples and counting under a light microscope. Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. ANOVA; F = 1.48; df = 4; P = 0.246; R2 = 0.228. Group means sharing the same letter are not significantly different. a a a a 43 Figure 11. Abundance of plant parasitic nematodes in 100 cm3 soil samples from plots with different cover crop treatments. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 5). Values were obtained from extracting nematodes from the soil samples and counting under a light microscope. Values were log transformed before analysis. Hollow red circles represent individual replicates, solid red dots and black lines represent mean and standard error. ANOVA; F = 1.70; df = 4; P = 0.189; R2 = 0.253. Group means sharing the same letter are not significantly different. a a a a 44 Figure 12. Effect of cover crop treatment on the PPN to FLN ratio. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 5). Data obtained using a mixed effect logistic regression model with plot number as a random factor. Blue points represent mean PPN to FLN ratio and pink lines represent standard error. ANOVA; χ2 = 13.87; df = 4; P = 0.008. Group means sharing the same letter are not significantly different. a b ab b 45 Figure 13. Odds ratio comparing the occurrence of PPN over FLN given exposure to a brassica cover crop treatment compared to the undisturbed control. Treatments: RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 5). Data obtained using a mixed effect logistic regression model with block as a random factor. Asterisks represent significance levels (* P ≤ 0.05, ** P ≤ 0.01, *** P ≤ 0.001). Lines represent 95% confidence limits. Species of Brassica RC SP TR WM 46 Figure 14. Scatterplot of the total nematodes counted and the estimated biomass of brassicas in each plot (n = 20). Black dots represent one plot, blue line represents the least squares line and gray shading represents confidence interval. Spearman rs = 0.49, P = 0.030. Estimated biomass (gm-2) 47 Figure 15. Scatterplot of the FLNs counted and the estimated biomass of brassicas in each plot (n = 20). Black dots represent one plot, blue line represents the least squares line and gray shading represents confidence interval. Spearman rs = 0.46, P = 0.043. Table 10. Spearman correlation analysis comparing estimated brassica cover crop biomass to each nematode measurement (n = 20). Includes all brassica treatments (tillage radish, white mustard, shepherd’s purse, and rockcress). PPN (Plant Parasitic Nematode), FLN (Free-living Nemtode). Nematode measurement P value rs Test statistic Total 0.030 0.49 684 PPN 0.197 0.30 929 FLN 0.043 0.46 723 PPN:FLN 0.830 0.05 1262 Estimated biomass (gm-2) 48 2.5 Discussion 2.5.1 Brassica cover crop growth Of the similar studies which looked at changes to the soil microbial community as a result of brassica living mulches (Larkin et al., 2010; Wen et al., 2017; Cloutier et al., 2020), only Wen et al. 2017 ended up measuring the brassica treatment biomass. Although the species of brassica used were different, the estimated biomass of brassicas from the current study are within the same range found in Wen et al. 2017. All brassica cover crops established successfully in the plots they were planted in. Additionally, each species grew at a similar rate and was the dominant plant species in their respective plots. This suggests there should not be significant variation between plots of the same species as a result of differences in plant growth. Although the growth within species was similar, there were differences in the amount of growth between different species. Both exotic species – white mustard and tillage radish – grew quickly and took up more space than the native species – shepherd’s purse and rockcress. This difference in growth may have resulted in changes to the community based on biomass production alone. Additionally, the life cycles of each brassica species had different lengths and were at different stages by the time of harvest. Shepherd’s purse grew in abundance in the first half of the season but died off before all the other plants starting in August. This may impact the effect it would have had on the soil microbial community since any changes may have had time to revert to be more similar to the control after the plants reached senescence. 2.5.2 Do intact brassica cover crops affect soil fungal diversity and community composition? If so, are there differences in effects among brassica species? When looking at diversity in terms of SDI, soil fungal community diversity was found to be lower in the control when compared to all brassica treatments, but this was not found to be the case for FPD. The reason SDI is different but FPD is not is because the calculation for SDI uses the species evenness value while FPD is based on species richness. I found significant 49 differences between the control and brassica treatments when looking at species evenness, but not species richness. In particular, the rockcress treatment was found to have significantly higher species evenness when compared to the control which goes against my expectation that tillage radish and white mustard would have a greater influence on the microbial community due to their exotic status in the Okanagan. In fact I found no differences in diversity or composition of the soil fungal community between native and exotic brassica species treatments. Control treatments having a lower fungal diversity was unexpected as I had predicted the opposite – a decrease in diversity for all brassica treatments when compared to the control. These findings are surprising in that the control plots had much higher plant diversity than our brassica plots. Thus, I would expect plots with lower plant evenness to be associated with soils with reduced fungal species evenness, since above and belowground diversity are often highly correlated (Steinauer et al., 2016; Thapa et al., 2021). Additionally, decreases in microbial community diversity as a result of brassica green manure or seed meal addition have been reported consistently in species richness, but not evenness (Hollister et al., 2012; Hu et al., 2015; Siebers et al., 2018). It is unclear why my brassica plots had higher fungal species evenness compared to the control. However, there is evidence that presence of GSLs and their hydrolysis products promote the growth of microbial organisms beneficial for brassicas while suppressing the growth of others (Mazzola et al., 2015; Hu et al., 2015; Siebers et al., 2018; Ren et al., 2018b). It is possible that the presence of GSLs in all the brassica treatment plots was beginning to promote the growth of certain fungal species that were already present in the soil. A higher species evenness has been found to help mitigate plant responses to environmental stressors (Chaparro et al., 2012), so these cover crops resulting in higher species evenness could translate to better overall soil health. It should be noted that the studies which report decreases in microbial community diversity as a result of brassica amendments were performed using brassicas in the form of green manure, seed meal, or ITC isolates which were added once in a high concentration (Hollister et al., 2012; 50 Hu et al., 2015; Siebers et al., 2018). There are far fewer studies looking at the effects of brassica “living mulch” effects on soil fungal communities. Low levels of GSLs and ITCs, such as those associated with living mulches, may be less likely to result in changes to the community (Wei et al., 2016; Ren et al., 2018a; Hu et al., 2019). Nevertheless, the studies are consistent in that they found no significant changes to fungal community diversity from brassica crops alone (Larkin et al., 2010; Wen et al., 2017; Cloutier et al., 2020) which is supported by the lack of changes to species richness found in my study. However, all of these studies still report some sort of change in the microbial community whether it be compositional (Cloutier et al., 2020), functional (Larkin et al., 2010), or in disease suppression (Wen et al., 2017). In the current study I did not analyze function changes or disease suppression, but I did analyze community composition and found no significant differences in species composition between control and brassica treatments or among all treatments. Although the differences between the soil fungal community in the control and brassica treatments was considered significant in both SDI and evenness, the differences between the means was small. Additionally, soil fungal community diversity did not differ in brassica treatments when compared to the control in community composition or species richness. To determine the reasons for the lack of changes to the fungal community, I examined the literature related to brassica crop effects on target organisms. The results from this perspective are mixed. There are studies using intact brassicas that have found levels of ITCs and GSLs produced to be high enough to result in suppressive effects on a target organism (Schreiner & Koide, 1993a; Kirkegaard et al., 2000; Rumberger & Marschner, 2004; Wen et al., 2017) while others report the levels of ITCs and GSLs to be too low for suppressive effects (Choesin & Boerner, 1991; Smith et al., 2004; Watt et al., 2006). The inconsistency of GSL and ITC abundance in soils around intact brassicas is likely related to the level of activation by myrosinase occurring naturally. In the case of plants with biofumigant properties, there are non-biocidal GSLs and their biocidal 51 hydrolysis products (usually ITCs). Although non-biocidal root exudates have been found to have a significant influence on the whole soil microbial community (Hanschen et al., 2015), GSL hydrolysis products are still essential to the suppression of certain species (Mocali et al., 2015). GSL production, as well as release of the myrosinase required for hydrolysis, is dependent on the species of brassica but is also induced by wounding, disease, and hormones (Bellostas et al., 2004). Given the no-till nature of the vineyard used for our experiment, GSL and myrosinase production would need to rely on cell damage occurring naturally. It is possible that the lack of differences in our soil fungal community are due to a low level of GSLs produced and/or activated by our brassica treatments. In addition to myrosinase produced from the brassica plant itself, there are also microbial organisms, including fungi, that produce myrosinase (Sakorn et al., 2002; Rakariyatham et al., 2005). It is possible that the difference in ITC levels is due to the presence or absence of these microbes in the soil (Borek et al., 1996). Aspergillus spp. and Fusarium spp. are some of the more commonly discussed fungi that are known to produce myrosinase (Smits et al., 1993; Sakorn et al., 1999; Rakariyatham et al., 2005) and were not detected in our soil communities. If our field lacked microorganisms that produce myrosinase, then it is not unexpected for abundance of ITC in the soil to be too low to result in changes to the soil fungal community. Lack of differences in the soil fungal community treatments may also be related to time of sampling as the developmental stage of the plant results in different concentrations and profiles of GSLs (Brown et al., 2003; Bellostas et al., 2004; Bhandari et al., 2015; Hanschen & Schreiner, 2017; Yang et al., 2020). In our study, soils were sampled when plants were at different developmental stages: rockcress had not flowered, tillage radish was flowering, and shepherd’s purse and white mustard were senescencing. GSL output is reduced in later developmental stages, especially during leaf senescence (Brown et al., 2003; Yang et al., 2020), thus our shepherd’s purse and white mustard may have had less biocidal activity than the rockcress and 52 tillage