Investigating the role of Pseudomonas sp. and Bacillus sp. as plant growth promoting inoculants

By Emily Claudia Ricci

A thesis submitted to the Faculty of Graduate and Postdoctoral Studies in partial fulfillment of the requirements of the degree of Masters of Science

Department of Plant Science

McGill University, Montreal

Quebec, Canada

December 2015

©Emily Claudia Ricci, December 2015

TABLE OF CONTENTS

TABLE OF CONTENTS ...... i

LIST OF TABLES ...... v

LIST OF FIGURES ...... vi

ABSTRACT ...... vii

RÉSUMÉ ...... viii

ACKNOWLEDMENTS ...... ix

Chapter 1 ...... 1

GENERAL INTRODUCTION ...... 1

1.1 Introduction ...... 1

1.2 List of Objectives ...... 4

1.3 List of Hypotheses ...... 5

Chapter 2 ...... 6

GENERAL LITERATURE REVIEW ...... 6

2.1 Plant microbe interactions...... 6

2.1.1 ...... 6

2.1.2 Plant growth promoting rhizobacteria (PGPR) ...... 7

2.2 Direct mechanisms ...... 9

2.2.1 ...... 9

2.2.2 Phosphate solubilisation ...... 10

2.2.3 production ...... 11

2.2.4 Phytostimulators ...... 12

2.3 Indirect mechanisms ...... 13

2.3.1 Antimicrobial activity ...... 13

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2.4 Issues with planktonic inoculants ...... 14

2.5 Biofilms...... 14

2.5.1 formation ...... 14

2.5.2 Biofilm properties ...... 17

2.5.3 Biofilms as and phytostimulators ...... 18

2.5.4 Biofilms as biocontrol agents ...... 19

Chapter 3 ...... 20

MATERIALS AND METHODS ...... 20

3.1 Bacterial Strains ...... 20

3.1.1 Bacterial strain isolation and screening ...... 20

3.1.2 16s rRNA sequencing and identification ...... 20

3.1.3 Optimal growth conditions of mono-species planktonic cultures ...... 21

3.1.4 Standardization of mono and dual – species planktonic cultures ...... 22

3.1.5 Optimal growth conditions of mono-species biofilm cultures ...... 22

3.1.6 Standardization of mono and dual – species biofilm cultures ...... 23

3.1.7 Stereo microscopy imaging ...... 24

3.2 Phenotypic characterization of planktonic and biofilms cultures...... 25

3.2.1 Nitrogen fixing ability ...... 25

3.2.2 Phosphate solubilisation activity ...... 25

3.2.3 Inodole-3-acetic acid production ...... 26

3.2.4 Siderophore production ...... 27

3.2.5 Ammonia production ...... 28

3.2.6 Hydrogen cyanide production ...... 28

3.2.7 Antimicrobial activity ...... 28

3.2.8 Minimal bactericidal concentration ...... 29

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3.3 Effect of planktonic and biofilm inoculants on crop growth ...... 29

Chapter 4 ...... 31

RESULTS ...... 31

4.1 Bacterial strain identification ...... 31

4.1.1 16s rRNA sequencing analysis ...... 31

4.2 Growth curves ...... 33

4.2.1 Optimal growth conditions of mono-species planktonic cultures ...... 33

4.3 Biofilm formation assay ...... 35

4.3.1 Optimal growth condition of mono-species biofilm cultures ...... 35

4.3.2 Stereo microscopy imaging ...... 37

4.4 Optimization of bacterial cultures ...... 39

4.4.1 Standardization of mono and dual – species cultures ...... 39

4.5 Phenotypic characterization of planktonic and biofilms cultures...... 44

4.5.1 Nitrogen fixing ability ...... 44

4.5.2 Phosphate solubilisation activity ...... 44

4.5.3 Ammonia production ...... 47

4.5.4 Siderophore production ...... 47

4.5.5 Hydrogen cyanide production ...... 50

4.5.6 Inodole-3-acetic acid (IAA) production ...... 50

4.5.7 Antimicrobial activity ...... 52

4.5.8 Minimal bactericidal concentration planktonic/biofilm (MBC P/B) ...... 52

4.6 Bacterial crop inoculation ...... 56

4.6.1 Effect of planktonic and biofilm inoculants on crop growth ...... 56

4.6.2 Stereo microscopy imaging of the tomato root system ...... 63

Chapter 5 ...... 65

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DISCUSSION ...... 65

Chapter 6 ...... 73

CONCLUSION ...... 73

Chapter 7 ...... 74

REFERENCES ...... 74

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LIST OF TABLES

Table 1. Molecular identification of bacterial isolates based on blastN queries in NCBI...... 32

Table 2. Standardization of mono species cultures ...... 42

Table 3. Standardization of dual species cultures ...... 43

Table 4. Biochemical attributes of mono and dual-species biofilm and planktonic plant growth promoting rhizobacteria ...... 45

Table 5. Antimicrobial activity of biofilm and planktonic plant growth promoting rhizobacteria against 4 plant pathogens ...... 54

Table 6. Minimal bactericidal concentration of mono and mixed planktonic and biofilm cultures ...... 55

Table 7. The effects of Bacillus sp. inoculants on tomato plant growth variables of plants grown under greenhouse conditions...... 57

Table 8. The effects of Pseudomonas sp. inoculants on tomato plant growth variables of plants grown under greenhouse conditions ...... 58

Table 9. The effects of mixed inoculants on tomato plant growth variables of plants grown under greenhouse conditions ...... 59

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LIST OF FIGURES

Figure 1. Plant growth promoting rhizobacteria (PGPR) mechanisms...... 8

Figure 2. The process of biofilm development...... 16

Figure 3. Mono-species planktonic growth curves...... 34

Figure 4. Biofilm formation assays...... 36

Figure 5. Bacillus sp. and Pseudomonas sp. stereo microscopy (SM) imaging...... 38

Figure 6. Bacterial colony morphology ...... 41

Figure 7. Phosphate solubilisation activity of mono and dual-species biofilm and planktonic plant growth promoting rhizobacteria ...... 46

Figure 8. Siderophore production by mono and dual-species biofilm and planktonic plant growth promoting rhizobacteria ...... 49

Figure 9. Indole acetic-acid production by mono and dual-species biofilm and planktonic plant growth promoting rhizobacteria ...... 51

Figure 10. Stereo microscopy (SM) of Pseudomonas sp., Bacillus sp. and mixed inoculants on tomato root systems ...... 64

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ABSTRACT

Beneficial free-swimming planktonic plant growth promoting rhizobacteria (PGPR) have long been used as and biocontrol agents. However, their effects in the field are inconsistent, which has limited commercial application. This is probably caused by the inoculants’ inability to compete with existing endogenous microbial communities (Gupta et al, 2015). To overcome this issue, the use of biofilms is being investigated as potential PGPR inoculants. Biofilms are defined as dense colonies of single or multi-species of microbial cells, adherent to either a biotic or abiotic surface, encased in a self-produced matrix composed of extracellular polymeric substances (Davey et al., 2000). The formation of biofilms not only enhances the survival of but also allows them to continue plant growth promotion in ways similar to that of planktonic PGPR. Moreover, biofilms withstand a wide range of physical conditions including extreme temperatures, salt levels and pH as well as the presence of commonly used antibiotics (Baty et al., 2000 Todar, 2008) making the use of biofilm-PGPR (B-PGPR) a promising prospect.

The objective of this study was to develop and elucidate the potential use of novel Pseudomonas sp. and Bacillus sp. mono- and dual-species biofilm inoculants. To address this issue, optimization experiments for bacterial inoculants were conducted to standardize cell density (colony forming units) levels of each treatment in order to compare and contrast B-PGPR traits against their planktonic counterparts. Results indicate that mono- and dual-species biofilms exhibited enhanced values for various biochemical attributes, compared to their planktonic counterparts. These include phosphorus solubilisation as well as the production of indole-3-acetic acid (IAA) and . Moreover, greenhouse trials with tomato plants reinforced our in- vitro findings, suggesting that B-PGPRs are more effective inoculants than their planktonic equivalents. Results, therefore, suggested that the use of innovative B-PGPR technologies could prove extremely advantageous as biocontrol and biofertilizer agents, thus potentially alleviating some of the dependence on agro-chemicals for effective crop production.

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RÉSUMÉ

Les rhizobactéries planctoniques favorisant la croissance des plantes ont longtemps été utilisées comme biofertilisants et agents de biocontrôle. Par contre, leurs effets dans les champs sont inconsistants, ce qui limite leur application commerciale. Ceci est probablement causé par l’incapacité de l’inoculant à survivre dans un environnement avec des micro-organismes endogènes existantes (Gupta et al, 2015). Afin de surmonter ce problème, notre laboratoire a décidé d’enquêter sur l’utilisation des biofilms comme inoculant potentiel. Les biofilms sont définis comme étant des colonies denses d’une ou plusieurs espèces de cellules microbiennes adhérentes à une surface biologique ou non biologique enfermées dans une matrice autoproduite composée de substances polymères extracellulaires (Davey et al., 2000). La formation de biofilms améliore non seulement la survie des bactéries mais leur permet également de continuer la promotion de la croissance de manière similaire à celle des cellules planctoniques. De plus, les biofilms tolèrent une gamme de conditions physiques comprenant la température extrême, la teneur de sel, le pH ainsi que la présence d’antibiotiques couramment utilisés (Baty et al., 2000; Todar, 2008) qui rend prometteuse leur utilisation.

L'objectif de cette étude était de développer et d'élucider l'utilisation potentielle de biofilms mono ou double-espèces à base de Pseudomonas sp. et Bacillus sp. comme inoculant. Pour résoudre ce problème, des expériences d'optimisation des inoculants bactériens ont été menées pour normaliser la densité cellulaire (unités formant des colonies) de chaque traitement afin de comparer et de contraster les caractères des biofilms par rapport à leurs homologues planctoniques. Les résultats indiquent que les biofilms mono- et double-espèces ont présenté des valeurs améliorées pour les divers attributs biochimiques par rapport à leurs homologues planctoniques. Ceux-ci comprennent la solubilisation de phosphore ainsi que la production d'acide indole-3- acétique (IAA) et de sidérophores. En outre, des essais en serre avec des plants de tomates ont confirmé nos résultats in vitro, ce qui suggère que les inoculants de biofilm sont plus efficaces que leurs équivalents planctoniques. Les résultats, par conséquent, suggérent que l'utilisation de technologies innovantes de biofilm pourrait se révéler extrêmement avantageux comme biofertilisants et agents de biocontrôle, ce qui pourrait donc potentiellement atténuer la dépendance sur les produits agro-chimiques pour une production agricole efficace.

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ACKNOWLEDMENTS

I would like to take this opportunity to thank numerous people who assisted me throughout my Masters research. First and foremost I would like to express my sincere gratitude to my research supervisor Professor Dr. Donald Smith for his academic guidance, continuous encouragement throughout the course of my studies and for this incredible opportunity.

Additionally, I would also like to express appreciation to all of the lab members with whom I had the opportunity to work with. I would especially like to thank Dr. Sowmya Subramanian, Dr. Alfred Souleimanov, Rachel Backer, Yoko Takishita and Surashri Shinde for their constant moral support, assistance and patience.

I wish to express my sincere gratitude to the members of my thesis advisory committee consisting of my co-supervisor Professor Dr. Valerie Gravel and Professor Dr. Sebastien Faucher for their valuable expert advice and guidance throughout my Masters studies. Furthermore I would like to thank Dr. Jean-Benoit Charron, Dr. Jacqueline Bede, Dr. Alan Watson and Dr. Kushalappa for allowing me to use their laboratory equipment and strains which was absolutely necessary to complete my studies.

Last but not least, I would like to express my deepest appreciation to my family and friends for their patience and support throughout these years. You allowed me to become the person that I am today and I thank you.

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Chapter 1

GENERAL INTRODUCTION

1.1 Introduction

Currently, global population is over 7 billion and this is projected to reach 9 billion by 2050 (FAO, 2009). One of the biggest challenges to overcome is to substantially increase the amount of food produced in a sustainable and environmentally friendly manner. In fact, studies have shown that in order to meet global food demand by 2050, we will need to increase food production by at least 70% (FAO, 2009). One of the limitations to increasing worldwide crop productivity is insufficient essential nutrient in most agricultural soils, resulting in suboptimal crop growth. Many studies have demonstrated that low soil fertility is one of the major constraints to production of food grains and vegetables; this is caused by annual harvests that deplete soils of natural soil reserves (GOA, 1998). This is particularity so for nitrogen and phosphorus, which are essential nutrients for plant growth and development (Hameeda et al., 2008). Other factors that contribute to decreased crop yields worldwide are the effects of weeds, pests and diseases. In fact, these components account for the loss of 20 to 40% of the world’s potential crop productivity annually (FAO, 1985; Oerke et al., 2006; FAO, 2012). A decrease in agricultural crop production can cause a tremendous impact on both global food prices and availability.

To overcome these challenges, agricultural practices have depended on the use of chemical fertilizers, insecticides, fungicides and herbicides to increase crop yields. However, these approaches are costly and have numerous negative environmental impacts. Negative environmental impacts include; greenhouse gas emissions, such as carbon dioxide (CO2) and nitrous oxide (N2O), depletion of non-renewable resources such as oil, natural gas and phosphorus as well as poor water quality due to nitrate (NO3) and phosphorus leaching into the groundwater, (Bohlool et al., 1992; Smil, 1999; Socolow, 1999). Moreover, these effects can cause irreversible and devastating impacts to flora and fauna biodiversity in surrounding areas (Vitousek et al., 1997). Likewise, only a small fraction of the applied fertilizers is actually used directly by the plants. On

1 average it is estimated that only 15 to 20% of the total amount of phosphorus fertilizer is used by plants in the year of application (Liu et al., 2008). Furthermore the overuse of pesticides can rapidly lead to evolved resistance, which not only has negative impacts on the environment but can have deleterious effects on natural beneficial agents which aid in keeping pests in check and on human health (Ali et al, 2014). In fact, when exposed to pesticides, health hazards can include neurological disruptions, birth defects, fetal death, neurodevelopmental disorders, leukemia and Ewing’s sarcoma (Reynolds et al., 2002; Valerey et al., 2002; Sanborn et al., 2007; Jurewicz et al., 2008). In order to meet the increased food demands projected for 2050, a tremendous increase in the production, distribution and application of fertilizers will be required, which is neither economically nor environmentally desired.

Current trends in agriculture are focused on searching for alternative sustainable and environmentally friendly approaches that will improve soil quality and crop production. One of these alternatives is the use of biological agents, such as (bacteria and fungi). The focus of this thesis will be on bacteria. Compared to chemical fertilizers, the production and application of microbial inoculants is cost effective, minimally depletes non-renewable resources, improves soil quality and does not cause negative impacts on either the environment or human health. Overall, various studies have proven that the use of planktonic-PGPR (P-PGPR) offers promising prospects in in vitro, growth chamber and greenhouse trials, demonstrating the potential to diminish the use of chemicals. Unfortunately, planktonic microbial inoculants have proven inconsistent under field conditions most certainly due to the inability of the planktonic cells to compete with the indigenous microflora (Gupta et al, 2015).

To address this issue, scientists have begun to investigate the use of biofilm PGPR (B- PGPR) as alternative inoculum. Biofilms are defined as dense colonies of single or multi-species microbial cells adherent to either a biotic or abiotic surface encased in a matrix of extracellular polymeric substances (EPS) (Davey et al., 2000). Interestingly, in natural environments microbial cells are predominantly found as biofilms, as opposed to their planktonic equivalents, which may prove to be a more suitable and durable option compared to the traditional application of P-PGPR inoculums. In fact, the formation of biofilms enhances the survival of the bacterial cells, which may allow the bacterial inoculants to survive the initial application into natural soils and allow

2 them to thrive in the long term. Although relatively new, initial investigations have provided encouraging results in regards to utilizing B-PGPR. These discoveries have led our laboratory to further investigate the use and roles of biofilms as PGPR delivery systems. Therefore the goal of this research project is to develop and elucidate the potential use of mono- and dual-species biofilm inoculants. More specifically, B-PGPR biochemical traits and their effects on plant growth will be compared and contrasted to that of their P-PGPR counterparts.

