Expanding the host range of Rhynchosporium alismatis, a potential biological control agent for Alismataceae weeds in Australian rice fields.
Wayne Maxwell Pitt BAppSci (Medical and Applied Biotechnology) Honours Class 1
A thesis submitted in partial fulfilment of the requirements for the degree of Doctor of Philosophy
School of Agriculture Faculty of Science and Agriculture Charles Sturt University Wagga Wagga, NSW, Australia April 2003
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I Wayne Maxwell Pitt
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Acknowledgements
ACKNOWLEDGEMENTS
• Peter and Jan Pitt. • The Cooperative Research Centre for Sustainable Rice Production for the provision of funding for this study (Project No. 2401). • Dr Laurie Lewin. • Dr Gavin J. Ash and Dr Eric J. Cother. • Dr Janet L. Taylor, Janet Condie, Carla Barber and the staff of the Plant Biotechnology Institute, NRC, Canada. • Dr Harsh Raman (NSW Agriculture, Wagga Wagga), Dr Rob Loughman and Christy R. Grime (University of Western Australia) for the provision of R. secalis samples. • Megan, Wendy and the staff at the Office for Research and Graduate Studies. • Robyn Lonard at interlibrary loans. • Dr. Karen Bailey and Dr. C. Y. Chen of Agriculture and Agri-Food Canada for supplying the pABC vector. • Eric Hines from CSIRO department of Entomolgy. • Ian Mason for the provision of A. plantago–aquatica plants. • Dr Jim Virgona, Helen Nicol and Dr Neil Coombes for statistical advice and consultation..
i Table of Contents
TABLE OF CONTENTS
STATEMENT OF AUTHENTICITY ...... i
ACKNOWLEDGEMENTS ...... i
TABLE OF CONTENTS ...... ii
LIST OF FIGURES...... iv
LIST OF TABLES...... v
LIST OF ABBREVIATIONS AND STATISTICAL SYMBOLS...... vi
ABSTRACT...... ix
1 Introduction...... 1 1.1 INTRODUCTION ...... 1 1.2 OBJECTIVES OF THIS STUDY ...... 2 2 Literature Review ...... 3 2.1 THE AUSTRALIAN RICE INDUSTRY...... 3 2.2 WEEDS...... 4 2.3 WEED MANAGEMENT IN RICE ...... 5 2.4 DEVELOPMENT OF THE MYCOHERBICIDE...... 6 3 Phylogenetics...... 12 3.1 INTRODUCTION ...... 12 3.2 TAXONOMY AND NOMENCLATURE ...... 13 3.3 CONCEPTS AND METHODOLOGY ...... 14 3.4 MATERIALS AND METHODS ...... 16 3.4.1 Fungal isolate collection, isolation, cultivation, storage and maintenance: ...... 16 3.4.2 Culture preparation and DNA extraction: ...... 18 3.4.3 Amplifying the ITS region (ITS−PCR):...... 18 3.4.4 Sequencing the ITS region:...... 19 3.4.5 Assembling the ITS database:...... 20 3.4.6 DNA sequence alignment and phylogenetic analysis:...... 23 3.5 RESULTS ...... 24 3.6 DISCUSSION ...... 28 4 Population Structure ...... 33 4.1 INTRODUCTION ...... 33 4.2 CONCEPTS AND METHODOLOGIES ...... 34 4.3 TOOLS AND TECHNIQUES ...... 36 4.4 MATERIALS AND METHODS ...... 38 4.4.1 Fungal isolates and DNA extraction:...... 38 4.4.2 Repetitive Repeat motif (ERIC/REP)−PCR:...... 40 4.4.3 Simple sequence repeat (SSR)−PCR: ...... 40 4.4.4 Data analysis:...... 41 4.5 RESULTS ...... 45 4.6 DISCUSSION ...... 50
ii Table of Contents
5 Infection Process ...... 55 5.1 INTRODUCTION ...... 55 5.2 FUNGAL ATTACHMENT AND/OR ADHERENCE TO THE PLANT SURFACE ...... 56 5.3 GERMINATION AND DIFFERENTIATION OF INFECTION STRUCTURES...... 57 5.4 FUNGAL PENETRATION AND PATHOGENESIS...... 59 5.5 PLANT DEFENCES ...... 62 5.6 MATERIALS AND METHODS ...... 67 5.6.1 Fungal isolates: ...... 67 5.6.2 Plant growth and leaf material preparation: ...... 67 5.6.3 Pathogenicity and selection for virulence ...... 67 5.6.3.1. Inoculum preparation:...... 67 5.6.3.2. Leaf disc inoculation and disease assessment: ...... 68 5.6.4 Infection of host and non host species and Microscopy ...... 68 5.6.4.1. Tissue preparation and inoculation:...... 68 5.6.4.2. Light microscopy:...... 68 5.6.4.3. Fluorescence microscopy: ...... 69 5.6.4.4. Electron microscopy:...... 70 5.6.5 Data analysis:...... 70 5.7 RESULTS ...... 71 5.8 DISCUSSION ...... 78 6 Transformation...... 90 6.1 INTRODUCTION ...... 90 6.2 FUNGAL CUTINASES AND HOST PENETRATION...... 91 6.3 TRANSFORMATION...... 92 6.4 TECHNIQUES ...... 93 6.5 PROTOPLAST REGENERATION ...... 96 6.6 SELECTION OF TRANSFORMANTS ...... 96 6.7 FATE OF TRANSFORMING DNA ...... 101 6.8 MATERIALS AND METHODS ...... 106 6.8.1 Fungal isolates: ...... 106 6.8.2 Vector construction and preparation: ...... 106 6.8.3 Preparation of Protoplasts:...... 108 6.8.4 Transformation and plating:...... 109 6.8.5 Molecular analysis of putative transformants:...... 111 6.8.6 Southern analysis of putative transformants and probe construction:...... 111 6.9 RESULTS AND DISCUSSION...... 114 7 General Discussion...... 129
8 Appendices...... 137 8.1 TABLES ...... 137 8.2 FIGURES...... 139 8.3 STATISTICAL DATA (CHAPTER 5)...... 141 9 Publications ...... 153
10 References...... 154
iii List of Figures
LIST OF FIGURES Figure 2.1: The rice growing region of southern New South Wales, Australia ...... 3 Figure 2.2: The Alismataceae family ...... 5 Figure 2.3: Typical appearance and morphology of propagules of R. alismatis...... 8 Figure 3.1: ITS–PCR of R. alismatis and R. secalis isolates...... 25 Figure 3.2: Phylogenetic relationships of R. alismatis and related species...... 27 Figure 4.1: Geographic origin of R. alismatis isolates used in this study...... 38 Figure 4.2: PCR fingerprint patterns of R. alismatis isolates ...... 49 Figure 5.1: Light and fluorescent micrographs during infection studies of R. alismatis ...... 73 Figure 5.2: Scanning electron micrographs during infection studies of R. alismatis...... 75 Figure 5.3: Macroscopic disease symptoms caused by R. alismatis ...... 77 Figure 6.1: Plasmid map of the pABC expression vector ...... 107 Figure 6.2: Plasmid map of the pBARGEM7–2 vector...... 108 Figure 6.3: PCR analysis of putative pABC transformants...... 118 Figure 6.4: Dot blot analysis of putative pABC transformants ...... 120 Figure 6.5: Southern analysis of putative pABC transformants...... 120 Figure 6.6: Confirmation of DNA transfer via southern analysis ...... 121 Figure 6.7: Southern analysis of putative pBARGEM7–2 transformants...... 123 Figure 8.1a: Conidial germination on Alismataceae species ...... 139 Figure 8.1b: Rate of conidial germination on Alismataceae species ...... 139 Figure 8.2a: Appressorium formation on Alismataceae species ...... 140 Figure 8.2b: Rate of appressorium formation on Alismataceae species ...... 140
iv List of Tables
LIST OF TABLES Table 2.1: The recorded host range of R. alismatis...... 10 Table 3.1: Collection information for Rhynchosporium.isolates...... 17 Table 3.2: Summary information for isolates included in the ITS sequence database...... 21 Table 4.1: Origin, host, population and collection date of R. alismatis isolates...... 39 Table 4.2: Primers used during population structure analysis of R. alismatis...... 41 Table 4.3: Allele frequencies in Australian populations of R. alismatis...... 46 Table 4.4: Nei’s measures of diversity in Australian populations of R. alismatis...... 48 Table 4.5: Genotype diversity and genetic distance among populations of R. alismatis ...... 49 Table 5.1: Conidial germination and appressoria formation on Alismataceae species ...... 71 Table 6.1: Plant pathogenic fungi transformed to hygromycin B resistance...... 99 Table 6.2: Primers used and constructed during transformation of R. alismatis...... 110 Table 6.3: Details of individual transformation experiments...... 117 Table 8.1: Pathogenicity of R. alismatis isolates against Alismataceae species...... 137
v List of Abbreviations and Statistical Symbols
LIST OF ABBREVIATIONS AND STATISTICAL SYMBOLS ABC ATP-binding cassette Ampr Ampicillin resistance ANOVA Analysis of variance ARS Autonomously replicating sequence ATMT Agrobacterium tumefaciens mediated transformation BLAST Basic local alignment search tool CIA Coleambally Irrigation Area CST Central and Southern Tablelands CWDE Cell wall degrading enzymes DM Dicroic mirror DNA Deoxyribonucleic acid EDTA Ethylenediaminetetraacetic acid ERIC Enterobacterial repetitive intergenic consensus f. sp. Formae specialis G418 Geneticin GFP Green fluorescent protein GUS B–glucuronidase HCl Hydrochloric acid HmB Hygromycin B hph/HPH Hygromycin B phosphotransferase IPTG Isopropylthio–β–D–galactosidase ISSR Inter-simple sequence repeat ITS Internal transcribed spacer LB Luria–Bertani LBA Lima bean agar LM Liquid medium LSD Least significant difference M Molar MCPA 2–Methyl–4–chlorophenoxyacetic acid MFS Major facilitator superfamily
MgSO4 Magnesium sulphate MIA Murrumbidgee Irrigation Area mL Millilitre
vi List of Abbreviations and Statistical Symbols mM Millimolar MPa Megapascals MVID Murray Valley Irrigation District NaCl Sodium chloride NaOH Sodium hydroxide ng Nanogram NIP Necrosis inducing protein nm Nanometres NSW New South Wales PAS Periodic acid Schiff's reagent PCNB Pentachloronitrobenzene PCR Polymerase chain reaction PDA Potato dextrose agar PEG Polyethylene glycol PPT Phosphinothricin r2 Correlation coefficient RAPD Random amplified polymorphic DNA rDNA Ribosomal DNA REMI Restriction enzyme mediated insertion REP Repetitive extragenic palindromic RNA Ribonucleic acid rpm Revolutions per minute SDS Sodium dodecyl–sulphate SEM Scanning electron microscopy SNF1 Serine threonine protein kinase SR Southern Riverina SSC Saline sodium citrate buffer SSPE Saline sodium phosphate–EDTA buffer SSR Simple sequence repeat TAE Tris–Acetate–EDTA TD Touchdown Tm° Temperature melting TMTC Too many to count µg Microgram
vii List of Abbreviations and Statistical Symbols
µL Microliter UV Ultraviolet Xgal 5–bromo–4–chloro–3–indoyl–β–D–galactopyranoside amount of gene flow between populations average gene diversity between subgroups
average gene diversity of all subgroups
chi-square statistic degrees of freedom
frequency of ith allele in population
Frequency of ith allele in population
frequency of ith genotype in population
genetic distance genetic identity
genotypic diversity of observed sample No. of alleles at each locus No. of populations No. of distinct genotypes in population No. of genotypes observed times in sample
overall gene diversity population differentiation coefficient
population gene diversity
population genotype diversity Probability of identity of a gene from population and a gene from population
Probability of identity of two randomly chosen genes in population
Probability of identity of two randomly chosen genes in population
sample size students -test statistic total gene diversity of all groups
viii Abstract
ABSTRACT The phytopathogenic fungus Rhynchosporium alismatis infects Alisma, Sagittaria and several other genera within the Alismataceae, and is currently under investigation as a potential biocontrol agent for weedy members of this family. Recently, the prominence of two species, S. graminea and S. montevidensis, previously exotic to the rice growing regions of southern New South Wales has increased, but to date no records of the fungus on either species have been reported. Because efforts to develop commercial weed management alternatives must endeavour to provide broad spectrum weed control in order to compete financially, attempts to expand the host range of this fungus to include these species are warranted. In attempting this goal several distinct studies were performed prior to attempts to develop reliable and efficient transformation techniques for the fungus. These studies included a comprehensive phylogenetic analysis, population structure analysis and a detailed study of the infection process of the fungus on both host and non–host Alismataceae species. Sequencing of internal transcribed spacer (ITS) regions of a large number of fungal isolates revealed close relationships between R. alismatis and the teleomorph genus Plectosphaerella as well as several anamorphic fungi including species of Verticillium and Nectria. Whilst intraspecific variation in R. alismatis was minimal, isolates of the fungus clustered with Plectosphaerella cucumerina in a monophyletic group with 100% bootstrap support and were clearly not related to the forme species R. secalis nor R. orthosporum, which formed a separate group also with high bootstrap support. The genetic structures of populations of R. alismatis also displayed minimal variation with low to moderate (0.1924) levels of gene diversity across three geographically isolated populations. The average genetic distance (D = 0.0265) and overall population differentiation (Gst = 0.1001) were also small. However, measures of both genotypic and allelic diversity were significant (P = 0.100) between populations. Populations structures appeared clonal supplemented with intermittent rounds of recombination or gene flow over considerable distances. Migration via infected seed and/or plant material transported by irrigation channels and watercourses is probable. However, little evidence for gene exchange was apparent and evolution of pathogenic ability was minimal. Further investigation of the infection process of the fungus on target weeds indicated that the rates of conidial germination and appressorium formation on these species were almost identical to that of A. plantago–aquatica, a known host. Germ tube elongation and appressorium formation occurred randomly over the leaf surface with no apparent stimulus and the fungus entered the host by way of direct penetration facilitated by the production of a penetration peg.
ix Abstract
Holes left by penetration pegs, 0.25–0.5 µm in diameter, were observed by scanning electron microscopy on both A. plantago–aquatica and S. graminea following the removal appressoria. However, no penetration of S. montevidensis was witnessed and penetration sites on S. graminea were accompanied by evidence of host defense responses. In order to facilitate fungal penetration of S. montevidensis transformation with constructs designed to increase the penetrative ability of the fungus was required. Protoplasts were released readily from pre–germinated spores of R. alismatis which underwent 4–24 hours enzymatic lysis. Transformation frequencies ranged between 6 to 187 transformants per 5 µg of DNA or 1 colony per 11 000 protoplasts and of 10 individual experiments 42 putative transformants were transferred. Molecular analysis confirmed the presence of vector DNA in transformed cells, however radioactive probes failed to hybridise to the membranes during southern analysis, and positive confirmation of transformation in the fungus was not obtained.
x Chapter 1. Introduction
1 Introduction
1.1 Introduction Rice is the world’s most important food crop (McDonald 1994). Despite dramatic increases in the productivity of rice including the development of new higher yielding varieties (McDonald 1994), rapid population growth and hence the demand for rice was predicted to exceed production by the turn of the century (David 1991). Whilst improvements to existing production procedures are required to achieve the levels of production necessary to meet future demands, improvements to weed management strategies to reduce yield losses resulting from weed infestation also have the potential to contribute to production (Cother 1996). Traditional methods of weed control involved purely physical means (Cox 1984). However, the advent of chemical herbicides drastically changed our approach to and success in weed control, but not without provoking an awareness of the potential disturbance an entirely chemical response to weed control places on the ecosystem of rice paddies and the surrounding environment (Cother 1996). In response to both more stringent regulatory requirements for chemicals and a growing awareness of the wider social costs of large scale pesticide use, alternative means of weed control are required. A modern trend in weed control is the utilisation of fungi as biological control agents, and the endemic fungal pathogen Rhynchosporium alismatis (Oudem.) Davis 1922 is being studied as a potential biological control agent for Damasonium minus (R. Br.) Buchenau. (Cother & Gilbert 1994a; 1994b; Jahromi et al. 1998; 2002), considered the most significant aquatic broadleaf weed of rice in Australia (McIntyre et al. 1991). Recently, the prominence of two species, Sagittaria graminea Michx and Sagittaria montevidensis Cham & Schlecht., previously exotic to the rice growing regions of southern New South Wales, has increased, and to date no records of the fungus on either species have been reported, despite several species in this genus being recorded hosts of the fungus elsewhere (Cother & Gilbert 1994a). Because efforts to develop commercial weed management alternatives must endeavour to provide broad spectrum weed control in order to compete financially, attempts to expand the host ranges of this fungus to include these species are warranted. By widening the host range of R. alismatis the feasibility of a broad spectrum fungal bioherbicide to control Alismataceae weeds in Australian rice crops would become a more viable option.
1 Chapter 1. Introduction
1.2 Objectives of this study This study aims to investigate the factors involved in expanding the host range of R. alismatis, a candidate mycoherbicide for the broad spectrum control of Alismataceae weeds in Australian rice crops. This investigation focused on four areas of research.
1. Phylogenetic analysis of the relationships between R. alismatis and other members of the genus Rhynchosporium. 2. Population structure analysis of R. alismatis collections from different geographic origins. 3. Investigation of the infection process of R. alismatis on both host and non–host Alismataceae species. Development of transformation techniques for the insertion of pathogenicity–related genes into R. alismatis.
2 Chapter 2. Literature Review
2 Literature Review
2.1 The Australian Rice Industry Rice (Oryza sativa) is a semi aquatic, annual cereal grass produced in over 110 countries (Webster & Gunnell 1992). Second only to wheat with regard to hectares harvested for nutritional purposes, rice is one of the most versatile and important food crops in the world supplying approximately 21% of the total caloric intake of the world’s population (McDonald 1994). Successful commercial cultivation of rice commenced in Australia in the early 1920s, with attempts being made to cultivate rice in Queensland, Western Australia and on the coastal plains east of Darwin (McDonald 1979). To date, the Australian rice industry is confined to southern New South Wales where it encompasses the Murrumbidgee (MIA) and Coleambally irrigation areas (CIA) and the Murray Valley irrigation districts (MVID) (Figure 2.1).
Figure 2.1: The rice growing region of southern New South Wales, Australia. Reproduced by permission of Greg Doran.
3 Chapter 2. Literature Review
2.2 Weeds In the rice growing regions of southern New South Wales, weeds surpass all other pests (McDonald 1994), and yield losses of up to 40% are common despite the implementation of control strategies (Watson 1991). Of particular importance to the Australian rice industry is the Alismataceae, a small family of monoecious aquatic marsh herbs, containing some of the most important aquatic broadleaf weeds of rice in this country (Cox 1984). Comprising the natives Alisma plantago–aquatica L. (waterplantain), Damasonium minus (starfruit) and three introduced species, Alisma lanceolatum With. (alisma), Sagittaria montevidensis (arrowhead) and Sagittaria graminea, the importance of the Alismataceae has been enhanced greatly by the introduction of aerial sowing, irrigation development and the emergence of herbicide resistance, such that all five species are now endemic to southern New South Wales, where they range from minor members of the rice weed flora to serious weeds (McIntyre & Newnham 1988). Presently, A. lanceolatum and D. minus are among the most important aquatic broadleaf weeds of rice in Australia with D. minus being present at more than 70% of locations throughout the rice growing districts of southern NSW (McIntyre et al. 1991). A third species, A. plantago–aquatica, is also increasing in importance, but having been discovered only recently in this region is not yet considered a significant weed (Sainty & Jacobs 1994). By contrast, the recent incursion of S. graminea and S. montevidensis, into rice growing regions has occurred at such a rate that these species now assume greater importance than D. minus at some locations. The most aggressive of these species, S. montevidensis, originally discovered in the MVID in 1988, has spread slowly southwards and is now well established in the CIA where it occurs sporadically in drainage channels and increasingly throughout rice crops (McIntyre & Newnham 1988). The second species, S. graminea, which is not currently considered a significant weed of rice in Australia or elsewhere in the world, is however regarded as a troublesome weed in northern Victoria where it obstructs water flow in irrigation channels (McIntyre & Newnham 1988). The distribution of S. graminea is expanding slowly northwards and is increasingly prominent in drainage channels throughout the Murrey Valley. A further species, Caldesia oligococca (F. Muell.) Buchenau., is also common in lagoons, billabongs and irrigation channels throughout tropical Queensland, Western Australia and the Northern Territory (Jacobs 2003), but thus far has not been cited in the Riverina. The appearance and distribution of the aforementioned species are shown in Figure 2.2.
4 Chapter 2. Literature Review
Figure 2.2: The Alismataceae family. Top row, A. lanceolatum, A. plantago–aquatica and S. montevidensis. Bottom row, S. graminea, D. minus and C. oligococca. Insert, distribution of each species in Australia (Individual photographs reproduced from Sainty and Jacobs (1994)). Data for weed distributions compiled from Aston (1973), Sainty and Jacobs (1981) and Jacobs (1993).
2.3 Weed Management in Rice During the last two decades weed growth and competition in rice has increased (Hill et al. 1994). The combined effects of new higher yielding semi–dwarf rice varieties with their short statured growth habit and the widespread use of aerial sowing in southern New South Wales has reduced the competitive ability of the rice (Hedditch 1984) and aquatic weeds now assume much greater importance.
5 Chapter 2. Literature Review
Under these cultural conditions high rates, and often multiple applications of herbicides have been necessary to maximise yield potential. Unfortunately, while the use of MCPA (Nott et al. 1974), and the release of Londax® (bensulfuron methyl) in the mid 1980s dramatically increased the effectiveness of aquatic weed control, the continued use of site specific herbicides led to the development of herbicide resistance in several aquatic weed species including Cyperus difformis, S. montevidensis and D. minus. (Fowler & McCaffery 1994). Combined with an increasing awareness of the hazards posed by the use of chemical pesticides, social and environmental concerns have slowed the development and registration of new herbicides for rice–based cropping (Hill et al. 1994). With fewer herbicides and a cultural system prone to losses from weeds, integrated weed management strategies encompassing alternative forms of weed control are clearly required One strategy for integrated weed management in rice which has gained momentum abroad, is inundative biological control using fungal pathogens. This process involves the application of infective propagules of an indigenous weed pathogen in a manner analogues to the application of chemical herbicides (Templeton 1982; TeBeest & Templeton 1985) and has currently produced favourable results for the potential control of water chestnut (Eleocharis kuroguwai) in South–east Asia (Tanaka et al. 1992), the perennial sedge Scirpus planiculmis in Korea (Kim 1992), barnyard grass (Echinochloa crus–galli) in Japan (Gohbara & Yamaguchi 1992), gooseweed (Sphenoclea zeylanica) in the Philippines (Watson 1994) and for the control of northern jointvetch (Aeschnomene virginica) in the United States (Bowers 1986). In this latter example, exploitation of this technology resulted in the registration of the mycoherbicide, Collego, which consists of a dry formulation of Colletotrichum gloeosporioides (Penz.) Sacc f. sp. aeschynomene applied aerially to rice fields once per season following the emergence of weed above the rice canopy (Templeton 1987). It has a success rate greater than 90% and a shelf life in excess of 12 months (Templeton 1985). In Australia, the endemic fungal pathogen R. alismatis is currently under investigation for the control of Alismataceae weeds in rice (Jahromi et al. 1998; 2002).
2.4 Development of the mycoherbicide In 1994, Cother and Gilbert (1994b) reported the effects of a naturally occurring fungus, R. alismatis, on the growth of A. lanceolatum and demonstrated its pathogenicity to, and growth suppression of, five other species in the Alismataceae, including A. plantago–aquatica, D. minus and three species of Sagittaria obtained from overseas (Cother & Gilbert 1994a).
6 Chapter 2. Literature Review
In field applications using conidial suspensions of the fungus, significant reductions in total plant leaf number, root dry weight, inflorescence development and biomass production were reported in Alisma and a number of other Alismataceae species recognised for their importance in rice cropping overseas. In one trial, inoculation of Sagittaria guyanensis H.B.K., an important rice weed in Malaysia, but also found throughout Central and South America and parts of Africa, resulted in a 60% reduction in plant height, and reduced leaf and root dry weights by 89% and 84% respectively (Cother & Gilbert 1994a). Spray–inoculations of seedlings resulted in a 62–87% reduction in plant biomass production per plot (Cother 1996), and equally significant reductions in various growth parameters were reported for Alisma canaliculatum A. Br. & Bouche, Echinodorus rostratus (Nutt.) Englem., Sagittaria brevirostra Mack & Bush and S. pygmaea Miq.(Cother & Gilbert 1994a). Discovered in Australia in 1941 causing leaf spots on A. plantago–aquatica, the hyphomycete R. alismatis, is a fungal pathogen endemic to Australia, which causes necrotic lesions on the leaves, petioles and inflorescence stalks of many species in the Alismataceae (Figure 5.3). Symptoms begin as small lens shaped dark brown necrotic spots with pale green to yellow halos (chlorosis), which eventually increase in diameter and coalesce to form larger elongated lesions, becoming visible on mature leaves within three to four days of infection (Cother et al. 1994). The taxon, R. alismatis, by which the fungus is commonly referred, was originally described as Septoria alismatis by Oudemans in 1875 (Punithalingam 1988). However, examination of several collections including type material failed to reveal a pycnidial fungus conforming to the description of Oudemans and eventually the combination R. alismatis was adopted with the basionym Septoria alismatis Oudem. 1875 and synonyms Ascochyta alismatis Ell. & Ev. 1889, Ramularia alismatis Fautrey 1890, and Didymaria aquatica Starbäck 1895 (Punithalingam 1988). Despite being reclassified to Spermosporina Braun in 1993 (Braun 1993), the currently recognised name for this hyphomycete is R. alismatis, and this is the name under which the only known cultures of the fungus are stored at the Agriculture and Scientific Collections Unit, New South Wales Agriculture, Orange, NSW, Australia. The fungus is characterised by ‘mycelium subcuticular (as well as ramifying the leaf tissue and becoming inter- and intra-cellular), composed of branched, septate hyphae forming stromata which are composed of short, broad hyphal cells from which conidia are produced from short conidiogenous cells. Conidiophores absent. Conidiogenous cells hyaline, acrogenous, percurrently proliferating (annellidic). Conidia holoblastic, hyaline, straight or slightly curbed towards the ends or slightly beaked, medianly 1–septate 16–19 (–20) x 2.5–3 (–3.5) µm, guttulate’.
7 Chapter 2. Literature Review
Cultures of the fungus are slow growing, compact, initially white but later pinkish/orange, sporulating after 3 days (Cother et al. 1994). The shape, size of conidia and appearance of culture of potato dextrose agar are shown in Figure 2.3.
Figure 2.3: (a) The typical white to pinkish–orange appearance of mycelium of R. alismatis on potato dextrose agar (PDA), and (b) size and shape of fungal propagules of R. alismatis (Source: Punithalingam 1988).
8 Chapter 2. Literature Review
Traditionally R. alismatis has been isolated solely from species of the Alismataceae. However, recent host range studies, which included 28 species of aquatic plants in the Alismataceae and 39 cultivars of 25 agricultural important species, identified a number of additional hosts from a range of plant families including the Aponogetonaceae, Hydrocharitaceae, Juncaginaceae and Marsileaceae (Cother 1999). In these studies R. alismatis caused typical lesions on several aquatic species including Aponogeton, Triglochin, Marsilea, and the broadleaf form of Vallisneria, which developed bleached light brown lesions from which the fungus could be reisolated. In both species of Triglochin, pepper spotting developed at the point of inoculation, whilst symptoms on Marsilea and Aponogeton ranged from non–specific browning of tissue to necrotic and chlorotic lesions. However, unlike Vallisneria, the fungus was not re–isolated from diseased lesions from any of these species (Cother 1999). In addition to several aquatic species identified as additional hosts of the fungus, scattered infrequent lesions were also observed on the leaves of barley, oats, triticale, lupin, soybean, lettuce and tomato. Whilst the fungus was reisolated only from lesions on bowyer soybean cultivars (Fabaceae), the random plating of leaf pieces from these and other species resulted in re–isolation of the fungus from additional species within the Cucurbitaceae, Gramineae and Solanaceae. Based on frequency of re–isolation, cucurbits and tomato were the most susceptible to infection despite being asymptomatic in appearance, with the highest rate of infection occurring on the Salad Bush cucumber cultivar. In addition to several other cucurbits including rockmelon, pumpkin, zucchini and squash, the only other species from which the fungus was re–isolated following inoculation were the Muir and Tahara triticale cultivars, however, this was demonstrated only at low frequency (Cother 1999). The recorded host range of R. alismatis is shown in Table 2.1. Whilst the identification of additional hosts refutes the previous suggestion that R. alismatis is confined to species of the Alismataceae, the testing of 28 species in the Alismataceae along with 39 cultivars of 25 species of agriculturally important plants suggests the fungus maintains a relatively narrow host range. Whilst this host specificity is often deemed an important characteristic of mycoherbicides, under some circumstances the ability to attack multiple target species may be an advantage (McRae 1988). However, it is clear from the studies of Cother (1999) that R. alismatis does not infect the two species of Sagittaria which are now considered prominent weeds of rice not only in Australia, but elsewhere in the world (Itoh & Miyahara 1988).
9 Chapter 2. Literature Review
Table 2.1: The recorded host range of R. alismatis. Re–isolation of Host Plant Reaction R. alismatis Alismataceae Alisma canaliculatum A. Br. & Bouche +A +B Alisma lanceolatum With. + + Alisma plantago–aquatica L. + + Alisma plantago–aquatica var. orientale nt Damasonium minus (R. Br.) Buchenau + + Echinodorus rostratus (Nutt.) Englem. ± + Sagittaria brevirostra Mack & Bush ± + Sagittaria guyanensis H.B.K. + + Sagittaria graminea Michx. – – Sagittaria heterophylla Bert. Ex Steud. + + Sagittaria natans Pallar + – Sagittaria pygmaea Miq. ± + Sagittaria rigida Pursh nt Sagittaria subcordatum nt Sagittaria subulata var subulata (L.) Buchenau + + Aponogetonaceae Aponogeton distachyos L. + – Cucurbitaceae Cucumis melo L. (rockmelon) – + Cucumis pepo L. (pumpkin) – + Cucumis pepo L. subsp. pepo (zucchini) – + Cucumis sativa L. (cucumber) – + Cucurbita sp. (squash) – + Fabaceae Glycine max L. (soybeans) Bowyer cult. – + Gramineae x Triticosecale Wittm. Ex A. Camus. (triticale) Muir – + x Triticosecale Wittm. Ex A. Camus. (triticale) Tahara – + Hydrocharitaceae Vallisneria sp. (broad leaves) + + Juncaginaceae Triglochin dubia R. Br. (+) – Triglochin multifractum Aston + – Marsileaceae Marsilea drummondii A. Braun + – Marsilea mutica Mett. + – Solanaceae Lycopersicum esculentum Mill. (tomato) – + Data compiled from Cother et al. (1994); Cother and Gilbert (1994a; 1994b); Cother (1999). +A = Necrosis or chlorosis at point of inoculation. +B = The fungus was re–isolated from inoculated tissue. ± = Reduction in plant growth/development, no lesion. ( ) = Pepper spotting on control. nt =. Not tested by Cother and colleagues. Data obtained from Punithalingam (1988), Farr et al. (1989) and Tai (1979).
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Nevertheless, R. alismatis does possess many other qualities that are deemed necessary of the active ingredient of a mycoherbicide against Alismataceae weeds. These include the ability to be cultured both in solid and liquid fermentation, the production of large quantities of infectious propagules (Jahromi et al. 1998; Cother & Van de Ven 1999), high levels of pathogenicity towards target plants (Jahromi et al. 2002) and poor dispersal mechanisms (Fox et al. 1999). However, because efforts to develop commercial weed management alternatives must attempt to provide broad spectrum weed control in order to compete financially, and the development of a broad spectrum alternative to chemical herbicides is warranted, the host range of this fungus requires expansion. By expanding the host range of R. alismatis to include the aforementioned species of Sagittaria, the feasibility of a fungal bioherbicide to control Alismataceae weeds in Australian rice crops would become a more viable option.
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3 Phylogenetics
A phylogenetic analysis of the genus Rhynchosporium – addressing the position of R. alismatis within the genus and evidence for possible reclassification
3.1 Introduction During the past 65 years, the position of R. alismatis within the genus Rhynchosporium has been debated (Caldwell 1937). Because fungi have traditionally been classified based on a particular niche, or in plant pathology, defined mainly with respect to host affiliation, the phylogeny of fungi has been inferred largely from various phenotypic characters and/or biochemical responses which mostly relate to functional or structural attributes. Yet despite the morphological characteristics which resulted in the exclusion of R. alismatis from the genus Rhynchosporium during earlier studies (Caldwell 1937) and more recent classifications by Braun (1995) which relegate the organism to the genus Spermosporina, recent publications continue to refer to this organism as a species of the genus Rhynchosporium (Cother & Van de Ven 1999; Lanoiselet et al. 2001). The introduction of molecular techniques coinciding with the advent of the polymerase chain reaction (PCR) (Saiki et al. 1988) improved drastically the study of taxonomic and phylogenetic relationships in fungi. In one of the first applications of PCR in mycology, White et al. (1990) amplified and sequenced ribosomal DNA (rDNA) regions from fungi, establishing a technique which has enabled researchers to study the relationships within (Kistler et al. 1991; Edel et al. 1995; Zhang, Hartman et al. 1997), and between, a diverse range of species (Lobuglio et al. 1993; Sreenivasaprasad et al. 1996; Goodwin & Zismann 2001), based purely on the analysis of nucleic acids. To date, however, these techniques have not yet been applied to studies in the genus Rhynchosporium, and neither Caldwell’s theory nor the placement of R. alismatis within this genus have been confirmed. In this chapter, the phylogeny of the genus Rhynchosporium will be investigated. To begin with, the taxonomy and nomenclature surrounding the genus Rhynchosporium and the methods pertinent to the study of fungal phylogenies will be presented. The phylogenetic relationships of Rhynchosporium species and closely related organisms will then be discussed. Finally, the position of the organism within the current genus and evidence for reclassification will be provided. 12 Chapter 3. Phylogenetics
3.2 Taxonomy and Nomenclature The description of the genus Rhynchosporium Heinsen has been modified several times since its creation by Frank in 1897. Although the author credited the naming of the genus to his associate, E. Heinsen, it was some years before Heinsen (1901) published this work. Unfortunately, neither Frank nor Heinsen provided formal descriptions of the genus or species, and many more years passed before these were provided by Saccardo (1906) and Lindau (1910) (references cited by Caldwell 1937). Historically, the taxonomic position of the genus Rhynchosporium is controversial since the genus possesses characteristics common to both the orders Moniliales and Melanconiales, and was further revised in 1922 to include all members of the Mucedinaceae (Davis 1922). Whilst the resemblance to the later order was shown to be erroneous by Brooks (1928), morphological studies of fructification structures showed a resemblance to the order Melanconiales. Whilst this resemblance was later dismissed by Caldwell (1937), who found little evidence to suggest that Rhynchosporium was closely related to genera in this order, the morphology of fructifying structures on the host was deemed an important distinguishing feature of the genus, placing it securely in the order Moniliaceae, which contains many genera with similar fructifying structures. At present, the genus Rhynchosporium comprises only two legitimate species, the barley scald pathogen R. secalis (Oudem.) Davis, and a second species R. orthosporum which produces disease symptoms identical to those caused by R. secalis, but on an alternate host, Orchardgrass (Dactylis glomerate L.) (Caldwell 1937). A third combination R. alismatis (Oudem.) originally proposed by Davis in 1922, and placed in synonymy with Septoris alismatis (Oudem.), Aschochyta alismatis Ell and Ev., Ramularia alismatis Fautrey, and Didymaria aquatica Starback, is also occasionally placed in this genus (Punithalingam 1988). However, because this species lacked the superficial fertile stroma of other members of the genus and bore conidia on short flask shaped conidiophores, Caldwell (1937) excluded this species both from the genus Rhynchosporium and the division Micronemeae. To date, only one study has been published on the phylogenetic relationships of species in the genus Rhynchosporium (Goodwin 2002). Whilst this study showed that the genus Rhynchosporium was monophyletic with regards to the recognised species, it did not include isolates of R. alismatis, and as a result the phylogenetic relationships between this species and the other members of the genus have not been established.
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3.3 Concepts and Methodology The polymerase chain reaction (PCR) is a powerful method in molecular biology, the applications of which are extensive in many fields of mycology including plant pathology, fungal genetics and systematics (Edel 1998). Whilst the major PCR–based techniques considered for species level definition in fungi include those developed from random amplified polymorphic DNA (RAPD) and other fingerprinting techniques (Bridge & Arora 1998), DNA sequence analysis has become the most popular method for inferring phylogenetic relationships (Takamatsu 1998). According to Bruns et al. (1992), the utility of sequence analysis lies in the ‘the large number of characters which can be compared’. However, determination of the mode of variation, whether it be transition or transversion, and confirmation of results through the deposition of sequences to online databases such as GenBank, EMBL and DDBJ, are also considerable advantages. Although several DNA regions have proven useful for phylogenetic studies, much of the activity in fungal systematics has been derived from the study of the DNA sequences which encode genes within the ribosomal RNA gene cluster (Takamatsu 1998). Found universally in all living cells, where it is thought that their evolution may reflect the evolution of the whole genome (Edel 1998), the ribosomal RNA gene cluster contains both conserved and divergent regions (Robb et al. 1993). Whilst the conserved or coding regions evolve relatively slowly, the interspersed non−coding regions known as internal transcribed spacers (ITS) can display considerable variation (Chen 1992; O’Donnell 1992). Although intraspecific variability within the ITS region is rare (Lee & Taylor 1992), variation is seldom uniform across species boundaries and can be sufficient to allow clear discrimination between related species and genera (Lanfranco et al. 1998). To date, PCR–derived phylogenetic studies based on the analysis of ribosomal RNA genes have been used to investigate the evolutionary relationships within a diverse range of fungi from many different orders, including the filamentous ascomycetes (Appel & Gordon 1995; 1996; Morales et al. 1995), basidiomycetes (Hibbett et al. 1995; Zambino & Szabo 1993), zygomycetes (Nagahama et al. 1995; Gehrig et al. 1996) and honorary oomycetic fungi such as Pythium (Briard et al. 1995) and Phytophthora (Cooke et al. 1996; Crawford et al. 1996). Whilst examples of the many organisms which make up these diverse groups cannot be summarised here, Takamatsu’s (1998) coverage of this topic in the recently published ‘Applications of PCR in Mycology’ provides a comprehensive summary of the contribution that PCR–based methods have made to the study of fungal phylogenetics.
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The shear volume of examples in this treatise is privy to the sensitivity and reliability of a technique that has contributed considerably to the progress of this field and proven superior to many other conventional methods (White et al. 1990). Because these regions also comprise the largest components of existing databases, they subsequently also provide the greatest likelihood of identifying closely related species. In view of the controversial classification of R. alismatis, investigation of the phylogenetic relationships between species in the genus Rhynchosporium is warranted. The phylogeny of a genus provides information on the taxonomic status and nomenclature of the organism, is essential for the interpretation of the literature, provides information on the relationships between the agent and pathovars that are known to cause disease on other species of agricultural importance (Weidemann & Te Beest 1990), and satisfies some of the requirements imposed by regulatory agencies prior to the release of an organism (Berthier et al. 1996). By reconstructing phylogenies between species currently assigned to the genus Rhynchosporium and other related organisms the true relationships between these species can be more adequately defined.
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3.4 Materials and Methods 3.4.1 Fungal isolate collection, isolation, cultivation, storage and maintenance: During the period January 1989 to March 1997, leaves from Alisma lanceolatum, A. plantago– aquatica and Damasonium minus infected with R. alismatis were collected from a number of sites throughout New South Wales by Dr E.J. Cother. As described previously by Cother et al. (1994), the margins of necrotic lesions were excised, surface sterilised in sodium hypochlorite solution (1% available chlorine) for 45 seconds, rinsed in sterile distilled water, blotted dry with sterile filter paper and transferred to 9 cm diameter plastic Petri dishes containing quarter–strength potato dextrose agar (PDA; Amyl Media Pty. Ltd., Dandenong, Victoria, Australia) supplemented with 10 mg/L rifampicin (Sigma−Aldrich, Castle Hill, NSW, Australia), 250 mg/L ampicillin (Sigma−Aldrich) and 100 mg/L pentachloronitrobenzene (PCNB; Sigma−Aldrich). Plates were incubated at 25°C in the dark for 3–5 days and, following the emergence of fungal growth, isolates were transferred to PDA, incubated at 25°C for a further 5–10 days and stored at 4°C. Additionally, cultures were lyophilised for long term storage. With the exception of isolate RH057 which was obtained from the leaves of S. montevidensis, artificially inoculated with spores of RH001 in a pot experiment conducted at Yanco in 1991, all leaf samples were collected from naturally infected hosts and conveyed to the laboratory under refrigeration. In addition to the large number of isolates collected from Australian sites, several fungal isolates were also obtained from other sources including several species of Alismataceae found in Malaysia and Japan. One isolate designated RH126, was collected from infected leaves of Potamogeton spp. obtained from a Sydney nursery, two isolates, RH147 and RH148, were collected from infections occurring in the glasshouse at Wagga Wagga and a single isolate of the scald pathogen R. secalis was provided by Dr. H. Raman at NSW Agriculture, Wagga Wagga, NSW, Australia. The origin, host and isolation date of all Rhynchosporium isolates collected for this study are shown in Table 3.1. With the exception of isolates RH047, RH055, RH080, RH126, RH136, RH137, RH140, RH141, RH147, RH148, RH149 and W12868B all cultures are lodged with the Agricultural Scientific Collections Unit at NSW Agriculture, Orange, NSW, Australia (Herb. DAR). Upon receipt of cultures from the herbarium, all isolates were passaged through their respective host plants, single spored and stored at 4°C on supplemented PDA. Where host plants were unavailable, isolates were single spored only. To maintain viability, cultures were transferred monthly. Spore suspensions of each culture were also stored for the duration of the study at −80°C by inoculating Protect tubes (Technical Service Consultants Limited, Heywood, Lancashire, UK) with spore suspensions prepared as described in section 3.4.2.