radish. However, my results do not support this as I found no difference in diversity among treatments. If the GSL exudates and their hydrolysis products are the main driver for changes in the fungal community, it is possible that the community may have changed significantly at the developmental stage where GSL production is highest, but when GSL exudates decrease the fungal community begins to revert to the equilibrium that existed before the brassica was grown. There is not a consensus in the literature about permenance of changes to the community resulting from addition of brassicas as they have been found to be transient in some studies (Rumberger & Marschner, 2004; Potgieter et al., 2013; Fouché et al., 2016) and persistent in others (Hollister et al., 2012; Mazzola et al., 2015; Hu et al., 2019). However, most previous studies were looking at the addition of concentrated brassica biomass (i.e., green manure, oilseed meal), while I am looking at brassica cover crops. It would be expected that with brassica cover crops, any changes occurring in the community would be more subtle and revert more easily as it is a less extreme alteration to the environment. Given this information, changes to the soil fungal community might occur during certain developmental stages of the cover crop but revert when no longer at that stage. It is also possible that the minimal differences seen in the fungal community could be related to the processing of the soil after sampling. After I took the soil samples on the day of harvest, I dried them in an oven at 40 °C for 48 hours. This was done to ensure a consistent amount of soil was measured for DNA extraction from each plot. Drying soil may have effected the fungal DNA and downstream sequencing results. However, a study by Castaño et al. 2016 found air drying soil before DNA extraction to not have any significant effects on the soil fungal community composition. Therefore, although possible, I do not think it is likely that the drying step in my methodology significantly altered the soil fungal community sequencing Even without significant changes to the fungal community, there may be benefits associated with using brassica cover crops. For example, there may be GSL production sufficient 53 to reduce disease incidence without a concomitant change in soil diversity. Studies have found growth with brassica cover crops to reduce disease incidence in neighbouring plants (Larkin et al., 2010; Larkin & Halloran, 2014; Hossain et al., 2015; Wen et al., 2017; Baysal-Gurel et al., 2020). Not all of these studies looked at changes to soil fungal diversity as well, but those who did found no differences (Larkin et al., 2010; Wen et al., 2017). Furthermore, brassicas may engender increased microbial activity, in the absence of microbial community changes (Cohen et al., 2005; Larkin et al., 2010; Potgieter et al., 2013; Galletti et al., 2015). It is possible that the brassica species were not producing enough GSLs to alter the soil fungal community but may have influenced enzymatic activities of the community already present. 2.5.3 Composition analysis The most abundant phylum in our soil communities was Ascomycota, which is typical of agricultural soils. The most abundant class in our soil communities was Sordariomycetes which is one of the largest classes of Ascomycota and contains a range of fungi including some plant pathogens (Maharachchikumbura et al., 2016). The Sordariomycetes in our soil were primarily represented by the family Nectriaceae which is known to contain some plant pathogens as well as several species used as biocontrol agents (Lombard et al., 2015). The Nectriaceae in this study were not identified past the family level so we cannot say what the functions were for the species found in this community. The next most common phylotype found in Sordariomycetes was the family Lasiosphaeriaceae which are saprobes (Maharachchikumbura et al., 2016). The genus Coniochaeta was also found in abundance in our soil. Species of this genus are known to be pathogens of woody hosts, including grapevines (Damm et al., 2010). There are also some species in this genus that are also known to be beneficial in that they are inhibitory to plant pathogenic fungi (Kokaew et al., 2011). The next most abundant class found in our soil communities was Dothideomycetes which is the largest class of Ascomycota and are mainly composed of saprobes but do contain some 54 plant pathogens (Hyde et al., 2013). This class was mainly represented by the genus Curvularia, most of those identified in this genus were identified to the species Curvularia ryleyi Y.P. Tan & R.G. Shivas. C. ryleyi is known to be a pathogen of the genus Saprobolus (Hetherington & Irwin, 1999). The class Agaricomycetes was the next most abundant. This class belongs to the Basidiomycetes phylum and consists of mushroom-forming fungi (Esser et al., 2014). This class was mainly represented by the order Agaricales which is the largest order of Agaricomycetes and contain mainly saprobes but also ectomycorrhizal fungi and plant pathogens (Esser et al., 2014). The reads found in our study were not identified further than the order Agaricales so I cannot confer what the main role in our community might be. 2.5.4 How do brassicas affect beneficial fungi AMF in the field? Glomeromycota read abundance was not significantly different between treatments and there was no significant difference in AMF spore count in the soil between treatments. Brassica cover crops did not appear to have any influence on the abundance of AMF in the soil. This was unexpected as I had hypothesized that AMF spores would be less abundant in brassica treatments as Brassica sp. are considered non-mycorrhizal and sometimes known to inhibit germination of AMF spores (Vierheilig & Ocampo, 1990; Schreiner & Koide, 1993a; Vierheilig et al., 2000). Greenhouse studies looking at overall Glomeromycota abundance in soils as a reaction to ITC amendments found a decrease in abundance (Siebers et al., 2018) including when applied as living mulch (Cloutier et al., 2020). However, interactions between AMF and brassicas may differ under field conditions. In a field study by Pellerin et al. 2007 they found no evidence of a negative effect of the incorporation of brassica residues on the colonization of crop roots by AMF (Pellerin et al., 2007). Another study by Tong et al. 2014 looked at the interactions between a brassica crop and a mycorrhizal crop when grown together (Tong et al., 2014). They found the presence of a mycorrhizal crop with the brassica stimulated the GSL production/activation through 55 constant invasion/contact with AMF. It is possible that growing the brassica crops in the presence of the mycorrhizal grapevine negated any negative biofumigant effects the brassica would have on AMF if grown alone. 2.5.5 Do intact brassica cover crops affect soil nematode abundance and community composition? If so, are there differences in effects among brassica species? The lack of differences in the abundance of nematodes in the soil among cover crop treatments is expected. I had predicted that the brassica cover crops would not result in a significant difference in abundance between treatments. This is because there is a minimum GSL/ITC requirement to reduce nematode abundance (Ramirez et al., 2009; Wu et al., 2011; Reardon et al., 2013). Although there are many examples in the literature of brassica green manure or seed meal treatment successfully suppressing nematodes in the soil (as reviewed in (Dutta et al., 2019)), a study by Zasada & Ferris (2003) found that the amount of GSLs added to the soil in field studies had to be 70% higher than that applied in in vitro experiments in order to achieve the same result. As cover crops that are not tilled into the soil produce less GSLs/ITCs than a seed meal or green manure application (Wen et al., 2017), it is possible that the cover crops in our experiment did not produce enough GSLs/ITCs to result in any significant changes to the abundance of nematodes in the soil. This is supported by the results of a field study by Stirling & Stirling (2003) who treated soil with brassica residues and found no changes to the population of the target nematode pathogen. They suggest the lack of changes may be due to the degree of tissue disruption being insufficient for producing ITCs at a level high enough to result in significant biocidal effects. Despite the lack of differences in overall abundance, when comparing ratio of PPN to FLN, both shepherd’s purse and white mustard had significantly lower ratio of PPN vs FLN when compared to the control. This was unexpected as I had hypothesized that there would be no changes to the nematode community composition due to low levels of GSLs/ITCs. If the brassica 56 crops were able to produce enough GSLs/ITCs to result in a change to the nematode community composition, the reasoning for the lower ratio of PPN in shepherd’s purse and white mustard may be due to their GSL profiles. Multiple studies have shown nematode communities react differently depending on the types of GSLs/ITCs added (Potter et al., 1999; Zasada & Ferris, 2003, 2004; Reardon et al., 2013; Mocali et al., 2015), including the different GSL profiles produced by different crops. The PPN present in this field may be particularly susceptible to the GSLs that are secreted by shepherd’s purse and white mustard, while some of the FLN present may be stimulated by it. A reason that rockcress may be less efficient in reducing PPN populations relative to FLN is that the rockcress plant is relatively small compared to the other brassicas and its root system does not reach as far into the soil. Many PPN of interest live deep in the soil which may be unaffected by rockcress root presence and exudates (Goodell & Ferris, 1980). Independent of GSL/ITC presence, the ratio of PPN to FLN may be expected to be driven by the amount of vegetation present – more roots to feed on or provide nutrients for nematodes present. However, the amount of brassica biomass in each plot did not appear to play a role in the ratio of PPN to FLN as there was no correlation between estimated brassica biomass and PPN to FLN ratio. Total nematode abundance as well as FLN abundance were found to be positively correlated with estimated brassica biomass while PPN abundance was not. This suggests the brassica biomass was affecting the FLN population more than the PPN population. Our results of nematode abundance being positively correlated with plant biomass aligns with results from a study by Lu et al., 2016 who found the same correlation when they compared plant biomass (mixed brassica and non-brassica) with nematode abundance. Higher plant biomass means increased nutrient cycling and could feasibly allow for an increased free-living nematode population. However, many other studies comparing total nematode population to non-brassica plant biomass have not found a correlation (De Deyn et al., 2004; Viketoft, 2008; Cortois et al., 2017); this may suggest the differences seen in the current study may be due to GSL production 57 (higher biomass suggesting higher GSL production). A study by Viketoft (2008) compared the effects of six non-brassica plant species on the abundance and composition of nematode communities. They found PPN to respond more strongly to changes in plant species rather than changes in plant biomass. They suggest this reaction is related to differences in plant secondary metabolites. PPN interact more directly with the plants present than FLN so it makes sense that they would respond more strongly to plant species as the plants host status or defense mechanisms would directly influence the PPN life cycle. 