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The effect of other rhizosphere microorganisms on the ability of Paenibacillus spp. to promote the growth… Bent, Elizabeth 2000

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THE EFFECT OF OTHER RHIZOSPHERE MICROORGANISMS ON THE ABILITY OF PAENIBACILLUS SPP. TO PROMOTE THE GROWTH OF LODGEPOLE PINE [PINUS CONTORTA VAR. LATIFOLIA (DOUGL. ENGELM.)] By ELIZABETH BENT B.Sc., University of Guelph, 1995 A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE STUDIES (Department of Soil Science) We accept this thesis as conforming to the required standard THE UNIVERSITY OF BRITISH COLUMBIA November, 2000 © Elizabeth Bent, 2000 I n p r e s e n t i n g t h i s t h e s i s i n p a r t i a l f u l f i l m e n t o f t h e r e q u i r e m e n t s f o r an advanced degree a t t h e U n i v e r s i t y o f B r i t i s h C o l u m b i a , I agree t h a t t h e L i b r a r y s h a l l make i t f r e e l y a v a i l a b l e f o r r e f e r e n c e and s t u d y . I f u r t h e r agree t h a t p e r m i s s i o n f o r e x t e n s i v e c o p y i n g o f t h i s t h e s i s f o r s c h o l a r l y p u r p o s e s may be g r a n t e d by t h e head o f my department or by h i s or h e r r e p r e s e n t a t i v e s . I t i s u n d e r s t o o d t h a t c o p y i n g o r p u b l i c a t i o n o f t h i s t h e s i s f o r f i n a n c i a l g a i n s h a l l not be a l l o w e d w i t h o u t my w r i t t e n p e r m i s s i o n . Department o f <^>[U i S G l ^ U C E r . The U n i v e r s i t y o f B r i t i s h C olumbia Vancouver, Canada Date M O v F A A f e f e K . 2-If . Q-CXDQ Abstract II The interaction between lodgepole pine and plant growth-promoting rhizobacteria (PGPR) strains Paenibacillus polymyxa L6 and Pw-2 was examined. Experiments were conducted to determine whether the extent of PGPR-mediated growth promotion seen under gnotobiotic conditions is influenced by the presence of co-inoculated rhizobacteria (Curtobacterium flaccumfaciens PF322 or Pseudomonas fluorescens M20) or a co-inoculated ectomycorrhizal fungus (Rhizopogon subcularescens Smith 11616), whether a change in the extent of PGPR-mediated plant growth promotion is related to (1) a quantitative change in the extent of PGPR population density on or within the root, (2) a qualitative change in PGPR colonization of the root surface (determined using immunofluorescence techniques and confocal laser scanning microscopy), (3) a change in levels of auxins and/or cytokinins within root tissue, and finally whether P. polymyxa L6 and Pw-2 vary in their ability to form spores in the rhizosphere. It was concluded that the extent of P. po/ymyxa-mediated growth promotion of lodgepole pine can be reduced by the presence of a single rhizobacterial co-inoculant. The decreases in growth promotion were not related to reductions in rhizospheric population densities of PGPR, and were not linked to qualitative changes in the ability of PGPR to colonize specific root surface microsites. Rhizobacteria had strain-specific effects on root hormone levels, and the possibility that decreases in growth promotion are related to the effects of combinations of bacteria on root hormones should be investigated. Pw-2 was found to produce significantly more spores than L6 under the experimental conditions, and the possiblity that Pw-2, but not L6, was identifed as an endophyte on chemically surface-sterilized root tissues due to this fact should be considered. Initial attempts to identify Pw-2 colonization sites in the root interior via confocal microscopy were not successful, and further attempts should be made to verify the endophytic status of Pw-2. Co-inoculating seedlings with P. polymyxa and R. subcularescens did not result in additive growth effects on pines. However, further work is needed to determine if additive growth improvements from P. polymyxa and ectomycorrhizal fungus co-inoculation will occur under different growing conditions, and/or in the presence of different ectomycorrhizal fungi. Ill Table of Contents List of Figures v List of Tables vii Chapter 1 Introduction 1.1 Introduction 1 1.2 PGPR and other plant-interacting rhizosphere microorganisms 3 1.3 What may inhibit PGPR-mediated plant growth promotion 4 1.4 The rhizosphere microhabitat 6 1.5 Root colonization and its importance in PGPR-mediated growth promotion 8 1.6 The effect of mycorrhizae on PGPR-plant interactions 10 1.7 Phytohormones and plant growth 10 1.8 Lodgepole pine and Paenibacillus polymyxa 11 1.9 Hypotheses 12 Chapter 2 Co-inoculation effects of Curtobacterium flaccumfaciens PF322 with Paenibacillus PGPR on extent of pine growth promotion and PGPR root colonization 2.1 Introduction 13 2.2 Materials and Methods 13 2.3 Results 18 2.4 Discussion 25 Chapter 3 Co-inoculation effects of Pseudomonas fluorescens M20 with Paenibacillus PGPR on extent of pine growth promotion, PGPR root colonization, and levels of root phytohormones 3.1 Introduction 28 3.2 Materials and Methods 29 3.3. Results 36 3.4 Discussion 42 Chapter 4 Spatial colonization of the root surface by Pseudomonas fluorescens M20 and Paenibacillus PGPR, applied as single- or co-inoculated treatments 4.1 Introduction 46 4.2 Materials and Methods 47 4.3 Results 56 4.4. Discussion 83 Chapter 5 Spore formation in the rhizosphere by Paenibacillus PGPR, and implications for identification of sporeforming bacteria as endophytes 5.1 Introduction 86 5.2 Materials and Methods 87 5.3 Results 88 5.4 Discussion 94 iv Chapter 6 Co-inoculation effects of a mycorrhizal fungus, Rhizopogon subcularescens Smith 11616, with Paenibacillus PGPR on extent of pine growth promotion and PGPR root colonization 6.1 Introduction 97 6.2 Materials and Methods 98 6.3 Results 102 6.4 Discussion 107 Chapter 7 Conclusions and recommendations for future work 111 Literature Cited 114 V List of Figures Chapter 2 Figure 1: Average rhizosphere population sizes of strains when inoculated singly and coinoculated. Page 19. Figure 2: Average root interior population sizes of strains when inoculated singly and coinoculated. Page 21. Figure 3: Average number of lateral roots per seedling for each treatment after 6, 9 and 12 weeks of incubation. Page 22. Figure 4: Average lengths of (a) primary and (b) combined lateral roots for each treatment after 6, 9 and 12 weeks of incubation. Page 23. Figure 5: Average (a) shoot and (b) root biomass accumulation for each treatment after 6, 9 and 12 weeks of incubation. Page 24. Chapter 3 Figure 6: Average rhizosphere population sizes of strains when inoculated singly and co-inoculated. Page 39. Figure 7: Average (a) root dry weight, (b) shoot dry weight, (c) root length and (d) number of lateral roots measured for seedlings in each treatment after 6, 9 and 12 weeks of incubation. Page 40. Chapter 4 Figure 8: Illustration of the process by which three two-dimensional images, each representing emitted light of different wavelengths, were combined into a single image and interpreted. Page 56. Figure 9: Staining controls, representative of controls viewed in each experiment (only one set of images is depicted to conserve space), from randomly sampled root segment surfaces, thirteen weeks after inoculation. Pages 61-64. Figure 10: Rhizobacterial colonization of randomly sampled root segment surfaces, thirteen weeks after inoculation. Pages 65-68. Figure 11: Rhizobacterial colonization of randomly sampled root segment surfaces, seven weeks after inoculation. Pages 69-73. Figure 12: Surfaces of root segments sampled thirteen weeks after inoculation, in six different locations: the root tip (1), the segment immediately after the root tip (2), from the middle of the root (3 and 4, with 3 being closer to the tip), to the base of the root (5 and 6, with 5 being closer to the root tip. Pages 74-80. Figure 13: Thermal print depicting a magnified, on-screen green channel image of bacterial cells on the root surface of a pine seedling inoculated with P. polymyxa strain Pw-2, next to a 2 urn scale bar. Page 78. vi Chapter 5 Figure 14: Recovery of Paenibacillus polymyxa strains L6 and Pw-2 from washes of lodgepole pine seedling roots before and after heat treatment (55°C, 30 min). Page 89. Figure 15: Representative cross-sections of lodgepole pine roots, examined thirteen weeks after inoculation. Pages 90-93. Chapter 6 Figure 16: Average (a) root dry weight, (b) shoot dry weight, (c) root length and (d) number of lateral roots measured for seedlings in each treatment after 9 and 12 weeks of incubation. Page 104. Figure 17: Percent of seedlings having bifurcated root tips in each treatment (data for week 9 and 12 observations were pooled). Page 105. Figure 18: Average rhizosphere population sizes of Paenibacillus polymyxa strains (a) L6 and (b) Pw-2 when inoculated singly and co-inoculated with Rhizopogon subcularescens Smith 11616. Page 106. Vll List of Tables Chapter 1 Table 1: Bacterial biocontrol products commercially available as of January 1, 1999. Page 2. Chapter 2 Table 2: Distinguishing characteristics used to differentiate Paenibacillus polymyxa L6, P. polymyxa Pw-2 and Curtobacterium flaccumfaciens PF322. Page 16. Table 3: Effects of PGPR (Paenibacillus polymyxa L6 and Pw-2) and six different putative DRMOs on lodgepole pine growth in screening tests, six weeks after inoculation. Page 18. Chapter 3 Table 4: Distinguishing characteristics used to differentiate Paenibacillus polymyxa L6, P. polymyxa Pw-2 and Pseudomonas fluorescens M20. Page 33. Table 5: Effects of nine different Gram negative soil isolates on lodgepole pine growth in screening tests, six, nine and twelve weeks after inoculation. Page 38. Table 6: Levels of indole-3-acetic acid (IAA) and dihydrozeatin riboside (DHZR) in lodgepole pine root tissue. Page 41. Chapter 4 Table 7: Binding of anti-M20 antiserum (A405) before and after purification by preabsorption and IgG collection, to cells of each bacterial strain (P. fluorescens M20 and B. polymyxa L6 and Pw-2). Page 59. Table 8: Binding of anti-L6 antiserum (A405) before and after purification by IgG collection, to cells of each bacterial strain (P. fluorescens M20 and B. polymyxa L6 and Pw-2). Page 60. Table 9: Binding of anti-Pw-2 antiserum (A405) before and after purification by IgG collection, to cells of each bacterial strain (P. fluorescens M20 and B. polymyxa L6 and Pw-2). Page 60. Chapter 6 Table 10: Effect of Rhizopogon subcularescens 11616 and growth media on growth of bacterial strains in vitro. Page 103. Table 11: Growth of bacteria, sampled from areas within and outside of zones of inhibition, on different media. Page 107. Chapter 1: Introduction 1 1.1 Introduction Plant-growth promoting rhizobacteria (PGPR) are a heterogenous group of soil bacteria that can be found in the rhizosphere, which improve the extent or quality of plant growth or health (Kloepper and Schroth 1978). Most PGPR are free-living, or associative, meaning they do not form symbiotic structures such as nodules with host plants. Throughout this document, to simplify discussion, the term "PGPR" will refer only to these associative organisms, and symbiotes such as rhizobia will be discussed separately. The rhizosphere is usually defined as "the region of soil exterior to the root which is subject to the influence of the root", meaning the region of soil into which root-produced soluble and volatile substances that can affect microbial activity diffuse (Curl and Truelove 1986). PGPR hold great promise as potential agricultural and forestry inoculants, and may even have bioremediation applications. The widespread use of effective PGPR-based, growth-enhancing bioinoculants could reduce or eliminate the use of toxic or environmentally damaging chemical fertilizers and pesticides. Biologically-mediated means of plant pathogen control, such as biological control of crown gall (Agrobacterium tumefaciens), can sometimes be more effective than chemical treatments (Cook 1993). Forest tree PGPR inoculants may enhance the growth and survival of young seedlings in nurseries, or older seedlings in stressful site environments, increasing the effectiveness of forest replanting operations. For example, avoidance of drought stress by faster establishment of an extensive root system in PGPR-inoculated seedlings may be possible. Cold-tolerant PGPR expressing antifreeze proteins may benefit plants growing in colder climates (Sun et al. 1995). Alternatively, rhizobacteria able to degrade chlorinated organic compounds or other toxins may assist the growth of plants at polluted sites (Yee et al. 1998), including heavy-metal hyperaccumulating plants used in phytoremediation (Brown 1995). Unfortunately, despite promising laboratory and greenhouse results, the results of field trials involving PGPR inoculants are often too variable for successful commercial application (e.g. Chanway and Holl 1993, de Freitas and Germida 1992, Frommel et al. 1993, Kloepper et al. 1980, Zhender et al. 1999). In order for PGPR to be useful as plant growth-stimulating inoculants, they must demonstrate a consistent benefit to the inoculated plants, preferably under a wide range of growing conditions. PGPR have been studied for forty years or more (Brown et al. 1964, Brown 1974, Burr et al. 1978, Cooper 1959, Rovira 1963) yet relatively few of these organisms have been developed into useful commercial products, in part because it is difficult to guarantee that the product will always have a desirable effect on plant growth (Kempf 2 etal. 1997, Kenney 1997). Currently, there are about 18 different biological control products incorporating one of seven different bacterial species commercially available in the United States (Table 1). Products incorporating plant-growth promoting microbes with no direct antimicrobial or suppressive activity against pathogens are even rarer, and may consist of a compost-like "microbial solution", containing a variety of soil bacteria and actinomycetes, and blended with soil amendments such as humus or micronutrients (i.e. SuperBio®, Advanced Microbial Solutions, Pilot Point, TX USA 76258). Table 1: Bacterial biocontrol products commercially available as of January 1,1999 (information provided by the USDA Biocontrol of Plant Diseases Laboratory) Biocontrol Agent Product Name Manufacturer Agrobacterium radiobacter Galltrol-A AgBioChem Inc., Orinda CA USA 94563 Nogall, Diegali Bio-Care Technology Pty. Ltd., Somsersby, NSW 2250, Australia Norbac 84C New BioProducts, Inc., Corvallis, OR USA 97330 Bacillus subtilis Epic, Kodiak Gustafson, Inc., Dallas TX USA 75266 Rhizo-Plus KFZB Biotechnik GmbH, D-12489 Berlin, Germany Serenade AgraQuest, Inc., Davis CA USA 95615 System 3 Helena Chemical Co., Memphis, TN USA 38119 Burkholderia (Pseudomonas) cepacia Intercept Soil Technologies Corp., Fairfield IA USA 52556 Deny (aka Blue Circle, Precept) Stine Microbial Products, Shawnee, KS USA 66216 Pseudomonas fluorescens BlightBan A506 Plant Health Technologies, Fresno CA USA 93704 Conquer, Victus Sylvan Spawn Laboratory, Kittanning PA USA 16201 Pseudomonas solenacearum (avirulent) PSSOL Natural Plant Protection, 64150 Nogueres, France Pseudomonas syringae Bio-save 100, 110 Eco Science Corp., Orlando FL USA 32802-3228 Streptomyces griseoviridis Mycostop Kemira Agro Oy, 00101 Helsinki, Finland 3 1.2 PGPR and other plant-interacting rhizosphere microorganisms There are numerous environmental factors which can influence plant and microbial metabolism, and therefore the ability of these organisms to interact with one another. PGPR are thought to promote plant growth via a variety of mechanisms, any or all of which might be influential in a given plant-PGPR interaction. Some PGPR aid in the formation of mycorrhizae (Garbaye 1994) or rhizobial nodules (Srinivasan et al. 1997), while others may promote plant growth more directly, by producing phytohormones, additional nutrients, or other growth-stimulating substances to the plant (Glick 1995). PGPR may also influence systemic plant defense mechanisms in an as-yet undetermined manner, resulting in more effective plant defense responses against plant pathogens (Ku6 1990, Tuzun and Bent 1999) or even herbivorous pests (Zhender ef al. 1997,1999). PGPR, notably pseudomonads, may also act as biocontrol agents, suppressing the growth of plant-pathogenic microorganisms by producing siderophores or antimicrobial substances in the rhizosphere (O'Sullivan and O'Gara 1992, Weller 1988) or within the plant root (Pan etal. 1997). It is speculated that plants may cultivate the growth of disease-suppressive bacteria on their root systems, which then protect the plant from soilbome pathogens, via root exudates (Cook ef al. 1995, Ryan and Jagendorf 1995). Other rhizosphere microorganisms include free-living fungi or bacteria which can have a negative effect on plant growth, known collectively as deleterious rhizosphere microorganisms (DRMO) or in the case of bacteria, deleterious rhizobacteria (DRB) (Nehl etal. 1996, Schippers ef al. 1987, Suslow and Schroth 1982). Instead of providing the plant with nutrients, DRMO may compete with the plant or with other, more beneficial, rhizosphere organisms for them, particularly scarce trace nutrients (Nehl et al. 1996, Schippers ef al. 1987). DRMO may prevent plants from absorbing nutrients by destroying phytosiderophores (van Wiren etal. 1995), or may produce metabolites that damage the plant or inhibit plant germination or growth, including volatile substances (Astrom 1991, Nehl ef al. 1996), toxins (Kennedy etal. 1991), ice-nucleating proteins (Ramstedt etal. 1994) or cyanide (Astrom 1991, Alstrom and Bums 1989, Gutierrez-Manero etal. 1996). DRMO may increase plant susceptibility to disease (Alstrom 1991) or produce excessive levels of auxins or other phytohormones that curb rather than enhance seedling growth and development (Barazani and Friedman 1999, Nehl etal. 1996, Schippers etal. 1987). DRMO have been called "minor pathogens" (Suslow and Schroth 1982), but it is not clear where the lines between virulent plant pathogens, DRMO and PGPR lie. An organism which promotes the growth of one plant species or variety may inhibit the growth of others (Yuen and Schroth 1986). Similarly, an organism which promotes plant growth under one set of environmental or experimental conditions may have no effect, or a deleterious effect, under others (Chanway and Holl 1992,1993, Chiarini et al. 1998, de Frietas and Germida 1992, Germida and Walley 1996, Frey-Klett etal. 1999, Nehl etal. 1996, Zhang et al. 1997). How can this be explained? Plant-beneficial symbionts, plant-beneficial associative microbes such as PGPR, plant-deleterious associative microbes (DRMO) and virulent pathogens may exist as part of a continuum, with no absolute boundaries between any of these categories. Plants inoculated with plant pathogens have demonstrated both increased growth and elevated plant defense responses, leading to a reduction in the incidence of plant disease (Tuzun et al. 1986,1992). These same phenomena have been recorded in a variety of plants inoculated with associative PGPR (e.g. Maurhoferef al. 1994, Wei etal. 1996, van Wees etal. 1997), and it has been speculated that free-living PGPR which induce plant disease resistance may also have evolved from plant pathogenic organisms (Tuzun and Bent 1999). A PGPR which stimulated Douglas-fir growth was identified as P. syringae, normally considered a plant pathogen (Chanway and Holl 1992). James etal. (1997) speculate that weakly pathogenic, nitrogen-fixing Herbaspirillum species in sugarcane might be in the process of evolving into endophytic symbiotes. Mycorrhizal fungi (Malloch 1987) and rhizobia (Spaink 1995) may have evolved from phytopathogens, and it is possible for a fungal plant pathogen to mutate into a nonpathogenic, endophytic mutualist (Freeman and Rodriguez 1993). Mycorrhizal fungi, generally thought of as plant-beneficial organisms, can inhibit plant growth (Francis and Read 1995, Sylvia and Sinclair 1983a) or even become parasitic (Aldea 1998, Beyrie ef al. 1995). 1.3 What may inhibit PGPR-mediated plant growth promotion What might prevent a PGPR from exerting a plant-beneficial effect? In order for this effect to occur, the following events must take place. Each of these events may be affected by a variety of interrelated environmental factors, and some examples are given. The events are: 1. The signal(s) which stimulate growth in the plant must be synthesized by the PGPR in sufficient quantity to exert an effect. The signal(s) may be attached to the surface of the microbe, or they may be diffusible substances produced constrtutively, or in response to a stimulus. For example, diffusible "quorum-sensing" signals, such as acetylated homoserine lactones, are produced in response to increased population density (Shapiro 1998). The synthesis of microbial signals may be affected by any factors which affect microbial growth and activity, including the presence of antagonists, the availability and quality of nutrients, soil pH, soil texture and structure, water content, redox potential, temperature and salinity (Sorensen 1997, Stotzky 1997). The availability and quality of nutrients will be influenced by the presence of other organisms in the soil which may compete for and sequester these nutrients and excrete other metabolites (Sorensen 1997), by the cation exchange capacity (CEC) of the soil, 5 which is in turn affected by the composition and quantity of clays and organic matter present (Stotzky 1997), and by the quantity and composition of photosynthate released by growing plant roots (Marschner 1991, Sorensen 1997). 2. The signal(s), if diffusible, must travel from the surface of the microbe via surface water films to plant root tissue capable of detecting the signal(s). The ability of a diffusible signal to travel from a microbe to a receptive plant cell will depend upon the distance between these two, the presence of water films able to carry the signal this distance, and the abundance of other soil organisms, exoenzymes or soil surfaces (such as clays or organic matter) that might assimilate, modify, destroy or immobilize the signal and so prevent it from reaching the root (Stotzky 1997). 3. The plant root must be able to perceive and respond to the signal(s), and the plant must be in a physiological state such that the response can result in detectable growth promotion. The ability of the plant to perceive and respond to a microbial signal will depend upon whether the plant receptors required for its detection are active, and whether the plant can react in a manner that will produce a detectable growth promotion response. It has been noted that PGPR sometimes exert a larger or more consistent growth promotion effect on stressed plants than on unstressed ones (Chanway and Holl 1993, Iswandi et al. 1987, Seong etal. 1991). Depending on the basis of the plant-microbe interaction, stressed plants might be more able to respond to microorganisms than unstressed ones. For example, temperature and nitrogen status were found to affect rhizobial nodule formation in legumes (Abaidoo et a/. 1990), and in plant-mycorrhizal fungus interactions, it has been demonstrated that phosphate-stressed plants are more receptive to infection by arbuscular mycorrhizal fungi, due to a reduction in the expression of defense-related plant genes that can inhibit mycorrhization (Lambais and Mehdy 1993,1995). Under normal conditions, a PGPR-inoculated seedling is exposed to numerous other soil organisms, which may themselves influence the growth of the plant. Different soils are likely to support different indigenous microbial communities (Latour et al. 1996, McArthur et al. 1988, Timonen etal. 1998). Indeed, disease-suppressive (Cook etal. 1995, Schroth and Hancock 1982, Yuen etal. 1985) and disease-prone (Aldea 1998, Alstrom 1992) soils have been described, and the ability of these to enhance or suppress disease were in each case linked to the composition of the soil microflora. PGPR-inoculated spruce were observed to grow more poorly in soil which yielded a greater number of spruce DRB (O'Neill etal. 1992). The release of endophytic DRB from ploughed red clover is thought to alter the composition of the soil microflora and cause 'clover-maize' syndrome, where soils previously cropped with clover inhibit the growth of subsequently planted maize (Sturz and Christie 1996), and whether an inoculated 6 bacterial strain acted as a PGPR or a DRMO on potatoes was found to depend upon its exposure to differently-cropped soils (Sturz and Christie 1995). Could the composition of the indigenous microflora in different soils be a major factor influencing the variability of PGPR-mediated growth promotion under field conditions? Perhaps the growth-impairing effects of a DRMO, for example, can partially obscure, or even completely mask the growth-promoting effects of PGPR by reducing the observed extent of growth promotion (Schippers etal. 1987). If this were so, PGPR-inoculated plants might grow more poorly at a site which supports a greater number of, or more effective, DRMO than at a similar site in which these deleterious organisms were not present. Given that PGPR and DRMO activities seem to shift with environmental or climactic conditions, this question can only be studied under controlled conditions, where most environmental variables that might affect microbial or plant metabolism are kept constant. 