Isolation of Rhizobacteria in Southwestern Québec, Canada: An

Isolation of rhizobacteria in Southwestern Québec, Canada: An investigation of their impact on the growth and salinity stress alleviation in Arabidopsis thaliana and crop plants

Di Fan

Department of Plant Science

Faculty of Agricultural and Environmental Sciences

Macdonald Campus of McGill University

21111 Lakeshore Road, Sainte-Anne-de-Bellevue, Québec H9X 3V9

December 2017

A thesis submitted to McGill University in partial fulfillment of the requirements of the

degree of DOCTOR OF PHILOSOPHY

Abstract ...... x

Résumé ...... xiii

Acknowledegments ...... xv

Preface ...... xviii

Contribution of authors ...... xviii

Chapter 1...... 1

Introduction ...... 1

Chapter 2...... 5

Literature Review ...... 5

2.1 What are root exudates? ...... 5

2.2 Roles of root exudates ...... 5

2.2.1 Manipulating the rhizosphere environment ...... 6

2.2.2 Plant-to-microbe signals ...... 7

2.2.2.1 Flavonoids ...... 7

2.2.2.2 Jasmonates ...... 8

2.2.2.3 Other nod gene inducers ...... 9

2.2.2.4 Strigolactones...... 9

2.2.3 Quorum sensing ...... 10

2.3 Plant growth promotion ...... 12

2.3.1 Root colonization ...... 12

2.3.2 Biofertilization ...... 13 ii

2.3.2.1 Nitrogen fixation ...... 13

2.3.2.2 Phosphate solubilization ...... 14

2.3.2.3 Iron availability ...... 15

2.4 Phytoremediation ...... 16

2.5 Phytostimulation ...... 17

2.5.1 Indole-3-acetic acid (IAA) ...... 17

2.5.2 Gibberellins ...... 18

2.5.3 Cytokinins ...... 18

2.5.4 Regulation of Ethylene ...... 19

2.5.5 Other plant growth regulators ...... 19

2.6 Biocontrol ...... 20

2.6.1 Antibiotics ...... 21

2.6.2 Bacteriocins ...... 21

2.6.3 Extracellular lytic enzymes ...... 22

2.6.4 Siderophores ...... 22

2.6.5 Phytoalexins ...... 23

2.6.6 Control of other biotic stresses ...... 23

2.6.7 Signal interference...... 24

2.6.8 Induced systemic resistance...... 24

2.7 Novel type of signaling ...... 26

2.7.1 Nod factors ...... 26

2.7.2 Bacteriocins ...... 27

2.8 Old-timer versus rising star ...... 29

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2.9 PGPR screening ...... 30

Connecting text ...... 33

Chapter 3 ...... 34 Isolation and diversity of culturable rhizobacteria associated with economically important crops and uncultivated plants in Québec, Canada ...... 34

3.1 Abstract ...... 34

3.2 Introduction ...... 35

3.3 Materials and Methods ...... 39

3.3.1 Study site and sample collection ...... 39

3.3.2 Isolation and preservation of cultivable rhizobacteria ...... 42

3.3.2.1 Isolation of ectophytic bacteria ...... 42

3.3.2.2 Isolation of cultivable endophytes ...... 43

3.3.2.3 Isolation of endophytic bacteria from soybean root nodules ...... 44

3.3.3 Taxonomic identification and diversity indices ...... 45

3.3.3.4 Statistical analysis ...... 46

3.4 Results ...... 46

3.4.1 Enumeration, isolation, and identification of rhizobacteria ...... 46

3.4.2 Diversity of culturable ectophytic bacterial ...... 51

3.4.3 Culturable endophytic bacterial community survey...... 55

3.5 Discussion...... 64

3.6 Conclusions ...... 69

Connecting text ...... 123

Chapter 4 ...... 124 Inoculation of Arabidopsis thaliana by novel rhizobacterial isolates improves plant growth under normal and stress conditions ...... 124

4.1 Abstract ...... 124

4.2 Introduction ...... 125

4.3 Materials and Methods ...... 129

4.3.1 Plant material, bacterial strains, and growth conditions ...... 129

4.3.2 Rapid bacterial screening assay ...... 131

4.3.3 Secondary bacterial screening...... 131

4.3.4 Petri plate assay for effects of bacterial inoculation on seedling growth ..... 132

4.3.4.1 Germination activities of Arabidopsis seeds after bacterization ...... 132

4.3.4.2 In Vitro osmotic stress ...... 132

4.3.4.3 In Vitro salt stress ...... 133

4.3.4.4 Root architecture analysis ...... 133

4.3.4.5 Quantitative measurement of gene expression via qRT-PCR analysis ...... 133

4.3.5 Split-plate Petri-dish study ...... 137

4.3.6 In planta bacterial impact on growth of A. thaliana in soil ...... 137

4.3.6.1 In Vivo assessment of plant-growth promotion ...... 137

4.3.6.2 Salinity tolerance assay ...... 137

4.3.6.3 Measurement of ROS scavenging activity ...... 138

4.3.6.4 Metabolite measurements ...... 139

4.3.6.5 Proline estimation ...... 140

4.3.7 Screening for bacterial PGP traits ...... 140

4.3.7.1 Ammonia production ...... 140

4.3.7.2 ACC deaminase activity measurement ...... 141

4.3.7.3 Indole-3-acetic acid (IAA) detection ...... 142

4.3.7.4 Nitrogen fixing ability ...... 143

4.3.7.5 Solubilization of insoluble phosphate ...... 145

4.3.7.6 Siderophore production ...... 146

4.3.7.7 Hydrogen cyanide production ...... 146

4.3.7.8 Evaluation of antimicrobial activities ...... 147

4.3.7.9 Exopolysaccharide production...... 148

4.3.7.10 Blood hemolytic assay ...... 148

4.3.8 Identification and characterization of bacterial isolates ...... 148

4.3.8.1 Morphological characterization ...... 148

4.3.8.2 Intrinsic antibiotic sensitivity/resistance ...... 149

4.3.8.3 In Vitro determination of bacterial abiotic stress tolerance ...... 149

4.3.8.4 Extracellular enzyme production ...... 150

4.3.8.5 Carbon source utilization ...... 151

4.3.8.6 Evaluation of other biochemical activities ...... 151

4.3.8.7 Phylogenetic analysis of 16S rDNA sequencing data for selected bacteria ...... 152

4.3.8.8 Growth rate ...... 153

4.3.9 Protection against phytopathogens by rhizobacteria in Arabidopsis ...... 153

4.3.9.1 Bacterial pathogenesis assay ...... 154

4.3.9.2 Protection against phytopathogenic fungi ...... 154

4.3.10 GFP-tagging for in Planta tracking ...... 155

4.3.10.1 Preparation of competent cells ...... 155

4.3.10.2 GFP plasmid transformation ...... 156

4.3.10.3 Microscope analysis of root colonization ...... 156

4.3.10.4 Bacterial re-isolation in Arabidopsis ...... 157

4.3.11 Statistical analysis ...... 157

4.4 Results ...... 157

4.4.1 Bacterial screening for growth promotion of Arabidopsis plants ...... 157

4.4.2 Phenotyping of the strains and their biocontrol and PGP traits ...... 162

4.4.2.1 Molecular taxonomy of candidate bacterial strains by 16S rDNA sequencing .... 162

4.4.2.2 Plant-growth-promotion traits of selected bacteria ...... 166

4.4.2.3 Primary characterization of selected bacteria ...... 173

4.4.2.4 Growth curves and growth kinetics...... 178

4.4.2.5 Minimal inhibitory concentration (MIC) test ...... 179

4.4.3 Spatial distribution of rhizobacteria in Arabidopsis...... 180

4.4.4 In vitro response of treated Arabidopsis to salt and osmotic stress ...... 184

4.4.5 Possible effect of volatile chemicals produced rhizobacteria ...... 188

4.4.6 Bacteria confer facilitated growth and increased salt resistance in planta .. 188

4.4.7 Impact of rhizobacteria on disease severity in Arabidospsis ...... 194

4.4.8 Effect of rhizobacteria on transcript levels of Arabidopsis ...... 198

4.5 Discussion ...... 200

4.6 Conclusions ...... 214

Connecting text ...... 217

Chapter 5 ...... 218 Effect of inoculation with bacterial endophytes on the growth of canola (Brassica napus) and maize (Zea mays) ...... 218

5.1 Abstract ...... 218

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5.2 Introduction ...... 219

5.3 Materials and Methods ...... 222

5.3.1 Bacterial strains and plant materials ...... 222

5.3.2 Effect of sodium chloride on bacterial growth ...... 223

5.3.3 Canola germination ...... 223

5.3.4 Maize germination ...... 223

5.3.5 In Vitro inoculation of maize ...... 224

5.3.6 Evaluation of root colonization ...... 224

5.3.7 Impact of bacterial inoculation on greenhouse growth of maize plants ...... 225

5.3.7.1 Evaluation of isolated bacteria effects on maize plant growth ...... 225

5.3.7.2 DPPH Radical Scavenging Assay ...... 226

5.3.7.3 Determination of total phenolic compound content ...... 226

5.3.7.4 Estimation of proline ...... 227

5.3.7.5 Estimation of carbohydrate ...... 227

5.3.7.6 Determination of nutrient uptake ...... 228

5.3.8 Impact of bacterial inoculation on chamber growth of canola ...... 228

5.3.9 Data analysis ...... 229

5.4 Results ...... 229

5.4.1 Effect of NaCl on bacterial growth ...... 229

5.4.2 Canola inoculation experiment ...... 230

5.4.2.1 Effect of seed inoculation on germination ...... 230

5.4.2.2 Pot experiment ...... 231

5.4.3 Maize inoculation experiment ...... 234

5.4.3.1 Effect of PGPR on in vitro germination and growth ...... 234

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5.4.3.2 Effect of PGPR on maize growth under pot (controlled environment) conditions 238

5.4.3.3 Total carbohydrate ...... 239

5.4.3.4 Antioxidant activities and proline content in leaves ...... 239

5.4.4 Endophytic colonisation ...... 244

5.5 Discussion ...... 244

5.6 Conclusions ...... 250

Chapter 6...... 252

Concluding remarks ...... 252

6.1 General discussion ...... 254

6.2 Future directions ...... 261

References ...... 264

Abstract

Rhizobacteria are important participants in biogeochemical cycles and affect many aspects of plant physiology. Wild plants, especially those grown on impoverished land, have been shown to host a great diversity of bacterial species, contributing to the fitness of their hosts. Since there is no holonomic analysis of the diversity of the established rhizobacteria for crops and wild plants of Southwestern Québec, Canada, the first goal of this project was to determine the culturable rhizobacterial communities associated with a selected set of crops and wild plants in the region of Sainte-Anne-de-Bellevue, Québec. A total of 446 rhizobacterial strains were isolated, which were classified into 90 bacterial genera, with Proteobacteria, Actinobacteria, Firmicutes and Bacteroidetes as the dominant phyla across all samples. The culturable bacterial community composition and diversity were strongly affected by plant species. In order to develop potential bioproducts for sustainable agriculture, two sets of experiments were conducted. First, the potential plant growth-promoting (PGP) rhizobacteria from the reservoir pool indicated above were screened for growth promotion effects on Arabidopsis thaliana ecotype Columbia (Col-0). Four out of 98 selected rhizobacterial strains (strains n, L, K and Y), from Phalaris arundinacea, Solanum dulcamara, Scorzoneroides autumnalis, and Glycine max, respectively, showed significant growth stimulation effects with regard to total leaf area, total rosette fresh and dry weight, root length, total antioxidant capacity, salinity stress attenuation ability, and total contents of proline and chlorophyll. The 16S rRNA gene sequencing and phylogenetic diversity of the strains were analyzed and indicated that these isolates belong to the genera Pseudomonas, Bacillus, Mucilaginibacter and Rhizobium, respectively. All the strains were characterized using classical methods of bacterial identification and using biochemical test kits (API20E, API20NE, API ZYM, and API 50CH), revealing the metabolic versatility of the four strains. Strains n and L also showed antagonistic effects against two phytopathogens, Pseudomonas syringae pv. tomato (Pst) DC3000 EV (virulent) and Botrytis cinerea wild-type isolate B191, both in vitro and in planta. Bacterial treatment caused an increase in the transcription of the genes related to plant growth, osmolyte accumulation, and defense response. A study on spatial distribution of the four strains, using either conventional Petri-plate counts or x

GFP-tagged bacteria, indicated that all the strains were able to colonize the endosphere of Arabidopsis tissue. The four selected strains were also characterized in pure culture for currently recognized PGP traits, to evaluate the involvement of the traits in PGP performance. All rhizobacterial isolates were positive for ACCD and IAA production and phosphorous solubilization. The amounts of soluble phosphate generated in liquid medium were 341, 69, 63 and 38 g L-1, for n, L, Y and K, respectively. PCR analysis confirmed the presence of the nifH gene in strains n, L and Y showing their N2-fixation potential. Surprisingly, one of the best plant growth promoters, strain K, did not display the largest number of currently recognized PGP abilities in vitro, but outperformed the strain possessing the most PGP traits, strain n, in planta. Secondly, the four selected strains were further tested on two crop plants, Brassica napus (canola) and Zea mays (maize), for their growth promotion and salt stress alleviation effects. Bacterial strain n improved Fe and P uptake into maize leaves, while the nitrogen content was increased by all four strains, with increases of up to 93 %. Strain L was the only endophyte to cause highly significant increases in N/C ratio. Thus, strains L, K and Y caused increases in the growth of canola and maize, likely through the integration of various mechanisms that improve plant performance, indicating the need for a more in-depth understanding of plant-bacteria interactions. In germination and early seedling growth experiments, the four strains, especially strains L, K and Y, increased the root length and seedling dry weight in canola under both unstressed and salt stressed conditions; while all four strains significantly improved the shoot length (up to 33 %) and fresh weight (up to 25 %) of maize under unstressed conditions. Strains L, K and Y increased the seed vigor index of maize up to 20 % over the control. In pot experiments, canola plant biomass and chlorophyll content were evaluated after 5-weeks of growth; all four strains significantly increased above-ground dry weight, by 14 to 21 %. A greenhouse growth experiment, under non-sterile conditions, evaluated effects of the four strains on maize plant biomass, nutrient uptake, chlorophyll content, total antioxidant capacity, total carbohydrates, accumulation of proline and total phenolics accumulation. Plant growth was promoted by all strains, especially L, K and Y, for root length, aerial biomass, and root dry weight. Only strains n and L increased total antioxidant and proline contents, while strains L, K and Y increased total plant carbohydrate. Nutrient uptake was also evaluated, with higher levels of Fe and P uptake in maize leaves of plants inoculated with strain n, and increased nitrogen content due to treatment with each of the four strains.

Overall, this research (1) highlighted for the first time, a rich reservoir of the diverse rhizobacteria from various crops and uncultivated plants in the region of Sainte-Anne-de-Bellevue, Québec, Canada; these are potential sources for the discovery of PGPR, (2) demonstrated that four isolates out of 98 from the reservoir significantly increased plant growth and salt stress resistance in A. thaliana, both in vitro and in planta and (3) determined that the same four strains also enhanced the growth of two tested crop plants, canola and maize, under both sterile and non-sterile conditions. This is the first study regarding the PGP effects of Bacillus sp. from Solanum dulcamara (bittersweet). To our knowledge, there has been no perviouse detailed study of PGP effects by Mucilaginibacter (stain K) in Arabidopsis. Our direct screening approach revealed this strain to be a fitness-enhancing rhizobacterium of Arabidopsis, as well as canola and maize, under either optimal or stressed (salinity) conditions. As such this is the first comprehensive report from Canada about the potential agricultural applications of Mucilaginibacter sp., in particular, from a wild plant, Scorzoneroides autumnalis (fall dandelion). This research will help in the development of bioproducts that can be utilized to increase crop growth and yield under both normal and stressful conditions, in an environmentally friendly manner for development of more sustainable, and possible climate change resilient, agricultural systems.

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Résumé

Les rhizobactéries sont des acteurs importants des cycles biogéochimiques et affectent de nombreux aspects de la physiologie des plantes. Il a été démontré que les plantes sauvages, en particulier celles qui poussent dans les terres arides, abritent une grande diversité d’espèces bactériennes contribuant au bon fonctionnement de leurs hôtes. Dans la mesure où aucune analyse holistique n’avait été faite jusqu’alors sur la diversité des rhizobactéries établies dans les espaces sauvages et agricoles du Sud-Ouest du Québec, le premier objectif de ce projet était d’identifier les communautés rhizobactériennes cultivables associées à différents types de plantes sauvages et cultivées de la région de Sainte-Anne-de-Bellevue. Un total de 446 souches de rhizobactéries ont été isolées et classées en 90 genres bactériens, les phyla dominants dans tous les échantillons étant Proteobacteria, Actinobacteria, Firmicutes et Bacteroidetes. La composition de la communauté bactérienne cultivable et sa diversité dépendaient de l’espèce végétale considérée. Dans le but de développer de nouveaux bioproduits pour l’agriculture durable, deux séries d’expériences ont été menées. Tout d’abord, les rhizobactéries provenant du réservoir indiqué ci-dessus ont été criblées pour identifier celles qui avaient le potentiel de favoriser la croissance des plantes. Leur capacité à promouvoir la croissance a été évaluée en utilisant l’écotype Columbia (Col-0) d’Arabidopsis thaliana. Quatre des 98 souches rhizobactériennes ainsi sélectionnées (souches n, L, K et Y), initialement associées à Phalaris arundinacea, Solanum dulcamara, Scorzoneroides autumnalis et Glycine max, respectivement, ont montré des effets significatifs sur la stimulation de la croissance, plus précisément en termes de surface foliaire totale et de poids sec, de longueur des racines et de capacité antioxydante totale, de capacité d’atténuation de la salinité et de teneur totale en proline et en chlorophylle. Les souches n et L ont également présenté, in vitro et in planta, des effets antagonistes contre deux phytopathogènes : Pseudomonas syringae pv. tomato (Pst) DC3000 EV (virulent) et l’isolat de type sauvage B191 de Botrytis cinerea. Le traitement bactérien a provoqué une augmentation de la transcription des gènes liés à la croissance, à l’accumulation d’osmolytes, à la réponse immunitaire et aux activités antioxydantes. La capacité des quatre souches sélectionnées à favoriser la croissance des plantes a également été caractérisée en culture pure. Étonnamment, la souche K n’a pas présenté une grande capacité à

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promouvoir la croissance in vitro, mais a surpassé la souche n in planta, malgré le fait que cette dernière était celle qui accumulait le plus de traits associés à la promotion de la croissance. Deuxièmement, les quatre souches sélectionnées ont été testées sur deux plantes cultivées, Brassica napus (le canola) et Zea mays (le maïs), pour leurs effets sur la promotion de la croissance et la réduction du stress salin. Dans les tests sur la germination et la croissance précoce des semis, les quatre souches, et en particulier L, K et Y, ont entraîné une augmentation de la longueur des racines et du poids sec des plantules du canola à la fois en absence et en présence de salinité. Les quatre souches ont aussi significativement amélioré la longueur des plantules (jusqu’à 33%) et le poids frais (jusqu’à 25%) du maïs en absence de stress. Dans les expériences en pot, la biomasse des plantes de canola et leur teneur en chlorophylle ont été augmentées par les quatre souches évaluées dans des conditions stériles. De même, la biomasse des plants de maïs et leur teneur en chlorophylle ont présenté un accroissement significatif avec toutes les souches dans des conditions non stériles en serre. Seules les souches n et L ont augmenté les teneurs totales en antioxydants et en proline, tandis que les souches L, K et Y ont augmenté la teneur totale en glucides dans la plante. L’absorption des nutriments a également été évaluée, avec des niveaux plus élevés d’absorption de fer et de phosphore dans les feuilles de maïs des plantes inoculées avec la souche n, et une teneur en azote accrue après le traitement par les souches n, L et Y. Dans l’ensemble, cette recherche (1) a mis en évidence, pour la première fois, la richesse du réservoir des diverses rhizobactéries associées aux plantes cultivées et non cultivées de la région de Sainte-Anne-de-Bellevue, Québec, Canada, ce qui ouvre la voie à la découverte de rhizobactéries favorisant la croissance des plantes cultivées, (2) a démontré que quatre isolats sur 98 ont significativement augmenté la croissance et la résistance au stress salin chez A. thaliana, in vitro et in planta et (3) a montré que les mêmes quatre souches ont amélioré également la croissance de deux plantes cultivées testées, le canola et le maïs, dans des conditions stériles et non stériles. Cette recherche aidera au développement de bioproduits qui pourront être utilisés pour augmenter, d’une manière respectueuse de l’environnement, la croissance et le rendement des plantes d’intérêt agricole dans des conditions normales et de stress. Ceci permettra de développer des systèmes agricoles plus durables et résilients aux changements climatiques.

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Acknowledegments Doing a Ph.D. is like preparing a challenging dish. From my personal point of view, I would say that the recipe takes 1 cup of ambition, 1 cup of perseverance, 1 cup of enthusiasm and approximately ½ cup of wits and luck, respectively. However, the people you meet along the journey are of invaluable importance, beyond almost everything else. They are the mysterious ingredient that will shape your mind and make a difference in creating your dish. Therefore, I would like to take this opportunity to acknowledge all of those who made contributions to this thesis. First, I am greatly indebted to my supervisor, Dr. Donald L. Smith, for offering me the opportunity to pursue my studies at McGill University and for allowing me to grow up in his lab. I would like to express my warmest thanks for his inspiring guidance, wisdom, support, encouragement, enormous patience and open-mindedness during each stage of the study, without which the project would not have been able to come to fruition. He has always been there whenever I have needed advice. The enthusiasm he has for his research was motivational for me in the Ph.D. pursuit. I am grateful for all the stories and experience he has shared with me, which has added color and spice to this journey. I would also like to thank the committee members, Drs. Jean-Benoit Charron (Department of Plant Science, McGill University) and Lyle Whyte (Department of Natural Resource Sciences, McGill University), for their advising and help, both academically and mentally, throught out my Ph.D. work. I am especially thankful for Dr. JB Charron for allowing me to use his lab facilities. My special thanks also go to Dr. Fazli Mabood (Inocucor Technologies Inc., Montréal, Canada) for his help with crop plant root sampling and generous guidance on rhizobacteria isolation. In addition, a big thanks to Fangwen (Institut de recherche en biologie végétale, Université de Montréal), for providing Pseudomonas syringae pv. tomato DC3000 and Botrytis cinerea B191, and for sparing no effort in giving me valuable advice on plasmid transformation, gene amplification and cloning. My gratitude is also extended to Gabriel Lambert-Rivest (Department of Plant Science, McGill University) for his guidance on conducting quantitative real-time polymerase chain reaction (qRT-PCR). I am grateful to all the members of Smith group at McGill, both past and present, for the advice, encouragement, friendship, and comic relief I have received, and in particular to Qianying Ruan, Uliana Shvyreva, Martyna Aleksandra Głodowska, Yoko Takashita, Rachel

Backer, Franziska Srocke, Tehmeena Mukhtar, Gayathri Ilangumaran, and Drs. Alfred Souleimanov, Dana Praslickova, Sowmyalakshmi Subramanian, Selvakumari Arunachalam, Timothy Schwinghamer, and Xiaomin Zhou. Special thanks must go to Yoko, who has helped me countless times, to water my plants in the lab. My sincere thanks go to Dr. Alfred Souleimanov for making the translation of the English abstract into French, and for providing advice on how to grow maize in the greenhouse. Special thanks also go to Valentin Joly (Département de Sciences Biologiques, Université de Montréal) for polishing my abstract (French version). I also gratefully acknowledge financial support from the Natural Sciences and Engineering Research Council (NSERC) of Canada during my Ph.D. candidature. I sincerely appreciate the Patricia Harney Scholarship from Nova Scotia Agriculture College (NSAC), Nova Scotia, Canada, for the years of 2012, 2013, and 2014. My stay at McGill has been enjoyable largely due to the friends that I have been lucky enough to have. I am grateful for the time spent with flat-mates and friends, for Selina’s generous help upon my arrival at Montréal, Amy’s hospitality as I finished up my first semester, for the funny trips and memorable get-togethers, and for many other people. I have been very fortunate to have Tina and Serella during their years of continued friendship, constant support, and infinite cheeriness. This Ph.D. thesis could not have been completed without help and support from the Department of Plant Science faculty and staff. I would like to express special thanks to Guy Rimmer, Lynn Bachand, and Carolyn Bowes. On a personal note, I would like to extend my heart-felt regards to my parents Chenghua and Ledu, and my paternal and maternal grandmothers Xianying and Shuyu, respectively, for their constant love, support, inspiration and understanding through all the ups and downs during my degree; although they have been thousands of kilometers away. I owe a special thanks to my lovely child, Dario, who is a precious gift and the joy of my life. I appreciate all your patience during mommy’s thesis writing. Moreover, I have been able to learn to manage a good balance between family care and quality research, which has allowed me to have a more productive strategy. I also greatly appreciate my mother-in-law, Shibi, who has helped a lot in taking care of my son during my times away from home. And most of all for my dearest and patient husband

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Fangwen Bai, whose faithful support, both scientifically and emotionally, during my thesis, was so much appreciated.

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Preface

Contribution of authors

This Ph.D. thesis consists of three experimental chapters (Chapters 3, 4, and 5), of which the candidate, Di Fan, was fully responsible for the experimental design, development and execution of the laboratory, growth chamber and green house research studies, collection and analysis of the data, discussion of the results, and writing of the thesis. However, this thesis could not have been achieved without the contribution of the supervisor, Dr. Donald L. Smith, who conceived the initial research idea and the concept of screening for plant growth-promoting rhizobacteria in Arabidopsis thaliana and testing in crop plants. Dr. Donald L. Smith provided guidance, advice and funding support during the preparation, laboratory work, interpretation of the findings, and writing of the thesis. Dr. Donald L. Smith also critically revised this thesis.

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Chapter 1 Introduction The rhizosphere is a biologically active soil zone surrounding plant roots in which complex relationships exist among plants, soil-borne organisms (i.e., algae, bacteria, fungi, protozoan and nematodes) and the soil itself (Ryan and Delhaize, 2001). Generally, plant-root associated microbes reside in three separate yet interacting niche spaces associated with the roots: the soil (rhizosphere), the rhizoplane (root surface), and the root itself. The microbial community associated with plant roots is much richer in microbes than is the surrounding bulk soil due to root-exuded metabolites that serve as nutrient reservoirs for the root associated component of the plant microbiome, or phytomicrobiome (Bais et al., 2006; Lundberg et al., 2012). Early terrestrial plants almost certainly had less sophisticated root systems than the species that eventually evolved from them, so that these early floral elements would have required microbial assistance in the acquisition of water and nutrients (Knack et al. 2015). A close association between microbes and terrestrial plants is thought to have existed for approximately half a billion years. Just as the microbes, including fungi, archaea, protists and bacteria, in human body, have been recognized as the human microbiome (Lloyd-Price et al., 2016), critical to human health and body functioning (Calo-Mata et al., 2016; Hugon et al., 2017), the plant-associated microbiome has now been recognized as the phytomicrobiome, containing a diversity of collaborating and competing species that are essential to the well being of their host plants (Smith et al., 2017). Arbuscular mycorrhiza (AM) constitute a near-ubiquitous and mutualistic symbioses between most terrestrial plants (70-90 %) and fungi that belong to the monophyletic phylum Glomeromycota. This relationship has been shown to be present at least 450 million years ago and probably played a role in establishing terrestrial plants (Porras-Alfato and Bayman, 2011). Nitrogen fixing root nodule symbioses evolved approximately 60 million years ago (Kistner and Parniske, 2002). It is likely that associations between plants and microbes evolved as plants developed and adapted to a range of diverse and often challenging environments. Even simple plants, such as Sphagnum sp., have highly complex associations with diazotrophs (Bragina et al., 2013). Hence, a plant growing in nature is not a single organism but a holobiont (phytomicrobiome plus plant – Hartmann et al. 2014), and the associated phytomicrobiome is a very well integrated and regulated community. Microbes within this community, also now considered as the second genome of the plant, can be either root- or phyllo-associated (Gkorezis et al., 2016); they actively interact with their hosts, forming mutually beneficial associations (Mueller and Sachs, 2015). Since roots are in constant contact with wet, nutrient-rich soil, the number and range of root-associated microbes exceeds the above-ground tissue-associated ones. Naturally, plants and rhizospheric microbes (in the rhizosphere or on the root surface) and endophytic microbes (inside plant tissues) have complex associations with each other, and these are critical for both plants and microbes, in terms of nutrients, survival, development, growth and reproduction. Root colonizing beneficial bacteria, known as plant growth promoting rhizobactera (PGPR), are known to include species of Pseudomonas, Azospirillum, Azotobacter, Klebsiella, Enterobacter, Alcaligen, Arthobacter, Burkholderia, Bacillus and Serratia, have been shown to assist plant growth and to control plant diseases (Kloepper et al., 1989; Glick, 1995; Bai et al., 2002; Rosenblueth and Martínez-Romero, 2006), either directly or indirectly, through nitrogen fixation (Goswami et al., 2016), antagonism to phytopathogens (Rashid and Chung, 2017), improving plant responses to abiotic stressors (Vurukonda et al., 2016), alteration/production of phytohormones, and nutrient mobilization (Lugtenberg and Kamilova, 2009); it seems likely additional mechanisms of plant growth promotion will be discovered as these relationships are further explored. Bacillus and fluorescent Pseudomonas spp. are the most studied and exploited bacteria, as biocontrol agents (Hass and Défago, 2005; Shafi et al., 2017). The communication between plants and soil microorganisms, through various signaling mechanisms regulate these mutually beneficial associations. A growing body of evidence indicates that plants secrete molecules that might act as signals initiating dynamic associations with soil microbes, and vice versa (Bais et al., 2006; Lugtenberg and Kamilove, 2009; Smith et al 2017; 2015a,b). The best understood example of such signaling occurs between leguminous plants and their corresponding

N2-fixing rhizobia, in which plant roots secret specific isoflavonoids to attract specific responsive rhizobia, which, in response, produce distinct lipo-chitooligosaccharides (LCOs), to signal back, initiating nodulation by the activation of nodulation-related genes within specific legumes (Mus et al., 2016). There is a clear need for increased crop plant production due to a growing world population, diminishing tillable land area and increasing meat consumption, all of which are driving the search for novel microbes for inclusion in inoculants that can be added onto both crop

plants and biomass-producing ones used to underpin the growing bioeconomy and allowing production of advanced biofuels and associated high-value bioproducts, e.g. Panicum virgatum (switchgrass) (Madakadze et al., 1999). Chemical fertilizers cause large increases in crop production but are expensive and can cause severe environmental damage (Babalola, 2010). Due to positive effects on plant yield (Vessey, 2003), indigenous PGPR have been developed as commercial inoculants, used to increase crop productivity (Hume and Shelp, 1990; Ravuri and Hume, 1992; Tabassum et al., 2017). Thus, it is interesting and necessary to continue the screening of rhizobacterial diversity for efficacious PGPR. Recent studies have revealed a clear signature of the host plant in determining its own microbiome community (Ai et al., 2015). Land use intensity also exerts pronounced effects on the distribution of bacterial communities, especially those associated with rhizosphere (Estendorfer et al., 2017). Collectively, these considerations led us to investigate the following areas: (1) the diversity of cultivatable rhizobacterial communities associated with the roots of selected plants in Southwestern, Québec, Canada, (2) the specific communities hosted by various plant species under the conditions present in Southwestern Quebec, (3) whether there are potential PGPR in this pool of microbes and, if there are, how they influence the growth and development of the model plant, Arabidopsis thaliana, and (4) whether the selected potential PGPR stimulate the growth and alleviate salt stress of the key Canadian crop plants, Brassica napus (canola) and Zea mays (maize). To address these questions, rhizosphere and endosphere bacterial communities of 5 crops and 21 uncultivated plant species were assessed using culture-dependent techniques and 16S rRNA gene sequencing. The obtained pool of bacteria was screened (evaluated for plant growth promotion) in A. thaliana, under optimal growth conditions, resulting in identification of promising bacterial strains, which were further assessed under salt stress conditions. The plant growth-promoting effects of selected isolated strains were evaluated both qualitatively and quantitatively. Their effects on the growth, development and gene expression of A. thaliana were determined, using a gnotobiotic Petri dish system and a non-sterile soil system, under both optimal conditions and salt stress. Root colonization was tracked using GFP-tagging of the strains most beneficial to plant growth. Finally, the chosen strains were further evaluated on two crop plants (canola and corn); they were characterized for effects on biomass production, nutrient uptake, secondary metabolite accumulation and enhancement of salt stress resistance. The main aims of this project were:

(1) Analysis of the root-associated microbial diversity of a selected set of plants (5 crop and 21 uncultivated plant species) using culture-dependent methods. (2) Identification of isolated rhizobacteria able to promote the growth of Arabidopsis thaliana under both optimal and salt stress conditions, and determination of mechanisms potentially involved. (3) The localization of the selected isolates in planta, and their effects on gene expression in A. thaliana. (4) The potential of the selected strains to act as PGPR for crop plants.

Chapter 2 Literature Review

2.1 What are root exudates? Typically, roots are underground organs of plants that grow through the soil with a complex architecture/anatomy (Uren, 2007). In addition to anchoring plants, providing water and nutrients to plants, and storing food and nutrients, roots exude a wide range of potentially valuable substances into the surrounding soil. Root exudation is essential for plant responses to changes in the local environment. The majority of the compounds secreted by plant roots are low-molecular-weight compounds such as amino acids, fatty acids, organic acids, phenolics, sugars, sterols, vitamins, and other carbon-containing primary and secondary metabolites (Ahemad and Kibret, 2014; Bertin et al., 2003; Uren, 2007). However, root exudates also include high-molecular-weight compounds such as mucilage (polysaccharides) and enzymes, few non-carbon compounds (inorganic ions, H+, water, etc.), and volatile organic compounds (e.g. carbon dioxide, certain secondary metabolites, alcohols and aldehydes) (Ahemad and Kibret, 2014; Uren, 2007). The amount and composition of exudates varies considerably with the type of soil, physiological status and species of plants and microorganisms (Kang et al., 2010), nutrient availability (Brimecombe et al., 2001), and the presence of microbes (Dutta and Podile, 2010). It has been estimated that between 5 and 21 % (Kuzyakov and Domanski, 2000) of the photosynthetically fixed carbon is found in root exudates of older plants while young seedlings may exude over 30 % (Marschner, 1995; Uren, 2007). Lipophilic compounds in root exudates may diffuse directly across the cellular membranes, while the release of most specialized metabolites and certain organic compounds are facilitated by vesicular transport (exocytosis) and/or membrane-bound transport proteins (e.g. the ATP-binding cassette family, the major facilitator superfamily, and the aluminum-activated malate transporter family) (Weston et al., 2012).

2.2 Roles of root exudates

Though the functions of many root exudates have not been identified, some have been shown to play important roles in interactions that determine the survival and growth of plants and soil microorganisms. Besides attracting soil microbes to the vicinity of plant roots, chemical components (e.g. defense proteins, phytoalexins, and other unidentified chemicals) of root exudates can also deter pathogenic microorganisms for the better survival of vulnerable root cells (Brigham et al., 1999). For example, phytotoxic root exudates (e.g. (±)-catechin (Bais et al., 2002); sorgoleone (Dayan et al., 2009) can mediate negative plant-plant interactions, causing allelopathy and autotoxicity (Yu et al., 2003; Perry et al., 2005; Khan et al., 2010), which strengthen a plant’s ability to ward off competition from plant neighbors among different species or within conspecific plants. Phytotoxic root exudates can also have positive effects in plant-plant interactions, such as induced herbivore resistance in plant neighbors (Glinwood et al., 2003).

2.2.1 Manipulating the rhizosphere environment Nutrients in root exudates stimulate microbial growth near or on plant roots, thus there is usually higher number (10-1,000-fold) of microbes that thrive in the rhizosphere than in bulk soil. Large range of organic and inorganic substances secreted by roots into the soil inevitably change the biochemical and physical properties of the soil surrounding the root system. Root mucilage secreted from continuously growing root cap cells can facilitate water storage and root-soil contact (McCully and Boyer, 1997). Ectoenzymes and organic acids such as citric, malic, and oxalic acid can increase the phosphorus availability (Dakora and Phillips, 2002; Uren, 2007). Phytosiderophores, as well as organic acids, can increase the availability of micronutrients, including manganese, iron, and zinc (Dakora and Phillips, 2002). These changes in moisture, nutrient availability and pH can affect plant growth, the surrounding microbe population (Dakora and Phillips, 2002; Castagno et al., 2008; Kuzmicheva et al., 2017), and the ability of root associated bacteria to synthesize antibiotics (Duffy and Défago, 1999). The age of host plants can also affect the biosynthesis of antimicrobial compounds by microbes in the rhizosphere; for example, only root exudates from older plants can induce 2,4-diacetylphloroglucinol (DAPG) production (Picard et al., 2000).

2.2.2 Plant-to-microbe signals To establish at least some mutualistic interactions, the corresponding partners have to secrete chemical signals to mediate the recognition and complex exchanges between them. From the viewpoint of plants, root exudates provide nutrients to microorganisms in their immediate vicinity. Signal compounds can allow plants and bacteria to recognize each other and prepare for establishment of symbiotic interactions, while avoiding attempts to enter into relationships with inappropriate organisms and the determents that could follow. Pseudomonas putida genes involved in amino acid uptake and metabolism of aromatic compounds were found to be up-regulated during interaction with maize roots, indicating the availability of specific nutrients in root exudates (Matilla et al., 2007). Plant roots initiate communication with soil microbes by secreting specific signal compounds that are attracted to and recognized by the microbes.

2.2.2.1 Flavonoids More than 10,000 flavonoids, with diverse physiological and ecological functions, have been identified in plants (Ferrer et al., 2008). The types and concentrations of flavonoids in root exudates vary extensively (Cesco et al., 2010; Nelson and Sadowsky, 2015). Flavonoids can be metabolized by some microorganisms as a carbon source; they also act as antioxidants and metal chelators (Cesco et al., 2010). Isoflavonoid and flavonoid compounds secreted by leguminous plant roots are among the best studied group of signal molecules which act as chemoattractants to rhizobia (genera Rhizobium, Bradyrhizobium, Azorhizobium, Mesorhizobium and Sinorhizobium), which form symbiotic associations with legumes and fix atmospheric dinitrogen into ammonia through activity of nitrogenase, while residing in nodules on the roots of legumes. Isoflavonoids, such as daidzein and genistein, are found only in members of the legume family (e.g. soybean (Glycine max), common bean (Phaseolus vulgaris), and pea (Pisum sativum)) (Kistner and Parniske, 2002). It is well documented that different legumes produce different isoflavonoids, which act as specific signals and interact with the transcriptional regulator nodD to induce nod genes in specific rhizobia (Peters et al., 1986); while most flavonoids lack this unique host-rhizobial specificity. The products of nod genes, so-called Nod factors, play a major role in the early stages of nodule organogenesis (Mabood et al., 2006a). Flavonoid exudation from host roots changes during symbiosis in several legumes. This alteration can fine-tune Nod factor synthesis (Schmidt et al., 1994) and, thus, could influence microbe colonization of nodules. 7

It has been shown that genistein application increases soybean crop yield and protein content (Smith and Zhang, 1999). The use of genistein, together with indigenous bradyrhizobial populations, to enhance nodulation in soybean, has been commercialized for over a decade and proven to increase grain yield by approximately 7 % (Leibovitch et al., 2001). Flavonoids may also play a role in the symbioses of some families of non-legumes (actinorhizal plants) able to form relationships with nitrogen-fixing actinomycetes, particularly Frankia species, though canonical nod genes are absent in those bacteria (Normand et al., 2007). Flavonoids exert a host-specific selection of compatible Frankia strains. This might be due to the exudation of phytoalexins by host plant roots which inhibit the growth of incompatible strains (Hassan and Mathesius, 2012). Flavonoids can also regulate a bank of other Rhizobium genes. For instance, exopolysaccharide synthesis, which is important for regulating defence responses in host plants, has been shown to be altered in Rhizobium fredii by genistein (Dunn et al., 1992); nevertheless, many of the proteins with expression altered in response to host flavonoids remain to be characterized (Guerreiro et al. 1997). In addition, flavonoids and other phenolic compounds have been shown to inhibit root pathogens, due to their role as antimicrobial toxins (Cushnie and Lamb, 2011) and anti- or pro-oxidants (Jia et al., 2010). Flavonoids can be induced by both symbionts and pathogens, which could be an additional factor in nodule promotion. Flavonoids can also be stored as phytoanticipins to exert quick defense responses against future attacks (Lattanzio et al., 2006)

2.2.2.2 Jasmonates Jasmonic acid is a hormone that occurrs ubiquitously in plants and plays a central role in plant growth, development and defense responses (Creelman and Mullet, 1997). Both legume and non-legume roots have been shown to exude jasmonates into the rhizosphere. It has been reported that jasmonates can induce nod gene expression (Mabood et al., 2005) and the production of N factors (Mabood et al., 2006b); moreover, jasmonates can also indirectly induce nod genes by inducing genes involved in (iso)flavonoid biosynthesis, such as flavone synthase FNSII-2 gene in Medicago truncatula (Zhang et al., 2007). The growth and yield promoting effects of jasmonates have been demonstrated under field conditions (Mabood et al., 2006c). Since (iso)flavonoids and jasmonates lead to synergistic effects on the production of Nod factors,

it is reasonable to speculate that these two inducers activate nod genes via different receptor proteins (Mabood et al., 2006b) and/or by different binding sites on the same receptors (Hause and Schaarschmidt, 2009).

2.2.2.3 Other nod gene inducers In addition to (iso)flavonoids and jasmonates, there are other compounds which have been shown to induce rhizobial nod genes, such as betaines and aldonic acids. Alfalfa seeds produce stachydrine and trigonelline (betaines) which activate nod genes in S. meliloti through NodD2 protein interaction (Phillips et al., 1995). Gagnon and Ibrahim (1998) found that erythronic and tetronic acids (aldonic acids), produced by lupin (Lupinus albus) roots, can induce nod genes in S. meliloti and Rahizobium fredii.

2.2.2.4 Strigolactones Strigolactones are versatile molecules that are synthesized from carotenoids, with at least 20 different natural canonical ones been structurally characterized, all of which consist of four rings (A, B, C and D rings) (Tokunaga et al., 2015). They can stimulate the germination of seeds of the parasitic plant Striga; this led to the original discovery of strigolactones and hence the name (Smith, 2014). They are well-known endogenous plant hormones that influence different physiological processes in planta, such as shoot branching, leaf shape and senescence, and environmental stress responses (Al-Babili and Bouwmeester, 2015). Strigolactones are also important plant-microbial signaling molecules in root exudates of diverse monocotyledonous and dicotyledonous plant species (Xie, 2016). Just like flavonoids, different plant species produce different strigolactones, indicating that specific modifications in structure allow specific mutualistic interactions among plant varieties and microbes. For example, strigolactones in root exudates of Lotus japonicus were found to induce extensive branching in germinating spores of the arbuscular mycorrhiza (AM) fungi (Akiyama et al., 2005); hyphal branching is a critical developmental step as it assists in contact with the host roots and the establishment of symbiosis (Giovannetti et al., 1994, 1996). Besides the role as a cue for arbuscular mycorrhizae, strigolactones were also discovered to play a role in the communication with rhizobia. Observations showed that rac-GR24, a synthetic strigolactone, had a stimulating effect in the nodule number in a concentration-dependent manner in Medicago

truncatula at low concentrations (De Cuyper, 2015). Though some discrepancies existed regarding whether strigolactones act on the plant side as a plant hormone in controlling nodule number, it has been confirmed that strigolactones can act as host detection cues for rhizobia by triggering the swarming motility of bacteria in a dose-dependent manner (Peláez-Vico et al., 2016). Although the ecological relevance of the swarming behavior needs to be further investigated, it is highly probable that, eventually, strigolactones can be used to stimulate the rhizobia-legume symbiosis. No function with other bacteria in the root has been observed for strigolactones so far (De Cuyper and Goormachtig, 2017).

2.2.3 Quorum sensing Many bacterial behaviors are controlled by a population-density-dependent mechanism, termed quorum sensing (QS). This was first described in the marine bacterium Vibrio fischeri, a mutualist with the bobtail squid (Eberhardt et al., 1981). QS activity in bacterial cell-cell communication is coordinated by small signal molecules, named autoinducers, of which N-acyl homoserine lactones (AHLs) effects on Gram-negative bacteria are the best studied ones (von Bodman et al., 2003). Gram-positive bacteria use peptide-signaling molecules for QS, whereas several Gram-negative bacteria utilize molecules like diketopiperazines (Holden et al., 2000). After their concentration exceeds a certain threshold in the extracellular milieu, in response to cell density, QS signals are perceived by AHL receptors (global transcription regulators LuxR or LuxR-like proteins) in bacteria and this then triggers the expression of numerous bacterial genes, many of which are involved in plant-microbe interactions (e.g. biofertilization, biocontrol, phytoremediation and phytostimulation) such as biofilm formation, nitrogen fixation, siderophore and volatile organic molecular production, extracellular hydrolytic enzymes production, synthesis of exopolysaccharides, toxins, as well as mobility (Gonzalez and Marketon, 2003; von Bodman et al., 2003; Müller et al., 2009). For example, two QS systems, LasI/R and RhlI/R, in Pseudomonas aeruginosa PUPa3 from rice rhizosphere positively regulate plant growth-promoting traits such as root colonization and antifungal activity (Steindler et al., 2009). It has been shown that B. subtilis biofilm improves plant growth, root proliferation and acts as biocontrol (Farrar et al., 2014). Recently, research on endophytism has extended to the concept of QS and its controls in regard to the community and species levels of endophytic fungi of grasses (Bacon and White, 2016). 10

The most widely studied QS signal mimics are halogenated furanones produced by the marine red alga Delisea pulchra (Givskov et al., 1996). The Delisea furanones are structurally similar to AHLs (Givskov et al., 1996) and have been shown to inhibit AHL-regulated behaviors in several bacteria by binding to AHL receptors and facilitating in the receptor degradation (Manefield et al., 2002). It has also been shown that several higher plants, including pea, tomato, Medicago truncatula and rice (Teplitski et al., 2000), exude QS signal mimics to interfere with QS regulation in bacteria, either inhibit or stimulate QS-dependent genes in corresponding bacteria. This implies that the QS signal-mimics might help plants to manipulate the diversity of bacteria that they encounter. Most of these compounds from plants remain unidentified and are structurally different from the AHLs, suggesting that they probably interact with the bacterial QS system through particular receptor proteins. The first QS signal mimic identified from plants was lumichrome, a riboflavin derivative (Rajamani et al., 2008). Other potential AHL mimics are phenolic compounds p-coumaric acid (Bodini et al., 2009), catechin (Vandeputte et al., 2010), and naringenin (Vikram et al., 2010). Plants can also perceive and respond to bacterial QS signals. Joseph and Phillips (2003) found that bean plants responded to homoserine lactone, a product of enzymatic degradation of AHLs by rhizobacteria, leading to increased transpiration in the shoots, which might potentially benefit nutrient flow. A proteomic study revealed that the accumulation of over 140 proteins including those that function in host defense responses from roots of Medicago truncatula was significantly affected by nanomolar concentrations of AHLs from two rhizobia (Mathesius et al., 2003). Treating the roots with AHLs also induced tissue-specific transcriptional activation of an auxin-inducible gene and three chalcone synthase genes. Moreover, the amounts or types of QS mimics exuded by Medicago truncatula changed upon root treatment with AHLs (Mathesius et al., 2003). It has been shown that nod gene-inducing flavonoids enhanced the expression of AHL synthesis genes in three legume-nodulating strains, leading to increased overall AHL production (Pérez-Montaño et al., 2011). All this evidence suggest a positive interaction might exist between the legume-rhizobia symbiosis and QS. Various types of AHLs have been shown to positively affect plant growth and systemic resistance in many plant species (Hartmann et al., 2014). It should be noted that some bacteria produce AHL-degrading enzymes to disrupt QS regulation in nearby bacteria, known as ‘quorum quenching’, which might also hold true for epiphytic or endophytic bacterial communications (Bauer and Robinson, 2002).

2.3 Plant growth promotion Bacteria thrive on nutrients in the vicinity of plant roots, and some of these rhizobacteria, referred to as plant growth-promoting bacteria (PGPR), can provide benefit to plants. Rhizospheric bacteria, or ectophytes, are those living in the rhizosphere or on the rhizoplane, whereas endophytic bacteria colonize inner plant tissues, without host damage or strong defense responses. The majority of PGPR belong to the genera Actinetobacter, Agrobacterium, Arthobacter, Azotobacter, Azospirillum, Bacillus, Bradyrhizobium, Burkholderia, Frankia, Pseudomonas, Rhizobium, Serratia, and Thiobacillus, (Vessey, 2003). Multitudinous mechanisms have been employed to promote plant growth and development by PGPR, including production of various substances (Ahemad and Kibret, 2014; Kloepper and Schroth, 1981), nitrogen fixation (Drogue et al., 2012), enhancement of mineral uptake, suppression of disease through antibiosis (Spence et al., 2014), induction of plant defense responses (Jung et al., 2008, 2011), etc. The cumulative effect of these complex interactions between host roots and rhizobacteria can lead to plant growth promotion (Fig. 2.1).

2.3.1 Root colonization Bacteria use cues in root exudates to locate plant roots. Chemotaxis, the directional motility toward root exudates, strongly contributes to rhizoplane competitiveness. Root exudates such as organic acids (De Weert et al., 2002) and carbohydrates (Bacilio-Jiménez et al., 2003) attract a range of PGPR, probably due to specific bacterial nutritional requirements. When moving towards roots in the rhizosphere, the metabolism in a competitive bacterium will be geared to enable optimal growth rate. Flagellar mobility, lipopolysaccharide (LPS) structure, and the bacterial Major Outer Membrane Protein (MOMP) all play an important role in early host recognition and adhesion (Dutta and Podile, 2010). After attaching to root surface, bacteria will establish microcolonies by increasing cell divisions, and then invasion of root tissue might occur at cracks formed at the emergence of lateral roots or at root tips. Enzymatic cell-wall-degradation might aid in this invasive process (Reinhold-Hurek et al., 2006). Once inside the roots, competent endophytes can further invade vascular tissue by passing the casparian strips, allowing systemic spreading into shoots (McCully,

2001). The competent endophytes will alter their cellular processes to facilitate dwelling (entering the endophytic life stage) or spreading (Reinhold-Hurek et al., 2006).

Figure 2.1 Mechanisms of plant growth promotion by rhizobacteria (Ahemad and Kibret, 2014).

2.3.2 Biofertilization A biofertilizer is “a substance which contains living microorganisms which, when applied to seed, plant surfaces, or soil, colonizes the rhizosphere or the interior of the plant and promotes growth by increasing the supply or availability of primary nutrients to the host plant” (Vessey 2003). As compared to chemical fertilizers, biofertilizer is cost-effective and environmentally friendly, and can improve soil structure, leaving no toxic effects.

2.3.2.1 Nitrogen fixation 13

Nitrogen (N) is the most vital nutrient for plants. Though it is abundant (78 %) in the atmosphere, atmospheric N is unavailable to plants. Biological nitrogen fixation has been shown to contribute 180×106 metric tons/year globally, out of which 80 % is from specific root/legume symbiotic association and the rest is from free-living systems (Graham, 1988). Nitrogen fixing bacteria are the most widely used bacterial fertilizers, which account for about 65 % of the nitrogen supply to crops worldwide. Bacteria and Archaea, including symbiotic nitrogen fixing forms, viz. Rhizobium, the obligate symbionts in leguminous plants (Zahran, 2001) and Frankia in non-leguminous trees, and non-symbiotic N2-fixing forms, have the ability to fix atmospheric nitrogen and enrich soil (Young, 1992). Bradyrhizobium species are slow growing in contrast to Rhizobium ones, which are fast growing. It has been shown that co-inoculation of Bradyrhizobium and PGPR enhances symbiotic nitrogen fixation by increasing root nodule number, dry weight of nodules, grain yield, and enhanced nitrogenase activity (Son et al., 2006) and increases nodulation and nitrogen fixation in Glycine max at suboptimal root zone temperature (Dashti et al., 1998; Zhang et al., 1996).

Free-living N2-fixing bacteria establish a very close relationship which is described as a non-specific symbiosis and is sufficiently close to the root such that the atmospheric nitrogen is fixed by the bacteria and is then taken up by the plant. These bacteria include Acetobacter, Azospirillum, Azotobacter, Azoarcus, Burkholderia, Gluconacetobacter diazotrophicus (Bhattacharyya and Jha, 2012), Bacillus, Herbaspirillum, and Paenibacillus (Goswami et al., 2015; Heulin et al., 2002). They can fertilize non-legume plants, including maize, sorghum, wheat, sugar cane (Boddey et al., 1991), rye grass (Gangwar and Kaur, 2009) and kaller grass (Reinhold et al., 1986). Oberson et al. (2013) reported that inoculation of Azotobacter chroococcum and Azospirillum brasilense in cereals resulted in significantly increased crop yields. Other than Azotobacter and Azospirillum, the most widely used nitrogen fixers in agricultural trails, genera Bacillus and Paenibacillus have also gained importance as they possess nitrogenase (nif) genes, which code for a nitrogenase complex that is required for nitrogen fixation (Ding et al., 2005). Kifle and Laing (2016a) found that all 8 diazotrophic bacterial strains from genera Bacillus, Pseudomonas, Burkholderia, Enterobacterm and Pantoea significantly stimulated plant growth and yield of maize, equal to that of a N fertilized control.

2.3.2.2 Phosphate solubilization

Phosphorous (P) is an essential macronutrient for biological growth and development, and is the second most limiting nutrient for plants after nitrogen. Though phosphorous reserves are abundant, it is not in the form that is readily available for plants, which can only uptake soluble forms of phosphate (mono- and dibasic phosphate) (Bhattacharyya and Jha, 2012). Some PGPR produce phosphatases to release soluble phosphorus from phosphate-rich organic compounds in the soil (mineralization) (Vassilev et al., 2006), while others produce organic acids (e.g. acetic acid, gluconic acid, 2-ketogluconic acid, lactic, isovaleric, isobutyric, and propionic acid) to solubilize phosphorus from mineral phosphate (solubilization) (Chen et al., 2006; Lipton et al., 1987; Patel et al., 2015; Zaidi et al., 2009), all of which make P available to plants and increase the P uptake by plants in the meantime. The most efficient phosphate solubilising bacteria (PSB) belong to genera Bacillus, Rhizobium and Pseudomonas (Govindasamy et al., 2011; Goswami et al., 2013; Goswami et al., 2015). The PSB Bacillus megaterium from tea rhizosphere has been shown to increase plant growth (Chakraborty et al., 2006). Application of PSB along with other PGPR reduced P application by 50 % without any significant reduction of grain yield in Zea mays (Yazdani et al., 2009). The use of PSB isolates as inoculants has been shown to enhance the growth of rice (Ashrafuzzaman et al., 2009). Moreover, PSB can be used for rapid revegetation of barren or disturbed land (Jeon et al., 2003), where soils are with low fertility in which nutrients, especially P, are scarce.

2.3.2.3 Iron availability Iron acts as a cofactor in many enzymes that are vital to important physiological processes such as respiration, phototsynthesis, and nitrogen fixation. The poor bioavailability of iron in rhizospheric soil foments a furious competition (Loper and Henkels, 1997). Some PGPR synthesize siderophores which can solubilize and sequester Fe3+ from the soil and then provide it to plants (Bloemberg and Lugtenberg 2001). Most of the siderophore-producing isolates belong to Gram-negative bacteria corresponding to the Pseudomonas and Enterobacter genera, and some Gram-positive ones, such as Bacillus and Rhodococcus (Tian et al., 2009). The major groups of siderophores are the catecholates, hydroxamates and carboxylates. Siderophore synthesis is modulated by a range of environmental factors, such as pH, the level of iron, the form of iron ions, the presence of other trace elements, and supply of carbon, nitrogen and

phosphorus (Duffy and Défago, 1999). It has been shown that siderophore-producing rhizobacteria increased early growth of soybean in non-sterile soil (Cattelan et al., 1999). The siderophore-producing rhizospheric bacteria, Bacillus megaterium, from tea, helped in plant growth and reduction of disease intensity (Chakraborty et al., 2006).