58 Chapter 3: Brassica species host status 3.1 Background When considering the soil fungal and nematode communities in a vineyard, there are specific species that are of interest. Regarding pathogens, a cover crop that can host them and allows for their reproduction would be a detriment to the vineyard. It is also important to consider interactions with beneficial organisms. If a cover crop can suppress pathogens but similarly suppresses beneficial organisms, it may still result in an overall net loss for the vineyard. Therefore, in this experiment I looked at the status of four brassica species (tillage radish, white mustard, shepherd’s purse, and rockcress) as hosts for two soil-borne grapevine pathogens: one nematode and one fungus, as well as one beneficial fungus. Species from the genus Ilyonectria are the main soil-borne fungal pathogens responsible for grapevine trunk diseases such as Petri disease and black-foot disease, which cause young vine decline and result in significant economic losses in the Okanagan as well as other wine grape regions around the world (Úrbez-Torres et al., 2014; Gramaje & Marco, 2015). Although most commonly found in woody plants (Tewoldemedhin et al., 2011; Probst et al., 2019; Farr & Rossman, 2021), Ilyonectria species have been found to colonize and produce disease symptoms in a wide range of non-woody plant hosts (Agustí-Brisach et al., 2011; Cabral et al., 2012; Aiello et al., 2017; Langenhoven et al., 2018; Farh et al., 2018). I wanted to know if the brassica species used in this study are susceptible to colonization by Ilyonectria liriodendri (Halleen, Rego & Crous) Chaverri & C. Salgado, an Ilyonectria species commonly responsible for young vine decline in grapevines. Therefore, I conducted a greenhouse experiment to determine if candidate brassica cover crops could act as a host for I. liriodendri. If these four brassicas are being used in a vineyard and they are able to host Ilyonectria to a degree where they are increasing the amount of the pathogen and allowing them to persist in the soil, this would make them a poor cover crop choice. I predicted that only tillage radish would be successfully colonized, as a previous study looking at 59 Ilyonectria root colonization in various weeds and rotation crops found colonization in the same tillage radish species but none in white mustard (Langenhoven et al., 2018). This same study leads me to believe I will not find any colonization in my white mustard either. There have been no previous studies specifically looking at colonization of Ilyonectria in the other three brassica species. However, although not ubiquitous, previous studies have found colonization of black foot disease pathogens in multiple brassica species (Agustí-Brisach et al., 2011; Langenhoven et al., 2018). This inconsistent ability to colonize brassica species, as well as the evidence in the literature of brassica green manure (Berlanas et al., 2018) and intact brassicas (Richards et al., 2020) successfully suppressing Ilyonectria, caused me to predict neither shepherd’s purse nor rockcress would be colonized by Ilyonectria. If colonization occurred in tillage radish as well as any of the other brassica species, I expected it to occur at a lower abundance when compared to grapevines. This is because grapevines are a known host for Ilyonectria (Pathrose et al., 2014; Holland et al., 2019; Probst et al., 2019) while brassicas are known for having a biofumigant defense mechanism against fungal pathogens (Walker et al., 1937; Sarwar et al., 1998; Tiznado-Hernandez & Troncoso-Rojas, 2008). Ring nematode (Mesocriconema xenoplax (Raski) Loof & Degrisse) is an ectoparasitic nematode that stunts vine growth and is widespread across vineyards in the Okanagan Valley (Forge et al., 2020). Although ring nematode is typically discussed as a parasite of woody plants, there are many common legumes, grains, and weeds that have been shown to be suitable hosts (Nyczepir & Bertrand, 1990; Zehr et al., 1990; Nyczepir, 2011). If a species of plant being used as a cover crop is a good host for ring nematodes, this would increase the abundance of ring nematodes in the soil and could result in unhealthy grapevines and economic losses for the vineyard. For this reason, I wanted to determine if the brassica cover crops studied in this experiment are able to act as hosts for ring nematodes. There is little in the literature on the host status of brassica plants for ring nematodes. One study by Kruger et al., (2015a) included some 60 brassica species when looking at the host status of various cover crops. They found brassicas to be poor hosts, at best, with much lower abundance of ring nematodes in the soil when compared to the known host. I predicted similar results for our own study. I expected grapevine to be a good host for ring nematodes (reproductive factor (RF) >1) and all brassica species to be resistant to ring nematodes (RF ≤1). AMF are obligate symbiotic fungi that penetrate the roots of plants and are associated with 80% of terrestrial plant species, including grapevines (Wang & Qiu, 2006). Because of the variety of advantages associated with AMF symbiosis, the presence of AMF in vineyard soils is of key concern. The literature is uncertain as to the role cover crops may play in AMF colonization of grapevines. Some studies have implicated the presence of mycorrhizal cover crops to influence the association between AMF and grapevines (Cheng & Baumgartner, 2004; Radić et al., 2012). However, another study by Baumgartner et al., (2010) found no correlation between mycorrhizal cover crops and the colonization of AMF in grapevines. Although there is evidence of brassica species being colonized by AMF (Vierheilig et al., 2000; Orłowska et al., 2002; Regvar et al., 2003; Vogel-Mikuš et al., 2006; Pongrac et al., 2007), brassica plants are largely considered non-mycorrhizal (Walker et al., 1937; Vierheilig & Ocampo, 1990; Sarwar et al., 1998). The AMF host status of cover crops may influence the interactions between AMF and grapevines; therefore, I wanted to determine if the brassica species in this study are able to be colonized by AMF. In particular, I used Rhizophagus irregularis (Błaszk., Wubet, Renker & Buscot) C. Walker & A. Schüßler, a species of AMF commonly used as an inoculant in agriculture including vineyards. Due to the low amount of brassica species reported to allow AMF colonization (Newman & Reddell, 1987; Demars & Boerner, 1996), I predicted that none of our brassica species will be colonized by AMF. 61 3.2 Methods 3.2.1 Grapevine propagation Vitis vinifera L. cv. Chardonnay clone 76 cuttings were collected from Tantalus Vineyards in November 2019 and placed in storage at 4 oC until propagation. Cuttings were removed from storage December 2019, trimmed to 2-3 nodes, and placed in buckets with water in a greenhouse at SuRDC. In February 2020, cuttings with a sufficient root system as well as leaves growing out of two nodes were planted in 7 cm pots in Sunshine All-Purpose Potting mix (Sun Gro, Agawam MA). The grapevines were fertilized with 20-20-20 after potting. They were moved to a greenhouse on the University of British Columbia Okanagan campus in Kelowna, BC in March 2020 and set to water on drip irrigation for 1 minute every day at 5 am. Grapevines were pruned, and roots were trimmed when they were transferred to the experimental pots in August 2020. 3.2.2 Greenhouse setup Treatments included four candidate brassica cover crops (tillage radish, white mustard, shepherd’s purse, and rockcress) as seeds or seedlings, and a control consisting of single grapevines from the previously propagated stock. In August 2020, each plant (brassica or grapevines) was inoculated with either a vine pathogen (ring nematode or I. liriodendri) or mutualist (AMF). Each inoculation group contained eight replicate pots of each plant treatment for a total of 40 pots in each group with 120 pots total. Pots from all treatments were completely randomized on a greenhouse bench and left to grow for three months. 3.2.3 Microbes 3.2.3.1 Fungal pathogen Ilyonectria liriodendri (SuRDC 60 and 393) (Halleen, Rego & Crous) Chaverri & C. Salgado was selected as a pathogen of interest as it is a known soil-borne grapevine pathogen 62 (Úrbez-Torres et al., 2014). I obtained cultures from SuRDC where they had been previously isolated from vineyards in British Columbia (Úrbez-Torres et al., 2014). The isolate was grown and maintained by inoculating 5% potato dextrose agar (PDA) plates by cutting a 1 cm2 slice of colonized agar and placing it upside on a new PDA plate. The spores used were extracted from plates that had been inoculated and incubated at room temperature for one month. Spores were extracted by flooding the plates with 1% Tween solution and gently scraping the agar surface with a metal utensil before pouring the liquid through a cheesecloth into a collection container. Spores were observed under a compound light microscope (Nikon Eclipse E800) to ensure there was no contamination. The stock spore solution was diluted to 106 spores/ml. To inoculate plants with Ilyonectria, 1 ml of the spore suspension (106 spores/ml) was pipetted directly either onto the soil where the brassica seeds or seedlings were planted, or, in the case of the grapevines, it was pipetted directly onto the roots before transplanting. 3.2.3.2 Nematode pathogen Ring nematode (Mesocriconema xenoplax (Raski) Loof & Degrisse) was chosen as the PPN of interest as it is a known grapevine pathogen (Forge et al., 2020). The population of nematodes used was originally isolated from a block of grapevines on the grounds of SuRDC in 2006. The population has since been maintained via serial transfer on rooted cuttings of grapevines in greenhouse pots. The inoculum was prepared by harvesting a pot, removing the roots, homogenizing the soil, and extracting nematodes from a subsample for enumeration using a modified wet sieving and sucrose flotation technique (Jenkins, 1964). The nematodes/cm3 of soil was determined to be 12 nematodes/cm3. Soil was stored at 4 oC until ready to be used as inoculant. To inoculate the plants with ring nematode, 200 ml (approximately 2400 nematodes) of infested soil was mixed by hand into the top 5 cm of soil in the pot before planting the seeds/seedlings/grapevines. 63 3.2.3.3 Plant symbiotic fungi AMF Rhizophagus irregularis (Błaszk., Wubet, Renker & Buscot) C. Walker & A. Schüßler was chosen as one that is commonly used as an inoculant in vineyards and has been found to have a positive impact on grapevine growth (Linderman & Davis, 2001). Inoculant powder (Premier Tech Biotechnologies, Rivière-du-Loup QC) was obtained as 6,000 spores/g and stored at 4 oC until use. To inoculate plants with AMF, 0.1 g of powdered inoculant (600 spores) was weighed out and added to the soil directly before adding the seeds/seedlings. For the grapevines, the powder was added directly to the roots before transplanting. 3.2.4 Setup and growing conditions Plants were planted and grown in 2 L pots in Sunshine Sphagnum Peat Moss, play sand (Sakrete, Atlanta GA), and vermiculite (Specialty Vermiculite, Enoree SC) (1:1:1 by volume) on August 20, 2020. Rockcress and shepherd’s purse seeds were planted in seeding trays (one pinch of seeds per cell) in play sand (Sakrete, Atlanta GA) and incubated in a growth chamber for approximately three weeks before experiment setup. This was done because rockcress and shepherd’s purse seeds take longer to establish. Tillage radish and white mustard seeds were sown in the experimental pots the day of setup (five seeds per pot). When planting shepherd’s purse and rockcress, all seedlings from one cell were used per pot. After setup, the pots were watered by hand daily for the first two weeks, then every other day until harvest in November 2020. Tillage radish was thinned to 1 plant per pot, white mustard 1-2 plants per pot, and shepherd’s purse and rockcress 2-4 plants per pot one week after planting. Number of plants per pot was decided by size of the plant (fewer individuals for larger species). Rockcress grows better when there are multiple individuals together. 