1.4 The rhizosphere microhabitat The physical size of the rhizosphere is dependent on the size of the root and the amount of material released from it, as well as the physical characteristics of the soil (water content, texture, porosity, etc.) which influence the rate of movement of gases and solutes through it (Curl and Truelove 1986, Sorensen 1997). It should also be noted that plant rhizospheres will overlap, particularly in natural situations (Westover et al. 1997). The increase in microbial populations and activity in the rhizosphere relative to the bulk soil, or edaphosphere, (a 10- to 10,000-fold increase) is known as the "rhizosphere effect" (Curl and Truelove 1986), and first led researchers to conclude that nutrients are abundant in the rhizosphere. It has since been demonstrated that the rhizosphere can be considered a more oligotrophic environment, where microorganisms exhibit symptoms of starvation, such as the formation of cells which are smaller than normal (Marschner and Crowley 1996) or viable but non-culturable (VBNC) cells (Hase et al. 1999). It has long been known that bacteria were present in what appeared to be a mucilaginous biofilm on the surface of roots (Foster 1986); more recently, the formation of heterogenous and randomly-spaced clusters, or microcolonies, of bacteria on the root surface or in cracks between epithelial plant cells has been described (Chin-A-Woeng etal. 1997, Dandurand etal. 1997, Hansen etal. 1997). The ecology of the rhizosphere ultimately depends upon the plant, which is the primary carbon source for all the various creatures inhabiting the areas around its roots. It has been well established that there are variations in the microbial communities supported by different plant species (Kloepper etal. 1991a, Latour etal. 1996, Orazova etal. 1999), combinations of plant species (Westover etal. 1997), and plant cultivars (Cattelan etal. 1998, Siciliano etal. 7 1998). Plants release nutrients in a variety of ways: via root exudation, leakage of cytoplasmic solutes from wounds or sites of lateral root emergence, and the sloughing of dead cells from the root surface (Curl and Truelove 1986, Marschner 1991, Sorensen 1997). Changes in the metabolism and health of the plant will be reflected in the quantity and quality of nutrients or defensive compounds released by the plant into the rhizosphere. Root exudation involves the release of low-molecular weight compounds from the root tip and cap (Hawes etal. 1998), the zone of elongation (Sorensen 1997) and root hairs (Hofer 1996), possibly to regulate the influx of water into cells by reducing the osmotic potential between the cell's cytoplasm and its exterior (Schwenke and Wagner 1992). Compounds may also leak from the root as a result of damage to root cells, due to wounding or the emergence of lateral roots (Charlton 1996, Hale etal. 1971); these latter can generally not be distinguished experimentally from root exudates and so are usually considered part of root exudate. The quantity or composition of root exudates has been found to vary between plant species (Guckert et al. 1991) and stages of development (Smith 1970), or when plants are exposed to various stimuli, including bacteria (Dakora etal. 1993, Prikryl and Vancura 1980), mechanical impedance (Boeuf-Tremblay etal. 1995), wounding (Hale etal. 1971), water stress (Reid and Mexal 1977), nutrient limitations (Hoffland etal. 1992, Marschener 1998) or metals (Basu etal. 1994, Zhang et al. 1991). As much as 10-40% of the photosynthate fixed by the plant is estimated be released into the rhizosphere as exudate (Cheng etal. 1994). Root exudate can contain many microbial nutrient sources, including sugars, organic acids, amino acids and phenolics (Marschner 1991, Vancura 1988), as well as chemoattractants such as nod factors (Hungria etal. 1991), phytosiderophores (van Wiren etal. 1995), or factors which stimulate fungal (Zhou and Paulitz 1993) or plant germination (Weerasuriya et al. 1993). Root exudate-inducible genes in Pseudomonas spp. have been identified that respond to amino acids or sugars found in exudate (Bayliss etal. 1997, van Overbeek and van Elsas, 1995). Antimicrobial substances, such as phytoalexins, hydrocyanic acid and antifungal metabolites or inhibitors, may also be present in root exudate (Bouillant etal. 1994, Fontenla etal. 1999, Stevenson etal. 1995, Vancura 1988). The continual sloughing and replacement of root cap cells may also affect the growth of rhizosphere microorganisms. Dead or dying root cap cells act as a food source, while living cells released from the root cap (known as root border cells) may release metabolites which attract microorganisms, and promote or inhibit the growth of different bacteria (Hawes et al. 1998). Root senescence and turnover can also contribute a great deal of organic matter to the soil (Bloomfield etal. 1996). Mucilaginous material, consisting mainly of polysaccharides and polyuronic acids, is produced both by soil bacteria and by the plant root cap and root hair cells 8 (Guckert etal. 1991, Marschner 1991, Watt etal. 1993), and is collectively known as 'mucigel' (Curl and Truelove 1986). These polysaccharides may lubricate younger plant tissues, protecting them from abrasion as the root tip pushes through the soil (Hawes et al. 1998), and may also enhance the rhizosphere effect: mucigel, in addition to serving as a carbon source for microorganisms, can adsorb water and soluble nutrients, keeping a ready supply of these near the root surface (Watt et al. 1993). The presence of mucigel has been shown to influence the composition of bacterial populations associated with different root regions in corn (Gochnauer et al. 1989). Mucigel may also allow bacteria to adhere to the root surface, preventing bacteria from being washed away, and perhaps assisting in the formation of microcolonies (Foster 1986, Pereg-Gerk et al. 1998). Herbivorous fauna (both above and below ground) and microfauna (such as protists, nematodes, and arthropods) can affect the composition of the rhizosphere community indirectly, by grazing upon the plant and perhaps altering the quantity or composition of plant root exudate (Bardgett etal. 1998, Denton etal. 1999). Fauna may also graze upon microorganisms directly (Kandeler et al. 1999, Lawrence and Snyder 1998). Microorganisms that are not able to grow solely upon the nutrients provided by a plant may still be able to grow in its rhizosphere, utilizing the metabolites of other microbes or preying upon them, or upon microfaunal inhabitants (Curl and Truelove 1986, Kloepper et al. 1991a, Sayre 1980). 1.5 Root colonization and its importance in PGPR-mediated growth promotion Rhizosphere competence, or the ability to colonize and persist in the rhizosphere, seems to be required for biological control of root diseases using PGPR. Inadequate root colonization is thought to explain the lack of consistent results in these systems (Weller 1988), although superior rhizosphere-colonizing ability by some bacteria was not associated with disease suppression or enhanced growth or yield of wheat (Milus and Rothrock 1993) or tomato (Pillay and Nowak 1997). Optimal disease suppression by Pseudomonas PGPR on radish may require a threshold density of PGPR in the rhizosphere; once the threshold is reached, continuing to increase the density of PGPR results in little improvement (Raaijmakers et al. 1995). More recently, studies of endophytic PGPR with biological control activity have suggested that the extent of PGPR colonization of the root interior, rather than the rhizosphere, is important in disease control (Duijff et al. 1997, Pan et al. 1997, Pillay and Nowak 1997, Troxleref al. 1997). Colonization of the rhizosphere or the root interior is considered to be essential for PGPR in general, whether they function indirectly as pathogen-attacking biological control agents or via some other mechanism(s) (Kloepper and Beauchamp 1992, Loper et al. 1984). 9 Root colonization by PGPR which do not directly attack pathogens can be positively correlated with plant growth (Frommel et al. 1993), although sometimes low populations of these PGPR are sufficient to generate a significant effect (Chiarini etal. 1998, Halverson etal. 1993), and large populations may even reduce the beneficial effect of some PGPR (Frey-Klett et al. 1999). Kloepper et al. (1991b) define PGPR as "efficient microbial competitors in the root zone which displace native root-colonizing microorganisms", and while this is true in some cases (Kloepper and Schroth 1978), it is perhaps simplistic to assume all PGPR will share this ability-particularly since most PGPR are identified by screening for an ability to promote plant growth, and the ability to displace native microflora often remains unexamined. Still, in most cases, PGPR inoculants are added with plants to soils already containing well-established microflora, and it is reasonable to assume that in order to exert an effect on the plant, these PGPR must establish themselves to some degree in or around the plant root. Rhizobacteria have been shown to vary in their ability to displace other bacteria from the rhizosphere (Hartel etal. 1993, Ikeda etal. 1998, Suslow and Schroth 1982), with introduced PGPR being at a disadvantage in some systems (Ikeda etal. 1998), but not in others (Milus and Rothrock 1993, Suslow and Schroth 1982). Genetically modified bacteria may also experience greater metabolic loads than wild-type organisms, and so be less able to colonize roots or persist in soil (Evguenieva-Hackenberg etal. 1994, Hartel etal. 1993, Tang etal. 1995, van Elsas et al. 1994), although this is not always the case (Hong et al. 1995). The majority of root colonization work in the literature has focused on bacterial population densities, but it is also possible that PGPR may not function unless they are able to colonize specific microsites on the root surface. A greater quantity of nutrients may be found in some regions, released as dead cells or root exudates by the root tip and the regions immediately following it, or leaked from the sites of lateral root emergence (Charlton 1996, Hawes ef al. 1998, Hofer 1996, Sorensen 1997). It may be that these are the only sites on the root where there is adequate nutrition for the synthesis and accumulation of PGPR growth promoting signal(s) (Chin-A-Woeng ef al. 1997, Dandurand etal. 1997). The ability of Azospirillum PGPR to flocculate, adhering to each other and to root surfaces, has been linked to the abilities to colonize the root surface and to promote plant growth (Pereg-Gerk ef al. 1998), suggesting that the formation of bacterial clusters or microcolonies on the root surface might be important to plant growth promotion by these bacteria. Alternatively, specific root tissues may be involved in anchoring PGPR, or in the perception of PGPR growth promoting signal(s). The ability of Pseudomonas PGPR to adhere to root surfaces has been linked to the presence of agglutinin, a root surface glycoprotein (Anderson ef al. 1988), although it is not known whether the expression of agglutinins is localized to specific root tissues. 10 1.6 The effect of mycorrhizae on PGPR-plant interactions Many plants form associations with mycorrhizal fungi under natural conditions (Wilcox 1996). Mycorrhizal fungi have been demonstrated to directly affect plant growth by increasing the absorptive surface area of roots and mobilizing or capturing sparingly available nutrient sources via the production of chelators or extracellular enzymes, thereby providing the plant with increased access to mineral nutrients (Leyval and Berthelin 1991, Paris et al. 1995, Marschner and Dell 1994) as well as fixed carbon or nitrogen, transferred via mycelial links from other plants (Arnebrant et al. 1993, Griffiths ef al. 1995). Mycorrhizal fungi may also protect plants from pathogens (Hwang etal. 1995, Strobel and Sinclair 1991, Sylvia and Sinclair 1983a,b), aluminum stress (Schierand McQuattie 1995), or attract plant-growth promoting microfauna (Jentschke etal. 1995). Auxins produced by ectomycorrhizalfungi have been demonstrated to stimulate rooting in cuttings (Normand et al. 1996), to promote fine root formation (Rudawska and Kieliszewska-Rokicka 1997) and increase conifer growth in the complete absence of mycorrhiza formation (Karabaghli-Degron ef al. 1998). Mycorrhizal fungi can also influence the ability of rhizobacteria to colonize roots (Belimov et al. 1999, Marschner and Crowley 1996), possibly through competition for carbon (Marschner and Crowley 1996) or the modification of root exudates that are released into the rhizosphere (Curl and Truelove 1986). Mycorrhizae might be able to affect PGPR-mediated growth promotion of mycorrhizal seedlings, by masking PGPR effect with their own growth-promoting effects, or by reducing the ability of PGPR to adequately colonize the root and exert a growth-promoting effect. PGPR-mediated growth promotion of mycorrhizal conifer seedlings has been reported (Shishido ef al. 1996a,b) but may depend upon the PGPR strain used and the plant species inoculated (Shishido etal. 1996b). 1.7 Phytohormones and plant growth Phytohorrnones have been described as organic substances other than nutrients (i.e., not a substance which supplies carbon and energy, nor an essential mineral) which are active in minute quantities, formed in one part of the plant and translocated to other sites, where they can evoke specific biochemical or physiological responses (Moore 1989). Commonly recognized classes include auxins, gibberellins, cytokinins, abscisic acid, ethylene, and brassinosteroids (Moore 1989). lndole-3-acetic acid (IAA) is the most abundant naturally occurring auxin, although a variety of lAA-related indoles have been found to occur naturally in 11 various plant species (Ernsten and Sandberg 1988, Normanly 1997). Naturally-occuring cytokinins include isopentenyl adenine and its numerous derivatives, including zeatin and zeatin riboside (Moore 1989). Exogenous auxins and cytokinins produced by rhizosphere microflora, including mycorrhizal fungi (Karabaghli-Degron etal. 1998, Normand etal. 1996, Rudawska and Kieliszewska-Rokicka 1997, Scagel and Linderman 1998a) and bacteria (Barazani and Friedman 1999, Glick 1995, Nehl etal. 1996, Nieto and Frankenberger 1989, Omay ef al. 1993, Schippers ef al. 1987) may exert a direct effect on plant physiology. Lateral root development in pine seedlings is governed by levels of auxin and cytokinin, where a high ratio of auxin:cytokinin in root tissues seems to promote lateral root development (Atzmon ef al. 1997). Exogenous applications of auxin have been shown to induce tumors, which resemble lateral root primordia, in roots (Christiansen-Weniger 1998). Cytokinins may control the ability of plants to respond to stimuli such as wounding or pathogen attack (Sano ef al. 1996), and root-produced hormones can modify the extent of shoot growth as well as increase leaf photosynthetic capacity (Aiken and Smucker 1996). Some PGPR have been found to influence plant responses to herbivory or pathogen attack (Tuzun and Bent 1999) or to directly influence photosynthetic rates (Zhang ef al. 1997), and it is tempting to speculate that they may do so in part through the production of cytokinins or other phytohormones. Alternatively, rhizosphere microorganisms may stimulate plants via some other kind of signal, resulting in endogenous changes in phytohormone levels or sensitivity to phytohormones, and altered plant growth. For example, elicitors (fragments of plant or fungal cell walls, produced by enzymatic hydrolysis) can increase the sensitivity of plant tissues to auxin (van der Krieken et al. 1997). 1.8 Lodgepole pine and Paenibacillus polymyxa The PGPR-plant interaction chosen for study was the Paenibacillus polymyxa (formerly Bacillus polymyxa; Ash et al. 1993) - lodgepole pine system. Lodgepole pine is the most widely distributed conifer in British Columbia (Brockley 1996), and a species of interest to the forestry industry for its ability to regenerate clearcuts (Daintith and Newsome 1996) and to provide veneer, lumber, and pulp for paper manufacture (Spelter etal. 1996). Paenibacillus polymyxa strain L6 was isolated from rhizosphere soil of a perennial ryegrass and white clover pasture (Holl et al. 1988). Strain Pw-2, tentatively identified as P. polymyxa, was isolated from surface-sterilized roots of a naturally regenerating lodgepole pine seedling (Shishido etal. 1995). Both strains are capable of promoting lodgepole pine seedling growth (Holl and Chanway 1992, 12 Shishido et al. 1995). P. polymyxa parental strains L6 and Pw-2 were used instead of their respective rifampicin-resistant derivative strains (L-16R and Pw-2-R) because the introduction of marker genes may alter the ability of bacteria to grow and colonize soil or roots (Tang et al. 1995, van Elsas etal. 1994). Both P. polymyxa strains L6 and Pw-2 have been demonstrated to produce IAA (Srinivasan etal. 1996), but it is not known whether this forms the basis of the PGPR-pine growth promotion interaction. P. polymyxa strains which produce cytokinins have also been described (Timmusk et al. 1999). 1.9 Hypotheses The following hypotheses were examined: 1. The extent of PGPR-mediated growth promotion seen under gnotobiotic conditions is influenced by the presence of a co-inoculated rhizobacterium. 2. A change in the extent of PGPR-mediated plant growth promotion is related to a quantitative change in the extent of PGPR population density on or within the root. 3. A change in the extent of PGPR-mediated plant growth promotion is related to a qualitative change in PGPR colonization of the root surface. 4. The extent of PGPR-mediated plant growth promotion is related to levels of auxins and/or cytokinins within root tissue. 5. PGPR strains Paenibacillus polymyxa L6 and Pw-2 vary in ability to form spores able to survive in soil and later germinate. 6. The extent of PGPR-mediated growth promotion seen under gnotobiotic conditions is influenced by the presence of a co-inoculated ectomycorrhizal fungus. 13 Chapter 2: Co-inoculation effects of Curtobacterium flaccumfaciens PF322 with Paenibacillus PGPR on extent of pine growth promotion and PGPR root colonization 2.1 Introduction These experiments were conducted to determine whether the presence of a competing rhizobacterium (strain PF322, identified as Curtobacterium flaccumfaciens) altered the ability of Paenibacillus polymyxa PGPR strains L6 and Pw-2 to (1) colonize lodgepole pine seedling roots and (2) promote lodgepole pine growth, during the first 12 weeks of seedling development. The P. polymyxa strains were described earlier (section 1.8). These results have been reported previously (Bent and Chanway 1998). 2.2 Materials and Methods Sources of bacterial strains The sources of Paenibacillus polymyxa PGPR strains L6 and Pw-2, and the rationale for using the unmodified parental strains instead of antibiotic-resistant derivative strains, were noted previously (section 1.8). Strain PF322 was isolated from the rhizosphere of a naturally-regenerating lodgepole pine seedling (ca. 3 years old) growing in a stand near Fort St. John, BC (M. Shishido, unpublished data). This strain was one of twenty-seven root-associated isolates originating from forest soils which had neutral or deleterious effects on pine growth in screening assays (M. Shishido, unpublished data). Six of these isolates (PF214, PF216, PF217, PF312, PF313, PF322) were chosen and screened further for DRMO activity. Source and surface sterilization of pine seeds Pinus contorta var latifolia (Dougl. Engelm.) seeds were collected from a stand in the Cariboo region of B.C. (52°06" Eat., 121°20' long.). Seeds were surface sterilized by immersion in 70% ethanol for 70 seconds, followed by a single rinse with sterile distilled water, then immersion in 3% NaOCI for 2 minutes, followed by three rinses with sterile distilled water. Surface sterility was verified by incubating several seeds on half-strength tryptic soy agar (TSA) plates (15 g tryptic soy broth, 15 g agar per L) for up to one week at room temperature and checking for microbial growth. Seeds were air-dried in a laminar flow hood overnight. Screening of soil isolates for DRMO activity Seedling growth assays were conducted in glass plant tubes (25 mm dia x 15 cm) sealed with translucent caps. Each tube contained 30 ml_ of a peat-vermiculite nursery mix (Sunshine Mix 4, Fisons Horticulture Inc. Vancouver B.C.) which had been moistened with 10 ml_ of distilled water and autoclaved for 30 minutes. No fungal or bacterial contaminants were observed after seven days of incubation at room temperature of similarly-treated nursery mix spread across TSA plates. Bacterial inocula were prepared as follows: 20 mL of half-strength tryptic soy broth (TSB, 15g/ L; Difco Laboratories, Detroit, Ml 48232) were inoculated with strain L6, Pw-2 or one of six isolates provided by M. Shishido (PF214, PF216, PF217, PF312, PF313 and PF322), and incubated overnight at room temperature on a shaker (150 rpm). Cultures were then diluted with fresh TSB, and the optical density (A^o) of each subculture was determined. After the Aeeo had doubled at least once, cells were collected by centrifugation at 4°C and 7700 x g, resuspended by gentle vortexing, and washed twice via centrifugation (also at 4°C and 7700 x g) in cold KP buffer (0.13 M KCI, 7 mM K 2 HP0 4 , 3 mM K H 2 P 0 4 , pH 7.2. Cells were resuspended and diluted to an A 6 6o of 0.1. Surface-sterilized pine seeds were placed in the bottom of sterile 10 mL beakers and soaked without agitation for 1 h in sterile KP buffer, or one of the bacterial suspensions. Treated seeds were aseptically sown in prepared plant tubes (one seed per plant tube) with sterilized forceps. Two experiments were conducted, with 36 tubes inoculated per treatment in the first experiment, and 15 tubes per treatment in the second. After sowing seed, tubes were placed in a growth chamber (Conviron CMP3244, Conviron Products Company, Winnipeg MB) for 6 weeks with a 16 h photoperiod (photosynthetic photon flux at canopy level ~ 155 umol s"1m"2) and a lightdark temperature regime of 23:15°C. After 6 weeks of incubation, seedlings were gently shaken from tubes and washed in tap water. Roots were severed from shoots and allowed to dry to constant weight at room temperature (about 5 days). The root and shoot weight data from both experiments were pooled and analyzed according to completely random design ANOVAs for unequally replicated treatments, with an error rate (a) < 0.05 (Steel and Torrie 1980). Multiple Student's t tests for unequally replicated means were used for further mean comparison (Steel and Torrie 1980). Inoculation and growth of pine seedlings for growth promotion/root colonization assays Seedling growth assays were conducted in glass plant tubes, prepared as indicated above for the DRMO screening experiments. Bacterial inocula for growth promotion experiments were also prepared as indicated above, with a minor modification to the procedure. Only strains L6, Pw-2 and PF322 were used, and after washing each prepared inoculum twice with cold KP buffer, each culture was diluted with KP buffer to an A 6 6o corresponding to an estimated cell density of 108 CFU/mL (0.8 for strains L6 and Pw-2, and 0.2 for PF322). Cultures were always harvested in the logarithmic phase of growth and resuspended, after washing, to the appropriate Aeeo-15 Bacterial colonization dynamics and seedling growth promotion were evaluated in five experiments. In the first and second experiments, pine seeds were placed in the bottom of sterile 10 mL beakers, and soaked without agitation for one hour in one of the four following treatments: (i) 4 mL of sterile KP buffer, (ii) 4 mL of strain L6 in KP buffer, (iii) 4 mL of strain PF322 in KP buffer, and (iv) a mixture comprising 2 mL of strain PF322 and 2 mL of strain L6, both in KP buffer. The third and fourth experiments were of similar design, with the exception that strain Pw-2 was used instead of strain L6. In the fifth experiment, strains L6 and Pw-2 were evaluated in pure culture and mixture following the methodology described above (i.e., seeds were soaked in (i) 4 mL of sterile KP buffer, (ii) 4 mL of strain L6 in KP buffer, (iii) 4 mL of strain Pw-2 in KP buffer, and (iv) a mixture comprising 2 mL of strain Pw-2 and 2 mL of strain L6, both in KP buffer). Two seeds were then taken from each beaker and vortexed in 2 mL of KP buffer with 0.01% Tween 20 for 10 seconds. The buffer was diluted and spread across TSA plates to estimate the number of bacteria present on each inoculated seed. Plates were incubated at room temperature for 2-3 days, after which colonies were picked or replica plated onto TSA containing nalidixic acid (30 ug/mL) or streptomycin (10 ug/mL). Treated seeds were aseptically sown in prepared plant tubes (one seed per plant tube) with sterilized forceps. Ninety plant tubes were used for each seed inoculation treatment in each experiment. After sowing seed, tubes were placed in a growth chamber (Conviron CMP3244, Conviron Products Company, Winnipeg MB) for 6, 9 or 12 weeks with a 16 h photoperiod (photosynthetic photon flux at canopy level ~ 155 umol s"1m"2) and a lighfdark temperature regime of 23:15°C. At week 10,1 mL of sterile distilled water was added to each tube to prevent desiccation. Verification of strain identities/contamination checks Characteristic profiles based on intrinsic antibiotic resistance, pectolytic activity, growth on modified TTC medium (Bochnerand Savageau 1977) containing soribitol, colony morphology and growth rate were developed for each bacterial strain (Table 2). To check for contamination during experiments, samples of bacterial inocula before and after seed bacterization, as well as all bacterial colonies growing on representative rhizosphere wash/ root macerate dilution plates, were routinely subjected to these tests, and results were compared with those from pure laboratory cultures of strains L6, Pw-2 and PF322. All bacterial cultures were stored in aliquots at -80°C as cell suspensions (from an overnight growth of culture in TSB) diluted 1:1 with 80% (w/v) glycerol. 16 Table 2: Distinguishing characteristics used to differentiate Paenibacillus polymyxa L6, P. polymyxa Pw-2 and Curtobacterium flaccumfaciens PF322 Medium Strain L6 Pw-2 PF322 Tryptic soy agar (TSA) + + + TSA + 10 ug/mL streptomycin - + -TSA + 30 ug/mL nalidixic acid - - + Pectolytic activity medium (PEC-YA) 3 P P NP Indicator medium with sorbitol (TTC-sorb)b w P w Phenotype Colony morphologyc B B Y Growth rate (days)d 2 2 4 Notes: +, growth (antibiotic resistance); -, no growth (anti biotic susceptibility); P, pectinolytic; NP, not pectinolytic; p, pale pink; w, white (after 2 days incubation only); B, light beige, matte, slightly serrate colonies 4-8 mm in diameter; Y, deep yellow, mucoid, entire colonies 1-3 mm in diameter. "Modified from Starr et al. (1977): contains 10 g sodium polypectate, 10 g yeast extract, 15 g agar and 1 mL of a cresol red solution (0.76% (w/v) in 0.002 N NaOH) per L, pH 7.8. A 1 % (w/v) aqueous hexadecyltrimethylammonium bromide solution was used to flood plates to precipitate polypectate, and visualize zones of clearing around pectinolytic colonies. bModified from Bochner and Savageau (1977): contains 10 g glycerol, 15 g agar, 7 g K 2HP0 4, 3 g KH 2P0 4, 2 g proteose peptone, 0.1 g MgS04, 2.5 g sorbitol and 2.5 mg 2,3,5-triphenyl tetrazolium chloride (TTC) per L. cColony morphology was determined on TSA plates 2-5 days after inoculation at room temperature. dGrowth rate refers to the number of days until visible colony morphology at room temperature. Growth promotion/root colonization assays at 6, 9 and 12 weeks Bacterial populations within the root and in the rhizosphere were enumerated as follows: thirty plant tubes were taken from each treatment 6, 9 and 12 weeks after seed inoculation. From these, five seedlings per treatment were chosen at random for shoot biomass determination and evaluation of root colonization. Seedlings were gently shaken from their tubes under aseptic conditions, and large soil fragments carefully removed with sterile forceps. Shoots were separated from roots, dried to a constant weight at room temperature (at least 5 days) and weighed. The root was transferred aseptically to a flask containing 5 mL of KP buffer with 0.01% Tween 20 and shaken (100 rpm) for 20 minutes. Roots were then removed and the length of 17 the primary and lateral roots measured before surface sterilization: 1 minute in 70% ethanol, 10 seconds in sterile distilled water, then 3 minutes in 3% sodium hypochlorite followed by two 10-second rinses with sterile distilled water. Roots were ground in 1 mL of chilled KP buffer containing 0.01% Tween 20. Root interior and rhizosphere populations were enumerated by diluting and plating root macerate and root washes onto TSA or TSA plus nalidixic acid plates, as described above. The minimum detectable population sizes for the root interior and rhizosphere were ~10 and 5000 CFU/ root, respectively. All colonies were assessed for their resistance to streptomycin or nalidixic acid by picking or replica plating onto the appropriate medium. Seedlings in the remaining test tubes were gently removed by shaking the tube, and rinsed under tap water. Root lengths (primary and lateral roots) were measured manually, roots were severed from shoots and each dried and weighed as described above. Statistical analysis The number of observations for each treatment varied with the number of healthy germinants. Dead or dying seedlings suffering from severe desiccation (containing brown, dried-out needles: 6.0% of the total) or symptoms of corky root disease (highly elongated, coiled stems and stubby roots: 1.3%, or 23 seedlings) were omitted from analyses. In addition, data from the Pw-2 treatment in the second Pw-2/PF322 experiment were also omitted from the analyses because this treatment was contaminated with strain PF322. In total, seven treatments were evaluated in the five experiments, i.e., seedlings inoculated with: (i) strain L6 alone, (ii) strain Pw-2 alone, (iii) strain PF322 alone, (iv) strains L6 + PF322, (v) strains L6 + Pw-2, (vi) strains L6 + PF322, and (vii) strains Pw-2 + PF322. Seedlings treated with the same bacteria in different experiments were observed to respond similarly, therefore data from the same treatment in different experiments were pooled to increase the sample size. Residual errors were generally well-distributed after pooling. Data were analyzed according to completely random design ANOVAs for unequally replicated treatments, with an error rate (P) < 0.05 (Steel and Torrie 1980). Rhizospheric and root interior bacterial population sizes (CFU/ mm root only) were analyzed according to randomized complete block design ANOVAs (P < 0.05) for unequally replicated treatments, where harvest dates (6, 9 and 12 weeks after sowing) constituted blocks. Multiple Student's t tests for unequally replicated means were used for further mean comparison (Steel and Torrie 1980). 