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Table 3.1: Collection information for Rhynchosporium isolates used in the phylogeny study. Isolate Herb. DAR Host Origin Collection accession number date Rhynchosporium alismatis RH001 DAR67515 A. lanceolatum Coleambally Jan. 89 RH005 DAR73786 A. lanceolatum Pericoota Dec. 89 RH021 DAR67510 A. plantago–aquatica Narranderra Jan. 91 RH024 DAR67517 A. lanceolatum Coleambally Jan. 91 RH025 DAR73158 A. lanceolatum Coleambally Jan. 91 RH037 DAR73787 A. lanceolatum Gambles Lane Jan. 91 RH038 DAR73788 A. lanceolatum Gambles Lane Jan. 91 RH039 DAR73789 A. lanceolatum Gambles Lane Jan. 91 RH041 DAR73790 A. lanceolatum Pericoota Jan. 91 RH046 DAR67511 D. minus Yanco Jan. 91 RH047 DAR76105 D. minus Yanco Jan. 91 RH054 DAR73791 A. lanceolatum Hanwood May 91 RH055 DAR76104 D. minus Yanco May 91 RH057 DAR67516 S. montevidensis Yanco May 91 RH058 DAR67508 A. plantago–aquatica Hobbys Yards Jan. 92 RH062 DAR67513 A. plantago–aquatica Khancoban Jan. 92 RH064 DAR73146 A. plantago–aquatica Rosewood Jan. 92 RH066 DAR67512 A. plantago–aquatica Tumut Jan. 92 RH069 DAR73148 A. plantago–aquatica Petfield Jan. 92 RH074 DAR73149 A. plantago–aquatica Rockley Jan. 92 RH080 DAR76092 S. pygmaea Japan Feb. 93 RH091 DAR73673 A. lanceolatum Hanwood May 91 RH095 DAR73150 A. plantago–aquatica Petfield Jan. 92 RH097 DAR73151 A. plantago–aquatica Rockley Jan. 92 RH100 DAR73860 D. minus Yanco May 91 RH108 DAR73861 D. minus Yallakool Feb. 93 RH111 DAR73793 A. lanceolatum Caldwell Feb. 93 RH118 DAR73672 A. lanceolatum Pericoota Dec. 89 RH121 DAR73794 A. lanceolatum Coleambally Jan. 91 RH122 DAR73795 A. lanceolatum Coleambally Jan. 91 RH123 DAR73796 A. lanceolatum Coleambally Jan. 91 RH124 DAR73797 A. lanceolatum Coleambally Jan. 91 RH126 DAR76095 Potamogeton sp. Sydney Oct. 94 RH127 DAR73798 A. lanceolatum Coleambally Jan. 91 RH133 DAR73799 A. plantago–aquatica Deniliquin Jan. 95 RH135 DAR73800 A. plantago–aquatica Deniliquin Jan. 95 RH136 DAR76107 D. minus Derrulaman Jan. 95 RH137 DAR76106 D. minus Derrulaman Jan. 95 RH138 DAR73862 D. minus Derrulaman Jan. 95 RH139 DAR73145 D. minus Derrulaman Jan. 95 RH140 DAR76093 S. guyanensis Malaysia Nov. 95 RH141 DAR76094 S. guyanensis Malaysia Nov. 95 RH143 DAR73152 D. minus Barmah Mar. 97 RH144 DAR73153 D. minus Barmah Mar. 97 RH145 DAR73154 D. minus Barmah Mar. 97 RH147 DAR76283 S. graminea Wagga Wagga May 99 RH148 DAR76284 A. plantago–aquatica Wagga Wagga May 99 RH149 DAR76285 D. minus Coleambally Jan. 91 Rhynchosporium secalis W12868B – Hordeum vulgare Wagga Wagga Jun. 98
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3.4.2 Culture preparation and DNA extraction: For DNA extraction, spore preparations of each isolate were prepared by transferring cultures to lima bean agar (LBA; Difco Laboratories, Detroit, Michigan, USA). After incubation at 25°C for 5–10 days under 12 hour light provided by two 40–W, 120cm long cool white fluorescent tubes placed 30 cm above the plates, spores were harvested by irrigating the cultures with 1 mL of sterile distilled water and scraping gently across the mycelium with a sterile glass rod. The resulting slurry of spores and mycelium was used to inoculate 250 mL conical flasks containing 150 mL of clarified 20% v/v V8 juice (Campbell’s Soup Company, Lemnos, Victoria, Australia), which were then incubated on a shaker at 25°C for up to 14 days. When sufficient growth had occurred, mycelium was harvested by filtration, lyophilised and ground in liquid nitrogen. A 2.0 mL microtube was filled to the top of the conical portion with ground, lyophilised mycelium and 800µL of lysis buffer (1% sodium dodecyl sulphate, 10mM Tris−HCl [pH 8.0], 0.5 M NaCl and 10mM EDTA) were added. The extraction was then performed according to the method of Sambrook et al. (1989) with the following modifications; the phenol: chloroform: isoamylalcohol (25:24:1) and phenol: chloroform (1:1) extraction steps were conducted in duplicate to maximise the yield of DNA, and the sample was treated for 2 hours with RNase (10 µg/mL) at 37°C, followed by the addition of 0.1 volumes of 3 M sodium acetate, prior to precipitation with isopropanol. The pellet was allowed to resuspend in 100 µL of TE buffer (10 mM Tris−HCl [pH 8.0] and 0.1 mM EDTA) at -20°C. DNA was visualized by electrophoresis on 1% agarose gel containing 1 × TAE (Tris acetate−EDTA [Sambrook et al. 1989]), stained with 0.3 µg/mL ethidium bromide and photographed over a 302 nm UV transilluminator (Ultra Violet Products Incorporated, Upland, California, USA) with a CompuScope CCD 1600 integrating digital camera (CompuScope Precision Instruments and Software, Santa Barbara, California, USA). DNA was quantified by the inclusion of an 800 ng/µL Lambda DNA quantity marker (Advanced Biotechnologies Ltd., Epsom, Surrey, UK) and calculated using Image−Pro Plus for Windows version 1.3 (Media Cybernetics, Silver Spring, Maryland, USA). DNA was adjusted to 10 ng/µL via the addition of TE buffer and stored at -20°C until required.
3.4.3 Amplifying the ITS region (ITS−PCR): The complete ITS region (ITS1, 5.8S rRNA gene, ITS2) of each isolate was amplified with the universal ITS oligonucleotide primers AB28 (GCG GAT CCA TAT GCT TAA GTT CAG CGG GT; Tm 58°C) and TW81 (GCG GAT CCG TTT CCG TAG GTG AAC CTG C; Tm 61°C) as described by Howlett et al. (1992). Primers are specific for the area between the 3’ end of the 18S rDNA gene and the 5’ end of the 28S rDNA gene. 18 Chapter 3. Phylogenetics
The primers were synthesised by Gibco BRL Life Technologies Pty. Ltd. (Mount Waverly, Victoria, Australia). PCR reactions were performed in a total volume of 50 µl by mixing 35 ng of purified genomic DNA with 100 pmol each of primers AB28 and TW81, 0.1 mM each of dATP, dCTP, dGTP and dTTP (Amersham Pharmacia Biotech, Castle Hill, NSW, Australia), 2.0 mM
MgCl2 and 2 units of Taq DNA polymerase (Promega Corporation, Annandale, NSW, Australia) in PCR reaction buffer (50 mM Tris−HCl [pH 8.0], 100 mM NaCl, 0.1 mM EDTA, 1 mM DTT, 50% glycerol and 1% Triton X−100). The reaction was performed with an initial denaturation of 5 min at 95°C followed by 35 cycles of denaturation (45 seconds at 94°C), annealing (30 seconds at 55°C) and extension (90 seconds at 72°C), and a final extension (5 minutes at 72°C). PCR was performed in a Corbett Research FTS 960 DNA thermal cycler (Corbett Research, Mortlake, NSW, Australia). The reaction mixture was supplemented with 0.4 volumes of bromophenol blue loading buffer (0.25% w/v bromophenol blue [Sigma−Aldrich]; 40% w/v sucrose) and 35 µl aliquots of each PCR product were analysed by electrophoresis as described in section 3.4.2. The inclusion of 0.5 µg of each of two molecular weight markers, pUC19/HpaII and SPP1/EcoRI (Bresatec, Thebarton, SA, Australia) with lower and higher registers respectively, enabled fragment sizes to be calculated accurately. All PCR amplifications were performed in duplicate to confirm that results were reproducible.
3.4.4 Sequencing the ITS region: Following electrophoresis, ITS products of all 49 isolates were excised in duplicate from agarose gel using a sterile scalpel blade. Products were purified using the Wizard PCR Preps DNA Purification System according to the manufacturers recommendations (Promega Corporation), and following the addition of 0.1 volumes of 3 M sodium acetate, DNA re−extracted with two volumes of 100% ethanol. DNA was quantified visually against a 1 µg Lambda DNA/Hind III marker (Promega Corporation), and the concentration adjusted to approximately 20 ng/µl by resuspension in the appropriate volume of TE buffer. ITS primers AB28 and TW81 were also adjusted to the appropriate concentration for subsequent sequencing reactions (approximately 50 pmol in 5µl per reaction). Sequencing of the purified PCR products was performed by Newcastle DNA Express Sequencing, Newcastle, NSW, Australia. PCR products were sequenced using double−stranded DNA with both the forward (AB28) and reverse (TW81) primers in an ABI PRISM 377 DNA thermocycle sequencer (Perkin Elmer Applied Biosystems Inc., Sydney, Australia) using fluorescent dideoxy (dye−labelled)−nucleoside triphosphate terminators. Sequences mismatches were corrected by eye using the ABI PRISM electropherograms in conjunction with the computer package Chromas version 1.56 (Technelysium Pty. Ltd., Helensvale, QLD, Australia). 19 Chapter 3. Phylogenetics
3.4.5 Assembling the ITS database: To identify species that may be closely related to R. alismatis, a BLAST (Altschul et al. 1997) search was performed on the ITS sequence from R. alismatis isolate RH01. Sequences from 100 accessions with high similarity to the R. alismatis sequence were downloaded from GenBank and used to compile an ITS database. In addition to the sequences downloaded from GenBank following a BLAST search, sequences for several isolates of R. secalis, the related species R. orthosporum and four species from the Arthrodermataceae family were also downloaded. The latter group of organisms, which was shown to cluster as a single monophyletic group outside the group containing the Rhynchosporium isolates in a previous study (Goodwin 2002) were included as an outgroup. Multiple sequences of the same species were removed for the final analysis unless their sequences differed or they were listed as separate species in the database. The final database contained sequences of 74 isolates representing 39 species and varieties in 10 anamorphic and 5 teleomorphic genera (Table 3.2).
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Table 3.2: Summary information for isolates included in the ITS sequence databasea. GenBank Species Isolate Family Order accession no. Arthroderma benhamiaeb UAMH7339 Arthrodermataceae Onygenales AF170467 A. ciferriic CBS272.66 Arthrodermataceae Onygenales AJ007844 A. incurvatumd CBS161.69 Arthrodermataceae Onygenales AF168129 Chaetosphaeria innumera MR1175 Chaetosphaeriaceae Sordariales AF178551 Cordyceps jezoensis – Clavicipitaceae Hypocreales AB027365 Cylindrocarpon spp. l 4/97–1 Nectriaceae Hypocreales AJ279490 Cylindrocarpon spp. l 5/97–12 Nectriaceae Hypocreales AJ279482 Fusarium solani f. sp. piperis NRRL22570 Nectriaceae Hypocreales AF178422 Hypocrea aureoviridis CCRC33594 Hypocreaceae Hypocreales AF414326 H. aureoviridis IMI355906 Hypocreaceae Hypocreales AF194016 H. aureoviridis IMI311745 Hypocreaceae Hypocreales AF194018 H. aureoviridis CBS138.79 Hypocreaceae Hypocreales AF194005 H. aureoviridis CBS245.63 Hypocreaceae Hypocreales Z48819 H. cf. koningii GJS96–30 Hypocreaceae Hypocreales Z95924 H. lactea CCRC33590 Hypocreaceae Hypocreales AF414325 H. muroiana GJS90–93 Hypocreaceae Hypocreales Z95927 H. muroiana GJS90–108 Hypocreaceae Hypocreales Z95928 H. nigrovirens GJS99–64 Hypocreaceae Hypocreales AF275335 H. rufa GJS90–125 Hypocreaceae Hypocreales AJ230677 H. schweinitzii ICMP1694 Hypocreaceae Hypocreales X93967 H. strictipilise CBS347.93 Hypocreaceae Hypocreales AF400263 Nectria inventaf CBS112.16 Nectriaceae Hypocreales AF324878 N. ventricosag NRRL20846 Nectriaceae Hypocreales L36657 Neonectria radicicola var. coprosmaeh GJS85–182 Nectriaceae Hypocreales AF220971 N. radicicola var. macroconidialisi GJS83–162 Nectriaceae Hypocreales AF220972 N. radicicola var. radicicolaj AR2553 Nectriaceae Hypocreales AF220968 N. radicicola var. radicicola CTR71–322 Nectriaceae Hypocreales AF220969 Plectosphaerella cucumerinak NRRL20430 Phyllachoraceae Phyllachorales AF176952 P. cucumerina 00017 Phyllachoraceae Phyllachorales AJ246154 P. cucumerina 380408 Phyllachoraceae Phyllachorales AJ492873 P. cucumerina – Phyllachoraceae Phyllachorales L36640 Rhynchosporium alismatis RH01 – – AY258151 R. alismatis RH62 – – AY258150 R. alismatis RH126 – – AY258149 R. orthosporum CBS698.79 – – AY140669 R. secalis 763 – – AF384677 R. secalis NKT12 – – AF384682 R. secalis RS020 – – AY140668 R. secalis W12868B – – AY247261 Tolypocladium parasiticum – Clavicipitaceae Hypocreales U19039 T. parasiticum CRCC32863 Clavicipitaceae Hypocreales Z54112 Trichoderma spp.l Cornell – Hypocreales AF400267 T. fasciculatum CCRC33565 – Hypocreales AF414311 T. fasciculatum CCRC33595 – Hypocreales AF414327 T. konilangbra IAA1 – Hypocreales AF400261 T. cf. harzianum CBS435.95 – Hypocreales AF400264 T. viride GJS90–20 – Hypocreales AJ230676 Trichophyton rubrum ATCC28188 Arthrodermataceae Onygenales AF170472 Verticillium albo–atrum UAMH5393 – Hypocreales AF108476 V. albo–atrum KRS1 – Hypocreales AF364008 V. albo–atrum 166 – Hypocreales AF364016 V. albo–atrum – – Hypocreales L19499 V. albo–atrum ATCC44943 – Hypocreales X60705 21 Chapter 3. Phylogenetics
Table 3.2: (cont.) GenBank Species Isolate Family Order accession no. V. albo–atrum 1776 – Hypocreales Z29509 V. albo–atrum 2 – Hypocreales Z29523 V. bulbillosum CBS145.70 – Hypocreales AJ292410 V. dahliae UAMH5360 – Hypocreales AF108478 V. dahliae 001 – Hypocreales AF363986 V. dahliae MD80 – Hypocreales AF364004 V. dahliae MD73 – Hypocreales AF364018 V. dahliae 2341 – Hypocreales Z29511 V. gonioides CBS891.72 – Hypocreales AJ292409 V. luteo–album IMI182719 – Hypocreales AJ292420 V. luteo–album IMI017438 – Hypocreales AJ292421 V. nigrescens UAMH6687 – Hypocreales AF108473 V. nigrescens IMI044575 – Hypocreales AJ292440 V. nubilum IMI130213 – Hypocreales AJ292463 V. theobromae IMI172699 – Hypocreales AJ292422 V. tricorpus 1988 – Hypocreales AF364017 V. tricorpus – – Hypocreales L28679 V. tricorpus 267 – Hypocreales Z29524 Volutella colletotrichoides BBA71246 Nectriaceae Hypocreales AJ301962 leaf litter ascomycete strain its016l 1000493338 – – AF502611 leaf litter ascomycete strain its380l 1000494694 – – AF502872 aTaxonomy according to GenBank taxonomy database. ATTC = American Type Culture Collection, CBS = Centraalbureau voor Schimmelcultures, CRCC = Cereal Rust Culture Collection, ICMP = International Collection of Microorganisms from Plants, NRRL = National Centre for Agricultural Utilisation Research (formerly, Northern Regional Research Laboratory), IMI = CABI Bioscience Genetic Resource Collection (formerly, International Mycological Institute), UAMH = University of Alberta Devonian Botanic Garden Microfungus Collection and Herbarium, GJS = G.J. Samuels Systematic Botany and Mycology Laboratory Collection, CTR = Clark T. Rogerson, New York Botanical Garden Collection. Anamorph: bMicrosporum gypseum, cChrysosporum georgiae, dTrichophyton mentagrophytes, eTrichoderma strictipilis, fVerticillium luteo–album, gFusarium ventricosum, hCylindrocarpon destructans var. coprosmae, iCylindrocarpon macroconidialis, jCylindrocarpon destructans, kPlectosporium tabacinum. lThese accessions were not identified to species in GenBank.
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3.4.6 DNA sequence alignment and phylogenetic analysis: All sequences downloaded from GenBank were trimmed to include the complete ITS region (ITS1, 5.8S rRNA gene, ITS2). Where available, up to 10 bases each of the 18S and 26S gene sequences were included at the beginning and end of most sequences, respectively, to aid in the alignment. Individual sequences were then entered separately into the Web−based computer software package WebANGIS version 2.0 (Australian National Genomic Information Service, Sydney, NSW, Australia), where the Pileup subprogram was used to create a single file (pileup.msf) containing sequences representing the entire ITS database. This file was then used as input to load sequences into the ClustalX program developed by Thompson et al. (1997). Once loaded, sequences were aligned via a multistep process, similar to that described by Goodwin et al. (2001). First, a multiple sequence alignment comprising all 74 sequences was performed to identify groups of closely related taxa. A preliminary dendrogram was produced using the Draw NJ–tree option to aid in the selection of groups. A separate alignment was then conducted on each group and the resulting alignments saved as different profiles. The individual profiles were then aligned to each other using this original dendrogram as a guide. Sequences that did not cluster with any of the others in the initial step were aligned as separate profiles. Following alignment, the Draw NJ– tree option was further utilised to produce a second, more accurate representation of the phylogenetic relationships between the isolates. The Draw NJ–tree function calculates genetic distances among isolates using Kimura’s two parameter method for estimating evolutionary distances (Kimura 1980) and prepares a phylogenetic tree based on the neighbour–joining algorithm of Saitou and Nei (1987), equivalent to the DNADIST and NEIGHBOR programs of the PHYLIP package (Felsenstein 1989; Goodwin & Zismann 2001). Statistical support for inferred groups was estimated by bootstrap analysis (Felsenstein 1985) using 1000 replications and was performed using the Bootstrap NJ–tree option. The final tree was displayed with NJplot (Perriere & Gouy 1996) and edited, where required, using MS Word.
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3.5 Results PCR using the oligonucleotide primers AB28 and TW81 permitted the amplification of a single DNA fragment between 501 bp and 710 bp in length for each of the R. alismatis isolates used in this study (Figure 3.1). Following sequence analysis and subsequent removal of terminal primer sequences, the length of the ITS region amplified from each of the R. alismatis isolates measured 501 bp, with the exception of RH126 which measured 503 bp. This stretch included the entire ITS region (ITS1, 5.8S ribosomal RNA gene, ITS2), 13 bp of the 18S ribosomal DNA subunit and 24 bp of the large 28S ribosomal DNA subunit, which were retained to aid alignment. The R. alismatis ITS region alone spanned 464 bases; 136 bases for ITS1, 159 for the 5.8S region and 169 bases for ITS2. An extra nucleotide in each of the ITS regions was observed in isolate RH126. Intraspecific sequence variation between the 48 R. alismatis isolates was minimal, with 46 of the sequences identical, and a further sequence (RH62) differing by only two nucleotides due to the presence of two transition mutations within the ITS1 region at positions 71(T→C) and 125(C→T). However, the remaining isolate, RH126, differed considerably with 15 transition (T↔C, A↔G), 5 transversion (T,C ↔ A,G) and 7 insertion/deletion (indels) mutations. Isolates RH62, RH126 and RH01 were chosen for inclusion in the final ITS database, with the latter isolate representing the 46 identical sequences. All three isolates were also submitted to the GenBank database and in decreasing numerical order were given the accession numbers AY258149–AY258151. In contrast to R. alismatis, PCR amplification of genomic DNA from R. secalis isolate W12868B yielded two fragments between 501 bp and 992 bp in length, corresponding to sequences 575 bp and 711 bp in length respectively, following sequence analysis and the removal of terminal primer sequences. Due to the amplification of multiple fragments, a BLAST search was conducted to determine the identity of these sequences. The shorter sequence returned several hits to other R. secalis isolates in the database, and was identical throughout the entire sequence to a region spanning bases 18–592 of R. secalis isolate RS020 (GenBank accession no. AY140668), published recently by Lee et al. (2001a). Using RS020 as a reference, the sequence of isolate W12868B therefore comprised 13 bp of the 18S ribosomal RNA gene, the entire ITS region, and 23 bp of the 26S ribosomal RNA subunit. The ITS region alone measured 539 bases, consisting of 230 bases for ITS1, 160 bases for the 5.8S gene and 149 bases for ITS2. This sequence differed by one base in comparison with a region spanning bases 342–917 of five identical R. secalis sequences submitted to GenBank by Goodwin (2002), and by a further nucleotide in comparison with a region comprising bases 342–918 bp of a sixth isolate, NKT12, also submitted to GenBank by Goodwin (2002).
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In contrast, the larger fragment had BLAST hits to a number of smut fungi and showed little resemblance to other sequences typical of R. secalis. Because the variation was considerable, this sequence was unlikely to represent a variant within the species (Goodwin pers. comm., 27th July 2001). Furthermore, it was unlike anything reported by the laboratories of Lee et al. (2001a) or Goodwin (2002), and hence, most likely arose from a second organism present as a contaminant. Subsequently, this sequence was omitted from future studies. The sequence for isolate W12868B was also submitted to the GenBank database and was assigned the accession number AY247261.
1 2 3 4 5 6 7 8 9 10 11 12 13 8557
4899 3639
2799
1953 1515 1412 1164 992
710
501 489 404 331 242
Figure 3.1: ITS–PCR of several of R. alismatis and R. secalis isolates. Lane 1: Lower register DNA molecular weight marker, pUC19 DNA digested with HpaII, Lanes 2–10: R. alismatis isolates RH74, RH57, RH41, RH25, RH05, RH64, RH143, RH100, RH122; Lane 11: R. secalis W12868B, Lane 12: pUC19/HpaII DNA molecular weight marker, Lane 13: Higher register DNA molecular weight marker, EcoRI digested DNA from the bacteriophage SPP1. Fragment sizes are shown in bps as indicated on the right.
A BLAST search of the GenBank database using the RH01 isolate identified strong matches with many species of Verticillium and Trichoderma, as well as the teleomorph genera Nectria, Neonectria and Hypocrea. The highest BLAST scores were obtained to isolates of Plectosphaerella cucumerina followed by Verticillium nigrescens and V. nubilum. All sequences 60 downloaded from GenBank had expected values of 2 × 10− or lower in the BLAST results. Based on the results of the BLAST search, 66 sequences representing 34 species, 11 genera, 5 families and 4 orders were downloaded from GenBank and added to the database.
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These included four species from the Arthrodermataceae (order Onygenales) included as an outgroup, and 5 accessions that were not identified to genus or species. With the addition of the ITS sequences of R. secalis and R. orthosporum from Lee et al. (2001a), two R. secalis sequences from Goodwin (2002) and one new R. secalis isolate and three R. alismatis isolates generated de novo, the final database contained 74 sequences representing 39 species in 15 genera (both meio– and mitosporic), 6 families and 4 orders (Table 3.2). The alignment of 74 sequences required 28 profile steps with the original multiple alignment dendrogram being used as a guide to the final alignment. The use of the profile mode of ClustalX to build the alignment ensured that accurate relationships among species within each group were maintained at each step. This yielded a better result with generally higher bootstrap support compared with the original multiple alignment. For all of the profile alignments the gap opening and extension penalties were left at the default setting of 15.00 and 6.66 respectively. To study the relationships between isolates in the ITS database, a phylogenetic tree was produced according to the neighbour–joining method of Saitou and Nei (1987). Confidence values for the inferred groups were estimated by bootstrap analysis (Felsenstein 1985). This analysis clearly indicated that R. alismatis is very closely related to P. cucumerina, Nectria inventa, its anamorph Verticillium luteo–album and several other species in this genus (Figure 3.2). The ITS sequence of P. cucumerina 00017 was identical to RH126 throughout the entire alignment region, spanning bases 4–489 of RH126 and clustered together with RH126 in 93% of bootstrap replications. The remaining P. cucumerina isolates 380408, NRRL20430 and a fourth undesignated isolate differed from RH126 by 5, 8 and 22 nucleotides, respectively, and together with RH126 formed a monophyletic group with 97% bootstrap support. The next most closely related species were those of R. alismatis isolates RH01 and RH62, which differed from RH126 by 27 and 29 nucleotides, respectively, and formed a lower branch with 100% bootstrap support. Together with isolate RH126 and the Plectosphaerella isolates, this lower group formed a larger monophyletic group with 100% bootstrap support and encompassed all 7 isolates from both species. The anamorphs V. nigrescens, V. theobromae and the teleomorphic species N. inventa were the next most closely related species. In addition to identifying close relatives of R. alismatis, this analysis suggests that R. alismatis is neither related to the forme species R. secalis, nor its close relative R. orthosporum. The R. secalis isolate W12868B studied de novo, formed a monophyletic group with the 3 other R. secalis isolates downloaded from the GenBank database, clustering with 100% bootstrap support both within this group and in a second group which also contained the related species, R. orthosporum, which differed from W12868B by 39 nucleotides over an alignment region spanning bases 4–562 of W12868B. 26 Chapter 3. Phylogenetics
Figure 3.2: Unrooted neighbour–joining tree of 74 sequences of the internal transcribed spacer (ITS) region of ribosomal DNA from species of R. alismatis and related anamorphs and teleomorphs. All bootstrap values of 70 or greater (percentage of 1000 replications) are indicated, rounded to the nearest integer. The ITS sequence of several species in the Arthrodermataceae were used as an outgroup. All species are indicated by anamorph name, if known, otherwise by teleomorph. If more than one isolate of a species was analysed, isolate designations are provided after the species name. Branch lengths are proportional to genetic distance, which is indicated by a bar at the upper left.
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3.6 Discussion The analysis of ITS sequences revealed clearly that R. alismatis is closely related to the teleomorph genus Plectosphaerella, but bears little resemblance to the anamorphic ascomycetes R. secalis and R. orthosporum that comprise the genus Rhynchosporium. Whilst the close relationship between R. alismatis and P. cucumerina was not completely unexpected, considering both the controversial classification of R. alismatis and Caldwell’s exclusion of the species from the genus in 1937, the degree of similarity between the organisms was not anticipated. Clearly, isolate RH126 which formed a monophyletic group with all four Plectosphaerella isolates was misidentified during isolation from its host and from a genetic standpoint does not resemble sufficiently the other R. alismatis isolates to be considered the same species. However, RH01 and RH62 which differ by only two nucleotides and together represent the 48 R. alismatis isolates sequenced for this study were also identical to sequences of P. cucumerina over greater than 95% of their length, indicating that R. alismatis is clearly not part of the genus Rhynchosporium and should be renamed. The isolation of RH126 from a non−Alismataceae host, the aquatic grass–like genus Potamogeton, obtained from a Sydney nursery and isolated in the glasshouse by Dr E. J. Cother at the Orange Agricultural Institute, supports this assumption. However, whilst this isolate was clearly misidentified and represents a different species, very likely P. cucumerina, the correct classification of R. alismatis remains somewhat of a mystery. Nevertheless, the similarity between R. alismatis and P. cucumerina suggests that they may represent different species within the same genus. Furthermore, the relatively short branch lengths between these species also suggest they shared a common ancestor relatively recently. For comparison, the sequences of R. secalis and R. orthosporum are 94% similar, suggesting that a similar level of divergence in this genera represents the delineation of species. Whilst the same level of variation cannot be assumed to be the equivalent determinant of species delineation in other genera, some of which are known to comprise species which have identical ITS sequences (Goodwin et al. 2001), this level of divergence provides a potential indicator of the variation one might assume appropriate for the delineation of species within some genera. Considering isolates of the P. cucumerina clustered with those of R. alismatis in this analysis and generated expected values of zero, indicating that there is a zero likelihood of obtaining a match of that magnitude purely by chance from the GenBank database, isolates of R. alismatis may very well represent a separate species within the teleomorphic genus Plectosphaerella.
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The systematics of P. cucumerina, like its counterpart R. alismatis, have been problematic. Although originally described as Venturia cucumerina by Lindfors in 1919, and later as Plectosphaerella cucumeris by Klebahn in 1929, application of the earliest designation, cucumerina Lindfors, was effected in 1972 with the transfer of the organism to the appropriate genus P. cucumerina (Lindfors) W. Gams (references cited by Uecker 1993). Placed in synonymy with several other combinations including Micronectriella cucumeris (Klebahn) (Booth 1971) and Monographella cucumerina (Lindfors) (Von Arx 1984), P. cucumerina and its anamorph, the recently renamed Plectosporium tabacinum (Van Beyma) M.E. Palm, W. Gams et Nirenberg (Palm et al. 1995) (syn. ≡ Cephalosporium tabacinum (Van Beyma 1933, cited by Seifert 1996), Fusarium tabacinum (van Beyma) W. Gams (Gams & Gerlach 1968) and Microdochium tabacinum (Van Beyma) (Von Arx 1984), are common constituents of the rhizosphere and decaying plant materials (Domsch et al. 1980). Although rarely an important pathogen, under the appropriate conditions Plectosporium blight, which is characterised by the appearance of small brown spots that coalesce to form large necrotic lesions on the leaves, stems and petioles of susceptible hosts, has caused severe diseases in a wide range of crop species including tobacco (Nicotiana tabacum) in Tanzania, Malawi and the Netherlands (Booth 1971; Palm et al. 1995), basil (Ocimum basilicum L.), sunflower (Helianthus annuus L.) and tomato (Lycopersicon esculentum Miller) in Italy (Matta 1978; Zazzerini & Tosi 1987), the latter also in Australia (Pascoe et al. 1984), groundnuts (Arachis hypogaea L.) in Nigeria (Odunfa 1979), cucurbits (Cucurbita pepo L., Cucumis sativa L.) in the United States, the former U.S.S.R and Bulgaria (Saad & Black 1981; Bost & Mullins 1992; Everts 2002; Elbakyan 1970; Koleva & Vitanov 1988), arabidopsis (Arabidopsis thaliana) in Spain (Berrocal–Lobo et al. 2002) and Belgium (Thomma et al. 2000; Tierens et al. 2001; 2002) and white lupins (Lupinus termis Forsk.) in Egypt (Youssef et al. 2001). Furthermore, the organism was identified by Alderman and Polglase (1985) as a gill parasite of crayfish (Austropotamobius pallipes) in the United Kingdom and has been isolated from muskmelon (Cucumis melo L.; Bruton & Miller 1997) potato (Solanum tuberosum L.), sugar beet (Beta vulgaris), stem rots in cauliflower (Brassica oleracea), taproot rot in parsnip (Pastinaca sativa), vascular discoloration in balsam (Impatiens balsanii; Pascoe et al. 1984) and wilt diseases in Italian bellflower (Campanula isophylla Moretti; Mygind 1986) and is believed to play a role in the health of soybeans (Glycine max (L.) Merr; Mengistu & Grau 1986; Chen et al. 1996; Hughes et al. 2002).
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Recently however, an additional role for P. tabacinum (the most common form) has been proposed with several groups now studying the organism as a potential biocontrol agent for economically important weed species including false cleavers (Galium spurium L.) in western Canada (Zhang et al. 2002; Zhang, Sulz & Bailey 2002), the invasive aquatic species hydrilla (Hydrilla verticillata (L.) C. Presl) in the United States (Smither–Kopperl et al. 1997; 1998; 1999) and Sagittaria spp. in rice growing areas of the Yusung province, South Korea (Chung et al. 1998). Not surprisingly, this latter study is particularly interesting, considering that the species R. alismatis, currently being considered as a biocontrol agent for Alismataceae weeds in Australian rice crops, now appears to be very closely related to this fungus. Although revised morphological studies (Uecker 1993) and a more recent phylogenetic analysis conducted by Rehner and Samuels (1995) now clearly distinguish P. tabacinum and its teleomorph from their earlier classifications, the taxonomic history and nomenclature of this species is somewhat intertwined with that of the genus Rhynchosporium. Particularly interesting is the history of R. oryzae (Hashioka and Yokogi) which was placed in synonymy with Microdochium oryzae (Hashioka & Yokogi) by Samuels and I.C. Hallett in 1983, and bears the teleomorph classification Monographella albescens (Von Thumen) V.O. Parkinson, Sivanesan et C. Booth (1981). The nomenclature afforded to this organism, which is synonymous with the classifications afforded to both the former meio– and mitosporic forms of Plectosphaerella/Plectosporium provides significant historical evidence that species from both genera were at one time considered not only morphologically similar, but ultimately part of the same genera, both anamorphic and teleomorphic. Coincidently, the descriptions of R. alismatis and P. tabacinum show many similarities. Macroscopically, both fungi are similar, with colonies ranging in colour from off–white to a peach– orange, with a felty or woolly appearance and often with radiating concentric rings. Microscopically, conidia are mostly ellipsoid, straight or slightly curved, appear solitary or in groups, are smooth, hyaline and taper to a broadly rounded apex and to a narrower truncate base. Conidiogenous cells are hyaline, smooth, phialidic in both species and arise from hyphal aggregations. Additionally these species have similar nutritional and environmental requirements during growth and sporulation (Jahromi et al. 1998; Zhang et al. 2001), produce similar disease symptoms on susceptible hosts and have overlapping host ranges that include members of the Alismataceae, specifically S. pygmaea (Chung et al. 1998; Cother & Gilbert 1994a), and possibly the Cucurbitaceae and Solanaceae, from which R. alismatis was isolated on several occasions following host range studies (Cother 1999). Species from both these families were also identified as potential hosts by Zhang, Sulz & Bailey (2002).
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However, in both cases P. tabacinum was shown to cause only low level non–progressive disease on cucurbits and tomato, similar to that of R. alismatis. Similarly, P. tabacinum caused only partial blight of jimson weed (Datura stramonium L.) when tested by Chung et al. (1998), suggesting that both R. alismatis and P. tabacinum are only weak pathogens of these hosts. However, whilst the conidia of R. alismatis are mainly 1–septate, those of P. tabacinum are mostly non–septate or less than 50% are septate (Palm pers. comm., 26th December 2001). Furthermore, conidia in P. tabacinum are produced from unbranched or infrequently branched conidiophores, a feature bearing little resemblance to Caldwell’s (1937) early descriptions of the equivalent structures in R. alismatis. Incidentally Caldwell (1937) described the presence of short flask–shaped conidiophores in this species, later suggesting their mere existence combined with a lack of superficial stroma, was clearly enough to exclude it from the genus Rhynchosporium. Strangely, the presence of conidiophores in this species remains controversial, with Punithalingam (1988) stating their absence in his 1988 treatise. Similarly, Braun’s (1995) interchangeable use of the terms conidiogenous cells and conidiophores during his recent monograph of Cercosporella, Ramularia and allied genera (Phytopathogenic Hyphomycetes) provided little resolution to this debate, making it difficult to determine similarities on this basis. Nevertheless, Braun’s (1995) recent statement that the conidiogenous cells of Spermosporina resemble those of some Microdochium species suggests a further link between what could be considered the past and present classifications of the species R. alismatis. In addition to the relationships between the aforementioned species, the phylogenetic analyses conducted during this study also indicate that other higher level classifications of the ascomycetes may need to be revised. For example, both the Nectriaceae and Clavicipitaceae were polyphyletic, with N. inventa, its anamorph V. luteo–album and other mitosporic Hypocreales forming sub– clusters proximal to the Phyllachoraceae (order Phyllachorales), and suggesting some degree of evolutionary divergence between the Phyllachorales and the Hypocreales. The was also observed by Rehner and Samuels (1995) who reported polyphyly in the genus Verticillium during a recent phylogeny study of the Hypocreales. These authors observed that V. dahliae clustered with P. cucumerina during analysis of 28S rDNA sequences. Domsch et al. (1980) suggested that the formation of microsclerotia was a characteristic that potentially excluded them from the Hypocreales, however this does not seem to be a characteristic of V. nigrescens (Pegg & Brady 2002), which clustered as a sub–group to Plectosphaerella and appeared to be more closely related to the Phyllachorales than to the other members of the Hypocreales.
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Similarly, members of the Clavicipitaceae were separated by the mitosporic Hypocreales, V. gonioides and V. bulbillosum, which clustered with both isolates of T. parasiticum under 100% bootstrap support and suggested that these isolates may well need to be revised, possibly to the Clavicipitaceae. This cluster also suggested that the leaf litter ascomycetes which were previously unidentified at both the genus and species level are related to the anamorphic genus Cordyceps, and are also members of the Clavicipitaceae. Unlike these clusters however, bootstrap support for many higher level relationships was often low, with exclusion of gaps and correction for multiple substitutions having little effect on tree topology, especially among highly supported groups. Difficulty in aligning sequences at greater evolutionary distances probably accounted for these results. Despite the morphological, developmental and epidemiological similarities between R. alismatis, P. tabacinum and its teleomorph P. cucumerina including the synonymy between former classifications of the two species, the question still remains as to the correct classification of R. alismatis. Although the current data is unlikely to provide all the evidence necessary to rename, or fully identify this species, it is clear that R. alismatis should no longer be considered under the genus Rhynchosporium. However, until such time as this species is officially reclassified, the current combination R. alismatis and the morphological comparisons with the forme species R. secalis that occur throughout the remainder of this thesis will be retained. Whilst these studies belie fears that R. alismatis may pose a similar threat to cropping regimes as the scald pathogen R. secalis, the close relationship between R. alismatis and the teleomorph genus, Plectosphaerella, introduces new questions about the risks and potentials of this organism as an agent for biological weed control. These factors are investigated via the application of population structure analyses (Kistler 1991).
32 Chapter 4. Population Structure
4 Population Structure
A population structure analysis of R. alismatis – evaluating the risks and potentials of a biocontrol control agent
4.1 Introduction Over the last decade or so, plant pathologists have begun to realise that more knowledge about the genetic structure of populations of plant pathogens is required to implement effective control strategies (Wolfe & Caten 1987). As a result, research on the genetic structure of fungal populations has mushroomed, and review papers that summarise these studies are numerous (Leung et al. 1993; McDermott & McDonald 1993; McDonald & McDermott 1993; McDonald et al. 1989; Milgroom 1995). Although the number of studies has increased greatly, the most comprehensive work has focused on a small number of agriculturally destructive fungal pathogens such as Phytophthora infestans, the casual agent of potato late blight, and the wilt disease pathogen Fusarium oxysporum (Goodwin 1997; Kistler 1997). To a lesser extent, Burdon (1992; 1993) has applied the theory, approaches and tools of population genetics to the study of pathogen populations in natural plant communities. However, thus far few studies have focused on determining the population structure of beneficial fungal pathogens such as those being investigated as potential biological control agents for weeds (Gosselin et al. 1999; Hintz et al. 2001; Ramsfield et al. 1996; 1999). In this chapter, a ‘populations genetics approach’ will be applied to the study of biological weed control. Firstly, a brief introduction will delineate the basic concepts and methodologies used in describing population structure. This will be followed by a detailed discussion of the population structure of R. alismatis, including the forces that give rise to and maintain genetic diversity within and between populations of the organism. Finally an interpretation of this knowledge as it applies to the discipline of biological weed control will be given. This will include an evaluation both of the risks and potentials of this organism, and an assessment of the suitability of the organism as a future agent for the control of Alismataceae weeds in rice.
33 Chapter 4. Population Structure
4.2 Concepts and methodologies Defining the ‘genetic structure’ of populations is the logical first step in studies of fungal population genetics. ‘Genetic structure’ refers to the amount and distribution of genetic variation within and among populations (Chen et al. 1995). For plant pathogenic fungi, the processes of mutation and recombination are responsible for introducing variation into a population. Existing variation is acted upon by selection, migration and random drift (Burdon & Silk 1997). Genetic approaches have not previously been applied to populations of the fungus R. alismatis, and descriptive questions about the amount and distribution of genetic variation must therefore be answered before mechanistic questions can be posed. It is not possible to address questions relating to selection, migration, mutation and genetic drift for example, until we have answered questions concerning allele frequencies, mating systems and the geographic scale over which variation is found. In many cases, the geographic distribution of sampling sites depends on population subdivision and allele frequencies, which cannot be estimated without a prior survey (Braverstock & Moritz 1990). Hence, the first analysis performed on a population quantifies variation and forms the basis of a preliminary study. Once the results of preliminary studies have been evaluated, iteration may be required to better define the boundaries of populations by sampling over larger or smaller spatial scales. In most population studies, gene frequencies are the basic units of measurement of variation (Leung et al. 1993). For organisms like fungi that undergo both sexual and asexual reproduction, it is necessary to differentiate between diversity at individual loci, ‘gene diversity’ and diversity based on the number of genetically distinct individuals in a population, otherwise known as ‘genotype diversity’(McDonald 1997). ‘Gene diversity’, based on the number and frequency of alleles at a locus, is affected by the age of the population, population size and selection (Welz et al. 1994). Populations that have evolved in one location over long periods generally have more alleles than populations that are more recent introductions, and hence have higher gene diversity (McDonald 1997), because there has been more time for mutation to introduce new variants and for genetic drift to increase the frequencies of new alleles to detectable levels (Slatkin 1987). Populations with low gene diversity on the other hand, may have been affected by large reductions in population size (bottlenecks) or by local extinction and recolonisation events, ‘founder effects’, that eliminate many alleles (Chen et al. 1994). In most cases, gene frequency is preferred over genotypic frequency because gene frequencies remain relatively stable over time and are independent of the mating system (Leung et al. 1993). Hence, fungi that reproduce exclusively through asexual reproduction may have as many alleles at individual loci as their sexual counterparts (McDonald 1997).