64 3.2.5 Sampling and harvest 3.2.5.1 Plant measurements All plants (shoots and roots) were removed and stored in plastic bags at 4 oC until use in downstream analysis (2-7 days). The wet weight of roots and shoots of all samples was taken within two days of harvest; roots were washed with reverse osmosis (RO) water before weighing. Tillage radish root weights were taken with the tuber as well as with only the adventitious roots. The grapevine stem was cut off before weighing the roots and shoots. A subsample of the roots from the AMF treatments were taken and stored in plastic bags with a small amount of RO water at 4 oC until ready for root staining. Shoots for all plants as well as roots from the ring nematode treatments and any roots remaining from the AMF treatments were stored in paper bags and dried at 45 oC for 48 hours. Once dried, dry weights were taken for roots and shoots. 3.2.5.2 Soil measurements Soil was homogenized by hand and a subsample was stored in a plastic bag at 4 oC for nematode extraction (ring nematode inoculated pots). Remaining soil was dried in a paper bag at 45 oC for 48 hours then transferred to a plastic bag and stored at -20 oC until use in DNA extraction. 3.2.6 Do these brassicas act as hosts for Ilyonectria liriodendri? If so, do some brassicas allow for higher levels of colonization than others? 3.2.6.1 DNA extraction Roots from Ilyonectria-inoculated samples were surface sterilized with 10% bleach for 5 minutes, then rinsed with RO water. This process was repeated 5 times for each sample. The sterilized roots were patted dry and stored in 50 ml Falcon tubes at 4 oC until DNA extraction. The surface sterilized roots were crushed using a mortar and pestle with liquid nitrogen. The weight of the mashed roots was recorded and in most cases all of the roots was used for 65 DNA extraction (to a maximum of 250 mg). For grapevine roots, parts of the roots were randomly cut from different areas to be used for the extraction. For tillage radish, only adventitious roots were used. The DNA extraction was performed using the kit DNeasy PowerSoil HTP 96 (QIAGEN, Hilden Germany). The manufacturer protocol was followed with the following change: centrifugation at 1850 x g instead of 4500 x g but for 40% longer. DNA was eluted in 100 µl DES and stored at -20 oC. 3.2.6.2 Digital droplet polymerase chain reaction (ddPCR) assay To assay the abundance of Ilyonectria in roots I used an assay designed by Holland et al. (2019). Briefly, this assay targets the highly conserved beta-tubulin region of the Ilyonectria genome using a reaction volume of 20 µL (10 µL Supermix (Supermix for probes no dUTP by Bio-Rad Inc., Hercules, CA, USA), 7 µL DNAse free water, 1 µL primer/probe, and 2 µL DNA. Droplets were created using the Bio-Rad QX100 Droplet Generator using the total reaction volume per sample and 70 µL of Bio-Rad Droplet Generator Oil for Probes. PCR was run in the C1000 Thermal Cycler (Bio-Rad) with the following conditions: initial heating at 95 ◦C for 10 min, 94 ◦C for 1 min, and annealing at 59 ◦C for 2 min × 44 cycles. Fluorescence was measured using the QX 100 Droplet Reader (Bio-Rad) and QuantaSoft Analysis Pro software (version 1.0.596. Bio-Rad). The threshold was set manually at 7108 using pure positive controls as a reference. For analysis, the copy number of each sample was back calculated to represent the number of copies per gram of root (cpgr) using a formula described in Kokkoris et al. (2019). 3.2.7 Do these brassicas act as hosts for Mesocriconema xenoplax (ring nematode)? Nematodes were extracted from 100 cm3 of soil from pots using a modified wet sieving and sucrose flotation technique (Jenkins, 1964). Ring nematodes were counted in a counting dish under an inverted light microscope (Olympus CK 2) by counting half the dish and multiplying by 2 to get nematodes/100 cm3 soil. 66 3.2.8 Do these brassicas act as hosts for AMF Rhizophagus irregularis? To determine if brassica species are able to be colonized by AMF, I detected AMF colonization in roots (no spores) using the following protocol, based on Vierheilig et al., (1988): roots were cleaned with RO water and loosely wrapped in a cheese cloth before being placed in a Fisherbrand TRUFLOW Tissue Cassette (Thermo Fisher Scientific, Pittsburgh PA). Cassettes were then placed in a jar filled with 10% (w/v) KOH (preheated to 90 oC) and placed in a 90 oC water bath for five minutes. The cassettes were then washed three times with RO water. The jar was filled with 5% ink-vinegar (black Sheaffer ink, noname vinegar (5% acetic acid)) solution and left for 4 minutes at 90o C. The ink-vinegar solution was then removed and the cassettes de-stained in RO water with a few drops of vinegar for 20 minutes. Cassettes were then rinsed with RO water until there was no more ink coming out and stored in RO water at 4 oC. For the grapevine roots this protocol was altered as follows: cassettes left in 10% KOH for three hours at 90 oC then overnight at room temperature in fresh 10% KOH, washed five times with RO water, left in 5% ink-vinegar solution at 90 oC for 15 minutes. Roots from each pot were then cut to approximately 20 mm lengths and 15 pieces were placed on a glass slide and preserved with polyvinyl alcohol-lactic acid-glycerol (PVLG). Roots were observed under a light microscope (Nikon Eclipse E800) to determine presence/absence of AMF colonization. Consistent presence of AMF structures (spores, arbuscules, and hyphae) in five or more root pieces was considered positive for presence of AMF. AMF inside the roots was not quantified. 3.3 Statistical analysis 3.3.1 Ilyonectria All statistical analysis was done in R statistical software (version 1.4.1106). As the results contained a high number of zeros, a zero inflated negative binomial regression was used for this 67 analysis. This model has two parts – a binary model to model which process the zero outcome is associated with, and a count model (negative binomial) to model the count process. For this model, the cpgr values were converted to integers. The function zeroinfl from the package pscl (version 1.5.5) (Zeileis et al., 2008) was used to create the model for cpgr as a function of cover crop species. To determine if, among the samples that tested positive, there was any difference among treatments in level of colonization, a regression model was done. First, all replicates with zero cpgr in the roots were removed from the analysis. To normalize the data, a boxcox transformation was performed using the boxcox function from the package MASS (version 7.3-53.1) (Venables & Ripley, 2002) with λ = -0.35. A one-way ANOVA was run with species as fixed factor using the anova command from the stats package (version 4.0.5) (R Core Team, 2021). In order to establish if level of Ilyonectria colonization was correlated with plant root weight, a correlation test was conducted. Root wet weight was first normalized by log transformation. The boxcox transformed cpgr was then compared with the log transformed root wet weight using the cor_test function from the package rstatix (version 0.7.0) (Kassambara, 2021) with the method pearson (Pearson, 1895) and a confidence level of 0.95. 3.3.2 Ring nematode Since ring nematode abundance data did not meet the equal variance assumption (Levene test; P < 0.05) (car package version 3.0-10) (Fox & Weisberg, 2019), a log transformation was performed. Using the log transformed data, an ANOVA was performed using the anova function from the stats package (version 4.0.5) (R Core Team, 2021) to compare abundance of ring nematodes as a function of species of plant (i.e., the four brassicas and the grapevine). This was followed by a Tukey-Kramer post-hoc test (function TukeyHSD from the stats package (version 4.0.5) (R Core Team, 2021)) to determine which groups were significantly different. The reproductive factor (RF = Final Population/Initial Population) was calculated to determine host 68 status of each treatment. ANOVA was performed on these values to determine differences in host status among treatments. Classification of host status was based on Oostenbrink (1966) with values >1 considered as susceptible hosts and <1 considered as poor to resistant hosts. 3.3.3 AMF No statistical analysis was done on AMF as only the grapevine treatment was found to have any positive colonization. 3.4 Results 3.4.1 Do these brassicas act as hosts for Ilyonectria liriodendri? If so, do some brassicas allow for higher levels of colonization than others? To determine if any of the plants used were able to act as hosts for I. liriodendra, I extracted DNA from the roots of each plant and ran it on ddPCR using a primer for the β-tubulin gene of I. liriodendra. First, I looked at how many plants from each treatment had a positive result for presence of Ilyonectria. Fifty percent of grapevines, fifty percent of white mustard, fifty percent of shepherd’s purse, twenty-five of rockcress, and 12.5% of tillage radish replicates were positive for Ilyonectria. Next, in order to determine if some species allowed for higher levels of colonization than others, I looked at the amount of β-tubulin gene cpgr as a proxy for amount of Ilyonectria in each plants roots. Values of Ilyonectria β-tubulin cpgr in positive samples ranged from 347 cpgr (low in grapevines) to 8150 cpgr (high in shepherd’s purse). To test whether any of the treatments were a significant predictor for predicting excessive zeros, as well as to test if there was a correlation between the amount of Ilyonectria in the roots and a treatments species, I ran a zero inflated negative binomial regression analysis. None of the treatments were found to be a significant predictor for predicting excessive zeros (i.e., lack of colonization) (Table 11). Among replicates that tested positive for colonization, I found no significant difference in abundance of Ilyonectria among treatments (P = 0.621) (Figure 16). 69 Table 11. Results of a zero inflation model for prediction of excess zeros (no Ilyonectria inside the roots) based on treatment species (tillage radish, white mustard, shepherd’s purse, rockcress, and grapevine) (n = 8). Estimate is binomial logit coefficient for predicting excess zeros. Log likelihood: -153.6. Plant P value Estimate z-value Rockcress 0.309 1.100 1.018 Shepherd’s purse 0.999 0.001 0.001 Tillage radish 0.129 1.947 1.518 White mustard 0.999 0.001 0.001 Grapevine 0.999 -0.001 -0.002 70 Figure 16. Abundance of Ilyonectria beta-tubulin copies per g root in roots of cover crop plants. Only samples with non-zero value are shown. Data was transformed (Box Cox) to achieve normality. Values were obtained by ddPCR. Treatments: RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard. ANOVA; F = 0.68; df = 4; P = 0.621; R2 = 0.214. 3.4.2 Do these brassicas act as hosts for Mesocriconema xenoplax (ring nematode)? To test whether any of the species of brassicas used in this experiment were able to act as hosts for ring nematodes, I extracted nematodes from the soil of each ring nematode inoculated pot and counted the ring nematodes under a microscope. I then compared all ring nematode counts from each treatment plant with each other using an ANOVA. Ring nematode abundance differed significantly among treatments (P < 0.001) (Figure 17). All brassica species had significantly lower nematode abundance than the grapevine control (Tukey-Kramer; P < 0.001). Tillage radish had significantly lower abundance of ring nematodes than rockcress (Tukey-Kramer; P = 0.055). 