18 2.3 Results Screening of soil isolates for DRMO activity The results of the DRMO screening experiments are summarized in Table 3. None of the putative DRMO isolates exhibited an ability to significantly retard lodgepole pine growth after a 6-week period of incubation. Insignificant decreases in both root and shoot biomass accumulation were noted in the PF322 treatment, and strain PF322 was chosen in the hope that these decreases would become significant with a longer period of incubation. Biomass accumulation was also not observed in the PGPR treatments after 6 weeks of incubation (Table 3). Table 3: Effects of PGPR {Paenibacillus polymyxa L6 and Pw-2) and six different putative DRMOs on lodgepole pine growth in screening tests, six weeks after inoculation Treatment Shoot biomass (mg) Root biomass (mg) Uninoculated control 4.9 1.1 L6 6.0 1.3 Pw-2 5.7 1.3 PF214 5.1 1.4 PF216 5.1 1.6 PF217 7.7* 1.0 PF312 5.0 1.5 PF313 5.3 1.4 PF322 4.8 0.99 Notes: * indicates a significant difference (P<0.05) from the uninoculated control treatment. Identification of strain PF322 Strain PF322 was identified as Curtobacterium flaccumfaciens on the basis of cell and colony morphology, substrate utilization (BIOLOG system, Bochner 1989) and fatty acid composition determined using the MIDI system at Auburn University, according to Kloepper et al. (1992) (data not shown). Bacterial population dynamics in the rhizosphere When singly-inoculated, strains L6 and Pw-2 maintained similar average population sizes that were lower than those of strain PF322 by at least an order of magnitude (Fig. 1). When co-inoculated with either strain L6 or Pw-2, the population size of strain PF322 was Weeks of incubation 0 3 6 9 12 Weeks of incubation Figure 1: Average rhizosphere population sizes of strains when inoculated singly and co-inoculated. At week 0, average CFU/seed is reported. For clarity of presentation, error bars were omitted and symbols depicting some treatment means were offset slightly at each time point, (a) Paenibacillus polymyxa L6, (b) P. polymyxa Pw-2 and (c) Curtobacterium flaccumfaciens PF322 populations in various treatments. Lowercase letters above graph symbols indicate significant differences (P<0.05) between two or more populations: a, L6 alone < L6 with PF322, b, Pw-2 alone > Pw-2 with L6; c. PF322 alone > PF322 with L6 and PF322 with Pw-2. 20 reduced relative to levels associated with plants that received only strain PF322 (Fig. 1c). Similarly, when co-inoculated with strain PF322, the density of strain L6 was reduced 6 and 12 weeks after treatments were administered, relative to levels associated with plants inoculated with strain L6 alone (Fig. 1a). Co-inoculation did not affect levels of strain Pw-2 (Fig. 1b). Rhizosphere population sizes of all three strains were stable throughout the experimental period with two exceptions: (i) when co-inoculated with strain PF322, strain L6 had a larger rhizosphere population 9 weeks after inoculation, compared to its population levels 6 and 12 weeks after inoculation (P< 0.05; Fig. 1), and (ii), when co-inoculated with strain L6, the rhizosphere population size of strain Pw-2 was significantly smaller 12 weeks after inoculation than it was at the previous two sampling times (P < 0.05; Fig. 1). Bacterial population dynamics in the rootintenor Paenibacillus polymyxa L6 was never observed in the root interior, while both P. polymyxa Pw-2 and C. flaccumfaciens PF322 were consistently found as endophytes. Variation in endophytic population densities between treatments and weeks was generally insignificant with two exceptions. When co-inoculated with strain L6, the size of internal root population of strain Pw-2 was significantly smaller 12 weeks after inoculation than it was six and nine weeks after treatments were established (P < 0.05; Fig. 2a). In addition, strain PF322 had a higher root endophytic population size at the six-week sampling time when it was inoculated alone compared to its population when co-inoculated with strain Pw-2 (Fig. 2b). Bacteriostatic effect of strain L6 on strain PF322 Strain PF322 was unable to grow in the vicinity of strain L6 colonies on TSA medium (zones of inhibition were 2-3 mm wide), but it grew normally near strain Pw-2 colonies. This phenomenon was consistently observed on all plates containing both strains L6 and PF322, whether incubated at room temperature or 30°C. When plates containing colonies of strains L6 and PF322 on TSA medium were replica plated onto fresh TSA containing 30 ug/mL nalidixic acid, colonies of strain PF322 which were not visible on the TSA medium (i.e., were within a zone of inhibition) were observed to grow (data not shown). Growth effects of strains Pw-2, L6 and PF322 on lodgepole pine Strain Pw-2 promoted an increase in root branching (Fig. 3), which was accompanied by an increase in combined lateral root lengths, that was early (6 weeks post-inoculation) compared to the noninoculated control and strain L6 treatment (Fig. 4b). When inoculated alone, strain Pw-2 increased shoot biomass after 9 and 12 weeks of seedling growth (Fig. 5a). 21 a ) 1000 o o E e o CN CN CO 100 •H" PF322 inoculated alone co-inoculated with L6 co-inoculated with Pw-2 Weeks of incubation Figure 2: Average root interior population sizes of strains when inoculated singly and co-inoculated. For clarity of presentation, error bars were omitted and symbols depicting some treatment means were offset slightly at each time point, (a) Paenibacillus polymyxa Pw-2 and (b) Curtobacterium flaccumfaciens PF322 populations. Lowercase letters above graph symbols indicate significant differences (P<0.05) between two populations: a, Pw-2 alone > Pw-2 with L6; b, PF322 alone > PF322 with Pw-2. 22 Weeks of incubation Uninoculated control VsWK L6 alone ESSS Pw-2 alone PF322 alone F===1 L6 + Pw-2 mm L6 + PF322 FfflTO Pw-2 + PF322 Figure 3: Average number of lateral roots per seedling for each treatment after 6, 9 and 12 weeks of incubation. Error bars indicate standard deviations. Treatments sharing the same letter are not significantly different at P<0.05. 23 E E o> c O 2 E •n a. 140 120 100 80 60 40 20 o 8 5 a> c 5 E o O 9 e,s 120 -i 100 A E E, <o jr o> 80 60 A 40 20 Weeks of incubation b,o c a,b • Uninoculated control UZZZZA L6 alone Pw-2 alone RSSgsga PF322 alone ^ 3 L6 + Pw-2 irrrrrn L6 + PF322 m m Pw-2 + PF322 Weeks of incubation Figure 4: Average lengths of (a) primary and (b) combined lateral roots for each treatment after 6, 9 and 12 weeks of incubation. Error bars indicate standard deviations. Treatments sharing the same letter are not significantly different at P<0.05. a ) 24 d A e d e '.A? a,c a c c Weeks of incubation Uninoculated control VZZZZA L6 alone Pw-2 alone KgggS PF322 alone f=\ L6 + Pw-2 rTTTTTI L6 + PF322 rrTfffl Pw-2 + PF322 Weeks of incubation Figure 5: Average (a) shoot and (b) root biomass for each treatment after 6, 9 and 12 weeks of incubation. Error bars indicate standard deviations. Treatments sharing the same letter are not significantly different at P<0.05. 25 In contrast, strain L6 alone did not stimulate root branching or promote biomass accumulation until week 12, when elongation of the primary root, and increases in root biomass accumulation and branching were observed (Figs. 3,4, 5). The competitor strain PF322 had a modest effect on lodgepole pine growth. Twelve weeks after inoculation, seedlings inoculated with PF322 had a similar number and length of lateral roots, and similar amounts of root and shoot biomass, as noninoculated control seedlings, despite an increase in primary root elongation (Figs. 3,4, 5). Effects of co-inoculation on PGPR-mediated growth promotion When strain PF322 was combined with the PGPR strain L6, co-inoculated seedlings did not grow differently (in terms of biomass accumulation or root elongation or branching) than seedlings inoculated with L6 alone (Figs. 3,4, 5), with one exception. Combined lateral root lengths were greater in 6-week old co-inoculated seedlings, but this difference was transient and could not be detected in subsequent harvests. When PF322 was combined with PGPR strain Pw-2, a decrease in primary root length (relative to the Pw-2-inoculated treatment) was observed in co-inoculated seedlings at 12 weeks after inoculation (Fig. 4). A significant decrease in shoot biomass was also observed (Fig. 5). However, the number and length of lateral roots in co-inoculated seedlings were similar to those in seedlings inoculated with PW-2 alone (Figs. 3, 4). Interestingly, in some cases co-inoculation with strains L6 and Pw-2 resulted in growth inhibition of seedlings after 12 weeks compared to treatment with their respective pure cultures (e.g, Figs. 3,4, 5b). 2.4 Discussion Curtobacterium flaccumfaciens strain PF322 significantly decreased Paenibacillus polymyxa Pw-2-mediated shoot biomass accumulation and primary root length in pine, but these effects were not linked to a concomitant decrease in the population size of strain Pw-2 in the rhizosphere or in the root interior of co-inoculated seedlings. This may indicate that colonization of specific root microsites, such as root hair zones, lateral root junctions or root cortical cells may be more important for Pw-2-mediated stimulation of plant growth than the maintenance of a large population along the entire root. Strain PF322 may have displaced strain Pw-2 from these microsites, but further work is required to evaluate this possibility. The growth inhibition that resulted from co-inoculation with Paenibacillus polymyxa strains L6 and Pw-2 presents an interesting conundrum regarding which of the two microoganisms was inhibitory. The beneficial effects of Pw-2 (e.g., increased root number and 26 lateral root length) were neutralized more often than those of L6 (e.g., increased primary root length), which suggests that Pw-2 was more sensitive to L6 than vice versa. The endophytic strain PF322 also reduced PGPR efficacy of strain Pw-2 (Fig. 4). Based on these results, it appears that competing microorganisms colonizing internal or external plant microsites may inhibit the activity of endophytic PGPR. Strain Pw-2 is able to colonize conifer seedling stems after pure culture inoculation of seeds or roots (Shishido 1997), which suggests that host plants don't perceive this strain to be a pathogen. We have not determined if strain Pw-2's growth promotion capabilities are linked to its endophytic capacity. If they are,'one way co-occurring strains such as PF322 or L6 could inhibit strain Pw-2 is by activating plant defense mechanisms, thereby excluding it from moving systemically in the plant. This possibility can not be evaluated in the present study because internal stem tissue colonization was not assessed. Alternatively, strain Pw-2 may simply be a poor competitor when other microorganisms are present with population sizes similar to its own. Pine growth promotion by strain L6 was restricted to increases in root number and biomass. These parameters were not affected by co-inoculation with strain PF322, despite a decrease in the L6 population size when co-inoculated with strain PF322. These results might indicate that endophytic PGPR may be more susceptible to losses of efficacy in response to microbial competition than external root colonizing PGPR. Strain PF322 was identified as Curtobacterium flaccumfaciens. Members of this species have been identified as plant pathogens (Komaga and Suzuki 1986), but strain PF322 did not exhibit any visible pathogenic effects on pine. Strain PF322 is not a deleterious rhizosphere bacterium (DRB) on pine (Nehl etal. 1996), at least under the experimental conditions used, because it had a neutral or positive effect on the measured pine growth parameters when inoculated alone. Therefore, if the presence of strain PF322 or the PGPR strain L6 can partially mask Pw-2-mediated growth promotion, it is reasonable to hypothesize that DRB may neutralize the positive effects of strain Pw-2 or similar PGPR completely. Various DRB have been isolated from forest soils and linked to decreases in tree growth. For example, Probanza et al. (1996) isolated a variety of alder DRB from the rhizosphere soil of a natural alder stand. In a comparative study of the effects of spruce growth responses to soils and soil microflora from ecologically distinct regions of British Columbia, spruce DRB were isolated from unpasteurized Mackenzie area soil, which was found to inhibit spruce growth (O'Neill et al. 1992). Could DRB naturally present in soils completely mask the ability of an introduced PGPR to promote plant growth and cause the characteristic variability in seedling growth responses? 27 The complexities of microbe-microbe interactions render the task of finding effective PGPR difficult, as the associated microflora in screening assays could influence the extent of plant growth promotion. It is also clear that the results of root colonization experiments will depend in part upon the experimental methods chosen (Kloepper et al. 1991c, Kloepper and Beauchamp 1992). Whether a rhizobacterium appears to be plant-beneficial or not seems to depend at least in part, on conditions of the growth-promotion assay. This does not mean that PGPR and DRB do not exist, but that the classification of a rhizobacterium into one of these groups is operational rather than absolute: a PGPR, for example, promotes the growth of a given range of plants, under a given range of conditions. Both Paenibacillus polymyxa strains L6 and PW-2 have been identified as PGPR in greenhouse experiments (Holl and Chanway 1992, Shishido et al. 1995), yet if one were to look only at the 6-week old seedlings in these experiments, strain L6 would probably be classified as a DRB. Interestingly, Chanway et al. (1991) observed rapid (i.e., after 6 weeks of incubation) L6-mediated root elongation in a gnotobiotic system similar to that used in this study, except that pine seedlings were well fertilized and seeds received a smaller direct dose of cells. Without a solid understanding of the mechanisms involved in a plant-microbe interaction, small changes in an experimental procedure can apparently influence results in unexpected ways. Paenibacillus polymyxa strains L6 and Pw-2 differed in their ability to colonize the root interior, had very different effects on root morphology and promoted seedling growth at different rates. Both these PGPR strains have been demonstrated to produce the auxin indole-3-acetic acid (IAA) in vitro (Srinivasan et al. 1996). Paenibacillus polymyxa strains L6 and Pw-2 might produce compounds which act, directly or indirectly, to alter levels or ratios of auxins, cytokinins or other phytohormones within pine roots and generate the observed changes in root morphology. However, the in vitro inhibitory effect of strain L6 on PF322 suggests that biocontrol may also be important in strain L6-mediated plant growth promotion under natural conditions. 28 Chapter 3: Co-inoculation effects of Pseudomonas fluorescens M20 with Paenibacillus PGPR on extent of pine growth promotion, PGPR root colonization, and levels of root phytohormones 3.1 Introduction The isolation, screening, identification and characterization of a Gram negative bacterium are part of the work described in this chapter. These activities were conducted to obtain a Gram negative isolate that could be used as a co-inoculant in microscopy experiments, which are described in Chapter 4. In order to clearly explain why a Gram negative isolate was needed, and why the already-characterized Gram positive rhizobacterium, Curtobacterium flaccumfaciens strain PF322, was not used as the co-inoculant instead, it is necessary to first describe some of the methods used in Chapter 4. One of the objectives of this dissertation was to determine whether a change in the extent of PGPR-mediated plant growth promotion is related to a qualitative change in PGPR colonization of the root surface. A suitable means of visually distinguishing between two different co-inoculated bacterial strains (one P. polymyxa strain, and one non-P. polymyxa strain) was required. Indirect immunofluorescence techniques, in which strain-specific antibodies bind to the bacterium, and are identified by a fluorescent compound conjugated to a secondary antibody which binds to the constant (Fc) region of the first, have been successfully used to identify various bacterial strains on or within plant roots (e.g. Chin-A-Woeng etal. 1997, Shishido etal. 1999, Schloter et al. 1993, Troxleref al. 1997). Antibodies, or immunoglobulins, are secreted glycoproteins that are able to recognize and bind to a unique epitope, or small, three-dimensional region of an antigen; antibodies can bind to more than one antigen if both antigens contain identical, or very similar, epitopes (Harlow and Lane 1999). When using immunofluorescence techniques, the accuracy of the image obtained depends largely upon how many of the antibodies present in the antiserum used bind to the target antigen of interest, and how many bind to other, non-target antigens. The polyclonal antiserum raised against bacterium A should not contain many antibodies that recognize epitopes on the surface of bacterium B, and vice versa. One way to minimize this cross-reaction phenomenon is to ensure that the surface of bacterium A is very different from the surface of bacterium B. The exterior surfaces of Gram positive and Gram negative species are distinct. Gram positive species may have one or more proteinaceous S-layers (Sara and Egelseer 1996) exterior to the peptidoglycan, while the outer membrane of Gram negative species contains lipopolysaccharide antigens as well as protein (Freer 1985). In order to minimize the possibility of cross-reactivity between polyclonal antibody preparations raised against Paenibacillus 29 polymyxa strains L6 and Pw-2, both Gram-positive species, and a coinoculated rhizobacterium, it was decided that the co-inoculated rhizobacterium should be a Gram negative species. In order to identify a Gram negative rhizobacterium, preferably with DRMO activity against lodgepole pine, Gram negative bacteria were isolated from soil that had previously yielded spruce DRMO (O'Neill etal. 1992) and screened for growth-inhibiting activity. The most inhibitory Gram negative isolate, strain M20, was chosen and identified as Pseudomonas fluorescens. Experiments to determine the ability of P. fluorescens M20 to affect the extent of Paenibacillus po/ymyxa-mediated pine growth promotion were conducted, so that the extent of root surface colonization observed (Chapter 4) could be related to the extent of growth promotion. Experiments to determine the ability of P. fluorescens M20 to affect the population densities of P. polymyxa PGPR were conducted simultaneously. Phytohormones regulate plant growth. If the growth of pines can be altered by inoculation with different rhizobacteria, this may be a result of altered levels of phytohormones in plant roots. Rhizobacteria may produce phytohormones that the plant absorbs, or alternately, may produce some other signal that results in an alteration in root hormone levels and subsequently in root and plant growth. In order to determine whether the extent of PGPR-mediated plant growth promotion is related to levels of auxins and/or cytokinins within root tissue, the levels of auxin and a cytokinin were measured in pine root tissues collected during the growth promotion experiments described above. 3.2 Materials and Methods Isolation of bacteria from Mackenzie soil Soil, consisting primarily of material from the Ah and LFH horizons, was collected from a planting site near Mackenzie, BC (55°N 123°W), using a shovel and container which had been rinsed briefly with 90% ethanol. The soil sample was kept on ice or refrigerated, and used within a week of collection. Small amounts of soil were removed from the collection container with an aseptic scoop and shaken across the surface of half-strength tryptic soy agar (TSA, 15 g tryptic soy broth, 15 g agar per L) plates containing 100 ug/mL cycloheximide (Cyc) to prevent fungal growth. Bacterial colonies were isolated by picking individual colonies and re-streaking them across fresh plates. Each isolate was re-streaked several times across Cyc media, until no contaminants were visually apparent, then picked and streaked several times across TSA. Pure cultures of twenty bacterial isolates, chosen on the basis of different colony morphologies, were obtained in this manner. 30 Gram stains were performed to verify the purity of each culture, and determine the cellular morphology of each isolated bacterium. Resistance to vancomycin was assessed by plating a small quantity of bacterial culture (grown overnight at room temperature in 10 mL TSB, with 150 rpm shaking) on TSA plates, then adding an antibiotic disc containing 30 ug vancomycin (BRL). The plates were incubated at room temperature, and examined for the formation of zones of inhibition. Screening of bacteria for DRMO activity Nine Gram-negative bacterial isolates with distinctive colony morphologies were chosen for screening (M1, M5, M6, M7, M11, M12, M13, M14 and M20). The screening assays were conducted in glass plant tubes, in a similar manner to the seedling growth promotion assays described in Chapter 2, with some adaptations. Each of the nine strains listed above was inoculated into a separate 50 mL flask containing 10 mL of TSB and allowed to grow overnight at room temperature, with 150 rpm shaking. PGPR strains Paenibacillus polymyxa L6 and Pw-2 were not used. The following day cells were pelleted by centrifugation, as described previously, washed twice with 10 mL volumes of cold KP buffer, then resusupended to an A660 of 0.1. Surface-sterilized pine seeds were soaked in 4 mL of sterile KP (uninoculated control treatment) or a bacterial suspension without agitation for one hour. Seeds were placed with sterile forceps into prepared test tubes (containing 30 mL of nursery mix, 11 mL distilled water, and autoclaved for 30 minutes), one seed per tube, and incubated for 6, 9 or 12 weeks in a growth chamber, with the settings described previously. No water was added to the tubes at week 10. Ninety test tubes were inoculated per treatment. After 6, 9 or 12 weeks, thirty test tubes were collected from each treatment. No attempts were made to determine bacterial colonization of the rhizosphere or root interior. Seedlings were gently shaken from each tube, rinsed in tap water, and the length of primary and lateral roots measured manually and recorded. Roots and shoots were severed and allowed to dry for at least 5 days at room temperature before weighing. Inoculation and growth of pine seedlings for growth promotion/ root colonization assays These assays were conducted in glass plant tubes, prepared and inoculated in a similar manner to the growth promotion assays described in Chapter 2, with some modifications. Tubes contained 30 mL of nursery mix and 12 mL of distilled water. Only bacterial strains L6, Pw-2 and M20 were used, and after washing each prepared inoculum twice with cold KP buffer, each culture was diluted with KP buffer to an Aeeo corresponding to an estimated cell density of 108 CFU/mL (0.8 for strains L6 and Pw-2, and 0.3 for strain M20). 31 Bacterial colonization dynamics and seedling growth promotion were evaluated in four experiments. In each experiment, pine seeds were placed in the bottom of sterile 10 mL beakers, and soaked without agitation for one hour in one of the six following treatments: (i) 4 mL of sterile KP buffer, (ii) 4 mL of strain L6 in KP buffer, (iii) 4 mL of strain Pw-2 in KP buffer, (iv) 4 mL of strain M20 in KP buffer, (v) a mixture of 2 mL each of L6 and M20, both in KP buffer, (vi) a mixture of 2 mL each of Pw-2 and M20, both in KP buffer. Two seeds were taken from each beaker and vortexed in 2 mL of KP buffer containing 0.01% Tween 20 for 10 seconds. The buffer was diluted and spread across TSA plates to estimate the number of bacteria present on each inoculated seed. Plates were incubated at 27°C for 2 days, after which colonies were picked or replica plated onto TSA containing streptomycin (10 ug/ mL), Pseudomonas Agar F (PAF, Difco Laboratories, Detroit, Ml 48232) or an indicator medium containing lactulose (MOPS-lac) (Table 4). Treated seeds were aseptically sown in prepared plant tubes (one seed per tube) with sterilized forceps. A scoopful of sterilized sand (about 1 cc) was added to each tube to cover the seed. One hundred and twenty plant tubes were used for each seed inoculation treatment in each experiment. After sowing seed, tubes were placed in a growth room illuminated by a fusion lamp (SolarRay), for 6, 9 or 12 weeks, with a 16 h photoperiod (photosynthetic photon flux at canopy level ~170 pmol s' 1 m'2) and a lightdark temperature regime of roughly 22:18°C. No water was added at week 10. Growth promotion/root colonization assays at 6, 9 and 12 weeks Bacterial populations in the rhizosphere were enumerated in a manner similar to the experiments described in Chapter 2, with several modifications. Forty plant tubes were taken from each treatment 6, 9 or 12 weeks after seed inoculation. Eight seedlings per treatment were chosen at each time point for an evaluation of root exterior colonization. Evaluations of root interior colonization were not made. Shoots were severed from roots, and shoots were placed on ice in pre-weighed plastic bags until they could be freeze-dried. For seedlings sampled after 6 or 9 weeks of incubation, the roots were washed in 50 mL flasks containing 5 mL of sterile KP with 0.01% (v/v) Tween 20; for seedlings sampled after 12 weeks, 8 mL of this solution was used. After sampling the root wash solution, flasks containing roots were stored on ice until root measurements could be made. Root wash solutions were diluted in 0.85% (w/v) NaCI and plated on TSA. Plates were incubated at 27°C for 2 days, and representative colonies from each sample were assessed for resistance to streptomycin, ability to grow produce fluorescent pigments on PAF, and colony colour on MOPS-lac medium (Table 4). Seedlings in the remaining test tubes were gently removed by shaking the tube, and rinsed in distilled water. Root measurements, including total length and number of lateral roots, 32 were made using a Hewlett-Packard flatbed scanner with a transparency lid at 600 dpi. Digital images of roots were scanned and analyzed with WinRhizo root analysis software (Regent Instruments, Inc., Quebec City, Quebec). All roots were rinsed, scanned, placed in pre-weighed plastic bags and kept on ice until they could be freeze-dried. Roots and shoots were frozen with liquid nitrogen, or by storage for at least 24 hours at -80°C. Multiple holes had been punched in the plastic bags previously, to allow samples to freeze-dry. Frozen samples were placed in a lyophilizer and freeze-dried for at least 16 hours. Upon removal from the lyophilizer, samples were placed in undamaged, airtight plastic bags and stored at 4°C until they could be weighed. Samples were allowed to come to room temperature in the sealed bags before weighing. After weighing, samples were stored in the airtight bags at -20°C. Verification of strain identities/ contamination checks Characteristic profiles based on intrinsic antibiotic resistance, ability to grow and produce water-soluble fluorescent pigment on PAF, and colony colour on indicator media containing different sugars, were developed for each strain (Table 4). To check for contamination during experiments, samples of bacterial inocula before and after seed bacterization, as well as all bacterial colonies growing on representative rhizosphere wash dilution plates, were routinely subjected to these tests. The results were compared with those from pure laboratory cultures of strains L6, Pw-2 and M20. All bacterial cultures were stored in aliquots at-80°C as cell suspensions (from an overnight growth of culture in TSB) diluted 1:1 with 80% (w/v) glycerol. Preparation of root extracts Root extracts were prepared from the freeze-dried root tissues kept from experimental replication 2, essentially as described in Vonderwell (1998), with some modifications to the procedure. Previously measured and weighed freeze dried roots were removed from storage at -20° C, thawed at 4°C, then brought to room temperature before use. Similar masses of root tissue were sampled for each treatment: around 18 mg of root tissue, or 9 to 16 roots, was sampled for week 6 seedlings. Around 79 mg (13 to 26 roots) was sampled for week 9 seedlings, and around 140 mg (14 to 22 roots) for week 12 seedlings. Lyophilized root samples from a single treatment and sampling time were pooled, suspended in extraction medium (80% methanol with 0.02% sodium diethyldithiocarbamate, an antioxidant) and homogenized on ice for 30 seconds with a Tissue-Tearor (Polytron, Inc.) homogenizer set at its maximum speed. 