PHENAZINE-1-CARBOXYLIC ACID-PRODUCING
PSEUDOMONAS SPP. OF THE INLAND
PACIFIC NORTHWEST (U.S.)
By
JAMES A. PAREJKO
A dissertation submitted in partial fulfillment of the requirements for the degree of
DOCTOR OF PHILOSOPHY IN MICROBIOLOGY
WASHINGTON STATE UNIVERSITY School of Molecular Biosciences
DECEMBER 2012
To the Faculty of Washington State University
The members of the Committee appointed to examine the dissertation of James A. Parejko find it satisfactory and recommend it be accepted.
Linda S. Thomashow, Ph.D., Chair
David M. Weller, Ph.D.
Michael L. Kahn, Ph.D.
William B. Davis, Ph.D.
ii
Acknowledgments
I am endlessly grateful to the support that the USDA-ARS Root Disease and Biological Control
Unit has provided for me in my years as a graduate student. I am thankful for the advisement and independence that Dr. Thomashow has given me and her unparalleled eye for the details has taught me to make every word and revision count. The encouragement from Dr. Weller to keep the big picture in mind has provided me with new perspectives on the microbial world and the shown me the importance of ‘keeping one foot in the lab and one foot firmly planted in the field’. The help, support and advice that Dr. Dmitri Mavrodi and Dr. Olga Mavrodi have given me over the last few years has helped me not only grow as a scientist, but also as a person.
I would also like to thank Dr. Kahn and Dr. Davis for serving on my thesis committee and providing me with advice and encouragement. Their help was crucial to successfully traversing the rolling hills of graduate school (metaphorically-speaking) from my first proposal as a second-year Ph.D. student to the Ph.D. candidate I am today.
I am extremely appreciative for the help provided by Karen Hansen, Dr. A. Kamil Mohd
Jaaffar, Jennifer Apple, Emilia Gan, Chelsea Stone, Irina Mavrodi and Nathalie Walter in completing the work described in this dissertation.
I am also grateful to the support provided by the School of Molecular Biosciences and
Department of Plant Pathology. Without the intellectual and finanacial support from both departments I wouldn’t have been able to complete the work described in this dissertation.
iii
PHENAZINE-1-CARBOXYLIC ACID-PRODUCING
PSEUDOMONAS SPP. OF THE INLAND
PACIFIC NORTHWEST (U.S.)
Abstract
by James A. Parejko, Ph.D. Washington State University December 2012
Chair: Linda S. Thomashow
Phenazine antibiotic-producing rhizobacterial strains belonging to the genus Pseudomonas are effective biocontrol agents against soilborne fungal plant pathogens. Beyond the well-studied model phenazine-producing (Phz+) biocontrol strain Pseudomonas fluorescens 2-79, there existed few studies demonstrating phenazine production in root-colonizing members of the ‘P. fluorescens’ lineage. Furthermore, our collective knowledge of these important biological control rhizobacteria lacked a comprehensive analysis of representatives of Phz+ populations indigenous to intensive agricultural regions, most notably the Inland Pacific Northwest (U.S.) dryland agroecosystem. In the first of the studies within this dissertation, a collection of 412 Phz+
Pseudomonas spp. isolates from three cereal fields of east-central Washington State was genotypically and phenotypically characterized. The results revealed four new groups of Phz+ strains, three of which were divergent from P. fluorescens 2-79. These groups were associated
iv with dryland cereal crops with seven different cropping histories, however cropping history shaped the community of genotypes present. Another collection of 497 Phz+ isolates from throughout the Inland Pacific Northwest dryland agroecosystem was taxonomically described by multi-locus sequence analysis and the phenazine biosynthesis gene phzF was cloned from winter wheat at nine sites. Using this approach, it was determined that the four groups of Phz+ strains from the first study are separate Pseudomonas species, two of which have never been described, and agroclimatic conditions influence Phz+ community diversity. The influence of phenazine-1- carboxylic acid (PCA) on biofilm formation under water limitation and biocontrol characteristics against Rhizoctonia solani AG-8 were also determined for selected strains from the Phz+ species.
Generally the strains developed high levels of biofilm and responded uniquely to simulated matric or osmotic stress. The importance of PCA production to biofilm formation was variable by strain, although biofilm architecture was significantly impacted in most Phz- mutants.
Representative Phz+ strains protected wheat plants from the Rhizoctonia root rot causal agent R. solani AG-8 in greenhouse bioassays with reduced plant protection from Phz- mutants of the same strains. These findings significantly increase our understanding of indigenous Phz+ populations in dryland agriculture and their involvement in an effective cropping system for a more sustainable dryland agroecosystem.
v
Table of Contents
Page
Acknowledgments...... iii
Abstract ...... iv
List of Tables ...... x
List of Figures ...... xiii
General Introduction
Sustainable dryland cereal production and root-associated bacteria and fungi ...... 1
The ‘Pseudomonas fluorescens’ species-complex...... 12
Phenazines and the ecological implications for phenazine-producing Pseudomonas spp. in the rhizosphere micro-environment ...... 14
Microbial population genetics and the emerging biogeography of indigenous phenazine- producing P. fluorescens strains ...... 25
Goals of this dissertation ...... 29
References cited ...... 32
Specific contributions to this work ...... 45
vi
Chapter 1. Population structure and diversity of phenazine-1-carboxylic acid producing fluorescent Pseudomonas spp. from dryland cereal fields of central Washington State (U.S.)
Abstract ...... 47
Introduction ...... 48
Materials & methods ...... 49
Results ...... 60
Discussion ...... 82
References cited ...... 89
Chapter 2. Taxonomy and distribution of phenazine-1-carboxylic acid-producing Pseudomonas spp. in the dryland agroecosystem of the Inland Pacific Northwest (U.S.)
Abstract ...... 98
Introduction ...... 99
Materials & methods ...... 103
Results ...... 113
Discussion ...... 127
References cited ...... 135
vii
Chapter 3. Significance of phenazine-1-carboxylic acid (PCA) to biofilm production and colony biofilm morphology of phenazine-producing Pseudomonas species under stress
Abstract ...... 142
Introduction ...... 143
Materials & methods ...... 147
Results ...... 154
Discussion ...... 167
References cited ...... 172
Chapter 4. Biological control of Rhizoctonia solani AG-8 by phenazine-1-carboxylic acid- producing Pseudomonas spp. isolated from the Inland Pacific Northwest (U.S.) dryland agroecosystem
Abstract ...... 180
Introduction ...... 181
Materials & methods ...... 184
Results ...... 190
Discussion ...... 198
viii
References cited ...... 202
General Conclusions ...... 207
ix
List of Tables
Page
Chapter 1
1. List of Phz+ Pseudomonas strains used in this study ...... 51
S1. Distribution of BOX-PCR genotypes throughout sampled sites ...... 52
2. Five alpha ecological biodiversity indices of sampled sites as a function of Phz+ genotypes ...66
3. Two tests conducted to detect recombination events in recA or phzF sequences within Phz+ clusters ...... 72
S2. Population-wide recA and phzF nucleotide sequence distance and diversity calculations .....73
S3. Sixty-two Biolog substrates used for cluster analysis (as determined by principle components analysis) ...... 78
S4. Utilization data for seventeen common wheat rhizosphere exudates extracted from the
Biolog GN2 microplate dataset ...... 79
S5. Production of phenazine-1-carboxylic acid and biosurfactants by 31 Phz+ genotypes ...... 82
Chapter 2
S1. Characteristics of 2009 and 2010 sites sampled for Phz+ strains ...... 106
1. Characteristics of sites sampled for total rhizosphere DNA ...... 109
x
2. Population-wide (n = 88) descriptive analyses of phzF and housekeeping genes from Phz+
Pseudomonas spp...... 117
3. The MLSA-based sequence dissimilarity comparision of Phz+ strain groups to species of the P. fluorescens lineage ...... 119
S2. Total number of Phz+ Pseudomonas spp. strains isolated from sampled sites ...... 121
S3. Distinct phenotypic differences between the four IPNW Phz+ Pseudomonas species identified in this study ...... 129
Chapter 3
1. Bacterial strains and plasmids used in this study ...... 150
Chapter 4
1. Bacterial strains and plasmids used in this study ...... 186
2. Biological control of Rhizoctonia solani AG-8 C-1 by PCA-producing Pseudomonas spp. in natural soil...... 194
3. Biological control of Rhizoctonia solani AG-8 C-1 by PCA-producing Pseudomonas spp. in pasteurized soil ...... 195
xi
4. Biological control of Rhizoctonia solani AG-8 C-1 by wild-type Phz+ and Phz- mutant
Pseudomonas spp. in pasteurized soil ...... 197
xii
List of Figures
Page
General Introduction
1. Agroclimatic zones (A) and mean annual precipitation (mm, yrs 1971 to 2000)
(B) of the Inland Pacific Northwest in the western contiguous United States ...... 5
2. Dryland winter wheat and fallow fields in late summer near Ritzville, WA ...... 6
3. Rhizoctonia bare patch of dryland wheat at the Ron Jirava Farm and Washington
State University rotation plots near Ritzville, WA ...... 11
4. Enzymatic reactions involved in phenazine biosynthesis in fluorescent Pseudomonas spp. and other phenazine-producing bacteria (from Mavrodi et al. 2012) ...... 20
Chapter 1
1. BOX-PCR dendrogram of 31 Phz+ genotypes and four major clusters ...... 64
2. Phylogeny inferred from data for 1,297-bp fragments of 16S rDNA of 31 genotype representatives and closely related members of the “P. fluorescens” complex ...... 69
3. Neighbor joining trees of recA sequence (518 nt positions); a) as compared to phzF
(389 nt positions); b) and translated phzF sequence (129 amino acids); c) for members of the four BOX-PCR clusters ...... 74
xiii
4. Split decomposition network analysis of Phz + clusters 1, 3, and 4 where significant
(P<0.05) evidence of recombination within phzF was found ...... 75
5. Dendrogram of ODM values for utilization of 62 Biolog GN2 carbon substrates by the 31 genotype members of the four main BOX-PCR clusters ...... 80
Chapter 2
1. Distribution of sites sampled for Phz+ populations throughout the Inland Pacific
Northwest (U.S.) ...... 105
2. Phylogenetic placement of Phz+ Pseudomonas spp. isolates in the ‘P. fluorescens’ and ‘P. gessardii’ sub-groups as defined by Mulet et al. (2010) ...... 118
3. Unrooted phylogenetic tree of partial phzF (389 nt) DNA sequence from MLSA- representative strains (ntotal = 113) and phzF clones (ntotal = 454) ...... 124
S1. Rarefaction curves for the Phz+ communities from the eight sites sampled for total rhizosphere DNA ...... 125
4. The effect of soil % silt and agroclimatic zone on the diversity of phzF-containing
Pseudomonas spp. communities at eight sampled IPNW sites ...... 126
xiv
Chapter 3
1. Growth of Phz+ and Phl+ Pseudomonas spp. and their Phz- and Phl- mutants on 1/3
KMB amended to simulate matric stress (A) or osmotic stress (B) ...... 156
2. Total biofilm formation by Phz+ wild-type and Phz- mutant Pseudomonas spp. strains under matric (PEG 8000, blue) and osmotic (NaCl, red) stress relative to unamended 1/3 KMB (grey) ...... 159
3. Total biofilm formation by Phl+ wild-type and Phl- mutant Pseudomonas spp. strains under (PEG 8000, blue) and osmotic (NaCl, red) stress relative to unamended
1/3 KMB (grey) ...... 160
4. Total biofilm formation by P. cerealis L1-45-08r and P. orientalis L1-3-08r and their Phz- mutants with 0, 25, 50 and 100 µg ml-1 of PCA in (A) unamended
1/3 KMB, (B) -0.25 MPa PEG amended 1/3 KMB and (C) -0.25 MPa NaCl amended
1/3 KMB ...... 162
5. Colony biofilm morphology of four Phz+ Pseudomonas spp. strains
(L1-45-08r, S36709r, L1-3-08r and 2-79RN10) and two Phl+ strains
(Q8r1-96 and Q2-87) and their Phz- or Phl- mutant derivatives ...... 166
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Chapter 4
1. In vitro inhibition of R. solani AG-8 C-1 by eight strains of Phz+ Pseudomonas spp. representing four different Pseudomonas species ...... 192
xvi
Dedication
To my loving parents, Ken and Judy, brother, John, and all of my family members:
I am forever indebted to you for the support and love that you have shown me through the some
of most trying times of my life thus far.
To my closest friend and loving partner of over four years, Lauren:
You have given me the strength to work through the hardest and most disheartening times and shown me your unconditional love. We have an endless number of unforgettable memories from
past times spent together and I am excited for the years ahead of us.
To my faithful canine friends, Duchess & Lyla:
Although you’ll never read this dissertation or understand the science described within, your
boundless excitement for the simplest things in life, from my return home to a walk down the
street, always warms my heart and puts a smile on my face.
And finally, to all of my teachers, friends, advisors and supporters:
The encouragement you have provided has helped me reach heights I would not have imagined
as a younger man. Every inspiring word has reassured me to continue being inquisitive as I
wander down life’s unexplored paths into the unknown.
xvii
General Introduction
Sustainable dryland cereal production and root-associated bacteria and fungi
Sustainability in dryland cereal production of the Inland Pacific Northwest
Globally, drylands (land in arid, semi-arid or dry subhumid climates) encompass at least 40% of total worldwide land area (44). Agricultural production in these climates is generally limited to crop production for one part of the year, or every other year, if annual precipitation is low (44).
Due to increasing water shortage frequencies and diminishing availability in many of these regions, irrigation is not economically or environmentally sustainable for most dryland farmers.
Rainfed dryland crop production is dependent upon annual precipitation patterns and has significantly decreased crop yields as compared to irrigated cropping. It relies upon stored soil moisture through fallowing, reduced or no till cultivation or mulching, to compensate for the lack of growing season precipitation needed in the growth and maturation of crops (44).
The Inland Pacific Northwest (IPNW) is an expansive region encompasses north central
Washington, northern Idaho and northeastern Oregon (Fig. 1). The Columbia Plateau is a
62,000-km2 scrub steppe and sagebrush dryland ecosystem of the IPNW that extends from easteran and central Washington State into northern Oregon (88). The Palouse region of eastern
Washington and northern Idaho, known for its rolling hills of deep fertile ancient flood-deposited loess soil, defines the eastern boundary of the Columbia Plateau. These geographic regions are intensively farmed for cereal crops, particularly the low precipitation zone (<350 mm annual precipitation) of the Columbia Plateau (>1.56 million cropland hectares) which has been defined as the ‘largest contiguous crop production region’ in the western U.S. (88).
1
There are six agroclimate zones that define the IPNW agroecosystem and are based upon soil depth, mean annual precipitation and cumulative growing degree days (22) (Fig. 1A). Zone
1, annual crop-wet-cold, is constrained to areas at high elevation bordering the primary cereal production areas of the Columbia Plateau and the Palouse. Zone 1 includes areas where farmers can plant an annual crop, precipitation is over 400 mm year per year, soil depth is not a defining factor and there are under 700 cumulative growing degree days per year. Zone 2, annual crop- wet-cool, differs from zone 1 only in the number of growing degree days (700 to 1000 growing degree days); however, the soil profile can be significantly deeper (greater than 60 m) and fertile.
The Palouse region, renowned for its deep fertile loess soils, is found in zone 2 and both zone 1 and zone 2 soils have relatively large amounts of organic matter. Zone 3, annual crop-fallow transition, occasionally receives too little annual precipitation to support an annual crop (350 to
400 mm mean annual precipitation) and has moderately deep loess-derived soils and less organic matter than zones 1 and 2. It defines the western edge of the Palouse toward the lower precipitation region of central Washington. Zone 4, annual crop-dry, generally receives between
250 to 400 mm annual precipitation, has shallow top soil (less than1 m deep; much of the zone is rocky scrubland and rangeland) and fewer than 1000 growing degree days. Zone 5, grain-fallow, defines much of the low annual precipitation (less than 350 mm) dryland agricultural region of
Washington state and northern Oregon (Fig. 1B). It is in this zone that cereal farmers must include an annual fallow to store enough moisture in the deep loess-derived soils for significant crop yields. Cropping in zone 6 requires the use of irrigation and is found deep in the rain shadow of the Cascades in west-central Washington.
Cultivable soils in agroclimatic zones 3 through 6 are predominantly loess-derived silt loam. Loess is composed of fine silt soil particles that were deposited predominantly during the
2 late Pleistocene (ca. 15,400-13,100 years ago) and early Holocene (ca. 13,100 years ago to present) by interglacial episodes of outburst flooding in the region and distributed by wind- erosion over time (93). Much of the region also was influenced by recent deposits of volcanic ash from the Mount St. Helens (32 years ago) and Mount Mazama (ca. 7,700 years ago) eruptions. Loess is generally characterized by less than twenty percent clay composition with sand and silt as the remainder. The silt loam soil texture found throughout the region is generally composed of 50-70% silt with less than 25% sand and defined by several soil types including
(west to east in Washington State) Quincy silt loam, Shano silt loam, Ritzville silt loam, Palouse silt loam and Thatuna silt loam. Generally the soil series range (west to east, dryland to wet-cold) from above or at neutral pH to more acidic pH, less than 1% organic matter to above 2% organic matter and higher percent sand with lower percent clay to lower percent sand and higher percent clay (74).
The dryland cereal crop grown on the Columbia Plateau and in the Palouse region of the
Inland Pacific Northwest is predominantly wheat, with barley grown in some areas as a rotation crop (89). Other potential rotation crops in the Palouse region are legumes, i.e. pea, lentil or chickpea, and brassicas such as winter or spring canola in regions with enough annual precipitation. Farmers in zones 1-4 have the option to plant spring wheat or winter wheat, depending on the rotation and management practices. In low precipitation regions such as those found in zone 5, farmers that do not have the capacity to irrigate are restricted to planting winter wheat every other year in rotation with a summer fallow (Fig. 2). Unfortunately, fallowing typically requires several passes with soil tilling equipment and often relies upon the use of a significant amount of chemical herbicides to reduce the growth of volunteer plants and aggressive weeds (89). Tilling and spraying are a major cost to the farmer and result in
3 significant environmental degradation through air pollution from wind erosion and the use of fossil fuels and non-target herbicides (14). ‘No till’ farming in dryland agroecosystems was developed to make crop production more sustainable by reducing economic hardships on farmers, retaining soil moisture, reducing fossil fuel consumption and reducing the plethora of problems associated with wind erosion and airborne particulates (14). Unfortunately a major obstacle to greater adoption of ‘no-till’ farming by cereal producers is yield losses due to damage caused by specific soilborne root pathogens.
4
5
6
The chemical warfare between root-associated bacteria and pathogenic fungi
Plants direct a significant amount of photosynthate to the roots, 20-30% of fixed carbon in cereal crops (45), and roughly 11% of net fixed carbon into the region directly surrounding and influenced by the plant roots (41). Generally the region influenced by the plant roots is termed the ‘rhizosphere’ while, more specifically, the actual root surface is termed the ‘rhizoplane’ (50).
The domain within the plant tissue and epidermal cells is termed the ‘endorhizosphere’ and the zone extending just beyond the rhizoplane is termed the ‘ectorhizosphere’ (50). However, because it is difficult to delineate the zones of plant influence in the soil, ‘rhizosphere’ is often used to describe the soil influenced by the root zone. The spectrum of compounds exuded into and deposited in the rhizosphere by the plant depends on plant age, plant species and soil type (6)
In addition, the sloughing of root cap cells (known as ‘border’ cells) on actively growing roots is constant as long as the plant is growing (35). Additionally, the plant can exude certain compounds or increase root exudation in response to temperature and light fluctuations and to deal with a wide range of abiotic stresses like soil compaction, hypoxia, soil moisture, and nutrient deficiencies (6). Root exudates include low molecular weight organic and amino acids, sugars, fatty acids, phenolic compounds and a wide spectrum of secondary metabolites as well as higher molecular weight proteins, enzymes and sloughed border cells that can compose mucilage
(6). Exudates are released over the entire length of the root, however specific sites of root exudation are found at the primary and lateral root tips (62) and the branchpoints of lateral roots where temporary wounds in the root surface leak exudates into the surrounding soil matrix (96).
Rhizodeposits like sloughed cells generally are released closer to the actively growing root apical meristem (19). Root cells also can utilize specific transporters to exude sugars and amino acids while higher molecular weight compounds are exuded using vesicle-trafficking (6).
7
The carbon-rich micro-ecosystem of the rhizosphere serves as a nutrient source for soil- dwelling organisms. Plant seeds and roots are colonized by soil-dwelling bacteria, fungi and microscopic eukaryotes and the composition of the associated ‘microbiome’ shifts as the plant develops from seed to seedling (73). This transition in the compositions of the microbiome is due largely to root exudation processes that initiate as seedling growth progresses. Motile soil- dwelling microbes can chemotactically migrate toward the root in response to a gradient in exudate concentration and other soil-dwelling microbes come into contact with the root as it grows through the soil profile. Depending on the relative competitiveness of the microbes, successful root colonization may occur (11, 49). The exact determinants of colonization likely depend in part upon soil type, plant root structure and exudate composition, but the production of surface adhesins like pili, fimbriae, flagella and O-antigen lipopolysaccharides contribute to the process of bacterial colonization by promoting reversible cellular attachment to the root (49, 97).
When successful colonization occurs, the competency of microbes in the rhizosphere is highly dependent on the utilization of root exudates (48). In certain microbes, the production of bactericidal or fungicidal antibiotic-like compounds, some of which may also increase resistance to abiotic stress which indicates adaptability to fluctuating soil conditions, also can contribute to greater rhizosphere competence (28, 61). Once cell-to-root attachment has occurred, polysaccharide biosynthesis initiates a phase of irreversible cellular attachment that structurally resembles a biofilm (16). Throughout both phases of attachment, social coordination by quorum sensing, a process of cell-to-cell communication via the production of chemical signals like acyl- homoserine lactones (AHL), allows cells to aggregate into microcolonies and regulate specific population-wide processes (16).
8
Soilborne fungal pathogens infect plant roots at various stages of plant growth, causing plant diseases. Fungi are ubiquitous in the soil and are also attracted to the root system by root exudates. Fungal hyphae may infect through tears in the root surface where nutrients are released, but they also can release enzymes to degrade the plant cell wall to enter the root tissue.
The primary soilborne fungal pathogens of dryland cereals in the IPNW are Gaeumannomyces graminis var. tritici, Fusarium spp. (phylum Ascomycota), Rhizoctonia solani and R. oryzae
(phylum Basidiomycota) and the oomycete Pythium spp. (82). Gaeumannomyces graminis var. tritici (Ggt) hyphae infect plant roots primarily in high precipitation regions (above 450 mm annual precipitation) and in irrigated dryland fields to cause the disease take-all. Ggt hyphae block vascular plant tissue, resulting in reduced water uptake by the plant, necrotic root tissue and stunted plant growth or death. Fusarium spp. and Pythium spp. generally are found in non- irrigated dryland fields and wet-cold climates, respectively. Fusarium culmorum and F. pseudograminearum are primarily responsible for Fusarium foot rot, which involves the loss of water conductance because of the decay of basal stem tissues, resulting in plant death and empty grain heads (82). At least six Pythium spp. in the IPNW (81) can cause root rot or damping-off diseases and generally are distributed as a function of climate, with the most severe root rot generally found in wetter climates with moist soil (82). Rhizoctonia solani is the most aggressive
Rhizoctonia species in dryland cereal fields and causes Rhizoctonia root rot, which infects seminal and crown roots, resulting in root lesions and stunted root growth (82). Often R. solani mycelia infect multiple plants, resulting in large bare patches of dead or stunted plants in dryland fields (Fig. 3). Disease caused by R. solani is prevalent in direct-seeded (‘no-till’) fields, but may decline with long-term no-till crop management (82). This effect may be the result of a build-up of beneficial root-associated bacteria and/or fungi.
9
Interactions between fungal phytopathogens and root-associated bacteria are likely the result of evolutionary forces driving competition-related traits when the two groups co-occur in the rhizosphere. For instance, fusaric acid produced by some pathogenic Fusarium spp. negatively impacts the expression of genes related to the biosynthesis of the antibiotic 2,4- diacetylphloroglucinol in root-associated bacteria (71). Conversely, non-pathogenic Fusarium spp. are known to interact with other indigenous root-associated bacteria to suppress pathogenic
Fusarium spp. (60). This interaction may be due to out-competing pathogenic Fusarium spp. in the rhizosphere, or result from a more complex interaction. Among the best studied and most ubiquitous root-associated bacteria that are highly competitive in the rhizosphere, can suppress a range of soilborne fungal phytopathogens and interact with other rhizosphere microbial community members are the γ-proteobacteria of the species complex Pseudomonas fluorescens.
10
11
The ‘Pseudomonas fluorescens’ species-complex
The genus Pseudomonas originally was described in 1894 by Walter Migula (68), and our understanding of its complexity has evolved over the years as new species have been described and new methods to reveal additional diversity within those species have been developed. The type species of the genus is Pseudomonas aeruginosa, although recent DNA-sequence-based analyses suggest that the genus contains at least eleven clades of closely related type strains with the P. aeruginosa lineage diverging markedly from another lineage, the Pseudomonas fluorescens lineage (70). Interestingly, although P. aeruginosa is the type species of the genus, it differs both genetically and phenotypically from most of the described Pseudomonas species (70,
92). For example, P. aeruginosa has low genome diversity compared to other Pseudomonas species (90), possibly due to the narrow range of environmental niches from which it has been isolated. In contrast the P. fluorescens lineage contains the largest number of species, and these have been isolated from a variety of ecosystems including bulk and rhizosphere soil. This lineage contains several sub-groups, one of which is the ‘P. fluorescens sub-group’ (70) that encompasses the rhizosphere bacteria on which this dissertation focuses.
From biovars to 16S rRNA to genomes: the P. fluorescens sub-group
Preceding Migula’s identification of Pseudomonas as a genus, in 1886 Carl Flügge had described two ‘biotypes’ of what was termed Bacillus fluorescens based on gelatinase production (P. fluorescens produces gelatinase and liquefies gelatin whereas the other biotype, later named
Pseudomonas putida, does not) (25). This likely represents the first recorded observation of a clear phenotypic difference in fluorescent pseudomonads. Forty years later, Den Dooren de Jong
(18) recognized that fluorescent pseudomonads were trophically flexible and diverse, an
12 observation that continues to confound species descriptions within the genus even today. Since then, it has been recognized that although all members of the P. fluorescens species-complex
(and some in other groups, i.e., P. putida) produce fluorescent yellow-green siderophore pigments known as pyoverdines under iron limited conditions, their production may be one of the few phenotypic characteristics that maintain P. fluorescens as a loosely cohesive group. A significant number of studies into the taxonomy of Pseudomonas have generally concluded that
‘P. fluorescens’ is better described as a species-complex or species group (5, 69, 70, 92). It is because of these poor delineations that ‘biotypes’ of Pseudomonas traditionally were used in bacteriology to describe groups of strains with a small number of shared biochemical or physiological features. Accordingly, ‘P. fluorescens’ strains were separated into seven homogeneous groups based on twenty-two phenotypic traits including pigment production, biochemical reactions and carbon assimilation (92). These biotypes later were re-classified into five ‘biovars’ by Palleroni (78), who removed biotypes D and E which defined P. chlororaphis and P. aureofaciens, respectively.
In addition to biotypes and biovars, other attempts were made to define P. fluorescens as a species containing closely related groups of strains. These attempts included siderotyping (65,
67) the species complex into ten ‘siderovars’ (66), analyzing intra-genus relatedness using DNA-
DNA hybridization (76), defining five rRNA homology groups based on rRNA-DNA hybridization (77) and conducting a comprehensive study of 16S rRNA sequence phylogeny (5).
These different techniques were used extensively, albeit inconsistently, by researchers until multilocus DNA sequence-based methods emerged. Recently, DNA sequence-based analyses have been used successfully to restructure the evolutionary history of the P. fluorescens species- complex (3, 69, 70), which remains highly diverse and heterogeneous even today.
13
Phenazines and the ecological implications for phenazine-producing
Pseudomonas spp. in the rhizosphere micro-environment
Phenazines are a diverse group of heterocyclic redox-active nitrogen-containing compounds that contribute to the physiology and ecology of the various bacteria that produce them (57). Since the original description in 1859 of the blue-green phenazine pyocyanin (PYO) produced by P. aeruginosa (26), approximately 100 naturally synthesized phenazine derivatives have been discovered, and these display a wide range of functional groups (46). Phenazines are detectable due to their two distinct absorption maxima in the UV range and at least one more in the visible range. Each phenazine also has a unique redox potential (Eh) (10) determined in part by the substituent groups on the phenazine tricycle. PYO, phenazine-1-carboxylic acid (PCA), phenazine-1,6-dicarboxylic acid, phenazine-1-carboxamide (PCN), 1-hydroxyphenazine (1-
OHPHZ), 2-hydroxyphenazine (2-OHPHZ) and 2-hydroxy-phenazine-1-carboxylic acid (2-
OHPCA) are all phenazines known to be produced by Pseudomonas spp. (55). PYO, PCA, PCN and 1-OHPHZ have redox reaction couplings that exchange two protons and two electrons and whose redox potentials vary with pH (99). Generally, these redox potentials range
PYO>PCA>PCN>1-OHPHZ with over 100 mV difference between PYO and 1-OHPHZ, indicating that PYO and PCA are particularly good electron acceptors and, subsequently, that they can function as electron shuttles (99).
It has been recognized that phenazines have several physiological roles in cells that produce them. In some Pseudomonas spp., phenazines allow anaerobic growth by providing a conduit for extracellular electron transfer when terminal electron acceptors, like oxygen or nitrate, are absent from the local environment (98, 99). Phenazines also may allow the cell to
14 recover Fe(II) from biologically unavailable Fe(III) in ferric hyr(oxides), via electron shuttling
(37, 99). The endogenous production of some phenazines also promotes the formation and structure of biofilms, possibly as a result of the extent to which each phenazine encourages anaerboic survival and Fe(II) recovery (51, 52, 100). In addition to these beneficial activities for the producing cell, phenazines also have been investigated for antitumor activity against human cancers (13) and for biotechnology applications (84, 87).
The phenazine biosynthesis pathway, regulation and transport in
P. fluorescens 2-79
All phenazine-producing bacteria contain orthologs of the core phenazine biosynthesis operon
(phzABCDEFG), which encodes seven enzymes involved in the production of the core tricyclic structure (54). Phenazine biosynthesis is linked to the shikimic acid pathway, the source of the essential aromatic amino acids tryptophan, tyrosine and phenylalanine used as precursors for other essential cellular products or in peptide assembly via tRNAs. The aromatic amino acid precursor chorismate is the branch-point compound linking the shikimate and phenazine pathways and enhanced chorismate production in phenazine-producers likely is due to the plant- like 3-deoxy-D-arabinoheptulosonate 7-phosphate (DAHP) synthase enzyme PhzC, encoded by phzC (54). PhzC is not as well characterized as the other phenazine biosynthesis proteins.
However, consistent with its 39.1% amino acid identity to plant DAHP synthases, PhzC is thought to direct additional phosphoenolpyruvate and erythrose-4-phosphate into the shikimate and phenazine pathways by catalyzing the synthesis of DAHP and consequently chorismate.
Plant DAHP synthases are not regulated by aromatic amino acid feedback inhibition (38), a characteristic of PhzC that enhances chorismate flow into the phenazine biosynthesis pathway.
15
Chorismate is directed into phenazine biosynthesis via the 140-kDa 2-amino-2- desoxyisochorismate (ADIC) synthase homodimer PhzE encoded by phzE in P. fluorescens 2-79
(47, 54) (Fig. 4). Mg2+ is required for the binding of chorismate to PhzE, which undergoes a structural transformation when both bind to the active site, followed by binding of glutamine to the type 1 glutamine amidotransferase (GATase1) domain (47). Glutamine is required for the reaction that cleaves a hydroxyl group from chorismate. Additionally, deprotonation of free NH3 through an ammonia channel formed during enzyme transformational changes is required for the addition of a primary amine group to produce ADIC, the precursor for the next enzymatic reaction (47). PhzE is feedback inhibited by divalent cations like Zn2+, Mn2+ and Ni2+ (47). The
ADIC product is transformed to the next phenazine precursor, trans-2,3-dihydro-3- hydroxyanthranilate (DHHA), by vinyl ether hydrolysis performed by the 49-kDa dimer isochorismatase PhzD (79), The DHHA is then isomerized to an enol product by the 42-kDa dimer isomerase PhzF encoded by phzF (8). This precursor can undergo tautomerization to a reactive ketone that dimerizes to form the phenazine tricycle precursor through PhzF-mediated catalysis (8). The nucleotide sequence of phzF near the PhzF active site is highly conserved in
Pseudomonas spp., allowing for its use as a target in discovering new environmental strains of
Phz+ Pseudomonas spp. (57). The two nearly identical genes phzA and phzB encode 163 and 162 amino acid protein monomers, respectively, and are the only ones in the core phz operon not required for efficient phenazine biosynthesis (1, 8, 54). PhzB forms a small homodimeric protein without close identity to any known protein and with fold-predicted similarity to members of the
Δ5-3-ketosteroid isomerase/nuclear transport factor 2 family (1, 2) that can serve a multitude of functions in the cell. PhzB catalyzes a condensation reaction that produces the phenazine tricycle however the reaction proceeds without the enzyme in vitro, albeit at a significantly slower
16 reaction rate (1). PhzA is not involved in this condensation reaction and its function in the phenaine biosynthesis pathway is not clear at the present time (58, 64). Therefore, the likely evolutionary advantage of maintaining phzB in the genome lies in the interconnected nature of the phenazine biosynthesis pathway with the essential aromatic amino acid-producing shikimate pathway via chorismate. Since the shikimate pathway requires a significant amount of chorismate, which is also the initial precursor of phenazines, the cellular concentration of chorismate available for conversion to phenazine precursors by PhzE, PhzD and PhzF is hypothesized to be below the threshold required for the spontaneous occurrence of the reaction catalyzed by PhzB (1). Also, the precursor of the PhzB condensation reaction, 6-amino-5- oxocyclohex-2-ene-1-carboxylic acid, can undergo side reactions with other cellular proteins, so its efficient consumption in the cell is crucial to phenazine production. Thus PhzB not only accelerates the conversion of 6-amino-5-oxocyclohex-2-ene-1-carboxylic acid to the tricyclic phenazine precursor, but this acceleration also increases potential phenazine tricyclic precursor yields (1). The conversion of the PhzB product to phenazine-1-carboxylic acid, the final product of the pathway, is thought to be catalyzed by the 51 kDa dimer flavin-dependent oxidase PhzG encoded by phzG (80). PhzG has been hypothesized to be paralogous to the pyridoxine-5’- phosphate oxidase (pdxH in E. coli) involved in the final reaction of pyridoxal-5'-phosphate biosynthesis, and may be the result of a past pdxH duplication event in the genome (80). It is hypothesized that PhzG deprotonates the penultimate phenazine precursor to produce PCA in P. fluorescens 2-79 (1). The suite of core phenazine biosynthesis enzymes encoded by the phz operon performs similar functions in all Phz+ Pseudomonas spp.; however, the final phenazine products and regulation differ somewhat among different species.