2.4 Phytoremediation Phytoremediation is the use of plants with their associated microorganisms, to remediate polluted soils and water (Weyens et al., 2009). Rhizospheric bacteria can degrade pollutants close to the root zone and thrive on root exudates and cell debris from plant roots, while endophytes can degrade organic contaminants absorbed or sequestered by plants, all of which form a synergistic system for the phytoremediation of organics (Weyens et al., 2009). For instance, Van Aken et al. (2004) showed that poplar plants infected with the endophyte Methylobacterium populum sp. nov., strain BJ001 could degrade 2,4,6-trinitrotoluene, hexahydro-1,3,5-trinitro-1,3,5-triazine, and octahydro-1,3,5,7-tetranitro-1,3,5-tetrazocine. In addition, when inoculated with a genetically tagged bacterial endophyte that naturally can degrade 2,4-dichlorophenoxyacetic acid (2,4-D), an herbicide for weed control, pea (Pisum sativum) plants showed an advanced ability to degrade 2,4-D from soil with no herbicide accumulation in their tissues (Germaine et al., 2006). Some PGPR exhibit metal resistance or sequestration ability and may reduce metal toxicity and affect metal translocation within plants (Khan, 2005), thus they can serve as growth-promoting bioinoculants for plants in metal-contaminated soils (Rajkumar and Freitas, 2008). Some PGPR mitigate toxic effects of heavy metals on plants via secretion of acids, proteins, and other chemicals (Denton, 2007). Others protect plants against the inhibitory effects of heavy metals through ACC deaminase activity (Amico et al., 2008). For instance, when inoculated with engineered nickel-resistant endophytic bacterium, Lupinus luteus L. showed an increase in nickel (Ni) accumulation in roots, however, the Ni concentration in shoots remained comparable with that in control plants (Lodewyckx et al., 2001). Endophyte-infected Alyssum bertolonii and fine fescues showed better growth under Ni (Barzanti et al., 2007) and aluminum (Zaurov et al., 2001) stress, repectively. Wu et al. (2006) reported that inoculation of heavy-metal resistant rhizobacteria led to stimulated plant growth (larger aboveground biomass) in Brassica juncea and protected plants from metal toxicity in Pb-Zn mine tailings. Li et al. 16

(2016) reported that an Enterobacter sp., isolated from roots of Sorghum sudanense grown in Cu mine wasteland soils, significantly increased the dry weight and root Cu accumulation of S. sudanense and significantly decreased the availibity of Cu in the rhizosphere soil. Some rhizobacteria, such as Pseudomonas putida, are tolerant to various kinds of heavy metals, which make P. Putida an excellent candidate for field application in metal-stressed soil (Chacko et al., 2009). Siderophores from Streptomyces acidiscabies E13 can bind nickel and promote cowpea growth under nickel stress (Dimkpa et al., 2008). Moreover, Wan et al. (2012) reported that infection of heavy metal resistant endophyte alleviated Cd-induced changes in Solanum nigrum L. and resulted in more biomass production, probably through improving antioxidant enzyme activities and enhancing uptake of essential mineral nutrition in infected plants.

2.5 Phytostimulation Besides nutrient supply, there are other plant growth promoting mechanisms in PGPR, either directly, such as production of plant growth regulators including phytohormones and certain volatilesor indirectly, through modulation of plant hormonal balance.

2.5.1 Indole-3-acetic acid (IAA) The best understood hormone in hormone-related activity among plant-associated bacteria is auxin, in which IAA is generally considered the most important native one (Ashrafuzzaman et al., 2009). IAA functions as an important signal molecule in the regulation of plant development such as cell expansion, division and differentiation, organogenesis, tropic responses, and gene regulation (Ryu and Patten, 2008). It has been speculated that about 80% of rhizobacteria produce IAA (Khalid et al., 2004). The synthesis and excretion of auxins into soil by PGPR make a great contribution to the plant-growth-promoting effect (Amara et al., 2015; Kaymak, 2011; Kidoglu et al., 2007; Steenhoudt and Vanderleyden 2000). IAA produced by rhizobacteria affect root systems by increasing branching number, weight, and root surface area, all of which lead to enhanced ability of roots to probe soil for nutrients, thus improving plant’s nutrition availability and growth capacity (Ramos-Solano et al., 2008). It has been shown that a microbial consortium comprising two species (Burkholderia sp. MSSP and Sinorhizobium meliloti PP3) with IAA production ability exerted significant increase in the growth of Cajanus

cajan seedlings (Pandey and Henkels, 2007). Mattos et al. (2008) showed that when inoculated with auxin-producing endophytic Burkholderia kururiensis, the growth of rice plantlets was promoted as revealed by an increase in IAA-induced gene expression. IAA is usually synthesized by rhizobacteria from tryptophan. It has been reported that the root weight of radish significantly increased upon inoculation with auxin-generating Pseudomonas fluorescens WCS365 (Kamilova et al., 2006). But this effect was not observed in cucumber, sweet pepper, or tomato, probably due to the much greater tryptophan concentration in radish root exudate than those vegetables (Kamilova et al., 2006), since the amino acid tryptophan in root exudates is used by PGPR as a precursor in the synthesis of the root-growth-promoting hormone IAA (Etesami et al., 2009; Karnwal, 2009). Though most of these PGPR utilize L-tryptophan for IAA production, very few PGPR, such as Azospirillum brasilense, can produce IAA by a L-tryptophan independent pathway, albeit most of the details in this route are yet unknown. The effects of IAA on plant growth have been shown to be concentration dependent (Arshad and Frankenberger, 1991) and species specific (Ahmad et al., 2005). The strains with the greatest amount of auxins led to maximum increase in growth and yield of wheat (Khalid et al., 2004), while those producing low concentrations of IAA, but with continuous release, could also improve plant growth (Tsavkelova et al., 2007).

2.5.2 Gibberellins PGPR also produce gibberellins (GAs), hormones involved in many developmental processes such as seed germination, stem elongation, shoot growth, flowering, and fruit setting of plants (Hedden and Phillips, 2000). There are as many as 136 GAs from 128 plant species and

4 GAs (GA1, GA3, GA4, and GA20) from 7 bacterial species have been identified (MacMillan, 2001). Plant growth promotion by GA-producing PGPR has been reported (Atzorn et al., 1988;

Gutierrez-Manero et al., 2001). It has been shown that a N2-fixing inoculant, Azotobacter chroococcum, produced GA3 and drastically influenced the growth of tomato seedling roots.

2.5.3 Cytokinins Production of cytokinins, hormones important in cell division, root growth, shoot formation, chloroplast development and bud formation, in several plant-associated microbes 18

(genera such as Pseudomonas, Azospirillum, and Bacillus) has been well characterized as early as the 1970s (Kaiss-Chapman and Morris, 1977; Persello-Cartieaux et al., 2001). Over all these years, more than 30 different growth-promoting cytokinins have been found in plants and plant-associated microorganisms (Amara et al., 2015). Cytokinins have been implicated in initiating nitrogen fixing nodule organogenesis (Tirichine et al., 2007). They also promote primary root growth and enhance branching (Ortiz-Castro et al., 2009). The complete abolishment of growth promotion effect by Bacillus megaterium in Arabidopsis thaliana triple mutant of cytokinin receptors indicated the important role of cytokinin perception in plant’s response to PGPR strain (Ortiz-Castro et al., 2008).

2.5.4 Regulation of Ethylene Ethylene is an essential plant regulator for normal growth and development of plants, e.g., breaking seed dormancy, increasing the number of roots, shoot and root growth differentiation, induction of flowering, leaf senescence, and fruit ripening (Babalola et al, 2007b). The sustained high level of ethylene after seed germination would impede plant root elongation and lead to abnormal root growth. Though they can synthesize ethylene in small amount (Babalola, 2010), PGPR are more widely able to produce 1-aminocyclopropane-1-carboxylate (ACC) deaminase to facilitate plant growth and development by decreasing the ethylene level in developing plants.

PGPR take up the ethylene precursor ACC and hydrolyze it into α-ketobutyrate and NH3 as carbon and nitrogen sources (Persello-Cartieaux et al., 2003). Under stress conditions, including flooding (Grichko and Glick, 2001), heavy metals (Belimov et al., 2001; Zhang et al., 2011), phytopathogen (bacteria, fungi, and viruses etc.) attack (Wang et al., 2000), drought and salinity (Nadeem et al., 2007; Yang et al., 2009; Zahir et al., 2008), extreme temperatures, bacterial ACC deaminase can reduce the elevated ethylene content and release plant stress, leading to the normal plant growth (Glick, 2014; Lugtenberg and Kamilova, 2009; Saleem et al., 2007). Actually, it has been suggested that ACC deaminase-production might be one of the major mechanism that PGPR utilize to offer promise for improvement of plant growth, especially under unfavourable environmental conditions (Hardoim et al., 2008).

2.5.5 Other plant growth regulators

It has become increasingly evident that rhizospheric bacteria can produce volatile compounds to mediate bacterium-plant interactions. For instance, hydrogen cyanide (HCN) produced by Pseudomonas fluorescen CHA0 has been shown to stimulate plant growth due to antifungal activity by suppressing black root-rot caused by the fungus Thielaviopsis basicola (Voisard et al., 1989), while for Pseudomonas aeruginosa PUPa3, HCN is the key factor for plant-killing effects in Arabidopsis thaliana (Blom et al., 2011). Some PGPR strains, such as Bacillus subtilis GB03 and Bacillus amyloliquefaciens IN937a, promote Arabidopsis growth and development by releasing volatile compounds, 2,3-butanediol and its precursor 3-hydroxy-2-butanone (acetoin) (Ryu et al., 2003), through the modulation of endogenous energy acquisition by the plant, such as increases in chlorophyll content and photosynthetic efficiency (Zhang et al., 2008). Banchio et al. (2009) reported that volatile compounds emitted from PGPR enhanced the accumulation of aroma compounds in sweet basil, which presents a novel strategy for food flavor enhancement. Though a well-known antimicribial compound produced by fluorescent pseudomonads (Couillerot et al., 2009), DAPG, when at lower concentrations, can also be a signal molecule in plants, inducing systemic resistance (Bakker et al., 2007), stimulating root exudation (Phillips et al, 2004), and enhancing root branching (Couillerot et al., 2011).

2.6 Biocontrol Plant diseases bring about annual crop-losses valued at more than $300 billion globally. There have been many efforts, such as conventional breeding, genetic engineering and agrochemicals, to control plant disease. The use of PGPR as biological control agents in living plants turns out to be an environmentally-friendly approach alternative to chemical pesticides (Lugtenberg and Kamilova, 2009). In order to effectively control plant disease, a bacterium must be able to compete successfully with other organisms for nutrients and have the potential to dominate the niches near or on the roots. Microbial control of plant diseases can indirectly benefit plant growth by preventing the growth or activity of plant pathogens through competition for space and nutrients, production of hydrolytic enzymes, antibiosis, and induction of plant defense mechanisms and inhibition of pathogen-produced enzymes or toxins (Lugtenberg and Kamilova, 2009).

2.6.1 Antibiotics Some rhizobacteria, especially antagonistic Gram-negative biocontrol bacteria, produce antimicrobial compounds that kill a wide range of pathogenic microorganisms; thus, these strains are efficient in controlling root and seedling diseases. Fluorescent pseudomonads produce various antibiotic substances, such as phenazines, pyoluteorin, pyrrolnitrin, DAPG, phenazine-1-carboxamide (PCN), cyclic lipopeptide, pseudomonic acid, and hydrogen cyanide (Haas and Défago, 2005). DAPG inhibits growth of phytopathogenic fungi (Nowak-Thompson et al., 1994). Although cyanide plays an important role in pseudomonad virulence affecting plant root growth (Rudrappa et al., 2008), positive correlations have been found between cyanide production in vitro and plant protection in the cucumber/Pythium ultimum and tomato/Fusarium oxysporum f. sp. radicis-lycopersici pathosystems (Ramette et al., 2003). Some Bacillus and Streptomyces spp. also produce a wide range of antibacterial and antifungal antibiotics such as subtilin, subtilosin A, sublancin, oligomycin A, zwittermycin A, kanosamine, and cyclic lipopeptide surfactin (Bais et al., 2004; Choudhary and Johri, 2009; Kim et al., 1999). It has been shown that phenazine and rhamnolipid biosurfactants act synergistically against soilborne diseases caused by Pythium spp. (Perneel et al., 2008).

2.6.2 Bacteriocins Some bacteria, mostly Gram-positive ones, can carry out ribosomal-synthesis of antimicrobial peptides known as bacteriocins, which kill other competitive bacteria, either in the same species or across genera, to facilitate in the underground warfare for nutrients and niches among rhizobacteria (Cotter et al., 2005; Drider et al., 2016). The bacteriocins of lactic acid bacreria (LAB) have received much interest in terms of biopreservatives due to their bactericidal effects to key Gran-positive pathogens such as Listeria monocytogenes and Staphylococcus aureus (Sobrino-López and Martín-Belloso, 2008). For example, one of LAB bacteriocins, nisin, is generally recognized as safe (GRAS) and is widely used as an effective food preservative (Sobrino-López and Martín-Belloso, 2008). Bacillus species also produce a diverse array of bacteriocins and represent one of the best characterized genera with regard to bacteriocin production and application (Abriouel et al., 2011). As compared to most LAB bacteriocins, Bacillus bacteriocins have somewhat broader spectra of inhibition including both Gram-positive and Gram-negative bacteria, yeasts, and/or fungi. 21

Bacillus thuringiensis, a Gram-positive spore-forming bacterium, has been found to produce a varied range of bacteriocins (Subramanian and Smith, 2015). Bacillus thuringiensis subsp. Kurstaki strain BUPM4 synthesizes a bacteriocin Bacthuricin (BF4), which can kill closely related bacteria in the rhizosphere (Kamoun et al., 2005). An antifungal substance, iturin, produced by Bacillus amyloliquefaciens CNU114001, showed antagonistic activity against cucumber scleotiorum rot, tomato gray mold, and cucumber and pumpkin powdery mildew (Ji et al., 2013). A novel bacteriocin, Thuricin 17, characterized by Gray (2005), is a low molecular weight peptide of 3162 Da, super strong in heat resistant, and is biologically active within a broad pH range (1.00-9.25). This bacteriocin inhibits not only the growth of related Gram-positive bacteria, but also a Gram-negative strain, Escherichia coli MM294(pBS42). It has also been shown to promote increased disease resistance in plants (Smith et al., 2008).

2.6.3 Extracellular lytic enzymes Besides antibiotics, some rhizobacteria also produce cell-wall-degrading enzymes such as chitinases, cellulases, glucanases, laminarinase and proteases to lyse and disrupt the structural integrity of the walls of target fungal cells, leading to antifungal activities (Kobayashi et al., 2002). For example, actinomycete isolates producing β-1,3-, β-1,4-, β-1,6-glucanases for glucan hydrolyzation in Phytophthora cell walls, were used for the control of raspberry root rot caused by Phytophthora fragariae var. rubi (Valois et al., 1996). Chitinase producing Paenibacillus illinoisensis has been shown to biologically control Phytopthora blight in pepper plants (Jung et al., 2005).

2.6.4 Siderophores Low-molecular-weight siderophores produced by PGPR can sequester iron from soil. For instance, fluorescent pseudomonads exude a diffusible pigment called pyoverdin (Pvd), which has a high affinity for Fe3+ ions. Upon binding the ion, ferripyoverdin (Pvd-Fe3+ complex) binds to a specific outer-membrane receptor at the bacterial cell surface and Fe3+ is subsequently transported into the cytoplasm where it is reduced to Fe2+ (Kloepper et al., 1980; Whipps, 2001). When Fe3+ concentration is low, e.g., in alkaline soils, PGPR actively synthesizing siderophores in the rhizosphere might exert biological control by depriving pathogens of iron, leading to growth inhibition in the competitors (Loper and Henkels, 1997; Shen et al., 2013). For instance, 22

siderophores produced by Pseudomonas fluorescens F113 has been shown to be involved in biocontrol of potato soft rot when soil iron is limited (Whipps, 2001).

2.6.5 Phytoalexins Arabidopsis, rice, corn, soybean and Medicago truncatula all produce different sources of antimicrobial substances, such as indole, terpenoid, benzaxazinone, and (iso)flavonoids. Antimicrobial molecules constitutively produced prior to attack are defined as phytoanticipins, while phytoalexins are a group of low molecular weight, antimicrobial compounds that are synthesized by plants upon microbe challenges. For example, rosmarinic acid (RA) in root exudates of sweet basil (Ocimum basilicum), elicited by fungal challenge, showed potent antimicrobial activity against several soilborne microbes, including an opportunistic plant pathogen Pseudomonas aeruginosa (Bais et al., 2003). Flavonoid synthesis in Medicago truncatula was elevated when plants were challenged by pathogen Phymatotrichopsis omnivora (Uppalapati et al., 2009). In addition, symbionts such as Pseudomonads and Bacillus spp. can enhance flavonoid production by plant roots; non-pathogenic strains of Fusarium oxysporum have been shown to increase phytoalexin production, leading to a delay on the onset of Fusarium wilt development (Landa et al., 2002). Camalexin, the major component of Arabidopsis phytoalexins, has been shown to play a major role in resistance in the powdery mildew fungus G. cichoracearum (Liu et al., 2016).

2.6.6 Control of other biotic stresses Except for pathogen control, some rhizobacteria are also active against weeds (Flores-Faragas and O’Hara, 2006), insects, and nematodes. For example, Pseudomonas fluorescens CHA0 and the related strain Pf-5, which release exoproducts that are toxic to pathogenic fungi, also exhibit potent insecticidal activity manifested by the production of insect toxin that is related to the insect toxin Mcf (Makes caterpillars floppy) (Péchy-Tarr et al., 2008). Morever, P. fluorescens CHA0 produces an extracellular EDTA-sensitive protease, AprA, which has been shown to contribute to the biocontrol activity against Meloidogyne incognita, a root-knot nematode (Siddiqui et al., 2005). Fungal endophytes have also been demonstrated to exert insect and nematode control, most of which is attributed to the production of alkaloid compounds by the fungal endophytes (Lewis, 2004; Mendoza et al., 2009). 23

2.6.7 Signal interference Many pathogens express virulence factors at high population densities, sensed when the extracellular concentration of quorum-sensing molecules reaches a threshold level. For instance, the synthesis of cell-wall-degrading enzymes in pathogenic Erwinia carotovora requires QS signal molecules. QS enhances virulence of pathogens and contributes to protection against antibiotics and the host immune responses. Bacterial response to QS signals and the subsequent distuption of bacterial QS molecules is one of the biocontrol mechanisms (Chang et al., 2017; Dong et al., 2002). Bacillus produces AHL lactonases that hydrolyze the lactone bond or AHL acylases that break the amide link, enabling these bacteria to compete more effectively with Gram-negative bacteria (Lee et al., 2002; Lin et al., 2003). Some potent but unidentified QS inhibitors were isolated from garlic extracts (Rasmussen et al., 2005). Interestingly, not like other treatments, QS disruption is neither bactericidal nor bacteriostatic and does not impose a strong selective pressure, thus bacterial resistance is less likely to develop. The identification of QS inhibitor agents is of considerable interest, especially in the development of biocontrol treatments against phytopathogens (Bacon et al., 2016).

2.6.8 Induced systemic resistance When a plant encounters pathogenic microorganism, its “immune system” is primed to defend against disease, a response called systemic acquired response (SAR), which was coined by Ross (1961). Salicylic acid (SA) (Vlot et al., 2009) and a regulatory protein NONEXPRESSOR OF PATHOGENESIS-RELATED (PR) GENES1 (NPR1) (Pajerowska-Mukhtar et al., 2013) seem to be major signals inducing SAR in plants (Sticher et al., 1997). Interactions of some PGPR with plants can result in systemic expression of a broad spectrum and long-lasting disease resistance to phytopathogens (i.e. fungi, bacteria and viruses), a phenomenon known as induced systemic resistance (ISR) (Kloepper et al., 1999; Verhagen et al., 2010). Moreover, the non-specific character of ISR provides an increase in the basal level of resistance to multiple pathogens simultaneously. This character is of benefit since there are usually several pathogens present under natural conditions (Thakker et al., 2011).

A wide varity of soilborne bacterial mutualists, including Bacillus, Pseudomonas, Trichoderma, can sensitize plant immune system for enhanced defense. Many bacterial compounds can induce ISR, such as antibiotics (e.g. 2,4-diacetylphloroglucinol), flagella, lipopolysaccharides (LPS), siderophores, cyclic lipopeptides, QS signal AHLs and volatile compounds (e.g. 2,3-butanediol, HCN and acetoin) (Choudhary and Johri, 2009; Lee et al., 2001; van Loon, 2007). With their side chain of the outer membrane lipopolysaccharides acting as an inducing agent, some Pseudomonas strains have been shown to induce ISR in Arabidopsis, carnation, and radish. Non-pathogenic rhizobacteria may induce defence response in plants in a similar way to pathogens, such as expression of pathogenesis-related proteins (PRs), reinforcement of plant cell walls, and production of phytoalexins (Hammond-Kosack and Jones, 1996). ISR is generally considered different from SAR in that 1) visible symptoms does not occur on host plants during ISR, 2) ISR does not involve PRs accumulation, 3) ISR involves jasmonic acid- and ethylene-mediated signaling pathway (Pieterse et al., 1998), while SAR is SA-dependent (Pieterse et la., 1998; Verhagen et al., 2004), and 4) the effectiveness of SAR and ISR against different attackers (e.g. bacteria, fungi, and insects) differ (Ton et al., 2002). Moreover, it has been found that some PGPR only specifically elicit ISR in certain plant species (van Loon, 2007). The complexities of SAR and ISR responses might be due to the specificity of the perception, by the plant, of bacterial signals (van Loon, 2007). Though most studies show that jasmonates and ethylene are central players in the PGPR-induced resistance in plants, there has been some research of PGPR demonstrating the triggering of SA-dependent type of ISR that resembles SAR as well. For instance, resistance induced by Bacillus subtilis FB17 against Pseudomonas syringae pv. tomato in Arabidopsis was shown to be associated with an increase in SA levels and enhanced PR-1 expression (Rudrappa et al., 2008). Moreover, the defense regulatory protein NPR1, which is an important transcriptional co-activator of PR gene expression upon SA-activation, has also been shown to be involved in jasmonate/ethylene-dependent ISR. For instance, in Arabidopsis npr1 mutant was not capable of mounting ISR upon Pseudomonas putida LSW17S root colonization (Ahn et al., 2007). Several PGPR strains, in which Pseudomonas fluorescens holds the largest number studied, have been shown to produce SA. It’s interesting that SA produced by some PGPR often becomes incorporated into SA-based siderophores thus making SA not the causal agent for

triggering SAR (Bakker et al., 2014). Mutants that do not produce SA and/or the SA-containing siderophore do not behave differently from the wild type. More recently, Feng et al. (2012) reported a novel molecular mechanism that genes related to biosynthesis and signaling of abscissic acid were involved in biocontrol agent-induced resistance to bacterial wilt in A. thaliana.

2.7 Novel type of signaling

2.7.1 Nod factors As described, plant-to-bacteria signals can induce the expression of the rhizobial nod genes, which leads to the production of lipo-chitooligosaccharides (LCOs), often referred to as Nod factors. LCOs generally consist of three to five 1-4, β-N-acetyl-glucosamine molecules (chitooligosaccharide backbone) and a fatty acyl chain attached to the N-terminal residue of the nonreducing end. Different types of LCOs occur due to the modification in the length of chitin backbone, attributes of the lipid side chain and the presense of various added groups at both reducing and non-reducing ends of the chitin backbone. Some nod genes required for the modification of chitin backbone or terminal sugar resides of LCOs play a role in host specificity determination (Gray and Smith, 2005). LCOs are key bacteria-to-plant signals required for the establishment of rhizobia-legume nitrogen fixing associations. They trigger a range of biochemical and morphological symbiotic responses in plant roots, such as calcium spiking (prerequisite to nodulation), ion fluxes (including H+, Cl－, and K+), root hair deformation, cortical cell division, changes in gene expression and hormone levels, leading to the nodule primordium formation (Geurts and Bisseling, 2002; Murray, 2011). After bacteria reach a receptive root hair, Nod factor-induced root hair deformation eventually trap the producing bacteria in a “shepherd’s crook” curl structure, within which a bacterial microcolony is established. Concomitantly, cortical cells are stimulated to divide, to form an original nodule meristem. Through actin cytoskeleton rearrangements (Yokota et al., 2009), infection threads elongate, carrying bacteria into the growing nodule. The enlarged and differentiated nitrogen fixing forms are known as bacteroids (Jones et al., 2007). It has been shown that LysM-RLK proteins (lysin motif proteins associated with a kinase-like domain) are key components of Nod

factor perception in plants, and the LysM domains recognize specific Nod factor structures characteristic of corresponding symbionts (Lohmann et al., 2010). Non-legume plants can also perceive Nod factors (Denarie and Cullimore, 1993), and our lab initially found that LCO application (nM level) can stimulate plant growth directly in both legume and non-legume plants (Souleimanov et al., 2002; Prithiviraj et al., 2003; Khan et al., 2008; Miransari and Smith, 2009; Khan et al., 2011; Wang et al., 2012). The direct growth stimulation effects of LCOs have been confirmed by root growth in Medicago truncatula (Oláh et al., 2005), and accelerated flowering and increased yield in tomato (Chen et al., 2007). It has been speculated that LCOs may have diverse enhancing effects on plant growth through, for instance, inducing seed germination, enhancing plant biomass (Chen et al., 2007), increasing photosynthesis (Khan et al., 2008), stimulating flavonoid genes (Inui et al., 1997), cell cycling and phytohormone-mimic functions (Dyachok et al., 2000, 2002). Nod factors have been shown to provoke defense resistance in plants, as Nod factors induce LjNFR1-dependent expression of some defense gene homologs in Lotus japonicus (Nakagawa et al., 2011), as well as elicit increases in total phenolic compounds, phenylalanine ammonia lyase (PAL), tyrosine ammonia lyse (TAL), and activities of peroxidase and catalase (Jung et al., 2008). But it should be noted that some Nod factor-induced defense is either transient or absent (Hogslund et al., 2009; Nakagawa et al., 2011), and the niche of defense-related reactions could be restricted to specific cells. How plants specifically respond to symbiotic or defense-responsive activation by LCOs requires further study. However, it cannot be overlooked that some chitinase-producing rhizobacteria, such as Paenibacillus illinoisensis KJA-424 and Bacillus thuringiensis subsp. pakistanin HD 395, can hydrolyze LCOs resulting in a substantial degradation of Nod factors and reduced nodule establishment (Jung et al., 2006). The effects of LCOs have been shown to be much greater when crops are exposed to stress (salt, drought, cold) (Schwinghamer et al., 2014; Subramanian et al., 2014; Prudent et al., 2015). Products based on LCO findings from our laboratory were used to treat seeds sown into several million ha of North American crop land in 2011. Though the downstream signaling pathways for nodulation are been studied (Gough and Cullimore, 2011), the mechanism by which LCOs exert direct plant growth enhancement remains largely unknown.

2.7.2 Bacteriocins 27

Bacteriocins are bacteria-produced peptides that are either bactericidal or bacteristatic to closely-related competitive bacteria (Jack et al., 1995). The first bacteriocin discoveried was colicin, produced by Escherichia coli in response to stress (Cascales et al., 2007). Many rhizobacteria such as Pseudomonas spp. (Parret and De Mot, 2002), Bacillus strains and nodulating endophytes (Oresnick et al., 1999) produce bacteriocins and these compounds have been implied in biocontrol and/or nodulation, thus leading to plant growth promotion. It is presumed that bacteriocins produced by PGPR exert competitive advantage to the producer strains (Wilson et al., 1998). Both lactic acid bacteria and Bacillus strains have three main classifications of bacteriocins, based on Post-Translational Modification (PTM) component, peptide size, heat and pH stability (Abriouel et al., 2011). When co-inoculated with Bradyrhizobium japonicum under nitrogen free conditions, Bacillus thuringiensis NEB17 promoted soybean growth, nodulation and grain yeild (Bai et al., 2003). Subsequently, a novel bacteriocin, thuricin 17, isolated from Bacillus thuringiensis NEB17 (Gray et al., 2006), showed stimulation in the growth of corn and soybean when applied to leaves or the root system (Lee et al., 2009), which comprise the first evidence of direct plant growth stimulation by a bacteriocin. Bacteriocin perception by leaf tissue results in enhancement of photosynthesis, nodulation and total biomass, indicating that this bacteriocin can promote plant growth directly (Lee et al., 2009). A protein produced by a PGPR Serratia proteomaculans shows similar results in soybean (Bai et al., 2002). The effects of thuricin 17 on plants are much stronger when stress (salt, drought, or cold) is present than under optimum conditions (Subramanian, 2014; Prudent et al., 2015). All these findings suggest a new mechanism of PGPR activity. There has been some research into the mechanisms by which bacteriocins directly stimulate plant growth. Jung et al. (2011) showed that the activity of defense-related enzymes such as PAL, guaiacol peroxidase (POS), ascorbate peroxidase (APX), superoxide dismutase (SOD), and polyphenol oxidase (PPO) were all significantly induced by thuricin 17 after 72 h of feeding through cut stems of soybean, which is in agreement with a previous report (Jung et al., 2008). These results indicate that a bacteriocin might exert its growth promotion effect via induced defense resistance in plants, which seems to be a common mechanism induced by PGPR, as previously described. Gray (2005) proposed an alternative mode of action, in which bacteriocins inhibit susceptible bacteria, hence reducing niche-space competition for the

producer strain, and also directly stimulating plant growth. It is noteworthy that generally thuricin 17 causes greater effects than do LOCs. In-depth research into the mechanisms by which thuricin 17 acts is imperative for further increases in crop production. The detailed signaling response mechanisms by bacterial molecules, e.g. thuricin 17, is currently under exploitation in our laboratory (Subramanian, 2013; Subramanian et al., 2016), and the manipulation of these mechanisms could help ncrease plant production.

2.8 Old-timer versus rising star

World food production needs to be increased to meet the projected population growth. Plant products serve as staple foods for humans and livestock. As technologies that increase crop production with minimal side effects on environment are preferred, inoculating plants with specific PGPR as biofertilizers and biocontrol agents to increase overall plant productivity is undoubtedly of great interest. The methods for PGPR treatments mainly include drench application, seed bacterization and seedling treatment (Babalola et al., 2007a,b). Commercial bacterial biofertilizers are prepared in various ways. Sporulating, Gram-positive bacteria can be prepared by desiccation, while Gram-negative bacteria with heat-resistant spores can be formulated into dry powder. Alternatively, suspension of bacteria in oil is also used (Honeycutt and Benson, 2001). Seed coating with commercial inoculants based on dry powder (lignite, charcoal, etc.) might interfere with shelf-life and cell viability. The viability of using biochar as a chronic-release delivery mechanism for PGPR has been examined (Glodowska, 2014), which demonstrated that biochar is a promising bacterial carrier and can improve soybean growth and reduce N fertilizer demand as well. Rhizobial inoculants have been used as commercial products to enhance nitrogen fixation in legumes (Vessey, 2003). Bradyrhizobium japonicum 532C has been commercilized and used for soybean nodulation (Hume and Shelp, 1990; Ravuri and Hume, 1992), and is also applied along with Nod factor inducers, e.g. genistein and daidzein (Leibovitch et al., 2001). Novozymes, now a part of BASF, has introduced a seed-applied product, Optimize®, which contains LCO and quality nitrogen inoculants such as B. japonicum for soybean (2003), peanut (2004), alfalfa and pea/lentil (2005) (Smith, 2005). McIver (2005) reported the release of products based on LCO-stimulation effects in a wide range of non-legume crops. Bacillus licheniformis SB3086 is 29

produced by Novozymes as a biofertilizer with phosphate solubilizing ability and antagonistic effect against Dollar spot disease of plants. There are some other bacterial strains that have been commercialized to act against disease of ornamental flowers and vegetable crops (Goswami et al., 2016). It should be noticed that the results from commercial PGPR inoculants sometimes are less consistent in the field than in controlled environments, and this has often been attributed to unpredictable climatic and environmental conditions, leading to the poor colonization of plant roots by inoculants (Zaidi et al., 2009). The unreliability in the field may also be due to the agonistic and antagonistic effects of indigenous rhizobacteria, host incompatibility, pathogens, rhizospheric conditions and climate. What should not be neglected is that, when plants are exposed to little stress or cultivated under optimal conditions (i.e., nutrient-rich soil and sufficient water supply), PGPR often exert little effect on plant growth. Energy costs are rising, and there are increased public concerns regarding the environmental impacts associated with crop production (for example, nitrogen contamination and production of greenhouse gases). Increasingly, consumers have concern over foods processed from genetically modified organisms. In addition, we are expecting crop plants to feed a growing population while also being capable of providing biofuels and other novel bioproducts (Ragauskas et al., 2006). It is imperative to develop environmentally friendly reliable technologies, such as plant growth stimulants, that can serve to promote overall plant productivity (Orrell and Bennett, 2013). Studies regarding the effects that PGPR molecules (i.e., bacteriocins) manifest on plant growth enhancement have begun to emerge, and recruiting highly effective yet economical compounds, such as bacteriocins from PGPR, for boosting plant productivity represent a promising strategy in modern agriculture. Nevertheless, we should bear in mind that certain metabolites produced by endophytes in vivo in order to compete for nutrients and niches or fine-tune the metabolism of host plants might be absent in axenic cultivation, such as in controlled environment research, a situation that could also hold true for ectophytes.

2.9 PGPR screening

Researchers usually utilize long, time consuming and tedious approaches to discover new PGPR strains. This entails isolating a great number of bacterial strains from rhizospheric soils of certain plants. These bacteria will then be tested using various biological assays (Ahmad et al., 30

2008; Wahyudi and Astuti, 2011) including assays for IAA and siderophore production, phosphate solubilization, and antipathogenic and antibiosis activity. These are only the methods for in vitro screening of bacterial isolates for their plant growth promotion (PGP) traits, thus typically, plant growth assay will be used to select from a large number of bacteria for their effects on crop plants, either in a gnotobiotic growth pouch or in soil, either of which will require large numbers of plants and is time consuming. Glick et al. (1995) developed a rapid and novel procedure for isolating new PGPR by testing the ability of bacterial strains to hydrolyze ACC and use ACC as a sole nitrogen source. Subsequently, the plant growth promoting activity is evaluated via a gnotobiotic root elongation assay using ethylene sensitive plant seeds such as canola. This view is supported by the work reported by others (Glick et al., 1998; Li et al., 2000). Long et al. (2008) reported that phytohormone manipulations by PGPR do not always result in positive growth responses in different plants, which might be due to the imbalance between phytohormones such as ethylene and IAA, and/or the PGPR’s incompatible interactions with non-hosts. Though ACC-deaminase activity, as well as IAA production, has been considered an efficient parameter for isolating promising PGPR (Khalid et al., 2004; Rahman et al., 2010; Shaharoona et al., 2007), other traits of PGPR are also important for selecting PGPR. If this were not so, the novel PGP agent bacteriocin thuricin 17 might be overlooked. More recently, Meldau et al. (2012) showed that a dramatically efficient PGPR, Bacillus sp. B55, improves Nicotiana attenuata growth neither through affecting IAA nor ethylene production in both wild type and ethylene insensitive (35S-etr1) seedlings, pointing to the possibility of other unexplored PGP mechanisms, which turned out to be the ability of B55 to reduce sulfur and produce dimethyl disulfide, which N. attenuata can use to alleviate sulfur deficiencies. An assay to screen for plant growth stimulating activity associated with rhizobacteria has recently been developed in our laboratory (Subramanian, 2011; personal communication), in which Arabidopsis thaliana is used as a model system (Zhang et al., 2008; Blom et al., 2011; Khan et al., 2011). A. thaliana, a dicotyledonous plant that belongs to the Family Brassicaceae (Cruciferae), though is not directly economically important, has been adopted in plant-microbe interactions studies due to its experimental advantages over crop species, such as small size, relatively short life cycle (3-5 weeks), and easy culture conditions. I have adopted this plant for initial testing as we have already worked out a growth promotion assay for it in our laboratory.

This assay is now quite well-developed and has been shared with industry partners. We have used this assay with a range of stress conditions, including low temperature, drought and salt. Of these three, salt stress, as well as drought, is the easiest to impose in a controlled manner and provides the most rapid results, hence, assay based on A. thaliana under salt stress is being deployed in this instance, along with an unstressed treatment, in case the growth stimulation mechanism of a particular bacterial strain does not involve stress. In addition, numerous mutants and transgenes are available in the Arabidopsis Biological Resource Center (ABRC) for research in the molecular and physiological mechanisms by which rhizobacteria modulate plant growth and development, thus making A. thaliana the most convenient plant system to work with for this project, as well as future works. The proposed method involves exposing A. thaliana seeds or seedlings (contained in Petri dishes) to bacterial suspension and monitoring early seedling growth. This assay (see section 4.3.2 for details) is inexpensive, quick and especially useful for the initial screening for effective PGPR.

Connecting text

Given the lack of the information regarding the culturable rhizobacteria community associated with crop and wild plants in southwestern Québec, we performed an isolation experiment to document the cultivable rhizobacteria community component of 6 crops and 20 wild plants in the region of Sainte-Anne-de-Bellevue, Québec, Canada.

The chapter was reformatted and is published as:

Fan, D., Schwinghamer, T., Smith, D.L., 2018. Isolation and diversity of culturable rhizobacteria associated with economically important crops and uncultivated plants in Quebec, Canada. Syst. Appl. Microbiol. https://doi.org/10.1016/j.syapm.2018.06.004, reprined with permission from the publisher

Chapter 3 Isolation and diversity of culturable rhizobacteria associated with economically important crops and uncultivated plants in Québec, Canada

3.1 Abstract With the increasing appreciation of studies that unravel the mutually beneficial interactions between flora and microbiota, the beneficial properties of rhizobacteria that improve plant fitness are increasingly valued. Plants, including medicinal species, are associated with microorganisms. often residing within their tissues. Holonomic analysis of the diversity of the established rhizobacteria in uncultivated plants is particularly scarce. Thus, the present study was conducted to access the rhizospheric and root-endospheric bacterial diversity of 6 crops and 20 uncultivated plant species in the region of Sainte-Anne-de-Bellevue, Québec, Canada, using pure-culture methods. On the basis of 16S rRNA gene sequence analysis, a total of 446 bacterial isolates (144 cultivable rhizospheric bacteria and 302 root endophytes) were distributed onto four phyla (Proteobacteria, Firmicutes, Actinobacteria and Bacteroidetes), 32 families and 90 bacterial genera. The phylum Proteobacteria constituted the largest group of the isolated rhizobacteria, with 40 and 61 % for ectophytic and endophytic bacteria, respectively (240 isolates). Representatives of the genera Bacillus and Pseudomonas were found to dominate in rhizosphere soil, while Microbacterium and Pseudomonas were the predominant endospheric bacteria. The bacterial community composition differed among plant species. Some genera were only present in association with specific plant species, such as Stenotrophomonas, Yersinia, Labrys and Luteibacter. Several members of the endophytic community were sometimes observed in the rhizosphere, and vice versa. Our results showed that the diversity of bacterial communities in the tested crops and uncultivated plants differed from each other, suggesting that plant species is particularly important in shaping the bacterial community component of the Sainte-Anne-de-Bellevue region. This is the first survey of culturable endophytic bacteria associated with uncultivated plants in Québec, Canada. Although the culturable bacterial community studies herein are assumed to represent a partial proportion of the whole

phytomicrobiome of the evaluated plants, the results from the present study confirmed that the crops and uncultivated plants of Québec, Canada represent an extremely rich reservoir of the diverse rhizobacteria. These are potential sources for the discovery of plant growth-promoting rhizobacteria. Future studies will determine the potential application of these isolates in plant growth promotion and/or biocontrol for agricultural purposes.

3.2 Introduction Humans began to cultivate plants in tropical and subtropical regions of the Middle East and Africa at least 20,000 years ago, aiming to improve food and fiber supply. The initial actual domestication of crop plants took place about 10,000 years ago in Southwest Asia, China, Mexico, and South America (Price, 2009). The estimated 7.6 billion people (http://www.worldometers.info/world-population/) now living in the world depend on cultivated plants for nourishment and quality life at an unprecedented level. Thus, it has been a major challenge to plant science to provide the agricultural productivity sufficient to feed the world’s growing population (Glick, 2014). Many agricultural practices were developed to meet this goal, including use of external chemical inputs such as fertilizers, fungicides, herbicides, pesticides, and genetically modified crops. These practices, however, were scrutinized publicly, and practioners were criticized for endangering the health of the environment and humans (Glick, 2014). Some people refuse to consume genetically modified produce related to concerns around potential health risks associated with altered genes (Zhang et al., 2016). Fertilizer application can lead to nitrogen and phosphate leakage into the environment (Adesemoye and Kloepper, 2009). Moreover, excess application of chemical herbicides, pesticides, and fungicides contributes to the development of weed and pathogen resistance, leading to higher chemical inputs and more economic investments in the development of new chemicals (Chavez et al., 2013). Thus, it is of vital importance to find alternative approaches. Microbiota associations are ubiquitous to animals and plants (Heckman et al., 2001; Rosenberg and Zilber-Rosenberg, 2016). This kind of relationship, developed between dissimilar organisms living together for mutual benefit, is referred to as symbiosis (Wallenstein, 2017). Roots of all plant species are associated with numerous microbial biota, some of which do not exert overt harmful effects and lead to improved growth and/or vigor of their eukaryotic hosts,

thus these are considered to be plant growth-promoting rhizobacteria (PGPR) (Gaiero et al., 2013; Rosenblueth and Martínez-Romero, 2006). It was demonstrated that, following rhizosphere occupation, bacteria may reside in interior parts of plants, and as such are recognized as endophytes (Chanway, 1996; Haedoim et al., 2015). Plant roots encourage colonization by PGPR by offering them with metabolizable organic matter and protective niches (Bais et al., 2006). In return, PGPR help their hosts take up water and nutrients (Kuan et al., 2016; Mumtaz et al., 2017; Oteino et al., 2015), grow faster (Imperiali et al., 2017; Zahid et al., 2015), produce more biomass and yield (Kudoyarova et al., 2017; Rudolph et al., 2015), withstand biotic stresses, such as pathogens (Haque et al., 2016; Yu et al., 2011) and insects (Rashid and Chung, 2017), and abiotic stresses, including salinity stress (Chatterjee et al., 2017; Mahmood et al., 2016), drought (Chen et al., 2017), low-temperature (Turan et al., 2012), heavy metals (Mallick et al., 2018) and organic pollutions (Pawlik et al., 2017). Plant-bacteria interactions have also been shown to play an important role in soil aggregation and stabilization around plant roots (Ndour et al., 2017), and exert critical impacts on the integrity, productivity and sustainability of agro-ecosystems (Welbaum et al., 2004). Therefore, perhaps one of the best ways to meet the afore mentioned challenges is to use microbial inoculants, an environmentally friendly alternative to chemical use, for sustainable agricultural systems. Distinct microbiomes have been found in association with specific plant species (Haedoim et al., 2015; Pawlik et al., 2017), indicating that microbiomes (phytomicrobiomes) are selected and supported by plants. Plant physiology, growth stage, soil properties (pH, temperature and moisture), plant nutrient access (Cai et al., 2017) and environmental factors, have also been found to play major roles in determining phytomicrobiome structure (Hardoim et al., 2012). However, many aspects of this mutually beneficial association are not well understood. For instance, the genetic and molecular mechanisms adopted during this process, by host plants to specifically select beneficial over pathogenic organisms, are not clear. Moreover, agricultural practices have been shown to alter the phytomicrobiome members recruited by plants (Pérez-Jaramillo et al., 2016; Wemheuer et al., 2017). It seems likely that, during the early period of crop domestication, plants with beneficial microbiomes were selected, as these microbiomes would result in favorable host plant traits, such as rapid growth, increased production, and disease resistance, all of which were in keeping with human needs. But there would be irreversible changes in microbial communities and soil physicochemical properties (Trivedi et al., 2016), due to the replacement of

indigenous flora with those associated with a relatively small group of crop species. For example, crop yield seems now to be less directly linked to the beneficial microbiomes associated with host plants. The widespread use of fertilizers may have jeopardized the selection of microbial species, as well as genotypes, that support microbial nitrogen fixation and mineralization, and phosphate solubilization (Hardoim et al., 2015; Schmidt et al., 2016; Yeoh et al., 2016). In addition, the variation in the soil bacterial community composition was shown to be greater for scrubland than agricultural systems, indicating reduced diversity due to agricultural practices (Ding et al., 2013). Xia et al. (2015) reported that endophytic bacterial species abundance and diversity was significantly higher in organically grown plants (tomato, melon, pepper and maize) than those grown under current conventional agricultural practices. Isolation and identification of native bacterial populations is required to understand the diversity and distribution of indigenous bacteria in the rhizosphere of plants, leading to a second “green revolution” with innovative and biological agricultural solutions to addressing the requirement for agricultural productivity and environmental sustainability (Majeed et al, 2015). The rhizospheres of many plant species have been well-studied and are known to be promising sources of fitness enhancing microbes (Philippot et al., 2013). A large body of literatures related to plant-growth promoting rhizobacteria exists in public databases, available at http://webofknowledge.com, with more than 3,500 articles over the past decade. However, tremendous effort has been put on research to isolate PGPR strains from fields that have been substantially modified by human activities (Kumar et al., 2016; Sharma et al., 2015), and more specifically, from rhizospheric soils (Zahid et al., 2015) and specimens (shoots, roots, seeds, etc.) of economic important plants (Abiala et al., 2015; Fan et al., 2016b; Xia et al., 2015). Plant species, and even specific genotype, has been shown to be a key determinant in shaping microbiome composition (Lundberg et al., 2012; Yang et al., 2016; Yeoh et al., 2017). Until now, most of the 16S rDNA sequencing-based root associated phytomicrobiome surveys were conducted on model plants and crops. In contrast, the root microbiomes of uncultivated plants have not been well characterized (El-Sayed et al., 2014; Zhao et al., 2011; Zinniel et al., 2002), but are beginning to receive increasing attention. There have been some studies in which the microbial community associated with wild plants, such as those of the Amazon (endophytic actinomycetes; Bascom-Slack et al., 2009) and tropical rain forests in China (endophytic bacteria; Qin et al., 2009), specifically associated with Ginkgo biloba (microfungi; Adamčíková and

Hrubík, 2015), Dodonaea viscosa (endophytic bacteria; Afzal et al., 2017), Commiphora wightii (rhizospheric bacteria; Patel et al., 2017), and Tamarix chinensis, Suaeda salsa and Zoysia sinica (ectophytic and endophytic bacteria; Fan et al, 2016a). Numerous studies have employed culture-independent molecular techniques, such as terminal restriction fragments length polymorphism (T-RFLP; Berg et al., 2005), denaturing gradient gel electrophoresis (DGGE) and pyrosequencing (Edwards et al., 2015; Schreiter et al., 2014; Ventorino et al., 2015; Yang et al., 2017), which can lead to a much improved perspective of obligate and difficult to culture rhizobacteria (Yang et al., 2017), and facilicate understanding regarding interactions among soil properties, microbiota, and host genetics, which could well be transformational in plant breeding and biotechnology work (Peiffer et al., 2013). Though the culture-based approach is laborious and might underrepresent some bacterial taxonomic units (Xia et al, 2013), culturing and identifying individual bacteria can yield pure isolates that are then available for examination and development into bioproducts for enhancement of plant growth and yield, or protection against diseases for economically important plants. With increasing concern around chemical-based agricultural practices, it is important to find more bacterial strains from a wider range of plant species and environments, in order to screen for potential PGPR and then develop successful bioproducts for development of sustainable agronomic practices. Researchers have turned their attention to a range of ecological niches, aiming to discover novel potential PGPR. Rhizobacteria were isolated from harsh climates (Kumar et al., 2017), poor soil conditions (Goudjal et al., 2016; Pradhan et al., 2017; Sankhla et al., 2015), extreme locations such as near active volcanos (Mishra et al., 2017) or areas untouched by human activities (Bascom-Slack et al., 2009; Huang et al., 2014). Nevertheless, the high functional trait diversity of indigenous rhizobacteria from unexplored and unmanaged sources, can facilitate the establishment and growth of plants in suboptimal or unfavorable conditions. The present study was therefore undertaken to analyze the structure of bacterial communities associated with the roots of crop plants (maize, canola, soybean, reed canarygrass, alfafa, and miscanthus), via isolation and identification of individual cultivable ectophytic and endophytic bacteria. Diversity studies are of great importance in understanding the ecological role of rhizobacteria in the holobiont. A great diversity of bacterial species contributes to the fitness of wild (uncultivated) host plants, especially those grown in impoverished land (El-Sayed

et al., 2014), but the endophytic bacterial diversities of many uncultivated plant species are largely unknown. In order to begin filling this gap, the primary objective of the present study was to evaluate, for the first time, the diversity of cultivable endosymbiotic bacteria that colonize the roots of 21 species of uncultivated plants in a field with low levels of human disturbance in Québec, Canada. Aspects of their classification were also presented.

3.3 Materials and Methods

3.3.1 Study site and sample collection Plant rhizosphere samples (root and rhizosphere soil) were collected from Sainte-Anne-de-Bellevue (Québec, Canada), which is an on-island suburb located at latitude 45°24′22″ N, longitude 73°56′44″ W, and an average elevation of 20.87 m above sea level in southwestern Québec (Fig. 3.1). The area is characterized by a cold and temperate climate with an annual rainfall of 932 mm distributed across the year and an average annual temperature of 6.3 °C. July is the warmest month with an average of 20.7 °C, while January is the coldest month (-9.7 °C) (https://en.climate-data.org/location/14946/). Various crops and wild plant species of southwestern Québec were surveyed for the presence of cultivatable rhizobacteria. Plants were selected based either on their importance to agriculture, on availability at sampling dates, on their vigorous growth habit, or on their biennial or perennial nature (potentially with more stable associated bacterial ecosystems). In the month of July 2012, intact root systems from 4 plants per agricultural species (Brassica napus, Medicago sativa, Glycine max, Phalaris arundinacea and Zea mays) were sampled from an experimental farm of McGill University, Québec, Canada (site A), by excavating the roots to a depth of 15-30 cm. The soil in close proximity to the roots of crop plants, which was considered as rhizospheric soil (von der Weid et al., 2000), was sampled concomitantly. For nodule sample collection, healthy, green soybean plants were excavated carefully. Adhering soil particles were removed, and large nodules with pink/red coloration, which indicates effective N2 fixation, were collected. Each nodule was excised along with a small portion of root. At least five nodules were placed in a preservation vial (Somasegaran and Hoben, 2012), which contained anhydrous CaCl2

as desiccant. The tightly-capped vials were kept at ambient temperature for long-term storage (up to several months) until further processing. It was postulated that, among the myriad ecosystems, those with great biodiversity would host a great diversity of endophytes (Strobel et al., 2004). Thus, an area of substantial wild plant biodiversity, located in Sainte-Anne-de-Bellevue (Québec, Canada), was chosen. In July 2013, the wild plants, including various species of grasses, legumes, medicinal herbs, and wildflowers, were photographed, and then the roots were collected, to a 5-15 cm depth, from a field site (site B). This study site was a small (less than 1 ha) field site which was quite botanically diverse, with the most common species being Poa pratensis. There was very minimal management of the site; aside from occasional herbage removal, there were no agricultural inputs (lime, fertilizer, herbicide, pesticide) for over 25 years and sampling did not involve endangered or protected species of plants, so no specific permission was required for sampling activity. Identified wild plants (authenticated by Dr. Donald L. Smith, Department of Plant Science, McGill University, Canada) included: Miscanthus × giganteus (family Poales) (at the edge of the area), Aster pilosus (family Asteraceae; reassigned to Symphyotrichum pilosum), Daucus carota (family Apiaceae), Taraxacum officinale (family Asteraceae), Scorzoneroides autumnalis (family Asteraceae), Ginkgo biloba (family Ginkgoaceae), Erigeron strigosus (family Asteraceae), Sonchus arvensis (family Asteraceae), Plantago major (family Plantaginaceae), Vicia cracca (family Fabaceae), Verbascum thapsus (family Scrophulariaceae), Tussilago farfara (family Asteraceae), Solanum dulcamara (family Solanaceae), Prunella vulgaris (family Lamiaceae), Euphrasia officinalis (family Orobanchaceae), Glechoma hederacea (family Lamiaceae), Campanula rotundifolia (family Campanulaceae), Lactuca canadensis (family Asteraceae), Myosotis arvensis (family Boraginaceae), Chrysanthemum leucanthemum (family Asteraceae) and Rumex crispus (family Polygonaceae) (Fig. 3.2). After collection, all the root samples were wrapped in plastic bags, carried to the laboratory immediately and stored at 4 °C and processed the following day.

Fig. 3.1 Geographical location of sampling sites for rhizobacteria isolation in Sainte-Anne-de-Bellevue, Canada. The figure was adopted from Google Map and modified.

Fig. 3.2 Representive pictures of 19 wild plant species harvested for endophytic bacteria isolation (July 2013; Canada; 45°24′22″ N; 73°56′44″ W).

3.3.2 Isolation and preservation of cultivable rhizobacteria 3.3.2.1 Isolation of ectophytic bacteria Excess soil was removed by shaking root samples, and then approximately 2 g of rhizosphere soil (including root-adhering soil fraction) were carefully collected from each root

sample and placed in a sterile tube (25 mL) and to each, 20 mL of sterile phosphate-buffered saline solution (PBS; per L ddH2O: 0.24 g KH2PO4, 1.44 g Na2HPO4, 0.20 g KCl, 8.00 g NaCl, pH 7.0) was added. The tubes were vigorously vortexed at least 5 times and then allowed to stand for at least 30 min at ambient temperature, to allow settling of large particles. The supernatants were serially-diluted to 10-5 with PBS. Triplicate 100 μL aliquots of the second to fifth diluents were plated on King’s B (KB; per L ddH2O: 20 g proteose peptone no. 3, 1.5 g TM K2HPO4, 1.0 g MgSO4, 10 mL glycerol, pH 7.2), Difco Nutrient broth (NB) and low-salt

Lysogeny broth (LB; per L ddH2O: 10.0 g tryptone, 5.0 g yeast extract, 5.0 g NaCl, pH 7.0) agar plates, which have often been used to isolate plant-associated bacteria (Chen et al., 2012; Fan et al., 2016b; Marasco et al., 2012). The isolation plates were incubated at 28 °C and a total count (colony forming units (CFUs) g-1 fresh weight soil) was performed each day until no further change occurred. Bacterial colonies were chosen based on differentiable colony morphological features such as color, elevation, opacity, shape, surface, margin, and diameter. The colonies were purified by repeated streaking of single colonies; they were maintained on agar plates at 4 °C or stored frozen in cryovials in the corresponding growth medium with 25 % (v/v) glycerol at -80 °C .

3.3.2.2 Isolation of cultivable endophytes Plant tissue samples were surface sterilized using a procedure adapted from Qin et al. (2009), with modifications. Fresh roots, and in some cases rhizomes, were brushed and then thoroughly washed with tap water to remove adhering soil and attached epiphytic bacteria. Any visibly damaged tissue was removed. Root segments were then rinsed in ddH2O several times, until the water was clear, followed by a further cleaning using 10 % SparkleenTM 1 TM (Fisherbrand ) solution (v/v), rinsed in ddH2O several times and then patted dry with sterile paper towels. Median parts of the root samples, from a pooled mixture within each plant species, were cut into 1-2 cm portions with sterile scissors, transferred into separate sterile tubes, surface sterilized with 70 % (v/v) ethanol (EtOH) for 10 min, rinsed three times with sterile ddH2O, and then shaken for 5-10 min in commercial bleach (3% available chlorine), followed by six changes of sterile ddH2O to remove the disinfectant. Surface disinfection was confirmed in two ways.