100 mL of extraction medium was used per gram dry 33 Table 4: Distinguishing characteristics used to differentiate Paenibacillus polymyxa L6, P. polymyxa Pw-2 and Pseudomonas fluorescens M20 Strain Medium L6 Pw-2 M20 Tryptic soy agar (TSA) + + ++ TSA + 10 ug/mL streptomycin - + + Pseudomonas agar F (PAF) +,nf +/-, nf ++,f Indicator medium with lactulose (MOPS-lac)a dark pink/cerise not mucoid light pink/ salmon mucoid white Indicator medium with sucrose (TTC-suc)b cerise salmon white Notes: + , growth; ++, profuse growth; -, antibiotic susceptibility; +/-, poor growth (barely visible); f, production of a water soluble greenish fluorescent pigment; nf, no pigment produced. ""Modified from Niedhardt etal. (1974): contains 100 mL of MOPS buffer (10X concentrate), 3.012 mg K2HP04, 15 g agar, 12.5 mg 2,3,5-triphenyl tetrazolium chloride (TTC), 5 g lactulose. "Modified from Bochner and Savageau (1977): contains 10 g sucrose, 15 g agar, 7 g K 2 HP0 4 , 3 g KH 2 P0 4 , 2 g proteose peptone, 0.1 g MgS0 4 and 12.5 mg TTC per L. Plates must be read before two days of incubation. weight of week 6 roots, while for week 9 and 12 roots, 20 mL of extraction buffer were used per gram dry root weight. After homogenization, samples were extracted at 4 °C for one hour. Samples were centrifuged in 14 mL Fisherbrand tubes (5 minutes, 3000 rpm, 4°C, Sorvall SS34 rotor with inserts) and the supernatant collected with a pipette. Fresh extraction medium was added to the pellet, and the homogenization, extraction and centrifugation process was repeated two more times. Supernatants were pooled and passed using mild suction through disposable Sep-Pak C18 vacuum columns (Waters, inc.) previously primed with 2 mL of extraction medium. One mL of extraction medium was passed through the column after passage of the sample. Samples were then reduced to a residual volume of 2 mL or less under vaccuum (using a Speed-Vac). Samples were each brought to a volume of exactly 2 mL by the addition of fresh extraction medium where necessary, then each sample was divided into two aliquots of one mL each. These aliquots were frozen at -80°C until use (3-4 days). 34 One aliquot from each sample was dried under vacuum and /or lyophilized, then resuspended in 1 mL 25 mM aqueous Tris-buffered saline, pH 7.5 (TBS), and used to determine levels of dihydrozeatin riboside (DHZR) in root tissues. The other aliquot would be methylated, then used to determine levels of methyl indole-3-acetic acid (melAA) in root tissues. The methylation procedure was as follows: Each one mL aliquot was diluted to a pH of 9.0 with cold 5% NaOH (0.5 mL added per mL sample) and partitioned twice with 3 mL volumes of ether, which was added to tubes and vortexed (setting 7) for 5 minutes. Ether phases were discarded. The aqueous phase was then adjusted to a pH of 2.5 with cold, dilute 0.01 M HCI (0.695 mL added to 1.5 mL diluted sample) and partitioned three times with 0.75 mL ether. Aqueous phases were discarded. The ether phase was collected, reduced to dryness under a stream of N 2 , then redissolved in 400 uL of methanol. Diazomethane was freshly prepared: 50 mg Diazald (N-methyl-N-nitroso-p-toluene sulfonamide) was added very slowly to a continuously stirred mixture of 10 mL deionized distilled water and 10 mL ether in an unscratched 25 mL flask, immersed in a bath of ice and ethanol. 100 uL of the ether phase (containing diazomethane) was added to each sample, and the methylation reaction was allowed to proceed for two minutes. Excess diazomethane was deactivated by the addition of 50 uL 0.2 M acetic acid in methanol to each sample. Ether was evaporated under a stream of N 2 , then samples were reduced to dryness under vaccuum, resuspended in 100 uL methanol, and diluted by the addition of 900 pL TBS. The diluted samples were stored at -80°C until they were used to determine levels of melAA in root tissues. Phytohormone analyses Commercially available Phytodetek ELISA kits (Sigma-Aldrich Chemical Co., St. Louis, MO 63178) were used in accordance with the manufacturer's instructions to determine levels of DHZR and melAA in root tissues. A standard curve which incorporated lower concentrations (a minimum of 0.75 pmol/0.1 mL) of melAA was used in melAA assays conducted on week 9 and 12 samples to improve the sensitivity of the assay, since results with week 6 tissues indicated that the levels of melAA in root tissues were too low to be calibrated using the standard curve suggested by the manufacturer. For each set (three sets in total: one from each of week 6, 9 and 12 root tissues) of six samples, two assays were conducted. In the first, the most appropriate dilution to use for each sample was determined by preparing a range of dilutions of each sample (undiluted to 1/1000 dilutions) and inoculating one well with each. In the second, the amount of phytohormone was quantified by inoculating 7-8 wells with the most appropriate dilution of each sample (usually undiluted, or a 1/10 dilution), as determined in the previous experiment. A405 was measured using a Dynatech MR 700 plate reader (Dynatech Laboratories). Data analyses Statistical analyses for the DRMO screening experiment were conducted according to completely random design ANOVAs for unequally replicated treatments, with an error rate (P) < 0.05 (Steel and Torrie 1980). Multiple Student's t tests for unequally replicated means were used for further mean comparison (Steel and Torrie 1980). For all other experiments in this chapter, statistical analyses were conducted using SAS v.6.12 software, with an error rate (P) < 0.05 unless otherwise indicated. Of the four growth promotion/ root colonization experiments conducted, data were only pooled and utilized from two of these (the second and third), owing to problems with contaminants in the first replication and a breakdown of the growth room lights during the fourth (10 days of no illumination were followed by 12 days of decreased illumination by six 100-watt plant grow lights, photosynthetic photon flux at canopy level averaging 130 pmol s"1 m'2; beginning during the fifth week after inoculation). Analysis of variance by the general linear models procedure was performed on all seedling variable data (pooled), and on pooled data from replications two and three. Analysis of variance by the general linear models procedure was also performed on rhizosphere CFU and density (CFU/root surface area) data from replications two and three. Comparisons were made between all treatments, and also between treatments incorporating a bacterial strain (L6, Pw-2 or M20) in order to estimate differences in the rhizosphere populations of each strain when singly- or when co-inoculated. Duncan's procedure for means separation was also applied to seedling variable and rhizosphere population data. Calculations of phytohormone levels were conducted according to the ELISA kit manufacturer's instructions. At least six A 4 05 readings from the microwell plates were used to calculate percent binding, and the concentration of phytohormone present in each sample was determined from the standard curve. Analysis of variance and Duncan's mean separation procedures were performed on calculated pmol hormone/ mg dry root weight values using SAS v. 6.12 software and an error rate (P) < 0.05. 36 3.3. Results Screening of soil isolates for Gram negative DRMO Strain M20 was chosen for further study since it was the only isolate to significantly impair seedling growth in terms of primary root elongation twelve weeks after inoculation, and seemed to be least stimulatory to pine growth (Table 5). Identification of strain M20 Strain M20 was identified as Pseudomonas fluorescens on the basis of cell and colony morphology, substrate utilization (BIOLOG system, Bochner 1989) and fatty acid composition determined using the MIDI system at Auburn University, according to Kloepper et al. (1992) (data not shown). Bacterial population dynamics in the rhizosphere When singly inoculated, Paenibacillus polymyxa PGPR strains maintained different average populations, with that of strain Pw-2 being about twice as large as that of strain L6 (Fig. 6a). The populations of both PGPR strains were lower, however, than those of strain M20 by at least an order of magnitude (Fig. 6b). When co-inoculated with either strain L6 or Pw-2, the population size of strain M20 was reduced relative to levels associated with plants that received only strain M20 (Fig. 6b), but only at 6 weeks post-inoculation. The population of M20 on plants co-inoculated with M20 and Pw-2 increased between weeks 9 and 12 (Fig. 6b). Interestingly, levels of strain Pw-2, which were initially lower on plants co-inoculated with Pw-2 and M20 than on plants inoculated with Pw-2 alone, also increased at 12 weeks after inoculation to a level similar to that of singly (Pw-2)-inoculated plants (Fig. 6a). When co-inoculated with strain M20, the density of PGPR strain L6 in the rhizosphere rose, but to a lesser extent than on singly (L6)- inoculated plants. The density of L6 on plants co-inoculated with L6 and M20 was decreased relative to the density found on singly (L6)-inoculated plants 9 and 12 weeks after inoculation (Fig. 6a). Growth effects of strains L6, Pw-2 and M20 on lodgepole pine When singly-inoculated onto lodgepole pine seeds, strains L6, Pw-2 and M20 all stimulated increases in seedling growth under normal experimental conditions. Rapid growth responses, i.e. observed at 6 weeks after inoculation, were seen in seedlings singly-inoculated with strain Pw-2, which stimulated significant increases (relative to the unbacterized control 37 treatment) in root biomass accumulation and total root elongation (Fig. 7a,c). Significant increases in shoot biomass accumulation and lateral root formation followed, at 12 weeks after inoculation (Fig. 7b,d). Rapid (at 6 weeks post-inoculation) growth responses were also seen in seedlings singly-inoculated with strains L6 (increased root elongation) and M20 (increased root elongation and lateral root formation) (Fig. 7c,d). Seedlings singly-inoculated with strain L6 also had increased root biomass accumulation and lateral root formation at 9 weeks after inoculation (Fig. 7a,d), and increased shoot biomass accumulation at 12 weeks after inoculation (Fig. 7b), relative to the unbacterized control treatment. Seedlings singly-inoculated with strain M20 demonstrated some slower growth responses as well, with a significant increase in lateral root formation visible at 9 weeks after inoculation (Fig. 7d), and increased shoot biomass accumulation visible at 12 weeks after inoculation (Fig. 7b), relative to the unbacterized control treatment. Effects of co-inoculation on PGPR-mediated growth promotion When strain M20 was co-inoculated with strain L6 onto lodgepole pine seeds, there was no significant difference in the growth of the seedlings relative to those singly-inoculated with strain L6 alone (Fig. 7). However, when strain M20 was co-inoculated with strain Pw-2 onto lodgepole pine seeds, the co-inoculated seedlings grew differently than seedlings inoculated with strain Pw-2 alone. There were some early improvements in the growth of co-inoculated seedlings vs. those inoculated with Pw-2 alone: at 6 weeks after inoculation, root elongation, root biomass accumulation and lateral root formation were all improved (Fig. 7a,c,d). However, at 9 weeks after inoculation, these increases in root biomass accumulation and lateral root formation disappeared (Fig. 7a,d), and at 12 weeks after inoculation, the increase in root elongation also vanished (Fig. 7c). At 12 weeks after inoculation, there were significant decreases in the growth of seedlings inoculated with Pw-2 and M20 versus those inoculated with Pw-2 alone: lateral root formation and root biomass accumulation were both impaired (Fig. 7a,d), the latter parameter to the point where it was no longer significantly different from the unbacterized control treatment (Fig. 7a). Levels of auxin and cytokinin in root tissues The levels of auxin (indole-3-acetic acid, IAA) and a cytokinin (dihydrozeatin riboside, DHZR) in extracts of root tissues sampled during replication 2 are presented in Table 6. When all treatment means were compared, no significant differences between treatments were observed in roots harvested 6 or 9 weeks after inoculation. Significant differences in root 38 o CM CO i CM CO CO CO Is-o o 00 CM CO to OO to CO c CD E " c o 9> CM CO to h o co CO CO |<0 T" m co CO If) CO to : o ' If) in CO CO c o IO CM CN CO CO CO CO CO CD CD to CD . 2 * I •5 I x : .31 I a 8 x : CO o 2 2 E i ! J E CD c E o O co CO 00 o o CO o 00 o CO m co to m co to m 00 T— d co o 2 2 . f i B =6 c o c o — < u i i E z I Si I O O |w Is-O O CD .a* I •o 1 x : CO in to in to co to to CM Is-Is-co to 00 to "5 S s< E • e Q . CM CM to to CM to CO |(0 CO CM CM leo O CM [</> 00 |03 CM CM to" S E CO f= •2 CO " S i , X> CD O o to to CM co iri to CO 06 |eo CO oS to 1^  o Is-' to Is-td CM to o s 1 c? "5 8 CD CD E 3 00 .St I # o o a: to Is-co to to co CO in CO m to Is-.31 I x : CO CO i E 0 . co 00 10 CO to CO to CO m CO o m to Is-CO O CO o p 2 E a > E CO ^ " — CO •o x : P X) CD E -o o to co to Is-to CO to Is-oS CO o p I f « =5 7 5 i i E z m o d v Q. c CD E CO CD •o CD 3 O o CD x : E 2 8 c I •D c s c D) '<0 c o CO I 8 c 39 a) 8.0e+5 A ^ 7.08+5 6.08+5 5.08+5 4.0e+5 A 3.0e+5 2.08+5 1.0e+5 A 0.0 4 b ) 2.5e+7 2.0e+7 A 1.5e+7 1.0e+7 5.08+6 A Weeks after inoculation a: Pw-2 > all other treatments at week indicated b: Only other significant difference at week 12 is between Pw-2 + M20 and L6 + M20 treatments C: Increases (L6) from week 6 to week 9 d: Increases (Pw-2 + M20) from week before to week after e: Increases (L6 + M20) from week 6 to week 9 L6 —®- L6 + M20 —V- Pw-2 Pw-2 + M20 a: M20 > other treatments at week 6 b: Increases (Pw-2 + M20) from week 9 to week 12 M20 -@- L6 + M20 — Pw-2 + M20 Weeks after inoculation Figure 6: Average rhizosphere population sizes of (a) Paenibacillus polymyxa strains L6 and Pw-2, and (b) Pseudomonas fluorescens strain M20 when inoculated singly and co-inoculated Symbols depicting some treatment means were offset slightly at each time point for clarity of presentation. Error bars represent standard deviations. Lowercase letters indicate significant differences (P<0.05) as indicated on the right. 40 Figure 7: Average (a) root dry weight, (b) shoot dry weight, (c) root length and (d) number of lateral roots measured for seedlings in each treatment after 6, 9 and 12 weeks of incubation. Treatments sharing the same letter are not significantly different at P<0.05. 41 hormone levels were observed in the largest (harvested 12 weeks after inoculation) roots. Seedlings which had been inoculated with either Paenibacillus PGPR strain had significantly higher levels of root IAA after 12 weeks of incubation, regardless of whether the Paenibacillus strain was singly-inoculated or co-inoculated with strain M20. Likewise, seedlings which had been inoculated with strain M20 had significantly higher levels of DHZR, regardless of whether M20 was singly-inoculated or co-inoculated with one of the Paenibacillus strains. Table 6: Levels of indole-3-acetic acid (IAA) and dihydrozeatin riboside (DHZR) in lodgepole pine root tissue Treatment pmol IAA/ mg dry root3 Significant differences (a<0.05)b pmol DHZR/ mg dry root Significant differences (a<0.05)b Ratio IAA/DHZR Week 6 roots Uninoculated control < 1.28 — 0.47 a — M20 < 1.36 — 0.84 ab — L6 < 1.08 — 0.62 a — Pw-2 < 1.13 — 0.75 ab — L6 + M20 < 1.25 — 0.90 ab — Pw-2 + M20 < 1.24 — 0.97 ab — Week 9 roots Uninoculated control 0.31 be 0.82 ab 0.38 M20 0.41 cd 0.96 ab 0.43 L6 0.37 c 0.80 ab 0.47 Pw-2 0.38 c 0.76 ab 0.50 L6 + M20 0.30 be 0.65 ab 0.46 Pw-2 + M20 0.30 be 0.91 ab 0.33 Week 12 roots Uninoculated control 0.17 a 0.59 a 0.29 M20 0.22 ab 1.62 cd 0.14 L6 0.50 de 0.67 ab 0.74 Pw-2 0.52 e 0.65 ab 0.79 L6 + M20 0.57 e 1.79 d 0.32 Pw-2 + M20 0.58 e 1.21 be 0.48 aIAA concentrations in week 6 root extracts could not be calculated since virtually all the data points did not fall within the range of the standard curve. A modified, more sensitive curve was used for week 9 and 12 root extracts. "Treatments sharing a lowercase letter in common are not significantly different. 42 When the ratio of IAA/DHZR was calculated for each treatment, differences were noted between treatments in roots collected 12 weeks after inoculation. Roots singly-inoculated with a Paenibacillus strain had an IAA/DHZR ratio of above 0.7, while roots inoculated with a Paenibacillus strain and M20 had hormone ratios of about half that amount (Table 6). Roots singly-inoculated with strain M20 had the lowest IAA/DHZR ratio (Table 6). 3.4 Discussion Under the conditions of the experiment, Pseudomonas fluorescens strain M20 significantly decreased the extent of Paenibacillus polymyxa strain Pw-2-mediated increases in lateral root formation and root biomass accumulation at 12 weeks post-inoculation, the latter parameter to the point where there was no significant difference in root biomass accumulation between unbacterized seedlings and those inoculated with Pw-2 and M20 (Fig. 7). This phenomenon occurred despite the fact that strain M20 itself appears to act as a PGPR, stimulating increases in shoot biomass accumulation, root elongation and lateral root formation (Fig. 7), and despite the fact that levels of Pw-2 were steadily increasing in the rhizosphere of seedlings co-inoculated with Pw-2 and M20 (Fig. 6a). The density of Pw-2 in the rhizosphere of seedlings co-inoculated with Pw-2 and M20 was lower at weeks 6 and 9 after inoculation than in the singly-inoculated Pw-2 treatment (Fig. 6a). If one assumes that increased growth in PGPR-inoculated plants is proportional to the density of PGPR colonizing the rhizosphere, one might explain this result: an early deficiency in the rhizospheric density of Pw-2 in the co-inoculated treatment could lead to a deficiency in the extent of growth which is only observed after a delay. Delays in growth promotion responses by Paenibacillus PGPR have been observed before in conifers (Holl and Chanway 1992). Unfortunately, this scenario does not explain the increased extent of growth promotion observed in seedlings co-inoculated with Pw-2 and M20, relative to the Pw-2 singly-inoculated treatment, at weeks 6 and 9 after inoculation, when Pw-2 levels are lower. PGPR which become ineffective at elevated densities have been described (Frey-Klett et al. 1999), but the data does not suggest that strain Pw-2 is similar to these, since the density of Pw-2 in the rhizosphere of pines singly-inoculated with Pw-2 or co-inoculated with Pw-2 and M20 was identical 12 weeks after inoculation. A similar result was seen in the experiments in Chapter 2, where Pw-2 was less able to promote shoot biomass accumulation in pines when Curtobacterium flaccumfaciens strain PF322 was present as a co-inoculant, despite the fact that no decreases in rhizospheric or endophytic root colonization by Pw-2 were observed in the co-inoculated treatment relative to the Pw-2 singly-inoculated treatment (Figs. 1,2,4 in Chapter 43 2). It seems that the extent of lodgepole pine growth promotion mediated by strain Pw-2 cannot be predicted solely from the average population density of Pw-2 in the rhizosphere of inoculated seedlings. It is odd that the initial increases in growth observed at weeks 6 or 9 in seedlings co-inoculated with Pw-2 and M20, relative to seedlings singly-inoculated with Pw-2, did not result in increased growth at week 12. It is possible that the conditions under which the seedlings are grown are responsible: each plant tube is a closed system containing a finite amount of mineral nutrients, water and space in which to grow. These resources are depleted as the seedlings grow, leaving older seedlings, which require more water, nutrients and space, with the least amount of these resources. It is possible that the effect of these potential stresses on the physiology of the plant may alter its ability to respond to various combinations of bacteria. P. fluorescens strain M20 had no effect on the ability of P. polymyxa strain L6 to promote plant growth. Seedling growth was identical for all measured parameters at all times in treatments where L6 was singly-inoculated, and those where L6 was co-inoculated with M20 (Fig. 7). This phenomenon occurred despite the fact that at 9 and 12 weeks after inoculation, the rhizosphere population density of strain L6 was significantly lower in the L6 + M20 co-inoculated treatment than in the L6 singly-inoculated treatment (Fig. 6a). A similar result was observed in Chapter 2, where there were no significant differences in root elongation, lateral root formation or root or shoot biomass accumulation between seedlings singly-inoculated with strain L6, and those co-inoculated with L6 and C. flaccumfaciens strain PF322, despite the fact that rhizosphere populations of L6 had decreased in the co-inoculated treatment. The extent of lodgepole pine growth promotion mediated by strain L6 cannot be predicted from the population density of L6 in the rhizosphere of inoculated seedlings. Effects of bacterial inoculants on root hormone levels Inoculation with P. polymyxa strains L6 or Pw-2 resulted in similarly elevated levels of IAA in lodgepole pine roots at 12 weeks after inoculation, regardless of whether strain M20 was present as a co-inoculant (Table 6). Inoculation with strain M20, on the other hand, resulted in similarly elevated levels of the cytokinin DHZR at 12 weeks after inoculation, regardless of whether strains L6 or Pw-2 were also present (Table 6). In contrast, the root IAA content of loblolly pine seedlings inoculated with Bacillus subtilis strain GB03 was not significantly greater than in unbacterized control seedlings, and only a transient increase (observed in seedlings harvested at 6 weeks after emergence, but not at 12 weeks after emergence) in root IAA was noted in seedlings inoculated with Bacillus pumilis strain INR7 (Vonderwell 1998). 44 Both P. polymyxa strains L6 and Pw-2 are known to produce IAA (Srinivasan et al. 1996), and while the ability of strain M20 to produce DHZR or other cytokinins has not been evaluated, many cytokinin-producing rhizobacteria, including P. polymyxa (Timmusk et al. 1999) and Pseudomonas spp. (Frankenberger and Arshad 1995) have been identified. It is possible that these bacteria affect root hormone levels by producing plant hormones which are then absorbed by the root. The ability of strains GB03 and INR7 to produce IAA is not known, nor has the mechanism by which these strains interact with plants been elucidated. It is possible that these strains differ from P. polymyxa strains L6 and Pw-2 in how they interact with plants and affect root IAA levels. It is also possible that different pine species respond differently to inoculation with Paenibacillus PGPR, or that pines will respond differently to bacterial inoculants under differing experimental conditions (the loblolly pine seedlings described above were grown under greenhouse conditions). Auxins and cytokinins have different effects on root development, with exogenously supplied auxin generally promoting lateral root formation, and exogenous cytokinin inhibiting it (Chariton 1996). Atzmon et al. (1997) found that a high ratio of IAA/ cytokinin resulted in the formation of lateral roots in Pinus pinea seedlings. Seedlings singly-inoculated with strain Pw-2 had a ratio of root IAA/DHZR roughly twice as large as seedlings co-inoculated with both Pw-2 and M20 at 12 weeks after inoculation (Table 6). It is possible that the poorer growth of seedlings co-inoculated with both Pw-2 and M20, compared to seedlings inoculated with Pw-2 alone (Fig. 7), may be explained in part by differences in the ratio of root IAA/DHZR observed in these treatments (Table 6). The alteration in IAA/DHZR ratio cannot entirely explain the difference in growth between these treatments, however, for seedlings inoculated with M20 alone possessed the lowest root IAA/DHZR ratio at 12 weeks after inoculation (Table 6), yet seedlings singly-inoculated with M20 often grew as well as seedlings inoculated with either Paenibacillus strain, treatments which resulted in high IAA/DHZR ratios (Fig. 7, Table 6). Growth of seedlings inoculated with L6 and co-inoculated with both L6 and M20 was also similar, in spite of the fact that root IAA/DHZR ratios were quite different in these seedlings at 12 weeks after inoculation (Fig. 7, Table 6). DHZR is only one of many different forms of cytokinins or cytokinin conjugates, which may be metabolized from one form to another (Eklof et al. 1996, Sembdner et al. 1994). While the ELISA assay used can detect some non-DHZR cytokinins or cytokinin conjugates (such as dihydrozeatin, c/s-zeatin riboside, c/s-zeatin and dihydrozeatin-O-glucoside) via cross-reactivity (Phytodetek DHZR kit instructions, Agdia Inc., Elkhart, IN 46514), the root DHZR levels measured in this study are not necessarily representative of the total cytokinin content or cytokinin activity in the roots. It is possible that an increase in the level 45 of a cytokinin not well detected by the assay (e.g. isopentenyl adenine) occurred, but was not detected. Not every cytokinin present is necessarily involved in a plant physiological phenomenon (e.g. Wagner and Beck 1993), and to gain a complete picture of the effects of bacterial inoculants on root hormone levels, a variety of cytokinins ought to be analyzed. Also, other signals that might be involved in determining root growth, including other phytohormones, carbohydrates or mineral nutrients (Charlton 1996, Aiken and Smucker 1996), were not monitored in these experiments. While auxins are known to affect root development (Charlton 1996), is thought that cytokinin synthesis in roots can promote protein and chlorophyll synthesis in the shoot, leading to improved shoot development and more carbon fixation via photosynthesis (Aiken and Smucker 1996), and levels of trans-zeatin cytokinins are thought to control biomass partitioning in nettles (Wagner and Beck 1993). Shoot biomass accumulation was numerically greatest in seedlings singly-inoculated with strain M20 at 12 weeks after inoculation (Fig. 7). It is possible that the low ratio of IAA/ DHZR in the roots of these seedlings resulted in improved shoot growth as a consequence of increased cytokinin availability, and better root growth as a consequence of increased photosynthate availability rather than elevated levels of root auxin. However, such a scenario would not explain why unbacterized control roots, which also had a low IAA/DHZR ratio, also had significantly smaller shoots than all the other treatments (Fig. 7, Table 6). Events in seedling and root development are determined by levels of a variety of stimulatory and inhibitory signals, plant hormones in particular. The inoculation of pine seeds with a rhizobacterium can result in significant changes in the root hormone content of those seedlings, regardless of whether another bacterium is also present (Table 6). When two rhizobacteria which affect root hormone levels are co-inoculated (i.e., strain M20 and Paenibacillus strains L6 or Pw-2), the effects of both are seen in the plant (Table 6). Under the conditions of these experiments, co-inoculation of a Paenibacillus PGPR with strain M20 may result in no change in seedling growth and development, or may actually inhibit seedling growth and development relative to treatments in which the PGPR is present alone. The basis of the inhibition does not seem to relate to the average population density of PGPR present in the rhizosphere. It may be related to changes in root hormone ratios, which were observed to differ between singly- and co-inoculated treatments (Table 6). Further work, involving the quantification of a variety of auxins and cytokinins, as well as other phytohormones and conjugates, in bacterially-inoculated root tissues would be required to answer this question satisfactorily. 