a
Ericaceae root associated fungi revealed by culturing and culture – independent molecular methods.
by
Damian S. Bougoure
BSc (Hons)
Thesis submitted in accordance with the requirements for the degree of Doctor of
Philosophy
Centre for Horticulture and Plant Sciences
University of Western Sydney
February 2006 2
ACKNOWLEDGEMENTS
Although I am credited with writing this thesis there is a multitude of people that have contributed to its completion in ways other than hitting the letters on a keyboard and I would like to thank them here. Firstly I’d like to thank my supervisor, Professor John
Cairney, whose knowledge and guidance was invaluable in steering me along the PhD path. The timing of John’s ‘motivational chats’ was uncanny and his patience particularly, during the writing stage, seemed limitless at times.
I’d also like to thank the Australian government for granting me an Australian
Postgraduate Award (APA) scholarship, Paul Worden from Macquarie University and the staff from the Millennium Institute at Westmead Hospital for performing DNA sequencing and the National Parks and Wildlife Service of New South Wales and
Environmental Protection agency of Queensland for permission to collect the Ericaceae plants. Thankyou to Mary Gandini from James Cook University for showing me the path to a Rhododendron lochiae population through the thick North Queenland rainforest.
Without her help and I’d still be pointing the GPS at the sky.
Thankyou to the other people in the lab studying mycorrhizas including Catherine
Hitchcock, Susan Chambers, Adrienne Williams and particularly Brigitte Bastias with whom I shared an office. Everyone mentioned was generally just as willing as I was to talk about matters other than mycorrhizas. I’d also like to thank David Midgley and
Nicole Sawyer, both former members of the research group who remained good friends after they’d moved on. Thanks particularly to Dave who I faced off against many times 3 on the squash court. This was useful in offsetting the fruits of our homebrew enterprise
(A science background has more than one use). Thanks also to my friends, George and
Peta, who, to my relief at times, knew absolutely nothing of the world of mycorrhizas.
They helped me become a resident expert on the clubs of Western Sydney and we enjoyed multiple concerts at such venues as well as the ritual band nights at Hyland
Road.
I’d also like to thank everyone at the Macaulay Institute in Aberdeen who made an
Aussie feel most welcome. Thankyou particularly to Ian Anderson who gave me the opportunity to work in his group and Pamela Parkin who ensured my work there went like clockwork. Thanks also to Ian Alexander from The University of Aberdeen for help with statistical analyses and whose enthusiasm for science was always refreshing. I also thank him for choosing the sunniest day of the year in Scotland to sample in the field.
Thankyou to all my family which has recently extended to include a set of in-laws. Dad and Mum have always supported my endeavours, including this PhD, and continue to offer unconditional help whenever they’re called upon, for that I am eternally grateful.
Bones and Mizz, thankyou for being model siblings that have always made life outside the lab interesting and worthwhile.
Special thanks to my wife, Bec. Without her this thesis would not have been completed.
Her love, support, friendship, understanding and patience have helped me every day, particularly the hard ones when doing a PhD in Science seemed like a bad idea. Going 4 home to Bec at Myrtle street after a day in the lab was always something to look forward to when pouring agar plates became too much. Funnily enough, for someone with no biology background, Bec has developed (by no choice of her own) a substantial understanding of ericoid mycorrhizas.
Finally, I’d like to dedicate this thesis to my Nana who passed away during my PhD. She once gave me a second chance when I probably didn’t deserve one. Without that chance there’d be no thesis to dedicate to her.
Damian Bougoure
March 2006
5
I hereby certify that this thesis is my own work and contains no material that has been submitted for the award of any other degree or diploma in any university or any other tertiary institution and, to the best of my knowledge and belief, contains no material previously published or written by another person, except where due reference has been made in the text.
Signature:…………………… Date:………………..