17
The phenazine biosynthesis pathway is tightly controlled, likely due to its connection with the shikimate pathway. The regulation of phenazine biosynthesis is often portrayed as a signal cascade involving three important interacting signal-based systems. The production of
PCA and numerous other extracellular, ecologically relevant metabolites including exoproteases, cyclic lipopeptides and alginate is regulated by the GacS/GacA global regulation system in
Pseudomonas spp. (36). GacS is a sensor histidine kinase that recognizes a specific unknown environmental signal and translates it via phosphoryl transfer to the posttranscriptional response regulator, GacA (36). The GacS/GacA system represents the initial step in a signaling ‘waterfall’ that cascades via GacA into the next regulation system, which is composed of small non-coding
RNAs (sRNA). GacA positively regulates highly structured and abundant sRNAs that function to posttranscriptionally control metabolite production by binding to a specific RNA protein chaperone, Hfq, along with the target mRNA that results in sRNA to target mRNA 5’ pairing
(29). Depending on the sRNA and the target gene, the result can be mRNA degradation, mRNA stabilization or translational stimulation. The signal cascade involved in phenazine biosynthesis also includes operon-specific regulation through a quorum sensing system, represented by phzI/phzR in the PCA-producing strain P. fluorescens 2-79 (42). The phzI/phzR quorum sensing system is a member of the LuxI/R response regulator family and requires high cell density for expression of regulated genes. In strain 2-79, PhzI synthesizes six acylhomoserine lactones
(HSL), with N-(3-hydroxy-hexanoyl)-HSL as the cognate HSL of PhzR, an ambidextrous activator of the phz operon (42). When quorum levels of HSL are present, PhzR binds to N-(3- hydroxy-hexanoyl)-HSL and likely undergoes a conformational change that allows it to bind to a phz DNA box element just upstream of the -35 phz operon promoter element of phzA (42). The binding of PhzR to the phz box element activates transcription of the phz operon in addition to
18 autoinducing phzI and phzR resulting in increased levels of HSL and PhzR and increased expression of the phz operon. mRNA encoding PhzI also can be the target of sRNA to regulate
HSL signal production until a specific environmental stimulus is ‘sensed’ by GacS and the signal cascade begins. Additionally, the RNA polymerase stationary phase sigma-38 may provide another level of transcriptional control for phenazine biosynthesis, as it does for other antibiotic synthesis genes (32), although this has yet to be shown in phenazine-producing P. fluorescens.
Transport of phenazines into the extracellular environment remains somewhat unclear in most Phz+ Pseudomonas spp. Once PCA is synthesized, there are likely several active transport systems for releasing it. Upregulation of the multidrug efflux proton-dependent pump mexGHI- opmD (20) occurs in the presence of phenazine via the transcription factor SoxR and PCA is selectively transported by the MexGHI-OpmD pump in P. aeruginosa (21). Additionally, in the presence of phenazine, the cell increases general enzymatic antioxidant (e.g. catalase), superoxide dismutase and monooxygenase expression to counter the production of reactive oxygen species (ROS) by endogenous phenazine (20). Increased resistance to ROS also could allow passive diffusion of uncharged phenazine across the cell membrane in addition to active transport by efflux pumps.
19
20
Pseudomonas fluorescens 2-79: a model PCA-producing Pseudomonas strain
P. fluorescens 2-79 was originally isolated in 1979 from the rhizosphere of wheat grown in soil from a twelve-year continuous monoculture wheat field suppressive to take-all disease on the
Washington State University Lind Dryland Research Station northeast of Lind, WA (U.S.) (101).
Strain 2-79 has gained prominence as a model Phz+ and competitive rhizosphere-colonizing
Pseudomonas strain. Interest in P. fluorescens 2-79 arose from observations of its biological control activity toward soilborne fungal pathogens. In field tests, 2-79 showed significant suppression of take-all disease in both fumigated and non-fumigated soils, and Weller and Cook
(101) hypothesized that antibiotic synthesis by 2-79 was a factor in Ggt inhibition. Initially, the antibiotic was described as phenazine-like (31) and was further characterized as a heterocyclic nitrogenous phenazine-1-carboxylic acid (PCA) compound (9) with pH dependent biological activity toward soilborne fungal phytopathogens in vitro. Studies by Thomashow, et al. showing the role of PCA production in situ by 2-79 in controlling Ggt in the wheat rhizosphere (94) and the accumulation of PCA in the rhizosphere as well as the antibiotic’s specific role in pathogen suppression (95) were the first to demonstrate the effect of PCA on a fungal pathogen and were fundamental to understanding the importance of phenazines in the environment.
In addition to PCA, P. fluorescens 2-79 produces other compounds that have putative biocontrol properties against soilborne phytopathogens including pyoverdine siderophores, anthranilic acid (34), and a viscosin-like cyclic lipopeptide biosurfactant (unpublished data). The relative importance of these compounds in controlling fungal phytopathogens appears to be subordinate to the effect of PCA, although their effectiveness likely depends on the fungal phytopathogen and environmental conditions that are encountered. For instance, PCA is the
21 dominant biocontrol compound against Ggt when compared with siderophore and anthranilic acid using transposon mutants defective in the biosynthesis of each compound (34).
PCA has significantly diminished antifungal activity in media with pH >7.0 (PCA has a pKa of 4.24) as it is fully deprotonated and in its ionic form, phenazine-1-carboxylate, which has no biological activity (9). The optimal inhibition of Ggt by purified PCA was demonstrated at pH
6.0 (75). Batch liquid-culture PCA production by 2-79 was optimal at pH 7.0 and 25-27°C, yet diminished at higher pH values and temperatures (91). This pH and temperature dependency suggests that regulation of the PCA biosynthesis pathway in 2-79 may be influenced by pH and temperature, environmental factors that fluctuate seasonally in the field. Other Phz+
Pseudomonas spp. also are known to regulate certain phenazine-related genes when temperature changes occur (40). Carbon source also had a profound effect on PCA production in
2-79, and PCA was produced to the greatest concentration (0.31 g PCA g-1 biomass) on glucose, indicating significant effects of specific root exudate components on cellular PCA production
(91). However, effects of pH and carbon source on the biosynthesis and biological activity of
PCA are not necessarily representative of effects in the soil matrix and the rhizosphere.
Phenazine-producing strains within the P. fluorescens lineage have only infrequently been reported, in contrast to strains of P. chlororaphis subsp. aureofaciens and P. aeruginosa
(92) in which phenazine production is the rule. Phz+ ‘P. fluorescens’ represent a group of relatively rare Pseudomonas spp. that are highly divergent from P. aeruginosa and Stanier’s biotypes D, E and F (92). Biotypes D, E and F define P. chlororaphis, P. aureofaciens and P. lemonnieri, respectively, and historically were considered to be the only groups of Phz+ ‘P. fluorescens’ outside of P. aeruginosa (92). It has become increasingly apparent that there exist
Phz+ phylogenetic ‘hotspots’ in the P. fluorescens species-complex and as more evolutionarily
22 distinct Phz+ strains are discovered in diverse ecological niches the ecological role of phenazines may become more apparent.
The ecology of indigenous rhizosphere phenazine-producing Pseudomonas spp. and
the role of PCA
Rhizosphere competence is associated with several essential traits. The ability to catabolize a wide range of carbon substrates (e.g. root exudates), acquire iron from the surrounding soil using iron-scavenging siderophores, reduce nitrates via denitrification and communicate with nearby cells via acyl-homoserine lactone (AHL) signal synthesis are all traits that provide certain root- associated bacteria with a distinct advantage over bulk soil-inhabiting species (28). Phenazine production also is a key trait for rhizosphere colonization competence, and Phz- mutants are significantly impaired in rhizosphere colonization (28, 61). Phenazine production represents a trait that has evolved to confer a distinct advantage in the highly competitive rhizosphere niche, and it also provides increased resistance to certain environmental stresses (e.g., low iron availability) (28, 61). The conferred resistance to abiotic stress allows Phz+ cells to out-survive other rhizobacteria and correlates to the unique redox properties of the phenazine molecule. For instance, as a direct result of their electron shuttling capacities, PCA, PYO and 1-OHPHZ allow
Δphz mutants to survive anaerobic conditions in an endogenous phenazine-specific manner (98).
Similarly, phenazines help the cell maintain redox homeostasis by passing electrons from intracellular NADH to oxidized extracellular phenazine in low oxygen conditions such as those encountered in a biofilm (85).
Biofilm formation in some phenazine-producing Pseudomonas spp. is dependent upon phenazine production (51), and different phenazines play unique roles in structuring biofilm architecture
23
(52). For instance, biofilms of P. chlororaphis subsp. aureofaciens, producing PCA are structured in a more heterogeneous and cell-dense manner less prone to dispersion forces, than biofilms composed of cells producing different ratios of PCA and 2-OH-PCA (52). Biofilms containing phenazine- producers also are able to influence the electrochemical properties of the surroundings by creating a redox gradient, or ‘electrocline’, which extends up to 400 µm beyond the surface of biofilms formed by P. aeruginosa (43). Nearly all phenazines are in the reduced form at 100 µm from the biofilm (43). The presence and size of the phenazine electrocline was a
- 3+ function of alternative electron acceptors such as NO3 or Fe , and resulted in increased soluble
Fe2+, albeit with a decreased electrocline size (43). Biofilms containing P. aeruginosa were able to reduce several mineralogical forms of ferric iron (37, 99), increasing levels of available ferrous iron that aid in biofilm formation (100). Ferric hydr(oxides) such as Fe(OH)3 were reduced to soluble ferrous iron by phenazines at pH values below 7 to 8 depending on the ratio of
Phzred/Phzox (37), suggesting that under normal soil conditions phenazines should increase bioavailable ferrous iron levels. Additionally, reduced PCA reacted with Fe(OH)3 at a higher rate than α-Fe2O3 at all pH values between pH 5 and pH 8, decreasing in reactivity as pH increased
(99). However, between pH 5 and pH 6, the reaction rate of reduced PCA with Fe(OH)3 was nearly zero, indicating a pH independence for reductive dissolution of Fe(OH)3 in this range
(99). Ignoring adsorption to soil particles, PCA would be more reactive toward both Fe(OH)3 and
α-Fe2O3 in dryland soils, which are generally slightly acidic to slightly alkaline (pH 6.5 to pH 8).
Cells of P. aeruginosa were able to modulate iron-uptake systems depending on phenazine concentrations. In the presence of exogenous PYO, cells decreased expression of iron transport receptors including the fpvA receptor for ferripyoverdine (20), in preparation for an influx of available iron. Other genes, including efflux- and redox-related genes, also were upregulated via
24 transcriptional regulators like SoxR independent of superoxide formation when the cells of P. aeruginosa were in the presence of PYO, indicating a specific signaling role for PYO in stationary phase (20). This specific cellular response to phenazine was also prominent in shaping the architecture of phenazine-producing bacterial communities (21) and may have profound effects on the physiology of a diverse range of prokaryotes. As a direct result of abiotic stress resistance, respiratory flexibility and phenazine-dependent biofilm tolerance, phenazine- producing bacteria may be particularly well-adapted to unsaturated conditions like those found in dryland soil.
Microbial population genetics and the emerging biogeography of indigenous
phenazine-producing P. fluorescens strains
Our understanding of the global distribution and diversity of indigenous phenazine-producing strains belonging to the P. fluorescens lineage is surprisingly limited. Detectable populations of indigenous Phz+ Pseudomonas spp. have only been reported in the rhizosphere niche, which is likely a result of the metabolic requirements for phenazine production, a key trait for root colonization and competition. The majority of research on Phz+ Pseudomonas spp. has relied upon culture-based isolations of a relatively small number of biocontrol strains from the rhizosphere. Although this has resulted in significant advances in biocontrol research, the disparate nature of isolation source data, isolation techniques and laboratory-specific practices makes studying the global distribution and ecology of phenazine-producing Pseudomonas spp. a nearly impossible endeavor.
Globally, there are very few examples of agricultural soils containing indigenous Phz+ populations of the ‘P. fluorescens’ lineage with specific biological roles. Phz+ strains related to
25 the ‘P. fluorescens’ species complex have been isolated from Chinese wheat fields (103) and the
Châteaurenard soil in southern France (4, 60). The Phz+ strains from these examples suppress take-all disease of wheat caused by Ggt and Fusarium wilt of tomato caused by pathogenic
Fusarium oxysporum, respectively. Suppressive soils can exhibit general and specific suppression. General suppression is a non-transferable phenomenon, suggesting endemic microbial populations that are particularly adapted to the soil ecology of the specific region from which the soil came (102). Specific suppression is transferable between soils and is often attributed to a single group of microorganisms that function to suppress a specific soilborne phytopathogen. Suppressive soils generally support a specific enrichment of Firmicutes and
Actinobacteria as well as Proteobacteria (63). The Châteaurenard soil contains indigenous Phz+
Pseudomonas spp. populations, some of which are closely related to 2-79, that colonize flax and tomato (103-105 CFU g. root-1) (60). Three Pseudomonas spp. isolates from the flax rhizosphere are closely related to P. fluorescens 2-79 in phzC sequence and likely play a significant role alongside non-pathogenic Fusarium oxysporum in suppressing Fusarium wilt of flax due to PCA production. In the same Châteaurenard soil, twenty-six phzC-containing strains were isolated from tomato rhizosphere and bulk soil that were related to P. aureofaciens PGS12 and P. chlororaphis PCL1391. None of the P. chlororaphis-related isolates were from the flax rhizosphere and all of them were slightly less effective than the P. fluorescens 2-79-like isolates at suppressing Fusarium wilt in the presence of non-pathogenic F. oxysporum. The
Châteaurenard study was one of the first to observe the beneficial ecological interactions of indigenous Phz+ with other root-colonizing microbial communities in suppressive soils.
Observing and elucidating the biogeographic patterns of microbes: what’s the
point?
26
The study of the ecological interactions of microorganisms with their physical, chemical and biological surroundings was begun by early microbiologists who observed variations in microbial spatial distribution, the effects of chemical or temperature gradients on microbes, and the spectrum of species inhabiting various microecosystems. Sergei Winogradsky (1856-1953), inarguably the founder of modern microbial ecology, made the very first conclusive observations that showed the essential role of indigenous soil bacteria in maintaining homeostasis in natural systems through nutrient cycles like nitrification (23). One of the foundational hypotheses that he accepted and advanced, with inspiration from Louis Pasteur, was that specific bacteria in the environment were the cause, not the result, of specific environmental chemical transformations
(23). This hypothesis represents the foundation of modern microbial ecology, particularly in determining the role specific strains play in nutrient cycling, biotransformations of contaminants and recalcitrant minerals, and biodegradation. Winogradsky’s studies of chemolithoautotrophy
(23), which modern-day students of microbial ecology often study using the ‘Winogradsky column,’ are microcosm studies of microbial biogeography at the microscopic scale as only certain conditions allow for chemolithoautotrophic bacteria to thrive. A classic example is
Beggiatoa, which is responsible for sulfur oxidation in sulfur-rich environments as observed by
Winogradsky. The contemporary microbiologists of Winogradsky’s time, Martinus Wilhelm
Beijerinck (1851-1931) and Lourens Baas Becking (1895-1963), worked to inspire and compose, respectively, the phrase “Everything is everywhere, but the environment selects” (17, 72), a hypothesis that has historically been accepted by microbial ecologists. The formulation of this hypothesis was largely based upon the assumption that the lower the organization of an organism
(it was assumed microbes were of the lowest organization), the more likely it was to be globally dispersed and distributed (72). Beijerinck’s observations of the selective effects of enrichment
27 cultures to coax out otherwise undetectable microbial species from a complex microbial community likely dominated by other species (17) also provided evidence that microbial species were globally distributed. The modern-day discipline of microbial biogeography has revolutionized our understanding of microbial distribution and ecology. It has been unable to disprove that ‘everything is everywhere,’ but it has revealed countless examples that the environment does indeed select (53).
The influence of biogeochemical and/or anthropogenic effects on microbial biogeographic patterns is profound and clearly has a selective force on the relative abundance, diversity and distribution of microbial communities (24, 30, 53). Studying the impact of these factors on root-associated microbial communities has been confounded by the complex nature of plant-microbe interactions, the elastic nature of plant physiology and the heterogeneity of soils from different geographic locations. Soil type, geographic location, plant host species and plant host habitat can all have a significant, direct influence on shaping the structure and function of soil and rhizosphere microbial communities (7, 12, 27, 33). By limiting the differences in soil characteristics and crop rotations to a single contiguous agroecosystem, it is possible to begin to examine potential environmental factors influencing indigenous rhizobacterial communities.
The impact of the environment on phenazine-producing P. fluorescens populations
Rhizosphere-associated microbial communities are generally impacted by plant host, edaphic characteristics and climatic conditions. However, an overarching factor in determining the distribution of the plant host is the regional climate, particularly in intensive dryland agriculture.
The climate has a profound effect on soil characteristics, in addition to precipitation patterns and agricultural crop growing degree days. Annual precipitation has a distinct effect on indigenous
28
Phz+ Pseudomonas spp. rhizosphere populations associated with cereal crops throughout the
IPNW (56). The populations are generally at biologically significant levels (105-107 CFU g.-1 root f.w.) as is the concentration of PCA (upwards of 1 µg g -1 root f.w.). However, the frequency of plant colonization decreases as the gradient of annual precipitation increases from the eastern slope of the Cascade Mountains east to the Palouse region (56). The distinct effect of soil moisture on wheat root colonization by Phz+ Pseudomonas spp. also can be observed in side-by- side irrigated versus non-irrigated wheat fields (59). Factors like wheat plant physiology may be influencing the frequency of these populations; however, Phz+ populations can readily be detected on the roots of native and invasive plants in undisturbed dryland soils (59). Together, these observations indicate that Phz+ Pseudomonas spp. are particularly adapted to rhizosphere conditions encountered in rainfed dryland cropping systems with low annual precipitation (<300 mm annually). It has been shown that soil matric potential affects the ability of 2-79 to colonize wheat roots; colonization was maximum at -140 kPa in non-sterile soil (39). Phz+ populations divergent from 2-79 in several molecular characteristics have been found in two dryland fields on the Columbia Plateau (57), indicating new groups of indigenous Phz+ Pseudomonas spp. that may be particularly adapted to drier conditions.
Goals of this dissertation
Preceding the discovery of significant Phz+ Pseudomonas spp. populations in dryland soils that receive <300 mm annual precipitation (57), indigenous populations were not detected in the dryland agroecosystem of the IPNW (86). By surveying non-irrigated, low precipitation dryland fields I was able to isolate and screen new rhizosphere-colonizing Phz+ Pseudomonas spp. from locations that had never been sampled. This collection of novel Phz+ strains provided the
29 opportunity to investigate strain diversity, molecular evolution, biofilm physiology and biocontrol capacity against R. solani AG-8.
The goals of Chapter 1 were a direct result of the discovery of large populations of Phz+
Pseudomonas spp. in three fields of the IPNW low-precipitation zone (57). The goals, founded upon the isolation of representative Phz+ strains from the rhizosphere populations of several common cereal crop rotations, were to 1) describe strain-level diversity by genotyping a representative collection of new isolates; 2) identify the most closely related type strains among
Phz+ isolates and explore inter-strain evolutionary relationships; 3) characterize strain phenotypes within the collection, particularly with regard to the phenazine production; and 4) determine if crop rotation history had an impact on the diversity of Phz+ genotypes isolated.
The primary goals of the studies in Chapter 2 were to further explore the diversity of Phz+
Pseudomonas spp. distributed throughout the IPNW and test the hypothesis, developed from the results of Chapter 1 and other studies (56, 59), that these populations contain new Pseudomonas species. The goals of the chapter were to 1) establish the taxonomic placement of Phz+
Pseudomonas spp. strains isolated from throughout the IPNW; 2) provide evidence for a new
Pseudomonas species based on DNA sequence analysis; and 3) explore the impact of certain abiotic environmental factors on the distribution and diversity of phzF alleles from distinct geographic locations.
The primary goals of the work in Chapter 3 were developed from the observation that
Phz+ population colonization frequencies were high in non-irrigated dryland fields in the low precipitation zone of the IPNW Columbia Plateau (56, 59). This observation suggested both that the populations had specific adaptations to low soil water availability and a possible role of PCA
30 in these adaptations. The goals were to 1) determine the effect of matric and osmotic stress on bacterial growth by representative Phz+ strains from Chapter 2; 2) determine the role PCA has in biofilm adhesion and biofilm structural properties by these strains; and 3) determine the impact
PCA has on biofilm formation under matric and osmotic stress in vitro.
The primary goals of the experiments in Chapter 4 were based upon observations that
Pseudomonas spp. that produce biosurfactants and phenazines are known be effective in suppressing soilborne fungal phytopathogens (15, 83), and that representative Phz+ strains from our collection may suppress Rhizoctonia solani AG-8 (56). My hypothesis is that these indigenous Phz+ strains may provide plant protection from Rhizoctonia root rot as a result of
PCA production. The goals of this chapter were to 1) determine whether representative Phz+ strains significantly inhibit R. solani AG-8 growth in vitro; 2) test that the same strains for control of R. solani AG-8 under greenhouse conditions and 3) examine the role of PCA in disease suppression by using PCA- mutants.
31
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44
Specific Contributions to this Work
Chapter 1: J. A. Parejko performed all experiments presented in this chapter. Dr. O. V. Mavrodi aided in the analysis of BOX-PCR banding patterns. Dr. D. V. Mavrodi aided in the analysis of
DNA sequence. Both Dr. D. V. Mavrodi and Dr. O. V. Mavrodi helped with the sampling effort at the Jirava farm, Kagele farm and WSU Dryland Research Station near Lind, WA. Dr. D. V.
Mavrodi, Dr. O. V. Mavrodi, Dr. D. M. Weller and Dr. L. S. Thomashow aided in the experimental design, data analysis and the preparation and revising of the manuscript.
Chapter 2: J. A. Parejko performed all experiments presented in this chapter. Dr. D. V. Mavrodi,
Dr. D. V. Mavrodi and Dr. D. M. Weller helped perform the 2008 and 2009 IPNW region-wide sampling effort. Dr. D. V. Mavrodi and Dr. L. S. Thomashow aided in data analysis and the preparation and revising of the manuscript.
Chapter 3: J. A. Parejko performed all experiments presented in this chapter. Dr. D. V. Mavrodi and Dr. L. S. Thomashow aided in data analysis and the preparation and revising of the manuscript.
Chapter 4: J. A. Parejko created phenazine-deficient mutants, with guidance from Dr. D. V.
Mavrodi, used in this study. Karen Hansen performed in vitro Rhizoctonia solani AG-8 C-1 inhibition assays. J. A. Parejko and Dr. A. K. Mohd Jaaffar performed all greenhouse
Rhizoctonia root rot biocontrol assays presented in this chapter. Dr. D. M. Weller and Dr. L. S.
Thomashow aided in data analysis and the preparation and revising of the manuscript.
45
Chapter 1
Population structure and diversity of phenazine-1-carboxylic acid
producing fluorescent Pseudomonas spp. from dryland cereal fields of
central Washington State (U.S.)
James A. Parejko1, Dmitri V. Mavrodi2, Olga V. Mavrodi2, David M. Weller3,
Linda S. Thomashow3*
1School of Molecular Biosciences, Washington State University, Pullman, WA 99164-4234
2Department of Plant Pathology, Washington State University, Pullman, WA99164-6430
3U.S. Department of Agriculture, Agricultural Research Service, Root Disease and Biological
`Control Research Unit, Pullman, WA 99164-6430
*For correspondence. Mail: USDA-ARS Root Disease and Biological Control Research Unit,
Washington State University, Pullman, WA 99163-6430. E-mail: [email protected]; Tel.:
(+1) 509-335-0930; Fax: (+1) 509-335-7674
Microbial Ecology 2012. 64(1):226-41
46
Abstract
Certain strains of the rhizosphere bacterium Pseudomonas fluorescens contain the phenazine biosynthesis operon (phzABCEDF) and produce redox-active phenazine antibiotics that suppress a wide variety of soilborne plant pathogens. In 2007 and 2008 we isolated 412 phenazine- producing (Phz+) fluorescent Pseudomonas strains from roots of dryland wheat and barley grown in the low-precipitation region (< 350 mm annual precipitation) of central Washington State.
Based on results of BOX-PCR genomic fingerprinting analysis these isolates, as well as the model biocontrol Phz+ strain P. fluorescens 2-79, were assigned to 31 distinct genotypes separated into four clusters. All of the isolates exhibited high 16S rDNA sequence similarity to members of the P. fluorescens species complex including P. orientalis, P. gessardii, P. libanensis, and P. synxantha. Further recA-based sequence analyses revealed that the majority of new Phz+ isolates (386 of 413) form a clade distinctly separated from P. fluorescens 2-79.
Analysis of phzF alleles, however, revealed that the majority of those isolates (280 of 386) carried phenazine biosynthesis genes highly similar to those of P. fluorescens 2-79. phzF-based analyses also revealed that phenazine genes were under purifying selection and showed evidence of intra-cluster recombination. Phenotypic analyses using Biolog substrate utilization and observations of phenazine-1-carboxylic acid production showed considerable variability amongst members of all four clusters. Biodiversity indices indicated significant differences in diversity and evenness between the sampled sites. In summary, this study revealed a genotypically and phenotypically diverse group of phenazine producers with a population structure not seen before in indigenous rhizosphere-inhabiting Phz+ Pseudomonas spp.
47
Introduction
Plant growth-promoting rhizobacteria (PGPR) are an expansive group of highly diverse bacteria that colonize plant roots and benefit from the unique microenvironment found in the plant rhizosphere. In return, PGPR provide the plant with certain compounds that directly or indirectly stimulate the plant’s growth [1]. The ‘Pseudomonas fluorescens species complex’ is one group of extensively studied and highly effective PGPR. Antibiotic production represents an essential mechanism by which some PGPR of the ‘P. fluorescens complex’ compete against other indigenous rhizobacteria, maintain significant rhizosphere populations, and subsequently provide effective suppression of soil-borne plant pathogens [2]. Redox-active phenazines are a structurally and functionally diverse group of potent antibiotics that are produced by several
PGPR and non-PGPR groups of bacteria, including members of Streptomyces, Pantoea,
Pectobacterium, Burkholderia, and strains of several fluorescent Pseudomonas species [3].
Phenazine-1-carboxylic acid (PCA) production by Pseudomonas fluorescens strain 2-79, isolated from the rhizosphere of wheat from near Lind, WA (U.S.) in 1979, was the first definitive example of suppression of take-all disease of wheat caused by Gaeumannomyces graminis var. tritici [4, 5]. Since this discovery, many studies have shown that P. fluorescens 2-79 and other phenazine-producing (Phz+) strains are effective biocontrol agents against common agriculturally significant soil-borne fungal pathogens (including Rhizoctonia, Fusarium, G. graminis var. tritici, and Pythium) when applied to seeds before planting [4, 6–12].
Although the application of Phz+ biocontrol strains to crops has been well studied, almost nothing is currently known about the structure, diversity, and ecology of indigenous Phz+ populations in agriculture. A recent study by Mazurier et al. [7] elucidated the population
48 structure and diversity of native Phz+ strains isolated from Fusarium wilt suppressive soils of the
Châteaurenard agricultural region of southern France. The study found that 29 indigenous Phz+
Pseudomonas strains could be differentiated into 11 BOX-PCR genotypes, two of which were highly similar to P. fluorescens 2-79 [7]. Strains of these genotypes maintained populations of
103 and 104 CFU g-1 root on flax and tomato, respectively, grown in the Châteaurenard wilt suppressive soils [7]. Intriguingly, the effectiveness of the Phz+ isolates in suppressing Fusarium wilt of flax was a function of the presence of non-pathogenic Fusarium oxysporum, indicating a complex interaction between suppressive agents [7]. The Phz+ isolates of the Châteaurenard are the first major indigenous population of Phz+ Pseudomonas species discovered to play a significant role in a natural biocontrol process.
The recent discovery of abundant Phz+ Pseudomonas spp. populations in dryland wheat and barley fields throughout the Columbia Plateau of central Washington State, U.S.A., represents the second global example (first in the U.S.A.) of a highly productive agricultural region replete with indigenous Phz+ biocontrol populations [3, 13]. Considering the geographic location of the sites sampled, the hypothesis of the present study was that the populations reported by Mavrodi et al. were genetically and phenotypically similar to P. fluorescens 2-79.
The aims of the current study were three-fold: 1) to determine the genotypic and genetic diversity, supported by phenotypic evidence, of the Phz+ Pseudomonas spp. isolated from these abundant populations, 2) to determine if there was clear Phz+ genotype selection or diversification under different common dryland crop rotations and 3) to look for recombination in phzF, which should help improve our understanding of how indigenous Phz+ populations diversify over evolutionary time. To address these aims, Phz+ Pseudomonas strains were isolated from three sites in central Washington State and were analyzed using several molecular
49 techniques. The effect of crop rotation on the population diversity was also investigated using several popular biodiversity indices. This study represents the first step toward characterization of the population structure of beneficial indigenous Phz+ Pseudomonas spp. colonizing the roots of commercially grown cereal crops in the Inland Pacific Northwest of the United States.
Materials & methods
Sampled sites. Fluorescent Pseudomonas spp. were isolated from rhizospheres of wheat and barley collected at three separate sites in central Washington (Table 1 and Table S1). The first site was a 9-ha parcel of land at the Ron Jirava farm 8.5 km west of Ritzville, WA (N 47°8' 37"
W 118°28' 5"). The site has been used by Washington State University for 12 years to investigate the effect of crop rotation, fallow, and tillage on yields, soil erosion and management of soil- borne pathogens in cereal-based cropping systems [14]. The site is characterized by average annual precipitation of <301 mm and Ritzville silt loam soil with pH 8.29 and organic matter content of 0.93%. The second sampled site was represented by a commercial irrigated wheat field at the Don Kagele farm 15.5 km west of Ritzville, WA (N 47°7' 44" W 118°34' 27"). The field is irrigated by a central-pivot system and is predominantly cropped to winter wheat.
Excluding irrigation water, the average annual precipitation and soil type are identical to those at the first site. The third sampled site was a commercial dryland field with an alternating winter wheat-fallow rotation near the WSU Dryland Research Station at Lind, WA (N 46°59' 56" W
118°33' 47"). The site receives an average of 242 mm of precipitation per year and is characterized by Shano silt loam soil with pH 5.63-6.27 and organic matter content of 1.13-
1.14%. Percent calcium carbonate (CaCO3) for Ritzville silt loam and Shano silt loam is below
50
5% for the first 33-36 inches of the soil profile according to the National Resources Conservation
Service Soil Data Mart for Adams County, WA (http://soildatamart.nrcs.usda.gov, accessed
April 27, 2011).
Table 1. List of Phz+ Pseudomonas strains used in this study
a Isolate group Site Cropping history Number of Phz+ isolates
R1-X-08 1 Commercial winter wheat 72 R2-X-07 1 Spring barley/spring wheat 14 R2-X-08B 52 R2-X-08W 1 Spring wheat/spring barley 64 R11-X-07 1 Winter wheat/summer fallow 2 R3-X-08 54 R4-X-07 1 Continuous spring wheat 6 R5-X-07 R14-X-07 R15-X-07 R4-X-08 74 R6-X-08 2 Irrigated winter wheat 28 L1-X-07/08 3 Winter wheat/summer fallow 46
a Sampled sites are: 1, crop rotation plots at the Ron Jirava farm at Ritzville, WA; 2, commercial irrigated winter wheat field at the Don Kagele farm at Ritzville, WA; 3, commercial dryland winter wheat field at Lind, WA
51
Table S1. Distribution of BOX-PCR genotypes throughout sampling sites
Site & Crop Rotationa BOX-PCR Total per Total in Genotype R1 R2W R2B R3 R4 R6 L1 Total Cluster cluster collection 1 4 4 2 19 19 3 1 1 1 3 4 6 6 1 45 5 10 1 11 6 1 1 7 1 1 8 1 1 9 5 23 2 30 10 2 2 11 8 8 12 2 2 11 7 22 2 106 13 8 8 16 14 1 1 15 24 24 16 2 2 413 17 9 1 9 1 20 18 1b 1 19 1 1 3 27 20 1 1 21 4 4 22 7 7 23 16 4 20 24 6 6 25 1 1 26 13 3 32 6 12 66 4 235 27 39 2 2 1 10 54 28 1 1 29 2 10 12 30 2 17 17 1 2 2 7 48 31 18 2 20 aOnly pertaining to 2008 isolate nomenclature but includes all 2007 and 2008 isolates (see Table 1) bP. fluorescens 2-79 (not isolated in 2007 or 2008)
52
Phz+ strain isolation & screening. Plant samples were processed and Phz+ strains were isolated according to Mavrodi et al. [13]. Briefly, individual plant rhizospheres were excised and placed into 50-mL Falcon tubes pre-filled with 10 mL of sterile water. The tubes were vortexed for 1 min, sonicated in an ultrasonic bath (1 min) and the resultant rhizosphere soil suspensions were serial-diluted and spread on Pseudomonas Agar F (PsF, MP Biomedicals, Solon, OH) supplemented with ampicillin (40 μg ml-1), chloramphenicol (13 μg ml-1), and cyclohexamide
(100 μg ml-1) (PsF+++). Plates were incubated at 27°C for 24-48 hrs and isolated colonies were picked, spotted onto a new PsF plate, and incubated at 27°C overnight.
Colony PCR with primers Ps_up1 and Ps_low1 that target a highly conserved sequence within the phzF gene (P. fluorescens nomenclature) of the phz operon [3] were used to identify
Phz+ isolates. Each 15-μl PCR reaction contained a small amount of bacterial cells suspended in
1X Green GoTaq Flexi buffer (Promega, Madison, WI) supplemented with 200 μM deoxynucleoside triphosphates, 1.5 mM MgCl2, 20 pmol of each Ps_up1 and Ps_low1 primers, and 1 unit of GoTaq Flexi DNA polymerase (Promega). Amplifications were performed with a
PTC-200 gradient thermocycler (Bio-Rad, Hercules, CA). PCR conditions were: 1 min initial denaturation at 94°C followed by 30 cycles of 94°C for 20 sec, 57°C for 30 sec, 72°C for 25 sec, and brought down to 10°C before being stored at -20°C. Amplicons (5 μl) were separated on 1% agarose gels at 3.5 V/cm for 30 minutes and visualized with ethidium bromide. phzF-positive isolates were streaked on PsF and incubated at 27°C for 24-48 hrs to ensure a pure culture was present and stored as glycerol stocks at -80°C.
Isolates were screened for 2,4-diacetylphloroglucinol (phlD), pyrrolnitrin (prnC) and pyoluteorin (pltB) biosynthesis genes using the primers B2BF and BPR4 [15], PrnC1 and PrnC3
53
[16], and PltBf and PltB [16], respectively. PCR reactions were the same as for phzF and amplification conditions were identical to those published for each primer pair.