34 Chapter 4. Population Structure
‘Genotype diversity’, based on the number and frequency of genotypes within a population (Chen et al. 1995), is also an informative parameter of the genetic structure within populations and is used to integrate the diversity from individual loci into multilocus genotypes, a process mediated primarily by mode of reproduction (Stoddart & Taylor 1988). Asexual reproduction leads to clonal lineages with few genotypes present at high frequencies (Zeigler et al. 1995), whilst random mating (sexual) populations are expected to display a high degree of genotypic diversity (Leung et al. 1993). In situations where the genetic control of phenotypes is unknown, such as in many asexual populations, DNA fingerprints (Chen et al. 1995) or multilocus haplotypes (composite genotypes) (Burdon & Jarosz 1992) may be used to differentiate genotypes and measure variation within and between populations. Taken together, gene and genotype diversity constitute genetic variation and are among the easiest and most informative parameters to quantify (Weir 1996). Because the evolution of a population is largely the result of gene frequency changes due to mutation, selection, migration and genetic drift (Leung et al. 1993), gene frequency comparisons between populations can be useful for defining population boundaries, sources of introduction and paths of migration among populations. Where the movement of individuals between populations is absent, the expectation is that genetic drift will lead to random changes in allele frequencies in different populations (McDonald 1997). If there is movement of genes among populations, comparisons of allele frequencies in different populations provides an estimation of population subdivision and gene flow (Boeger et al. 1993). Differences in mating systems are reflected by measures of genotype diversity (Leung et al. 1993). The distribution and spatial analysis of genotypes within and between populations provides evidence of dispersal. If non−random clustering of genotypes provides evidence of restricted dispersal, the processes of migration, selection and genetic drift become important as factors maintaining similarities or contributing to genetic divergence between populations (Burdon et al. 1990) and the chances of significant differentiation between individual populations increases (Wright 1969). Where dispersal is unrestricted, the force of gene flow may be counteracted by genetic drift and the differentiation between populations may be minimal. In these cases, the balance between gene flow and genetic drift provides a backdrop against which to consider the effects of different kinds of selection (Slatkin 1987).
35 Chapter 4. Population Structure
4.3 Tools and Techniques Unlike plants and animals, fungi have few morphological characters that can be used to study their population structure. Fortunately, PCR amplification using DNA markers has facilitated the testing of hypothesis about structure and relationships within phenetically defined species (Hawkesworth et al. 1995). By measuring genotype, rather than phenotype, molecular markers provide excellent tools for assessing and quantifying genetic variation (Huff et al. 1994; McDermott et al. 1994; Kistler 1997), defining genetic relationships (O’Hanlon et al. 2000) and distinguishing among the forces that generate and maintain genetic variation (Slatkin 1987; Goodwin et al. 1994; Chen & McDonald 1996, McDonald et al. 1996; Burdon & Silk 1997). Whilst the advantages and disadvantages of different molecular markers have been reviewed elsewhere (Michelmore & Hulbert 1987; Kohn 1992; Burdon 1993; McDonald & McDermott 1993; Rosewich & McDonald 1994; Jones et al. 1997; McDonald 1997), one strategy that has gained recognition more recently is PCR−fingerprinting with primers directed against repetitive elements including simple sequence repeats (SSR), enterobacterial repetitive intergenic consensus (ERIC) sequences and repetitive extragenic palindromic (REP) sequences (Meyer et al. 1993; Versalovic et al. 1991). The technique of PCR−fingerprinting is an inexpensive and rapid technique for genetic structure analysis of phytopathogenic fungi, recognised for its relative accuracy, reliability and reproducibility (Welsh & McClelland 1990; Caetano−Anolles et al. 1991). In PCR−fingerprinting, primers are directed against repetitive elements, arbitrarily chosen or defined, including microsatellites (Meyer et al. 1993), prokaryotic and eukaryotic repeat motifs (Versalovic et al. 1991) or at the regions between them. In cases where two distinct flanking primers are typically required and the application of PCR−fingerprinting is limited by insufficient knowledge of the sequences flanking the locus of interest (Godwin et al. 1997), a single arbitrary primer that is complementary to nucleotide sequences that are present within an amplifiable distance and in an inverted orientation will prime amplification of the intervening DNA segment between adjacent motifs (Zietkiewicz et al. 1994). Termed inter–simple sequence repeat–PCR or ISSR–PCR (Godwin et al. 1997), polymorphisms are detected based on the number of repeat motifs and the characteristic pattern of PCR products thus obtained can be considered a fingerprint of the DNA template (Williams et al. 1990). The utility of microsatellites or SSRs, which are tandemly repeated motifs 2−10 bp in length, results from their inherent variability, wide range of repeats and Mendelian inheritance (Groppe et al. 1995). The hypervariability of SSR loci is a consequence of unusually high mutation rates that are thought to be the result of slipped−strand mispairing during DNA replication, adding or subtracting repeat units from the locus (Tautz 1989; Strand et al. 1994).
36 Chapter 4. Population Structure
As a result of high mutation rates, SSR loci may be polymorphic even in species otherwise characterised by low levels of genetic diversity (Peakall et al. 1998) and hence are considered ideal markers for population genetics (Jarne & Logoda 1996). Repetitive repeat motifs such as the prokaryotic ERIC and REP sequences have been examined extensively in bacterial genomes (Versalovic et al. 1991). PCR−fingerprinting with primers matching these regions has been used extensively to characterise a number of agriculturally important bacterial species including several species of Rhizobium (De Bruijn 1992) and Bradyrhizobium (Judd et al. 1993). Recently, it has been confirmed that ERIC− and REP−like sequences are also present in the genomes of diverse fungal species including Aspergillus (Van Belkum et al. 1993), Fusarium (Edel et al. 1995), Verticillium (Arora et al. 1996), the rice blast pathogen Magnaporthe grisea (George et al. 1998) and Leptosphaeria maculans, causal agent of blackleg disease of oilseed rape (Jedryczka et al. 1999). These sequences contain highly conserved palindromic inverted repeats which have the potential to form stem−loop structures, believed to play important roles in the organization of bacterial genomes (Krawiec & Riley 1990). Because genome organization is thought to be shaped by selection, the dispersion of ERIC and REP sequences may reflect the structure and evolution of the genome (Krawiec 1985). Furthermore, because the relative positions of these sequences in the genome appear to be conserved in closely related strains and distinct in diverse species (Versalovic et al. 1991; Lupski & Weinstock 1992), they too represent useful markers for studies in fungal population genetics. To more effectively assess the risks and potentials of R. alismatis as a candidate mycoherbicide for Alismataceae weeds in Australian rice fields, knowledge of the genetic structure and hence, the evolutionary history of the organism, would be useful. By determining the genetic structure of field populations of R. alismatis from different geographic locations, we may be able to gain some insight into the reproductive strategies, pathogenicity attributes and dispersal potential of the fungus in the environment. Furthermore, the potential risk that unexpected dispersal poses to non−target species (Watson 1994), the likelihood of evolution of pathogenicity towards other species and the potential for unexpected genetic exchange with closely related fungal species present within its natural range could be evaluated (Weidemann 1992). By studying these relationships, population structure analyses enable researchers to make educated decisions about the suitability of organisms as candidates for biological weed control prior to their release in the field.
37 Chapter 4. Population Structure
4.4 Materials and Methods 4.4.1 Fungal isolates and DNA extraction: Fungal isolates were collected, isolated, cultivated, stored, maintained, prepared and subsequently extracted for DNA by the methods described in section 3.4.1. The three geographic regions encompassing 17 locations from which isolates of R. alismatis were obtained are shown in Figure 4.1. Several isolates used in the previous study were excluded from this analysis. These included isolates RH80, RH140 and RH141 which were omitted due to their non–Australian origin, isolates RH147 and RH148 obtained from glasshouse infections, and RH126 which appeared to be more closely related to the teleomorph species P. cucumerina. The scald pathogen isolate W12868B was also omitted from this analysis. Isolates used in this study are listed in Table 4.1.
Figure 4.1: The three geographic regions, Coleambally Irrigation Area (CIA: 1–4), Southern Riverina (SR: 5–11) and Central and Southern Tablelands (CST: 12–17) encompassing a total of 17 different locations, from which isolates of R. alismatis were collected for use in this study.
38 Chapter 4. Population Structure
Table 4.1: Origin, host, population and collection date of R. alismatis cultures used in this study. Isolate name and Herb. DAR Host Origin Isolation population accession number date designation Coleambally Irrigation Area (CIA) RH001 DAR67515 A. lanceolatum Coleambally Jan. 89 RH021 DAR67510 A. plantago–aquatica Narranderra Jan. 91 RH024 DAR67517 A. lanceolatum Coleambally Jan. 91 RH025 DAR73158 A. lanceolatum Coleambally Jan. 91 RH046 DAR67511 D. minus Yanco Jan. 91 RH047 DAR76105 D. minus Yanco Jan. 91 RH054 DAR73791 A. lanceolatum Hanwood May 91 RH055 DAR76104 D. minus Yanco May 91 RH057 DAR67516 S. montevidensis Yanco May 91 RH091 DAR73673 A. lanceolatum Hanwood May 91 RH100 DAR73860 D. minus Yanco May 91 RH121 DAR73794 A. lanceolatum Coleambally Jan. 91 RH122 DAR73795 A. lanceolatum Coleambally Jan. 91 RH123 DAR73796 A. lanceolatum Coleambally Jan. 91 RH124 DAR73797 A. lanceolatum Coleambally Jan. 91 RH127 DAR73798 A. lanceolatum Coleambally Jan. 91 RH149 DAR76285 D. minus Coleambally Jan. 91 Central and Southern Tablelands (CST) RH058 DAR67508 A. plantago–aquatica Hobbys Yards Jan. 92 RH062 DAR67513 A. plantago–aquatica Khancoban Jan. 92 RH064 DAR73146 A. plantago–aquatica Rosewood Jan. 92 RH066 DAR67512 A. plantago–aquatica Tumut Jan. 92 RH069 DAR73148 A. plantago–aquatica Petfield Jan. 92 RH074 DAR73149 A. plantago–aquatica Rockley Jan. 92 RH095 DAR73150 A. plantago–aquatica Petfield Jan. 92 RH097 DAR73151 A. plantago–aquatica Rockley Jan. 92 Southern Riverina (SR) RH005 DAR73786 A. lanceolatum Pericoota Dec. 89 RH037 DAR73787 A. lanceolatum Gambles Lane Jan. 91 RH038 DAR73788 A. lanceolatum Gambles Lane Jan. 91 RH039 DAR73789 A. lanceolatum Gambles Lane Jan. 91 RH041 DAR73790 A. lanceolatum Pericoota Jan. 91 RH108 DAR73861 D. minus Yallakool Feb. 93 RH111 DAR73793 A. lanceolatum Caldwell Feb. 93 RH118 DAR73672 A. lanceolatum Pericoota Dec. 89 RH133 DAR73799 A. plantago–aquatica Deniliquin Jan. 95 RH135 DAR73800 A. plantago–aquatica Deniliquin Jan. 95 RH136 DAR76107 D. minus Derrulaman Jan. 95 RH137 DAR76106 D. minus Derrulaman Jan. 95 RH138 DAR73862 D. minus Derrulaman Jan. 95 RH139 DAR73145 D. minus Derrulaman Jan. 95 RH143 DAR73152 D. minus Barmah Mar. 97 RH144 DAR73153 D. minus Barmah Mar. 97 RH145 DAR73154 D. minus Barmah Mar. 97
39 Chapter 4. Population Structure
4.4.2 Repetitive Repeat motif (ERIC/REP)−PCR: Oligonucleotide primer pairs, ERIC–1R/–2 and REP1R−1/REP2−1, described by Versalovic et al. (1991) and synthesised by Gibco BRL Life Technologies, were used for PCR amplification of genomic DNA from R. alismatis isolates. Details of primers used in this study are shown in Table 4.2. Amplification was performed in PCR reaction buffer, the constituents of which were described in section 3.4.3, with the addition of 1.25 mM each of dATP, dCTP, dGTP and dTTP (Amersham Pharmacia Biotech); 50 pmol of each primer; 2.0 mM MgCl; with 50 ng of purified genomic DNA and 2 units of Taq DNA polymerase (Promega Corporation) per 25 µL reaction. DNA amplifications were performed as described by Edel et al. (1995), with an initial denaturation (ERIC: 7 minutes at 95°C; REP: 6 minutes at 95°C) followed by 30 cycles of denaturation (1 minute at 94°C), annealing (ERIC: 1 minute at 52°C; REP 1 minute at 40°C), and extension (8 minutes at 65°C) with a final extension (16 minutes at 65°C).
4.4.3 Simple sequence repeat (SSR)−PCR: One hundred simple sequence repeat primers were screened in preliminary experiments (UBC801−900, Nucleic Acid Protein Service Unit, University of British Columbia, Vancouver, Canada). Sixteen primers were screened for the entire population of isolates. Of these, primers SSR–807, –808, –809, –810, –835, –842, –857 and –890 (Table 4.2) generated polymorphic banding patterns. PCR reactions were carried out in a 25µL volume containing the DNA template (35 ng of purified DNA); 0.15 µM of each primer; 2.0 mM MgCl; 0.1 mM each of dATP, dCTP, dGTP and dTTP (Amersham Pharmacia Biotech); and 1 unit of Taq DNA polymerase (Promega Corporation) in a reaction buffer as described in section 3.4.3. After an initial denaturation of 1 minute at 95°C, 40 cycles of denaturation (1 minute at 93°C), annealing (2 minutes at 47°C [primers 807 and 810], 48°C [890], 49°C [808 and 809] or 52°C [835, 842 and 857]), and extension (3 minutes at 72°C) were performed, followed by a final 5 minutes extension at 72°C. PCR for both repeat motif and SSR–PCR was performed essentially as described in section 3.4.3. However, 15 µl aliquots of each PCR product, as opposed to the 35 µl ITS aliquots, were mixed with loading buffer and analysed by electrophoresis. Electrophoresis was performed as described in section 3.4.2 and all ERIC–, REP– and SSR–PCR amplifications were performed in duplicate to confirm that results were reproducible.
40 Chapter 4. Population Structure
Table 4.2: Primers used during population structure analysis of R. alismatis. Primer name Nucleotide sequencea Size Tm (bp) (°C) Repetitive repeat motif (ERIC/REP)–PCR ERIC–1R ATG TAA GCT CCT GGG GAT TCA C 22 50 ERIC–2 AAG TAA GTG ACT GGG GTG AGC G 22 52 REP1R–1 III (ICG)2 ICA TCI GGC 18 43 REP2–1 ICG ICT TAT CIG GCC TAC 18 43 Simple sequence repeat (SSR)–PCR SSR–807 (AG)8T 17 50 SSR–808 (AG)8C 17 52 SSR–809 (AG)8G 17 52 SSR–810 (GA)8T 17 50 SSR–835 (AG)8YC 18 55 SSR–842 (GA)8YG 18 55 SSR–857 (AC)8YG 18 55 SSR–890 VHV(GT)7 17 51 aSingle letter abbreviations for mixed base positions are as follows: Y stands for pYrimidine, H for non−G, V for non−T and I for Inosine (Nomenclature Committee of the International Union of Biochemistry 1986).
4.4.4 Data analysis: PCR products were analysed using visual pairwise comparisons of adjacent lanes by reading horizontally across the gel from the bottom to the top. All definitive bands were scored regardless of their intensity and the presence of single, dominant bands in each lane allowed alignment of PCR products across non−contiguous lanes. DNA fragments or combinations of fragments with different sizes were treated as loci for each primer. Only loci that were polymorphic across all isolates were included in the analysis. Isolates having the same loci for each of the single primers were assigned a multilocus haplotype, resulting from the combination of all primers used in the analysis. Isolates with the same multilocus haplotype were assumed to be individual members of the same clone (McDonald & Martinez 1991). Genetic variation in different populations was measured using Nei’s (1973) measure of gene diversity for individual loci and across loci. Nei introduced the concept of gene diversity to describe allelic variation that is applicable to both sexual and asexual populations. Gene diversity
(Hi) is defined as the probability of obtaining two different alleles at a locus when two haploid individuals are sampled from a population, where xi is the frequency of the ith allele at a locus. The method of Nei was modified according to Selander et al. (1985), where n/(n−1) is a correction for bias in small samples.
(1)
41 Chapter 4. Population Structure
A gene diversity of one indicates that any two alleles at a locus sampled from a population are different. A genetically uniform population (with no allelic variation at the loci sampled) will have a diversity of zero since any two individuals sampled will be identical. The χ2 statistic was used to test for differences in allele frequencies between populations (Workman & Niswander 1970; Hudson et al. 1992);
(2)
in which R is the number of populations in the total sample, V is the number of alleles at each locus, ni is the sample size in location i, nij is the observed number of isolates with allele j from location i, and pj is the frequency of allele j in the population. The degrees of freedom for this test is (v−1)(r−1), where v is the number of alleles at each locus and r is the number of populations. Nei’s (1973) population subdivision approach to haploids and asexual populations was used to partition the total gene diversity into component diversities according to subgroups from different geographic locations. The average gene diversity between subpopulations including the comparison of subpopulations with themselves is Dst;
(3)
where Ht is the total genetic diversity over all groups as defined in equation (1) and Hs is the average genetic diversity of all subgroups. The individual Hs estimates are calculated according to equation (1) based on the gene or multilocus haplotype frequencies found within a particular subpopulation. The population differentiation coefficient Gst is defined as;
or or (4)
The amount of gene flow between populations (Nm), where N is the population size and m is the fraction of individuals in a population that are immigrants, was estimated by substituting Nei’s Gst for Fst in Wright’s island model of gene flow (Wright 1951; 1969). Gst is useful because it is not contingent upon any assumption about the breeding history of the population. According to Wright’s model;
(5)
42 Chapter 4. Population Structure
The formula used to calculate Nm was;
(6)
The 4 was replaced by a 2, because R. alismatis is haploid (Braun 1995). If two populations are similar in size, Nm estimates the average number of individuals that migrate between populations per generation. If Nm < 1, then local populations will differentiate; if Nm ≥ 1 , then there will be little differentiation among populations (Slatkin 1987). Nei’s (1972) pairwise measures of genetic identity (I) and genetic distance (D) were also calculated. The normalised identity of genes between populations X and Y with respect to individual loci is defined as;
(7)
where xi and yi are the frequencies of the ith alleles in populations X and Y, respectively, and the 2 2 probability of identity of two randomly chosen genes is jX = ∑xi in population X and jY = ∑yi in population Y. The probability of identity of a gene from X and a gene from Y is jXY = ∑xiyi. The genetic distance between X and Y is then defined as;
(8)
In addition to Nei’s measures of gene diversity, genotypic diversity in a population, based on comparisons of the number of isolates with different DNA fingerprints or multilocus haplotypes, was calculated using the measure proposed by Stoddart and Taylor (1988);
(9)
where N is the sample size and fx is the number of genotypes observed x times in the sample. The maximum possible value for Ĝ, which occurs when each individual in the sample has a unique genotype, is the number of individuals N in the sample.
43 Chapter 4. Population Structure
To compare Ĝ in populations with different sample sizes, Ĝ from each population was divided by its sample size to calculate the percentage of maximum possible diversity that was obtained. Normalised measures of genotypic diversity were compared using a t−test for differences between the percentage of maximum possible diversity determined for each population. Stoddart and Taylor (1988) showed that;
(10)
where G is the population genotypic diversity, K is the number of genotypes in the sample, and pi is the frequency of the ith genotype in the sample. Ĝ is the maximum likelihood estimator for G in this formula. The t−test for the significance of differences between genotypic diversities was calculated as;
(11)
where Ĝ1 and Ĝ2 were the genotypic diversities observed in populations 1 and 2 which had sample sizes of N1 and N2, respectively. The degrees of freedom for this t−test is N1 + N2 – 2. The freeware computer package POPGENE − Population Genetics Analysis Version 1.32 was used intermittently during this study (Yeh & Boyle 1997; Yeh et al. 1997).
44 Chapter 4. Population Structure
4.5 Results A total of 42 R. alismatis isolates from 17 locations within three Australian geographic regions (populations) were analysed in this study. The number of loci per primer ranged from one to four with an average of 2.7 polymorphic loci per primer. Allele frequencies, sample sizes and the χ2 statistic (Workman & Niswander 1970; Hudson et al. 1992) for differences in allele frequencies at each loci among populations are shown in Table 4.3. Despite significant differences in the frequency of alleles at several loci, Nei’s measure of genetic diversity across all loci was similar for the three regions, ranging from 0.1159 for the CIA to 0.2682 in the CST region. Nei’s measures of gene diversity (Hi), pairwise comparisons of genetic identity (I), genetic distance (D), population differentiation (Gst), gene flow (Nm) and the distribution of diversity into components between (Dst) and within (Hs) regions are shown in Table 4.4. On average the total gene diversity across all regions was moderate to low (0.1924), with diversity between and within regions accounting for 9% and just over 90% of the total diversity, respectively. Gene diversity was approximately one third of that reported by McDonald et al. (1999) and Salamati et al. (2000) for the scald pathogen
R. secalis. The average genetic distance (D =0.0265) and overall gene differentiation (Gst =0.1001) between regions were relatively small, and Nei’s normalised genetic identity (I) was 0.97 for all regions across all loci, compared to the theoretical maximum of 1.00, which is reached when the same alleles are present at the same frequencies. Stoddart and Taylor’s (1988) measures of genotypic diversity were also very similar among the three regions. In the CIA, there were 10 different DNA fingerprints among the 17 isolates (Figure 4.2). One DNA fingerprint was observed four times, one three times, two twice and the remaining six only once. Stoddart’s measure of genotype diversity in this sample was 7.41± 1.84, which was 43.59% of the maximum possible value of 17. In the CST area there were seven different DNA fingerprints among the eight isolates. With the exception of one DNA fingerprint, all were observed only once. Stoddart’s measure was 6.40±1.57 or 80.00% of the theoretical maximum. Stoddart’s measure for the 17 isolates comprising the SR region was 7.41±2.78 or 43.59%. Of the twelve DNA fingerprints observed, 10 were found to be unique. Five and two isolates respectively, shared the remaining two genotypes. Pairwise t−tests between the three populations identified significant differences in genotype diversities between the CST and the other regions (Table 4.5).
45 Chapter 4. Population Structure
Table 4.3: Allele frequencies at 24 loci in Australian populations of R. alismatis collected from three regions in New South Wales during the period January 1989 to March 1997. Sample sizes (N) used to calculate allele frequencies are shown at the bottom of the table. A χ 2–test (degrees of freedom in parenthesis) for differences in allele frequencies is shown at the end of each locus. Locus Alleles CIA CST SR χ2–test
1 0 0.2941 0.0000 0.0000 1 0.7059 1.0000 1.0000 8.35(2)* 2 0 0.0000 0.2500 0.0588 1 1.0000 0.7500 0.9412 5.19(2) 3 0 0.3529 0.3750 0.2941 1 0.6471 0.6250 0.7059 0.21(2) 4 0 0.9412 1.0000 1.0000 1 0.0588 0.0000 0.0000 1.51(2) 5 0 0.2353 0.2500 0.2941 1 0.7647 0.7500 0.7059 0.16(2) 6 0 0.7647 0.7500 0.7647 1 0.2353 0.2500 0.2353 0.01(2) 7 0 1.0000 0.8750 1.0000 1 0.0000 0.1250 0.0000 4.35(2) 8 0 0.0000 0.2500 0.4706 1 1.0000 0.7500 0.5294 10.38(2)** 9 0 1.0000 0.7500 0.5294 1 0.0000 0.2500 0.4706 10.38(2)** 10 0 1.0000 0.8750 1.0000 1 0.0000 0.1250 0.0000 4.35(2) 11 0 0.0000 0.1250 0.0000 1 1.0000 0.8750 1.0000 4.35(2) 12 0 0.8235 0.7500 0.7647 1 0.1765 0.2500 0.2353 0.25(2) 13 0 0.1765 0.2500 0.2353 1 0.8235 0.7500 0.7647 0.25(2) 14 0 1.0000 0.8750 1.0000 1 0.0000 0.1250 0.0000 4.35(2)
46 Chapter 4. Population Structure
Table 4.3: (cont.) Locus Alleles CIA CST SR χ2–test
15 0 0.8824 0.7500 1.0000 1 0.1176 0.2500 0.0000 4.11(2) 16 0 0.0000 0.1250 0.0000 1 1.0000 0.8750 1.0000 4.35(2) 17 0 1.0000 0.7500 1.0000 1 0.0000 0.2500 0.0000 8.93(2)* 18 0 0.8235 1.0000 0.9412 1 0.1765 0.0000 0.0588 2.41(2) 19 0 0.0000 0.2500 0.0000 1 1.0000 0.7500 1.0000 8.93(2)* 20 0 0.0000 0.2500 0.0000 1 1.0000 0.7500 1.0000 8.93(2)* 21 0 0.0000 0.2500 0.0000 1 1.0000 0.7500 1.0000 8.93(2)* 22 0 0.0000 0.0000 0.0588 1 1.0000 1.0000 0.9412 1.51(2) 23 0 1.0000 1.0000 0.9412 1 0.0000 0.0000 0.0588 1.51(2) 24 0 1.0000 0.7500 1.0000 1 0.0000 0.2500 0.0000 8.93(2)* N 2 17 8 17 * and ** indicate significance at P = 0.025 and 0.010, respectively.
There were three cases where isolates possessing the same DNA fingerprint were found in more than one region. In two of these cases, a clone was sampled in each of two populations, whilst in the remaining case eight isolates, five from the SR area, two from the CST region and one from the CIA, were found to have identical multilocus haplotypes and hence the same genotype. In one case two isolates from the CIA formerly thought to be identical, were found to have different genotypes after a multilocus haplotype was constructed.
47
Chapter 4. Population Structure
Table 4.4: Nei’s (1973) measures of gene diversity for each collection (Hi), total gene diversity over all collections (Ht), gene diversity within collections (Hs), and average gene diversity between collections (Dst) for 24 loci in Australian populations of R. alismatis collected from three regions in New South Wales during the period January 1989 through March 1997. Nei’s (1972) normalized measures of genetic identity (I), genetic distance (D), pairwise comparisons of population differentiation (Gst), and amount of gene flow (Nm) (Nei 1973) between collections are also shown for each locus. Hi I D Gst Locus Overall Nm # CIA CST SR Ht Hs Dst CIA/CST CIA/SR CST/SR CIA/CST CIA/SR CST/SR CIA/CST CIA/SR CST/SR Gst 1 0.4152 0.0000 0.0000 0.1769 0.1384 0.0385 0.9231 0.9231 1.0000 0.0800 0.0800 0.0000 0.1724 0.1742 0.0000 0.2176 1.8000 2 0.0000 0.3750 0.1107 0.1847 0.1619 0.0228 0.9487 0.9981 0.9666 0.0527 0.0019 0.0340 0.1429 0.0303 0.0700 0.1234 3.5538 3 0.4567 0.4688 0.4152 0.4492 0.4469 0.0023 0.9992 0.9945 0.9894 0.0008 0.0055 0.0106 0.0005 0.0040 0.0073 0.0051 95.8686 4 0.1107 0.0000 0.0000 0.0384 0.0369 0.0015 0.9981 0.9981 1.0000 0.0019 0.0019 0.0000 0.0303 0.0303 0.0000 0.0391 12.0000 5 0.3599 0.3750 0.4152 0.3846 0.3834 0.0012 0.9997 0.9954 0.9973 0.0003 0.0046 0.0027 0.0003 0.0044 0.0025 0.0031 153.4038 6 0.3599 0.3750 0.3599 0.3650 0.3649 0.0001 0.9997 1.0000 0.9997 0.0003 0.0000 0.0003 0.0003 0.0000 0.0003 0.0003 1898.2500 7 0.0000 0.2188 0.0000 0.0799 0.0729 0.0070 0.9899 1.0000 0.9899 0.0101 0.0000 0.0101 0.0667 0.0000 0.0667 0.0876 5.2000 8 0.0000 0.3750 0.4983 0.3650 0.2911 0.0739 0.9487 0.7474 0.9191 0.0527 0.2912 0.0843 0.1429 0.3077 0.0528 0.2025 1.9691 9 0.0000 0.3750 0.4983 0.3650 0.2911 0.0739 0.9487 0.7474 0.9191 0.0527 0.2912 0.0843 0.1429 0.3077 0.0528 0.2025 1.9691 10 0.0000 0.2188 0.0000 0.0799 0.0729 0.0070 0.9899 1.0000 0.9899 0.0101 0.0000 0.0101 0.0667 0.0000 0.0667 0.0876 5.2500 11 0.0000 0.2188 0.0000 0.0799 0.0729 0.0070 0.9899 1.0000 0.9899 0.0101 0.0000 0.0101 0.0667 0.0000 0.0667 0.0876 5.2500 12 0.2907 0.3750 0.3599 0.3439 0.3418 0.0021 0.9939 0.9962 0.9997 0.0061 0.0038 0.0003 0.0081 0.0053 0.0003 0.0061 84.6786 13 0.2907 0.3750 0.3599 0.3439 0.3418 0.0021 0.9939 0.9962 0.9997 0.0061 0.0038 0.0003 0.0081 0.0053 0.0003 0.0061 84.6786 14 0.0000 0.2188 0.0000 0.0799 0.0729 0.0070 0.9899 1.0000 0.9899 0.0101 0.0000 0.0101 0.0667 0.0000 0.0667 0.0876 5.2500 15 0.2076 0.3750 0.0000 0.2151 0.1942 0.0209 0.9821 0.9912 0.9487 0.0180 0.0088 0.0527 0.0292 0.0625 0.1429 0.0972 4.6555 16 0.0000 0.2188 0.0000 0.0799 0.0729 0.0070 0.9899 1.0000 0.9899 0.0101 0.0000 0.0101 0.0667 0.0000 0.0667 0.0876 5.2500 17 0.0000 0.3750 0.0000 0.1528 0.1250 0.0278 0.9487 1.0000 0.9487 0.0527 0.0000 0.0527 0.1429 0.0000 0.1429 0.1819 5.2500 18 0.2907 0.0000 0.1107 0.1446 0.1338 0.0108 0.9778 0.9890 0.9981 0.0255 0.0111 0.0019 0.0968 0.0333 0.0303 0.0747 6.2143 19 0.0000 0.3750 0.0000 0.1528 0.1250 0.0278 0.9487 1.0000 0.9487 0.0527 0.0000 0.0527 0.1429 0.0000 0.1429 0.1819 2.2500 20 0.0000 0.3750 0.0000 0.1528 0.1250 0.0278 0.9487 1.0000 0.9487 0.0527 0.0000 0.0527 0.1429 0.0000 0.1429 0.1819 2.2500 21 0.0000 0.3750 0.0000 0.1528 0.1250 0.0278 0.9487 1.0000 0.9487 0.0527 0.0000 0.0527 0.1429 0.0000 0.1429 0.1819 2.2500 22 0.0000 0.0000 0.1107 0.0384 0.0369 0.0015 1.0000 0.9981 0.9981 0.0000 0.0019 0.0019 0.0000 0.0303 0.0303 0.0391 12.0000 23 0.0000 0.0000 0.1107 0.0384 0.0369 0.0015 1.0000 0.9981 0.9981 0.0000 0.0019 0.0019 0.0000 0.0303 0.0303 0.0391 12.0000 24 0.0000 0.3750 0.0000 0.1528 0.1250 0.0278 0.9487 1.0000 0.9487 0.0527 0.0000 0.0527 0.1429 0.0000 0.1429 0.1819 2.2500 All loci 0.1159 0.2682 0.1396 0.1924 0.1746 0.0178 0.9753 0.9739 0.9761 0.0255 0.0295 0.0246 0.0759 0.0427 0.0612 0.1001 4.9104 St. dev 0.1650 0.1561 0.1878 0.1291 0.1254 0.0209 0.0244 0.0715 0.0271 0.0252 0.0822 0.0281 0.0619 0.0896 0.0545 0.0744 384.9570
48
Chapter 4. Population Structure
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19
4899
3639
2799
1918
1515 1412
1164
992
710
501
489 404
331 242 147 111
Figure 4.2: PCR fingerprint patterns of genomic DNA of R. alismatis isolates from the Coleambally Irrigation Area (CIA). Lanes 3–6 represent the 4 isolates RH001, RH57, RH123 and RH124 of genotype 1, lanes 7–12 represent the six isolates RH46, RH54, RH100, RH121, RH122 and RH149 with unique genotypes 2–7, lanes 13–14 represent the 2 isolates RH21 and RH91 of genotype 8, lanes 15–17 represent the 3 isolates RH24, RH25 and RH47 of genotype 9, and lanes 18–19 represent the two isolates RH55 and RH127 of genotype 10. Lanes 1 and 2 contain DNA molecular weight markers SPP1/EcoRI and pUC19/HpaII respectively; fragment sizes are shown in bps as indicated on the left.
Table 4.5: Pairwise comparisons of genotype diversity (Stoddart & Taylor 1988, above diagonal) and genetic distance (Nei 1972, below diagonal) among R. alismatis populations from different geographic regions of New South Wales. Population CIA CST SR CIA ….. 1.6258a* 0.0000 (23)b (32) CST 0.0255 ….. 1.4265* (23) SR 0.0295 0.0246 ….. aStudents t–test values. bdegrees of freedom. * indicates significance at P = 0.100. CIA, Coleambally Irrigation Area; CST, Central and Southern Tablelands; SR, Southern Riverina.
49 Chapter 4. Population Structure
4.6 Discussion Three populations of R. alismatis originating from different geographic locations throughout New South Wales and separated by distances of up to 300 km, were surprisingly similar by all measures used for comparison. The average population differentiation (Gst = 0.1001) and genetic distance (D = 0.0265) between populations were surprisingly small. The average gene diversity within (Hi = 0.1746) and between (Dst = 0.0178) populations were moderate to low, and normalised genetic identities (I) between populations across all loci averaged greater than 0.97, compared with the theoretical maximum of 1.00 that is reached when two populations have the same alleles present at the same frequencies. Despite these similarities, significant differences were observed in the frequencies of alleles at eight of the 24 loci. However, at more than half of these loci the differences in allelic diversity were solely attributed to the CST population and probably resulted from differences in sample size and sampling strategy between the populations. The larger spatial area over which infected leaves were sampled and larger distances between sampling points in this population probably decreased the likelihood of sampling the same clone as a result of splash dispersal of conidia. Alternatively, the small sample sizes in our study, especially for the CST population, whilst appearing large enough to detect low frequency alleles, may have underestimated the frequency of such alleles in the population and increased the overall measure of diversity at several loci. Because the primary inoculum of R. alismatis consists of conidia dispersed by rainsplash (Fox et al. 1999), and the teleomorph has not yet been identified (Braun 1995), it was expected that populations of the fungus would display a low level of genetic diversity, but a large degree of population differentiation because of limited potential for long−distance dispersal of conidia (Fitt et al. 1986; 1989). However, little evidence of significant population subdivision was observed, with overall Gst values across the 24 loci averaging only 10% between the three populations and < 8% for all pairwise comparisons. Two hypotheses were considered to explain the high degree of genetic similarity between these populations. One hypothesis is that natural selection has resulted in the same alleles achieving similar frequencies in all populations. This hypothesis assumes that selection for DNA sequences at loci themselves or for genes that are linked to each locus occur similarly in all populations, such that the same alleles are present in each population at the same frequencies (McDonald et al. 1996). However, this would require that the same mechanisms of selection occur in each population despite differences both in climate and host specificity, which is considered unlikely for three isolated populations.
50 Chapter 4. Population Structure
An alternative hypothesis is that substantial gene flow has occurred between the three geographic populations. Under this hypothesis the island model of gene flow (Wright 1951; 1969) can be used to estimate the number of individuals that would be required to have successfully migrated between the populations to account for their degree of similarity. Estimates of Nm under this model ranged from approximately one to 1898 for individual loci (Table 4.4), which on the basis of an average Gst of 0.1001 across all 24 loci, would require the movement of five individuals per generation to account for the degree of genetic similarity between the populations. Because the movement of only one individual per generation is adequate to prevent populations from diverging significantly by genetic drift (Wright 1931), this level of gene flow is sufficient to make these geographically separated populations a coevolving unit. Several mechanisms could facilitate gene flow between populations of R. alismatis. According to Osbourn et al. (1986), efficient migration is common in fungi even with splash dispersed conidia. However, since conidia dispersed by rainsplash are not expected to travel long distances, it is unlikely that populations are of R. alismatis are linked by the movement of conidia. Similarly, airborne dispersal of ascospores possibly from an undescribed teleomorph seems unlikely considering the sexual stage has not yet been identified in R. alismatis (Braun 1995) or the forme species R. secalis (McDonald et al. 1999). In a recent host range study, Cother (1999) suggested a role for alternate hosts in the epidemiology of R. alismatis. During this study 28 species of aquatic plants in the Alismataceae and related families and 39 cultivars of 25 agricultural plant species were tested for their reaction to inoculation with conidial suspensions of R. alismatis. However, whilst scattered infrequent lesions were reported on several species, no evidence of disease progression or influence on plant growth and development was observed. Furthermore, although the pathogen was reisolated from several members of the Cucurbitaceae and from the soybean cultivar Bowyer, the fungus failed to sporulate on many of the potential hosts likely to be found in the vicinity of rice fields, indicating that alternative species likely played little role in facilitating gene flow in this organism. Recently, the role of infected seed and plant material during the dissemination of R. alismatis was considered, when Fox et al. (1999) demonstrated that infection of D. minus leaves and inflorescence stalks by R. alismatis, weakens the scape, usually at water level, causing it to collapse before seed maturation. Fox (1995) also detected the pathogen at low levels both on and within seed of D. minus, suggesting that given the movement not only of people and farm machinery, but also water via irrigation, gene flow between populations could be facilitated over considerable distances up to hundreds of kilometres through dispersal of infected seed and plant materials.
51 Chapter 4. Population Structure
Coincidentally a recent publication by Lee et al. (2001b), in which the asymptomatic infection of barley seed by R. secalis is reported, now suggests this may be a common mechanism of gene flow within this genus. Unfortunately indirect measures of gene flow cannot be used to determine the time frame over which gene flow has occurred and these data do not indicate whether gene flow occurs over the course of decades, centuries or millennia (Slatkin & Barton 1989). Whilst the results of this study indicate that gene flow occurs over long distances, most likely through the dissemination of infected seed or plant materials, they provide no indication of the historical context by which this occurs. However, because Riverina populations of R. alismatis had fewer private alleles and lower levels of gene diversity, it is likely that these populations undergo regular reductions in population size (bottlenecks, as a result of seasonal host distribution) that eliminate many alleles, and that gene flow through regular extinction and recolonisation events (Maruyama & Kimura 1980; Olivieri et al. 1990; Wade & McCauley 1988) is the most likely explanation for the genetic similarity of R. alismatis populations in New South Wales. To test this hypothesis further, the spatial distribution of genotypes within and between populations, which can provide information about the dispersal potential of the organism in the field (Kohli et al. 1995; Chen et al. 1994), were investigated. Although populations showed few differences with respect to their genes, there were significant (P = 0.100) differences in pairwise comparisons of normalised measures of genotype diversity between populations (Table 4.5). Whilst both the CIA and SR populations displayed relatively low levels of genotype diversity, the CST population displayed high levels of genotypic diversity, with seven genotypes identified among eight isolates, which is 80% of the theoretical maximum. In most cases isolates with the same genotype were sampled within the same population, suggesting that asexual spores rarely travel more than a few kilometres over the course of a season. However, several DNA fingerprints were shared among isolates from different populations indicating that some genotypes are widely distributed over distances of at least several hundred kilometres and are present at higher frequencies. In one case eight isolates with identical genotypes were identified. Five of these isolates originated from the SR population, whilst the remaining three were distributed between the CIA and CST populations. The presence of identical genotypes or clones within all three populations, confirmed that gene flow over significant distances occurs between the populations. However, the directional movement of individuals between populations remains unknown.