71 To determine the host status of each treatment plant, I compared the abundance of ring nematodes counted at the end of the experiment to the amount inoculated at the beginning to find the ring nematode RF for each plant species. I compared the RF among treatments and found ring nematode RF differed significantly among treatments (P < 0.001). Grapevine was significantly higher than all brassica treatments (Tukey-Kramer P < 0.001). Tillage radish had a significantly lower RF when compared to rockcress (Tukey-Kramer; P = 0.055). The mean RF for grapevines was >1 while all brassica treatments had RF <1 (Table 12). Figure 17. Abundance of ring nematodes (nematodes per 100 ml soil) in soil samples taken from pots grown with different plants. Treatments: RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 8). Values were obtained from extracting nematodes from soil samples and counting under a light microscope. Data were log transformed before statistical analysis. Individual replicates are represented by black dots while means are represented by red dots with red lines as standard error. ANOVA; F = 17.90; df = 4; P < 0.001; R2 = 0.672. Group means sharing the same letter are not significantly different according to the Tukey-Kramer post-hoc test. a bc bcd bd bcd 72 Table 12. Mean reproductive factor (RF) of ring nematodes grown with different plant species (n = 8). Treatments: tillage radish, white mustard, shepherd’s purse, rockcress, and grapevine). 3.4.3 Do these brassicas act as hosts for AMF Rhizophagus irregularis? To determine if and of the brassica species used were able to act as hosts for AMF R. irregularis, I stained the roots for fungal structures. I then looked at the roots under a microscope to see if any AMF structures could be seen. None of the brassica treatments were colonized by AMF, while six of the eight replicates for grapevines were positive for AMF root colonization (Figure B1). Although no AMF structures were identified in rockcress roots, four of the eight replicates appeared to be colonized by another fungal endophyte (Figure B2). 3.5 Discussion 3.5.1 Do these brassicas act as hosts for Ilyonectria liriodendri? I found evidence that all our brassica species were able to act as Ilyonectria hosts in some capacity. This was unexpected as I had predicted only tillage radish would show any level of Ilyonectria inside the roots as it was the only one that has shown colonization in previous literature. I was surprised to find Ilyonectria in white mustard in particular, as a previous study by Langenhoven et al., (2018) studied the roots of white mustard plants exposed to Ilyonectria and Plant Mean RF Standard error Rockcress 0.43 0.07 Shepherd’s purse 0.31 0.06 Tillage radish 0.17 0.02 White mustard 0.23 0.07 Grapevine 1.85 0.29 73 other black foot related pathogens found no evidence of root colonization. The same study found evidence of Ilyonectria colonization in tillage radish at a similar rate as our study (Langehoven et al. 2018). Although all brassica species in our study contained some replicates positive for Ilyonectria, none of the treatments had more than 50% of replicates positive for Ilyonectria, including the grapevine control. This result is unusual for grapevine, as Ilyonectria is a serious pathogen of grapevines (Pathrose et al., 2014; Holland et al., 2019; Probst et al., 2019). Such inconsistent colonization across all plant species in our study may be related to other factors beyond host suitability. For instance, Ilyonectria requires a wound in the root to be able to enter. All cover crop species were grown with the inoculum from seed or from intact seedlings. The chances of colonization may have been reduced due to the treatment plants not being purposefully macerated. They would have had to rely on maceration occurring naturally due to insects, nematodes, or abrasion with soil particles. However, this would not apply to the grapevines as they were trimmed at the time of inoculation. Further, inoculant abundance may have been too low for consistent colonization. Compared to previous studies inoculating grapevines with Ilyonectria, I inoculated with half or less Ilyonectria per pot (Vukicevich et al., 2018; Holland et al., 2019; Richards et al., 2020). The lower level of inoculation in my study was due to a miscalculation during spore extraction. A study by Probst et al. (2019) conducted a threshold propagule experiment with I. liriodendri that correlated disease incidence and severity with the concentration of propagules. 3.5.2 Do some species of brassica allow for higher levels of colonization than others? Presence of Ilyonectria in the roots alone is not sufficient evidence for host suitability. If a plant is a good host for Ilyonectria, it would be expected to proliferate within roots. Overall, the abundance of Ilyonectria found in our roots was low relative to previous studies measuring abundance of Ilyonectria in grapevine roots. In the study by Holland et al., (2019), where the 74 primers used were designed, their grapevine controls contained an average amount of Ilyonectria 95 times that of the average found in our grapevine roots. Another study by del Pilar Martínez-Diz et al. (2020) used ddPCR to detect I. liriodendri in environmental grapevine samples. They found, at a minimum, 150 times the amount of Ilyonectria inside the roots compared to our study. However, in another study by Richards et al. (2020), using the same primers as the current study, the grapevine roots contained similar levels of Ilyonectria as our own. As previously mentioned, the amount of Ilyonectria added to plants in our study was less than that of previous studies. An overall lower abundance available for colonization is a likely reason for us to be seeing lower abundance in the roots compared to other studies. That said, the abundance of Ilyonectria in grapevine roots in our study was similar to that of Richards et al. (2020) who added more inoculant but had the same incubation times. This suggests that another possible reason for this discrepancy in Ilyonectria abundance in roots between studies may be due to length of incubation. Holland et al. (2019) had grown their grapevines for six months, while Richards et al. (2020) and the current study’s grapevines were grown for three months. Additionally, with del Pilar Martínez-Diz et al. (2020) grapevines originating from vineyards that were five to seven years old, they would have been exposed to the pathogen for a long time. If our plants were allowed to grow for longer, it is possible that we would start seeing higher levels of colonization within the roots. The low abundance of Ilyonectria inside the roots suggests the Ilyonectria can penetrate roots but are not able to reproduce or are remaining latent. The reasoning for this may be related to the fact that our plants were grown in a relatively stress-free environment. Previous literature has linked disease incidence in plants to stresses imposed on those plants (i.e. poor nutrition, drainage, water stress, etc.) (Halleen et al., 2006). Severity and spread of black foot disease is known to be enhanced by environmental stress (Gramaje & Armengol, 2011; Probst et al., 2012; Úrbez-Torres et al., 2014). Stress-free plants may have been able to devote more resources to their defense mechanisms resulting in overall lower susceptibility to colonization. It is possible 75 that if the plants that were successfully colonized as hosts were transferred to a stressful environment, the number of Ilyonectria in the roots would increase. I did not detect differences in the abundance of Ilyonectria among brassica species, including grapevine. I predicted that grapevines would have the highest abundance of Ilyonectria due to their status as a known host for this isolate. I had predicted all brassicas to be resistant to fungal colonization or suppressive of sporulation due to their anti-fungal properties (Walker et al., 1937; Sarwar et al., 1998; Tiznado-Hernandez & Troncoso-Rojas, 2008) and have been found to reduce disease incidence related to Ilyonectria spp. (Berlanas et al., 2018; Richards et al., 2020). Even if colonization was to occur, I expected a lesser amount in brassica roots compared to grapevine. A possible explanation for the lack of differences between treatments in terms of both colonization and abundance of Ilyonectria is that the Ilyonectria inoculant used had lost some of its ability to penetrate roots and proliferate inside them. Since the isolate was maintained on PDA plates for several months, it may have mutated and lost some of this ability. As the pathogen is being grown outside of a host with plenty of nutrients, there would be no selection pressure to keep genes relating to colonization or pathogenicity, or to remove those who have mutated to reduce these abilities. There is evidence in the literature that some plant pathogens contain genes that when lost or mutated, would alter their pathogenicity, but still allow them to colonize the plant (Idnurm & Howlett, 2001; Hatta et al., 2002; Kanda et al., 2003). Genes specifically relating to pathogenicity have been noted in Ilyonectria spp. (Guan et al., 2020). Additionally, studies have found that when culturing a pathogenic organism for long periods of time outside of their preferred host, they can lose some of their pathogenic potential (Köhsler et al., 2008; Moreira et al., 2012). If the inoculant had mutated in a way for it to reduce its ability to reproduce in roots, then we would not see any differences in abundance among treatments. The ability to colonize the roots of potential cover crops is the first step to determining host potential of the plant, however, it does not tell us whether the presence of that crop would 76 influence the abundance or functionality of the pathogens in the field. If the brassica roots are not only able to be colonized by Ilyonectria but are also susceptible to showing disease symptoms, then it would be a poor choice for suppressing the pathogen. Similarly, if Ilyonectria can reproduce inside the brassica roots, the crop would be acting as a spore bank which would allow the pathogen to persist, and result in more spread within the soil. Further studies on the host capabilities of these plants are needed to confirm whether they would be able to host Ilyonectria in a capacity that would be truly detrimental to grapevines in a vineyard setting. 3.5.3 Do these brassicas act as hosts for Mesocriconema xenoplax (ring nematode)? Grapevines in our study had significantly more nematodes than all brassicas, suggesting that the brassica cover crops used for this experiment are not good hosts for ring nematodes. Whereas numbers of ring nematodes doubled over the course of our experiment in grapevines, they decreased for all four brassica treatments. Our analysis of the ring nematode RF for each of our treatments resulted in the same pattern with the grapevines being significantly higher than all brassica treatments. All brassica species had RF <1, suggesting they are poor hosts for ring nematodes while the grapevines had RF >1, suggesting they are good hosts (Oostenbrink, 1966). These results are expected as previous studies have confirmed the host status of grapevines for ring nematodes. Additionally, the brassica treatments having the lowest abundance of ring nematodes aligns with our prediction. Our results align with a similar greenhouse study by Kruger et al., 2015a looking at the host status of different cover crops, including brassicas, for ring nematodes. They found tomatoes to have significantly higher levels of ring nematodes compared to all other plant species. However, they did not find any significant differences between the brassica treatments and the soil only control, suggesting the brassicas to be poor hosts at best. Among our brassica treatments, rockcress had the highest abundance of ring nematodes while tillage radish had the lowest. It is well known that different species of brassica produce different types of GSLs (Fahey et al., 2001; Rangkadilok et al., 2002; Bellostas et al., 2004). It is 77 possible that tillage radish was able to produce more nematocidal compounds or the compounds produced were more efficient at suppression of ring nematodes. Nematode reactions to ITCs have been reported to be very dependent on the type of ITC added as well as the species of nematode and species of plant involved (Reardon et al., 2013; Mocali et al., 2015). In the case of rockcress, it may be less effective because it co-occurs naturally with the nematodes used in this study. Thus, the nematodes may have developed more resistance to the GSL profile of rockcress and would therefore be less affected by its presence. Analysis of the abundance of GSLs in the soil as well as the types produced would be needed to determine the cause for the discrepancy seen between brassica species. 