46 Chapter 4: Spatial colonization of the root surface by Pseudomonas fluorescens M20 and Paenibacillus PGPR, applied as single- or co-inoculated treatments 4.1 Introduction Lodgepole pine seedlings co-inoculated with P. polymyxa PGPR strain Pw-2 and P. fluorescens strain M20 tend to be smaller than seedlings singly-inoculated with strain Pw-2 alone, and the basis of this inhibition does not relate to the average population density of Pw-2 present in the rhizosphere, as measured via conventional dilution plating techniques (Chapters 2,3). The root surface is not uniformly colonized by bacteria (e.g. Chin-A-Woeng et al. 1997, Dandurand et al. 1997, Hansen et al. 1997, Troxler et al. 1997). It is possible that a PGPR such as strain Pw-2 cannot function well unless it is able to colonize specific areas, or microsites, on the root surface, such as areas near the root tip or at sites of lateral root emergence. The presence of other rhizobacteria, which may compete with strain Pw-2 for nutrient resources or space on the root surface, may reduce the ability of Pw-2 to colonize these specific microsites, and thereby reduce the ability of Pw-2 to promote plant growth. Indirect immunofluorescence techniques, in which strain-specific antibodies bind to the bacterium, and are identified by a fluorescent compound conjugated to a secondary antibody which binds to the constant (Fc) region of the first, have been successfully used to identify various bacterial strains on or within plant roots (e.g. Chin-A-Woeng etal. 1997, Shishido etal. 1999, Schloter et al. 1993, Troxler et al. 1997), and were used in conjunction with confocal scanning laser microscopy (CSLM). Samples viewed using CSLM may remain in aqueous solutions, which means that tissues do not need to be dehydrated and fixed in a hydrophobic matrix, a procedure which often results in damage or distortion of tissues (Flicker and White 1992). CSLM has an advantage over epifluorescence microscopy in that emitted light which is not in focus is eliminated, allowing clearer images to be recorded. This is of especial importance when dealing with plant tissues, which tend to produce autofluorescence which, when out of focus, can obscure details or objects of interest (Fricker and White 1992). The spatial colonization of areas near the root tip or lateral root junctions of lodgepole pine roots, inoculated with P. polymyxa strains L6 or Pw-2, P. fluorescens strain M20, or combinations of strains L6 with M20, or Pw-2 with M20, was observed via immunofluorescence techniques, and confocal scanning laser microscopy (CSLM). The objective of these experiments was to determine whether a change in the extent of PGPR-mediated plant growth promotion (monitored in Chapter 3) is related to a qualitative change in PGPR colonization of the root surface. 47 4.2 Materials and Methods Antigen preparation Duplicate sterile 250 mL Erlenmeyer flasks containing 50 mL of half-strength tryptic soy broth (TSB, 15 g/L) were inoculated with one of the following strains: P. polymyxa L6, P. polymyxa Pw-2, or P. fluorescens M20. Flasks were incubated at room temperature overnight with 150 rpm shaking. Cells were transferred to 50 mL oak ridge tubes (25 mL per tube) and pelleted via centrifugation (8000 rpm for 10 mins, at 4°C). Each pellet was washed twice in 25 mL of cold 0.85% (w/v) NaCI (saline) solution, using the centrifuge settings described above, then pellets were pooled and resuspended in 10 mL of cold saline solution. A strain test was conducted at this point to verify the identity of each strain and the lack of contamination in each suspended pellet: loopfuls of cell suspension were spread across TSA, Str, PAF and TTC-suc plates (see Chapter 3, Table 1 for details) and the plates incubated at room temperature and examined daily for up to 4 days. A small sample (0.5 mL) of each cell suspension was mixed with 0.5 mL sterile glycerol in a cryovial and kept at -80°C. The remainder of each suspension was thoroughly mixed with 10 mL cold 0.6% paraformaldehyde-saline solution (15 mL 4% paraformaldehyde in 0.1 M P 0 4 (pH 7.4) solution + 85 mL saline), to kill the cells without destroying surface antigens. Each suspension was allowed to sit for 2 days at 4°C, with occasional vortexing to re-mix the suspension. After two days, each suspension was centrifuged to pellet the cells, and each cell pellet was washed 2-3 times in 30 mL of saline in 50 mL oak ridge tubes at 8000 rpm for 10 minutes at 4°C in a Sorvall SS34 rotor. Cells were resuspended in 10 mM phosphate-buffered saline (PBS) solution (Shishido 1997) and adjusted to a cell density of 108 CFU7 mL ( A ^ of 0.3 for strain M20, and 0.8 for strains L6 and Pw-2) or 109 CFU/mL. The death of cells was verified by streaking loopfuls of each suspension across a TSA plate and incubating that at room temperature for one week. Killed cell suspensions were kept frozen in aliquots at -80°C until use. Polyclonal antiserum from rodents Polyclonal antisera were generated from rabbits and mice using standard methods (Ball etal. 1990). Blood samples (pre-bleeds) were collected from two female rabbits, which were then each injected with 1.0 mL of thawed M20 killed cell suspension (108 CFU/mL) without adjuvant on Day 0. On Day 15, each rabbit was inoculated with 0.6 mL of thawed M20 killed cell suspension (109 CFU/ mL) emulsified with 0.6 mL of Freund's incomplete adjuvant. On Days 26 and 39, blood samples (test bleeds) were collected from each rabbit. Serum was 48 immediately collected from each blood sample: blood was allowed to clot for 1 hr at 30°C, then clots were separated from the serum via centrifugation in a microfuge (2000 rpm for 5 min at 4°C). The serum was collected and kept frozen at -20°C until use. The sensitivity of serum from each test bleed to M20 killed cells as antigens was determined via an enzyme-linked immunosorbent assay (ELISA) within a few days of collection (see next section). On Day 43, the rabbits were killed and their blood was collected (final bleeds). The sensitivity of the serum collected from each rabbit to M20 killed cells as antigens, and the cross-reactivity of each serum with killed cells of P. polymyxa strains L6 and Pw-2 as antigens, was estimated via ELISAs. Blood samples (pre-bleeds) were collected from each of six female mice on Day 0, three of which were then injected subcutaneously with 0.2 mL of a 1:1 mixture of thawed L6 killed cell suspension (108 CFU/mL) emulsified with Freund's complete adjuvant. The remaining three mice were injected subcutaneously with 0.2 mL of a 1:1 mixture of thawed Pw-2 killed cell suspension (108 CFU/mL) emulsified with Freund's complete adjuvant. On Days 10, 24 and 37, each mouse received a 0.2 mL intraperitoneal injection of a 1:1 mixture of thawed L6 or Pw-2 (as appropriate) killed cell suspension (108 CFU/mL) emulsified with Freund's incomplete adjuvant. Serum was collected from mouse blood samples as indicated above for rabbits. On Days 17 and 33, blood samples (test bleeds) were collected from each mouse. The anti-L6 or anti-Pw-2 (as appropriate) antibody content of serum from each test bleed was determined via ELISAs within a few days of collection. On Day 45, the mice were killed and their blood was collected (final bleeds). The anti-L6 or anti-Pw-2 antibody titres were estimated via ELISAs in serum collected from mice injected with heterologous cells as antigens. The cross-reactivity of each serum with killed cells of the heterologous P. polymyxa strain and P. fluorescens M20 as antigens was also evaluated via ELISAs. Titration of polyclonal antisera Indirect ELISA analyses were conducted (McLaughlin and Chen 1990). Immulon 4 96-microwell plates (Dynex Technologies, Inc., Chantilly VA 20151) were coated with killed bacterial cells as follows: each well was filled with 100uL of killed bacterial cells suspended in 50 mM carbonate-bicarbonate buffer (coating buffer, pH 9.6), or with 100 pi of coating buffer only in "blank" wells. For P. polymyxa strains L6 and Pw-2, a 10 6 CFU/mL cell suspension provided optimal results. For P. fluorescens M20, a 105 CFU/mL cell suspension provided optimal results. Plates were allowed to dry uncovered overnight in a 37°C incubator. Coated plate wells were rinsed three times with 250 uL of 10 mM PBS and blotted dry on paper towels. Plates were then blocked by adding 200 pi of a 2% (w/v) powdered milk solution in PBS to each well. Plates were covered and incubated 1 hr at 37°C. Plates were subsequently rinsed and blotted dry as described previously. Plate wells were then filled with primary antisera, 100 uL per well. Serum samples were diluted in a 0.1% (w/v) powdered milk solution in 10 mM PBS immediately before use. ELISA analyses of first test bleeds included both pre-bleed and test bleed serum samples from each rodent, with the pre-bleed sera used to indicate levels of antigen-binding antibodies in each animal prior to inoculation with bacteria. Analyses of subsequent test bleeds included serum samples from each previous test bleed performed, as well as from pre=bleeds, kept separate for each rodent. An analysis of each final bleed was also conducted once in this manner. Rabbit serum samples were never combined. Mouse serum samples were not combined until after the sera had been analyzed and found to contain relatively similar amounts of cell-binding antibodies. In these tests, primary serum dilutions usually ranged from 1/500 -1/2000 for pre-bleeds and initial test bleeds, to 1/2000 -1/8000 for later test bleeds and final bleeds. The cell-binding antibody titre was estimated for each final bleed antiserum in two separate experiments. In the first, a range of serum dilutions (1/2000 - 1/32000) were used on a single plate coated with the same number of bacterial cells/ well (106 cells/mL for P. polymyxa, and 105 cells/ mL for P. fluorescens). In the second, a single serum dilution (1/4000) was used on a plate coated as described above, with a range of bacterial cell suspensions (from 105 or 10 6 cells/mL down to 5 cells/ mL) diluted in coating buffer. For each of these tests, pre-bleed antisera was included as a basis for comparison, and cells from all three bacterial strains were present on each plate so that heterologous binding could be evaluated. Aliquots of each dilution of each antiserum sample tested were added to three to eight wells. Plates were covered and allowed to incubate for 2 hr at 37°C. Plates were then rinsed three times with 10 mM PBS and blotted dry, as described earlier. Wells were then filled with 100 uL of secondary antisera, diluted immediately before use in 0.1% (w/v) powdered milk solution in 10 mM PBS. For wells filled with rabbit sera, goat anti-rabbit IgG secondary antibodies conjugated to horseradish peroxidase (HRPO) were used (1/4000 dilution). For wells filled with mouse sera, goat anti-mouse IgG antibodies conjugated to HRPO (1/3000 dilution) were used. Plates were covered and allowed to incubate for 1 hr at 37°C. Plates were then rinsed three times with 10 mM PBS and blotted dry, as described earlier. A tablet of 3,3'5,5'-tetramethyl benzidine (TMB) (Sigma Chemical Co., St. Louis, MO 63178) was dissolved in a mixture of 1 mL dimethylsulfoxide + 9 mL citrate=sodium phosphate buffer (pH 5.0). Each well was filled with 200 pi of this solution, then plates were covered and allowed to incubate at 37°C until significant colour development was observed (usually 20 min). 50 HRPO was inactivated by the addition of 50 ul of 2.5 M H 2 S 0 4 to each well. Colour development was measured at 450 nm. Evaluation of cross-reactivity between final bleeds and heterologous bacteria These tests were conducted via indirect ELISA analyses, as described above. Cross-reactivity was estimated for each final bleed antiserum by using a smaller range of antiserum dilutions (1/2000 - 1/8000) on a plate coated with each of the three bacterial cell suspensions (106 cells/mL for P. polymyxa L6 and Pw-2, and 105 cells/ mL for P. fluorescens M20). Preabsorption of rabbit antisera Whole killed P. polymyxa L6 and Pw-2 cells (10 mL of 108 CFU/ mL suspensions) were collected separately by centrifugation in 50 mL oak ridge tubes in a Sorvall SS34 rotor (8000 rpm for 10 min at 4°C). Each cell pellet was resuspended in 10 mL of a 2% (w/v) powdered milk solution in 20 mM NaP0 4 buffer (blocking solution, pH 7.2). Six 1 cm-long formaldehyde-treated (placed in 10 mL of 4% (v/v) formaldehyde in10 mM PBS at 4°C for 3 days, then rinsed three times in 8 mL PBS) segments of a lodgepole pine seedling root were added (two per tube) to three 1 mL aliquots of blocking solution. The bacterial cells and root pieces were incubated separately for 1 hr at 37°C in sealed Eppendorf tubes. The cell mixtures were then centrifuged as described previously and the cell pellets washed twice in 10 mL cold 20 mM NaP0 4 (wash buffer, pH 7.2), using the same centrifugation settings. Each cell pellet was resuspended in cold wash buffer to a cell density of 1.25 X 108 CFU/mL (or an A ^ o f 1.1), and kept at 4°C until use (same or next day). The root pieces were each transferred with forceps to fresh 1 mL aliquots of wash buffer in Eppendorf tubes, and gently rinsed by inversion of each tube for 30 seconds. This rinsing procedure was conducted three times for all roots. Root pieces were collected and kept in 2 mL wash buffer at 4°C until use (same or next day). 0.8 mL of rabbit polyclonal antiserum was mixed with 0.4 mL of each blocked bacterial cell suspension, the two blocked root pieces and 40 pL of blocking solution (for a final concentration of 0.1% (w/v) powdered milk), and allowed to sit for 22 hr at 4°C. Debris were separated from unbound antiserum by centrifugation for 10 min at 4°C in a Hemle microfuge (14000 rpm). Approximately 1.5 mL of supernatant was transferred to a clean tube. The protein content of the supernatant was determined spectrophotometrically (A 2 8 0) and compared to the protein content of a sample of wash buffer, and a sample of 0.1% (w/v) powdered milk in wash buffer. The supernatant was then subdivided into two 0.7 mL portions, and to each portion 2 blocked root pieces and 0.2 mL each of the L6 and Pw-2 blocked bacterial cell suspensions were added. Each suspension was mixed and allowed to sit at 4°C for 24 hr. Debris were separated from unbound antiserum as described previously. Supernatant samples were pooled, and the protein content of the serum (supernatant) was determined as described previously. The serum was then filtered with a 0.2 urn Millipore syringe filter. A sample (200 pi) of the preabsorbed serum was reserved and kept frozen at -20°C until use. The rest was subjected to IgG purification, as described below. Mouse antisera were not subjected to preabsorption since a high level of cross-reactivity was expected to occur between antisera raised against L6 or Pw-2, and antigens in cells of the heterologous P. polymyxa strain. It was feared that the antibody titre of each antiserum would be too low after preabsorption to be useful for detection of cells during confocal microscopy. Since an insufficient quantity of serum was available to both test the effects of the preabsorption procedure and reserve enough for use in microscopy, the preabsorption procedure was omitted. This was not expected to negatively influence the experimental results since Pw-2 and L6 were never present in the same treatment, and neither of the antisera raised against the P. polymyxa strains demonstrated a significant amount of cross-reactivity with cells of P. fluorescens M20 in preliminary tests (data not shown). Purification of immunoglobulin G (IgG) from rabbit and mouse antisera Purification of IgG from polyclonal antisera was conducted using 1 mL HiTrap Protein A affinity columns (Pharmacia Biotech), according to the manufacturer's instructions. A separate column was used for each antiserum. All antisera and buffers were filtered with 0.22 pm Millipore filters prior to use, to remove debris or undissolved particles. 0.5 mL samples of each mouse polyclonal antiserum, and around 1 mL of preabsorbed rabbit antiserum, were used. Each serum was diluted by 1/5 in 20 mM NaP0 4 (wash buffer, pH 7.2) prior to injection into the equilibrated columns, and samples were passed through the column 10-12 times (eluent was saved each time, and passed through the column again). After the final pass, the eluent was reserved. The Am of a sample of wash buffer ("baseline") was determined spectrophotometrically. Wash buffer was then passed through the column, the eluent collected in fractions, and the protein content (A28o) of each fraction determined spectrophotometrically. Wash buffer was passed through the column until the A280 of the eluent had decreased to the baseline value (about 8-10 mL). All the wash fractions were reserved. Elution buffer (0.1 M citric acid, pH 3, 3.5 mL) was passed quickly through the column, which was rinsed immediately by passing 10 mL of wash buffer through the column. Elution fractions (0.5 mL each) were added to Eppendorf tubes already containing 150 uL 1 M Tris-HCI (pH 9). Each elution fraction, and the first five post-elution wash fractions (1 mL each), were reserved. The A 2 8o of each elution fraction and reserved post-elution wash fraction were determined spectrophotometrically. Elution fractions containing significant amounts of protein (IgG) were frozen, lyophilized, pooled and resuspended in 0.5 mL 0.02% aqueous sodium azide. The 52 purified IgG fractions, and each sample reserved during the purification process, were stored at -20°C until use. Titration of polyclonal IgG samples In order to determine whether preabsorption and IgG purification procedures altered the titre of antibodies that bind homologous antigens in each polyclonal serum, indirect ELISAs were conducted. The procedure was as described previously, with the following modifications. Primary antisera samples used included: (1) unpurified antiserum, (2) antiserum sampled immediately after preabsorption (rabbit antiserum only), (3) pre-elution wash fractions, (4) post-elution wash fractions and (5) the purified (eluted) IgG samples. Dilutions ranged from 1/1000 -1/4000 for samples expected to contain IgG (treatments 1,3 and 5 above) to 1/500 - 1/1 (no dilution) for wash samples. These samples were used with plates coated with homologous antigens (10 5CFU/ mL of M20 killed cells, or 106 CFU/mL of L6 or Pw-2 killed cells, as described previously). Wells that contained mouse primary antisera were subsequently filled with 100 pL aliquots of a 1/3000 dilution of goat anti-[mouse IgG Fab' fragment] conjugated to alkaline phosphatase (PhoA). Wells that contained rabbit primary antisera received a 1/3000 dilution of goat anti-[rabbit IgG] conjugated to PhoA. The substrate used was p-nitrophenyl phosphate (pNpp), freshly prepared from Sigma Fast™ tablets (Sigma Chemical Co., St. Louis, MO 63178) according to the manufacturer's instructions. Plates were incubated covered at room temperature in the dark until significant colour development had occurred (20-30 min). The enzymatic reaction was stopped by the addition of 50 pL of 3 M NaOH, and the colour development in each well measured at 405 nm with a Dynatech MR700 plate reader. Cross-reactivity of polyclonal IgG preparations with heterologous bacteria These tests were conducted as described above, to determine whether preabsorption and IgG purification procedures decreased the number of antibodies that would cross-react with heterologous antigens except that plates were coated with both heterologous and homogenous antigens (same concentrations as above), and only the serum samples which were expected to contain IgG (unpurified antiserum, pre-elution wash fractions, and purified (eluted) IgG samples) were tested. Samples were diluted as described above. CSLM experiments Bacterial colonization of the root surface was examined in three different experiments. In the first, seedlings harvested 7 weeks after inoculation were used. Root segments (~ 1 cm long) containing root tips or lateral root junctions were randomly sampled from 3-4 seedlings per treatment. Root tip segments from the seedlings in the same treatment were pooled to 53 economize on reagents, and stained. Segments containing lateral root junctions were likewise pooled and stained. The second experiment was identical to the first, only seedlings harvested 13 weeks after inoculation were used. In the third experiment, two seedlings from each treatment, harvested 13 weeks after inoculation, were used. Roots were photographed prior to segmenting, and six segments, from areas ranging from the tip to the base of the root, were labeled on the photograph and then sampled. These root segments (72 in total) were handled individually. Preparation of slides for CSLM Lodgepole pine seedlings were grown from surface-sterilized, bacterized seeds in plant tubes under gnotobiotic conditions, as described for the growth promotion/ root colonization assays in Chapter 3. Seeds were treated with the following: (i) sterile buffer (ii) P. polymyxa strain L6, (iii) P. polymyxa strain Pw-2, (iv) P. fluorescens strain M20, (v) a mixture of strains L6 and M20, and (vi) a mixture of strains Pw-2 and M20. Seedling roots were first fixed in formaldehyde: plants were gently shaken from their tubes, rinsed in tapwater, and the roots were severed from the shoots. Roots were immediately submerged in 10-15 mL of a 3% formaldehyde solution in 0.1 M P 0 4 buffer (wash buffer, pH 7.4) in a petri dish. Dishes were covered and sealed with parafilm, and roots were allowed to sit in this solution overnight in a fume hood. The following day, roots were rinsed three times in 10-15 mL cold wash buffer, photographed (in the third experiment only), and segmented by hand into sections roughly 1 cm long with new, thin shaving razor blades. Root segments were blocked and immunologically stained using 24-well tissue culture trays. Segments (from one up to five per well) were placed into wells containing 1 mL of a 2% (w/v) powdered milk solution in wash buffer, and allowed to sit without agitation for 1 hr at room temperature. Plates were covered to prevent evaporation. Segments were then removed from the milk solution with forceps, or a small scoop made from a circle of wire mesh glued to a wire handle, and rinsed by placing them in 1 mL fresh wash buffer and gently tapping the plates. Segments were rinsed twice more in this fashion. At this point, at least two root segments (one root tip and one segment containing lateral root junctions) were set aside from the unbacterized control treatment seedlings. These segments would be used to evaluate the amount of background fluorescence in untreated root tissue, and were not treated further before mounting onto slides. At least two root segments (one root tip and one segment containing lateral root junctions) were set aside from each treatment. These segments would be exposed to secondary antisera only, in order to evaluate the degree of non-specific binding of secondary antibodies to root tissues. The remaining root segments were transferred to solutions containing primary antisera, which were diluted immediately before use in 0.1% (w/v) powdered milk in wash buffer. Segments from seedlings inoculated with single bacterial strains (L6, Pw-2 or M20) were suspended in 0.5 mL (for multiple segments, in wells) or 0.2 mL (for single segments, in Eppendorf tubes) of diluted homologous unpurified (for randomly sampled root segments taken from roots grown for 13 weeks only) or purified polyclonal IgG (1/500 dilution for anti-L6 or anti-Pw-2 sera, 1/1000 dilution for anti-M20 serum). Segments from seedings co-inoculated with two bacterial strains were suspended as indicated above in a 1:1 mixture of the homologous diluted antisera, each diluted as indicated above. At least two segments (one root tip and one segment containing lateral root junctions) from unbacterized seedlings were suspended in one of each of the three primary antiserum dilutions: anti-L6, anti-Pw2, and anti-M20, diluted as indicated above. Segments were allowed to sit without agitation for 2-3 hr at room temperature. When tissue culture plates were used for staining, they were covered and sealed in plastic baggies to prevent evaporation. Segments were then rinsed three times in wash buffer, as described previously. All segments exposed to primary antisera, and the segments set aside for exposure to secondary antisera only, were transferred to solutions containing secondary antisera. These antisera were diluted (1/100) immediately before use in 0.1 % (w/v) powdered milk in wash buffer, and protected from exposure to light as much as possible. Monoclonal goat anti-[mouse IgG] conjugated with fluorescein isothiocyanate (FITC, Rockland Inc., Gilbertsville PA 19525) was used for the following: all root segments from seedlings bacterized with only strain L6, or only strain Pw-2 (whether exposed to primary antisera or not), segments from unbacterized seedlings exposed only to antNL6 or anti-Pw-2 antisera, and some segments from unbacterized seedlings not exposed to primary antisera. Monoclonal goat anti-[rabbit IgG] conjugated with Cyanine 5.29-Osu (Cy5™, Rockland Inc., Gilbertsville PA 19525) was used for the following: all root segments from seedings bacterized with only strain M20, all segments from unbacterized seedlings exposed only to anti-M20 antiserum, and the remaining segments from unbacterized seedings not exposed to primary antisera. A 1:1 mixture of each secondary antiserum was used for the following: all root segments from seedlings bacterized with a combination of bacteria, and segments from unbacterized seedlings exposed to a 1:1 mixture of an\\-Paenibacillus strain: anti-M20 antisera. Segments were incubated in the dark without agitation at room temperature for at least one hour, in either 0.5 mL of solution (in tissue culture plate wells) or 0.2 mL of solution (in Eppendorf tubes). When plates were used, they were covered and placed in sealed baggies to 55 prevent evaporation. Segments were then rinsed three times in wash buffer, as described previously. Segments were placed in the wells of 1 mm deep single-well glass slides, covered with a glycerol-based mounting solution containing antifade reagents (4% (w/v) n-propyl gallate (3,4,5-trihydroxybenzoic acid n-propyl ester) and 2.5% (w/v) DABCO (1,4 diazabicyclo [2.2.2] octane) in a mixture of 80% (v/v) glycerol and 20% (v/v) 50mM carbonate-bicarbonate buffer (pH 9.6)) and a cover slip. Care was taken to minimize exposure of root segments to light. Finished slides were stored at 4°C in the dark until use. Slide viewing Slides were viewed using a BioRad MRC 1024 3-channel confocal scanning laser imaging system and BioRad COMOS software (v. 7.0/7.1) (BioRad Microscience Ltd., Hemel Hempstead HP2 7TD UK). The preprogrammed COMOS method for 3*channel (green, red and far-red) excitations was used. This method adjusts the microscope's filters for the detection of three different, widely used fluorochromes: FITC (green channel, absorption maximum at 495 nm and emission at 528 nm), Texas Red™ (red channel, absorption maximum at 596 nm and emission at 620 nm) and Cy5™ (far-red, absorption maximum at 650 nm and emission at 667 nm). Plant tissues naturally autofluoresce (Fricker and White 1992). While they absorb and re* emit light at a variety of wavelengths, they strongly absorb yellow (around 596 nm) light and re-emit it as red (around 620 nm) light, allowing them to be clearly viewed with the red channel. The channel settings were generally as follows: for green, the gain was 1120, blacklevel -15, and the iris 2.0. For red, the gain was 1250, blacklevel - 5 and iris 2.5. Forfar red, the gain was 1500 (maximum), the blacklevel 0 o r -5 , and the iris 2.5. The vertical distance between sections (z-step) was usually 1.08 urn, although for thick samples a larger z-step (up to 8 pm) was sometimes used, to reduce collection time and image file size. Images were collected using a Kalman filter set (n = 3). The image from the far-red channel was collected first, since Cy5 emissions tend to be fairly dim, and would fade most quickly during prolonged exposure to light. The image from the green channel was collected next, then the image from the red channel. Collection parameters were kept the same for the images in all three channels. Images from all three channels were recorded for each control slide. For experimental slides, images were recorded only from the channels observed to contain information, in order to save time and conserve disk space. Scale was determined by noting the height and width of the image in pixels, then superimposing a scale bar (10-100 pm) over the onscreen image and printing a copy of the image with a Codonics thermal video printer (Seikosha Co, Ltd., Tokyo, Japan). 