52 Chapter 4. Population Structure
The lower estimated Gst values between the CIA and SR populations compared to other pairwise comparisons, however, indicated that the populations from the Riverina were genetically more isolated, despite their closer geographic origins, than the CST population. An additional consequence of regular extinction and recolonisation processes is that propagules originating from a founding population act to reduce local differentiation (Slatkin 1977). Because Gst estimates between Riverina populations seem to reflect this restriction, it is likely under the adopted hypothesis that propagules ‘founding’ these populations originate from outside the Riverina area, possibly contributing to the diversity of these populations on a seasonal basis in conjunction with annual agricultural practices. Additionally, because gene diversity is based on the frequency of alleles at each locus, and is affected by the age of the population, population size and selection, populations that have evolved over long periods at one location are expected to have more unique alleles and thus higher levels of gene diversity than populations that have moved into an area more recently (Goodwin 1997). Because the CST population has a higher proportion of unique alleles, higher gene diversity and is outside agricultural regions where it is likely to be affected by agricultural practices, it is most likely also older in origin than those in the Riverina. Whilst the possibility that gene flow through sharing of genotypes between Riverina populations either through the movement of people or machinery cannot be eliminated, no explanation for the presence of clones some 300 km east of this region is evident from this scenario. Hence current data suggests that these populations may share a common source of inoculum. Moreover the locations of both the CIA and SR populations, which are associated with major sources of irrigation directly downstream of the CST region, provide overwhelming evidence that watercourses disperse infected seed and/or infected plant materials sufficiently to represent effective sources of long distance gene flow. For the most part the results of this study are particularly favourable with respect to the suitability of R. alismatis as a candidate for control of Alismataceae weeds in Australian rice fields. Because a teleomorph has not yet been identified in R. alismatis, population structures of this organism are thought to result exclusively from asexual reproduction. In fact the identification of several clonal lineages and relatively low levels of gene and genotype diversity within R. alismatis populations supports this assumption. As a consequence of this purely asexual lifestyle, clonal populations often also have limited spectrums of virulence (Zeigler et al. 1995). However, unlike their sexually derived counterparts, this spectrum, as reflected by measures of genetic distance, is often a reliable indicator of the likely pathogenic variation between a population’s individuals (Leung et al. 1993; Francis & St. Clair 1997).
53 Chapter 4. Population Structure
Hence, it maybe conceived that the relatively small genetic distances between populations of R. alismatis, which averaged less than 3% for all pairwise comparisons across the 24 loci, may be reliably reflected by equally insignificant differences in the pathogenicity attributes of individuals. Consequently, pathogenicity tests of 40 isolates against both host and non−host Alismataceae species, conducted during a later part of this project revealed negligible differences in the pathogenicity of individual isolates (Chapter 5), and similar associations between pathogenicity and genotype due to asexual reproduction have also been reported in rust fungi (Burdon & Roelfs 1985) and in the scald pathogen R. secalis (Goodwin et al. 1992). For biocontrol candidates such as R. alismatis, unexpected dispersal, which poses potential risks to non−target plant species (Watson 1994), and the likelihood of genetic exchange with fungal species found within the intended range (Weidemann 1992) are major risk components. However, the limitations imposed by splash dispersal of conidia, combined with the limited host range and evolutionary potential of R. alismatis as regards to pathogenicity, suggest that the application of infective propagules of this agent pose negligible risks to crops grown adjacent to, or in rotation with rice in this country. Additionally, the absence of a sexual cycle and a population structure that may be contrived through regular rounds of clonal reproduction suggests there is little diversity from which genes may be acquired to contribute to an expansion in the pathogenic ability of this fungus even in the presence of moderate gene flow. Furthermore, despite the opportunity for asexual gene exchange via heterokaryosis or mitotic recombination remains, neither of which has been reported in this organism, there is little evidence of prior recombination between individuals. With a population structure that is more indicative of regular clonal reproduction supplemented by low levels of gene exchange from a ‘founding’ population, the opportunity for genetic exchange with other fungal species native to the release environment is minimal. Unfortunately, whilst R. alismatis poses a minimal threat to non–target species and is limited in its capacity for long distance dispersal, the minimal evolutionary potential of this organism also suggests that it is unlikely to evolve pathogenicity towards Sagittaria montevidensis and S. graminea, which are becoming increasingly widespread throughout the rice crops of southern New South Wales. Nevertheless, recent demonstrations of the pathogenicity of R. alismatis to several species of Sagittaria, recognised as important weeds of rice elsewhere in the world (Cother 1999), suggests that R. alismatis has the ability to infect species within this genus. Hence, more detailed studies of the interactions between R. alismatis and these species will be required before factors limiting the progression of disease can be identified.
54 Chapter 5. Infection Process
5 Infection Process
The behaviour of R. alismatis during interactions with host and non−host Alismataceae species and host structural responses
5.1 Introduction Due to the complexity of the plant structures, many fungi differentiate specialised infection structures in order to infect their hosts. In general, a spore lands on a host surface, attaches to the cuticle, produces a germ tube which recognises suitable penetration sites, and a bulbous structure called an appressorium is formed, during which the synthesis of polymer−degrading enzymes is often initiated. The deposition of additional wall layers in some fungi also suggests that the appressorium supports considerable pressure during the penetration process. Eventually, a narrow penetration hypha develops below the appressorium which exhibits additional features that serve to pierce the plant cuticle and cell wall. Following invasion, compatible and incompatible interactions between pathogen and potential host are established. In the latter, characteristic resistance reactions may take place, and subsequent mycelial proliferation within the plant results in a predictable destructive and observable effect. This chapter considers the mechanisms and processes by which fungal pathogens adhere to plant surfaces, germinate to form specialized infection structures and subsequently penetrate and colonise plant tissues. Initially, literature pertaining to the infection process will be considered in three sections; fungal attachment and/or adherence to the plant surface, germination and differentiation of infection structures, and fungal penetration and pathogenesis. A brief discussion of the common reactions elicited by hosts and non−hosts in response to fungal infection will then be provided. Finally, a detailed analysis of the infection process of R. alismatis on the host species A. plantago– aquatica and two non–host species, S. graminea and S. montevidensis will be presented. Where appropriate, factors which may affect disease progression in non–host species are discussed.
55 Chapter 5. Infection Process
5.2 Fungal attachment and/or adherence to the plant surface Attachment of fungal spores and germ tubes to the host surface is universally recognised as an essential pre−infection event that determines the success of infection (Nicholson & Epstein 1991; Mendgen & Deising 1993). Adhesion is believed to be involved in the initial recognition of the host surface and probably contributes to preparation of the infection court (Kunoh et al. 1988; Nicholson 1990; Nicholson & Epstein 1991; Nicholson & Kunoh 1995). Traditionally, fungal attachment to the plant host was believed to result from entrapment of spores on the leaf surface (Nicholson 1984). However, several researchers (Caesar−TonThat & Epstein 1991; Deising et al. 1992; Braun & Howard 1994) suggested that spores of fungal pathogens adhere to the plant surface through the secretion of an adhesive matrix. For example, Pascholati et al. (1993) identified and characterised several extracellular enzymes including proteases, cellulases, pectinases and non−specific esterases in an adhesive discharge produced by spores of Colletotrichum graminicola during infection of maize leaves. Despite their suspected role in fungal pathogenesis, Deising et al. (1992) showed that esterases and cutinases found in spores of the broad bean rust fungus, Uromyces viciae−faba, are essential for adhesion of infection structures to the leaves of broad bean. Prior to penetration cutinase and other serine esterases are localised on the surface of spores. Consequentially if the enzymes are removed from the spore surface or if a serine esterase inhibitor is added, adhesion is significantly reduced. On the other hand, addition of cutinase and the serine esterases to autoclaved spores restored their ability to adhere to the plant surface (Deising et al. 1992). Additionally, Hamer et al. (1988) demonstrated that spores of the rice blast fungus Magnaporthe grisea possess a definitive mechanism of attachment to the host surface prior to germination. Using phase contrast microscopy Hamer et al. (1988) identified a preformed mucilaginous substance released from the conidial apex of freshly harvested spores of M. grisea. The adhesive material, called spore tip mucilage is released following hydration of spores prior to the emergence of germ tubes and facilitates the attachment of the spore tip to the hydrophobic leaf surface. The composition and function of fungal adhesives are heterogenous, but many consist of water insoluble glycoproteins, lipids and polysaccharides (Xiao, Ohshima et al. 1994; Nicholson 1996; Sugai et al. 1998). The chemical composition and function of a wide range of fungal adhesives has been summarised by Jones (1992) and includes citations from relevant literary sources. More elusive adhesive materials which have received considerable attention recently (Wessels 1996; 1997; 1999; Talbot 1999; Kershaw & Talbot 1998) and which may allow fungal pathogens to interact with hydrophobic plant surfaces, are hydrophobins, small hydrophobic proteins that appear to prepare sites on the leaf surface for attachment of the appressorium (Beckerman & Ebbole 1996).
56 Chapter 5. Infection Process
Talbot et al. (1996) constructed a hydrophobin negative M. grisea through mutation of the MPG1 gene which encodes a 15 kDa hydrophobin protein produced during appressorium formation. Mutants failed to attach to leaf surfaces and loss of hydrophobin activity affected both the production of appressoria and their ability to penetrate the plant epidermis. To date, fungal adhesion has been demonstrated in many species of Colletotrichum (Mercure et al. 1994), Botrytis (Doss et al. 1993; Doss et al. 1995), Nectria (Jones & Epstein 1989; 1990), rusts (Mendgen 1978; Gold & Mendgen 1984; Staples & Hoch 1987), and powdery mildew fungi (Kunoh et al. 1988). Additionally, numerous reports have arisen regarding adhesion of spores to artificial surfaces (Young & Kauss 1984; Beckett et al. 1990; Read et al. 1992), and the role of adhesion in fungal pathogenesis is the subject of constant review (Kunoh et al. 1991; Nicholson & Epstein 1991; Jones 1992; Hardham 1992; Braun & Howard 1994; Mendgen 1996; Epstein & Nicholson 1997; Tucker et al. 2001).
5.3 Germination and differentiation of infection structures The first event both in the formation of a new fungal colony and infection of a plant by a fungal pathogen is the germination of a spore. Spores are equipped not only with provisions for germination and continued growth and development, but also with the means for detecting and responding to external stimuli whereby a favourable environment may be encountered (Allen 1976). Once a propagule has germinated, the germ tube extends to form a highly specialised infection structure called an appressorium (Jones 1992). Similarly, appressoria, differentiated by many fungal species as a means to gain access to the nutrient sources of the plants they parasitise, are also induced in response to specific physical and chemical cues provided by the host (Deising et al. 2000). Coined in 1893, the term ‘appressorium’ originally referred to the ‘adhesive organs’ first observed by Frank (1893) as ‘swellings’ at the apical regions of germ tubes of C. lindemuthianum. Extended by Emmett and Parberry (1975) to include ‘all structures adhering to the host surface to achieve penetration’ regardless of morphology, appressoria have long been accepted as the structures by which many fungi mediate penetration (Frank 1893, cited by Deising et al. 2000). Unfortunately, whilst appressoria are known to facilitate penetration of the host by mechanical force (Emmett & Parberry 1975) or with assistance by chemical or enzymatic action (Cooper et al. 1988), the role of the appressorium in adhesion to the host is often overlooked. Adhesion, irrespective of the mode of penetration is crucial to the success of the infection and is necessary to counteract the force of the emerging penetration peg as it protrudes from the appressorial base (Deising et al. 2000).
57 Chapter 5. Infection Process
Pathogens respond to numerous signals to recognise, adhere to and invade their plant hosts including cues that regulate germ tube growth and appressorium development. The fact that appressoria often form equally well on susceptible and resistant hosts and non−hosts, artificial and plant surfaces and both living and dead tissue indicate that specificity may be unimportant (Parberry & Blakeman 1978). Many researchers have contributed to an understanding of these processes (Johnson 1934; Dickinson 1949; Hoch & Staples 1987; 1991; Hoch, Staples & Bourett 1987; Macko et al. 1978; Maheshwari et al. 1967; Staples et al. 1983; Wynn 1976; 1981; Wynn & Staples 1981; Allen, Hazen et al. 1991), but for the most part the signals that induce germination and appressorium formation during plant–pathogen interactions are poorly understood. Nevertheless, some important cues have been identified. For example, urediospores of many rust fungi respond to physical (thigmotropic) signals, forming appressorium over stomatal pores (Hoch, Staples, Whitehead et al. 1987; Terhune et al. 1991) and maximizing the likelihood of locating stomates by extending across the leaf surface at right angles to leaf veins and other structures in response to morphological and ultrastructural aspects of the leaf surface (Read et al. 1992). The ridges created by stomatal guard cell lips (Wynn 1976) and the wax crystal lattice have been implicated in this response (Lewis & Day 1972). The frequency and height of the ridges on the leaf surface also appear significant (Hoch, Staples, Whitehead et al. 1987). Similarly, surface hardness, hydrophobicity and perhaps porosity have been implicated as physical signals in the induction of appressorium formation in Colletotrichum and Magnaporthe (Lapp & Skoropad 1978; Jelitto et al. 1994; Lee & Dean 1994; Xiao et al. 1994). Surface thickness was also found to greatly influence appressorial formation in C. gloeosporioides during infection of avocado (Coates et al. 1993). Naturally occurring components of the plants surface including waxes, cutin derived fatty acids, phenols and other volatiles have been recognised as chemical inducers (chemotropic signals) of appressorium formation (Deising et al. 2000). For example, chemical exudates washed from the surface of pepper fruit have been shown to stimulate germ tube differentiation in C. piperatum (Grover 1971). Similarly, surface waxes from host avocado fruits, rich in long chain fatty alcohols, stimulate both germination and appressorium formation in C. gloeosporioides (Podila et al. 1993), and epicuticular waxes in rice leaves promote appressorium formation in Pyricularia oryzae (Kumar & Sridhar 1987). Non−host surface waxes were found to be inhibitory in both cases. Kolattukudy et al. (1995) suggested that leaf exudates often contain both inducers and inhibitors of germination, the balance of which is likely responsible for the selective signaling during host−parasite relationships.
58 Chapter 5. Infection Process
Other chemical signals include simple sugars, K+ (Staples 1985), Ca2+ (Warwar & Dickman 1996; Kim et al. 1998), cAMP (Lee & Dean 1993; Mitchell & Dean 1995; Yang & Dickman 1999; Kronstad 1997) endogenous (Grover 1971) and exogenous compounds (Emmett & Parberry 1975) and hormones (Flaishman & Kolattukudy 1994). Recently, a role for protein kinase A (PKA) in the mobilization of carbohydrates and synthesis of osmoticums such as glycerol, accumulated during appressorial melanisation has been suggested (De Jong et al. 1997). The role of thigmotropic and chemotropic responses in fungal pathogenesis is considered in more detail in several recent reviews (Kolattukudy et al. 1995; Mendgen et al. 1996; Read et al. 1997; Hahn et al. 1997; Dean 1997; Knogge 1998; Hamer & Talbot 1998). Additionally, the role of hydrophobins as recognition signals during appressorium differentiation in addition to adhesion, considered earlier, has been suggested (Beckerman & Ebbole 1996). Talbot et al. (1996) speculated that polymerisation of such surface proteins could induce appressorium morphogenesis. Environmental stimuli including nutrient status (Xiao, Ohshima et al. 1994), free moisture, temperature, light (Emmett & Parberry 1975) and pH (Edwards & Bowling 1986) have also been considered, but freedom from internal and external constraints in the presence of a favorable environment seems like an oversimplification.
5.4 Fungal penetration and pathogenesis Subsequent to the differentiation of infection structures, fungal pathogens can infect their hosts by direct penetration of the outer cell layer or by entering natural openings such as stomata or wounds. Whether direct penetration is accomplished by the physical force of the growing hyphal tip or through enzymatic degradation of the plant cuticle and cell wall is unresolved (Schafer 1994). However, a combination of both physical and enzymatic means has been proposed (Kolattukudy 1985; Walton 1994; Money & Howard 1996). To date, the role of enzymes during fungal pathogenesis remains controversial and few studies provide conclusive evidence for their involvement during plant infection. Nevertheless, all the major groups of cellular plant pathogens are known to produce extracellular enzymes capable of degrading plant structures (Walton 1994), and as such the role of cell wall degrading enzymes (CWDE) in fungal pathogenesis has been reviewed extensively (Bateman & Basham 1976; Kolattukudy 1985; Walton 1994; De Lorenzo et al. 1997; Annis & Goodwin 1997; Ten–Have et al. 2002).
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Some enzymes identified to date include protease (Porter 1969; Ball, Ashby et al. 1991), cellulase (Muller et al. 1997; Pryce–Jones et al. 1999), endo− and exopolygalacturonase (Walton & Cervone 1990; Yao et al. 1996; Zhang et al. 1997) invertase (Louis & Cooke 1985b), pectinase (Cleveland & McCormick 1987), non−specific esterase (Nicholson & Moraes 1980), β−glucosidase (Ramadoss et al. 1985) β–1,3–glucanase (Schaeffer et al. 1994; Tenberge et al. 1999) catalase (Van der Vlugt– Bergmans, Wagemakers, Dees et al. 1997), pectate lyase (Wattad et al. 1997), xylanase (Giesbert et al. 1998) and a range of others (Wubben et al. 1999; 2000). Until very recently proof of the involvement of CWDE during fungal pathogenesis was lacking even despite the reports of researchers like Mendgen and Deising (1993) who demonstrated that enzymes in the extracellular matrix of Colletotrichum species were responsible for altering the plant surface underneath conidial germ tubes and appressoria. However, in 1997 Shieh et al. (1997) confirmed the involvement of CWDE during fungal pathogenesis when they provided unequivocal evidence of the involvement of endopolygalacturonase in the infection of cotton bolls by Aspergillus flavus, and became the first of a select group to demonstrate a phenomenon that had previously only been observed in bacteria (Roeder & Collmer 1985; Hugouvieux–Cotte–Pattat et al. 1996). To date, only six studies including the pioneering work of Shieh et al. (1997) have been published in which the definitive role of CWDE in fungal pathogenesis has been demonstrated. In addition to A. flavus the fungi involved are Botrytis cinerea (Ten–Have et al. 1998), Nectria haematococca (Rogers et al. 2000), Alternaria citri (Isshiki et al. 2001), C. gloeosporioides (Yakoby, Beno–Moualem et al. 2001) and Claviceps purpurea (Oeser et al. 2002). For many phytopathogens however, the first association with plant organs involves contact with the plant cuticle, which is composed of a structural polymer called cutin (Kolattukudy 1984a). Responsible for degrading this compound is the enzyme cutinase, a lipolytic enzyme produced by a range of phytopathogens (Walton 1994). Unlike the role of CWDE however, the role of cutinase in host infection was confirmed very early in the study of host pathogen interactions with several groups demonstrating the involvement of this enzyme during host penetration (Dickman et al. 1989; Kolattukudy et al. 1989). Whilst the role of cutinase is pivotal to any discussion of plant pathogenesis, it will not be dealt with here in detail. Instead a detailed discussion will be provided in the ensuing chapter, in which the involvement of this enzyme in host penetration will be more adequately demonstrated.
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Whilst experimental proof of the involvement of enzymes remains elusive (Schafer 1993), it has been apparent since the pioneering work of Miyoshi (1895) that some fungi can penetrate plant cuticles physically. Light microscope studies revealed that some fungi including species of Colletotrichum (Kubo et al. 1982; 1984; 1987; Wolkow et al. 1983a), Magnaporthe (Howard et al. 1991; Howard & Valent 1996; Chumley & Valent 1990) and Pyricularia (Okuno et al. 1983; Woloshuk et al. 1980; Yamaguchi et al. 1983) produce appressoria with tough melanin−pigmented cell walls. Melanins are heavily pigmented polymers that primarily function to protect the fungal mycelium from UV radiation, oxidants and antifungals (Bell & Wheeler 1986; Butler & Day 1998; Henson et al. 1999; Butler et al. 2001). However, melanin biosynthesis also plays a role in the penetration of host surfaces by allowing appressoria to establish and maintain high internal hydrostatic pressures (Howard & Ferrari 1989). Pressure is generated through the influx of the cytoplasmic osmoticum, glycerol, which is generated rapidly during fungal germination and germ tube elongation (De Jong et al. 1997). Because melanised appressorial walls are largely impermeable to glycerol, the maintenance of enormous glycerol concentrations within appressoria is likely to be a consequence of this reduced permeability (Money 1997). Appressoria produced by melanin−deficient mutants of Colletotrichum and Magnaporthe were extremely permeable to glycerol, failed to generate turgor (De Jong et al. 1997) and were non−pathogenic (Suzuki et al. 1982; Kubo et al. 1985; Kubo & Furusawa 1991; Chumley & Valent 1990; Howard et al. 1991; Howard & Valent 1996; Mendgen et al. 1996). Despite these studies, however, direct evidence that appressoria can produce enough force to penetrate plant cuticles has been lacking until very recently when Bechinger et al. (1999) measured the force exerted by C. graminicola, the causal agent of anthracnose of maize. This force was found to be approximately 5.4 MPa, which was slightly less than the 6−8 MPa reported by Howard, Ferrari et al. (1991) for M. grisea, but still equivalent to 30 or 40 times the pressure of an average car tire. Whilst this is a colossal force for such a miniscule cell and an equally impressive technical effort on behalf of Bechinger et al. (1999), the mechanisms by which turgor is translated to physical pressure and applied at the penetration peg remain unknown (Money 1997). However, fortunately one thing seems clear and that is the innumerable requirement of melanin deposition during appressoria formation where a purely mechanical induced penetration of plant cuticles is initiated. Money (1995) and Frederick et al. (1999) agree that fungal turgor pressure may be an obligatory requirement for penetration and ramification through plant tissues, perhaps where the absence of sufficient levels of lytic enzymes fail to liquefied the cuticle, cell walls and other obstructions.
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5.5 Plant defences In general, the vast majority of plants are resistant to fungal invasion. Resistance (used here synonymously with non−susceptibility and incompatibility) is usually shown by all members of a particular species, which are regarded as non−hosts, but may be exhibited by varieties and cultivars of otherwise susceptible plants regarded as hosts (Heath 1980, 1981b). In non−susceptible hosts and non−hosts, resistance to pathogenesis is usually accompanied by some form of reaction from the plant. Such reactions have been described in whole or in part by numerous authors (Currier 1957; Kosuge 1969; Kuc 1972; Aist 1976; Vance et al. 1980; Nicholson & Hammerschmidt 1992; Bennett & Wallsgrove 1994; Dixon et al. 1994; Heath 1998; 2000b; Kombrink & Schmelzer 2001), but the reviews by Heath (1980, 2000a), Ride (1992) and Rioux & Biggs (1994) which deal specifically with non−host systems are particularly interesting, and the following section will be devoted to characteristics of non−hosts that appear to be implicated in their resistance to disease. A short discussion of the ‘specific accommodation’ exhibited by susceptible hosts will also be included. A non−host species avoids disease not only through resistance factors, but also due to evasion (Heath 1984). Because every plant is unlikely to possess a characteristic specific defence to defend itself from the assortment of potential invaders, plants likely possess a range of defence mechanisms including both preformed deterrents and an armoury of induced responses that they can initiate either singularly or as a combined arsenal during invasion (Heath 1981c; 1981e). Furthermore, because the cuticular structure is similar in most plants (Martin 1964), many such defence mechanisms may be non−specific, passive, constitutive properties of the potential host. Alternatively, defence mechanisms may be induced by fungal invasion. Consequently, the pathogen must avoid, inhibit or tolerate resistance mechanisms in its hosts (Heath 1987; 2000a). Constitutive or putative defence mechanisms are the most common form of non−host resistance (Heath 1984). Preformed defences generally involve surface features of the plant which may trigger ‘miscues’ or incorrect behaviour on behalf of the pathogen, inhibiting or preventing critical stages in pathogenesis. Defence may depend on the absence of specific characteristics essential for differentiation and development in the pathogen, as opposed to inhibitory attributes. For example, rust fungi respond primarily to the physical topography of the leaf surface (Wynn 1976). The absence of suitable topographical signals may prevent germ tubes from locating the stomates they need to enter the leaf. This type of physical recognition is reviewed in detail by Read et al. (1992).
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Alternatively, structural features may act as physical barriers. For example, the plant cuticle represents a substantial barrier to wound pathogens such as Mycosphaerella that are unable to breach the intact cuticle due to a failure to produce the enzyme, cutinase (Dickman et al. 1989). In some cases, cuticle thickness has been correlated with increased passive resistance (Koller 1991). Additionally, lignified tissues can prevent fungal growth through resistance to enzymes typically produced to degrade plant cell walls (Heath 1984). Thus far few chemical constitutive defences have been identified with any certainty. Break down products of cutin and suberin (Kolattukudy & Espelie 1985), and a multitude of enzymes (Hoagland 1990) have been implicated. Nutrient availability seems appropriate, but has rarely been considered essential (Kessler 1966), and the resistance of unripe fruit in relation to the susceptibility of ripe tissues seems to indicate the presence of fungistatic compounds, at least in a transient sense (Prusky et al. 1981; Prusky & Plumbley 1992). Unfortunately, such preformed defences rarely restrict all members of a pathogen population and other defence mechanisms, usually induced by the fungus, must also contribute to the resistance of the plant. Conveniently, induced defences can be grouped into three main types; cell wall alternations, antifungal proteins and hypersensitive reactions. As indicated terminologically ‘induced’ defence mechanisms rely on recognition of pathogens by host cells and involve specific fungal molecules called ‘elicitors’ (Keen 1982). Whilst several extensive reviews have been published on these topics (Daly 1984; Hoch & Staples 1991; Ouchi 1984; Ride 1992; Dixon et al. 1994; Kolattukudy et al. 1995), including the recent review by Dean (1997), much of this research, although relevant, is beyond the scope of this thesis and hence will not be dealt with here in detail. Briefly however, elicitors are substances that induce resistance responses in plants by acting as recognition signals between host and parasite. Elicitors fall into two categories, cell wall components and secreted enzymes (Ride 1992). Chitin, chitosan, β−glucans, glycoproteins and poly unsaturated fatty acids extracted from fungal cell walls have been implicated (Ouchi 1984; Heath 1984). Because they are essential structural components of cell walls of most fungi they presumably represent a source of recognition signals to plants, probably released from fungal cell walls through the action of plant enzymes (Yoshikawa et al. 1981). Chitosan specifically has been shown to induce the synthesis of callose (Kohle et al. 1985) and phytoalexins (Hadwiger & Beckman 1980; Keen et al. 1983). In addition to cell wall products, enzymes secreted by fungi appear also to act as resistance inducers through the release of endogenous elicitors from plant cell walls (Ride 1992). Pectic enzymes (West et al. 1985) and also xylanases have been implicated. However, the action of the latter appears to depend on structure rather than an ability to release cell wall components (Lotan & Fluhr 1990).
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As with constitutive defences, some induced changes have been assumed to act as mechanical barriers. One of the most conspicuous signs of infection in non−hosts following fungal invasion is the thickening of plant cell walls due to the deposition of materials between the plant cell wall and cell membrane (plasmalemma). These wall appositions referred to as papillae and synonymous with callosity, lignituber and callus (Aist 1976), are essentially wound plugs that form in response to injury but may serve as mechanisms of resistance to fungal penetration (Bracker & Littlefield 1973; Bushnell & Bergquist 1975; Griffiths 1971), by conforming to and encasing fungal structures (Aist 1976). Papillae vary from small, dense, amorphous deposits to large structures which contain vesicular and membranous elements (Politas & Wheeler 1973), and may be elicited by both non−pathogenic and pathogenic fungi (Sherwood & Vance 1980; Johnson et al. 1982). Papillae formation is often accompanied by the appearance of ‘haloes’ around penetration sites. Haloes are characterised by a circular area at the site of attempted or successful penetration and result from changes in cell wall chemistry that are visible after staining with histological reagents (Sherwood & Vance 1980). Several groups insist the processes of papillae formation and the appearance of haloes are metabolically related (Ride & Pearce 1979; Zeyen & Bushnell 1979); however, detailed information on the structure and chemical composition both of papillae and haloes is numerous and heterogenous. Callose (Aist & Williams 1971; Mercer et al. 1974; Sargent et al. 1973) and lignin (Fellows 1928; Sherwood & Vance 1976) are presumed to be the principal components and feature prominently following inoculation and wounding with pathogens and non−pathogens. Lignin, a biopolymer of coniferyl and sinapyl alcohols, is laid down in cell walls in response to microbial attack, forming an interpenetrating network resistant to enzymatic degradation and mechanically impenetrable to most microorganisms (Ride 1975; Vance et al. 1980). Similarly, callose, an aniline blue−fluorescing polysaccharide composed of (1−3) β−glucans forms an impermeable network, inhibiting the passage of small ions and molecules in papillae, implicated in sequestration of toxins and phenolics resulting in resistance both of papillae and ultimately the host to fungal penetration (Smart 1991; Skalamera & Heath 1996). The accumulation of cytoplasm at sites of penetration, presumably responsible for deposition of papilla material, has also been implicated in the death of cells and necrosis through the discharge of materials into underlying cells (Heath 1984). Additionally, several other substances have been localised in papillae and haloes including suberin, phenols, silicon and enzymes (Carver et al. 1998; Edwards 1970; Sherwood & Vance 1976; Aist 1976; Kunoh & Ishizaki 1975; Hargreaves 1982). Cellulase in particular has been implicated in the chemical dissolution of cellulose around penetration sites of C. lagenarium (Suzuki et al. 1981; 1983), resulting in haloes.
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Studies suggest fungal−induced cellulase plays a significant role in supplying nutrients necessary for the formation of the emerging penetration hyphae by chemically dissolving host cell wall components around infection sites (Suzuki et al. 1982). A universal response of resistant plants to fungal attack is the accumulation of stress−induced metabolites or phenolic compounds, some of which are referred to as phytoalexins (Heath 1984). Phytoalexins are low molecular weight, broad spectrum antimicrobial compounds induced in plants by stress and fungal invasion (Ride 1992) and, as with phenolics, their synthesis and role during the expression of ‘resistance’ has been reviewed extensively (Nicholson & Hammerschmidt 1992; Bennett & Wallsgrove 1994; Smith 1996). The demonstration that phytoalexins accumulate in the tissues both beneath and adjacent to sites of invasion by fungal pathogens (Snyder & Nicholson 1990) has strengthened views of the importance of phytoalexins in plant defence against fungal invasion. However, their infrequency in monocotyledonous species (Nicholson & Hammerschmidt 1992) puts this view in doubt. Nevertheless, studies show strong correlation between rapid phytoalexin biosynthesis and resistance to fungal infections in many plant species, including grapevines (Dercks & Creasy 1989), carnation (Baayen et al. 1991), citrus (Afek & Szetejnberg 1988) and soybeans (Hahn et al. 1985; Graham et al. 1990). The accumulation of phenolics and phytoalexins in infected tissues is also controversially suspected to be closely related to cell necrosis observed in incompatible plant−fungus interactions (Heath 1984). Other pathogenesis– related proteins exhibiting antimicrobial and antifungal activity include lectins, thionins and proteinase inhibitors, which have been reviewed in detail by Bowles (1990). In addition to the role of cell death in relation to phytoalexin accumulation, necrosis as a resistance mechanism has been debated continuously under the guise of hypersensitivity−related cell death, the much publicised plant based equivalent of programmed cell death common in animals (Heath 2000b). The hypersensitive response (HR) is the subject of much controversy, and perhaps ‘unhealthy focus’ according to Grant & Mansfield (1999), but has been debated feverishly since its inception almost a century ago and is the subject of continual review (Mittler et al. 1997; Gilchrist 1998; Heath 1998; 2000b; Kombrink & Schmelzer 2001). Extremely common in incompatible reactions, the HR results in cell death around infection sites and pathogen limitation, generally recognised by the appearance of brown, dead cells at infection sites (Heath 2000b), but may be expressed prior to fungal entry into cells, as miscues from the plant surface resulting in avoidance (Heath 1997). Unfortunately, while necrosis as a defence mechanism may be possible against biotrophic fungal pathogens, for non−biotrophic pathogens, cell death alone cannot restrict pathogen ingress (Heath 2000b). For necrotrophs this role must be ascribed to other processes inhibitory to fungal growth, some of which have been described above.
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Regardless of the mechanisms of non−host resistance, the ability of fungal pathogens to successfully parasitise their hosts relies on a different set of events that collectively result in ‘specific accommodation’ (basic compatibility) between the two organisms, and which ultimately reduces or eliminates the activity of defence responses (Wood 1976), through avoidance, inhibition or tolerance (Heath 1981d; 1987; 2000a). For example, experimental evidence for the production of enzymes (Schonbeck 1976), host selective toxins (Daly 1972) and other suppressors (Oku et al. 1979), that neutralise toxic compounds and/or kill cells prior to the expression of resistance, have been identified. By meddling with defence mechanisms, either through ‘passive’ or ‘active’ responses, the pathogen effectively induces susceptibility in its host, establishing ‘specific accommodation or compatibility and negating the presence of preformed or induced mechanisms’ (Heath 1981a). Why there is an absence of visible responses in hosts under invasion by compatible pathogens is unknown (Heath 1980; 2000b), but increasing evidence points strongly to some activity of the pathogen, as outlined above, rather than mere absence of induction of response mechanisms (Heath 1980). To date, many environmental and cultural aspects affecting fungal growth, sporulation and conidial germination and infectivity of R. alismatis have been documented (Jahromi et al. 1998). However, despite infecting a number of species in the Alismataceae including several species of Sagittaria from abroad (Cother & Gilbert 1994a), there are no records of R. alismatis on species of Sagittaria in Australia and detailed studies of the infection process of R. alismatis have been documented only during interactions with the native species D. minus (Jahromi et al. 2002). In order to identify constraints that limit disease initiation and progression in S. graminea and S. montevidensis an investigation of the infection process of R. alismatis on these species is required. Because the fungus may undergo differences in the modes of infection during each interaction, a better understanding of the factors affecting germ tube elongation, appressorial formation, host penetration and diseased progression in individual species will provide the knowledge necessary to identify specific constraints to infection and instigate methods to overcome them (Van Dyke 1989). Additionally, a knowledge of the infection process of biocontrol agents on target weeds is required to facilitate commercial registration of the product and differences in the capacity of pathogens to infect target species are likely to influence resulting improvement procedures. The native host A. plantago–aquatica was included in the study to provide both a positive control and informative comparisons between host and non−host species.
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5.6 Materials and Methods 5.6.1 Fungal isolates: Fungal isolates were collected, isolated, cultivated, stored and maintained by the methods described in section 3.4.1. The isolates used throughout this study were essentially the same as those listed in Table 4.1 however, isolates RH005 and RH149 failed to sporulate during inoculum preparation and hence were discarded from ensuing experiments.
5.6.2 Plant growth and leaf material preparation: All D. minus, A. plantago−aquatica, S. montevidensis and S. graminea plants used in this study were collected from field sites located within the rice−growing region of Southern New South Wales, transported to Charles Sturt University in plastic tubs and transplanted into 1100 mm diameter stock water troughs (Riverina Cooperative Society Ltd., Wagga Wagga, NSW, Australia) maintained in a temperature controlled glasshouse (day/night, 30/25°C) with diurnal light regime. The water level in the troughs was maintained at approximately 20 cm above the soil surface. Fully expanded leaves were excised from healthy adult plants and cut into spherical discs with a 2 cm diameter cork borer. Leaf discs were surface sterilised in a sodium hypochlorite solution (1% available chlorine) for 45 seconds, rinsed in sterile distilled water, blotted dry with filter paper and immediately transferred to 20 cm diameter plastic Petri dishes containing 1.5% Technical agar (Amyl Media Pty. Ltd.) supplemented with 1 µg/mL benzylaminopurine (Sigma−Aldritch).
5.6.3 Pathogenicity and selection for virulence 5.6.3.1. Inoculum preparation: Prior to the commencement of infection studies, fungal isolates were passaged through the host plant from which they were originally isolated, as indicated by Table 4.1, reisolated as described in section 3.4.1 and transferred to 9 cm diameter plastic Petri dishes containing LBA (Difco Laboratories). Following incubation at 25°C for 5−10 days under the conditions described in section 3.4.2, spores and mycelium were transferred to additional LBA plates to produce confluent stock spore cultures of each isolate. Plates were then incubated for an additional 4 days and spores harvested as described in section 3.4.2. Conidial suspensions were enumerated using a Weber haemocytometer (Crown Scientific Pty. Ltd., Burwood, Victoria, Australia), centrifuged at 10 000g for 30 seconds, the supernatant discarded and the concentration of spores adjusted to 1.0 × 106/mL by the addition of sterile distilled water.
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5.6.3.2. Leaf disc inoculation and disease assessment: The leaf disc bioassay method (Jahromi 2000) was employed to assess the virulence of 40 R. alismatis isolates against the four Alismataceae species. Because isolates were not expected to cause significant levels of disease on the non−host species S. graminea and S. montevidensis, the native host species D. minus, which was the topic of earlier studies by Jahromi (2000), was included in the pathogenicity trials to provide a better indication of the spectrum of virulence of individual isolates. Petri dishes containing leaf discs for each plant species were inoculated to ‘run–off’ with a conidial suspension distributed via a hand held atomiser, wrapped with Parafilm ‘M’ laboratory film (American National Can, Chicago, Illinois, USA) to avoid dehydration, and incubated at 25°C for 14 days under the conditions described in section 3.4.2. Control discs were inoculated with sterile distilled water. Note: each Petri dish contained only leaf discs of a single plant species. The percentage of leaf disc area affected by disease was determined visually following the conclusion of the experimental period. Lesion development was given a score ranging from 1 to 11 according to the Horsfall and Barrett (1945) disease severity scale. Three replicates were conducted for each plant/pathogen interaction and the scores from both methods were combined into a single numerical score for each replicate and averaged across the three experiments (Table 8.1). Based on these observations, isolate RH097 (DAR 73151) was the most pathogenic against all four of the plant species and was used throughout the remainder of the study.
5.6.4 Infection of host and non host species and Microscopy 5.6.4.1. Tissue preparation and inoculation: Leaf discs and inoculum were prepared as described in sections 5.6.3.1 and 5.6.3.2 respectively. Petri dishes containing leaf discs of A. plantago−aquatica, S. montevidensis and S. graminea were ‘point–inoculated’ with 10 µl of spore suspensions of isolate RH097 and incubated for 36 hours at 25°C under the conditions described in Section 3.4.2. The germination of conidia of R. alismatis and the process of infection on both host and non−host species was studied using light and electron microscopy.
5.6.4.2. Light microscopy: Samples were removed after 6, 12, 18, 24, 30 and 36 hours incubation. Six leaf discs for each species at each of the six intervals were fixed and cleared in 1:2 acetic acid/ethanol solution, stained with lactophenol cotton blue (phenol 20% w/v; lactic acid 20% v/v; glycerol v/v; cotton blue 0.1% w/v) and mounted on 40 × 20 mm glass slides with glycerol.
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Observations were made at 100× and 400× magnification using a Nikon Labophot Episcopic microscope fitted with EPlan objectives (Nikon Corporation; Nippon, Kogaku, K.K., Chiyoda−ku, Tokyo, Japan). One hundred conidia were examined on each of the six leaf discs comprising each treatment. The number of conidia that had germinated, the number of germinated conidia that had formed appressoria and the number of un−germinated conidia were determined. Areas where conidia were pooled in large concentrations were avoided to ensure consistency during the counting process. Conidia were considered to have germinated if the length of the germ tube was equal to, or greater than, half the length of the conidium (Lacy 1994), or if an appressorium was present. Appressoria were recognised by their globose, sometimes slightly lobed structure formed terminally on a germ tube (Emmett & Parberry 1975). The periodic acid−Schiff (PAS) reagent (Sigma−Aldrich) was used to test for appressorial melanisation. The protocol of Hotchkiss (1948) was employed. Tissue was oxidised in PA (~1% for 10−30 minutes), rinsed in distilled water and immersed in Shiff’s reagent for 10−30 minutes. Tissue was then de–stained in sodium metabisulphite, NaHSO3 or Na2SO3 (~0.5%). In addition to the photographs taken using the aforementioned microscopes, some additional photographs were taken at 40× magnification using a Zeiss Axioplan 2 MOT Imaging microscope equipped with an AxioCam high resolution digital camera and universal semi−apochromatic Plan−NEOFLUAR objectives (Carl Zeiss Microscopy, Göttingen, Germany).
5.6.4.3. Fluorescence microscopy: Six leaf discs of each species were removed at each of the six intervals and fixed and cleared by boiling in 1:2 lactophenol/ethanol solution for 2 minutes. Leaves were washed twice in 50% ethanol for 15 minutes, twice in 0.05M NaOH for 15 minutes and three times in distilled water before being immersed in a 0.1M Tris HCl buffer, pH 8.5 for 30 minutes. Leaves were stained with a 0.1% solution of Leucophor BMB (Sandoz, Australia Pty. Ltd.) in this buffer for 5 minutes, washed four times in distilled water for 10 minutes and once in 25% glycerol for 30 minutes (Rohringer et al. 1977). Leaves were mounted in glycerol containing a trace of lactophenol as preservative and examined with a Nikon Labophot Episcopic microscope with fluorescence attachment EF−D and equipped with a 100 watt mercury lamp and a HB−10101AF super high pressure mercury lamp power supply. A UV−2A filter block containing a DM400 dicroic mirror, BA420 barrier filter and a EX330−380 nm ultraviolet excitation filter was employed. Some additional photography was performed using BV−2A and B−2A filter blocks, containing DM455 and DM510 dicroic mirrors, BA470 and BA520 barrier filters and EX400−440 and EX450−490 excitation filters, respectively.
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Fungal structures and plant responses associated with sites of infection were visualised both at 100× and 400× magnification using Fluor objectives 341405 and 333843, respectively, and photographed at 400× with a Nikon FX−35WA 35 mm camera operated by a Nikon UFX−IIA camera mount and exposure control unit (Nikon Corporation).
5.6.4.4. Electron microscopy: Leaf discs and inoculum were prepared as described in sections 5.6.3.1 and 5.6.3.2 respectively. Leaf discs were inoculated 24 and 36 hours prior to observation. Un−inoculated leaf discs were also prepared for observation in order to view differences in the surface structure of the different plant species. Samples were prepared for electron microscopy via the method of Craig and Beaton (1996). However, samples were frozen in liquid nitrogen prior to insertion into the cold stage. Sites of direct penetration by R. alismatis were investigated by removing conidia, germ tubes and appressoria from the leaf surface by coating the leaf surface with molten gelatin, which was peeled off after it hardened (Preece 1971; Wheeler 1975). Gelatin (20% aqueous w/v) was heated to 35°C prior to application, and appeared to cause minimal distortion of the leaf cuticular surface. Clear Scotch brand sticky tape was also used to remove inoculum from the leaf surface. Observations were made at 15 kV using a JOEL 6400 Scanning electron microscope equipped with a Bio−Rad E7400 cryotrans system, under the operation of Mr E. Hines at the Department of Entomology, CSIRO, Canberra, Australia.