First, aliquots of ddH2O from the last wash were plated on KB, NB and LB plates, and they were examined for contaminant colonies following a 3 to 7-d incubation at 28 °C. Second, the

surface-sterilized tissue was imprinted onto the corresponding agar, incubated at 28 °C, and then checked for microbial growth. Following sterilization, the samples were thoroughly dried under a laminar flow and were divided into 3 subsamples. The root fragments (1 g) from each subsample, from each plant species, were then were macerated aseptically in PBS with a sterile mortar and pestle and the supernatants were diluted in 10-fold series in PBS to 10-3 and subsequently aliquots of 100 μL of the first to third diluents were plated in triplicate onto appropriate agar media to recover any bacterial endophytes present in the plant root tissue. The agar plates were incubated at 28 °C for 3-5 days, after which CFU g-1 were determined. In some cases, the roots were too hard to crush; in these cases, 1 cm root segments were placed separately on the corresponding medium and then incubated at 28 °C for 3-5 days (Chen et al., 2013). The endophytic bacteria growing from the cut ends of the root specimens, onto the agar plates, were selected and sub-cultured separately (Long et al., 2008). Morphologically different colonies, based on their rate of growth and differences in colony characteristics on the plates, such as size, shape, texture (e.g., shriveled, mucoid), and color, were chosen and purified three times on the original medium by single-colony streaking. Individual colonies were sub-cultured and frozen in the appropriate growth medium, containing 25 % glycerol at -80 °C, until use.

3.3.2.3 Isolation of endophytic bacteria from soybean root nodules Intact nodules from 11 soybean root samples were carefully severed from the root by cutting the root about 0.5 cm on each side of the nodule. Desiccated nodules were treated according to Somasegaran and Hoben (2012), with some modifications. Briefly, nodule samples were rehydrated by imbibing in cool tap water at 4 °C overnight. Following rehydration, nodule samples were gently rinsed several times in ddH2O until the water was clear. They were then immersed in 95 % EtOH for 15 s, surface sterilized for 2 min in commercial bleach (3 % available chlorine), followed by 5 changes of sterile ddH2O. The nodules were transferred by sterile forceps to clean sterile tubes and were kept at 4 °C overnight. To each tube, 1 mL of sterile 0.9 % NaCl soultion was added and mixed by gently tapping the tubes. The efficacy of tissue surface disinfection was verified by plating a 0.9 % NaCl soultion of the last washing step on Yeast-Mannitol medium (YEM; per L ddH2O: 10 g manitol, K2HPO4 0.5 g, MgSO4·7H2O 0.2 g, NaCl 0.1 g, Yeast Extract 0.4 g, pH 6.8, and 15 g agar for solid medium), and bacterial growth was monitored following a 2-7-day incubation at 28 °C. Following sterilization, the nodules (root

portion discarded) were squashed aseptically in 3 mL 0.9 % NaCl, using a mortar and pestle and the supernatants were serially-diluted in 0.9 % NaCl to 10-6. Triplicate 200 μL aliquots of the dilutions were plated on YEM agar medium and incubated at 28 °C in a dark incubator for 7 days. Colonies that varied in morphology (shape, appearance, margin and color) were chosen and re-streaked on YEM medium three times, until the colony morphology of each isolate was homogenous. Individual colonies were sub-cultured and preserved on plates at 4 °C for temporary storage. Pure-culture isolates were preserved at -80 °C in YEM medium containing 25 % (v/v) glycerol for long-term use. Note that although Bradyrhizobium sp. were isolated from soybean nodules, these slow-growing rhizobacteria were not pursued further in this study.

3.3.3 Taxonomic identification and diversity indices A standard PCR protocol was used for presumptive taxonomic identification of all the isolated rhizobacteria. Bacterial DNA was extracted using a TE (10 mM Tris-Cl, 1 mM EDTA, pH 7.5) boil extraction method (Yang et al., 2008). All isolated strains were grown separately in tubes containing 5 mL KB medium at 28 °C on a rotary shaker at 150 rpm. Then, 100 µL of the bacterial culture was collected and transferred to sterile 1.5-mL tubes and incubated at 95 °C. After 5 min, the tubes were centrifuged for 5 min at 8,000 g and were then placed on ice. The supernatants contained the DNA crude extract (OD260/OD230 was more than 1.8, and

OD260/OD280 between 1.8 and 2.0) and these were collected and used for PCR amplication reactions of the 16S rRNA gene. PCR of 16S rDNA and sequencing were conducted using the Genome Québec Innovation Centre (McGill University) service. In brief, genomic DNA (1 μL) was used as the template and the bacterial 16S rRNA gene was amplified with the KAPA2G Fast HotStart DNA Polymerase (Kapa Biosystems), using universal primers 27F (5’-AGRGTTYGATYMTGGCTCAG-3’) and 1064R (5’-CGACRRCCATGCANCACCT-3’) (Rubje et al., 2014; Winsley et al., 2012). The amplicons were purified with ampure beads, out of which 3 μL of purified PCR product was used for the Sanger sequencing using Big Dye Terminator V3.1 (Applied Biosystems) with the same primers mentioned above. Conditions consisted of an initial denaturation at 96 °C for 1 min, followed by 25 cycles of 96 °C for 10 s, 50 °C for 5 s, 60 °C for 4 min, hold at 4 °C . Sterile ddH2O was used as a negative control. The PCR amplified products (ca. 1350 bp) were separated by agarose gel electrophoresis and sequenced on a 3730XL DNA analyzer system (Applied 45

Biosystems). The obtained sequences were assembled, analysed and edited using DNAMAN 8.0. Pairwise similarity values were retrieved from the EzTaxon-e server (access time: November 2017, Yoon et al., 2017), by comparing the resulting 16S rRNA gene sequences to type strains, supplemented with the nucleotide Basic Local Alignment Search Tool (BLASTN) of the National Centre for Biotechnology Information (NCBI), to search for the closest matching sequence. Diversity of the ectophytes and endophytes in the plant samples was analyzed using the following equations.

S (1) Shannon diversity index (H´) = -  piln pi, where S is the total number of species i=1

obtained in the community, pi is the proportion of S made up of the ith species in a sample (MacDonald et al., 2017).

(2) Species evenness (EH) = H´ / ln S, where S is species richness. The value of species evenness ranges from 0 to 1, with 0 representing no evenness and 1, a completely even distribution in abundance among species (Zhang et al., 2012).

3.3.3.4 Statistical analysis Data were analyzed by one-way analysis of variance (ANOVA) and differences between control and bacteria-treatments were considered statistically significant at the P ≤ 0.05 level using Tukeys Honestly Significant Differences (HSD) test of the COSTAT ® statistical software (CoHort Software, Monterey, CA, USA). Non-metric multidimensional scaling (NMDS) was performed using the metaMDS function from the vegan (Oksanen et al., 2017) package in R (R Core Team 2017).

3.4 Results

3.4.1 Enumeration, isolation, and identification of rhizobacteria

For selection of bacteria that might be useful for plant inoculation, only cultivable microbial cultures can be used. To assess the rhizobacterial populations and diversity in the rhizosphere of 5 crop and 21 wild plant species, rhizobacteria, either from the rhizoplane, healthy roots or root nodules, were isolated. The water used to wash surface-sterilized root tissues or direct imprinting of the sterilized-surface root tissue failed to transfer any microbial colonies onto agar medium, proving the efficacy of the decontamination procedures, and indicating that the microbes were eradicated from the root surface. Thus, cultivation of the endophytic bacteria was successful. All the rhizobacterial strains were isolated by the standard streaking method. Based on morphological differences, individual colonies were picked from agar plates that had been inoculated with the supernatant of crude extract of rhizosphere soil or surface-sterilized roots. Keeping in view the morphological features and growth rates, the isolates were purified by at least three restreakings onto fresh corresponding agar medium. All purified isolates were propagated three times on KB agar medium and were eventually frozen in KB liquid medium containing 25 % (v/v) glycerol at -80 °C for long-term preservation. Generally, the bacterial cell numbers for the rhizospheric soil were, as expected, higher than the endosphere, except for Medicago sativa. Statistically higher numbers of cultivable bacteria, expressed as colony-forming units (CFUs) g-1 fresh weight of sample, occurred in the rhizospheric soil (109) of maize and decreased to 107 in alfafa (data not shown). The root samples had significantly lower population densities for culturable endophytes, varying from 104 to 106 g-1 FW of root. Specifically, the total endospheric microbial flora was found to be minimumal, at 1.2 × 104, with V. thapsus and maximum of 690 × 104 in D. carota (Table 3.1). Initially, based on colony morphological characteristics of isolated strainsa, a total of the 446 pure rhizobacterial strains were obtained. Of this number, 144 isolates (32.4 %) originated from the rhizospheric soil of 6 crop plant species, 48 strains (10.8 %) were recovered as endophytes from roots of crop plant species (canola, alfafa and rees canarygrass), and 252 strains (56.8 %) were obtained from within root tissues of 21 wild plants species (Table 3.1). The distribution of culturable endophytic bacterial isolates from crops and wild plants is given in Fig. 3.3. The highest percentage of culturable endophytes (12.9 %) occurred in root samples of D. carota, 9.9 % from M. sativa, and 6.3 % from T. farfara. While the lowest percentage of culturable endophytes was 2.0 % for V. cracca and 1.7 % for S. arvensis. Individually, the number of

endophytic bacterial isolates from biennial/perennial plants, as well as a medicinal tree (ginkgo), were comparable to that of crop plants. All the isolates were subjected to partial sequencing of the 16S rDNA and the sequences obtained were BLASTN-aligned with the sequences of genera available in the GenBank database, which provided maximum identity of these isolates and allowed presumptive genetic identification (Table 3.2). Based on the analyses of 16S rDNA sequences, all isolates were within the Domain Bacteria. These morphologically distinct isolates (446 strains) were divided into 6 bacterial divisions: Gammaproteobacteria (25 %), Alphaproteobacteria (20 %), Betaproteobacteria (9 %), Firmicutes (14 %), Actinobacteria (25 %) and Bacteroidetes (7 %) (Fig. 3.4a). These isolated rhizobacteria could be then be assigned to 90 different bacterial genera (Table 3.2), dominated by Pseudomonas (in Gammaproteobacteria), which contained 72 strains of the 446 isolates (16.1 % of the total isolates), Bacillus (9.0 %) in the Bacilli (13.5 %), Microbacterium (7.2 %) in the Actinobacteria, Phyllobacterium (6 %) in the Alphaproteobacteria and Variovorax (4.5 %) in the Betaproteobacteria. The phylogenetic analyses revealed that the majority of the rhizobacterial isolates shared high sequence similarities (greater than 98.7 %) with recognized species (Table 3.2). However, there were 39 strains showing pairwise similarities less than 98.7 %, to the closest reference type strains, indicating that potential novel species might have been isolated (Stackebrandt and Ebers, 2006).

Fig. 3.3 Distribution percentages of culturable endophytic bacterial isolates among the examined crops and wild plants.

Table 3.1. Number of rhizobacteria isolated from agronomic crops, prairie and wild plants.

No. of No. of Plant species (common name) CFUa ectophytes endophytes Agronomic crops Brassica napus (canola) 17 7 12 g Glycine max (soybean) 19 0 − Zea mays (maize) 50 0 − Miscanthus × giganteus (giant miscanthus) 23 11 66 e Prairie plants Medicago sativa (alfafa) 13 30 490 b Phalaris arundinacea (reed canary grass) 22 11 70 e Wild plants Daucus carota (wild carrot) − 39 690 a Aster pilosus (frost aster) − 9 61 e Taraxacum officinale (common dandelion) − 16 340 b Scorzoneroides autumnalis (fall dandelion) − 11 110 d Ginkgo biloba (ginkgo) − 13 99 d Erigeron strigosus (daisy fleabane) − 15 180 c Sonchus arvensis (field milk thistle) − 5 1.4 i Plantago major (broadleaf plantain) − 8 17 g Vicia cracca (blue vetch) − 6 11 h Verbascum Thapsus (great mullein) − 7 1.2 i Tussilago farfara (coltsfoot) − 19 230 bc Solanum dulcamara (bittersweet) − 8 9.2 h Prunella vulgaris (common self-heal) − 10 70 e Euphrasia officinalis (eyebright) − 11 58 e Glechoma hederacea (ground-ivy) − 9 34 f Campanula rotundifolia (harebell) − 10 29 f Lactuca canadensis (Canada lettuce) − 7 8.7 h Myosotis arvensis (field forget-me-not) − 12 98 d Chrysanthemum leucanthemum (ox-eye daisy) − 14 120 d Rumex crispus (curly dock) − 14 140 cd Sum 144 302 −, not determined a CFU, mean number of endospheric bacterial isolates from plant roots (104 g-1 fresh weight; n = 3). Different letters are significantly different at a probability level of 5 %.

Fig. 3.4 The classification of isolated rhizobacteria of crop and wild plants (a) total, (b) ectophytic and (c) endophytic bacteria to the Alpha-, Beta-, Gammaproteobacteria, Actinobacteria, Bacteroidetes or Firmicutes. The percentages indicate the relative abundance of isolates that were present in all the tested plant species to the total 446 isolates.

3.4.2 Diversity of culturable ectophytic bacterial The microbiota of rhizopheric soil associated with six crop plant species that were grown in an experimental farm (B. napus, G. max, Z. mays, M. sativa, and P. arundinacea) and an unattended field (M. × giganteus), was characterized by partial 16S rDNA sequencing of pure colonies of culturable bacterial strains. A total of 144 ectophytic bacterial isolates were obtained

and found to display considerable diversity. Details of the 16S rRNA gene-based identification of the bacterial isolates are given in Table 3.2. A large proportion (77 %) of the ectophytes formed a cluster of Gram-positive bacteria. The ectophytic bacteria could be classified into 35 distinct genera, belonging to 19 families and 4 bacterial phyla (Figs. 3.5 and 3.6). Among the 144 isolates, 50 were isolated from maize, 23 from miscanthus, 22 from reed canarygrass, 19 from soybean, 17 from canola and 13 from alfafa (Fig. 3.6). The most common isolates were in the phylum Proteobacteria, which included 16 genera and 57 isolates and constituted ca. 39.6 % of the total species types in this study, followed by the phylum Firmicutes, with 6 genera and 34 isolates (23.6 %). Members of the phylum Actinobacteria were the third most abundant group with 7 genera. Isolates in the Bacteroidetes were less abundant than those in the other three phyla and were in rhizospheric soils of all tested plants, except for alfafa. Isolates from the class Gammaproteobacteria, Actinobacteria and Firmicutes were obtained from all the rhizopheric soil samples. Gammaproteobacteria was dominant and accounted for 25 % of isolates from rhizospheric soil. Analysis at the genus level showed that Pseudomonas was the most frequently isolated genus (17.4 %, 25 strains) and at least 19 Pseudomonas species were isolated. Bacillus was the second most frequently isolated genus (13.9 %, 20 strains) and 11 Bacillus species were isolated. Other genera, such as Sphingomonas, Raoultella, Herbaspirillum, Rummeliibacillus and Nocardia were isolated much less frequently and were represented by minimum populations for only one or two strains, constituting 0.7 to 1.4 % of the total population. Among the isolates, Bacillus spp. were isolated from rhizospheric soils of all 6 crop plant species. Flavobacterium spp. and Variovorax spp. were the second-most common ectophytic bacteria. Certain genera were unique in certain plant species. Buttiauxella, Novosphingobium and Nocardia bacteria were only isolated from soil associated with roots and rhizomes of reed canarygrass and were represented by only one strain for each, whilst Clavibacter, Chryseobacterium and Acidovorax bacteria were only obtained from miscanthus associated soil. Rhizobium sp. was only isolated from soil associated with soybean roots (Fig. 3.6). Maize soil harbored the largest number of culturable ectophytic bacteria from 23 bacterial genera, while alfafa bore the smallest number of bacteria, belonging to only 6 genera (Fig. 3.6).

Fig. 3.5 Microbial distribution in the collection of culturable bacterial isolates based on 16S rRNA gene sequence similarity associated to alfafa, canola, reed canarygrass, soybean, miscanthus and maize. (a) Percentage composition of different phyla of ectophytic bacteria present in the tested plants, (b) Phylum level classification of ectophytic rhizobacteria. Individual bacteria were categorized into Phylum and compared among different plant species to illustrate distribution.

Fig. 3.6 (a) Diversity of individual ectophytic isolates (n = 144) broken into a genus level classification (n = 35) and (b) Bar chart showing the relative abundance (%) of ectophytic bacterial phyla, compared among six crops species.

3.4.3 Culturable endophytic bacterial community survey To obtain insight into how the endophytic bacterial communities differ among plant species, we attempted to isolate bacteria from the same microhabitate (root endosphere tissue) of 6 crop plants and 21 different wild plant species, with successes in 4 crops and 20 wild plant species (Table 3.1). The abundances of bacterial taxa were examined at the level of phyla and genera to determine whether there were any apparent shifts in the composition of the bacterial communities across plant species. The endophytic bacteria isolated in this study displayed considerable diversity. We sequenced the 16S ribosomal-RNA genes from 302 independent endophytic colonies, and these genes fell into a total of four principle bacterial phyla, 40 families and 72 genera, ca. 27 % of which were in the classes Alphaproteobacteria and Actinobacteria, followed by Gamaproteobacteria (24.3 %) (Fig. 3.4). Proteobacteria and Actinobacteria comprised the majority of the isolated strains and this was the predominant phylum group in all the investigated plant species. The most commonly isolated phylum of endophytic bacteria was Proteobacteria: there were 39 species (~54 % of the total endophytic bacterial types) and 183 isolated strains (~61 % of the total endophytic bacterial isolates) from this phylum (Figs. 3.4 and 3.7; Table 3.2). The genera (ie. Pseudomonas, Variovorax, Sphingomonas, Ensifer) included in the phylum Proteobacteria are shown in Fig. 3.7c. The second-most abundant phylum of the isolated endophytic bacteria was Actinobacteria: there were 19 species (~26 % of the total endophytic bacterial types) and 82 isolates (~27 % of the total endophytic bacterial isolates), out of which Microbacterium was the most frequently isolated genus (34 %, 26 strains) and at leaset 14 Microbacterium species were isolated (Fig. 3.7c). Other phyla identified included Firmicutes and Bacteriodetes. Firmicutes constituted approximately 9 % of the total isolates numbers, out of which only 4 genera (Paenibacillus, Bacillus, Rummeliibacillus and Staphylococcus) were obtained (Fig. 3.7d). There were 17 wild plant species, such as T. officinale, E. strigosus, E. 55

officinalis and C. rotundifolia, which did not have associated members of the phylum of Firmicutes. The lowest abundance was observed for the phylum Bacteriodetes. Examination of the distribution pattern among all the endophytic bacterial isolates showed that 9 genera comprised ca. 51 % of the 302 isolates, with the most predominantly isolated genera being Pseudomonas (15.6 %), Microbacterium (8.6 %), Bacillus (6.3 %) and Rhizobium (4.6 %). The frequencies of certain genera, such as Novosphingobium, Ensifer, Rhodococcus, Luteibacter, Pararhizobium, Glutamicibacter and Achromobacter falls between 0.7 and 2 %. There were some genera, such as Jatrophihabitans, Starkeya, Serratia, Neorhizobium, Rahnella, Curtobacterium and Glycomyces that were isolated rarely and were represented by only one or two strains. The distribution of root endosphere bacterial community members differed with rhizospheric microbiome. Even within the same family (8 wild plant species), the root endophytic bacterial community showed diverse compositions (Table 3.2). A comparative analysis indicated that, for S. arvensis, all the endophytes (5 strains) were from the Phylum Firmicutes, with one Paeniacillus sp. and 4 strains of Bacillus. For E. strigosus and L. canadensis, endophytic bacteria were restricted to groups of Alphaproteobacteria and Actinobacteria. Moreover, some genera showed a specific distribution; only Miscanthus × giganteus harboured Flavobacterium and Chryseobacterium spp. (Flavobacteria class). A small number of wild plants contained a limited number of strains in the class Sphingobacteria, including S. autumnalis with Mucilaginibacter and Chitinophaga spp., P. major with Chitinophaga spp., and D. carota with Pedobacter and Mucilaginibacter spp. Pseudomonas and Microbacterium were retrieved from various tested plants (each associated with 15 plant species out of 24), while some genera were confined to a very limited number of sampled plant species. For example, Herbaspirillum was found only in M. giganteus and T. farfara, Aminobacter only in M. sativa and E. strigosus, Nocardioides only in D. carota and E. strigosus, and Glycomyces and Rahnella only in P. vulgaris and R. crispus. Strains of the Achromobacter were mainly restricted to G. biloba (5 out of 6 isolates). Moreover, some genera were only found in one plant species, such as Agromyces, Rummeliibacillus and Arthrobacter in P. vulgaris, V. cracca and T. officinale, respectively (Fig. 3.8; Table 3.2). In general, M. sativa (a crop) and D. carota (a wild plant) showed the highest endophytic diversities among the tested 24 plant species (Fig. 3.7b).

16S rDNA sequences of most bacterial isolates showed 98.7 to 100 % similarity with available sequences in EzTaxon-e server. Among the different endophytic bacterial isolates, strains 6K7 and K46 exhibited 96.55 and 97.1 % 16S rRNA gene sequence homology with Roseomonas aquatica TR53T and Mucilaginibacter lappiensis ANJLI2T, respectively, strongly indicating that these two strains could be new members of the respective genera. It should be noted that, with the exception of Medicago sativa, Miscanthus giganteus, Phalaris arundinacea and Brassica napus, the sampled plants harbored restricted endospheric bacterial communities, while their rhizospheric soil fractions were inhabited by a more variable and complex microbiome (Fig. 3.6 and 3.7b), which is in agreement with the findings of Marasco et al. (2012) for Capsicum annum L. plants from a traditional farm. Species richness of the root endosphere sample was determined, and the index values according to Shannon, with species evenness indicating the diversity of bacterial endophytes, are represented in Table 3.3. There were significant differences amongst plant species, with the maximum diversity of bacterial endophytes observed in D. carota (3.00), which also showed equitable distribution of individual endospheric bacteria, and the minimum diversity in S. arvensis. Evenness index varied due to unequal distribution of individual species (Maheswari and Rajagopal, 2003). The relative lower evenness index might be the cause of frequent isolations of the same species and/or scarce or no appearance of other bacteria isolates. All the indices shown in Table 3.3 suggested a relatively high diversity of the endophytic bacterial community in most plant species.

Table 3.3. Species diversity in terms of Shannon index and taxa evenness of endophytic rhizobacterial assemblage from different plant species (a different letters indicate significantly different at a probability level of 5 %; n = 3)

Shannon index Species evenness Plant species (common name) (H)a (E)

Agronomic crops

Brassica napus (canola) 1.55 e 0.96

Miscanthus × giganteus (giant miscanthus) 2.02 d 0.97

Prairie plants

Medicago sativa (alfafa) 2.70 b 0.95

Phalaris arundinacea (reed canary grass) 1.16 g 0.72

Wild plants

Daucus carota (wild carrot) 3.00 a 0.97

Aster pilosus (frost aster) 1.89 d 0.97

Taraxacum officinale (common dandelion) 1.92 d 0.88

Scorzoneroides autumnalis (fall dandelion) 2.15 c 0.98

Ginkgo biloba (ginkgo) 1.84 de 0.88

Erigeron strigosus (daisy fleabane) 2.39 c 0.96

Sonchus arvensis (field milk thistle) 0.50 i 0.72

Plantago major (broadleaf plantain) 1.91 d 0.98

Vicia cracca (blue vetch) 1.26 f 0.91

Verbascum thapsus (great mullein) 1.95 d 1.00

Tussilago farfara (coltsfoot) 1.72 de 0.83

Solanum dulcamara (bittersweet) 0.90 h 0.82

Prunella vulgaris (common self-heal) 2.30 c 1.00

Euphrasia officinalis (eyebright) 2.15 c 0.98

Glechoma hederacea (ground-ivy) 2.04 cd 0.98

Campanula rotundifolia (harebell) 1.36 ef 0.84

Lactuca canadensis (Canada lettuce) 1.95 d 1.00

Myosotis arvensis (field forget-me-not) 1.98 d 0.95

Chrysanthemum leucanthemum (ox-eye daisy) 1.82 d 0.83

Rumex crispus (curly dock) 2.30 c 0.96

Fig. 3.7 Microbial distribution in the collection of culturable endophytic bacterial isolates on the basis of 16S rRNA gene sequence similarity associated to 3 crops and 21 wild plant species. (a) Percentage composition of different phyla of bacteria. Numbers indicate the relative abundance, expressed as a percentage of the total number of isolates, (b) Phylum level classification of endophytic bacteria. Individual isoaltes (n = 302) were grouped at the phylum level classification, and compared among plant species, (c) Distribution of strains at the genus level was examined for phylum Proteobacteria and (d) Actinobacteria, in which bars represent the occurrence of strains (n = 183) among 39 and 19 different bacterial genera, respectively. 62

Fig. 3.8 Abundance of individual endophytic isolates (n = 302) grouped at the genus level (n = 69) and compared among four crops and 20 wild plant species. They were organized into a meta-analysis bar chart, which reveals the abundance of specific species shifting among plant species. The different colors indicate different taxa.

3.5 Discussion

There have been numerous studies investigating the relationship between bacterial communities associated with specific plant species and agricultural management regimes, revealing large diversities of the bacteria associated with the roots of host plants (Xia et al., 2015). The large variety observed, and the variation between plant species, has been attributed to the effect of root exudates of the individual plants on the soil microbial biomass, community composition (Steinauer et al., 2016) and other aspects of plant-microbial interactions (Zhou et al., 2016). In many cases, specific rhizobacteria have been isolated and identified worldwide, from single plant species or under limited environmental conditions, especially agro-ecological niches (Kang et al., 2017; Kifle and Laing, 2016b). Wild plants are likely to harbor specific bacterial communities that differ from those associated with cultivated plants, which have been extensively bred and grown under intensive agrochemical inputs. Although plant associated rhizobacteria have been isolated from a variety of crop plants as well as some wild plant species (Wei et al., 2017), scant data are available on the identification of cultivable rhizobacteria from crops and wild plants in the region of Québec, Canada. The present work is the first report on the isolation and genetic diversity of culturable endophytic bacteria from wild plants in southwestern Québec. In this study, a culture-dependent approach was used to isolate rhizobacterial community members present in the rhizospheric soil and the roots of various crops (farm land) and wild plant species (relatively undisturbed field), most of which were perennial plants. It should be mentioned that, although the root specimens in the present study were surfaced disinfested and confirmed to be surface sterile on control agar medium, the green fluorescent protein (GFP)-tagged microscopic proof has not been provided, to show conclusively that so-called endophytic bacteria should be considered as true endophytes. In addition, only fast- to medium-fast-growing bacteria were selected from soybean nodules, the conventional symbiotic composition of nodule-associated bacteria was excluded from analysis. Agricultural practices have been shown to influence the root-associated microbiome composition (Egamberdieva et al., 2016), but this conclusion could not be reached for this study, since no regularly cultivated crop plant species was available from site B, where a group of wild plant species thrived. In our study, we sought culturable bacteria, rather than mega-analysis approach, to obtain a view of the bacterial community profile, aiming to build a rich reservoir for PGP screening in planta. While knowledge of the unculturable strains is extremely interesting, there is little that 64

can be done with these strains, in terms of utilization as plant growth promoting technologies, simply because they are unculturable. We used four different culture media, attempting to isolate specific groups of bacteria: LB medium was used to enrich for Bacillus, KB for Pseudomonas, NB for other fastidious bacteria, and YEM for Rhizobium. Numerous rhizobacterial strains were isolated from extracts of rhizospheric soil and surface-sterilized root tissues, which is consistent with previous reports of bacterial enumeration (Marasco et al., 2012; Xia et al, 2013). The root samples supported a total of 446 rhizobacteria, indicating the richness of the niche for plant-microbe interactions. These bacterial isolates were also analyzed for their phylogenetic diversity and distribution in association with specific host plants. In general, the population density of rhizospheric bacteria isolated from 3 plant species, Phalaris arundinacea, Miscanthus giganteus and Brassica napus, in the present study, was higher than the endospheric bacterial communities, which was consistent with results from Edwards et al. (2015). Rhizospheric soils have been considered as mesotrophic environments, due to plant root exudates (Dessaux et al., 2016). In addition, specific and rigorous selection by the host plants for mutualistic associates that are competent in inciting specific aspects of plant physiology that favor growth and stress alleviation might lead to a significantly lower density of the microbiome inside the roots. Phylogenetic analysis indicated that soil bacteria (144 isolates) from 6 plant species could be assigned to 7 classes within four bacterial phyla: Proteobacteria, Actinobacteria, Bacteroidetes, and Firmicutes. The dominant bacterial phyla identified in this study were not significantly different from those in other soil types, e.g., tropical agricultural land (Sul et al., 2013), German forest soils (Nacke et al., 2011) and Chinese forest land (Wei et al., 2017), all of which suggested that dispersal limitation was less important in determining bacterial community, at least at this classification level (Finlay, 2002). The majority of these rhizospheric bacteria belonged to the Proteobacteria, which was consistent with most of the previous observations from various crop plant species (Mendes et al., 2013; Spain et al., 2009). Isolated members of the phylum Proteobacteria had a high taxonomic diversity, which included 16 genera. The genus Pseudomonas predominated in all rhizospheric soil samples and has generally been reported as the most abundant rhizobacteria in the soil, and also as endophytes. The second and third most dominant genera were Bacillus and Flavobacterium, respectively. Thirty-five genera were shared by all soil samples; maize harbored 23, while alfafa was associated with only 6. The appearance

of unique genera in specific rhizospheric soil samples might be the result of effects exerted by factors, such as soil pH, nutrient content and root excreta. We also identified 302 endophytic bacterial strains from 4 crops and 20 wild plant species that could also be divided into the four-above-mentioned phylum, with members of Bacillus, Pseudomonas, and Microbacterium being dominant. This is similar to the results of Santoyo et al. (2016). Pseudomonas, again, displayed the most promising levels of colonization, and in the endophytic niche, this may have been due to its wide spectrum of carbon and fatty acid utilization (Ryan et al., 2008). Genera in the phylum Proteobacteria, including Pseudomonas, in phylum Firmicutes including Bacillus, in phylum Actinobacteria including Microbacterium, were conserved across almost all host plant species. Based on such data, we propose that these bacteria are the dominant culturable rhizobacteria in both the rhizosphere and endosphere, at least for the plants included in this study, which is in general agreement with Sood et al. (2008). Members of Pseudomonas, Microbacterium, and Bacillus are ubiquitous and frequently obtained as endophytes through culture-dependent approaches (Santoyo et al., 2016); thus, it was not surprising that bacteria from these groups were identified in association with most of the plants sampled. Though these three genera were absent for certain plant species, the proportion of rhizobacteria belonging to them in the present study suggested they are highly competent in the rhizosphere and endosphere, compared with those of other genera, and are well adapted to the near-root soil environment and other plant-affected niches, indicating there might be a core microbiota conserved across plant species. For all isolated genera from crop plants, there were overlaps between ectophytes and endophytic bacteria, while some (e.g., Raoultella) were exclusively found in the rhizosphere soil and some others (e.g. Xanthomonas) were only present as endospheric bacteria, indicating that compartment-specific enrichment might exist. The comparison between rhizosphere- and endosphere-bacteria of most wild plant species could not be achieved because of the lack of rhizospheric bacteria community data. Moreover, the endophytic bacterial load of wild plant species varied significantly, with some bacterial types only existing in association with specific plant species while others were spread relatively widely. For example, distinct Mesorhizobium sp. existed in the root-endosphere of only two uncultivated plant species, D. carota and E. strigosus. For the tested crop plants, two different Mesorhizobium strains colonized the interior roots of M. sativa. Plant species specificity was also shown by Lumactud et al. (2016) in the bacterial endophytic communities of four plant

species, Achillea millefolium, Solidago canadensis, Trifolium aureum, and Dactylis glomerata. Thus, our data implied that plant species is one of the driving forces shaping the structure of endophytic communities. Bacteria may be better able to adapt to environmental conditions through influences by root exudates from their hosts, the quantity and chemical composition of which varied greatly among plant species, as did their preference for specific groups of bacteria (Dennis et al., 2010; Kuzmicheva et al., 2017). Research regarding the molecular determinants and the genetic basis for the interactions in rhizobacteria colonization are still lacking. The culturable endophytic bacterial communities of a few wild plant species were characterized by a low diversity, suggesting that more specific selective media might be used to enrich specific taxonomic groups, or media containing root extracts to simulate natural environments (Vartoukian et al., 2016) could be used for endobacterial isolation. M. × giganteus is a C4 grass and a very good candidate for bioenergy production because of its high productivity and adaptation to harsh conditions. There has been some research regarding its association with endophytic bacteria, especially diazotrophs, such as Herbaspirillum and Azospirillum spp. (Davis et al., 2010). In the present study, we found that Herbaspirillum hiltneri existed in the roots of miscanthus, along with two Rhizobium sp., which probably play important roles in supplying biologically fixed nitrogen for miscanthus plants in nutrient impoverished fields (Keymer and Kent, 2013). The same Herbaspirillum speices was also found in the rhizosphere of maize and soybean and in the endosphere of T. farfara from an unmanaged field, as was the case with miscanthus, suggesting that this strain might have a long history of association with specific host plant species. P. arudinacea is a C3 forage and biofuel crop growing well on nutrient poor and contaminated soils. A few studies have been conducted on the influence of P. arundinacea on the composition of fungal (Heděnec et al., 2014) and bacterial communities (Espenberg et al., 2016), but these did not involve in depth bacterial identification, as in the present study. For the tested crop plants, the rhizobacterial community, at the phylum level, was similar to previous reports for various crop species. Proteobacteria was found to be one of the most prominent phyla in soil samples. Some unique genera in this phylum form mutualistic associations with plants. For example, a Tardiphaga robiniae strain was exclusively isolated from the rhizosphere soil of maize. It has also been isolated from legume plant nodules such as Vavilovia formosa (Safronova et al., 2015) and was recently described as a genus of rhizobia, but

little is known about the role of T. robiniae in plants. A Stenotrophomonas isolate was also obtained from maize associated soil. Interestingly, a nitrogen-fixing Stenotrophomonas was recently reported as associated with Saccharum officinarum (Ramos et al., 2011) and switchgrass (Xia et al., 2013), both of which are C4 grasses. However, among our tested plants, the wild plant T. officinale, a C3 perennial, also harbored Stenotrophomonas in its roots. Moreover, a Buttiauxella sp. was isolated from the rhizospheric soil of reed canary grass only. Further common isolated bacteria from tested crop plants included Pseudomonas, Streptomyces, Variovorax, Paenibacillus, and Bacillus. Chinese Ginkgo biloba is a valued plant recognized for its longevity, sturdiness and resistance to plant diseases (Huang et al., 2000). The associated endophytes might have special functions such as resistance to pathogens of G. biloba (Li et al., 2011), though this was not explored in the current study. There have been a few studies around the endophytic microbes of G. biloba cortical root cells (Kumar et al., 2009), barks (Zhan et al., 2017), leaves (Yang et al., 2015), branches and trunks (Leff et al., 2015). To the best of our knowledge, the present study is the first report on the occurrence of root-associated endophytic bacteria for G. biloba in Canada. The endobacterial communities contained representatives of 3 bacterial phyla and 8 genera, which was less diverse than a previous report for above-ground tissue-associated bacteria in G. biloba. Since older plant tissues have been shown to harbor more diverse communities than younger ones in G. biloba (Leff et al., 2015) and the plants collected here were seedlings, we postulate that the relatively low diversity observed herein was partially due to the young age of the sampled plants. Daucus carota is widely used for both consumptional and medicinal purposes (da Silva Dias, 2014). Only one paper about its bacterial endophytes has been published to date (Surette et al, 2003), in which the population densities (106) recovered from root tissues of D. carota was comparable to our results. There were 28 bacterial genera identified, 6 more than our result. The most common genera in the Surette study were Pseudomonas, Staphylococcus, and Agrobacterium, while ours were Psedomonas, Streptomyces, and Sphingopyxis. The apparent differences in bacterial composition might be due to soil properties and environmental conditions, since Surrette et al. (2013) conducted their work on farm lands, verus the minimally managed field of our study.

For the leguminous plants, M. sativa (from farm land) and V. cracca (from unattended field), the root endospheric bacterial community differed significantly from each other, with Bacillus and Actinobacteria being dominant with V. cracca and Protobacteria dominating for M. sativa. Since they were from two different sampling sites, it is not clear whether these large differences were due to species specificity or soil properties. In previous studies, only the nodule bacteria from these two species were examined (Cauwenberghe et al., 2014; Laslo et al., 2017). While the culturable root-endospheric bacterial population, in most of the other uncultivated plant species, including aromatic and medicinal plants, grasses and wildflowers, in the present study has not been previously reported.

3.6 Conclusions To the best of our knowledge, this is the first study to provide insight into the culturable bacterial communities of crops and wild (uncultivated) plants present in the region of Saint-Anne-de-Bellevue, Québec, Canada. We have demonstrated that crops and wild plants harbor a great variety of ectophytic and endophytic bacteria. In general, our results suggested that the compositional structure of bacterial communities in the Saint-Anne-de-Bellevue area was, in meaningful part, determined by plant species. The characteristic environmental fitness of uncultivated plants that grow in impoverished soil conditions may be, at least partially, due to the presence of beneficial endophytes. The metabolic versatility of bacterial ectophytes and endophytes may increase host plant fitness, providing them a competitive advantage over other indigenous (Foster et al., 2017). Since agrochemicals may have harmful impacts on ecosystems, there is an urgent need to develop biological agents, such as PGPR, as a better alternative for crop production and to assist in maintaining overall ecosystem stability (Gupta et al., 2015). The present study has provided a set of versatile rhizobacterial strains that are promising for exploration as potential PGPR, with a wide range of possible applications; they could be used to increase the health and production of food and energy crops, and to develop more sustainable agriculture practices. If these rhizobacteria (members of the overall phytomicrobiome) improve the ability of crop plants to manage stress, their deployment could result in a more climate change resilient agricultural sector. This was an initial attempt to assess bacterial diversity with culture-dependent techniques; culture-independent approaches, such as pyrosequencing are needed for analyzing the full extent of rhizobacterial diversity and its association with aspects of 69

local environment, such as soil conditions and plant species. However, only culturable bacteria have the potential to be useful as crop productivity enhancing technologies.

Table 3.2. Molecular identification of culturable rhizobacterial isolates using 16S rDNA as query sequences. Isolation source, species/strain and 16S rDNA-based taxonomical identification for each taxonomic unit examined are shown (a isolate name with only a number means it was from rhizosphere soil).

Isolate name Host plant Result of identify Similarity (%) Phylum Class analysis

2a Zea mays Sphingomonas 100.00 Proteobacteria Alphaproteobacteria kyeonggiensis (KC252615)

4 Z. mays Arthrobacter sp. 98.52 Actinobacteria Actinobacteria

5 Z. mays Arthrobacter pascens 99.46 Actinobacteria Actinobacteria (AF235091)

6 Z. mays Rhodococcus 100.00 Actinobacteria Actinobacteria globerulus (BCWX01000023)

8 Z. mays Microbacterium 99.78 Actinobacteria Actinobacteria testaceum (X77445)

9 Z. mays Variovorax 99.47 Proteobacteria Betaproteobacteria boronicumulans (AB300597)

10 Z. mays Flavobacterium 98.48 Bacteroidetes Flavobacteria panaciterrae (JX233806)

11 Z. mays Arthrobacter oryzae 99.68 Actinobacteria Actinobacteria (CLG_48533)

12 Z. mays Streptomyces 99.25 Actinobacteria Actinobacteria cinereoruber subsp. fructofermentans (AB184647)

13 Z. mays Aminobacter 99.89 Proteobacteria Alphaproteobacteria aminovorans (AJ011759)

14 Z. mays Paenibacillus 99.27 Firmicutes Bacilli xylanexedens (CLG_48537)

18 Z. mays Bacillus 100.00 Firmicutes Bacilli butanolivorans (LGYA01000001)

19 Z. mays Lysobacter soli 98.76 Proteobacteria Gammaproteobacteria (EF623862)

20 Z. mays Pseudomonas 99.77 Proteobacteria Gammaproteobacteria caspiana (LOHF01000033)

21 Z. mays Flavobacterium 99.02 Bacteroidetes Flavobacteria frigidimaris (jgi.1107687)

22 Z. mays Bacillus 100.00 Firmicutes Bacilli zhangzhouensis

(JOTP01000061)

Bacillus safensis (ASJD01000027)

50 Z. mays Herbaspirillum 99.06 Proteobacteria Betaproteobacteria hiltneri (CP011409)

65 Z. mays Caballeronia sp. 98.94 Proteobacteria Betaproteobacteria (CP014505)

67 Z. mays Bacillus megaterium 99.90 Firmicutes Bacilli (JJMH01000057)

68 Z. mays Stenotrophomonas sp. 99.65 Proteobacteria Gammaproteobacteria

71 Z. mays Bacillus aryabhattai 100.00 Firmicutes Bacilli (EF114313)

72 Z. mays Bacillus simplex 100.00 Firmicutes Bacilli (BCVO01000086)

Brevibacterium frigoritolerans (AM747813)

96 Z. mays Brevibacillus brevis 99.48 Firmicutes Bacilli (AB271756)

121 Z. mays Paenibacillus 99.66 Firmicutes Bacilli polysaccharolyticus (EU912452)

Paenibacillus

cucumis (KU201962)

124 Z. mays Agromyces salentinus 99.41 Actinobacteria Actinobacteria (AY507129)

Agromyces allii (DQ673873)

125 Z. mays Ensifer morelensis 100.00 Proteobacteria Alphaproteobacteria (AY024335)

126 Z. mays Rhodococcus 100.00 Actinobacteria Actinobacteria pedocola (KT301938)

128 Z. mays Flavobacterium 97.18 Bacteroidetes Flavobacteria branchiicola (HE612102)

129 Z. mays Solibacillus 100.00 Firmicutes Bacilli isronensis (AMCK01000046)

Solibacillus kalamii (KT763359)

130 Z. mays Bacillus gibsonii 99.77 Firmicutes Bacilli (X76446)

133 Z. mays Bacillus wiedmannii 99.79 Firmicutes Bacilli (LOBC01000053)

Bacillus proteolyticus

(KJ812418)

134 Z. mays Flavobacterium 99.79 Bacteroidetes Flavobacteria endophyticum (KJ873109)

136 Z. mays Rummeliibacillus 99.10 Firmicutes Bacilli pycnus (AB271739)

137 Z. mays Pseudomonas 100.00 Proteobacteria Gammaproteobacteria migulae (AF074383)

141 Z. mays Pseudomonas moorei 100.00 Proteobacteria Gammaproteobacteria (AM293566)

183 Z. mays Variovorax 98.62 Proteobacteria Betaproteobacteria ginsengisoli (AB245358)

184 Z. mays Staphylococcus 100.00 Firmicutes Bacilli pasteuri (AF041361)

185 Z. mays Rhodococcus 100.00 Actinobacteria Actinobacteria wratislaviensis (BAWF01000105)

186 Z. mays Rhodococcus sp. 99.53 Actinobacteria Actinobacteria (CP019572)

187 Z. mays Raoultella planticola 99.76 Proteobacteria Gammaproteobacteria (JMPP01000074)

190 Z. mays Microbacterium 99.64 Actinobacteria Actinobacteria trichothecenolyticum

(JYJA01000006)

191 Z. mays Flavobacterium 99.75 Bacteroidetes Flavobacteria aquidurense (jgi.1107986)

192 Z. mays Brevibacillus 100.00 Firmicutes Bacilli laterosporus (AB112720)

210 Z. mays Rhodococcus 100.00 Actinobacteria Actinobacteria erythropolis (BCRM01000055)

214 Z. mays Streptomyces 100.00 Actinobacteria Actinobacteria stelliscabiei (LBNW01000338)

219 Z. mays Bacillus sp. 99.77 Firmicutes Bacilli

220 Z. mays Tardiphaga robiniae 98.43 Proteobacteria Alphaproteobacteria (FR753034)

221 Z. mays Bacillus asahii 99.39 Firmicutes Bacilli (AB109209)

224 Z. mays Pseudomonas lini 99.77 Proteobacteria Gammaproteobacteria (AY035996)

226 Z. mays Streptomyces sp. 100.00 Actinobacteria Actinobacteria

83 Glycine max Streptomyces sp. 100.00 Actinobacteria Actinobacteria

84 G. max Flavobacterium sp. 97.85 Bacteroidetes Flavobacteria

85 G. max Pedobacter 99.89 Bacteroidetes Sphingobacteriia panaciterrae (AB245368)

106 G. max Pseudomonas 99.58 Proteobacteria Gammaproteobacteria koreensis (AF468452)

173 G. max Pseudomonas 99.53 Proteobacteria Gammaproteobacteria kilonensis (LHVH01000037)

Pseudomonas corrugata (D84012)

174 G. max Flavobacterium 99.53 Bacteroidetes Flavobacteria saccharophilum (jgi.1107675)

175 G. max Variovorax 99.53 Proteobacteria Betaproteobacteria boronicumulans (AB300597)

176 G. max Pseudomonas 99.30 Proteobacteria Gammaproteobacteria frederiksbergensis (AJ249382)

177 G. max Flavobacterium 97.38 Bacteroidetes Flavobacteria branchiicola (HE612102)

Flavobacterium ginsenosidimutans

(GU138377)

178 G. max Herbaspirillum 99.08 Proteobacteria Betaproteobacteria hiltneri (CP011409)

179 G. max Brevibacterium 100.00 Firmicutes Bacilli frigoritolerans (AM747813)

180 G. max Bacillus gibsonii 99.88 Firmicutes Bacilli (X76446)

181 G. max Pedobacter 98.38 Bacteroidetes Sphingobacteriia ginsengisoli (AB245371)

182 G. max Bacillus simplex 99.79 Firmicutes Bacilli (BCVO01000086)

254 G. max Variovorax 100.00 Proteobacteria Betaproteobacteria paradoxus (BCUT01000013)

258 G. max Rhizobium 99.87 Proteobacteria Alphaproteobacteria mesosinicum (DQ100063)

Rhizobium alamii (AM931436)

260 G. max Bacillus sp. 100.00 Firmicutes Bacilli

261 G. max Pseudomonas 100.00 Proteobacteria Gammaproteobacteria brassicacearum

subsp. neoaurantiaca (EU391388)

262 G. max Polaromonas sp. 100.00 Proteobacteria Betaproteobacteria (AKIV01000055)

B2 Daucus carota Rhizobium sp. 100.00 Proteobacteria Alphaproteobacteria

B3 D. carota Pedobacter humicola 98.64 Bacteroidetes Sphingobacteriia (KT032156)

B4 D. carota Caulobacter 99.74 Proteobacteria Alphaproteobacteria rhizosphaerae (KX792139)

B7 D. carota Mucilaginibacter 97.77 Bacteroidetes Sphingobacteriia sabulilitoris (JQ739458)

Mucilaginibacter lappiensis (jgi.1095764)

B8 D. carota Streptomyces 99.77 Actinobacteria Actinobacteria alboniger (LIQN01000245)

B69 D. carota Mesorhizobium 100.00 Proteobacteria Alphaproteobacteria caraganae (EF149003)

B70 D. carota Sphingomonas sp. 100.00 Proteobacteria Alphaproteobacteria (JOOE01000006)

B71 D. carota Nocardioides caeni 97.74 Actinobacteria Actinobacteria (FJ423551)

B72 D. carota Roseomonas lacus 99.61 Proteobacteria Alphaproteobacteria (AJ786000)

B73 D. carota Sphingopyxis 99.37 Proteobacteria Alphaproteobacteria fribergensis (CP009122)

B75 D. carota Bosea thiooxidans 99.38 Proteobacteria Alphaproteobacteria (jgi.1048902)

K1 D. carota Pseudoxanthomonas 99.54 Proteobacteria Gammaproteobacteria sacheonensis (EF575564)

K4 D. carota Sphingobium 99.71 Proteobacteria Alphaproteobacteria czechense (KQ130434)

K5 D. carota Pedobacter humícola 99.52 Bacteroidetes Sphingobacteriia (KT032156)

K6 D. carota Streptomyces 100.00 Actinobacteria Actinobacteria caeruleatus (KQ948975)

Streptomyces ciscaucasicus (KQ948339)

K7 D. carota Gordonia sp. 100.00 Actinobacteria Actinobacteria

(CP002907)

K8 D. carota Streptomyces 100.00 Actinobacteria Actinobacteria lavendulae subsp. lavendulae (JOEW01000098)

K9 D. carota Polaromonas sp. 99.77 Proteobacteria Betaproteobacteria (AKIV01000055)

K10 D. carota Pseudomonas 99.58 Proteobacteria Gammaproteobacteria brassicacearum subsp. neoaurantiaca (EU391388)

K12 D. carota Sphingobium 100.00 Proteobacteria Alphaproteobacteria herbicidovorans (JFZA01000008)

K13 D. carota Ensifer adhaerens 99.89 Proteobacteria Alphaproteobacteria (JNAE01000171)

K14 D. carota Microbacterium 99.46 Actinobacteria Actinobacteria trichothecenolyticum (JYJA01000006)

K15 D. carota Novosphingobium sp. 98.56 Proteobacteria Alphaproteobacteria

K18 D. carota Bosea vaviloviae 100.00 Proteobacteria Alphaproteobacteria (KJ721000)

K20 D. carota Caulobacter segnis 99.74 Proteobacteria Alphaproteobacteria (CP002008)

Caulobacter vibrioides (AJ009957)

K21 D. carota Sphingopyxis 99.28 Proteobacteria Alphaproteobacteria bauzanensis (NISK01000001)

L1 D. carota Variovorax 99.58 Proteobacteria Betaproteobacteria boronicumulans (AB300597)

L2 D. carota Phyllobacterium 99.89 Proteobacteria Alphaproteobacteria trifolii (AY786080)

L3 D. carota Phyllobacterium 100.00 Proteobacteria Alphaproteobacteria brassicacearum (AY785319)

L4 D. carota Pseudomonas 99.68 Proteobacteria Gammaproteobacteria frederiksbergensis (AJ249382)

L6 D. carota Novosphingobium 98.24 Proteobacteria Alphaproteobacteria subterraneum (JRVC01000007)

L11 D. carota Variovorax 99.37 Proteobacteria Betaproteobacteria ginsengisoli (AB245358)

L12 D. carota Sphingopyxis 98.45 Proteobacteria Alphaproteobacteria panaciterrae

(AB245353)

L16 D. carota Pseudomonas mohnii 99.01 Proteobacteria Gammaproteobacteria (FNRV01000001)

Pseudomonas baetica (FM201274)

L17 D. carota Rhizobium altiplani 100.00 Proteobacteria Alphaproteobacteria (LNCD01000038)

Rhizobium cauense (JQ308326)

L18 D. carota Pseudomonas 99.41 Proteobacteria Gammaproteobacteria kilonensis (LHVH01000037)

Pseudomonas corrugate (D84012)

L53 D. carota Sphingomonas 98.56 Proteobacteria Alphaproteobacteria insulae (EF363714)

L54 D. carota Bacillus 99.63 Firmicutes Bacilli paralicheniformis (LBMN01000156)

L55 D. carota Nocardioides albus 99.78 Actinobacteria Actinobacteria (AF004988)

K11 Aster pilosus Sphingopyxis 98.56 Proteobacteria Alphaproteobacteria panaciterrae

(AB245353)

B67 A. pilosus Mycobacterium 99.64 Actinobacteria Actinobacteria frederiksbergense (AJ276274)

K23 A. pilosus Novosphingobium 98.80 Proteobacteria Alphaproteobacteria subterraneum (JRVC01000007)

K24 A. pilosus Pseudoxanthomonas 99.54 Proteobacteria Gammaproteobacteria sacheonensis (EF575564)

K26 A. pilosus Microbacterium 99.88 Actinobacteria Actinobacteria natoriense (AY566291)

K61 A. pilosus Achromobacter 98.79 Proteobacteria Betaproteobacteria marplatensis (EU150134)

K62 A. pilosus Pantoea sp. 98.18 Proteobacteria Gammaproteobacteria (AKIT01000060)

L20 A. pilosus Microbacterium lacus 98.63 Actinobacteria Actinobacteria (AB286030)

L23 A. pilosus Pseudoxanthomonas 100.00 Proteobacteria Gammaproteobacteria indica (jgi.1118276)

B10 Taraxacum Streptomyces 100.00 Actinobacteria Actinobacteria officinale caeruleatus

(KQ948975)

Streptomyces canus (KQ948708)

B12 T. officinale Microbacterium sp. 98.77 Actinobacteria Actinobacteria (JHET01000001)

B13 T. officinale Microbacterium 99.88 Actinobacteria Actinobacteria natoriense (AY566291)

B17 T. officinale Variovorax 99.40 Proteobacteria Betaproteobacteria ginsengisoli (AB245358)

B19 T. officinale Pseudomonas 99.77 Proteobacteria Gammaproteobacteria granadensis (LT629778)

Pseudomonas koreensis (AF468452)

B20 T. officinale Pseudomonas 99.30 Proteobacteria Gammaproteobacteria kilonensis (LHVH01000037)

Pseudomonas corrugate (D84012)

B21 T. officinale Microbacterium 99.88 Actinobacteria Actinobacteria yannicii (FN547412)

B22 T. officinale Sphingomonas 100.00 Proteobacteria Alphaproteobacteria kyeonggiensis (KC252615)

K27 T. officinale Stenotrophomonas sp. 100.00 Proteobacteria Gammaproteobacteria (MUYX01000273)

K30 T. officinale Glutamicibacter 99.53 Actinobacteria Actinobacteria halophytocola (JX993762)

K30II T. officinale Arthrobacter 98.83 Actinobacteria Actinobacteria endophyticus (KM114214)

K33 T. officinale Pseudoxanthomonas 100.00 Proteobacteria Gammaproteobacteria indica (jgi.1118276)

K37 T. officinale Pseudomonas 99.76 Proteobacteria Gammaproteobacteria granadensis (LT629778)

Pseudomonas koreensis (AF468452)

L22 T. officinale Microbacterium 99.51 Actinobacteria Actinobacteria trichothecenolyticum (JYJA01000006)

L25 T. officinale Pseudomonas 98.12 Proteobacteria Gammaproteobacteria paralactis

(KP756923)

L27 T. officinale Pseudomonas putida 99.77 Proteobacteria Gammaproteobacteria (AP013070)

B27 Scorzoneroides Ancylobacter sp. 99.38 Proteobacteria Alphaproteobacteria autumnalis (JNLC01000015)

B27II S. autumnalis Starkeya novella 98.39 Proteobacteria Alphaproteobacteria (CP002026)

B30 S. autumnalis Rhizobium 100.00 Proteobacteria Alphaproteobacteria mesosinicum (DQ100063)

Rhizobium alamii (AM931436)

B31 S. autumnalis Ancylobacter defluvii 98.64 Proteobacteria Alphaproteobacteria (KC243678)

K40 S. autumnalis Sphingobium 99.76 Proteobacteria Alphaproteobacteria yanoikuyae

K44 S. autumnalis Chitinophaga 98.26 Bacteroidetes Sphingobacteriia pinensis (CP001699)

K46 S. autumnalis Mucilaginibacter 97.21 Bacteroidetes Sphingobacteriia lappiensis (jgi.1095764)

K47 S. autumnalis Mycobacterium 99.40 Actinobacteria Actinobacteria frederiksbergense

(AJ276274)

K49 S. autumnalis Pseudoxanthomonas 99.43 Proteobacteria Gammaproteobacteria sacheonensis (EF575564)

K50 S. autumnalis Sphingomonas sp. 98.95 Proteobacteria Alphaproteobacteria (JOOE01000006)

L30 S. autumnalis Ancylobacter defluvii 98.72 Proteobacteria Alphaproteobacteria (KC243678)

L32 S. autumnalis Sphingobium qiguonii 98.00 Proteobacteria Alphaproteobacteria (EU095328)

L33 S. autumnalis Rhizobium 99.87 Proteobacteria Alphaproteobacteria cellulosilyticum (DQ855276)

L34 S. autumnalis Rhizobium 100.00 Proteobacteria Alphaproteobacteria metallidurans (JX678769)

B38 Plantago major Chitinophaga 100.00 Bacteroidetes Sphingobacteriia pinensis (CP001699)

B43 P. major Luteibacter anthropi 98.90 Proteobacteria Gammaproteobacteria (FM212561)

K17 P. major Novosphingobium 98.76 Proteobacteria Alphaproteobacteria subterraneum (JRVC01000007)

K19 P. major Polaromonas sp. 99.77 Proteobacteria Betaproteobacteria (AKIV01000055)

K66 P. major Phyllobacterium 100.00 Proteobacteria Alphaproteobacteria sophorae (KJ685937)

Phyllobacterium brassicacearum (AY785319)

K84 P. major Mycobacterium 99.57 Actinobacteria Actinobacteria fluoranthenivorans (AJ617741)

Mycobacterium frederiksbergense (AJ276274_

K85 P. major Mycobacterium 99.52 Actinobacteria Actinobacteria aichiense (X55598)

L61 P. major Pseudomonas 99.16 Proteobacteria Gammaproteobacteria helmanticensis (HG940537)

B40 Rumex crispus Microbacterium 99.77 Actinobacteria Actinobacteria maritypicum (AJ853910)

B80 R. crispus Glycomyces 100.00 Actinobacteria Actinobacteria lechevalierae (AY462041)

B81 R. crispus Micromonospora 99.89 Actinobacteria Actinobacteria chokoriensis (LT607409)

Micromonospora saelicesensis (AJ783993)

K56 R. crispus Cellulosimicrobium 99.68 Actinobacteria Actinobacteria sp.