56 Digital imaging ' Digital images (BioRad PIC files, consisting of a three-dimensional "stack" of two-dimensional slices) were temporarily saved to Zip discs and archived for long-term storage on CD-ROMs. Flattened, two-dimensional "projections" (in which the slices in the image stack are superimposed atop one another) were made of each image, which were then saved as tagged image format (.TIF) graphic files using Confocal Assistant (v. 4.02) software. Two-dimensional images from the red, far-red and green channels were superimposed in different colours (red channel information is shown in blue, green channel information in green, and farmed information in red) using Adobe PhotoShop (v.6.0) software. This procedure is illustrated in Figure 8. Fluorescence associated with FITC or Cy5 labels could generally be distinguished from plant tissue autofluorescence due to the fact that plant tissue autofluorescence was usually noted in more than one wavelength. In Figure 8, this is illustrated by the circled object (probably a plant cell nucleus) that re-emits both red and far-red light, and is tinted a purplish colour in the final picture as a result. Flattened projections and finished colour pictures were also archived for long-term storage on CD-ROMs. 4.3 Results Purification of polyclonal antisera After optimizing the assay conditions (number of cells used to coat each well, coating buffer, etc.) and verifying that target-specific antibodies remained in each preparation following the various purification procedures (data not shown), a series of ELISA tests were conducted to determine the cross-reactivity of each polyclonal antiserum before and after preabsorption/IgG purification treatments (Tables 7-9). Results indicate that the titre of unwanted antibodies reacting with heterologous antigens decreased after these procedures, but that the titre of desired antibodies reacting with homologous antigens also decreased by roughly the same amount, indicating that the overall specificity of each antiserum was not improved by the purification methods used. For antiserum raised against strain M20, the titre of M20-binding antibodies dropped by roughly 70% after preabsorption and IgG purification, and the titre of antibodies binding heterologous strains L6 or Pw-2 by about the same amount (Table 7). For antiserum raised against strain L6, the situation was similar: the titre for homologous and heterologous bacterial cells dropped by 80% or more, a decrease perhaps larger than it should be given that some IgG from this serum was lost in a spill (Table 8). For antiserum raised against strain Pw-2, the titre of Pw-2-binding antibodies dropped by about 70%, while the titre of L6-binding antibodies dropped by an estimated 90% (Table 9). It should be noted that decreases in antibody titre reported here and in Tables 7-9 do not reflect the fact that each 57 Figure 8: Illustration of the process by which three two-dimensional images, each representing emitted light of different wavelengths, were combined into a single image and interpreted. Far-red, green and red light images were collected as described in the text, and the information in each assigned a different colour: red, green and blue, respectively. The images were then combined to produce a final, full-colour image. Plant tissue autofluorescence, brightest in the red light image, was distinguished from fluorescence produced by FITC or Cy5 labels on the basis of whether bright objects observed in the far-red or green light images were also present in the red light image. The circled object is an example of plant autofluorescence in more than one wavelength: it is observed in both the red and, more faintly, the far-red images. In the final image, it is tinted purple as a consequence. The white arrows indicate bright objects in the far-red or green images that are not present in the red image, and unlikely to be produced by autofluorescence. 59 antiserum was concentrated 3X following IgG purification. The concentrated IgG preparations were used in experiments. Confocal microscopy Figure 9 depicts a series of control slides, from the first experiment conducted, in which root segments were randomly sampled from seedlings harvested 13 weeks after inoculation. Similar results were observed for the control slides from the remaining two experiments, in which root segments were randomly sampled from seedlings harvested 7 weeks after inoculation, and in which root segments harvested 13 weeks after inoculation, from different locations along the root, were sampled (data not shown). Figures 10 and 11 depict randomly-sampled root segment surfaces from each treatment, on roots taken thirteen and seven weeks after inoculation, respectively. Figure 12 depicts surfaces of root segments sampled thirteen weeks after inoculation, in six different locations, ranging from the root tip to areas close to the root base. Figure 13 is a reproduction of a thermal print depicting an on-screen, magnified green channel image of bacterial cells from a root inoculated with P. polymyxa strain Pw-2 next to a 2 pm scale bar. Table 7: Binding of anti-M20 antiserum (A405) before and after purification by preabsorption and IgG collection, to cells of each bacterial strain (P. fluorescens M20 and P. polymyxa L6 and Pw-2). M20 cells3 L6 cellsb Pw-2 cells'3 Serum dilution Before After0 Before After0 Before After0 1.379 0.987 0.125 0.050 0.503 0.389 1/125 1.487 0.508 0.062 0.071 0.229 0.109 1/250 1.518 0.508 0.039 0.012 0.121 0.082 1/500 1.513 0.494 0.022 0.010 0.068 0.030 1/1000 1.383 0.412 0.017 0.006 0.038 0.016 1/2000 1.010 0.362 0.011 0.004 0.017 0.008 1/4000 0.702 0.232 0.007 0.003 0.007 0.005 1/8000 0.393 0.145 0.008 0.002 0.003 0.003 1/16000 aPlate wells coated with bacteria cell suspension of 105 CFU/ mL. "Plate wells coated with bacterial cell suspension of 106 CFU/mL. °Antisera were concentrated 3X following purification. For purposes of comparison to unpurified sera, A405 after purification are reported at 1/3 the actual values. 60 Table 8: Binding of anti-l_6 antiserum (A405) before and after purification by IgG collection, to cells of each bacterial strain {P. fluorescens M20 and P. polymyxa L6 and Pw-2). M20 cells8 L6 cells15 Pw-2 cells" Serum dilution Before After0 Before After0'* Before After0 0.015 0.003 1.460 0.361 0.601 0.100 1/125 0.009 0.002 1.359 0.207 0.468 0.060 1/250 0.009 0.001 1.044 0.117 0.317 0.038 1/500 0.009 0.003 0.863 0.062 0.218 0.029 1/1000 0.008 0.003 0.517 0.031 0.112 0.017 1/2000 0.003 0.003 0.243 0.017 0.069 0.013 1/4000 0.007 0.005 0.113 0.010 0.046 0.009 1/8000 0.009 0.004 0.069 0.008 0.039 0.007 1/16000 aPlate wells coated with bacterial cell suspension of 10 CFU/ mL. "Plate wells coated with bacterial cell suspension of 106 CFU/mL. °Antisera were concentrated 3X following purification. For purposes of comparison to unpurified sera, A405 after purification are reported at 1/3 the actual values. dSome of the purified IgG was lost during a spill, which may partially account for the low titre. Table 9: Binding of anti-Pw-2 antiserum (A405) before and after purification by IgG collection, to cells of each bacterial strain (P. fluorescens M20 and P. polymyxa L6 and Pw-2). M20 cells3 L6 cells" Pw-2 cells" Serum dilution Before After0 Before After0 Before After0 0.001 0.001 0.253 0.016 1.524 0.510 1/125 0.000 0.001 0.145 0.009 1.472 0.480 1/250 0.005 0.000 0.100 0.003 1.299 0.345 1/500 0.001 0.000 0.056 0.003 0.986 0.217 1/1000 0.000 0.000 0.031 0.003 0.749 0.128 1/2000 0.000 0.001 0.015 0.000 0.460 0.067 1/4000 0.000 0.001 0.004 0.000 0.271 0.033 1/8000 0.000 0.001 0.004 0.000 0.148 0.018 1/16000 3Plate wells coated with bacteria cell suspension of 105 CFU/ mL. "Plate wells coated with bacterial cell suspension of 10 s CFU/mL. cAntisera were concentrated 3X following purification. For purposes of comparison to unpurified sera, A405 after purification are reported at 1/3 the actual values. Colonization patterns on singly-inoculated roots Paenibacillus cells were easier to locate on younger (taken 7 weeks after inoculation) root segments than on older ones (taken 13 weeks after inoculation). When Paenibacillus spp. cells were identified on the root surface, they were generally sparse and scattered, with the majority of green signal being associated with debris (sloughing cells, humic substances and so on still attached to the root), with transparent, mucilaginous material at or slightly above (2-6 61 urn) the root surface, or with clusters in the grooves between epidermal cells. It was difficult to discern whether Paenibacillus spp. were root tip colonizers since sometimes there was a great deal of green autofluorescence from plant cell nuclei and organelles in these areas (e.g. Fig. 10j). Fig. 11a depicts L6 cells associated with sloughing root cells or other debris on the root surface, while Figs. 10a, 10b, 11b, 11c, 12a and 12b depict L6 cells on a lateral root (detail, Fig. 11c) or close to lateral root junctions. Figs. 11d and 11e depict Pw-2 cells on the primary root surface, found mostly in grooves between epithelial cells (this is more visible in 11e). Figs. 10c and 10d show Pw-2 cells on a lateral root and between two root hairs, respectively. Fig. 12e depicts Pw-2 cells on the tip of a lateral root, while Figs. 12f and 12g (detail in 12g) depict Pw-2 on the lateral root surface, in a region about 1 cm above the root tip. P. fluorescens strain M20 seemed to be more abundant than the Paenibacillus spp. on the root surface, and was easier to locate. No differences were observed in strain M20 colonization of roots from seedlings harvested at 7 or at 13 weeks. On roots singly-inoculated with strain M20, M20 cells were usually associated with areas at (Fig. 11f) or near root tips (Figs. 10e and 11g). At lateral root junctions, M20 was uniformly absent (e.g. Figs. 10f, 12m). Figs. 12j and 12k also depict M20 cells on or near root hairs. Colonization patterns on co-inoculated roots On roots co-inoculated with strains L6 and M20, cells of both species could be found co-existing in the same microsites (Figs. 10g, 10h, 11 h-11k, 12n, 12o) on the root surface. On roots co-inoculated with strains Pw-2 and M20, the same phenomenon was noticed (Figs. 10i, 111-110,12r, 12s) although it was somewhat more difficult to find Pw-2 cells on these roots than it had been to locate L6 cells. Interestingly, strain M20 could be found at lateral root junctions of co-inoculated roots (e.g. Figs. 10g-10i, 11h-11m, 12q), where it had been absent from these sites on roots that had been singly-inoculated with M20. No improvements to the extent of Paenibacillus spp. root colonization were noted in the co-inoculated seedling treatments, relative to treatments in which a single Paenibacillus strain had been applied. Colonization patterns on root segments of different ages The images depicted in Fig. 12 suggest that strains L6, Pw-2 and M20 all colonize younger tissues (areas closest to the root tip) more abundantly than older tissues. In each treatment, fewer or no bacteria were observed on older root segments, although one root co-inoculated with L6 and M20 seemed to be colonized fairly heavily in an older section (Fig. 12q). Fig. 12u depicts some putative M20 cells on a lateral root junction of a root from a seedling that had been co-inoculated with strains Pw-2 and M20. No Pw-2 cells seem to be present in Fig. 62 Figure 9: Staining controls, representative of control images viewed for each experiment (only one set of images is depicted to conserve space), from randomly sampled root segment surfaces, thirteen weeks after inoculation. These controls are used to determine the extent of non-specific binding (i.e. binding of primary or secondary antibodies directly to plant tissues, or binding of secondary antibodies to bacteria) in the samples. Emissions of red, green and far-red light Were collected for each image as described in the text and in Figure 8. Plant tissue (red autofluoresence) is depicted in blue, far-red light (including Pseudomonas fluorescens strain M20 and far-red autofluorescence) in red, and green light (including Paenibacillus polymyxa strains and green autofluoresence) in green. The absence of a red or green colour in these images indicates that no light emissions in the far red or green (respectively) wavelengths were recorded by the equipment. (a) Unbacterized root, no antibodies (b) unbacterized root, anti-L6 primary antibodies (1° abs) and secondary antibodies (2° abs) (c) unbacterized root, anti-Pw-2 1° abs and 2° abs (d) unbacterized root, anti-M20 1° abs and 2° abs (e) L6 singly-inoculated root, 2° abs only (f) Pw-2 singly-inoculated root, 2° abs only. The arrow indicates an object that might represent non-specific binding of these antibodies to Pw-2 cells. (g) M20 singly-inoculated root, 2° abs only (h) (h) root co-inoculated with L6 and M20, 2° abs only (i) root co-inoculated with Pw-2 and M20, 2° abs only. 50 Mtn ' * 0 66 Figure 10: Rhizobacterial colonization of randomly sampled root segment surfaces, thirteen weeks after inoculation. Plant tissue (red autofluoresence) is depicted in blue, far-red light signals (including Pseudomonas fluorescens strain M20 and far-red autofluorescence) in red, and green light signals (including Paenibacillus polymyxa strains and green autofluoresence) in green, as described in the text and illustrated in Figure 8. Appropriate primary and secondary antibodies were used in each case. Roots were singly-inoculated with: (a,b) P. polymyxa strain L6, (c,d) P. polymyxa Pw-2 or (e,f) Pseudomonas fluorescens M20, or co-inoculated with mixtures of (g,h) strains P. polymyxa L6 with P. fluorescens M20, or (i,j) strains P. polymyxa Pw-2 with P. fluorescens M20. White arrows indicate sample objects identified as bacterial cells or cell clusters (not all such objects are indicated on each image), unless the arrow is followed by a question mark, in which case the identification is tentative. Other symbols are as follows: L = lateral root, rh = root hair, and a = an example of an area of bright autofluorescence that does not represent FITC- or Cy5-associated signals. 100 urn 50 iim 68 ) "V, 50 p i n 70 Figure 11: Rhizobacterial colonization of randomly sampled root segment surfaces, seven weeks after inoculation. Appropriate primary and secondary antibodies were used in each case. Plant tissue (red autofluoresence) is depicted in blue, far-red light signals (including Pseudomonas fluorescens strain M20 and far-red autofluorescence) in red, and green light signals (including Paenibacillus polymyxa strains and green autofluoresence) in green as described in the text and illustrated in Figure 8. White arrows indicate sample objects identified as bacterial cells or cell clusters (not all such objects are indicated on each image). Other symbols are as follows: L = lateral root. (a,b,c) Roots singly-inoculated with P. polymyxa strain L6. The image in (c) is a magnification of the lateral root in the lower left in (b); scale bar = 20 pm. (d,e) Roots singly-inoculated with P. polymyxa strain Pw-2 (f,g) Roots singly-inoculated with P. fluorescens strain M20. (h,i,j,k) Roots co-inoculated with a mixture of P. polymyxa L6 and P. fluorescens M20. The images in (i) and (k) are magnifications of areas in (h) and (j), respectively; unmarked scale bars = 20 pm. (l,m,n,o) Roots co-inoculated with a mixture of P. polymyxa Pw-2 and P. fluorescens M20. The images in (m) and (o) are magnifications of areas in (I) and (n), respectively; unmarked scale bars = 20 pm. 73 75 Figure 12: Surfaces of root segments sampled thirteen weeks after inoculation, in six different locations: the root tip: (1), the segment immediately after the root tip (2), from the middle of the root (3 and 4, with 3 being closer to the tip), to the base of the root (5 and 6, with 5 being closer to the root tip). All segments were approximately 1 cm long. Plant tissue (red autofluoresence) is depicted in blue, far-red light signals (including Pseudomonas fluorescens strain M20 and farmed autofluorescence) in red, and green light signals (including Paenibacillus polymyxa strains and green autofluoresence) in green, as described in the text and illustrated in Figure 8. White arrows indicate sample objects identified as bacterial cells or cell clusters (not all such objects are indicated on each image). Other symbols are as follows: L = lateral root, rh ~ root hair. (a, b) L6- inoculated root near middle of root (3). The image in (b) is a magnification of a section of the lateral root junction on the lower right in (a); scale bar = 20 pm. (c) L6- inoculated roots near root base (5) (d) L6=inoculated root, near root base (6). (e) Pw-2-inoculated root near root tip (1). The image is of a lateral root tip, not the primary root tip. (f, g) Pw-2-inoculated root immediately after root tip (2). The image in (g) is a magnification of a section taken from the center of the image in (f); scale bar = 20 pm. (h) Pw-2-inoculated root near root base (5). (i) M20*inoculated root near root tip (1), (j, k) M20-inoculated root near middle of root (3), showing root hairs on a lateral root. The image in (k) is a magnification of the root hair on the right in (j); scale bar = 20 pm. (I) M20-inoculated root between middle and base of root (4) (m) M20-inoculated root near root base (5). (n, o) L6 & M20 co-inoculated root near root tip (1). Note that this image is not of the root tip, but of a section distal to it. The image in (o) is a magnification of an area on the lower left in (n); scale bar = 20 pm. (p) L6 & M20 co-inoculated root near middle of root (3) (q) L6 & M20 co-inoculated root near root base (5). (r) Pw-2 & M20 co-inoculated root near root tip (1) (s) Pw-2 & M20 co-inoculated root immediately after root tip (2) (t) Pw-2 & M20 co-inoculated root between middle and base of root (4) (u) Pw-2 & M20 co-inoculated root near root base (5). Figure 13: Thermal print depicting a magnified, on-screen digital green channel image of bacterial cells on the root surface of a pine seedling inoculated with Paenibacillus polymyxa strain Pw-2, next to a 2 pm scale bar. 83 12u, which was an unusual observation; usually where one strain was present on a co-inoculated root, the other strain was present as well. 4.4. Discussion Tables 7, 8 and 9 indicate that purification of IgG from polyclonal antisera, whether or not it was combined with a cross-absorption procedure, decreased the titre of desired antibodies that bound to the homologous bacterial strain, as well as undesired antibodies that bound to heterologous strains, and that these decreases were roughly equivalent. The cross-absorption procedure did not have the intended effect of removing only antibodies that bound P. polymyxa strains L6 or Pw-2, but leaving behind antibodies that bound P. fluorescens M20. This may have been due to inadequate blocking of non-specific protein binding sites on the surface of the cells and tissues added to the cross-absorption reaction mixture, or conversely, the destruction or obscuration of epitopes on the surface of the P. polymyxa cells so that antibodies specific for these epitopes could not bind them. It is also possible that there are epitopes of sufficient similarity between the P. polymyxa strains and strain M20 that removal of antibodies specific for these epitopes resulted in equivalent decreases in both wanted (homologous antigen-binding) and unwanted (heterologous antigen-binding) antibodies. However, since non-IgG serum components (including lipids, cells or cell fragments, other, non-IgG immunoglobulins, proteases or other proteins) that may interfere with IgG antibody stability or binding were absent in the anti-L6 and anti-Pw-2 purified antisera, these antisera were used for microscopy work. Again, it should be noted that each antiserum was concentrated to 3X the levels reported in Tables 7 to 9 prior to storage and use, to compensate for anticipated decreases in antibody titre. The control slides (Fig. 9) demonstrate that there was little or no nonspecific antibody binding {i.e., binding of primary or secondary antisera, used to stain the slides, to root tissues absent of bacteria) in these experiments. The object indicated in Fig. 9f is unlikely to represent nonspecific binding of FITC-labeled secondary antibodies to plant tissues as it was not observed on unbacterized tissues exposed to this antibody (Fig. 9c). The object might represent nonspecific binding of FITC-labelled secondary antibodies to a dense cluster of Pw-2 cells, or it may be the result of a precipitated cluster of FITC-labelled antibodies. The latter interpretation seems most likely, since no other green specks are visible in Fig. 9f, nor were they visible in other, similar control slides (data not shown), and if secondary antibodies were binding Pw-2 cells, presumably they would bind Pw-2 cells in more than one location. The small, rod-shaped objects identified as putative Paenibacillus or Pseudomonas ceils were of an appropriate size to be bacteria, generally about 2 urn long and 1 um wide (Fig. 84 13), and were not observed on unbacterized root samples. The small, rod-shaped objects identified in Figs. 10-13 will therefore be considered bacterial cells. In these experiments, rhizobacteria have been identified in the grooves between epithelial root cells, and as strings or patches of cells on the surface of lodgepole pine roots (Figs. 10-12). Similar observations of uneven surface colonization, and the preferential colonization of the grooves between root cells, have been made of bacteria colonizing the surfaces of wheat (Assmus etal. 1995, Dijkstra etal. 1987, Ruppel etal. 1992, Schloter et al. 1993), barley (Hansen et al. 1997) tobacco (Troxler et al. 1997), pea (Dandurand et al. 1997) and tomato (Chin-A-Woeng etal. 1997) roots. Foster (1986) describes a layer of mucigel, 1-10 pm above the root surface, that is often densely colonized by bacteria and is not removed by washing. In these experiments, rhizobacteria have often been identified apparently "floating" in a transparent matrix, most likely to be mucigel, 2-6 pm above the surface of the root. This is particularly visible in Figs. 12 n and 12 r, but was generally observed in most slides. Colonization of root mucigel by bacteria has also been directly observed by others (Chin-A-Woeng etal. 1997, Hansen etal. 1997). There seem to be a greater number of M20 cells present on the root surface than P. polymyxa L6 or Pw-2 cells, both from seedlings singly-inoculated with one of the bacterial strains (M20, L6 and Pw-2) and from seedlings co-inoculated with strain M20 and one of the P. polymyxa strains (Figs. 10-12). This observation supports the previous observation that average population densities of P. fluorescens M20 recovered from roots in various treatments was greater by at least an order of magnitude than the densities of P. polymyxa strains L6 and Pw-2 (Fig. 6, in Chapter 3). The ability of strain M20 to colonize areas near sites of lateral root emergence was clearly improved in the presence of either Paenibacillus spp, while the reverse does not seem to be the case. It may be that the roots leak compounds that are inhibitory to the growth of P. fluorescens M20 at these sites, which Paenibacillus are not affected by and can neutralize. Alternately, Paenibacillus may produce metabolites that serve as chemoattractants, drawing strain M20 to areas it would not ordinarily colonize. It is possible, but unlikely, that the effect is due to improvements in the nutritional quality of compounds leaked from roots at these sites by Paenibacillus spp., since Pseudomonas sp. are known for their ability to feed on diverse nutrient sources (Montie 1998). P. polymyxa strains L6 and Pw-2 were able to colonize the same microsites on seedling roots co-inoculated with a P. polymyxa strain and P. fluorescens strain M20, as on roots singly-inoculated with strains L6 or Pw-2 alone. Lack of competition between Pseudomonas fluorescens and Bacillus subtilis strains for colonization sites on the root surface has been described before (Dijkstra et al. 1987). It is not clear whether root microsites were colonized by the same quantity of P. polymyxa cells in each treatment, since background autofluorescence in the far-red and green channels, which could not be separated from the light emitted from tagged bacterial cells, made the quantification of light emitted solely from tagged bacterial cells impossible. Further work, in which background autofluorescence is largely eliminated by the use of bright fluorophores with excitation/emission spectra different from most root tissues may answer this question. The images in Fig. 12 support the idea that the root surface is an evolving environment. As the root develops physiologically, and the exudation and water permeability characteristics of a particular segment change, so too does the micro-environment on the root surface of that segment, and the ability of bacteria to proliferate in that environment. It seems reasonable that rhizobacteria would thrive in an environment where nutrients are most plentiful, and the release of nutrients is generally associated with young tissues. Root exudates are released from root tips (Hawes et al. 1998), root hairs (Hofer 1996) and epithelial cracks at sites of recent lateral root formation (these cracks are eventually sealed) (Charlton 1996). Cells are also continually being sloughed from the root cap, which also produces mucilage that may anchor an/or feed bacterial cells (Guckert etal. 1991, Marschner 1991, Watt etal. 1993). It should be remembered that each root segment imaged was subject to a great deal of washing, and rhizobacteria may simpty adhere less tightly to older root segments where the mucilage is older and thinner (as the root diameter increases, the thickness of the mucilage coating would be expected to diminish, unless replenished by a source other than the root tip). Gradients in rhizobacterial colonization along the length of seed-inoculated roots, based on the recovery of viable cells from washed root segments, have been described where a majority of bacteria colonize the root base, and diminish in number toward the root tip (Boelens ef al. 1994, Hebbar etal. 1992, Howie etal. 1987, Liu and Sinclair 1993, Weller 1984). However, this may be an effect of competition between the inoculated rhizobacterium and other, unspecified organisms for plants grown in nonsterile soils (Rattray et al. 1995). 