5.6.5 Data analysis: Replications consisted of the 6 leaf discs per time interval for each of the three plant species, and experiments were repeated in triplicate. Data were analysed as a split plot design, with replicates and species as main effects. Because variance for all experiments were similar, data for each plant species were pooled before analysis. Because the residuals were normally distributed, transformation of data was unnecessary, and an analysis of variance (ANOVA) was performed at each time interval for both germination and appressorium formation. The means at each interval were separated using the least significant difference (LSD) method. The computer package Genstat 5.0 was used for all statistical analyses (Lawes Agricultural Trust, Rothamsted Experimental Station, AFRC Institute of Arable Crop Research, Harpenden, Hertfordshire, UK).
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5.7 Results Conidial germination and appressorium formation were observed on all three Alismataceae species examined in this study. Regardless of species, germination commenced within 6 hours of inoculation with greater than 50% of applied conidia germinating by the 12 hours mark. Whilst significant differences were observed in the percentage of conidia that germinated on the leaves of different species at all but one of the examination times (Table 5.1), the majority of differences were reflected by host and non−host affiliations with four of the intervals separating the species on this basis. Despite these differences, however, there was no significant difference in the percentage of germinated conidia on the leaves of the 3 species after 36 hours, with greater than 90% of conidia elongating to form germ tube structures (Table 5.1). The rate of germination on each species was also nearly identical (Figure 8.1a, b).
Table 5.1: Percentage of conidia of R. alismatis that germinated and percentage of germinated conidia that formed appressoria on the leaves of A. plantago−aquatica, S. graminea and S. montevidensis. Figures represent the pooled data from six replicates and three experimental treatments. Time after inoculation A. plantago− S. graminea S. montevidensis LSD 5% (hours) aquatica Conidial germination (G)% 06 21.1 29.7 56.5 17.77 12 52.5 69.5 63.9 12.48 18 55.7 79.8 89.6 13.14 24 72.9 89.9 89.2 10.72 30 79.9 94.9 92.1 7.88 36 91.1 97.0 91.9 6.54 Appressoria formation (A)% 06 0.4 3.0 21.8 8.29 12 9.1 35.9 48.0 13.09 18 23.3 57.0 53.5 12.16 24 38.4 67.4 68.3 15.79 30 35.9 57.6 91.6 12.32 36 39.8 77.7 89.0 13.44 %G = G+A/TOT × 100; %A = A/G × 100; TOT = U+G+A (U = un−germinated conidia). LSD = Least significant difference between means (P=0.05).
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The formation of appressoria also commenced within 6 hours of inoculation, but did not reach an appreciable level on A. plantago−aquatica and S. graminea until approximately 12 hours after inoculation. Significant differences were observed in the percentage of germinated conidia that formed appressoria on the leaves of different species at all of the examination times (Table 5.1). Once again differences were evident between host and non−host species with rates of appressoria formation on the leaves of A. plantago−aquatica differing significantly from those on Sagittaria species at four of the six time intervals. Appressoria formation continued to increase throughout the examination period on both S. graminea and S. montevidensis, but remained relatively stable on the host species during the latter part of the experimental period (Table 5.1). Although the total number of conidia that formed appressorium on A. plantago–aquatica was only about half that of the non– host species, the rate of appressorium formation was almost identical on both host and non−host species (Figure 8.2a, b). Interestingly the greatest numbers of conidia progressing to appressoria formation were observed on species of Sagittaria. Two stained areas separated by an internal septum were visible in most of the conidia of R. alismatis prior to germ tube emergence (Figure 5.1a). Usually only a single unbranched germ tube was produced by each conidium. The length of germ tubes was variable, but most produced a terminal appressorium of variable size and shape (Figure 5.1b). Frequently the contents of the spore were observed to migrate from the conidium along the germ tube to the appressorium (Figure 5.1c). On several occasions multiple germ tubes were observed originating from a single conidium. In most of these cases a germ tube extended from either end of the conidium but on some occasions up to four germ tubes were observed per conidium. In situations where multiple germ tubes were observed, germ tube elongation was often lengthy, and frequently progressed into highly branched networks where appressorium formation was less frequent and appeared to be associated with terminal and intercalary branches (Figure 5.1d). Multiple germ tube production, extensive elongation and the branching of germ tubes into massive networks were frequently observed during the later stages of infection of S. montevidensis, but were not observed during infections of S. graminea nor the host species A. plantago−aquatica. Where multiple germ tubes where observed, the expectation that multiple appressorium would eventuate was not realised, and multiple appressoria were observed infrequently. In cases where multiple appressoria were observed, never more than two per conidium eventuated, and for the most part even in the presence of extensive germ tube elongation and branching, only a single terminal appressorium was produced per conidium.
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a b c
d e
f g h
Figure 5.1: Light and fluorescent micrographs following inoculation of leaves with R. alismatis; (a) septate un–germinated conidium, (b) germinated conidium showing germ tube and terminal appressorium, (c) conidium displaying cytoplasmic migration, (d) multiple germ tube formation and extensive branching, 36 hours after inoculation of S. montevidensis, (e) necrosis and cell death around infection sites on S. graminea 24 hours after inoculation, (f) ‘halo’ formation around infection sites on S. graminea 24 hours after inoculation (g) penetration sites (holes) on S. graminea 24 hours after inoculation, (h) fungal structures associated with infection sites on S. graminea 36 hours after inoculation. Bars = 10 µm, except insert photos f and g, Bars = 1µm.
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Germ tube elongation and appressorium formation seemed to occur randomly on the leaf surface. No indication of directional growth or common stimulus for appressorium formation was observed. Appressoria were either sessile on conidia (Figure 5.2a), formed at the junctions between adjacent epidermal cells, near the edges of stomatal guard cells (Figure 5.2b) or occasionally associated with stomates (Figure 5.2c). Some stomatal infiltrations were also observed (Figure 5.2d) but germ tubes frequently passed over stomata without forming infection structures (Figure 5.2b). The development of appressoria and subsequent direct penetration through the cuticle and epidermis of Alismataceae species appeared to be the most common method by which the fungus entered the leaf. Sites of appressorium formation by the fungus were frequently accompanied by evidence of changes in the structure of the leaf surface. Fluorescent micrographs showed evidence of necrosis in epidermal cells of S. graminea beneath sites of appressorial formation (Figure 5.1e). Light micrographs revealed the presence of blue, disc shaped anomalies or ‘haloes’ in association with infection sites around appressoria of S. graminea (Figure 5.1f). Haloes first appeared after 18 hours, and were visible by SEM as areas of increased electron density (Figure 5.2e). These irregularities appeared to result from the release of some form of exudate onto the host tissue surrounding the sites of appressorium formation. An additional form of exudate was also often present in conjunction with fungal structures. Unlike the former material, which seemed to furrow the leaf surface, this release appeared to form a coating around fungal structures, but often appeared cracked and deformed (Figure 5.2c). Penetration pegs from developing appressoria were not observed with any certainty on any of the species examined. However, sites of penetration were observed both on A. plantago−aquatica and S. graminea both by fluorescent microscopy (Figure 5.1g) and under SEM (Figure 5.2f) following the removal of inoculum of R. alismatis and other leaf microflora. Fluorescent microscopy allowed visualisation of both the fungus and infection holes without the removal of fungal material from the leaf surface (Figure 5.1h). Holes from penetration pegs were observed within 24 of inoculation. Infection sites were further investigated by SEM 48 hours after inoculation. In most cases perfectly spherical holes approximately 0.25 − 0.5 µm in diameter were observed, but occasionally smaller holes were observed accompanied by depressions or deformations of the leaf surface (Figure 5.2f insert). In some cases the attempted removal of inoculum appeared to be only partially successful, and remnants of the appressorium remained on the leaf surface surrounding the point of invasion (Figure 5.2g). On one occasion the cuticle of S. graminea was torn away from lower leaf tissue beneath the appressorium (Figure 5.2h), exposing a clearly visible penetration peg hole in the underlying leaf tissue (Figure 5.2h insert).
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a b c
d e f
g h i
j k l
m n o
Figure 5.2: SEM micrographs, captions next page.
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Figure 5.2: SEM micrographs following inoculation with R. alismatis. (a) Sessile appressoria, (b) germs tubes passing over open stoma on S. montevidensis 48 hours after inoculation, (c) appressorium formation above open stoma on S. montevidensis 60 hours after inoculation including appearance of extracellular matrix or ice crystal damage, (d) stomatal infiltration on A. plantago−aquatica 24 hours after inoculation, (e) appressorium formation on S. graminea 48 hours after inoculation including ‘halo’ appearance associated with increased electron density, (f) penetration sites (holes) 42 hours after inoculation of S. graminea, insert: depressions around penetration sites (g) penetration sites and remnants of partially removed appressorium on A. plantago−aquatica 42 hours after inoculation, (h) appressorium and damaged cuticle following attempted removal of inoculum 48 hours after inoculation of S. graminea, insert: subcuticular penetration site, (i) appressorium and associated penetration peg following attempted removal of inoculum, including penetration site and cuticular depression on A. plantago−aquatica 48 hours after inoculation, (j) adherence of spores to the leaf surface of S. montevidensis 24 hours after inoculation, with evidence of physical stress exerted on stomal structure, (k) collapsed spore and germ tubes resulting from cytoplasmic migration, (l) host reaction (swelling) after inoculation of D. minus (Figure 5.2l, Photograph reproduced by permission of Dr F. G. Jahromi), leaf surface and associated wax deposition of un–inoculated leaves of (m) A. plantago−aquatica, (n) S. graminea and (o) S. montevidensis. Bars = 10µm, except photos f, g, i and insert photos f and h, Bars = 1µm.
On another occasion the attempted removal of the fungus exposed a penetration peg hole and what appeared to be evidence of a penetration peg still attached to the appressorium (Figure 5.2i). Neither penetration peg holes or any other indication of infection was observed on S. montevidensis leaves. Observations of this nature suggested that the fungus adhered tightly to the leaf surface. Another example of this was evident where a spore caused significant stress to an opening stomate (Figure 5.2j). Successful invasion of the plant cuticle appeared to be accompanied by collapse of conidia and germ tubes and was mirrored the movement of cellular contents from the conidia into the appressorium, similar to that observed on earlier light micrographs (Figure 5.2k, 5.1c). Visible symptoms of invasion usually appeared within four to six days of inoculation of A. plantago−aquatica and S. graminea but were observed infrequently on S. montevidensis. The first macroscopic symptoms of disease include ‘pepper spots’ (< 0.5 mm) that develop into dark brown necrotic spots 1−3 mm in diameter with or without a pale green to yellow halo. As disease progression continued individual lesions often coalesced to form larger lens shaped lesions (Cother et al. 1994). Examples of macroscopic disease symptoms as they appear on the leaves of A. plantago−aquatica and S. graminea are shown in Figures 5.3a, b. A symptomatic photograph of D. minus was included for comparison (Figure 5.3c). No disease symptoms were observed on whole leaves of S. montevidensis.
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a b
c
Figure 5.3: Macroscopic disease symptoms following inoculation of leaves with R. alismatis; (a) A. plantago–aquatica, (b) S. graminea, (c) D. minus (Figure 5.3c, Photograph reproduced by permission of Dr. F. G. Jahromi).
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5.8 Discussion This study represents the first critical investigation of the events in the infection process of R. alismatis on the leaves of A. plantago–aquatica, S. graminea and S. montevidensis. The results of this study indicated that the leaf surface of the host species A. plantago−aquatica was less conducive both to germination and the production of infection structures by R. alismatis than the leaves of either of the non−host species S. graminea and S. montevidensis. Whilst this was reflected by significant differences both in the numbers of germinating conidia (P ≤ 0.05) and germinated conidia that formed appressoria (P = 0.001; Table 5.1), the rate of germination and appressorium formation by R. alismatis did not differ significantly between the three species (P = 0.05; Figures 8.1 & 8.2), indicating that R. alismatis can form infection structures on the leaves of non−host species at similar rates to host species. It was observed that spores often pooled on various parts of the leaf surface, especially along leaf veins, resulting in high spore densities but low germination rates. Several groups have demonstrated that ‘crowding’ of spores causes this phenomenon (Allen 1976; Louis & Cooke 1985a; Bailey et al. 1992). Bartnicki−Garcia (1984) reported that urediospores of rust fungi, when placed in a ‘crowded’ situation, release substances that inhibit their own germination. When spores spread out, these inhibitors are diluted and spores are free to germinate. In addition to rusts (Staples & Wynn 1965; Macko 1981; French 1992) self inhibitors have been washed from the spores of Colletotrichum (Lax et al. 1985; Leite & Nicholson 1992; Tsurushima et al. 1995; Weng & Chuang 1997) and R. secalis (Ayres & Owen 1970). Self inhibitors prevent premature germination of spores under less than optimal conditions, where competition for space and nutrients reduces the probability of successful disease initiation (Macko et al. 1972). In light of these reports, and previous research that demonstrates that spores of Rhynchosporium produce such substances, areas where conidia congregated were avoided during the enumeration process. Conidial germination by R. alismatis was similar to that described for both R. secalis (Ayesu−Offei & Clare 1970; Lyngs−Jorgensen et al. 1993; Jones & Ayres 1974) and R. orthosporum (Perez−Fernandez & Welty 1991) on their respective hosts, with one or both conidial cells elongating to form germ tube structures (Figure 5.1b). Multiple germ tube formation was reported frequently by many groups studying R. secalis and R. orthosporum, and in previous studies conducted by Jahromi et al. (2002) on interactions between D. minus and R. alismatis. However, few reported the emergence of three or more germ tubes from a single conidium (Ayesu−Offei & Clare 1970; Lyngs−Jorgensen et al. 1993; Jahromi et al. 2002), which was a feature of this study.
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Mount and Slesinski (1971) showed that deviations from optimal environmental conditions result in an increase in the number of germ tubes per conidium. These authors along with many others stressed the importance of inoculum quality and uniformity if accurate and quantitative observations and determinations of host−parasite interactions are to be made (Graf−Marin 1934; Nair & Ellingboe 1965; Ellingboe 1968; 1972; Emmett & Parberry 1975). In circumstances where multiple germ tubes were observed, germ tube elongation appeared to continue until a stimulus for appressorium formation was received. In the absence of such a stimulus branching networks often resulted, and these were frequent during the later stages of infection on S. montevidensis, but have not been previously reported by other researchers (Figure 5.1d). Ayesu−Offei and Clare (1970) cited branching during their study of the infection process of R. secalis on barley leaves but described branching of germ tubes as infrequent, never elaborate. Branching and extensive germ tube elongation, however, is not an uncommon observation in histological studies of fungal−plant interactions (Staub et al. 1974). With few exceptions (Bonde et al. 1976; 1982; Koch & Hoppe 1988), urediospore infections by rust fungi involve entry through stomates (Royle 1976; Dodman 1979). The infection process of rusts, therefore, results in marked differences in the lengths of germ tubes, as spores may be deposited at various distances from these entry points. Van Dyke and Mims (1991) added that germ tube length of C. truncatum appeared to be related to moisture conditions, with wetter conditions favouring longer germ tubes. Branching is commonplace in germ tubes of Cochliobolus and Pyrenophora (Emmett & Parberry 1975). Appressorium formation by R. alismatis did not seem to be affected by germ tube length, nor was it associated with melanisation. Mostly, appressoria formed at the ends of both short and long germ tubes and terminally on branched networks (Figure 5.1b, d). However, sessile appressoria (Figure 5.2a) were also extremely common both in this study and in studies conducted on R. secalis (Caldwell 1937; Hosemans & Branchard 1985; Ayesu−Offei & Clare 1971; Lyngs−Jorgensen et al. 1993; Xi et al. 2000; Jahromi et al. 2002), but were not reported during infection of orchard grass by R. orthosporum (Perez−Fernandez & Welty 1991). Appressorial formation was accompanied by the movement of the cytoplasmic contents from the conidium to the appressorium (Figure 5.1c). This phenomenon was easily observed under light microscopy, and was reflected by a collapsing of germ tubes under SEM (Figure 5.2k). Roderick and Thomas (1997) reported that the cytoplasm of P. graminis spores was observed to migrate into the appressoria and subsequently into the sub−stomatal vesicle during infection of ryegrass leaves, resulting in appressoria that often had a collapsed appearance under SEM. Jones and Ayres (1974) also observed similar arrangements in their studies on the barley−R. secalis pathosystem, whilst Van−Dyke and Mims (1991) reported similar occurrences on dialysis membranes infected with C. truncatum.
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The significance of cytoplasmic migration is largely speculative but, within the cytoplasm, diverse structures and organelles undergo independent motility possibly providing the energy and components necessary for elongation of the growing germ tube (McKerracher & Heath 1987). Interestingly, the movement of cytoplasmic contents from the conidium into the appressorium did not appear to be restricted to species in which a successful infection had been accomplished, and was observed during interactions with all species including S. montevidensis. The formation of an appressorium on branched networks seemed to signal cessation of elongation, and multiple appressoria were observed infrequently. Appressoria varied in size and shape depending on their position on the leaf and often formed at the junctions between adjacent epidermal cells or near the edges of guard cells (Figure 5.2b). Martin and Juniper (1970) reported that these regions correspond closely to those which, in many plants, are rich in pectic substances. However, other reports indicate that yeast and bacteria live and divide in the junctions between epidermal cells in preference to other sites, and according to Preece et al. (1967), comments in the literature are numerous which suggest parasitic fungi are no exception. Whilst Preece et al. (1967) warned that a single example of such an event depicted in an accompanying micrograph is often enough for such an occurrence to become accepted as typical, their studies confirmed this phenomenon in red clover and cauliflower and strengthened very early reports by De Bary (1887) which suggested fungi belonging to at least three widely different taxonomic groups made their entrance via these routes. More recently, appressoria formation at these sites has been confirmed in species of Colletotrichum (Pantidou & Schroeder 1955; Politas & Wheeler 1973) and Alternaria (Green et al. 2001). Willmer (1983) suggested that the morphological and anatomical differences between guard cells and other epidermal cells also contributes to this phenomena. For example, some areas of the guard cell cuticle are known to be thinner than the cuticle of surrounding epidermal cells and may facilitate more rapid or more successful penetration of the cuticle. Additionally, guard cells are known to accumulate large quantities of starch (Fahn 1982) and Willmer (1983) speculated that these sites may represent nutrient rich regions which encourage appressorium formation and fungal penetration. Earlier researchers (Bartels 1928; Mackie 1929) suggested that fungal germ tubes of R. secalis may also enter the host through stomates. Stomatal penetrations are common in many species of fungi including some well known pathogens such as Colletotrichum (McRae 1989), but are best known for their role in the infection process of rust fungi (Staples & Macko 1980; Staples 1985; Hoch & Staples 1987), where they often represent the principal ports of entry into the plant.
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In this study, several stomatal infiltrations were observed (Figure 5.2d), consistent with earlier studies by Jones and Ayres (1974) on R. secalis and recently published work by Jahromi et al. (2002). More frequently, germ tubes were observed to travel over stomates without initiating infection (Figure 5.2b). Occasionally an appressorium also formed over an open stomate (Figure 5.2c). Caldwell (1937) and Ayesu−Offei and Clare (1970) strenuously opposed the suggestions of Bartels (1928) and Mackie (1929) that stomatal openings represent sites of infection for Rhynchosporium. Ayesu−Offei and Clare (1971) confirmed that hyphae aggregated above guard cells and effected penetration between the end walls of guard cells and contiguous epidermal cells, but argued that there is no evidence that infection can be established in this manner, suggesting that these observations may have given the impression of direct stomatal penetration. The production of penetration pegs by appressoria was not observed with any certainty on any of the species under examination. On one occasion following the attempted removal of inoculum, a fungal structure was observed and photographed that gave the impression of a penetration peg (Figure 5.2i). Unfortunately this type of structure was observed only once during the study and despite evidence of a depression and a penetration hole in the background of the micrograph, the two fixtures could not be proven to be related. For the most part it was not expected that these structures would be observed without sectioning and staining because viewed from above the appressorium would interfere with viewing of pegs. However, it was possible to observe penetration peg holes in the leaf surface under fluorescent and electron microscopy, and these structures were observed both on A. plantago−aquatica and S. graminea (Figures 5.1g and 5.2f). The penetration holes were round, had smooth edges and were approximately 0.25 − 0.5 µm in diameter. Occasionally some smaller holes were observed in conjunction with significant depressions around the outer edges, indicating signs of considerable physical stress (Figure 5.2f insert). These depressions suggested that the fungus adhered tightly to the leaf surface, at least during the initial stages of penetration, but their absence around many holes suggested that this force dissipated following successful penetration of the leaf surface. On one occasion the attempted removal of inoculum resulted in only partial removal of fungal structures, and a portion of an appressorium remained adhered to the leaf surface around the penetration peg hole (Figure 5.2g). On a second occasion a spore was observed to adhere so tightly to the surface of a stomatal guard cell that the function of the stoma appeared to be compromised, and this was manifested by the appearance of stress ridges (Figure 5.2j). Furthermore, attempted removal occasionally resulted in the complete removal of fungal structures including the cuticle area to which they were attached. In one instance this provided a subcuticular view of the resulting penetration hole (Figure 5.2h).
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These observations further strengthened the argument that not only do fungal species adhere tightly to leaf surfaces but that attachment occurs both prior to and during infection, with appressoria, germ tubes and spores structures all adhering to the host surface. The different sizes of holes and evidence of depressions indicated that the fungus may have been removed at different stages during the infection process. Occasionally, fungal structures appeared to be coated in some form of extracellular matrix, or mucilaginous material (Figure 5.2c). Many authors have suggested that adhesion involves the secretion of fluids or adhesive materials that secure the fungus to the plant surface prior to infection (Lapp & Skoropad 1978; Epstein et al. 1987; Latge 1991; Chaubal et al. 1991; Schuerger & Mitchell 1993; Doss 1999). For example, Colletotrichum species are known to be embedded in a water−soluble mucilage, which enables the spore to adhere to the hydrophobic leaf surface (Nicholson & Epstein 1991). However, the so called ‘exudate’ material in this study often had a cracked or torn appearance and may have merely represented artefacts that commonly originate during the cryofixation process of sample preparation. Read (1991) reported that low temperature SEM produces several specific artefacts which are associated with frozen−hydrated specimens. These include ice crystal damage and surface rupturing and tearing. In some cases, however, materials seemed to be solely associated with fungal structures (Figure 5.2a) and possibly contribute to the infection process through deposition of enzymes. For example, analyses of mucilaginous materials from Colletotrichum revealed the presence of a variety of enzymes that have since been shown to alter the plant surface underneath conidial germ tubes and appressoria (Mendgen & Deising 1993). Hence, in addition to the role of adhesion, the conidial matrix may play a part in pathogenicity of some species of fungi (McRae & Stevens 1990). Appressorium formation and sites of penetration by R. alismatis were frequently accompanied by the presence of blue disc–shaped zones or ‘haloes’, presumably representing the sites of penetration by the fungus (Figure 5.1f). The margins and centres of haloes appeared to have a greater affinity for stain than the areas in between (Figure 5.1f insert). A darker staining central spot possibly indicated the presence of papillae formation. Papillae and haloes have been described frequently during interactions between barley and the scald pathogen R. secalis, both in resistant and susceptible cultivars and during successful and unsuccessful penetration attempts (Caldwell 1937; Ayesu−Offei & Clare 1970; Hosemans & Branchard 1985; Lehnackers & Knogge 1990; Lyngs−Jorgensen et al. 1993; Xi et al. 2000). In this study, haloes and papillae were observed frequently on the leaves of S. graminea as early as 18 hours after inoculation and were similar in size, approximately 10−15 µm, to those reported for the barley−R. secalis pathosystem (Ayesu−Offei & Clare 1970).
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Host cells containing papillae appeared dead when viewed under fluorescence and fungal development appeared to be restricted to a single epidermal cell (Figure 5.1e). Comparable anomalies were evident in electron micrographs of sites of appressorium formation in S. graminea (Figure 5.2e). These sites were accompanied by an increase in electron density, similar to that reported by Stavely et al. (1969) during studies on Trifolium host cell walls surrounding E. polygoni papilla, and have been suggested to be the result of fungal enzymes. Unlike interactions reported for the R. secalis−barley pathosystem, however, no evidence of haloes or papillae formation in susceptible hosts was observed during interactions between Alismataceae species and R. alismatis, and neither Jahromi (2000) nor Perez–Fernandez and Welty (1991) observed this phenomenon during studies of other compatible Rhynchosporium pathosystems. Symptoms of successful invasion were apparent on the leaves of both A. plantago–aquatica and S. graminea within four to six days of inoculation (Figure 5.3a, b). Small dark brown necrotic spots, with or without pale green to yellow haloes (chlorosis), were common. Occasionally disease symptoms were also observed on the leaves of S. montevidensis. However, considering there was no evidence of penetration of this species, it is unlikely these symptoms are the result of direct infections established through the cuticle. During inoculation of the leaf discs, it was noted that the surface of S. montevidensis was less hydrophobic than the other species. Inoculum droplets frequently were observed to bead on the surface of both A. plantago–aquatica and S. graminea, but this was rarely the case with leaves of S. montevidensis and droplets frequently diffused out over the leaf surface. When viewed under electron microscopy, the leaf surface of S. montevidensis was significantly different from that of A. plantago–aquatica (Figure 5.2m) and S. graminea (Figure 5.2n), being devoid of wax crystals (Figure 5.2o). These morphological differences possibly aided the distribution of inoculum and increased the probability of sporadic infections initiated through wound sites, explaining the occasional disease symptoms observed on leaf discs of S. montevidensis. The chlorotic appearance of lesions also suggests that R. alismatis produces toxins during the infection process. At present there is no data to support such assumptions, however, several low molecular weight toxins, the rhynchosporosides (Auriol et al. 1978; Rafenomananjara et al. 1983), and a toxic glycoprotein (Mazars et al. 1984; 1989), have been isolated from culture filtrates of R. secalis. The process of infection of Alismataceae by R. alismatis bears many similarities to that observed in R. secalis, including the descriptions of Jones and Ayres (1972) that it adopts a hemibiotrophic lifestyle, relying heavily on a biotrophic relationship with the host during the early stages of infection.
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While the results of this study support this notion, the organism appears to establish both biotrophic and necrotrophic relationships with the host, depending on compatibility or incompatibility, similar to that reported during interactions between Sorghum bicolor and C. sublineolum (Wharton & Julian 1996; Perfect et al. 1999; Wharton et al. 2001) and many other Colletotrichum pathosystems including several that are currently being investigated in a biocontrol context (Auld et al. 1988; McRae & Stevens 1990; Makowski & Mortensen 1998). During the colonisation of plants many species of Colletotrichum exhibit both biotrophic and necrotrophic modes of nutrition (Perfect et al. 1999). In compatible reactions, infected cells remain alive for considerable periods through the establishment of a biotrophic interaction in the first colonised cell. Incompatible reactions result in rapid death of cells as a result of cytoplasmic aggregations that become pigmented, lose their shape and release their contents into the cytoplasm, killing the cells, restricting further fungal development and limiting the progression of disease and macroscopic disease symptoms (Snyder & Nicholson 1990; Snyder et al. 1991). During the biotrophic phase, infection vesicles are surrounded by an interfacial matrix containing glycoproteins that separate the fungal cell wall from invaginated host plasma membranes (O’Connell 1987) and the fungal parasite avoids triggering, or suppresses host defence responses such as papillae formation, hypersensitive cell death and the release of antifungal compounds (Heath & Skalamera 1997). This initial biotrophic phase probably enables the fungus to become established within sufficient tissue to lessen the inhibitory effects of such defence compounds (O’Connell & Bailey 1991). The absence of haloes and papillae–like appositions in the host A. plantago–aquatica is unlike reactions observed during interactions in the barley pathosystem, where both resistant and susceptible cultivars and successful and unsuccessful penetration attempts were accompanied by such appositions, but may simply reflect the duration over which observations were recorded. Wharton and Julian (1996) reported that the biotrophic phase of Colletotrichum sublineolum persisted for at least 24 hours following inoculation, but lost structural integrity shortly thereafter, which correlated with a failure of cells to accumulate vital stains. Hence, the plausible suppression of host defence mechanisms during this period as a result of a similar biotrophic phase is not unlikely, and observations over longer durations may result in the appearance of similar haloes and wall appositions in A. plantago−aquatica. There is considerable evidence that papillae formation in hosts occurs more slowly than in non−hosts (Lyngs−Jorgensen et al. 1993) and Waterman et al. (1978) have suggested that the speed of papillae formation is a resistance determining factor. Additionally, antifungal metabolites such as phytoalexins, which are synthesised by plants in response to fungal ingress, are believed to accumulate more slowly and reach lower concentrations in susceptible as opposed to resistant hosts (Nicholson & Hammerschmidt 1992). 84 Chapter 5. Infection Process
Furthermore, the compounds that do accumulate in susceptible hosts are often less fungitoxic than those that accumulate in resistant species (Bennett & Wallsgrove 1994; Hahn et al. 1985; Graham et al. 1990; Lo et al. 1999). Also some fungi may suppress or prevent host responses through the secretion of ‘blockers’ (Loegering 1978; Heath 1981a). Hence, suppression, slower initiation or absence of wound related responses in A. plantago−aquatica may account for the lack of observed host related responses in this species. However, it is more likely that the interaction between R. alismatis and the host species A. plantago−aquatica represents a compatible relationship in which the ability of the fungus to successfully parasitise the plant is due to ‘specific accommodation’ of the pathogen by the host that renders defence mechanisms inactive or ineffective (Ward & Stoessl 1976; Heath 1981d). This relationship is termed ‘basic compatibility’ (Heath 1981c). During incompatible interactions with the resistant host S. graminea, pigmented appositions or ‘haloes’, possibly accompanied by papillae, appeared at the sites of appressorium formation within the first 24 hours of infection and were accompanied by rapid death of infected cells which then adopted a necrotic appearance. During this necrotrophic phase, fungal development is restricted and the progression of disease and macroscopic disease symptoms are limited to the epidermal cells which have been invaded by the fungus. There is considerable evidence that suggests ‘haloes’ and associated papillae−like appositions play a significant role in arresting disease progression during these incompatible interactions. Firstly, haloes in barley epidermal cell walls are considered to be closely associated with the response of the host to R. secalis attack (Lyngs−Jorgensen et al. 1993). That similar responses should be observed during interactions between R. alismatis and Alismataceae is not unreasonable. Secondly, infection by R. secalis causes extensive cell wall breakdown in barley, presumably associated with a cellulase complex that has been demonstrated in culture filtrates (Olutiola & Ayres 1973). Suzuki et al. (1982) present considerable evidence that halo formation is caused by partial digestion of cellulose around penetration sites. Finally, the appearance of smooth edges around infection holes is also suggestive of enzymatic involvement. For example, a penetration process effected by physical pressure might be expected to leave signs of tearing around penetration holes (Staub et al. 1974). However, enzymatic degradation of surrounding leaf tissue would be expected to provide a depression. The accumulation of dye in such depression during the staining process may provide the appearance of blue discs or ‘haloes’. Unfortunately, the occurrence of papillae often seem to be indicators of resistance rather than barriers to fungal invasion (Heath 1984), and although there is a good correlation between development of lignified papillae and resistance of epidermal cells to fungal invasion (Sherwood & Vance 1980), other mechanisms may be involved in the expression of incompatibility exhibited by S. graminea during this study.
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One mechanism which results in rapid death of cells around infection sites and limitation of disease progression is the hypersensitive response (HR) (Grant & Mansfield 1999). In this study, however, it is suspected that the pathogen undergoes a necrotrophic relationship with incompatible plant species. Hence, necrosis per se is unlikely to result in the limitation of disease progression common in this species. Under these circumstances additional induced responses including the release of phenolics, phytoalexins or other antifungal compounds are likely to play significant roles in disease resistance (Heath 1984). Histological studies, however, are clearly needed before definite conclusions can be made about the role of these and other defence mechanism during the interactions between R. alismatis and Alismataceae species. During interactions with the resistant host, S. montevidensis, there was a distinct lack of activity following appressorial formation. Neither haloes nor papillae−like structures were observed and there was no evidence of penetration by the fungus. While this latter point may not be significant in that insufficient leaf area may have been observed during the examination, there was no evidence to suggest that either a compatible or incompatible reaction developed between the two organisms. The reasons for this are unclear. There was a distinct correlation between germ tube length and inability to penetrate the host, however, and Niks (1990) reported similar correlations between germ tube length and establishment of sporelings of Puccinia hordei on barley cultivars. Niks (1990) suggested that the formation of long germ tubes, necessary for some sporelings to reach a stoma, decreased the amount of energy available to the sporelings to complete the infection cycle. Unfortunately, Niks’s (1990) study was related to the establishment of infection structures rather than penetration of the host, and germ tube length in this study did not seem to result in reduced appressorium formation. In fact appressoria formed readily on the surface of S. montevidensis at rates equivalent to the other species, and at several times during the experimental period was found to be significantly greater on S. montevidensis than the other species, despite the fact the differentiation of infection structures failed to result in successful penetration of the plant. Several groups have suggested that topographical stimuli such as nutrient status, especially exogenous compounds may be an important stimulus for germination and appressorium formation (Parberry & Blakeman 1978; Lucas & Knights 1987; Staples & Hoch 1997). For example, the need for nutrients at the plant surface for full development of appressoria in R. solani and E. cruciferum has been demonstrated (Emmett & Parberry 1975) and chemicals exogenous to fungi have been implicated in the development of appressorium of Sclerotinia, Colletotrichum and several other fungal species (Purdy 1958; Agnihotri 1969).
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Similarly, nutrient deprivation may also be associated with initiation of germination and appressorium development (Dunkle et al. 1970; Blakeman & Parberry 1977) and several groups suggest that absence of conducive conditions is a potent stimulus for germ tube differentiation and appressorial formation (Hamer et al. 1988; Howard et al. 1991). The fact that high numbers of appressoria were formed on the leaves of S. montevidensis, a seemingly resistant species with which neither a compatible or incompatible relationship is evident, suggests that appressorial formation by R. alismatis may too be stimulated in response to non−conducive topographical cues. While this may not affect the number of appressorial initials, non−host topographical stimuli that differ significantly from the host leaf may mean that germ tubes make ‘mistakes’ in locating or recognising conducive sites for development, considerably reducing the number of individuals that are able to enter the host, or preventing their entry altogether. The fact that the leaf surface of S. montevidensis is noticeably different from that of the other species examined in this study, as described previously, is significant. In such cases where stimuli are absent, appressorial differentiation may take place in response to depletion of endogenous energy supplies or as a survival or panic mechanism, resulting in considerable appressorium formation but little or no disease initiation. The infection process of R. alismatis bears many similarities both to that of the form species R. secalis and to the compatible and incompatible interactions observed during infections associated with anthracnose fungi. The responses of host and non−host plants to invasion by the fungus varied greatly. Whilst this was expected, the response of known host plants A. plantago−aquatica and D. minus also differed significantly, with swelling reported around infection sites in the latter species (Figure 5.2l). Likewise non−host infections differed dramatically with significant defence mechanisms being triggered during invasion of S. graminea. While several groups (Coates et al. 1993; O’Connell et al. 1985) have noted variation in the behaviour of different hosts to the same pathogen the stimulus for such differences remains a mystery despite the fact that differences almost certainly result from variation in host morphology. Whilst much is known about the responses of compatible and incompatible hosts to fungal invasion, little is known about the mechanisms that trigger these reactions. For rusts, these signals are largely physical as demonstrated by Allen et al. (1991) and summarised by Ride (1992), and recently it has been demonstrated that solid surface recognition is essential for infection structure formation in Magnaporthe (Xiao et al. 1997). However, thus far the signals that stimulate germ tube differentiation and appressorium formation in Rhynchosporium, not to mention the vast majority of fungal pathogens, have not been investigated.
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For R. alismatis spore germination and appressorium formation is most likely a non−specific response that requires little or no stimulus from the host. This concept was first suggested by Emmett and Parberry (1975) and probably explains why such a large proportion of fungal pathogens appear to be able to form infection structures on such a wide range of surfaces. Host recognition almost certainly occurs after the formation of appressorium, possibly after successful penetration of the plant cuticle and epidermis, which explains the lack of symptoms on S. montevidensis. Because cytoplasmic migration is evident in spores forming appressoria on the leaves of S. montevidensis, penetration peg formation may also be non−specific. Penetration of the host surface likely involves the secretion of enzymes, resulting in smooth penetration points and is evidenced by both the secretion of fungal exudates and the appearance of haloes thought to involve dissolution of cellulose around infection sites. Mechanical pressure is unlikely to be involved during the penetration process as few records indicate that mechanical penetration of the host surfaces is accomplished in the absence of appressorial melanisation. However, the depressions that were observed and attributed to attachment mechanisms earlier, may actually represent evidence of mechanical penetration and Muirhead and Deverall (1981) have reported previously that unmelanised appressoria of C. musae are capable of penetrating the cuticle of banana peel. During compatible reactions the fungus is able to inhibit plant defences through the establishment of ‘specific accommodation’ with the plant. This relationship is likely a biotrophic one similar to that observed in Colletotrichum, and reviewed recently by Perfect et al. (1999). Incompatible reactions quickly elicit host defence mechanisms that seek to limit the progression of disease through cell death and decompartmentalisation of infection structures. Haloes and commonly associated papillae, themselves responsible for rapid thickening of cell walls through deposition of new materials and compartmentalisation of infection structures, appear likely. Hypersensitivity reaction−related symptoms are also apparent, but the relationship appears to be a necrotrophic one that complements both the hemibiotrophic nature of Rhynchosporium as suggested by Jones and Ayres (1972) and the earlier comparisons to similar growth phases of Colletotrichum (Wharton & Julian 1996; Wharton et al. 2001). Hence this response is unlikely to limit the growth of the pathogen. Because this model does not correspond with the observations obtained from the field, where no prior records of the fungus on S. graminea have been reported, other defence mechanisms are likely. In R. alismatis however, these mechanisms most likely occur in addition to host cell wall thickening (papillae formation) which is considered an indication of resistance rather than a barrier to fungal growth (Heath 1984).
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Where neither compatible or incompatible reactions were observed, as in the case of S. montevidensis, it is likely that infection is limited through the inability of the fungus to penetrate the plant cuticle. Miscues, mistakes and surface topography may all play key roles. Additionally, Ellingboe’s (1968) and Mount and Slesinski’s (1971) suggestions on inoculum quality and uniformity are certainly relevant in light of the frequency of multiple and extensively branched germ tubes observed during infection of S. montevidensis. Whilst the situation is most likely a complex one, Niks (1990) assumption that spores may merely ‘run out of steam’ could be a relevant one. This study was critical to the understanding of disease initiation and development by R. alismatis on both host and non–host species and was pivotal in identifying major barriers during the disease cycle of R. alismatis that will facilitate future experiments to expand the host range of this potential biocontrol pathogen. Unfortunately, because of the stability of non−host resistance, pathogens rarely if ever spontaneously alter their host ranges (Heath 2000a). Hence, despite the ability of the fungus to penetrate the leaves of S. graminea under laboratory and glasshouse conditions, no records of the fungus have thus far been reported in the field. Nevertheless, the results of this study suggest that R. alismatis possesses the mechanisms required to invade S. graminea and as such it is likely through the application of formulation procedures that the constraints to disease progression can be overcome in this species. One approach that has been successful in improving the efficacy of R. alismatis in previous studies is the integration of low doses of chemical herbicides in conjunction with the fungus (Jahromi 2000). During this study, which involved the two most important herbicides in rice, Londax and MCPA, a synergistic effect was observed where fungal application followed the use of sublethal doses Londax. Whilst this represents but one example, clearly formulation procedures offer much hope for increasing the efficacy of R. alismatis on this species. In contrast, R. alismatis was unable to breach intact surfaces of S. montevidensis and as such the cuticular barrier of S. montevidensis imposed considerable constraints to the disease initiation and progression in this species. Whilst formulation procedures provide avenues through which the efficacy and host range of agents may be improved, these procedures require effective disease initiation in the target species. Hence, in the absence of host penetration, which is paramount to the ensuing success of the agent, alternative methods will be required to incite disease in this species. One approach which has provided considerable success is genetic transformation with genes encoding cuticle degrading enzymes.
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6 Transformation
The development of transformation techniques for the insertion of pathogenicity–related genes into R. alismatis
6.1 Introduction For many phytopathogens the first association with plant organs involves contact with the plant cuticle and carbohydrate–rich cell wall (Kolattukudy 1984b). Although it is universally accepted that mechanical force contributes to the pathogenesis of some fungi, many phytopathogens produce enzymes which are capable of degrading these structures (Walton 1994). To date, the major obstacle in addressing the function of many enzymes during pathogenesis has been the lack of efficient and reliable transformation procedures. Transformation systems provide the tools necessary to manipulate the genomes of fungal pathogens through the introduction of new or additional copies of existing genes or by mutation of genes by replacement and/or disruption, thereby identifying factors associated with pathogenicity and virulence. For example, Dickman et al. (1989) isolated a gene responsible for the production of cutinase, an enzyme which degrades cutin, the insoluble polymeric compound comprising the plant cuticle, from the vascular wilt pathogen Fusarium solani f. sp. pisi. By transforming species of the wound–pathogen Mycosphaerella with constructs containing this gene, these authors generated transformants with the capacity to infect intact papaya fruits, providing conclusive evidence of the role of enzymes during fungal penetration and demonstrating the utility of transformation procedures in determining pathogenicity attributes in fungi. In light of these studies, and the results of the previous chapter in which R. alismatis failed to effect penetration of S. montevidensis, this chapter considers the role of the cuticle–degrading enzyme cutinase during the initial stages of plant infection, and our attempts to the develop transformation procedures for the rice weed pathogen R. alismatis. Firstly a comprehensive treatise of the literature pertaining to genetic transformation in plant pathogenic fungi will be presented. This will then be followed by a sequential presentation and discussion of our attempts to effect transformation in R. alismatis, including a detailed breakdown of modifications that were initiated to improve the likelihood of success. Finally, speculative discussions regarding the outcomes of this study will be presented.