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1.2 List of Objectives

The research project is focused on determining whether or not isolated plant growth promoting bacterial strains are able to form mono- and dual-species biofilms and to characterize their B-PGPR traits as potential inoculants. The general objectives of the research project are as follows;

1) Characterize the bacterial strains

2) Determine if isolated bacterial strains can form mono-species biofilms in vitro

3) Standardize colony forming units (CFU) of planktonic and biofilm mono- and dual-species cultures

4) Elucidate and compare the biochemical plant growth promotion attributes of mono- and dual-species biofilm and planktonic PGPR

5) Establish if mono- and dual-species B-PGPR are more resistant to antibiotics then their planktonic counterparts

6) Determine if mono- and dual-species B-PGPR are more effective at enhancing crop growth, under greenhouse conditions, than their planktonic equivalents

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1.3 List of Hypotheses

The overall hypothesis of the project is that B-PGPR are more effective plant growth promoters than their planktonic counterparts. The specific hypotheses are as follows;

1) Plant growth promoting bacterial strains isolated from field conditions can form mono- and dual-species biofilms

2) Biofilm forms of PGPR are more effective than their planktonic counterparts at enhancing crop growth.

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Chapter 2

GENERAL LITERATURE REVIEW

2.1 Plant microbe interactions

2.1.1 Rhizosphere

The rhizosphere, a narrow region of soil surrounding the root system which is affected by microorganisms and root secretions, plays an important role in plant-microbe interactions (Walker et al., 2003). Numerous microorganisms can be found in this region and include bacteria, fungi and protozoa. However, bacteria are by far the most abundant. In fact it is estimated that there are 108 to 109 bacterial cells per gram of rhizosphere soil (Schoenborn et al., 2004). However, in environmentally stressed soils, these numbers can be as low as 104 cells per gram of rhizosphere soil (Timmusk et al., 2011). Environmentally stressed soils can arise for numerous reasons, including drought stress, soil salinity and nutrient deprivation (Timmusk et al., 1999; Mayak et al., 2004; Yang et al., 2009). Within the rhizosphere, plants and microbes can communicate with other each other by releasing a variety of signal molecules, which ultimately leads to the appropriate physiological response(s) (Mabood et al., 2006). More specifically, compounds secreted by plants, termed root , modify the physical and chemical properties of soil thus influencing the microbial community in the immediate proximity of root surface (Dakora et al., 2002). In fact, certain root exudates can act as repellents while some function as attractants. These interactions are completely dependent on the species of plant and involved. As a specific example, certain species of rhizobia can associate with numerous plants and therefore have a broad host range, while others are more specific and therefore have a narrower host range (Perret et al., 2000). Interactions between soil bacteria and plants can be beneficial, harmful or neutral. However, the focus of this research project is to investigate beneficial interactions.

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2.1.2 Plant growth promoting rhizobacteria (PGPR)

Bacteria that are beneficial to plants are referred to as plant growth promoting rhizobacteria (PGPR). Rhizobacteria is a collective termed used to describe bacteria capable of colonizing the rhizosphere (Zablotowicz et al., 1991). PGPR can be divided into two groups based on their vicinity to the roots and intimacy of association (Gray et al., 2005). These include intracellular PGPR (iPGPR) and extracellular PGPR (ePGPR). iPGPR colonize the inside of the root cell leading to the formation of a new specialized plant organ termed the nodule, while ePGPR are capable of colonizing the rhizosphere (zone of soil around the roots), the rhizoplane (root surface) or the spaces between cells of the root cortex (Gray et al., 2005; Figueiredo et al., 2011). PGPR can be further classified according to their modes of action, which include both direct and indirect mechanisms. More specifically, PGPR can promote plant growth directly by acting as (1) biofertilizers that supply nutrients such as nitrogen, phosphorus and essential minerals directly to the plants and as (2) phytostimulators which directly stimulate the production of plant hormones such as (Zahir et al., 2003; Somers et al., 2004; Glick et al., 2007), shown in Figure 1. Moreover, PGPR can promote plant growth indirectly by functioning as (3) biocontrol agents which control or inhibit the deleterious effects of numerous pathogenic agents by releasing an array of compounds such as antibiotics and antifungal metabolites (Somers et al., 2004; Antoun et al., 2006), shown in Figure 1. However it is important to note that a single PGPR will frequently display numerous modes of action (Kloepper et al., 2003; Vessey, 2003). In return, the PGPR will receive photosynthate (reduce carbon) from the plant, which is a crucial energy source required for the bacteria to survive, as well as numerous compounds such as carbohydrates, amino acids and organic acids which serve as nutrients (Gnanamanickam, 2006). Thus, beneficial PGPR can have profound effects on both crop health (by recycling soil nutrients and providing them to plants) and soil quality (Sturz et al., 2003; Glick, 2012).

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Figure 1. Plant growth promoting rhizobacteria (PGPR) mechanisms. PGPR can function as biofertilizers by providing nitrogen, phosphate, iron and hormones to plants as well as biocontrol agents by scavenging iron in addition to producing ammonia, hydrogen cyanide and antimicrobial agents.

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2.2 Direct mechanisms

2.2.1 Nitrogen fixation

Nitrogen plays a crucial role in the synthesis of , nucleic acids and chlorophyll and is therefore a crucial element required for plant growth. However, nitrogen is the most frequently limiting essential macronutrient. Although the Earth’s atmosphere is 79% dinitrogen gas, plants cannot assimilate this form on their own, due their inability to fix nitrogen (Burris et al., 1993; Rees et al., 2005; Robertson et al., 2009). In fact, nitrogen fixation can only be accomplished by prokaryotic microorganisms (Rubio et al., 2008). Therefore plants must rely on nitrogen fixing bacteria, also termed diazotrophs, to convert atmospheric dinitrogen (N2) to ammonia (NH3) (Kennedy et al., 1992; Robertson et al., 2009).

Diazotrophs are able to fix nitrogen through the presence of nif , which code for , an enzyme capable of catalyzing the N-fixing reaction (Kim et al., 1994). Diazotrophs have been shown to have a profound effect on a variety of important crops. In a study conducted by Polonenka et al. (1987), treatment of soybean with either a japonicum and or a B. japonicum and P. fluorescens inocula resulted in an increased number of nodules and nodule dry weight (compared to the B. japonicum alone control) under growth chamber conditions. Moreover, in a study conducted by Bai et al. (2003) soybean (Glycine max) plants treated with either a B. japonicum and Bacillus thuringeinsis or a B. japonicum and B subtilis inocula led to enhanced growth in greenhouse and field conditions. In fact, results show an increase in number of nodules, nodule weight, shoot weight, root weight, total biomass, total nitrogen and grain yield (Bai et al., 2003). Furthermore symbiotic Rhizobia can be beneficial for non- plants, such as cereals, by directly transferring fixed nitrogen to non-legume crops growing in intercrop systems with N2-fixing , or when non-legumes are rotated with symbiotic legumes (Eaglesham et al., 1981; Dakora et al., 1997).

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2.2.2 Phosphate solubilisation

Phosphorus, another essential nutrient required by plants, plays a key role in the synthesis of nucleic acids as well as the storage and distribution of energy for plants. Phosphorus is also one of the major limiting essential macronutrients. Although the amount of phosphorus (both organic and inorganic) found in soils is quite high (between 400 and 1,200 mg kg -1) the majority is insoluble and therefore unavailable to support plant growth (Glick, 2012). In fact, plants must rely on phosphate solubilising microorganisms such as bacteria and mycorrhizal fungi located in the rhizosphere to solubilise both organic and inorganic phosphate into dihydrogen phosphate (H2PO4) or hydrogen phosphate (HPO4) ions (Bhattacharyya et al., 2012). More specifically, the solubilisation of inorganic phosphorus is achieved when microbes synthesize low molecular weight organic acids such as gluconic, oxalic, citric and ketoglucanic acid (Kucey et al., 1983; Rodriguez et al., 1999; Bolan et al., 1994; Zaidi et al., 2009). On the other hand, the solubilisation of organic phosphorus is achieved through the synthesis of enzymes such as phosphatases, phytase and phosphonoacetate hydrolase, which induce the hydrolysis of phosphoric esters (Rodriguez et al., 1999; Rodriguez et al., 2006). While phosphate solubilising bacteria represent 1-50% of the total population of microbes in the soil, phosphate solubilising fungi represent only 0.1-0.5% soil fungi; these fungi have been shown to exhibit greater solubilising activity due to their ability to transverse long distances more effortlessly than bacteria and due their capacity to produce a greater amount of acids required to solubilise phosphorus (Kucey et al., 1983; Venkateswarlu et al., 1984). It is important to note that certain bacterial strains can exhibit both organic and inorganic phosphate solubilisation characteristics (Sharma et al., 2011). Some of the best-known phosphates solubilising bacteria include the genera Rhizobium, Azobactor, Erwinia, Serratia, Burkholderia, Pseudomonas and Bacillus (Sturz et al., 2000; Sudhakar et al., 2000; Mehnaz et al., 2006).

Although the effects of phosphate solubilising PGPR on plant growth have shown promising results, it is important to note that the majority of phosphate solubilising bacteria do not solely use phosphate solubilisation as their mechanism of plant growth enhancement. In fact, most phosphate solubilising bacteria utilize an array of mechanisms (nitrogen fixing, production of hormones and/or production of pathogen antagonists as well), making it difficult to interpret the effects of phosphate solubilisation alone on plant growth. In a study conducted by Vyas et al.

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(2009) germinated seeds treated with Pseudomonas trivialis or a P. poae inocula showed enhanced corn growth under growth chamber conditions, causing significant increases in shoot weight, root weight and nitrogen and phosphorus contents compared to the control (Vyas et al., 2009). Possible mechanisms of action responsible for these increases are thought to include phosphate solubilisation and the production of phytohormones (Vyas et al., 2009). Furthermore, in a study conducted by Yasmin et al. (2011), corn seeds treated with rhizobacteria inoculum under greenhouse conditions showed increases in shoot (43%) and root length (41%) compared to the control through mechanisms involving phosphate solubilisation as well as the production of bacteriocins and siderophores.

2.2.3 Siderophore production

Iron, another vital and sometimes limiting nutrient in agricultural soils, is required by both plants and microorganisms for both cellular growth and . In aerobic environments, iron predominantly exists as ferric chloride (Fe3+), which tends to form insoluble hydroxides and oxyhydroxides making it inaccessible to both plants and microorganisms (Rajkumar et al., 2010). In order to overcome this, plants often rely on bacteria to solubilise iron. In fact, bacteria secrete low-molecular weight iron chelators, termed siderophores, which have high association constants for complexing iron thereby functioning as an iron solubilising agent (Neilands, 1995; Miethke et al., 2007). More specifically bacterial isolates can produce various types of siderophores, which are classified according to the ligands used to chelate the ferric iron such as catecholates, hydroxamates and carboxylates (Miethke et al., 2007). Plants can then assimilate the iron they require from bacterial siderophores by a number of mechanisms, including the direct uptake of siderophore-iron complexes, or by a ligand exchange reaction, or by means of chelating and releasing iron (Schmidt et al., 1999).

Siderophore secreting bacteria include the genera Pseudomonas, Bradyrhizobium, Bacillus and Enterobactor and have been shown to enhance plant growth (Ahemad et al., 2014). For instance, in a study conducted by Vansuyt et al. (2007), the application of a Pseudomonas fluorescens isolate showed improved iron acquisition in plant tissue and improved plant growth of Arabidopsis thaliana. Moreover, maize seeds treated with a Pseudomonas strain showed higher

11 root and shoot length in addition to less severe chorotic patches in comparison to the uninoculated control (Sharma et al., 2003). Additionally, when a Bacillus inoculant was applied to cassava plants, there was an increase in height, branching and shoot biomass in comparison to the control (Freitas et al., 2015). More specifically, results showed enhanced levels of iron in the aerial portions of the plant (Freitas et al., 2015). In yet another study, Arabidopsis thaliana seedlings exposed to a Bacillus inoculum showed higher whole-plant iron content in comparison to the water treated control (Zhang et al., 2009). Also, results showed more abundant lateral roots, higher chlorophyll content and greater photosynthetic efficiency (Zhang et al., 2009).

2.2.4 Phytostimulators

In addition to supplying nutrients directly to the plant, PGPR can synthesize and secrete plant hormones, termed phytohormones, which are chemical signals that play a crucial role in regulating almost every aspect of plant growth and development, including defense responses against both biotic and abiotic stresses. Phytohormones, or plant growth regulators (PGR) include auxins such as indole-3-acetic acid (IAA). Interestingly, it is estimated that 80% of microorganisms isolated from the rhizosphere possess the ability to synthesize and release auxins as secondary metabolites (Patten et al., 1996). Interestingly, IAA has been shown to enhance the production of root hairs and lateral roots, which are directly involved in nutrient uptake (Patten et al., 2002). Although certain microbial isolates can produce IAA independently of tryptophan, the majority of rhizobacteria require tryptophan, an amino acid found in root exudates, which functions as a precursor for IAA biosynthesis (Patten at al., 1996). Furthermore several IAA biosynthesis pathways have been elucidated for PGPR.

Microbes proficient in IAA production include strains that belong to the genera Rhizobium, Pseudomonas and Bacillus (Dey et al., 2004; Zaidi et al., 2006; Poonguzhali et al., 2008). In fact, several studies have shown that PGPR can directly enhance plant growth through this mechanism. More specifically, in a study conducted by Egamberdiyeva et al. (2002) the application of IAA producing Pseudomonas spp to corn plants enhanced growth under controlled environment chamber conditions. Results indicated significant increases in both shoot and root length (Egamberdiyeva et al., 2002). In yet another report, the application of a Pseudomonas inoculant to

12 canola seedlings led to 35 to 50% increases in root length in comparison to uninoculated seeds and to mutants deficient in IAA production (Patten et al., 2002). Furthermore when an IAA producing Bacillus inoculant was applied to wheat seeds there was a significant increase in plant root and shoot elongation compared to the control (Mohite, 2013).

2.3 Indirect mechanisms

2.3.1 Antimicrobial activity

Biocontrol of phytopathogens is yet another mechanism employed by PGPR to indirectly enhance plant growth. PGPR can employ an array of mechanisms, which include general competition for nutrients, niche exclusion, induced systemic resistance and the production of antimicrobial metabolites, all of which suppress pathogen development (Kloepper et al., 1999; Bloemberg et al., 2001; Lugtenberg et al., 2009). Antimicrobial compounds released by PGPR include agrocin, herbicolin, surfactins, hydrogen cyanide, ammonia, phenazines, pyrrolnitrin, pyoluteorin, 2,4-diacetylphloroglucinol, viscosinamide, tensin as well as thuricin 17, a relatively new class of antibiotics termed bacteriocins (Howell et al., 1988; Voisard et al., 1989; Picard et al., 2000; Ennahar et al., 2000; Bhattacharyya et al 2012). Competition for nutrients and niche exclusion involves the production of siderophores, which renders iron unavailable to other microorganisms (Haas et al., 2005). Some of the best-known PGPR biocontrol agents include bacteria from the genera Bacillus and Pseudomonas (Raaijmakers et al., 2010).