K57 R. crispus Luteibacter anthropic 98.69 Proteobacteria Gammaproteobacteria (FM212561)

K58 R. crispus Pseudomonas sp. 99.15 Proteobacteria Gammaproteobacteria

K59 R. crispus Rahnella sp. 100.00 Proteobacteria Gammaproteobacteria (CP002505)

K60 R. crispus Pseudomonas cedrina 99.68 Proteobacteria Gammaproteobacteria

L36 R. crispus Xanthomonas melonis 100.00 Proteobacteria Gammaproteobacteria (Y10756)

Xanthomonas translucens (CAPJ01000550)

L38 R. crispus Yersinia sp. 98.69 Proteobacteria Gammaproteobacteria (CQBW01000017)

L39 R. crispus Ancylobacter defluvii 99.38 Proteobacteria Alphaproteobacteria (KC243678)

L41 R. crispus Pseudomonas cedrina 99.88 Proteobacteria Gammaproteobacteria subsp. fulgida (AJ492830)

Pseudomonas cedrina subsp. cedrina (AF064461)

B41 Prunella vulgaris Rahnella sp. 100.00 Proteobacteria Gammaproteobacteria (CP002505)

B82 P. vulgaris Glycomyces 100.00 Actinobacteria Actinobacteria lechevalierae (AY462041)

K45 P. vulgaris Sphingobium qiguonii 98.01 Proteobacteria Alphaproteobacteria (EU095328)

K48 P. vulgaris Rhizobium 100.00 Proteobacteria Alphaproteobacteria metallidurans (JX678769)

K120 P. vulgaris Agromyces 98.99 Actinobacteria Actinobacteria iriomotensis (AB546308)

L19 P. vulgaris Variovorax 99.60 Proteobacteria Betaproteobacteria boronicumulans (AB300597)

L50 P. vulgaris Bacillus sp. 99.88 Firmicutes Bacilli (CP013984)

n97 P. vulgaris Labrys 100.00 Proteobacteria Alphaproteobacteria methylaminiphilus (AY766152) n102 P. vulgaris Ensifer meliloti 100.00 Proteobacteria Alphaproteobacteria (X67222) n116 P. vulgaris Paenibacillus 99.78 Firmicutes Bacilli glucanolyticus (AB073189)

B42 Verbascum thapsus Pseudomonas cedrina 98.99 Proteobacteria Gammaproteobacteria subsp. fulgida (AJ492830)

B83 V. thapsus Sphingomonas 100.00 Proteobacteria Alphaproteobacteria aquatic (KT309085)

K68 V. thapsus Phyllobacterium 100.00 Proteobacteria Alphaproteobacteria sophorae (KJ685937)

Phyllobacterium brassicacearum (AY785319)

K81 V. thapsus Bacillus simplex 100.00 Firmicutes Bacilli (BCVO01000086)

Brevibacterium frigoritolerans (AM747813)

L9 V. thapsus Microbacterium 99.46 Actinobacteria Actinobacteria trichothecenolyticum 92

(JYJA01000006)

L15 V. thapsus Novosphingobium 98.79 Proteobacteria Alphaproteobacteria subterraneum (JRVC01000007)

6N4 V. thapsus Herbiconiux flava 99.47 Actinobacteria Actinobacteria (AB583921)

B44 Ginkgo biloba Paenibacillus 99.78 Firmicutes Bacilli xylanilyticus (AY427832)

B45 G. biloba Rhodococcus jostii 99.36 Actinobacteria Actinobacteria (FNTL01000001)

B46 G. biloba Achromobacter 99.89 Proteobacteria Betaproteobacteria marplatensis (EU150134)

B47 G. biloba Pseudomonas 99.37 Proteobacteria Gammaproteobacteria koreensis (AF468452)

B48 G. biloba Inquilinus limosus 99.53 Proteobacteria Alphaproteobacteria (AUHM01000026)

B49 G. biloba Achromobacter sp. 100.00 Proteobacteria Betaproteobacteria

B50 G. biloba Pantoea sp. 99.87 Proteobacteria Gammaproteobacteria (AKIT01000060)

B53 G. biloba Luteibacter anthropic 98.26 Proteobacteria Gammaproteobacteria

(FM212561)

K63 G. biloba Labrys 100.00 Proteobacteria Alphaproteobacteria methylaminiphilus (AY766152)

K64 G. biloba Achromobacter 99.79 Proteobacteria Betaproteobacteria marplatensis (EU150134)

K65 G. biloba Pantoea sp. 99.89 Proteobacteria Gammaproteobacteria (AKIT01000060)

L43 G. biloba Achromobacter 100.00 Proteobacteria Betaproteobacteria marplatensis (EU150134)

L44 G. biloba Achromobacter sp. 100.00 Proteobacteria Betaproteobacteria

B55 Erigeron strigosus Rhizobium 99.88 Proteobacteria Alphaproteobacteria mesosinicum (DQ100063)

Rhizobium alamii (AM931436)

B56 E. strigosus Curtobacterium 100.00 Actinobacteria Actinobacteria flaccumfaciens (AJ312209)

B57 E. strigosus Streptomyces 99.17 Actinobacteria Actinobacteria pulveraceus (AJ781377)

B59 E. strigosus Inquilinus limosus 99.75 Proteobacteria Alphaproteobacteria (AUHM01000026)

B60 E. strigosus Pararhizobium 100.00 Proteobacteria Alphaproteobacteria giardinii (ARBG01000149)

B61 E. strigosus Microbacterium 99.17 Actinobacteria Actinobacteria yannicii (FN547412)

Microbacterium paraoxydans (BCRH01000180)

B79 E. strigosus Cellulomonas 99.51 Actinobacteria Actinobacteria humilata (X82449)

B92 E. strigosus Nocardioides 99.39 Actinobacteria Actinobacteria ganghwensis (AY423718)

K67 E. strigosus Mesorhizobium 97.89 Proteobacteria Alphaproteobacteria albiziae (jgi.1055318)

K69 E. strigosus Streptomyces 100.00 Actinobacteria Actinobacteria caeruleatus (KQ948975)

Streptomyces canus (KQ948708)

K70 E. strigosus Phyllobacterium 100.00 Proteobacteria Alphaproteobacteria sophorae (KJ685937)

Phyllobacterium brassicacearum (AY785319)

K73 E. strigosus Streptomyces 100.00 Actinobacteria Actinobacteria xanthophaeus (JOFT01000080)

Streptomyces spororaveus (AJ781370)

L46 E. strigosus Leifsonia soli 99.23 Actinobacteria Actinobacteria (EU912483)

L48 E. strigosus Aminobacter 99.67 Proteobacteria Alphaproteobacteria anthyllidis (FR869633)

L56 E. strigosus Phyllobacterium 99.89 Proteobacteria Alphaproteobacteria trifolii (AY786080)

B64 Sonchus arvensis Bacillus cereus 99.80 Firmicutes Bacilli (AE016877)

K75 S. arvensis Bacillus mobilis 100.00 Firmicutes Bacilli (KJ812449)

K76 S. arvensis Bacillus aryabhattai 100.00 Firmicutes Bacilli (EF114313)

K86 S. arvensis Paenibacillus catalpa 99.76 Firmicutes Bacilli (HQ657320)

Paenibacillus glycanilyticus (AB042938)

L49 S. arvensis Bacillus niacin 99.18 Firmicutes Bacilli (AB021194)

B65 Vicia cracca Bacillus simplex 99.79 Firmicutes Bacilli (BCVO01000086)

K78 V. cracca Bacillus sp. 100.00 Firmicutes Bacilli (CP013984)

K79 V. cracca Microbacterium 99.13 Actinobacteria Actinobacteria yannicii (FN547412)

K80 V. cracca Rummeliibacillus 100.00 Firmicutes Bacilli stabekisii (DQ870754)

L51 V. cracca Mycobacterium 99.47 Actinobacteria Actinobacteria frederiksbergense (AJ276274)

L52 V. cracca Agromyces 99.37 Actinobacteria Actinobacteria iriomotensis (AB546308)

B102 Tussilago farfara Xanthomonas sp. 99.89 Proteobacteria Gammaproteobacteria

B108 T. farfara Variovorax 99.16 Proteobacteria Betaproteobacteria ginsengisoli (AB245358)

B110 T. farfara Cupriavidus 100.00 Proteobacteria Betaproteobacteria campinensis (AF312020)

B111 T. farfara Pseudomonas sp. 99.47 Proteobacteria Gammaproteobacteria

B116 T. farfara Herbaspirillum 99.04 Proteobacteria Betaproteobacteria hiltneri (CP011409)

K91 T. farfara Xanthomonas sp. 99.90 Proteobacteria Gammaproteobacteria

K92 T. farfara Pseudomonas baetica 99.48 Proteobacteria Gammaproteobacteria (FM201274)

K93 T. farfara Pseudomonas 99.58 Proteobacteria Gammaproteobacteria nitritireducens (HM246143)

K94 T. farfara Lelliottia sp. 99.89 Proteobacteria Gammaproteobacteria (CP000653)

K96 T. farfara Pseudomonas 99.89 Proteobacteria Gammaproteobacteria granadensis (LT629778)

K97 T. farfara Pseudomonas 99.37 Proteobacteria Gammaproteobacteria frederiksbergensis (AJ249382)

K98 T. farfara Pseudomonas mohnii 99.79 Proteobacteria Gammaproteobacteria (FNRV01000001)

L70 T. farfara Pseudomonas baetica 98.95 Proteobacteria Gammaproteobacteria

(FM201274)

L74 T. farfara Cupriavidus pauculus 98.64 Proteobacteria Betaproteobacteria (AF085226)

L75 T. farfara Cupriavidus 99.37 Proteobacteria Betaproteobacteria yeoncheonensis (KF915797)

L77 T. farfara Pseudomonas sp. 99.06 Proteobacteria Gammaproteobacteria

L79 T. farfara Bacillus luciferensis 98.76 Firmicutes Bacilli (AJ419629)

L80 T. farfara Xanthomonas sp. 100.00 Proteobacteria Gammaproteobacteria

L81 T. farfara Rhizobium sp. 100.00 Proteobacteria Alphaproteobacteria (CP002249)

K110 Solanum dulcamara Bacillus siamensis 99.79 Firmicutes Bacilli (AJVF01000043)

K111 S. dulcamara Bacillus siamensis 98.72 Firmicutes Bacilli (AJVF01000043)

K112 S. dulcamara Pseudomonas sp. 99.16 Proteobacteria Gammaproteobacteria

K114 S. dulcamara Serratia myotis 99.78 Proteobacteria Gammaproteobacteria (KJ739884)

K115 S. dulcamara Pseudomonas 99.68 Proteobacteria Gammaproteobacteria helmanticensis (HG940537)

K116 S. dulcamara Bacillus toyonensis 99.58 Firmicutes Bacilli (CP006863)

K117 S. dulcamara Bacillus sp. 99.79 Firmicutes Bacilli

L62 S. dulcamara Bacillus mobilis 100.00 Firmicutes Bacilli (KJ812449)

KB1 Euphrasia Rhodococcus 100.00 Actinobacteria Actinobacteria officinalis qingshengii (LRRJ01000016)

KB2 E. officinalis Mycobacterium 98.86 Actinobacteria Actinobacteria hodleri (X93184)

KB3 E. officinalis Microbacterium 100.00 Actinobacteria Actinobacteria phyllosphaerae (AJ277840)

KB4 E. officinalis Pseudomonas 99.89 Proteobacteria Gammaproteobacteria extremaustralis (AHIP01000073)

NB1 E. officinalis Caulobacter henricii 99.34 Proteobacteria Alphaproteobacteria (AJ227758)

NB2 E. officinalis Methylobacterium 99.76 Proteobacteria Alphaproteobacteria bullatum (GU983169)

NB4 E. officinalis Mycobacterium 99.20 Actinobacteria Actinobacteria helvum (KM216314)

100

NB5 E. officinalis Microlunatus 99.78 Actinobacteria Actinobacteria aurantiacus (EF601828)

NB6 E. officinalis Variovorax 99.37 Proteobacteria Betaproteobacteria ginsengisoli (AB245358)

NB7 E. officinalis Pseudomonas 99.68 Proteobacteria Gammaproteobacteria extremaustralis (AHIP01000073)

NB11 E. officinalis Bradyrhizobium sp. 98.24 Proteobacteria Alphaproteobacteria (BADD01000237) n1 Brassica napus Bacillus sp. 100.00 Firmicutes Bacilli (CP013984) n40 B. napus Rhizobium lusitanum 99.22 Proteobacteria Alphaproteobacteria (jgi.1052907) n41 B. napus Variovorax 99.89 Proteobacteria Betaproteobacteria paradoxus (BCUT01000013) n42 B. napus Microbacterium 99.76 Actinobacteria Actinobacteria xylanilyticum (AJ853908) n84 B. napus Bacillus sp. 99.64 Firmicutes Bacilli (CP013984) n85 B. napus Staphylococcus 100.00 Firmicutes Bacilli

101

warneri (L37603) n86 B. napus Variovorax 100.00 Proteobacteria Betaproteobacteria paradoxus (BCUT01000013)

86 B. napus Pseudarthrobacter 100.00 Actinobacteria Actinobacteria sp.

92 B. napus Bacillus 100.00 Firmicutes Bacilli zhangzhouensis (JOTP01000061)

94 B. napus Flavobacterium 98.51 Bacteroidetes Flavobacteria piscis (LVEN01000016)

162 B. napus Arthrobacter 100.00 Actinobacteria Actinobacteria ginsengisoli (KF212463)

163 B. napus Rhodococcus 99.88 Actinobacteria Actinobacteria corynebacterioides (AF430066)

165 B. napus Pseudomonas 98.97 Proteobacteria Gammaproteobacteria chlororaphis subsp. chlororaphis (BCZX01000031)

Pseudomonas brassicacearum subsp. neoaurantiaca

102

(EU391388)

167 B. napus Flavobacterium 99.21 Bacteroidetes Flavobacteria frigidimaris (jgi.1107687)

168 B. napus Bacillus aryabhattai 100.00 Firmicutes Bacilli (EF114313)

169 B. napus Variovorax 100.00 Proteobacteria Betaproteobacteria paradoxus (BCUT01000013)

171 B. napus Staphylococcus 100.00 Firmicutes Bacilli warneri (L37603)

172 B. napus Bacillus drentensis 100.00 Firmicutes Bacilli (AJ542506)

247 B. napus Paenibacillus 99.89 Firmicutes Bacilli liaoningensis (KF997862)

Paenibacillus algorifonticola (GQ383922)

248 B. napus Pseudomonas 98.96 Proteobacteria Gammaproteobacteria yamanorum

249 B. napus Flavobacterium 99.74 Bacteroidetes Flavobacteria piscis (LVEN01000016)

103

250 B. napus Pedobacter borealis 99.65 Bacteroidetes Sphingobacteriia (JAUG01000180)

251 B. napus Polaromonas aquatic 99.20 Proteobacteria Betaproteobacteria (AM039830)

253 B. napus Flavobacterium 99.21 Bacteroidetes Flavobacteria aquidurense (jgi.1107986) n10 Medicago sativa Variovorax 100.00 Proteobacteria Betaproteobacteria ginsengisoli (AB245358) n11 M. sativa Ensifer adhaerens 100.00 Proteobacteria Alphaproteobacteria (JNAE01000171) n12 M. sativa Phyllobacterium 100.00 Proteobacteria Alphaproteobacteria ifriqiyense (AY785325) n15 M. sativa Rhizobium sp. 100.00 Proteobacteria Alphaproteobacteria (CP002249) n17 M. sativa Rhodococcus 100.00 Actinobacteria Actinobacteria globerulus (BCWX01000023) n19 M. sativa Novosphingobium 99.50 Proteobacteria Alphaproteobacteria lindaniclasticum (ATHL01000125) n20 M. sativa Microbacterium 99.88 Actinobacteria Actinobacteria natoriense 104

(AY566291) n23 M. sativa Microbacterium 99.65 Actinobacteria Actinobacteria maritypicum (AJ853910)

Microbacterium paraoxydans (BCRH01000180) n24 M. sativa Pseudoxanthomonas 100.00 Proteobacteria Gammaproteobacteria indica (jgi.1118276) n54 M. sativa Phyllobacterium loti 100.00 Proteobacteria Alphaproteobacteria (KC577468) n57 M. sativa Devosia neptuniae 99.75 Proteobacteria Alphaproteobacteria (AF469072) n58 M. sativa Caulobacter 99.34 Proteobacteria Alphaproteobacteria rhizosphaerae (KX792139) n59 M. sativa Phyllobacterium 99.78 Proteobacteria Alphaproteobacteria trifolii (AY786080) n61 M. sativa Pseudoxanthomonas 99.57 Proteobacteria Gammaproteobacteria japonensis (AB008507) n62 M. sativa Rhodococcus 100.00 Actinobacteria Actinobacteria pedocola (KT301938)

105

n63 M. sativa Pseudomonas 99.79 Proteobacteria Gammaproteobacteria frederiksbergensis (AJ249382) n64 M. sativa Rhizobium altiplani 100.00 Proteobacteria Alphaproteobacteria (LNCD01000038) n65 M. sativa Pseudoxanthomonas 99.57 Proteobacteria Gammaproteobacteria sacheonensis (EF575564) n67 M. sativa Mesorhizobium 100.00 Proteobacteria Alphaproteobacteria japonicum (BA000012)

Mesorhizobium erdmanii (AXAE01000021) n71 M. sativa Paenibacillus 100.00 Firmicutes Bacilli provencensis (EF212893) n94 M. sativa Aminobacter sp. 97.82 Proteobacteria Alphaproteobacteria (KB890024) n95 M. sativa Terrabacter 99.89 Actinobacteria Actinobacteria ginsenosidimutans (EU332827) n96 M. sativa Variovorax 99.54 Proteobacteria Betaproteobacteria boronicumulans (AB300597)

106

n99 M. sativa Bacillus sp. 100.00 Firmicutes Bacilli (CP013984) n100 M. sativa Ensifer meliloti 100.00 Proteobacteria Alphaproteobacteria (X67222) n103 M. sativa Microbacterium 99.28 Actinobacteria Actinobacteria ginsengiterrae (EU873314) n104 M. sativa Pseudomonas 99.68 Proteobacteria Gammaproteobacteria brassicacearum subsp. neoaurantiaca (EU391388) n105 M. sativa Pseudomonas 99.47 Proteobacteria Gammaproteobacteria caspiana (LOHF01000033) n106 M. sativa Agromyces cerinus 99.89 Actinobacteria Actinobacteria subsp. cerinus (jgi.1107671)

Agromyces cerinus subsp. nitratus (AY277619) n108 M. sativa Pseudomonas 100.00 Proteobacteria Gammaproteobacteria koreensis (AF468452)

43 M. sativa Pseudomonas 99.46 Proteobacteria Gammaproteobacteria caspiana

107

(LOHF01000033)

44 M. sativa Variovorax 99.58 Proteobacteria Betaproteobacteria boronicumulans (AB300597)

46 M. sativa Bacillus sp. 99.69 Firmicutes Bacilli

47 M. sativa Pseudomonas 99.58 Proteobacteria Gammaproteobacteria prosekii (LT629762)

48 M. sativa Pseudomonas moorei 99.40 Proteobacteria Gammaproteobacteria (AM293566)

153 M. sativa Pseudomonas 99.43 Proteobacteria Gammaproteobacteria kilonensis (LHVH01000037)

Pseudomonas corrugate (D84012)

154 M. sativa Bacillus megaterium 100.00 Firmicutes Bacilli (JJMH01000057)

155 M. sativa Variovorax 100.00 Proteobacteria Betaproteobacteria paradoxus (BCUT01000013)

157 M. sativa Pseudomonas lini 99.76 Proteobacteria Gammaproteobacteria (AY035996)

159 M. sativa Arthrobacter 100.00 Actinobacteria Actinobacteria humicola

108

(AB279890)

Arthrobacter pascens (X80740)

160 M. sativa Microbacterium 99.29 Actinobacteria Actinobacteria ginsengiterrae (EU873314)

161 M. sativa Streptomyces 100.00 Actinobacteria Actinobacteria zhihengii (KU936048)

244 M. sativa Arthrobacter sp. 100.00 Actinobacteria Actinobacteria n28 Miscanthus × Staphylococcus 99.79 Firmicutes Bacilli giganteus pasteuri (AF041361) n29 M. giganteus Staphylococcus 100.00 Firmicutes Bacilli warneri (L37603) n31 M. giganteus Flavobacterium 99.30 Bacteroidetes Flavobacteria hercynium (MUGW01000033)

Flavobacterium saccharophilum (jgi.1107675) n34 M. giganteus Herbaspirillum 99.19 Proteobacteria Betaproteobacteria hiltneri (CP011409) n72 M. giganteus Chryseobacterium 98.09 Bacteroidetes Flavobacteria ginsenosidimutans

109

(GU138380) n73 M. giganteus Microbacterium 99.67 Actinobacteria Actinobacteria laevaniformans (Y17234) n74 M. giganteus Rhizobium alamii 99.89 Proteobacteria Alphaproteobacteria (AM931436) n81 M. giganteus Flavobacterium 98.80 Bacteroidetes Flavobacteria saccharophilum (jgi.1107675) n112 M. giganteus Phyllobacterium sp. 100.00 Proteobacteria Alphaproteobacteria n113 M. giganteus Rothia terrae 99.89 Actinobacteria Actinobacteria (DQ822568) n114 M. giganteus Rhizobium 99.62 Proteobacteria Alphaproteobacteria oryziradicis (KX129901)

26 M. giganteu Chryseobacterium 98.96 Bacteroidetes Flavobacteria luteum (JPRO01000001)

27 M. giganteus Flavobacterium 98.70 Bacteroidetes Flavobacteria pectinovorum (jgi.1107681)

28 M. giganteus Pseudomonas 99.27 Proteobacteria Gammaproteobacteria chlororaphis subsp. aurantiaca

110

(DQ682655)

31 M. giganteus Flavobacterium 99.35 Bacteroidetes Flavobacteria saccharophilum (jgi.1107675)

32 M. giganteus Pseudomonas lini 99.79 Proteobacteria Gammaproteobacteria (AY035996)

33 M. giganteus Pseudomonas 99.79 Proteobacteria Gammaproteobacteria brassicacearum subsp. neoaurantiaca (EU391388)

34 M. giganteus Pseudomonas mohnii 99.57 Proteobacteria Gammaproteobacteria (FNRV01000001)

36 M. giganteus Pseudomonas 99.36 Proteobacteria Gammaproteobacteria frederiksbergensis (AJ249382)

37 M. giganteus Pseudomonas moorei 99.35 Proteobacteria Gammaproteobacteria (AM293566)

39 M. giganteus Variovorax 100.00 Proteobacteria Betaproteobacteria paradoxus (BCUT01000013)

41 M. giganteus Pseudomonas sp. 99.24 Proteobacteria Gammaproteobacteria

111 M. giganteus Pedobacter alluvionis 99.53 Bacteroidetes Sphingobacteriia (jgi.1096603)

111

112 M. giganteus Paenibacillus vini 99.43 Firmicutes Bacilli (KJ005124)

113 M. giganteus Bacillus 100.00 Firmicutes Bacilli zhangzhouensis (JOTP01000061)

115 M. giganteus Pseudomonas 100.00 Proteobacteria Gammaproteobacteria extremaustralis (AHIP01000073)

118 M. giganteus Acidovorax wautersii 100.00 Proteobacteria Betaproteobacteria (jgi.1068022)

119 M. giganteus Pseudomonas 100.00 Proteobacteria Gammaproteobacteria donghuensis (AJJP01000212)

120 M. giganteus Arthrobacter oryzae 99.76 Actinobacteria Actinobacteria (CLG_48533)

200 M. giganteus Microbacterium 99.15 Actinobacteria Actinobacteria oxydans (Y17227)

202 M. giganteus Flavobacterium 98.82 Bacteroidetes Flavobacteria tructae (MUHH01000012)

Flavobacterium spartansii (MUHG01000041)

203 M. giganteus Clavibacter 99.89 Actinobacteria Actinobacteria

112

michiganensis

208 M. giganteus Microbacterium 100.00 Actinobacteria Actinobacteria hatanonis (AB274908)

209 M. giganteus Arthrobacter 99.14 Actinobacteria Actinobacteria humicola (AB279890) n37 Phalaris Pararhizobium 100.00 Proteobacteria Alphaproteobacteria arundinacea herbae (GU565534) n38 P. arundinacea Pseudomonas 99.79 Proteobacteria Gammaproteobacteria koreensis (AF468452) n43 P. arundinacea Xanthomonas melonis 99.89 Proteobacteria Gammaproteobacteria (Y10756)

Xanthomonas translucens (CAPJ01000550) n45 P. arundinacea Streptomyces 100.00 Actinobacteria Actinobacteria lavendulae subsp. lavendulae (JOEW01000098) n47 P. arundinacea Pseudomonas baetica 99.27 Proteobacteria Gammaproteobacteria (FM201274) n48 P. arundinacea Variovorax 99.48 Proteobacteria Betaproteobacteria

113

boronicumulans (AB300597) n49 P. arundinacea Pseudomonas 100.00 Proteobacteria Gammaproteobacteria brassicacearum subsp. neoaurantiaca (EU391388) n51 P. arundinacea Pseudomonas 99.77 Proteobacteria Gammaproteobacteria granadensis (LT629778)

Pseudomonas koreensis (AF468452) n87 P. arundinacea Pseudomonas 99.76 Proteobacteria Gammaproteobacteria kilonensis (LHVH01000037)

Pseudomonas corrugate (D84012) n88 P. arundinacea Pseudomonas 98.23 Proteobacteria Gammaproteobacteria chlororaphis subsp. aureofaciens (BBQB01000031) n92 P. arundinacea Pseudomonas 99.41 Proteobacteria Gammaproteobacteria turukhanskensis (KP306892)

56 P. arundinacea Pseudomonas lini 99.76 Proteobacteria Gammaproteobacteria

114

(AY035996)

57 P. arundinacea Pseudomonas 99.76 Proteobacteria Gammaproteobacteria kilonensis (LHVH01000037)

Pseudomonas corrugate (D84012)

59 P. arundinacea Paenibacillus 99.75 Firmicutes Bacilli glucanolyticus (AB073189)

61 P. arundinacea Pseudomonas 100.00 Proteobacteria Gammaproteobacteria migulae (AF074383)

62 P. arundinacea Flavobacterium 99.29 Bacteroidetes Flavobacteria hercynium (MUGW01000033)

Flavobacterium saccharophilum (jgi.1107675)

99 P. arundinacea Buttiauxella gaviniae 99.16 Proteobacteria Gammaproteobacteria (LXEP01000074)

100 P. arundinacea Pseudomonas sp. 99.88 Proteobacteria Gammaproteobacteria

101 P. arundinacea Streptomyces 99.76 Actinobacteria Actinobacteria gulbargensis (DQ317411)

115

102 P. arundinacea Ensifer morelensis 100.00 Proteobacteria Alphaproteobacteria (AY024335)

143 P. arundinacea Bacillus mobilis 100.00 Firmicutes Bacilli (KJ812449)

144 P. arundinacea Arthrobacter 99.57 Actinobacteria Actinobacteria globiformis (BAEG01000072)

145 P. arundinacea Arthrobacter 100.00 Actinobacteria Actinobacteria ginsengisoli (KF212463)

148 P. arundinacea Pseudomonas 99.47 Proteobacteria Gammaproteobacteria frederiksbergensis (AJ249382)

229 P. arundinacea Pseudomonas 99.77 Proteobacteria Gammaproteobacteria caspiana (LOHF01000033)

230 P. arundinacea Novosphingobium 99.26 Proteobacteria Alphaproteobacteria rhizosphaerae (KM365125)

Novosphingobium capsulatum (D16147)

232 P. arundinacea Flavobacterium sp. 97.87 Bacteroidetes Flavobacteria (AM110992)

233 P. arundinacea Nocardia coubleae 99.42 Actinobacteria Actinobacteria

116

(BDBD01000030)

234 P. arundinacea Bacillus sp. 100.00 Firmicutes Bacilli

236 P. arundinacea Lysobacter caeni 97.56 Proteobacteria Gammaproteobacteria (KJ008918)

238 P. arundinacea Agromyces salentinus 99.48 Actinobacteria Actinobacteria (AY507129)

240 P. arundinacea Pseudoxanthomonas 98.28 Proteobacteria Gammaproteobacteria suwonensis (AY927994)

242 P. arundinacea Flavobacterium sp. 99.65 Bacteroidetes Flavobacteria (AKJZ01000102)

1B1 Glechoma Jatrophihabitans 98.45 Actinobacteria Actinobacteria hederacea huperziae (KR184574)

1B2 G. hederacea Mycobacterium sp. 99.44 Actinobacteria Actinobacteria

1K1 G. hederacea Sphingomonas 98.63 Proteobacteria Alphaproteobacteria crusticola (KT346426)

1K2 G. hederacea Acinetobacter sp. 100.00 Proteobacteria Gammaproteobacteria

1K3 G. hederacea Plantibacter flavus 99.03 Actinobacteria Actinobacteria (jgi.1118344)

1N1 G. hederacea Pseudomonas moorei 98.93 Proteobacteria Gammaproteobacteria (AM293566)

117

1N2 G. hederacea Microbacterium 99.89 Actinobacteria Actinobacteria hatanonis (AB274908)

1N3 G. hederacea Microbacterium sp. 99.23 Actinobacteria Actinobacteria (CBVQ010000169)

1N4 G. hederacea Herbiconiux flava 98.83 Actinobacteria Actinobacteria (AB583921)

3K1 Campanula Pseudomonas sp. 99.05 Proteobacteria Gammaproteobacteria rotundifolia

3K2 C. rotundifolia Herbiconiux flava 98.83 Actinobacteria Actinobacteria (AB583921)

3K3 C. rotundifolia Pseudomonas baetica 99.01 Proteobacteria Gammaproteobacteria (FM201274)

3K4 C. rotundifolia Pseudomonas 99.79 Proteobacteria Gammaproteobacteria mandelii (AF058286)

3K5 C. rotundifolia Pseudomonas moorei 98.93 Proteobacteria Gammaproteobacteria (AM293566)

3K6 C. rotundifolia Pseudomonas baetica 99.47 Proteobacteria Gammaproteobacteria (FM201274)

3N2 C. rotundifolia Rhizobium 99.89 Proteobacteria Alphaproteobacteria radiobacter (AJ389904)

3N3 C. rotundifolia Neorhizobium sp. 99.55 Proteobacteria Alphaproteobacteria

118

3N4 C. rotundifolia Rhizobium sp. 99.44 Proteobacteria Alphaproteobacteria

3N5 C. rotundifolia Pararhizobium 99.43 Proteobacteria Alphaproteobacteria herbae (GU565534)

4K1 Lactuca canadensis Agromyces 98.94 Actinobacteria Actinobacteria iriomotensis (AB546308)

4K2 L. canadensis Bosea lupine 99.56 Proteobacteria Alphaproteobacteria (FR774992)

4L1 L. canadensis Microbacterium 99.89 Actinobacteria Actinobacteria hatanonis (AB274908)

4L2 L. canadensis Caulobacter 99.66 Proteobacteria Alphaproteobacteria vibrioides (AJ009957)

4N1 L. canadensis Sphingomonas sp. 99.78 Proteobacteria Alphaproteobacteria (JOOE01000006)

6N3 L. canadensis Herbiconiux flava 99.68 Actinobacteria Actinobacteria (AB583921)

5K1 Myosotis arvensis Pseudomonas sp. 99.16 Proteobacteria Gammaproteobacteria

5K3 M. arvensis Cupriavidus 99.51 Proteobacteria Betaproteobacteria campinensis (AF312020)

5K4 M. arvensis Frigoribacterium 99.46 Actinobacteria Actinobacteria

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faeni (Y18807)

5K5 M. arvensis Variovorax 99.37 Proteobacteria Betaproteobacteria ginsengisoli (AB245358)

5K6 M. arvensis Agromyces ulmi 98.50 Actinobacteria Actinobacteria (AY427830)

5L1 M. arvensis Pseudomonas 99.27 Proteobacteria Gammaproteobacteria helmanticensis (HG940537)

5N1 M. arvensis Microbacterium 99.89 Actinobacteria Actinobacteria proteolyticum (KM359785)

5N2 M. arvensis Caulobacter 98.48 Proteobacteria Alphaproteobacteria vibrioides (AJ009957)

5N4 M. arvensis Bacillus 98.09 Firmicutes Bacilli oceanisediminis (GQ292772)

5N5 M. arvensis Microbacterium 99.03 Actinobacteria Actinobacteria yannicii (FN547412)

5N6 M. arvensis Caulobacter 99.78 Proteobacteria Alphaproteobacteria vibrioides (AJ009957)

5N7 M. arvensis Microbacterium sp. 99.57 Actinobacteria Actinobacteria

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(CBVQ010000169)

6K6 Chrysanthemum Sphingomonas 98.89 Proteobacteria Alphaproteobacteria leucanthemum crusticola (KT346426)

6K7 C. leucanthemum Roseomonas aquatic 96.55 Proteobacteria Alphaproteobacteria (AM231587)

6K8 C. leucanthemum Microbacterium 99.03 Actinobacteria Actinobacteria proteolyticum (KM359785)

6K10 C. leucanthemum Pseudoxanthomonas 99.14 Proteobacteria Gammaproteobacteria spadix (AM418384)

6K13 C. leucanthemum Ensifer sesbaniae 99.78 Proteobacteria Alphaproteobacteria (JF834143)

6K14 C. leucanthemum Herbiconiux 99.35 Actinobacteria Actinobacteria moechotypicola (FJ828659)

6K17 C. leucanthemum Staphylococcus 99.79 Firmicutes Bacilli warneri (L37603)

6L1 C. leucanthemum Herbiconiux flava 99.85 Actinobacteria Actinobacteria (AB583921)

6L5 C. leucanthemum Microbacterium 98.92 Actinobacteria Actinobacteria trichothecenolyticum (JYJA01000006)

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6N2 C. leucanthemum Herbiconiux solani 99.05 Actinobacteria Actinobacteria (BCST01000017)

6N2II C. leucanthemum Schumannella luteola 97.42 Actinobacteria Actinobacteria (AB362159)

6N5 C. leucanthemum Microbacterium 100.00 Actinobacteria Actinobacteria hatanonis (AB274908)

6N7 C. leucanthemum Herbiconiux flava 99.04 Actinobacteria Actinobacteria (AB583921)

6N8 C. leucanthemum Mycobacterium 98.61 Actinobacteria Actinobacteria hodleri

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Connecting text

Chapter 3 described the isolation and identification of rhizobacteria that associate with 6 crops and 20 wild plants in the Saint-Anne-de-Bellevue, Quebec area. It showed that the tested plants harbored a great variety of ectophytic and endophytic bacteria. Each plant species was associated with a specific microbial community. It was thus concluded that the crops and wild plants of Québec, Canada represent a rich reservoir of versatile rhizobacterial strains that is promising for exploration with the goal of identifying potential plant growth-promoting rhizobacteria (PGPR). This led us to hypothesize that there are potential PGPR from this pool that could promote the growth and stress resistance of Arabidopsis thaliana, a model plant, as well as the crop plants, canola (Brassica napus) and maize (Zea mays). In Chapter 4, we screened for potential PGPR that cause growth promotion and salt stress alleviation in A. thaliana. The chapter will be reformatted and submitted for publications as a pair of papers in Frontiers in Microbiology and Soil Biology and Biochemistry.

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Chapter 4 Inoculation of Arabidopsis thaliana by novel rhizobacterial isolates improves plant growth under normal and stress conditions

4.1 Abstract Plant growth promoting rhizobacteria (PGPR) are a functionally diverse group of microbes having immense potential as biostimulants and biopesticides. Their exploitation in agro-ecosystems as an eco-friendly and cost-effective alternative to traditional chemical inputs may positively affect agricultural productivity and environmental sustainability. The present study describes the detailed polyphasic characterization of PGPR, from a range of origins, having plant growth promoting potential under controlled conditions. A total of 98 isolates (ectophytic or endophytic) from various crop and wild plants were screend, out of which four endophytes (n, L, K and Y) from Phalaris arundinacea, Solanum dulcamara, Scorzoneroides autumnalis, and Glycine max, respectively, were selected in vitro for their growth stimulating effects on Arabidopsis col-0 seedlings with regard to leaf surface area and shoot fresh weight. All the strains were characterized using classical methods of bacterial identification and using biochemical test kits (API20E, API20NE, API ZYM, and API 50CH), revealing the metabolic versatility of the four strains. Various PGP traits in selected strains were also determined to evaluate the involvement of the traits in PGP performance. All rhizobacterial isolates were positive for ACCD and IAA production and phosphorous solubilization. Their amounts of soluble phosphate in liquid medium were 341, 69, 63 and 38 g L-1, for n, L, Y and K, respectively. PCR analysis confirmed the presence of the nifH gene in strains n, L and Y showing their N2-fixation potential. The 16S rRNA gene sequencing and phylogenetic diversity of the strains were analyzed and indicated that these isolates belong to the genera Pseudomonas, Bacillus, Mucilaginibacter and Rhizobium. The potential of the strains as biocontrol agents was also tested. In vitro dual culture methods and bacterial infestation in planta demonstrated that strains n and L excreted material antagonistic to Pseudomonas syringae pv tomato DC3000 and Botrytis cinerea 191, and provided protection to Arabidopsis plants against both phytopathogens.

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Strains were then further tested for their effects on abiotic stress alleviation and gene expression in Arabidopsis under Petri plate or pot conditions. Results from Petri-dish assays indicated strains L, K and Y alleviated salt stress in Arabidopsis seedlings, while strains K and Y conferred increases in fresh weight and leaf area under osmotic stress. Results from subsequent in vivo trials indicated all the isolates, especially strains L, K and Y, distinctly increased Arabidopsis growth under both normal and high salinity conditions, as compared to control plants. At the molecular level, short-term treatment revealed rapid changes in transcript levels of some genes annotated to stress-response and hormone metabolism in plants. The expression of several of these genes remained stable in shoots over weeks. Particularly, the expression of stress responsive genes in Arabidopsis showed an up-regulation under salt stressed conditions. A study on spatial distribution of the four strains, using either conventional Petri-plate counts or GFP-tagged bacteria, indicated that all the strains were able to colonize the endosphere of Arabidopsis tissue. Over all, the study revealed that the four selected rhizobacteria are good candidates to be explored as biofertilizers, among which, strain L showed a marked plant growth-promoting attributes, and the potential to be developed as functional biostimulants and biopesticides for sustainable agriculture. Moreover, the study is the first report of a Scorzoneroides autumnalis (fall dandelion) and Solanum dulcamara (bittersweet) associated endophytes with PGP effects.

4.2 Introduction The concept of the rhizosphere was first coined by pioneer German agronomist and plant physiologist Lorenz Hiltner in 1904 (Hartmann et al., 2008). After more than a half-century of relative dormancy, importance of the rhizosphere began to attract attention and appreciation, and now has become one of the most intensely investigated areas of plant science (Brink, 2016). Many studies have reported that microbes associated with root systems are generally abundant and manifest beneficial activity as plant growth promoting rhizobacteria (PGPR) by stimulating plant growth, reducing pathogensis, and allievating abiotic stresses, through a set of underlying mechanisms; this understanding has been widely examined (Bacon and White, 2016). In order to provide adequate food and nutrition worldwide, agricultural production faces challenges, in which chemical fertilizers as well as pesticides have become one of the limiting factors, due to their increasing costs, limited availability and negative impact on agro-ecosystems (Savci, 2012). 125

Moreover, increasing desire for food free of chemical residues has boosted demand for biofertilizers and biopesticides as a sustainable option for improving plant growth and yields in an environmentally friendly manner (Vessey, 2003). Use of microbial consortia for reducing chemical inputs without compromising yield is now an important feature of research in agriculture, microbiology and biotechnology (Minorsky, 2008). Thus, the screening and identification of PGPRs has gained considerable attention. Strains of root associated bacterial genera including Azospirillum, Arthrobacter, Azotobacter, Bacillus, Burkholderia, Erwinia, Enterobacter, Klebsiella, Paenibacillus, Pantoea, Pseudomonas, Serratia, and Xanthomonas are among the main rhizobacteria investigated for promotion of plant growth (Bhattacharyya and Jha, 2012; Kasa et al., 2015; Xing et al., 2016; Li et al., 2017); some of these have already been deployed as biofertilizers (De Souza et al., 2015). The fossil record shows that mycorrhizal interactions with plants have occurred for more than 400 million years (Hata et al., 2010); this relationship has been studied extensively and molecular biology and ‘omics’ have provided approaches into the detailed mechanisms regarding the beneficial physiological and molecular interactions (Bonfante and Genre, 2010). Likewise, interactions between plants and nitrogen-fixing bacteria, especially legumes and rhizobia have been very well investigated (Maróti and Kondorosi, 2014; Westhoek et al., 2017). Further, various beneficial interactions between plants and PGPR (those that colonize either the rhizosphere, rhizoplane or are within plant root tissues) are involved in a wide range of PGP activities (Anwar et al., 2016; Bulgarelli et al., 2013), such as nutrient solubilization, N2 fixation, production of siderophores, volatile compounds and plant-like hormones, induction of systemic resistance, as well as direct biocontrol of disease organisms (Bhattacharyya and Jha, 2012; Vejan et al., 2016). Recently, some additional PGP traits have been discovered, such as bacteriocin production, production of microbe-to-plant signals (Subramanian and Smith, 2015) and sulfur deficienciy alleviation (Meldau et al., 2012). Plants are multi-cellular organism that cope with various environmental stresses, the most common being soil salinity, cold temperature and drought, all of which can elicit some common gene responses related to nearly every aspect of plant morphology, physiology and metabolism (Abreu et al., 2013; Zhu, 2001), leading to inhibition of seed germination, seedling growth, flowering and seed set (Sairam and Tyagi, 2004). Soil salinity adversely affects plant growth via both osmotic and ionic stresses and has become a major limiting factor in agricultural production

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worldwide, leading to as much as $27 billion US loss per year (Qadir et al., 2014). Mechanistically, high salinity results in high concentrations of sodium (Na+) and chlorine (Cl-) ions and low potassium (K+) levels, after prolonged exposure (Munns, 2002; Xiong et al., 2002). Salinity stress also disrupts the cellular osmotic balance by lowering the water potential inside cells in a similar fashion to all three abovementioned abiotic stresses (Krasensky and Jonak, 2012). Stressfully high salt conditions subsequently induce oxidative stresses by generating reactive oxygen species (ROS; Chawla et al., 2013; Pang and Wang, 2012) within cells, resulting in oxidative damage of membrane lipids, proteins and nucleic acids (Gill and Tuteja, 2010). To cope with salt stress, stressed plants tend to activate various mechanisms through conserved signal transduction pathways (Xiong et al., 2002), resulting in the production and accumulation of diverse functional components such as osmolytes (i.e., proline and glycine betaine; Qureshi et al., 2013) and non-enzymatic (i.e., phenolics, flavonoids, and glutathione) and enzymatic antioxidants (i.e., peroxidase, catalase, as well as the enzymes involved in ascorbate-glutathione cycle) (Karuppanapandian et al., 2011; Talbi et al., 2015), all of which mitigate the oxidative damage caused by high salinity (Xiong et al, 2002; Zhu, 2002). There have been numerous reports of PGPR-mediated salt tolerance in plants (Bharti et al., 2016). For example, salt stress in tomato was ameliorated by Achromobacter piechaudii ARV8 producing 1-aminocyclopropane-1-carboxylate (ACC) deaminase (ACCD) (Mayak et al., 2004). Bacillus subtilis GB03 conferred salt tolerance via tissue-specific regulation of the ion transporter high-affinity K+ transporter (HKT) 1 in Arabidopsis (Zhang et al., 2008). In a recent report, a carotenoid producing halotolerant PGPR Dietzia natronolimnaes STR1 boosted salt tolerance in wheat (Bharti et al., 2016). Mechanistically, such tolerance is explained by means of hormone homeostasis, production of ACCD and volatile compounds, stimulation of expression of stress related genes, and sodium uptake/transport (Yang et al., 2009). An ABA-signalling cascade and innate immunity enhancement were mainly involved in the stress resistance (Bharti et al., 2016; Kim et al., 2014). Ethylene is important for plant growth and development, but excessive amounts of ethylene can decrease root growth (Saravanakumar and Samiyappan, 2007). The production of ACCD is linked to stress resistance, because abiotic stressors increase ethylene production from ACC in plants (Glick et al., 1998; Yang et al., 2009). During the last few decades, chemical pesticides have been the main strategy to manage phytopathogens. However, like chemical fertilizers, the extensive use of chemical pesticides can

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lead to negative effects on environment and food safety. Biopesticides, usually inherently less toxic, very targeted and biodegradable, have generated enormous interest as promising alternatives to chemical control (Olubukola, 2010). Over the last several decades, a great diversity of rhizobacteria, such as species within the genera Bacillus and Pseudomonas, have been reported to have antipathogen activities and effectively ward off a broad spectrum of phytopathogens in plants (Giorgio et al., 2015; Khabbaz and Abbasi, 2014; Lugtenberg and Kamilova, 2009). The mechanisms employed by these bacteria for disease suppression in plants may be a function of their ability to control niche-space competition, nitrogen fixation, phosphate solubilization, and production of antibiotics, siderophores, volatile compounds, hydrolytic enzymes and phytohormones (Santoyo et al., 2012). More recently, there have been clear demonstrations of specific signal compounds exchanged between plants and rhizosphere bacteria (members of the phytomicrobiome) that control each others’ gene expression, protein production, physiology and growth (Smith et al. 2017). Moreover, rhizobacteria-induced systemic resistance (ISR; Pieterse et al., 2014) has been reported in many plant species (i.e., rice, bean, cucumber, tobacco, tomato, and Arabidopsis) and is effective against a broad range of phytopathogens including fungi, bacteria, viruses, and insect herbivores. A prior infection, such as microbial symbiosis, that is effective for a period of time can induce resistance to pathogenic attacks that occur after symbiosis establishment (Fu and Dong, 2013). ISR requires the plant defensin 1.2 (PDF1.2) gene and is largely mediated by jasmonic acid (JA) and ethylene (ET) signaling pathways (Balmer et al., 2013). Previous studies reported that rhizobacterial strains trigger robust ISR against pathogens in various plant species via either JA/ET, ET, or salicylic acid (SA) signaling pathways, or both (Niu et al., 2016; Pieterse et al., 2012; Rudrappa et al., 2010). Benefical rhizobacteria trigger ISR by priming plants for a potentiated activation of various cellular defense responses, such as oxidative burst, cell-wall reinforcement (Nie et al., 2017), defense-related enzyme accumulation, and phytoalexin production (Kottb et al., 2015). With increasing awareness regarding environmental and economic effects of chemical inputs, it is important to search for novel isolates of beneficial rhizobacteria with multiple modes of action and to develop effective bioinoculants for plant growth promotion thereafter. In search of efficient PGPR strains, multiple traits related to PGP have been tested together during the initial screening process (Ambrosini et al., 2012; Li et al., 2017). These tests are usually in vitro screenings for PGP traits and are always tedious and time-consuming. A. thaliana has been

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developed as a model dicot plant for plant molecular biology (Meyerowitz, 1989) as well as plant-microbe interactions studies (Contesto et al., 2010; Kwon et al., 2016; Nie et al., 2017). Valuable information has been obtained about growth promoting interactions between Arabidopsis and PGPR isolated from a wide variety of plant species (Cartieaux et al., 2008). Moreover, there has been an increasing focus on the extent to which plant metabolism is altered by PGPR and the expression of which genes, measured as either transcriptional or translational expression, are changed by the beneficial interactions (Du et al., 2016; Kwon et al., 2016; Mortel et al., 2012; Poupin et al., 2013; Schwachtje et al., 2011; Spaepen et al., 2014; Stijin et al., 2014). Thus, in this study, Arabidopsis thaliana was used to screen for plant growth stimulating activity associated with rhizobacteria from a wide range of plant species (Chapter 3). In our previous study, 446 ectophytic and endophytic rhizobacteria were isolated from various crop and uncultivated plant species. The major goals of this work were to: (1) in-vitro screen bacterial isolates through direct plant growth-promoting effects on Arabidopsis Col-0 at an early growth stage; (2) characterize the most effective bacterial isolates on the basis of morphological and physiological attributes as well as by 16S rRNA gene sequence analysis; (3) to assess the PGP traits of selected isolates in vitro in order to identify them as potential biofertilizers; (4) investigate their effects on Arabidopsis under unstressed and salt stressed conditions (long-term growth) as stress relievers; (5) assess antagonistic ability against phytopathogens; (6) investigate the interaction between Arabidopsis and selected strains through a GFP technique and (7) investigate the possible PGP mechanisms of selected strains by using qRT-PCR to examine expression of selected genes. Our results demonstrated that all four tested rhizobacterial strains are able to colonize Arabidopsis roots promoting their growth in different stages. To the best of our knowledge, this is the first description of endophytic rhizobacteria associated with bittersweet (Solanum dulcamara) and fall dandelion (Scorzoneroides autumnalis) and able to promote plant growth in Arabidopsis.

4.3 Materials and Methods

4.3.1 Plant material, bacterial strains, and growth conditions

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The wild type Arabidopsis thaliana Columbia (Col-0 accession) was obtained from the Arabidopsis Biological Resource Center (ABRC, Ohio State University, Columbus, OH, USA). Seeds were aliquoted into 1.5 mL tubes and surface-sterilized for 5 min in 70% ethanol, followed by thorough rinses with sterile ddH2O (3 times), and then dipped in sterilized ddH2O, stored at 4 °C in the dark for 48 h to ensure stratification. Seed sterility was verified by incubating 20 seeds on DifcoTM Nutrient broth (NB) agar at 28 °C for 3 d. For agar medium experiments, seeds were sown on 10-cm Petri dishes containing one-half-strength (½) Murashige and Skoog (MS) (pH 5.7, adjusted with 10 mM KOH) supplemented with 0.8 % (w/v) agar. All plates were covered and sealed with Parafilm and were arranged in a germination chamber maintained at 22 ± 2 °C with a 16 h light/8 h dark photoperiod, at a photon flux density of 100 μmole m-2 s-1. The plates were relocated to a new position in the germination chamber twice a week to control for position-dependent variation. Petri dish experiments were organized following a completely randomized design and repeated three times, unless otherwise stated. For plant-growth chamber experiments, Arabidopsis seeds were either sown in individual Jiffy-7 peat pellets (Jiffy products, Plant Products Ltd., Brampton, ON, Canada) or at the soil surface at the rate of 4 seeds per pot filled with nutrient-containing vermiculite-potting soil mixture that was autoclaved twice for 20 min at 121 °C each time, with a 24 h interval between autoclavings. Thinning of seedlings to 1 per pellet or pot was conducted after the emergence of the first two true leaves (full germination, at 7 d). Plants were maintained in a plant growth chamber (Conviron PGR15, Winnipeg, MB, Canada) programmed for a 16 h photoperiod, day-night cycle with 200 μmol m-2 s-1 photosynthetic photon flux density at plant height and a constant temperature at 22 °C , and 60 % relative humidity. Plants were irrigated from a bottom reservoir with half-strength Hoagland nutrient solution (Hoagland and Arnon, 1950) twice a week and with ddH2O every 48 h. To avoid position effects in the soil-based experiment, plants were relocated each week until harvested. All the experiments were established following a completely randomized design and were repeated independently at least three times with consistent results, unless otherwise stated. Isolated bacterial strains (from ultra-cold storage) used for initial and secondary screening were routinely maintained on King’s B medium (KB). Single colonies were inoculated in liquid KB and grown aerobically on a rotary shaker (150 rpm) in the dark at 28 °C for up to 72 h, to reach the exponential phase of growth. Just before inoculation, bacterial cells were pelleted

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by spinning them down (6,000 g, 10 min, 4 °C ), washed free of the growth medium 4 times with sterile 10 mM MgSO4, and then the pellets were resuspended in 10 mM MgSO4 to a final density of 109 CFU mL-1, as determined by optical density (OD) at 600 nm and serial dilutions with plate counts. These suspensions were the standardized bacterial suspensions used for bacterization experiments. Control Arabidospsis plants of all bacterial treatments were mock-inoculated with sterile 10 mM MgSO4 and kept in parallel. The experiments related to phenotypic characterization of selected rhizobacteria were conducted three times each with four replicates, unless otherwise stated.

4.3.2 Rapid bacterial screening assay The effect of isolated bacteria on seedling growth in vitro was monitored as described by Subramanian (2013), with modifications. Ninety-eight newly isolated bacterial strains were used in this initial screening of potential plant growth promoting rhizobacteria. Sterilized Arabidopsis seeds were incubated for 1 h at room temperature with a given bacterial cell suspension prepared as mentioned above, or sterile 10 mM MgSO4 as the control (Schwachtje et al., 2012). Thereafter, 6 seeds were sown on ½ MS medium plates, with 8 plates for each treatment. The seedlings, 21 d after stratification (DAS), were collected by cutting the hypocotyls to separate the above- and within-agar parts (whole rosette) from Petri dishes, after which the rosette fresh weight (FW) was recorded.

4.3.3 Secondary bacterial screening Potential PGPR were selected through an initial screening (section 4.3.2) for a second-round screening. In the in vitro assays on ½ MS medium, two application methods were employed. For seed treatment, surface sterilized Arabidopsis seeds were incubated for 1 h at room temperature in 2 mL of bacterial suspension in 10 mM MgSO4, with sterile 10 mM MgSO4 as the control. For root tip inoculation (Mortel et al., 2012), 2 μL of bacterial cell suspension was pipetted onto root tips of 7-day-old seedlings, using equivalent amounts of sterile 10 mM MgSO4 as an axenic control. There were 10 replicate plates for each treatment and 6 seedlings in the same plate. For total leaf surface area quantification, photographs were taken of each plate at 11, 15, and 20 DAS. Images were analyzed using the free Java image processing program ImageJ (http://rsb.info.nih.gov/ij; developed by Wayne Rasband, U. S. National Institute of Health, 131

Bethesda, Maryland, USA). At 21 DAS, the seedlings, inoculated or not with bacteria, were harvested and the whole rosette FW was determined. This experiment was conducted in quadruplicate.

4.3.4 Petri plate assay for effects of bacterial inoculation on seedling growth 4.3.4.1 Germination activities of Arabidopsis seeds after bacterization Four isolates of rhizobacteria named as (n, L, K and Y) were selected from secondary screening (section 4.3.3) on the basis of their stimulation effect on early seedling growth of Arabidopsis. These finally chosen rhizobacterial isolates were used for all the following experiments, unless otherwise stated. Studies have indicated that Arabidopsis seedlings could not germinate or grow normally on MS medium with 200 mM NaCl or 400 mM mannitol (Huang et al., 2008; Wang et al., 2013). Based on the results of our pre-experiments, 100 and 150 mM NaCl and 100 and 300 mM mannitol were chosen for treating Arabidopsis. Sterilized and stratified seeds were imbibed in the cell suspension of the chosen bacteria and were sown on ½ MS medium or medium supplemented with specific concentrations of NaCl and mannitol. Plates were transferred to a germination chamber with a light and dark cycle of 16 and 8 h at 100 μmole m-2 s-1 light intensity, at 22 ± 2 °C . Germination was defined as 1 mm protrusion of radicle from the seed coat (Chen et al., 2012a). The number of germinated seeds was counted on a daily basis for a 10-d period. The percentage of germinated seeds was scored in four independent experiments, with 50 seeds per treatment.