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6.2 Fungal cutinases and host penetration Because plant cuticles are the first defensive barriers encountered by directly penetrating plant pathogenic fungi, the role of fungal cutinases during plant pathogenesis has received considerable attention (Van den Ende & Linskens 1974; Kolattukudy 1985; Chasan 1992; Schafer 1993; Koller et al. 1995; Schafer 1998). Encouraged largely by the early work of Purdy and Kolattukudy (1975a; 1975b), who isolated the first cutinase isozymes from the pea pathogen Fusarium solani f. sp. pisi, the belief that this enzyme played a crucial role in the early stages of host infection began to gather considerable momentum within the scientific community. However, it was not until later that decade when Shayhk et al. (1977) demonstrated the presence of cutinase around infection sites of F. solani f. sp. pisi that conclusive evidence began to accumulate in favour of a role for cutinase during fungal penetration. In the years that followed, numerous publications arose which provided ongoing support for this hypothesis. Woloshuk and Kolattukudy (1986) and Podila et al. (1988) demonstrated a mechanism by which contact with the plant cuticle triggered cutinase activity and gene expression in fungal spores. Evidence arose that fungal infection could be prevented in a number of plant species through specific inhibition of fungal cutinases by phosphoorganic pesticides (Koller et al. 1982a; 1982b; Dickman et al. 1983; Koller et al. 1991), esterase inhibitors such as ebelactone (Umezawa et al. 1980; Koller et al. 1990; Chun et al. 1995) and di–isopropylfluorophosphate (Dickman et al. 1982; Muller & Ishii 1997), or through the preparation of antibodies against cutinase (Maiti & Kolattukudy 1979; Coleman et al. 1993). The construction of cutinase–deficient mutants in a number fungal species, the virulence of which was reduced or abolished but could be restored through the addition of exogenous cutinases, was demonstrated (Dantzig et al. 1986; Dickman & Patil 1986). Most significantly however, Dickman et al. (1989) converted the former wound pathogen Mycosphaerella to a directly penetrating pathogen of papaya by transforming this organism with a cutinase gene isolated from Fusarium. Kolattukudy et al. (1989) paralleled this work, demonstrating increased virulence in a cutinase–deficient strain of F. solani, through transformation with the same gene construct. Unfortunately, not all studies supported these earlier concepts, and while more recent research demonstrated additional roles for cutinase in tissue specificity (Trail & Koller 1990), and pre– penetrative spore–surface adhesion (Pascholati et al. 1992; 1993), the advancement of molecular biology and gene disruption via transformation failed to support the involvement of a number of previously characterised cutinases during plant infection (Sweigard et al. 1992b; Stahl & Schafer 1992).
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Conflicting reports in which the virulence of gene–disrupted transformants of F. solani f. sp. pisi was reduced or completely unaltered (Rogers et al. 1994; Stahl et al. 1994; Van Kan et al. 1997; Crowhurst et al. 1997), combined with evidence that chemical inhibitors of cutinase failed to prevent penetration and infection during interactions with several important crop species including beans, rice and cucumbers (Wolkow et al. 1983b; Woloshuk et al. 1983; Bonnen & Hammerschmidt 1989), all questioned the role of cutinase during host infection. Additionally, a publication by Yao and Koller (1995) in which the expression of different cutinases was reported during saprophytic and pathogenic growth of Alternaria brassicicola, further highlighted the complexity of the situation. Nevertheless, cutinases have been isolated and characterised from a large range of fungal pathogens including A. brassicicola (Trail & Koller 1993; Yao & Koller 1994; Fan & Koller 1998), Venturia inaequalis (Koller & Parker 1989), Phytophthora capsici (Munoz & Bailey 1998), Magnaporthe grisea (Sweigard et al. 1992a), and species of Colletotrichum (Ettinger et al. 1987; Liyanage et al. 1993) and Fusarium (Soliday & Kolattukudy 1976; Lin & Kolattukudy 1980; Koller et al. 1982c). Despite the controversial interpretation of data, many recent studies provide ongoing evidence for the role of cutinase during infection by Erysiphe graminis (Francis et al. 1996), Aspergillus flavus (Guo et al. 1996), Botrytis cinerea (Van der Vlugt–Bergmans et al. 1997), Ascochyta rabiei (Tenhaken et al. 1997), and Pyrenopeziza brassicae (Davies et al. 2000).
6.3 Transformation Genetic transformation of filamentous fungi is a relatively new, but rapidly developing area, that has been reviewed extensively (Mishra 1985; Hynes 1986; Fincham 1989; Timberlake & Marshall 1989; Finkelstein 1992; Goosen et al. 1992; Hynes 1996; Ruiz–Diez 2002). Historically, the basis for fungal transformation can be attributed to the early protoplast technology of Hutchinson and Hartwell (1967), who dissolved the cell walls of Saccharomyces cerevisiae with commercial enzyme preparations. Although these authors studied macromolecular synthesis, Hinnen et al. (1978) showed that protoplasts of S. cerevisiae produced by this method and treated with exogenous DNA, readily transformed in the presence of calcium chloride and polyethylene glycol. At around the same time, Beggs (1978) developed the first Escherichia coli shuttle vectors for this organism, and together these discoveries marked the beginning of a new era in fungal genetics.
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Ultimately, this technology led to the transformation of filamentous fungi, Neurospora crassa (Case et al. 1979) and Aspergillus nidulans (Tilburn et al. 1983), and finally to a whole range of fungal species including several higher basidiomycetes (Burrows et al. 1990; Randall & Reddy 1991; Marmeisse et al. 1992; Hirano et al. 2000; Saito et al. 2001) and oomycetes (Judelson et al. 1993; Mort–Bontemps & Fevre 1997). To date, transformation systems have been developed for a multitude of filamentous fungi, including several species pathogenic to humans (Messina et al. 1995; Peng et al. 1995; Hogan & Klein 1997; Hua et al. 2000). Whilst phytopathogens constitute the greatest percentage of such fungi, and are the only group that will be considered here, rarely have plant pathogenic fungi been considered and reviewed in detail (Wang & Leong 1989; Hargreaves & Turner 1992; Mullins & Kang 2001). Therefore, whilst the role of yeasts, lower eukaryotes and other filamentous fungi is acknowledged where relevant during this review, examples where possible refer to fungal pathogens of plants.
6.4 Techniques Typically, transformation protocols involve the preparation of fungal protoplasts, delivery of transforming DNA, and the regeneration and selection of transformants. The first step involves the release of protoplasts from starting cells. Germinated asexual spores, mycelium fragments or basidiospores are often utilised depending on species and convenience and under osmotically stable conditions, the cell wall is removed through the addition of hydrolytic enzymes. Young cells propagated in liquid culture are preferred and according to Vollmer and Yanofsky (1986) protoplasts can be stored for extended periods at –70°C with minimal loss of competence. Several factors affecting the release of protoplasts in filamentous fungi, including the nature and molarity of osmotic stabilisers, pH of lytic medium and the age and amount of starting material have been considered in detail (Peberdy et al. 1976). However, often the most critical factor involved in protoplast preparation is the concentration and duration of exposure to enzymes during lytic digestion. As a result, numerous enzyme preparations have been used to generate fungal protoplasts (Peberdy 1989). Historically, researchers employed mycolytic enzymes obtained as by–products from other microorganisms, but more recently several commercial preparations have been utilised including Helicase (Biological Industries, France), Zymolase 100T (Kirin Brewery Company, Japan) and Driselase (Sigma Chemical Company, USA). Currently Novozyme 234, a mixture obtained from the fungus Trichoderma viride (Novo Enzyme Products, UK) is the most popular preparation, but batch to batch variation has prompted some workers to construct better defined enzyme preparations (Binninger et al. 1987).
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Periods of exposure range from a few minutes to several hours depending on the starting material, enzyme preparation and fungal species involved. Workers should remove cell debris and enzyme residues through successive washing and centrifugation following digestion, taking care to avoid protoplast lysis during resuspension. Osmotic stabilisers such as sodium chloride, potassium chloride, magnesium sulphate, mannitol, sorbitol or sucrose may be used to stabilise protoplasts during these procedures. The isolation of fungal protoplasts is considered in more detail by Perberdy (1991) and Hashiba (1992). Despite numerous modifications, only three basic methods have been utilised to induce uptake of incoming DNA by competent cells. Adapted from the work of Yelton et al. (1984), Ballance and Turner (1985) and Vollmer and Yanofsky (1986), these protocols are presented in a review by Hargreaves and Turner (1992) and differ almost solely on the size and concentration of polyethylene glycol (PEG) employed. In general, µg quantities of transforming DNA are incubated in the presence of large numbers of protoplasts and calcium ions. After durations ranging from 10 to 30 minutes, several high concentration aliquots of polyethylene glycol (PEG) are administered. Protoplasts aggregate, fuse and become temporarily permeable to DNA, during which time molecular transformation may occur. Where possible, exposure of protoplasts to transforming solutions should be minimised in order to reduce the lethal side effects associated with prolonged PEG treatment. As an alternative to chemical methods of DNA induction, electroporation has shown considerable promise (Chakraborty & Kapoor 1990). The application of high amplitude electric pulses in short bursts renders the biomembranes of protoplasts temporary permeable to incoming DNA and, therefore, amenable to molecular transformation (Watts & Stacey 1991). To date, few examples apart from those involving species of Aspergillus (Ward et al. 1989; Ozeki et al. 1994; Kwon–Chung et al. 1998) and Neurospora (Kothe & Free 1996; Vann 1995) are available in the literature, and evidence for significant application of electroporation–mediated transformation to fungal pathogens of plants is minimal (Marek et al. 1987; Bej & Perlin 1988; Richey et al. 1989; Chakraborty et al. 1991; Redman & Rodriguez 1994). Nevertheless, some workers continue to develop new and innovative methods of transformation. For example, Sanchez & Aguirre (1996) adapted an electroporation method originally developed for yeast cells of Kluyveromyces lactis (Sanchez et al. 1993) to germinated conidia of Aspergillus nidulans, thus circumventing completely the requirement for protoplast preparation. Recently, this technique was also utilised by Dantas– Barbosa et al. (1998), during the first successful transformation of the thermophilic fungus Humicola grisea var. thermoidea.
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Furthering the tradition of protoplast–free techniques, several additional whole cell techniques including lithium acetate treatment, biolistic transformation and Agrobacterium tumefaciens– mediated transformation (ATMT) have been developed and adopted for use with phytopathogens. To date, the use of high concentrations of lithium ions, which avoids protoplast formation by rendering the cell wall permeable to incoming DNA (Dhawale et al. 1984), has not found widespread use, and few examples of phytopathogens transformed via this method are available (Dickman 1988; Soliday et al. 1989; Bej & Perlin 1989; 1991; Leslie & Dickman 1991). In contrast, biolistic transformation (particle bombardment) and ATMT, whereby whole cells are either bombarded with DNA–containing particles or co–incubated with DNA–carrying strains of A. tumefaciens, have been adopted extensively. In the former, tungsten or gold particles are coated with transforming DNA and accelerated at high velocity into starting cells (Klein et al. 1987). Particles generally penetrate both the cell wall and underlying cell layers piercing intact cells non–lethally without the trauma of chemical or electrical methods, and avoiding the problems associated with protoplast regeneration (Watts & Stacey 1991). To date, biolistics has been applied successfully to a wide range of agriculturally significant pathogens including Botrytis cinerea (Hilber et al. 1994), Venturia inaequalis (Parker et al. 1995), Cercospora caricis (Aly et al. 2001), and several species of Phytophthora (Bailey et al. 1993). Most significantly however, has been the transformation of a range of obligate parasites for which systems were previously unavailable. These include the rusts Uromyces appendiculatus (Bhairi & Staples 1992) and Puccinia graminis f. sp. tritici (Schillberg et al. 2000), and powdery mildew fungi Uncinula necator (Smith et al. 1992) and Erysiphe graminis f. sp. hordei (Christiansen et al. 1995; Chaure et al. 2000). Unfortunately, the cost of equipment and often only transient expression of incoming DNA may limit the adoption of this technique in some laboratories. Transformation of several fungi, including Magnaporthe grisea (Rho et al. 2001) and species of Fusarium (Covert et al. 2001; Mullins et al. 2001), Calonectria (Malonek & Meinhardt 2001), Colletotrichum (De Groot et al. 1998) and Mycosphaerella (Zwiers & De Waard 2001) has also been mediated through the plant pathogenic bacterium A. tumefaciens, which has long been used to transfer genes to a wide variety of plants (Zupan & Zambryski 1995). In plants, the interaction between the two organisms results in the integration of a section of the bacterial plasmid DNA, termed T–DNA, into the host plant genome. Bound by left and right border sequences, and containing the virulence region essential for infection, the T–DNA segment effectively engineered to incorporate foreign genes, can be employed to transfer such genes to the host of choice (Sheng & Citovsky 1996). In fungi, this process currently requires further investigation, but it is speculated that similar mechanisms are involved (Dunn–Coleman & Wang 1998; Mullins & Kang 2001). 95 Chapter 6. Transformation
More recently, A. rhizogenes, a second Agrobacterium species capable of T–DNA transfer has been identified and subsequently used to transfer genes to several Brassica species (Henzi et al. 2000; Cogan et al. 2002).
6.5 Protoplast Regeneration Subsequent to protoplast–mediated transformation, putative transformants are generally transferred to solid media and shortly thereafter exposed to suitable selection pressures. In order to obtain growing colonies, osmotically stabilised media similar to those described for protoplast preparation are required to enable protoplast cell walls to regenerate. If transformation involves complementation of auxotrophic mutations, selection to prototrophy through adjustment of plating media to specific nutritional specifications is straight forward. However, where selection is based on expression of drug resistance, a period of recovery in the absence of selective pressure is essential to allow gene expression to develop to levels conferring resistant phenotypes (Ward et al. 1986). For this purpose many groups employ non–selective basal media as sources for regeneration, after which selective overlays are applied to obtain the desired type of transformant (Wang et al. 1988). Although a large proportion of protoplasts will produce cell wall material during regeneration, only a fraction will progress to form functional cells and hyphae (Peberdy 1991).
6.6 Selection of Transformants In order to identify transformants it is essential that a gene conferring a selective advantage to the transformed cell be included in the transforming DNA. Presently, a wide range of systems incorporating numerous antibiotic, fungicide and herbicide resistance genes are available including several systems employing visual selection of transformants. However, prior to the introduction of dominant and semi–dominant drug resistance markers, studies focused solely on the rescue of auxotrophic mutants by complementation with genes corresponding to certain nutritional deficiencies. For example, Unkles, Campbell, Carrez et al. (1989) transformed chlorate resistant Aspergillus niger and A. oryzae mutants (niaD–) defective in nitrate reductase, to prototrophy using the niaD+ gene from A. nidulans. Hahm & Batt (1988) transformed argB mutants of A. oryzae with genes from A. nidulans. Buxton et al. (1989) complemented selenate resistance mutants of A. niger with an ATP sulphurylase gene (sC+) isolated from A. nidulans and Gouka et al. (1993) successfully transformed fluoroacetate–resistant mutants of Penicillium chysogenum using acetyl CoA synthetase genes (facA) obtained from sensitive strains.
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Additionally, acetamide utilisation conferred by the amdS gene from A. nidulans (Hynes et al. 1983), ornithine carbamoyltransferase (OCTase) activity (Penttila et al. 1987) and fluoro–orotic acid resistance have been used as selectable markers, the latter producing auxotrophs selectable through complementation with specific pyrimidine biosynthesis genes (Ballance et al. 1983; Alic et al. 1989; 1990). Unfortunately, the difficulty or impossibility, in many cases, of isolating recipient strains for the transforming DNA limited auxotrophic complementation as a means of selection for many fungal species and to date only a handful of phytopathogens have been transformed by this technique. Examples include C. heterostrophus to acetamide utilisation (Turgeon et al. 1985), U. maydis (Banks & Taylor 1988; Kronstad et al. 1989) and Claviceps purpurea to fluoro–orotic acid resistance (Smit & Tudzynski 1992), F. oxysporum (Malardier et al. 1989; Langin et al. 1990; Diolez et al. 1993), F. moniliforme (Sanchez–Fernandez et al. 1991) and B. cinerea to chlorate resistance (Levis et al. 1997), and F. solani to ArgB+ through complementation of OCTase mutations in the arginine biosynthesis pathway (Rambosek & Leach 1987). In some cases model pathogens such as Magnaporthe and Colletotrichum have proven amenable to several different systems (Parsons et al. 1987; Rodriguez & Yoder 1987; Daboussi et al. 1989), but, this seems to be the exception rather than the rule for many phytopathogens. A more comprehensive list of auxotrophic markers used during transformation of filamentous fungi is provided by Van den Hondel and Punt (1991). As an alternative, genes conferring antibiotic or fungicide resistance are increasingly employed as selectable markers during fungal transformations. Once integrated into the genome of the recipient species, these genes provide the resulting transformants with the ability to grow in the presence of compounds otherwise toxic to untransformed wild–type cells. Provided recipient fungi are susceptible to the drug of choice, have low levels of spontaneous mutation to drug resistance and expression of such genes can be garnered through coupling to fungal promoters, the range of potential markers is extensive. Selection based on drug resistance is usually conferred by genes encoding resistance to glycopeptide antibiotics phleomycin and bleomycin, aminoglycosides such as geneticin (G418) or the aminocyclitol antibiotic hygromycin B. Phleomycin and bleomycin are closely related broad spectrum antibiotics produced by a number of bacterial species including Streptomyces verticillus (Austin et al. 1990), Streptoalloteichus hindustanus (Drocourt et al. 1990), Streptococcus hindustanus (Mattern et al. 1988) and Staphylococcus aureus (Semon et al. 1987). Effective against both prokaryotes and eukaryotes, glycopeptide antibiotics kill susceptible cells through site specific cleavage of single and double stranded DNA (Ueda et al. 1985; Hertzberg et al. 1985).
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Resistance, conferred by the ble gene, is in the form of a protein product which binds reversibly to the antibiotic resulting in drug sequestration (Gatignol et al. 1988). Despite functioning equally well as agents for selection, bleomycin tends to find application in plants (Hille et al. 1986), and is used rarely in fungi (Van Engelenburg et al. 1989; Kinal et al. 1993). In contrast, phleomycin has been used extensively in yeasts (Gatignol et al. 1987; Glumoff et al. 1989) and filamentous fungi (Punt & Van den Hondel 1992) and is also preferred by those working with plant pathogenic fungi (Ball et al. 1991; Pilgeram & Henson 1990; Gold et al. 1994). Resistance to the aminoglycoside antibiotic geneticin, also known as G418, has also been employed, albeit on a limited basis, as a selectable marker during transformation of yeasts (Gmunder & Kohli 1989; Chen et al. 1989) and some pathogenic fungi (Marek et al. 1989; Churchill et al. 1990; Aragona & Porta–Puglia 1993). G418 inhibits protein synthesis both in prokaryotes and eukaryotes (Jimenez & Davies 1980). Resistance is conferred by genes encoding two aminoglycoside phosphotransferases, APH(3’)–I and –II, enzymes that detoxify the antibiotic through site directed phosphorylation (Haas & Dowding 1975). Unfortunately, high levels of natural resistance, variability in antifungal activity of commercial preparations and the accompanying high cost of the compound have resulted in geneticin being replaced in some cases by less expensive aminoglycosides such as neomycin (Woestemeyer et al. 1987; Banks 1983; Judelson 1993; Noel et al. 1995) and kanamycin (Reiss et al. 1984; Arnau et al. 1988). By far, the most popular marker for fungal transformation studies is the aminocyclitol hygromycin B, which has been employed for selection of transformants of many plant pathogens (Table 6.1). Hygromycin B (HmB) is produced by Streptomyces hygroscopicus (Pettinger et al. 1953) and inhibits protein synthesis in prokaryotes and eukaryotes by impeding enzyme translocation (Gonzales et al. 1978), causing mistranslation (Singh et al. 1979). Plasmid–borne genes encoding hygromycin B phosphotransferase (hph), an enzyme which inactivates HmB, have been described by Kaster et al. (1983) and Gritz and Davies (1983), and vectors containing the E. coli hph gene under the control of regulatory sequences from yeasts (Kaster et al. 1984) and filamentous fungi have been constructed (Punt et al. 1987; Carroll et al. 1994; Orbach 1994). Other less popular markers include resistance to blasticidin, (Kimura et al. 1995) and oligomycin C (Ward et al. 1988; Bull et al. 1988).
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Table 6.1: Some plant pathogenic fungi transformed to hygromycin B resistance. Fungal species1 Host/s Reference Alternaria alternata Tomato Shiotani & Tsuge 1995
Botryotinia fuckelianaa Grape Hamada et al. 1994 Botryotinia squamosa Onion Huang et al. 1989
Claviceps purpurea Rye Comino et al. 1989 Cochliobolus carbonum Maize Scott–Craig et al. 1990 C. heterostrophusb Maize Turgeon et al. 1987 C. sativusc Barley, wheat Liljeroth et al. 1993 Cryphonectria (syn. Endothia) parasitica Chestnut Kim, Rigling et al. 1995
Fulvia fulva (syn. Cladosporium fulvum) Tomato Oliver et al. 1987 Fusarium culmorum Wheat, cereal Curragh et al. 1992 F. graminearum Wheat, barley Wiebe et al. 1997 F. oxysporum f. sp. conglutinans Cabbage Powell et al. 1987 F. oxysporum f. sp. erythroxyli Coca Bailey et al. 2002 F. oxysporum f. sp. lycopersici Tomato Powell et al. 1987 F. oxysporum f. sp. niveum Watermelon Kim et al. 1995 F. oxysporum f. sp. raphani Radish Kistler & Benny 1988 F. solani f. sp. pisi Pea Dickman & Kolattukudy 1987 F. subglutinans Mango Freeman et al. 1999
Gibberella fujikuroid Rice, maize Fernandez–Martin et al. 2000 Gibberella pulicarise Wheat, maize Salch & Beremand 1988 Glomerella cingulataf Avocado, legume Rikkerink et al. 1994
Leptosphaeria maculansg Brassica spp. Farman & Oliver 1992 Leptosphaeria nodorumh Wheat Cooley et al. 1991
Magnaporthe griseai Rice Leung et al. 1990 Mycosphaerella graminicolaj Wheat Pnini–Cohen et al. 1996 Mycosphaerella pinik Coniferous spp. Bradshaw et al. 1997
Nectria haematococcal Cucurbita spp. Crowhurst et al. 1992
Phytophthora capsici, P. parasitica Apple, pepper Bailey et al. 1991 P. infestans Potato, tomato Judelson et al. 1991 Pseudocercosporella herpotrichoides Wheat, rye, barley Blakemore et al. 1989 Pyrenopeziza brassicaem Brassica spp. Ashby & Johnstone 1993
Thanatephorus cucumerisn Potato Robinson & Deacon 2001
Ustilago hordei, U. nigra Barley Holden et al. 1988 U. maydis Maize Kamper et al. 1995
Verticillium dahliae Potato, tomato Dobinson 1995 1Anamorph: aBotrytis cinerea, bBipolaris maydis, cBipolaris sorokiniana, dFusarium moniliforme, eFusarium sambucinum, fColletotrichum gloeosporioides, gPhoma lingam, hSeptoria nodorum, iPyricularia oryzae, jSeptoria tritici, kDothistroma pini, lFusarium solani f. sp. cucurbitae, mCylindrosporium concentricum, nRhizoctonia solani.
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In addition to antibiotics, the use of fungicides or herbicides as selectable markers has received considerable attention. Mutations in genes encoding β–tubulin have been demonstrated to confer resistance to the benzimidazole fungicide benomyl and related compounds, in turn serving as efficient selectable markers (Orbach et al. 1986; Payne et al. 1998). Insensitive forms of the β– tubulin gene have reduced affinity for benzimidazole–containing compounds and transformation of sensitive strains with genes cloned from benomyl–resistant mutants has been shown to confer resistance in a wide range of fungal plant pathogens such as Gaeumannomyces graminis (Henson et al. 1988), Colletotrichum graminicola (Panaccione et al. 1988), Aspergillus flavus (Seip et al. 1990), Cercospora kikuchii (Upchurch et al. 1991), Septoria nodorum (Cooley & Caten 1993) and Magnaporthe grisea (Kachroo et al. 1997). More recently, the herbicide bialaphos has received attention as a versatile selectable marker for transformation of filamentous fungi including N. crassa (Avalos et al. 1989), Pleurotus ostreatus (Yanai et al. 1996), Paecilomyces fumosoroseus (Cantone & Vandenberg 1999) and several phytopathogens (Straubinger et al. 1992; Upchurch et al. 1994; Leung et al. 1995). Bialaphos contains phosphinothricin (PPT), an analogue of glutamate which is an inhibitor of glutamine synthase, an enzyme responsible for detoxifying poisonous plant by–products such as ammonia (De Block et al. 1987). The gene bar, isolated from S. hygroscopicus and characterised by Thompson et al. (1987), confers resistance to bialaphos by encoding the enzyme phosphinothricin acetyltransferase responsible for detoxifying PPT through acetylation (Murakami et al. 1986). Originally used to engineer herbicide resistance in plants (De Block et al. 1987), a series of more compact vectors specifically designed for use in fungi have since been constructed by Pall and Brunelli (1993). As an adjunct to the wide range of selectable markers conferring drug resistance, several reporter or gene fusion systems have been developed to study gene expression in fungi. The most frequently used reporter gene is probably the lacZ gene from E. coli which encodes β–galactosidase (Van Gorcom et al. 1985). Expression and regulation of genes fused to the lacZ gene can be assayed quantitatively by measuring β–galactosidase activity and expression can be detected in vivo using the chromogen X–Gal (Kolar et al. 1988). Unfortunately, endogenous enzyme levels in some systems are sufficient to make detection of chimeric β–galactosidase difficult by enzymatic methods. Coupled with the large size of the gene, in vitro construction and analysis of transforming vectors is rarely ideal. Fortunately, several other systems are available. The GUS gene fusion system described by Jefferson et al. (1986) utilises the E. coli uidA gene which codes for β– glucuronidase. β–glucuronidase catalyses the cleavage of a wide variety of substrates, many of which can be assayed by fluorimetry, spectrophotometry or via histochemical analysis.
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Transformation with vectors containing GUS fusions has enabled researchers to study gene expression in a wide range of plant pathogens including Cochliobolus (Monke & Schafer 1993), Pseudocercosporella (Bunkers 1991), Fulvia (Roberts et al. 1989), Leptosphaeria (Oliver et al. 1993; Chen & Seguin–Swartz 1997), and several species of Fusarium (Couteaudier et al. 1993; Doohan et al. 1998; Yates et al. 1999; Olivain & Alabouvette 1999). One of the most popular applications however, involves the use of GUS–fusions to monitor isolates of the biocontrol fungus Trichoderma harzianum, which is currently under investigation as a biocontrol agent for a range of both aerial and soilborne fungal plant pathogens (Green & Jensen 1995; Thrane et al. 1995; Bowen et al. 1996). A more versatile system involves the use of green fluorescent protein (GFP), a chromophore isolated from the jellyfish Aequorea victoria. Similar gene fusions are involved and expression can be detected in planta using anti–GFP antibodies or fluorescent microscopy (Chalfie et al. 1994). To date, researchers have used GFP fusions to study pathogenic development in fungi during infection of banana, plantain (Balint–Kurti et al. 2001), beans (Dumas et al. 1999), maize (Spellig et al. 1996; Maor et al. 1998) and canola (Sexton & Howlett 2001). Liu & Kolattukudy (1999) have also used GFP technology to study in vitro gene expression during appressorium formation in the rice blast pathogen Magnaporthe grisea.
6.7 Fate of transforming DNA In contrast to yeast and bacteria, transformation in filamentous fungi generally occurs through integration of vector sequences into the host genome (Esser & Mohr 1986). In general three types of integration events are recognised (Hinnen et al. 1978). Types one and three both involve homologous integration of the transforming DNA. In the former, termed homologous additive integration, crossing over results in tandemly arranged alleles separated by plasmid sequences. However, in the latter gene conversion replaces crossing over and homologous interaction between the plasmid gene and chromosome results in gene replacement at the expense of plasmid sequences. Finally, during ectopic or type two integration, single crossovers between plasmid DNA and non– homologous chromosome sites result in random or non–homologous integration of plasmid DNA at variable locations (Fincham 1989). Similar to mammalian cells, ectopic integration in fungi is believed to result from ligation of incoming DNA into nicks or breaks in chromosomal sequences (Asch et al. 1992; Schiestl et al. 1993; 1994).
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On some occasions multiple copy integration may also be observed both during ectopic and homologous recombination (Wernars et al. 1985). During homologous integration recombination between circular plasmids may result in the appearance of tandem arrays through integration of circular oligomers. Alternatively, successive rounds of recombination into a single locus or into previously integrated plasmid copies may occur. Similarly multi–copy ectopic integration may take place through a single round of non–homologous integration followed thereafter by multiple events of homologous recombination (Fincham 1989). The frequency of individual recombination events is dependent upon a multitude of factors, notwithstanding the methods and conditions for transformation, which have been studied in detail by Specht et al. (1988; 1991). To date, approaches to increase such events in filamentous fungi have focused on two main areas; linearisation of vector and/or host sequences by restriction enzyme digestion, and vectorial incorporation of DNA sequences homologous to the recipient chromosome. The role of DNA conformation during transformation first received attention when Orr–Weaver et al. (1981) demonstrated that linear DNA molecules transformed strains of S. cerevisiae 10– to 1000–fold more efficiently than their circular counterparts. Shortly thereafter Dhawale and Marzluf (1985) and Skatrud et al. (1987) demonstrated similar results in the filamentous fungi Neurospora crassa and Cephalosporium acremonium. By digesting circular plasmids with restriction enzymes the interaction between highly recombinogenic DNA ends and homologous chromosomal sequences is increased (Struhl 1983). An extension of this technique referred to as REMI or restriction enzyme–mediated integration involves the addition of restriction enzymes to facilitate linearisation both of the introduced plasmid and of the host genome at analogous sites. Reviewed extensively (Brown & Holden 1998; Riggle & Kumamoto 1998; Kahmann & Basse 1999; Maier & Schafer 1999), this process further increases the probability of transformation by providing large numbers of potential integration sites within the genome. First reported in yeast by Schiestl and Petes (1991), REMI has been used successfully to increase the transformation efficiency in the slime mold Dictyostelium discoideum (Kuspa & Loomis 1992), the model filaments Neurospora crassa (Garnand & Nelson 1995) and Aspergillus nidulans (Sanchez et al. 1998) and both the rice blast and maize pathogens, Magnaporthe grisea and Colletotrichum graminicola (Shi et al. 1995; Thon et al. 2000). More recently, numerous papers have been published that describe the application of REMI to gene functionality studies in phytopathogens. Examples of pathogenicity related genes cloned by REMI include the AAL and PM–toxin synthesising genes of Alternaria alternata and Mycosphaerella zeae–maydis, respectively (Akamatsu et al. 1997; Yun et al. 1998), the polyketide synthase gene required for T–toxin biosynthesis in Cochliobolus heterostrophus (Lu et al. 1994), and the victorin biosynthesis gene of C. victoriae (Churchill et al. 1995). 102 Chapter 6. Transformation
As a rule, REMI favours single vector integration and results in transformants that generally possess only single copies of the integrated DNA. These qualities enable researchers to rescue the integrating DNA in conjunction with portions of the tagged gene (Mullins & Kang 2001). In some circumstances however, the double stranded breaks created by the addition of restriction enzymes are not always repaired effectively in the absence of vector incorporation or chromosomal religation, and the occurrence of deletions or rearrangements during transformation may result (Bolker et al. 1995; Turgeon et al. 1995; Linnemannstons et al. 1999). Nonetheless, REMI has proven invaluable in dissecting the mechanisms of pathogenesis in several important pathogens including Cochliobolus heterostrophus (Yang et al. 1996), Magnaporthe grisea (Sweigard et al. 1998; Balhadere et al. 1999; Fang & Dean 2000) and several species of Colletotrichum (Epstein et al. 1998; Redman et al. 1999; Yakoby et al. 2001). If REMI is to employed however, the conditions, especially the temperature at which protoplasts are prepared and regenerated following transformation, is critical to the success of the procedure (Cockram & Sealy–Lewis 2000). The incorporation of sequences within transforming DNA which are homologous to regions within the genome of the recipient species are often employed to increase transformation frequencies in fungi. By targeting plasmid integration to regions known to be present at multiple sites within the fungal genome it is assumed that plasmid copy numbers and hence transformation efficiencies can be improved. Although this approach was not successful in Neurospora crassa (Russell et al. 1989), vectors harbouring repetitive ribosomal DNA sequences have been used to transform Alternaria alternata (Tsuge et al. 1990), the human pathogenic fungus Scedosporium prolificans (Ruiz–Diez & Martinez–Suarez 1999), and the oil producing fungus Mortierella alpina at high frequencies (MacKenzie et al. 2000). In addition to homologous elements, vectors containing ARS or autonomously replicating sequences have been developed for some fungi. ARS sequences promote autonomous replication of the plasmids on which they are carried, are maintained extrachromosomally and replicate to high copy numbers in the cytosol, by virtue of their action as origins of replication (Esser & Mohr 1986). Because they can be propagated both in prokaryotic and eukaryotic cells they also facilitate the development of efficient cloning systems often referred to as shuttle vectors (Paietta & Marzluf 1985). In yeast and lower fungi, vectors containing ARS sequences significantly increased transformation efficiencies in S. cerevisiae (Struhl et al. 1979) and Phycomyces blakesleeanus (Revuelta & Jayaram 1986), and similar sequences have since been identified in a wide range of fungal species including Mucor circinelloides (Roncero et al. 1989), Pleurotus ostreatus (Peng et al. 1993), Aureobasidium pullulans (Thornewell et al. 1995) and Penicillium chrysogenum (Fierro et al. 1996).
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To date however, few sequences conferring replicative transformation in plant pathogenic fungi have been identified. Tsukuda et al. (1988) isolated and characterised a 383 bp sequence, UARS1, from Ustilago maydis. When inserted into an integrative vector, the ARS–containing fragment directed autonomous replication of the plasmid and generated large numbers of unstable transformants containing up to 25 copies of the plasmid per cell. Samac and Leong (1989) also demonstrated replicative transformation in U. maydis by ligating a 1211 bp terminal inverted repeat obtained from Nectria haematococca into an integrative vector. More recently autonomous replicating plasmids have been described for the maize pathogens U. maydis (Kinal et al. 1991) and Fusarium moniliforme (Bruckner et al. 1992), the grey mold pathogen Botrytis cinerea (Santos et al. 1996) and the tomato pathogen F. oxysporum f. sp. lycopersici (Garcia–Pedrajas & Roncero 1996). Fotheringham and Holloman (1992) have also described replicative transformation in U. maydis by linear DNA lacking ARS sequences. Centromeric and telomeric sequences, which contain ARS–like elements, have also been investigated for development of replicating vectors in plant pathogenic fungi. The discovery that some fungi add terminal telomeric repeats to transforming DNA in vivo, resulting in the production of linear extrachromosomal elements capable of autonomous replication was first demonstrated by Powell and Kistler (1990). By exploiting this phenomenon, these authors created high efficiency, autonomously replicating vectors in Fusarium oxysporum. Thought to represent replication origins for chromosomes, the addition of terminal telomeric repeats to transforming DNA resulted in a greater than 1000–fold increase in the transformation efficiency in this organism, and also transformed several other fungal species including the plant pathogens F. solani and Cryphonectria parasitica at high frequencies (Powell & Kistler 1990). In similar studies Woods and Goldman (1992; 1993) identified and later rescued telomeric sequences from Histoplasma capsulatam. By incorporating telomeric sequences in transforming DNA these authors were able to construct high efficiency shuttle vectors in H. capsulatum and provide the first demonstration of foreign gene expression in this organism (Woods et al. 1998). So far, only two other fungal species have been identified which are capable of in vivo conversion of transforming DNA to linear extrachromosomal elements. These include the basidiomycetous yeast Cryptococcus neoformans (Edman 1992) and the taxol–producing fungus Pestalotiopsis microspora (Long et al. 1998). While an explanation for this phenomenon is not currently available, both the diverse phylogenetic relationship between these organisms and the results of a recent study that exploits human telomeric sequences to direct autonomous vector replication in fungi (Aleksenko & Ivanova 1998), may suggest that the practice is more common than currently recognised.
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In addition to the many features outlined above, high transformation frequencies can often be obtained in filamentous fungi through co–transformation, which is particularly useful in situations where transforming genes cannot be selected directly. In such cases researchers may pursue the desired gene in correspondence with a more readily selectable marker, introduced separately on an alternative vector, thus circumventing the construction of additional vectors (Fincham 1989). Although only a small fraction of cells appear to be competent for transformation (Grotelueschen & Metzenberg 1995), those that are amenable to transformation readily admit multiple DNA molecules (Maio et al. 1995) and extremely high frequency integration of unselected plasmids can be obtained in some species (Wernars et al. 1987; Austin & Tyler 1990). In plant pathogenic fungi, transformation frequencies are generally more conservative. Cooley and Caten (1988) and Cooley et al. (1990) reported co–transformation frequencies of approximately 50% in Septoria nodorum, whilst Vaillancourt and Hanau (1994) demonstrated frequencies between 30% and 87% in Colletotrichum graminicola depending on selection method. Recently, co–transformation has found particular application in the transformation of fungi being investigated as potential biocontrol agents for agricultural pests including locusts, grasshoppers (St. Leger et al. 1995; Inglis et al. 2000) and several mycoparasites (Bae & Knudsen 2000; Jones et al. 1999). This chapter represents our attempts to develop a transformation system for R. alismatis. In addition to the development of techniques both for the production of protoplasts and efficient transformation in this species, we aspired to increase the penetrative ability of R. alismatis by transforming the fungus with a vector containing a cutinase gene construct from F. solani f. sp. pisi. Expression of this gene is essential for cuticular penetration in some plant pathogens and transformation and expression in R. alismatis may facilitate penetration in S. montevidensis by inocula previously unable to breach the plant cuticle, as demonstrated by Dickman et al. (1989) in the Mycosphaerella–papaya pathosystem. Additionally, if entry level resistance mechanisms observed during invasion of S. graminea could be overcome by encouraging more rapid penetration of the host, for example by increasing gene copy number and ultimately expression of pathogenicity–related genes such as cutinase, the current constraints to field infection of S. graminea may be reduced.
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6.8 Materials and Methods 6.8.1 Fungal isolates: The R. alismatis isolate RH097 (DAR 73151), selected in chapter 5 based on its pathogenicity to the three plant species, was also used during this study. The collection, isolation, cultivation, storage, maintenance of this culture was as described in section 3.4.1. The fungus was cultivated on LBA, spores were harvested, enumerated as described in section 3.4.2 and approximately 1 × 109 spores were transferred to each of several 20 cm diameter plastic Petri dishes containing clarified 20% v/v V8 juice. The final volumes was adjusted to 50 mL, and dishes were incubated at 28°C with gentle agitation for between 4 and 24 hours, or until spores appeared to have germinated. Germination was associated with an increase in spore diameter following hydration and a woolly appearance.
6.8.2 Vector construction and preparation: An expression vector (pABC) containing the Fusarium solani f. sp. pisi (fsp) cutinase gene with its promoter and terminator, and the E. coli hygromycin–B phosphotransferase (hph) gene driven by the Aspergillus nidulans gpd promoter and trpC terminator constructed by Dr C. Y. Chen was obtained from Agriculture and Agri–Food Canada, Saskatoon, Saskatchewan, Canada. The vector (Figure 6.1), which also contains the ampicillin resistance (amp) gene for selection in bacteria, was constructed as follows. The superlinker from pSL 1180 (Amersham Pharmacia Biotech) was released by restriction with HindIII–EcoRI and cloned into the HindIII–EcoRI site of pBluescript II KS (Stratagene, Heidelberg, Germany); the resultant plasmid was designated pSUPERBKS II. The cutinase gene with its promoter and terminator from F. solani f. sp. pisi was released from pU5–11 (Dickman et al. 1989) by AgeI and SalI restriction, and cloned into pUC–19 (New England Biolabs, Beverly, Massachusetts); the resulting plasmid was designated pAUDY. The plasmid pAUDY was digested with KpnI and PstI to release the 1.7 kb cutinase construct, and cloned into the NruI–PstI site of pSUPERBKS II; the resulting plasmid was designated pCHEN1. Finally, the cutinase construct (fsp) was released from pCHEN1 by digestion with BamHI–SfiI and cloned into the corresponding BamHI–SfiI site of pAN26 (Taylor & Borgmann 1996; GenBank accession number U09715); the resultant plasmid was designated pABC.
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Figure 6.1: Plasmid map of the pABC expression vector, containing the F. solani f. sp. pisi cutinase gene construct (fsp cut) with its promoter and terminator, and the E. coli hygromycin B phosphotransferase (hph) gene driven by A. nidulans promoter (gpd) and terminator (trpC) for selection in fungi. The vector also contains the ampicillin resistance (Ampr) for selection in E. coli. Plasmid map constructed using BioEdit version 5.0.9 (Hall 1999).