Several studies have shown promising results regarding use of PGPR as biocontrol agents to promote plant growth. For instance, in a study conducted by Cavaglieri et al. (2005) application of a Bacillus subtilis inoculum to maize seeds protected the crop against the phytopathogen Fusarium vertililliodes, responsible for seedling decay, stalk rot and ear rot. More specifically, results indicate that the B. subtilis inoculum significantly inhibited pathogen growth under both in vitro and greenhouse conditions (Cavaglier et al., 2005). In a study conducted by Khanuchiya et al. (2012) both Pseudomonas fluorescens and B. subtilis were able to inhibit the phytopathogens Fusarium oxysporum and Aspergillus niger under both in vitro and growth chamber conditions. In

13 fact, results indicate that castor seeds treated with either of these inoculants showed significant root length enhancement compared to the control (Khanuchiya et al., 2012).

2.4 Issues with planktonic inoculants

The use of planktonic PGPR inoculants has shown promising results in regards to promoting plant growth, both directly and indirectly under in vitro, growth chamber and greenhouse conditions, alleviating dependence on chemical fertilizers. Unfortunately, planktonic microbial inocula have proven inconsistent in field applications, therefore limiting their widespread commercial application. This is almost certainly caused by the inability of the microbial inocula to compete with the indigenous microflora and establishing itself in the rhizosphere (Gupta et al, 2015). To overcome this issue, scientists have begun to investigate the use of biofilms as potential inoculants.

2.5 Biofilms

2.5.1 Biofilm formation

Bacterial cells can exhibit two general growth modes: either as free-swimming planktonic cells or as surface attached communities termed biofilms. Biofilms are defined as dense colonies of single or multi-species of microbial cells adherent to either a biotic or abiotic surface encased in a self-produced matrix composed of extracellular polymeric substances (EPS) (Davey et al., 2000). Biofilms in the environment date to at least 3.25 million years ago, as indicated by the fossil record, and occur across a wide range of organisms both in the Archaea and Bacteria lineages (Rasmussen et al., 2000; Westall et al., 2001). Biofilms can form in virtually any environment so long as a surface is sufficient in both nutrients and water. In most clinical, industrial and natural settings, including the rhizosphere, bacteria are predominantly found as biofilms, as opposed to their planktonic state (Campbell et al., 1990; Costerton et al., 1995; Davey et al., 2000). More specifically, plant associated biofilms have been shown to establish themselves on various parts of plants such as leaves, roots, seeds and internal vasculature (Ramey et al., 2004; Danhorn et al., 2007; Eberl et al., 2007). In fact, various Pseudomonas isolates have been shown to form biofilms

14 on leaves, roots and in the soil (Ude et al., 2006). The ability to form biofilms not only enhances bacterial survival but also enhances plant growth through the various mechanisms described above.

Numerous stages are required for biofilm formation, as illustrated in Figure 2. In response to environmental cues such as nutrient availability, free-swimming planktonic cells move towards an appropriate surface (through ) and initially attach to a surface via their fimbria and/or pili through weak interactions such as van der Waal forces or hydrogen bonding, making the attachment reversible (Allison et al., 2000). Over time, the attachment becomes essentially irreversible, as a result of an accumulation of weak interactions and changes in expression, which is when the bacterial cells begin to secrete EPS (De Weger et al., 1987; O’Toole et al., 2000; Hoiby et al., 2001). When enough bacterial cells have aggregated in this growing “pre-biofilm” due to bacterial cell division and other microbes joining in, microcolonies develop which ultimately leads to the formation of macrocolonies or a “mature biofilms” (O’Toole et al., 2000). Finally when the biofilm is large enough, bacterial cells begin to detach from the biofilm, with these bacterial cells either resuming their planktonic growth mode or establishing their own biofilms.

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1. Initial Attachment 5. Dispersal 4. Macrocolony Formation 3. Microcolony Formation

2. Irreversible attachment

Figure 2. The process of biofilm development. Stages of biofilm development: 1. Initial reversible attachment, 2. Irreversible attachment, 3. Microcolony formation, 4. Macrocolony formation, and 5. Dispersal (O’Toole et al., 2000).

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2.5.2 Biofilm properties

Biofilms exhibit numerous properties that are different from their planktonic counterparts. This includes the fact that biofilms are encased in a self-produced EPS matrix composed of polysaccharides, proteins, deoxyribonucleic acid (DNA) and ribonucleic acid (RNA) (Allison et al., 2000). The EPS compromises 70-80% of the biofilm mass while the bacteria only constitute 10-20% of the mass (Allison et al., 2000; Hoiby et al., 2001). However, the exact composition of the EPS depends entirely on the microbes involved. The EPS plays a crucial role in both the attachment process and the overall structure of the biofilm. Furthermore, the EPS gives bacterial cells protection from sudden environmental changes. In fact, biofilms have an increased tolerance to a wide range of potentially stressful physical conditions including temperature, extreme pH, nutrient and waste levels, salts, antiseptics, commonly used antibiotics and desiccation (Baty et al., 2000; Todar, 2008). Furthermore, the EPS not only enhances bacterial survival but improves soil aggregation, which ultimately improves the stability of soil water supply, which generally improves plant survival and growth (Davey et al., 2000).

Another distinct feature of biofilms is that they are a structurally, chemically and biologically heterogeneous mixture, both spatially and temporally, meaning that individual bacterial cells express different genes over time and depending on their location within the biofilm (Stewart et al., 2008). Hence, at any stage of development, bacterial cells located near the edge of the biofilm express different genes than cells at the core of the biofilm. Furthermore, cells express different gene sets within different stages of biofilm development. This is caused by the heterogeneity in metabolic activities and diffusion processes which leads to concentration gradients of nutrients, signaling compounds and bacterial waste inside the biofilm, thus allowing the bacterial cells to respond to environmental changes quickly (Stewart et al., 2008).

Finally, cells in biofilms are much more resistant to antibiotics than their planktonic counterparts, improving their chance of survival in a competitive environment. More specifically, when free-swimming planktonic bacteria form biofilms, they can become up to 1,000 times more resistant to antibiotics (Mah et al., 2001). Numerous biofilm-specific antibiotic resistance mechanisms have been identified and include physical or chemical barriers provided by the EPS,

17 slow growth, stress response, heterogeneity and biofilm-specific phenotypes (Mah et al., 2001). However it is important to note that multiple mechanisms are required for overall antimicrobial resistance (Mah et al., 2001). In fact, studies have shown that the EPS prevents diffusion of antimicrobial agents through the biofilm, hence preventing the agent from reaching its site of action, (Nichols et al., 1988; Anderl et al., 2000; Donlan et al., 2002; Walters et al., 2003; Zhang et al., 2008). Although, this is only the case for certain antimicrobial agents. In addition, it has been suggested that a lower growth rate (seen in biofilms in response to oxygen, nutrition deprivation and environmental stress) can explain some level of biofilm-specific resistance, simply because many antibiotics are only active against fast growing cells (Costertan et al., 1999). Finally, the gene expression profile of biofilms is different than that of planktonic cells. In fact, a number of biofilm specific antibiotic genes are upregulated under biofilm conditions, and responsible for an increase in antibiotic resistance, compared to planktonic cells (Whiteley et al., 2001; Resch et al., 2005; Beaudoin et al., 2012).

2.5.3 Biofilms as biofertilizers and phytostimulators

Biofilms, like their planktonic counterparts, can directly enhance plant growth in a number of ways including supplying nutrients to the plant and stimulating the production of phytohormones, as previously discussed. In a study conducted by Triveni et al. (2012) a Bacillus subtilis and Trichoderma viride as well as Pseudomonas fluorescens and Trichoderma viride dual- species biofilm showed greater ammonia production, IAA production and phosphate solubilisation than the planktonic inoculum. In addition, results indicate that biofilms have a more pronounced effect on cotton germination than planktonic cultures (Triveni et al., 2012). Moreover, in a study conducted by Mohd (2014) a P. fluorescens biofilm was able to enhance plant growth in wheat (Triticum aestivum) by increasing IAA production, phosphate solubilisation and siderophore production. Additionally, wheat seeds treated with the biofilm inoculant showed increases in root and shoot length (Mohd, 2014). In yet another study, Jayasinghearacchi et al. (2004) found that a dual-species biofilm containing Bradyrhizobium sp and a Penicillium sp showed increased nitrogenase activity compared to their individual planktonic counterparts, which had gone undetected due to values being below the detection limit. Additionally, the dry weights of the shoots, roots, nodules and nitrogen accumulation in the shoots and roots of soybean

18 inoculated with the dual-species biofilm were greater than those inoculated with the cultures of just one planktonic species (Jayasinghearacchi et al., 2004). Furthermore, in a study conducted by Buddhika et al. (2014) maize treated wither either an Aspergillus spp and Azorhizobium spp or Aspergillus spp and Acetobactor spp dual-species biofilm resulted in increased seed germination and longer roots than their individual planktonic monocultures.

2.5.4 Biofilms as biocontrol agents

An alternative mechanism used by biofilms to enhance plant growth is through biocontrol of disease organisms, such as competitive colonization of the rhizosphere and the production of antimicrobial compounds, as previously discussed for planktonic PGPR. This impedes pathogenic microbes from establishing themselves, preventing disease initiation (Lugtenberg et al., 2009). In a study by Bais et al. (2004) a B. subtilis biofilm was used to protect Arabidposis roots from the phytopathogen Pseudomonas syringae, which causes cankers. Results show that the biofilm reduced plant mortality from 85 to 10% both in vitro and in sterile soil conditions through the secretion of surfactin, a lipopeptide antimicrobial agent (Bai et al., 2004). In fact, there was a 2- fold increase in surfactin production when P. syringae was administered to the Aradidopsis-B. subtilis biofilm system (Bai et al., 2004). In yet another study, Chen et al. (2013) used B. subtilis biofilms to protect tomato (Solanum lycopersicum) plants against wilt disease caused by the plant pathogen Ralstonia solanecearum under greenhouse conditions. In fact, seven of the biofilm isolates retrieved from various locations in China were able to achieve more than 50% biocontrol activity through the production of surfactins, which act as an antimicrobial agent and a signalling compound that stimulates biofilm formation (Chen et al., 2013). Moreover, the biofilms showed promising in vitro antagonistic activities against numerous other plant pathogens, including Pythium ultimum, Fusarium graminearum, Phytophthora capsici, Fusarium oxysporum, Scleotinia sclerotiorum and Ralstonia solanacearum (Chen et al., 2013). Finally, in a study conducted by Wei et al. (2006) a P. fluorescens biofilm was shown to successfully protect wheat roots from the phytopathogens Rhizoctonia solani and Geaumannomyces graninis responsible for damping-off disease, by secreting antimicrobial compounds such as 2,4-diacetylphloroglucinol, hydrogen cyanide and siderophores.

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Chapter 3

MATERIALS AND METHODS

3.1 Bacterial Strains

3.1.1 Bacterial strain isolation and screening

Bacterial strains were isolated from roots and rhizospheric soils of field conditions from a field site in Ste-Anne-de-Bellevue, Québec, Canada (42°28°N 73°45°W) by Di Fan (PhD student in Dr. Smith’s laboratory). Bacterial isolates were purified by three successive rounds of single- colony streaking on Petri plates where single colonies were sub-cultured and frozen at -80°C in their respective medium containing 25% glycerol. Isolated strains were then subjected to a bacterial screening bioassay in order to confirm plant growth promoting abilities by testing their effects on Arabidopsis thaliana (Fan et al., personal communication).

3.1.2 16s rRNA sequencing and identification

The identification of two PGPR strains was determined by conducting16s rRNA sequencing. Genomic DNA was extracted and purified from the rhizobacteria via the DNeasy Blood & Tissue kit according to the manufactures manual (Qiagen, Canada). The purity of the genomic DNA was assessed by the A260/A280 and A260/A230 extinction ratio using a NanoDrop (Thermo Scientific, U.S.A.). The entire 16S rRNA region (1465 bp) was amplified by polymerase chain reaction (PCR) by using the forward and reverse primers S-D-Bact-0008-c-S-20 (5’- AGRGTTYGATYMTGGCTCAG-3’) and S-D-Bact-1046-a-A-19 (5’- CGACRRCCATGCANCACCT-3’), respectively. The entire PCR reaction (50 µL) included 1 µL of genomic DNA, 1 µL of each primer (10 µM), 1 µL of each deoxynucleotide tripshosphate

(dNTP) (10 µM), 5 µL 10xPCR buffer, 3 µL of 25 mM MgCl2, 1 µL Taq DNA polymerase (5 units) and dH20 (Bioshop, Canada). The reactions were run in a PT-100 Programmable Thermal

Controller (MJ Research Inc, U.S.A.) as follows; 94°C for 3 minutes (min) for initial denaturing,

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40 cycles at 94°C for 30 seconds (sec) for complete denaturing, 55°C for 30 sec for primer annealing, 72°C for 1 min for primer extension and 72°C for 10 min for complete elongation. The PCR products (5 µL) were analyzed by electrophoresis (Bio-Rad, Canada) on a 1% (w/v) agarose gel with 1X Tris acetate EDTA (TAE) buffer, stained with ethidium bromide (0.5 mg L-1) and visualized using an ultraviolet (UV) transilluminator (Bio-Rad, Canada). Bands were then purified using a QIAquick PCR purification kit (Quiagen, Canada), following the instructions of the manufacturer’s manual. Bands were then sent out for DNA sequencing, which was performed by the Genome Quebec Innovation Center (McGill University) on a 3730XL DNA analyzer system (applied biosystems). The 16S rRNA sequence retrieved was then analyzed by nucleotide basic alignment search tool (BLASTN) and compared against the GenBank database.

3.1.3 Optimal growth conditions of mono-species planktonic cultures

The optimal growth conditions (temperature and media) for mono-species planktonic bacteria were determined by assessing growth curves. Overnight bacterial cultures (1 mL) were diluted into 100 mL of either fresh Luria-Bertani broth (LB; 10 g tryptone, 5 g yeast extract and 5 -1 g NaCl L ), King´s B broth (KB; 20 g protease peptone #3, 1.5 g K2HPO4, 10 mL glycerol -1 supplemented with 0.006 M MgSO4) or Tryptic soy broth (TSB; 30 g L ) each supplemented or not with 0.5% glucose. The optical density (OD, at 600 nanometers) was taken immediately after inoculation (t = 0). Bacterial cultures were grown at either 30 °C or 37 °C in an Ecotron incubator (Infors HT, Switzerland) with shaking set at 150 revolutions per minute (rpm). Every hour, a 1 mL aliquot was removed and the OD600 was recorder with a Ultrospec 4300 Pro UV/Visible spectrophotometer (Fisher Scientific, Canada) until the stationary phase was reached, meaning the OD was constant for at least three time points. This experiment was conducted three times with three replicates for each treatment. The average OD at each time point was calculated and plotted.

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3.1.4 Standardization of mono and dual – species planktonic cultures

To quantify mono-species planktonic preparations of the bacterial strains, serial dilution plating was conducted. Overnight stationary phase bacterial cultures (1 mL) were diluted into 100 mL of TSB supplemented with 0.5% glucose, at 30°C with shaking set at 150 RPM. The quantity of each bacterial strain was determined by re-suspending 100 µL of bacterial cultures into 900 µL TSB supplemented with 0.5% glucose and streaking 100 µL of bacterial culture onto KB agar plates. Serial dilution plating was conducted when bacterial strains reached the mid-exponential (OD 0.5) and early stationary phase (OD 1) of growth as well as at 24 h of growth, in order to establish the number of viable colony forming units (CFUs). The optimal growth conditions of dual-species planktonic bacteria were determined with slight modifications to the method previously described. Bacterial cultures were tested at various ratios (Bacillus sp. to Pseudomonas sp. in mL; 1:1, 0.5:10, 0.25:15, 0.1 to 20 and 0.05 to 30) following 24 h of growth. After 24 h serial dilution plating was conducted on KB agar plates as previously described. Plating on KB agar determined if both bacterial species co-existed (based on morphology of colonies on the agar plate) and quantified the number of CFUs for each bacterial strain. This experiment was conducted three times with three replicates for each treatment and the average CFU at each time point was calculated.