4.3.4.2 In Vitro osmotic stress To test the effect of bacterial treatment on tolerance to osmotic stress, surface-sterilized

Arabidopsis Col-0 seeds were incubated in a bacterial suspension for 1 h in 10 mM MgSO4 or sterile 10 mM MgSO4, and were sown on ½ MS and allowed to grow for 3 days with the plates in a vertical orientation. Seedlings were then transferred, using sterile forceps, to ½ MS media or ½ MS media with osmoticum (100 mM mannitol) added, and allowed to grow for 11 days. After this 11-d stress treatment, the seedlings were observed and photographed. Rosette areas were measured using ImageJ (NIH). There were 8 replicate plates for each treatment and 6 seedlings in each plate. The seedlings were collected and the whole plant and rosette FW was measured and recorded. 132

4.3.4.3 In Vitro salt stress To evaluate salt tolerance during early seedling growth (Subramanian et al., 2016), surface-sterilized Arabidopsis Col-0 seeds were incubated in a bacterial suspension for 1 h, after which seeds were imbibed on ½ MS agar plates comprised of control and 100 mM NaCl treatment, and allowed to grow for 20 days, after vernalization at 4 °C in darkness for 3 d. There were 8 replicate plates for each treatment and 6 seedlings in the same plate. For total leaf surface area quantification, photographs were taken of each plate at 14 DAS. The seedlings were then collected and the whole plant (percentage growth reduction due to salinity, relative to control conditions) and rosette FW were recorded. Chlorophyll content in leaves of control and rhizobacteria-treated Arabidopsis was quantified according to the method of Fan et al. (2011). Fresh leaf tissue (50 mg) was extracted in the dark with 1 mL of pure methanol until the leaf material lost all its green pigment and became white. The homogenates were centrifuged at 10,000 g at 4 °C for 10 min and the corresponding supernatants were used for determining chlorophyll content. The absorbance of chlorophylls a and b were determined at 645, 663 and 750 nm. The amount of chlorophyll was calculated using the extinction coefficient, as indicated by Ritchie (2008). Concentrations of chlorophyll were expressed as μg mg-1 FW.

4.3.4.4 Root architecture analysis Root system architecture was assessed on 14-d-old Arabidopsis seedlings. In brief, surface-sterilized seeds were treated with chosen bacteria and sown on ½ MS medium supplemented or not with 100 mM NaCl, while in a vertical orientation. The seedlings were grown for 2 weeks, and then images of root systems were recorded and analyzed using the ImageJ software (NIH) with the NeuronJ plug-in (http://www.imagescience.org/meijering/; developed by Erik Meijering, University Medical Center Rotterdam, NL. The following root growth variables were measured: primary and lateral root length.

4.3.4.5 Quantitative measurement of gene expression via qRT-PCR analysis For quantitative real-time PCR, total RNA was isolated from 21-d-old seedlings, inoculated or not with bacterial suspension. For each treatment, 15 plants were randomly

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withdrawn (rosettes from 5 plants were pooled for each of 3 replicates) and were ground in liquid nitrogen using a pre-chilled mortar and pestle. Total RNA was extracted using an RNeasy Plant Mini Kit (Qiagen) following the manufacturer’s recommendations. The quality and quantity of the recovered total RNA were estimated by agrose (1%, w/v) gel runs and NanoDrop ND-1000 spectrophotometer (NanoDrop Technologies, Inc.), respectively. Genomic DNA was removed, and first strand cDNA were synthesized from 10 µg of total RNA using the QuantiTect Reverse Transcription kit (Qiagen), according to the manufacturer’s instructions, and stored at -20 °C. Expression analysis of key genes known to be involved in hormone signaling and production, ISR, phytoalexin production, and accumulation of osmolytes, were performed on a CFX Connect Real Time system (BioRad) using SYBR green Sso-advanced Supermix (BioRad), according to the manufacturer’s recommendations. Each reaction was performed on 3 µL of a 1:20 (v/v) dilution of the first cDNA strands (30 ng), in a final reaction volume of 15 µL. Negative controls (no template cDNA) were included. Gene specific primers used in this study were designed and verified using Primer-Blast (NCBI) (Ye et al., 2012) and TAIR BLAST (http://www.arabidopsis.org), respectively. Sequences of all primers are reported in Table 4.1. The following cycling parameters were used to perform qRT-PCR: 95 °C for 30s, and 40 cycles at 95 °C for 5s; 59 °C for 30s, followed by a melting cycle from 65 to 95 °C. The specificity of the amplification was checked by melting curve analysis, and then gel electrophoresis analysis to measure amplicon sizes. We found that expression of the constitutive SAND (At2g28390) was very stable; thus, the expression level of target genes was normalized using transcript abundance of SAND in each RNA preparations, as an internal control. The normalized value for the control was subtracted from each experimental value at each time point and the result inserted as ΔΔCt into the equation n=2-(ΔΔCt). This calculation provided the fold difference in gene expression of our experimental set in relation to the control (Livak and Schmittgen, 2001). Gene expression analyses were performed 21 days after treatment (seed-treated) or 12 h after bacterial treatment (3-week-old Arabidopsis seedlings). The gene expression levels were also determined during exposure to salt stress. All experiments were performed with three biological and three technical replicates. The experiment was performed twice, using independently grown plant tissues, to confirm the reproducibility of the results.

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Table 4.1. Primer sets used for real-time PCR in the present study

AGI Gene Primer sequences (5’-3’) Reference Amplicon size (bp) number CAGACAAGGCGATGGCGATA At2g28390 SAND Chen et al. (2016) 244 GCTTTCTCTCAAGGGTTTCTGGGT CTTGTTCTCTTTGCTGCTTTCGAC At5g44420 PDF1.2 current study 106 ATGCATTACTGTTTCCGCAAACC CCAAATCTCAAACCACGGCG At1g15550 GA3ox1 current study 81 ACAGGTAGCCCGAAGAGACT TGACGTTTGACCCCAACAGA At3g45640 MPK3 Kim et al. (2014) 146 CTGTTCCTCATCCAGAGGCTG CCGACAGTGCATCCTTTAGCT At2g43790 MPK6 Kim et al. (2014) 92 TGGGCCAATGCGTCTAAAAC GTGCTAGATGGAATCTGTCATGT At2g39800 P5CS1 current study 108 CGCATTACAGGCTGCTGGAT ATAATCAGTTGCAACTATGATCCTC At2g14610 PR1 Kim et al. (2014) 200 AAATAGATTCTCGTAATCTCAGCTC GCGTTGTCTCAAGCAGCATC At4g36110 SAUR9 current study 140 TAGCGACTTCGGTGTTGACC

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GTGGAGTCGCTGGCATAACA At3g26830 PAD3 current study 101 GTCCCCAAGTGTTGTCCGAA ACAAAACACACATAAACATCCAAAGT At5g52310 RD29A Kim et al. (2014) 98 ATCACTTGGCTGCACTGTTGTTC GAATCAAAAGCTGGGATGGA At5g42300 RD29B Kim et al. (2014) 197 TGCTCTGTGTAGGTGCTTGG

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4.3.5 Split-plate Petri-dish study In order to assess the possible effect of rhizobacterial volatile emissions on Arabidopsis growth, a split-plate Petri-dish study was conducted according to Ryu et al. (2004), with some modifications. Surface-sterilized Arabidopsis Col-0 seeds were placed on one side of the partitioned Petri dishes containing ½ MS solid media. Four-d-old Arabidopsis seedlings, selected for uniformity of size and development, were inoculated with 20 μL of a given bacterial strain (10-9 CFU mL-1) applied to the non-plant side of the partitioned Petri dish, which contained KB solid media. Non-growth-promoting Escherichia coli DH5α and 10 mM MgSO4 was used as negative controls (Zhang et al., 2008). There were 8 replicate plates for each treatment and 5 seedlings in the same plate. The seedlings were collected at 21 d after stratification (DAS). The whole rosette fresh weight was recorded.

4.3.6 In planta bacterial impact on growth of A. thaliana in soil 4.3.6.1 In Vivo assessment of plant-growth promotion The ability of isolated rhizobacteria to promote plant growth was assessed in a controlled environment (growth chamber) with 18 plants per treatment using a completely randomized design, in two steps, seedlings and seed production. Briefly, surface-sterilized seeds were placed on the surface of sterilized potting soil in each pot, after which each seed was inoculated with 1 mL of a bacterial suspension in 10 mM MgSO4. For the mock treatment

(control), seeds were incubated with an equal amount of sterile 10 mM MgSO4. At 28 days after bacterial innoculation, Arabidopsis plants were collected for measurement of shoot FW. Photographs were taken of each plant at harvest for total leaf surface area quantification. Fresh rosettes were dried in an oven at 85 °C for 48 h, and DW was recorded as mg g-1 FW. Rosette areas were measured using ImageJ (NIH). The effect of bacterial treatment on the growth of Arabidopsis was also tested under non-sterile conditions (peat pellets). Furthermore, the long-term growth effects of bacteria on Col-0 were determined following a 63-d growth period. Rosette diameter and stalk length was measured.

4.3.6.2 Salinity tolerance assay

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Prelimiary experiments showed that the Arabidopsis plants could tolerate 200 mM NaCl, while at 250 mM, stress symptoms were very severe (e.g. retarded growth, leaf senescence and loss of turgor) and plants did not recover well after the imposition of salt stress, thus 200 mM NaCl was chosen for use in this work. The ability of bacterial treatment to enhance plant resistance to salt stress was evaluated according to the methods of Zhang et al. (2010), with some modification. Briefly, developmentally uniform 6-d-old healthy seedlings were aseptically transferred from ½ MS agar plates, using tweezers, to pots filled with sterile potting soil, with one seedling per pot. Three days later, plants were root-treated with 10 mL of a given bacterial suspension, followed up by root-flooding with 200 mM NaCl (at 25 mL plant-1), 10 d post bacterization. Then, the plants were allowed to recover for a week by irrigating with sterile ddH2O, after which the plants were assessed for visual symptoms of salt stress. Relative chlorophyll content (or leaf greenness) was measured at the end of the experiment for all plants in each treatment, using a hand-held chlorophyll meter (SPAD-502, Konica Minolta), as the optical density. All plants were well irrigated before this measurement was taken. The fresh and dry weight of the aerial parts of plants were also measured.

4.3.6.3 Measurement of ROS scavenging activity The antiradical activity of Arabidopsis leaves (under normal growth conditions or salt stress) was determined using a DPPH (2,2-diphenyl-1-picrylhydrazyl hydrate) assay following Fan et al. (2011) with modifications. The antioxidants react with DPPH• and convert it to 1,1-diphenyl-2-picryl hydrazine with decoloration (from deep violet to light yellow). The degree of decoloration indicates the scavenging potentials of the antioxidants in terms of hydrogen donating capabilities. Fresh leaf tissue (1.5 g) was homogenized in 15 mL pure methanol (MeOH) using a mortar and pestle. After centrifugation at 10,000 g for 10 min, the supernatant was recovered. The pellet was re-extracted with 10 mL MeOH. Supernatants were combined, and the total volume was made up to 25 mL. Each extract (150 μL) was added to 2850 μL fresh DPPH methanolic solution (0.11 mM) and incubated in the dark for 1 h at 22 °C . Absorbance of the reaction mixture was then read at 515 nm, against an MeOH blank. The scavenging activity was calculated using the equation: Inhibition % = [(Ab−As)/Ab] 100, where Ab is the absorption of the blank sample and As is the absorption in the presence of a test sample. The results were

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expressed as μM ascorbate equivalents (AE, μM ascorbic acid) g-1 FW through comparison against an ascorbate standard curve (0.01–0.8 mg mL-1). For enzyme assays, approximately 0.2 g of leaves were ground into fine powder in the presence of liquid N2. The ground powder was then collected, using a spatula, into a microfuge tube with 1.5 mL of ice-cold 50 mM potassium phosphate buffer (pH 7.5) containing 1 mM ascorbate. The homogenate was vigorously vortexed for 10 min and then centrifuged at 10,000 g for 20 min at 4 °C . The supernatant (crude enzyme) was transferred to a new microtube, kept on ice, and analyzed right away. Protein concentration was determined by measuring the absorbance at 595 nm 5 min after mixing the solution with Bradford’s reagent. Bovine serum albumin (200–900 μg mL-1) was used as a reference standard. The activities of ascorbate peroxidase (APX, EC 1.11.1.11) were determined by following the decrease of ascorbate according to Nakano and Asada (1981) with modifications. The enzymatic reaction for APX was initiated by adding 50 μL of crude extract to the reaction mixture (1372 μL of 50 mM potassium phosphate, 75 μL of 10 mM ascorbate, 3 μL of 100 mM

H2O2). After 1 min at 25 °C , the H2O2-dependent oxidation of ascorbate to dehydroascorbate was monitored by measuring the decrease in absorbance at 290 nm. Nonenzymatic oxidation of ascorbate without enzyme extract was used as control. The amount of APX that can oxidize 1 μmol of ascorbate at 25 °C within 1 min is defined as 1 unit of enzyme. Catalase (CAT, EC 1.11.1.6) activity was determined as a decrease in absorbance at 240 nm at 25 °C following the decomposition of H2O2. The assay mixture (1 mL) contained 50 mM potassium-phosphate buffer (pH 7.5), 20 mM H2O2 and 20 μL of enzyme extract. Phosphate buffer without H2O2 was used as the control. The activity (1 unit of CAT) was calculated as the amount of enzyme catalyzing the decomposition of 1 μmol of H2O2 for 1 min (Piero et al., 1980). For determination of peroxidase (POD, EC 1.11.1.7) activity, the reaction mixture (2 mL) consisted of 100 mM potassium-phosphate buffer (pH 6.0), 0.1 mM pyrogallol, 5 mM H2O2, and 10 μL of crude enzyme. After incubation at 25 °C for 5 min, the reaction was stopped by adding

1.0 mL of 2.5 N H2SO4. The absorbance (indigo color) formed by purpurogallin was read at 420 nm against a blank (Milli-Q water). One unit of POD forms 1.0 mg of purpurogallin from pyrogallol in 20 seconds at pH 6.0 and 20 °C.

4.3.6.4 Metabolite measurements

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Shoots from bioprimed or control plants were harvested after 28-d of growth and were immediately frozen in liquid N2. Each bacterial treatment was measured with three independent samples of four plants (whole rosettes) pooled for one replicate. For total soluble sugar estimation (Singh and Jha, 2016), 50 mg leaf tissue were homogenized in 3 mL 80 % ethanol, followed by centrifugation at 10,000 g at 4 °C for 5 min to remove insoluble material. To 100 μL of sample was added 3 mL of freshly prepared anthrone reagent (200 mg of anthrone in 100 mL of concentrated sulphuric acid), and the reaction mixture was incubated in a boiling water bath for 10 min. After cooling on ice, absorbance was read at 620 nm against 80 % ethanol as a blank. The total soluble sugar was expressed as glucose equivalent by comparison with a glucose standard curve (0-20 mg). The extraction of total soluble protein was conducted according to Fan et al. (2011).

Fresh rosette (0.5 g) was extracted in 2 mL of 0.1 M KH2PO4/K2HPO4 buffer (pH 8.0), containing 5.0 mM β-mercaptoethanol and 2 % (w/v) PVP, and centrifuged at 10,000 g for 20 min at 4 °C. The supernatant was collected as the crude enzyme. Protein concentration was determined as described in section 4.3.6.3.

4.3.6.5 Proline estimation Proline content in the fresh leaves of 4-week-old plants were determined following Singh and Jha (2016) with modificatioins. Fresh leaf tissues (100 mg) were homogenized in 3 mL of 3% sulfosalicyic acid at 95 °C for 15 min. After centrifugation at 8,000 g for 10 min, the resulting supernatant was combined with a 1:1:1 mixture of glacial acetic acid, and 2 % acid ninhydrin (1.25 g ninhydrin in 30 mL glacial acetic acid) and 20 mL 6 M orthophosphoric acid. The mixture was boiled for 30 min in a boiling water bath. After cooling at room temperature for 30 min, 1 mL of toluene was added to the mixture, to extract the red chromophore, with vigorous shaking for 30 s. The absorbance of the upper toluene phase was determined spectrophotometrically at 520 nm against toluene as a blank. Free proline content was measured by comparison with a pure L-proline standard curve prepared by the same method and calculated on a fresh weight basis expressed as mg 100 g-1 FW.

4.3.7 Screening for bacterial PGP traits 4.3.7.1 Ammonia production 140

The ability of bacterial strains to produce ammonia was assessed using the method described by Marques et al. (2010). Bacterial culture (0.2 mL) was added to 5 mL peptone water

(per 100 mL ddH2O: 1 g Bacto-peptone, 0.5 g NaCl) and incubated at 28 °C for 5 days, after which 0.5 mL Nessler’s reagent (alkaline solution of potassium tetraiodomercurate (II)) was added to each tube and development of an orange yellow to brown color indicated a positive result (presence of ammonia) while the intensity of color was indicative of amount of ammonia produced by the test strain. The control tube showed a pale yellow, indicating no reaction.

4.3.7.2 ACC deaminase activity measurement Quantitative measurement of ACC deaminase activity was estimated by measuring the amount of α-ketobutyrate produced from the cleavage of ACC (Glick et al., 1995; Penrose and Glick, 2003; Smyth et al., 2011), by comparing the absorbance at 540 nm of a sample with a standard curve of α-ketobutyrate. Bacterial strains were cultured in 5 mL KB liquid medium at 30 °C to reach the stationary phase. After centrifugation at 6,000 g at 4 °C for 10 min, the supernatant was removed and cell pellets were washed twice with 0.1 M Tris-HCl (pH 7.6), after which the cells were suspended in 2 mL of DF minimal salts medium (per L ddH2O: 4.0 g

KH2PO4, 6.0 g Na2HPO4, 0.2 g MgSO4·7H2O, 2.0 g glucose, 2.0 g gluconic acid and 2.0 g citric acid; 10 mL trace elements (per L ddH2O: 100 mg FeSO4·7H2O, 1 mg H3BO3, 1.12 mg

MnSO4·H2O, 12.46 mg ZnSO4·7H2O, 7.82 mg CuSO4·5H2O and 1 mg MoO3, pH 7.2) in a fresh tube. The stock 0.5 M ACC solution (12 μL) was added to the tube immediately prior to incubation, to get a final ACC concentration of 3.0 mM, as the sole nitrogen source. DF salts minimal medium without supplemented ACC, served as the negative control. The tubes were incubated at 30 °C to induce ACC deaminase activity, after which the cells were harvested (10,000 g at 4 °C for 10 min) and washed twice with 0.1 M Tris-HCl (pH 7.6). The pellets were flash freezed in liquid nitrogen (N2) and stored at -80 °C until ACC deaminase activity determination. The pellet of induced bacterial cells was re-dissolved in 200 μL of 0.1 M Tris-HCl (pH 8.5), to which 10 μL of toluene (5 %, v/v) was added and then vortexed for 30 s. The tube was kept at 4 °C for 1 h and then centrifuged at 12,000 g for 10 min at 4 °C , after which the thin layer of toluene was aspirated gently. Part of the toluenized cells was used for the determination of total protein concentration by the dye binding method of Bradford (1976), using the Bio-Rad

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protein reagent (Bio-Rad Lab., USA) according to the manufacturer’s instructions. Briefly, a 26.5 μL aliquot of the labilized cell sample was diluted with 173.5 μL of 0.1 M Tris-HCl (pH 8.5) and boiled with 200 μL of 0.1 N NaOH for 10 minutes. After the sample was cooled to room temperature, the protein concentration was determined by measuring the absorbance at 595 nm 5 min after mixing the solution with 200 μL of Bradford’s reagent. Bovine serum albumin (BSA) was used to establish a standard curve. A stock solution of BSA (10 mg mL-1) was diluted with Milli-Q water to specific final concentrations of 200 to 900 μg mL-1. The remaining labilized cell suspension was immediately assayed for ACC deaminase activity. Briefly, 50 μL of the toluenized mixture was transferred to a fresh Eppendorf microcentrifuge tube, to which 5 μL of 0.5 M ACC was added and gently vortexed. The negative control for this assay was 50 μL of labilized cell suspension without ACC, while the blank was 50 μL of 0.1 M Tris-HCl (pH 8.5) with 5 μL of 0.5 M ACC. After incubation at 30 °C for 30 min, 0.5 mL of 0.56 N HCl was added to completely terminate the reaction and then the mixture was votexed and centrifuged at 12,000 g at 4 °C for 10 min to remove the cell debris. An aliquot of the supernatant (500 μL) was transferred to a 13×100 mm glass test tube and mixed throughly with 400 μL of 0.56 N HCl; thereupon, 150 μL of the 2,4-dinitrophenylhydrazine reagent (DNPH; 0.2 % 2,4-dinitrophenylhydrazine in 2N HCl) was added; and the mixture was vortexed and incubated at 30 °C for 30 min. After the addition of 1 mL of 2 N NaOH for α-ketobutyrate-derivatized phenylhydrazone color development, the absorbance of the mixture was read at 540 nm. To create a standard curve, 500 μL of α-ketobutyrate solutions at known concentrations (0.1 to 0.5 mM) were mixed with 400 μL of 0.56 N HCl and 150 μL of DNPH solution. The absorbance was measured at 540 nm after adding 1 mL of 2 N NaOH to the mixture. The enzyme activity was defined as μmol α-ketobutyrate h-1 mg-1 protein.

4.3.7.3 Indole-3-acetic acid (IAA) detection The in vitro ability of bacterial strains to produce IAA like substances was quantitatively assayed according to the colorimetric method of Gordon and Weber (1951) with modifications. In brief, bacterial cultures (0.1 mL, in exponential growth) were used to inoculate 5 mL KB medium supplemented with L-tryptophan (200 or 500 μg mL-1; as an IAA precursor) or not in the case of the control and incubated at 28 °C for 2-5 d in the absence of light. These cultures were centrifuged at 10,000 g for 5 min at 20 °C , and to 1 mL supernatant, 1 drop of

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orthophosphoric acid and 2 mL freshly prepared Salkowski’s reagent (98 mL of 35 % perchloric acid plus 2 mL of 0.5 M FeCl3) was added and incubated at room temperature in the dark for 30 min. Salkowski’s reagent can detect IAA and its intermediates (Glickmann and Dessaux, 1995) and then produce a proportionate amount of tric-(indole-3-acetato) iron (III) complex, which is pink in color. Development of reddish to pinkish color indicates IAA production, while control tubes show a pale yellow or colorless response. Equivalent mixtures without inoculation served as blanks. Bacterial cells were separated by centrifugation at 10,000 g for 20 min and the amount of IAA produced in the supernatant was measured spectrophotometrically at 530 nm and expressed as mg μL-1 by comparison with a pure IAA standard curve within the range of 5-100 μg mL-1.

4.3.7.4 Nitrogen fixing ability

Nitrogen-free bromothymol blue malate (NFb) media (per L ddH2O: 5.0 g L-malic acid

(malate), 4.0 g KOH, 0.5 g K2HPO4, 0.2 g MgSO4·7H2O, 0.1 g NaCl, 0.02g CaCl2·2H2O, 2 mL 0.5% bromothymol blue (BTB) in 0.2 M KOH, 4 mL 1.64% Fe EDTA, pH 6.8 and 1.5 % agar; 2 mL sterile, filtered micronutrient solution (per L ddH2O: 0.235 g MnSO4·H2O, 0.2 g

Na2MoO4·2H2O, 0.024 g ZnSO4·7H2O, 0.04 g CuSO4·5H2O, 0.28 g H3BO3); 0.5 mL filter-sterilized vitamin solution (per L ddH2O: 0.01 g biotin, 0.02 g pyridoxol-HCl) was used for visual detection of nitrogen-fixing activities of bacterial strains (Baldani et al., 2014). Each bacterium was placed in triplicate onto the agar medium using sterile toothpicks. The plates were incubated at 30 °C for up to 7 d. The bacteria were re-streaked to new NFb media plates and incubated for another 7 d at 30 °C . Green color changing to blue color indicated that the isolate probably had nitrogen fixing activity in the solid culture conditions. Second, to test the presence of nifH gene (marker gene which encodes nitrogenase protein Componet II), polymerase chain reaction (PCR) specific for nifH gene segments was attempted using four sets of universal primers (19F/388R, IGK3/DVV, ZehrF/ZehrR, PolF/PolR) (Table 4.2). Primers were synthesized and desalted by Integrated DNA Technologies. These universal primers were supposed to amplify a 350 to 400 bp fragment of the nifH gene (Table 4.3). Total bacterial genomic DNA was extracted with the DNeasy Blood & Tissue Kit (Qiagen) according to the manufacturer’s instruction. The purity was assessed from the A260/280 and A260/230 extinction ratios using a NanoDrop (Thermo Scientific). A blank/negative control (PCR mixture

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without DNA template) was maintained in PCR reaction. PCR reaction was performed in a 50 μL final volume including 2 μL of total DNA, 5 μL of universal forward (FWD) primer (10 μM), 5 μL of universal reverse (REV) primer (10 μM), 5 μL of deoxynucleotide triphosphates (dNTPs)

(2 mM each), 5 μL 10×PCR buffer, 5 μL of 25 mM MgCl2, 1 μL Taq DNA polymerase (5U) and nuclease-free water. The amplification reaction was carried out in a PTC-100TM Programmable Thermal Controller (MJ Research, Inc., MN, USA): initial denaturation for 3 min at 94 °C ; 30 s at 94 °C, 30 s at 58 °C , 1 min for 72 °C (30 cycles); and a final elongation at 72 °C for 5 min. The PCR products were seperated by running 10 μL of the PCR reaction mixture in 1.2 % (w/v) agarose (Bio-Rad, ON) gels buffered with 0.5×TAE (Tris Acetate EDTA) and staining the bands with SYBR Safe DNA gel stain (1:10000 dilution in TAE) for 45 min at 100 volts and 500 miliamps. Molecular weight of PCR products was estimated by comparison with Fast DNA ladder (New England Biolabs). Bands were visualized and photographed using a digital camera (GIS-1000; Tanon) under UV light. Target bands were purified using QIA quick Gel Extraction Kit (Qiagen), cloned into the vector pDONR/Zeo (Invitrogen) using ClonaseTM GatewayTM BP Clonase II enzyme mix (Invitrogen), and then subjected to sequencing. The sequences were compared against GenBank databases using the BLASTN search algorithm (http://blast.ncbi.nlm.nih.gov), on the basis of identity percentage, E-value and Match score.

Table 4.2. PCR Primer sets used for nifH gene amplification.

Primers Sequences (5’-3’) Amplicon Reference (bp) 19F GCIWTYTAYGGIAARGGIGG 390 [a] 388R AAICCRCCRCAIACIACRTC IGK3 GCIWTHTAYGGIAARGGIGGIATHGGIAA 390 [b] DVV ATIGCRAAICCICCRCAIACIACRTC ZehrF TGYGAYCCIAARGCIGA 360 [c] ZehrR ADNGCCATCATYTCNCC PolF TGCGAYCCSAARGCBGACTC 350 [d] PolR ATSGCCATCATYTCRCCGGA

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a Ueda et al (1995); b Gaby and Buckley (2012); c ZehrR and McReynolds (1989); d Poly et al (2001)

4.3.7.5 Solubilization of insoluble phosphate The dicalcium phosphate (inorganic phosphate) solubilizing ability of selected strains was first qualitatively tested by plate assay using National Botanical Research Institute's phosphate (NBRIP) growth medium (per L ddH2O: 10 g glucose, 5 g MgCl2 · 6H2O, 0.25 g

MgSO4 · 7H2O, 0.2 g KCl and 0.1 g (NH4)2SO4; pH 7.0 and 1.5 % agar) (Nautiyal, 1999), with the addition of Ca3(PO4)2 (5 g) to form a milky appearance on plates. Ten microliters of the bacterial suspension in 10 mM MgSO4 was spotted on the agar plates and was allowed to dry completely followed by incubation at 28 °C . The zone of clearance around the colony was considered to be a positive indication of P solubilization. The halo (area of turbid medium cleared) and colony diameters were measured after a 14-d incubation. Halo size was calculated by subtracting colony diameter from the total diameter. Mineral phosphate solubilizing ability was also tested under buffering conditions, using

Tris phosphate medium (per L ddH2O: 100 mM Tris-HCl, 100 mM glucose, 25 μM MgSO4, 10 mM NH4Cl, 5 g Ca3(PO4)2 and 15 g agar; 10 mL sterile, filtered micronutrient cocktail (per L ddH2O: 0.35 g FeSO4·7H2O, 0.016 g ZnSO4·7H2O, 0.008 g CuSO4·5H2O, 0.05 g H3BO3, 0.003 g CaCl2·2H2O, 0.04 g MnSO4·4H2O); pH 8.0) (Singh et al., 2014) with 0.01 % methyl red as pH indicator to check phosphate solubilization or acid production. The Tris-buffered Methyl red agar plates represent a much more stringent condition of screening for phosphate solubilization microorganisms (Gyaneshwar et al., 1998). P solubilization was monitored by the formation of a red zone (from pH=8.0 to pH<5), which indicates a drop in pH of the buffered medium. The effect of buffering on mineral phosphate solubilization was determined by measuring the diameter of the red zone around the colony after 14 d of incubation at 28 °C. The in vitro phosphate-solubilizing potential of selected strains was quantitatively estimated (Halder et al., 1990) by inoculating a 1 % (v/v) bacterial suspension (in 10 mM MgSO4) into 100 mL NBRIP medium broth in Erlenmeyer flasks (500 mL) and incubating at 28 °C for 14 d at 150 rpm. Autoclaved uninoculated broth incubated under same conditions served as the control. To ensure no contaminating P was carried over, all glassware was throughly cleaned with 6 M HCl and ddH2O prior to the preparation of NBRIP medium. An aliquot of 2 mL was withdrawn periodically from each flask on days 1, 3, 5, 7, 10 and 14. The samples were then

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centrifuged at 10,000 g for 10 min at room temperature and the P content in the cell-free supernatant of each culture was estimated using the Fiske and Subbarow method (1925), by adding Barton’s reagent (2.5 mL) to a 0.5 mL aliquot of the supernatant and the volume was made up to 50 mL with ddH2O in a volumetric flask. After a 10-min incubation, the intensity of vanado-molybdate-yellow color was spectrophotometrically determined at OD430 and expressed as equivalent phosphate (μg mL-1) by comparison with a potassium dihydrogen orthophosphate -1 (KH2PO4) standard curve (0-900 μg mL ).

4.3.7.6 Siderophore production The production of siderophores by the four bacterial strains of interest was qualitatively assayed using the chrome azurol S (CAS) method (Alexander and Zuberer, 1991) with modifications, in which the tertiary complex (CAS/Fe3+/hexadecyltri-methylammonium bromide (HDTMA)) served as an indicator dye. To ensure no contaminating iron was carried over, all glassware was throughly cleaned with 6 M HCl and ddH2O prior to media preparation. Briefly, a fresh blue Fe-CAS indicator solution (Blue Dye solution) was prepared (per 100 mL: 60 mg

CAS, 2.43 mg FeCl3·6H2O, 73 mg HDTMA) and autoclaved and stored in a plastic bottle. The buffer solution containing 32.24 g piperzine-1,4-bis (2-ethanesulfonic acid) (PIPES), 0.3 g

KH2PO4, 0.5 g NaCl, 1.0 g NH4Cl, and 15 g agar in 800 mL ddH2O was pH-adjusted to 6.8 with 50 % NaOH and autoclaved and cooled to 50 °C , after which 30 mL filter-sterilized 10 % (w/v) casamino acids and 10 mL of sterile 20% glucose was added. Finally, 100 mL of Blue Dye solution was slowly added, and the CAS agar plates were then poured aseptically. Bacterial single colonies were stabbed in triplicate on CAS agar medium and incubated in the dark at 28 °C for up to 10 d. When the iron was removed from the ferric complex of CAS by bacterial siderophores, which have a higher affinity for Fe3+, a clear halo around the colony was formed and a color change, from blue to yellow-orange developed, which was the result of siderophore production.

4.3.7.7 Hydrogen cyanide production hydrogen cyanide (HCN) is a broad spectrum antimicrobial compound that plays an important role in the suppression of root diseases by rhizobacteria, especially many plant-associated Pseudomonas isolates. The ability of isolates to produce HCN from glycine was

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tested by adapting the method of Ahmad et al. (2008). Briefly, isolated bacteria were streaked on LB agar medium amended with glycine (4.4 g L-1). A sterile Whatman filter paper No. 1 soaked in 2 % sodium carbonate (Na2CO3) in 0.5 % picric acid solution was placed on the underside of the plate lid. Non-inoculated plates and inoculated media with only Na2CO3 impregnated filter paper were used as controls. The plates were then sealed with parafilm and incubated at 28 °C for up to 7 d. A color change from yellow to orange or reddish-brown indicated cyanogenic potential.

4.3.7.8 Evaluation of antimicrobial activities The in vitro antagonistic abilities of bacterial isolates against other microorganisms were evaluated using a dual culture method as described by Shehata et al. (2016), with modifications. Pseudomonas syringae pv. tomato (Pst) DC3000 EV (virulent) (Xin and He, 2013), a biotrophic plant pathogenic bacterium, was maintained on LB agar plates supplemented with kanamycin (50 μg mL-1) and rifampicin (10 μg mL-1) at 28 °C . A necrotrophic fungus, Botrytis cinerea wild-type isolate B191 (Oirdi and Bouarab, 2007), was maintained on potato dextrose agar (PDA) amended with streptomycin (20 mg L-1) at 22 °C . Routine subculture was conducted by transferring an agar plug from a stock plate onto the center of a fresh PDA plate. Antibacterial activity was determined through two approaches. In the first, KB agar plates 8 -1 were inoculated with Pst DC3000 (10 CFU mL ) in sterile ddH2O prepared from overnight culture. Wells were then punched into each plate with a cork borer (5 mm size) and 50 μL of overnight broth cultures of each rhizobacterium (108 CFU mL-1) was applied to each well. In a second approach, 5 μL aliquots of rhizobacterial suspension from overnight culture were spotted on Pst DC3000 lawn. All plates were then incubated at 28 °C for 3 d, after which, the diameter of the inhibition zone, which was clear of pathogenic bacterial growth, was measured. For antifungal assay, a 5-mm agar plug from the colony margin of a 7-d-old PDA culture of Botrytis cinerea 191 at 22 °C was recuperated and deposited firmly in the center of a fresh PDA plate, with a 5-mm well filled with 50 μL of overnight culture of bacterial isolates approximately 2 cm apart. Plates were incubated at 22 °C and examined daily for antagonism by measuring the distance between the edges of the bacterial isolates and the fungal mycelium (zone of inhibition). All in vitro tests of antagonism were performed three times with six replicates for each bacterial isolate.

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4.3.7.9 Exopolysaccharide production Exopolysaccharide (EPS) production was detected macroscopically on RCV medium modified from Santaella et al. (2008) (per L ddH2O: 4.4 mM KH2PO4, 5.2 mM K2HPO4, 0.1 g yeast extract, 4.0 g glucose, 0.1 g MgSO4·7H2O, 0.1 g CaCl2·2H2O, 22 mg FeSO4·7H2O, 20 mg

EDTA, 0.43 mg ZnSO4·7H2O, 2.8 mg H3BO3, 1.30 mg MnSO4·H2O, 26 μg CuSO4·5H2O, 0.75 mg Na2MoO4, 70 μg CoSO4; pH 6.8 and 1.5 % agar). Single colonies were streaked on RCV-agar medium and incubated at 28 °C for 3 d. Strains whose colonies showed a mucoid phenotype or watery surface having a glistening and slimy appearance on agar plates probably produce EPS.

4.3.7.10 Blood hemolytic assay The four selected strains were investigated for their possible activity on biological membranes via in vitro hemolysis assay according to Liu (2015). Briefly, a colony of each bacterium was transferred from KB plates to blood agar plates (sheep blood plates, MP1301, HiMedia) by using sterile toothpicks. After incubating for 72 h at 28 °C , hemolysis was visualized by the development of a clear hemolytic halo (β-hemolysis) around the colonies, indicating biosurfactant production. Dection of α-hemolysis was indicated by the production of greenish and dark color in the agar under the colony, which is usually caused by hydrogen peroxide produced by the bacterium, indicating partial damage of erythrocytes; γ-henolysis, or so-called non-hemolytic, resulted in no alteration of color or opacity in the agar medium.

4.3.8 Identification and characterization of bacterial isolates 4.3.8.1 Morphological characterization Gram reactions were determined using the bioMérieux Gram-stain kit according to the manufacturer’s instructions. The four strains were grown on KB agar at 28 °C and were characterized according to their colony shape (i.e., circular, irregular or pointed), elevation (i.e., flat, spread, convex, high convex, umbilicate, umbonate or raised), edge (i.e., complete, undulate, lobate, dented, firbriate or ciliate), consistency (mucoid, friable, firm or butyrous), aspect of colony surface (i.e., smooth, granular or rough), opacity (i.e., transparent, translucent or opaque), color, and growth speed (i.e., very fast: visible to the naked eye after less than 24 h of incubation; 148

fast: visible within 24-48 h; intermediate: visible within 48-72 h; slow: visible within 72-96 h; or very slow: visible only after 96 h) (Nagoba and Pichare, 2016; Zarpelon et al., 2016).

4.3.8.2 Intrinsic antibiotic sensitivity/resistance Though a wide variety of techniques, such as broth microdilution and disk diffusion, have been used for antimicrobial susceptibility testing (AST), many of them are cost-ineffective, time-consuming or labor-intensive. Thus, the antimicrobial gradient diffusion method using Etest® (bioMérieux, La Balme-les-Grottes, France) (Jorgensen and Ferraro, 2009) was chosen in this study to quantitatively determine the susceptibility of isolated bacteria to various antibiotics (ampicillin, erythromycin, kanamycin, chloramphenicol, gentamicin, streptomycin, tetracycline, rifampicin, vancomycin, and tobramycin). Briefly, selected rhizobacteria were grown for 12-48 h in 4 mL KB medium, with shaking (150 rpm) at 28 °C in 15 mL culture tubes and diluted to 108 CFU mL-1 in sterile 0.8 % NaCl. Bacterial suspension (100 μL) was spread-inoculated on Müeller-Hinton (Sigma) agar medium, which is routinely used for susceptibility testing, and plastic Etest strips with predefined stable antimicrobial gradient were dispensed firmly onto the inoculated agar surface with sterile forceps. The plates were incubated at 28 °C and showed pure confluent growth. Since strain K showed no growth on Müeller-Hinton medium, this bacterium was tested on KB agar. After 12-72 h of incubation, the Minimum Inhibitory Concentration (MIC) for individual antimicrobial agents, in μg mL-1, of the antibiotics were determined by measurement of the symmetrical ellipse shaped growth inhibition area. This experiment was performed 3 times with 4 strips per antibiotic for each bacterium each time to confirm the reproducibility of the results.

4.3.8.3 In Vitro determination of bacterial abiotic stress tolerance Bacterial isolates were evaluated for salt stress and pH tolerance (Roman-Ponce et al., 2017). For salinity analysis, single colonies of bacterial strains were streaked onto the surface of KB agar amended with increasing concentrations of NaCl at 0, 0.5, 1, 2, 3, 4, 5, 6, 7, and 10 % (w/v). The pH ranges for bacterial growth were determined by inoculating each isolate onto KB plates adjusted to various pH values in the range of 4.0 to 10.0 in 1.0-unit increments. Plates were incubated at 28 °C for a 7-d period. The ability to grow in corresponding medium with different concentrations of NaCl or pH value was examined. Similarly, the growth characteristics

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of isolates across a range of temperatures (30, 35, 37, 40 and 42 °C) in KB liquid medium was recorded every 24 h for 7 d of culture.

4.3.8.4 Extracellular enzyme production To test proteoloytic activity (protease), qualitative analysis was performed on Lysogeny broth (LB) supplemented with 1 % (w/v) skim milk (Oxoid). Individual strains were streaked on agar plates and incubated at 28 °C for up to 4 d. A positive result (casein hydrolysis) was indicated by a whitish, opaque halo (coagulated casein) zone of clearance around grown colonies due to casein degradation (hydrolysis) and formation of soluble nitrogenous compounds. The presence of the enzyme catalase was evaluated by aseptically transferring a loopful of fresh cultures with the help of a sterile glass rod onto a glass slide and adding 2 drops of 3 %

H2O2. Immediate bubbling (O2 formed) was considered as a positive indication for catalase. Oxidase activity was determined using oxidase reagent (bioMérieux) according to the manufacturer’s instructions. Hydrolysis of starch was determined based on a color reaction of non-hydrolysed starch with Gram’s iodine (per 300 mL ddH2O: 1.0 g iodine, 2.0 g potassium iodine). When starch is broken-down, as hydrolysis progresses, it results in a brownish-red color and is, finally, colorless.

Starch agar plates (per L ddH2O: 3.0 g beef extract, 10.0 g soluble starch; pH 7.5 and 1.2 % agar) was streak-inoculated and incubated at 28 °C for 3 d. The surface of each plate was flooded with Gram’s Iodine. A clear, colorless zone of degradation around the colony indicated a positive result (detection of an amylolytic strain), while a negative result was indicated by a deep blue color throughout the medium. The cellulolytic activities of the isolates were measured by a quantitative agar spot method. To determine the production of endoglucanases, the cells (108 CFU mL-1) were spotted on agar medium (Mandels and Reese, 1957) supplemented with carboxymethylcellulose sodium salt (CMC-Na; per L ddH2O: 2.0 g KH2PO4, 1.4 g (NH4)2SO4, 0.3 g MgSO4 ·7H2O, 0.3 g CaCl2,

0.4 g yeast extract, 0.005 g FeSO4·7H2O, 0.0016 g MnSO4, 0.0017 g ZnCl2, 0.002 g CoCl2, 5.0 g CMC-Na; pH 7.0 and 1.5 % agar), and incubated at 28 °C for 7 d, after which all the plates were stained with 0.1 % (w/v) Congo-red, which binds to intact β-D-glucans, for 30 min and then washed and discolored with 1 M NaCl for 30 min (Teather and Wood, 1982). The presence of

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yellow-colored halo zones visible around the bacterial colonies indicated endo-glycolytic activities. Similarly, the presence of the exoglucanase that release cellobiose from the reducing and nonreducing ends, was estimated on a synthetic medium containing avicel (microcrystalline cellulose; per L ddH2O: 0.5 g KH2PO4, 1.4 g (NH4)2SO4, 0.3 g MgSO4 ·7H2O, 2.0 g yeast extract, 3.0 g Avicel; pH 7.0 and 1.5 % agar). Clear halo development by bacterial colonies after 10-d of incubation at 28 °C indicated exo-cellulase activity. Positives showed a clear halo against a milky background. There were 3 independent experiments with 4 replicates per isolated strain.

4.3.8.5 Carbon source utilization API (Appareils et Procédés d'Identification, bioMérieux) 50 CH galleries were used, as part of the approach employed for the study of carbohydrate metabolism of bacteria, along with the HiCarbohydrateTM kit (KB009, HiMedia), to test the ability of bacterial isolates to metabolize various substrates as sole carbon source (through acid production from various carbohydrates). Bacterial inoculation was conducted according to the manufacturer’s instructions. For the API 50 CH system, fresh cultures of isolates on KB agar medium were washed off the plates and resuspended in sterile 0.85 % NaCl and then diluted in API 50 CHB medium (bioMérieux). Homogenized suspensions were transferred into the wells on the API 50 CH strips. For HiMedia kits, bacterial suspension in 0.85 % NaCl with an OD600 of 0.6 was transferred into each of the 36 wells on HiMedia strips. API or HiMedia strips were covered, as recommended by the manufacturer, and incubated at 28 °C. Sugar fermentation results in the formation of acid, which changes the pH of the medium, which is expressed as change in color of the indicator from red to yellow. Color changes were monitored after 1, 2 and 4 days. Specifically, esculin hydrolysis was presented as positive with a color change to black. For HiMedia kit, citration and malonate utilization was considered positive with a color change to blue. Complete experiments were performed twice with highly reproducible results.

4.3.8.6 Evaluation of other biochemical activities Activities of constitutive enzymes and other physiological features of the bacteria were generated using the API 20E and API 20NE (bioMérieux). The activities of the 19 enzymes involved in the main nutrient biogeochemical cycles: carbon (β-glucosidase, α-glucosidase, 151

α-galactosidase, β-galactosidase, β-glucuronidase, α-mannosidase, α-fucosidase, esterase lipase, and lipase activities), phosphorous (acid and alkaline phosphomonoesterase and Naphthol-AS-BI-phosphohydrolase activities), and nitrogen (protease, leucine-arylamidase, valine arylamidase, N-acetyl-glucosaminidase activities), were tested according to the API ZYM assay (Martínez et al., 2016). All API tests were conducted according to the manufacturer’s instruction. Briefly, fresh-prepared bacterial suspensions in 0.85 % NaCl solution were added to the API reaction strips and incubated at 28 °C , in the dark, for up to 3 d. The results were read according to the color change following manufacturer's instructions. Each strain was tested twice with the API kits to ensure reproducibility of results. Aerotolerance of selected bacteria was determined in thioglycollate broth (Sigma), in which sodium thioglycolate serves as a reducing agent and creates a redox potential in the tube. In brief, freshly prepared thioglycolate broth was inoculated with fresh bacterial culture using a long toothpick. Tubes were then incubated statically in the dark at 28 °C for up to 7 d, after which the growth characteristics were recorded for each strain.

4.3.8.7 Phylogenetic analysis of 16S rDNA sequencing data for selected bacteria To determine the phylogenetic positions of the four strains, 16S rRNA gene sequneces for them were amplified using three sets of universal 16S rRNA gene primers: P0 (GAGAGTTTGATCCTGGCTCAG) and P6 (5′-CTACGGCTACCTTGTTACGA), 27F (5’-AGRGTTYGATYMTGGCTCAG-3’) and 1064R (5’-CGACRRCCATGCANCACCT-3’), and 27F (5’-AGRGTTYGATYMTGGCTCAG-3’) and 1391R (5′- GACGGGCGGTGWGTRCA-3′). Of these primers, primers P0 and P6 were designed on the basis of the conserved bacterial sequences at the 5′ and 3′ ends of the 16S rRNA gene (positions 27f and 1495r in Escherichia coli rDNA, respectively) and this allowed amplification of almost the entire gene (Grifoni et al., 1995). The four selected isolated rhizobacteria were cultivated in KB medium at 28 °C for 12-72 h. Genomic DNA was extracted with the DNeasy Blood & Tissue Kit (Qiagen) according to the manufacturer’s instruction. DNA sequencing of PCR products were carried out using the Genome Quebec Innovation Centre (McGill University) service on a 3730XL DNA analyzer systems (Applied Biosystems). Retrieved DNA sequences were edited by Seqman Pro (DNASTAR Lasergene package). DNA sequences were compared with other 16S rRNA gene sequences available by the BLASTN program 152

(http://blast.ncbi.nlm.nih.gov/Blast.cgi) and with the sequences held in the EzTaxon-e server (Yoon et al., 2017), and then were aligned with similar sequences by using the CLUSTX program. All reference sequences were obtained from the National Centre for Biotechnology Information (NCBI) and Ribosomal Database Project (RDP) databases (Maidak et al., 1997). Phylogenetic trees based on 16S rRNA gene sequences were constructed by applying the neighbor-joining method using MEGA version 7.0 (Hall, 2013) based on the Kimura 2-parameter model with 1000 replicates of bootstrap values (Tamura et al., 2013). The partial 16S rDNA gene nucleotide sequences of the four isolated bacterial strains n, L, K and Y have been deposited in the NCBI nucleotide sequence database and can be retrieved under the accession nos. MG669245 to MG669248, respectively.

4.3.8.8 Growth rate The growth rate of selected rhizobacteria was determined by assessing growth curves using a CytationTM 5 imaging reader. Firstly, bacteria were cultivated in tubes containing liquid KB medium and incubated at 28 °C on a rotary shaker at 150 rpm. Fresh bacterial cultures at the exponential growth phase were then diluted in KB broth to an initial OD600 of ~0.001, which was then filled in each well of a 96-well microplate. The microplates were incubated at 28 °C in the dark and assayed spectrophotometrically at 600 nm at the indicated time points (every 2 h) to establish the growth curves. The CFU was also analysed by serial dilutions with plate counts. The mean generation time was calculated from growth in the logarithmic growth phase (Stanier et al., 1970).

4.3.9 Protection against phytopathogens by rhizobacteria in Arabidopsis The potential of the four isolated bacteria to be utilized in reducing disease severity of phytopathogens on A. thaliana was investigated, according to Ingle and Roden (2014), with modifications. To determine the role of induced resistance as a mechanism in pathogenic control, sterilized Arabidopsis Col-0 seeds were treated with bacterial suspension of each isolate and then plated on ½ MS agar medium (in vitro) or sown in moistened Jiffy-7 peat pellets (soil assay). For the control, seeds were treated with sterilized 10 mM MgSO4. Plants were grown in a growth chamber maintained at 22°C with a 12-h photoperiod and 60 % relative humidity. The

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experiment was established following a completely randomized design with 16 plants per treatment. Each experiment was carried out three times, with comparable results.

4.3.9.1 Bacterial pathogenesis assay Overnight culture of Pst DC3000 in liquid LB medium was sub-cultured and grown at

28 °C until OD600 reached 0.8. Bacteria were centrifuged (8,000 g, 10 min) and re-suspended in

10 mM MgSO4. For soil assay, Arabidopsis leaves of 4-week-old plants were pressure infiltrated with a Pst inoculum (106 CFU mL-1) containing 0.004 % (v/v) Silwer L-77 to facilitate infiltration (Panchal et al., 2016), using a needeless 1 mL syringe. Mock inoculation was performed with a solution of 10 mM MgSO4, and this acted as a control. After infiltration, all plants were covered with plastic wrap to maintain high humidity. At 1 h after challenge inoculation and day 4, bacterial populations in leaf tissue were determined. There were three replicates, each with nine plants for each treatment. Briefly, inoculated leaves from three plants were pooled, weighed and surface sterilized with 70 % ethanol for 1 min, after which the leaves were rinsed in sterile water, blot dried on tissue paper, and homogenized in 0.8 % NaCl using a ceramic mortar and pestle. To titer the bacterial growth, a dilution series from 10-1 to 10-4 was plated individually on LB agar plates containing appropriate antibiotics. Colonies were counted after 2 d of incubation at 28 °C, and the number of CFU per mg of infected leaf tissue was determined. For in vitro assay, 14-d-old seedlings were inoculated with 2 μL of Pst suspension (109 CFU mL-1) in the center of the rosette. Seven days after inoculation, disease incidence was determined as the proportion of diseased leaves (necrotic or lesions surrounded by chlorotic tissue), expressed as the percentage of the total number of leaves per seedling. Colonization levels of Pst were determined as described above (Pieterse et al., 1996).

4.3.9.2 Protection against phytopathogenic fungi Botrytis cinerea B191 was stored as conidial suspension in 20 % glycerol at -80 °C and was reactivated on PDA medium amended with streptomycin (20 mg L-1) at 22 °C and a 12-h photoperiod. Spores were harvested in sterile water from a culture grown for 10 d, after which the suspension was filtered through glass wool to remove mycelium. Spores were then pelleted 3 times in ddH2O (2,000 g, 2 min) and re-suspended in potato dextrose broth (PDB) to a spore 154

density of 105 CFU mL-1 after counting using a hemocytometer. Fully developed leaves of 4-week-old Arabidopsis plants grown in peat pellets were drop-inoculated with 5 μL of B. cinerea spore suspension. Twelve plants for each treatment were incubated at 22 °C and covered with transparent plastic bags, to retain moisture and to facilitate disease development. It has been suggested that the inoculation drop should not dry out within the first 2 d, in order to allow for the formation of rot lesions (Muckenschnabel et al., 2002). Four days post challenge, the disease was assessed by measuring the necrosis diameter induced by B. cinerea spores, using the free Java image processing program ImageJ (NIH). In planta fungal growth was examined by simultaneous analysis of the transcript levels of B. cinerea actin gene (BcActin) with primers BcActin-1F (5’-TCCAAGCGTGGTATTCTTACCC-3’) and BcActin-1R (5’-TGGTGCTACACGAAGTTCGTTG-3’) and the Arabidopsis actin gene (AtActin2), an internal control, using primers AtActin2-1F (5’-GGCGATGAAGCTCAATCCAAACG-3’) and AtActin2-Ar (5’-GGTCACGACCAGCAAGATCAAGACG-3’) (Nie et al., 2017). Total RNA extracted from Arabidopsis leaves and cDNA synthesis determination was done according to section 4.3.4.5. Relative fungal growth was determined by ratios of BcActin/AtActin (Nie et al., 2017).

4.3.10 GFP-tagging for in Planta tracking 4.3.10.1 Preparation of competent cells Electro-competent cells of strains n and Y were prepared according to the standard protocol for Escherichia coli with some modifications. Briefly, 10 mL of KB broth was inoculated with 100 μL of overnight culture and grown to an OD600 of 0.5. The cells were first chilled on ice for 15 min then centrifuged and washed repeatedly with ice-cold 10 % sterile glycerol, and were then re-suspended in chilled water and pelleted and washed two times, after which the pellets were re-suspended in ice-cold 10 % glycerol, divided into 40 μL aliquots, and stored at -80 °C. For the Gram-positive strain L, chemo-competent cells were prepared following the method of Anagnostopoulos and Spizizen (1961), with some modifications. In brief, overnight culture of strain L in LB medium were diluted 50-fold in a growth medium (per 100 mL ddH2O:

0.54 g KH2PO4, 1.26 g K2HPO4·H2O, 0.09 g trisodium citrate·2H2O, 2 g glucose, 0.2 g potassium glutamate, 1.1 mg ferric-ammonium-citrate, 5 mg L-tryptophan, 36 mg MgSO4) and 155

grown at 37 °C (200 rpm) until OD600 reached 1.0, from which 400 μL aliquots were made and frozen at -80 °C .

4.3.10.2 GFP plasmid transformation Broad scale GFP tagging was attempted with a wide-host promoter plasmid, pDSK-GFPuv (obtained from Dr. Kiran Mysore at The S. R. Noble Foundation). This binary plasmid was used to transform chemically competent cells of E. coli DH5α (Invitrogen), which were then stored at -80 °C . Strains n and Y were electroporated with the plasmid extracted from E. coli via standard protocol, as described by Wang et al. (2007). Briefly, 40 μL of cold competent cells were mixed with 4 μL of plasmid DNA (250 ng μL-1) and then electroporated using an Eppendorf Multiporator®. For strain L, competent cells were transformed by mixing with 600 ng of plasmid DNA in a transformation mixture (per 100 mL ddH2O: 2.5 g yeast extract, 4 g casamino-acids, 0.025 g L-tryptophan) and incubating at 37 °C, 150 rpm for 1 h. Transformants were selected on kanamycin (35 μg mL-1) KB agar plates. The expression of GFP was analyzed by visualization under fluorescence microscopy.

4.3.10.3 Microscope analysis of root colonization The overnight cultures of GFP-tagged strains were diluted and grown in KB liquid medium at 28 °C to the exponential growth stage. Cells were collected by centrifuging at 9 -1 5,000 g and pellets washed twice in 10 mM MgSO4. The viable cells (10 CFU mL ) for each strain was determined by plant counting on KB agar plates containing 50 mg L-1 of kanamycin. To verify root-endophyte habit, that is, the ability to endophytically colonize in Arabidopsis, seeds of A. thaliana Col-0 were surface sterilized and treated with GFP-labelled bacteria as described above. Seeds treated with 10 mM MgSO4 were used as a negative control. After growing vertically in ½ MS agar medium for 14 d, plantlets were washed with ethanol and sterile water to remove unbound bacteria, after which the entirety of the roots were examined along the whole root material. Light and fluorescence microscopy were performed with a Leica fluorescent microscope and Northern Eclipse software. Observations of unlabeled seedlings provided information on the auto-fluorescence level of the sample. The experiment was repeated twice and

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a representative image of at least 20 roots of each treatment (inoculated or non-inoculated) was collected. Final pictures were generated through ImageJ (NIH) by merging the two channels.