The vector pABC was obtained within E. coli strain DH5 (Clontech), streaked on Luria–Bertani (LB) agar containing 50 µg/mL ampicillin and incubated at 37ºC for 24 hours. A single colony was transferred to LB broth containing 50 µg/mL ampicillin and incubated for a further 24 hours with agitation at 37ºC. Plasmid DNA was extracted using a HiSpeed plasmid midi kit according to the manufacturers protocol (QIAGEN; Mississauga, Ontario, Canada). DNA concentration was determined spectrophotometrically at 260 nm. To confirm the presence of the cutinase insert (fsp cut), the pABC vector was sequenced. Because the pABC vector comprised largely of the cosmid vector pAN26 designed by Taylor and Borgmann (1996), primers matching specific sites on the pAN26 vector were used to initiate the sequencing process. Additional primers were designed and constructed from data obtained thereafter, and from the original GenBank accession (M29759) made by Soliday et al. (1989), from which Dr C. Y. Chen at Agriculture and Agri–Food Canada obtained the original pU5–11 vector containing the ~1.8 kb cutinase construct. Primer construction and vector sequencing were completed at the Plant Biotechnology Institute, National Research Council of Canada, Saskatoon, Saskatchewan, Canada by B. Panchuk and D. Schwab, respectively.
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All primers constructed including size, nucleotide sequence, and sites on the pABC vector to which they correspond are shown in Table 6.2. Several co–transformations using the vector pBARGEM7–2 (Pall & Brunelli 1993) that contains the bar gene as a fungal selectable marker conferring resistance to the herbicide bialaphos or ‘Ignite’ (Figure 6.2) were also performed (Table 6.3).
Figure 6.2: Plasmid map of the pBARGEM7–2 vector, which carries the gene bar, under the control of A. nidulans regulatory sequences, as a selectable marker for basta/Ignite resistance in fungi. Additionally, the vector contains both an ampicillin resistance gene (Ampr) and pUC type origin for growth and selection in E. coli. Source: Pall and Brunelli (1993).
6.8.3 Preparation of Protoplasts: Protoplasts were prepared essentially as described by Yelton et al. (1984). Freshly harvested spores of R. alismatis were incubated with agitation at room temperature in 10% V8 juice. Germinated spores were transferred to 50 mL Falcon tube(s) and centrifuged at 4°C for 20 minutes at 5000 rpm. The supernatant was discarded and spores were re–suspended in 1 mL of 0.6 M
MgSO4, transferred to Eppendorf tubes and centrifuged for 3 minutes at 5000 rpm. The supernatant was again discarded and spores were re–suspended in 1 mL of osmotic medium (1.2 M MgSO4; 10 mM sodium phosphate (Na2HPO4/NaH2PO4) buffer, pH5.8). This suspension was transferred to a 10 cm diameter plastic Petri dish containing 4 mL of osmotic medium and 8 mg/mL lysing enzyme, 6.4 mg/mL glucuronidase (type B–1, bovine liver), and 50 µg/mL chitinase and incubated at 28°C overnight (16–24 hours) with gentle agitation. The protoplasting medium was previously sterilised by passing the solutions through membrane filters (0.2 µm).
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The reaction was arrested prior to the appearance of cell wall debris, which indicated over exposure to enzymes. Protoplasts were sedimented by centrifugation (20 minutes, 6000 rpm) at 4°C, following the addition of 5 mL of ice–cold trapping buffer (0.6 M sorbitol; 100 mM Tris–HCl, pH 7.0) via overlay at a rate of 2.5 mL per minute. Cell wall debris and spores pelleted while protoplasts formed a thin band between the trapping buffer and osmotic buffer. Protoplasts were transferred to a 50 mL Falcon tube. A further 5 mL of trapping buffer was added to the above suspension and the procedure repeated to increase the yield of protoplasts. An additional 5 mL of osmotic medium (underlay) and 5 mL of trapping buffer (overlay) were added to the Falcon tube containing protoplasts and again centrifuged at 5000 rpm, 4°C for 20 minutes. The protoplast suspension was transferred to Eppendorf tubes(s), centrifuged at 10 000 rpm, 4°C for 5 minutes, the supernatant discarded and pellet resuspended in 1 mL of ice–cold STC buffer (1.2 M sorbitol; 10 mM CaCl2; 10 mM Tris–HCl, pH 7.0). Protoplasts were washed twice by consecutive centrifugation and re–suspension in STC buffer. Finally, the protoplast titre was enumerated using a Weber haemocytometer (Crown Scientific Pty. Ltd.) and adjusted to between 1 × 106 and 1 × 107/100 µL via the addition of sterile distilled water.
6.8.4 Transformation and plating: Based on the method of Andrianopoulos and Hynes (1988). The protoplast suspension (100 µL) was transferred to a 15 mL Falcon tube containing 5 µg of circular or linearised (NotI or SfiI) plasmid DNA (Table 6.3) and an equal volume of 2 × STC (2.4 M sorbitol; 20 mM CaCl2: 20 mM Tris–HCl, pH 7.0). The suspension was mixed thoroughly and an ice–cold 25 µL aliquot of PTC
(60% PEG 4000 (high quality); 10 mM CaCl2; 10 mM Tric–HCL, pH 7.0) was incorporated by pipetting. The suspension was incubated for 30–60 minutes on ice. A further 1 mL aliquot of room temperature PTC was then added and the mixture allowed to stand at room temperature for an 10– 30 minutes. Finally, 4 mL of liquid medium (1.2 M sucrose; clarified 10% v/v V8 juice, pH 6.0) was added. The resulting protoplast and transformation suspension (5 mL) was plated immediately or cells were allowed to regenerate overnight (16–24 hours) (Table 6.3). Plating was conducted according to the method of Farman and Oliver (1988) and involved the addition of 10 mL of molten ‘top agar’ (1.2 M sucrose; 1 % agar; clarified 10% v/v V8 juice, pH 6.0) to the transformation suspension prior to the division of the suspension into three 5 mL aliquots, each of which was transferred to a separate 10 cm plastic Petri dish containing 5 mL of solid ‘bottom agar’ (0.6 M sucrose; 2% agar; clarified 10% v/v V8 juice, pH 6.0).
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–2 –4 –2 Controls comprising (1) protoplasts in STC (10 and 10 dilutions), (2) protoplasts in H2O (10 dilution), (3) a 25 µL aliquot of the transformation mix, and (4) a ‘positive’ control (10–2 protoplasts in STC), were plated in a similar manner onto ‘bottom agar’ plates following the addition of 5 mL of molten ‘top agar’ (all plates contained a total of 10 mL). Plates were incubated overnight at 25°C. Selective pressure was applied to putative transformants and the ‘positive’ control via the addition of 5 mL of molten ‘top agar’ containing a sufficient concentration of the selective agent to provide a final concentration of 50–75 µg/mL of HmB or 20 µg/mL of Ignite (Table 6.3). Plates were wrapped with Parafilm ‘M’ (American National Can) and incubated at 25°C. Putative transformants appeared to the naked eye within a week to 10 days. Transformants were permitted to grow for two weeks before being transferred to fresh selective plates.
Table 6.2: Primers used and constructed during transformation of R. alismatis. Primer name Nucleotide sequence Tm Size Position on (°C) (bp) vector pABC sequence analysis and fsp cut probe constructiona T3 ATT AAC CCT CAC TAA AGG GA 64 20 178–197 T3+1 CCT ACT TCT AAC CCT GC 57 17 681–697 T3+2 GGC TAC ACC AAG AAC CT 59 17 1171–1187 CutmRNA–N GAG AAT TTT AGC AGG CG 61 17 1382–1398 CutmRNA–N+1 ACC TTT CCT CGA TAT CC 59 17 1893–1909 SP6bc ATT TAG GTG ACA CTA TA 45 17 2091–2075 SP2b ACA CTA TAG AAC CGC GG 61 17 2082–2066 CutmRNA–Nrevb GGC AAA GCG ATA TGT AG 59 17 1538–1522 CutmRNA–Cb CGC CTG CTA AAA TTC TC 61 17 1398–1382 T3+1revbc CTA CGA ACC AAG TTG CC 61 17 829–813 T3rev1b ATG AAG ATG ACA TCG GC 60 17 787–771 T3rev2b ATC ATC GTC CAA GCT CT 59 17 493–477 Hph PCR analysis and probe construction HPH–L GAG CCT GAC CTA TTG CAT CT 67 20 4517–4536 HPH–L2 GCA AGG AAT CGG TCA ATA 64 18 4676–4693 HPH–Rb ACT TCT ACA CAG CCA TCG GT 67 20 5233–5214 HPH–R2b CGT CAA CCA AGC TCT GAT 64 18 5103–5086 SNF1 probe construction SNF–L CAY CCN CAY ATH ATH AA TDd 17 – SNF–Rb TCN GGN GCN GCR TAR TT TD 17 – M13RP CAG GAA ACA GCT ATG AC 55 17 205–221 M13UPb TGT AAA ACG ACG GCC AGT 67 18 824–807 acutinase insert range 216–2056 (1840 bp). bposition on reverse strand. cfailed to provide sequence information. dTD = Touchdown PCR (Don et al. 1991). Single letter abbreviations for mixed base positions are as follows: Y stands for pYrimidine, R for puRine, H for non−G and N for A, T, G or C (Nomenclature Committee of the International Union of Biochemistry, 1986).
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6.8.5 Molecular analysis of putative transformants: For PCR analysis, spore suspensions of putative transformants were prepared as described in section 3.4.2, and used to inoculate 250 mL conical flasks containing 150 mL of clarified 20% v/v V8 juice. Inoculated flasks were incubated at 25°C for 14 days. Mycelium was harvested by filtration, lyophilised and ground in liquid nitrogen. DNA was extracted as described in section 3.4.2 and adjusted to 20 ng/µL by the addition of TE buffer. PCR was performed in a volume of 25µL using 50–100 ng of DNA as template, 0.2 mM each of dATP, dCTP, dGTP and dTTP, 2.0 mM of MgCl2, 1.5 units of Taq DNA polymerase, and 50 pmol of each primer. Two primer pairs synthesised by Dr. J. L. Taylor; HPHL/R and HPHL2/R2 amplifying a large 697 bp and smaller 410 bp fragment of the hph gene respectively, were used during various analyses (Table 6.2). DNA amplifications were performed with an initial denaturation of 5 minutes at 95°C, followed by 35 cycles of denaturation (30 seconds at 95°C), annealing (30 seconds at 62°C (HPHL2/R2) or 65°C (HPHL/R)) and extension (1 minute at 72°C), and a final extension of 5 minutes at 72°C. Reaction products were analysed on 1% agarose gels cast in 1 × TAE and visualised by ethidium bromide staining (0.3 µg/mL). A 1 kb plus DNA ladder (Gibco BRL Life Technologies) was used.
6.8.6 Southern analysis of putative transformants and probe construction: For Southern analysis (Southern 1975), DNA was obtained and isolated as described in section 3.4.2 or extracted from lyophilised powdered mycelium using the DNeasy plant mini kit according to the manufacturers protocol (QIAGEN). DNA was adjusted to 3 µg/µl by the addition of TE buffer and approximately 5 µg of genomic DNA from putative transformants or the untransformed controls (wild type DNA, TC1 and TC2) digested with SacI or HindIII (New England Biolabs). DNA was analysed by electrophoresis on 0.8% agarose gel cast in 1 × TAE buffer stained with 2.5 µL of a 10 mg/mL solution of ethidium bromide and visualised under ultraviolet (UV) light. DNA was capillary transferred, fixed to Hybond N nylon membranes (Amersham Pharmacia Biotech) under a source of ultraviolet radiation (254 nm), and hybridised with a 32P–labelled probe by the method of Sambrook et al. (1989). Hybridisation was at 40°C overnight in pre–hybridisation buffer containing 5 × SSPE, 5 × Denhardt’s solution, 0.5% SDS, 50% formamide and 100 µg/mL of denatured fragmented salmon sperm DNA (Sigma–Aldrich). For slot–blot analyses, DNA was denatured in 0.2 M NaOH for 10 minutes, chilled on ice, diluted to 200µL with 10 × SSC and applied to the membrane via a 48–well slot–blot manifold. Gentle suction was applied after 2 hours, followed by several rinses of 10 × SSC. The membrane was washed with 2 × SSC and fixed as described above. For dot–blot analyses the procedure was essentially as described above, although DNA was applied directly to the membrane without the use of a suction manifold.
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Where hybridisation with multiple probes was conducted, probes were removed from the membrane using either the alkali method, which involved incubation of membranes in a 0.2 M NaOH solution at 42°C with constant agitation for a total 20 minutes, followed by a 15 minute rinse in 2 × SSC, or the hot SDS procedure. In the latter, membranes were immersed in boiling solution of 0.1% SDS and allowed to cool to room temperature before being rinsed thoroughly in 2 × SSC. Probes were constructed against both the cutinase construct (fsp cut) and the hygromycin phosphotransferase (hph) gene. Cutinase probes were constructed by double digestion of the pABC vector with enzymes NotI and SfiI. Products were analysed on 1% agarose gel cast in 1 × TAE and visualised by ethidium bromide staining. In correspondence with a 1 kb plus DNA ladder, a ~1.8 kb fragment was excised from agarose gel and purified using a QIAEX II agarose gel extraction kit according to the manufacturers protocol (QIAGEN). A second cutinase probe was constructed by PCR amplification of an ~1.2 kb fragment using the primers T3 and CutmRNA–C (Table 6.2). Hygromycin probes were also constructed by PCR amplification. Two fragments, 410 bp and 697 bp in length were amplified from the pABC vector as described in section 6.8.5 using the HPHL2/R2 and HPHL/R primer pairs respectively. These fragments were also visualised by agarose gel electrophoresis, and extracted and purified from the gel as described above. The pBARGEM7–2 probe was constructed by HindIII digestion of vector DNA. Following digestion, restriction enzymes were removed using the micropure–EZ centrifugal system (Millipore Corporation, Bedford, Massachusetts, USA). Probes were labelled with radioactive [α32P] dCTP (3000 Ci/mmole; Amersham Pharmacia Biotech) using the Random Primers DNA labelling method developed by Feinberg and Vogelstein (1983; 1984). The High Prime solution developed by Roche Diagnostics (Mannheim, Germany) was utilised, according to the manufacturers protocols, to reduce pipetting steps and increase the convenience and reproducibility of the procedure. Unincorporated labelled nucleotides were removed using MicroSpin G–25 columns (Amersham Pharmacia Biotech). An additional probe constructed by PCR amplification of a 417 bp fragment from R. alismatis using the degenerate oligonucleotide primer pair SNF–L/R designed by Tonukari et al. (2000) based on the conserved regions of the maize pathogen C. carbonum SNF1 gene was also utilised during this study. ‘Touchdown’ (TD)–PCR (Don et al. 1991) was performed with 2 pmol of each primers, 50 ng R. alismatis genomic DNA as template, 0.2 mM each of dATP, dCTP, dGTP and dTTP, 2.0 mM of MgCl2 and 2 units of Taq DNA polymerase, in a volume of 25µL, under the following conditions: initial denaturation at 94°C for 3 minutes, followed by 30 cycles of denaturation at 94°C for 30 seconds, annealing for 30 seconds, and extension at 72°C for 2 minutes.
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The annealing temperature ranged from 60°C to 45°C with a decrease of 0.5°C every cycle. This was followed by 10 cycles of denaturation at 94°C for 30 seconds, annealing at 45°C for 30 seconds, and extension at 72°C for 2 minutes. The resulting fragment of approximately 400 bp was visualised, extracted and purified from agarose gel as described above and cloned into a pCR2.1– TOPO TA cloning vector (Invitrogen Corporation, Faraday, Carlsbad, California, USA), which was then used to transform One shotTM chemically competent E. coli cells. Transformed cells are resistant to ampicillin and appear on LB plates (0.5% yeast extract, 1.0% tryptone, 1.0% NaCl, 1.5% agar, pH 7.0) containing 50 µg/mL of the agent after 16–24 hours incubation at 37°C. Positive clones which possess the SNF1 insert appear white on LB plates after the incorporation of 200 mg/mL IPTG (isopropylthio–β–D–galactoside) and 20 mg/mL X–Gal (5–bromo–4–chloro–3– indolyl–β–D–galactopyranoside), courtesy of a disruption in the αlacZ gene, which inhibits their ability to utilise galactose in the medium. Transformed colonies which do not contain the SNF1 insert possess a functional gene that enables them to utilise galactose and results in the production of by–products which assume a blue appearance. Several white colonies were chosen at random, transferred to LB broth and incubated at 37°C overnight (16–24 hours). Plasmid DNA was isolated using the QIAGEN Plasmid minikit (QIAGEN), restricted with EcoRI (New England Biolabs) and visualised by gel electrophoresis. Several intensely stained fragments approximately 400 bp in length were excised from the gel, purified as described above and sequenced using the M13RP/UP primer pair. Comparison of one such sequence with data published by Tonukari et al. (2000) revealed an 86–92% homology with the SNF1 gene isolated from the maize pathogen C. carbonum (GenBank accession number AAD43341). Consequently, this fragment was re–amplified from genomic DNA, excised from the gel, purified and labelled with α32P as described above. Although this fragment was originally isolated as part of another experiment, not discussed here in detail, the value of this fragment as a probe for the present study proved considerable.
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6.9 Results and Discussion Although several methodologies have been used to transform filamentous fungi, including electroporation, biolistic transformation and ATMT, fusionogenic methods whereby protoplasts are incubated with DNA and calcium chloride in an osmotic stabilised medium and induced to take up DNA through the addition of PEG, remain the most popular and consequently were the methods employed during this study. To date, protoplasts have been obtained from all the major taxonomic groups of fungi (Peberdy 1989). For R. alismatis, the physiological state of the starting cells and the duration of enzymatic exposure had the greatest influence on protoplast yield. Although no attempt was made to further optimise the conditions outlined previously by Taylor (unpublished data), freshly harvested and germinated spores as opposed to un–germinated spores or spores harvested and stored cryogenically for periods prior to experimentation, produced the greatest quantities of protoplasts, often in excess of 107/mL, after 16–24 hours of enzymatic lysis. The age of starting cells is deemed a critical factor in determining the likely yield of protoplasts for many fungi. In some studies younger cultures (10– to 20–hour cultures) produced the greatest yields (Lynch et al. 1985; Harling et al. 1988). However, 24– to 48–hour cultures (Tanaka et al. 1981; Hashiba & Yamada 1982), and cultures up to 10 days old have also been demonstrated to produce appreciable quantities of protoplasts (Bartnicki–Garcia & Lippmann 1966; Hocart et al. 1987). In this study older cells could be germinated albeit over longer periods, but enzymatic exposure for periods equivalent to younger cells failed to produce significant quantities of protoplasts from these cells. Increasing the duration of enzymatic exposure failed to improve protoplast yields, and instead was frequently accompanied by the appearance of large quantities of cell debris that may have resulted from the lytic system attacking newly formed protoplasts (Kitamoto et al. 1988). As Bos (1985) suggested, un–germinated spores were relatively resistant to mycolytic enzymes and failed to release appreciable quantities of protoplasts, which may reflect changes in the cell wall structure as the cell population ages, since the growing hyphal tip is believed to be the part of the cell most vulnerable to enzymatic lysis (Peberdy 1991). In order for protoplasts to be useful during transformation experiments, successful regeneration is critical. In the present study regeneration frequencies varied between 0.5 and 15%, which is within the range reported for majority of filamentous fungi (Stasz et al. 1988). Following treatment with DNA in the presence of PEG, regeneration frequencies were between 2 and 3%.
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Although protoplasts not treated with DNA generally failed to regenerate on selective media, large numbers of colonies were observed on control plates containing the selective agent on two occasions, both in experiments #1 and #5. Regeneration and growth of un–transformed protoplasts on control plates containing hygromycin suggested either spontaneous mutation or experimental error, casting into doubt the legitimacy of putative transformants. While high levels of spontaneous mutation to several other selective agents including the antibiotic phleomycin (Gaillardin & Ribet 1987; Nicaud et al. 1989), and the PPT–containing herbicide bialaphos (Ignite) (Pall 1993; Leung et al. 1995) have been reported, spontaneous mutation to HmB resistance is considered rare (Cordero– Otero & Gaillardin 1996). Instead, many of these colonies were thought to have arisen through uneven exposure to the selective agent. Because the temperature and volume of agar were minimal, solidification of the selective component occurred quickly following contact with the basal layer. As a result this layer was often uneven and may have permitted untransformed cells to escape the constraints applied by the antibiotic. In subsequent experiments the volume and temperature of liquefied agar was increased slightly, and these problems were eliminated.
On one occasion large numbers of regenerated protoplasts were also observed on H2O control plates. Because the osmotic stability of water results in the destruction of protoplasts, high numbers of colonies on these plates are suggestive of low protoplasting efficiencies indicating insufficient exposure to, or activity of, enzyme suspensions used during protoplast preparation. Hence, colonies on such plates may reflect the growth of spores which retained their cell coats in the absence of sufficient enzyme activity. Because optimal rates of enzyme exposure had been established previously, the preparation of fresh batches of enzyme had little effect. However, the use of freshly sporulating cultures for protoplast preparation alleviated such problems. Prior to transformation of fungal protoplasts, a suitable selection strategy for transformants must be developed. For transformation of R. alismatis the bacterial hygromycin B (HmB) phosphotransferase (hph) gene and the phosphinothricin (PPT) resistance–encoding (bar) gene, both under the control of A. nidulans regulatory sequences, were employed as selectable markers on two different plasmids, pABC and pBARGEM7–2, respectively (Bailey & Chen unpublished data; Pall & Brunelli 1993). Antibiotic resistance studies revealed that the wild–type R. alismatis isolate RH097 is sensitive to these compounds at concentrations approaching 25 µg/mL and 15 µg/mL respectively. These concentrations are similar to those employed during transformations respectively, of the barley pathogen R. secalis (Rohe et al. 1996) and Colletotrichum gloeosporioides f. sp. aeschynomene, the formulation registered for the control of northern jointvetch (Brooker et al. 1996).
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Putative transformants became visible by microscopic examination 4–5 days after plating of the transformation suspension on selective media containing either 50 µg/mL of HmB or 20 µg/mL of PPT. Unlike reports by Leung et al. (1995) and unpublished data from Pall (cited by Leung et al. 1995) however, there did not seem to be an observable difference in the rates of emergence of transformants on the two different selective media. During transformations of M. grisea and N. crassa both to HmB and PPT resistance, these researchers observed more rapid emergence of transformants on media containing PPT as opposed to HmB, and suggested that expression of the bar gene occurs considerably more rapidly than that of the hph gene. In the present study however, potential transformants became visible on both media after similar durations. Transformation frequencies fluctuated during ten individual experiments, but ranged from 6 to 187 transformants per 5 µg of DNA during four experiments conducted to fruition (marked #2, #3, #4 and #6 in Table 6.3). These rates were equivalent to up to 37 colonies/µg of DNA, or approximately 1 colony per 11 000 protoplasts and were once again within the range reported for the barley scald pathogen R. secalis (Rohe et al. 1996). Following plating, two types of transformants were observed on selective media. These included a small number of large rapidly growing colonies and a greater number of smaller slower growing entities. Additionally, a significant number of very small, rather stationary colonies were observed in proximity to the most aggressive transformants. Whilst these colonies are likely to represent un– transformed cells that arise in response to degradation of the active ingredient by their transformed counterparts, the smaller more isolated colonies are likely to represent ‘abortive transformants’. Reported by numerous researchers in a large range of fungal pathogens including Septoria (Cooley et al. 1988) and Fusarium (Kistler & Benny 1988), abortive transformants show only transient resistance to the selective agent and are thought to represent instances in which DNA enters the cell but fails to become integrated into the genome (Yap & Schiestl 1995).
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Table 6.3: Details of Individual transformation experiments. # putative Controls Transformation # transformants Experiment # Vector details protoplasts + selective (1) (2) (3) (4) (5) abc –2 –2 –4 agent trans. mix H2O 10 STC 10 STC 10 – control 1 PABC uncut 1.6 × 106 1402(0)a 802 150 TMTC 265 58a 2 PABC uncut 2.1 × 106 187(24)a 237 5 102 0 0a 3 PABC uncut 1.5 × 106 7(3)b 156 0 149 0 0b 4 PABC uncut 7.0 × 106 6(3)b 1002 1 TMTC 64 0b 5d pABC/pBARGEM7–2 8.3 × 106 0c TMTC 0 TMTC 61 52c 6d pABC/pBARGEM7–2 1.3 × 106 113(12)c 185 0 188 6 0c 7e PABC uncut 1.9 × 106 0b 934 0 TMTC 27 0b 8e PABC uncut 2.0 × 106 0b 121 0 160 1 0b 9f PABC/SfiI 4.9 × 105g 29(0)a 13 2 18 1 0a 10f PABC/NotI 4.9 × 105g 66(0)a 13 2 18 1 0a ahygromycin concentration = 50 µg/mL. bhygromycin concentration = 75 µg/mL. ctransformants plated on agar containing 20 µg/mL Ignite. () putative transformants analysed by PCR and/or Southern analysis. TMTC = too many to count. dco–transformation with the pBARGEM7–2 vector conferring basta/Ignite (phosphinothricin) resistance. eovernight (16–24 hours) protoplast regeneration in LM prior to plating. fvector linearised by digestion with SfiI or NotI prior to transformation. g4.9 × 105/2 = 2.45 × 105 protoplasts per enzyme digestion. Controls: (1) trans. mix = transformation mixture (effects of transformation), no antibiotic; –2 –2 (2) H2O 10 = protoplasts in H2O 10 (protoplasting efficiency), no antibiotic; (3) STC 10–2 = protoplasts in STC 10–2 (survival of protoplasts), no antibiotic; (4) STC 10–4 = protoplasts in STC 10–4 (survival of protoplasts), no antibiotic; (5) – control = protoplasts in STC 10–2 + selection agentabc.
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Of the colonies that grew rapidly on selective media, a total of 42 were transferred to fresh plates. Of these, 30 colonies arose from three experiments employing HmB as a selective agent. Antibiotic resistance in these individual transformants appeared stable under both selective and non–selective conditions for at least 4–5 generations, and a total of 24 colonies from experiment #2 were subjected to PCR and southern hybridisation analyses. Using oligonucleotide primer pairs HPHL2/R2 and T3+1/CutmRNA–C that were synthesised according to sequences derived from the hph and fsp cut genes respectively, amplification products of the expected sizes of 410 bp and 701 bp, were obtained with DNA from all transformants, a subset of which are shown in Figure 6.3.
1 2 3 4 5 6 7 8 9 10 11 12
12000
5000
2000
1650
1000 850
650
500
400
300
200
100
Figure 6.3: PCR analysis of putative pABC transformants. Lanes: 1; 1 Kb DNA ladder, 2–6; putative transformants T1, T6, T14, T20 and T24, 7; wild type R. alismatis DNA control, TC2, 8– 12; Control samples; 8; T3+1 primer only, 9; CutmRNA–C primer only, 10; HPHL2 primer only, 11; HPHR2 primer only, 12; Positive control, pABC vector DNA. Arrows indicate positions of bands.
Unfortunately, amplification fragments of equivalent sizes were also obtained in the un– transformed wild type control colonies. Because the analysis included primers against both hygromycin and cutinase however, the probability of obtaining both fragments of the correct proportions purely by chance would be extremely low. Therefore it appeared that the wild type control had been contaminated with transformant or vector DNA during sample preparation.
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Regrettably, attempts to confirm this hypothesis, which involved ultraviolet treatment of equipment, re–extraction of DNA from additional control samples not previously used in the analysis and single primer reactions to eliminate the possibility of fragment amplification through inversion of primers on opposing strands proved unsuccessful, and neither the source of the suspected contamination or the appearance of amplification fragments in control samples could be eliminated. Because primers prepared against the fsp cut gene may have sufficient homology to amplify native cutinase genes from R. alismatis, the appearance of fragments proportional to cutinase in both transformed and control samples was not particularly troubling. However, the persistence of amplification products in response to the hygromycin primers was cause for concern. Nevertheless, southern analysis of all 24 putative transformants was conducted. DNA was digested with SacI, which has four cleavage sites within the pABC vector at nucleotide positions 170, 234, 3225 and 3388. One of these sites (234) was introduced into the vector during construction and although present within the final BamHI–SfiI fragment cloned into the pAN26 vector, it is not part of the cutinase insert originally released from pU5–11 through AgeI–SalI digestion. This site occurs 23 bp upstream of the SfiI restriction site (211). The SacI restriction site at nucleotide position 170 occurs 41 bp downstream of the SfiI site. The remaining two SacI restriction sites both occur within the promoter (gdp) region of the hygromycin resistance gene (hph). Because there are no restriction sites within the coding or flanking regions of the cutinase gene nor within the coding region of the hph gene, against which probes were constructed, and SacI is recognised as an efficient digester of fungal DNA, this enzyme was an ideal choice for restriction analysis of DNA prior to southern blotting and hybridisation. Numerous slot–, dot–blot and southern analyses utilising one or more of the four probes outlined previously were utilised during these analyses. A small amount of vector DNA was included in each analysis as a positive control and to indicate successful hybridisation between the probe and DNA. In all analyses however, probes failed to hybridise to lanes containing DNA from putative transformants, and ultimately further analysis of these isolates was abandoned. Examples of dot– blot and southern analysis are shown in Figures 6.4 and 6.5.
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Figure 6.4 (left) : Dot blot analysis of putative pABC transformants. Dots represent transformants T1–T24 loaded left to right in consecutive rows. Untransformed wild type control samples TC1 and TC2 appear in the last row. A small amount of pABC vector DNA, which acts as a positive control, was placed in the bottom right hand corner of the membrane. Figure 6.5 (right) : Southern analysis of putative pABC transformants. Lanes: 1; 1 Kb DNA ladder, 2–10; putative transformants T7, T8, T9, T12, T13, T14, T17, T22 and T24, 11–12; wild type R. alismatis DNA controls TC1 and TC2, 13; positive control, pABC vector DNA, removed prior to probing.
Unfortunately, during slot–, dot–blot and southern analyses vector DNA was applied to the membrane following blotting, which meant that successful hybridisation of the probe to the membrane was not indicative of successful transfer of DNA from the gel to membrane. To ensure DNA was being transferred to the membrane during blotting, the remaining 6 putative transformants selected from HmB plates, consisting of two subsets of 3 isolates derived from experiments #3 and #4, were first analysed via hybridisation with an additional probe constructed via PCR–amplification of a 417 bp fragment from R. alismatis genomic DNA. This fragment, which has greater than 86% homology to the SNF1 gene isolated and characterised from C. carbonum by Tonukari et al. (2000) and found to be responsible for the expression of multiple CWDE in this fungus, was isolated from R. alismatis using the degenerative primer pair SNF–L/R during a related experiment. The objectives of this experiment included isolation and characterisation of homologues of this gene from R. alismatis, construction of transforming vectors containing SNF1 homologues and transformation of the fungus. Because the SNF1 gene has been found to be involved in the regulation of expression of multiple CWDE in C. carbonum, a similar role may be conferred in R. alismatis. By increasing the gene copy number it was envisioned that the expression and ultimately the production of CWDE by R. alismatis could be increased, factors which may enhance the pathogenicity, virulence or host range of the fungus.
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Unfortunately we were unable to isolate the entire gene from R. alismatis, and ultimately few of the objectives of this experiment were achieved. Nevertheless, this original 417 bp fragment that is known to be present in the genome of R. alismatis provided an additional probe by which the efficiency of DNA transfer between the gel and the blotting membrane could be gauged. The results of this experiment which provide confirmation of the transfer of DNA from the gel to the blotting membrane are shown in Figure 6.6.
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Figure 6.6: Confirmation of DNA transfer via southern analysis of putative pABC transformants. Lanes: 1; 1 Kb DNA ladder, removed, 2–7; putative transformants A1–A6, 8–9; empty, 10; wild type R. alismatis DNA control TC1, 11; empty, 12; positive control, Leptosphaeria maculans transformed with hph containing vector, 13; positive control, pABC vector DNA, removed prior to probing. Arrow indicates position of band.
The confirmation that DNA was being transferred successfully to the blotting membrane was encouraging, but all succeeding attempts to hybridise additional probes to regions within the hph and fsp cut genes of potential transformants A1–A6 once again proved unsuccessful. Because marker and control lanes frequently hybridised strongly to the probe, producing bands of high intensity, these lanes were removed prior to probing in all ensuing analyses. Concerns that these lanes sequestered large amounts of the probe, reducing the sensitivity of the analysis and raising the background noise arose, but proved unsubstantiated when probes once again failed to hybridise to the membrane to produce a visible signal. New membranes, increased DNA concentrations, reconstruction of probes and hybridisation of membranes often with multiple probes, also proved ineffectual.
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In response to the numerous southern hybridisation analyses which failed to confirm the presence of vector sequences in putative transformants, a subset of 12 putative transformants that arose from several co–transformation experiments involving a second plasmid, pBARGEM7–2, conferring resistance to the PPT–containing herbicide bialaphos (Ignite) were analysed. Because co–transformation efficiencies may approach frequencies up to 90% in some fungi (Vaillancourt & Hanau 1994), and the inclusion of a second plasmid in the transformation mix may promote uptake of the first (Specht et al. 1991), co–transformations utilising this second plasmid and a different selectable marker may reduce the appearance of false positives and increase the uptake of the former plasmid, pABC. Whilst the first of these experiments (#5) failed to produce any putative transformants and resulted in the appearance in a considerable number of false positives, most likely in response to low protoplasting efficiencies which was reflected by a large number of colonies on the negative control plate, the latter transformation, designated experiment #6, produced a total of 113 putative colonies following selection on media containing 20 µg/mL of Ignite. Of these, 30 of the most vigorous colonies were selected and transferred to plates containing hygromycin at a concentration of 50 µg/mL. Unfortunately, none of the colonies selected were able to grow in the presence of this compound. Despite these drawbacks, a dozen colonies were selected and analysed by southern hybridisation using probes directed towards the pBARGEM7–2 vector. Genomic DNA was digested with HindIII, which has one site within the polylinker region of the vector. Prior to probing, the marker lane was removed, then realigned prior to exposure to the photographic film. The intensity of the signal generated by the marker lane as described previously, is demonstrated in Figure 6.7. However, probes once again failed to hybridise to the putative transformant DNA. Because colonies once again grew in the presence of the selective agent, presumptive transformants appeared to be abortive, and coincidently, many colonies developed a grainy appearance and failed to sporulate following transfer. The literature suggests that the highest rates of co–transformation (80–90%) occur when the selected gene is used at very low levels, and the non–selected gene used at high levels (Austin & Tyler 1990). However, Roberts et al. (1989) reported that equimolar ratios of selected and unselected plasmids produced co–transformation efficiencies close to 50% in Aspergillus species. Hence, the optimisation of vector ratios may form an important step during co–transformation studies in filamentous fungi. The addition of plasmid DNA at a ratio of 1:3µg of pABC: pBARGEM7–2 respectively, in the present study may therefore explain the absence of colonies with the ability to grow in the presence of hygromycin.
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1 2 3 4 5 6 7 8 9 10 11 12 13 14 15
Figure 6.7: Southern analysis of putative pBARGEM7–2 transformants. Lanes: 1; 1 Kb DNA ladder, removed and probed separately then realigned prior to photographic exposure, 2–13; putative transformants B1, B2, B5, B6, B10–B14, B16, B20, B23, 14; empty, 15; wild type R. alismatis DNA control TC1.
Some groups suggest that the period of recovery prior to application of the selective agent may be critical to the recovery and hence expression of drug resistance genes following transformation (Avalos et al. 1989). For example, in the absence of a recovery period, Sivan et al. (1992) noted that neither transformed or un–transformed protoplasts of Trichoderma spp. were able to regenerate. Successful selection of putative transformants was possible only after a recovery period of 6–8 hours prior to exposure to the selective agent. During the present study, selective pressure was applied as an overlay, 16–24 hours after plating of the transformation suspension on non–selective underlays. Although putative transformants were obtained during most individual experiments, numbers obtained during experiments #3, #4, #5, #6, #9 and #10 were considerably lower than those obtained during the experiments #1 and #2, and it was perceived that by extending the period of recovery, gene expression and ultimately transformation and survival of putative transformants may be improved (Hargreaves & Turner 1992). Accordingly, protoplasts in experiments #7 and #8 were allowed to recover overnight in LM prior to plating. Unfortunately, this modification failed to produce any putative transformants, a situation reflected by Farman & Oliver (1988) who reported that extended periods of recovery in excess of 18 hours resulted in the appearance solely of ‘abortive’ transformants during studies on blackleg fungus L. maculans.
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Because only limited numbers of putative transformants were obtained during the majority of experiments in this study, for which the recovery period was consistently greater than 16 hours, it is possible that recovery durations employed in this study were responsible for the limited numbers of putative transformants generated throughout the project. The complete absence however, of colonies in experiments #7 and #8 is likely to be a result of prolonged exposure to PEG present in the transformation suspension, the toxicity of which is known to contribute to cellular mortality (Hargreaves & Turner 1992). Additionally, the concentration of the selective agent employed during plating may influence the rate of expression of genes introduced through transformation. For example, Schafer et al. (1988) demonstrated that by increasing the selective pressure on recovering colonies, the level of expression of the pisatin demethylase gene transformed into Nectria haematococca could be increased. Hence, during experiments #3 and #4 the concentration of HmB employed during selection was increased to 75 µg/mL. In comparison with some of the other experiments, this modification produced lesser numbers of putative transformants, with only 13 being recovered. Although six were analysed by southern analysis, as described previously (designated A1–A6), it is difficult to speculate as to whether the increase in selective pressure or the timing of selective application described previously affected recovery rates in these experiments. Whilst this technique almost certainly reduced the number of false positives, which seemed to plague many of the experiments thus far, the failure of ensuing southern analyses to produce audible signals meant that banding intensities could not be consulted to provide an indication of the effectiveness of Schafer’s earlier demonstrations. To date, the protocols utilised in this study, including the modifications discussed above, have failed to provide concrete evidence of successful transformation in R. alismatis. Nevertheless, PCR analysis combined with the continued prosperity of transformants on the selective medium suggested that the expression of antibiotic resistance genes introduced during the transformation process was being accomplished in some capacity, despite the inability to detect such sequences during southern analysis. These properties suggest the possibility that putative transformants harbour autonomously replicating plasmids. While extrachromosomal maintenance of transforming plasmids is unusual in filamentous fungi (Fincham 1989), a number of species have been identified in which this phenomenon occurs. To date, only two species, A. nidulans (Gems et al. 1991; Gems & Clutterbuck 1993; Aleksenko 1994) and the filamentous basidiomycete Phanerochaete chrysosporium (Randall et al. 1989; 1991; Randall & Reddy 1992) have been studied in detail.
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However, research in the latter indicates that such plasmids are maintained in copy numbers too low to be detected by southern hybridisation, unless sequences are amplified by PCR analysis (Randall et al. 1991). While this may not be the case for all autonomously replicating sequences, this process may be indicative of the results experienced in the present study. Additionally, because plasmids undergo extrachromosomal maintenance, they may be lost during DNA extraction, since plasmid DNA is not readily isolated in the total DNA fraction. Hence, while a small amount of plasmid DNA amplified by PCR analysis may produce audible bands, quantities sufficient to produce visible signals during southern analyses may not persist during the DNA extraction process, even in the presence of high plasmid copy numbers. More efficient recovery of plasmid DNA through the application of extraction protocols such as those described by Garber and Yoder (1983) may help to alleviate these problems. Research suggests that plasmids may undergo extrachromosomal maintenance due to recombination with endogenous plasmids (Randall & Reddy 1992), the presence of which have been documented in number of plant pathogens including C. purpurea (Tudzynski et al. 1983; Gessner–Ulrich & Tudzynski 1994), G. graminis (Honeyman & Currier 1986), R. solani (Hashiba et al. 1984; Jabaji et al. 1994), C. heterostrophus (Garber & Yoder 1984), C. musae (Freeman et al. 1997), F. solani (Samac & Leong 1988), and Tilletia spp. (Kim et al. 1990). Although it is not yet known whether species of Rhynchosporium harbour such plasmids, their presence in a wide range fungi suggest that although unlikely, this is a distinct possibility. Conceivably, recombination with such plasmids, if they do exist in this species, may render the status of autonomous replication to transforming DNA. To date, the precise function of many fungal plasmids remains unknown, although roles in growth senescence (Bertrand et al. 1986), toxin production (Meinhardt et al. 1990) and host specificity (Kistler & Leong 1986) have been demonstrated. Non–integrative transformation in fungi characterised by slowly growing transformants, is responsible for episodes of transient transformation reported in many fungal species (Judelson & Michelmore 1991; Peng et al. 1992; Durand et al. 1997) and indicates that integration, as opposed to the uptake of DNA, may be a limiting step during transformation. Furthermore, because plasmid integration in fungi occurs frequently by illegitimate or ectopic recombination, often resulting in the generation of tandem repeats, target sites for integration become increasingly important in these species (Itoh & Scott 1997). To date, it is almost universally accepted that linearisation of transforming vectors improves transformation efficiencies in filamentous fungi (Kim & Marzluf 1988; Unkles et al. 1989; Van de Rhee et al. 1996). Whilst some groups suggest this practice increases DNA uptake into the nucleus, with the entry of linear and circular DNA forms into the cell and subsequently the nucleus occurring at different rates (Royer et al. 1991), others suggest that vector linearisation increases the percentage of integration events (Skatrud et al. 1987). 125 Chapter 6. Transformation
Hence, by increasing the availability of integration sites, transformation efficiencies may be enhanced. Additionally, gene copy numbers and expression levels of some genes may be increased, improving the detection of transforming sequences through southern analysis. The addition of restriction enzymes SfiI and NotI to linearise vector DNA during experiments #9 and #10 respectively, produced a total of 95 putative transformants in this study. Although these recovery rates were higher than those obtained during some of the earlier experiments, colonies failed to grow after transfer to fresh selective medium, and hence no further analysis was conducted. Because the behaviour of putative transformants in these experiments closely resembled the nature of abortive transformants described previously, vector linearisation (despite increasing the regeneration rate) failed to increase the frequency of integration events. Regrettably, periods of transient expression once again were suggestive of extrachromosomal maintenance of transforming DNA, but the rapid loss of drug resistance in these isolates suggests that transforming DNA in this case failed to recombine with endogenous elements possibly present in the cytoplasm. In contrast, it is equally likely that the results obtained during PCR analysis resulted from contamination both of putative transformants and controls during sample preparation, most likely with plasmid DNA. Under this scenario however, untransformed cells must escape the constraints of the antibiotic agent applied following the transformation process. One mechanism by which fungi may facilitate this process is through the activity of fungal membrane transporters, a large family of proteins which include members with active roles in the secretion of pathogenicity factors, and in protection against plant defence compounds during pathogenesis (Urban et al. 1999; Fleiner et al. 2002). In fungi, two classes of proteins play a major role in these processes, the ATP–binding cassette (ABC) and the major facilitator superfamily (MFS) transporters (Wolfger et al. 2001; Higgins et al. 2001). Although the latter group of transporters are largely associated with the efflux of virulence factors that include both the specific and non–specific toxins produced by Cochliobolus carbonum (Pitkin et al. 1996), Fusarium sporotrichioides (Alexander et al. 1999) and Cercospora kikuchii (Callahan et al. 1999), the former class are largely responsible for protecting fungi against fungicides and toxic plant metabolites (De Waard 1997; Zwiers & De Waard 2000). Members of this class include almost a dozen families of eukaryotic efflux pumps (Saier & Paulsen 2001), responsible for many examples of multidrug resistance (MDR) in yeast (Balzi et al. 1987; 1994; Alarco et al. 1997) and filamentous fungi (Hayashi et al. 2001; Taylor 2002). By binding and hydrolysing nucleotide triphosphates such as ATP, ABC transporters generate the necessary energy required to transport solutes across cell membranes, often against the electrochemical gradient.