BOX-PCR genotyping. For whole-cell BOX-PCR amplifications, Phz+ isolates were grown overnight at 27°C in 100 μl LB broth in 96-well microtiter plates. Microtiter plates of overnight cultures were gently vortexed and optical density (OD) was read spectrophotometrically at 600 nm. Another 96-well microtiter plate was used to dilute isolates in sterile water to an OD600nm of
0.1. Plates were frozen at -20°C for at least an hour and thawed to room temperature to facilitate cell lysis. For DNA-based BOX-PCR amplifications, isolates were grown in 2 ml LB broth overnight at 27°C with shaking and total DNA was extracted from the overnight cultures using a cetyltrimethylammoniumbromide (CTAB) miniprep procedure [17]. DNA concentrations were determined using a Fluorescent DNA Quantitation Kit (Bio-Rad, Hercules, CA) and samples were diluted to 200 ng μl-1. The diluted DNA samples were profiled by BOX-PCR in duplicate using amplification conditions outlined below.
BOX-PCR amplifications were carried out with the BOXA1R primer [18]. Individual 25-
μl reactions contained 1x Gitschier buffer [19], 4.2 μg of bovine serum albumin, 5% (v/v) dimethyl sulfoxide, 12.5 μM deoxynucleoside triphosphates, 50 pmol of BOXA1R primer, 1.7 units of GoTaq Flexi DNA polymerase (Promega), and 0.5 μl thawed cell lysate or diluted DNA.
The cycling conditions were: an initial 2 min denaturation at 95°C followed by 30 cycles of 94°C for 1 min, 55°C for 1 min, 65°C for 8 min, and brought down to 4°C before storage at -20°C.
PCR products were separated on a 1.5% agarose gel in 0.5X Tris-borate-EDTA (TBE) buffer at
4°C and 4.5V/cm for just over 5 hr. Individual gels were stained with ethidium bromide and digitized using the Kodak 1D Image Analysis system (Eastman Kodak Co., Rochester, NY).
BOX-PCR banding patterns were analyzed using GelCompar 4.0 software (Applied Maths,
54
Kortrijk, Belgium) by the Pearson product-moment correlation coefficient and the average linkage (UPGMA) clustering method [20]. Genotypes were defined by the 95th-percentile similarity coefficient of replicate assays [20].
Genotype data analysis. Four commonly used diversity indices were calculated and interpreted according to Hill, et al [21] using the BOX-PCR genotyping results. The genotype diversity of each sample was represented by the Shannon-Wiener (H’ or eH’), log alpha (log α) indices and
Simpson’s index of diversity (1-D). The Pielou’s evenness (J) index was used to represent the genotype evenness of each sample. All four indices were calculated for R1, R2W, R2B, R3 and
R4 using the Vegan package v.2.0-1 [22] implemented in the BiodiversityR package v.1.6 [23] of the R statistical language v.2.13.2 (http://www.r-project.org). The irrigated (R6) and Lind,
WA sites (L1) had too few Phz+ isolates and therefore were not included in calculation of the four diversity indices described above. Rarefaction analysis of all seven samples was conducted in Analytic Rarefaction 1.3 (http://www.uga.edu/strata/software/win/aRarefactWin.exe) and rarefaction curves and the Margalef index (DMg) [21] were plotted and calculated in MS Excel,
H’ respectively. Increases in e , log α, 1-D or DMg values correlate to an increase in genotype diversity within the sampled Phz+ populations. A value of J closer to 1.0 indicates a sample with more evenly distributed genotypes among the total number of genotypes found.
A statistical test for genotype selection was performed in a similar manner to that described by Bergsma-Vlami, et al [24]. To test genotype selection, the frequency of genotypes present from each sample was calculated (number of genotype isolates/total number of isolates per sample) and then the percentage was arcsine transformed and tested using the nonparametric
Kruskal-Wallis test in XLSTAT (Addinsoft, New York, NY).
55
16S rDNA, recA, and phzF sequencing. The small-subunit ribosomal RNA (i.e. 16S rDNA,1455 bp), the housekeeping DNA repair recombinase gene recA (598 bp), and the phenazine biosynthesis enzyme gene phzF (427 bp) were amplified and sequenced essentially as described by Mavrodi et al. [3] using oligonucleotide primer sets 8F and 1492R (Tann = 55°C)
[25], recAf1 and recArps (Tann = 61°C) [3], and Ps_up1 and Ps_low1 (Tann = 57°C) [3], respectively. Two additional internal primers, 16Sps2 and r16Sps1 [3], were used to obtain the sequence of the entire 16S rDNA amplicon. Amplification products were cleaned with QIAquick
PCR purification spin columns (QIAGEN, Valencia, Calif., USA) and sequenced with a BigDye
Terminator v.3.1 Cycle Sequencing Kit (Applied Biosystems, Foster City, Calif., USA) according to the manufacturer’s recommendations. Sequence data were assembled and trimmed using Vector NTI Advance v.10 (Invitrogen, Carlsbad, CA).
Phylogenetic & molecular evolution analysis. 16S rDNA, recA, and phzF sequences were aligned using ClustalW implemented in Molecular Evolutionary Genetics Analysis software
(MEGA) version 5.03 [26] with default settings except for pairwise alignment gap opening penalty = 4 and multiple alignment gap opening penalty = 5. For the recA and phzF protein- coding sequences, the nucleotide sequences were translated in MEGA 5.03 and compared to those from P. fluorescens 2-79 to ensure codons were in the correct frame [3]. Gamma distribution shape parameters were also calculated in MEGA 5.03 using the same methods as the phylogeny reconstruction for each gene. Phylogenetic relationships were tested for confidence by bootstrapping with 1000 replications. The 16S rDNA phylogeny was constructed in MEGA 5.03 and recA, phzF, and translated phzF trees were constructed in SplitsTree4 [27].
The preliminary taxonomic placement of Phz+ isolates used in this study was established by comparing 16S rDNA sequences to those deposited in the Ribosomal Database Project (RDP;
56 release 10). The RDP blast database was restricted to type strains with 16S rDNA sequences of good quality and greater than 1200 nucleotides. 16S rDNA phylogeny was inferred using neighbor-joining (NJ) of genetic distances and evolutionary distances were computed using the
Kimura 2-parameter model with gamma distribution (α-parameter = 0.05) [28].
To study Phz+ population structure specific to genotype isolates defined by BOX-PCR and to compare to the phzF sequences, a NJ tree of the conserved housekeeping gene recA (518 nt) was constructed in SplitsTree4 [27] using the BIONJ method [29] and Kimura 2-parameter model with gamma distribution (α-parameter = 0.05) [28]. Interpopulation phylogeny of phzF
(389 nt) was inferred in the same manner as the recA phylogeny (phzF α-parameter = 0.19).
Translated phzF sequences (129 amino acid residues) were modeled using the Jones-Taylor-
Thornton (JTT) matrix amino acid substitution model with gamma distribution (α-parameter =
0.11) [30].
Population genetics of Phz+ isolates. To understand the extent of divergence between clusters of Phz+ isolates, nucleotide sequence distance and diversity calculations were done in MEGA
5.03 [26] for recA and phzF using the same parameters that were used for phylogeny inference.
Standard errors of distance and diversity values were calculated using the bootstrap method with
1000 repetitions. To test for recombination, split decomposition trees (same parameters as phylogeny inference) for recA and phzF were constructed for each separate cluster and tested for recombination using the φw-test [31] in SplitsTree4 [27]. Additionally, the clusters were tested using a different recombination test (coalescent simulation test) in DnaSP v5 [32]. To determine the method to use for detecting neutral, positive, or purifying selection, the transition/transversion ratio was first calculated in MEGA 5.03 [26]. The presence of neutral, positive, or purifying selection within the Phz+ clusters was tested using the codon-based Z-test
57 of selection in MEGA 5.03 [26] with the proportion-based Nei-Gojobori method [33] and bootstrapped with 1000 replications. To evaluate population structure and cluster-assignments, and to further investigate possible evidence for recombination events between clusters, Bayesian
Analysis of Population Structure (BAPS version 5.2) was implemented using population mixture and admixture analyses [34]. For population mixture analysis, sequence data were studied using the ‘clustering with linked loci’ function with K = 20. For population admixture analysis, the number of clusters generated from the population mixture analysis was used as the minimum population size analyzed with 100 iterations of simulated allele frequencies for the individuals,
200 reference individuals from each population, and 15 iterations of simulated allele frequencies for reference individuals. The genetic shapes of the population mixture results were graphed using the ‘changes of log likelihood’ function and genetic exchange within the available gene pool was visualized using the ‘plot gene flow’ function in BAPS.
Nucleotide sequence accession numbers. Sequence data for 16S rDNA were retrieved from
NCBI GenBank for ‘P. fluorescens complex’ strains for phylogenetic inference. Sequence data generated in a previous study for recA (in addition to 16S rDNA) were obtained from NCBI
GenBank for Pseudomonas spp. R14-24-07, R5-90-07, R11-45-07, R11-23-07, R2-7-07, R4-35-
07, R4-34-07, R5-89-07, L1-11-07, and P. fluorescens 2-79 [3]. Additionally, phzF sequence data were obtained for the same strains as well as Pseudomonas spp. R2-6-07, R2-10-07, R2-12-
07, R2-13-07, R2-30-07 and R15-2-07 [3].
Sequence data generated in this study have been deposited in the NCBI GenBank database under the following accession numbers: 16S rDNA sequence data were assigned numbers JQ361817 to JQ361844, phzF sequence data were assigned numbers JQ361845 to
JQ361872 and recA sequence data were assigned numbers JQ361873 to JQ361900.
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Biolog carbon substrate utilization assays. Isolates representing the unique genotypes were subjected to carbon substrate utilization profiling. Prior to analysis, the strains were pre-grown on Biolog Universal Growth (BUG) agar (Biolog, Inc., Hayward, CA) at 27°C for 24-48 hrs.
Several colonies of the same morphology were scraped off the plate and suspended in sterile water to produce an OD600 reading of 0.1. Then 1 ml of each cell suspension was added to 20 ml of GN/GP Inoculating Fluid (Biolog), mixed, and 150 μl aliquots were dispensed into wells of a
GN2 microplate (Biolog). Plates were incubated at 27°C and color development was measured after 4 and 6 days at 600 and 750 nm using a Safire microplate spectrophotometer (Tecan Group,
Ltd., Männedorf, Switzerland). Each strain was assayed twice. The difference of the two values was used for substrate utilization scoring with an OD600-750 of <0.1 as no respiration.
Analysis of the Biolog data was performed according to Alisi, et al. [35]. Briefly, the maximum OD (ODM) was an average of the duplicate measurements at four days and six days.
The ODM values for all 95 carbon substrates were consolidated into twelve groups based on structural similarities (carboxylic acids, carbohydrates, amino acids, polymers, nucleic acids, phosphorylated substrates, aromatics, alcohols, amides and amines, esters, brominated substrates, and miscellaneous substrates). Any substrate that had an ODM value below 0.1 for all isolates was removed from further analysis. The remaining substrates (65 total) were then analyzed by principal components analysis to determine the substrates that showed significant ODM variability across all of the genotype-representatives. Principal components were retained to account for at least 70% of the cumulative variability (essentially the first three or four components). The final dataset consisted of 62 substrates and was analyzed by multivariate clustering with Ward linkage and Pearson correlation coefficient distance measure. Clustering was considered significant at a percent similarity value at or above 75%. The principal
59 components analysis and the clustering of Biolog data were performed using Minitab 15
(Minitab, Inc., State College, PA, USA).
Drop collapse assay. Isolates were also tested for the capacity to produce biosurfactants. Liquid cultures in LB supplemented with 2% glucose and King’s medium B (KMB) [36] were grown at
28°C with shaking for 48 hrs and 25-μl droplets of each culture were pipetted onto a 12 cm x 10 cm rectangle of parafilm. After 5 to10 min the droplets were scored (+ or -) for droplet collapse as compared to a droplet of distilled water.
Phenazine extraction & detection. A liquid-state solvent-based extraction of phenazines from cultures was done according to Bonsall et al. [37]. Liquid cultures (4 ml) in KMB or LB supplemented to 2% with glucose were inoculated in a standard way and incubated for 48 hrs at
28°C with shaking. Cultures were acidified with 45 μl of 10% trifluoroacetic acid and extracted with 10 ml of ethyl acetate. The organic phase was separated by centrifugation and dried at room temperature for 3-5 days. The extracts were analyzed by either thin-layer chromatography (TLC) or reverse phase HPLC. For TLC, samples were dissolved in 20 μl of methanol and 2-μl aliquots were spotted on a Uniplate Silica Gel GHLF plate (Analtech, Inc, Newark, DE, USA) alongside a control of purified PCA. The separation was carried out in a 95:5 benzene:acetic acid system and detection was by UV at 254 nm [5]. HPLC was carried out on a Waters 2695 liquid chromatograph equipped with a 996 photodiode array and separated on a Nova-Pak C18 Radial-
Pak cartridge (4 μm, 8x100 mm) (Waters Corp., Milford, MA, USA). The photodiode array detection range was from 180 to 470 nm with screening for PCA at 248 nm. The HPLC gradient profile included a 2 min initiation at 10% acetonitrile/0.1% trifluoroacetic acid followed by a 20 min linear gradient to 100% acetonitrile/0.1% trifluoroacetic acid.
60
PCA was also extracted in the same manner as described above from colonies grown on tryptic soy broth (TSB) (BD Biosciences, Franklin Lakes, NJ) and potato dextrose agar (PDA)
(BD Biosciences, Franklin Lakes, NJ). Control and test PDA plates were spotted with 10 l of a
104 CFU ml-1 dilution (thus, ca. 102 – 103 cells were spotted) of an isolate grown overnight in 1 ml LB in quadruplicate. P. fluorescens 2-79 and phenazine-deficient mutants 2-79Z and 2-79IZ
[38] were spotted in the middle of separate test plates in the same manner as the isolates and used as controls. Results were recorded after growth at 27°C for 3 d, 6 d, and 8 d and considered positive for PCA production if dark green crystals were observed within the isolate spot. PCA extractions and TLC were performed as before to confirm PCA production.
Results
BOX-PCR genotyping. A total of 413 Phz+ isolates were subjected to BOX-PCR analysis.
Among these, 388 isolates were collected during the 2008 field season. An additional 24 Phz+ isolates were collected from the same sites during a preliminary study performed in 2007 and whose genetic relationships were included in a previous study [3]. P. fluorescens 2-79, a model
PCA-producing strain isolated from wheat grown in soil from a site in Lind, WA adjacent to one site sampled in this study, was also included in the analysis [4]. The comparative analysis of
BOX-PCR banding patterns was carried out in three stages. First, isolates from each location were profiled separately by whole cell BOX-PCR and the dendrogram topography and banding patterns were examined by GelCompar 4.0 software and confirmed by eye. Isolates with unique banding patterns and typical representatives of groups of isolates sharing highly similar patterns were identified and selected for further work. At the second stage, the selected representative
61 whole cell BOX-PCR fingerprints were combined in a single analysis to determine unique isolates across all three sampled locations. Unique fingerprints were determined in the same manner as described above. Finally, the resultant set of strains (n = 77) was subjected to genomic
DNA-based BOX-PCR amplification in which each strain was profiled independently twice.
The BOX-PCR profiling of genomic DNA resulted in distinct banding patterns ranging in size from 5 kb down to 400 bp (Fig. 1). Correlation-based cluster analysis revealed four large distinct clusters of BOX-PCR banding patterns at approximately 30% similarity. However, when unique BOX-PCR patterns were defined using the 95th-percentile definition of identity based on replicate assays for identical strains (corresponding to 72.5% similarity), the original four clusters were further divided into 31 unique genotypes. Of these, 30 genotypes corresponded to
Phz+ isolates from the 2007 and 2008 samplings, and the 31st genotype (genotype 18) corresponded to P. fluorescens 2-79.
BOX-PCR cluster 4 corresponded to the largest and most genotypically diverse group of
Phz+ isolates. The cluster encompassed ca. 57% (235 of 413) of the total isolates and ten clearly defined genotypes analyzed in this study. Of these ten genotypes, genotypes 26, 27, and 30 were most abundant and encompassed ca. 71% of the cluster 4 isolates. Considering all ten genotypes,
BOX-PCR cluster 4 is the only group of Phz+ isolates that are ubiquitous in the sampled sites, with high BOX-PCR profile similarities amongst genotypes (Fig. 1). The majority of cluster 4 isolates originated from the rotation plots near Ritzville, WA and only 17 isolates of genotypes
29 and 30 were isolated from Lind, WA.
BOX-PCR cluster 2 represented the second largest cluster at 106 Phz+ isolates. The cluster is dominated by genotypes 9, 12, and 15, which contain 76 of the 106 cluster 2 isolates.
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The majority of cluster 2 Phz+ isolates (83 of 106) were from the continuous spring wheat and winter wheat/fallow plots at the Ron Jirava farm. None of the isolates belonging to cluster 2 were isolated from the irrigated field or from the commercial winter wheat field near Ritzville, WA
(Table S1). The similarity of cluster 2 genotypes was low due to isolate R5-89-07 (genotype 16, cluster retained at similarity ca. 35%), which likely represents another BOX-PCR cluster entirely
(Fig. 1).
Cluster 1 was the third most frequently isolated BOX-PCR cluster at 45 Phz+ isolates.
Over half of BOX-PCR cluster 1 isolates (25 isolates) were found in the commercial winter wheat/fallow field near Lind, WA and 19 of those isolates belong to genotype 2. Like those of cluster 2, the BOX-PCR banding profiles of cluster 1 isolates were similar except for isolate L1-
44-08. L1-44-08 represents genotype 7 and the cluster is retained at ca. 60% similarity. No cluster 1 strains were isolated from the Ritzville commercial winter wheat field, the spring barley rotation, or the irrigated wheat fields.
The least isolated cluster was BOX-PCR cluster 3 with 27 Phz+ isolates. It contained Phz+ isolates that are genotypically closest to P. fluorescens 2-79 (cluster retained just below 50% similarity), although none of the isolates was found to be a member of the same genotype as P. fluorescens 2-79. Only one isolate of genotype 17 was isolated from Lind, WA and 20 of the 27 isolates belonging to cluster 3 were isolated from the two spring wheat plots at the Ron Jirava farm. The four isolates belonging to genotype 21 of cluster 3 were isolated exclusively from the irrigated field. As in cluster 1, no cluster 3 strains were isolated from the Ritzville commercial winter wheat field or the irrigated winter wheat field.
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Distribution and diversity of Phz+ BOX-PCR genotypes from different crop rotations. To determine if we had sampled a sufficient portion of the Phz+ rhizosphere communities in each sample, rarefaction curves were calculated. The curves for R1, R6 and R2B showed a distinct plateau; however the R2W, R3, R4 and L1 rarefaction curves were still approaching a plateau
(data not shown). Using the rarefaction analysis, the number of Phz+ isolates sampled was regressed to 27 isolates to correspond with the least sampled site (R6). Genotype richness was determined using the Margalef index (DMg) and revealed the same order of index values as seen with eH’ (see below), except R6 (0.91) and L1 (2.03) which were ranked higher than R1 (0.79) and R3 (1.85), respectively (Table 2).
The four diversity indices calculated for R1, R2B, R2W, R3 and R4 are shown in Table
2. The indices were calculated for R1, R2B, R2W, R3 and R4 as the sample sizes for all five are relatively close and they represent a wide range of crop rotations from a single site. The
Shannon-Wiener index (H’), represented here as its exponent (eH’) and moderately sensitive to sample size, was lowest for R1 (2.97), followed by R2B (3.91), R3 (5.48), R2W (6.88) and R4
(9.76). The values for the Simpson’s index of diversity (1-D) and log α index, which is not as sensitive to sample size, showed the same trends as eH’. The Pielou’s evenness index (J) ranged from R3 with the lowest value (0.739) to R4 with the highest (0.841).
The non-parametric Kruskal-Wallis statistical test of sample genotype selection showed that there was likely no genotype selection due to differences in the samples, as there was not significant evidence for population differences between sampled sites (p = 0.069, α = 0.05).
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Table 2. Five alpha ecological biodiversity indices of sampled sites as a function of Phz+ genotypes
Shannon- Pielou’s Simpson’s Margalef Rotation Wiener Log α (J) (1-D) (D )a (eH’) Mg
Commercial winter wheat (R1) 2.97 0.91 0.78 0.611 0.79
Spring barley/spring wheat (R2B) 6.88 1.60 0.76 0.676 1.27
Spring wheat/spring barley 3.91 3.82 0.80 0.817 2.15 (R2W)
Winter wheat/summer fallow 5.48 3.54 0.74 0.754 1.85 (R3)
Continuous spring wheat (R4) 9.76 5.45 0.84 0.858 2.88
Irrigated winter wheat (R6) - - - - 0.91
Winter wheat/summer fallow (L1) - - - - 2.03
a Calculated using rarefaction analysis E values regressed to 27 Phz+ isolates
Sequence-based phylogenetic analyses. We explored the taxonomic placement of genotype representatives (n = 31) by amplifying and sequencing 16S rDNA and comparing the sequences to those of Pseudomonas type strains available through the Ribosomal Database Project (release
10). Based on 16S rDNA identity, the majority of Phz+ isolates featured in this study were closely related to P. orientalis, whereas Phz+ isolates of genotypes 5, 6, 17, and 19-21 had 16S rDNA sequences nearly 100% identical to that of P. libanensis. The NJ phylogeny inferred from
1291-bp fragments of 16S rDNAs of the 31 representative Phz+ isolates and type strains representing different Pseudomonas species revealed that phenazine producers from Ritzville and Lind form two divergent and well-supported clades within the ‘P. fluorescens’ species
66 complex (Fig. 2). The first clade was larger than the second one and encompassed 24 Phz+ strains with 16S rDNA similar to that of P. orientalis. The clade contained two distinct clusters formed by sequences from Phz+ strains of BOX-PCR clusters 2 and 4, with the former being most similar to 16S rDNA of P.orientalis. Additionally, the first clade also contained the majority of strains belonging to BOX-PCR cluster 1 (Fig. 2). This clade contained Phz+ strains whose 16S rDNA sequences grouped closely with those from type strains of P. synxantha, P. gessardii, and P. libanensis. The second clade encompassed two genotypes (5 and 6) of BOX-
PCR cluster 1 and all strains of BOX-PCR cluster 3, including P. fluorescens 2-79.
The NJ phylogeny inferred from recA sequences was highly congruent with that derived from 16S rDNA, but had higher resolution of taxa within clades (Fig. 3A). Overall, the results of the recA-based analysis reinforced groupings observed through analysis of corresponding 16S rDNAs. Again, two well-supported major clades were observed, with clade 1 being larger and incorporating three distinct (also well-supported) clusters comprised of recA genes from Phz+ strains of BOX-PCR clusters 1, 2 and 4, respectively. The second clade contained sequences from BOX-PCR cluster 1 strains (genotypes 5 and 6) and all BOX-PCR cluster 3 strains (Fig.
3A).
We also established a phylogeny based on sequences of the key phenazine biosynthesis gene phzF. The overall topology of the resultant NJ phylogenetic tree was different from that of the recA-based phylogeny, but the main clustering was retained and had robust bootstrap support
(Fig. 3B). Four major clusters were observed that perfectly matched the corresponding groupings observed in the recA-based phylogeny. Like the 16S rDNA and recA-based phylogenies, the clade formed by phzF genes of P. fluorescens 2-79 and other strains of BOX-PCR cluster 3 also contained strains R14-24-07 and R2-4-08W, and R2-54-08W that belong to genotypes 5 and 6,
67 respectively, of BOX-PCR cluster 1. The placement of R5-89-07 (genotype 16) close to cluster
4, while its closest recA-based relatives were in cluster 2 suggested that R5-89-07 may represent an entirely different cluster (Fig. 3B). Another major difference from the recA phylogeny is the increased evolutionary divergence of cluster 2 from the rest of the clusters. To explore PhzF amino acid sequence within clusters, NJ phylogeny of translated phzF was built (Fig. 3C). The majority of the PhzF tree mirrored that of the phzF tree. There were two exceptions: L1-44-08
(genotype 7) was placed in cluster 4 rather than in its recA and phzF assignment in cluster 1, and
R5-89-07 (genotype 16) was placed into cluster 4 based on phzF, but placed into cluster 1 based on PhzF. However, the internal bootstrap support of clusters 1 and 4 was low compared to that of the phzF phylogeny (Fig. 3C).
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Analysis of Phz+ population structure. Clusters preliminarily assigned based on the recA and phzF phylogenetic analyses were tested for significance in BAPS [34]. The a priori cluster analysis found significant evidence for four clusters in both recA and phzF sequence data (data not shown). The four clusters mirrored the phylogeny results and placed isolate R5-89-07 in cluster 1 based on phzF sequence (which differs from phzF phylogeny but not PhzF phylogeny,
Fig. 3B & 3C). When discussing population structure, recA clusters will be used unless otherwise stated.
To determine how substantial the molecular evolutionary differences between clusters were, evolutionary distance and diversity calculations were done using MEGA 5.03 according to
Nei & Kumar [39]. Considering the placement of genotypes 5 and 6 in the second clade by 16S rDNA (Fig. 2), recA (Fig. 3A), and phzF (Fig. 3B) phylogenies, both genotypes were placed in clade 2 for intra- and interpopulation nucleotide sequence distance and diversity calculations.
The results of the divergence and diversity calculations (Table S2) can largely be visualized in the corresponding phylogenetic trees for recA and phzF (Fig. 3).
The presence of recombination events in recA and phzF was tested using split decomposition networks, pairwise homoplasy index (φw) [27], a coalescent simulation [32], and population admixture analysis and gene flow plots in BAPS [34]. Evidence for recombination events was essentially absent in recA according to all of the methods, as should be expected for a fairly well-conserved housekeeping gene in P. fluorescens (Table 3). Minimal signals for recombination in the network and coalescent simulation were detected for clusters 2 and 3, although the most robust test, the φw-test, did not detect statistically significant evidence for recombination (Table 3). However, significant recombination was detected for phzF within clusters 1, 3, and 4 (Fig. 4) by all three tests and the highest recombination rate (133) seen in
70 cluster 4. Intercluster gene flow was investigated using gene flow plots generated by BAPS [34].
No exchange of recA genetic information was found (data not shown). However, the plots showed the possibility for gene flow from clusters 2, 3, and 4 to cluster 1 in the phzF gene pool
(data not shown). The isolate (R5-89-07) involved in this one-way gene flow was identified using the population admixture partitions result. Cluster 3 showed the highest contributing proportion of phzF sequence (0.034) to R5-89-07, but its closest recA relative (cluster 2) showed the lowest phzF contribution (0.0057).
To detect the influences of selection on recA and phzF, the codon based Z-test for selection was used [26]. The null hypothesis of neutral selection for both recA and phzF was rejected (at a level of significance of 5%) in favor of purifying selection (dN < dS), but not positive selection (dN > dS) for all four clusters. In pairwise comparisons of isolates, intermittent neutral selection was detected (dN = dS) between strains within each of the clusters but not between clusters (data not shown).
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Table 3. Two tests conducted to detect recombination events in recA or phzF sequences within Phz+ clusters
recA (n = 37) b a Coalescent sim. φw-test R Rm Cluster 1 - 10.7 0 Cluster 2 -* 4.6 3 Cluster 3 -* 29.5 1 Cluster 4 - 14.7 0
phzF (n = 36c) Coalescent sim. φw-test R Rm Cluster 1 + (p = 0.0090)* 73.7 3 Cluster 2 - 14 0 Cluster 3 + (p = 0.027)* 32.1 5 Cluster 4 + (p = 0.032)* 133 4
*Networked structure (split decomposition analysis) detected in SplitsTree4 [27] aCalculated in SplitsTree4 bR = recombination rate per generation between most distant sites & Rm = minimum number of possible recombination events calculated using DnaSP v5 [32] cR5-89-07 not included in analysis due to its ambiguous phzF phylogeny and detection of population-wide admixture in BAPS [34]
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Table S2. Population-wide recA and phzF nucleotide sequence distancea and diversity calculationsb
recA (n = 37) Cluster 1 Cluster 2 Cluster 3 Cluster 4 Cluster 1 2.20 ± 0.8 7.00 ± 2.9 16.2 ± 7.2 6.50 ± 2.5
Cluster 2 5.30 ± 2.7 1.20 ± 0.4 10.2 ± 4.1 6.50 ± 2.7 Cluster 3 14.2 ± 6.7 8.70 ± 3.9 1.90 ± 0.6 8.50 ± 3.6 Cluster 4 5.10 ± 2.3 5.60 ± 2.5 7.20 ± 3.4 0.60 ± 0.3
πS = 1.5 ± 0.3 πT = 6.9 ± 2.2 δST= 5.4 ± 2.0 NST = 78.5 ± 5.0 phzF (n = 37) Cluster 1 Cluster 2 Cluster 3 Cluster 4 Cluster 1 c 6.7 ± 1.2 21.4 ± 4.8 15.3 ± 3.2 11.6 ± 2.3 Cluster 2 17.8 ± 4.6 0.4 ± 0.2 26.9 ± 6.9 20.5 ± 4.8 Cluster 3 10.6 ± 2.8 25.3 ± 7.0 2.80 ± 0.7 15.5 ± 3.4 Cluster 4 7.0 ± 1.8 19.1 ± 4.7 12.9 ± 3.3 2.5 ± 0.5
πS = 3.1 ± 0.4 πT = 14.7 ± 2.7 δST= 11.6 ± 2.3 NST = 78.9 ± 2.8 a Avg number of nucleotide (nt) substitutions per site within cluster (dX) are on the diagonal, avg number of nt substitutions per site between clusters (dXY) are above diagonal, and avg number of net nt substitutions per site between clusters (dA) are below diagonal. All values were multiplied by 100. bStandard errors for all values were calculated using the bootstrap method (1000 repetitions). cIncludes R5-89-07, as assigned by BAPS [34]
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74
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Biolog carbon substrate utilization profiling. We also explored the phenotypic diversity of the
Phz+ isolates by studying the differences in carbon substrate utilization profiles using Biolog
GN2 microplates. The principal components analysis applied to results of the metabolic profiling revealed that of the 95 carbon substrates present on the GN2 microplate, 62 accounted for at least
70% of the observed variance between the ODM of all 31 strains (Table S3). Some examples of key differences in substrate utilization were in the respiration of the natural sugar alcohol i- erythritol by all strains except for those in genotypes 1 to 4, the utilization of the deoxy sugar L- rhamnose by only genotypes 1 to 4, the utilization of succinamic acid restricted to all genotypes of cluster 2 and cluster 4 strains except genotype 12, and the utilization of the carboxylic acid itaconic acid restricted to the strains with close 16S rDNA relation to P. fluorescens 2-79
(clusters 1 and 3).
The results for the utilization of several substrates by the Phz+ isolates also indicate distinct differences from the described species. As reported by Dabboussi, et al. [40], P. orientalis does not utilize i-erythritol for its growth; however, the closely related (by 16S rDNA)
Phz+ cluster 2 isolates respire i-erythritol (Table S3). Additionally, members of the same cluster do not respire malonic acid, whereas P. orientalis does assimilate it for growth. The other very closely related group by 16S rDNA that contained most of the Phz+ cluster 1 isolates was significantly different from P. orientalis based upon Biolog data as compared to literature values
[40]. Based on Biolog substrate utilization profiles, cluster 3 consists of strains very closely related to P. libanensis, differing from this species only in utilization of malonic acid [41].
Additionally, P. gessardii, another close relative, grows on D-sorbitol, while P. libanensis and members of cluster 3 do not.
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The 62 substrates determined significant in variability by principal components analysis were then used for multivariate clustering to determine groups of Phz+ strains with similar carbon substrate utilization profiles. Results of the analysis revealed four clearly defined clusters of strains that closely matched the groupings observed in the BOX-PCR and sequence-based analyses (Fig. 5). The only difference was observed with strain L1-44-08 (genotype 7 of BOX-
PCR cluster 1), which had a metabolic profile similar to that of strains R2-4-08W and R2-54-
08W (genotypes 5 and 6, BOX-PCR cluster 1) and strains of cluster 3.
We also analyzed the utilization of carbon sources known to be present in wheat root exudates [42, 43]. There were no clear trends in the data like those seen in the overall dataset, however there did appear to be significantly stronger respiration of many of the common root exudate components by the strains from clusters 2 and 4 as compared to those from clusters 1 and 3 (Table S4). For instance, most of the cluster 2 and 4 strains had ODM values >1.5 times higher than the cluster 1 and 3 strains for the utilization of the amino acids L-histidine, L- glutamic acid, and L-asparagine. Overall, cluster 2 strains appeared to respire α-D-glucose (a common plant-derived simple sugar) to a lesser extent than did many of the other isolates.