4.3.10.4 Bacterial re-isolation in Arabidopsis Five individual Arabidopsis Col-0 plants treated with GFP-marked bacterial isolates (strains n, L and Y) or original bacterium (strain K) were harvested for collecting the shoot (strains n, L and Y) or root samples (strain K) 14 d post inoculation. Control seedlings were seed-treated with 10 mM MgSO4. Endophytic bacteria were isolated following the procedure described in section 3.3.2.2. Briefly, after appropriate sterilization to remove possible epiphytes, rosette or root samples were mixed and ground in sterile 0.8 % NaCl and vortexed for 10 min. Serial dilutions of the supernatant from rosette or root tissues were plated on KB medium supplemented with appropriate antibiotics. After incubation for 24-72 h at 28 °C, inoculated agar plates were observed and GFP fluorescence was detected under UV light. Colony-Forming Units (CFUs) on these plates were counted thereafter. Identity of each isolate was confirmed by using the EzTaxon-e server (http://eztaxon-e.ezbiocloud.net/; Yoon et al., 2017) based on 16S rDNA sequencing.

4.3.11 Statistical analysis For all experiments, the overall data were analyzed by one-way analysis of variance (ANOVA) and differences between control and bacteria-treatments were considered statistically significant at the P ≤ 0.05 level using Tukeys Honestly Significant Differences (HSD) test of the COSTAT® statistical software.

4.4 Results

4.4.1 Bacterial screening for growth promotion of Arabidopsis plants In order to find potential PGPR from newly isolated rhizobacteria (Chapter 3), we conducted an in vitro study to screen for the plant growth promoting effects of these isolated strains by measuring various physiological parameters of A. thaliana Col-0. A total of 98 isolates were selected based on their origins and taxonomical diversity for the initial screening on

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Arabidopsis (Fig. 4.1). While some isolates (e.g. B58, K2, Y9 and L1) showed no significant stimulation effect on plant growth, others promoted shoot growth (Fig. 4.1). Based on the above-ground fresh weight (FW) of Arabidopsis seedlings after 21-d of growth, 12 potential isolates (Fig. 4.1) which caused significant enhancement of shoot growth (from 1.3 to 2.5-fold increase over the control), were chosen for a second screening test. Only those that clearly showed plant-growth-promoting effects in two independent experiments were selected for further research. Four out of 12 selected isolates showed consistent and significant growth promoting effects compared to controls and other isolates. In vitro bioassays showed that treatment of seeds or root tips of Arabidopsis with rhizobacteria n, L, K and Y significantly increased seedling growth relative to controls after 21 d of incubation (Fig 4.2). For example, the rosette fresh weight (root-tip treated) was significantly increased (by approximately 20, 43, 30, and 60 %, respectively) by strains n, L, K and Y, respectively, as compared to control plants that were not exposed to the bacteria. Inoculation of root-tip by strains L, K and Y, respectively, led to a 77 % increase in the total leaf surface area versus the mock-inoculated control.

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Fig. 4.1 Effects of seed bacterization on Arabidopsis Col-0 seedling growth. Results are expressed as shoot fresh weight relative to untreated controls. Pictures are phenotype of Col-0 plants treated with the rhizobacteria after a 3-week growing period on vertical agar plates. Asterisks indicate statistically significant differences from the control treatment (* P < 0.05).

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SEED ROOT TIP

Control K

Fig. 4.2 Effects of seed or root tip treatment of Arabidopsis wild-type Col-0 with rhizobacteria on early seedling growth. The total leaf surface area (a, b) and rosette fresh weight (c, d) of Col-0 Arabidopsis plants inoculated with the 4 selected rhizobacteria strains after a 3-week growing period on agar plates. Pictures are phenotype of Col-0 plants treated with strain K after a 3-week growing period on vertical agar plates. Different letters indicate statistically significant differences between treatments (P < 0.05). Bars indicate mean ± standard error from three independent experiments, with 20 seedlings per treatment.

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4.4.2 Phenotyping of the strains and their biocontrol and PGP traits The 4 representative endophytic isolates (n, L, K and Y) were selected for the characterization described in this study based on their best appearance in plant growth-promoting effects in Arabidopsis on Petri plates revealed by variance (ANOVA) and Tukey’s test (P < 0.05). They were subjected to numerous tests using conventional morphological, physiological, and phenotypic techniques or API systems when possible, along with 16S rDNA identification.

4.4.2.1 Molecular taxonomy of candidate bacterial strains by 16S rDNA sequencing To determine the relatedness of strains at the genetic level, the selected bacterial isolates n, L, K and Y were characterized by partial 16S rDNA sequencing. Results were from the comparison of the nucleotide sequences with deposited sequences in GenBank using the Mega BLAST algorithm. Similarities of the 16S rDNA sequences revealed that strains n, L and Y were from the phylum Proteobacteria and were from the families Pseudomonadaceae, Bacillaceae and Rhizobiaceae, respectively. Strain n showed high identity with Pseudomonas and clustered with P. koreensis Ps 9-14T, based on 99.8 % similarity, and strain K was from the phylum Bacterioidetes and from the family Sphingobacteriaceae and grouped with Mucilaginibacter lappiensis ATCC BAA-1855T, with identity value > 97 %. Strain L clustered with Bacillus mobilis 0711P9-1T and strain Y with Rhizobium jaguaris CCGE525T. These later two strains had 100 % identity with the 16S rRNA gene sequences of their corresponding type strain (Fig 4.3). The identification of selected endophytic rhizobacteria encoded n, L, K and Y most closely resembled Pseudomonas sp., Bacillus sp., Mucilaginibacter sp. (97.2 % confidence) and Rhizobium sp., respectively (summarized in Table 4.3). A neighbour-joining phylogenetic tree of the isolated strains n and K was constructed by combining the sequences of the specific strain with its closest relatives (Fig. 4.3). The data from phylogenetic tree suggested their relatedness with several other strains of Pseudomonas, Bacillus, Mucilaginibacter and Rhizobium sp., respectively (Fig 4.3). All the four isolates showed 97 % to 100 % similarities with type strains of two or more species within the corresponding genera (data not shown), thus the species affiliation for all the 4 isolated strains was not defined.

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Table 4.3. Molecular identification of rhizobacterial isolates using 16S rDNA as query sequences.

Strain Closest type strain (accession Closet relative (NCBI) % Similarity (EzTaxon) Origin code number) Pseudomonas koreensis Pseudomonas sp. 99.76 Phalaris arundinacea n (AF468452) L Bacillus sp. Bacillus mobilis (KJ812449) 100.00 Solanum dulcamara Sphingobacteriaceae bacterium Mucilaginibacter lappiensis 97.21 Scorzoneroides autumnalis K kmd_018 (jgi.1095764) Y Rhizobium sp. Rhizobium jaguaris (JX855169) 100.00 Glycine max

163 Table 4.4. In vitro biochemical characterization of bacterial isolates on the plant growth promotion traits.

Antimicrobial activity 3 ACC deaminase activity P solubilization Siderphores Nitrogen HCN Ammonia Strains IAA (mg L-1) 1 Pseudomonas Botrytis (μmol mg-1 h-1) 1 (mm) 2 on CAS agar fixation production production syringae cinerea n 0.79a 8.10a 5.67a +++ +++ +++ ++ 2.00b 5.00a L 0.10b 2.01b 2.83b + + − +++ 6.33a 2.67b K 0.83a 0.98c − − − − − − − Y 0.86a 4.09b 2.17b − ++ − − − − IAA, indole acetic acid; HCN, Hydrogen cyanide Means followed by different letters within a row are significantly different (P ≤ 0.05). +, low activity; ++, moderate activity; +++, strong activity; −, no activity. 1 Values are presented as means of 6 replicates. 2 Means of diameter of solubilization measured on National Botanical Research Institute’s phosphate (NBRIP) agar plates (average of 6 replicates). 3 Values are the means of diameter of inhibition zones (mm) (average of 4 replicates).

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Fig. 4.3 Phylogenetic tree generated using neighbor-joining algorithm based on 16S rRNA gene sequence showing inter-relationship of representative strains with the closely related type strains (accession number in parentheses). The numbers at branch nodes are percentages of bootstrap support based on 1000 resamplings. Bar, 0.01 or 0.05 substitutions per nucleotide position.

165 4.4.2.2 Plant-growth-promotion traits of selected bacteria The four selected strains were thoroughly examined for several traits that are often associated with biocontrol and plant growth promotion.

Nitrogen fixation Two methods were deployed to evaluate the ability of the selected strains to fix atmospheric nitrogen: traditional microbiological method (Nfb medium) coupled to a molecular probe-directed approach (PCR specific for the nifH gene). The Nfb medium methodology revealed that strains n, L and Y probably had nitrogen fixing capacity in the solid culture conditions used, since the color of medium was changed from green to blue (Fig. 4.4a). It has been shown that nitrogen-fixing (nif) genes include subunit genes nifH, nifD, and nifK and various additional genes (Dai et al., 2014); whilst the nifH (encoding the nitrogenase reductase subunit) is the most widely sequenced and utilized marker for nitrogen fixation. Here, we used four sets of universal PCR primers (Table 4.2) to target the nifH gene as a proxy for potential nitrogen fixation activity in the selected rhizobacteria. 19F/388R was preferred over the other three pairs because these latter primers showed ineffective for non-specific products on gels. The expected fragment, of about 390 bp, was obtained from isolated strains n, L and Y, using primers 19F and 388R, further confirming the nitrogen-fixing capacity of these three isolates (Fig. 4.4b). The amplified fragments of nifH gene were cloned, sequenced, analyzed, and identified by BlastN. They were found to show 92 to 93 % sequence similarity with other nifH sequences available in the NCBI GenBank.

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Fig. 4.4 Nitrogen-fixing capacity of selected strains as determined by (a) Nfb solid medium (inset; strain Y) and (b) PCR-amplification using primer pairs Ueda19F and 388R of nifH gene from genomic DNA of selected strains. Electrophoretic patterns of PCR products of the nifH gene showed that 3 out of the 4 selected strains harbor the nitrogen-fixing gene. Lanes: 1-4, n, L, Y and K respectively; (−), negative control (water); M, Fast DNA Ladder.

Phosphorous solubilization In order to evaluate whether the four selected strains can solubilize the insoluble form of

P, simple tri-calcium phosphate (Ca3(PO4)2; TCP) was chosen and tested in NBRIP agar medium (Nautiyal, 1999). After a 2-week incubation, clear dissolution halos were observed around the

167 colonies of strains n, L and Y (Fig. 4.5a). These clear zones indicated the excretion of enzymes or organic acids into the surrounding medium that solubilize TCP. Strain n (Pseudomonas sp.) was capable of higher amounts of solubilisation (5.67 mm) than were strains L (Bacillus sp.) and Y (Rhizobium sp.) (2.83 and 2.17 mm, respectively) (Table 4.4). While strain K showed no phosphate solubilisation activity on NBRIP agar plates. Since the buffering capacity of soil could limit the ability of bacteria to solubilize soil P, phosphate agar (Gyaneshwar et al., 1998), which was buffered with 100 mM Tris-HCl (pH 8.0) to mimic the buffering capacity of alkaline vertisols, was chosen to ascertain the efficacy of PSB. The PS phenotype was indicated by formation of a red zone around the microbial colony, which indicate that the bacteria secreted sufficient organic acids to overcome the buffering, resulting in color change of the pH indicator Methyl Red, usually with a drop in the medium pH from 8.0 to < 5.0. Using this medium, only strain n, a Pseudomonas, showed a pink coloration zone (Fig. 4.5b). The quantity of inorganic phosphate solubilisation was determined for a 14-d period, utilizing liquid NBRIP broth containing 1000 μg P mL-1 in the form of TCP (Fig. 4.6). Strain n (Pseudomonas sp.) showed significantly (P < 0.001) higher amount of solubilized P (maximized at 350 μg mL-1 on the 7th d of incubation) throughout the entire period; whilst strains L (Bacillus sp.) and Y (Rhizobium sp.) showed comparable but much lower activities, with the highest amount of solubilized P (69 and 63 μg mL-1) on the 14th d of incubation. Strain K showed very low activity, which was not detectable on agar plates.

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Fig. 4.5 Representative P-solubilizing phenotype of strain n on different screening media. (a) NBRIP agar (b) Buffered medium. The ability to solubilize and use Ca3(PO4)2 as a sole source of phosphate was evident as a clearing zone around the colony of strain n.

n 400 L K Y

)

-1 300

200

P solubilization L (mg 100

0 0 2 4 6 8 10 12 14

Incubation period (d)

Fig. 4.6 Concentration of solubilized phosphate by strains n, L, K, and Y during incubation in NBRIP broth at different time intervals.

Indole-3-acetic acid (IAA) production IAA is the major auxin in plants and plays a multifaceted role in many important physiological processes, such as cell elongation and cell division, tissue differentiation and responses to light (Patten and Glick, 2002). Exogenous IAA can also stimulate the growth of plant root systems, including root branching and lateral root development (Patten and Glick, 2002). Thus, in vitro auxin production by selected isolates was investigated colorimetrically as IAA equivalents in the presence and absence of L-tryptophan. When grown with 500 mg mL-1 of

169 L-tryptophan in KB liquid medium, strain n, a Pseudomonas, produced an average of 0.79 μg mL-1 of IAA (or its intermediates), which was comparable to those produced by strain K and Y (Table 4.4). The Bacillus sp. (strain L) produced significantly lesser amounts of IAA (0.1 μg mL-1) than strains n, K and Y. The production of IAA (or its intermediates) produced by the selected strains in media without L-tryptophan was significantly less (P<0.01) (data not shown).

ACC deaminase activity The ability to use ACC as a sole nitrogen source indicates the activity of the enzyme ACCD. The ACC-deaminase activity in the four selected strains were determined quantitatively by monitoring the amount of ammonia generated by the hydrolysis of ACC. It was observed that rhizobacterial isolate n had maximum ACCD activity (8.10 µmol mg-1 h-1) while strain K exhibited lowest ACCD activity (0.98 µmol mg-1 h-1) (Table 4.4).

Production of ammonia, siderophore and HCN The ability of bacteria to sequester iron confers a competitive advantage. Only strains n and L formed a distinct orange halo around the colonies on the CAS agar medium indicative of hydroxamate type siderophore production (Fig. 4.7). The Pseudomonas sp. (strain n) exhibited much higher levels of siderophore activity (+++) than strain L (Bacillus sp.), which showed only a minor production of siderophore (+) (Table 4.4). Moreover, only strain n showed positive results for HCN production, as shown by the development of color of the filter paper from yellow to dark orange (Fig. 4.7). The ability of selected rhizobacteria to produce ammonia was assessed with Nessler reagent. All the strains, except for strain K, were capable of producing ammonia, as shown by the development of a color change from light yellow to orange (Fig. 4.7). Based on the intensity of orange color, the amount of ammonia produced from highest to lowest was as follows: strain L (+++), followed by strain n (++) and finally the Rhizobium strain Y (+).

170

Fig. 4.7 Representative pictures showing the production of (a) siderophore, (b) HCN, and (c) ammonia by selected strains. The black arrows indicate the halo zone around the colonies.

Antagonistic activity against microbial pathogens The antagonism assay was performed using regular dual culture technique for both pathogenic bacterium and fungus. Each isolated bacterium was tested in three independent replicates. Preliminary tests showed that all the four selected bacterial isolates exhibited (visually) good growth in PDA. Strains n and L showed clear zones of growth suppression of B. cinerea B191 (Fig. 4.8a). The inhibition with strain n was more prominent than that of strain L, with an inhibition zones of 5 and 2.67 mm, respectively (Table 4.4). The inhibition of mycelial growth was stable, indicating that strains n and L released compounds that interfered with the growth of this fungus. Conversely, strains K and Y showed no inhibition effects, such that in their presence

171 B. cinerea manifested growth that was not different from control plates (uninoculated with rhizobacteria). Of the tested bacteria, strains n and L demonstrated growth antagonism towards P. syringae DC3000, with an inhibition zone of 2 and 6.33 mm, respectively (Table 4.4). Strains K and Y exhibited no antagonistic activity against this phytopathogen; in their presence, P. syringae DC3000 manifested confluent growth similar to that in control medium plates (without rhizobacterial inoculation) (Fig. 4.8b).

Fig. 4.8 Antagonism assessments of PGPR strains against selected phytopathogens using dual-culture techinque. (a) Bacteria were spot-applied on side of the plate and Botrytis cinerea B191 was placed on the other side as agar plugs so that the mycelium and the bacteria were not in direct contact. The distance between tested bacteria with fungal hyphae shows inhibition of fungal growth by strains n and L at 5 d post-inoculation (dpi). (b) Pseudomonas syringae pv. tomato (Pst) DC3000 was streaked on KB plates, after which tested strains were spot-inoculated to the middle of the plate. The zones of inhibition indicated antagonistic activity of strains n and L against Pst at 4 dpi. Tests were done three times with six replicates; only representative plates are shown.

172

4.4.2.3 Primary characterization of selected bacteria The primary characterization of the 4 selected isolates (strains n, L, K and Y) was done by specific microbiological and biochemical tests. There were clear visual differences in morphology among the 4 strains when grown on KB agar medium. Strain L, a Bacillus, formed white opaque, circular, and dry colonies with a flat surface and undulate margins, while the appearance of the colonies of strain n, a Pseudomonas, was translucent, intensely yellow-colored, mucoid, and round with high convex elevation and complete edges. White, moist, translucent circular colonies with a complete edges and convex elevation were observed for strains K and Y. Strain K underwent a color change as the cultures aged, with an initial color of off-white and light pinkish after 2-3 d. Both strains n and L grew very fast on KB agar plates at 28 °C ; while strain Y grew fast; the colonies were only visible after 24 h. Strain K was a slow-growing rhizobacterium. A general microscopic view of the four isolates showed them to be rod-shaped cells. All 4 strains were positive for catalase and negative for oxidase. Three of the tested isolates, strains n, K and Y, were found to be Gram-negative and positive for Voges-Proskauer (VP) reaction (acetoin production), while strain L was Gram-positive and showed positive tests for nitrate reductase, amylase and gelatinase. Of all the bacteria, only strains L and Y were motile. Strain n tested positive for protease, arginine dihydrolase and tryptophane deaminase, whereas it tested negative for cellulase and urease production. Strain L was positive for cellulase, protease, arginine dihydrolase and tryptophan deaminase production and negative for urease. Strain K was negative for urease and arginine dihydrolase and positive for tryptophan deaminase, cellulase and protease production. For strain Y, there were positive reactions for cellulase and urease, but negative reactions for protease, arginine dihydrolase and tryptophan deaminase production. All these morphological and biochemical characteristics are tabulated (Table 4.5). All 4 of the strains grew well on KB, LB, tryptic soy agar (TSA) and R2A agar medium, but only strain n showed growth on MacConkey agar (all from Difco) with a pink colony color, indicating that strain n is a lactose fermenting Pseudomonas. Only strains K and Y showed a mucoid phenotype on RVC-agar medium, indicating potential EPS production by the two isolates. The selected strains used a large variety of carbohydrates as sole carbon and energy sources. None of the isolated bacteria changed the color (red) in control wells lacking carbon

173 sources. Therefore, any color change observed was attributed to assimilation of the specific carbohydrate. Carbohydrate utilization efficacy of the isolates has been summarized in Table 4.6. Carbohydrate utilization pattern of the 4 tested strains varied extensively. Specifically, Pseudomonas strain n produced acid from glycerol, arabinose, xylose, galactose, glucose, mannose, mannitol, trehalose, fucose, ribose, melibiose, caprate, adipate, maliate, and dextrose. Rhizobium strain Y assimilated arabinose, xylose, galactose, mannitol, adonitol, erythritol, fucose, adipate, and maliate. Strain L was found to utilize glucose, maltose, trehalose, ribose, adipate, naliate, and only strain L was positive for the utilization of arbutin, salicin, starch, glycogen, N-acetylglucosamine, and phenyl-acetate. For strain K, acid was produced from fermentation of galactose, glucose, mannose, maltose, trehalose, melibiose, sucrose, cellobiose, and dextrose. All strains acidified fructose. All except for strain n were found to hydrolyze esculin, while only strain L was negative with citrate and malonate utilization. Only strain K produced acid from raffinose. Thus, in total, strain n, followed by L, showed utilization of the broadest range of carbon sources (58 % of those tested) (Table 4.6).

Table 4.5. Some phenotypic characteristics of isolated rhizobacteria

Strains Characteristic n L K Y

Colony color on KB yellow milk white light pink white Grams’ reaction N P N N Voges proskauer + + + − Motility + + − +

Oxygen requirement obligate aerobe facultative anaerobe aerobe microaerophiles Catalase + + w + Oxidase − − − − Urease − − − + Fermentation/oxidation + − − − (glucose)

174 Nitrates reduction to nitrites − + − − Arginine dihydrolase + + − − Tryptophan deaminase + + + −

Hydrolysis of: Starch (amylase) − + − − CMC (cellulase) − + − + Casein (protease) + + + − Gelatin (gelatinase) − + − −

Growth tolerance: 37 °C + + − + 40 °C − + − − NaCl 6% 7% 1.2% 1.5% pH range 5.0-10.0 5.0-10.0 5.0-8.0 5.0-8.0

Enzyme activities (API ZYM):

Alkaline phosphatase − + + − Esterase lipase + + − − Leucine-arylamidase + + + + Valine arylamidase + − + − Cystin arylamidase − − + − Acid phosphatase − + + + Naphtol-AS-BI-phosphohyd + − − − rolase α-galactosidase − − + − β-galactosidase − − + + α-glucosidase − − + + β-glucosidase − + + + N-acetyl-β-glucosaminidase − − + −

175 α-fucosidase − − + −

+: indicates a positive reaction (color change due to substrates consumption) while − indicates a negative reaction. N: negative Gram reaction; P: positive Gram reaction.

Table 4.6. Carbon source utilization by isolated rhizobacteria a

Acid fermentation Strains n L K Y Glycerol + − − − L-Arabinose + − − + D-Xylose + − − + D-Galactose + − + + D-Glucose + + + − D-Fructose + + + + D-Mannose + − + −

D-Mannitol + − − + Arbutin − + − − Esculin hydrolysis − + + + Salicin − + − − Maltose − + + − Trehalose + + + − Adonitol − − − + Erythritol − − − + Starch − + − − Glycogen − + − − D-Fucose + − − + Ribose + + − − N-Acetylglucosamine − + − − Melibiose + − + − Sucrose − − + − D-Cellobiose − − + −

176 Caprate + − − − Adipate + + − + Maliate + + − + Phenyl-acetate − + − − Citrate utlization + + − + Dextrose + + + − Raffinose − − + − Malonate utilization + + − + a This experiment was tested in API CH strips and HiMedia kits, respectively. The color change was observed after 1-3 days of incubation at 28 °C . +: positive result corresponds to acidification revealed by the phenol red indicator changing to yellow.

The selected isolates were also tested for extracellular enzymatic profiles involved in the breakdown of peptides, phosphomonoesters, lipids, mucopolysaccharides, polysaccharides, chitin, cellulose, starch, and galactans, using the API ZYM system. The API ZYM activities evaluated for the four isolated rhizobacteria are shown in Table 4.5. All four strains exhibited strong leucine arylamidase activity. The remaining tests produced differentiating patterns among the four genera examined. All but strain n showed strong acid phosphatase activity, but only strain n was found to hydrolyze the substrate for phosphoamidase. Strain K (Table 4.5) possessed a wide range of enzyme activities; it was positive for galactosidase, N-Acetyl-β-glucosaminidase, and α-fucosidase, and was found to efficiently hydrolyze the glucosidase. Tolerance of abiotic stressors by the selected strains was also studied. Strains n, L, K and Y were restricted to KB medium at different temperature, pH, and salt levels. High concentrations of NaCl repressed bacterial growth, where strains n and L tolerated higher concentrations of NaCl than K and Y (Table 4.5). All tested strains showed high tolerances to NaCl, from 12 (strain K) to 70 (strain L) g L-1. Strain L tolerated up to 40 °C , but showed no growth at 42 °C . At 37 °C , strain K showed no growth on KB agar. The sensitivity of selected isolates to extreme pHs was also examined. All the isolates grew well at pHs of 5, 6, 7 and 8, but only strains n and L were able to grow at pHs 9 and 10. Overall, strain L was the most tolerant to these stressors.

177 All in all, characterization of biochemical/physiological traits (Tables 4.5 and 4.6) confirmed the classification of these strains, but further assignment, to specific species, was not possible.

4.4.2.4 Growth curves and growth kinetics To establish the growth rate for the four selected bacterial strains, growth curves were developed (Fig. 4.9). The Pseudomonas sp. and Bacillus sp. (strains n and L) grew relatively fast at 28 °C in KB medium. When comparing the growth rates, strains K and Y exhibited a slow-growth and intermediate-growth phenotype, respectively (Fig. 4.9). The mean growth rates noted for strains n, L, K and Y were 0.6716, 0.1181, 0.0146 and 0.0293 generations per hour, respectively.

n 2.0 L Y K

1.5

1.0

Absorbance (600 nm) Absorbance .5

0.0

0 20 40 60 80 100 120 Incubation time (h)

Fig. 4.9 Growth curves of strains n, L, K and Y based on absorbance measured at a wavelength of 600 nm. Strains were grown on KB medium under aerobic conditions at 28 °C, with agitation at 150 rpm.

178 4.4.2.5 Minimal inhibitory concentration (MIC) test In order to test for intrinstic antibiotic resistance, fresh cultures of the 4 isolates were streaked on KB agar medium and the intrinsic antibiotic resistance was determined using Etest strips, which created an antimicrobial gradient on heavily inoculated agar plate and resulted in a clear elliptical zone of growth inhibition. Data in Table 4.7 indicated that strains n, L and Y were highly sensitive to kanamycin while strain K was very resistant (up to 900 μg mL-1). The same trend occurred for tobramycin, gentamicin and tetracycline in that strain K demonstrated the greatest resistance, while the other bacteria were sensitive to these antiobiotics. Regarding ampicillin, the Rhizobium sp. (strain Y) was very sensitive, while the other bacteria showed strong resistance (> 256 μg mL-1). For erythromycin and chloramphenicol, strain n showed greater resistance than strain K, while strains n and Y were susceptible to them. For vancomycin, strains n and Y demonstrated substantial resistance while strains L and K were sensitive. Overall, there was a mixed pattern observed for all the tested antibiotics (Table 4.7). The antibiotic sensitivity profile can aid in identification of bacterial strains, as well as avoid contamination or strain misuse (Zarpelon et al., 2016).

Table 4.7. Susceptibility (MICs) of bacterial isolates to various antibiotics (μg mL-1) as determined by Etest method a

Treatment Isolate n L K Y

Ampicillin >256 >256 >256 0.064

Erythromycin >256 3 24 0.5

Kanamycin 1.5 2.5 >256 0.38

Chloramphenicol >256 2 16 12

Gentamicin 0.38 0.38 16 0.125

Streptomycin 3 1.5 48 8

Tetracycline 3 0.38 16 0.75

Rifampicin 1.5 0.38 >32 8

179 Treatment Isolate n L K Y

Vancomycin >256 2 24 >256

Tobramycin 1.0 1.5 >256 0.125

a Experiments were repeated three times with same results.

4.4.3 Spatial distribution of rhizobacteria in Arabidopsis Fusing reporter genes, such as the green fluorescent protein (GFP), to plasmids which are used to transform selected bacteria, has considerably advanced our knowledge of the localization of rhizobacteria within plants. In vivo microscopy gives more precise insights into the cellular localization of rhizobacteria. Thus, in this study, the pattern of root and shoot colonization (Col-0) was investigated by using recombinant bacterial strains expressing GFP and fluorescent microscopy, in order to evaluate whether the selected bacterial strains originally isolated from the interiors of roots of different crop and wild plants could colonize the internal tissues of the model plant Arabidopsis. Attempts were made to GFP tag all four isolates; unfortunately, only three isolates were successfully transformed with the GFP-harbouring plasmid (pDSK-GFPuv) (Fig. 4.10). Though GFPuv produces 45-fold brighter green fluorescence in E. coli than other GFP variants, and pDSK-GFPuv is shown to express GFPuv at high levels (Wang et al., 2007), we could clearly see that transformed strain L (Bacillus sp.) showed apparently lower green fluorescence than strains n and Y; the underlying reason is not known. When Arabidopsis was treated with wild-type strains n, L, and Y, no fluorescent signal was detected throughout the entire root system (data not shown). Two weeks after seed treatment with bacterial suspension of GFP-expressing strains n, L and Y, seedlings exhibited GFP signals in various tissues (i.e. apoplastic regions and the cytosolic compartment) of the maturation zone of the roots (Fig. 4.11), demonstrating that these species are root associated. Extensive screenings of roots from 20 seedlings inoculated with GFP-carrying bacteria n and Y revealed abundant individual bacteria and/or bacterial colonies inside the roots (Fig. 4.11), thus these strains can colonize inside the roots of Arabidopsis and become true endophytes. It should be noted that, only a small number of GFP-labeled cells was observed inside the roots of Arabidopsis treated with GFP-labelled strain L, indicating that strain L only sparsely colonized inside the roots.

180 Epiphytic growth of all four strains along root tissues after one week of co-cultivation was visualized on agar medium (data not shown). Re-isolation experiments indicated that there was transport throughout the entire plantlets and bacterial occurrence of strains n, K and Y appeared to be in roots (Fig. 4.11). Plate culturing confirmed that these four strains were present in both roots and leaves; with the endosphere of Arabidopsis roots harboring approximately 103 more bacteria than the rosette endosphere (data not shown). This again confirmed the endosymbiotic origin of the four selected rhizobacteria from their host plants and demonstrated their ability to colonize the interior parts of both roots and shoots of Arabidopsis seedlings and then exhibit endospheric properties.

181

Fig. 4.10 Visualization of rhizobacteria labeled with pDSK-GFPuv plasmid. The 3 isolates (Pseudomonas sp. (n), Bacillus sp. (L) and Rhizobium sp. (Y)) indicated were successfully tagged with GFP (pDSK-GFPuv, KanR) (out of 4 isolates attempted). Transformed bacteria were cultured in liquid KB medium supplemented with kanamycin overnight. Pictures show GFP-labeled bacteria under a fluorescent microscope (B) and GFP-labeled bacteria in white light (A).

182

Fig. 4.11 Root persistence of recombinant rhizobacteria in roots of Arabidopsis wild-type Col-0. The transformed bacteria were inoculated onto Arabidopsis seeds before plating on ½ MS agar medium. Fluorescent microscope of a root colonized endophytically by GFP-tagged bacteria. Roots were surfaced-sterilized with ethanol and sterile water before being observed. Cells of (a) Pseudomonas sp. (strain n), (b) Bacillus sp. (strain L), and (c) Rhizobium sp. (strain Y) were observed inside of roots 14 days after primed with selected strains: focus at maturation zone of the root as seen by the focused xylem strand. (d) control roots, from uninoculated seedlings. For each image set, the first panel (1) refers to phase contrast microscopy of Arabidopsis root; the second panel (2) shows the corresponding image acquired under fluorescent light; the third (3) results from the merge of both images (1st and 2nd).

183 4.4.4 In vitro response of treated Arabidopsis to salt and osmotic stress Firstly, to check whether bacterial treatments affected the germination activities of Arabidopsis, the seeds were germinated on the ½ MS medium containing varying concentrations of NaCl or mannitol and the germination rates were compared. There were no significant differences among the control and treated plants when germinating under normal conditions (Fig. 4.12). In the presence of exogenous NaCl or mannitol, the germination of both control and bacterium-treated seeds was inhibited significantly. For example, at the 4th d after sowing on ½ MS agar medium supplemented with 150 mM NaCl, only approximately 10 % of the control and treated seeds germinated (Fig. 4.12). We also observed that the germination of treated seeds was comparable to the controls under stress conditions (Fig. 4.12). These results indicated that rhizobacterial treatment of Arabidopsis did not affect plants at the seed germination stage. Secondly, growth variables such as root length, chlorophyll content, and fresh weight were measured and analyzed to observe the effect of biopriming on the early seedling growth of Arabidopsis plants under salt stress. When Arabidopsis seeds were primed with bacterial strains n, L, K and Y, the seedlings grew better than controls during the first 14-d of growth (data not shown), which is consistant with our earlier study (section 4.4.1) that the total rosette fresh weight was increased by 20-50 % following inoculation with the selected four strains. The growth of both treated and control seedlings was significantly repressed after a 14-d growth on ½ MS media containing 100 mM NaCl, but the growth of the treated plants was much less inhibited compared to the controls (Fig. 4.13). Inoculation with all four strains led to 23.6 to 122.36 % increases in whole plant biomass (Fig 4.14a). The average lateral root lengths of the seedlings treated with strains n, L, K and Y were significantly increased under normal conditions (Fig. 4.13d), showing 1.36-, 2.71-, 3.61-, and 2.33-fold longer than the control, respectively (Fig. 4.13d); while under 100 mM NaCl treatment, the lateral root length of strains L, K and Y was significantly longer than the control. Bacterial inoculation resulted in no significant change in the primary root length under either normal or salinity stress conditions (Fig. 4.13c). Rosette fresh weight was maximum where the inoculation of strains L, K and Y was employed (Fig.4.13b). Likewise, we observed that NaCl caused obvious decreases in total chlorophyll in control plants as compared to that of treated seedlings (Fig. 4.13e), but total chlorophyll was increased significantly in plants inoculated with strains L, K and Y compared to non-inoculated salinized plants. Overall, these results suggested that, in our culture conditions, bacterial treatments

184 enhanced tolerance to salt stress and stimulated the lateral root growth rate during the early seedling growth of Arabidopsis on agar plates.

Fig. 4.12 Comparison of germination rate of Arabidopsis under normal, salt, and osmotic stress conditions. The control and treated seeds were germinated on ½ MS agar plates (a) supplemented with 100 mM (b) and 150 mM (c) of NaCl and mannitol (100 and 300 mM) (d), respectively. The germinated seeds were scored regularly up to 10 d. The results are mean ± standard error from three independent experiments, with 100 seeds per treatment.

185

Fig. 4.13 The growth of Arabidopsis plants under salt stress conditions. Surface-sterilized seeds were treated with bacteria and sown onto ½ MS agar medium supplemented with 100 mM NaCl. Total fresh weight (a), rosette fresh weight (b), and root length (c, d) of Arabidopsis seedlings were measured after 14 d of co-cultivation with the indicated bacterial treatment. (e) Assay of chlorophyll content in leaves of bacteria-treated seedlings. (f) Phenotype of Col-0 plants treated with the rhizobacteria after a 3-week growing period vertically. For root length measurement, the average value for 20 plantlets per treatment was determined for each replicate. Vertical bars represent the mean and error bars represent standard error based on three independent experiments. Different letters indicate statistically significant differences (P < 0.05).

186

Fig. 4.14 The growth of Arabidopsis plants under osmotic stress. Surface-sterilized seeds were treated with bacteria and sown on ½ MS agar medium supplemented with 100 mM mannitol. Bacterial treatment triggered enhancement of plant growth as characterized by (A and B) total fresh weight and rosette fresh weight and (C) total leaf surface area per plant. (D) Representative images of plants with bacterial treatment or 10 mM MgSO4 as control. The results are mean ± standard error from three independent experiments. Different letters indicate statistically significant differences (P < 0.05).

187 As it is known that NaCl imposes both ion toxicity and osmotic stress in plants, here we used mannitol in ½ MS medium to check for the osmotic stress effect on seedling growth in Arabidopsis. At the early stages of observation after 14 d of growth, we found that the growth of control seedlings was more hampered than bioprimed ones (Fig. 4.14). The average leaf area of seedlings treated with strains L, K and Y were 1.3, 1.8 and 2.2-fold higher than the control under 100 mM mannitol, respectively (Fig. 4.14c). Another measurement showed that seed treatment with strains K and Y not only resulted in higher leaf surface areas but also caused higher seedling fresh weight as compared to controls (Fig. 4.14), indicating that plants treated with strains K and Y showed better adaptation to the dehydration stress.

4.4.5 Possible effect of volatile chemicals produced rhizobacteria During the last two decades, certain volatile compounds have been found to play a role in plant morphogenetic processes and subsequent growth and development (Ryu et al., 2003; Gutiérrez-Luna et al., 2010). Thus, in this study, divided Petri dishes were used to determine whether or not microbial volatile compounds have an important role in plant growth promotion in Arabidopsis. Rosette fresh weight of 21-d-old seedlings was measured after inoculation of the bacterial strains for 16 d on the other side of the divided plate. Our results showed no significant differences in fresh biomass production in seedlings inoculated with the four selected isolates and controls (10 mM MgSO4 and E. coli) (data not shown). Since plants and bacteria were grown separately in the Petri dish, it is not possible that bacterial volatiles are involved in plant growth promotion by these rhizobacteria.

4.4.6 Bacteria confer facilitated growth and increased salt resistance in planta Previous results showed that pre-incubation with selected rhizobacteria significantly increased early seedling growth on MS agar plates, with full nutrition, under sterile conditions (section 4.4.3). However, the in vitro assay is not a good measure of long-term growth determinations as the seedlings are within an enclosed environment, which could induce other stresses over the long term. Thus, the effect of bacteria on plants’ long-term growth in pots was tested. In brief, 10-d-old seedlings of Arabidopsis Col-0 were inoculated with one of the four selected rhizobacteria and were then grown for a 28-d period in a growth chamber. The four tested strains presented different PGP abilities and nearly all of them had considerable impact on

188 different growth parameters of A. thaliana compared with the negative control. Growth promotion was first assayed using sterilized potting mixture. Under stress-free conditions, the lowest amount of rosette fresh weight was found in the control treatment, while the total leaf surface area was clearly increased (by ca. 57 %) by bacteria L, K and Y, as compared to control plants (data not shown). Moreover, the rosette fresh and dry weights were also significantly increased by the same treatments (Fig. 4.15a). These results were in line with those from the in vitro experiments (section 4.4.1). Secondly, plant growth was assayed on non-sterile peat substrates. Again, the bacteria significantly enhanced growth of 28-d-old plants in terms of rosette fresh and dry weight as well as total leaf area (data not shown). To study whether the stimulated growth after 4 weeks is reflected in the carbohydrate and protein status of the plants, total soluble sugar and total protein levels in leaves of soil-grown Arabdidopsis Col-0 plants seed-treated with selected strains were measured. The content of total soluble sugar and total protein was not altered upon bacterization with all four strains (data not shown). As for long-term effects, bacteria significantly enhanced growth of 9-week-old plants. Stalk length of Arabidopsis plants exposed to bacterial strains L, K and Y was significantly increased, by 51 % after 63 days, as were rosette diameters, by 3.4 % (data not shown). Taken together, these results demonstrate that bacteria efficiently boosted the vegetative growth of Arabidopsis Col-0 on various substrates under both sterile and non-sterile conditions (stress-free).

189

Fig. 4.15 The effect of bacteria on growth and salt tolerance of Arabidopsis. The average rosette fresh weight (a), dry weight (b), chlorophyll content (c) of Col-0 Arabidopsis plants root-inoculated with the rhizobacteria strains after a 4-week growing period in soil (0, 200 mM NaCl). (d) Reduction of DPPH by Arabidopsis seedling extracts. Protein extracted from the leaves of 4-week-old plants were used in DPPH scavenging activity. (e) Representative plants grown in Peat pellet (non-sterile) treated or untreated with bacteria, and (f) in sterile soil with salt stress. Arabidopsis speciments were photographed after one month. The results are mean ± standard error from three independent experiments. Different letters indicate statistically significant differences (P < 0.05). Six-d-old Arabidopsis seedlings germinated on ½ MS plantes were transferred into pots and then treated with bacterial cell suspension or 10 mM MgSO4. After 3 d, seedlings were irrigated with either 0 mM or 200 mM NaCl. After three weeks in pots, the Arabidopsis specimens were photographed. The aerial parts were then taken for fresh weight and dry weight (85 °C for 2 d) measurement.

190 To further test whether bacteria modulated plant responses against abiotic stress in vivo, we infected Arabidopsis seedlings and monitored the growth behavior after a salt shock with 200 mM NaCl by capillary process from the bottom of the pots. Then, the plants were allowed to recover for 7 d by normal irrigation with sterile ddH2O. Under these moderate salt stress conditions, all plants survived. After one-month growth in pots, the most prominent effect of inoculation in the presence of NaCl was observed in terms of fresh and oven dry weight of rosettes (Fig. 4.15a, b). Fresh weight was increased significantly in response to inoculation with all four strains (1.5-2-fold higher than the uninoculated control). A similar effect was observed in the case of oven dry weight of rosette, in which inoculation of strains L, K and Y resulted in rosette dry weight ca. 1.67-fold greater than other treatments (Fig. 4.15a, b). The combined results from both stress-free and salt stress conditions strongly suggested that the selected bacteria, especially strains L, K and Y promoted growth and alleviated salinity stress in Arabidopsis. The total chlorophyll contents were also measured on the 28th DAS. The results showed that the chlorophyll content in wild-type Arabidopsis plants decreased due to salt stress, but its content in both control and treated plants under normal or salt stress conditions were not significantly different (Fig. 4.15c). The accumulation of various osmolytes were also analyzed. There was no difference between control and bacteria-treated plants with regard to proline content. Under stress conditions, proline contents increased in both control and treated plants. However, we observed a significant increase (32-51 %) in proline in treated plants due to salt stress (Fig. 4.16a). Since salt stress exposure can cause oxidative stress and produce reactive oxygen species (ROS), i.e., superoxide, siglet oxygen, and hydrogen peroxide, the beneficial effects of bacteria on stress tolerance of Arabidopsis was further explored. To study the metabolic mechanism behind the enhancement of salt-stress tolerance in bacteria treated plants, the accumulation of certain osmoprotectant molecules and ROS scavenging systems in plants was measured under both normal and salt stress conditions. We first measured the total antioxidant capacity (the DPPH assay). The ability of the leaf extracts of Arabidopsis, treated with bacteria, or not, to act as donors of hydrogen to transform the DPPH radical (deep purple) into DPPH-H (decolorization) was investigated. The total antioxidant capacity of A. thaliana is given in Fig. 4.16d, showing that DPPH was neutralized more in plants treated by strain L (ca. 50 %) than in untreated controls, under salt stress. Interestingly, treatments by strains n, K and Y (with a non-significant

191 trend to increase) did not exert significantly increased total antioxidant capacity under either normal or stress conditions. Secondly, the activities of antioxidant enzymes were measured, showing significant enhancement in the activities of APX, CAT, and POD in inoculated plants, compared to respective control plants. The level of APX increased by 79 % in salt-stressed plants inoculated with the bacterial strain L, and 63 % by strain Y (Fig. 4.16a). In addition to APX, the level of the other two antioxidant enzymes also showed significant increases in plants treated with selected strains under salt stress. Specifically, in the presence of bacterial inoculation, CAT activity was significantly increased by 19.8 (strain n) and 29.6 % (strain L) at 200 mM NaCl stress, compared to the controls (Fig. 4.16b). Bacterial inoculation with strains n, L, and K led to increases (33, 49 and 27 %, respectively) in POD activity, as compared to uninoculated plants under salt stress (Fig. 4.16c). There were no significant differences in the activities of all three enzymes, under normal conditions, among treatments and the respective control (data not shown). Osmotolerance is induced by accumulation of compatible solutes to protect cells against salt-induced cell injury. Thus, the content of the osmoprotectant molecule proline within leaf tissues was measured, showing comparable results in both inoculated and control plants but significant augmentation after salt stress in plants treated with strains L (89 %) and Y (46 %) (Fig. 4.17).

192

Fig. 4.16 Effect of bacteria inoculation on the activities of (a) APX, (b) CAT and (c) POD in Arabidopsis supplemented with 200 mM NaCl. Results are expressed as enzyme activity relative to untreated controls, are given as mean from three independent experiments. Asterisks (*) indicates statistically significant differences from control plants (P < 0.05).

193

Fig. 4.17 Effect of bacteria inoculation on the production of proline in Arabidopsis supplemented with 200 mM NaCl. Values are given as mean ± standard error from three independent experiments. Asterisks (*) indicates statistically significant differences from control plants (P < 0.05).

4.4.7 Impact of rhizobacteria on disease severity in Arabidospsis We previously found that strains n and L could inhibit the growth of Pst DC3000 and Botrytis cinerea on agar plates (section 4.4.2.2). Since protection against pathogens has been considered to be one of the core components of PGP traits of PGPR (Persello-Cartieaux et al., 2003), experiments were designed to test the protective potential of the four selected strains by challenging the bioprimed Arabidopsis with P. syringae pv. tomato DC3000 and B. cinerea B191. Leaves of 4-week-old Arabidopsis plants were challenged with the virulent strain of Pst DC3000 and showed typical yellowing symptoms on the 3rd d after inoculation. Visual observations showed that plants (peat pellet assay) treated with strains n and L strongly restricted disease spread in comparison to other treatments (Fig. 4.18). Pathogenic bacteria colonizing

194 Adabidopsis leaves were extracted from surface-sterilized leaves 4 d after inoculation and enumerated on a selective medium that allowed growth of Pst only. In the peat substrate bioassay, extraction of Pst DC3000 immediately after infiltration resulted in approximately 3×103 CFU mg-1 in both control and treated plants. Compared with immediate extraction, the number of virulent bacteria from control leaves and those treated with strains K and Y was increased from 3×103 to 5.5×106 mg-1, whereas treatment with strains n and L significantly inhibited the growth of Pst to 4.3×104 mg-1. Significant reductions in Pst disease severity were also observed for Arabidopsis seedlings grown in vitro on ½ MS agar plates under the same bacterial treatments (Fig. 4.18). Consistent with this macroscopic observation, the population density of Pst in Arabidopsis rosettes was reduced significantly for seedlings treated with either strain n or L. Before B. cinerea infection, Arabidopsis was first seed-treated with selected bacteria and grown for 4 weeks. The fungal pathogen used in this work exhibited pathogenicity and high virulenece on Arabidopsis plants, resulting in typical symptoms such as rapid expansion of visibly infected regions and water-soaked lesions at the inoculation loci, leaf yellowing, and then severe necrosis around the spotting site in control leaves 2 d after infection. The lesion diameter (mm) in 4-week-old plants was measured on the 4th d after inoculation. We observed that the fungus caused spreading disease necrotic lesions and tissue damage in the leaves of Arabidopsis. The disease severity was similar among the control plants and plants that were pre-treated with strains K and Y. While strains n and L showed significant improvement in resistance to B. cinerea over untreated control plants, showing significantly decreased disease necrosis compared with controls (Fig. 4.19a). The lengths of lesions were significantly reduced in plants treated with strains n and L when compared with other treatments (Fig. 4.19b). Moreover, fungal growth, which is used as an indicator of the severity of pathogen infection, was measured 3 d after inoculation by real-time PCR. In line with the reduced disease symptoms observed on infected leaves, fungal growth was significantly inhibited in leaf tissue from plants treated with strains n and L when compared with other treatments, indicating that these two strains effectively protected Arabidopsis from B. cinerea infection (Fig. 4.19c).

195

Fig. 4.18 Effects of treatment of Arabidopsis wild-type Col-0 with isolated rhizobacteria on resistance against a bacterial pathogen. In planta experiment: 4-week old plants were challenge-infected with Pst DC3000. (a) The growth of Pst DC3000 and (b) disease symptoms in control and rhizobacteria-treated plants was investigated. An in vitro experiment showed significant reduction in disease symptoms in seedlings treated with rhizobacteria (d), as confirmed by the inhibition of Pst growth (c). The number of bacteria in the leaves was determined at 7 d after drop inoculation with a suspension at 109 CFU mL-1 in a plate assay. Different letters indicate statistically significant differences among treatments (P < 0.01). Bars indicate mean ± standard error from three independent experiments, with 12 inoculated leaves per treatment.

196

Fig. 4.19 Effects of treatment of Arabidopsis wild-type Col-0 with isolated rhizobacteria on resistance against Botrytis cinerea. Four-week old plants were challenge-inoculated with spores of B. cinerea at 105 CFU mL-1. (a) Visual comparison of disease symptoms on Arabidopsis leaves 3 d after inoculation. (b) At 7 d postinoculation, the average diameter of the expanding lesions formed in Arabidopsis leaves was measured. Disease severity measured as the lesion diameter in the rhizobacterial treatments were expressed as a percentage of the lesion diameter in the control treatments (set at 100 %). (c) In planta growth of B. cinerea as measured by simultaneous quantification of the expression levels of B. cinerea Actin gene (BcActin) and the Arabidopsis Actin gene (AtActin). Relative fungal growth was determined by ratios of BcActin/AtActin. Bars indicate mean ± standard error from three independent experiments, with 12 inoculated leaves per treatment (** P < 0.01 versus control).

197 4.4.8 Effect of rhizobacteria on transcript levels of Arabidopsis To assess the molecular mechanisms underlying the phenotypic effects observed in plants inoculated with selected strains under both normal and salt stress conditions, the expression levels of several chosen genes were measured at 12 h (short term) and 21 days (long term) after inoculation, using the qRT-PCR technique. For testing the early transcriptional changes in salt-stressed A. thaliana plants, 14-d-old seedlings grown vertically on MS agar plates were transferred to fresh plates supplemented with 200 mM NaCl, and RNA extractions were performed 12 h after transplanting. The expression of marker genes in ABA-dependent and/or independent pathways in A. thaliana were analyzed. Twelve hours after salt stress, the expressions of RD29A and RD29B were increased in seedlings treated with strains L, K and Y, compared to the uninoculated controls; increases in the transcript levels of these two genes in bioprimed A. thaliana were salt-stress dependent (Fig. 4.20). The stress alleviation strategies employed by A. thaliana involve the accumulation of compatible solutes. We observed that the expression of the P5CS1 gene, which encodes rate limiting steps in proline synthesis, was significantly up-regulated in plants treated with strains n, L and Y (Fig. 4.20); however, this stimulation effect was not present under optimal growth conditions. The same was true for At4g36110 (SAUR-like auxin-responsive protein), where the transcriptional level was induced by strains L and K only under salt stress (Fig. 4.20). As shown in Fig. 4.20b, inoculation with strains L, K and Y, under normal conditions, resulted in a slight induction (short term; data not shown) and a significant up-regulation (long term) of AtGA3ox1 (Gibberellin 3-beta-dioxygenase, At1g15550), which is one of the key genes regulating the synthesis of bioactive gibberellins. The effect of PGPR on plant defense priming was also investigated. PR-1 expression, which is a salicylic acid (SA)-induced marker, showed 20 and 17-fold increases following treatment with strains n and L (long term), respectively (Fig. 4.21). Interestingly, under salt stress, plants inoculated with strains n, L and Y showed a clear up-regulation of PR-1; a minor but significant effect was also exerted by strain K inoculation (Fig. 4.20). PDF1.2 gene expression was induced by strains n and L with 2.6 and 4.8-fold increases, respectively, at 12 h post inoculation (Fig. 4.21). We also examined the expression levels of MPK3 and MPK6. The transcript levels of the two genes were marginally, but significantly, increased by strains L, K and Y, at 12 h post inoculation (Fig. 4.21). However, salt treatment did not induce any

198 differences between bioprimed and uninoculated conditions, at least within the first 12 h. At 21 d after inoculation of Arabidopsis with strains n and L, the transcript level of the camalexin synthetase gene PAD3 was significantly higher than in the control plants (Fig. 4.21).

Fig. 4.20 Quantitative measurements of the gene expression levels (fold differences) in Arabidopsis under salt stress. Fourteen-d-old seedlings (seed-bioprimed) were treated with 200 mM NaCl. After 12 h, rosettes were collected and subjected to RNA isolation, following by quantitative RT-PCR.

199

Fig. 4.21 Quantitative measurements of the gene expression levels (fold differences) in Arabidopsis rosette under normal growth conditions, after bacterial treatment at 21 d and 12 h.