3.5.4 Do these brassicas act as hosts for AMF Rhizophagus irregularis? None of the brassicas in this study were successfully colonized by AMF. This result aligns with our expectations that only the grapevines would be colonized by AMF. Brassicas are largely considered non-mycorrhizal due to the fungicidal properties of the GSLs they produce (Walker et al., 1937; Vierheilig & Ocampo, 1990; Sarwar et al., 1998). A study by Vierheilig & Ocampo (1990) found ground Brassica oleracea roots applied to AMF spores successfully inhibited germination. Similarly, Schreiner & Koide, 1993 incubated intact roots of Brassica kaber (DC.) Wheeler and Brassica nigra L. with AMF spores and found that living, intact brassica roots were enough to inhibit the germination of AMF spores. A more recent study by Cantor et al., 2011 found soils with higher levels of the brassica Alliaria petiolate (and its associated GSL/ITC) had low levels of fungal hyphae. The same study showed that ITC levels similar to field conditions lowered AMF spore germination by 57%. Despite evidence of brassicas suppressing AMF, there are several examples in the literature of brassicas serving as AM hosts (Vierheilig et al., 2000; Orłowska et al., 2002; Regvar et al., 2003; Vogel-Mikuš et al., 2006; Pongrac et al., 2007). Early studies estimated between 8% (Newman & Reddell, 1987) and 18.9% (Demars & Boerner, 1996) of brassica species are able to 78 host AMF. For these studies, the presence of AMF structures inside the roots was enough to consider it a host. In more recent studies that have found AMF colonization in brassicas, there has been doubt as to whether the associations result in an exchange of nutrients as typically expected from AMF symbiosis (Orłowska et al., 2002; Regvar et al., 2003; Veiga et al., 2012; Cosme et al., 2018). In such cases the association could be beneficial (in ways aside from nutrient exchange) or detrimental to the host plant. If AMF colonization occurs without the presence of the structures used for nutrient exchange (arbuscules and vesicles), there may still be beneficial effects on the host plant. For instance, AMF colonization has been associated with plant protection against pathogens, which is not necessarily correlated with nutrient exchange (Powell et al., 2009; Fernández et al., 2014). There is evidence in the literature that not all AMF associations result in benefits for the host plant (Johnson et al., 1997; Rinaudo et al., 2010; Veiga et al., 2011). A study by Veiga et al., 2013 found brassica species Arabidopsis thaliana to have reduced growth when colonized by AMF. Similarly, a study by Rinaudo et al., 2010 found colonization in the roots of several agricultural non-host weeds to be associated with reduced biomass. If the colonization by AMF would be detrimental in our brassica species, it would be expected for them to have defense mechanisms to resist colonization. It is possible that the conditions of our study were not conducive to AMF colonization in our brassica species. For instance, there is evidence in the literature that suggests time of year or stage of plant development may play a role in susceptibility to colonization (Orłowska et al., 2002; Regvar et al., 2003; Vogel-Mikuš et al., 2006), suggesting if our plants could grow for longer, they may have shown signs of colonization. In particular, the flowering stage tends to increase the likelihood of colonization, likely due to increased nutrient requirements (Orłowska et al., 2002; Vogel-Mikuš et al., 2006). Typically, the main benefit of AMF symbiosis for the host plant is acquisition of additional nutrients (Clark & Zeto, 2000; Cheng & Baumgartner, 2006). Thus, nutrient availability is an important factor in determining AMF root colonization (Carrenho et al., 79 2007; Oliveira & Oliveira, 2010; Khakpour & Khara, 2012). AMF colonization has been found to be reduced in plants grown in highly fertile soils (Schreiner, 2005; Khakpour & Khara, 2012). Although our crops were not fertilized, there is no evidence to suggest they were lacking in any nutrients and therefore would not benefit from AMF symbiosis. Additionally, there is evidence to suggest AMF colonization is more likely to occur in plants that are typically considered non-hosts when they are in the presence of host plants as signals released by AMF host plants into the soil to trigger gene expression in AMF that are absent in non-hosts (Giovannetti & Sbrana, 1998; Delaux et al., 2014; Bravo et al., 2016). In our greenhouse experiment all brassica species were grown without the presence of a mycorrhizal host. The choice of AMF isolate in our study may have affected the ability of our brassicas to act as hosts. Studies have found different species of host plants to react differently depending on species of AMF present (van der Heijden et al., 1998; Avio et al., 2006; Yao et al., 2008; Fernández et al., 2014). The isolate chosen for this experiment, R. irregularis, is a commercial produced strain commonly used in agriculture (Thonar et al., 2014; Egan et al., 2018; Kokkoris et al., 2019). In particular, a study by Vogel-Mikuš et al., (2006) found native AMF species to be more efficient at colonization in the brassica species they used. Thus, we might expect different results with a wider variety of AMF isolates and/or brassica hosts. It is unlikely that our experimental conditions suppressed mycorrhization as grapevine controls were highly colonized by AMF structures (spores, arbuscules, hyphae) (Figure B1). There was evidence of non-AM root colonization in our rockcress (Figure B2). Closely related species Arabis alpina have been found to contain non-mycorrhizal fungal endophytes (Almario et al., 2017; Sharma et al., 2019). Recent studies have looked towards genetics to determine the reasoning behind AMF incompatibility in plants and found many brassicas are missing genes related to AMF symbiosis (Delaux et al., 2014; Favre et al., 2014; Bravo et al., 80 2016). If this is the case, their resistance to AMF colonization may not necessarily translate to other fungal endophytes. 81 Chapter 4: Conclusion I had asked how brassicas influence the soil fungal and nematode communities under field conditions. For application in a vineyard, the desired effect would be disease suppression with minimal influence on the rest of the community, especially beneficials. Overall results from this study suggest that after one season of growth, the brassica cover crops used in this study do have a minor but significant difference on the diversity of the fungal community in a vineyard; however, they do not not have significant influence on the fungal community composition or the beneficial AMF. The minimal changes to the fungal community may also suggest that brassicas are not enough to achieve the desired disease suppression effects after only one season of growth. However, there are factors aside from species composition and diversity that should be measured to determine if the brassica plants have any influence on the community. Similarly, the results suggest brassica plants have no negative effects on FLN populations. Additionally, in this study I was able to determine that our brassica species are not hosts for ring nematode. On the other hand, all brassica treatments were shown to have potential as hosts for Ilyonectria; however, more studies should be done to confirm this. Finally, our brassica species were found not able to be colonized by AMF. Since the results from the field suggest they are not suppressing AMF in the soil, further studies would need to be done to determine if the lack of mycorrhization would have any influence on grapevines in the field. 4.1 Do intact brassica cover crops affect soil fungal diversity and community composition? If so, are there differences in effects among brassica species? Based on the results from this study, brassica cover crops used increase the soil fungal diversity compared to the control through increasing species evenness, but made no changes in species richness. Among brassica species, there is no significant differences in soil fungal diversity. There is no effect on fungal community composition between brassicas and the control 82 or among brassica species. I had expected fungal diversity to decrease in all brassica treatments and for changes in the species composition to occur when compared to the control. The increase in fungal diversity does not align with other studies that looked at fungal community changes in response to brassica cover crops which found no changes (Larkin et al., 2010; Wen et al., 2017; Cloutier et al., 2020). Regardless of diversity changes, compositional changes were still expected based on the results of a similar study by Cloutier et al. 2020. The lack of changes to the fungal community composition could be a result of the GSL secretion and/or the activation level being too low. The most common way to increase GSL activation is to mulch the brassica cover crops into the soil, which is not an option in no-till systems. One possible method to resolve the low levels of GSL activation would be inoculation of the soil with myrosinase producing microbes. If the addition of these microbes can increase GSL activation to a level that results in changes to the soil fungal community, it could result in a self-perpetuating low maintenance system. This is because changes to the soil microbial community as a result of biofumigation tend to result in an increase in microbes that are beneficial to brassicas, which would include myrosinase producers (Mazzola et al., 2015; Hu et al., 2015; Siebers et al., 2018; Ren et al., 2018b). This could be confirmed with a follow-up experiment in which ITC levels in the soil are measured and compared between plots inoculated with myrosinase producing organisms, and those without. I would also be interested to run the soil DNA samples though ddPCR using a primer for the myrosinase producing gene to determine if there is any correlation between ITC levels and microbial myrosinase production. Higher SDI was detected when comparing all brassica treatments to the control. This was found to be due to a change in species evenness with the control plot having lower evenness than all brassica species. Rockcress had the highest evenness. It is uncertain as to why brassicas had higher evenness, but it is possible that the brassica treatments were having some effect on the soil community, but not enough to stimulate compositional changes. If GSL activation was 83 stimulated, we might see a more pronounced difference in our results. As for why rockcress had higher evenness than the other brassica species, the GSL profile might be reacting differently with the soil community. It would be beneficial to compare the evenness with the different GSL profiles produced by each treatment. Finally, lack of changes to the fungal community does not necessarily mean there were no functional changes. Brassicas have been found to increase microbial activity in the soil even in the absence of microbial community changes (Cohen et al., 2005; Larkin et al., 2010; Potgieter et al., 2013; Galletti et al., 2015). Additionally, growing brassica cover crops has been found to reduce disease incidence in neighbouring plants (Larkin et al., 2010; Larkin & Halloran, 2014; Hossain et al., 2015; Wen et al., 2017; Baysal-Gurel et al., 2020), a result that has been found to be independent of diversity changes (Larkin et al., 2010; Wen et al., 2017). Therefore, it would be beneficial to determine if our treatments had any functional effects on the soil microbial community. A follow-up study looking at disease incidence or changes to the fungal community inside the roots of the grapevines would be beneficial in determining if these brassica cover crops play any role in suppression of disease regardless of soil community diversity. Running transcriptomics on the soil samples from the same experiment would help us determine if there were any changes in the gene expression of the organisms already present. The functional changes because of treatment with brassica cover crops could be the more important factor for determining its benefits. 