3.1.5 Optimal growth conditions of mono-species biofilm cultures

To determine if isolated bacterial strains can form mono-species biofilms, crystal violet (CV) assays were conducted as described by Merrit et al. (2005). Overnight stationary phase bacterial cultures were inoculated into a 96-well microtiter plate (100 µL per well) (Fisher Scientific, Canada) at 1:50 and 1:100 dilutions in LB, KB, TSB, TSB supplemented with 0.5% -1 glucose, Murashige and Skoog (MS; 4.4 g L ), Bushnell Haas (BH; 1 g K2HPO4, 1 g KH2PO4, 1 -1 g NH4NO3, 0.083 g FeCl3, 0.02 g CaCl2·6H2O and 0.2 g MgSO4·7H2O L ) supplemented with

0.2% glucose and 0.5% tryptone in addition to both monobasic minimal media (M63; 10 g K2HPO4 -1 and 2 g (NH4)2SO4 L ) and dibasic minimal media (M63; 3 g KH2PO4, 7 g K2HPO4 and 2 g -1 (NH4)2SO4 L ) supplemented with 1 mM MgSO4 and 0.4% arginine. Plates were incubated at 30

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°C under static conditions for 5 days and supplied with their respective fresh media every 24 h. Plates were removed from the incubator every 24 h, the contents of the plates were dumped into a waste container and then thoroughly rinsed with distilled water (dH2O) to remove planktonic cells. To determine if biofilms formed after 5 days, 125 µL of either 0.1% or 1% crystal violet (Fisher Scientific, Canada) was added to each well and incubated at room temperature (RT) for 1 h. The dye was then rinsed off 4 times with dH2O and allowed to air dry. To quantify biofilm formation, 200 µL of a 95% ethanol solution was added to each well and incubated at RT for 30 min to solubilise the dye. One hundred and twenty five microlitres of each well was then transferred to a new microtiter plate and the absorbance was recorded with the aid of an Infinite M200 Pro

Microplate Reader (Tecan, Switzerland) set at OD550 nm. This experiment was conducted three times with three replicates for each treatment. The average OD was measured every 24 h for 5 days.

3.1.6 Standardization of mono and dual – species biofilm cultures

To determine the number of CFU’s within the biofilm, biofilm cells were prepared in a 6 well microtiter plate (Fisher Scientific, Canada). Briefly, 3.5 mL of overnight stationary phase bacterial cultures were inoculated at a 1:100 dilution in TSB supplemented with 0.5% glucose. Plates were incubated at 30 °C under static conditions for 5 days and supplied with fresh media every 24 h. Plates were removed from the incubator every 24 h and the spent media was removed from each well, followed by the addition of 3.5 mL of fresh media. To harvest biofilm biomass, the bottom and sides of each well was vigorously scraped with a plastic pipette tip, transferred into a 1.5 mL Eppendorf tube and spun down at 13,000 RPM for 1 min. The supernatant was discarded and the wells were scraped again. The cells were added to the same tubes and spun down at 13,000 RPM. The procedure was repeated once again to ensure that all biofilm cells were collected.

Biofilm pellets were resuspended in 10mM MgSO4 and subject to serial dilution plating on KB agar as previously described. To determine if isolated bacterial strains can form dual-species biofilms serial dilution plating was conducted with slight modifications to the procedures previously described. Overnight stationary phase bacterial cultures were inoculated into a 6 well microtiter plate (Falcon) at a dilution of 1:100 for each bacteria in TSB supplemented with 0.5% glucose. Plates were removed from the incubator every 24 h and biofilms were harvested and

23 subject to serial dilution plating on KB agar plates, as previously described, in order to determine if both bacterial species can co-exist (based on morphology of colonies on the agar plate) and to quantify the CFU of both bacterial strains. This experiment was conducted three times with three replicates for each treatment and the average CFU at each time point was calculated.

3.1.7 Stereo microscopy imaging

To establish if biofilms could be distinguished from their planktonic counterparts in subsequent plant growth experiments, a F10318 FilmTracer™ SYPRO® Ruby biofilm matrix stain (Thermo Scientific, U.S.A.) was tested which specifically targets matrices of biofilms. Briefly, 100 µL of overnight stationary phase bacterial cultures were inoculated into a 96-well microtiter plate with sterilized tabs at a 1:100 dilution in TSB supplemented with 0.5% glucose. Plates were incubated for 24 h as previously described in section 3.1.5. To determine if biofilms formed, 200 µL of the SYPRO® stain was added to each well and incubated at RT for 30 min in the dark. Each tab was then removed with sterilized forceps and rinsed twice with 500 µL dH2O in order to remove excess stain. Tabs were visualized one at a time by placing the tab on a sterile slide with 8 µL dH2O to prevent immediate drying. Planktonic cells were utilized as the controls since these cultures do not possess a matrix and will therefore remain unstained. Planktonic bacteria were prepared by inoculating an overnight stationary phase culture at a 1:100 dilution in TSB supplemented with 0.5% glucose. Flasks were incubated as previously described in section 3.1.4. Pseudomonas sp. was incubated for 24 h whereas Bacillus sp. was incubated to the mid exponential stage in order to have a standard number of CFU between the planktonic and biofilm cultures. To visualize planktonic cells, 1 mL of bacterial culture was mixed with 1mL of the SYPRO® stain and incubated as previously described. 8 µL of this solution was then placed on a sterile slide and visualized immediately. Stereo Microscopy (SM) images of each tab were acquired using a SteREO Discovery V20 microscope (Zeiss, Germany) equipped with fluorescence illumination. For the FilmTracer SYPRO® Ruby biofilm matrix stain, the fluorophores were excited at wavelengths of 450/610 nm. This experiment was conducted three times with three replicates for each treatment.

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3.2 Phenotypic characterization of planktonic and biofilms cultures.

For all phenotype characterization experiments, mono-species biofilms and planktonic inoculums were prepared as previously described in section 3.1.7. Dual-species biofilm were grown at a 1:100 dilution for each bacteria for 48 h under static conditions at 30°C whereas the planktonic inoculants were grown for 24 h at a 0.05 mL: 30 mL ratio of Bacillus sp. to Pseudomonas sp. and incubated as previously described in section 3.1.4 in order to have a standard number of CFU between the planktonic and biofilm cultures.

3.2.1 Nitrogen fixing ability

The ability of planktonic and biofilm cultures to fix nitrogen was determined by the method of Ker (2011). Once appropriate CFU were reached, 10 µL bacterial cultures were spotted on N- free solid LG medium (10 g sucrose, 0.5 g K2HPO4, 0.2 g MgSO4·7H2O, 0.2 g NaCl, 0.001 g -1 MnSO4·H2O, 0.001 g FeSO4, 0.001 g Na2MoO4·2H2O, 5 g CaCO3 and 15 g Bacto-agar L ) and incubated at 30 °C until colonies were apparent. This experiment was conducted three times with three replicates for each treatment. The non-inoculated media served as the negative control.

3.2.2 Phosphate solubilisation activity

Phosphate solubilisation by planktonic and biofilm cultures was assessed by the method developed by Johri et al. (1999) and Nautiyal (1999). Bacterial cultures (10 µL) were spotted on

Pikovskaya (PVK) agar plates (10 g glucose, 5 g Ca3(PO4)2, 0.5 g (NH4)2SO4, 0.2 g NaCl, 0.1 g

MgSO4·7H2O, 0.2 g KCL, 0.5 g yeast extract, 0.002 g MnSO4·H2O, 0.002 g FeSO4·7H2O and 15 g Bacto-agar L-1) and National Botanical Research Institute’s phosphate (NBRIP) agar plates (10 g glucose, 5 g Ca3(PO4)2, 5 g MgCl2·6H2O, 0.25 g MgSO4·7H2O, 0.2 g KCl, 0.1g (NH4)2SO4 and 15 g Bacto-agar L-1) until a halo surrounding the colony was apparent. Halo width was calculated after 9 days by subtracting colony diameter from the total diameter. The method described by Fiske et al (1925) was applied to quantify phosphate solubilisation (µg mL-1) activity. Bacterial cultures (0.1 mL) were inoculated into 10 mL NBRIP broth supplemented with 1000 µg mL-1 of tri-calcium

25 phosphate (TCP) and incubated at 30 °C for 11 days at 180 RPM. Bacterial cultures were then centrifuged at 10,000 RPM for 20 min at room temperature. Barton’s reagent (2.5 mL) was then added and the volume was made up to 50 mL. After 10 minutes the intensity of yellow colour was determined spectrophotometrically at OD430 nm and compared to a standard curve generated from potassium dihydrogen orthophosphate (KH2PO4) with a concentration range of 0 to 900 µg/ml (Fiske et al, 1925). This experiment was conducted three times with three replicates for each treatment. The non-inoculated media served as the negative control.

3.2.3 Inodole-3-acetic acid production

Indole acetic acid (IAA) production by biofilms was analyzed and compared to planktonic cultures by the method of Hartmann et al. (1983). Planktonic and biofilm cultures were prepared to their appropriate CFU with the addition of sterilized tryptophan (200 µg mL-1) to the growth medium. Planktonic and biofilm supernatant (0.5 mL) was transferred to a sterile falcon tube, thoroughly mixed with 1 mL of Salkowski reagent (98 mL 35% perchloric acid and 2 mL 0.5 M

FeCl3) and incubated at room temperature for 45 min in the dark to allow colour development (from yellow to pink). The quantity of IAA produced (µg mL-1) was measured spectrophotometrically at OD530 nm and compared to a standard curve generated from commercial Indole-3-acetic acid with a concentration range of 0 to 10 µg/ml (Sigma-Aldrich, Canada). This experiment was conducted three times with three replicates for each treatment. The non-inoculated media served as the negative control.

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3.2.4 Siderophore production

Siderophore production by planktonic and biofilm cultures was detected by the chrome azurol assay (CAS) as described by Schwyn et al. (1987). CAS agar plates were prepared in 3 steps: (1) CAS indicator; (2) Basal agar medium and (3) CAS agar plates. (1) 100 mL of CAS indicator was prepared as follows; 0.06 g chrome azurol S (CAS) in 50 mL dH2O, 0.0027 g FeCl3-

6H2O in 10 mL of 10 mM HCl and 0.073 g hexadecyltrimetyl ammonium bromide (HDTMA) in

40 mL dH2O autoclaved. (2) Basal agar medium was prepared as follows; 100 mL MM9 salt solution stock (15 g KH2PO4, 25 g NaCl and 50 g NH4Cl in 500 mL dH2O) to 750 mL dH2O, 32.24 g piperzine-1,4-bis (2- ethanesulfonic acid) (PIPES) supplemented with 15 g agar adjusted to pH 6.5 with NaOH and autoclaved. (3) CAS agar plates were prepared as follows; 30 mL of sterile Casamino acid, 10 mL 20% glucose and 100 mL of the CAS indicator were added to the basal agar medium and mixed. Bacterial cultures (10 µL) were spotted on CAS agar plates and incubated in the dark at 30°C for 9 days until the development of a halo was apparent. Positive siderophore production resulted in a halo with a colour development from blue to purple-orange. In addition a modified overlay CAS (O-CAS) agar assay (60.5 mg CAS, 72.9 mg HDTMA, 30.24 g PIPES,

1mM FeCl3·6H20 in 10 mL of a 10 mM HCl solution supplemented with 0.9% agarose adjusted to pH 6.5 autoclaved and cooled) was performed which entails overlaying the CAS solution onto bacterial colonies grown on KB agar plates (Perez-Miranda et al., 2007). Plates were incubated in the dark at 30°C for 2 days until the development of a halo was apparent. To quantify siderophore production (g L-1), 0.5 mL of bacterial supernatant was mixed with 0.5 mL CAS solution (same as the O-CAS solution previously described) supplemented with 10 µL shuttling solution (0.2 M sulfosalicylic acid), incubated for 30 min at room temperature and quantified spectrophotometrically at OD630 nm. Moreover both a blank (TSB supplemented with 0.5% glucose) and a reference solution (CAS solution with TSB supplemented with 0.5% glucose and the shuttling solution) were used during the quantitative determination. Percent siderophore production was calculated as follows:

27 where Ar represents the reference solution (CAS solution + shuttling solution + non-inoculated media) and As represents the sample solution (CAS solution + shuttling solution + bacterial supernatant) (Schwyn et al., 1987). This experiment was conducted three times with three replicates for each treatment. The non-inoculated media served as the negative control.

3.2.5 Ammonia production

Ammonia production by planktonic and biofilm cultures was assessed by the method described by Dye et al. (1962). Bacterial cultures (0.5 mL) were added to 10 mL of peptone water and incubated at 30°C for 4 days. Nessler´s reagent (1 mL) was then added to each suspension and colour development was noted (yellow to brown indicates positive results).

3.2.6 Hydrogen cyanide production

Hydrogen cyanide production of planktonic and biofilm cultures was assessed by the method described by Alstrom et al. (1989). Planktonic and biofilm cultures (100 µL) were spread on KB agar plates in order to form bacterial lawns. Sterile filter papers soaked in 2% sodium carbonate and 0.5% picric acid was placed on the upper lid of Petri plates and incubated at 30°C for up to 2 days. Positive hydrogen cyanide production resulted in a colour development from yellow to reddish-brown. This experiment was conducted three times with three replicates for each treatment. The non-inoculated media served as the negative control.

3.2.7 Antimicrobial activity

The antagonistic activities of mono- and dual-species biofilm cultures were analyzed and related to planktonic cultures by performing diffusion assays as described by Berg et al. (2005), with slight modifications. A 5-mm disk of fungal pathogen mycelia was placed in the center of a Potato Dextrose Agar (PDA) (39 g L-1) and the plate was inoculated with the planktonic or biofilm cultures (10 µL) at a distance of 2 cm. For pathogenic bacteria, a lawn was streaked on KB agar plates and inoculated with the planktonic or biofilm culture on sterile filter disks in the center of the agar plates. The plates were incubated at 30 °C for up to 4 days and the zones of inhibition

28 were measured as previously described. Phytopathogens tested are Clavibacter michiganansis, Pseudomonas syringae, Schlerotinia minor and Fusarium graminearum.

3.2.8 Minimal bactericidal concentration

Antibiotic resistance was analyzed by minimal bactericidal concentration (MBC) assays for both planktonic (P) and biofilm (B) cultures as described by Mah et al. (2003). Assays were performed in a 96 well microtiter plate (Falcon). For the MBC-B assay, 100 µL of overnight bacteria was inoculated at a 1:100 dilution in TSB supplemented with 0.5% glucose and incubated at 30 °C under static conditions for 24 h. Spent media was then replaced with 90 µL of fresh media and serial dilutions of 10 µL of antibiotics of interest (kanamycin, gentamycin, streptomycin and tetracycline) were supplied. Plates were then incubated for 24 h at 30°C. Spent media was then replaced with 100 µL fresh media to remove planktonic cells and bacterial cells which survived the antibiotic treatment were incubated for 24 h at 30°C. Surviving biofilms cells were then plated onto KB plates by a replica plater for 96 well plates (Sigma-Aldrich) and grown at 30°C for 24 h, in order to establish the concentration of drug which completely killed biofilm cells. For the MBC- P assay, 90 µL of overnight bacteria was inoculated at a 1:100 dilution in TSB supplemented with 0.5% glucose with serial dilutions of 10 µL of antibiotics of interest and incubated at 30 °C under static conditions for 24 h. Surviving planktonic cells were then plated onto KB plates and grown at 30°C for 24 h in order to establish the concentration of drug which completely killed planktonic cells.