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0.14 0.29 0.03 0.06 0.28 0.29 0.38 0.26 0.16 0.21 0.28 0.89 0.40 0.55 0.62 0.39 0.58 0.72 0.00 0.14 0.30 0.51 0.33 0.57 0.66 0.00 0.83 0.71 0.62 0.01 0.00 0.70 0.63 0.28 1.14 1.06 0.39 0.00 0.38 0.25 0.30 0.48 0.22 0.34 0.09 0.40 0.16 0.29 0.18 0.41 0.37 0.04 0.06 0.24 0.29 0.39 0.21 0.15 0.36 0.31 0.06 0.19
31 R1-75-08 0.33 0.23 0.07 0.10 0.32 0.28 0.33 0.50 0.11 0.25 0.38 0.87 0.29 0.42 0.67 0.42 0.59 0.58 0.07 0.13 0.35 0.34 0.47 0.65 0.60 0.00 0.75 0.61 0.63 0.10 0.00 0.85 0.48 0.47 0.83 1.12 0.33 0.00 0.53 0.30 0.27 0.46 0.24 0.35 0.09 0.26 0.27 0.36 0.22 0.34 0.47 0.11 0.19 0.07 0.13 0.22 0.24 0.09 0.25 0.30 0.04 0.06 30 R2-48-08W 0.23 0.39 0.00 0.15 0.42 0.23 0.30 0.40 0.17 0.33 0.33 1.01 0.47 0.53 0.79 0.49 0.49 0.67 0.01 0.10 0.35 0.39 0.53 0.84 0.69 0.01 1.10 0.76 0.88 0.06 0.04 1.03 0.75 0.44 1.10 1.11 0.96 0.03 0.25 0.26 0.31 0.43 0.23 0.34 0.15 0.31 0.25 0.30 0.26 0.33 0.65 0.21 0.14 0.15 0.29 0.31 0.15 0.11 0.31 0.27 0.15 0.20 29 L1-36-08 0.31 0.62 0.03 0.16 0.43 0.44 0.36 0.34 0.05 0.40 0.42 1.17 0.59 0.72 0.72 0.59 0.63 0.83 0.05 0.07 0.46 0.18 0.51 0.77 0.93 0.01 1.35 0.94 0.79 0.02 0.16 0.46 0.79 0.36 1.06 1.10 0.06 0.16 0.36 0.27 0.39 0.43 0.21 0.34 0.18 0.37 0.23 0.43 0.27 0.42 0.83 0.18 0.17 0.01 0.44 0.36 0.35 0.17 0.49 0.48 0.17 0.13 28 R3-52-08 0.53 0.24 0.04 0.08 0.37 0.29 0.52 0.56 0.08 0.35 0.43 1.12 0.53 0.66 0.86 0.58 0.65 0.91 0.04 0.05 0.38 0.12 0.57 0.90 0.59 0.00 0.97 0.54 0.82 0.00 0.00 0.63 0.77 0.46 0.95 1.17 0.92 0.00 0.56 0.42 0.35 0.48 0.25 0.42 0.29 0.28 0.23 0.36 0.21 0.53 0.73 0.08 0.18 0.13 0.22 0.24 0.22 0.13 0.31 0.47 0.08 0.04 27 R1-43-08 0.43 0.25 0.01 0.00 0.00 0.34 0.41 0.16 0.10 0.33 0.33 0.77 0.48 0.48 0.65 0.44 0.51 0.76 0.00 0.07 0.39 0.30 0.46 0.53 0.77 0.00 0.62 0.53 0.40 0.00 0.00 0.43 0.59 0.15 0.82 0.71 0.29 0.01 0.44 0.29 0.35 0.40 0.23 0.31 0.10 0.26 0.20 0.37 0.21 0.42 0.25 0.00 0.17 0.11 0.23 0.31 0.25 0.13 0.35 0.41 0.18 0.03 26 R2-37-08W 0.44 0.38 0.09 0.14 0.25 0.43 0.35 0.34 0.09 0.38 0.39 0.91 0.24 0.52 0.58 0.52 0.56 0.68 0.16 0.15 0.36 0.26 0.55 0.50 0.70 0.14 1.17 0.64 0.44 0.01 0.08 0.59 0.62 0.28 1.00 0.97 0.25 0.11 0.37 0.33 0.35 0.43 0.24 0.34 0.15 0.40 0.30 0.33 0.30 0.37 0.65 0.14 0.25 0.14 0.31 0.26 0.20 0.11 0.43 0.37 0.16 0.20 25 R4-35-07 0.27 0.18 0.02 0.16 0.29 0.30 0.40 0.41 0.05 0.26 0.32 1.02 0.44 0.00 0.74 0.43 0.58 0.64 0.00 0.10 0.34 0.07 0.69 0.76 0.64 0.00 1.10 0.67 0.64 0.10 0.00 0.49 0.70 0.28 0.98 1.13 0.49 0.08 0.33 0.26 0.29 0.40 0.16 0.26 0.11 0.24 0.24 0.28 0.19 0.41 0.67 0.07 0.17 0.15 0.19 0.19 0.23 0.12 0.30 0.34 0.14 0.18 24 R4-39-08 0.46 0.41 0.01 0.14 0.36 0.43 0.56 0.67 0.06 0.45 0.42 1.20 0.73 0.75 0.78 0.62 0.72 0.98 0.02 0.09 0.59 0.21 0.58 0.63 0.79 0.00 0.70 0.75 0.50 0.08 0.00 0.26 0.60 0.40 1.00 0.98 0.30 0.01 0.44 0.34 0.33 0.54 0.22 0.31 0.13 0.43 0.34 0.53 0.25 0.51 0.79 0.08 0.20 0.23 0.09 0.14 0.38 0.19 0.54 0.48 0.26 0.25 23 R2-60-08W 0.29 0.52 0.07 0.16 0.37 0.45 0.45 0.31 0.14 0.34 0.36 1.15 0.68 0.47 0.68 0.46 0.58 0.81 0.05 0.14 0.46 0.16 0.50 0.58 0.83 0.07 0.69 0.73 0.49 0.03 0.02 0.46 0.73 0.32 1.07 0.99 0.55 0.03 0.41 0.40 0.33 0.50 0.24 0.39 0.15 0.33 0.25 0.43 0.28 0.38 0.60 0.15 0.24 0.12 0.23 0.25 0.30 0.17 0.46 0.44 0.25 0.06 22 R3-18-08 0.07 0.10 0.05 0.16 0.22 0.15 0.40 0.85 0.17 0.33 0.32 1.03 0.46 0.47 0.61 0.34 0.57 0.58 0.00 0.16 0.39 0.36 0.95 0.92 0.61 0.02 1.26 0.86 0.68 0.00 0.65 0.62 0.63 0.17 0.99 1.10 0.15 0.18 0.29 0.16 0.08 0.32 0.11 0.22 0.05 0.20 0.09 0.25 0.04 0.23 0.39 0.00 0.09 0.05 0.13 0.13 0.17 0.11 0.33 0.30 0.14 0.00 21 R6-28-08 0.00 0.06 0.00 0.11 0.20 0.16 0.35 0.37 0.15 0.26 0.23 0.65 0.24 0.36 0.34 0.31 0.25 0.49 0.00 0.10 0.23 0.36 0.58 0.66 0.48 0.00 0.37 0.45 0.44 0.02 0.54 0.57 0.62 0.19 0.57 0.64 0.27 0.24 0.35 0.28 0.32 0.31 0.21 0.31 0.12 0.25 0.17 0.24 0.21 0.22 0.35 0.02 0.12 0.18 0.31 0.25 0.23 0.13 0.15 0.32 0.21 0.13 20 R4-30-08 0.04 0.08 0.01 0.15 0.11 0.06 0.22 0.75 0.05 0.14 0.24 0.79 0.30 0.26 0.31 0.18 0.41 0.37 0.02 0.06 0.21 0.21 0.61 0.63 0.41 0.02 0.85 0.41 0.35 0.02 0.45 0.41 0.52 0.15 0.41 1.01 0.09 0.10 0.27 0.18 0.22 0.20 0.13 0.18 0.05 0.12 0.07 0.17 0.09 0.28 0.23 0.01 0.00 0.00 0.03 0.06 0.10 0.09 0.22 0.17 0.00 0.00 19 R4-42-08 18 0.02 0.10 0.22 0.14 0.13 0.11 0.19 0.36 0.12 0.19 0.23 0.47 0.22 0.22 0.28 0.27 0.20 0.35 0.13 0.10 0.22 0.20 0.41 0.50 0.36 0.02 0.77 0.53 0.35 0.00 0.21 0.62 0.51 0.30 0.47 0.64 0.11 0.17 0.19 0.14 0.21 0.23 0.06 0.20 0.02 0.18 0.07 0.20 0.09 0.18 0.33 0.00 0.09 0.11 0.11 0.14 0.13 0.06 0.21 0.17 0.09 0.00 2-79 0.05 0.26 0.05 0.25 0.36 0.25 0.54 0.83 0.29 0.50 0.46 1.21 0.82 0.61 0.77 0.58 0.73 0.92 0.10 0.38 0.56 0.55 0.71 0.76 0.57 0.03 0.39 1.02 0.74 0.01 0.65 0.79 0.79 0.29 0.94 1.22 0.09 0.27 0.34 0.23 0.37 0.36 0.19 0.38 0.12 0.42 0.29 0.37 0.22 0.39 0.89 0.00 0.10 0.08 0.27 0.25 0.24 0.14 0.45 0.51 0.38 0.02 17 R4-36-08 0.78 0.51 0.05 0.14 0.32 0.35 0.67 0.29 0.11 0.45 0.52 1.28 0.81 0.79 0.81 0.64 0.78 0.92 0.03 0.08 0.58 0.39 0.44 0.66 0.64 0.00 0.99 0.78 0.59 0.00 0.80 0.54 0.78 0.42 0.99 0.99 0.36 0.40 0.43 0.42 0.36 0.54 0.22 0.46 0.16 0.40 0.22 0.45 0.15 0.43 0.75 0.13 0.24 0.09 0.37 0.27 0.31 0.14 0.38 0.51 0.18 0.06 16 M R5-89-07 OD 0.76 0.49 0.08 0.45 0.58 0.27 0.52 0.60 0.04 0.43 0.51 1.34 0.91 0.76 0.86 0.66 0.77 1.03 0.02 0.13 0.64 0.67 0.63 0.79 0.81 0.00 0.86 0.77 0.81 0.20 1.04 0.77 0.96 0.52 1.06 1.09 0.30 0.00 0.33 0.30 0.35 0.48 0.22 0.40 0.08 0.48 0.33 0.46 0.22 0.41 1.00 0.03 0.14 0.08 0.39 0.33 0.34 0.15 0.54 0.55 0.18 0.00 15 R4-7-08 0.58 0.51 0.45 0.21 0.42 0.32 0.71 0.36 0.03 0.47 0.45 0.89 0.47 0.69 0.78 0.70 0.64 0.89 0.00 0.34 0.49 0.39 0.40 0.46 0.65 0.00 0.61 0.40 0.48 0.18 0.54 0.53 0.86 0.46 0.65 0.56 0.27 0.52 0.51 0.41 0.38 0.55 0.22 0.34 0.09 0.31 0.27 0.57 0.21 0.56 0.75 0.00 0.20 0.04 0.29 0.32 0.43 0.19 0.36 0.57 0.22 0.21 14 Genotype Representative & Genotype Representative Genotype R2-45-08W 0.46 0.40 0.02 0.24 0.43 0.22 0.32 0.54 0.07 0.22 0.35 1.12 0.40 0.51 0.57 0.39 0.52 0.60 0.04 0.10 0.46 0.70 0.38 0.60 0.66 0.00 0.88 0.74 0.59 0.15 0.93 0.74 0.92 0.82 0.98 0.98 0.21 0.26 0.27 0.16 0.30 0.34 0.12 0.23 0.04 0.25 0.19 0.32 0.19 0.23 0.74 0.00 0.12 0.12 0.22 0.24 0.24 0.10 0.36 0.55 0.11 0.05 13 R2-57-08B 0.03 0.33 0.21 0.20 0.40 0.32 0.60 0.41 0.08 0.41 0.42 1.18 0.49 0.74 0.92 0.65 0.57 1.07 0.05 0.13 0.63 0.47 0.38 0.58 0.58 0.02 0.66 0.68 0.50 0.35 0.59 0.57 0.77 0.37 0.74 0.68 0.33 0.54 0.39 0.25 0.34 0.52 0.23 0.29 0.00 0.29 0.31 0.44 0.25 0.36 0.62 0.06 0.25 0.09 0.38 0.32 0.34 0.18 0.42 0.76 0.21 0.05 12 R2-6-08B Sixty-two Biolog substrates used for cluster analysis (as determined by principle componenets analysis) componenets principle by determined (as analysis cluster for used substrates Biolog Sixty-two 0.50 0.45 0.20 0.10 0.36 0.25 0.52 0.40 0.00 0.43 0.46 1.20 0.66 0.70 0.69 0.57 0.51 0.89 0.00 0.00 0.33 0.48 0.50 0.60 0.69 0.00 0.81 0.64 0.57 0.02 0.59 0.58 0.90 0.40 0.88 0.80 0.29 0.39 0.31 0.20 0.33 0.40 0.16 0.25 0.04 0.26 0.25 0.37 0.19 0.36 0.78 0.00 0.13 0.13 0.45 0.47 0.23 0.08 0.27 0.74 0.21 0.25 11 R4-65-08 Table S3. Table 0.12 0.70 0.20 0.31 0.29 0.35 0.67 0.58 0.13 0.42 0.51 1.37 0.54 0.77 0.85 0.56 0.47 1.01 0.00 0.31 0.53 0.62 0.66 0.75 0.62 0.00 0.85 0.73 0.78 0.03 0.82 0.62 1.01 0.65 0.78 0.85 0.26 0.59 0.44 0.31 0.30 0.43 0.22 0.29 0.11 0.33 0.20 0.52 0.22 0.52 0.79 0.00 0.17 0.08 0.44 0.41 0.34 0.19 0.39 0.84 0.22 0.15 10 R2-64-08W 0.46 0.41 0.19 0.27 0.45 0.27 0.66 0.45 0.05 0.39 0.46 1.12 0.58 0.60 0.67 0.49 0.66 0.82 0.01 0.18 0.53 0.42 0.45 0.52 0.51 0.04 0.57 0.62 0.51 0.17 0.67 0.59 0.70 0.60 0.77 0.79 0.36 0.26 0.33 0.27 0.31 0.57 0.24 0.34 0.11 0.33 0.30 0.35 0.25 0.45 0.79 0.02 0.22 0.13 0.29 0.27 0.28 0.16 0.44 0.62 0.20 0.03 9 R2-2-08B 0.58 0.57 0.25 0.30 0.39 0.37 0.66 0.40 0.03 0.40 0.42 0.88 0.18 0.73 0.79 0.62 0.51 0.99 0.00 0.20 0.39 0.33 0.31 0.50 0.54 0.00 0.67 0.59 0.51 0.15 0.57 0.48 0.78 0.48 0.69 0.69 0.21 0.00 0.34 0.22 0.27 0.46 0.17 0.29 0.06 0.29 0.20 0.45 0.22 0.34 0.65 0.07 0.17 0.04 0.23 0.29 0.28 0.14 0.41 0.62 0.20 0.04 8 L1-45-08 0.02 0.20 0.14 0.20 0.34 0.16 0.38 0.62 0.09 0.20 0.22 0.93 0.47 0.36 0.32 0.29 0.34 0.48 0.14 0.19 0.23 0.51 0.31 0.59 0.46 0.01 0.93 0.22 0.52 0.24 0.51 0.24 0.51 0.15 0.65 1.04 0.19 0.01 0.24 0.19 0.20 0.24 0.11 0.20 0.05 0.19 0.10 0.29 0.10 0.24 0.38 0.05 0.04 0.12 0.18 0.27 0.19 0.12 0.19 0.23 0.08 0.10 7 L1-44-08 0.03 0.12 0.13 0.18 0.28 0.16 0.32 0.54 0.13 0.22 0.26 0.90 0.45 0.37 0.37 0.30 0.34 0.52 0.15 0.11 0.29 0.41 0.23 0.56 0.47 0.02 0.93 0.41 0.50 0.25 0.49 0.30 0.49 0.17 0.70 1.00 0.19 0.13 0.28 0.24 0.24 0.29 0.14 0.22 0.09 0.20 0.11 0.23 0.17 0.24 0.45 0.00 0.09 0.09 0.26 0.28 0.15 0.09 0.25 0.23 0.10 0.04 6 R2-54-08W 0.11 0.14 0.15 0.20 0.22 0.15 0.39 0.71 0.19 0.25 0.32 1.00 0.42 0.37 0.45 0.33 0.51 0.50 0.11 0.23 0.27 0.25 0.62 0.60 0.44 0.00 0.87 0.48 0.45 0.00 0.54 0.39 0.47 0.35 0.53 1.08 0.17 0.17 0.22 0.20 0.21 0.23 0.14 0.20 0.09 0.21 0.15 0.21 0.15 0.25 0.36 0.02 0.10 0.00 0.09 0.10 0.13 0.10 0.27 0.20 0.09 0.02 5 R2-4-08W 0.09 0.41 0.08 0.21 0.32 0.18 0.15 0.41 0.04 0.27 0.28 0.71 0.73 0.35 0.41 0.33 0.48 0.45 0.07 0.11 0.29 0.21 0.00 0.90 0.50 0.60 1.10 0.41 0.80 0.04 0.78 0.00 0.65 0.00 0.88 0.96 0.38 0.01 0.38 0.30 0.24 0.35 0.20 0.25 0.13 0.26 0.25 0.25 0.16 0.34 0.98 0.06 0.20 0.07 0.24 0.23 0.22 0.14 0.28 0.22 0.14 0.03 4 R4-35-08 0.07 0.21 0.02 0.18 0.29 0.19 0.17 0.36 0.05 0.25 0.22 0.54 0.57 0.33 0.50 0.38 0.38 0.48 0.05 0.15 0.33 0.53 0.00 0.71 0.58 0.75 0.99 0.66 0.56 0.02 0.53 0.00 0.64 0.00 0.83 0.97 0.35 0.00 0.37 0.25 0.22 0.33 0.13 0.19 0.07 0.24 0.10 0.25 0.17 0.28 0.88 0.03 0.14 0.08 0.23 0.26 0.17 0.14 0.38 0.23 0.06 0.07 3 R2-66-08W 0.05 0.27 0.10 0.19 0.23 0.21 0.01 0.23 0.08 0.30 0.35 0.95 0.47 0.48 0.56 0.39 0.43 0.58 0.06 0.22 0.41 0.14 0.02 0.78 0.59 0.67 0.88 0.68 0.61 0.51 0.61 0.00 0.83 0.01 0.78 0.93 0.44 0.29 0.46 0.36 0.29 0.47 0.24 0.24 0.12 0.26 0.23 0.31 0.23 0.57 0.62 0.05 0.22 0.15 0.26 0.34 0.30 0.14 0.34 0.27 0.17 0.04 2 L1-12-08 0.02 0.18 0.03 0.17 0.16 0.20 0.23 0.26 0.09 0.27 0.29 1.05 0.49 0.53 0.58 0.36 0.57 0.62 0.03 0.11 0.70 0.17 0.02 0.83 0.53 0.65 1.04 0.77 0.78 0.64 0.94 0.00 0.73 0.00 0.77 0.89 0.33 0.22 0.32 0.22 0.26 0.40 0.14 0.20 0.08 0.32 0.15 0.33 0.16 0.36 0.80 0.19 0.14 0.13 0.26 0.22 0.18 0.13 0.36 0.37 0.17 0.02 1 L1-3-08 Substrate Succinamic Acid Succinamic Glucuronamide L-Alaninamide Putrescine 2-Aminoethanol L-Histidine Hydroxy-L-Proline L-Leucine L-Ornithine D-Alanine L-Alanine L-Proline L-Alanyl-Glycine Acid L-Pyroglutamic L-Asparagine L-AsparticAcid L-Serine Acid L-Glutamic L-Threonine Acid Glycyl-L-Glutamic γ-AminobutyricAcid N-Acetyl-D-Glucosamine i-Erythritol D-Fructose D-Galactose L-Rhamnose α-D-Glucose D-Sorbitol m-Inositol Sucrose D-Trehalose Adonitol L-Arabinose Xylitol D-Arabitol D-Mannitol D-Mannose p-HydroxyphenylaceticAcid Acid Citric α-KetoglutaricAcid Acid Lactone D-Galactonic Acid D-Galacturonic Acid D,L-Lactic Acid D-Gluconic MalonicAcid Acid D-Gluconsaminic PropionicAcid Acid D-Glucuronic Acid Quinic Acid D-Saccharic β-HydroxybutyricAcid γ-HydroxybutyricAcid AcidSuccinic Dextrin Tween40 Tween80 Inosine Uridine Glycerol UrocanicAcid PyruvicAcid Methyl Ester AcidSuccinic Mono-Methyl Ester
78
0.02 0.22 0.38 0.06 0.62 0.72 0.40 0.00 0.21 0.28 0.89 0.26 0.00 0.00 0.58 0.29 0.83 31 R1-75-08 0.02 0.24 0.53 0.19 0.67 0.58 0.29 0.07 0.25 0.38 0.87 0.50 0.03 0.02 0.59 0.28 0.75 30 R2-48-08W 0.04 0.23 0.25 0.14 0.79 0.67 0.47 0.01 0.33 0.33 1.01 0.40 0.00 0.01 0.49 0.23 1.10 29 L1-36-08 0.06 0.21 0.36 0.17 0.72 0.83 0.59 0.05 0.40 0.42 1.17 0.34 0.02 0.06 0.63 0.44 1.35 28 R3-52-08 0.02 0.25 0.56 0.18 0.86 0.91 0.53 0.04 0.35 0.43 1.12 0.56 0.04 0.03 0.65 0.29 0.97 27 R1-43-08 0.05 0.23 0.44 0.17 0.65 0.76 0.48 0.00 0.33 0.33 0.77 0.16 0.00 0.00 0.51 0.34 0.62 26 R2-37-08W 0.13 0.24 0.37 0.25 0.58 0.68 0.24 0.16 0.38 0.39 0.91 0.34 0.01 0.08 0.56 0.43 1.17 25 R4-35-07 0.03 0.16 0.33 0.17 0.74 0.64 0.44 0.00 0.26 0.32 1.02 0.41 0.01 0.02 0.58 0.30 1.10 24 R4-39-08 0.06 0.22 0.44 0.20 0.78 0.98 0.73 0.02 0.45 0.42 1.20 0.67 0.02 0.02 0.72 0.43 0.70 23 R2-60-08W 0.07 0.24 0.41 0.24 0.68 0.81 0.68 0.05 0.34 0.36 1.15 0.31 0.02 0.06 0.58 0.45 0.69 22 R3-18-08 0.00 0.11 0.29 0.09 0.61 0.58 0.46 0.00 0.33 0.32 1.03 0.85 0.00 0.00 0.57 0.15 1.26 21 R6-28-08 0.03 0.21 0.35 0.12 0.34 0.49 0.24 0.00 0.26 0.23 0.65 0.37 0.00 0.00 0.25 0.16 0.37 20 R4-30-08 0.00 0.13 0.27 0.00 0.31 0.37 0.30 0.02 0.14 0.24 0.79 0.75 0.00 0.00 0.41 0.06 0.85 19 R4-42-08 18 0.00 0.06 0.19 0.09 0.28 0.35 0.22 0.13 0.19 0.23 0.47 0.36 0.00 0.00 0.20 0.11 0.77 2-79 0.03 0.19 0.34 0.10 0.77 0.92 0.82 0.10 0.50 0.46 1.21 0.83 0.00 0.03 0.73 0.25 0.39 17 R4-36-08 0.05 0.22 0.43 0.24 0.81 0.92 0.81 0.03 0.45 0.52 1.28 0.29 0.02 0.00 0.78 0.35 0.99 16 M R5-89-07 OD 0.01 0.22 0.33 0.14 0.86 1.03 0.91 0.02 0.43 0.51 1.34 0.60 0.00 0.00 0.77 0.27 0.86 15 R4-7-08 0.02 0.22 0.51 0.20 0.78 0.89 0.47 0.00 0.47 0.45 0.89 0.36 0.00 0.00 0.64 0.32 0.61 14 Genotype Representative & Genotype Representative Genotype R2-45-08W 0.00 0.12 0.27 0.12 0.57 0.60 0.40 0.04 0.22 0.35 1.12 0.54 0.00 0.00 0.52 0.22 0.88 13 R2-57-08B 0.05 0.23 0.39 0.25 0.92 1.07 0.49 0.05 0.41 0.42 1.18 0.41 0.02 0.01 0.57 0.32 0.66 12 R2-6-08B 0.00 0.16 0.31 0.13 0.69 0.89 0.66 0.00 0.43 0.46 1.20 0.40 0.00 0.00 0.51 0.25 0.81 11 Utilization data for seventeen common wheat rhizosphere exudates extracted from the Biolog GN2 microplate dataset. microplate GN2 Biolog the from extracted exudates rhizosphere wheat common seventeen for data Utilization R4-65-08 0.02 0.22 0.44 0.17 0.85 1.01 0.54 0.00 0.42 0.51 1.37 0.58 0.00 0.01 0.47 0.35 0.85 10 Table S4. Table R2-64-08W 0.02 0.24 0.33 0.22 0.67 0.82 0.58 0.01 0.39 0.46 1.12 0.45 0.00 0.00 0.66 0.27 0.57 9 R2-2-08B 0.00 0.17 0.34 0.17 0.79 0.99 0.18 0.00 0.40 0.42 0.88 0.40 0.00 0.00 0.51 0.37 0.67 8 L1-45-08 0.01 0.11 0.24 0.04 0.32 0.48 0.47 0.14 0.20 0.22 0.93 0.62 0.00 0.00 0.34 0.16 0.93 7 L1-44-08 0.01 0.14 0.28 0.09 0.37 0.52 0.45 0.15 0.22 0.26 0.90 0.54 0.02 0.03 0.34 0.16 0.93 6 R2-54-08W 0.01 0.14 0.22 0.10 0.45 0.50 0.42 0.11 0.25 0.32 1.00 0.71 0.01 0.04 0.51 0.15 0.87 5 R2-4-08W 0.02 0.20 0.38 0.20 0.41 0.45 0.73 0.07 0.27 0.28 0.71 0.41 0.02 0.02 0.48 0.18 1.10 4 R4-35-08 0.02 0.13 0.37 0.14 0.50 0.48 0.57 0.05 0.25 0.22 0.54 0.36 0.02 0.02 0.38 0.19 0.99 3 R2-66-08W 0.01 0.24 0.46 0.22 0.56 0.58 0.47 0.06 0.30 0.35 0.95 0.23 0.02 0.01 0.43 0.21 0.88 2 L1-12-08 0.01 0.14 0.32 0.14 0.58 0.62 0.49 0.03 0.27 0.29 1.05 0.26 0.05 0.00 0.57 0.20 1.04 1 L1-3-08 Substrate AceticAcid Acid D,L-Lactic Acid Citric Acid Succinic L-Asparagine Acid L-Glutamic L-Alanyl-Glycine L-Threonine D-Alanine L-Alanine L-Proline L-Leucine L-Phenylalanine D-Serine L-Serine L-Histidine α-D-Glucose
79
80
Production of phenazines and screening for other antibiotics. As a part of phenotypic profiling we examined culture conditions that favor in vitro production of PCA in strains representing unique genotypes of Phz+ isolates that were defined in the study. Of 31 tested strains, 19 produced significant quantities of PCA when grown on LB supplemented with 2% glucose, KMB, or both media (Table S5). No other phenazine derivatives were detected in studied strains by using TLC or HPLC analyses. Unexpectedly, twelve phzF-carrying isolates failed to produce PCA under the aforementioned culture conditions, as well as during growth on tryptic soy broth (TSB). We succeeded in detecting small amounts of PCA in all twelve genotypes only after >8 days of growth on potato dextrose agar (PDA).
The 31 unique Phz+ genotypes were screened for the production of biosurfactants by the drop collapse assay. Biosurfactants were detected in 16 of the 31 tested isolates after growth on
LB amended with 2% glucose, KMB, or both media (Table S5). We also screened all Phz+ isolates for the presence of biosynthesis pathways for other antibiotics known to contribute to the ability of Pseudomonas spp. to control soilborne plant pathogens. However, probing by PCR with primers targeting pathways for production of 2,4-diacetylphloroglucinol, pyrrolnitrin and pyoluteorin revealed the absence of corresponding target genes in genomes of all Phz+ strains used in this study.
81
Table S5. Production of phenazine-1-carboxylic acid and biosurfactants by 31 Phz+ genotypes
Biosurfactant PCA production production Genotype and representative isolate LB + LB + 2% KMB PDA KMB 2% glc glc 1 Pseudomonas sp. L1-3-08 - + n.d.a +/- +/- 2 Pseudomonas sp. L1-12-08 - - + + +/- 3 Pseudomonas sp. R2-66-08W - + n.d. + - 4 Pseudomonas sp. R4-35-08 + + n.d. + - 5 Pseudomonas sp. R2-4-08W - + n.d. - - 6 Pseudomonas sp. R2-54-08W + + n.d. - - 7 Pseudomonas sp. L1-44-08 - - + + +/- 8 Pseudomonas sp. L1-45-08 - - + + +/- 9 Pseudomonas sp. R2-2-08B - - + + + 10 Pseudomonas sp. R2-64-08W - - + + +/- 11 Pseudomonas sp. R4-65-08 - - + + +/- 12 Pseudomonas sp. R2-6-08B - - + + +/- 13 Pseudomonas sp. R2-57-08B - - + + +/- 14 Pseudomonas sp. R2-45-08W - - + + +/- 15 Pseudomonas sp. R4-7-08 - - + + +/- 16 Pseudomonas sp. R5-89-07 - + n.d. + +/- 17 Pseudomonas sp. R4-36-08 - - + - - 18 P. fluorescens 2-79 + + n.d. + - 19 Pseudomonas sp. R4-42-08 + - n.d. - - 20 Pseudomonas sp. R4-30-08 - - + - - 21 Pseudomonas sp. R6-28-08 - + n.d. - - 22 Pseudomonas sp. R3-18-08 + + n.d. - - 23 Pseudomonas sp. R2-60-08W + + n.d. - - 24 Pseudomonas sp. R4-39-08 + + n.d. - - 25 Pseudomonas sp. R4-35-07 + + n.d. +/- +/- 26 Pseudomonas sp. R2-37-08W - + n.d. +/- +/- 27 Pseudomonas sp. R1-43-08 - + n.d. - - 28 Pseudomonas sp. R3-52-08 + + n.d. - - 29 Pseudomonas sp. L1-36-08 - + n.d. - - 30 Pseudomonas sp. R2-48-08W - + n.d. - - 31 Pseudomonas sp. R1-75-08 + + n.d. - - a Not determined
82
Discussion
Very little research has been done to characterize natural populations of plant-associated phenazine-producing bacteria in comparison to the significant emphasis that has been placed on understanding the mechanism, efficacy, and applicability of selected Phz+ strains as biological control agents in agriculture. Our discovery of Phz+ Pseudomonas spp. in wheat and barley rotation plots near Ritzville, WA and in a wheat field near Lind, WA in 2007 [3] provided the foundation for this study. The close proximity of the sites sampled for Phz+ isolates and the relative homogeneity of site characteristics provided a unique opportunity to study the diversity, population genetics and evolution of these Phz+ populations. These highly enriched Phz+
Pseudomonas rhizosphere populations are the first example of such a phenomenon in the United
States [13].
We found significant diversity at the genotype level within the sampled Phz+ population of 31 BOX-PCR genotypes in four clusters that was generally supported by sequence and phenotypic differences (with a few important exceptions). The BOX-PCR genotyping of the entire Phz+ collection revealed cluster 4 as a dominant and ubiquitous cluster of genotypes. The three other Phz+ clusters comprised less than half of the total number of isolates and appeared less widely distributed than members of cluster 4. We found no evidence for specific genotype selection based on the sampled site but there were significant differences in BOX-PCR genotype diversity and evenness between crop rotations as indicated by five common diversity indices.
Genotype diversity in the samples from the Jirava farm, as indicated by all five diversity indices
(Table 2), was highest in continuous spring wheat (R4) and lowest in commercial winter wheat
(R1). R4 also showed the greatest genotype evenness, as indicated by its J value (0.841). This
83 suggests that continuous spring wheat production may select for a wide range of Phz+ genotypes in relatively equal abundances as compared to commercial winter wheat production that may have been in rotation with another dryland crop. A similar phenomenon has been shown in a take-all suppressive soil continuously cultivated to wheat which contained a greater diversity of
2,4-diacetylphloroglucinol (Phl+) producing Pseudomonas spp. than the conducive soil in rotation [24].
The winter wheat/summer fallow fields at Lind, WA (L1) and Ritzville, WA (R3) showed comparable DMg values (2.03 & 1.85, respectively); suggesting that annual fallow positively influences the diversity of Phz+ populations as compared to R1 but not to the extent of continuous wheat production. We had previously found an inverse correlation between precipitation and Phz+ population size [13], however it appears that relative soil moisture also
+ affects Phz diversity on wheat as shown by the low DMg value of the irrigated site (R6, 0.91).
We are currently further investigating the link between low soil moisture and the Phz+ populations at Lind, WA.
The Phz+ populations described in this study are significantly different from the dominant population of Phz+ strains isolated from the Châteaurenard wilt suppressive soils. The majority of the Châteaurenard Phz+ isolates were genetically related to Phz+ P. chlororaphis strains 30-84,
PGS12, and PCL1391, which are relatively distantly related to the P. fluorescens subgroup [7].
Only three out of twenty-nine of the Châteaurenard Phz+ isolates showed almost identical phzC
RFLP patterns, however different BOX-PCR profiling, from that of P. fluorescens 2-79 [7]. The genetic difference in Phz+ populations between this study and the Châteaurenard study can likely be ascribed to biotic and edaphic factors, as each have been shown to play key roles in shaping rhizosphere microbial communities [24, 44, 45]. Consequently, the soil pH and CaCO3 of Shano
84 silt loam at the Lind, WA site was much lower (pH 5.63-6.27, >5% CaCO3) than the wilt suppressive Châteaurenard soil (pH 7.9, 37.4% CaCO3) and closer to the wilt conducive
Carquefou soil (pH 5.3, 8.4% CaCO3) [7]. However, the Ritzville silt loam has a pH of 8.29 and is much closer in pH to the Châteaurenard soil, but its % CaCO3 of >5% is more similar to Shano silt loam [7]. It is possible that pedology aside, Châteaurenard suppressive soil cropped to wheat or barley (rather than tomato or flax) would result in selection of the P. fluorescens 2-79 related phzC RFLP-genotype VII population. Plant host, in cooperation with soil type, has been suggested to play a key role in selecting and shaping specific indigenous antibiotic-producing rhizobacterial populations [24, 44]. Therefore, it is likely that a combination of cropping history and abiotic factors have shaped the two geographically and genotypically distinct Phz+ populations of Châteaurenard and Ritzville and Lind, WA.
The Châteaurenard Phz+ populations suppress Fusarium wilt of tomato and flax in cooperation with non-pathogenic Fusarium oxysporum. However, currently, the biological control effectiveness of the Phz+ populations discovered by Mavrodi et al. [3, 13], and described in this study, is largely unknown. Three isolates from BOX-PCR cluster 4 have been tested in the greenhouse and show comparable, and in some cases better, biological control of Rhizoctonia solani AG8 than P. fluorescens 2-79 [13]. Additionally, one study has shown that at the Jirava farm near Ritzville, WA a reduction in Rhizoctonia bare patch of wheat has been occurred over an eight year period when in rotation with barley [14]. In our study, an annual spring wheat/spring barley rotation appeared to inversely affect Phz+ population diversity (Table 2), decreasing in the barley-cropped year (R2B) and increasing in the wheat-cropped year (R2W).
This rotation effect may play a role in selecting for BOX-PCR cluster 4 genotypes, as 51 of 66
85 isolates from R2B are in cluster 4. Thus the reduction in Rhizoctonia bare patch with a barley rotation may be attributable to an enrichment of BOX-PCR cluster 4 strains.
The variability in PCA production between genotypes was an unexpected result observed in this study. Although 19 of 31 genotype representatives readily produced PCA in vitro, 12 of the genotypes that represent 147 out of 412 total Phz+ isolates did not initially produce PCA under the conditions that we provided for growth. Traditionally, KMB and LB + 2% glucose represent optimal media for PCA production; however when grown on PDA the ‘non-PCA producing’ isolates began to show production after several days. A possible explanation for this slow production could lie in the complex plant-derived nutrient composition of PDA as compared to that of the other media tested (KMB, LB + 2% glucose, or TSB). Additionally, environmental stimuli or specific rhizospheric conditions may promote PCA synthesis that we are unable to reproduce under laboratory conditions. We also tested for the production of biosurfactants, which are membrane-solubizing compounds like cyclic lipopeptides that have been shown to act synergistically with phenazines to control Pythium spp. and Rhizoctonia root rot of bean [11, 46]. The same isolates reluctant to initially produce PCA in vitro appeared to readily produce biosurfactants as measured by the drop collapse assay (Table S5). The variability in PCA and biosurfactant production suggests complex conditional biological control traits that will require further study to understand their importance to biological control efficacy and ecology.