4.2 How do brassicas affect AMF in a field? There were no differences in AMF abundance when looking at either the spore count or the molecular data. I had expected the brassica treatments to have lower abundance of AMF when compared to the control due to evidence that intact brassica plants have been able to successfully suppress AMF germination in the soil (Schreiner & Koide, 1993b; Cantor et al., 2011). The reason for the lack of changes to AMF abundance may be related to the presence of 84 nearby mycorrhizal plants in the field. Studies reporting a reduction in AMF abundance as a result of biofumigation are typically greenhouse studies or field studies with no indication of nearby mycorrhizal plants (Schreiner & Koide, 1993b; Siebers et al., 2018; Cloutier et al., 2020) (Cantor et al. 2011 being the exception; however, this study was not done in an agricultural setting). Other field studies looking at the effect of incorporation of brassicas have found no evidence of a negative effect on cash crop root colonization or AMF abundance (Pellerin et al., 2007; Tong et al., 2014). If brassica cover crops are not suppressing the AMF population in the soil, this is good news for farmers. To determine if the lack of AMF suppression is a result of growth with a mycorrhizal plant (grapevines), a greenhouse study could be conducted. We could compare the AMF populations in the soil between pots grown with brassicas alone and pots grown with both brassicas and grapevines. At the same time, we could determine whether the presence of the brassica is influencing AMF root colonization of the neighbouring grapevine. 4.3 Do intact brassica cover crops affect soil nematode abundance and community composition? If so, are there differences in effects among brassica species? I had found no differences in nematode abundance between treatments. Similar to the lack of changes to the fungal community, this is thought to be due to low levels of GSL/ITC from the intact brassica plants. The results were as expected as there is evidence in the literature to suggest GSL/ITC levels need to be much higher under field conditions to result in a suppression effect on nematodes (Stirling & Stirling, 2003; Zasada & Ferris, 2003). As previously mentioned, it would be beneficial in future studies to measure the levels of GSLs and ITCs in the soil to determine if there is a correlation with nematode abundance. Although there were no significant differences in overall abundance among cover crop treatments, total nematode abundance was positively correlated with brassica biomass. The same positive correlation was found in FLN abundance, but not in PPN abundance. This correlation between total nematode abundance and brassica biomass could be related to an 85 increase in biomass resulting in an increase in nutrient availability for the soil community. However, multiple other studies using non-brassica plants do not see the same correlation (De Deyn et al., 2004; Viketoft, 2008; Cortois et al., 2017), which may suggest what I saw in my study may be a result of brassicas being used. Shepherd’s purse and white mustard treatments tended to have lower population densities of both PPN and FLN than controls. Those cover crops also had significantly lower ratios of PPN to FLN when compared to the control. This result suggests that presence of shepherd’s purse and white mustard may have suppressed PPN more than FLN. A follow-up analysis could be done to determine which species of PPN or FLN are being affected by these brassicas to cause the shift in relative abundance. Ultimately, the result of a decrease in relative abundance of PPN is a beneficial result for vineyards. Rather than suppressing all nematode populations, it is ideal to suppress the parasites while keeping FLN populations. That said, the FLN population contains a wide range of nematode species and this data alone cannot tell us how that population is changing. To get a better idea of how the FLN population is changing, it would be beneficial to count community members by trophic group. 4.4 Do these brassicas act as hosts for Ilyonectria liriodendri? All brassica species as well as the control were found to contain Ilyonectria in the roots. However, the colonization of Ilyonectria was not ubiquitous in any of the treatments and there were no differences between treatments. The lack of consistency among replicates may be attributed to insufficient inoculation as I applied less inoculant than previous studies. We could determine if insufficient inoculant is the reason for the inconsistent colonization as well as the low levels of colonization by repeating the experiment with increased volume of inoculum. Although inconsistent, all treatments contained some replicates with Ilyonectria in the roots. This suggests all species are susceptible to colonization of Ilyonectria and may act as hosts. Abundance in the roots were relatively low; however, this may be attributed to the relatively short incubation time as well as the stress-free conditions of the plants. If the plants were allowed to 86 grow for longer or were subject to stress, it is possible the levels of Ilyonectria in the roots would increase. To determine if this is the case, a similar experiment could be performed during a longer period of time where half the replicates would be subject to stress. Potential cover crops being hosts for Ilyonectria has two potential detriments. For one, they can become infected and display disease symptoms resulting in losses of the cover crop and therefore a reduction in whatever benefits they might bring. The second is that they can become infected and participate in spreading the disease as well as becoming spore banks which keep the pathogen in the soil. Thus, a follow-up study should be done to determine if our brassica species are susceptible to expression of disease symptoms after entering the roots, as well as if the effect they have on Ilyonectria abundance in the surrounding soil. 4.5 Do these brassicas act as hosts for Mesocriconema xenoplax (ring nematode)? I found that all our brassica species were resistant to ring nematodes. As all the brassica species had RF < 1, they may also be active in suppressing the ring nematode population. To determine if this is the case, another study could be done using a soil only negative control. If the abundance of ring nematodes in the brassica treatments end up being significantly less than the abundance in the soil only control, I can conclude that they are active in suppressing ring nematodes. At the very least, our results suggest that any of these brassica species would be a better cover crop choice than plants that host ring nematodes. Although the effects in this greenhouse experiment were clear, there is no guarantee that the same results would be seen under field conditions. The level of GSLs/ITCs needed under field conditions to result in suppression has been found to be much higher than what is needed for the same results under greenhouse conditions (Stirling & Stirling, 2003; Zasada & Ferris, 2003). An experiment using these brassica species as cover crops in a ring nematode infested vineyard would help us to determine if intact brassicas are enough to result in suppression under field conditions. 87 4.6 Do these brassicas act as hosts for AMF Rhizophagus irregularis? None of the brassicas were found to contain AMF in their roots. Conditions of the greenhouse experiment were stress-free and so the brassicas would have not needed the benefits from a symbiosis with AMF which may have contributed to the lack of colonization seen. However, I believe our brassicas are simply non-mycorrhizal and do not have the genes necessary for AMF colonization and symbiosis. This is because less than 20% of brassica plants have been found to host AMF (Newman & Reddell, 1987; Demars & Boerner, 1996). That said, it would be beneficial to see if the presence of a mycorrhizal plant nearby would cause a different result. There is evidence in the literature to suggest the presence of a mycorrhizal plant increases the odds of AMF colonization in non-hosts (Giovannetti & Sbrana, 1998; Delaux et al., 2014; Bravo et al., 2016). It would be beneficial to conduct another study with these brassica species grown in the presence of grapevines in AMF inoculated soil to see if their ability to host AMF improves. At the same time, we could see how the presence of brassicas might influence AMF colonization of neighbouring grapevines. Whether the brassicas can act as hosts or not, an ideal grapevine cover crop would not interfere with the symbiosis between AMF and the grapevines. 4.7 Strengths and limitations With the growing demand for sustainable agriculture practices, this study gives insight into a potential low cost, organic method of disease suppression. There are few studies that consider the effects of intact brassicas on the soil fungal community and only one that uses high-throughput sequencing techniques to do so. This study is the first to use high resolution techniques to look at changes that occur in both the fungal and nematode communities as a result of the addition of intact brassica cover crops under field conditions in a vineyard. Many studies tend to focus on the effects of brassica treatments on specific organisms of interest (usually pathogens). When selecting cover crops, it is important to consider their effect on the soil community as a whole, but also how they might interact with organisms of interest. I was able to do both in this study by 88 looking at how these brassicas might influence the soil microbial community under field conditions, as well as their host status for organisms of interest. In particular, the rockcress and shepherd’s purse used have not been studied to this extent in any other study to date. Although this study considered changes to the soil microbial community from multiple angles in regards to fungi and nematodes, I did not include analysis of the bacterial community or functional changes. When determining beneficial or detrimental effects on the soil, the bacterial community should also be considered. Similarly, not all significant changes to the microbial community occur through shifts in community structure. Functional changes may be important in understanding the effects of brassica cover crops on soil health. To better understand functional changes related to the nematode community, it would have been beneficial to conduct a community analysis at the trophic level to see which groups are changing among FLN (bacteriovores, fungivores, predators, etc.). Another limitation of this study is that I did not measure GSL or ITC production from my brassicas under field conditions. Brassica application is typically done in the form of green manure or seed meal to maximize GSL activation. In order to determine if there is any correlation between changes detected in the soil microbial community and amount of GSLs/ITCs produced by the intact brassicas, GSL/ITC levels in the soil should be measured. 