3.3 Effect of planktonic and biofilm inoculants on crop growth

To determine if mono- and dual-species biofilms are more effective at enhancing crop growth than their planktonic counterparts, plant growth assays were conducted. The experiment was conducted in 15.24 cm (6 inch) diameter plastic pots filled with 1,200 mL sterile vermiculite. Each pot was seeded with three surface sterilized seeds of tomato (Stokes, U.S.A.) placed 6.35 cm

(2.5 inches) below the rooting medium surface. Each pot received 100 mL of dH2O and was placed in the greenhouse at 25 ± 2 °C with a relative humidity of 60-80% and a 16 h:8 h (day: night) cycle. Plants were then thinned to 1 plant per pot. Once the first true leaves developed each pot

29

7 -1 received either; 35 mL 10mM MgSO4 (non-inoculated) or 35 mL (10 cfu mL of Bacillus sp. 9 -1 and/or 10 cfu mL Pseudomonas sp. resuspended in sterile 10 mM MgSO4) of the appropriate bacterial inoculants. Bacterial inoculants were prepared as described in section 3.2. Tomato plants were given either ½ strength Hoagland Solution (HS), ½ strength HS with insoluble phosphorus

(HS-P) in the form of tricalcium phosphate (Ca3(PO4)2), ½ strength HS with insoluble iron (HS-

Fe) in the form of ferric chloride (FeCl3) or ½ strength HS with insoluble phosphorus and iron (HS-FeP) once a week until harvested (45 days total). This experiment was conducted twice with 6 replicates per treatment. At harvest, data was collected on shoot dry weight (DW), root (DW), root length, leaf area and plant height. Plant DW’s were determined by drying plant material at 60°C for 3 days.

In addition, SM observations were performed on the tomato root system. Briefly, tomato roots were harvested and placed in a sterile falcon tube. To determine if and where biofilms formed, 30 mL of the SYPRO® stain was immediately added to each tube and incubated as previously described in section 3.1.7. The roots were then rinsed twice with dH2O in order to remove excess stain and visualized under the microscope. After viewing the entire root system, root sections which were representative of the overall system were cut into small sections and placed on a sterile slide with 8 µL of dH2O in order to prevent drying.

30

Chapter 4

RESULTS

4.1 Bacterial strain identification

4.1.1 16s rRNA sequencing analysis

Isolates which showed significant plant growth promotion in the bacterial screening bioassay previously conducted by Di Fan (PhD student in Dr. Smith’s laboratory) were chosen for further analysis. Efficient extraction of DNA was important in order to proceed with downstream 16s rRNA analysis. The concentration and purity of the DNA extracted from bacterial cells was determined. The A260/280 ratio of each sample was above 1.8, whereas the A260/230 ratio was above 2, indicating that pure genomic DNA was successfully extracted. The purity of the DNA samples was verified via electrophoresis on a 1% weight/volume agarose gel. A single band with a molecular weight above 1 kilobase (Kb) was obtained for each isolate, signifying that pure whole genomic DNA was effectively retrieved. Sequences for 16s rRNA were retrieved from the Genome Quebec Innovation Center and were subject to nucleotide BLAST analysis (Table 1). Both isolates sequenced were from the phylum Proteobacteria and were from the families Bacillaceae and Pseudomonadaceae. More specifically, isolate 1 was identified as a Bacillus sp. while isolate 2 was identified as a Pseudomonas sp. However the exact species of the isolates was undetermined as multiple hits, with the same query coverage, max identity score and e value, were obtained. Future work should include sending the material for full sequencing.

31

Table 1. Molecular identification of bacterial isolates based on blastN queries in NCBI.

Isolate Highest scoring 16s Query Max Accession rRNA BLAST match coverage identity number % % 1 Bacillus sp. 74 98 LN680102 2 Pseudomonas sp. 82 97 KJ534477

32

4.2 Growth curves

4.2.1 Optimal growth conditions of mono-species planktonic cultures

To establish the optimal growth conditions required for the two bacterial strains, growth curves were developed (Figure 3). The Bacillus sp. grew equally well at 30 and 37 °C and showed no preference regarding type of medium (Figure 3a). Conversely, the Pseudomonas sp. grew faster at 30 °C than 37 °C and there was also no difference in growth rate among the evaluated media (Figure 3b). When comparing the growth rates of Pseudomonas sp. to Bacillus sp., it was evident that the Pseudomonas sp. grows much more slowly than the Bacillus sp. The Pseudomonas sp. took 6 h to reach the stationary phase at 30 °C compared to 4 h for the Bacillus sp. At 37 °C the Pseudomonas sp. took 10 h to reach the stationary phase versus 4 h for the Bacillus sp. Therefore all subsequent experiments were conducted at 30 °C.

33

A Bacillus sp.

1.4 1.2 LB -> 30 °C 1 TSB -> 30 °C 0.8 KB -> 30 °C 0.6 TBS+glucose -> 30°C 0.4 LB -> 37 °C 0.2 0 TSB -> 37 °C 0 1 2 3 4 5 6 7 8 9 10 KB -> 37 °C

Optical density (OD) at nm 600 at (OD) density Optical TBS+glucose -> 37°C Time (hours)

B Pseudomonas sp.

LB -> 30 °C 1.4 1.2 TSB -> 30 °C 1 KB -> 30 °C 0.8 TBS+glucose -> 30°C 0.6 LB -> 37 °C 0.4 0.2 TSB -> 37 °C 0 KB -> 37 °C 0 1 2 3 4 5 6 6.5 7.5 8.5 9.5 10 TBS+glucose -> 37°C Optical density (OD) at nm 600 at (OD) density Optical Time (hours)

Figure 3. Mono – species planktonic growth curves. Bacillus sp. (Figure 3a) and Pseudomonas sp. (Figure 3b) cultures were grown in various growth media at two different temperatures until the stationary phase was reached.

34

4.3 Biofilm formation assay

4.3.1 Optimal growth condition of mono-species biofilm cultures

To determine the capacity of both bacterial strains to form biofilms, crystal violet assays were conducted (Figure 4). The Bacillus sp. was only able to form biofilms in the TSB supplemented with 0.5% glucose media (Figure 4a). By contrasts, the Pseudomonas sp. was capable of forming biofilms, to varying degrees, in all 8 media (Figure 4b). More specifically, weak biofilms formed in MS, LB, TSB and BH media, whereas strong biofilms formed in KB, TSB supplemented with 0.5% glucose and both monobasic and dibasic M63 media. Furthermore there was only a slight difference between the 1/50 and 1/100 dilutions and the use of 0.1 and 1 % CV for both bacterial species. Since both bacterial species can form biofilms in TSB supplemented with 0.5% glucose, all subsequent experiments were conducted with this media.

35

e e inoculation at a

broth (KB), Murashige and Murashige (KB), broth

4b) 4b) cultures wer

ure

(Fig

sp. sp.

Pseudomonas

4a) and

ure

Bertani (LB), Trypsin soy broth (TSB), King’s King’s (TSB), broth soy Trypsin (LB), Bertani

-

(Fig

sp. sp.

Bacillus

B

A

Skoog Skoog (MS), TSB supplemented with glucose, monobasic minimal media (M63 mono), dibasic minimal media (M63 di) and 1/100 and 1/50 ratio in various growth media: Luria media: growth various in ratio 1/50 and 1/100 solution violet (CV) 0.1% and crystal and 1% a with stained (BH) Haas Bushnell Figure 4. Biofilm formation assays.

36

4.3.2 Stereo microscopy imaging

To determine if the SYPRO® stain can distinguish between biofilm and planktonic cells for future plant experiments, stereo microscopy (SM) imaging with both light and fluorescent microscopy was conducted (Figure 5). Light microscopy of the Bacillus sp. and Pseudomonas sp. planktonic cells (Figures 5a and Figure 5e, respectively) in addition to the biofilm cells (Figures 5c and Figure 5g, respectively) were visualized on tabs of the 96 well plates, and placed on sterilized slides. However fluorescent imaging of the Bacillus sp. and Pseudomonas sp. planktonic cells did not show any florescent staining with the SYPRO® stain (Figures 5b and Figure 5f, respectively) whereas the Bacillus sp. and Pseudomonas sp. biofilms were able to produce a staining pattern with the SYPRO® stain (Figures 5d and Figure 5h, respectively).

37

Bacillus sp. Pseudomonas sp.

Planktonic Light microscopy

A E

Planktonic SYPRO® Ruby stain

B F

Biofilm Light microscopy

C G

Biofilm SYPRO® Ruby stain

D H

Figure 5. Bacillus sp. and Pseudomonas sp. stereo microscopy (SM) imaging. Assessment of the SYPRO® stain via SM, to distinguish between planktonic and biofilm formation. Phase contrast imaging of Bacillus sp. planktonic (Figure 5a) and biofilm (Figure 5c) conditions, in comparison to fluorescent imaging of planktonic (Figure 5b) and biofilm (Figure 5d) cells, are shown above. Phase contrast imaging of Pseudomonas sp. planktonic (Figure 5e) and biofilm (Figure 5g) conditions, in comparison to fluorescent imaging of planktonic (Figure 5f) and biofilm (Figure 5h) conditions, are also shown above.

38

4.4 Optimization of bacterial cultures

4.4.1 Standardization of mono and dual – species cultures

When bacterial isolates were grown on KB agar plates there was a clear visual difference in morphology between the two strains (Figure 6). Specifically, the Bacillus sp. formed white colonies with undulate margins (Figure 6a), whereas the Pseudomonas sp. formed yellow/greenish circular colonies (Figure 6b). Therefore all proceeding standardization experiments were conducted on KB agar plates, in order to determine if both bacterial species co-exist, in potential dual-species culture, and to determine the quantity of each strain. The CFU mL-1 of each bacterial culture was recorded in order to establish a constant number of cells for the biochemical analysis and plant growth assay, which compared planktonic and biofilms cells (Tables 2 and 3). For the mono-species cultures at a 1/100 dilution, the Pseudomonas sp. planktonic culture had 1.30E+08 to 1.10E+09 CFU mL-1 for the mid exponential phase to the 24 h mark indicating a constant increase in the number of cells (Table 2). Meanwhile, the Bacillus sp. planktonic culture displayed 3.90E+07 to 3.40E+08 CFU mL-1 for the mid exponential phase to the 24 h mark displaying a continuous increase in CFU (Table 2).

As for the Pseudomonas sp. biofilm culture, there were 1.69E+09 to 6.53E+09 CFU mL-1 for the day 1 to day 5 growth stages indicating that the biofilm contains a constant number of cells (Table 2). The Bacillus sp. biofilm culture showed 4.47E+07 to 1.60E+08 CFU mL-1 for the day 1 to day 5 growth stages suggesting that the biofilm was relatively steady (Table 2). For the dual- species biofilm culture at a 1/100 dilution for both bacterial strains, the CFU range of Bacillus sp. was from 3.53E+06 to 3.27E+07 while the CFU range of Pseudomonas sp. was from 6.50E+08 to 4.17E+09 for the day 1 to day 5 growth stages. These values signify that both bacterial strains can in fact co-exist with each other and that the biofilm is in a stable state (Table 3).

For the planktonic culture, various volumes (mL) of each strain, Bacillus sp. to Pseudomonas sp., respectively, were tested at the 24 h mark. The CFU of each bacterial species were as follows. The ratio of 1:1 (1 mL of Bacillus sp. culture to 1 mL Pseudomonas sp. culture) resulted in 5.43E+08 to 4.13E+07, the ratio of 0.5:10 resulted in 3.50E+08 to 4.37E+08, the ratio

39 of 0.25:15 resulted in 2.73E+08 to 6.07E+08, the ratio of 0.1:20 resulted in 1.07E+08 to 8.27E+08 and the ratio of 0.05:30 resulted in 1.70E+07 to 1.03E+09 indicating that a substantial amount of the Pseudomonas sp. (in comparison to the Bacillus sp.) is required to obtain comparable CFU to its mono biofilm counterpart (Table 3).

In order to compare potential differences between the resulting bacterial mixes, for subsequent experiments, a constant number of CFU for each treatment was established. More specifically, the mono-species Pseudomonas sp. planktonic and biofilm cultures were grown for 24 h in order to obtain a CFU of 109. The mono-species Bacillus sp. planktonic culture was grown to the mid exponential growth stage whereas the mono-species Bacillus sp. biofilm culture was grown for 24 h, in order to acquire a CFU of 107 for both the planktonic and biofilm inoculants. Furthermore, the dual-species biofilm culture was grown for 48 h while the dual-species planktonic culture was grown at a 0.05:30 ratio for 24 h in order to obtain a CFU of 107 and 109 for Bacillus sp. and Pseudomonas sp., respectively.

40

A B A Figure 6. Bacterial colony morphology. Bacillus sp. (Figure 6a) and Pseudomonas sp. (Figure 6b) grown on KB agar plates show distinct differences in colony morphology allowing for the determination to mixed cultures.

41

diluted

Day 5 Day

1.13E+09

4.47E+07

-1

Day 4 Day

2.07E+09

7.37E+07

Day 3 Day

1.93E+09

1.33E+08

Biofilm CFU Biofilm mL CFU

Growth stage Growth

Day 2 Day

6.53E+09

1.60E+08

Day 1 Day

1.69E+09

8.50E+07

Strain

Bacillus sp. Bacillus

Pseudomonassp.

1/100

1/100

Dilution

24 hours 24

1.10E+09

3.40E+08

-1

4.10E+08

1.30E+08

Early stationary Early

Growth stage Growth

1.30E+08

3.90E+07

Planktonic CFU mL CFU Planktonic

Midexponential

Strain

Bacillus sp. Bacillus

Pseudomonassp.

1/100

1/100

Dilution

Mono - Mono species

Table 2. Standardization of mono species cultures species of TableStandardization 2. mono

Mono species planktonic and biofilm cultures were inoculated at a 1/100 ratio, grown to different growth stages and serially serially and stages growth different to grown ratio, 1/100 a at inoculated were cultures biofilm and planktonic species Mono units (CFU) forming colony standardize to

42

iluted iluted

Day 5 Day

1.37E+09

4.40E+06

-1

Day 4 Day

4.17E+09

4.00E+06

Day 3 Day

1.53E+09

3.53E+06

Biofilm CFU mL

Day 2 Day

1.60E+09

1.37E+07

in in mL; 1:1, 0.5:10, 0.25:15, 0.1 to

Day 1 Day

6.50E+08

3.27E+07

Growth stage Growth

Strain

Pseudomonas Pseudomonas sp.

Bacillus sp.

to to

Pseudomonas sp.

sp. sp.

1/100

1/100

Dilution

Bacillus

-1

24 hours 24

1.03E+09

1.70E+07

8.27E+08

1.07E+08

6.07E+08

2.73E+08

4.37E+08

3.50E+08

4.13E+07

5.43E+08

Growth stage Growth

Planktonic CFU mL Planktonic

Strain

n of dual species cultures nof species dual

Pseudomonas sp.

Bacillus sp.

Pseudomonas sp.

Bacillus sp.

Pseudomonas sp.

Bacillus sp.

Pseudomonas sp.

Bacillus sp.

Pseudomonas sp.

Bacillus sp.

1

1

30

20

15

10

0.1

0.5

0.05

0.25

Dual - species - Dual species

Volume of bacteria (ml) of Volume

TableStandardizatio 3.

20 20 and 0.05 to 30) whereas biofilm cultures were inoculated at a 1/100 ratio, grown to different growth stages and serially d units (CFU) forming colony standardize to Dual species planktonic cultures were grown at various ratios (

43

4.5 Phenotypic characterization of planktonic and biofilms cultures.

4.5.1 Nitrogen fixing ability

The plant growth promoting rhizobacteria (PGPR) were assessed for their ability to fix nitrogen by plating them on nitrogen-free agar plates as described by Ker (2011). None of the bacterial cultures were able to fix nitrogen (Table 4).