86 Chapter 5 Spore formation in the rhizosphere by Paenibacillus PGPR, and implications for identification of sporeforming bacteria as endophytes 5.1 Introduction Many spore-forming PGPR have been identified (e.g. Chanway et al. 1991, Halverson et al. 1993, Hwang et al. 1995, Wei et al. 1996). Bacterial spores are able to survive harsh environmental conditions that would kill vegetative cells, including exposure to heat, desiccation, UV radiation and sterilizing chemicals (Fairhead et al. 1994, and references therein). The ability of spores to survive harsh environmental conditions may explain why bacilli tend to dominate microbial communities in the more oligotrophic regions of the soil (Timonen et al. 1998). The ability of a sporeforming bacterium to persist in soil will depend upon its ability to form spores when conditions are unfavorable for growth, as well as the ability of its spores to germinate. Bacterial spore germination increases with the availability of nutrients (West et al. 1985) and decreases with the presence of other microorganisms, which tend to suppress the proliferation of bacilli in soil (West etal. 1985, Young etal. 1995). Endophytic bacteria have been identified as such on the basis of their isolation from chemically surface-sterilized root segments (Fisher et al. 1992, Misaghi and Donndelinger 1990) or macerates (Mahafee and Kloepper 1997, Shishido et al. 1995, van Peer et al. 1990). However, bacterial spores present on the root surface can resist chemical sterilization (Fairhead etal. 1994, and references therein), and when root macerates are plated on nutrient media, will germinate and form colonies along with any true endophytes that had been living in the root interior. Endophytes may be isolated with greater success from plant roots that are large enough to dissect, so that exterior tissues may be completely removed using aseptic techniques (e.g. Jacobs et al. 1985, Sturz 1995). Endophytic bacteria have also been identified on the basis of their isolation from surface-sterilized shoot tissues (Mclnroy and Kloepper 1995, Sturz and Christie 1995, Tester 1992), but the problem as to whether sporeforming bacteria isolated this way actually came from inside the shoot may still remain. Bacteria on seed coats or in soils may colonize the aerial parts of plants during the emergence of the radicle from the seed coat, or the seedling from the soil, (Raaijmakers et al. 1995). Isolation of a bacterial species from shoot tissue will only indicate that the bacterium is definitely an endophyte if the seedling has not been grown from a seed coated with bacteria, or a seed germinated in soil containing bacteria. When a bacterium is identified as an endophyte on the basis of its isolation from chemically surface-sterilized, macerated roots, or its isolation from shoot tissues of seedlings grown from inoculated seeds or germinated in inoculated soils, it is always best to verify the identification microscopically. Oversized cells, or ovoid cell clusters (5-7 urn long and 2.5-3 urn wide) of strain Pw-2R, a rifamycin-resistant derivative of P. polymyxa strain Pw-2, have been detected inside the vascular tissues of spruce stems (Shishido et al. 1999), but neither of P. polymyxa strains Pw-2 or L6 have been visualized within the root interior. The objectives of this study were to determine whether P. polymyxa strains Pw-2 and L6 can form spores in the rhizosphere that will germinate under the conditions used to assess endophytic and/or rhizospheric pine root colonization by these strains in Chapters 2 and 3. Attempts were also made to microscopically assess the ability of these strains to colonize the root interior, via indirect immunofluorescence staining techniques and confocal scanning laser microscopy (CSLM). 5.2 Materials and Methods Spore formation of Paenibaciilus sp. in the rhizosphere Lodgepole pine seedlings were seed-inoculated with either P. polymyxa strain L6 or strain Pw-2, or were left unbacterized, and grown in plant tubes under gnotobiotic conditions, as described for the growth promotion/ root colonization assays in Chapter 3. After 6, 9 or 12 weeks of incubation, four seedlings per treatment (twelve in total) were randomly sampled. Colonization of the rhizosphere by Paenibacillus sp. was assessed for each seedling as described in Chapter 3. The standard test for spore formation is a determination of whether a bacterial culture can survive heating (Claus and Berkeley 1986). Formation of spores in the rhizospheres of these seedlings was assessed by placing 1 mL samples of each root wash into sterile Eppendorf tubes and immersing in a 55°C water bath for 30 mins. Heat-treated root wash samples were then diluted in 0.85% (w/v) aqueous NaCI and plated onto half-strength tryptic soy agar (TSA, 15g tryptic soy broth, 15g agar/ L), and incubated at room temperature for up to three days. Representative samples of the colonies formed on each plate were tooth picked onto the foilowing media, to verify the strain identity of colonies: TSA, Str, PAF, and either MOPS-lac or TTC-suc plates (see Table 4 in Chapter 3 for details). These plates were incubated at room temperature and examined daily for up to 4 days. Spore formation in the rhizosphere was assessed five times, and the results of each experiment pooled. Analysis of variance and Duncan's mean separation procedures were performed on calculated values for the mean CFU/ root of cells recovered before and after heat treatment, using SAS v. 6.12 software and an error rate (P) < 0.05. 88 Confocal microscopy of the root interior Lodgepole pine seedlings were seed-inoculated with (i) P. polymyxa strain L6, (ii) strain Pw*2, or (iii) were left unbacterized, and grown in plant tubes under gnotobiotic conditions, as described for the growth promotion and root colonization assays in Chapter 3. Two experiments were conducted. In the first, cross-sections were prepared from randomly chosen areas of roots of seedlings (2-3 per treatment) harvested 7 weeks after inoculation. In the second, each seedling (2 per treatment, harvested 13 weeks after inoculation) was photographed prior to sectioning, and areas to be sectioned were indicated on the photograph. Cross-sections were prepared from (i) an area about 1 cm from the root tip and (ii) an area of the root near the middle of the root or the root base. Preparation of slides for microscopy was conducted as described in Chapter 4, except that cross sections instead of segments were prepared. Cross sections were made by hand by slicing formaldehyde-treated and rinsed roots in a droplet of 0.1 M P 0 4 (ph 7.4) buffer. Cross sections were handled using a small artist's paintbrush. Instead of transferring sections between different solutions in the wells of a 24-well tissue culture plate, they were transferred from 100 pL aliquots of solutions in small (1 x 2") plastic baggies (week 7 roots) or Eppendorf tubes (week 13 roots). Cross sections were rinsed by carefully pipetting out the old solution and replacing it with a fresh aliquot of wash buffer. Slides were made by placing severat cross-sections in a droplet of mounting solution on a flat glass slide and covering them with glass cover slip. Microscope slides were viewed, and the recorded digital images were processed, as described in Chapter 4. 5.3 Results Spore formation Both P. polymyxa strains formed spores in the rhizosphere, but strain Pw-2 formed a greater number or spores than strain L6 throughout the experiment (Fig. 14). The number of spores recovered increased in parallel with rhizosphere populations recovered from each strain (Fig. 14). Endophytic colonization of the root interior Two attempts were made to visualize P. polymyxa cells within pine roots. Results for thriteen week-old roots are depicted in Fig. 15, and were similar to the results obtained for seven week-old roots (not shown). On many root sections, a ring of green/red autofluorescence was observed between the cortex and the stele (Fig. 15b,c,e,g,i), most likely due to fluorescent compounds within the endodermis. 89 Figure 14: Recovery of Paenibacillus polymyxa strains L6 and Pw-2 from washes of lodgepole pine seedling roots before and after heat treatment (55°C, 30 min), from roots taken after 6, 9 or 12 weeks of incubation. Bars with asterisks are significantly larger than other bars within the same treatment (P<0.05). 90 Figure 15: Representative cross-sections of lodgepole pine roots, examined thirteen weeks after inoculation. Plant tissue (red autofluoresence) is depicted in red and green light signals (including Paenibacillus polymyxa strains L6 and Pw-2 and green autofluorescence) in green, as described in the text. Symbols are as follows: c = cortex, s = stele, e = epidermis, L = lateral root. White arrows indicate the green specks mentioned in the text. Control slides: (a) Unbacterized root, no antibodies, (b) unbacterized root, anti-L6 primary antibodies (1° abs) and secondary antibodies (2° abs) (c) unbacterized root, anti Pw-2 1° abs and 2° abs (d) L6 singly-inoculated root, 2° abs only. This figure is a composite of three different micrographs. (e) Pw-2 singly-inoculated root, 2° abs only. Treatment slides: (f,g) L6-inoculated roots, anti-L6 1° abs and 2° abs, sections taken (f) -1 cm. from root tip or (g) from middle of roots. (h,i) Pw-2-inoculated roots, anti-Pw-2 1° abs and 2° abs, sections taken taken (h) ~1 cm. from root tip or (i) from middle of roots. • 1 s • c ) i I v . . s 50 um 100 \itr\ 94 Unfortunately, it was not possible to identify with certainty either of the two P. polymyxa strains within the root interior. Some small green specks were observed in the stele tissue of young (7 week) L6=inoculated pines, but these specks could also be observed in the control tissues of L6-inoculated young pines which had not been exposed to the primary anti-L6 antiserum, (data not shown) suggesting these specks are a result of either nonspecific binding of the secondary antibody, or inadequate rinsing of the secondary antibody from the samples. Similar specks were observed in older (13 week) pine roots (Fig. 15f), but not in the control slides of L&-inoculated pine roots not exposed to primary antiserum (Fig. 15d). 5.4 Discussion Fig. 14 clearly indicates that P. polymyxa strain Pw-2 forms a greater number of spores in the rhizosphere of lodgepole pine than strain L6, under the experimental conditions used. This may be because strain L6 is better adapted to living in the lodgepole pine rhizosphere, and so experiences a lesser degree of physiological stress than strain Pw-2 in that environment. The results may also be explained by possible physiological differences between the two strains. Strain L6 may form spores that have specific nutritional or temperature requirements for germination not met under the experimental conditions used, and which therefore may exist but go undetected by the plating assay. Some strains of Bacillus are known to have specific nutritional requirements for endospore formation (e.g. manganous ions; Claus and Berkeley 1986) so it is possible that P. polymyxa strain L6 has a nutritional requirement for endospore formation that was not met in these experiments. Spores of bacilli contain unique proteins, known as small acid-soluble proteins (SASPs) (Cucchi and Rivas 1998, Fairhead et al. 1994), and it may be possible to determine whether P. polymyxa strain L6 forms spores that do not germinate under the experimental conditions used here by determining the amount of SASPs produced in the rhizosphere. The differential ability of these two P. polymyxa strains to form germinating spores in these experiments might suggest that strain Pw-2 was recovered from chemically surface-sterilized lodgepole pine roots, while strain L6 was not (Shishido etal. 1995, Bent and Chanway 1998) because strain Pw-2 forms more spores, which are able to withstand the chemical treatments and germinate later on plates, than strain L6 (Fig. 14). The ability of bacterial spores to resist chemical sterilization has been well documented (Fairhead et al. 1994), and may even be used as a basis for the determination of endospore formation in bacterial cells (Claus and Berkeley 1986). In 1995 experiments, P. polymyxa strain Pw-2 was unexpectedly recovered from glass hockey sticks which had been exposed to a root wash containing Pw-2, left sitting in ethanol for one week, then flamed (data not shown), 95 demonstrating the resistance of Pw-2 spores to dehydration with ethanol. P. polymyxa strain L6 could not be recovered in this way (data not shown). Previously, a rifamycin-resistant derivative of P. polymyxa strain Pw-2 was isolated from root and shoot tissues of seed-inoculated spruce, and oversized cells or cell clusters of this derivative detected via immunofluorescence techniques and epifluorescent microscopy within vascular tissues of shoots (Shishido etal. 1999), indicating that the parental strain, Pw-2, is likely to be a systemic endophyte. Attempts were made here to see whether individual cells of strain Pw-2 could be visualized within the root interior, using immunofluorescence and confocal scanning laser microscopy techniques. It is possible to produce more sharply focused images using confocal scanning laser microscopy than epifluorescence microscopy since out-of-focus light is excluded from confocal images (Fricker and White 1992), and it was hoped that strain Pw-2 could be localized within the root interior using this technique. Unfortunately, neither strain L6 nor Pw-2 were observed within the root interior (Fig. 15). For strain Pw-2, this could be due to the apparently low average population densities of this strain present in the lodgepole pine root interior (Fig. 2a, Chapter 2), and the fact that each root cross-section sampled only a tiny fraction of the root interior. The experimental technique used to fix and stain the roots may also be to blame: unless bacterial cells within a root were firmly adhered to plant cell walls, they might be washed away during the many rinsing steps. Studies of bacterial endophytes suggest that many of these preferentially colonize, and appear to live freely in, the apoplast (James etal. 1997, M'Piga etal. 1997, Pan etal. 1997, Quadt-Hallman et al. 1997). If P. polymyxa strain Pw-2 also colonizes the apoplast of roots, formaldehyde fixation may not be adequate to fix unbound or loosely bound Pw-2 cells in place. It may be advisable in the future to use a technique in which samples are embedded in a matrix, immobilizing cells which might wash away, or in which there is low background signal, and a single cell can be detected with greater certainty. Use of electron microscopy, bright fluorophores that re-emit light at wavelengths different from most root tissues (i.e. Pacific Blue™, Molecular Probes, Eugene, OR), or P. polymyxa derivatives expressing green fluorescent protein (Ramos et al. 2000), might be useful alternative techniques. Troxler et al. (1997) also report that a Pseudomonas fluorescens endophyte colonized the interior of maize roots to a greater extent during late ripening than at flowering, suggesting that the age or developmental status of a plant may be a factor in endophytic root colonization. It is possible that, with older and better-developed pine seedlings, endophytic colonization of the root interior may be easily visualized. Troxler et al. (1997) reported unusually large (7 pm long) P. fluorescens strain CHAO cells within the maize root interior, which agrees with the previously cited report of unusually large cells or cell clusters of a P. polymyxa strain Pw-2 derivative in spruce shoots (Shishido etal. 1999). It should also be noted, however, that in the 96 latter work, the P. fluorescens cells imaged in spruce shoot tissues were normal in size (Shishido et al. 1999). It is also possible that P. polymyxa strain Pw-2 colonizes the interior tissues of some gymnosperm genera more readily than others, or differs in its expression of epitopes in the tissues of different plant genera. Both this study and Shishido et al. (1999) relied upon immunofluorescence techniques to detect strain Pw-2, but Shishido et al. were examining 5-month old hybrid spruce (P/cea glauca X P. engelmannii) seedlings, while 7- and 13-week old Pinus contorta seedlings were examined here. The last possibility is that neither P. polymyxa strains L6 nor Pw-2 are endophytes, although this contradicts the microscopic observations made previously by Shishido et al. (1999). Unfortunately, the experiments conducted in this study cannot confirm the endophytic status of strain Pw-2, nor deny it. Further experiments using different techniques are required before conclusions can be drawn concerning whether strain Pw-2 colonizes the root interior of lodgepole pine. 97 Chapter 6 Co-inoculation effects of a mycorrhizal fungus, Rhizopogon subcularescens Smith 11616, with Paenibacillus PGPR on extent of pine growth promotion and PGPR root colonization 6.1 Introduction The majority of the world's plants form associations with mycorrhizal fungi (Wilcox 1996), and conifers are no exception. Ectomycorrhizae can have a variety of positive effects on conifer growth (reviewed in section 1.6), and may also influence the ability of PGPR to interact with plant roots since (i) the mantle may physically prevent soil bacteria from gaining access to the root surface, (ii) the positive effects of the mycorrhizal symbiosis may obscure any positive PGPR-mediated growth effects, and (iii) the ectomycorrhizal fungus may, via metabolism of root exudates and/or synthesis and release of its own metabolites, alter the composition of exudates to which rhizobacteria are chemotactically attracted, or upon which rhizobacteria feed (Christensen and Jacobsen 1993, Curl and Truelove 1986, Danell etal. 1993, Timonen etal. 1998). Previous greenhouse studies have indicated that the growth of mycorrhizal lodgepole pine seedlings may be improved by the application of Pseudomonas PGPR (Shishido ef al. 1996b) and rifamycin-resistant derivatives of P. polymyxa strains L6 and Pw-2 (Shishido et al. 1996a). In order to determine if these observations were due solely to interactions between the fungus, the PGPR and the plant, and not other microorganisms that may be present and affecting the growth or ability of any one of these to interact with the others, the abilities of P. polymyxa strains L6 and Pw-2 to colonize the rhizosphere, and promote the growth of, lodgepole pine seedlings exposed to an ectomycorrhizal fungus were evaluated under gnotobiotic conditions. The fungus, Rhizopogon subcularescens Smith 11616, was chosen primarily on the basis its ability to form ectomycorrhizae on lodgepole pine seedlings in a screening test, and partly due to the prevalence of mycorrhizae formed by Rhizopogon species on pines under natural conditions in the Pacific northwest. Rhizopogon, a genus with about 150 described species, is most abundant in western North America, and Rhizopogon species are found in association with several Pinaceae hosts, including Douglas fir (Pseudotsuga menziesii) and Pinus species (Molina ef al. 1997). 98 6.2 Materials and Methods Screening of ectomycorrhizal fungi Five ectomycorrhizal isolates were screened for their ability to form mycorrhizae under gnotobiotic conditions within a short period of time (after 7 weeks of incubation). Cultures of Rhlzopogon subcularescens Smith 11&1&, Rhizopogon albescens Luoma 1039 and SuiHus luteus (FT.) S. F. Gray 9013 were obtained from D. McKay (U.S. Forestry Service, Corvallis, OR 97331). A culture of Rhizopogon vulgaris 11564 was obtained from L. Paul (University of British Columbia, Vancouver BC), and a culture of Wilcoxina rehmii 1404-2 was obtained from K. Egger (University of Northern British Columbia, Prince George, BC). Cultures were maintained on potato dextrose agar (PDA, Difco Laboratories, Detroit, Ml 48232) plates, which were incubated at room temperature until fungal growth was evident, then stored at 4°C. Fresh PDA plates were inoculated with plugs of fungal cultures (obtained asepticatly using a #2 cork borer sterilized in 90% ethanol for at least 5 minutes) roughly every 3 months. Six Pyrex culture jars (100 X 80 mm) with glass covers were filled to a depth of about 2.5 cm with a peat-vermiculite nursery mix (Sunshine Mix 4, Fisons Horticulture Inc. Vancouver B.C), and 100 mL of distilled water was added to each jar. Jar lids were sealed firmly with masking tape and the jars were autoclaved 30 mins. Lodgepole pine seeds were surface-sterilized as described in Chapter 2, soaked for 1 hour in sterile distilled water, then 6 evenly-spaced seeds were planted in each jar. Previous experiments in which agar plugs had been used to inoculate plant growth tubes had provided uniformly disappointing results, so portions of fungal mycelium grown in nursery mix were used as inoculum: 125 mL Pyrex flasks containing 60 mL of Sunshine Mix 4 nursery mix and 20 mL potato dextrose broth (PDB, Difco Laboratories, Detroit, Ml 48232) were autoclaved 30 minutes, then inoculated in triplicate with fungal plugs (one per flask) from cultures maintained on PDA plates. Flasks were incubated at room temperature for at least two months. Fof each of five of the jars, six portions of nursery mix colonized by one fungal culture (~ 0.5 mL) were aseptically removed from flasks and placed directly atop the seeds, one chunk of fungus per seed. Each jar therefore contained multiple samples of one fungus. For the sixth jar, chunks of nursery mix from uninoculated 125 mL Pyrex flasks, prepared as described above, were used. The lids of the jars were affixed firmly with tape in a manner which would still allow for the movement of air into the jars, then placed in a growth room illuminated by a fusion lamp (SolarRay), with a 16 h photoperiod (phtfdsyritfietrc plidtdri ftux at cartdpy level ~170 prftot m"2) arid a lightrdark temperWre regime of roughly 22:18°C. After 7 weeks, seedlings were removed and the unstained roots examined using a dissecting microscope for the occurrence of bifurcated or swollen root tips, indicative of mycorrhizal infection (Massicotte et al. 1999). 99 Preparation ofRhizopogon subcularescens Smith 11616 inoculum Fungal inoculum for experiments was prepared as follows: Sixteen Magenta GA-7 tissue culture boxes (Sigma Chemical Co., St. Louis, MO 63178) containing 120 mL of Sunshine Mix 4 nursery mix and 45 mL of PDB were sealed with foil and autoclaved 30 minutes. Chunks of nursery mix colonized by hyphae were aseptically removed from 125 mL flasks inoculated with R. subcularescens 11616 as described above, and placed in the center of fourteen tissue culture boxes. The remaining two boxes were left as a contamination check. The boxes were then incubated at room temperature for 8-10 weeks. Prior to use as inoculum, chunks of nursery mix colonized by hyphae were aseptically removed from each tissue culture box and combined in the top of a sterilized 1 L vacuum filtration apparatus containing a 0.45 pm filter. To leach unused nutrients from the PDB added to the nursery mix in the fungal inoculum, 100 mL of sterile distilled water was thoroughly mixed with the fungal inoculum and as much of this as possible (50-70 mL) was subsequently removed by vacuum filtration. The resulting fungahpeat slurry was used to inoculate plant growth tubes as described below. Inoculation and growth of pine seedlings for growth promotion/root colonization assays Six hundred sterile plant growth tubes containing 30 mL Sunshine 4 nursery mix and 11 mL distilled water were prepared as described in Chapter 3. Half the tubes (300) were aseptically inoculated with a small scoopful (~ 0.5 mL) of leached fungal inoculum, placed atop the sterile nursery mix in the tube. Plant growth tubes were allowed to sit for 1-3 days at room temperature before the addition of seeds. To verify the absence of microbial contaminants in the fungal inoculum, a small scoopful of the fungal inoculum was smeared across one tryptic soy agar (TSA, 15 g tryptic soy broth, 15 g agar/L) and one PDA plate. Plates were incubated at 27°C for up to two weeks, and examined for the presence of bacteria or fungi morphologically distinct from R. subcularescens 11616. Lodgepole pine seeds (200 per treatment) were inoculated with either P. polymyxa strain L6 or strain Pw--2, or were left unbacterized, as described for the growth promotion/ root colonization assays in Chapter 3. For each seed treatment, half (100 seeds) were placed in plant growth tubes containing fungal inoculum, directly atop the inoculum, and the remaining 100 seeds placed in sterile plant growth tubes. A scoopful (~ 1 mL) of sterile sand was aseptically added to each growth tube after the addition of the seed. The resulting six treatments were as follows: (i) uninoculated control, (ii) R. subcularescens 11616, (iii) P. polymyxa L6, (iv) P. polymyxa Pw-2, (v) R. subcularescens 11016 + P. polymyxa L6, and (vi) R. subcularescens 1161& h^P, polymyxa Pw-= 2. Plant growth tubes were placed in a growth room illuminated by a fusion lamp (SolarRay), 100 with a 16 h photoperiod (photosynthetic photon flux at canopy level ~170 pmol s"1 m"2) and a lightdark temperature regime of roughly 22:18°C, and incubated for 9 or 12 weeks. The experiment was conducted twice. Growth promotion/root cotonizationassays at & and 12 weeks Bacterial populations in the rhizosphere were enumerated in a manner similar to the experiments described in Chapter 2, with several modifications. Fifty plant tubes were taken from each treatment at 9 or 12 weeks after seed inoculation. Eight seedlings per treatment were chosen at each time point for an evaluation of bacterial colonization of the root exterior. Evaluations of root interior colonization were not made. Shoots were severed from roots, and shoots placed on ice in pre-weighed plastic bags until they could be freeze-dried. For seedlings sampled after 9 weeks of incubation, the roots were washed in 50 mL flasks containing 5 mL of sterile KP with 0.01% (v/v) Tween 20; for seedlings sampled after 12 weeks, 8 mL of this solution was used. After sampling the root wash solution, flasks containing roots were stored on ice until root measurements could be made. Root wash solutions were diluted in 0.85% (w/v) NaCI and plated on TSA. Plates were incubated at 27°C for 2 days, and representative colonies from each sample were assessed for resistance to streptomycin, ability to grow on PAF, and colony colour on MOP&4ac medium (Table 4). Seedlings in the remaining test tubes were gently removed by shaking the tube, and rinsed in distilled water. Root measurements, including total length and number of lateral roots, were made using a Hewlett-Packard flatbed scanner with a transparency lid at 600 dpi. Digital images of roots were scanned and analyzed with WinRhizo root analysis software (Regent Instruments, Inc., Quebec City, Quebec). All roots were rinsed, scanned, placed in pre-weighed plastic bags and kept on ice until they could be freeze-dried. The number of macroscopic bifurcated root tips on each root was determined by visual examination. Roots and shoots were frozen with liquid nitrogen, or by storage for at least 24 hours at -80°C. Multiple holes had been punched in the plastic bags previously, to allow samples to freeze-dry. Frozen samples were placed in a lyophilizer and freeze-dried for at least 16 hours. Upon removal from the lyophilizer, samples were placed in undamaged, airtight plastic bags and stored at 4°C until they could be weighed. Samples were allowed to come to room temperature in the sealed bags before weighing. After weighing, samples were stored in the airtight bags at -20°C. \ 101 Verification of bacterial strain identities/contamination checks Characteristic profiles based on intrinsic antibiotic resistance, ability to grow and produce water-soluble fluorescent pigment on PAF, and colony colour on indicator media containing different sugars, were developed for each strain (Table 4, Chapter 3). To check for contamination during experiments, samples of bacterial inocula before and after seed bacterization, as well as all bacterial colonies growing on representative rhizosphere wash dilution plates, were routinely subjected to these tests. The results were compared with those from pure laboratory cultures of P. polymyxa strains L6 and Pw-2. AH bacterial cultures were stored in aliquots at -80°C as cell suspensions (from an overnight growth of culture in TSB) diluted 1:1 with 80% (w/v) glycerol. Determination of antimicrobial effects of fungus To determine whether R. subcularescens 11616 affected the growth of P. polymyxa strains L6 and Pw-2, and P. fluorescens strain M20, a simple plating experiment was performed. Loopfuls of stored frozen bacterial suspensions were freshly streaked across TSA medium and incubated 48 hr at 27°C. The fresh bacterial colonies which formed were sampled and suspended in 0.85% (w/v) NaCI solution to an A^o corresponding to an estimated cell density of 5 X10 7 CFU/mL (0.4 for strains L6 and Pw-2, and 0.15 for strain M20). For each strain, aliquots of 0.1 mL were plated on each of two PDA plates, and two PDA/TSA (19.5 g PDA, 7.5 g TSA, 7.5 g agar / L) plates. Uninoculated controls included plates upon which nothing was spread, and plates upon which only sterile NaCI solution was spread. A 5 mm dia. plug, taken from the perimeter of an P. subcularescens 11616 colony growing on PDA, was placed using aseptic techniques in the center of each plate, and plates were inverted, incubated at 27°C in the dark, and examined every other day for a period of up to 10 days. Each bacterial suspension was also streaked across strain test media to verify the identity of each strain, as described in the section above and in Table 4 (Chapter 3). To determine whether zones of inhibition which were observed around fungal plugs had a toxic or static effect on the bacteria, areas (1) within the zone of inhibition and (2) outside the zone of inhibition, near the margin of the inoculated plate, were sampled from each plate and streaked on each of two TSA plates, one PDA plate, and one PDA/TSA plate. Plates were incubated at 27°C in the dark and examined daily for a period of up to 7 days. Data analyses Statistical analyses were conducted using SAS v.6.12 software, with an error rate (P) < 0.05 unless otherwise indicated. Data from the two growth promotion/ root colonization 102 experiments were pooled for analysis. Analysis of variance by the general linear models procedure was performed on all seedling growth variable data. Analysis of variance by the general linear models procedure was also performed on rhizosphere CFU and density (CFU/root surface area) data. Comparisons were made between all treatments, and also between treatments incorporating a bacterial strain (P. polymyxa L6 or Pw-2) in order to estimate differences in the rhizosphere populations of each strain when singly- or when co-inoculated with R. subcularescens 11616. Duncan's procedure for means separation was also applied to seedling growth and rhizosphere population data. The percentage of seedlings having bifurcated root tips was found to vary between experimental repetitions, but not between weeks, so data from each week for each treatment in an experimental repetition were pooled for further analysis. Comparisons were made between all treatments, and between pooled data for those treatments incorporating R. subcularescens 11616, and those not, using Duncan's procedure for means separation. 