To date there has been no thorough examination of the molecular ecology within a large indigenous population of phenazine-producing rhizobacteria. The isolates examined in this study formed two distinct 16S rDNA clades, indicating that the two clades represent at least two separate species (Fig. 2). The carbon substrate utilization profiles of several isolates also seem to
86 separate each clade from its closest 16S rDNA relative and, in a similar manner to sequence data, separate clade 1 into three distinct clusters (BOX-PCR clusters 1, 2, and 4) with a carbon substrate utilization clustering similarity below 50% (Fig. 5). This suggests that the populations described by the four unique clusters could potentially represent four different Pseudomonas species. To delve deeper into the differences between the clusters, we studied the population genetics and molecular evolution of the Phz+ isolates that were collected. The four distinctly cohesive, yet highly diverse recA-based clusters of Phz+ isolates showed evolutionary histories that differed in the degree of divergence as well as diversity in both recA and phzF loci (Fig. 3 and Table S2). The diversity of phzF and PhzF (Fig. 3C) among the four clusters and the detection of purifying selection on that locus suggest that stable adaptations have occurred to allow colonization of particular niches within the rhizosphere. The importance of PCA production for competitive success in the rhizosphere has been shown previously [47]; thus the detection of purifying selective pressures on phzF is particularly relevant. The diverse results from Biolog substrate utilization (Tables S3 and S4) also hint at specific phenotypic traits that may provide a distinct advantage in the rhizosphere for each cluster.
To our knowledge, the significant evidence for recombination within phzF of a single natural population of Phz+ fluorescent Pseudomonas species has not been reported before. We were able to show that recombination in phzF has likely occurred over evolutionary time within recA-based clusters 1, 3 and 4 (Table 3). In addition, the split decomposition networks for clusters 1, 3 and 4 (Fig. 4) showed a variety of pathways for genetic exchange. This may suggest different mechanisms for the evolution of phzF. This exchange of genetic material, possibly through natural transformation [48], could play a role in how phzF and other phz genes evolve to fill a niche or rapidly respond to environmental conditions that may deleteriously affect
87 phenazine production. As more Phz+ isolates from this dryland agroecosystem are analyzed, it will be interesting to investigate if horizontal gene transfer has occurred between clusters.
It is possible that a certain amount of sampling bias has affected our interpretation of the real Phz+ populations in the field; however, cloning and sequencing of phzF directly from rhizosphere total DNA seems to support our culture-based findings in the present study [3].
Using the phzF sequence data from this study, we will be able to develop real-time PCR primers to assess relative cluster-specific population sizes from total rhizosphere DNA. Additionally, future high-throughput sequencing efforts of phzF from rhizosphere DNA can be put into the context of the findings from this study.
In this study, we described the population structure, genetics and diversity of a new group of PCA-producing fluorescent Pseudomonas species isolated from dryland wheat and barley rhizospheres in central Washington State (U.S.). The influence of our Phz+ isolates on soilborne pathogens of cereals in the dryland fields of the entire Inland Pacific Northwest is currently unknown, but preliminary results from a recent survey show widespread high Phz+ rhizosphere populations and biologically significant in situ rhizosphere PCA concentrations in the dryland region [13]. This widespread phenomenon shows that the diverse, yet distinct, Phz+ clusters of genetically similar isolates described here likely impact a large region of dryland agricultural fields and may play a role in the suppression of dryland soilborne pathogens like Rhizoctonia.
88
Acknowledgments
The project described was supported by Award Number T32GM083864 from the National
Institute of General Medical Sciences. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of General Medical
Sciences or the National Institutes of Health.
89
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Chapter 2
Taxonomy and distribution of phenazine-1-carboxylic acid-producing
Pseudomonas spp. in the dryland agroecosystem of the Inland Pacific
Northwest (U.S.)
James A. Parejko1, Dmitri V. Mavrodi2, Olga V. Mavrodi2, David M. Weller3,
Linda S. Thomashow3*
1School of Molecular Biosciences, Washington State University, Pullman, WA 99164-4234
2Department of Plant Pathology, Washington State University, Pullman, WA99164-6430
3U.S. Department of Agriculture, Agricultural Research Service, Root Disease and Biological
Control Research Unit, Pullman, WA 99164-6430
*For correspondence. Mail: USDA-ARS Root Disease and Biological Control Research Unit,
Washington State University, Pullman, WA 99163-6430. E-mail: [email protected]; Tel.:
(+1) 509-335-0930; Fax: (+1) 509-335-7674
97
Abstract
The majority of commercially grown cereals in the Inland Pacific Northwest (IPNW) of the
United States are colonized by high indigenous populations (105-107 colony forming units per gram of root) of phenazine-producing (Phz+) fluorescent Pseudomonas spp. The diversity of
Phz+ pseudomonads was studied previously in three IPNW wheat fields, but the regional distribution and taxonomy of these microorganisms remained unclear. The primary aim of this study was to investigate the taxonomic placement of Phz+ strains from dryland fields of the
IPNW by using multi-locus sequence analysis (MLSA) of four housekeeping genes (16S rRNA, gyrB, rpoB and rpoD). The secondary aim of the study was to explore abiotic factors that may affect the distribution of the four species throughout the IPNW. Results of the MLSA analyses revealed the presence of four large and distinct groups of Phz+ pseudomonads. Two of these were represented by new species that were provisionally named ‘Pseudomonas aridus’ and ‘P. cerealis’. The other two groups were comprised of the first described Phz+ strains of P. orientalis and P. synxantha. Analysis of the abiotic factors revealed that the genetic diversity within Phz+ populations correlated with the type of agroclimatic zone (coefficient t-test P-value = 0.0188,
ANOVA F-test P-value = 0.02167) and the soil silt content (coefficient t-test P-value = 0.0220,
ANOVA F-test P-value = 0.02334). Collectively, our findings demonstrate the presence of new species of Phz+ Pseudomonas in the dryland agroecosystems of the IPNW, and further clarify the impact of agronomic practices and edaphic factors on the abundance and diversity of these beneficial microorganisms.
98
Introduction
For over 135 years, non-irrigated dryland wheat (Triticum aestivum) has been the dominant crop cultivated in the Inland Pacific Northwest (IPNW) of the United States (37), which includes the areas of east and central Washington, northern Idaho and northcentral Oregon. This area begins deep in the rain shadow of the Cascade Mountains and is divided into the low, intermediate and high precipitation zones which receive <300, 300 to 450 and >450 mm of annual precipitation, respectively (37). In the eastern part of the IPNW, the ‘Palouse region’ is defined by rolling loess-covered hills and precipitation ranges from 450 to 600 mm, allowing farmers to plant rotation crops that include spring wheat, spring barley (Hordeum vulgare), brassicas (Brassica spp.) and legumes like pea, lentil (Lens culinaris) and chickpea (Cicer arietinum). Throughout the IPNW, the soil texture is characterized as silt loam and originates from postglacial sediments and ancient volcanic eruptions (37). The top soil of large expanses of the region is quite homogeneous, with minor variability in certain soil characteristics. Consequently, climatic effects like annual precipitation and growing-degree days have an overwhelming impact on agricultural viability.
Wheat plants exude approximately 11% of net fixed carbon into the region directly surrounding the roots (the ‘rhizosphere’) in the form of low or high molecular weight compounds like organic acids, sugars, and proteins (rhizodeposition) (16). Sloughed cells that are deposited by the actively growing apical meristem are also found in the rhizosphere (16). As a direct result of this active and passive photosynthate exudation, the plant provides a carbon and energy source for root-colonizing bacteria (‘rhizobacteria’) and exerts a selective force on the soil-dwelling microbial community (4). The composition of the resident microbial community is also strongly influenced by the soil type in which the plant is growing (4). One ubiquitous soil-dwelling 99 rhizobacterium is Pseudomonas fluorescens, classically described as a single species but more recently recognized as a species-complex by comparative genome and multi-locus sequence analysis (MLSA) (20, 30, 39). Many P. fluorescens-like rhizobacterial strains produce a multitude of metabolites that promote plant growth either directly (e.g. phytohormones and volatiles) or indirectly by suppressing soilborne pathogens through metabolites that act as antibiotics (23, 41). Among the antibiotics produced by certain rhizobacteria are the structurally and functionally diverse heterocyclic nitrogen-containing compounds called phenazines (25).
The simplest of the phenazine derivatives is phenazine-1-carboxylic acid (PCA). It has been extensively studied in Pseudomonas fluorescens 2-79, a model phenazine-producing (Phz+) strain that originally was isolated from a twelve-year continuous wheat plot near Lind, WA
(U.S.) in 1979 (45). The core enzymes involved in the biosynthesis of PCA in P. fluorescens 2-
79 are encoded by the seven gene phz operon (phzABCDEFG) (22, 25), and homologues of these genes are found in other Phz+ bacteria (25). Phenazines play multiple physiological roles for the cell including increasing the thickness and density of cells in biofilms (21), reducing Fe and Mn- oxides while acting as an electron shuttle (13, 34, 43) and functioning as a molecular signal (9).
Some phenazines also play a role in the plant induced systemic response, likely through the production of oxygen radicals (2). Finally, phenazines possess well-documented biocontrol activity, and PCA has been implicated as a key metabolite responsible for the capacity of P. fluorescens 2-79 to inhibit Gaeumannomyces graminis var. tritici (Ggt), the causal agent of take- all disease of wheat (41).
Several studies have provided evidence that indigenous Phz+ pseudomonads are abundant in certain agricultural soils. A 2009 study by Mazurier et al. (27) of the Châteaurenard Fusarium wilt-suppressive soil in France was the first to implicate indigenous populations of Phz+
100
Pseudomonas spp. in plant disease suppression. Some of these Phz+ strains were genetically related to P. fluorescens 2-79, but functioned synergistically with non-pathogenic Fusarium oxysporum to suppress pathogenic F. oxysporum in the Châteaurenard soil. The Phz+ pseudomonads and the non-pathogenic Fusarium were not present in the neighboring non- suppressive Carquefou soil. The Châteaurenard soil had higher pH and CaCO3 content and subsequent lower extractable iron content than the Carquefou soil, indicating that edaphic factors may influence Phz+ communities in the rhizosphere (27). More recently, Mavrodi et al. (25) developed primers targeting the key phenazine biosynthesis gene phzF and demonstrated the presence of high population sizes of Phz+ Pseudomonas spp. in the rhizosphere of dryland wheat grown in the IPNW (U.S.) near Ritzville and Lind, WA. Representatives of these Phz+ populations inhibited the prevalent dryland cereal root rot disease causal agent Rhizoctonia solani AG-8 in vitro and protect plant roots in greenhouse assays against root rot ((24), unpublished data). Parejko et al. (33) further characterized the Phz+ Pseudomonas spp. populations from this area and identified 31 BOX-PCR genotypes and four recA- and phzF- based groups with distinct phenotypic traits suggesting that new Pseudomonas species might be present. Subsequent studies revealed that indigenous Phz+ rhizobacteria were present at levels of
105-107 CFU g-1 root throughout commercial cereal fields of the IPNW region (24) and that Phz+ rhizobacteria also colonize the roots of native vegetation growing in uncultivated soils (26). The amount of PCA in the rhizosphere of field-grown wheat positively correlated with Phz+
Pseudomonas spp. population size (24). Interestingly, the frequency of plant colonization was inversely related to precipitation, suggesting that soil moisture directly influences the colonization competency or competitiveness of Phz+ Pseudomonas spp. in the rhizosphere (24,
26). However, it remained unclear as to what extent the composition of the Phz+ rhizobacterial
101 community varied between precipitation zones and if Phz+ rhizobacteria were affected by abiotic factors other than soil moisture.
The current study was designed to address the taxonomic placement and regional distribution of potentially novel Phz+ Pseudomonas spp. in the IPNW. Using partial gene (16S rRNA, gyrB, rpoB and rpoD) multi-locus sequence analysis (MLSA) (30), we found that two of the four previously described Phz+ strain clusters had less than 97% multi-locus sequence similarity with the closest described species which is a pre-requisite for identification of new
Pseudomonas species. These two clusters have been provisionally separated into new species named ‘P. aridus’ and ‘P. cerealis’. The other two Phz+ groups clustered with P. orientalis and
P. synxantha. Members of these groups are the first Phz+ strains to be described for either of these Pseudomonas species. We also found a highly divergent strain that had less than 95.5% sequence similarity to any previously described species of pseudomonads. By analyzing the distributions and frequencies of phzF alleles amplified from total rhizosphere DNA at different
IPNW sites, we found that agroclimate zone and soil percent silt had a significant effect on Phz+ community diversity. To our knowledge, this is the first study to examine extrinsic abiotic factors influencing the composition of indigenous rhizosphere Phz+ Pseudomonas spp. on a regional scale. The results of this study provide new insights into the prevalence of P. fluorescens-related phenazine-producing species in the environment and will serve as a foundation for more comprehensive studies of novel phenazine-producing Pseudomonas species in global agroecosystems.
102
Materials & methods
Sample collection, processing, Phz+ strain isolation and rhizosphere DNA extraction.
Winter and spring wheat plants from GPS-tagged locations throughout Washington State and northern Oregon, U.S. (Figure 1A) were sampled randomly as described in a previous study (24).
Characteristics of the locations are listed in Table S1. Phz+ strains were isolated from rhizosphere suspensions according to Parejko et al. (33). Rhizosphere DNA samples were extracted from winter wheat plants randomly sampled from fields listed in Table 1. From each site, four replicate samples of plants between stages 8 and 10 on the Feeke’s scale of development were collected in a manner similar to Mavrodi et al (24). Briefly, four replicate samples from each site were placed in separate plastic bags, transported back to the laboratory and stored at 4°C for no more than 24 hrs before processing. The roots of four randomly selected plants from each replicate bag were excised and placed in a 50-ml Falcon tube. Each tube received 20 ml of sterile dH2O, was placed on ice, and then was vortexed and sonicated for 1 min. Each suspension was diluted ten-fold in sterile dH2O and 100 µl was transferred to 200 µl sterile dH2O in a 96-well microtiter plate and serially diluted for the terminal endpoint-dilution assay (25, 28). Fifty microliters of each dilution were used to inoculate two microtiter plates filled, respectively, with 200 µl of 1/10 strength tryptic soy broth (TSB) amended with cycloheximide (100 µg ml-1) or with 1/3 strength King’s medium B (KMB) (18) containing ampicillin (40 µg ml-1), cycloheximide (100 µg ml-1) and chloramphenicol (13 µg ml-1). The 1/10
TSB was used to culture total aerobic heterotrophic soil bacteria, whereas 1/3 KMB amended with antibiotics was used to selectively grow fluorescent pseudomonads (28). The inoculated plates were incubated for 72 hours in the dark at room temperature, after which optical density at
600 nm (OD600) was measured for each well. All wells with an OD600 ≥ 0.1 were considered
103 positive for bacterial growth. The readings from 1/10 TSB plates were used to calculate populations levels of total culturable bacteria. The cultures in 1/3 KMB plates were screened by
PCR using the primer set Ps_up1/Ps_low1 and terminal dilution wells that tested positive for phzF were used to calculate the population size of Phz+ Pseudomonas spp. (25).
104
105
Table S1. Characteristics of 2009a and 2010 sites sampled for Phz+ strains
Phz+ Pseudomonas
spp. Mean No. Phz+ Date Site ID GPS coordinates Plant sampled Pop. density precip. yr-1 strains sampled (logCFU g-1 (mm)b isolated C.f. root f.w. ± SD) 46°15′59′′N, S16X09 03/17/09 Winter wheat 6.2 ± 2.1 0.3 478 14 118°11′15′′W
46°17′48′′N, S17X09 03/17/09 Winter wheat 5.6 ± 1.2 0.3 352 24 118°26′23′′W
46°9′49′′N, S21X09 03/24/09 Winter wheat 6.4 ± 0.9 1.0 235 38 119°42′23′′W
46°7′58′′N, S22X09 03/24/09 Winter wheat 6.5 ± 1.7 0.8 230 33 119°42′21′′W
46°7′37′′N, S23X09 03/24/09 Winter wheat 6.1 ± 0.8 0.8 246 9 119°44′20′′W
46°7′38′′N, S24X09 03/24/09 Winter wheat 5.6 ± 1.3 0.9 246 27 119°51′21′′W
46°1′36′′N, S25X09 03/24/09 Winter wheat 7.0 ± 1.3 1.0 348 41 120°15′53′′W
46°6′8′′N, S26X09 03/24/09 Winter wheat 5.5 ± 0.8 0.8 226 11 119°36′20′′W
46°9′27′′N, S27X09 03/24/09 Winter wheat 6.9 ± 0.9 0.9 232 22 119°36′7′′W
46°48′39′′N, S31X09 04/21/09 Winter wheat 5.5 ± 1.8 0.8 448 12 117°37′12′′W
46°51′15′′N, S33X09 04/21/09 Winter wheat 5.2 ± 1.3 0.9 309 18 118°20′10′′W
47°0′4′′N, S34X09 04/21/09 Winter wheat 5.7 ± 1.6 1.0 290 16 118°20′48′′W
47°7′26′′N, S35X09 04/21/09 Winter wheat 5.7 ± 2.0 0.8 275 4 118°36′4′′W
46°54′45′′N, 261 S36X09 04/21/09 Winter wheat 6.2 ± 1.3 1.0 42 118°33′58′′W
106
46°47′41′′N, S37X09 04/21/09 Winter wheat 6.0 ± 1.5 0.9 259 4 118°33′28′′W
46°47′46′′N, S38X09 04/21/09 Winter wheat 6.4 ± 0.9 0.9 243 2 118°40′19′′W
46°47′35′′N, S39X09 04/21/09 Winter wheat 5.9 ± 1.4 0.8 237 19 118°43′48′′W
47°1′49′′N, S41X09 05/04/09 Winter wheat 6.0 ± 1.3 0.4 520 30 117°21′47′′W
47°25′12′′N, S42X09 05/04/09 Winter wheat 4.5 ± 1.2 0.6 475 1 117°21′52′′W
47°39′28′′N, S44X09 05/04/09 Winter wheat 4.0 ± 0.9 0.6 441 34 117°56′54′′W
47°43′43′′N, S46X09 05/04/09 Winter wheat 6.9 ± 1.1 0.8 356 5 118°43′23′′W
47°10′39′′N, S49X09 05/04/09 Winter wheat 4.1 ± 0.6 0.3 336 11 117°52′12′′W
45°51′0′′N, S63X09 06/09/09 Winter wheat 5.2 ± 1.1 0.9 470 6 118°43′7′′W
45°50′31′′N, S64X09 06/09/09 Winter wheat 5.1 ± 0.9 0.9 405 2 118°47′47′′W
45°47′29′′N, S65X09 06/09/09 Winter wheat 5.6 ± 0.8 0.9 374 17 118°48′28′′W
46°37′40′′N, S611X09 06/09/09 Winter wheat 4.1 ± 0.6 0.8 250 4 118°43′54′′W
Blue wildrye 47°8′28′′N, 1X09 06/15/09 (Elymus 5.8 ± 1.1 1.0 295 2 118°29′40′′W glaucus Buckley) Wild grass 47°8′28′′N, 3X09 06/15/09 (Agropyron 4.9 ± 1.4 0.8 295 2 118°29′40′′W cristatum) Cheatgrass
47°8′28′′N, (Bromus 4X09 06/15/09 4.2 ± 1.1 1.0 295 3 118°29′40′′W tectorum L.)
Needle & 46°47′54′′N, 41X10c 06/15/10 thread grass 3.7 ± 0.9 0.6 264 10 118°29′32′′W (Stipa comata L.)
107
46°47′54′′N, 42X10c 06/15/10 Spring wheat 5.6 ± 1.0 1.0 264 18 118°29′32′′W
Common 46°47′54′′N, 43X10c 06/15/10 yarrow 4.9 ± 1.0 0.8 264 16 118°29′32′′W (Achillea millefolium L.) Total 497 strains a Site characteristics are extracted from Mavrodi, et al (2011) and Mavrodi, et al (2012). b Approximate mean annual precipitation (1971-2000) from PRISM Climate Group data mapped using ArcGIS v.9.3 (Esri, Inc.) c 2010 sites were sampled as a side-by-side comparison of strains colonizing spring wheat and two native plants bordering the field in undisturbed soil
108
Table 1. Characteristics of sites sampled for total rhizosphere DNA
Phz+ population Mean Total Geographic density Phz+ plant annual Sample phzF Agroclimate zoneb coordinates (log CFU g-1 colonizationa precip. clones root f.w. ± SD ) (mm) c
46°47′49′′N, S1.1 51 5.4 ± 0.6 1.0 Grain/Fallow 259 118°32′57′′W 46°46′53′′N, S1.2 54 5.6 ± 1.0 0.7 Grain/Fallow 297 118°21′28′′W 46°46′50′′N, S1.3 52 5.3 ± 0.8 0.8 Ann. Crop: Dry 371 117°56′58′′W 46°48′40′′N, S1.4 55 6.3 ± 0.6 0.6 Grain/Fallow 430 117°40′22′′W 47°16′33′′N, S2.1 52 5.4 ± 1.3 0.6 Grain/Fallow 273 118°41′34′′W 47°4′20′′N, S2.2 52 5.7 ± 0.9 0.6 Grain/Fallow 265 118°38′7′′W 46°56′26′′N, S2.3 46 5.3 ± 0.8 0.8 Grain/Fallow 258 118°34′26′′W 46°50′9′′N, Ann. Crop: Fallow- S2.5 44 4.8 ± 0.5 0.2 512 117°22′18′′W Trans.
46°45′20′′N, Ann. Crop: Wet- S2.8 48 5.0 ± 0.6 0.3 581 117°6′41′′W Cool a Sixteen plants were randomly sampled from each field b Agroclimatic zones defined by Douglas et al. (10) c A 30 yr (1971-2000) mean annual precipitation data from the PRISM Climate Group
109
Cloning and sequencing of phzF from total rhizosphere DNA. Four random rhizosphere suspensions from each of samples S1.1 to S2.8 (Fig. 1B, Table 1) with high populations of Phz+
Pseudomonas spp. were chosen for total rhizosphere DNA extraction. The extractions were carried with a PowerSoil DNA isolation kit (MoBio, Carlsbad, CA) using the manufacturer’s protocol for wet soil samples. The rhizosphere suspensions from four plants per site were pooled and concentrated by centrifugation. Total pooled suspension volume was reduced to one-quarter of the total volume to concentrate the pooled suspensions. Two milliliters of each suspension per site were used for total rhizosphere DNA isolation. Amplification of phzF was performed using
Ps_up1/Ps_low1 primers essentially as described elsewhere (25), except for the addition of 10%
(final volume) of molecular-grade dimethylsulfoxide to PCR reactions. The reactions were separated by electrophoresis in 1% agarose extracted by using a Zymoclean Gel DNA Recovery kit (Zymo Research Corp., Irvine, CA). The recovered phzF amplicons were then ligated into the pGEM-T Easy vector (Promega, Madison, WI) and into E. coli DH5α Subcloning Efficiency competent cells (Life Technologies, Grand Island, NY) according to the manufacturer’s recommendations. Individual transformants were re-plated and screened by PCR with phzF- specific primers. The resultant amplicons were purified by using MinElute 96 UF plates (Qiagen,
Valencia, CA) and sequenced with Ps-up1 and Ps_low1 primers by Elim Biopharm (Hayward,
CA).
DNA fingerprinting by BOX-PCR. To reduce the number of isolates for multilocus sequence analysis (MLSA), all phzF-positive isolates were subjected to profiling by BOX-PCR (42). A procedure that was previously used to genotype Phz+ Pseudomonas spp. was used without modification (33). Isolates of the same banding profile from each location were grouped and one representative isolate was selected for MLSA.
110
Amplification and sequencing of phzF and MLSA loci from bacterial isolates. To isolate total genomic DNA, representative Phz+ strains were grown overnight at 28 ºC in Luria-Bertani
(LB) broth and used for DNA extractions with the GenElute Bacterial Genomic DNA kit
(Sigma-Aldrich Co., St. Louis, MO). The purified DNA was used to amplify partial 16S rRNA, gyrB and rpoB with primer sets 8F/1492R (44), Up-1G-/ Up-2G-(25) and rpoBup1/rpoBlow1
(25), respectively. Two different primer sets, LAPS/LAPS27 (1) and PsEG30F and PsEG790R
(29), were used to amplify rpoD and amplification of phzF was performed using primers
Ps_up1/Ps_low1 (25). The amplifications were performed with a PTC-200 thermal cycler (Bio-
Rad, Hercules, CA) using GoTaq DNA polymerase (Promega, Inc., Madison, WI) as described elsewhere (25, 29, 30). PCR products were cleaned with QIAquick PCR purification spin columns (Qiagen, Valencia, CA), DNA concentrations were measured using a DNA quantitation kit (Bio-Rad, Hercules, CA) and the amplicons were sequenced by Elim Biopharm (Hayward,
CA).
Sequence analysis, phylogenetic inference and evolutionary estimation. All DNA sequences were initially trimmed of primers and preliminarily analyzed for quality using CLC bio Main
Workbench v.6.6.2 (CLC bio, Aarhus, Denmark). Sequences were aligned in MEGA5 (40) using
MUSCLE (11). The MLSA gamma-parameter was estimated using the Jukes-Cantor substitution model and phylogenetic distances were estimated using the Jukes-Cantor substitution model with the estimated gamma-parameter of 0.1. The significance of the phylogenetic distances was tested by using boot-strapping with 1000 repetitions. Pairwise proportions of dissimilar nucleotide sites
(p-distance) were also calculated in MEGA with both transitional and transversional differences and are represented as ‘DNA sequence similarity values’ using 1-(p-distance).
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Simultaneous Alignment and Tree Estimation v. 2.2.2 (SATé-II) software (19) was used for MLSA phylogenetic tree construction using the ‘Multi-Locus Data’ settings best suited for the MLSA data suggested by the software. The settings were the following: the initial alignment was used for the initial tree, the maximum subproblem size was 51, the aligner used was
MAFFT, merger was MUSCLE, maximum-likelihood (ML) tree estimation and Shimodaira-
Hasegawa test (SH)-like local support values (1000 repetitions) calculated using FASTTREE and the model was GTR+G20. All other settings were default. The estimated phylogenetic tree was visualized in MEGA5 (40) and only local support scores over 50 were retained.
The 389 nt-long phzF sequences from Phz+ Pseudomonas strains featured in this (n = 76) and previous studies (n = 37) (25, 33) as well as those amplified from rhizosphere DNA (n =
454) were aligned using the ‘Very Accurate’ proprietary progressive alignment algorithm in
CLC bio Main Workbench v.6.6.2 (CLC bio, Aarhus, Denmark). The resulting alignment was used for phylogenetic tree estimation using default settings in SATé-II (19). phzF-based taxonomy classification was performed using ‘classify.seqs’ in Mothur v.1.26.0 (38) with the
Bayesian method and a taxonomy database constructed from the reference phzF alleles (n =
113). The SATé-II-derived phzF phylogenetic tree was visualized in Dendroscope v.3.2.2 (15).
An alignment without reference phzF alleles was used to calculate the inverse Simpson diversity
(1/D) estimate (14) in Mothur v.1.26.0 (38). 1/D indicates the relative diversity in a sample with
1.0 being a sample containing only a single OTU. Mothur also was used to produce rarefaction curves for the eight sites analyzed for Pseudomonas spp. phzF. All calculations in Mothur used a phylogenetic distance cut-off of 0.04 nt substitutions site-1.
Environmental variables and statistics. A matrix of various environmental variables (pH, % organic matter, % clay, % sand, % silt, CaCO3) obtained from the USDA Soil Data Mart
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(http://soildatamart.nrcs.usda.gov/) for Adams, Whitman and Lincoln counties in Washington
State (U.S.) was constructed. Additionally, mean annual precipitation (years 1971-2000) was obtained from the PRISM Climate Group (http://www.prism.oregonstate.edu/) and agroclimatic zones were used as defined by Douglas et al. (10). All data were visualized geographically in
ArcGIS v.9.3 (Esri, Inc) and sampled sites were identified and located using GPS coordinates.
Appropriate data were extracted from each data layer for each sampled site. Exploratory analyses were run in BiodiversityR (17) using the values from inverse Simpson’s diversity index and the environmental variables matrix. A two-sample F-test (α = 0.05) to determine equal or unequal variances was performed on needle and thread grass, spring wheat and common yarrow pairwise log-transformed Phz+ Pseudomonas spp. population data in Excel 2010 (Microsoft, Corp.). Two- tail t-tests (α = 0.05) were then performed on the same data in Excel 2010 (Microsoft, Corp.).
Nucleotide sequence accession numbers. The nucleotide sequences used in this study were deposited in NCBI GenBank.
Results
Taxonomy of Phz+ Pseudomonas spp. isolates. The descriptive sequence analyses for the four loci used for MLSA are listed in Table 2. As expected, the partial 16S rRNA sequence (1385 nt) was the least polymorphic (34 polymorphic sites) and had the lowest nucleotide diversity (π =
0.005). Partial gyrB (794 nt) was the most polymorphic (196 polymorphic sites), had the highest nucleotide diversity (π = 0.057) and the lowest ratio of non-synonymous to synonymous nt substitutions site-1 (dN/dS, 0.011), which identifies the presence or absence of selective forces on a nt sequence (31). Partial rpoB (903 nt) was the least polymorphic of the three protein-encoding
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MLSA loci (145 polymorphic sites) and also had the lowest nucleotide diversity of the protein- encoding sequences (π = 0.027). Partial rpoD (697 nt) had 161 polymorphic sites and π = 0.048 with the highest dN/dS (0.099). Partial phzF (389 nt) had the highest %G+C of the DNA sequences analyzed at 64.1%G+C and significantly higher nucleotide diversity (π = 0.082) with fewer polymorphic sites than the three MLSA protein-encoding genes (120 polymorphic sites).
All dN/dS values indicated that gyrB, rpoB, rpoD and phzF are under varying levels of purifying selection. Purifying selection is represented by values of dN/dS < 1.0 which indicate that most amino acid substitutions are in some way deleterious to the cell and are removed by natural selection (31).
All BOX-PCR site-representative Phz+ Pseudomonas spp. isolates (n = 76) and twelve representatives of major BOX-PCR genotype clusters defined previously (33) were analyzed for taxonomic placement in the P. fluorescens lineage using the four partial gene MLSA scheme
(16S rRNA, gyrB, rpoB and rpoD) proposed by Mulet, et al (30). The clade provisionally named
“Pseudomonas aridus” was established using MLSA data from eighteen Phz+ strains isolated from wheat rhizospheres in 2009 and 2010 and the strain R1-43-08 that was a 2008 isolate, which represents a large group of strains in BOX-PCR genotypes 21 through 31 (33). This clade conntained the largest number of Phz+ strains isolated throughout the IPNW as defined by BOX-
PCR (n = 177). The mean multi-locus sequence similarity within the P. aridus group was 0.9%.
The closest described species to the group were P. grimontii, P. marginalis, P. orientalis, P. panacis, P. synxantha and P. veronii at between 4.1% and 4.5% sequence dissimilarity (Table 3).
The clade provisionally named “Pseudomonas cerealis” was established with MLSA data from thirty strains collected between 2009 and 2010, and strains L1-45-08 and R4-65-08 isolated in 2008 represented BOX-PCR genotypes 8 through 15 (33) (Fig. 2). The P. cerealis clade
114 encompassed a total of 156 isolates that have been collected throughout the IPNW. The clade included a defined sub-group of strains that shared ca. 2% multi-locus sequence dissimilarity with the rest of P. cerealis isolates. The sub-group included strain R5-89-07 an unusual strain identified in previous studies and characterized by the BOX-PCR genotype 16 (25, 33).
Pseudomonas cerealis was most closely related to strains of P. orientalis (ca. 3.7% dissimilarity) and P. aridus. It was also the most cohesive group at 0.7% dissimilarity and was highly clonal in evolutionary relatedness among the main cluster of strains (Fig. 2). The MLSA-derived phylogeny also suggested that the P. cerealis and P. aridus groups shared a common ancestor with P. orientalis (Fig. 2).
The P. orientalis clade encompassed seventeen newly isolated Phz+ Pseudomonas strains, the type strain of the species, P. orientalis CFML-170T, and strains L1-3-08 and R2-66-08W from the 2008 study (33) that represent BOX-PCR genotypes 1 through 4. The group in its entirety included 140 Phz+ strains characterized by MLSA and/or BOX-PCR analyses and was smaller than the P. cerealis group. The P. orientalis clade was also the most divergent internally at 1.3% sequence dissimilarity, and contained a sub-group with two strains, L1-44-08 (BOX-
PCR genotype 7) and S21609. The sub-group could potentially represent a distinct subspecies because it was 2.8% dissimilar from the type strain CFML-170T (Table 3).
The P. synxantha clade contained the fewest number of Phz+ strains isolated from the
IPNW soils (n = 22) in addition to the type strain of the species LMG 2335T (Fig. 2). The Phz+ P. synxantha strains characterized by MLSA included six newly isolated and four previously defined strains (R2-4-08W, R2-54-08W, R4-36-08 and R6-28-08), as well as P. fluorescens 2-79 that represents BOX-PCR genotypes 17 through 21 (33). The clade had an internal sequence dissimilarity of 0.8%, and strains 2-79 and 1109 were ca. 1.4% divergent from the rest of the
115 clade members. Our results also revealed that P. synxantha was so closely related to P. libanensis that the two may actually represent sub-species of a single species or a species complex. At the same time, the P. synxantha clade was distinctly divergent from any other species (Fig. 2) with the closest being P. cedrina (Table 3).
Pseudomonas sp. 1209 was a single outlier and was not related to any group of Phz+ strains analyzed in this study. The strain was isolated from the rhizosphere of blue wild rye
(Elymus glaucus) growing in uncultivated soil near Ritzville, WA (26) and had a distinct morphology (produces copious amounts of mucoid exopolysaccharide). Pseudomonas sp. 1209 was most closely related to P. merdiana and P. proteolytica at ca. 4.8% and 4.9% dissimilarities, respectively (Table 3).
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Table 2. Population-wide (n = 88) descriptive analyses of phzF and housekeeping genes from Phz+ Pseudomonas spp.