4.5.2 Phosphate solubilisation activity

PGPR strains were evaluated for their ability to solubilise phosphorus on Pikovskaya (PVK) and National Botanical Research Institute’s phosphate (NBRIP) agar plates as described by Johri et al (1999) and Nautiyal (1999). Both bacterial strains, with the exception of the Bacillus sp. planktonic culture, were able to solubilise inorganic phosphate from the PVK and NBRIP agar plates (Table 4). Zones of phosphate solubilisation from the PVK and NBRIP agar plate assays were comparable, but not identical. The Pseudomonas sp. biofilm was capable of higher amounts of solubilisation (4.78 and 4.0 mm) than its planktonic counterpart (3.33 and 2.55 mm) for the PVK and NBRIP assays, respectively. Additionally, the Bacillus sp. biofilm showed greater solubilisation ability (4.33 and 3.22 mm) than the planktonic culture, which did not show any activity on the PVK or NBRIP plates. Moreover, the mixed dual-species biofilm showed a higher capacity for solubilisation (4.77 and 4.0 mm) than the mixed planktonic culture (3.11 and 2.33 mm) for the PVK and NBRIP assays. The quantity of inorganic phosphate solubilisation was determined utilizing liquid NBRIP solution supplemented with 1000 µg P mL-1 in the form of tri- calcium phosphate (TCP) and quantified spectrophotometrically with a standard curve as described by Fiske et al (1925) (Figure 7). A student t-test was performed in order to determine if biofilms are more effective than planktonic cells. In the presence of TCP, the Bacillus sp. biofilm performed significantly (p < 0.0001) better (343.96 µg mL-1) than its planktonic culture (162.69 µg mL-1). Similarly, the Pseudomonas sp. biofilm (550 µg mL-1) showed a significant (p < 0.0001) increase over its planktonic counterpart (291.26 µg mL-1). Finally, the mixed dual-species biofilm showed significantly (p < 0.0001) more activity (624.12 µg mL-1) than the planktonic culture (296.34 µg mL-1).

44

Table 4. Biochemical attributes of mono and dual-species biofilm and planktonic plant growth promoting rhizobacteria

Biochemical characteristics

Treatment Nz PVKy NBRIPy Ammoniax CASw O-CASw HCNv fixing (mm) (mm) Bacillus sp. biofilm - 4.33±0.03 3.22±0.32 +++ ++ ++ ++ Bacillus sp. planktonic - - - ++ - + +

Pseudomonas sp. - 4.77±0.09 4.00±0.47 ++++ ++++ ++++ ++++ biofilm Pseudomonas sp. - 3.33±0.02 2.55±0.23 +++ +++ +++ +++ planktonic

Mixed biofilm - 4.77±0.09 4.00±0.47 +++++ ++++ ++++ ++++ Mixed planktonic - 3.11±0.09 2.33±0.15 ++++ +++ +++ +++

Z: The ability of each bacterial culture to fix nitrogen was determined by observing the presence (+) or absence (-) of colonies on N-free agar plates.

Y: Radius of solubilisation measured on both National Botanical Research Institute’s phosphate (NBRIP) and Pikovskaya (PVK) agar plates.

X: The degree of activity for ammonia production was determined by observing a colour change from light yellow (-) to dark orange (+++++)

W: The degree of activity for siderophore production (++++ > +++ > ++ > + > -) was determined by observing a halo around the colony on CAS and O-CAS agar plates

V: The degree of activity for hydrogen cyanide (HCN) was determined by observing a colour change from light yellow (-) to dark brown (++++)

45

Figure 7. Phosphate solubilisation activity of mono and dual-species biofilm and planktonic plant growth promoting rhizobacteria. Within each pair of columns biofilm against planktonic treatments with asterisks are significantly different at p < 0.001.

46

4.5.3 Ammonia production

The PGPR ability to produce ammonia was assessed with Nessler reagent (Dye et al., 1962). All bacterial strains were capable of producing ammonia, as shown by the development of colour from light yellow to dark orange (Table 4). The amount of ammonia produced from highest to lowest was as follows: the mixed dual-species biofilm (+++++), followed by the Pseudomonas sp. biofilm as well as the mixed-species planktonic culture (++++), then the Bacillus sp. biofilm in addition to the Pseudomonas sp. planktonic culture (+++) and finally the Bacillus sp. planktonic culture (++).

4.5.4 Siderophore production

Siderophore production by the evaluated PGPR was detected by the CAS and O-CAS methods (Schwyn et al., 1987; Perez-Miranda et al., 2007). All strains produced siderophores with the exception of the Bacillus sp. planktonic culture, corresponding to the CAS assay as shown by the development of a colour change from blue to orange (Table 4). Assessment of siderophore production by the CAS and O-CAS methods were similar and comparable for the Pseudomonas sp. but not comparable for the Bacillus sp. The results for the CAS method are as follows. The Pseudomonas sp. biofilm exhibited higher siderophore production (++++) than its planktonic counterpart (+++). Furthermore, the Bacillus sp. biofilm had greater levels of siderophore activity (++) than the planktonic culture, which showed no production of siderophore (-). Finally, the mixed dual-species biofilm had enhanced siderophore activity (++++) compared to the planktonic culture (+++).

In addition, the O-CAS assay was performed and the results were as follows. The Pseudomonas sp. and dual-species mixed biofilm were able to produce slightly higher levels of siderophore activity (++++) than the Pseudomonas sp. and dual-species mixed planktonic culture (+++). Likewise, the Bacillus sp. biofilm strain showed enhanced siderophore activity (++) compared to its planktonic counterpart (+) (Table 4). The quantity of siderophores produced by PGPR culture was established by means of liquid CAS solution (Figure 8) as described by Schwyn et al (1987). A student t-test was performed in order to determine if biofilms are more effective

47 than planktonic cells. The Bacillus sp. biofilm strain showed significantly (p = 0.0003) higher levels of siderophore (6.08 %) than its planktonic counterpart (0.25%). Additionally, the Pseudomonas sp. biofilm showed a significantly (p = 0.0018) enhanced siderophore activity (9.59 %) compared to the planktonic culture (2.77 %). Finally, the mixed dual-species biofilm showed significantly (p = 0.0032) more siderophores activity (9.77 %) than its planktonic equivalent (3.29 %).

48

Figure 8. Siderophore production by mono and dual-species biofilm and planktonic plant growth promoting rhizobacteria. Within each pair of columns a significant difference in the biofilm against planktonic cultures comparison is indicated with asterisks; these differences were significant at ** p < 0.01 and *** p < 0.001.

49

4.5.5 Hydrogen cyanide production

The production of hydrogen cyanide by bacterial cultures was evaluated by the method of Alstrom et al. (1989). All bacterial strains showed positive results for hydrogen cyanide production, as shown by the development of colour from yellow to reddish-brown (Table 4). The highest amount of hydrogen cyanide produced was for the Pseudomonas sp. and mixed dual- species biofilms (++++) followed by their planktonic equivalents (+++). Lower amounts were produced by the Bacillus sp. biofilm (++) followed by its planktonic counterpart (+).

4.5.6 Inodole-3-acetic acid (IAA) production

PGPR cultures were assayed for their ability to produce IAA in the presence of the IAA precursor L-tryptophan (200 µg mL-1) as described by Hartmann et al (1983). All strains demonstrated the ability to produce IAA (Figure 9). A student t-test was performed in order to determine if biofilms are more effective than planktonic cells. In the presence of L-tryptophan, the Bacillus sp. biofilm produced significant (p < 0.0001) amounts of IAA (0.45 µg mL-1) compared to its planktonic counterpart (0.07 µg mL-1). In addition, the Pseudomonas sp. biofilm produced significantly (p = 0.0152) higher amounts of IAA (2.37 µg mL-1) than the planktonic isolate (1.29 µg mL-1), similarly to the dual species biofilm which showed significantly ((p = 0.0006) higher IAA levels (3.10 µg mL-1) than the planktonic dual-species mixture (1.17 µg mL-1).

50

Figure 9. Indole acetic-acid production by mono and dual-species biofilm and planktonic plant growth promoting rhizobacteria. Within each pair of columns biofilm against planktonic treatments with asterisks are significantly different at * p < 0.05 and *** p < 0.001.

51

4.5.7 Antimicrobial activity

The antagonistic activity of mono- and dual-species B-PGPR and P-PGPR against 4 phytopathogens were evaluated, and was indicated by a zone of inhibition around the colonies (Table 5) (Berg et al., 2005). Pseudomonas sp. biofilm (15 mm) caused a greater zone of inhibition against Clavibacter michiganensis than its planktonic counterpart (10 mm). Correspondingly, the mixed dual-species biofilm (15 mm) showed activity levels similar to its planktonic culture (10 mm). However, the Bacillus sp. biofilm (1 mm) and its planktonic counterpart (1 mm) did not show any effect. In addition, the Pseudomonas sp. biofilm had greater inhibition against Pseudomonas syringae (9 mm) than its planktonic equivalent (7 mm). Equally, the mixed dual- species biofilm had level of activity (9 mm) similar to the planktonic culture (7 mm). Conversely, the Bacillus sp. biofilm and its planktonic counterpart did not have any effect on this phytopathogen. Moreover, the Pseudomonas sp. and mixed dual species biofilms (3.83 mm for each isolate) performed better than their planktonic counterparts (2.66 and 2.83 mm, respectively) against Schlerotinia minor, whereas there was no difference between the Bacillus sp. biofilm (3.33 mm) and planktonic (2.66 mm) forms. Finally the Pseudomonas sp. and mixed dual species biofilms (5.77 mm for each isolate), in addition to the Bacillus sp. biofilm (4.10 mm) exhibited more antimicrobial activity than their planktonic counterparts (3.66 mm for the mixed and Pseudomonas sp. culture and 0.11 mm for the Bacillus sp. culture) against Fusarium graminearum.

4.5.8 Minimal bactericidal concentration planktonic/biofilm (MBC P/B)

The minimal bactericidal concentration (MBC) of bacterial isolates against 4 antibiotics was established by determining the minimal amount of antibiotic required to completely kill the bacterial colonies as described by Mah et al (2003) (Table 6). In regards to gentamycin, the Pseudomonas sp. (200 µg), Bacillus sp. (60 µg) and mixed (480 µg) biofilms were much more resistant than their planktonic counterparts (60, 10 and 10 µg, respectively). Similarly for kanamycin, all three biofilms (Pseudomonas sp. 120 µg, Bacillus sp. 200 µg and mixed 250 µg) were also more resistant to their planktonic equivalents (60, 30 and 30 µg, respectively). The same trend occurred for streptomycin biofilms (Pseudomonas sp. 120 µg, Bacillus sp. 200 µg and mixed 240 µg) in that they demonstrated greater resistance than their planktonic partners (60, 30 and 60

52

µg, respectively). Finally for tetracycline, the Bacillus sp. biofilm (60 µg) showed enhanced resistance in comparison to the planktonic culture (1 µg). However, the MBC for the Pseudomonas sp. and the mixed planktonic and biofilm cultures could not be determined since the antibiotic dilution for this treatment exceeded the maximum stock solution (10 mg/ml) that could be produced.

53

Table 5. Antimicrobial activity of biofilm and planktonic plant growth promoting rhizobacteria against 4 plant pathogens

Inhibition zone (mm)z

Treatment Clavibacter Pseudomonas Schlerotinia Fusarium michiganensis syringae minor graminearum

Bacillus sp. Biofilm 1.00 a -y 3.33 a 4.10 a Bacillus sp. Planktonic 1.00 a - 2.66 a 0.11 b

Pseudomonas sp. Biofilm 15.22 a 9.96 a 3.83 a 5.77 a Pseudomonas sp. Planktonic 10.00 b 7.00 b 2.66 b 3.66 b

Mixed Biofilm 15.88 a 9.61a 3.83 a 5.77 a Mixed Planktonic 10.00 b 7.00 b 2.83 b 3.66 b

Within a column biofilm against planktonic treatments with different letters are significantly different at p < 0.001.

z: Values are means of 3 replicates. y: Hyphens (-) indicate that no inhibition was observed.

54

Table 6. Minimal bactericidal concentration of mono and mixed planktonic and biofilm cultures

MBC (µg mL-1)z

Treatment Gentamycin Kanamycin Streptomycin Tetracycline Py Bx P B P B P B

Bacillus 10 60 30 200 30 200 -w -

Pseudomonas 60 200 60 120 60 120 1 60

Mixed 10 480 30 250 60 240 - -

Z: Values are means of 3 replicates Y: Planktonic culture X: Biofilm culture W: Hypens (-) indicates that the MBC could not be established

55

4.6 Bacterial crop inoculation

4.6.1 Effect of planktonic and biofilm inoculants on crop growth

In order to determine if mono- and dual-species B-PGPR are more effective at enhancing crop growth in comparison to their planktonic equivalents, greenhouse experiment were conducted with tomato plants. All the data were verified for the assumptions of normality and homosdasticity.

4.6.1.1 Plant height

Plant height of the non-inoculated plants given Hoagland Solution (HS) (7.12 cm) showed enhanced values of the measured variables in comparison to the non-inoculated plants given HS- FeP (3.45 cm) and HS-P (3.91 cm) (Tables 7, 8 and 9). Plants inoculated with either Bacillus sp. planktonic (6.83 to 10.79 cm) or Bacillus sp. biofilm (10.08 to 14.41 cm) and provided with HS, HS-F, HS-P or HS-FeP were taller (3.45 to 7.12 cm) than the control plants (Table 7). Similar increases resulted from treatment with both the planktonic and biofilm Pseudomonas sp. (7.04 to 10.70 cm and 11.75 to 16.66 cm, respectively) and mixed, Bacillus sp. plus Pseudomonas sp., inoculants (7.50 to 10.70 cm and 11.25 to 14 cm, respectively) (Tables 8 and 9, respectively). In addition, Bacillus sp. biofilm-treated plants performed better than plants inoculated with Bacillus sp. planktonic cells when given HS-F, HS-P or HS-FP (Table 7). However, the Pseudomonas sp. and mixed biofilm treated plants were taller when provided with the full HS than their planktonic cell inoculated equivalents (Tables 8 and 9, respectively). Moreover the Bacillus sp., Pseudomonas sp. and mixed biofilm treated plants given either HS-FeP or HS-P showed an increase in plant height in comparison to the non-inoculated control given HS, whereas treatment with their planktonic counterparts did not result in any plant height differences (Tables 7, 8 and 9, respectively).

56

Table 7. The effects of Bacillus sp. inoculants on tomato plant growth variables of plants grown under greenhouse conditions.