6.3 Results Lodgepole pines inoculated with the mycorrhizal fungus R. subcularescens 1161& accumulated a greater amount of root and shoot biomass, had longer roots and a greater number of lateral roots compared to uninoculated control seedlings, throughout the experiment (Fig. 16). There were no significant differences between seedlings inoculated with 11646 alone, and those inoculated with a combination of 11616 and a PGPR strain (Fig. 16). PGPR strains P. polymyxa L6 and Pw-2 also had growth-promoting effects: seedlings inoculated with strain L6 alone had significantly longer roots than uninoculated control seedlings by week 9, and greater root and shoot biomass accumulation and lateral root formation by week 12 (Fig. 16). Seedlings treated with strain Pw^2 alone had significantly longer roots than the uninoculated control seedlings by week 9, and a greater number of lateral roots by week 12; however, these seedlings were not observed to have heavier roots or shoots (Fig. 16). More seedlings had bifurcated root tips in the first experiment than in the second, but the percentage of seedlings with bifurcated root tips were similar for all treatments in each experiment (Fig. 17), with no significant differences observed between seedlings harvested in weeks 9 and 12. No significant differences between treatments were observed when treatment data for the two experimental repetitions were pooled, even when data for the three treatments containing fungus, and the three that did not contain fungus, were pooled and compared. In this study, bifurcated "tuning fork shaped" root tips were considered to be mycorrhizal, or at least potentially mycorrhizal. 103 Preliminary attempts to verify that bifurcated tips were indeed ectomycorrhizal were made. A procedure for the specific staining of ectomycorrhizal root tip mantles in mature oak roots (Daughtridge et al. 1986) was found to be unsatisfactory for young pine seedlings, as the entire root system of each seedling was stained. Other preliminary attempts to visualize mantle and Hartig net formation via toluidine blue staining (Peterson 1994) in 9- and 12-week old pine seedling root tips were likewise unsatisfactory: while fungal mycelia were often observed on the exterior surface of root tips that were suspected of being mycorrhizal, Hartig nets were never clearly identified in any root tip. Attempts to measure the level of ergosterol, a more sensitive indicator of fungal infection in ectomycorrhizae than chitin (Johnson and McGill 1990), in seedlings with bifurcated root tips using the method of Martin et al. (1990), were likewise unsuccessful. This was most probably because the level of ergosterol detected in 35 mg (dry weight) samples of ft subcularescens 11616 hyphae scraped from an inoculated PDA plate was near the lower limit of detection for the HPLC equipment used (data not shown). Under these circumstances, small infected pine roots (dry weight 2-15 mg) would be unlikely to contain detectable levels of ergosterol. ft subcularescens 1161& appeared to reduce the population density of both PGPR strains in the rhizosphere (Fig. 18), although strain L6 was not apparently affected at 9 weeks after inoculation. In vitro, ft subcularescens was observed to have an antibacterial effect on both P. polymyxa strains Pw-2 and L6, but not P. fluorescens M20 (Table 10). This effect was limited to media upon which the bacteria grew poorly. When areas within observed zones of inhibition were sampled, bacterial growth was observed (Table 11), suggesting that ft subcularescens 11616 has a bacteriostatic rather than a toxic effect on P. polymyxa. ft subcularescens grew equally well on PDA as on PDA/TSA in the absence of bacteria. Table 10: Effect of Rhizopogon subcularescens 11616 and growth media on growth of bacterial strains in vitro P, fluorescens M20 P. polymyxa L6 Pr polymyxa Pw-2 Uninoculated controls Growth3 on PDA ++ +/- +/- — Zone0 radius on PDA 0 mm 3.5 mm 5.5 mm Growth3 on PDA/TSA ++ + + — Zone 0 radius on PDA/TSA 0 mm 0 mm 0 mm a Growth of bacteria: ++ = profuse, + = average (visible), +/- = weak ( aarely visible), - = no growth, observed 10 days after inoculation. bZone of inhibition, observed 10 days after inoculation. 104 Figure 16: Average (a) root dry weight, (b) shoot dry weight, (c) root length and (d) number of lateral roots measured for seedlings in each treatment after 9 and 12 weeks of incubation. Treatments having a lowercase letter in common were not significantly different (P<0.05). 105 Myc. fungus Control i i Experiment 1 W77A Experiment 2 ! L6 L6 + Fungus Pw-2 Pw-2 + fungus Treatment Figure 17: Percentage of seedlings having bifurcated root tips in each treatment (data for week 9 and 12 observations were pooled). Mean percentages for each experiment were divided by two and stacked, so that the height of the combined bar represents the overall average from both experiments. No significant differences (P< 0.05, P<0.1) were observed between treatments within each experiment, or when data from both experiments were pooled. 106 a ) CO £ CO s 3 in w E E % S 3 u. O % in c <u •a 7.0e+5 6.0e+5 5.0e+5 4.0e+5 3.0e+5 2.0e+5 1.0e+5 0.0 a 9 T 1 12 Weeks after inoculation b ) L6 L6+ 11616 CO 9> 8 cn E E 3 U-O in c (U "O 1.6e+6 1.4e+6 1.2e+6 1.0e+6 8.0e+5 6.0e+5 4.0e+5 2.0e+5 b 12 Weeks after inoculation Pw-2 - V - Pw-2 + 11616 Figure 18: Average rhizosphere population sizes of Paenibacillus polymyxa strains (a) L6 and (b) Pw-2 when inoculated singly and co-inoculated with Rhizopogon subcularescens Smith 11616. Error bars indicate standard deviations. Significant differences between treatments (P<0.05) are indicated by lowercase letters: a, L6 alone > L6 with 11616; and b, Pw-2 alone > Pw-2 with 11616. 107 Table 11: Growth a of bacteria, sampled from areas within and outside of zones of inhibition, on different media Within zone of inhibition Outside zone of inhibition (plate margin) Bacterium TSA PDA/TSA PDA TSA PDA/TSA PDA L6 ++ + +/* ++ + +/-= Pw-2 ++ + +/- ++ + +/-Growth of bacteria: ++ = profuse, + = average (visible), +/- = weak (barely visible), - = no growth, observed 5 days after inoculation. Growth on strain test media verified strain identities (not shown). 6.4 Discussion Bifurcated root tips The presence of bifurcated root tips on seedlings that were not exposed to R. subcularescens 11616 suggest that (i) non-mycorrhizal root tips were in some cases mistaken for mycorrhizal ones, or (ii) there was a mycorrhizal fungus present as a contaminant in the non-fungal treatments. Attempts were made to rule out the first possibility by determining the level of ergosterol in roots not exposed to the fungus, but owing to the small size of the roots (2-15 mg dry weight) and the barely detectable levels of ergosterol detected in a 35 mg (dry weight) sample of pure R. subcularescens 11616 hyphae (data not shown), ergosterol could not be quantified via HPLC reverse-phase chromatography in these roots. More sensitive quantification methods, involving gas chromatography and mass spectrometry (e.g. Axelsson etal. 1995), orfluorodensitometry and thin-layer chromatography (Bailly et al. 1999) could be useful for future studies. In addition, while some root tips in the fungus-inoculated treatments appeared to have a mycelial mantle when viewed unstained under a dissecting microscope, or sectioned and stained with toluidine blue (data not shown), it was unclear whether mycorrhizae had formed on all bifurcated root tips. It is possible that pine seedling lateral roots sometime appear to emerge at an angle, owing to growing conditions or to handling during an experiment, which to the unaided eye makes them resemble a bifurcated root tip. The fact that all experimental treatments, whether or not they included inoculation with R. subcularescens, experienced the same percentage of roots with bifurcated tips makes this possibility very likely. Careful examination of root tips with a dissecting microscope (which was not considered feasible for these experiments, due to time constraints in the processing of samples- each root had to be examined and frozen as quickly as possible, to minimize the degradation of root hormones) would eliminate this problem in future studies. In regards to possible contamination with a mycorrhizal fungus, one would expect a greater number of bifurcated tips in those treatments inoculated with the fungus, if bifurcated tips indicated mycorrhizal infections. However, it is possible that, despitetheuseof aseptic 108 technique, R. subcularescens 11616 was introduced to some of the non-fungal treatments tubes during the inoculation of test tubes with seeds. It is also possible that a mycorrhizal fungus present in the soil mix or on the seed coats somehow survived the sterilization methods employed, and was not detected during tests conducted to verify the sterility of seed coats and of autoclaved nursery mix. On average, only 8-20% of seedlings in a given treatment were found to have bifurcated root tips (Fig. 17). Given the lack of germination in some treatments, this could represent as few as two seedlings in a treatment during a particular harvest. Since it is not certain that the bifurcated root tips represent a fungal contaminant, no fungal contaminants were observed during dilution plating tests, the majority of seedlings in each treatment did not possess bifurcated root tips, and the percentage of seedlings with bifurcated root tips was statistically similar for ail treatments in both replications, the majority of seedlings in each treatment will be considered to be uncontaminated and treatments will be discussed as such. Fungus^mediated growth promotion in the absence of mycorrhizae Exposure to a mycorrhizal fungus can significantly improve the growth of lodgepole pine under gnotobiotic conditions, in spite of the fact that the majority of fungus-treated seedlings are not visibly mycorrhizal (Figs. 16 and 17). Growth promotion of lodgepole pine by R. subcularescens 11616, at least initially, is not reliant on the formation of mycorrhizal structures. Similar conclusions have been drawn for Norway spruce seedlings inoculated with a strain of Laccaria bicolor, in which the fungus was physically prevented from contacting the root and forming mycorrhizae (Karabaghili-Degron et al. 1998). It was hypothesized that growth promotion, in the latter case, was due to plant uptake of fungally-produced indoleacetic acid (Karabaghili-Degron et al. 1998). The ability of R. subcularescens 11616 to produce IAA or other phytohormones is not known. An in vitro ability to produce indole acetic acid or ethylene by different mycorrhizal fungi has been correlated to the endogenous root IAA content of infected lodgepole pine, Englemann spruce or Douglas-fir (Scagel and Linderman 1998b), and an ability to form mycorrhizae and improve the growth of Scots pine (Rudawska and Kieliszewska-Rokicka 1997), lodgepole and ponderosa pines, and Douglas-fir (Scagel and Linderman 1998a), suggesting that plant hormones produced exogenously by fungi may be absorbed by conifers, and might subsequently affect plant growth. Experiments to determine the in vitro ability of R. subcularescens 11616 to produce IAA, and to compare the root IAA levels of lodgepole pine seedlings exposed to R. subcularescens 11616 with those not inoculated with fungi, have been postponed until the ELISA kits to test for IAA become available. 109 Bacterial growth in the presence of R. subcularescens 77676 The growth of P. polymyxa was generally inhibited in treatments where it was co-inoculated with R. subcularescens 11616 (Fig. 16). Similar results were reported for bacteria growing in the rhizosphere of mycorrhizal cucumber (Christiensen and Jakobsen 1993). It is unlikely that bacterial colonization of the root surface was physically prevented by ectomycorrhizal mantles, since visible mycorrhizae failed to form on the majority of fungus-inoculated roots (Fig. 17), including those sampled for the determination of bacterial colonization of the rhizosphere. It is probable that R. subcularescens 116-16-and P. polymyxa competed directly for limiting nutrients in the rhizospshere, since the gnotobiotic system used here was relatively nutrient-poor. These results are supported by the fact that the growth of P. polymyxa strains were inhibited in the vicinity of the fungus, but only on the medium which these strains grew more poorly upon (Table 10), and that this inhibition appears to be bacteriostatic in nature (Table 11). Alternatively, the fungus may produce antibacterial metabolites under rhizosphere conditions that were not produced, or able to have a noticeable effect, in vitro. Shishido et al. (1996a) reported that the populations of rifamycin-resistant derivatives of P. polymyxa strains L6 and Pw-2 were not detectable in the rhizosphere of lodgepole pine grown under greenhouse conditions in different forest soils, further suggesting that P. polymyxa strains L& and Pw-2 may not be able to extensively colonize the rhizosphere in the presence of other microorganisms, including mycorrhizal fungi. Growth promotion effects of PGPR and mycorrhizal fungus are not additive , There was no additive effect of inoculation with R. subcularescens 11616 and either P. polymyxa strain on lodgepole pine growth: throughout the experiment, in every parameter measured, the growth of seedlings inoculated with R. subcularescens alone, and those dually-inoculated with R. subcularescens + P. polymyxa, was identical (Fig. 16). Shishido etal. (1996a) reported that rifamycin-resistant derivatives of P. polymyxa strains L6 or Pw-2 were able to increase the growth of mycorrhizal and non-mycorrhizal lodgepole pines grown under greenhouse conditions in a variety of forest soils, to a similar extent, although the increase in growth of mycorrhizal pines was statistically insignificant when compared to uninoculated mycorrhizal control seedlings. It was speculated that these PGPR strains were able to stimulate pine growth through a mechanism unrelated to mycorrhizal fungi. It is unlikely that the lower rhizosphere populations of P. polymyxa in treatments dually-inoculated with R. subcularescens + P. polymyxa (Fig. 18) are responsible, since earlier experiments with P. polymyxa strains L6 and Pw-2 demonstrated that the extent of growth promotion observed under gnotobiotic conditions was not related to rhizosphere populations of each strain (Chapter 2, Chapter 3). Shishido et al. (1996a) also reported that levels of 110 rifamyciri-resistant derivatives of P. polymyxa strains L6 and Pw-2 were not detectable in the rhizosphere of pine, yet were able to mediate improvements in pine biomass accumulation. The ability to improve the growth of mycorrhizal pines appears to be present under greenhouse conditions, and vanishes under gnotobiotic conditions. Pines grown under gnotobiotic conditions were subjected to a variety of stresses, including limitations on nutrients and growing space. It is possible that, under better growing conditions, pines inoculated with a combination of R. subcularescens + P. polymyxa would have grown even larger than those inoculated with the mycorrhizal fungus alone. It is also possible that, under non-gnotobiotic conditions, P. polymyxa is able to influence plant growth indirectly via interactions with other rhizosphere microorganisms; for example, biocontrol effects on deleterious rhizosphere microorganisms. Experiments conducted under gnotobiotic conditions can only evaluate the direct effects of PGPR on inoculated plants, since the majority of soil microorganisms that would normally be present are excluded. Under non-gnotobiotic conditions, even though a mycorrhizal fungus may prevent a PGPR from exerting a direct effect on plant growth, the PGPR may still be able to improve plant growth indirectly. The bacteriostatic effect of P. polymyxa strain L& on Curtobacterium flaccumfaciens strain PF322 in vitro has been noted (Chapter 2). It should also be noted that the mycorrhizal inoculum used by Shishido et al. (1996a) consisted of forest soil, and that a variety of different mycorrhizal associations developed. Different mycorrhizae interact differently with plants (Francis and Read 1995), and may have different effects on PGPR, or vice versa (Christiensen and Jakobsen 1993, Danell etal. 1993, Garbaye 1994, Frey-Klett et al. 1999, Vonderwell 1998). An additive effect due to PGPR inoculation might be possible on lodgepole pines infected with one mycorrhizal fungus, but not pines exposed to or infected by another. 111 Chapter 7 Conclusions and recommendations for future work This work was conducted primarily to determine whether the extent of lodegepole pine growth promotion mediated by Paenibacillus polymyxa PGPR is influenced by the presence of other rhizosphere microorganisms, including rhizobacteria and mycorrhizal fungi. This work also attempted to shed some light on the mechanism by which other rhizosphere microorganisms may affect PGPR-mediated growth promotion, by examining whether interference by other rhizosphere microorganisms is linked to altered PGPR population densities in the rhizosphere, or an altered ability by PGPR to colonize particular microsites (root tips, lateral root junctions) on the root surface, whether PGPR affect root hormone levels, and whether the two P. polymyxa strains used varied in the ability to produce spores able to survive in soil and later germinate. The presence of other rhizobacteria (Curtobacterium flaccumfaciens PF322 or Pseudomonas fluorescens M20) was demonstrated to reduce the extent of growth promotion of lodgepole pine mediated by P. polymyxa PGPR, with strain Pw-2 being more severely affected than strain L6. The extent of PGPR-mediated growth promotion observed in lodgepole pines was not related to the endophytic or rhizospheric population densities of each PGPR strain under gnotobiotic conditions. Bacterial inoculants seemed to alter levels of root hormones on pine seedlings after 12 weeks of incubation, although further work must be conducted to verify these observations. Further work also should determine (1) the full spectrum of hormones and their conjugates that are affected, and (2) whether these changes indicate an absorption of hormones produced exogenously by the bacteria, or an active synthesis of hormones in response to bacterial inoculation. Qualitative colonization of particular microsites on the lodgepole pine root surface did not seem related to the extent of growth promotion mediated by P. polymyxa. Colonization was observed to be heterogenous, with scattered clusters of bacteria located preferentially on younger root tissues. P. polymxya colonies and P. fluorescens colonies were always found co-existing in the same root areas on seedling roots co-inoculated with both bacteria. Colonization of lateral root junctions by P. polymyxa enabled P. fluorescens to also colonize those areas, but no qualitative changes to P. polymyxa surface colonization in the presence of P. fluorescens were observed. Examination of entire root systems, rather than individual segments, may provide further insight into spatial distributions of bacteria on root surfaces. The use of fluorochromes with excitation/emission spectra differing from plant compounds may also improve the specificity of detection, and allow for an accurate, quantitative assessment of bacterial colonization of particular root surface areas. 112 P. polymyxa strains L6 and Pw-2 were found to vary in the production of spores in the rhizosphere that were able to germinate under the experimental conditions used. This was an interesting observation, since the organism that produced more spores (strain Pw-2) has also been identified as an endophyte of conifers. It is not known whether these traits are related, or whether spores were produced by strain L6 that were unable to germinate. It is also not known whether enhanced spore production improves the ability of a PGPR strain to promote plant growth over an extended period of time (i.e. over one or several growing seasons). These questions are of practical as well as ecological significance for PGPR-conifer interactions. R. subcularescens was demonstrated to improve the growth of young lodgepole pines, the majority of which were not mycorrhizal. The effect of R. subcularescens on pine growth in systems where the fungus is physically prevented from contacting the plant (i.e. by a cellophane membrane) should be evaluated to confirm that R. subcularescens is indeed able to promote pine growth via non-nutritional means. Karabaghli-Degron etal. (1998) reported that the growth promotion of non-mycorrhizal spruce seedlings was due to auxin production by an ectomycorrhizal fungus. It may therefore be useful to investigate whether there are changes in root levels of auxin or other phytohormones in non-mycorrhizal seedlings exposed to R. subcularescens, and if so, whether hormones are exogenously produced by the fungus and absorbed by the plant, or whether hormones are produced endogenously in response to some stimulus. It would also be interesting to see whether the effects of bacterial inoculants on root hormone levels, observed previously, are also observed in seedlings exposed to a mycorrhizal fungus. P. polymyxa PGPR did not alter the growth of lodgepole pine exposed to R. subcularescens under gnotobiotic conditions. Other experiments, conducted under non-gnotobiotic conditions, indicated that there is an additive effect of P. polymyxa inoculation on mycorrhizal seedlings (Shishido ef al. 1996a). It is possible that P. polymyxa was able to improve the growth of these latter seedlings via interactions with other rhizosphere organisms, or that the conditions of the experiments reported here limited the extent of plant growth. Further experiments should be conducted to verify these conclusions, under the experimental conditions used here as well as under less restrictive conditions, perhaps in gnotobiotic microcosms to which a sterile nutrient solution is routinely added to alleviate drought and nutrient stresses. R. subcularescens also reduced the density of P. polymyxa populations in the rhizosphere. It is possible that P. polymyxa was affected in its ability to colonize of root surface, but in the absence of established mycorrhizae it is not clear how such an interaction would occur. Microscopy experiments to evaluate this possibility may provide interesting results. 113 It is important to understand interactions involving bacterial inoculants, host plant(s) and other soil microorganisms if PGPR are to be used effectively as practical agricultural inoculants. Without understanding how and when a growth-promotion stimulus is produced by a PGPR and perceived by the host, it will be difficult to predict which edaphic, climatic, or biotic factors are most influential in a given PGPR-plant interaction. Recent work by Fuchs et al. (2000) demonstrated that the ability of a PGPR to function as a biocontrol agent was influenced by the kind of growth media the inoculant was grown in, suggesting that physiological state of a bacterium is vital to its ability to interact with plants or other organisms. It would be useful to identify exactly what changes occur in bacteria that may affect its interactions with a plant-expression of specific surface proteins or other compounds or structures, for example, or the secretion of extracellular compounds. Interaction with a live bacterium may not be required for plant growth promotion in every plant-bacterial interaction. When this is the case, it would be prudent to test use of bacterial constituents in the absence of the bacterium, and avoid the variability inherent with dealing with a living bacterial inoculant. It should also be remembered that the rhizosphere is a complex system, one in which variables are interdependent and exist in positive and negative feedback mechanisms. For example, root and microbial respiration will increase levels of carbon dioxide in the soil atmosphere, decreasing the pH of the soil solution and altering (1) the solubility of ions in the soil solution and (2) the ability of soil organisms to tolerate the rhizosphere environment. Both items affect root and microbial respiration: item (1) will alter the exposure of plants and rhizosphere organisms to inorganic nutrients and heavy metals, indirectly affecting the growth and therefore respiration of roots and microbes in the rhizosphere. The rate of respiration, and whether overall respiration increases or decreases, is dependent on many other variables, too numerous to mention. 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