Sequence Nucleotide # of # of # of non- length # of diversity Locus %G+C polymorphic synonymous synonymous dN/dS analyzed alleles per site sites sites sites (nta) (π)
16S rRNA 1385 53.6 30 34 NA NA NA 0.005
gyrB 794 56.0 54 196 186.07 605.93 0.011 0.057
rpoB 903 59.6 49 145 226.66 676.34 0.022 0.027
rpoD 697 61.0 48 161 167.49 525.51 0.099 0.048
phzF 389 64.1 53 120 97.35 289.65 0.049 0.082 a Excluding sites with gaps or missing data NA: not applicable
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Table 3. The MLSA-based sequence dissimilarity comparison of Phz+ strain groups to species of the P. fluorescens lineage
P. aridus P. cerealis P. orientalis P. synxantha Pseudomonas Species group group group group sp. 1209
P. brenneri 5.3 ± 0.3a 5.4 ± 0.3 5.8 ± 0.4 5.0 ± 0.3 5.2 ± 0.3 P. cedrina 5.2 ± 0.3 5.0 ± 0.3 4.6 ± 0.3 4.5 ± 0.3 6.3 ± 0.4 P. gessardi 5.5 ± 0.3 5.4 ± 0.3 5.7 ± 0.4 5.0 ± 0.3 5.0 ± 0.3 P. grimontii 4.1 ± 0.3 4.2 ± 0.3 4.5 ± 0.3 4.6 ± 0.3 5.6 ± 0.4 P. libanensis 4.8 ± 0.3 4.9 ± 0.3 5.0 ± 0.3 1.5 ± 0.2 6.2 ± 0.4 P. marginalis 4.3 ± 0.3 4.3 ± 0.3 4.6 ± 0.3 4.7 ± 0.3 5.7 ± 0.4 P. meridiana 4.8 ± 0.3 4.8 ± 0.3 5.2 ± 0.3 5.3 ± 0.3 4.8 ± 0.3 P. mucidolens 6.2 ± 0.4 6.0 ± 0.4 6.8 ± 0.4 5.8 ± 0.4 5.5 ± 0.3 P. orientalis 4.1 ± 0.3 3.7 ± 0.3 2.4 ± 0.2 4.8 ± 0.3 6.6 ± 0.3 P. panacis 4.5 ± 0.3 4.5 ± 0.3 4.5 ± 0.3 4.8 ± 0.3 5.6 ± 0.4 P. proteolytica 5.0 ± 0.3 5.0 ± 0.3 5.4 ± 0.3 4.9 ± 0.3 4.9 ± 0.3 P. synxantha 4.5 ± 0.3 4.5 ± 0.3 4.6 ± 0.3 0.6 ± 0.1 6.2 ± 0.4 P. veronii 4.5 ± 0.3 4.5 ± 0.3 4.5 ± 0.3 4.8 ± 0.4 5.2 ± 0.3 a The sequence dissimilarity values (% ± SE) were calculated as p-distance in MEGA5 using 1000 bootstrap repetitions.
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Comparison of Phz+ Pseudomonas spp. from spring wheat and uncultivated vegetation. To identify the affinity of Phz+ Pseudomonas spp. towards different species of dryland plants, we isolated Phz+ strains from a field with spring wheat (n = 18) and from an adjacent undisturbed area covered with needle and thread grasses (n = 10) and common yarrow (n = 16) (see site
‘NP10’ in Fig. 1A). Needle and thread grasses supported 3.7±0.9 log Phz+ CFU g-1 root (fresh weight) of Phz+ rhizobacteria with plant colonization frequency (c.f.) = 0.6, whereas in common yarrow the Phz+ population was at 4.9±1.0 log Phz+ CFU g-1 root f.w. and c.f. = 0.8. The population of Phz+ rhizobacteria on spring wheat was estimated at 5.6±1.0 log Phz+ CFU g-1 root f.w. and c.f. = 1.0, and was not significantly different from common yarrow (P = 0.14).
However, Phz+ populations colonizing spring wheat were significantly greater than needle and thread grasses (P = 0.00045), and population levels on needle and thread grass were significantly different from those on common yarrow (P = 0.031).
The bulk of Phz+ isolates recovered from spring wheat belonged to P. aridus (16 of 18 strains) and were genetically close to strain 42110 (Fig. 2, Table S2). Common yarrow Phz+ isolates were dominated by P. orientalis (12 of 16 strains) (Table S2) and were close to strain
43110 (Fig. 2). All Phz+ isolates (n = 10) from needle and thread grass were identified as P. cerealis and were represented by strains 41210, 41310 and 41810 (Fig. 2). The four different plant hosts were also colonized by very closely related (ca. 99.7% multi-locus similarity) Phz+ strains of P. cerealis (Fig. 2). BOX-PCR genotyping indicated that these Phz+ strains likely belong to the same genotype (data not shown).
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Table S2. Total number of Phz+ Pseudomonas spp. strains isolated from sampled sites
No. strains belonging to Phz+ species isolated No. of unique BOX- Site or host P. P. P. P. PCR genotypes detected Total aridus cerealis orientalis synxantha
S16X09 1 0 0 14 0 14 S17X09 2 0 24 0 0 24 S21X09 3 0 37 1 0 38 S22X09 2 23 0 10 0 33 S23X09 3 0 0 6 3 9 S24X09 5 1 10 16 0 27 S25X09 4 18 9 0 14 41 S26X09 3 0 4 7 0 11 S27X09 2 14 8 0 0 22 S31X09 1 0 0 12 0 12 S33X09 2 17 1 0 0 18 S34X09 1 16 0 0 0 16 S35X09 3 3 1 0 0 4 S36X09 4 38 4 0 0 42 S37X09 2 2 0 2 0 4 S38X09 1 0 0 0 2 2 S39X09 5 16 3 0 0 19 S41X09 2 0 3 27 0 30 S42X09 1 0 0 1 0 1 S44X09 4 0 4 30 0 34 S46X09 1 5 0 0 0 5 S49X09 3 8 2 1 0 11 S63X09 2 0 6 0 0 6 S64X09 1 0 0 0 2 2 S65X09 2 0 17 0 0 17 S611X09 1 0 4 0 0 4 1X09 2 0 0 0 1 2a 3X09 2 0 0 2 0 2 4X09 3 0 3 0 0 3 41X10 4 0 10 0 0 10 42X10 2 16 2 0 0 18 43X10 2 0 4 12 0 16 a Pseudomonas sp. strain 1209 was also isolated from this site.
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Effect of climatic and edaphic factors on the distribution of Phz+ Pseudomonas spp. of the
IPNW. Amplicons containing a 427-nt fragment of phzF were PCR-amplified from total rhizosphere DNA isolated from winter wheat plants. The amplicons were cloned to produce a set of small libraries of 44 to 55 clones per sampled site that were sequenced and the phzF alleles were assigned to Pseudomonas species defined by MLSA (Table 1, Fig. 3). The majority of alleles were classified as one of the four dominant Phz+ species (i.e., P. orientalis, P. synxantha,
P. aridus or P. cerealis) (Fig. 3). The largest percentage of phzF alleles (45%) were classified as belonging to P. cerealis while 20% and 22% belonged to P. aridus and P. orientalis, respectively. A significant portion (20%) of P. cerealis-like alleles originating from sample S1.4 were assigned to the sub-group exemplified by R5-89-07 (Fig. 3). A slightly smaller percentage of phzF alleles from S1.2 and S2.5 were also assigned to the sub-group of P. orientalis (19%).
Finally, almost 10% of the phzF alleles (45 of 454) were classified as Pseudomonas sp. 1209- like and 44 of these originated from sample S2.8. No phzF alleles were assigned to P. synxantha, and a small group of sequences from samples S1.3 and S2.8 did not match any of the known
Phz+ species (Fig. 3).
The results of phzF-based rarefaction analysis revealed that the Phz+ communities were nearly completely or completely sampled at seven of the eight sites (Fig. S1). The only under- sampled population was located at site S2.5. The rarefaction curve for sample S2.8 showed the greatest OTU richness at four phzF-based OTUs, while plant roots sampled at sites S1.1 and S2.1 were colonized by Phz+ bacteria sharing a single phzF allele. We also estimated the diversity of phzF alleles by calculating the inverse Simpson’s index (1/D). The index indicates the relative diversity in a sample with 1.0 being a sample containing phzF alleles belonging to a single species or taxonomic unit. Although the Phz+ community from sample S2.5 was under-sampled,
122 it had the highest 1/D diversity at 2.47. Sites S2.8 and S1.4 had the next the next highest 1/D values at 2.01 and 1.58, respectively, whereas sites S1.1 and S2.1 had the lowest diversity of phzF alleles with 1/D at 1.00.
We also attempted to establish correlation between the 1/D values for sampled Phz+ communities and environmental variables (i.e., mean annual precipitation, agroclimatic zone, pH, CaCO3, cation exchange capacity (CEC), soil percent sand, soil percent silt, soil percent clay and soil percent organic matter) at the sampled locations. ANOVA testing of linear, Poisson and quasi-Poisson models showed that the residuals were not normally distributed and the quasi-
Poisson model provided the best fit for the data. Mean annual precipitation showed only a slight effect on 1/D diversity (coefficient t-test P-value = 0.056, ANOVA F-test P-value = 0.05929), however soil percent silt (coefficient t-test P-value = 0.0220, ANOVA F-test P-value = 0.02334) and agroclimatic zone (coefficient t-test P-value = 0.0188, ANOVA F-test P-value = 0.02167) showed a much greater effect (Fig. 4). Environmental interpretation using canonical correspondence analysis of soil percent silt and agroclimatic zone and phzF communities did not show a specific effect of either variable on any of the phzF-based groups.
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Discussion
The primary aims of this study were to provide evidence for new Phz+ species within the populations of indigenous Phz+ Pseudomonas spp. of the Inland Pacific Northwest (U.S.) and attempt to define environmental variables influencing the diversity and distribution of these microorganisms in the rhizosphere of dryland wheat. In the low precipitation zone of the IPNW, phenazine-producing Pseudomonas spp. colonized the roots of dryland cereal crops and native and invasive plants growing in uncultivated soils (26). They also protected dryland wheat from soilborne fungal pathogens G. graminis var. tritici and Rhizoctonia solani AG-8 (44, unpublished data). Our previous analysis of Phz+ Pseudomonas spp. from the IPNW using BOX-
PCR and recA and phzF sequence-based analyses revealed the presence of four distinct groups of strains (33). In this study we used the MLSA scheme developed for the identification of
Pseudomonas species by Mulet et al. (30) to show that two of these groups represent novel species of the P. fluorescens complex provisionally named ‘Pseudomonas aridus’ and
‘Pseudomonas cerealis’ (Fig. 2). The DNA sequence similarities of these groups were below
97%, which is the MLSA-defined cut-off for Pseudomonas species (30). In addition to genetic differences, representative members of P. aridus and P. cerealis differed in the dynamics of PCA production in vitro (33). P. aridus strains readily produced PCA on KMB and Luris-Bertani broth supplemented with 2% glucose whereas P. cerealis strains slowly accumulated PCA on potato dextrose agar only after 48 hrs of growth. In vitro production of biosurfactants, amphipathic molecules that disrupt cellular membranes and synergistically interact with phenazines to inhibit the growth of some fungal phytopathogens (6, 8, 35), also differed between the two provisional species (33).
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We also tested these two species using Biolog GN2 carbon substrate utilization microtiter plates (33) and identified three substrates differentially utilized by P. aridus and P. cerealis. L- alaninamide, D-trehalose and p-hydroxyphenylacetic acid were all utilized by the majority of P. cerealis strains while no strains of P. aridus grew on these carbon substrates. The closest described species, P. orientalis, utilizes D-trehalose but does not utilize p-hydroxyphenylacetic acid (7). Additionally, both P. aridus and P. cerealis utilize erythritol whereas P. orientalis does not. Finally, P. orientalis utilizes malonate and variably utilizes L-rhamnose whereas both new
Phz+ species do not utilize either substrate for growth or respiration (7, 33). These phenotypic differences strengthen our hypothesis that P. aridus and P. cerealis are novel Pseudomonas species.
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Table S3. Distinct phenotypic differences between the four IPNW Phz+ Pseudomonas species identified in this study
PCA production Substrate utilization
Hydroxyphenylacetic Hydroxyphenylacetic
Succinamic acid Succinamic
L
L
D
-
L
Alaninamide
-
-
- a Bio- Rhamnose
Trehalose Species ornithin surfactant acid LB+2% b p KMB PDA -
glucose
e
P. aridus - + + n.d. - - - - + v.
P. cerealis + - - + + - + + + -
Phz+ P. + + - n.d. - + + v. - - orientalis
Phz+ P. -c v. d v. n.d. v. - + + - + synxantha a No. of strains tested for each phenotype: P. aridus = 10 strains, P. cerealis = 9 strains, P. orientalis = 5 strains and P. synxantha = 7 strains (Parejko et al. 2012). b n.d., not determined. c P. fluorescens 2-79 produces a viscosin-like biosurfactant (unpublished data). d v. = variable result (one strain positive).
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The other two Phz+ groups that were described in this study were composed of strains belonging to Pseudomonas orientalis and Pseudomonas synxantha (Fig. 2). Little is known about the biology of these two species but both appear to be rhizosphere colonizers with biocontrol activity. Isolates of P. orientalis have been previously recovered from the potato endosphere and rhizosphere of cereal crops in the U.S. and Germany (3, 12, 25, 33). P. synxantha includes one strain, P. synxantha BG33R, with a recently published genome sequence (20). BG33R was originally isolated from the rhizosphere of peach and has been studied as a biocontrol agent of plant pathogenic nematodes. Interestingly, results of our MLSA profiling revealed that BG33R does not fall within the same cluster as the IPNW Phz+ P. synxantha strains and occupies an intermediate position between P. synxantha and P. cedrina (data not shown).
Our results provide the first evidence for phenazine production in both P. orientalis and
P. synxantha. The type strain of P. orientalis was described as phenazine-negative (7) but the absence of phenazine production was established solely on the basis of bacterial cultivation on
King’s A medium, which does not definitively prove the lack of phenazine biosynthesis genes.
The capacity to produce phenazines and biosurfactants was previously described in P. libanensis, a species closely related to P. synxantha (Fig. 2). P. libanensis M9-3, a strain originally isolated from a greenhouse hydroponic system was shown to produce PCA and the biosurfactant viscosin
(36). Interestingly, the partial 16S rRNA sequence of strain M9-3 is 100% identical to that of P. fluorescens 2-79, which also produces PCA and a viscosin-like biosurfactant.
The analysis of our 2009 and 2010 collection of strains revealed that over 67% of all Phz+ isolates belonged to P. aridus and P. cerealis (35.7% and 31.5% respectively). Both provisional species were isolated from the majority of sites throughout the IPNW (Table S2). P. orientalis strains represented 28.4% of the isolates and P. synxantha strains represented only 4.4% of the
130 isolates. There were no clear geographic patterns of Phz+ strain isolation however the majority of
P. orientalis strains were isolated from higher precipitation zones in eastern Washington State or from field sites in south central Washington State (Fig. 1A & Table S2). Interestingly, P. aridus strains were only isolated from the low-precipitation zone and P. cerealis strains were more cosmopolitan throughout the region. The cosmopolitan nature of P. cerealis was exemplified by the co-occurrence of P. cerealis strains on different plant hosts (spring wheat and two native plants, see Results section). As a result of the observed patterns, we assigned these new Phz+ groups the provisional species names ‘P. aridus’ and ‘P. cerealis’ referring to the unique aridity of the low precipitation zone and the cereal crop production throughout the IPNW, respectively.
In addition to the four Phz+ species described in this study, Phz+ strain Pseudomonas sp. 1209 was found to be less than 95.2% similar in MLSA to any previously described species (Table 3).
Based on MLSA, strain 1209 could represent an additional previously unobserved Phz+ species found in the IPNW, although we have yet to describe phenotypic characteristics needed to differentiate it from other related Pseudomonas species.
To investigate the influence of different environmental factors on the distribution of the four Phz+ Pseudomonas species in the IPNW, we also used a culture-independent approach and compared the diversity of phzF alleles in the rhizosphere of winter wheat grown in different precipitation and agroclimatic zones (Fig. 1B). A general effect of mean annual precipitation on colonization frequency (c.f.) was observed with lower than expected c.f.’s at S2.1, S2.2 and S2.4 and higher than expected c.f.’s at S2.8 (Table 1). A significant effect of precipitation on c.f. was previously shown by Mavrodi et al. (24). Rarefaction curves indicated that even with only 44 to
55 phzF clones sequenced, we were able to almost completely sample the Phz+ rhizosphere community at each site (Fig. S1). Taken together with the population densities of 105 to 106 Phz+
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CFU per gram of root, these results indicated very low overall diversity of the Phz+ community at most sites (Table 1). Although we were unable to show a major effect of any environmental variable on a particular Phz+ species, we did make several interesting observations about the general abundance of Phz+ species in the sampled area. Our results revealed that P. synxantha- like strains were not present on any of the roots samples for rhizosphere DNA and likely are rare in the IPNW. The P. orientalis-like strains were found in both low- and high-precipitation zones, whereas the majority of P. aridus-like strains originated from the low precipitation zone. In the wet-cool annual crop agroclimatic zone (site S2.8) most sequenced phzF alleles were similar to that of Pseudomonas sp. 1209, indicating that it is potentially a Phz+ species adapted mostly to higher precipitation areas.
The overall diversity of the Phz+ community appeared to be significantly affected by soil silt percent (Fig. 4). Soil silt is known to have a considerable impact on soil structure, soil bacterial community structure and biocontrol efficacy by PCA-producing bacteria. For example,
Carson et al. (5) compared mineralogically homogenous soils with different amounts of silt and clay and found that it affected the structure of the soil bacterial community. Additionally,
Ownley et al. (32) showed that soil percent silt was positively correlated with disease severity on roots infected with Ggt in the presence of P. fluorescens 2-79 in the greenhouse however the reason for this result remained unclear. Further analysis of the specific effect of soil type on the root colonization and PCA production by the four naturally occurring Phz+ Pseudomonas species described in this study will need to be done under controlled greenhouse conditions.
The other environmental variable that most significantly affected the Phz+ rhizosphere community was agroclimatic zone. The overall diversity of Phz+ Pseudomonas spp. was lowest under ‘grain/fallow’ management (Fig. 4). It has been shown that cereal rotation history can
132 influence the genotypic diversity of Phz+ communities and winter wheat/fallow cropping generally results in low Phz+ genotype community evenness but high genotype diversity (33).
The low precipitation grain/fallow agroclimatic zone is highly specific for P. aridus and P. cerealis with each field likely supporting one of these novel species. The results of the present study show that the diversity of Phz+ species is not affected by grain/fallow, but at the subspecies-level it is.
Results of this study provide new insights into the evolutionary ecology and global distribution of beneficial plant-associated phenazine-producing bacteria. Until this study was conducted, it has been thought that in fluorescent Pseudomonas spp. the capacity to produce phenazines was limited to just a few species (25). The results of this work show that phenazine genes may be shared by a greater number of pseudomonads than previously thought. We demonstrated the presence of an extended complex of phenazine-producing species in the agricultural and non-cropped soils of the Inland Pacific Northwest of the United States. The newly described P. aridus and P. cerealis are particularly abundant in this semi-arid region and may have unique traits allowing them to thrive in the rhizosphere of cereals grown under dryland conditions. By correlating the distribution and frequencies of phzF alleles with the agroclimatic and edaphic factors at different areas of the IPNW, we also demonstrated that, in addition to mean annual precipitation, the type of agroclimate zone and soil silt content significant affect the diversity of indigenous Phz+ communities. These results provide new insights into environmental factors affecting the distribution and activity of phenazine-producers and further contribute to the rational exploitation of beneficial microbial communities for the purpose of improvement of plant health and management of soilborne pahogens in the agroecosystems.
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Acknowledgments
The project described was supported by Award Number T32GM083864 from the National
Institute of General Medical Sciences. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of General Medical
Sciences or the National Institutes of Health.
The authors are grateful to Bellevue College summer interns Jennifer Apple, Emilia Gan, and
Chelsea Stone for their diligent and excellent help performing BOX-PCR and genomic DNA extractions. We also thank Irina Mavrodi for her help in purifying PCR amplified DNA from phzF clones.
134
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Chapter 3
Significance of phenazine-1-carboxylic acid (PCA) to biofilm
production and colony biofilm morphology of phenazine-producing
Pseudomonas species under stress
James A. Parejko1, Dmitri V. Mavrodi2, Linda S. Thomashow3*
1School of Molecular Biosciences, Washington State University, Pullman, WA 99164-4234
2Department of Plant Pathology, Washington State University, Pullman, WA99164-6430
3U.S. Department of Agriculture, Agricultural Research Service, Root Disease and Biological
`Control Research Unit, Pullman, WA 99164-6430
*For correspondence. Mail: USDA-ARS Root Disease and Biological Control Research Unit,
Washington State University, Pullman, WA 99163-6430. E-mail: [email protected]; Tel.:
(+1) 509-335-0930; Fax: (+1) 509-335-7674
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Abstract
Soil-dwelling bacteria, which are exposed to fluctuating levels of water availability that can affect survival and metabolic activity, often form surface-attached biofilms that allow cells to cope with these adverse conditions. Dryland agricultural soils in the low-precipitation zone of the
Inland Pacific Northwest (IPNW) receive less than 350 mm precipitation per year and commonly have significantly reduced soil water potential (below -0.25 MPa) from late spring through fall.
In non-irrigated agricultural fields of this region, populations of root disease-suppressive phenazine-producing (Phz+) fluorescent Pseudomonas spp. have been shown to be present at densities of 105 to 107 colony forming units (CFU) per gram of root of most dryland cereal plants, with high concentrations of the antibiotic phenazine-1-carboxylic acid (PCA) accumulated on those same roots. In irrigated fields Phz+ populations are often undetectable, whereas take-all root disease-suppressive 2,4-diacetylphloroglucinol (2,4-DAPG) producing fluorescent Pseudomonas spp. are abundant under those conditions. Phenazines are known to play a role in biofilm formation by other Pseudomonas species however their effect on biofilm formation by strains of the IPNW Phz+ Pseudomonas spp. has yet to be demonstrated. The present study aimed to identify the effect of PCA and 2,4-DAPG on biofilm formation under reduced water potentials, as a function of matric and osmotic potential. We used PCA biosynthesis-deficient mutants and Phz+ wild-type strains of several species to show that the ability to produce PCA influenced biofilm formation by some, but not all, Phz+ strains studied, and that its impact varied with matric or osmotic stress levels. Upwards of 100 µg ml-1 of exogenously added PCA restored Phz- mutant biofilm formation to wild-type levels in P. orientalis L1-3-08r, but not P. cerealis L1-45-08r, under control and stress conditions. Ultra- structural differences in colony biofilms grown under control and stress conditions also varied
142 among strains and distinct differences were observed between some wild-type Phz+ strains and their Phz- mutants. 2,4-DAPG-producing strains generally were reduced in biofilm levels compared to 2,4-DAPG-deficient mutants and biofilm structure was minimally impacted by the lack of 2,4-DAPG. The results of this study showed that not only do these groups of agriculturally significant antibiotic-producing Pseudomonas spp. differ substantially in biofilm production, but that PCA biosynthesis is important to biofilms formed by some of the IPNW
Phz+ Pseudomonas spp. in reduced water potentials.
Introduction
In the environment, bacteria often form assemblages of structured surface-attached mixed communities known as ‘biofilms’ which form on abiotic and biotic surfaces as the result of cell- to-cell signaling and/or as a response to environmental stimuli. Complex genetic networks involved in biofilm formation have been identified in numerous species, although generally biofilm formation follows a defined pathway (reviewed in (18, 41)). Biofilm development begins with reversible attachment of planktonic cells to a surface due to simple surface attraction forces and the action of outer membrane proteinaceous assemblages (i.e. pili or fimbrae). The second stage of biofilm development defines the process of irreversible attachment and results from the production of various exopolysaccharides, alginate and other secreted polymers that begin to construct the biofilm matrix. Later stages involved biofilm maturation and are characterized by structural and architectural changes in the biofilm matrix. The final stage is dispersion, which results in a release of planktonic cells into the environment. A mature biofilm can provide bacterial cells with increased resistance to external stresses and tolerance to harsh conditions.
143
Members of the genus Pseudomonas are prolific biofilms producers and plant root-associated
Pseudomonas spp. produce biofilm-like microcolonies on the root surface and in adhering rhizosphere soil that serve to provide protection from environmental stresses while allowing the cells to compete and persist in the highly competitive rhizosphere environment (6).
Fluorescent Pseudomonas spp. encompass a large group of rhizosphere-dwelling bacteria some of which produce compounds that promote plant growth and protect the plant from soilborne phytopathogens. One such group of compounds is comprised of heterocyclic nitrogen- containing redox-active phenazines that are produced by certain fluorescent pseudomonads.
Phenazines are multifunctional compounds that are required for rhizosphere competence (33), can serve as signaling molecules (14), can promote microbial-mediated mineral reduction (21) and help some pseudomonads survive anaerobically through electron shuttling (47).
Furthermore, it has been shown that phenazine biosynthesis in P. chlororaphis subsp. aureofaciens and P. aeruginosa plays a role in developing and structuring mature biofilms (15,
25, 26, 40, 48). In P. chlororaphis subsp. aureofaciens 30-84, phenazine production was linked to biofilm formation on abiotic surfaces as a function of total biofilm adhesion (25). Biofilm architecture is also differentially influenced by the phenazine derivative that is produced. For example, a mutant of 30-84 (30-84PCA) that produced only phenazine-1-carboxylic acid (PCA) formed thicker and more structured biofilms compared to the wild-type 30-84 that produced
PCA and 2-hydroxyphenazine-1-carboxylic acid (2-OH-PCA) (26). In contrast, biofilm adhesion in P. aeruginosa PA14 was not affected by phenazine biosynthesis (40). However, P. aeruginosa
PA14 phenazine-deficient mutants exhibited pronounced ultra-structural changes in colony morphology (15, 40). The colonies formed on solid media by phenazine-deficient mutants of P. aeruginosa PA14 were wrinkled and grew vertically with increased surface coverage, as
144 compared to smooth colonies of the wild-type strain that produced both PCA and pyocyanin
(PYO) (15, 40). The addition of 0.1M exogenous PCA resulted in a return to wild-type colony morphology and surface coverage whereas 0.2M PYO was required for complete reversion to the wild-type phenotype (40). Exogenous PYO added to phenazine-deficient mutants also expanded colony surface coverage levels by 35% in excess of that of wild-type PA14 (40), which indicated distinct roles for different phenazines in determining colony morphology.
Our previous surveys of dryland crops grown in the low-precipitation zone of the Inland
Pacific Northwest (IPNW) demonstrated the presence of large populations of Phz+ fluorescent
Pseudomonas spp. in the rhizosphere (105-107 CFU g-1 root fresh weight) that belong to the P. fluorescens species complex (29, 32, 37). These populations are largely endemic to non-irrigated cereal crops and areas with undisturbed soils, and have been shown to produce biologically significant amounts of PCA in situ (up to 1 µg g-1 root fresh weight) (32). Interestingly, the Phz+
Pseudomonas spp. were significantly less abundant in irrigated wheat fields of the same region, which were dominated by Phl+ Pseudomonas spp. that produce the polyketide antibiotic 2,4- diacetylphloroglucinol (2,4-DAPG) (32). As a result of these observations, it has been hypothesized that PCA may help the Phz+ populations survive soil conditions in dryland soils with reduced water availability through the promotion of stress-resistant biofilms in the rhizosphere (29, 30).
Biofilms resistant to stress imposed by low water availability are rich in alginate, expolysaccharides and bacterial cellulose that help cells survive water-limited conditions (3–5).
These biofilm components each have a distinct function in resistance to water limitation, biofilm formation and rhizosphere colonization by Pseudomonas putida mt-2 (35), and homologous compounds likely are produced by other Pseudomonas spp. found in the environment (16, 27).
145
The amount of water available for cells in the environment is a function of the water potential
(Ψ), which is expressed as a negative value, where Ψ = 0 is a fully hydrated environment with complete hydration. Water potential is the sum of six components that include a reference correction (Ψ0), the solute potential (Ψπ), the pressure potential (Ψp), the gravimetric potential
(Ψs), the potential due to humidity (Ψv) and the matric potential (Ψm) (36). On plant roots growing in most non-saline soils, the overwhelming component contributing to the water potential is the effect of Ψm with some positive influence from Ψπ in saline soils and a negative influence from Ψp in the root zone due to hydraulic pull of water into the plant roots.
Furthermore, in unsaturated soil, as water content decreases, the influence of Ψm and Ψπ on bacterial cells increases (36). Past studies of P. putida have investigated the effects of simulating water-limitation on biofilm and cell growth and physiology by amending media with membrane non-permeating solutes like polyethylene glycol (PEG, molecular weight 8000) to decrease Ψm or membrane permeating solutes like sodium chloride (NaCl) to decrease Ψπ (3, 5, 19, 44). Some strains of P. putida are able to survive a decrease in water availability down to -1.5 MPa by forming thick biofilms in the presence of low Ψm created with PEG 8000 (3). Decreasing simulated Ψm and Ψπ can cause significant oxidative stress within biofilms and mechanisms for coping with reactive oxygen species (ROS) in the biofilm were found to be mediated by the production of the exopolysaccharide alginate (4). The antioxidant activity of alginate was also found to be present in P. aeruginosa and P. syringae (4). A recent transcriptomic study of P. putida KT2440 found that simulating the Ψm using PEG 8000 does match transcriptome-wide changes observed in cultures subjected to specific Ψm on a pressurized porous surface model
(PPSM) (17). However, the authors speculated that bacterial cells may sense decreased water
146 availability using distinct pathways. As a result, understanding the effect of Ψπ as well as Ψm on bacterial cells could provide substantial insight into the Ψ effects on bacteria in the environment.
The aim of this study was to determine the importance of PCA in biofilm development under simulated matric or osmotic stress in strains representing major groups of Phz+
Pseudomonas spp. from the IPNW. Additionally, we wanted to compare the capacity of Phz+ pseudomonads to survive under matric and osmotic stress as compared to that of the 2,4-DAPG- producing (Phl+) strains P. fluorescens Q8r1-96 and Q2-87, which are adapted to the rhizosphere of irrigated wheat. Results of our study revealed that all tested Phz+ and Phl+ strains coped with matric and osmotic stresses, although most grew slower and to lower cell density under the conditions of matric stress. The comparison of wild type-strains to their isogenic Phz- mutants revealed that each group of phenazine producers has a unique response to matric and osmotic stress, as as manifested by biofilm formation. Furthermore, in some Phz+ species biofilm formation was significantly influenced by the presence/absence of PCA. This study represents the first step towards understanding the physiological implications of phenazine production to the biofilm lifestyle of indigenous Phz+ Pseudomonas spp. thriving in the rhizosphere of dryland cereal crops grown in the IPNW.
Materials & methods
Strains, growth conditions and plasmids. Bacterial strains and plasmids used in this study are described in Table 1. All strains were streaked for isolation from glycerol stocks (kept at -80°C) and grown for 24-48 hrs at 27°C on Pseudomonas Isolation Agar F (PsF, Difco) before use.
Spontaneous rifampicin-resistant (Rifr) derivatives of all strains were produced by growing
147 strains in King’s B Medium (KMB) with 15 µg/ml rifampicin for 8-12 hours. Cultures (100 µl) were spread onto PsF containing 100 µg/ml rifampicin and 40 µg/ml ampicillin, wrapped in aluminum foil and incubated for 48 hrs at 27°C. Any isolated colonies that grew were tested for exoprotease production on skim milk agar (to indicate a functioning GacA/GacS global regulatory system), siderophore production on CAS media (to indicate functioning pyoverdine biosynthesis), growth kinetics on 1/3 KMB and Luria Bertani broth (LB), and morphology on
PsF agar as compared to the non-Rifr wild type. Rifr derivatives that matched the original strain in all four tests were retained and used in place of wild-type strains for the construction of Phz- mutants and in all biofilm assays. Antibiotics were used when appropriate at the following concentrations: rifampicin, 100 µg ml-1; tetracycline, 15 µg ml-1; ampicillin, 40 µg ml-1; and gentamycin, 10 µg ml-1.
The techniques used to isolate the stains used in this study have been previously described (37, 38). P. brassicacearum Q8r1-96 and P. fluorescens Q2-87 were originally isolated from wheat grown in soil from a take-all decline irrigated field in central Washington State near
Quincy, WA (39, 46). In contrast, all of the Phz+ strains (except P. chlororaphis 30-84) used in this study were isolated from non-irrigated dryland wheat plants (except strain 41210) that average less than 350 mm of annual precipitation. During late spring through the summer growing season, the water potential of the soils of these fields can regularly decrease to < -0.5
MPa.
Strains L1-45-08 and 41210 are representatives of a new species of Pseudomonas
(provisionally named ‘P. cerealis’) that produces PCA as well as an unknown biosurfactant.
Strains of this species can be found on the roots of non-irrigated dryland cereal crops and native plants in undisturbed soil throughout the IPNW. Pseudomonas cerealis L1-45-08 was originally
148 isolated from a non-irrigated dryland winter wheat field near Lind, WA (U.S.) whereas P. cerealis 41210 originated from needle and thread grass (Stipa comata) growing in uncultivated soil 15 miles west of Washtucna, WA (37, 38).
Strain S36709 is a representative of a second new species of Pseudomonas (provisionally named ‘P. aridus’) that produces PCA but no biosurfactants in vitro and is predominantly found throughout the low precipitation zone of the IPNW. Pseudomonas aridus S36709 was isolated from a non-irrigated dryland winter wheat field that is situated 10 miles southeast of Lind, WA
(38).
Pseudomonas orientalis L1-3-08 was isolated from the same winter wheat field as P. cerealis L1-45-08 near Lind, WA whereas P. orientalis R2-66-08W was isolated from a spring wheat/spring barley rotation plot at the Jirava Farm northwest of Ritzville, WA. Both strains produce an orange pigment on PsF and LB agar, produce an unknown biosurfactant and produce small amounts of PCA on potato dextrose agar (37).
P. fluorescens 2-79RN10 is a spontaneous rifampicin resistant derivative of strain 2-79, a model biocontrol strain closely related to Pseudomonas synxantha (38). Strain 2-79 produces
PCA and a viscosin-like cyclic lipopeptide biosurfactant and was originally isolated in 1979 from roots of wheat grown in soil from a take-all decline field near Lind, WA (U.S.) (50).
Pseudomonas sp. 1209 was isolated from wild-rye roots (Elymus glaucus) growing in undisturbed soil near Ritzville, WA. Strain 1209 produces PCA but does not clearly produce biosurfactants, however it does produce highly mucoid colonies on PsF media. It is divergent from the described Pseudomonas species, P. aridus and P. cerealis (38).