4.5 Discussion

200 Plant microbiome communities consist of large numbers of bacteria (ectophytes and endophytes) providing critical and sustainability benefits of improved soil health and nutrient utilization and increased plant growth and development. Although numerous bacterial strains are already at least reasonably well-understood in this capacity (Vessey, 2003), until the previous study (chapter 3) the rhizobacterial diversity of different crop plants as well as wild plants in unmanaged fields in Québec remained largely unknown. In the present study, a total of 98 randomly selected rhizobacterial strains were first screened for short-term PGP effects in Arabidopsis seedlings in vitro and four of them were found to show excellent growth stimulating effects in initial screening; these promising strains were then selected for further studies. The 16S rRNA gene sequences of these four strains (designated as n, L, K and Y) showed maximum sequence similarity with members of the genera Pseudomonas, Bacillus, Mucilaginibacter and Rhizobium, respectively. Bacterial strain K showed 97 % similarity with M. lappiensis type strain ANJLI2T, which has been reported as a plant-associated bacterium isolated from a decaying lichen in Scots pine forests (Männistö et al., 2010). Strain K grew at up to 33 °C, indicating it was more adapted to cooler environments, which is in agreement with the fact that most members of the family Sphingobacteriaceae are isolated from cold areas (Männistö and Häggblom, 2006). PGPR have been found in a wide range of environments. Pseudomonads are among the most promising groups of rhizobacteria, as they are aggressive colonizers (Weller, 2007) and usually manifest a wide range of PGP traits, such as antibiotic production, phosphate solubilization, nitrogen fixation, ACCD activity, siderophore production, EPS, IAA, HCN, and ammonia production, stress alleviation, and plant hormone production (Ahemad and Khan, 2012a,b; Bhattacharyya and Jha, 2012; Saber et al., 2015; Shilev et al., 2012). The most important species are P. aeruginosa, P. fluorescens, P. putida and the plant pathogen P. syringae (Scarpellini et al., 2004). Bacillus are omnipresent in nature, having immense potential for agricultural applications (Medeiros et al., 2011; Singh et al., 2009), as they are able to enhance crop yield by direct and indirect mechanisms, most of which are quite similar to those in Pseudomonas, in addition to the characteristics of antibiotic production (Medeiros et al., 2011; Ongena and Jacques, 2008). On the other hand, bacteria belonging to the genus Rhizobium are well-known to establish symbioses with legumes and make atmospheric nitrogen available to the host plant (Long, 1989). However, they have also been shown to associate with non-legumes and have potential as PGPR in this context (Ahemad and Kibret, 2014; Vessey, 2003). The genus

201 Mucilaginibacter, belonging to the family Sphingobacteriaceae, is known to hydrolyze organic matter such as xylan and pectin, and produce enormous amounts of extracellular polymeric substances (Han et al., 2012). Though possessing various PGP traits, there are very few papers discussing their PGP effects in plants (Ritpitakphong et al., 2016). Madhaiyan et al. (2010) reported that two novel Mucilaginibacter strains increased root length of tomato and canola seedlings in a gnotobiotic growth pouch assay. ACC deaminase-producing rhizobacteria promote plant growth by decreasing inhibitory effects of higher ethylene concentrations of various crops (Zahir et al., 2003). Inoculation with rhizobacteria having ACCD activity resulted in a boost of root development, leading to better shoot growth (Glick et al., 1998). Shaharoona et al. (2006) also showed a significant positive correlation between root elongation in maize and in vitro ACCD activity of bacterial cells. The genera Burkholderia and Pseudomonas are known to have the greatest number of ACCD containing bacteria (Onofre-Lemus et al., 2009). All four tested strains showed ACCD activity ranging between ca. 1.0 and 8.0 μmol mg-1 h-1, with the highest activity observed for strain n; the levels for all strains were higher than the 20 nmol of α- ketobutyrate mg-1 h-1 threshold (Penrose and Glick, 2003), indicating that ACCD activity is likely to be involved in the mechanisms of plant growth stimulation by these strains. However, strains L and Y, with lower ACCD activity, caused greater root length enhancement than other strains (data not shown). The PGP effect was not strictly consistent with the ACCD activity level, indicating that ACCD activity is not a major PGP mechanism used by these strains. It is also possible that these strains performed differently as to their ability to hydrolyze the ACC in plant roots. An additional experiment to study the ethylene emission by Arabidopsis plantlets under both normal and salt stress conditions might confirm this deduction. Most PGPR synthesize the phytohormone IAA (Patten and Glick, 1996), which plays a multifaceted role in plant growth, that is, IAA increases root growth which is essential for plant nutrient and water uptake, while increased root exudates in turn benefit the associated bacteria (Kim et al., 2011). IAA has also been implicated in plant defense responses. In our study, all four strains could produce IAA, at levels ranging between 0.10 (strain L) and 0.86 mg L-1, which is lower than some previous reports, demonstrating values of ca. 10 to 100 μg mL-1 for the same genus (Li et al., 2017). Indeed, IAA production by rhizobacterial strains varied greatly within species and/or strains of the same species (Verma et al., 2014), and was dependent

202 on culture conditions (Cassán et al., 2013). The potential to produce IAA is indicative of the capability of these strains to be used in plant growth stimulation and mitigation of pathogenesis. Nitrogen application to crops is very high worldwide, which not only raises crop production cost but also exerts adverse effects on the environment. Use of PGPR or PGPR combined with N fertilizer may optimize N uptake and reduce N fertilizer losses of crop production systems (Arif et al., 2017). Biological nitrogen fixation is catalyzed by a complex enzyme, nitrogenase, in which a dinitrogenase reductase subunit is encoded by nifH, a biomarker widely used to indicate nitrogen fixers and to study the evolution of nitrogen-fixing bacteria (Raymond et al. 2004). The results for nitrogen fixation, as confirmed by successful amplification of the nifH gene, indicated that strains n, L and Y are potentially nitrogen-fixing bacteria and may be beneficial in improving the nitrogen level of host plants. Though approximately 80 % of all biological nitrogen fixation is accomplished through the symbiotic interaction between legumes and α-proteobacteria (Garg and Geetanjali, 2007), non-specific nitrogen-fixing bacteria also exit in relationships with a much wider range of plant species, including plants of agricultural importance. Several Pseudomonas (Mirza et al., 2006), Bacillus (Ding et al., 2005), and Paenibacillus (Górska et al, 2015; Puri et al., 2015) strains have been identified as nitrogen fixers. Phosphorous (P), is an essential macronutrient for plant growth and is a limiting factor in crop production; it is highly unavailable in soil and is largely in soil reservoirs as insoluble complexes (Wang et al., 2009). Mineral phosphate in chemical fertilizers is easily immobilized via precipitation in soil and soil erosion can lead to P losses (Rodrı́guez and Fraga, 1999). Fixed P can be rendered available by P-solubilizing rhizobacteria. Bacteria-mediated P solubilization is considered one of the most important PGP traits in rhizobacteria. Many rhizobacteria belonging to the genera Achromobacter, Azospirillum, Bacillus, Burkholderia, Flavobacterium, Paenibacillus, Pseudomonas and Rhizobium etc. have been reported to be P solubilizing bacteria (PSB). Field work has demonstrated that inoculation of plants with some P solubilizing micro-organisms increased the concentration of available P in the soil and enhanced yield and phosphate uptake by the plants. In this study, we found that all four strains were positive for P solubilization, being able to convert insoluble tricalcium phosphate to the soluble forms (H2PO4 or HPO4 ions) in liquid medium, which is concordant with previous studies (Ali et al., 2015; Han et al., 2012). But strains Y (Rhizobium) and L (Bacillus) solubilized much less P than indicated

203 in previous reports. The maximum phosphate solubilization activity was detected in strain n (Pseudomonas sp.) (350 mg L-1), a higher level than previously reported (Ali et al., 2015), but lower than most Pseudomonas isolates in a previous study (Oteino et al., 2015). Moreover, strain n could solubilize P in buffered medium, indicating that it is probably an efficient P-solubilizing bacterium, secreting strong or high amounts of organic acids, and may also perform well under soil conditions as a PSB. Rodrı́guez and Fraga (1999) reported that various organic acids, i.e., gluconic acid, lactic acid, acetic acid, oxalic acid, citric acid, were produced by P solubilizing microbes. Another important PGPR attribute, which may function directly or indirectly to influence plant growth, is the production of siderophores. The bioavailability of iron is often extremely limited because of the low solubility of ferric iron under the conditions of most natural habitats (Wandersman and Delepelaire, 2004). A great number of PGPR can produce ferric chelating compounds, siderophores, to assimilate iron; this exerts niche competition, in the rhizosphere, with other rhizobacteria including pathogens, leading to enhanced competitive fitness of PGPR, providing an improved ability to colonize roots and enhance plant growth. In the present investigation, strains n and L displayed siderophore production ability; siderophore production by Pseudomonas and Bacillus strains are well known (Liu et al., 2011; Yu et al., 2011). Wensing et al. (2010) suggested that siderophore production is a key factor for phytopathogen antagonism. A greenhouse experiment showed that strains n and L significantly increased plant growth for tomato under P and iron deficiency conditions, indicating at least partially that the traits of P solubilization and siderophore production are involved in growth stimulation of plants by these two strains (Ricci, 2015). Alternative indirect PGP traits, such as ammonia and HCN production and hemolytic activity, were also evaluated qualitatively. Ammonia synthesized by rhizobacteria can be used as a nitrogen source by host plants (Marques et al., 2010). Both HCN and ammonia are volatile compounds with antimicrobial activity and function as biocontrol agents for suppression of plant diseases (Brimecombe et al., 2001). Our results showed that only strain n, belonging to the genus Pseudomonas, produced HCN, while strains n and L could produce ammonia; HCN and ammonia might play an important role in biocontrol activity. More recently, Rijavec and Lapanje (2016) suggested a novel role of HCN in increasing phosphate availability in plants, suggesting that strain n might also deploy this mechanism for plant growth promotion. Hemolytic activity

204 came to our attention because it is considered an indicator of possible activity on biological membranes, leading to potential antifungal activity of rhizobacteria (Giorgio et al., 2015). Moreover, some hemolytic activities have been related to antibiotics, produced by bacteria (Leclère et al., 2005), which can form pores in the cell walls of bacteria or fungi. Thus, hemolytic activity has been associated with biocontrol of phytopathogens (Leclère et al., 2005). In this study, only strain L was able to lyse red blood cells, indicating potential antimicrobial activity for this strain. Frikha-Gargouri et al. (2017) showed that the antibacterial activity of a Bacillus strain against pathogenic Agrobacterium tumefaciens strains was correlated with hemolytic activity. Overall, strain n (Pseudomonas sp.) was the most numerous PGP possessor among the selected strains. Some genera of bacteria, such as Azotobacter, Azospirillum, Bacillus, Enterobacter, Flavobacterium, Rhizobium and Pseudomonas, have been shown to act potentially as biocontrol agents against plant pathogens (Ahemad and Kibret, 2014). To test strains for antimicrobial activity, two phytopathogens (P. syringae pv. tomato DC3000 and B. cinerea B191) were used; antagonistic activities were observed for strains n and L. With regard to Pst DC3000, a bacterial pathogen with broad host and symptom ranges, the most promising result was obtained for the Bacillus sp. (strain L). While for B. cinerea, a plant necrotrophic fungus that causes grey mould on over 200 food plant species, Pseudomonas sp. (strain n) showed more encouranging abilities, similar to a previous report (Rafikova et al., 2016). The Pseudomonas and Bacillus genera are able to produce a broad array of antibiotics, HCN, siderophores, hydrolytic enzymes (chitinase, protease, β-1,3-glucanase), and many other secondary metabolites with antibacterial and antifungal effects (Gorlach-Lira and Stefaniak, 2009; Raaijmakers and Mazzola, 2012). Strains n and L reported in this work, are siderophore producers and exhibit protease activity, and strain n produced HCN, all of which may be implicated in the antagonistic effects against phytopathogens. Ricci (2015) reported that strains n and L showed clear antagonist abilities in response to three other phytopathogens, Clavibacter michiganensis, Sclerotinia minor, and Fusarium graminearum, suggesting the two strains have a relatively broad spectrum of biological activity and are promising potential biocontrol agents. Further analyses, to identify and characterize the antimicrobial compounds produced by these two strains, will help to elucidate the specific roles in biocontrol mechanism. Further studies should be conducted to address the effects of strains n and L on Arabidopsis plants already infected by pathogens, an

205 area which has not been widely studied (Tran et al., 2007). PGPR have been shown to protect host plants from stresses by provoking induced systemic resistance (ISR). Therefore, some of the present experimentation investigated the ability of the four strains to prevent infection and to control the development of infections induced by B. cinerea and P. syringe tomato DC3000 in plant assays. The role of systemic resistance in Arabiopsis plants induced by rhizobacteria was determined by physically separating the inducing agents from the pathogen. Pre-inoculation of Arabidopsis with strains n and L resulted in significant reduction of disease symptoms in plants infested with P. syringae in both plate and pot experiments; in addition, they were effective for fungal resistance in the soil, confirming the in vitro results and suggesting that pre-treatment with isolated rhizobacteria n and L induced plant disease resistance against pathogens and reduced symptom development during disease susceptibility to B. cinerea and Pst DC3000, a possibly useful capability in competing against other rhizospheric microorganisms. A Pseudomonas, previously isolated from semi-arid soil, showed potential biocontrol ability against M. phaseolina on mung bean (Minaxi and Saxena, 2010). Bacillus subtilis CAS15 reduced the incidence of Fusarium wilt in pepper (Yu et al., 2011). Nie et al. (2017) reported that root drench with a Bacillus strain significantly reduced grey mold symptoms in Arabidopsis through enhanced PR1 protein expression, hydrogen peroxide accumulation and callose deposition, in a JA/ET-signaling pathway and NPR1-dependent manner. Overall, these findings suggest that the antagonistic activities of strains n and L were probably due to competition, direct antagonism (i.e., siderophore and ammonia production), or induced systemic resistance in host plants. There have been reports regarding activation of plant anti-pathogen related genes by rhizobacteria (Beneduzi et al., 2012; Kim et al., 2015). Thus, future studies should focus on the exploration of tissue-specific expression of pathogenesis genes, which will ultimately shed light on the mechanisms involved in resistance against pathogens following inoculation with strains n and L. There were differences in the behaviors of A. thaliana in response to infection by B. cinerea and Pst DC 3000. Caution should be exercised in extrapolating results from A. thaliana to the behavior of other plant species. The four selected strains were also subjected to metabolic profiling, comprising API 20E, API 20NE, API 50CH, and API-ZYM tests and analyses of tolerance to various stresses and antibiotic resistance. The traits of a bacterium associated with its fitness and metabolic assets contribute to a particular adaptation, providing support for its root colonization (Mazur et al.,

206 2013). The phenotypic patterns obtained revealed the substrate richness of these strains. The more metabolically flexible strains are generally more successful competitors in host plant colonization (Wielbo et al., 2007). Intriguingly, in this study, strains L and K were the most metabolically diverse strains among four tested bacteria. Phenotypic profiling is of great importance for understanding genotype difference, stress responses, and environmental condition effects on rhizobacteria (Chojniak et al., 2015). We also investigated the fate of bacterial root colonization in Arabidopsis plantlets by using GFP-tagged derivatives, since efficient root colonization by inoculated bacteria is a critical step in the initiation of beneficial interactions between bacteria and their host plants (Shehata et al., 2017; Weng et al., 2013). An endophyte, Burkholderia phytofirmans PsJNT, from onion roots, successfully colonized the roots of various plant species, such as Arabidopsis (Pinedo et al., 2015), maize (Naveed et al., 2014), and switchgrass (Kim et al., 2012). Although strains n, L and Y, respectively, were identified as endophytes from surface-sterilized roots of reed canary grass, bittersweet, and soybean, transformed strains colonized the interior part of root tissues of Arabidopsis 14 DAS. The cellulose secretion (strains L and Y) and motility of rhizobacteria (strains n, L, and Y) may help in the process of plant colonization (Pereira et al., 2016). Uninoculated seedlings and seedlings inoculated with wild-type strains did not show any fluorescence. Results obtained from this study indicated that strains n, L and Y were better colonizers of Arabidopsis. Plate counting showed that strain K efficiently colonized root tissue. Endophytic bacteria of the four strains were detected in the range 105-106 CFU mg-1 FW (data not shown). GFP has been used to determine the colonization pattern of Enterobacter sp. (Kim et al., 2014) and Rhizobium sp. (Glaeser et al., 2016) in Arabidopsis, and Bacillus sp. in rice (Liu et al., 2006) and Chinese cabbage (Yi et al., 2017). The present study shows that the GFP technique is effective in evaluation of colonizing ability of different species of rhizobacteria. Thus, all the above-mentioned abilities involved in plant growth promotion make our strains good candidates to perform in planta experiments, which should include various production steps and under a range of growth conditions. Salinity is a very common abiotic plant stressor, severely affecting plant growth and crop production worldwide (Al-Karaki, 2006). Thus, in the present study a Petri dish assay was performed using Arabidopsis seeds inoculated with the selected strains, and showing significantly enhanced plant growth under salt stress (100 mM NaCl) with regard to rosette fresh weight, seedling root length, and total chlorophyll. Their

207 ability to increase plant growth and ameliorate salt stress was further tested in a pot experiment. Inoculation with the isolated strains, especially L and K, significantly increased shoot and root growth and total antioxidant capacity in leaves of Arabidopsis under 200 mM NaCl. Leaf chlorophyll concentration is an indicator of salt tolerance. Chlorophyll is destroyed due to ions (Na and Cl) or reactive oxygen species (ROS), resulting in the degeneration of cell organelles, especially in leaf tissue. The increase in enhanced chlorophyll content in bacteria-treated plants under salt stress on Petri plates signified that inoculation with the PGPR identified in the current work counteracted the negative effects of salinity stress on photosynthetic activity, resulting in a positive effect on growth and plant development. This might be partially due to bacterial ACCD activity (Ali et al., 2014) or ISR in plants, both of which would have led to reduced ethylene biosynthesis. It should be noted that the total chlorophyll content was not significantly changed in treated Arabidopsis plants in the pot experiment, which was not in accordance with previous results from the in vitro experiment, suggesting that the pattern of beneficial effects exerted by rhizobacteria is at least partially affected by growth conditions and salt stress levels. A recent report by Ricci (2015) showed that under greenhouse conditions, strains n and L significantly increased tomato plant growth for all the growth variables measured (plant height, root length and dry weight, leaf area, and shoot dry weight) under normal and P and iron deficient conditions. The observed increase in plant growth due to inoculation of strains n, L, K and Y might be attributed to the direct, indirect or synergistic contributions of PGP traits, including IAA and ammonia production, nitrogen fixation, and P solubilization, although specific growth condition will affect the relative contribution of each mechanism. Some PGPR were reported to promote plant growth under both normal and stressed conditions (Chen et al., 2017; Gagné-Bourque et al., 2016), while others were effective only under stressed conditions, exerting no growth promotion effect under optimal conditions (Rolli et al., 2015), suggesting that the PGP activity of some rhizobacteria depend on stress. Our results showed the growth stimulating effects of the four tested bacteria existed in soil-free sterile unstressed conditions, suggesting direct effects of these strains, rather than indirect effects on other rhizosphere microbes. Indeed, there have been other reports of growth promotion effects following bacterial inoculation of plants under optimal conditions (sterile and full nutrient) (Poupin et al., 2013; Weselowski et al., 2016), thus, the plant growth-promoting effects observed in the present study probably more rely on the production of plant growth regulators (auxins,

208 gibberellins, cytokinins, etc.), either emitted by the bacteria themselves or produced by plants upon biopriming. Moreover, the non-sterile conditions (peat pellets) used in our in-planta assay obviously resulted in a complex microbe-interaction network in the rhizosphere (Cardinale et al., 2015), which was more natural than the sterile conditions. In the case of long-term growth, the inoculation effects of strains L, K and Y were also positive; the same results have been reported in Arabidopsis and various crops by PGPR (Shahzad et al., 2013; Zahid et al., 2015). Though commonly, PGPR with multi-functional traits are better than single traits (Imran et al., 2014); it is worth noting that, among the four selected strains, strain K was the least recognized PGP trait possessor but still caused significant growth promotion in Arabidopsis. ACC-deaminase activity was expected to be one of the best indicators of effective PGPR under salinity stress, but our results showed that strain K had significantly lower ACCD activity than strain n, which had the highest level of ACCD activity, as well as IAA production, among the four tested strains, yet did not show stronger or comparable salt alleviation effects, compared to the other three. Of the four tested bacteria, strains L, K and Y were the best performers, in the case of plant growth promotion and salinity alleviation, while strains n and L possessed the greatest number of PGP traits, indicating factors other than common PGP traits must be employed in their plant growth stimulating effects. However, the specific attribution of each PGP trait should be tested using knockout mutants. Thus, it appears that rhizobacteria do not always have predictable effects on the growth and well-being of host plants, with regard to common PGP traits (Long et al., 2008), the same phenomenon was also observed by Cardinale et al. (2015). This interesting result suggested a possibile explanation for the observation that, a number of supposed “promising” bacterial strains selected from traditional PGP screening, in fact, eventually failed in planta, while the best candidates would have been discarded simply because of not performing well in in vitro PGP assays. The common PGP traits may not actually be totally responsible for plant growth promotion. Recent studies have shown that bacterial molecules (i.e., oligosaccharides, proteins), are direct plant growth elicitors (Subramanian, 2013). Salinity stress imposes oxidative stress and osmotic stress, especially ion (Na+ and Cl− ion) toxicity, and reduces plant growth (Shabala et al., 2012). However, plants alleviate salt stress effects through various mechanisms such as synthesis of compatible solutes ‘osmolytes’, induction of an antioxidant defense system, and adaptive regulation of stress-related hormones (Ashraf et al., 2012). There was a 2.6-fold increase in proline concentration in plants subjected to

209 salt stress, indicating that proline was accumulated in response to salt stress. There were no differences in proline content between inoculation treatments and controls under normal conditions, which is consistent with the result published by García et al. (2017), indicating that PGPR inoculation did not influence proline content under un-stressed conditions. Inoculation with strains L or Y significantly enhanced the contents of both proline and soluble sugars in salt stressed Arabidopsis, indicating their ability to improve salt tolerance in plants, which correlated well with the phenotypic performance of Arabidopsis exposed to salt stress. Proline has been reported to reduce the injury induced by abiotic stresses through several functions, including osmotic adjustment (to maintain turgor pressure), molecular chaperones, signal transduction, and ROS scavenging (to protect cells against oxidative damage) (Verslues and Sharma, 2010). Under stress conditions, proline itself reacts with and detoxifies ROS, thus its accumulation may be an important factor to maintain high phtotsynthesic rates (Sharma and Dietz, 2009). It is plausible that increases in proline content following inoculation under salt stress is employed by strains L and Y to contribute to plant growth under salt stress conditions, while for strain K, it is probably the total soluble sugars. It is worth noting that EPS production by strains K and Y might play a role in alleviating salt stress in Arabidopsis since EPS have been shown to bind Na+, thus decreasing the content of Na+ available for plant uptake. Since ROS homeostasis is essential in protecting normal metabolism in plants, alleviation of oxidative damage by ROS-scavenging antioxidant enzymes, such as APX, CAT and POD, is an important stress tolerance enhancement strategy in plants (Jiang et al., 2007). The CAT catalysis pathway has been considered as a key ancillary component of photosynthesis that can efficiently remove the photorespiratory H2O2 produced by plants under drought and salinity stress (Willekens et al., 1995). APX also plays a key role in the conversion of H2O2 into H2O. It is of particular importance in maintaining the homeostasis of two non-enzymatic antioxidants, ascorbate and glutathione, and is involved in the prevention of the over-reduction of photosynthetic rates under stressed conditions (Sofo et al., 2015). While the DPPH scavenging activity was only significantly increased by stain L inoculation, it is very complex with regard to how the anti-oxidative enzyme activities were regulated in Arabidopsis exposed to different bacterial isolates under salinity stress, indicating that metabolic adjustments of Arabidopsis in response to rhizobacteria under salt stress conditions are dynamic and multifaceted; the same hold true for growth response. However, the detoxification of ROS is an important strategy.

210 In order to further decipher the pathways that allowed enhanced plant vigor and induced defense machinery in Arabidopsis, molecular investigations were conducted using qPT-PCR technique. Microarray, metabolomics and proteomic analyses have shown that PGPR inoculation changed the expression pattern of genes in Arabidopsis plants; most of these genes are involved in signal transduction, defense response, and metabolism (Kwon et al., 2016; Poupin et al., 2013; Mortel et al., 2012). One of the PGP mechanisms by PGPR is the production of hormones, such as IAA, cytokinins and gibberellins. The gibberellin hormones (GAs) can regulate almost every stage of growth and development in plants. GA3ox1 (At1g15550), which is involved in GAs biosynthesis and the determination of total leaf areas and flowering time (Hu et al., 2008), was up-regulated by strains L, K and Y, after 21 d treatment, under normal growth conditions, and, in agreement, these plants produce greater leaf areas than uninoculated controls. A similar study has shown that GA3ox1 gene expression was up-regulated in 14-d-old Arabidopsis seed-treated with a PGPR, Burkholderia phytofirmans (Poupin et al., 2013). In our study, the total number of leaves was not significantly affected by the 4 PGPR (data not shown), but the rosette area was, indicating that, at least under our experimental growth conditions, these strains might act as growth accelerators. The auxin-induced protein gene At4g36110 was not affected by any of the 4 PGPR inoculations of this study, indicating that the plant growth-promoting activity of these bacteria was not caused by the production of IAA in plants under normal conditions. However, plants treated by strains L and K showed upregulated expression levels after salt exposure, indicating that, the effect of salt stress alleviation by these two strains might be caused by qualitatively altering endogenous IAA levels in Arabidopsis. It would provide more knowledge if the IAA content in different plant tissues could be studied. A recent report showed that IAA content in wheat was increased by PGPR exposure under salt stress conditions (Barnawal et al., 2017). Upon salt stress exposure, plants can deploy various mechanisms to mitigate the salt effect, and osmolyte accumulation is one of the mechanisms which has been studied. Proline can act as both as an osmoprotectant molecule and a ROS scanvenger. We observed that, the expression level of P5CS1 was not altered in plants under normal growth conditions, but was augmented significantly 12 h after salt stress treatment, in plants bioprimed by strains n, L and Y. This gene is one of rate-limiting factors in proline biosynthesis (Yoshiba et al., 1997), and its transcription was associated with abiotic stress adaptation. Indeed, proline content was increased by strains L and Y under salt stress in planta. Kim et al. (2014) also reported that the expression

211 of P5CS1 was up-regulated by PRPG in response to salt stress. In the present study, we used above-ground tissues for qRT-PCR analysis; future experiments should include the analysis of gene expression patterns in root tissues (spatialization) (Pinedo et al., 2015). Moreover, it would be interesting to use mutant strains of the 4 bacterial isolates, with mutations in several genetic pathways, to study the colonization ability and plant growth-promotion performance in Arabidopsis, since studies have shown that original and fully-functional strains were needed for growth promotion ability (Zuniga et al., 2013). Plants have evolved a sophisticated surveillance system to sense environmental abiotic stresses, to recognize the attacking pathogens, and to induce effective defense responses. Resistance is often controlled by plant resistance (R) genes which recognize the pathogen-derived avirulence effectors and lead to hypersensitive response (HR) that stops the spreading of pathogen from the initial attack site (Heath, 2000). The other innate immune response is the pathogen-associated molecular pattern (PAMP)-triggered immunity (PTI), which is activated by a number of PAMPs such as flagellin and chitin (Segonzac and Zipfel, 2011). Upon the initiation of the innate immune responses, plant cells trigger a series of signaling cascades that lead to diverse cellular responses such as synthesis of the defense-related hormones and production of reactive oxygen species (ROS). It has been suggested that resistance associated with a salicylic acid (SA)-dependent signaling pathway acts against biotrophs, whilst some other plant-defense responses dependent on ethylene (ET) and/or jasmonates (JA) act against necrotrophs (Mur et al., 2006). Salicylic acid and JA can inhibit the expression of many genes (Mur et al., 2006). For instance, JA can suppress the expression of pathogenesis-related (PR)-genes that are SA-dependent (Niki et al., 1998). While genes that are involved in JA signaling, such as PDF1 and lipoxygenase, are compromised by SA (Spoel et al., 2003). Mitogen-activated protein kinase (MAPK) signaling networks are highly conserved signaling modules that represent important means to perceive environmental cues and transduce these extracellular stimuli into downstream intracellular responses by the way of phosphorylation (Meng and Zhang, 2013). Two entire MAPK cascades in Arabidopsis, MEKK1-MKK4/MKK5-MPK3/MPK6 and MEKK1-MKK1/2-MPK4, have been established and shown to act as regulators of signaling pathways involved in immune responses (Pitzschke et al., 2009), as well as biological processes, such as differentiation, proliferation, hormone signaling, osmotic stress, etc. (Lu et al., 2015). Thus, genes involved in plant resistance to biotic or abiotic

212 stresses were also studied here. Our data here showed that the accumulation of mRNA of MPK3 and MPK6 was slightly, but statistically, increased by short-term (12 h) PGPR treatment (strains L, K and Y), which supported the hypothesis that bacterial bioprime might induce the pre-activation of stress-responsive machineries. Nie et al. (2017) showed that bioprimed Arabidopsis presented an accelerated up-regulation of MPK3 and MPK6 after B. cinerea challenge. However, the post-translational modification should be monitored to give a better insight into the how these three strains initiate priming processes in A. thaliana. Camalexin, 3-thiazol-2’-yl-indole, is one of the major phytoalexins produced by A. thaliana, that can be induced by both biotic and abiotic stress (Liu et al., 2016). Reduction in camalexin levels in Arabidopsis led to compromised resistance in the powdery mildew fungus G. cichoracearum (Liu et al., 2016). Here, we found that treatment with strains n and L led to significantly increased expression levels of CYP71B15/PAD3 in Arabidopsis at day 21 (seed-treated). This gene encodes a cytochtome P450 enzyme, involved in the camalexin biosynthesis pathway. A microarray study by Mortel et al. (2012) also showed that, gene expression of PAD3 was increased by Pseudomonas fluorescens SS101. This finding was in accordance with our results, reinforcing the involvement of camalexin in biocontrol ability of PGPR. The determination of camalexin content in plants using spectrofluorometer would facilitate the confirmation of the hypothesis. The expression of PDF1.2 (encoding an ethylene and jasmonate-responsive plant defensin) and PR1, which are involved in JA and SA signaling pathway, was also analyzed. Strains n and L significantly induced PDF1.2 expression after 12 h of treatment. The highest gene expression also achieved by strains n and L, but at day 21 after seed treatment. Interestingly, under salt stress, all four strains induced PDF1.2 expression, by up to 6-fold (strain L). It has been reported that some PGPR did not affect PR1 and PDF1.2 expression under unstressed conditions (Pinedo et al., 2015), while Hong et al. (2016) showed that the transcript level of both genes was induced by Paenibacillus polymyxa AC-1 after short-time exposure. It can be postulated here, that, strains n and L activated both JA and SA signaling pathways in A. thaliana, which is not an uncommon phenomenon (Sharifi and Ryu, 2016), while SA signaling pathways were mainly associated with strains K and Y. Additionally, we examined expression of some stress-response marker genes (RD29A and RD22), which are induced by salt and drought stress. Expression of these marker genes in

213 bioprimed plants (with strains L, K and Y) under salt stress conditions, was significantly higher than those in controls, suggesting that bacteria may affect expression of these stress-related genes. This implied that moderate exposure to biotic stressors, in this case, PGPR, might accelerate the activation of signaling machineries governing multiple stress adaptation processes. All-in-all, this study contributes to our understanding of PGPR involved in plant defense responses to abiotic stress. It can be postulated that a combination of mechanisms might be explored for the four selected strains, allowing them to exert growth promotion (all four together) and biocontrol abilities (strains n and L). Induced resistance is generally expressed against a broad spectrum of organisms, resulting in a wide range of disease prevention in plants, thus, it would be valuable to investigate the induced resistance elicited by strains n and L against various crop pathogens, since under field conditions there are usually a range of pathogens present. It would be interesting to conduct transcriptome analysis in bioprimed A. thaliana after biotic or abiotic challenges, to see whether there are primed ISR specific gene expressions (Cartieaux et al., 2008). These four PGPR not only promoted plant growth, but also reduced the detrimental effects induced by salt stress. Climate change has already caused significant negative impacts on crop production. These negative effects are largely attributable to drought, flooding, reduced availability of irrigation water, soil salinization and temperature fluctuations. The four PGPR strains of this study impart salt stress tolerance in Arabidopsis, allowing them to grow under stressed conditions; this will be of great potential in sustainable agricultural practices and in developing more climate change resilient crop production practices.

4.6 Conclusions

In this study we analyzed quite a number of isolated rhizobacteria from Chapter 3 using an easy and labor- and time-saving in vitro screening method in Arabidopsis that made the discovery of potential PGPR possible. Four bacterial strains termed n, L, K and Y from different species of crop and wild plants stood out as possible PGPR as confirmed in Arabidopsis. 16S rDNA sequencing revealed them as species of Pseudomonas, Bacillus, Mucilaginibacter and Rhizobium, respectively. From the results of the present study, all four endophytic rhizobacteria promoted plant growth partially because of their intensified PGP activities such as IAA

214 production, phosphate solubilization, siderophore, and HCN production. Strains n and L peformed well in biocontrol, while strains L, K and Y resulted in a better response against NaCl stress by effectively inducing the physiological and biochemical parameters in Arabidopsis, allowing them to cope with salinity-induced toxicity. The mechanisms by which isolates exert growth promotion effects in Arabidopsis were also studied. The increased anti-oxidative defense machinery and expression of genes related to hormone signaling, ion-homeostasis, and osmolyte production under both normal and salt stressed conditions suggested an induction of systemic response in plants. Moreover, these bacteria promoted plant growth in a nonsterile system on various substrates with different nutrient conditions, indicating that these selected rhizobacterial strains are capable of enhancing plant growth and salinity protection under a range of conditions. GFP-signals confirmed the association of bacteria with plant roots. This is of great importance in case the growth of beneficial strains be compromised by abiotic soil conditions or they are outcompeted by other antagonistic bacteria in the soil environment. These selected isolates promoted the growth of Arabidopsis and increased tolerance to salt stress, and they could be re-isolated from inoculated plants, with a bacterial density within the range typical for endophytic bacteria (Hallmann et al., 1997). Thus, all four strains reported in this study could be defined as endophytes and are promising candidates to be developed as commercial biofertilizer formulation (strains n, L, K and Y) as well as biopesticides (strains n and L) for agricultural production. Based on the phenotypic characteristics of selected isolates, strain L, a Bacillus, can tolerate extreme environmental conditions, such as salinization (7 % NaCl) and high temperature (40 °C), and possesses the second widest range of PGP traits which differentiates this strain from the other three bacteria. In addition, the nifH gene amplification result confirmed that strain L is a potential nitrogen fixer. This leads to important implications for biofuel production where reducing inputs is highly desirable for production of biomass crops on more marginal lands. This strain could be applied as an inoculum that will significantly improve nitrogen fixation and may lead to increase in plant biomass production. This is the first study about the PGP effects of Bacillus sp. from bittersweet. To our knowledge, there is no detailed study about PGP effects by Mucilaginibacter (stain K) in Arabidopsis. Our direct screening approach revealed this new taxon as a fitness-enhancing rhizobacterium of Arabidopsis under either optimal, salinity or semi-natural conditions. As such this is the first comprehensive report about the potential agricultural applications of Mucilaginibacter sp. from a wild plant. The use of environmentally

215 friendly, safe agents to increase agricultural production would be ideal, thus it would be beneficial to study the potential of all four isolates to enhance plant growth in crops and at the field level.

216 Connecting text

In Chapter 4, we found four particularly promising rhizobacteria, able to cause marked growth promotion and salt stress alleviation and biocontrol effects for A. thaliana. They were phenotypically characterized and were shown to be true endophytes. This led us to wonder if these four selected strains could stimulate the growth of crop plants and attenuate NaCl-stress responses in them. In Chapter 5, we tested the hypothesis that the four selected bacterial strains could enhance root and shoot growth in maize and canola, as well as nutrient uptake in maize. We also set out to determine if they could alleviate the salt stress during germination and early seedling growth by both crops. To understand the colonization pattern of these four strains, a plate count method was adopted and used to measure the number of viable cells in root samples. The chapter will be reformatted and submitted for publication in Environmental Microbiology.

217 Chapter 5 Effect of inoculation with bacterial endophytes on the growth of canola (Brassica napus) and maize (Zea mays)

5.1 Abstract Plant growth-promoting rhizobacteria (PGPR) have beneficial effects on the growth of most crops. The previous chapter demonstrated that four selected endophytes significantly increased the growth performance in Arabidopsis. The current study assessed the potential of these candidate for growth promotion of two key Canadian crop plants, canola (Brassica napus) and maize (Zea mays). Canola and maize seeds were treated with bacterial solutions and the germination and growth tests were carried out in Petri dishes and plastic pots, respectively. Strains L, K and Y increased root length and seedling dry weight of canola under both unstressed and salt stressed conditions. In pot experiments, canola plant biomass and chlorophyll content were evaluated after 5-weeks of growth; all four strains significantly increased above-ground dry weight by 14 to 21 %. In maize Petri-dish assays, all four strains significantly improved the shoot length (up to 33 %) and fresh weight (up to 25 %), and strain Y increased root length (30 %). Strains L, K and Y increased the seed vigor index up to 20 % over the control. A greenhouse growth, under non-sterile conditions, evaluated effects of the four strains on maize plant biomass, nutrient uptake, chlorophyll content, total antioxidant capacity, total carbohydrates, and accumulation of proline and total phenolics were evaluated. Plant growth was promoted by all strains, especially L, K and Y, for root length, aerial biomass, and root dry weight. Total antioxidant capacity and content of polyphenol and proline in maize leaves were enhanced by inoculation with strains n and L, while total carbohydrate was increased by strains L, K and Y. Bacterial strain n improved Fe and P uptake into maize leaves, while the nitrogen content was increased by all four strains, with increases of up to 93 %. Strain L was the only endophyte to cause highly significant increases in N/C ratio. Thus, strains L, K and Y caused increases in the growth of canola and maize, likely through the integration of various mechanisms that improve the plant performance, indicating the need for an in-depth understanding of plant-bacteria interactions. Hence, this study provides evidence that the four tested PGPR, particularly strains L, K and Y, could have application to a broad range of crop plants and inoculation with these PGPR

218 may represent an important biotechnological approach to boosting plant growth and crop yield thereafter.

5.2 Introduction Chemical inputs to crop production systems (fertilizers and pesticides) are generally used to supply essential nutrients and protection from pathogens in soil-plant systems throughout the world. However, because of the prices, availability, and the ecological concerns, there has been an increased need to curb the use of chemicals in the modern agriculture. In order to cope with the increasing demand for food, due to population expansion, decreases in arable land, and environmental deterioration, a number of transgenic crop plants have been developed with both agronomic and economic benefits; however, there has been debate regarding health and ecological risks associated with modified gene(s) (Zhang et al., 2016), limiting their use for intensive agricultural production. Microbial flora associated with plant species are pivotally important regulators of terrestrial ecosystems (O’Callaghan, 2016). A key group of rhizosphere microbiome bacteria are plant growth-promoting rhizobacteria (PGPR), a heterogeneous group that can proliferate aggressively in the rhizosphere, at the rhizoplane or in plant roots, and are beneficial to plant physiological and developmental processes, such as enhancing a host plant’s nutrient uptake and resistance to biotic and abiotic stressors (Mendes et al., 2013; Vessey, 2003). These PGPR also affect soil conditions. While their mechanisms of action are far from fully elucidated, based on our current understandings these beneficial microbes can be categorized into five groups: (1) biofertilizers: for their ability to fix atmospheric nitrogen (Kuan et al., 2016) and to solubilize mineral phosphate (Oteino et al., 2015) and potassium (Saha et al., 2016), etc., (2) biopesticides: for their antagonistic effect against phytopaghogens (Haque et al., 2016) through the synthesis of cell-wall-degrading enzymes, siderophores, antibiotics, fungicidal compounds and induction of plant systemic resistance against pathogens (Ahmad et al., 2008; Yu et al., 2011; van Loon, 2007), (3) rhizoremediators: for their organic-degrading and heavy-metal-binding capabilities (Morales-Guzmán et al., 2017; Wu et al., 2006), (4) phytostimulators: for their ability to produce plant hormones such as indole acetic acid (IAA), cytokinins, gibberellins (Khan et al., 2016; Marques et al., 2010) and (5) microbe-to-plant signaling molecules, such as oligosaccharides and

219 bacteriocins (Subramanian, 2013): for their direct effects on plant growth and stress alleviation, thought the underlying mechanisms have not been fully studied. Crops from the genus Brassica are very ancient cultivated plants, dating back to at least 1500 BC (Prakash, 1980). Canola (Brassica napus L.) is the third most cultivated oilseed crop, following soybean and oil palm. Canola is also an option for ground cover plant in crop rotation systems and is important for protein meal production (Schuchardt et al., 1998). In 2016-2017, the world produced more than 563 million metric tonnes of oilseeds, of which rapeseed accounted for ca. 68 million metric tons (FAO; US Department of Agriculture). Maize (Zea mays L.) is thought to have originated in southwestern Mexico 7000 years ago (Matsuoka et al., 2002). It has well adapted to various climates and soil types and is geographically widely distributed from approximately 50 ºN to 40 ºS (Leff et al., 2004). Maize is the third most important cereal crop, after rice and wheat, for meeting increasing global food demand; it is used primarily as livestock feed, and also for biofuel production, with almost 1,070 million tonnes being produced in 2016-2017 (http://www.worldofcorn.com/#world-corn-production). Maize provides high economic returns but is accompanied by high production costs, for example, it requires large amounts of fertilizers. Besides their pre-eminence as food, canola and maize have been regarded as biofuel crops, which provide renewable and cleaner-burning alternatives to fossil fuels (Health Canada, 2012). The production of canola and maize has been constrained by environmental stresses, including soil salinity (Farooq et al., 2015; Long et al., 2015). There is a need to improve the salinity stress tolerance of canola (Ashraf and McNeilly, 2004) and maize (Farooq et al., 2015). Beneficial rhizobacteria, associated with crop plants, have attracted a great deal of attention in recent years (Egamberdieva, 2008; García-Fraile et al., 2012; Khande et al., 2017; Maeques et al., 2010; Singh and Jha, 2016; Yasmin et al., 2016). Several studies have demonstrated the beneficial effects of PGPR on the growth and yield of maize (Rojas-Tapias et al., 2012; Rudolph et al., 2015) and canola (Farina et al., 2012; Pallai, 2005). Inoculation with Herbaspirillum frisingense EMA-117 increased maize shoot and root growth, by 45.78 and 17.65 %, respectively, over un-inoculated plants (Montañez et al., 2012). Kuan et al. (2016) conducted experiments on pot-grown maize, to examine the effect of PGPRs maize growth under reduced fertilizer-N input and found that Bacillus pumilus S1r1 had the highest N2 fixing capacity, supplying 30.5 and 25.5 %, respectively, of the total N requirement of maize prior to

220 anthesis and at ear harvest. More recently, seed inoculation with a combination of three strains, Lysinibacillus sphaericus, Paenibacillus alvei, and Bacillus safensis, significantly increased maize yield, from 24 to 34 %, over two field seasons (Breedt et al., 2017). A field experiment showed that application of Azospirillum brasilense, with a half-dose of nitrogen fertilizer, resulted in the largest yield increases for canola variety Serw-4 (El-Howeity and Asfour, 2012). Another study demonstrated significantly increased 1,000 seed weight and seed protein content with Azospirillum treatment, while maximum increase in seed oil content and significantly lower glucosinolate in canola resulted from Azotobacter treatment (Nosheen et al., 2013). A more recent field study indicated that foliar application of two N2-fixing bacteria Azotobacter chroococcum and Azospirillum lipoferum, alone or together, significantly improved agronomic traits of canola, such as plant height, seed number per silique, and yield of oil and protein (Ahmadi-Rad et al., 2016). PGPR have been used to ameliorate salt stress in crop plants (Fu et al., 2010; Hahm et al., 2017; Kang et al., 2014; Mahmood et al., 2016; Martínez et al., 2015; Yao et al., 2010; Yue et al., 2007), including maize (Chen et al., 2016; Rojas-Tapias et al., 2012) and canola (Noorieh et al., 2013; Oskuei et al., 2017; Siddikee et al., 2010). There has been increasing interest in environmentally friendly and safe agricultural practices that can reduce potential negative effects associated with conventional food production. In this case, ‘Organic Agriculture’ stands out as a production system that minimizes the use of synthetic chemicals and promotes soil productivity, based on sustainable agricultural practices such as crop rotation, biocontrol, and biofertilization. Numerous PGPR inoculants, as commercially formulated products, have been adopted as sustainable agronomic practices leading to improved crop growth and yield or providing protection of the crop plants from diseases (Nakkeeran et al., 2005; Tabassum et al., 2017). Bacteria from genera such as Bacillus, Pseudomonas, Burkholderia, Streptomyces, Rhizobium, etc., have been used as biofertilizers and/or biocontrol agents (Souza et al., 2015; Tabassum et al., 2017). Intriguingly, at this time there is no published information regarding effects of Mucilaginibacter spp. on the growth and development of crop plants. We have recently shown that four selected rhizobacterial strains, one within each of the genera Pseudomonas, Bacillus, Mucilaginibacter and Rhizobium, elicit specific systemic physiological responses, leading to improved growth and salinity stress resistance in Arabidopsis (Chapter 4). However, their effect on crop plants is uninvestigated. In Canada, both canola (a close relative to Arabidopsis) and maize (a monocot) are widely

221 cultivated and of large economic importance, being Canada’s second and third most valuable crops, with anticipated production of 18.2 and 13.6 million tonnes for 2017, respectively (Statistics Canada, August 31, 2017). Growing crop plants in less fertile and/or salt affected soils may become successful if a relatively high economic yield could be produced with relatively small input costs. With this in mind, the present study evaluated the effects of the four selected PGPR on: (1) in vitro canola seed germination, with regard to root length and dry weight of seedlings, (2) in vivo biomass production and accumulation of chlorophyll in canola, (3) in vitro germination of maize seeds, and (4) biomass production, uptake of nutrients, total antioxidant capacity, accumulation of proline, total production of phenolic compounds, and chlorophyll in maize under greenhouse conditions.

5.3 Materials and Methods

5.3.1 Bacterial strains and plant materials In this study, four presumptive rhizobacterial strains of Pseudomonas sp., Bacillus sp., Mucilaginibacter sp., and Rhizobium sp., respectively: strains n, L, K and Y, were studied. They were isolated from roots of reed canary grass, bittersweet, fall dandelion, and soybean, respectively, in Sainte-Anne-de-Bellevue, Québec, Canada (45°24′22″ N lat, 73°56′44″ W long, elevation 20.87 m) (Chapter 3) and were characterized in Chapter 4. These strains were selected for their potential as biofertilizers and stress (salt) alleviators. After revivification on King’s medium B (KB) agar plates, isolates of these strains were grown aerobically in liquid KB on a rotary shaker with agitation at 150 rpm in darkness at 28 °C for 12 to 72 h, accordingly, to reach the exponential growth phase. Bacterial cells were pelleted by centrifugation (6,000 g, 10 min,

4 °C ), washed free of the growth medium 4 times with sterile 10 mM MgSO4, and resuspended in 9 -1 10 mM MgSO4 to a final density of 10 CFU mL , as determined by optical density and serial dilutions with plate counts. For the pot experiment, bacterial inocula were adjusted to 108 CFU mL-1, based on spectrophotometer estimates of cell density. Canola seeds (Invigor 5440), selected for uniformity of size, were surface sterilized in 70 % ethanol for 1 min followed by 3 % (w/v) sodium hypochlorite. After 10 min, the seeds were thoroughly rinsed 5 times with sterilized ddH2O, in order to remove excess bleach (Penrose and Glick, 2003). Uniform maize seeds (var. 19K19), procured from Doug Shirray Seeds and Ag

222 supplies (Tavistock, ON, Canada) were surface sterilized with 70 % ethanol for 10 min and were subsequently thoroughly washed twice with sterilized ddH2O (Marques et al., 2010). The seeds were then air dried in a laminar flow hood, and were sown in pots or treated at room temperature for 1 h in the absence of light with dense (109 cfu mL-1) suspensions of the bacterial strains at the exponential growth phase in sterile 10 mM MgSO4 or sterile 10 mM MgSO4 (as a negative control), while a small subset of the seeds were placed onto NB agar plates and incubated at 28 C for 7 d, to check for any microbial contamination.

5.3.2 Effect of sodium chloride on bacterial growth Tolerance of the selected strains to NaCl was studied in KB broth (pH 7.0), supplemented with increasing NaCl concentrations ranging, from 0 and 200 mM, at 50-mM increments. For each value studied, 200 µL of the KB medium, of the selected NaCl concentration, and 106 CFU mL-1 of inoculum were added into 96-well microplates, and incubated at 28 C for 72 h. Bacterial growth was monitored by measuring the optical density (OD) at 600 nm with a CytationTM 5 imaging reader.

5.3.3 Canola germination The effect of bacterial treatment on canola germination variables were studied under optimal and salt stress conditions. There were 10 replications of each treatment with 10 bacterially treated canola seeds in each 9-cm sterile Petri plate, on two layers of Fisherbrand P8 filter paper wetted with 7 mL sterile ddH2O (control) or saline water solution at 50, 100, 150 mM NaCl, and incubated at 20 °C in a germination chamber in darkness. Each plate was wrapped in parafilm to prevent contamination and drying out. Seed germination (a radicle > 1 mm) was evaluated daily up to 7 days. Seed germination percentage (GP) for each treatment was computed on the 7th day by the following formula GP = (total number of normal germinating seeds/total number of experimental seeds, in all replicates) ×100. The length of roots and dry weight of each seedling were measured at the end of day 7.

5.3.4 Maize germination

223 Seedling vigor was assayed to test the bacterial isolates for their plant growth promoting ability, specifically with regard to their effects on seed germination, root and hypocotyl growth (Abdul-Baki and Anderson, 1973). According to the treatments, inoculated maize seeds were arranged in an equidistant manner in a Petri dish previously lined on the bottom with two layers of Fisherbrand P8 filter paper moistened with 10 mL of sterile ddH2O or saline water solution at 150 mM NaCl. The plates were sealed with Parafilm and incubated at a constant temperature of 25 °C in a germination chamber. Seed germination was evaluated daily for up to 7 days. A seed was considered germinated when its radicle was at least 1 mm in length. By the end of the 7th day, germination percentage (GP) was calculated as described above. Seedling vigor characteristics (lengths of roots and hypocotyls), as well as seedling fresh and dry weight were recorded. The vigor index (VI) was calculated using the formula VI = % germination × (mean root length plus mean hypocotyl length). There were 10 replications each with 10 bacterially treated seeds, or in the case of the control, 10 untreated seeds. The experiment was conducted twice (repeated).

5.3.5 In Vitro inoculation of maize

Maize seeds were surface sterilized, incubated in bacterial suspension or 10 mM MgSO4 for 4 h and were then germinated in Petri plates containing 1.5 % (v/w) agar in water for 4 d in the dark, after which the germinated seeds were transferred aseptically to glass tubes (20 × 2.5 cm diameter) filled with 25 mL of ½ HS supplemented, or not, with 100 mM NaCl, and maintained at 25 °C in a growth chamber under a 16 h photoperiod with a 350 µmol m-2 s-1 photon flux density. Each endophyte was tested in 12 replicate tubes, randomly distributed in the growth chamber. The experiment was repeated twice. After 3 weeks, plants were removed from the tubes and allowed to air dry for 20 min (Shehata et al., 2017). Roots and shoots were then dissected from each plant for physiological index measurements.

5.3.6 Evaluation of root colonization Root colonization potential of inoculated bacteria was determined via serial dilution plating, using seedlings from the germination assay. To evaluate the population of endophytic bacteria, 6 randomly chosen seedlings of maize or canola, from each treatment, were harvested,

224 rinsed in sterile ddH2O several times and then patted dry with sterile paper towels. The root samples were surface sterilized with 70 % ethanol for 10 min, rinsed three times with sterile ddH2O, and then shaken for 5 min in commercial bleach (3 % available chlorine), followed by 5 changes of sterile ddH2O. To confirm tissue surface disinfection, a 100 μL sample of the sterile ddH2O water from the final rinse were spread on KB plates, in triplicate, and bacterial growth was assessed following a 7-day incubation period at 28 °C. Following sterilization, the root samples were cut into 1-cm segments and 1 g of segments was crushed aseptically in 5 mL of sterilized PBS using a mortar and pestle and the supernatants were serially-diluted, spread on KB agar plates and incubated at 28 °C. After 1-4 days, number of viable cells was estimated as colony forming units (CFUs) g-1 root (FW). This experiment was repeated three times. Identity of the isolated strains were confirmed by 16S rDNA sequencing.

5.3.7 Impact of bacterial inoculation on greenhouse growth of maize plants 5.3.7.1 Evaluation of isolated bacteria effects on maize plant growth To evaluate the efficiency of rhizobacterial inoculants on development and growth variables of maize, a greenhouse experiment was conducted. The potting mixture (1 Turface: 2 sand, v/v) for growing maize was weighed (1,800 g) for each pot (top diameter of 150 mm), after which the pot was moistened with Hoagland’s nutrient solution (Hoagland and Arnon, 1950), diluted to 25 % in ddH2O, the day before sowing. Each pot received 4 sterilized maize seeds of equivalent size and shape, sown at a depth of 2.5 cm. The seeds were then immediately inoculated with 10 mL of bacterium-MgSO4 suspension. Pots were organized following a completely randomized design and maintained under greenhouse conditions with average day and night temperatures of 26 and 18 °C, respectively, and natural light. Seven days after sowing (DAS), seedlings were thinned to one per pot and received a second inoculum dose at 10 mL per plant. Uninoculated control plants of all treatments were mock-inoculated with 10 mM MgSO4 and kept in the same growth conditions. During the growth period, plants were given equal amounts of ½ Hoagland’s solution twice a week. Chlorophyll content index of maize leaves was recorded using a portable, hand-held chlorophyll meter (SPAD-502, Konica Minolta), measured as the optical density. Four leaves, from each pot of eight plants per treatment, were used for measurement and two readings per leaf

225 (one on each side of the main vein) were recorded. All plants were well irrigated before this measurement was taken. The data on measured growth variable were collected from 35 DAS. After the vegetative growth was harvested, the height and circumference were measured. For root length measurement, samples were washed in ddH2O to removed debris, after which an 8-bit greyscale image of each root sample was acquired by digital scanning at a 400 dots per inch (dpi) resolution using a flatbed image scanner (Modified Epson Expression 10000XL; Epson America, Inc., San Jose, CA, USA) and saved in TIFF format, in which the root lengths were measured using commercial WinRHIZO software (Regent Instruments Inc., Montreal, Québec, Canada). A length-to-diameter ratio of 6:1 was used as a threshold to distinguish root from non-root materials (Backer et al., 2017). Total leaf area was measured by LI-3100C Area Meter (LI-COR®), according to manufacturer instructions. The above-ground and root biomass were collected and weighed; aboveground biomass dry weight was obtained after drying in an oven at 85 °C for 72 h.

5.3.7.2 DPPH Radical Scavenging Assay The total antioxidant capacity of maize leaves was determined using a DPPH (2,2-diphenyl-1-picrylhydrazyl hydrate) assay following Awika et al. (2003) with minor modifications. The antioxidants react with DPPH and convert it to 1,1-diphenyl-2-picryl hydrazine, resulting in decoloration (from deep violet to light yellow). Dried maize leaf tissue was homogenized in pure methanol (MeOH). After centrifugation at 10,000 g for 10 min, the supernatant was recovered. The pellet was re-extracted with 10 mL MeOH. Supernatants were combined, and the total volume was made up to 25 mL. Each extract (100 µL) was added to 2,850 μL fresh DPPH solution (0.11 mM) and incubated for 1 h at 22 °C in the dark. The absorbance of the samples was then read at 515 nm, against a MeOH blank. The results were expressed in mg Trolox equivalents (TE, mg Trolox)/100 g DW through comparison against a Trolox standard curve (25–800 μM), which was conducted in parallel.

5.3.7.3 Determination of total phenolic compound content Toal phenolic compounds was analyzed as described by Fan et al. (2011), with slight modifications. Fresh maize leaves (0.5 g) were frozen in liquid N2 and extracted twice with 70 % (v/v) aqueous MeOH at 40 °C for 2 h, and the extracts were centrifuged (10,000 g, 10 min) and

226 combined. The extract was then pipetted into 2 mL glass tubes and to each 1.58 mL water and 100 μL 2 N Folin-Ciocalteu reagent were added. After an 8 min-incubation, 300 μL 20 % (w/v) sodium carbonate (Na2CO3) was added to stop the reaction. The vortexed mixture was left at room temperature in darkness for 2 h and the absorbance was read at 760 nm against a blank (70 % MeOH). A calibration curve (20–500 mg L-1) was prepared using the procedure described above. Total phenolics were expressed as gallic acid equivalents (GAE, mg gallic acid g-1 FW).

5.3.7.4 Estimation of proline Proline (Pro) content in the fresh leaves of 35-d-old maize plants was determined by the ninhydrin method, described by Singh and Jha (2016). Briefly, 500 mg of tissues were frozen in liquid N2 and homogenized in 3 % sulfosalicyic acid, followed by centrifugation at 10,000 g for 10 min. To 500 µL of supernatant 500 µL of acetic acid were added and then gently vortexed with 500 µL of 2 % ninhydrin. The mixture was boiled for 30 min in a boiling water bath. After cooling at room temperature for 30 min, 1.25 mL of toluene was added to the mixture, to extract red products. The absorbance of the upper toluene phase was determined spectrophotometrically at 532 nm and the free Pro content in the supernatant was measured by comparison with a pure L-proline standard curve prepared by the same method.

5.3.7.5 Estimation of carbohydrate The total sugar content was estimated using the phenol-sulphuric acid method of Dubois et al. (1956), with minor modifications. Briefly, 50 mg of the dried leaf tissue was weighed into a glass tube with 2.5 mL of 2.5 N HCl and kept in a boiling water bath for 3 h. After neutralization with solid sodium carbonate, the volume was made up to 50 mL and centrifuged at 8,000 g for 10 min, to remove insoluble material. To 100 μL of sample was added, consecutively, 100 μL of

5 % phenol water solution and 500 μL of concentrated sulfuric acid (H2SO4); the result was mixed thoroughly. After 10 min of shaking at 100 rpm, the tubes were placed in a water bath at 25 °C for 20 min. The absorbance of the green colored product, formed by hydroxumethyl furfural reaction with phenol, was read at 490 nm against water as a blank. The total soluble sugar was expressed as glucose equivalent by comparison with a glucose standard curve (0-20 mg).

227 5.3.7.6 Determination of nutrient uptake At the end of the experiment, maize plants were washed three times with distilled water. All the shoots or roots within each replicate of each treatment were dried in an over (85 °C for 72 h) to obtain a constant weight. They were then combined and ground to a fine powder in a handheld electric coffee grinder. The concentrations of minerals: sodium (Na), potassium (K), calcium (Ca), magnesium (Mg), iron (Fe), zinc (Zn), copper (Cu), and phosphorous (P), were determined according to Nassar et al. (2012), by Kebs Technologies Inc. (Laval, Canada).

Samples were incubated overnight in a fume hood in 3.0 mL of nitric acid (HNO3) in a 10 mL Oak Ridge centrifuge tube (Thermo Scientific, NY, USA). The following day, samples were digested using a heating block (Thermolyne heater type 16500 dri-bath model DB16525; Thermolyne, Dubuque, IA 52001, USA) to 105 °C until no nitrous oxide gases (brown gases) are evolved, allowed to cool and diluted 4 times with Milli-Q water and injected into the ICP-OES apparatus for analysis. An elemental stock standard solution mixture (SCP Science, Baie-D’urfe, QC) was used to calibrate the instrument before sample injection. An inductively coupled argon plasma optical emission spectrometer was used for mineral analysis (model VISTA-MPX CCD, ICPOES, Varian, Australia PTY Ltd., Australia). The settings were: power 1.2 kW, plasma flow 15 L min-1, argon pressure 32 L min-1 (600 kPa), nebulizer flow 0.75 L min-1, auxiliary flow 1.5 L min-1, pump rate 15 rpm, viewing height 10 mm, replicate reading time 10 s, and instrument stabilization delay 15 s. The concentrations of P, Mg, Ca and K were expressed as mg g-1 of sample dry matter, whereas the Fe, Na and Cu concentrations were expressed as µg per g-1 of sample dry matter. Total carbon (C) and nitrogen (N) was analyzed with a Shimadzu TOC-V analyzer (Shimadzu Corporation, Kyoto, Japan), and expressed as % (concentration initially in 100 g of dry matter of leaf or root tissue).