Treatment Height (cm) Root DW (g)Root length (cm) Shoot DW (g) Leaf area (cm2) Type Biofilm 12.18 a 1.07 a 29.20 a 0.62 a 82.86 a Planktonic 8.85 b 0.36 b 23.95 b 0.38 b 59.84 b Non-inoculated 5.29 c 0.09 c 18.66 c 0.12 c 21.92 c

Nutrient HSz 10.19 a 0.66 a 25.58 a 0.54 a 78.66 a HS-Py 6.94 b 0.33 b 25.30 a 0.23 b 33.6 b HS-Fex 10.62 a 0.64 a 23.97 a 0.50 a 72.71 a HS-FePw 7.34 b 0.39 b 20.19 b 0.22 b 34.53 b

Type x Nutrient Planktonic HS 10.79 a 0.50 c 26.00 bc 0.47 bc 82.03 b Planktonic HS-P 6.83 c 0.23 d 23.22 cd 0.25 de 36.03 e Planktonic HS-Fe 10.29 b 0.48 c 24.41 bc 0.59 b 83.46 ab Planktonic HS-FeP 7.55 c 0.24 d 22.08 de 0.22 e 37.85 ed Biofilm HS 13.16 a 1.31 a 31.00 a 0.86 a 109.15 a Biofilm HS-P 10.08 b 0.75 b 30.00 ab 0.38 cd 53.73 cd Biofilm HS-Fe 14.41 a 1.34 a 29.67 ab 0.82 a 111.55 a Biofilm HS-FeP 11.08 b 0.88 b 26.16 bc 0.42 c 57.03 c Non-inoculated HS 7.12 c 0.17 e 21.33 ed 0.21 e 44.80 cde Non-inoculated HS-P 3.91 d 0.02 f 18.58 e 0.07 f 11.04 f Non-inoculated HS-Fe 6.66 c 0.12 e 20.25 e 0.17 ef 23.23 f Non-inoculated HS-FeP 3.45 d 0.03 f 14.50 f 0.03 f 8.72 f P value Type <0.001 <0.001 <0.001 <0.001 <0.001 Nutrient <0.001 <0.001 <0.001 <0.001 <0.001 Type x Nutrient NSv 0.0002 NS <0.001 0.0001 Different letters within a column annd for a given treatment indicate significant differences (P <0.05). v Non-significant with P>0.05

Z: Hoagland solution

Y: Hoagland solution with insoluble phosphate

X: Hoagland solution with insoluble iron

W: Hoagland solution with insoluble iron and phosphate

57

Table 8. The effects of Pseudomonas sp. inoculants on tomato plant growth variables of plants grown under greenhouse conditions

Treatment Height (cm) Root DW (g) Root length (cm) Shoot DW (g) Leaf area (cm2) Type Biofilm 13.60 a 1.54 a 28.58 a 0.75 a 128.07 a Planktonic 8.67 b 0.48 b 22.75 b 0.41 b 71.06 b Non-inoculated 5.29 c 0.08 c 18.45 c 0.12 c 21.17 c

Nutrient HSz 11.50 a 0.955 a 25.83 a 0.066 a 98.37 a HS-Py 7.83 c 0.57 b 22.05 b 0.25 c 52.66 c HS-Fex 9.93 b 0.84 c 25.11 a 0.57 b 89.69 b HS-FePw 7.50 c 0.45 d 20.05 b 0.23 c 52.99 c

Type x Nutrient Planktonic HS 10.70 c 0.81 d 26.50 a 0.77 b 98.64 b Planktonic HS-P 7.83 e 0.30 f 20.58 bc 0.23 d 43.70 c Planktonic HS-Fe 9.12 d 0.53 e 25.58 a 0.43 c 92.76 b Planktonic HS-FeP 7.04 ef 0.29 f 18.33 c 0.22 d 49.15 c Biofilm HS 16.66 a 1.89 a 29.66 a 1.11 a 154.59 a Biofilm HS-P 11.75 c 1.37 b 27.00 a 0.46 c 103.73 b Biofilm HS-Fe 14.00 b 1.87 a 30.33 a 1.00 a 151.51 a Biofilm HS-FeP 12.00 c 1.01 c 27.33 a 0.45 c 102.45 b Non-inoculated HS 7.12 ef 0.16 g 21.33 b 0.21 d 44.98 c Non-inoculated HS-P 3.91 g 0.02 h 18.58 bc 0.07 e 10.56 d Non-inoculated HS-Fe 6.66 f 0.12 g 19.41 bc 0.16 d 21.74 d Non-inoculated HS-FeP 3.45 g 0.03 h 14.50 d 0.03 e 7.38 d P value Type <0.001 <0.001 <0.001 <0.001 <0.001 Nutrient <0.001 <0.001 <0.001 <0.001 <0.001 Type x Nutrient NSz <0.001 0.0031 <0.001 <0.001 Different letters within a column annd for a given treatment indicate significant differences (P <0.05). zNon-significant with P>0.05

Z: Hoagland solution

Y: Hoagland solution with insoluble phosphate

X: Hoagland solution with insoluble iron

W: Hoagland solution with insoluble iron and phosphate

58

Table 9. The effects of mixed inoculants on tomato plant growth variables of plants grown under greenhouse conditions

Treatment Height (cm) Root DW (g)Root length (cm) Shoot DW (g) Leaf area (cm2) Type Biofilm 13.02 a 1.09 a 28.25 a 0.68 a 98.56 a Planktonic 8.88 b 0.37 b 23.35 b 0.34 b 50.76 b Non-inoculated 5.29 c 0.08 c 18.45 c 0.12 c 21.17 c

Nutrient HSz 10.94 a 0.69 a 24.22 a 0.56 a 83.41 a HS-Py 7.55 c 0.46 c 23.83 a 0.29 c 41.29 c HS-Fex 10.08 b 0.54 b 23.69 ab 0.43 b 58.70 b HS-FePw 7.68 c 0.45 c 21.66 b 0.24 c 43.90 c

Type x Nutrient Planktonic HS 10.70 b 0.49 d 23.85 bc 0.61 ab 84.13 bc Planktonic HS-P 7.50 d 0.31 e 24.41 bc 0.23 de 32.81 ef Planktonic HS-Fe 9.41 c 0.36 e 24.16 bc 0.34 cd 50.60 d Planktonic HS-FeP 7.91 d 0.30 e 21.00 cd 0.18 ef 35.50 e Biofilm HS 15.00 a 1.41 a 29.50 a 0.86 a 121.13 a Biofilm HS-P 11.25 b 1.01 c 28.01 ab 0.57 b 80.51 c Biofilm HS-Fe 14.16 a 1.19 b 27.51 ab 0.79 a 103.77 ab Biofilm HS-FeP 11.66 b 1.03 c 27.57 ab 0.50 bc 88.82 bc Non-inoculated HS 7.12 d 0.16 f 21.33 cd 0.21 def 44.98 de Non-inoculated HS-P 3.91 e 0.02 g 18.58 de 0.07 g 10.56 gh Non-inoculated HS-Fe 6.66 d 0.12 fg 19.41 d 0.17 efg 21.74 fg Non-inoculated HS-FeP 3.45 e 0.03 g 14.50 e 0.03 g 7.38 h P value Type <0.001 <0.001 0.0006 <0.001 <0.001 Nutrient <0.001 <0.001 <0.001 <0.001 <0.001 Type x Nutrient NSz 0.0009 <0.001 0.0031 NS Different letters within a column annd for a given treatment indicate significant differences (P <0.05). z Non-significant with P>0.05

Z: Hoagland solution

Y: Hoagland solution with insoluble phosphate

X: Hoagland solution with insoluble iron

W: Hoagland solution with insoluble iron and phosphate

59

4.6.1.2 Plant dry weight

Non-inoculated plants given HS (0.21 g) had greater shoot DW values than the non- inoculated plants given HS-FeP (0.03 g) and HS-P (0.07 g) (Tables 7, 8 and 9). Inoculation of tomato plants receiving HS growth medium with Bacillus sp. planktonic (0.22 to 0.59 g) or biofilm (0.38 to 0.86 g) resulted in shoot DWs (0.03 to 0.21 g) larger than the controls (Table 7). Similar results followed from inoculation of tomato plants with planktonic and biofilm Pseudomonas sp. (0.22 to 0.77 g and 0.45 to 1.17 g, respectively) and the cell mixture (0.18 to 0.61 g and 0.50 to 0.86 g, respectively) (Tables 8 and 9). In addition, the Pseudomonas sp. biofilm-treated plants performed better than their planktonic counterparts for any of the HS treatments (Table 8). However, the mixed biofilm inoculants performed better when given HS-Fe, HS-P and HS-FeP, while the Bacillus sp. biofilm performed better when given any of the HS nutrient media, with the exception of HS-P (Tables 7 and 9, respectively). Moreover, all biofilm treated plants provided with either HS-FeP or HS-P nutrient solution showed an increase (Tables 7, 8 and 9) in plant dry weight in comparison to the non-inoculated control given HS, whereas their planktonic counterparts did not result in differences.

4.6.1.3 Root length

Root length of non-inoculated plants given HS (21.33 cm) was greater than the non- inoculated plants given HS-FeP (14.50 cm) (Tables 7, 8 and 9). When comparing non-inoculated plants (14.50 to 21.33 cm) to all the planktonic and biofilm Bacillus sp. inoculated plants (22.08 to 26.0 cm and 26.16 to 31.0 cm, respectively) and mixed cell type inoculated plants (21.0 to 24.41 cm and 27.5 to 29.5 cm, respectively) treated plants given any of the HS treatments had greater root length (Tables 7 and 9, respectively). On the other hand, the Pseudomonas sp. planktonic cell treatments resulted in greater root lengths when given HS, HS-Fe or HS-FP (13.33 to 26.5 cm) than the non-inoculated equivalents, whereas the biofilm treatments caused greater root lengths than any of the HS treatments (27.0 to 30.33 cm) (Table 8). In addition, the Bacillus sp. biofilm- treated plants performed better than planktonic treated plants when given HS, HS-P or HS-Fe, while the Pseudomonas sp. biofilms inoculants caused an increase when the plants were provided with HS-P or HS-FeP, and finally, the mixed biofilms caused increases (p<0.001) when given HS

60 or HS-FeP (Tables 7, 8 and 9, respectively). Bacillus sp. biofilm treated plants given HS-FeP or HS-P had greater root lengths than the non-inoculated control given HS, whereas the planktonic counterparts did not cause any effects, relative to the controls (Table 7). Pseudomonas sp. biofilm treated plants given HS-P had greater root lengths than the non-inoculated control given HS, whereas treatment with the planktonic counterparts did not result in any differences (Table 8). Finally, the mixed biofilms caused increases in root length when given any of the HS solutions, whereas the corresponding planktonic inoculants did not cause any differences, in comparison to the non-inoculated HS control (Table 9).

4.6.1.4 Root dry weight

The non-inoculated plants given HS (0.17 g) had greater root DW values than the non- inoculated plants given HS-FeP (0.03 g) and HS-P (0.02 g) (Tables 6, 7 and 8). When comparing non-inoculated (0.02 to 0.17 g) to either planktonic or biofilm Bacillus sp. (0.23 to 0.50 g and 0.75 to 1.34 g, respectively) treatments, Pseudomonas sp. (0.29 g to 0.81g and 1.01 to 1.89 g, respectively) or mixed (0.30 to 0.49 g and 1.01 to 1.41 g, respectively) treated plants given any of the HS solutions, there was an increase in root DW (Tables 7, 8 and 9, respectively). Moreover, all biofilm treated plants had greater root DW than planktonic cell treated plants, when given any of the HS treatments (Tables 7, 8 and 9).

4.6.1.5 Leaf area

The non-inoculated plants provided with HS (44.8 cm2) had greater leaf areas than the non- inoculated plants given HS-FeP (8.72 cm 2), HS-P (11.04 cm2), HS-Fe (23.23 cm2) (Tables 7, 8 and 9). Non-inoculated (8.72 to 44.8 cm2), relative to Bacillus sp. planktonic or biofilm-treated plants (36.03 to 84.46 cm2 and 53.73 to 111.55 cm2, respectively), Pseudomonas sp.-treated plants (43.70 to 98.64 cm2 and 102.45 to 154.59 cm2, respectively) or mixed cell-treated plants (35.50 to 84.13 cm2 and 80.51 to 121.13 cm2, respectively) provided with any of the HS treatments resulted in greater leaf area (Table 7, 8 and 9 respectively) than the controls. The Bacillus sp. biofilm- treated plants performed better than their planktonic counterparts when given HS-P, HS or HS- FeP whereas there was an increase for the Pseudomonas sp. and mixed cell treatment when given

61 any of the HS nutrient solutions (Tables 6, 7 and 8, respectively). The Pseudomonas sp. and mixed biofilm-treated plants given HS-FeP or HS-P had greater leaf area than the non-inoculated control given HS, whereas their planktonic counterparts did not cause and leaf area difference (Tables 8 and 9, respectively).

62

4.6.2 Stereo microscopy imaging of the tomato root system

To distinguish the location and presence/absence of biofilm formation, with the aid of the SYPRO® stain, on tomato plant root systems, SM imaging with both light and fluorescent microscopy was conducted (Figure 10). Light microscopy of the Pseudomonas sp., Bacillus sp. and mixed planktonic cell inoculants (Figures 10a, 10e and 10i, respectively) in addition to the biofilm inoculants (Figures 10c, 10g and 10k, respectively) were visualized from root sections placed on sterilized slides. Fluorescent imaging of the Pseudomonas sp., Bacillus sp. and mixed planktonic inoculants revealed that biofilms do, in fact, form on specific sections of the tomato roots (Figures 10b, 10f and 10j, respectively), while the biofilm inoculants yielded much more pronounced biofilms, which formed everywhere on the root system (Figures 10d, 10h and 10l, respectively). Furthermore, fluorescent microscopy imaging of the non-inoculated treatment revealed that no biofilms formed for this treatment (Figures 10n).

63

Figure 10. Stereo microscopy (SM) of Pseudomonas sp., Bacillus sp. and mixed inoculants on tomato root systems. Phase contrast imaging of planktonic inoculant of Pseudomonas sp. (Figure 10a), Bacillus sp. (Figure 10e), mixed (Figure 10i) and non-inoculated control (Figure 10m). Fluorescent imaging of planktonic inoculant of Pseudomonas sp. (Figure 10b), Bacillus sp. (Figure 10f), mixed (Figure 10j) and non-inoculated control (Figure 10n). Phase contrast imaging of biofilm inoculant of Pseudomonas sp. (Figure 10c), Bacillus sp. (Figure 10g) and mixed (Figure 10k). Fluorescent imaging of biofilm inoculant of Pseudomonas sp. (Figure 10d), Bacillus sp. (Figure 10h) and mixed (Figure 10l).

64

Chapter 5

DISCUSSION

Biofilms are surfaced attached microbial communities which provide advantages to the microbes they contain, as compared to their planktonic counterparts. In fact, the formation of biofilms not only enhances the survival of bacteria but may also result in greater plant growth promotion by PGPR than that of planktonic PGPR. This can be explained by the high cell population density associated with biofilms, which may allow the achievement of several processes that single cells cannot accomplish efficiently (Danhorn et al., 2007). Furthermore, biofilms are able to withstand a wider range of physical conditions than planktonic cells, making the use of biofilm-PGPR (B-PGPR) a promising prospect (Baty et al., 2000; Todar, 2008). The purpose of this study was to investigate the role of biofilms as potential plant growth promoting inoculants. The specific goals of this research project were to determine if isolated bacteria can form biofilms, to determine if a stable dual-species biofilms could be developed, and to elucidate if biofilms are more effective PGPR than their planktonic counterparts.

To maximize the potential for biofilm formation, eight types of media were tested. Results from this study clearly demonstrate that the bacterial strains, identified as Bacillus sp. and Pseudomonas sp. on the basis of 16s rRNA sequencing, can form mono-species biofilms. This was expected since in most natural settings bacteria are predominantly found as biofilms, which represent a protected mode of growth allowing bacteria to survive and thrive in competitive environments (Cosertan et al., 1995; Davey et al., 2000). Results suggest that the Pseudomonas sp. formed biofilms in various types of growth media whereas the Bacillus sp. was more restricted. Importantly, the bacterial isolates shared a common biofilm formation media, TSB supplemented with 0.5% glucose, allowing for the development of a stable dual-species biofilm. This part of the experiment was crucial since most biofilms are composed of multiple species which interact with each other (Davey et al., 2000). Similarly, other researchers have reported the development of dual-species biofilms which includes; a Bradyrhizobium elkanii-Penicillium spp., a Trichoderma viride- Bacillus subtilis spp and a Trichoderma viride-Pseudomonas fluorescens spp (Jayasinghearacchi et al., 2004; Triveni et al., 2012). However, to the best of our knowledge, this

65 is the first report regarding the development of a novel Pseudomonas sp.-Bacillus sp. dual-species biofilm. In order to further investigate the potential use of biofilms as inoculants, as compared to their planktonic counterparts, optimization experiments were conducted to standardize CFU of each treatment. Planktonic and biofilm cultures having 109 CFU mL-1 of Pseudomonas sp. and 107 CFU mL-1 of Bacillus sp. were generated.