149
Table 1. Bacterial strains and plasmids used in this study
Reference or Strain or plasmid Description source
Pseudomonas spp. Wild type, dryland winter wheat rhizosphere isolate, Rifr, P. cerealis. L1-45-08r (37) Phz+; produces PCA Wild type, uncultivated dryland soil (needle & thread P. cerealis 41210r (38) grass- Stipa comata), Rifr, Phz+; produces PCA Wild type, dryland winter wheat rhizosphere isolate, Rifr, P. aridus S36709r (38) Phz+; produces PCA Wild type, dryland winter wheat rhizosphere isolate, Rifr, P. orientalis L1-3-08r (37) Phz+; produces PCA Wild type, dryland spring wheat rhizosphere isolate, Rifr, P. orientalis R2-66-08Wr (37) Phz+; produces PCA P. fluorescens 2-79RN10 Wild type, Rifr, Phz+; produces PCA (42)
- P. fluorescens 2-79Z Phz derivative of 2-79RN10, phzA::lacZ (23) L1-45-08rPHZ- to phzCD::Tetr, PCA-, Rifr, Tetr This study R2-66-08WrPHZ- Wild type, wild-rye rhizosphere isolate, Rifr, Phz+; Pseudomonas sp. 1209r (38) produces PCA Wild type, wheat rhizosphere isolate, Rifr, Phl+; produces P. brassicacearum Q8r1-96Rif (39) 2,4-DAPG P. brassicacearum Q4C5 2,4-DAPG- derivative of Q8r1-96Rif (9) Wild type, wheat rhizosphere isolate, Rifr, Phl+; produces P. fluorescens Q2-87Rif (46) 2,4-DAPG P. fluorescens Q2-87:Tn5 Phl- derivative of Q2-87Rif (28)
Escherichia coli rpsL (Strr) thr leu thi-1 lacY galK galT ara tonA tsx dam JM110 dcm supE44 Δ(lac-proAB) [F´ traD36 proAB (52) lacIqZΔM15], Dam- and Dcm- thi pro hsdR hsdM recA rpsL RP4-2 (Tcr::Mu) (Kmr::Tn7) SM17-1 (λ-pir) (8) λ-pir F– Φ80lacZΔM15 Δ(lacZYA-argF) U169 recA1 endA1 DH5α Invitrogen hsdR17 (rK–, mK+) phoA supE44 λ– thi-1 gyrA96 relA1 Plasmids
pEX18Gm Gene replacement vector; Gmr oriT sacB (22)
pGEM–T Easy SP6 and T7 promoters; Apr f1ori lacZα Promega
p34S-Tc Donor of Tcr cassette; Apr Tcr (12)
150
Construction of Phz- mutants. Phenazine biosynthesis-deficient mutants (Phz-) (except 2-79) were constructed by gene replacement mutagenesis (51). Briefly, a ca. 1.4-kb fragment spanning the phzC and phzD phenazine biosynthesis genes was amplified using the primer pair
PHZ1/PHZ2 (11), ligated into the pGEM-T Easy vector (Promega, Madison, WI) and electroporated into E. coli JM110. A unique ClaI restriction endonuclease site (converted to blunt with the Klenow fragment of DNA polymerase I) within phzC was used to insert a tetracycline-resistance (Tcr) cassette excised from p34S-Tc (12) with SmaI. The resultant plasmids were purified, digested with EcoRI, and the phzCD+Tcr insert was sub-cloned into the
EcoRI site of pEX18Gm (22). The pEX18Gm+ phzCD+Tcr plasmids were electroporated into E. coli S17-1 (λpir) and mated with the Rifr Pseudomonas strains. Double-crossover events were selected and screened by PCR for the absence of plasmid-borne sacB, tetracycline and gentamycin resistance genes as described elsewhere (31).
The absence of phenazine production in the selected Phz- mutants was confirmed by growing cultures in KMB for 72 hrs and extracting phenazines with ethyl acetate. The extracts were screened for the presence/absence of PCA by thin-layer chromatography in a 95:5 benzene:acetic acid solvent system (2, 43). Additionally 10 µl of overnight LB broth culture of
Phz- mutants and Rifr wild-type strains adjusted to an optical density at a wavelength of 600 nm
(OD600nm) of 0.1 were spotted onto potato dextrose agar. Culture spots were grown for ca. 7 to 14 days at 27°C. The presence/absence of dark green or yellow pigmentation within and around the edges of the culture indicated the presence/absence of PCA (37).
Matric or osmotic stress conditions. To simulate reduced water availability, a non-permeating solute (polyethylene glycol (PEG), m.w. 8000) and a permeating solute (NaCl) were added to
1/3-strength King’s medium B (1/3 KMB) (24). The following additions of PEG 8000 or NaCl
151 were made to simulate the corresponding matric or osmotic potential reduction, respectively: for a reduction of 0.125 MPa, 1.6 or 63.6 g liter-1; for a reduction of 0.25 MPa, 3.2 or 100 g liter-1; for a reduction of 0.5 MPa, 6.4 or 150 g liter-1 (19).
Growth kinetics measurements. Growth kinetics of Phz+ or Phl+ wild-type strains and their
Phz- or Phl- mutants were determined in unamended 1/3 KMB and 1/3 KMB amended by -0.25
MPa with PEG 8000 or NaCl. Strains were cultured on PsF agar for 48 hrs at 27°C and a loopful of cells was resuspended in 1.0 ml sterile dH2O. The cells were centrifuged and resuspended in fresh 1.0 ml sterile dH2O. One microliter of the washed cells was inoculated in triplicate into a
96-well microtiter plate prefilled with 200 µl of unamended media or media supplemented with
PEG/NaCl. Measurements were taken every 30 min for 48 hrs on a Safire microtiter plate reader
(Tecan, Männedorf, Switzerland) and the average of two wells of uninoculated media was used as a blank and subtracted from the inoculated wells for each respective medium. The OD600nm values of inoculated triplicate wells were averaged.
Microtiter plate biofilm assay. Static biofilm formation was quantified using a modified version of the biofilm microtiter plate assay described by Merritt et al. (34). Strain inocula were prepared and adjusted in the same manner as those used for growth kinetics. A volume of 200 µl of 1/3 KMB or 1/3 KMB amended with PEG 8000 or NaCl was added to sterile non-tissue treated flat bottom polystyrene 96-well microtiter plate. Each microtiter plate represented one treatment and was inoculated in triplicate with 1.0 µl of adjusted cell suspension. All assays were repeated three times. Biofilms were grown for 48 hrs at 27°C in a sealed box containing moistened paper towels to maintain relative humidity. After 48 hrs, culture liquid was discarded into a waste container and microtiter plate wells were washed with ca. 200 µl dH2O to remove remaining planktonic cells. Attached biofilms were stained for 10 min with 200 µl 0.1% crystal
152 violet solution, stain was discarded and wells were submerged in two consecutive dH2O washes to remove residual crystal violet stain. Microtiter plates were inverted and allowed to air-dry and once dry, stained biofilms were solubilized with 200 µl 95% ethanol for 15 min. Contents of each well were mixed three times by pipetting and 125 µl of each well was transferred to a new
96-well microtiter plate for determination of total mature biofilm adhesion at OD600nm.
To observe the effect of exogenous PCA on the Phz- mutants L1-45-08rPHZ- and L1-3-
08rPHZ-, 200 µl unamended media and media at -0.25 MPa containing PEG 8000 or NaCl was inoculated in the same manner as in the previous microtiter plate biofilm assays. Wild-type L1-
45-08r and L1-3-08r were included in the same plate in addition to the Phz- mutants without added PCA. Purified PCA dissolved in 5% NaHCO3 was added at a concentration of 25, 50 and
100 µg ml-1 to the appropriate wells inoculated with the Phz- mutants.
Colony biofilm formation assay. To analyze ultra-structural differences in biofilm formation and growth of biofilms on solid media, a modified protocol by Dietrich et al. was used (15).
Briefly, a flask of 1/3 KMB solid media was made containing 3.3 g proteose peptone #3 (Difco,
Inc), 3.3 g trypsin (Difco, Inc.), 3.3 g glycerol, 0.5 g K2HPO4 and 4 g. Phytagel (Sigma-Aldrich
Co.) with 40 µg ml-1 Congo Red and 20 µg ml-1 Brilliant Blue R and brought up in 500 ml
Nanopure deionized water. A second flask contained 0.5 l of deionized water with varying quantities of PEG 8000 or NaCl (Nanopure deionized water without solutes was used for unamended media). The flasks were autoclaved and 4.15 ml 1M MgSO4 was heated to at least
80ºC before being added to the first flask and mixed on a stir plate while the PEG 8000 or NaCl solution was slowly added. Plates were immediately poured and allowed to dry under a laminar flow hood before inoculation. Plates were inoculated in five spots with 10 µl of an adjusted cell suspension. Inoculated plates were incubated at 27ºC and placed into a plastic bag after one day
153 of incubation to retain relative humidity. Images were taken two, four and six days after inoculation using a Canon DR-1210C flatbed scanner.
Results
Growth kinetics of Phz+ strains and their isogenic Phz- derivatives under conditions of matric and osmotic stress. To determine the physiological effect of matric or osmotic stress on the growth kinetics of Phz+ Pseudomonas spp. and their Phz- mutant derivatives, a representative from each of the four Phz+ Pseudomonas species found in the low-precipitation zone of the
IPNW dryland agroecosystem was grown in 1/3 KMB amended with PEG 8000 (matric stress) or NaCl (osmotic stress). In unamended 1/3 KMB, most strains had similar growth kinetics with the exception of strains S36709r and 2-79RN10 and their corresponding Phz- mutants that reached late log phase around 25 hrs and 30 hrs after inoculation, respectively (Fig. 1). All other strains reached late log phase between 35 hrs to 40 hrs after inoculation (Fig.1). Under -0.25
MPa of simulated matric stress most strains reached a lower final OD600nm after 48 hrs of growth and had extended log and late log phases (Fig. 1A). Under the matric stress of -0.5 MPa, all strains exhibited further reduction in final OD600nm and significantly longer extension in log phases (Fig. 1A). Strains L1-3-08r and L1-3-08rPHZ- were less impacted by the decreasing matric potential, although these strains formed a noticeable pellicle at the liquid-air interface of the microtiter wells, which may have interfered with OD600nm measurements (Fig. 1A). The final
- OD600nm values for strains S36709r and S36709rPHZ under -0.5 MPa of matric stress were reduced to about half of those observed in the unamended 1/3 KMB (Fig. 1A). Finally, the phenazine-deficient mutant of 2-79 had lower OD600nm readings in both unamended 1/3 KMB
154 and under -0.25 MPa of matric stress, but had approximately the same OD600nm readings as 2-
79RN10 after 48hrs of growth at -0.5 MPa of matric stress (Fig. 1A).
Under simulated osmotic stress of -0.25 MPa, most strains reached log phase after approximately the same amount of time as in unamended 1/3 KMB (Fig. 1B). Strains S36709r,
2-79RN10, L1-45-08r and L1-3-08r and their corresponding Phz- mutants grew under the -0.25
MPa of osmotic stress to higher final OD600nm values than in the unamended 1/3 KMB. Upon further reduction of osmotic potential to -0.5 MPa, S36709r and its Phz- mutant grew to a final
OD600nm value slightly higher than that attained at -0.25 MPa (Fig. 1B). Finally, strain L1-45-08r reached late log phase significantly faster than its Phz- mutant and reached a significantly higher final OD600nm, and Pseudomonas sp. 1209 was largely unaffected by increasing osmotic stress
(Fig. 1B).
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Growth kinetics of Phl+ strains and Phl- mutants under stress. In unamended 1/3 KMB, strain Q2-87 grew at a significantly slower rate than strain Q8r1-96 and reached a slightly lower
- final OD600nm after 48 hrs (Fig. 1). The Phl mutant of strain Q2-87 (Q2-87:Tn5) grew significantly faster than the wild-type under unamended and both stress conditions (Fig. 1). Both
Q8r1-96 and Q2-87 had similar growth kinetics to Phz+ and Phz- strains under matric stress (Fig.
1A). Both strains and their Phl- mutants were mostly unaffected by osmotic stress imposed at -
0.25 MPa and -0.5 MPa osmotic potential (Fig. 1B).
Static biofilm formation of Phz+ or Phl+ strains and their isogenic mutants under simulated matric and osmotic stress. Biofilm formation of representatives of four different Phz+
Pseudomonas species (P.orientalis, P. synxantha and two new Pseudomonas species), two Phl+ strains (P. fluorescens Q8r1-96 and Q2-87) and their Phz- and Phl- mutant derivatives was quantified in 96-well microtiter plates after 48 hrs of growth. Biofilm formation by wild-type strains under control growth conditions in the unamended 1/3 KMB varied significantly within and between species (Fig. 2 & 3). For Phz+ wild-type strains and their Phz- isogenic mutants, biofilm formation varied significantly by strain (Fig. 2). Within the same species, biofilm formation by P. cerealis L1-45-08r was approximately twice as great as by P. cerealis 41210r, and biofilm formation by P. orientalis R2-66-08Wr was approximately 1.5 times as great as by
P. orientalis L1-3-08r (Fig. 2A, 2B, 2D, 2E). L1-45-08rPHZ- showed the most significant reduction in biofilm formation as compared to its respective wild-type, whereas 41210rPHZ- was not significantly different from its wild-type under any of the conditions tested (Fig. 2A & Fig.
2B). For Phl+ wild-types and their Phl- mutants, under control conditions wild-type Q8r1-96 was significantly reduced as compared to its 2,4-DAPG biosynthesis deficient mutant Q4C5 (Fig.
3A) whereas wild-type Q2-87 was not (Fig. 3B).
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Biofilm formation under simulated matric stress showed a general decrease as a function of decreasing matric potential (Fig. 2 & 3). Pseudomonas sp. 1209 was the only strain that demonstrated progressively increased biofilm formation as matric potential decreased from the control conditions to -0.5 MPa (Fig. 2G). In strain L1-45-08r, biofilm formation of the wild-type strain decreased under the conditions of matric stress whereas the biofilm formation of its isogenic Phz- mutant remained unaffected (Fig. 2A). At -0.125 MPa both Q8r1-96 and Q2-87 were significantly reduced in biofilm levels as compared to their respective Phl- mutants, however at lower matric potentials the difference was less clear (Fig. 3).
Biofilm formation under simulated osmotic stress was mostly strain-specific with no clear trend as osmotic potential decreased (Fig. 2 & 3). Similar to matric stress, strain L1-45-08r was significantly higher in biofilm formation than its Phz- mutant (Fig. 2A), however as osmotic stress increased, biofilm formation of L1-45-08r was not significantly affected. Biofilm formation of R2-66-08W increased significantly as compared to the control at -0.125 MPa and below (Fig. 2E) and at -0.5 MPa the Phz- mutant of this strain was significantly impaired in biofilm formation. At -0.5 MPa, strain 2-79RN10 also formed biofilms better than its phenazine non-producing derivative 2-79Z (Fig. 2F), and a similar trend was observed at -0.125 MPa for
L1-3-08r and its Phz- mutant (Fig. 2D). Q8r1-96 formed less biofilm than its Phl- mutant Q4C5 under all osmotic stress conditions tested and under control conditions (Fig. 3A). Similarly, Q2-
87 was only reduced in biofilm when compared to its Phl- mutant, but the effect was only apparent at -0.125 MPa and -0.25 MPa (Fig. 3B).
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The effect of exogenous PCA on static biofilm formation in P. cerealis L1-45-08rPHZ- and
P. orientalis L1-3-08rPHZ- under conditions of matric and osmotic stress. The effect of PCA on biofilm formation was investigated in detail in strains of two divergent species, P. cerealis
L1-45-08r and P. orientalis L1-3-08r that displayed distinct differences in biofilm formation between wild-type and their Phz- mutants. The effect of exogenously added PCA was determined by comparing the biofilm formation in the control medium and in medium supplemented with
PCA at 25, 50 and 100 µg ml-1. Under control conditions, both L1-45-08r and L1-3-08r formed quantifiably more biofilm as compared to the corresponding Phz- mutants (Fig. 4A). However, when the mutants were supplemented with 25 µg ml-1 PCA, biofilm formation increased significantly and the trend continued as more PCA was added (Fig. 4A). In L1-3-08rPHZ-, the addition of 25 µg ml-1 PCA fully restored biofilm formation, whereas in L1-45-08rPHZ- the addition of PCA significantly enhanced biofilm formation compared to the untreated control but never returned it to the wild-type level.
Under -0.25 MPa of matric and osmotic stress, both Phz- mutants also demonstrated significantly reduced biofilm formation as compared to the corresponding wild types (Fig. 4B).
In matric and osmotic stress conditions, both Phz- mutants responded in a similar manner to the control conditions when PCA was added. Biofilm formation returned to wild-type levels when
25 µg ml-1 PCA was added to L1-3-08rPHZ- under -0.25 MPa matric and osmotic stress conditions (Fig. 2B & 2C). Exogenously added PCA did not return biofilm formation in L1-45-
08rPHZ- to wild-type levels under -0.25 MPa matric or osmotic conditions (Fig. 2B & 2C).
However, the addition of PCA to L1-45-08rPHZ- under -0.25 MPa matric stress did not have a significant effect compared to control conditions or -0.25 MPa osmotic stress with no PCA added (Fig. 2).
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162
Colony biofilm morphology of Phz+ or Phl+ strains and Phz- or Phl- mutants under stress.
To determine if structural changes occur in wild-type versus mutant biofilms, colony biofilms were grown on solid 1/3 KMB plates amended with PEG 8000 or NaCl to -0.25 MPa and -0.5
MPa matric or osmotic potential, respectively. Strain L1-45-08r exhibited pronounced architectural difference under all conditions compared to its Phz- mutant (Fig. 5). Wild-type L1-
45-08r was more mucoid than the mutant and covered noticeably less area than the mutant under control and -0.25 MPa osmotic stress after six days of growth, however wild-type and mutant were equal in surface area under other conditions (Fig. 5). The middle of the Phz- mutant colony was also rougher in texture than the wild-type L1-45-08r and contained less Congo Red-bound to biofilm material. Wild-type and Phz- mutant colonies appeared to spread equally well under -
0.25 MPa and -0.5 MPa of matric stress (Fig. 5).
Strain S36709r and its Phz- mutant were equal in surface area coverage under all conditions; however, its Phz- mutant had distinct differences in colony biofilm texture and morphology under control conditions (Fig. 5). Under control conditions and both levels of osmotic stress, wild-type S36709r bound a noticeably larger amount of Congo Red than the Phz- mutant (Fig. 5). Under -0.25 MPa of matric stress the Phz- mutant of S36709r appeared smoother and slightly more mucoid than the wild-type (Fig. 5).
Under control conditions and -0.25 MPa matric stress, strain L1-3-08r spread more than its Phz- mutant after six days of growth, however the wild-type colony contained less bound
Congo Red than the mutant under control conditions but more than the mutant under osmotic stress (Fig. 5). Colonies were significantly smaller under -0.5 MPa of matric stress as compared to all other conditions (Fig. 5). Architectural differences were apparent under osmotic stress beginning at two days after inoculation, whereas distinct differences were only observed under -
163
0.25 MPa of matric stress after six days of growth, with lines radiating from the colony center in the Phz- mutant (Fig. 5).
The colony biofilm formed by strain 2-79RN10 was noticeably larger in surface coverage compared to its Phz- mutant over six days of growth under control conditions only (Fig. 5).
Additionally, the wild-type colony accumulated more Congo Red than the Phz- mutant. Under -
0.5 MPa of osmotic stress, the Phz- mutant colony appeared slightly larger in size than the wild- type strain after four days of growth but equal in size after six days of growth (Fig. 5). There were no pronounced differences in colony morphology or size under matric stress and both wild- type and mutant appeared to be able to spread equally well under -0.25 MPa and -0.5 MPa of matric stress (Fig. 5).
The colony biofilm formed by Q8r1-96 was not noticeably different in size between the wild-type and Phl- mutant, however the mutant appeared to accumulate slightly more Congo Red in a circular pattern under control conditions and osmotic stress (Fig. 5). The Phl- mutant accumulated a significantly greater amount of Congo Red in the center of the colony after six days of growth at -0.5 MPa of osmotic stress (Fig. 5).
Wild-type strain Q2-87 spread slightly less than its Phl- mutant under control conditions and -0.25 MPa of osmotic stress (Fig. 5). Both wild-type and mutant formed clear circles of
Congo Red with uncolored centers under control conditions which were reduced under -0.25
MPa of osmotic stress (Fig. 5), whereas the red circles were filled with red under -0.5 MPa of osmotic stress. After two days of growth under -0.25 MPa and -0.5 MPa of osmotic stress, the
Phl- mutant began breaking down the solidifying agent, Phytagel, in the medium (Fig. 5). This phenomenon was only observed in the wild-type Q2-87 after four days of growth under -0.5 MPa
164 osmotic stress. Under -0.25 MPa and -0.5 MPa of matric stress, the wild-type strain was noticeably rougher in texture than the Phl- mutant and neither wild-type or mutant were smaller in size than control conditions after four days of growth (Fig. 5).
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Discussion
In this study, we used several in vitro techniques to explore the role that PCA biosynthesis plays in surface-attached growth by strains of IPNW Phz+ fluorescent Pseudomonas species under matric or osmotic stress. These PCA-producing strains are of special interest because they are thought to contribute to the natural protection of dryland wheat and barley grown in the IPNW against certain soilborne pathogens. Additionally, their proliferation in soils with low soil moisture, in contrast with the proliferation of 2,4-DAPG-producing populations in areas with high soil moisture, suggests adaptations in Phz+ populations that allow them to survive these conditions. One potential adaptation is through the production of stress resistant biofilms.
Understanding the mechanisms behind these adaptations could aid in applications of rhizobacteria in controlling soilborne pathogens affecting dryland agroecosystems.
To date, the role of phenazines in biofilm formation has only been evaluated in the opportunistic human and animal pathogen P. aeruginosa and in one saprophytic plant-associated species, P. chlororaphis (15, 25, 26, 40). Both of the aforementioned organisms produce more than one phenazine and are highly divergent from the four Phz+ Pseudomonas species assayed in the current study and none of the phenazine-producing Pseudomonas spp. have been investigated for a link between phenazine-mediated biofilm formation and matric or osmotic stress. The
IPNW phenazine-producers formed biofilms better than the well-studied PCA and 2OH-PCA- producing strain P. chlororaphis 30-84 under our conditions and the overall capacity for surface- attached biofilm formation varied significantly among individual IPNW Phz+ strains (Fig. 2 &
3). Under matric stress, biofilm formation by wild-type strains was not clearly different from that of Phz- mutants except for strain L1-45-08r (Fig. 2A). However, under -0.125 MPa and -0.5 MPa of osmotic stress, biofilm formation by L1-3-08r and R2-66-08Wr, respectively, was
167 significantly greater than that of their Phz- mutants (Fig. 2D & 2E). These findings indicate a strain-specific biofilm response to reduced osmotic potential that may be linked to the presence of PCA. Although we didn’t have a Phz- mutant for Pseudomonas sp. 1209r available, its increase in biofilm levels at upwards of -0.5 MPa of matric stress, yet decreased growth kinetics under the same conditions, indicated the strain had adaptations that resulted in a greater amount of biofilm matrix components under matric stress but not osmotic stress (Fig. 1 & Fig. 2G).
Pseudomonas sp. 1209r produced highly mucoid colonies on KMB plates that are similar in morphology to P. aeruginosa strains with mutations in the mucA regulatory gene that result in constitutively produced alginate, a “mucoidy” phenotype, and altered biofilm architecture with greater protective characteristics (13, 20). In P. putida mt-2 biofilms, the amount of total exopolysaccharides produced increased with reduced matric potential (i.e., higher amounts of
PEG 8000) and alginate was specifically present in PEG 8000 but not in NaCl treated biofilms of
P. putida mt-2, P. aeruginosa PAO1 and P. syringae pv. syringae B728a (5). Moreover, an upregulation in alginate biosynthesis and regulatory genes has been observed in P. putida
KT2440 under matric stress applied using a pressurized porous surface model as well (17).
Interestingly, strain 1209 binds a significant amount of Congo Red under all stress conditions around the edge of a colony biofilm with an off-white colony center, which could indicate biofilm spatial differences in exopolysaccharides like alginate compared to non-Congo Red binding compounds like cellulose (data not shown). These findings suggest that Pseudomonas sp. 1209r increased production of alginate-like polysaccharides to cope with higher matric stress, whereas other strains like strain R2-66-08Wr responded uniquely to osmotic stress but not matric stress and may produce other biofilm components to deal with solute stress (Fig. 2E). Further analysis of specific compounds produced by these strains, and the effect of PCA on expression of
168 the corresponding biosynthesis genes, under matric or osmotic stress is an experiment that could reveal intriguing results.
In strains of two of the IPNW phenazine-producing species, P. cerealis and P. orientalis, the formation of biofilms was strongly affected by PCA biosynthesis (Fig. 4). Biofilm formation by the Phz- mutant of strain L1-3-08r recovered to wild-type levels with 25 µg ml-1 exogenous
PCA (Fig. 4). We didn’t test lower PCA concentrations, but in P. chlororaphis 30-84 only 12.5
µg ml-1 exogenous phenazine (not PCA) was required for Phz- mutant biofilm recovery to wild- type levels (25). However, P. cerealis biofilm formation was not recovered by adding up to 100
µg ml-1 of exogenous PCA. This finding indicates a unique role for PCA in biofilm formation by these two species and may indicate that higher PCA concentrations are required for recovering biofilm formation in the L1-45-08r Phz- mutant. In addition to higher PCA concentrations, the pH of the extracellular environment may also influence PCA-mediated biofilm formation by strain L1-45-08r, as pH influences the redox state of PCA. However, based on wild-type L1-45-
08r versus mutant biofilm formation levels and colony biofilm morphology, clearly phenazine biosynthesis is important to biofilm development and structure in this strain. At present, it is unclear as to the mechanism(s) by which phenazines influence biofilm formation and architecture but clearly it is a mechanism that uniquely influences biofilms produced by divergent Pseudomonas spp.
Wild-type strains of P. fluorescens Q8r1-96 and Q2-87, examples of two 2,4-DAPG- producing strains that are associated with the roots of irrigated wheat in the low-precipitation zone of the IPNW, were generally reduced in biofilm adhesion as compared to their Phl- mutants.
This was especially the case for Q8r1-96 and Q2-87 under matric stress and intermediate osmotic stress (Fig. 3). Thus, based on colony biofilm development of wild-type versus mutant strains,
169 the antibiotic 2,4-DAPG does not play as significant of a role as PCA in the morphology and structure of biofilms (Fig. 5). In fact, these findings indicate that 2,4-DAPG has a negative impact on surface adhesion and potentially biofilm development. Our findings provided new evidence to formulate a hypothesis that 2,4-DAPG negatively impacts biofilm formation in Phl+
Pseudomonas spp. under different matric or osmotic stress conditions. Interestingly, the 2,4-
DAPG biosynthesis deficient mutant of P. fluorescens Q2-87 (and wild type after four days of growth) broke down the solidifying agent in the media only under osmotic stress. We used 0.8%
Phytagel to solidify the media as a substitute to agar, as suggested by Halverson and Firestone
(19) for amending solid media with PEG 8000 or NaCl. Phytagel is composed of glucoronic acid, rhamnose and glucose derived from Sphingomonas elodea (45). The observation that Q2-
87:Tn5 breaks down Phytagel only under osmotic stress may indicate that 2,4-DAPG could play a role in regulation of certain genes under solute stress. The role of 2,4-DAPG in cellular physiology and multi-cellular assemblages is not clear at present compared to redox-active phenazines but the physiological role of 2,4-DAPG in the cell should be pursued further.
Due to technical limitations, our study only explored relatively young (ca. 48 hrs old) biofilms on abiotic surfaces (e.g. polystyrene microtiter plate wells and 1/3 KMB solid media) in pure culture. Biofilm formation on biotic surfaces is potentially different in the presence of competing rhizobacteria and plant-derived compounds. Moreover, biofilms in nature are known to be composed of diverse bacterial species. In the case of root-colonizing bacteria, the wheat root continuously releases high and low molecular weight compounds that can provide rhizobacteria with a plentiful carbon source and shape the rhizosphere biochemical conditions like pH (1). To our surprise, PCA biosynthesis does not appear to be clearly involved in biofilm adhesion in strains of P. aridus and P. synxantha (P. fluorescens 2-79RN10 is a member of the P. synxantha
170 species) (Fig. 2). Subtleties in biofilm formation dynamics and structure cannot be pinpointed by using the total biofilm formation assays we used in this study. In order to better understand the biofilm-related mechanisms in these two species, flow cell experiments could reveal intricate details about the earlier and later stages of biofilm development with continuous flow of new nutrients and removal of waste.
The current study did not aim to determine the effect of amphipathic biosurfactants produced by some of the Phz+ strains. However in P. aeruginosa the biosurfactant rhamnolipid played an essential role in structuring biofilms by creating and maintaining open channels that allowed the flow of nutrients and oxygen in mushroom-like clustered biofilms. Rhamnolipid also increased cellular migration along surfaces and into biofilms and played a role in the transition from a sessile to a planktonic lifestyle (7). Correspondingly, biosurfactants produced by dryland crop rhizosphere-colonizing bacteria would help cells move along the root surface under matric potentials that are generally considered highly inhibitory toward flagellar-mediated cell motility
(10, 36).
PCA and 2,4-DAPG-producing fluorescent Pseudomonas species represent two agriculturally significant microorganisms that control soilborne fungal phytopathogens which significantly reduce crop yields worldwide (28, 49). This is the first study to investigate the effect of PCA and 2,4-DAPG biosynthesis on surface-attached biofilm formation and architecture under matric or osmotic stress. As we begin to understand the molecular mechanisms involved in antibiotic biosynthesis and cell survival in these strains at the matric and solute potentials in unsaturated dryland soils, we will significantly improve our understanding of rhizosphere colonization and competency by PCA or 2,4-DAPG-producing Pseudomonas species on plant roots in the dryland agroecosystem.
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Chapter 4
Biological control of Rhizoctonia solani AG8 C-1 by phenazine-1-
carboxylic acid-producing Pseudomonas spp. isolated from the Inland
Pacific Northwest (U.S.) dryland agroecosystem
James A. Parejko1†, Ahmad Kamil Jaaffar2†, David M. Weller3,
Linda S. Thomashow3*
1School of Molecular Biosciences, Washington State University, Pullman, WA 99164-4234
2Department of Plant Pathology, Washington State University, Pullman, WA99164-6430
3U.S. Department of Agriculture, Agricultural Research Service, Root Disease and Biological
`Control Research Unit, Pullman, WA 99164-6430
*For correspondence. Mail: USDA-ARS Root Disease and Biological Control Research Unit,
Washington State University, Pullman, WA 99163-6430. E-mail: [email protected]; Tel.:
(+1) 509-335-0930; Fax: (+1) 509-335-7674
† These authors contributed equally to this work
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Abstract
Direct-seed (i.e., reduced or no-till) crop management provides substantial improvements in sustainability for dryland cereal production in the Inland Pacific Northwest (IPNW) dryland agroecosystem of the western United States. However, the root disease Rhizoctonia root rot and bare patch, caused by Rhizoctonia solani AG-8, result in more severe crop yield losses in direct- seeded wheat compared to conventional management practices and few solutions exist, which has been a major barrier in further adoption of sustainable direct-seed cropping by IPNW dryland farmers. Root-colonizing bacteria of the P. fluorescens lineage that produce phenazine antibiotics are abundant in IPNW dryland cereal fields. Phenazine-producing (Phz+)
Pseudomonas spp. are known to suppress several soilborne fungal phytopathogens, although the protection of wheat roots from R. solani AG-8 by strains of Phz+ Pseudomonas spp. found in the dryland region of the IPNW was largely unknown. Furthermore the significance of phenazine production to the suppression of Rhizoctonia root rot of wheat had yet to be demonstrated. In this study, we show that strains belonging to all four Phz+ Pseudomonas spp. found in IPNW dryland fields (P. aridus, P. cerealis, P. orientalis and P. synxantha) inhibit R. solani AG-8 C-1 in vitro to levels at or above the model biocontrol strain P. fluorescens 2-79. In Rhizoctonia root rot greenhouse bioassays with the same Phz+ strains, we observed significant plant root protection with some strain-level variability in the extent to which root rot was suppressed in both raw and pasteurized soil. Finally, we used Phz- isogenic mutants of the Phz+ Pseudomonas species in the greenhouse bioassays and found that PCA production plays an important role in the biocontrol of
Rhizoctonia root rot by all strains.
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Introduction
Wheat has been cultivated in the low precipitation zone (<300 mm annual precipitation) of the
Columbia Plateau of the Inland Pacific Northwest (IPNW) without irrigation for over 125 years.
The low-precipitation zone is the largest contiguous crop production region in the western
United States (>1.56 million cropland hectares) and wheat exports alone bring over $300 million to the regional economy (25). Traditionally, winter wheat has been grown with a two-year soil moisture management regimen (wheat-summer fallow rotation) which is dependent upon multiple passes with soil tilling equipment. More recently, sustainable reduced and no-till farming practices have been developed to retain soil moisture, reduce agricultural fossil fuel consumption and reduce soil erosion, but soilborne fungal root pathogens like Rhizoctonia solani anastomosis-group 8 (AG-8) causes more severe disease in no-till cropping as compared to conventional cropping systems (20, 27).
R. solani AG-8, causal agent of Rhizoctonia root rot and bare patch, has been an important pathogen in Australia for decades, but it was first discovered in the United States in
1984 in the IPNW (30) and is thought to be ubiquitous there. The fungus infects the plant seminal and crown roots, degrading the root cortex and resulting in root lesions and severed roots that resemble “spear tips” (30). Patches of stunted and dead and dying plants ranging in size from several centimeters to many meters in size develop in wheat and barley fields, significantly reducing plant vigor and yield (27). In a transition from conventional tillage to no-till wheat, disease caused by R. solani reaches a maximum level in the third and fourth years, with an 18 to
41% reduction in wheat yield in no-till as compared to conventionally tilled fields (28). It is hypothesized that conventional tilling mechanically breaks up R. solani mycelial networks
181 thereby reducing the density and inoculum potential of the infectious agent and also reducing soil moisture, making it less conducive to root infection by R. solani (21). Unfortunately there are currently few viable solutions for controlling this pathogen in no-till cropping systems and there is a lack of genes in wheat for resistance to R. solani (1). One promising solution is the biological control of R. solani AG-8 by indigenous rhizosphere-colonizing Pseudomonas spp. that produce antibiotics including phenazines.
Phenazines are heterocyclic nitrogen-containing compounds that exhibit antibiotic activity many soilborne fungal phytopathogens (12, 29). Phenazine-1-carboxylic acid (PCA), produced by the model biocontrol strain Pseudomonas fluorescens 2-79, is highly inhibitory to the growth of a wide range of soilborne fungal phytopathogens in vitro, with activity against
Gaeumannomyces graminis var. tritici, several Pythium spp. (P. aristosporum, P. heterothallicum and P. volutum) and R. solani at 1 µg PCA ml-1 minimum inhibitory concentration (5). However, R. solani AG-8 is the least sensitive of the R. solani AGs to PCA at a 50% effective dose of 16 µg ml-1, although not significantly less sensitive than R. solani AG-2-
1 (8). Moreover, a recombinant strain of P. fluorescens Q8r1-96 that produced both the polyketide antibiotic 2,4-diacetylphloroglucinol (2,4-DAPG) and PCA showed enhanced biological control of R. solani AG-8 as compared to the P. fluorescens Q8r1-96 wild-type that only produced 2,4-DAPG (7). Indigenous phenazine-producing (Phz+) Pseudomonas spp. are essential to suppression of Fusarium wilt in the Châteaurenard suppressive soils found in France, and the suppressive nature of the soil requires an interaction with non-pathogenic Fusarium oxysporum (14). Clearly, Phz+ Pseudomonas spp. have significant potential for natural suppression of soilborne fungal phytopathogens like R. solani AG-8.
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Phz+ Pseudomonas spp. that synthesize cyclic-lipopeptides (CLPs) in addition to phenazines are particularly effective at controlling Pythium splendens on bean (Phaseolus vulgaris L) and Pythium myriotylum on cocoyam (Xanthosoma sagittifolium L Schott) (22) as well as Rhizoctonia solani AG 2-2 and AG 4 HGI on bean (2). It was shown that P. aeruginosa
PNA1controlled damping-off of bean by P. splendens and degraded P. myriotylum mycelium only when both rhamnolipid CLP and phenazine were present (22). Also, Pseudomonas
CMR12a, a member of the P. putida species complex, controlled Rhizoctonia root rot of bean through the production of two different non-rhamnolipid CLPs and phenazines (CMR12a produces PCA and phenazine-1-carboxamide, PCN). However, the biocontrol activity of
CMR12a was still present in both Phz- and CLP- mutants as compared to the double Phz-/CLP- mutant or a GacA regulatory mutant (2). Both studies suggested a clear synergy between CLPs and phenazines in controlling Pythium spp. and R. solani. Both studies also revealed distinct mechanisms of action by different CLPs and varying degrees of CLP and phenazine synergism in biological control of soilborne fungal phytopathogens.