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In doing so, concentrations of these compounds are reduced in the cell, protecting the organism (Del Sorbo et al. 2000). Although antibiotics are among the vast array of substrates exported by ABC transporters, so far documented cases of antibiotic resistance through the expression of ABC transporter genes largely involve organisms of medical significance including Candida maltosa (Sasnauskas et al. 1992) and C. albicans (Prasad et al. 1995). Nevertheless, several cases have been documented outside this arena, including several examples of antibiotic resistance to common antibiotics utilised during many transformation procedures. For example, expression of the MDR gene, YOR1 confers oligomycin resistance in S. cerevisiae (Katzmann et al. 1995), whilst the imaA gene of A. nidulans confers resistance to neomycin following induction through the accumulation of the azole fungicides (Del Sorbo et al. 1997). Significantly, evidence suggests that resistance to some toxic metabolites such as azoles may have pleiotropic effects, whereby mutations for resistance to some compounds confer resistance to other unrelated toxicants (Bauer et al. 1999). Hence, the administration of toxic compounds can induce high level expression of ABC transporter systems resulting in unexpected resistance to many structurally dissimilar compounds including antibiotics. Whilst these proteins have not yet been identified in Rhynchosporium, their documentation in other fungal pathogens including Nectria haematococca (Akallal et al. 1998), Penicillium digitatum (Nakaune et al. 1998), Mycosphaerella graminicola (Stergiopoulos et al. 2000; 2001), Botrytis cinerea (Schoonbeek et al. 2001; Vermeulen et al. 2001) and Leptosphaeria maculans (Taylor & Condie 2000), suggest they are also likely to be present in this organism. Hence, colonies may take on the appearance of transformants, continuing to prosper on selective media despite being untransformed and may grow as a result of efflux pumps which reduce internal concentrations of antibiotics sufficiently to allow the survival of the organism. The hypotheses outlined above provide some insight into processes which may have been occurring during attempts to transform R. alismatis. Whilst the protocol previously outlined by Taylor (unpublished data), but adapted from Yelton et al. (1984), proved suitable for the production of large quantities of protoplasts from R. alismatis, thus far attempts to transform the fungus based on the methods outlined above have proven unsuccessful. Nevertheless, a gene transfer system has been developed for the barley scald pathogen R. secalis (Rohe et al. 1996), and genetic complementation of virulent strains with genes encoding the race–specific elicitor, NIP1, which determines avirulence on some host plants (Hahn et al. 1993), has been demonstrated (Rohe et al. 1995).
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Hence, whilst future transformation–based manipulation of R. secalis is expected to identify other genes involved in pathogenicity and virulence of this organism, continued experimentation with R. alismatis is likely to yield protocols and techniques that will facilitate similar studies of pathogenicity related genes in this organism. The research for this chapter was conducted during a nine month period at the Plant Biotechnology Institute, National Research Council of Canada, Saskatoon, Saskatchewan, Canada, under the supervision of Dr. Janet L. Taylor.
128 Chapter 7. General Discussion
7 General Discussion
The principal objective of this project was to expand the host range of Rhynchosporium alismatis, a fungal pathogen currently being developed as a mycoherbicide for the control of Alismataceae weeds in Australian rice fields. In attempting this goal several distinct studies were performed prior to attempts to develop reliable and efficient transformation techniques for the fungus. These studies included a comprehensive phylogenetic analysis, population structure analysis and a detailed study of the infection process of the fungus on both host and non–host Alismataceae species. In 1994 Cother et al. (1994) hypothesised that R. alismatis may have expanded its host range from A. plantago–aquatica to A. lanceolatum and finally to D. minus, speculating that given sufficient time the fungus may also evolve pathogenicity to Sagittaria species. However, reports by Heath (1980; 1981c) indicate that ‘pathogens rarely if ever undergo spontaneous or distinct changes in host range’. In fact the author reported that a move by a pathogen to a new host species, had to her knowledge, not been convincingly demonstrated over recorded history’ (Heath 1980). To date, no records of the fungus have been reported on species of Sagittaria in Australia, despite reports that species in this genus are hosts of the fungus elsewhere in the world (Cother & Gilbert 1994a). Because mutation alone cannot spawn the necessary elements required to establish ‘specific accommodation’ with a new host, particularly in the presence of multiple defences (Heath 1981c; 2000a), genetic manipulation was considered a logical alternative. Some of the major concerns regarding the release of biocontrol agents are the potential risks that unexpected dispersal poses to non–target species and the potential for unexpected genetic exchange with closely related microorganisms present within the environment. For the endemic pathogen R. alismatis, ample opportunity already exists for such exchange and the development of undesirable hybrids is certainly no greater of a threat than that which occurs naturally (Weidemann 1992). However, for genetically modified organisms, dispersal and gene exchange could become a risk factor if the introduced genes are exchanged among related crop pathogens with the propensity for extensive long distance dispersal.
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Phylogeny studies and population structure analyses provide detailed assessment of the risks associated with potential agents and fulfil some of the requirements of regulatory agencies prior to the release of candidate in the field. These studies clarify the nomenclature, identify relationships to closely related pathovars and provide information about the evolutionary history of the pathogen including the contribution of different modes of reproduction, likely pathogenicity attributes and potential for dispersal. In this study data showed that the hyphomycete fungus currently classified under the combination R. alismatis is very closely related to the teleomorph genus Plectosphaerella and highlighted similarities between the use of this organism and a recent study conducted in Korea, in which Alismataceae weeds were the target of biological control using the anamorph P. tabacinum (Chung et al. 1998). Clearly R. alismatis is not related to the scald pathogen R. secalis, which is renowned for its pathogenic variability, nor other species in the genus Rhynchosporium and should be renamed. Instead R. alismatis was found to possess minimal evolutionary capacity and limited dispersal potential by virtue of a clonal reproductive strategy and the absence of airborne ascospores generated via sexual reproduction. Pathogenicity studies supported assumptions that a clonal population structure may be reflected by limited spectrums of virulence and earlier host range testing by Cother (1999) confirmed this by demonstration of a limited host range. With minimal evidence of prior recombination, R. alismatis appears to be genetically isolated on its respective hosts and major shifts in host specificity through the transfer of genetic material are unlikely. Hence, genetic modification of R. alismatis is considered to pose a negligible threat to crops grown adjacent to, or in rotation with, rice in southern New South Wales. Whilst the correct classification of this fungus remains to be determined, some of the most significant discussions which speculated on the identity of this fungus, centred around the overlapping host ranges of these pathogens, most specifically with species from the Cucurbitaceae and Solanaceae. However, these discussions may have been premature considering host range testing under controlled environmental conditions may predispose plants to infection and thus broaden the host range artificially (Watson 1985; Dale 1999). Under this premise, members of the Cucurbitaceae and Solanaceae, to which R. alismatis was only lowly pathogenic, may be susceptible to infection in the glasshouse yet resistant to infection in the field where lower inoculum rates are applied. Hence, neither species may be a legitimate host of this fungus, dismissing much of the potential overlap in host range between the two phytopathogens.
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Furthermore, host range tests conducted by Cother (1999) failed to result in the infection of hydrilla, a host of Plectosporium now the target of biocontrol studies in southern Florida (Smither– Kopperl et al. 1999) and Potamogeton, the species from which isolate RH126, shown to be genetically identical to P. cucumerina, was obtained. Hence, only one species to date, S. pygmaea, is a definitive host of both R. alismatis and P. tabacinum (Cother & Gilbert 1994a; Chung et al. 1998). Nevertheless, individual populations or isolates of Plectosporium appear to have very different host specificities, especially where populations are separated by large distances such as those imposed continentally. For example, isolates of Plectosporium used by Zhang, Sulz & Bailey (2002), during biocontrol studies on false cleavers in Canada, produced little evidence of infection or disease progression when used to inoculate cucurbits or tomato, despite the fact that this organism is a recognised pathogen of both these species (Saad & Black 1981; Bost & Mullins 1992; Everts 2002; Pascoe et al. 1984). But whilst P. tabacinum appears to have a significant capacity to evolve in pathogenicity towards a range of hosts, a situation most likely enhanced by the presence of a sexual cycle that has the capacity to contribute to the virulence of the fungus through favourable gene exchange, R. alismatis appears to have a very narrow host range and limited genetic diversity. Combined with the absence of sexual spores and a population structure that may be contrived through regular rounds of clonal reproduction, there appears to be little diversity from which genes may be acquired to contribute to an expansion in the pathogenic ability of this fungus, even in the presence of moderate gene flow. Hence, current data indicates that the population dynamics of these organisms are very different and therefore, despite being similar genetically, are likely to be members of different, although closely related genera. Whilst, confirmatory morphological studies are now underway in the laboratories of both Dr Walter Gams and Dr Uwe Braun, mycologists accredited as authorities on Plectosphaerella, Rhynchosporium and allied genera including Spermosporina, the absence of genetic studies for species in the latter genera may inhibit further attempts to identify this species. Because the classifications which resulted in the relocation of R. alismatis to Spermosporina in 1993 were based solely on morphological features, errors that resulted in the earlier placement of R. alismatis in the genus Rhynchosporium may be mimicked in its relocation to the new genus Spermosporina. For instance, the genus Spermosporina currently contains 9 species, only two of which cause leaf spots on members of the Alismataceae (Braun 1993; 1994; 1998). However, like the former genus Rhynchosporium, which comprises only pathogens of the Poaceae, Spermosporina contains two designated species which are considered pathogens of grasses and cereals.
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Hence, there may be some overlap between these genera that requires further study to ensure that the placement of R. alismatis into this genus does not result in further comparisons to organisms which may themselves be wrongly classified. Such a situation could be realised by a grouping which comprises other pathogens of cereals and grasses, such as S. graminella (Hohn) U. Braun (1993) and S. graminicola (Peck) U. Braun (1993), considering studies to date have already demonstrated that R. alismatis although morphologically similar, is unrelated to several other pathogens of the Poaceae with which it was formerly classified. Hence, further phylogenetic studies may be required to determine species boundaries between these genera before an accurate revision of the nomenclature of R. alismatis can be made. In addition to the 9 species classified under the genus Spermosporina, these additional studies should take into account a number of other extant and excluded species such as Plectosporium himantia, P. melaena (Fr.) Kirschst., P. silenes, Rhynchosporium dryopteridis Sawada, R. oryzae Hashioka & Yokagi and two additional forms of the barley scald fungus, R. secalis f. phalaridis Iwata & Kajiw and R. secalis f. agropyri Iwata & Kajiw, all of which are cited in the CAB International Bioscience fungal database (http://www.indexfungorum.org). To date, many fungal pathogens assessed for use as mycoherbicides have been shown to require several hours of free moisture in order to germinate, form infection structures and penetrate their hosts (TeBeest et al. 1978; Templeton & Heiny 1989; Boyette et al. 1996). Hence, this free moisture requirement, or dew period, is often a major obstacle to disease initiation in semi–arid environments (Boyette 1994). Accordingly, the vast majority of research on improvement of bioherbicides has focused on reducing this free moisture requirement, with researchers devoting considerable resources to the study of adjuvants and formulations believed to have the capacity to improve the pathogenicity of potential candidates. However, the requirement for free moisture is unlikely to be a limiting factor in aquatic environments such as rice paddies, where a relatively high humidity is maintained (Auld 1992), and Jahromi (2000) has demonstrated that dew period is not a limiting factor for disease development in the R. alismatis–D. minus pathosystem under rice field conditions. Instead, the host range of R. alismatis has been a limiting factor in its development as a broad spectrum alternative to chemical herbicides. Nevertheless many formulations have been described that improve the efficacy of mycoherbicides (Boyette et al. 1996), and the formulation approach has met with some success with regard to expanding the host ranges of potential candidates (Amsellem et al. 1991; Boyette et al. 1991; 1992; Boyette & Abbas 1994). Unfortunately, whilst R. alismatis was observed to attach, germinate, and form infections structures on both S. graminea and S. montevidensis, penetration was observed only on the leaves of S. graminea and often in the accompaniment of strong host reactions which appeared to inhibit further disease progression. 132 Chapter 7. General Discussion
Some researchers have suggested that variability in appressoria formation and penetration by fungi may be linked to the heterogeneity of plants used in the study (Green et al. 2001). In a genetically diverse weed population there may be resistant biotypes (Auld & Morin 1995), and variation within a weed population may strongly influence the infection process. In studies to date, populations of D. minus collected from a number of sites throughout the rice growing regions of southern New South Wales, have not displayed phenotypic (Graham et al. 1996), nor appreciable levels of genotypic variation (Jahromi 2000), although several populations of Alisma lanceolatum collected from similar locations displayed considerable variation (G. Ash, pers. comm., 3rd March 2003). Because there is general agreement that the success of biological control may be limited by high levels of genetic variation in the target weed (Nissen et al. 1995; TeBeest et al. 1992), these factors should be investigated, specifically with reference to populations of S. montevidensis for which variation of this nature may contribute to entry level constraints of the fungus. The failure of R. alismatis to penetrate the leaf surface of S. montevidensis suggests that avenues to expand the host range of the fungus should take into factors which affect the entry requirements of the fungus. Constructs containing the cutinase gene from Fusarium solani f. sp. pisi were employed during ensuing studies which focused on the objective of host range expansion by inserting additional copies of the gene into the fungal genome. To date, the literature has been discussed with regard both to the selection and activity of this gene construct and to the mechanisms which may have prevented its uptake and expression in R. alismatis. However, the close relationship between R. alismatis and the teleomorphic genus Plectosphaerella may indicate that there are sufficiently important differences in gene structure between the transforming species and the species from which expression elements are derived which limit the ability of transformed colonies to recovery on the selective medium. Although promoters from one member of higher fungi frequently function in others (Judelson et al. 1992), including members of different phyla (Smith et al. 1990; Barrett et al. 1990), there are some exceptions. For example, the hsp70 promoter from Ustilago maydis failed to function in the related basidiomycete Coprinus cinereus (Casselton & De La Fuente–Herce 1989), and the promoter sequences of some yeasts are not readily interchangeable with those from filamentous ascomycetes (Ballance 1986). The pABC vector used in this study contains the E. coli hph gene under the control of A. nidulans promoter and terminator sequences. These regulatory sequences are the same as those carried by the pAN7–1 vector which has been shown to function in a large range of plant pathogens including the form species R. secalis (Rohe et al. 1996). Hence, although unlikely it is possible that the promoter and terminator sequences from Aspergillus failed to function in R. alismatis. To date, transformation studies have not been reported for Plectosporium.
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Alternatively, transforming DNA may be inactivated or deleted during integration into the genome of some species (Casselton & De La Fuente–Herce 1989). In Neurospora crassa for example, tandemly integrated plasmid DNA is deleted and/or methylated during meiosis in a process termed RIP or repeat–induced point mutation (Selker et al. 1987). Synonymous with the former acronym Rearrangement Induced Premeiotically, and analogous to processes reported in several other fungi including Ascobolus immerses (Goyon & Faugeron 1989) and Gibberella pulicaris (Salch & Beremand 1993), the RIP process mutates DNA sequence duplications introduced during transformation, resulting in rearrangement of restriction sites and methylation of cytosine residues (Selker 1991). RIP acts specifically on duplicated sequences, producing transition mutations in which G:C pairs are replaced by A:T residues (Cambareri et al. 1989). Because plasmid integration in fungi occurs frequently by illegitimate or ectopic recombination, often resulting in the generation of tandem repeats, transformed sequences in fungi are particularly amenable to this process and the presence of tandem repeats frequently leads to both mitotic and meiotic instability of integrated plasmids during fungal transformation (Keller et al. 1991; Tooley et al. 1992). Fortunately, this processes seems to occur only in haploid nuclei of specialised cells formed during the sexual phase, and hence is unlikely to effect DNA transformed into organisms which reproduce exclusively through asexual recombination, and do not enter into a meiotic phase. Nevertheless, the close relationship between R. alismatis and the teleomorph genus Plectosphaerella provides some evidence for the consideration of this phenomenon. Although these studies were not undertaken to instigate penetration in S. graminea and it is likely other methods will prove more effective in accentuating disease progression in this species, transformation may still have the capacity to improve the pathogenic ability of the fungus on S. graminea. For instance, there is a correlation between gene copy number and gene expression in some species (Durrens et al. 1986; Kelly & Hynes 1987; Turcq & Begueret 1987; Scorer et al. 1994). Increasing the gene copy number via transformation may heighten fungal gene expression, that may in turn promote more rapid penetration of the host. In basidiomycete fungus Coprinus cinereus for example, Mellon and Casselton (1988) reported enzyme levels close to four times that of the wild type in transformants containing multiple copies of the isocitrate lyase gene, acu–7, and increased gene expression through high gene copy transformation has been employed successfully by several groups to increase chitinase production in species of Trichoderma longibrachiatum, a mycoparasite currently being developed as a biocontrol agent for the cucumber pathogen Pythium ultimum (Sanchez–Torres et al. 1994; Migheli et al. 1998).
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Interestingly, Rohe et al. (1996) suggested that the interaction between R. secalis and its host (Hordeum vulgare L.) complies with the gene–for–gene hypothesis proposed by Flor (1971), in which a gene for virulence in the pathogen complements a host resistance gene, and it was initially presumed that this may also be the case with R. alismatis. Whilst phylogeny experiments now suggest that R. alismatis is not related to scald pathogen, leaf infection studies in the resistant host S. graminea suggested that the infection process occurs in two parts with initial infections arising through a biotrophic interaction, which may then switch to a necrotrophic phase following host recognition of the organism. This pattern of disease progression suggests that the coordinated expression of a large number of genes is involved, and in fact non–host resistance is reportedly not governed by the gene–for–gene relationship which controls many examples of cultivar resistance (Ellingboe 1976), such as that displayed commonly in R. secalis–barley pathosystems (Ayres & Owen 1971; Ali & Boyd 1974; Habgood 1977; Robinson et al. 1996; Cselenyi & Friedt 1998). Hence, future manipulative approaches to improve infection of this species should take into account the hemibiotrophic lifestyle of the pathogen. For example, biotrophic pathogens must avoid triggering the host’s defence responses through the elicitation of degradation products of enzymes or host cell wall constituents acted upon by enzymes during the initial stages of infection. However, necrotrophs are likely to benefit from the elicitation of host defences that might trigger plant cell death. Recently, the processive activity and concerted action of exo– and endopolygalacturonases has been shown to fulfil both functions with the expression of different forms during pathological and saprophytic phases of infection (Ten–Have et al. 2002). Future genetic manipulation of R. alismatis to improve disease symptoms on S. graminea may benefit from similar studies. In contrast, rather than focusing on the insertion of specific pathogenicity–related genes to improve infection in S. montevidensis, the likelihood of success may be improved by isolating and characterising genes involved during interactions with the host. Specifically, genes involved in the elicitation of host responses (pathogen avirulence genes) may provide appropriate targets for gene knock–out or disruption experiments by homologous transformation or REMI, described in detail in section 6.7, such that the host’s capacity to recognise the pathogen is reduced or eliminated. Experiments of this nature would be in direct contrast to those conducted by Rohe et al. (1995) in which the host response of barley cultivars was improved through transforming the pathogen.
135 Chapter 7. General Discussion
This thesis has addressed the risks, potentials, target sites and techniques for expanding the host range of R. alismatis for future control of Alismataceae weeds of rice in Australia. Whilst experiments have yet to yield transformed colonies with increased pathogenic ability, this study has provided avenues through which future research may address additional factors that affect the host range and virulence of this fungus.
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Chapter 8. Appendices
8 Appendices
8.1 Tables Table 8.1: Pathogenicity of R. alismatis isolates against host and non–host Alismataceae species. Isolate D. minus A. plantago–aquatica S. graminea S. montevidensis
Replicates 1 2 3 1 2 3 1 2 3 1 2 3 RH001 6 8 4 6.00 8 2 6 5.33 1 1 1 1.00 2 2 3 2.33 RH021 6 7 8 7.00 8 6 0 4.67 1 1 1 1.00 2 2 4 2.67 RH024 8 8 6 7.33 8 7 2 5.67 1 1 2 1.33 2 2 0 1.33 RH025 8 9 8 8.33 8 8 8 8.00 1 2 2 1.67 2 6 2 3.33 RH037 0 0 0 0.00 1 0 0 0.33 1 1 1 1.00 0 2 2 1.33 RH038 2 6 0 2.67 4 2 3 3.00 2 1 1 1.33 0 2 2 1.33 RH039 8 9 7 8.00 8 2 4 4.67 2 1 1 1.33 3 2 3 2.67 RH041 6 6 7 6.33 1 0 0 0.33 1 1 1 1.00 2 2 4 2.67 RH046 0 0 0 0.00 1 0 0 0.33 1 1 1 1.00 1 2 4 2.33 RH047 9 9 8 8.67 8 7 6 7.00 2 1 1 1.33 2 4 4 3.33 RH054 8 7 8 7.67 8 6 8 7.33 1 2 1 1.33 2 2 2 2.00 RH055 6 8 8 7.33 8 5 7 6.67 1 3 1 1.67 2 2 2 2.00 RH057 0 9 6 5.00 8 2 6 5.33 2 1 1 1.33 0 2 2 1.33 RH058 6 9 4 6.33 8 2 8 6.00 1 4 1 2.00 2 2 3 2.33 RH062 4 6 6 5.33 6 0 7 4.33 1 1 1 1.00 0 2 2 1.33 RH064 4 8 0 4.00 6 2 4 4.00 1 1 1 1.00 0 2 4 2.00 RH066 7 8 7 7.33 6 7 7 6.67 1 1 1 1.00 2 2 3 2.33 RH069 0 0 0 0.00 1 0 0 0.33 1 1 1 1.00 2 0 3 1.67
137
Chapter 8. Appendices
Table 8.1: (cont.) Replicates 1 2 3 1 2 3 1 2 3 1 2 3 RH074 8 8 7 7.67 6 6 6 6.00 1 1 6c 2.67 4 3 2 3.00 RH091 8 8 8 8.00 8 8 7 7.67 3 2 2 2.33 2 3 3 2.67 RH095 8 8 7 7.67 6 6 6 6.00 1 1 1 1.00 3 3 4 3.33 RH097 9 8 8 8.33 8 6 8 7.33 3 1 1 1.67 7 6 6 6.33 RH100 7 8 7 7.33 8 6 6 6.67 2 1 1 1.33 2 0 3 1.67 RH108 8 9 8 8.33 6 5 6 5.67 1 2 2 1.67 2 2 3 2.33 RH111 6 8 6 6.67 4 2 4 3.33 2 1 1 1.33 2 4 2 2.67 RH118 0 0 0 0.00 1 0 0 0.33 2 1 1 1.33 2 2 2 2.00 RH121 8 8 0 5.33 4 0 0 1.33 2 1 2 1.67 2 4 2 2.67 RH122 6 4 0 3.33 7 7 7 7.00 1 1 1 1.00 2 4 2 2.67 RH123 7 8 4 6.33 6 6 6 6.00 1 1 1 1.00 2 2 2 2.00 RH124 8 8 6 7.33 8 2 6 5.33 1 1 1 1.00 0 0 1 0.33 RH127 8 7 8 7.67 4 5 2 3.67 1 1 1 1.00 0 2 2 1.33 RH133 8 9 7 8.00 8 6 6 6.67 1 1 2 1.33 0 2 4 2.00 RH135 7 8 7 7.33 4 2 4 3.33 1 1 1 1.00 2 2 2 2.00 RH136 7 9 8 8.00 4 7 4 5.00 2 1 2 1.67 2 0 1 1.00 RH137 8 8 0 5.33 6 7 6 6.33 1 2 6a 3.00 2 2 4 2.67 RH138 0 0 0 0.00 1 0 0 0.33 1 1 2 1.33 0 0 2 0.67 RH139 8 9 8 8.33 8 2 8 6.00 2 2 1 1.67 10a,b 7 7 8.00 RH143 6 8 8 7.33 4 6 4 4.67 1 1 1 1.00 2 2 2 2.00 RH144 0 0 0 0.00 1 0 2 1.00 1 1 1 1.00 2 2 2 2.00 RH145 8 8 8 8.00 4 6 6 5.33 2 1 1 1.33 0 1 2 1.00 aLeaf disc cleared rather than diseased bControl disc has identical appearance to inoculated disc
138
Chapter 8. Appendices
8.2 Figures
Figure 8.1a: Percentage of conidia of R. alismatis that germinated on the leaves of A. plantago– aquatica (–●–), S. graminea (–▲–), and S. montevidensis (–×–). Error bars represent LSD of means at each time interval (P= 0.05).
× y = 37.0 ln(x) – 44.7 R2 = 0.9733
▲y = 37.2 ln(x) – 31.2 R2 = 0.9572
● y = 22.5 ln(x) + 15.5 R2 = 0.8731
Figure 8.1b: Percentage of conidia of R. alismatis that germinated on the leaves of A. plantago– aquatica (–●–), S. graminea (–▲–), and S. montevidensis (–×–). Error bars represent LSD of means at each time interval (P= 0.05).
139 Chapter 8. Appendices
Figure 8.2a: Percentage of germinated conidia of R. alismatis that formed appressoria on the leaves of A. plantago–aquatica (–●–), S. graminea (–▲–), and S. montevidensis (–×–). Error bars represent LSD of means at each time interval (P= 0.05).
× y = 24.2 ln(x) – 45.4 R2 = 0.9346
▲y = 38.9 ln(x) – 62.6 R2 = 0.9276
● y = 38.9 ln(x) – 50.4 R2 = 0.9432
Figure 8.2b: Percentage of germinated conidia of R. alismatis that formed appressoria on the leaves of A. plantago–aquatica (–●–), S. graminea (–▲–), and S. montevidensis (–×–). Error bars represent LSD of means at each time interval (P= 0.05).
140 Chapter 8. Appendices
8.3 Statistical Data (Chapter 5)
Time=1 (6 hours post inoculation)
***** Analysis of variance *****
Variate: germination
Source of variation d.f. s.s. m.s. v.r. F pr. rep stratum 2 15165.1 7582.6 49.32 rep.disc stratum 15 2306.1 153.7 0.22 rep.disc.*Units* stratum species 2 12273.6 6136.8 8.92 <.001 Residual 34 23393.1 688.0
Total 53 53137.9
* MESSAGE: the following units have large residuals. rep 1.00 disc 1.00 14.3 s.e. 6.5 rep 1.00 disc 5.00 -13.4 s.e. 6.5
***** Tables of means *****
Variate: germination
Grand mean 35.8
species 1.00 2.00 3.00 21.1 29.7 56.5
*** Least significant differences of means (5% level) ***
Table species rep. 18 d.f. 34 l.s.d. 17.77
Species 1= A. plantago-aquatica Species 2= S. graminea Species 3= S. montevidensis
141 Chapter 8. Appendices
***** Analysis of variance *****
Variate: appressorium formation
Source of variation d.f. s.s. m.s. v.r. F pr. rep stratum 2 923.1 461.5 14.47 rep.disc stratum 15 478.4 31.9 0.21 rep.disc.*Units* stratum species 2 4920.6 2460.3 16.44 <.001 Residual 34 5087.5 149.6
Total 53 11409.6
* MESSAGE: the following units have large residuals. rep 1.00 disc 1.00 -6.1 s.e. 3.0 rep 1.00 disc 3.00 *units* 3 22.7 s.e. 9.7
***** Tables of means *****
Variate: appressorium formation
Grand mean 8.4
species 1.00 2.00 3.00 0.4 3.0 21.8
*** Least significant differences of means (5% level) ***
Table species rep. 18 d.f. 34 l.s.d. 8.29
Species 1= A. plantago-aquatica Species 2= S. graminea Species 3= S. montevidensis
142 Chapter 8. Appendices
TIME=2 (12 hours post inoculation)
***** Analysis of variance *****
Variate: germination
Source of variation d.f. s.s. m.s. v.r. F pr. rep stratum 2 23716.7 11858.4 141.36 rep.disc stratum 15 1258.3 83.9 0.25 rep.disc.*Units* stratum species 2 2705.0 1352.5 3.99 0.028 Residual 34 11535.0 339.3
Total 53 39215.0
* MESSAGE: the following units have large residuals. rep 2.00 disc 5.00 9.8 s.e. 4.8 rep 2.00 disc 6.00 *units* 1 33.5 s.e. 14.6 rep 3.00 disc 2.00 *units* 1 -33.2 s.e. 14.6
***** Tables of means *****
Variate: germination
Grand mean 62.0
species 1.00 2.00 3.00 52.5 69.5 63.9
*** Least significant differences of means (5% level) ***
Table species rep. 18 d.f. 34 l.s.d. 12.48
Species 1= A. plantago-aquatica Species 2= S. graminea Species 3= S. montevidensis
143 Chapter 8. Appendices
***** Analysis of variance *****
Variate: appressorium formation
Source of variation d.f. s.s. m.s. v.r. F pr. rep stratum 2 3295.9 1648.0 6.79 rep.disc stratum 15 3643.1 242.9 0.65 rep.disc.*Units* stratum species 2 14223.8 7111.9 19.04 <.001 Residual 34 12700.5 373.5
Total 53 33863.4
* MESSAGE: the following units have large residuals. rep 1.00 disc 1.00 -19.9 s.e. 8.2 rep 2.00 disc 5.00 *units* 3 -36.4 s.e. 15.3 rep 2.00 disc 6.00 *units* 3 -34.7 s.e. 15.3
***** Tables of means *****
Variate: appressorium formation
Grand mean 31.0
species 1.00 2.00 3.00 9.1 35.9 48.0
*** Least significant differences of means (5% level) ***
Table species rep. 18 d.f. 34 l.s.d. 13.09
Species 1= A. plantago-aquatica Species 2= S. graminea Species 3= S. montevidensis
144 Chapter 8. Appendices
TIME=3 (18 hours post inoculation)
***** Analysis of variance *****
Variate: germination
Source of variation d.f. s.s. m.s. v.r. F pr. rep stratum 2 6328.1 3164.1 24.31 rep.disc stratum 15 1952.1 130.1 0.35 rep.disc.*Units* stratum species 2 10952.4 5476.2 14.54 <.001 Residual 34 12802.2 376.5
Total 53 32034.8
* MESSAGE: the following units have large residuals. rep 2.00 disc 1.00 -12.3 s.e. 6.0 rep 2.00 disc 6.00 12.7 s.e. 6.0 rep 2.00 disc 1.00 *units* 2 -35.8 s.e. 15.4 rep 2.00 disc 3.00 *units* 2 -35.4 s.e. 15.4
***** Tables of means *****
Variate: germination
Grand mean 75.1
species 1.00 2.00 3.00 55.7 79.8 89.6
*** Least significant differences of means (5% level) ***
Table species rep. 18 d.f. 34 l.s.d. 13.14
Species 1= A. plantago-aquatica Species 2= S. graminea Species 3= S. montevidensis
145 Chapter 8. Appendices
***** Analysis of variance *****
Variate: appressorium formation
Source of variation d.f. s.s. m.s. v.r. F pr. rep stratum 2 27.8 13.9 0.07 rep.disc stratum 15 2933.9 195.6 0.61 rep.disc.*Units* stratum species 2 12389.7 6194.8 19.22 <.001 Residual 34 10959.8 322.3
Total 53 26311.2
* MESSAGE: the following units have large residuals. rep 2.00 disc 6.00 21.7 s.e. 7.4 rep 2.00 disc 1.00 *units* 1 34.0 s.e. 14.2 rep 2.00 disc 3.00 *units* 1 31.3 s.e. 14.2
***** Tables of means *****
Variate: appressorium formation
Grand mean 44.6
species 1.00 2.00 3.00 23.3 57.0 53.5
*** Least significant differences of means (5% level) ***
Table species rep. 18 d.f. 34 l.s.d. 12.16
Species 1= A. plantago-aquatica Species 2= S. graminea Species 3= S. montevidensis
146 Chapter 8. Appendices
TIME=4 (24 hours post inoculation)
***** Analysis of variance *****
Variate: germination
Source of variation d.f. s.s. m.s. v.r. F pr. rep stratum 2 6425.1 3212.6 37.30 rep.disc stratum 15 1291.8 86.1 0.34 rep.disc.*Units* stratum species 2 3349.1 1674.6 6.69 0.004 Residual 34 8508.9 250.3
Total 53 19575.0
* MESSAGE: the following units have large residuals. rep 2.00 disc 2.00 -10.1 s.e. 4.9 rep 3.00 disc 3.00 10.8 s.e. 4.9 rep 2.00 disc 2.00 *units* 2 -32.6 s.e. 12.6 rep 3.00 disc 2.00 *units* 1 -28.2 s.e. 12.6
***** Tables of means *****
Variate: germination
Grand mean 84.0
species 1.00 2.00 3.00 72.9 89.9 89.2
*** Least significant differences of means (5% level) ***
Table species rep. 18 d.f. 34 l.s.d. 10.72
Species 1= A. plantago-aquatica Species 2= S. graminea Species 3= S. montevidensis
147 Chapter 8. Appendices
***** Analysis of variance *****
Variate: appressorium formation
Source of variation d.f. s.s. m.s. v.r. F pr. rep stratum 2 792.2 396.1 2.46 rep.disc stratum 15 2420.0 161.3 0.30 rep.disc.*Units* stratum species 2 10428.8 5214.4 9.60 <.001 Residual 34 18476.4 543.4
Total 53 32117.4
* MESSAGE: the following units have large residuals. rep 1.00 disc 4.00 -15.1 s.e. 6.7
***** Tables of means *****
Variate: appressorium formation
Grand mean 58.0
species 1.00 2.00 3.00 38.4 67.4 68.3
*** Least significant differences of means (5% level) ***
Table species rep. 18 d.f. 34 l.s.d. 15.79
Species 1= A. plantago-aquatica Species 2= S. graminea Species 3= S. montevidensis
148 Chapter 8. Appendices
TIME=5 (30 hours post inoculation)
***** Analysis of variance *****
Variate: germination
Source of variation d.f. s.s. m.s. v.r. F pr. rep stratum 2 4730.3 2365.1 28.19 rep.disc stratum 15 1258.3 83.9 0.62 rep.disc.*Units* stratum species 2 2298.3 1149.1 8.50 0.001 Residual 34 4595.1 135.1
Total 53 12881.9
* MESSAGE: the following units have large residuals. rep 3.00 disc 2.00 13.6 s.e. 4.8 rep 3.00 disc 6.00 *units* 1 -22.3 s.e. 9.2
***** Tables of means *****
Variate: germination
Grand mean 89.0
species 1.00 2.00 3.00 79.9 94.9 92.1
*** Least significant differences of means (5% level) ***
Table species rep. 18 d.f. 34 l.s.d. 7.88
Species 1= A. plantago-aquatica Species 2= S. graminea Species 3= S. montevidensis
149 Chapter 8. Appendices
***** Analysis of variance *****
Variate: appressorium formation
Source of variation d.f. s.s. m.s. v.r. F pr. rep stratum 2 2722.1 1361.1 19.39 rep.disc stratum 15 1052.8 70.2 0.21 rep.disc.*Units* stratum species 2 28354.5 14177.2 42.84 <.001 Residual 34 11252.1 330.9
Total 53 43381.5
* MESSAGE: the following units have large residuals. rep 3.00 disc 5.00 -10.0 s.e. 4.4 rep 3.00 disc 6.00 *units* 2 35.1 s.e. 14.4
***** Tables of means *****
Variate: appressorium formation
Grand mean 61.7
species 1.00 2.00 3.00 35.9 57.6 91.6
*** Least significant differences of means (5% level) ***
Table species rep. 18 d.f. 34 l.s.d. 12.32
Species 1= A. plantago-aquatica Species 2= S. graminea Species 3= S. montevidensis
150 Chapter 8. Appendices
TIME = 6 (36 hours post inoculation)
***** Analysis of variance *****
Variate: germination
Source of variation d.f. s.s. m.s. v.r. F pr. rep stratum 2 1453.44 726.72 34.65 rep.disc stratum 15 314.56 20.97 0.23 rep.disc.*Units* stratum species 2 370.11 185.06 1.99 0.153 Residual 34 3165.89 93.11
Total 53 5304.00
* MESSAGE: the following units have large residuals. rep 3.00 disc 3.00 -6.8 s.e. 2.4 rep 3.00 disc 3.00 *units* 2 17.0 s.e. 7.7 rep 3.00 disc 3.00 *units* 3 -19.9 s.e. 7.7 rep 3.00 disc 4.00 *units* 3 -24.6 s.e. 7.7
***** Tables of means *****
Variate: germination
Grand mean 93.3
species 1.00 2.00 3.00 91.1 97.0 91.9
*** Least significant differences of means (5% level) ***
Table species rep. 18 d.f. 34 l.s.d. 6.54
Species 1= A. plantago-aquatica Species 2= S. graminea Species 3= S. montevidensis
151 Chapter 8. Appendices
***** Analysis of variance *****
Variate: appressorium formation
Source of variation d.f. s.s. m.s. v.r. F pr. rep stratum 2 8960.9 4480.5 62.23 rep.disc stratum 15 1079.9 72.0 0.18 rep.disc.*Units* stratum species 2 23927.5 11963.7 30.41 <.001 Residual 34 13375.6 393.4
Total 53 47343.9
* MESSAGE: the following units have large residuals. rep 2.00 disc 4.00 9.6 s.e. 4.5 rep 2.00 disc 1.00 *units* 2 -35.9 s.e. 15.7 rep 2.00 disc 2.00 *units* 2 -36.8 s.e. 15.7
***** Tables of means *****
Variate: appressorium formation
Grand mean 68.8
species 1.00 2.00 3.00 39.8 77.7 89.0
*** Least significant differences of means (5% level) ***
Table species rep. 18 d.f. 34 l.s.d. 13.44
Species 1= A. plantago-aquatica Species 2= S. graminea Species 3= S. montevidensis
152 Chapter 9. Publications
9 Publications
Ash, G.J., Cother, E.J., Jahromi F.G., Pitt, W.M., Lanoiselet, V.M. and Cliquet, S. 2003. Status and future for biological control of aquatic weeds of rice in Australia. International Bioherbicide Group Workshop, Canberra, Australia, April 2003.
Ash, G.J., Cother, E.J., Jahromi F.G., Pitt, W.M., Lanoiselet, V.M. and Cliquet, S. 2003. Biological control of aquatic weeds of rice in Australia using Rhynchosporium alismatis. International Congress of Plant Pathology, Christchurch, New Zealand, February 2003.
Ash, G.J., Cother, E.J., Jahromi F.G., Pitt, W.M., Lanoiselet, V.M. and Cliquet, S. 2003. Developing Rhynchosporium alismatis for biological control of aquatic weeds of rice in Australia – past, present and future. Bioherbicide Workshop, Christchurch New Zealand, February 2003.
Pitt, W.M., Cother, E.J., Ash, G.J. 2002. Status Of Alismataceae Weeds in Australian Rice Crops and Potential for their biological Control. 2nd Temperate Rice Conference, Sacramento, California, USA, June 1999. Published Proceedings, p. 691.
Cother, E.J., Jahromi, F.G., Pitt, W.M., Ash, G.J., Lanoiselet, V.M. 2002. Development of the Mycoherbistat Fungus Rhynchosporium alismatis for Control of Alismataceae Weeds in Rice. 2nd Temperate Rice Conference, Sacramento, California, USA, June 1999. Published Proceedings, p. 509–513.
Jahromi, F.G., Pitt, W.M., Lanoiselet, V., Cother, E.J., Ash, G.J. 2001. Improving the activity of the mycoherbistat fungus, Rhynchosporium alismatis, for the biocontrol of aquatic weeds of rice in Australia. NATO Advanced Workshop on ‘Enhancing biocontrol agents and handling risks’, Florence, Italy.
Pitt, W.M., Jahromi, F.G., Ash, G.J., Cother, E.J. 1999. Alismataceae weeds in Australian rice crops: current control methods and the potential for biological control. 12th Australian Weeds Conference, Hobart, Tasmania, Australia, September, 1999. Published Proceedings, p. 664.
Pitt, W.M. 1999. Expanding the host range of Rhynchosporium alismatis, a potential biological control agent of Alismataceae weeds in rice fields – an update. Cooperative Research Centre for Sustainable Rice Production Symposium, Yanco Agricultural Institute, Yanco, New South Wales, Australia, August, 1999.
153 Chapter 10. References
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