To determine if biofilms are more effective PGPR than their planktonic counterparts well established biochemical tests were performed. The present study clearly demonstrated that the bacterial cultures were positive for numerous beneficial plant growth promoting attributes. The first direct plant growth promoting mechanism tested was phosphate solubilisation. Phosphate solubilising microbes play a crucial role in the rhizosphere by converting insoluble forms of phosphate into H2PO4 or HPO4 ions by synthesizing organic acids and/or enzymes which can then be readily assimilated by plants (Rodriguez et al., 2006; Zaidi et al., 2009; Bhattacharyya et al., 2012). Numerous reports have shown that inoculation with phosphate solubilising bacteria has resulted in increased phosphate availability which ultimately led to enhanced crop yields (Vyas et al., 2009; Yasmin et al., 2011). Results from this study indicated that both Pseudomonas sp. and Bacillus sp. were capable of solubilising phosphate, which is concordant with previous studies (Rodriguez et al., 1999; Triveni et al., 2012). In fact, the Bacillus sp. planktonic mono- and dual- species cultures produced 162 to 296 µg mL-1 phosphate solubilisation activity, which is similar to previous studies on the same genus, which exhibited values from 145 to 450 µg mL-1 (Nautiyal et al., 1999; Ali et al., 2015). Interestingly the mono and dual-species biofilms were capable of solubilising 343 to 624 µg mL-1 phosphate, representing a 1.8 to 2.1 fold increase, which is comparable to studies conducted by Jayasinghearachchi et al. (2006) suggesting that biofilms are more effective phosphate biosolubilisers than their planktonic counterparts. Therefore the application of biofilm inoculums as biofertilizers could lead to higher crop growth by rendering phosphorus more readily accessible for absorption (Glick, 2012).

Bacterial cultures were also evaluated for IAA production, which is an additional direct plant growth promoting mechanism. IAA producing bacteria have been shown to have positive effects on plants in numerous ways including plant growth, development and defense responses. More specifically, one of the most prominent effects of IAA on plants is its enhancing effect on

66 root hairs and lateral roots which allows for increased access to nutrients (Patten et al, 2002). In fact numerous studies have shown that IAA producing PGPR can have beneficial effects on plants (Zaidi et al., 2006; Poonguzhali et al., 2008). In this study we found that both Pseudomonas sp. and Bacillus sp. were IAA producers when the growth medium was supplemented with L- tryptophan, the precursor for synthesis. Under natural circumstances L-tryptophan is released into plant rooting media in root exudates. More specifically, the Bacillus sp. mono and mixed-species planktonic cultures produced 0.0785 to 1.29 µg mL-1 IAA, which is in agreement with previous reports demonstrating values of 0.04 to 3.98 µg mL-1 for the same genus (Triveni et al., 2012). In my study the amount of IAA associated with biofilms ranged from 0.0452 to 2.37 µg mL-1, which is more than their planktonic counterparts. Specifically there were 5, 1.8 and 2.6 fold increases for Bacillus sp., Pseudomonas sp. and the mixed-dual culture, respectively. Others have reported similar observations when comparing biofilms to their planktonic equivalents, implying that phytostimulating biofilms may provide an advantage over planktonic forms, particularly since IAA has been associated with enhanced plant growth (Jayasinghearacchi et al., 2004; Triveni et al., 2012).

Another important trait of PGPR is the production of siderophores. Siderophores can function both directly and indirectly to promote plant growth. More specifically, PGPR can function directly by solubilising insoluble forms of iron by secreting iron chelators termed siderophores, which have high association constants for complexing iron (Miethke et al., 2007). On the other hand, siderophore producing bacteria can function indirectly by competing with other microbes for iron thereby preventing the initiation of pathogenicity (Lugtenberg et al, 2009). Indeed, there is ample evidence suggesting that PGPR capable of synthesising siderophores enhance plant growth (Freitas et al., 2005; Zhang et al., 2009). The current investigation found that Pseudomonas sp. showed positive results on CAS agar plates while the Bacillus sp. did not show any growth on this substrate. This was most likely caused by the HDTMA detergent used in the CAS medium, which has been shown to be toxic to gram positive microbes (Schwyn et al., 1987). Therefore a modified assay, O-CAS, was employed, which showed that both bacterial strains were indeed positive for siderophore production, in accordance with previous studies (Ahmad et al., 2008; Ahemad et al., 2014). The mono and dual-species planktonic cultures exhibited 0.25 to 3.29% siderophore production. These values are somewhat lower than those reported in current

67 literature, possibly due to factors such as culture and/or medium conditions, strain and/or species of the isolates in addition to growth stage (Sayyed et al., 2005; Shobha et al., 2012). On the other hand, the Bacillus sp., Pseudomonas sp. and mixed- species biofilms displayed values between 6.08 to 9.77% siderophore production, representing 24.3, 3.45 and 2.96 fold increases, respectively, compared to their planktonic counterparts. This is in agreement with current reports (Triveni et al., 2012). In fact, it would seem that iron plays a vital role in biofilm formation. A study by Harrisson et al. (2009) found that Pseudomonas siderophore deficient mutants were less able to form biofilms than the wild type, suggesting a reason why biofilms may be associated with higher levels of siderophores. Consequently B-PGPR inocula could prove imperative as a biofetilizers, by rendering iron complexes more available for plant absorption, in addition to functioning as biocontrol agents by competing for iron and thereby hindering pathogenic microbes from establishing themselves in disease initiation (Lugtenberg et al., 2009).

Alternative indirect plant growth promoting mechanisms, such as hydrogen cyanide and ammonia production, were also evaluated for each strain. Both hydrogen cyanide and ammonia are volatile compounds with antimicrobial activity which have been shown to function as biocontrol agents against vital phytopathogens (Howell et al., 1988; Voisard et al., 1989). All cultures showed positive results for both attributes, which is in agreement with previous reports (Ahmad et al., 2006; Shobha et al., 2012; Triveni et al., 2012). The Pseudomonas sp., Bacillus sp. and mixed-species biofilm showed enhanced activity in comparison to their planktonic counterparts, in agreement with the study of Triveni et al. (2012). These results therefore suggest that biofilms may be better suited biocontrol agents than their planktonic equivalents.

To further investigate the potential use of these inoculants as biocontrol agents, cultures were tested against 4 common phytopathogens. All cultures showed antagonistic effects against pathogens, with the exception of the Bacillus sp., which did not show any effect on the Pseudomonas syringae strain. In regards to Clavibacter Michiganensis, a detrimental bacterial pathogen that causes cankers on tomato crops, the most promising results were obtained for the Pseudomonas sp. and dual-species culture, which is concordant with a previous study conducted by Lanteigne et al. (2012). Similarly, these bacterial cultures also showed prominent effects against P. syringae, a bacterial pathogen with a broad host and symptom range, in addition to Fusarium

68 graminearum, a fungal pathogen which causes head blight in a variety of cereal crops, which is also in accordance with previous findings (Bai et al., 2004; Chen et al., 2013; Dhanya et al., 2014). Moreover both the Bacillus sp. and Pseudomonas sp. showed encouraging antagonist abilities in response to Schlerotinia minor, a devastating fungal pathogen that infects numerous plant hosts and causes crown rot. These findings are in general agreement with recently published literature on this genus (Fischer et al., 2010; Chen et al., 2013). These effects were probably a direct result of siderophore, hydrogen cyanide and ammonia production. The biofilms showed enhanced biocontrol activity compared to their planktonic counterparts, in agreement with the study of Triveni et al. (2012). Possible mechanisms of action responsible for these increases include biochemical attributes described above, since it has been proven that biofilms are more effective PGPR than planktonic cells. Furthermore, enhanced biocontrol activity may have been triggered by the EPS matrix encasing the colonies of microbial cells. Intriguingly, several reports have shown that purified EPS can strongly inhibit plant pathogens (Pandey et al., 2010; Tewari et al., 2014).

Additionally the resistance of these prospective inoculants to antibiotics was determined by conducting minimal bactericidal concentration (MBC) assays against four antibiotics produced by Streptomyces commonly found in soil (Alexander, 1977; Mah et al., 2003). All cultures showed various levels of resistance to all antibiotics. In regards to gentamycin the mono- and dual-species planktonic cultures exhibited 10 to 60 µg mL-1 resistance, in agreement with previous literature (Singh et al., 2015). Alternatively, the mono- and dual-species biofilm cultures showed resistance up to the 60 to 480 µg mL-1 level, representing a 3.33 to 48 fold enhancement over their planktonic forms. Moreover, for kanamycin and streptomycin the mono- and dual-species planktonic cultures demonstrated resistance to levels as high as 30 to 60 µg mL-1, in agreement with previous reports (Yasmin et al., 2010; Singh et al., 2015). The mono- and dual-species biofilm cultures displayed resistance up to 120 to 250 µg mL-1, representing a 2 to 8.33 fold increase. Furthermore, the Bacillus sp. planktonic culture was resistant to tetracycline at a 1 µg mL-1 concentration, while biofilms were resistant at concentrations of up to 60 µg mL-1, demonstrating a 60-fold enhancement, which is similar to previous findings (Singh et al., 2015). Potential mechanisms involved in enhanced antibiotic resistance associated with biofilms include the physical or chemical barriers provided by the EPS, slow growth, stress response, heterogeneity and biofilm-

69 specific phenotypes (Mah et al., 2001). Results therefore suggest that B-PGPR inoculants may prove more advantageous than their planktonic counterparts, since biofilms are more resistance to antibiotics, which may aid the cells in surviving and establishing themselves in the rhizosphere, a competitive environment.

To determine if biofilms are more effective PGPR than their planktonic counterparts, plant growth experiments were conducted. When the Bacillus sp., Pseudomonas sp. or mixed species planktonic or biofilm inoculants were applied as root treatments of tomato plants, there was an enhancement in various plant growth variables in comparison to the non-inoculated treatments, suggesting that these bacterial treatments are, in fact, acting to promote plant growth and that the two evaluated bacteria are PGPR. More specifically, bacterial treatments enhanced shoot DW, root DW, root length, leaf area and plant height, which is concordant with previous studies conducted with the Bacillus and Pseudomonas genera (Goteti et al., 2013; Freitas et al., 2015). Remarkably, biofilms showed superior PGP activity when compared to their planktonic equivalents in numerous cases, suggesting that biofilm forms of PGPR are in fact more effective at enhancing crop growth than their planktonic counterparts under certain conditions which are in accordance with previous reports (Mohd et al., 2014 and Jayasinghearachhi et al., 2004).

For instance, when non-inoculated plants were given Hoagland Solution (HS) they had greater plant height, shoot DW and root DW values than the non-inoculated plants given Hoagland Solution with insoluble phosphate (HS-P) or Hoagland Solution with insoluble iron and phosphate (HS-FeP), which suggests that there was a phosphorus deficiency for plant receiving the HS-P nutrient solutions. Since all HS-P and HS-FeP planktonic and biofilm bacteria-treated plants performed better than the corresponding non-inoculated plants, the results suggest that PGPR were probably able to solubilize phosphate and produce IAA, which is in agreement with the in-vitro findings. Moreover, since all HS-P and HS-FeP biofilm-treated plants (with the exception of the Bacillus sp. HS-P treatment for shoot DW) showed enhanced growth in comparison to their planktonic counterparts, results which indicated that biofilm forms of PGPR are greater phosphate solubilisers and IAA producers than planktonic forms, and which further validated the previous in-vitro findings of this work. For the Bacillus sp. HS-P biofilm treatment, it is possible that differences may have been significant if plants had been grown for a longer period of time.

70

When non-inoculated plants given HS where compared to non-inoculated plants given HS- FeP it becomes apparent that nutrient deficiencies for phosphorus and iron, collectively, affected root length. Since all HS-FeP planktonic and biofilm bacterial treatments resulted in enhanced growth, when compared to their corresponding non-inoculated plants, the results suggested that PGPR mechanisms most likely involved phosphate solubilisation in addition to siderophore and IAA production, which is in agreement with the in-vitro findings. Additionally, since all HS-FeP biofilm treated plants showed increased root lengths, in comparison to their planktonic counterparts, my results indicated that B-PGPR are better phosphate solubilisers, IAA producers and siderophore producers, again, reinforcing the in-vitro findings.

When non-inoculated plants given HS nutrient solution where compared to non-inoculated plants given HS-FeP, HS-P or Hoagland Solution with insoluble iron (HS-Fe), it was apparent that phosphorus and iron deficiency were affecting leaf area development. All analogous planktonic and biofilm bacterial treatments increased values of the measured growth variables when compared to the corresponding non-inoculated plants, indicating that potential PGPR mechanisms include phosphate solubilisation as well as siderophore production, again, in agreement with the in-vitro findings. Furthermore, since all equivalent biofilm treated plants showed increased root length in comparison to their planktonic-cell treated counterparts (with the exception of the Bacillus sp. HS-Fe treatment), the results indicated that B-PGPR are more adequate phosphate solubilisers and siderophore producers than planktonic cells which further reinforces the in-vitro findings. Overall, phosphorus affected numerous plant growth variables, which was anticipated since this macronutrient plays a crucial role in plant growth. Overall, no iron deficiencies alone affected plant growth variables, which may be because iron is a micronutrient or simply because the plant did not require a substantial amount of this nutrient during the first 6 weeks of growth.

Finally, the stereo microscopy (SM) imaging made it clear that planktonic inoculants produced biofilms on the tomato plant root systems, which was expected since this system is static. However, it is important to note that planktonic inoculants did not produce robust biofilms, particularly in comparison to the biofilm inoculants, and this was probably caused by bacterial

71 cells in the biofilms derived from planktonic cells detaching from the biofilm to form new biofilms in different locations of the root system.

Future work should focus on determining the effects of B-PGPR when plants reach the fruiting stage, in order to fully grasp the final crop-productivity outcome of this potentially crucial technology. In addition it would be vital to determine if B-PGPR inoculants show promising results when applied under field conditions. Further, it would be of interest to identify the genes associated with these PGP attributes and to perform q-PCR to analyze the gene expression of B-PGPR and P-PGPR, in order to further understand the molecular mechanisms responsible for the growth enhancements described above. It would also be fascinating to test the biocontrol attributes of PGPR previously described for crops under both greenhouse and growth chamber conditions, in order to determine if B-PGPR are, in fact, more effective at biocontrol than their planktonic counterparts. Finally, it would be extremely interesting to determine if B-PGPR are more tolerant to abiotic stresses (such as salt, heavy metal and extreme pH) than P-PGPR.

72

Chapter 6

CONCLUSION

Overall I have demonstrated that PGPR strains isolated from the field, identified as a Bacillus sp. and Pseudomonas sp., via molecular techniques, can form mono- and dual-species biofilms. These findings are in agreement with preceding reports, which also successfully developed mono- and dual-species biofilms with alternative microbial partners.

Additionally, we demonstrated that biofilm forms of PGPR are more effective than their planktonic counterparts at manifesting potential plant growth promoting attributes under in vitro conditions, in agreement with previous findings. More specifically, I was able to show that mono- and dual-species biofilms are notably better at solubilising phosphate, producing siderophores and synthesizing IAA than their planktonic equivalents. Furthermore, I have established that biofilms showed enhanced antagonistic activity against plant pathogenic bacteria and fungi in addition to increased ammonia and hydrogen cyanide production.

Finally, I validated that biofilms are more resistant to antibiotics commonly found in soil than to their planktonic counterparts, which corroborates current literature. Thus, if appropriate formulation methods can be developed, B-PGPR could offer an important improvement over P- PGPR inoculants, which have often been inconsistent delivery systems, perhaps due to inability of the cells to compete with existing microbial communities. This remains to be tested under controlled environment and, most importantly, field conditions.

Therefore, the use of innovative B-PGPR technologies could prove extremely advantageous as biocontrol and biofertilizer agents, while alleviating dependence on chemical inputs such as fertilizers and pesticides.

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Chapter 7

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