4.8 Concluding remarks In conclusion, my study found brassica cover crops to increase species evenness which is inconsistent with current literature. I found no significant difference in effect on diversity metrics among different brassica species. There was no effect on the soil fungal community composition, soil nematode abundance, or AMF spore abundance between brassicas and the control or among brassica species. From the results of this study, I can suggest there would be no detriments to using these brassica species as cover crops in a vineyard. Although the brassica species used are non-mycorrhizal, they do not appear to suppress the AMF population. Additionally, they are 89 not able to host ring nematodes. Shepherd’s purse and white mustard in particular show potential for controlling the PPN population. Regarding other benefits expected from using plants with biofumigant properties, more research should be done on the functional changes in the soil rather than only species diversity and composition. The only possible negative result is that all brassica species used appear capable of being colonized by Ilyonectria. However, more research needs to be done to understand this interaction and if it would be detrimental in a vineyard. The results from this study provides evidence that brassica cover crops are a safe cover crop choice for vineyards. Although the beneficial effects may be smaller than desired, there is potential for suppression of soil-borne pathogens using brassica cover crops without the need for incorporation. 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Journal of Nematology 41: 221–227. 106 Zehr EI, Aitken JB, Scott JM, Meyer JR. 1990. Additional Hosts for the ring nematode, Criconemella xenoplax. Journal of nematology 22: 86–89. Zeileis A, Kleiber C, Jackman S. 2008. Regression models for count data in R. Journal of Statistical Software 27: 1–25. 107 Appendices Appendix A – Supporting material for chapter 2 A.1 Figures and tables Figure A1. Principal coordinates analysis of fungal communities from soil of plots treated with growth of different brassica cover crop species. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Bray curtis dissimilarity. Axis 2 (11.01 %) Axis 1 (13.67 %) Control Rockcress Shepherd’s purse Tillage radish White mustard 108 Axis 2 (17.67 %) Axis 1 (27.64 %) Control Rockcress Shepherd’s purse Tillage radish White mustard Figure A2. Principal coordinates analysis of fungal communities from soil of plots treated with growth of different brassica cover crop species. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Weighted unifrac. 109 Figure A3. Principal coordinate analysis of fungal communities from soil of plots treated with growth of different brassica cover crop species. Treatments: C, undisturbed control; RC, Rockcress; SP, Shepherd’s purse; TR, Tillage radish; WM, White mustard (n = 3 for control, n = 5 for all other treatments). Unweighted unifrac. Axis 2 (7.794 %) Axis 1 (11.52 %) Control Rockcress Shepherd’s purse Tillage radish White mustard 110 Table A1. Number of reads retained at each step in the DADA2 pipeline. From left to right are initial read counts (input), reads after filtering (filtered), reads after denoising (denoised), and number of reads after chimera removal (non-chimeric). Sample Treatment species input filtered denoised non-chimeric 1 TR 4103 3306 3150 3150 2 SP 7321 6221 5968 5968 3 C 7777 6320 6129 6129 4 RC 7587 6210 6007 6007 5 WM 6891 5506 5305 5305 7 TR 3526 2987 2842 2842 8 SP 6698 5780 5637 5604 9 WM 6859 5810 5627 5627 10 RC 4560 3797 3612 3612 11 RC 3479 2929 2769 2769 12 SP 11983 10179 9961 9941 13 TR 4189 3602 3421 3421 14 WM 2983 2547 2365 2365 16 TR 13308 11633 11418 11418 17 RC 5061 4406 4258 4258 18 WM 7911 6853 6699 6699 19 C 8592 7266 7071 7071 20 SP 8353 6855 6666 6666 21 C 4537 3855 3678 3678 22 RC 4531 3967 3778 3778 23 TR 3136 2763 2602 2602 24 WM 12443 10980 10695 10695 25 SP 8937 7766 7538 7538 111 Table A2. Pairwise PERMANOVA results of fungal community beta diversity using Bray-Curtis distance. Permutations = 999. Group 1 Group 2 Sample size pseudo-F p-value q-value C RC 8 1.412 0.053 0.265 C SP 8 1.418 0.032 0.265 C TR 8 0.946 0.555 0.793 C WM 8 1.085 0.311 0.654 RC SP 10 0.962 0.520 0.793 RC TR 10 0.812 0.894 0.894 RC WM 10 0.857 0.677 0.846 SP TR 10 1.106 0.227 0.654 SP WM 10 1.059 0.327 0.654 TR WM 10 0.820 0.818 0.894 Table A3. Pairwise PERMANOVA results of fungal community beta diversity using Unweighted UniFrac. Permutations = 999. Group 1 Group 2 Sample size pseudo-F p-value q-value C RC 8 1.248 0.124 0.638 C SP 8 1.231 0.166 0.638 C TR 8 0.913 0.628 0.698 C WM 8 1.141 0.240 0.638 RC SP 10 1.019 0.421 0.674 RC TR 10 0.982 0.539 0.674 RC WM 10 1.071 0.255 0.638 SP TR 10 0.802 0.938 0.938 SP WM 10 1.014 0.399 0.674 TR WM 10 0.964 0.538 0.674 112 Table A4. Pairwise PERMANOVA results of fungal community beta diversity using Weighted UniFrac. Permutations = 999. Group 1 Group 2 Sample size pseudo-F p-value q-value C RC 8 1.772 0.107 0.475 C SP 8 2.105 0.078 0.475 C TR 8 1.566 0.188 0.475 C WM 8 1.515 0.190 0.475 RC SP 10 1.124 0.272 0.544 RC TR 10 0.781 0.664 0.787 RC WM 10 1.043 0.365 0.608 SP TR 10 0.870 0.517 0.739 SP WM 10 0.732 0.708 0.787 TR WM 10 0.456 0.868 0.868 Table A5. Correlation analysis of all measured continuous variables with different alpha diversity metrics. Pielou's evenness Index, Shannon Diversity Index (SDI), species richness (observed OTUs), and Faith's Phylogenetic Diversity (FPD). Variables include soil pH, AMF spore count (per 20 g soil), nematode abundance (per 100 ml soil), and estimated biomass of treatment plant per plot (excluding control). Spearman correlation calculated in QIIME2. (n = 23). Variable Diversity metric Test statistic p-value pH Evenness 0.378 0.08 pH SDI 0.073 0.74 pH Richness -0.221 0.31 pH FPD -0.365 0.09 Spore count Evenness -0.063 0.78 Spore count SDI 0.055 0.80 Spore count Richness 0.115 0.60 Spore count FPD 0.233 0.29 Nematodes Evenness 0.007 0.98 Nematodes SDI 0.305 0.16 Nematodes Richness 0.225 0.30 Nematodes FPD 0.210 0.34 Biomass Evenness -0.050 0.83 Biomass SDI -0.141 0.55 Biomass Richness -0.229 0.33 Biomass FPD -0.253 0.28 113 A.2 R code A.2.1 Fungal community analyses Shannon diversity: shannon.lm <- lm(shannon ~ species, data = field2a) shannon.lm.anova <- anova(shannon.lm) shannon.lm.anova shannon.lm.summary <- summary(shannon.lm) shannon.lm.summary$r.squared Faith’s phylogenetic diversity: faith.lm <- lm(faith_pd ~ species, data = field2a) faith.lm.anova <- anova(faith.lm) faith.lm.anova faith.lm.summary <- summary(faith.lm) faith.lm.summary$r.squared Pielou’s evenness index: even.lm <- lm(pielou_e ~ species, data = field2a) even.lm.anova <- anova(even.lm) even.lm.anova even.lm.summary <- summary(even.lm) even.lm.summary$r.squared Observed OUT’s (species richness): otus.lm <- lm(observed_otus ~ species, data = field2a) 114 otus.lm.anova <- anova(otus.lm) otus.lm.anova otus.lm.summary <- summary(otus.lm) otus.lm.summary$r.squared Planned comparisons (example with evenness): f.tan$species.f <- as.factor(f.tan$species) #convert species to factors (5 levels) mat.c.b <- c(1, -1, -1, -1, -1) # control vs. brassica treatments contrasts(f.tan$species.f) <- mat.c.b model.even <- aov(pielou_e ~ species.f, data = f.tan) summary(model.even) summary.lm(model.even) #get r squared value summary.aov(model.even, split = list(species.f=list("Control vs. Brassicas"=1))) PERMANOVA for the phylum level a3.dist <- vegdist(a3, method = "bray") adonis2(a3 ~ species, data = level2, permutations = 999, method="bray", by = NULL) PERMANOVA for the class level b3.dist <- vegdist(b3, method = "bray") adonis2(b3 ~ species, data = level3, permutations = 999, method="bray", by = NULL) 115 PERMANOVA for the species level c3.dist <- vegdist(c3, method = "bray") adonis2(c3 ~ species, data = level7, permutations = 999, method="bray", by = NULL) A.2.2 Nematode analyses Total nematodes: totalnema.lm <- lm(total_nema ~ species, data = field2) totalnema.lm.anova <- anova(totalnema.lm) totalnema.lm.anova totalnema.lm.summary <- summary(totalnema.lm) totalnema.lm.summary$r.squared FLN: fln.lm <- lm(fln ~ species, data = field2a) fln.lm.anova <- anova(fln.lm) fln.lm.anova fln.lm.summary <- summary(fln.lm) fln.lm.summary$r.squared PPN: ppn.lm <- lm(logppn ~ species, data = field2a) ppn.lm.anova <- anova(ppn.lm) ppn.lm.anova 116 ppn.lm.summary <- summary(ppn.lm) ppn.lm.summary$r.squared Mixed effect logistic regression: m1 <- glmer(cbind(ppn, fl_nema) ~ species + (1 | plot), data = nema.field, family = binomial("logit")) summary(m1) 117 Appendix B – Supporting material for chapter 3 B.1 Plant growth In the greenhouse all but one pot had seedlings successfully established or seeds planted successfully germinated. The one pot that did not have a plant successfully establish was a rockcress with AMF treatment. The grapevines were inflicted with powdery mildew at the time of transplanting. The greenhouse was treated with sulphur once a week for a month until the powdery mildew was no longer visible, then once every two weeks until harvest. The sulphur application method used was vaporized sulphur using a Nivola sulphur evaporator and Greenhouse Essentials brand sulphur pellets (99.5%). The vaporizer was run for approximately 24 hrs. per use. At the time of harvest, all plants were at maturity and most shepherds purse and white mustard were flowering. 118 B.2 Figures and tables Figure B1. Microscopy images of stained grapevine roots containing AMF structures. 119 Figure B2. Microscopy images of stained rockcress roots containing spores of an unknown fungus. 120 B.3 R code B.3.1 Ilyonectria analyses Zero-inflated binomial regression: #convert cpgr into integers greenhouse_ilyo$i.cpgr <- as.integer(greenhouse_ilyo$cpgr) #convert species into factors greenhouse_ilyo$fspecies <- as.factor(greenhouse_ilyo$species) zm1 <- zeroinfl(formula = i.cpgr ~ fspecies, data = greenhouse_ilyo, dist = "negbin") summary(zm1) ANOVA for pots positive for Ilyonectria including box cox transformation: #removed the zeros onlypos <- dplyr::filter(greenhouse_ilyo, cpgr > 0) #fit an additive multiple regression model with ddpcr +1 as the response and species as the predictor pos_model = lm(cpgr ~ species, data = onlypos) pbc <- boxcox(pos_model, plotit = TRUE) #lambda is -0.35, transformation onlypos$bc.cpgr <- (((onlypos$cpgr) ^ -0.35) - 1) / -0.35 pos_cox = lm(((((cpgr) ^ -0.35) - 1) / -0.35) ~ species, data = onlypos) anova(pos_cox) 121 B.3.2 Ring nematode analysis variance.check <- leveneTest(ringnema ~ species, data = greenhouse_ring) variance.check leveneTest(logringnema ~ species, data = greenhouse_ring) #log transformed passes equal variance. #anova for transformed ring nema, data is normal and has equal variance rn.lm <- lm(logringnema ~ species, data = greenhouse_ring) rn.lm.anova <- anova(rn.lm) rn.lm.anova rn.lm.summary <- summary(rn.lm) rn.lm.summary$r.squared #post hoc rn.aov <- aov(logringnema ~ species, data = greenhouse_ring) TukeyHSD(rn.aov, conf.level = 0.95)
Thesis/Dissertation
eng
Biology
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Hart, Miranda
Do brassica cover crops influence soil fungal and nematode communities in a vineyard?
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