Phz+ Pseudomonas strains of Pseudomonas orientalis, P. synxantha and two new
Pseudomonas species colonize the roots of cereal crops throughout the IPNW dryland agroecosystem (19). Three of the four Pseudomonas species also produce biosurfactants in vitro
(17). With the knowledge that 1) PCA is highly effective at inhibiting R. solani, 2) PCA synthesis is important to suppression of Rhizoctonia root rot and 3) interactions between biosurfactants and phenazines enhance inhibition of R. solani, we hypothesized that the four indigenous Phz+ Pseudomonas species from dryland fields would be effective at controlling
Rhizoctonia root rot on wheat as a result of PCA biosynthesis. In this study, we investigated the biocontrol activity of representatives of these Pseudomonas species isolated from the IPNW
183 dryland agroecosystem against the highly aggressive Rhizoctonia root rot and bare patch wheat pathogen R. solani AG-8 (16). Inhibition assays in vitro against R. solani revealed that the strains differ in their inhibitory capacity and that the level of inhibition is not species-specific, although most were significantly better at inhibiting R. solani than P. fluorescens 2-79. Greenhouse bioassays showed that all strains provided significant protection to wheat plants in both raw
(natural) and pasteurized soil. Phenazine deficient mutants (Phz-) were significantly reduced in plant protection compared to their Phz+ wild-type parent strains in pasteurized soil.
This is the first study to demonstrate biological control of R. solani AG-8 by indigenous
Phz+ Pseudomonas spp. on wheat. It is also the first study to demonstrate the importance of PCA to the suppression of Rhizoctonia root rot of wheat in these new Phz+ Pseudomonas species from the IPNW dryland agroecosystem. The biological control of R. solani by these indigenous Phz+
Pseudomonas species could represent a promising solution to the problem of reduced yields in dryland no-till wheat fields, leading to a greater adoption of sustainable no-till crop practices by wheat farmers in the IPNW.
Materials & methods
Organisms, growth conditions and plasmids. Bacterial strains and plasmids used in this study are described in Table 1. All strains were streaked for isolation from glycerol stocks maintained at -80°C and grown for 24-48 hrs at 27°C on Pseudomonas Isolation Agar F (PsF, Difco) before use. Bacterial strains and plasmids used in this study are described in Table 1. All strains were streaked for isolation from glycerol stocks (kept at -80°C) and grown for 24-48 hrs at 27°C on
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Pseudomonas Isolation Agar F (PsF, Difco) before use. Spontaneous rifampicin-resistant (Rifr) derivatives of all strains were produced by growing strains in King’s B Medium (KMB) (10) with rifampicin (15 µg ml-1) for 8-12 hours. Cultures (100 µl) were spread onto PsF containing rifampicin (100 µg ml-1) and ampicillin (40 µg ml-1), wrapped in aluminum foil and incubated for 48 hrs at 27°C. Any isolated colonies that grew were tested for exoprotease production on skim milk agar (to indicate a functioning GacA/GacS global regulatory system), siderophore production on CAS media (to indicate functioning pyoverdine biosynthesis), growth kinetics on
1/3-strength KMB and Luria Bertani broth (LB) and morphology on PsF agar as compared to the non-Rifr wild type. Rifr derivatives that matched the original strain in all four tests were retained and used in place of wild-type strains for the construction of Phz- mutants and in all biocontrol assays.
The sampling and isolation techniques used to isolate the strains in this study have been described previously (18, 19). The sources of the majority of the strains used in this study were described in the Materials & Methods of Chapter 3 of this dissertation. Two of the strains not described in Chapter 3 are P. aridus R1-43-08 and P. synxantha R6-28-08. P. aridus R1-43-08 and P. synxantha R6-28-08 were isolated from a commercial winter wheat field and an irrigated wheat field near Ritzville, WA, respectively (18). Both strains produce PCA but do not produce biosurfactants in vitro.
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Table 1. Bacterial strains and plasmids used in this study
Reference or Strain or plasmid Description source
Pseudomonas spp. Wild type, dryland winter wheat rhizosphere isolate, Rifr, Phz+; P. cerealis. L1-45-08r (18) produces PCA Wild type, uncultivated dryland soil (needle & thread grass- Stipa P. cerealis sp. 41210r (19) comata) isolate, Rifr, Phz+; produces PCA Wild type, dryland winter wheat rhizosphere isolate, Rifr, Phz+; P. aridus sp. S36709r (19) produces PCA Wild type, dryland winter wheat rhizosphere isolate, Rifr, Phz+; P. orientalis L1-3-08r (18) produces PCA Wild type, dryland spring wheat rhizosphere isolate, Rifr, Phz+; P. orientalis R2-66-08Wr (18) produces PCA P. fluorescens 2-79RN10 Wild type, Rifr, Phz+; produces PCA (29)
- P. fluorescens 2-79Z Phz derivative of 2-79RN10, phzA::lacZ (9) L1-45-08rPHZ-, S36709rPHZ-, This study phzCD::Tetr, PCA-, Rifr, Tetr L1-3-08rPHZ- & Chapter 3 Escherichia coli rpsL (Strr) thr leu thi-1 lacY galK galT ara tonA tsx dam dcm JM110 supE44 Δ(lac-proAB) [F´ traD36 proAB lacIqZΔM15], Dam- and (31) Dcm- SM17-1 (λ-pir) thi pro hsdR hsdM recA rpsL RP4-2 (Tcr::Mu) (Kmr::Tn7) λ-pir (3)
F– Φ80lacZΔM15 Δ(lacZYA-argF) U169 recA1 endA1 hsdR17 DH5α Invitrogen (rK–, mK+) phoA supE44 λ– thi-1 gyrA96 relA1 Plasmids
pEX18Gm Gene replacement vector; Gmr oriT sacB (6)
pGEM–T Easy SP6 and T7 promoters; Apr f1ori lacZα Promega
p34S-Tc Donor of Tcr cassette; Apr Tcr (4)
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Phz- mutant construction and confirmation. Phenazine biosynthesis deficient mutants (Phz-) were constructed as described in the Materials & Methods of Chapter 3 of this dissertation. The
Phz- mutant P. fluorescens 2-79Z was previously constructed by Khan et al. (9).
Bacterial seed treatments. For greenhouse biocontrol studies (see below), bacterial strains were grown on PsF agar (Difco, Inc.) for 48 hours at 27ºC. Two to three loopfuls of each isolate were suspended in 1.0 ml of sterile distilled water (dH2O), washed twice by centrifugation for 3 min at
14,000 rpm in an Eppendorf centrifuge and the washed cell suspension was adjusted to an
OD600nm of 5. For each 4.68 g of seed, 173 µl of the washed cell suspension, 107 µl of sterile dH2O and 280 µl of sterile 2% (w/v) methylcellulose (MC; Sigma Chemical Co., St. Louis, MO) were mixed in a test tube by vortexing briefly and shaking by hand for 3 min. The seeds were air dried in a hood for at least an hour until dry and stored at 4ºC until sown. The density of bacteria per seed was determined by using ten seeds in triplicate from each treatment with the terminal dilution endpoint assay (15) in 1/3-strength KMB supplemented with cycloheximide (100 µg ml-
1), ampicillin (40 µg ml-1), chloramphenicol (13 µg ml-1) and rifampicin (100 µg ml-1).
Generally, the final density of the bacteria applied to the seeds was 106 to 107 CFU seed-1. Two controls were performed alongside the bacterial treatments, one using no bacterial treatment and no MC and one using no bacterial treatment with MC. A control with MC but without bacterial treatment and R. solani AG-8 was performed in initial biocontrol assays.
In vitro R. solani inhibition assays. Inoculum of R. solani AG-8 C-1 for use in in vitro inhibition assays was grown for four to six days on full strength PDA before a mycelial plug (4 mm diameter) was removed from the plate and placed mycelial side down in the center of a fifth- strength PDA. Bacterial inoculum was grown overnight in 5 ml Luria-Bertani (LB) broth at 28ºC with 200 rpm shaking, standardized to an OD600nm of 0.12 to 0.14 in sterile dH2O and deposited
187 in two 7 µl spots on opposite sides 1 cm from the edge of a plate inoculated in the center with a mycelial plug and incubated at 27ºC. A control plate with only a mycelial plug and no bacterial treatment was also set up. Once the mycelium on the control plate reached the edge of the plate
(four to five days of growth), bacterial treatment inhibition measurements were recorded.
Inhibition was measured as the distance from where the mycelial plug was placed to the edge of mycelial growth and also to the edge of the bacterial treatment colony. The inhibition index was determined as the distance of mycelial growth divided by the distance to the bacterial colony.
The two measurements were averaged per plate and each treatment was repeated at least once.
Preparation of oat kernel inoculum. Inoculum of R. solani AG-8 C-1 for use in greenhouse biocontrol assays was prepared essentially as described by Kwak et al. (11). Briefly, 250 ml of whole oat grains and 300 ml of sterile dH2O were mixed in a one-liter Erlenmeyer flask and autoclaved for 90 min on two consecutive days. Small pieces of agar from a 7-day-old culture of
R. solani AG-8 C-1 were added to the flasks and then incubated for three to four weeks at room temperature. Colonized oat kernels were dried for two days under a stream of sterile air and stored at 4ºC.
Greenhouse R. solani AG-8 C-1 root rot biocontrol assay. To assess the biological control potential of Phz+ Pseudomonas spp. against R. solani, tube assays were conducted in a growth chamber at 16ºC with a 12-hour photoperiod essentially as described by Ownley et al. (17) with the following modifications. Briefly, tapered plastic tubes (2.5 cm wide at the top x 16.5 cm long) were plugged at the bottom with a cotton ball and supported in a hanging position in plastic racks (200 tubes per rack). Each tube was filled with 10 cm of sterilized vermiculite topped with a 2-cm-layer of air-dried and sieved Quincy virgin soil (Shano silt loam). For some experiments, the soil was pasteurized (60ºC for 30 min) to reduce the interference from or interactions with
188 indigenous soilborne microorganisms and then air dried. The soil was mixed with 0.7% (wt/wt) oat kernel inoculum of R. solani AG-8 C-1 that had been ground and sieved to obtain particle sizes of 0.25 to 0.5 mm prior to addition to the soil.
Three wheat seeds (cv. Penawawa) coated with Phz+ Rifr wild-type bacteria (see above) were placed on the soil surface and covered with a 2-cm-thick layer of sterilized vermiculite. The control tube without pathogen or bacterial treatment was prepared in a separate room with the same soil, vermiculite and 2% methylcellulose. Five tubes served as a treatment replicate and each treatment was replicated five times. Treatments were arranged in a randomized complete block design. Each tube received 10 ml of H2O containing metalaxyl (0.075 g wettable powder liter-1 of tap water, Syngenta, Wilmington, DE) to suppress indigenous Pythium spp. that can cause damping-off and interfere with root rot disease ratings. The racks of tubes were covered with transparent plastic for two days before being moved into a growth room at 16ºC with a 12- hour photoperiod (17). The plastic was removed after emergence and plants were watered twice weekly with 12 ml of H2O and once weekly with 1/3-strength Hoagland’s solution
(macroelements only) as previously described (17). Four weeks after planting, plants were removed from the tubes and washed under a stream of water. To determine the root rot disease suppression of each bacterial strain, the severity of disease was rated on a scale of 0 to 8 (7), where 0 is a healthy or uninfected (no disease evidence) plant and 8 is a dead or nearly dead plant and shoot length was measured. Treatment replicates were split in half and each half was measured and scored by an independent researcher. Using both measurements, it is possible to determine the overall biocontrol activity of each bacterial treatment on plant growth and root protection for Rhizoctonia root rot.
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A Phz- mutant of a representative strain from each Phz+ Pseudomonas species was also tested using the same greenhouse Rhizoctonia root rot bioassay. Wild-type Phz+ strains and their
Phz- mutants were tested in one experiment using pasteurized Quincy virgin soil.
Statistical analysis. Data from greenhouse biocontrol assays were analyzed using STATISTIX
8.0 software (Analytical Software, St. Paul, MN). Differences in root disease (disease ratings) and plant height among treatments were determined by standard analysis of variance and mean comparisons among treatments were performed by using Fisher’s protected least significant difference test (P = 0.05) or by the Kruskal-Wallis test (P = 0.05).
Results
In vitro inhibition of R. solani AG-8 C-1 by Phz+ Pseudomonas spp. Two separate experiments of in vitro Phz+ Pseudomonas spp. inhibition assays against Rhizoctonia solani AG-
8 C-1 were performed to test the inhibitory capacity of eight strains representing four Phz+
Pseudomonas species isolated from the low-precipitation zone of the IPNW. All eight strains showed significant inhibition (Fig. 1). The assays included a pair of strains from the provisionally named ‘Pseudomonas aridus’ and ‘Pseudomonas cerealis’ as well as Pseudomonas orientalis and Pseudomonas synxantha. Colony morphologies of the strain pairs were largely homogenous with distinct differences between species (Fig. 1). Distinct green and yellow crystals of PCA were visible in strains of P. aridus (R1-43-08r and S36709r) and P. synxantha
(R6-28-08r and 2-79RN10) with yellow pigmentation in P. cerealis colonies (Fig. 1). Colonies of P. orientalis strains (L1-3-08r and R2-66-08Wr) produced a diffusible orange pigment and
190 spread significantly from the point of inoculation (Fig. 1) that was not present in their Phz- mutant derivatives (data not shown).
Inhibition indices of P. cerealis strains were not significantly different from each other and only strain 41210r differed somewhat from 2-79RN10 (Fig. 1). P. aridus R1-43-08r was significantly more inhibitory than 2-79RN10 (t-test p value = 0.0012) and P. aridus strain
S36709r was not different from 2-79RN10 (Fig. 1). The two P. orientalis strains (L1-3-08 and
R2-66-08W) were more inhibitory to R. solani than 2-79RN10 (Fig. 1), although they were not statistically significant (t-test p values = 0.50 and 0.34, respectively). The two P. synxantha strains (R6-28-08 and 2-79RN10) inhibited R. solani equally well (Fig. 1).
191
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Greenhouse R. solani AG-8 C-1 root rot biocontrol assay using wild-type Phz+ strains. The same strains were tested for their capacities to suppress Rhizoctonia root rot caused by R. solani
AG-8 C-1 in both raw and pasteurized Quincy virgin soil using a greenhouse biocontrol assay.
Root disease ratings and plant height data for two separate experiments in raw and pasteurized soil are shown in Tables 2 and 3, respectively.
All eight Phz+ strains suppressed Rhizoctonia root rot in both raw and pasteurized soil on the basis of disease ratings as compared to the untreated controls. In raw soil, mean disease ratings were generally higher than plants grown in pasteurized soil (Tables 2 and 3).
The various bacterial treatments did not result in consistent increases in plant height than the controls containing AG-8 C-1 and AG-8 C-1 + MC (Tables 2 and 3). However, in both pasteurized soil experiments plant height was significantly greater in the P. aridus strains (R1-
43-08r and S36709r) than the AG-8 C-1 control and the S36709r treatment was significantly greater than both controls in both experiments (Tables 2 and 3).
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Table 2. Biological control of Rhizoctonia solani AG-8 C-1 by PCA-producing Pseudomonas spp. in natural soil a
Disease rating Plant height Strain c Treatment (Mean 0-8 scale ± SD) (Mean cm ± SD) sourceb Exp. 1 Exp. 2 Exp. 1 Exp. 2
P. cerealis L1-45-08r Lind 3.05 ± 0.92 BC 2.61 ± 0.75 B 22.09 ± 3.83 BCD 22.83 ± 3.62 ABCD
41210r Washtucna 2.94 ± 1.04 CD 2.70 ± 0.94 B 21.39 ± 4.17 CD 22.34 ± 3.75 BCDE
P. aridus R1-43-08r Ritzville 2.76 ± 0.84 CDE 2.33 ± 0.74 B 21.94 ± 4.06 BCD 22.21 ± 3.36 BCDE
S36709r Lind 2.60 ± 0.72 CDE 2.23 ± 0.83 B 21.89 ± 3.30 BCD 22.02 ± 3.09 DE
P. orientalis L1-3-08r Lind 2.72 ± 0.76 CDE 2.47 ± 0.85 B 22.36 ± 2.74 BC 23.19 ± 3.49 AB
R2-66-08Wr Ritzville 2.81 ± 0.85 CDE 2.51 ± 0.68 B 21.32 ± 4.15 CD 22.63 ± 3.08 ABCD
P. synxantha R6-28-08r Ritzville 2.24 ± 0.755 E 2.36 ± 0.72 B 21.94 ± 3.38 BCD 23.12 ± 3.00 ABC
2-79RN10 Lind 2.32 ± 0.66 DE 2.33 ± 0.71 B 21.79 ± 3.39 BCD 22.92 ± 3.25 ABCD
Controls
+MC 0.16 ± 0.37 F 0.19 ± 0.47 C 24.15 ± 2.85 A 23.48 ± 2.99 A AG-8 C-1 + MC 3.57 ± 0.76 AB 3.42 ± 0.84 A 20.96 ± 3.04 D 21.24 ± 3.23 E AG-8 C-1 - MC 3.89 ± 0.94 A 3.50 ± 0.86 A 21.68 ± 3.70 BCD 22.49 ± 3.33 ABCD
a Experiments were conducted in Quincy virgin soil. b All strain sources are in the low precipitation zone of the IPNW in Washington State. c Means in the same column followed by the same letter are not significantly different at P = 0.05 according to Fisher’s protected least significant difference (plant height) or Kruskal-Wallis all-pairwise comparison (disease rating).
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Table 3. Biological control of Rhizoctonia solani AG-8 C-1 by PCA-producing Pseudomonas spp. in pasteurized soila
Disease rating Plant height Strain c Treatment (Mean 0-8 scale ± SD) (Mean cm ± SD) source b Exp. 1 Exp. 2 Exp. 1 Exp. 2
P. cerealis L1-45-08r Lind 2.44 ± 0.76 BC 2.30 ± 0.67 B 21.58 ± 2.81 BCD 25.13 ± 3.50 ABC
41210r Washtucna 2.43 ± 0.66 BC 2.34 ± 0.56 B 21.49 ± 2.46 BCD 25.44 ± 2.64 AB
P. aridus R1-43-08r Ritzville 2.22 ± 0.54 C 2.37 ± 0.63 B 22.24 ± 2.37 B 24.51 ± 3.31 BCD
S36709r Lind 2.46 ± 0.69 BC 2.37 ± 0.54 B 21.99 ± 3.00 B 25.97 ± 2.10 A
P. orientalis L1-3-08r Lind 2.45 ± 0.56 BC 2.32 ± 0.53 B 21.72 ± 2.38 BCD 25.27 ± 2.88 AB
R2-66-08Wr Ritzville 2.26 ± 0.72 BC 2.47 ± 0.87 B 21.78 ± 2.62 BCD 24.00 ± 3.65 CDE
P. synxantha R6-28-08r Ritzville 2.47 ± 0.60 BC 2.47 ± 0.76 B 21.98 ± 2.93 BC 25.44 ± 3.43 AB
2-79RN10 Lind 2.34 ± 0.59 BC 2.35 ± 0.67 B 20.99 ± 2.73 D 25.61 ± 2.58 A
Controls
+MC 0.01 ± 0.11 D 0.01 ± 0.12 C 23.41 ± 2.00 A 24.53 ± 4.07 BCD AG-8 C-1 + MC 3.50 ± 0.67 A 3.50 ± 0.89 A 20.92 ± 3.11 D 23.71 ± 3.92 DE AG-8 C-1 - MC 3.68 ± 0.64 A 3.70 ± 0.80 A 21.13 ± 2.29 CD 23.16 ± 4.59 E
a Experiments were conducted in Quincy virgin soil. b All strain sources are in the low precipitation zone of the IPNW in Washington State. c Means in the same column followed by the same letter are not significantly different at P = 0.05 according to Fisher’s protected least significant difference (plant height) or Kruskal-Wallis all-pairwise comparison (disease rating).
195
Greenhouse R. solani AG-8 C-1 root rot biocontrol assay using Phz- mutants. Site-specific mutagenesis was used to create Phz- isogenic mutants by inserting a tetracycline resistance cassette into the phenazine biosynthesis gene phzD. One representative Phz+ strain from P. cerealis, P. aridus and P. orientalis was selected for mutagenesis. P. fluorescens 2-79Z (also
Phz-) was created in a previous study and has lacZ fused to phzA (9). Wild-type and their Phz- mutants were tested in the same greenhouse Rhizoctonia root rot biosassay as previously described in this study.
All Phz+ wild-type strains were significantly better at protecting the plant from
Rhizoctonia root rot than their Phz- mutants in pasteurized soil (Table 4). The disease ratings of the roots treated with wild-type Phz+ strains were essentially the same as the first pasteurized soil experiment (Table 3), except for P. aridus S36709r which was slightly better at protecting the roots in this experiment (Table 4). The difference in protection provided by P. cerealis L1-45-
08r compared to its mutant was most striking as disease on roots treated with L1-45-08rPHZ- were actually significantly worse than either untreated control (Table 4). The Phz- mutant of P. aridus S36709r was not significantly different from the other Phz+ wild-type strains and the methylcellulose (MC) treated control (Table 4). Disease severity on the plant roots of seeds treated with the Phz- mutant of P. orientalis L1-3-08r was not significantly different from the control not treated with MC, while P. fluorescens 2-79Z was not significantly different from either untreated control (Table 4).
Plant height was not significantly different for most treatments compared to the controls
(Table 4). Plants treated with the Phz- mutant of P. cerealis L1-45-08r, L1-45-08rPHZ-, were significantly shorter than the rest of the treatments and the controls at a mean height of 20.71 cm
196
± 2.92 cm compared to mean heights of 22 cm to 23 cm in other treatments and the controls
(Table 4).
Table 4. Biological control of Rhizoctonia solani AG-8 C-1 by wild-type Phz+ and Phz- mutant Pseudomonas spp. in pasteurized soil a
Disease rating Plant height Treatment (Mean 0-8 scale ± SD) b (Mean cm ± SD)
P. cerealis L1-45-08r 2.35 ± 0.72 E 22.92 ± 2.13 A P. cerealis L1-45-08rPHZ- 3.43 ± 0.73 A 20.71 ± 2.92 B P. aridus S36709r 1.90 ± 0.77 F 23.05 ± 2.15 A P. aridus S36709rPHZ- 2.42 ± 0.66 DE 23.46 ± 2.32 A P. orientalis L1-3-08r 2.33 ± 0.92 E 22.60 ± 3.33 A P. orientalis L1-3-08rPHZ- 2.92 ± 0.78 B 22.86 ± 3.10 A P. fluorescens 2-79RN10 2.20 ± 0.69 E 23.14 ± 1.91 A P. fluorescens 2-79Z 2.71 ± 0.72 BC 22.56 ± 2.32 A Controls AG-8 C-1 + MC 2.65 ± 0.61 CD 22.73 ± 2.34 A AG-8 C-1 - MC 2.85 ± 0.82 BC 22.22 ± 2.70 A
a Experiments were conducted in Quincy virgin soil. b Means in the same column followed by the same letter are not significantly different at P = 0.05 according to Fisher’s protected least significant difference (disease rating) or Kruskal-Wallis all-pairwise comparison (plant height).
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Discussion
Rhizoctonia root rot is the most severe soilborne disease infecting direct-seeded (i.e., no-till) dryland cereal crops grown in the Inland Pacific Northwest (IPNW) dryland agroecosystem of the western United States. The causal agent, Rhizoctonia solani, can reduce no-till wheat yields by 18 to 41% compared to conventionally tilled wheat and few sustainable solutions for controlling the pathogen exist. In this study, we studied the capacity of strains of Phz+ fluorescent Pseudomonas spp. isolated from plants growing in IPNW dryland soil to inhibit R. solani AG-8 C-1 in vitro and protect plants in the greenhouse. We also used Phz- isogenic mutants to identify the relative importance of PCA to biocontrol of R. solani AG-8 C-1. We found that all Phz+ strains tested, some of which also produce biosurfactants, provided significant plant protection from Rhizoctonia root rot (Table 2 & Table 3). We also found that
PCA production was important to root protection (Table 4). These findings provide substantial evidence that the natural Phz+ populations observed in dryland fields contribute to the suppression of Rhizoctonia root rot. Mavrodi et al. (13) suggested that phenazine producers may contribute to the phenomenon of Rhizoctonia root rot decline recently described in some fields in the low-precipitation zone.
Phenazines and amphipathic cyclic lipopeptide biosurfactants are known to interact to suppress Pythium spp. (22) and R. solani (2). Previously, it was shown that P. orientalis L1-3-08,
P. cerealis L1-45-08 and P. fluorescens 2-79 produce biosurfactants in vitro, whereas P. aridus strains do not (18). High performance liquid chromatography (HPLC) of extracted biosurfactants from P. fluorescens 2-79 indicated that the strain produces an unknown compound with a similar
HPLC chromatogram to the viscosin produced by P. fluorescens SBW25 (unpublished data).
Viscosin and PCA are also produced by P. libanensis M9-3, a strain closely related to 2-79 (24). 198
Viscosin and viscosin-like lipopeptides are potent surfactants by themselves that lyse oomycete zoospores and directly affect the growth of R. solani mycelia (22). The combination of PCA and viscosin in protecting wheat roots from Rhizoctonia root rot has never been demonstrated. The combinatorial action of the two compounds may be crucial to enhancing biocontrol in some Phz+ species, which was shown with rhamnolipids and phenazines in P. aeruginosa PNA1 against
Pythium spp. (22). As was shown in Chapter 3, most Phz+ strains of P. cerealis, P. aridus, P. orientalis and P. synxantha have unique biofilm characteristics as compared to their phenazine- deficient mutants. Similarly surfactants also influence biofilm development and surface motility
(23), which are both important traits in successful bacterial colonization of the rhizosphere.
Considering the variability in biosurfactant and PCA production dynamics of these Phz+ species, it would be interesting to test root rot suppression and biofilm formation with biosurfactant deficient and biosurfactant/PCA deficient double mutants.
The general levels of disease observed in the Phz+ wild-type and Phz- mutant experiment was lower relative to the first Phz+ wild-type experiments in pasteurized soil as a function of the disease observed in both untreated controls (Table 3 and Table 4). The disease severity of
Rhizoctonia root rot can be impacted by temperature and soil moisture that can be difficult to control in the greenhouse for month-long experiments. However it was surprising that seed treatment with the Phz- mutant of P. cerealis L1-45-08r resulted in more disease to the plant roots and significantly stunted plants than both of the untreated controls (Table 4). One potential explanation could be due to biosurfactant production by L1-45-08r. Some cyclic lipopeptides are known to cause increased plant pathogenicity in some bacteria by forming ion channels in plasma membranes, directly solubilizing host membranes, and promoting plant cell wall degrading enzyme access (23). It is conceivable that biosurfactants produced by L1-45-08r, in
199 the absence of PCA, increase disease in plant roots by providing greater access to the root cortex for degradation by R. solani AG-8 C-1 hyphae.
Interestingly, plant protection provided by P. aridus S36709r was not completely dependent upon PCA production in pasteurized soil as the mean disease severity on roots of seeds treated with its Phz- mutant was not significantly different from the other Phz+ wild-type strains (Table 4). This finding is particularly surprising because strain S36709r does not produce measureable biosurfactant levels in vitro (18) and did not inhibit R. solani in vitro at the same levels as the other Phz+ strains (Fig. 1). Additionally, strains of P. aridus appear to produce higher levels of PCA in vitro than other Phz+, as can be seen in the dense accumulation of PCA crystals on PDA media (Fig. 1). Novel Rhizoctonia root rot biocontrol or plant-promoting traits expressed in the rhizosphere by this strain should be investigated in the future.
In direct-seeded cereals, a wheat-barley rotation reduces the severity of Rhizoctonia bare patch (26) which is surprising considering that barley is a better host for Rhizoctonia and it develops more severe root rot as compared to wheat (20). As a corollary, strains of all four Phz+
Pseudomonas species tested for biocontrol activity in this study were found on the roots of spring wheat sampled from an annual spring wheat-spring barley rotation plot (18). However only P. aridus and P. cerealis were isolated from spring barley of the same rotation and they generally had a less robust diversity of Phz+ genotypes than the spring wheat (18). Based on these observations, I suggest that specific Phz+ genotypes of P. aridus or P. cerealis, which exhibit unique PCA production-dependent biocontrol activities, are selected for by spring barley in this rotation and these strains provide substantial plant protection from dryland root rot that can develop into bare patches. Although this specific rotation effect hasn’t been observed before with these Phz+ Pseudomonas species, we are currently testing this hypothesis by measuring
200 relative levels of these Phz+ Pseudomonas spp. in field samples from rotation plots using phz- specific qPCR.
Clearly, the results of this study show that PCA production by representative strains of these four Phz+ Pseudomonas species found throughout the IPNW dryland agroecosystem is important to protecting wheat plants from Rhizoctonia root rot. Considering the Phz+ populations of 105 to 106 CFU g-1 root previously observed, the levels of phenazine that have already been detected in rhizospheres of IPNW dryland cereals (13) and the plant protection provided as a result of PCA production by representative strains of the IPNW populations, it is realistic to hypothesize that these Phz+ populations suppress Rhizoctonia root rot on dryland wheat across the entire region. To what extent the implementation of direct-seeded cropping affects the suppressive capacity of Phz+ Pseudomonas spp. and a specific rotation effect on root rot suppression by these populations remains to be determined.
Acknowledgments
The authors are grateful to Karen Hansen for performing the inhibition assays and taking down greenhouse Rhizoctonia root rot bioassays. The authors are also grateful to Zachary Day for his help in setting up and taking down greenhouse bioassays, watering plants and entering greenhouse data into spreadsheets for statistical analyses.
201
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General Conclusions
A collection of Phz+ fluorescent Pseudomonas spp. strains (n = 412 isolates) from cereal crop rhizospheres from the Jirava and Kagele farms near Ritzville, WA (U.S.) and near Lind, WA
(U.S.) were highly diverse at the sub-strain genotype level. Representatives of the genotypes formed four distinct groups based on genetic and phenotypic differences. Based on partial 16S rRNA sequence, a small divergent group of strains including the model PCA-producing
Pseudomonas fluorescens 2-79 clustered closely with Pseudomonas libanensis, Pseudomonas synxantha and Pseudomonas gessardii while the majority of isolates clustered close to
Pseudomonas orientalis. Partial recA and phzF DNA sequence analysis revealed slightly different results than the genotype clusters. phzF was confirmed to stably maintained in the chromosome as no evidence of horizontal gene transfer was observed. However, evidence for intra-group past recombination events in phzF DNA sequence suggests that closely related strains exchange phz-related genes and, with evidence that phzF is under purifying selection, are evolutionarily flexible in maintaining certain phz alleles under the high selective pressure in the rhizosphere. Carbon substrate utilization patterns confirmed the recA and phzF analysis and showed that the groups likely utilize different root exudates in the rhizosphere or colonize highly divergent rhizosphere niches. Seven different crops were sampled and the genotype diversity and evenness was greatest under continuous spring wheat which may signify a greater spectrum of rhizosphere community traits related to plant growth promotion. This observation reinforces the hypothesis that long term crop monoculture can develop a rhizosphere community that can suppress soilborne fungal phytopathogens.
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A region-wide collection of Phz+ fluorescent Pseudomonas spp. strains (n = 497 isolates) from winter wheat, spring wheat and native plant rhizospheres revealed that the four Phz+ groups found in the first study were distributed across the entire IPNW dryland agroecosystem.
Taxonomic placement of representative strains belonging to the four groups using multi-locus sequence analysis showed that each group was a distinct Pseudomonas species belonging to the
P. fluorescens species complex. Two of these species were < 97% similar in multi-locus sequence, the cut-off for putative new Pseudomonas species, and were provisionally named
Pseudomonas aridus and Pseudomonas cerealis. The other two clades were strains of P. orientalis and P. synxantha and are the first Phz+ strains to be described of these two species.
Another Phz+ strain, Pseudomonas sp. 1209, was < 96% similar to described Pseudomonas species. Significant differences in Phz+ population sizes were observed between spring wheat and native needle and thread grass plants. However spring wheat, common yarrow and needle and thread grass plants all shared strains of P. cerealis, but the majority of Phz+ population strains on each plant were separate Phz+ Pseudomonas species. Small libraries of phzF alleles (n
= 454 clones) from eight IPNW winter wheat fields provided evidence that agroclimatic zone and soil percent silt had a significant impact on Phz+ community diversity in this dryland agroecosystem. The limited species-level diversity in the small phzF libraries, as seen in rarefaction curves, suggested that the Phz+ communities in wheat fields generally contain one or two Phz+ Pseudomonas species per dryland field.
Biofilm formation assays of representative Phz+ wild-type strains belonging to P. aridus,
P. cerealis, P. orientalis and P. synxantha and their Phz- isogenic mutant derivatives demonstrated that PCA production was an important trait for biofilm formation in some of the test strains. As little as 25 µg ml-1 of exogenous PCA added to a Phz- mutant of P. orientalis L1-
208
3-08r caused recovery of biofilm formation to wild type levels, but did not do the same for the
Phz- mutant of P. cerealis L1-45-08r at concentrations of exogenous PCA up to 100 µg ml-1.
PCA production may be important to biofilm formation under matric and osmotic stress in Phz+
P. synxantha and P. aridus strains, but not in control conditions without matric or osmotic stress.
The wild-type 2,4-DAPG-producing strains Pseudomonas brassicacearum Q8r1-96 and P. fluorescens Q2-87 were both reduced in biofilm formation under intermediate matric stress
(-0.125 MPa and -0.25 MPa) compared to their 2,4-DAPG non-producing mutants. A comparison of colony biofilm morphology over six days of growth showed that the Phz+ strains generally grew larger and more structured colony biofilms than the 2,4-DAPG producers. All colony biofilms formed by Phz+ strains were significantly impacted in biofilm architecture under one of the conditions tested (matric stress, osmotic stress or control conditions) when deficient in
PCA production whereas there was little or no contrast in colony biofilm architecture in mutants deficient in 2,4-DAPG production compared to the wild-type strains.
Studies of the biocontrol activity of representative strains from the Phz+ P. aridus, P. cerealis, P. orientalis and P. synxantha against the Rhizoctonia root rot causal agent Rhizoctonia solani AG-8 C-1 demonstrated that all Phz+ species have significant capacities at suppressing this soilborne fungal plant pathogen. Greenhouse bioassays using wild-type Phz+ strains of each species in raw and pasteurized Quincy virgin soil showed wheat root protection at levels roughly equivalent to the model biocontrol strain P. fluorescens 2-79. Phz- mutants of the same strains tested in pasteurized soil were significantly reduced in biocontrol capacity compared to their
Phz+ wild-type strain. However, the Phz- mutant of P. aridus S36709r, although significantly reduced compared to the wild-type, still provided some plant protection.
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