5.3.8 Impact of bacterial inoculation on chamber growth of canola For canola growth evaluation, the experiment was carried out in a controlled environment chamber (Conviron Model No. PGR15, Controlled Environments Ltd, Winnipeg, MB, Canada) operating at 22 °C with 8-h-light/16-h-dark photoperiod provided by fluorescent lighting providing 350 µmol-2 s-1 photons and 60 % relative humidity. Briefly, surface-sterilized canola seeds of uniform size and shape were soaked in bacterial suspensions at room temperature for 1 h, after which the seeds were sown at the depth of 2 cm in pots containing a mixture of sterilized

228 perlite and vermiculite (1:1, v/v) and 25 % Hoagland solution (HS). Five seeds were sown per pot, with subsequent thinning, 7 d after sowing, to establish two plants per pot. Plants were kept under growth chamber conditions, equally irrigated with ½ HS twice a week, and then harvested at 35 DAS. The following variables were measured: fresh and dry weight of shoots and roots, total leaf area and root length. Briefly, fresh weight was measured on an electronic scale, while the dry weight was obtained by drying the plant tissue to a constant weight at 85 °C . Leaf area was measured using an LI-3100C Area Meter (LI-COR®), while root length was measured using a WinRhizoTM system (Regent Instruments, Inc.), as described above. Chlorophyll content index of canola leaves was recorded using a hand-held chlorophyll meter (SPAD-502, Konica Minolta).

5.3.9 Data analysis For greenhouse and chamber trails, there were 12 and 16 plants, respectively, arranged in a randomized complete block design, per each treatment group. Each experiment was carried out three times to confirm treatment effects and the combination results from each representative trial of each experiment are reported in the “Results” section. Experimental data were subjected to statistical analysis using one-way analysis of variance (ANOVA) and comparisons between treatment means were calculated at the 0.05 significance level using Tukeys Honestly Significant Differences (HSD) test of the COSTAT® statistical software. Graphics were drawn with SigmaPlot version 11.0.

5.4 Results

5.4.1 Effect of NaCl on bacterial growth High levels of NaCl repressed bacterial growth, however, strains n and L tolerated a much higher content of NaCl than K and Y (section 4.4.2.3). In order to determine their performance under slight and moderate NaCl concentrations, growth curves were constructed for the four selected strains. Bacterial growth kinetics were evaluated at 0, 50, 100, 150, and 200 mM NaCl. The results demonstrated that NaCl did not exert any negative effect on the growth of

229 strains n and L; whilst strain Y was negatively affected by 150 mM NaCl and strain K by 100 and 150 mM. At a concentration of 200 mM NaCl, strain K barely grew.

Fig. 5.1 Growth rate of strains n, L, K and Y in KB medium versus NaCl (0-200 mM) at 28 °C . Three independent experiments were carried out. Bars represent standard error of the mean (SEM).

5.4.2 Canola inoculation experiment 5.4.2.1 Effect of seed inoculation on germination Results from this study revealed that all four strains could significantly increase seedling root length, by up to 57 %, as compared to the control (Fig 5.2a). Seed germination percentage was not affected by either bacterial treatments or salt stress (data not shown). Salinity concentration of the media significantly affected other canola seed germination variable, that is, each increase in salinity concentration was associated with marked reduction in seedling root

230 length. While treatments with bacterial isolates, especially strains L, K and Y, significantly lowered the reduction at 50 mM NaCl. Strains L and K retained this feature when the concentration of salt was 100 mM. At 200 mM NaCl, strains n and K caused greater root length than other treatments (Fig. 5.2a). Strains L, K and Y were able to increase whole seedling dry weight at 0 and 50 mM NaCl (Fig. 5.2b). At 100 mM NaCl strains L and K increased dry weight by 30 %, while strain n did not exhibit any effect. At 150 mM NaCl, only strain K increased dry weight, by 9 %, with respect to the non-inoculated treatment (Fig. 5.2b).

Fig. 5.2 Effect of bacterial treatment on canola germination traits under various concentration of NaCl. Sterilized canola seeds were treated with bacteria and were then placed on filter papers in Petri plates. The length of roots (a) and dry weight (b) of all individual seedlings were measured at the end of day 7. Each histogram bar represents the mean ± standard error based on three biological experiments, with 100 seeds per treatment. Different letters above the bars indicate statistically significant differences between treatments (P < 0.05). An asterisk indicates a statistically significant difference from controls at P < 0.05.

5.4.2.2 Pot experiment Experiments with initially sterile soil in pots showed an extraordinary capacity of strains to increase measured variables, relative to the non-treated control in canola (Fig. 5.3). Specifically, inoculation with strains L and Y significantly increased total leaf area to 56 cm3 plant-1, as compared to 40 cm3 for control plants (Fig. 5.4a). Increases in chlorophyll content

231 were only caused by strain Y (Fig. 5.4b). Inoculation with strains L, K and Y increased above-ground fresh weight and dry weight, as well as root fresh weight by ca. 21, 18, and 33 %, respectively (Fig. 5.4c,f,e). Strain n increased above-ground dry weight by 14 % (Fig. 5.4f), while all the other variables were not different from the uninoculated control.

Fig.5.3 Response of canola seedlings to inoculation with the four tested rhizobacterial strains. The seedlings were 35-d-old grown in sterile soil. Seedlings were inoculated with different strains with a control plant not treated with any of the bacteria (control).

232

Fig. 5.4 Effect of bacterial treatment on canola growth. Sterilized canola seeds were treated with bacteria and were then sown in pots. The chlorophyll index (a), total leaf area (b), shoot fresh weight (c), root length (d), root fresh weight (e) and shoot dry weight of individual plants were measured at the end of day 35. Each histogram bar represents the mean ± standard error from three biological experiments. Different letters above the bars indicate statistically significant differences among treatments (P < 0.05).

233 5.4.3 Maize inoculation experiment 5.4.3.1 Effect of PGPR on in vitro germination and growth In this study, we tested the influence of all four selected bacterial strains on germination variables of maize seeds under unstressed and salt stressed conditions. Inoculation of maize seedlings with selected rhizobacteria had significant effects on root elongation, shoot length, whole seedling fresh weight and vigor index (Table 5.1). When under optimal conditions, all rhizobacteria increased shoot length up to 1.28-fold over the uninoculated control. Rhizobacterial isolates K and Y were the most effective and caused 1.28 and 1.30-fold increases, respectively, in shoot length, over the uninoculated control, followed by strain L (1.10-fold). Note that only strain Y induced an increase of root length, by 29.3 %. The fresh weight of seedlings was significantly increased in bioprimed maize seeds (1.10 to 1.25-fold), while the dry weight was unaffected. Strains K and Y induced the highest seed vigor index with an increase up to ca. 20 % compared to controls, while strain L caused a 1.12-fold increase. In the glass tube experiment, significant increases in the shoot and root dry weight of seedlings, in response to inoculation, over uninoculated controls, were measured (Fig. 5.5). Under unstressed conditions, seedlings inoculated with strains L and Y caused 1.6- and 1.8-fold increases in dry biomass of seedling shoot, respectively, along with significantly enhanced root dry weight due to inoculation of all four strains (Fig. 5.6), over uninoculated controls. Under salinity conditions (100 mM NaCl), seed germination percentage was not significantly affected by bacterial treatments or salt stress, but seedlings inoculated with strains L and K showed a significant increase in the dry weight of shoot biomass (data not shown).

234

Fig.5.5 Effect of inoculation by the four selected rhizobacterial strains on maize seedlings at 28 d after growing in a glass tube experiment under gnotobiotic conditions in a growth chamber.

235

Fig.5.6 Boxplot for biomass (dry weight; mg seedling-1) of 21-d-old maize seedling inoculated with the four selected endophytic rhizobacterial strains, under controlled conditions. The total range, interquartile range (boxes) and means (dots) were displayed. Asterisks indicate the statistically significant differences compared to controls (P < 0.05).

236 Table 5.1. Effect of isolated rhizobacteria on the maize germination parameters

Strains Germination (%) Shoot length (cm) Root length (cm) Vigor index Fresh weight (g)1 Dry weight (g)1

Control 83.3 6.4c 14.0b 1769c 0.83c 0.28 n 80.0 7.0b 13.7b 1718c 0.91b 0.30

L 80.0 7.4b 15.2b 1987b 0.99a 0.32

K 88.3 8.2a 13.2b 2062ab 1.04a 0.32

Y 88.3 7.9a 18.1a 2362a 1.03a 0.33

In a column, the means with different letters are significantly different than the probability level of 5%

1 Values are presented as seedling weight.

237 Table 5.2. Effect of isolated rhizobacteria on the growth parameters of maize plants at 35 DAS.

Strains Height (cm) Circumference (cm) Root length (cm) Leaf area (cm2)

Control 73 3.02 571c 663.46b n 69 3.24 689.73b 670.51b

L 83.8 3.42 784.25a 848.36a

K 76.6 3.30 778.10a 814.57ab

Y 82.4 3.36 785.96a 910.13a

Within a column, means with different letters are significantly different at the probability level of 5 %

Table 5.3. Effect of isolated rhizobacteria on biomass produced by maize plants at 35 DAS.

Aerial dry Strains Aerial biomass (g) Aerial dry weight (g) Root dry weight (g) matter (%)

Control 33.06b 3.8b 11.74 1.228b n 34.08b 3.6b 10.53 1.232b

L 39.92a 4.86ab 12.09 2.338a

K 37.86ab 4.44ab 11.66 2.129a

Y 44.1a 5.2a 11.78 2.103ab

Within a column, means with different letters are significantly different at the probability level of 5 %

5.4.3.2 Effect of PGPR on maize growth under pot (controlled environment) conditions The effect of four selected rhizobacteria on the greenhouse growth of maize plants under initially non-sterile conditions, at 35 DAS, was variable. All the tested strains had considerable

238 impact on growth parameters. There were no significant differences in the height of maize plants (Table 5.2). The plants inoculated with any of the four strains had numerically larger circumferences, though not statistically significantly different from controls. Treatments with strains L, K or Y improved leaf area per maize plant by, respectively, 27.87, 22.78 and 37.25 %, compared to the control. The same treatments also resulted in longest root length, with an increase of about 37 % (Table 5.3). Strain n promoted root length to a lessor extent, but still significantly greater than the control. Results indicated that highest values for growth variables were achieved by inoculation with strains L, K and Y, as compared to strain n and the un-inoculated control. A significant difference was also observed between the effects induced by rhizobacterial strains for biomass production by maize plants at 35 DAS under greenhouse conditions (Table 5.3). Strains L and Y induced greater production of fresh aerial biomass, with respective increases of 20.75 and 33.39 %, compared to controls. The plants treated with strain Y produced the highest above-ground dry weight, inducing an increase of 36.84 %, compared to the control. Strains L and K increased root dry weight by 90.39 and 73.37 %, respectively. Contrary to the three above mentioned variables, rhizobacterial inoculation had no significant effect on the production of aerial dry matter by the maize plants. The maximum values for biomass production in our case were, again, obtained following the application of strains L, K and Y. Variation in efficiency among these three strains was not apparent for maize growth variables or biomass production.

5.4.3.3 Total carbohydrate The effect of the four selected rhizobacteria on total carbohydrate in greenhouse-grown maize plants under non-sterile conditions at 35 DAS was investigated during the current study (Fig. 5.7a). Carbohydrate content was 4.2 mg g-1 in the control plant, PGPR treatment increased carbohydrate content in maize leaves from 37.62 (5.8 mg g-1) to 102 % (8.5 mg g-1).

5.4.3.4 Antioxidant activities and proline content in leaves Generally, inoculation with PGPR significantly increased the antioxidant activities in maize leaves (Fig. 5.7b). Among the treatments, inoculation with strains n and L resulted in significantly higher scavenging effect (1.36- and 1.33-fold increase, respectively) against DPPH

239 radicals. The same two treatments also resulted in significantly higher total phenolics (32 and 27 % greater than the uninoculated control, respectively) in maize leaves. Our findings indicated that plants synthesized proline to a greater extent (76 and 84 % increases) when inoculated with strains n and L, respectively. Strains K and Y had no effect on proline content of maize leaves, relative to the non-inoculated control treatment (Fig. 5.8a). The total phenolic compound contents in leaves were enhanced by NaCl. But an increase under salt stress in the content of polypehnols was observed when bacteria (strains n and L) were present.

Fig. 5.7 Effect of bacterial inoculation on the production of carbohydrate (a) and ROS scavenging activity (b) of maize. Sterilized maize seeds were sown into pots and treated with bacteria on days 0 and 7. Leaves were harvested at 35 DAS. Different letters associated with columns indicates statistical difference by t test analysis with P value < 0.05, as compared to control. The experiment was repeated twice.

240

Fig. 5.8 Effect of bacterial inoculation on the production of proline content (a) and total phenolics (b) of maize. Sterilized maize seeds were sown into pots and treated with bacteria on days 0 and 7. Leaves were harvested at 35 DAS. Different letters associated with columns indicates statistical difference by t test analysis with P value < 0.05, as compared to control. The experiment was repeated twice.

5.4.3.5 Acquisition of nutrient content by maize Inoculation of PGPR not only enhanced maize growth, but also influenced nutrient uptake by maize plants at 35 DAS (Table 5.4). Compared to untreated controls, maize plants treated with strain n had significantly higher leaf Fe and P contents, by 72.09 and 64.36 %, respectively. Each of the four strains notably increased the N levels in the aerial biomass of maize plants, by 40.31, 56.59, 28.68, and 93 %, over the non-inoculated control. However, only strains L and Y enhanced N/C ratios over the uninoculated control. The other macro- and micro-nutrients were not significantly influenced by bacterial treatments, as compared with untreated controls, except that strains L and Y significantly lowered the amount of K and Mg, respectively.

There were huge differences in the concentrations of nutrients in the roots, when compared to those in shoots (Table 5.5). Interestingly, all the tested minerals in the roots of maize plants were not significantly influenced by bacterial treatment, except that the N content was increased up to 173 and 112 %, with treatment by strains L and K, respectively, but only strain L showed a highly significant increase in N/C ratio, as compared to all the other treatments.

241 Table 5.4. Influence of rhizobacteria on nutrient content and C and N composition (dry weight basis) of maize leaves at 35 DAS.

P K Mg Ca Na Fe Cu Carbon (%) Nitrogen (%) N/C (mg (mg (mg (mg Treatment (μg g-1) (μg g-1) (μg g-1) g-1) g-1) g-1) g-1)

Control 1.01b 27.1ab 1.99a 2.7ab 8.13 40.59b 2.76 50.4 1.29c 0.026b n 1.66a 29.64a 2.06a 3.3a 7.12 69.85a 2.68 47.4 1.81b 0.038ab

L 1.05b 22.13c 1.72ab 2.53ab 6.09 40.26b 1.92 49.4 2.02a 0.041a

K 1.20ab 26.1abd 1.76a 2.56ab 6.25 32.46b 2.12 50.6 1.66b 0.033ab

Y 1.04b 22.62bc 1.38b 2.18b 6.36 46.30b 1.75 50.3 2.49a 0.050a

Within a column, means associated with different letters are significantly different at a probability level of 5 %

242 Table 5.5. Influence of rhizobacteria on nutrients content and C and N composition (dry weight basis) of maize roots at 35 DAS.

P K Mg Ca Na Fe Cu Carbon (%) Nitrogen (%) N/C (mg (mg (mg (mg Treatment (μg g-1) (μg g-1) (μg g-1) g-1) g-1) g-1) g-1)

Control 0.43 8.96 0.71 1.2 245.14 149.09 5.48 47.2 1.19c 0.025b n 0.48 8.72 0.87 1.6 318.16 195.8 7.71 46.2 2.2abc 0.049ab

L 0.38 8.47 0.76 1.03 246.58 109.89 4.19 44.4 3.26a 0.073a

K 0.45 9.22 0.88 1.48 374.17 182.9 4.82 46 2.52ab 0.055ab

Y 0.44 6.85 0.82 1.93 242.77 233.9 9.52 46.8 1.47bc 0.032b

Within a column, means associated with different letters are significantly different at a probability level of 5 %

243 5.4.4 Endophytic colonisation Petrie plate counts from roots of all inoculated plants resulted in numerous bacterial colonies on selective media plates, but none were observed from plates of control plants. The roots were macerated and spread on KB plates to obtain inoculated bacteria. All four strains were recovered from inside the roots, indicating that they had colonized the canola and maize roots endophytically. Strain L colonized the root interior at 1.5×105 CFU g-1 FW, while strains n, K and Y colonized the root endosphere in both crops at ca. 4.5×106 CFU g-1 FW. Seven-d-old seedlings were used for this experiment; hence, population size at plant maturity may well have been larger.

5.5 Discussion Maize and canola can establish a wide range of rhizospheric and/or endophytic associations with various indigenous PGPR (Ahmad et al., 2016; Arruda et al., 2013); many of which may provide beneficial effects to the associated plants, ultimately contributing to better growth of the entire holobiont. For instance, Montañez et al. (2012) reported that numerous putative endophytes were isolated from the root, stem and leaf tissues of maize cultivars and indicated beneficial associations of maize plants with some of these bacteria, such as Pseudomona fluorescens EMA-38 and Herbaspirillum frisingense EMA-117. In another study, interaction of intrinsic diazotrophic bacteria, including Pseudomonas spp. and Bacillus sp., with maize significantly increased biomass and chlorophyll content (Kifle and Laing, 2016a). More recently, Ghavami et al. (2017) showed that two siderophore-producing rhizobacteria, Micrococcus yunnanensis and Stenotrophomonas chelatiphaga, from the canola rhizosphere, significantly increased shoot and root biomass production and iron uptake by canola. There have also been numerous studies indicating the beneficial association of maize and canola with non-host PGPR, including species within the genera of Pseudomonas, Bacillus, Azospirillum, Sphingobacterium, etc. (Ghavami et al., 2017; Marques et al., 2010; Oskuei et al., 2017; Puri et al., 2015). The object of the present study was to determine if the four PGPR strains (Pseudomonas sp., Bacillus sp., Mucilaginibacter sp. and Rhizobium sp.), previously isolated from reed canary grass, bittersweet, fall dandelion, and soybean, respectively, are able to colonize internal tissues of canola and maize and promote seed germination and growth.

244 Enhanced plant growth in canola and maize as a result of seed treatment with these four strains in the current study is comparable to the findings for A. thaliana reported in Chapter 4. In the first part of our investigation, we determined seed germination effects of the four selected strains for canola and maize under in vitro conditions. All four strains induced better germination of canola, increasing root length and seedling dry weight, under both normal and salt stressed conditions (50 to 200 mM NaCl) in Petri dishes, while strains L (Bacillus sp.), K (Mucilaginibacter sp.), and Y (Rhizobium sp.) outperformed strain n, a Pseudomonas. The promotion of canola seed germination, as a result of seed treatment with the selected strains was similar to the findings for maize, in which shoot and root length, seedling fresh weight, and vigor index were significantly increased after inoculation with strains L, K and Y. Similar improvement of seed germination variables by PGPR has been reported in pearl millet (Niranjan et al., 2004) and maize (Nezarat and Gholami, 2008). These findings may be due to better synthesis of endogenous plant growth regulators (e.g. auxins and cytokinis) which can stimulate cell division and elongation (Su et al., 2011). Inoculation with selected strains also decreased negative salinity effects on canola seed germination and in vitro maize seedling growth. NaCl in germination media causes osmotic and other toxicity effects and altered processes of water uptake, which may retard and reduce final germination percentage. There was no reduction in canola seed germination rate even under salinity stress (up to 150 mM). Al-Thabet et al. (2004) reported that reduction of seed germination was obvious when NaCl concentration was increased to 150 mM. This may indicate that the canola cultivar used in this study was more salt tolerant than most. Since the seed germination rate was reduced by increasing salt concentrations, 100 mM NaCl was used for maize work (data not shown). High salinity can affect the growth and physiological performance of bacteria (Yan et al., 2015). Bacteria accumulate osmolytes, mainly proline, glycine betaine, sugars and potassium cations, as part of adaptation to osmotic stress and ion toxicity (García et al, 2017; Yan et al, 2015). We observed that growth and PGP traits, in the case of ACC deaminase activity and IAA production, associated with strains n and L, were not negatively influenced by salinity (100 and 150 mM NaCl), strain Y showed non-significant reduction in these, while the PGP traits tested (data not shown) and growth of strain K were reduced significantly. Nevertheless, our results indicated that strains n and L are more tolerant to salt stress than strains K and Y. Intriguingly, strain K stood out as one of the most efficient PGPR in the promotion of

245 canola and maize seed germination and early seedling growth of Arabidopsis under salinity stress, indicating that as a true endophyte, strain K might be protected from abiotic stresses when inside plant root tissue, and suggests that there are other mechanism(s) adopted by this strain to alleviate salt stress, other than the commonly recognized PGP traits. In pot experiments, it was observed that inoculation with strains L, K and Y significantly increased leaf area, above-ground biomass and root length of canola, by ca. 40, 20, and 98 %, respectively, while strain n only caused greater above-ground dry weight than the uninoculated control. Again, the three selected strains L, K and Y induced the best growth of maize plants, with increases in leaf area and root length of about 37 and 36 %, respectively. In general, inoculation resulted in enhanced early seedling growth, similar to the findings of Nezarat and Gholami (2009) and Pérez-Montañ et al. (2014). There have been several reports showing that PGPR inoculation induced synthesis of carbohydrate in maize (Abd El-Ghany et al., 2015; Subramaniyan et al., 2012), in agreement with the current study in that we found inoculation with strains L, K and Y resulted in higher total carbohydrate contents (ca. 7.0 mg g-1) than -1 control plants (ca. 4.0 mg g ). Some of the bacterial traits, such as N2-fixation, phosphate solubilization, siderophore production, and IAA, cytokinin and gibberellin synthesis, have been shown to exert positive influences on plant growth by increasing nutrient availability, root and shoot development and overall plant growth (Glick, 2010; Kiani et al, 2016; Mahmood et al., 2016). Therefore, growth promotion in canola and maize by the tested strains may be mediated by one or more of these traits, through the colonization of the root cells and enhanced root surface area, and consequently, more acquisition of nutrient and plant hormones. IAA is one of the mechanisms PGPR use to promote plant growth; this mechanism regulates several aspects of plant growth and development such as lateral root initiation, cell enlargement, cell division and increased root surface (Zhao, 2010), leading to increased N-uptake due to hormonal effects on root morphology and activity. Though all four bacterial isolates studied here produce IAA (Chapter 4), their levels were quite a bit lower than those produced by other PGPR reported to promote growth of various crop plants (Majeed et al., 2015; Sharma et al., 2016; Zahid et al., 2015), leading to uncertainty regarding the importance of this mechanism. Nitrogen fixation also contributes to plant growth promotion, as evidenced through a radiolabeling experiment with Herbaspirillum seropedicae and Azospirillum brasilense, which promoted weed growth by incorporating biologically fixed N into major metabolic processes

246 (Pankievicz et al., 2015). In our work total N content was statistically superior for all four evaluated strains (increases of 40 to 93 % compared to the control). Strains n, L and Y were proven to be nitrogen fixers while strains L, K and Y performed best at growth promotion of canola and maize, indicating that agronomically significant levels of N were probably not primarily supplied through the N2-fixation ability of PGPR, or at least not for strains n and K, suggesting that there may be alternate mechanisms that are more important than contributions through N2-fixation. It would be helpful to elucidate the contribution by N2-fixation using 15 atom % N excess in foliage, to analyze the percentage of foliar N derived from atmospheric N2 (Puri et al., 2015). The increase in root length due to PGPR inoculation may also have contributed to the increased N content in maize leaves as the two variables were significantly correlated (R2 = 0.71). Chlorophyll is absolutely critical in photosynthesis for production of food. Kandasamy et al. (2009) reported that Pseudomonas fluorescens enhanced expression of ribulosebisphosphate carboxylase large chain precursor, which plays a principle role in chlorophyll accumulation and photosynthesis. Chlorophyll content index of canola following inoculation with strain Y and maize inoculated with strains L and Y was increased over control levels, indicating that bacterial treatment might improve photosynthetic activity/efficiency of canola and maize, which could be a mechanism used by the two strains to cause plant growth stimulation. Similar observations have been reported in earlier studies (Glick et al., 1998; Kumar et al., 2016; Lee et al., 2009; Singh and Jha, 2016; Zhang et al., 2008), in which an increase in total chlorophyll content of 23.7 to 33 %, compared to non-bacterized controls, was reported. Ricci (2015) reported that root treatment with strains n and L significantly enhanced some growth variables of tomato plants in growth chamber experimentation, as compared to the non-inoculated controls, such as shoot and root dry weight, root length, leaf area and plant height. This is not in accord with the present study, in which strain n did not show as much growth promotion effect in canola or maize as other bacterial treatments, suggesting that this strain might be somewhat plant species specific, and probably perform better under biotic stress conditions. Researchers, for example, Breedt et al. (2017) and Egamberdiyeva (2007), found that heavier and more fertile soils led to poor PGPR performance whereas lighter and poorer soils resulted in better performance. The beneficial effects of PGPR in maintaining adequate levels of mineral nutrients such as P and Fe in crops have been reported by Majeed et al. (2015) and Ghavami et al. (2017),

247 respectively. In our study, strain n significantly enhanced Fe and P levels in maize leaves, as compared to all other treatments, under pot conditions, indicating that this strain probably facilitated the uptake of these two nutrients, in agreement with the in vitro findings. When tomato plants received insoluble iron and phosphate, plants inoculated with strain n had greater growth enhancement (i.e., plant height and shoot and root dry weight) than the non-inoculated controls (Ricci, 2015) in a pot experiment, indicating that this strain might perform better under nutrient deficient conditions. Bacterial-treatment of maize decreased Na shoot concentration, along with significantly lower Na/K ratios than non-inoculated plants (data not shown), indicating that these strains might assist in reducing salt toxicity through lower Na/K ratios in shoots; this is considered as a predominant mechanism to enhance plant growth under salt stress conditions (Munns and Tester, 2008). Similar observations were reported for crops such as inoculated peanut (Sharma et al., 2016; Shukla et al., 2012), maize (Abd El-Ghany et al., 2015) and wheat (Ramadoss et al., 2013). Internal tissue colonization of canola and maize roots was observed under gnotobiotic conditions, with population densities similar to those observed in previous reports of these bacterial strains in Arabidopsis (Chapter 4), indicating that these four strains are versatile root colonizers. These bacteria readily oxidized about 11 to 18 carbon substrates, some of which are found naturally in plants, such as glucose, sucrose, raffinose and maltose, indicating that these strains might interact mutualistically with host plants by supplying nutrients and/or plant hormones in return for access to a nutrient (reduced carbon) rich environment following colonization of the plant. Enhanced ROS scavenging ability has been considered as an important strategy to alleviate oxidative damage in plants exposed to abiotic and/or biotic stresses. Synthesis of phenolic compounds by plants is one adaptive mechanisms for reducing oxidative damage (Hichem et al., 2009). Proline has been reported to be both a compatible solute, crucial for maintaining the osmotic potential of tissues (Wang et al., 2012), and a hydroxyl radical scavenger leading to the stabilization of proteins, membranes and cellular functions (Ozden et al., 2009). In this study, the increases in DPPH scavenging capacity, total polyphenol content, and proline content in maize plants treated by strains n and L under non-stressed conditions indicated that bioprimed plants might be better equipped to withstand abiotic stressors, as shown in germination tests under saline stress, and/or phytopathogens. Sing and Jah (2016) reported that

248 Serratia marcescens induced systemic resistance to Fusarium graminearum and enhanced salt tolerance of wheat, along with increased production of antioxidant enzymes, soluble sugars and proline. Inoculation with two Azotobacter strains increased the accumulation of proline and total polyphenols, leading to the amelioration of saline stress in maize (Rojas-Tapias et al., 2012). However, there has been some controversy about proline as an indicator of stress responses. Proline content in wheat was increased significantly under salinity stress by inoculation with Dietzia natronolimnaea (Bharti et al., 2016) but reduced by Pantoea alhagi (Chen et al., 2017). Moreover, inoculation with PGPR significantly enhanced the proline content of wheat (Singh and Jha, 2016) under both non-saline and salt stress conditions. Rojas-Tapias et al. (2012) showed an increase in proline contents due to treatment with PGPR under normal conditions, along with a decrease in proline concentration in maize leaves under saline stress, all of which indicated that PGPR might invoke some biotic stress responses, which triggered proline biosynthesis in plants or perhaps suppressed the NaCl-induced damage through an alternative strategy for the activation of enzymatic and non-enzymatic organic antioxidants/osmolytes such as glycine betaine, SOD, CAT, etc. (Kim et al., 2017). The four tested strains did not influence proline content in Arabidopsis leaves under normal conditions, while strains L and Y significantly increased proline content under salinity stress (Chapter 4). Thus, it would be ideal to elucidate the changes in proline accumulation following inoculation of selected strains onto canola and maize, under saline conditions. More intriguingly, some researchers observed a decrease in the activities of antioxidant enzymes such as superoxide dismutase (SOD) and ascorbate peroxidase (APX) in wheat (Shaik et al., 2011) and SOD, APX, and guaiacol peroxidase (GPX) in tobacco (Kumar et al., 2016) as a result of PGPR priming under salt stress and virus infection, respectively. All these findings suggested that PGPR might better control the intracellular redox with lowered proline and ROS in bioprimed plants under stress conditions. There are many known ways that PGPR might promote plant growth. Bacteria within the species of Pseudomonas, Bacillus, and Rhizobium were reported to be involved in biocontrol (i.e., ISR induction, production of HCN, exo-polysaccharides, and anti-microbial secondary metabolites), biofortification (i.e., solubilization P, K and zinc, and production of siderophores), production of plant hormones and ammonia, ACCD activities, N2-fixation, etc.) (Ahemad and Kibret, 2014). Only one study was previously conducted with Mucilaginibacter sp.; it showed that two Mucilaginibacter strains possessing ACCD activity and able to oxidize sulfur enhanced

249 tomato and canola root length in gnotobiotic growth pouch assays (Madhaiyan et al., 2010). In the present work, the different response patterns of canola and maize following inoculation with the four PGPR strains indicated that the plant growth promoting effects induced by the four tested PGPR were dependent on plant species and bacterial strain (Brusamarello-Santos et al., 2017), and possible as yet to be identified alternative mechanisms; in addition complex feedback loops might well be used and exploided by PGPR to enhance plant growth under normal and/or salt conditions (Sharma et al., 2016). Until now, there is still uncertainty regarding the specific role of PGP traits (i.e., P solubilization, production of phytohormones, antibotics, and siderphores) in plant growth stimulation by rhizobateria of the phytomicrobiome (Rodrıgueź and Fraga, 1999). Though indigenous/plant specific bacterial strains are typically preferred due to their adaptation to local environments and competitiveness as compared to foreign strains (Bhattarai and Hess, 1993); the four tested bacteria here have growth promotion effects on Arabidopsis plants, as well as canola, maize and tomato, all of which are non-host plants for these isolates. Further studies, such as light and electron microscopic research on leaves, stems, and roots, plus yield evaluation, biophysical and biochemical studies (i.e. structural integrity and activities of stress related enzymes) as well as genomic, transcriptomic, proteomic and metabolomics studies using ‘Omic’ technologies are needed for a more detailed assessment of the effect of the four PGPR on canola and maize.

5.6 Conclusions We reported in Chapter 4 that all four selected rhizobacterial strains, independent from the origin, exhibited a variety of direct and/or indirect PGP traits, and significantly promoted Arabidopsis growth. However, bacterial growth promotion is sometimes species or genotypic specific. The current study demonstrates that, inoculation with three out of four selected strains resulted in significant plant growth promotion effects, not only in Arabidopsis (Chapter 4) but also in canola and maize, indicating these strains might have a broad spectrum of growth promotion in a wider group of plant species, with possible enhancement of salt stress tolerance, and making development and commercialization of these strains for sustainable agricultural production a viable option. Moreover, results from experiments with non-sterile soil, mimicing greenhouse conditions, showed similar growth promotion effects to those observed in sterile conditions, implying that these PGPR are able to compete with other flora in rooting media and

250 promote the growth of inoculated plants. Taken together, these beneficial endophytic bacteria, especially strains L, K and Y, are promising for use as alternative to synthetic chemicals to enhance crop growth (Dutta et al., 2014; Farrar et al., 2014). As this was an initial study of the association of the four selected strains with agricultural crops, additional study should be undertaken to analyze the performance of these inoculants on the yield under field conditions, on a range of soil types of soils and under nutrient deficiency and biotic and abiotic stress conditions, potentially leading to further savings to growers through reductions in chemical fertilizer and/or biopesticide use. These strains, and others like them are also of great potential for enhancing the efficiency of biomass production for sustainable and profitable production of biofuels from canola and maize, particularly if the negative impacts of environmental stressors such as salinity, drought, heavy metals, and infertility can be mitigated by PGPR inoculation. However, a more comprehensive and detailed study is needed to ascertain the extent to which these PGPR strains promote host plant production and resistance to salt stress. Numerous publications regarding growth promotion effects of Pseudomonas spp, Bacillus spp, and Rhizobium spp on canola (Kamal et al., 2015; Noel et al., 1996; Wang et al., 2014) and maize (Kuan et al, 2016; Mumtaz et al., 2017; Piromyou et al., 2011) have been published. However, to the best of our knowledge, there is no detailed study regarding PGP effects of Mucilaginibacter (stain K) on crop plants, though Mucilaginibacter has been shown to be one of the most abundant genera in agricultural soils (Quadros et al., 2012). As such, this is the first comprehensive report from Canada about the potential agricultural applications of Mucilaginibacter sp., in particular, from a wild plant, fall dandelion.

251 Chapter 6 Concluding remarks

Food security, climate change, and energy security are three inter-connected challenges that humankind must face in the 21st century (Lal, 2010). Agriculture, a fairly recent human invention, emerged about 10,000 years ago (Price, 2009). For quite a long time, the need for food, as well as fiber, biofuel and other products, have been met largely through traditional and then more modern agricultural production practices, i.e., deforestation to produce arable land, acknowledgments, use of high-yield hybrid seeds of a single crop and genetically modified crops, technologically advanced equipment, energy subsidies in the form of irrigation, chemical fertilizers, pesticides, etc., all of which are of high social and environmental costs, i.e., decreased biodiversity, greenhouse gas emissions, and eutrophication of waterways. Though modern agriculture uses very innovative practices to produce enough outputs for a growing world population, it also attempts to minimize its environmental footprint, leading to the emergence of the sustainable agriculture concept, an integrated system of plant and animal production practices that, over the long run, meets three goals of sustainability: (1) preserve or enhance environmental quality and natural resources, (2) meet human food, fibre and fuel needs and (3) be economically viable for both farm operators and consumers. There have been some regenerative agricultural practices adopted, such as crop rotation, polyculture, crop spatial organization, fallowing and mulching, biocontrol, and methods to increase soil fertility and reduce soil erosion. These transformational suites of practices can lead to fewer chemical inputs and greater profits. With the rapidly growing human population, projected for approximately nine billion by 2050, and the higher living standards (generally associated with increased meat consumption), the need to produce more food while devising better social, environmental, and economic strategies for agriculture, is growing. All-in-all, our primary challenge is to increase agricultural yields while reducing the need for chemical fertilizers, pesticides, water and other practices with negative environmental effectors. Bacteria have long been associated with other organisms. The most profound and intimate symbiosis is probably the plasmids and mitochondria, becoming organelles around 1.5 billion year ago, which might be the earliest evidence of bacteria assisting their eukaryotic hosts to survive, develop, and then thrive (Wang et al., 2015). Most eukaryote-microbe relationships

252 do not progress all the way to the microbes becoming organelles of their hosts; the predominant type of symbiosis is establishment of mutually beneficial unions, as is the case for symbioses between plants and microorganisms. The term “rhizosphere” was first coined at the beginning of the twentieth century by pioneer plant physiologist Lorenz Hiltner (Hartmann et al., 2008); it is a humid, nutrient-rich soil zone surrounding the plant roots and harbors a huge variety of plant microbiome (phytomicrobiome) members in a wide range of complex interactions with plants (Berendsen et al., 2012). The phytomicrobiome, especially the rhizomicrobiome, and more specifically, rhizobacteria, are the most diverse communities associated with plants as the soil surrounding plant roots is humid and contains high levels of organic nutrients. These rhizobacteria not only initiate complex associations with roots as free-living rhizobacteria, some of them can progress onto the root surface (ectophytic), between the cells, or even inside the cells (intracellular) of the plant roots (Gray and Smith, 2005). After more than a half-century of relative dormancy, the importance of the rhizosphere has begun to attract attention and appreciation, and now it has become one of the most intensely investigated areas of plant science (Brink, 2016). There have been numerous reports of plant growth promotion and yield stimulation by beneficial soil microorganisms (Adesemoye et al., 2008; Kuan et al., 2016; Pırlak and Köse, 2009). Plant growth-promoting rhizobacteria (PGPR) are the most effective and best studied soil microorganisms able to exert highly beneficial effects on plant fitness through direct or indirect mechanisms (Glick, 2012). They have been demonstrated to increase growth, productivity, or disease resistance of many commercial crops (Gupta et al., 2015; Paul and Lade, 2014). Microbial inocula have been shown to reduce movement of agricultural chemical inputs into the environment, such as fertilizers, especially nitrogen, and pesticides; they can also alleviate biotic (Rashid and Chung, 2017) and abiotic stresses (Paul and Lade, 2014; Vurukonda et al., 2016) in plants. Commercial development of microbe-based products, such as rhizobial inoculants (biofertilizers) and asymbiotic Bacillus thuringiensis (biopesticides), was initiated more than 100 years ago (Bashan, 1998) and has thrived since the 1950s (Timmusk et al., 2017). Development of a PGPR inoculant with commercialization potential is a complex task, involving isolation, screening for efficiency both in vitro and in vivo, and field trails (Timmusk et al., 2017). To add more pieces of knowledge to this rapidly growing body of research, this thesis investigated (1) the diversity of culturable root-associated bacteria from various plant species in

253 southwestern Québec, Canada and (2) their PGP effects in Arabidopsis thaliana and crop plants, Brassica napus (canola) and Zea mays (maize). Three underlying questions were addressed: (1) Can a diversity of root-associated plant-beneficial bacteria be isolated from the roots of crops and uncultivated plant species in southwestern Québec, using culture-dependent methods? (2) Are any of these isolatable rhizobacteria able to promote the growth of Arabidopsis thaliana under both optimal and salt stress conditions? (3) Do these microbes also promote the growth of selected key crop plants?

6.1 General discussion

The first goal of this thesis (Chapter 3) was to study the diversity of rhizobacteria from the rhizosphere and endosphere of various crops and uncultivated plant species in southwestern Québec (in and around the Macdonald Campus of McGill University) via culture-dependent techniques. Bacteria have evolved to be able to colonize essentially all the plants, in nearly every type of environment. They can inhabit in suboptimal environmental conditions and have developed specific biochemical and physiological properties to help their hosts survive. We aimed to sample (isolate and identify) this diversity of rhizobacteria from farm land, and also from an environment with far less human disturbance. We identified a field site with a good range of botanical diversity, as well as minimal human invention, with no chemical inputs for at least 25 years. Dr. Donald L. Smith and I sampled (in duplicate) the roots of each replicate of each uncultivated plant (21 species in total), including the rhizomes of one crop species (miscanthus). For the rest of the crop species, Dr. Fazli Mabood and I sampled (in quadruplicate) the roots of each replicate of each species, from an experimental farm site, at the Emile A. Lods Agronomy Research Centre in Sainte-Anne-de-Belleuve. From these, over 400 rhizospheric and endospheric bacterial isolates were obtained through growth on nutrient-rich solidified media. Based on the morphology on King’s B agar plates and 16S rRNA sequences, these isolates were shown to be 446 strains with high genetic diversity, belonging to 90 genera; among the most often identified from the rhizospheric soil were Pseudomonas (17 %), Bacillus (14 %), Flavobacterium (13 %), and Variovorax (6 %), while the endophytic bacteria were principally Pseudomonas (16 %), Actinobacteria (9 %), Bacillus (6 %), and Rhizobium (5 %). Though some

254 ubiquitous bacterial genera, such as Bacillus and Pseudomonas, have been identified across most of the samples explored (Chapter 3), there were also some specific bacterial communities involved with specific plant species, indicating that the root-associated microbiome component is, at least in part, determined to an important extent by the host plants. Among the isolated bacteria were some with 16S rRNA gene sequence pairwise similarities lower that 98.7 %, indicating they might be new species; other phylogenetic markers, such as fatty acids membrane profile, DNA-DNA hybridization with closest relative strains, multilocus sequencing with housekeeping genes, and almost complete 16S rDNA sequencing must be employed for reliable new taxa descriptions. Though we attempted to isolate root-associated bacteria as broadly as we could, it is likely that if different culture media had been used, different rhizobacteria might have been isolated and identified (Qin et al., 2009). As compared with previous scientific reports (Ramesh et al., 2012; Xia et al., 2015; Zinniel et al., 2002), the results reported in Chapter 3 are very promising with regard to the root-associated microbiome diversity, particularly given the relatively small sampling size and limited number of niches sampled in this initial effort and given that we did not address the fungal diversity. Moreover, since herbal medicinal plants were found to be associated with diversified and unique endophytes (Chapter 3), these coevolved bacterial strains might have exciting potential for improving the potency of medicinal plants, with regard to essential oils and bioactive phytochemicals that might help in diet or disease control (Ghodsalavi et al., 2013). Though researchers have shown that some bacterial phyla, such as Acidobacteria, are generally under-represented in culture (Wei et al., 2017), leading to more strains being present in the total bacterial communities than in the culturable microbiome (Mesa et al., 2017), the bacterial genera most abundant in our culturable assemblage coincided with the frequently detected genera in the total bacterial assemblages using pyrosequencing (Truyens et al., 2016). The culturable bacterial isolates obtained in Chapter 3 offered a rich reservoir of the diverse rhizobacteria as a potential source of novel taxa, and more importantly, for PGPR discovery; this set the stage for the studies in the remainder of this thesis, allowing us to screen for PGPR from the obtained bacterial pool. The choice was then made to use Arabidopsis thaliana to screen isolated bacteria for plant growth promotion, as described in previous reports (Huang et al., 2015), rather than to only screen for various common PGP traits (i.e., biological nitrogen fixation potential, phosphate solubilization, pathogen antagonist activities, and siderophore production), as we hypothesized that traditional PGP screening might discard

255 promising strains that did not perform in pure culture in vitro assays, as implied by Long et al. (2008). In addition, the effect of a PGPR will depend on its root colonization ability and interactions with other rhizobacteria (Compant et al., 2010). In Chapter 4, bacterial screening bioassays were initially conducted on A. thaliana Col-0 on MS agar plates and yielded four strains (out of 98) that consistently enhanced seedling growth as compared to controls. This is very intriguing given the small amount of sampling. We initially screened rhizobacteria under unstressed conditions, rather than abiotic or biotic stress conditions, based on two thoughts. Firstly, screening under stress conditions has often been linked to the natural biotic and abiotic status of host plants. For example, potential PGPR, isolated from plants grown under salinity stress conditions, were screened under salt stress in planta (Cardinale et al., 2015). Secondly, far more PGPR might be able to promote plant growth under stress conditions than un-stressed conditions, as has been previously observed, which will make strains initially screened under optimal conditions of particular interest. Thus, we hypothesized that there might be some rhizobacteria which can exert plant growth promotion even under more optimal growth conditions, and this turned out to be true, as was also evidenced by Huang et al., (2015). Seed treatment, as well as root inoculation, with the four selected bacterial isolates, led to consistent performance in either Petri dish assays (sterile) or soil experiments (initially sterile or non-sterile) in the growth chamber, which fortified our understanding of the PGP effects of these four isolates. The analysis of 16S rRNA gene sequences revealed that strain n is a Pseudomonas, L a Bacillus, K a Mucilaginibacter, and strain Y a Rhizobium. This is the first study of the PGP effects of a Bacillus from bittersweet, and a PGP strain, Mucilaginibacter sp., associated with Scorzoneroides autumnalis (wild dandelion). Moreover, root colonization ability in non-sterile growth medium, confirmed by GFP-labelling and plate counting, indicated that these bacterial isolates are efficient root colonizers for A. thaliana, indicating they are competitive with environmental bacteria and/or intrinsic endophytes. Though these isolates were shown to possess some PGP traits in vitro (i.e., P solubilization, ACCD activities, production of IAA-like substances, siderphore production, and N2-fixation), strain K, which possessed the least number of PGP traits (3 out of 8 tested), exerted the strongest growth promotion effects in A. thaliana, while strain n, the strain having the most PGP traits (100 % of tested), displayed a lower level of plant growth promotion. This proved the wise counsel by Dr. Donald L. Smith that I should directly look into the growth promotion effects of bacterial inoculation in planta, and not simply

256 search for already recognized methods of PGP. The same was found reported recently by Cardinale et al. (2015). The above-mentioned results have led to the conclusion that PGPR increased plant growth through a range of mechanisms, either by multi-activities of common PGP traits (Cardinale et al., 2015), through induced systematic resistance (Bakker et al., 2013), or by untested PGP activities, such as sulfur oxidization (Gahan and Schmalenberger, 2014), production of peptides and oligosaccharides (Subramanian, 2013), and quite possibly through currently unrecognized PGP activities. Besides being efficient in plant growth promotion, strains n and L also exerted antagonistic effects against two important phytopathogens, Pseudomonas syringae pv. tomato DC3000 and Botrytis cinerea B191, both in vitro and in planta. Moreover, quantitative real-time PCR revealed that these selected strains significantly changed the transcript levels of genes annotated to energy-, protein- and hormone metabolism in A. thaliana. All these results suggested that each isolate possessed multiple PGP activities, and indicated that combined application of these four PGPR, with specific sets of complementary benefits, might result in a more effective plant growth promotion than single inoculation, as shown by Delshadi et al. (2017) and Kumar et al. (2014). Some PGPR have been shown to promote plant growth under both normal and stressed conditions (Chen et al., 2017; Gagné-Bourque et al., 2016), thus, the effects of the four chosen isolates were evaluated in A. thaliana under salt stress in both soil-free sterile and soil non-sterile conditions, showing stress alleviation and growth promotion effects as compared with uninoculated control plants. This suggested that the beneficial effects of the four selected strains were stress-independent, while many of previous studies showed that beneficial effects of bacterial inoculation were stress-dependent (Cardinale et al., 2015; Rolli et al., 2015). The four bacterial strains identified in my work exerted growth promotion effects on A. thaliana under optimal conditions (full water and nutrient), indicating that the stimulation effect was probably through the production of plant growth regulators (auxins, gibberellins, cytokinins, etc.), either emitted by bacteria themselves or produced by plants upon biopriming. Having said this, we recognize that there could also be currently unknown mechanisms at work. All-in-all, the results from Chapter 4 suggested that the four rhizobacterial strains n, L, K and Y could stimulate plant growth and alleviate salinity stress in A. thaliana, leading to investigation of how they perform in crop plants. Since some PGPR induce plant growth enhancement in specific hosts, or a limited number of plant species (Zeller et al., 2007), while others exert growth-promotion effects on a

257 wide range of plant species (Batista et al., 2018), In Chapter 5, growth chamber and glasshouse experiments were set up using two crop plant species, Brassica napus (canola) and Zea mays (maize), to further test for growth promotion and salt stress alleviation effects of the four selected strains. The results of this investigation were presented, including effects on germination and seedling growth (in canola and maize), as well as the accumulation of some metabolites and nutrient uptake (in maize). Canola and maize were chosen because they are both important food and biofuel crops, whose production can be constrained by environmental stresses, including soil salinity. They are also among the three most economically important field crops in Canada. Though there are many methods of bacterial inoculation, we utilized the protocol described in Chapter 4. Results of seed inoculation onto the crop plants were significant and comparable to those with A. thaliana, indicating that the positive effects of the four isolated bacterial strains on plant growth that we observed with the model plant species, Arabidopsis, were also active with economically important crop plants. Inoculated maize and canola plants had enhanced growth, as detected through several measured growth variables and, overall, greater biomass production than uninoculated plants under both initially-sterile and non-sterile conditions, in growth chamber and greenhouse conditions, respectively. This is the first report about the potential agricultural applications of a Mucilaginibacter sp. from a wild plant, fall dandelion. It should be noted that, although strain K was not shown to be able to fix nitrogen, it increased the N content in the maize leaf tissue, along with the other three putative N2-fixers, who were equipped with a key gene involved in N2-fixation, reinforcing that the mechanisms under which PGPR stimulate plant growth can be very sophisticated. Thus, we felt that the positive growth effects observed under controlled and semi-controlled conditions resulted, at least partially, from the multiple PGP effects, as all of the tested bacteria possessed three or more of the plant growth promoting effects tested for, such as N2-fixation, P solubilization and auxin production. Interestingly, strain n was not the best growth promoter among the four tested strains, while it increased the contents of P and Fe, and did show some small, but statistically significant, growth stimulation effects on crops. This further strengthened our assumption that there could be great potential in deploying these strains as a consortium with synergistic positive effects on crop growth enhancement. The plants were given ½ strength Hoagland’s nutrient solution twice a week, which has been commonly used in a wide range of research studies, and generally thought to be more than adequate to meet plant growth requirements. Some greenhouse studies have shown that plants

258 grew better when the fertilization was combined with PGPR inoculation; in addition, bacterial root colonization was also increased under a medium rate of chemical fertilizers (Baldotto et al., 2012). This indicates that a promising biotechnological strategy for improving crop growth could be developed by biopriming the host plant in the presence of chemical fertilizer, which seems to be more reasonable under the real cultivation environment. PGPR inoculations have been shown to improve the flavor (Matsuoka et al., 2016) and nutrient contents (Banchio et al., 2009; Flores-Félix et al., 2015) of economic plants; in this thesis, the leaf content of nutrients such as N, P, Fe, as well as some secondary metabolites, including proline, total phenols and carbohydrates, were also influenced by bacterial inoculation, indicating the potential of these isolated strains to be used as efficient mechanisms for nutrient enhancement in food crops and the development of functional foods (Karakurt and Aslantas, 2010). In addition to the unstressed conditions, we also tested the effect of PGPR inoculation for salt stress alleviation for crop plants. More specifically, seeds of canola were incubated in bacterial suspension and were then exposed to different concentrations of NaCl for 7 d. The evaluated bacteria caused increased root length and seedling dry weight for canola, as compared to uninoculated plants. For maize, the germinated seeds were grown in agar tubes with 100 mM NaCl for 3 weeks, and inoculated seedlings had greater dry weight and root length than the controls. The conclusions we presented in Chapter 5 were that inoculation of canola and maize with the four selected endophytic bacterial strains enhanced early seedling growth and contributed salinity stress mitigation. The use of these four strains could be an effective strategy for improving the establishment and growth of crop plants, especially under salt stressed conditions. The results presented in Chapter 5, combined with the findings of Chapter 4, indicated the enormous potentials of these four strains in the context of crop production. These strains had multiple PGP traits, showing plant growth promotion and salt stress mitigation in A. thaliana, canola, and maize. All of them effectively colonized the roots of bacterized plants; two of them had effects in disease suppression. Given that the strains identified promoted the growth of both

C3 dicots and a C4 monocot, they are likely to be effective on a wide range of plant species. The use of PGPR products as sustainable agricultural inputs has been quite effective in many developing countries, as well as the developed world, where an important focus for this work has been organic agriculture (Timmusk et al., 2017).

259 There has been high-level use of agro-chemicals, especially fertilizers and pesticides, to improve the growth and yield of crops, which has led to ecological imbalances, and environmental deterioration, as well as risks to human health (Smith and Siciliano, 2015; Yadav et al., 2015). Crop productivity is regularly affected by a range of environmental stresses. Moreover, climate change (Chazdon, 2017) and its associated impacts on the environment, such as warming, increased precipitation, and depletion of renewable fresh water, threaten current biological niches, may devastate the precious biodiversity (https://www.ipcc.ch/report/ar5/wg2/) and could well diminish the current level of global food security. From this point of view, plants and cropping systems must adapt to the coming environmental changes and the related challenges, such as hot and cold stresses, drought, flooding, salinity, pests and diseases, all of which currently bring about daunting challenges to our ability to sustain adequate food supply for humanity, and all of which will be worsened by climate change. Thus, the use of PGPR as bio-inoculants to increase growth and yield and to suppress diseases, under various agricultural environments, and even in more extreme circumstances, contributing to sustainable agriculture, in which the use of various external chemical inputs is limited, is gaining momentum. It should also be noted here that potentially pathogenic rhizobacteria should be excluded from use as bioproducts, to maximize crop production while not increasing human (both crop producers and consumers) exposure to health risks (García-Fraile et al., 2012). If novel PGPR can be found to help crop plants tolerate stresses (Yang et al., 2009), such as drought, salt, and temperature changes, they will play essential roles in the development of climate change resilient agriculture. The results from this thesis show that there is an immense potential of bioprospecting the phytomicorbiome to increase crop productivity. However, there are challenges to widespread application of PGPR. Besides the specificity and selectivity of PGPR, the complex field environment, such as soil types and plant roots exudates (abiotic factors) and resident microbial community (biotic factors), can lead to inconsistent quality and efficacy of PGPR on plants. In order to enhance the likelihood for the effectiveness of PGPR, mathematical simulation based “tailored” inocula would facilitate the reproductability of PGPR and maximize the expected efficacy of PGPR in increasing crop production (summarized in Timmusk et al., 2017).

260 6.2 Future directions

This initial assessment of rhizobacteria from Québec has identified four very promising PGPR strains, however, there are still many gaps in our knowledge, in part, revealed by our findings; future experiments are needed to expand the work reported here and to further study the underlying mechanisms employed by these PGPR to promote plant growth.

1. Expand the range of bioprospecting of phytomicrobiomes and the methods for isolation and screening: • Sample root tissues from various environments, including agricultural lands, impoverished and/or unmanaged fields, contaminated soils, and forest soils. • Sample plant root tissues from wild progenitors of domesticated plants and also at centers of origin. • Sample roots over the course of plant development as the composition of the phytomicrobiome has been shown to change in a plant-controlled way as plants develop. • Isolate microbial strains from a wider range of plant species, such as crops, species related to crops, weeds/uncultivated plants, medicinal and ornamental plants. • Isolate both rhizobacteria and root-associated fungi, using various selective media, including those supplemented with plant root extracts. • Conduct initial screening for potential PGPR using stressful conditions (eg., salinity and drought stresses). • Check pollen and seed microbiota, along with root-associated bacteria, to study the possible transmission of bacteria from somatic to gametes/zygote.

2. Phenotypic and molecular microbiology: • Further characterize the four promising bacterial isolates, and any other identified, in vitro, such as for production of other plant hormone-like molecules (i.e., cytokinins and gibberellins), heavy metal resistance, and zinc and potassium solubilization. • Conduct DNA-DNA hybridization (DDH) to establish the identity of the four isolates to the species level, especially strain K. • Whole genome sequencing of these four strains, and any other identified, to expand our

261 understanding of them and their mechanisms of plant growth promotion.

3. Microbe-to-plant signaling: • Test for the possibility that microbial signaling molecule(s) might be responsible for some part of the observed plant growth promotion, using high-performance liquid chromatography (HPLC) techniques (for molecule fractions) and A. thaliana (for in vitro screening). • Isolate and identify these signaling compounds, should initial work show them to exist.

4. Additional microscopic, physiological and molecular work on A. thaliana: • Conduct light and electron microscopic research on the effects of the four strains, and any others identified, on development of leaves, stems, and roots. • Investigate the effects of the four strains, and any others identified, on flowering, seed setting and seed development, as well as overall yield. • Utilize Arabidopsis mutants to find genes critical to PGPR performance. • Conduct proteomics and metabolomics studies using Arabidopsis (both rosettes and roots), at various developmental stages, to assess effects of the four PGPR identified here in greater detail.

5. Agronomy related: • Light and electron microscopic research regarding effects of the four strains, and any others subsequently identified, on the development of leaves, stems and roots of canola, maize and other crops. • Investigate the in planta biocontrol activities of the four strains, and any others identified, against phytopathogens affecting maize, canola and other crops. • Initiate pre-field trial to confirm the PGP effects of selected rhizobacterial strains before the micro-environment become much more complicated (field conditions). • Study the effects of the four strains, and any others identified, appliled singly or as a mixture, on yields of canola, maize and other crops, grown on a range of soil types and under a range of climatic conditions (including climate-related abiotic stresses) and under nutrient deficiencies, to determine whether or not the growth promotion

262 observed during early growth stages can result in higher-yielding, lower input agricultural systems. • Study the effects of the four isolated plant growth-promoting strains, and any others identified, on the nutritional quality of maize, canola and other crops. • Investigate proteomic and metabolomic responses to the four strains, and any others identified, when inoculated onto maize, canola and other crops.

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