Damian Bougoure
i
TABLE OF CONTENTS
Table of Contents ...... i
Abbreviations ...... vi
Abstract...... ix
CHAPTER 1: General Introduction ...... 1
1.1. The plant family Ericaceae...... 1
1.2. Fungal endophytes ...... 3
1.3. Ericoid mycorrhizas in Ericaceae...... 4
1.4. Other fungal root endophytes of Ericaceae ...... 7
1.5. Community biology of Ericaceae endophytes ...... 12
1.6. Analysis of Ericaceae fungal root endophytes using directly extracted DNA ...... 15
1.7. Research Aims ...... 17
CHAPTER 2: Assemblages of ericoid mycorrhizal and other root-associated fungi from Epacris pulchella (Ericaceae) as determined by culturing and direct DNA extraction from roots ...... 18 2.1. Introduction...... 18
2.1.1 Diversity of ericoid mycorrhizal fungi ...... 18
2.1.2 Identification of cultured endophytes ...... 21 2.1.3 Synthesis of ericoid mycorrhizas in gnotobiotic culture ...... 23
2.1.4 Are unculturable ericoid mycorrhizal fungal endophytes present in Ericaceae ii
roots?...... 26
2.1.5 Identification of fungal root endophytes using DNA extracted directly from
environmental samples...... 27
2.2. Experimental Procedures ...... 34
2.2.1. Collection of plants and isolation of fungal cultures ...... 34
2.2.2. Testing for ericoid mycorrhiza formation ...... 35
2.2.3. DNA extraction...... 36
2.2.4. ITS amplification, cloning, RFLP analysis and sequence analysis...... 36
2.2.5. ITS amplification for DGGE analysis ...... 39
2.2.6. DGGE analysis...... 40
2.3. Results ...... 41
2.3.1. ITS-RFLP analysis of cultured fungal assemblages and clones from directly
extracted DNA ...... 41
2.3.2. Sequence analysis of ITS RFLP-types ...... 49
2.3.3. Ericoid mycorrhiza formation ...... 50
2.3.4. Phylogenetic analysis of RFLP-types...... 53
2.3.5. DGGE analysis...... 57
2.4. Discussion...... 59
CHAPTER 3: Chitinolytic activities of ericoid mycorrhizal and other root- associated fungi from Epacris pulchella ...... 68
3.1. Introduction...... 68 iii
3.1.1 Edaphic stresses in habitats of Ericaceae and their associated ericoid
mycorrhizal fungi...... 68 3.1.2 Abilities of ericoid mycorrhizal fungi to utilise organic nitrogen sources...... 69
3.1.3 Abilities of ericoid mycorrhizal fungi to produce cell-wall-degrading enzymes ...... 72 3.1.4 Abilities of ericoid mycorrhizal fungi to produce chitinase enzymes...... 76 3.2. Experimental Procedures ...... 77
3.2.1. Fungal isolates and growth conditions ...... 77
3.2.2. Assay for extracellular chitinase activities...... 79
3.3. Results ...... 80
3.4. Discussion...... 84
CHAPTER 4: Fungi associated with hair roots of Rhododendron lochiae in an
Australian tropical montane cloud forest...... 87
4.1. Introduction...... 87
4.1.1. Habitat diversity of Ericaceae and associated ericoid mycorrhizal fungi...... 87
4.1.2. Ericoid mycorrhizas in tropical montane cloud forest...... 90
4.1.3. TMCF in Australia ...... 92
4.2. Experimental Procedures ...... 94
4.2.1. Collection of plants and isolation of fungal cultures ...... 94
4.2.2. DNA extraction...... 95
4.2.3. ITS amplification, cloning, RFLP analysis and sequence analysis...... 95
4.2.4. Testing for ericoid mycorrhiza formation ...... 96
4.2.5. ITS amplification for DGGE analysis ...... 96
4.2.6. DGGE analysis...... 97 iv
4.3. Results ...... 97
4.3.1. ITS-RFLP analysis of cultured fungal assemblages...... 97
4.3.2. Sequence analysis of ITS RFLP-types ...... 99
4.3.3. Ericoid mycorrhiza formation ...... 105
4.3.4. Phylogenetic analysis of ITS RFLP-types ...... 108
4.3.5. DGGE analysis...... 118
4.4. Discussion...... 122
CHAPTER 5: Assemblages of ericoid mycorrhizal and other root associated fungi from Ericaceae across a moorland : forest gradient in North-East Scotland ...... 130
5.1. Introduction...... 130
5.2. Experimental Procedures ...... 137
5.2.1. Field site description and sampling ...... 137
5.2.2. DNA extraction...... 138
5.2.3. ITS amplification ...... 139
5.2.4. DGGE analysis...... 140
5.2.5. T-RFLP analysis...... 141
5.2.6. Cloning and sequence analysis ...... 142
5.2.7. Data analysis...... 143
5.3. Results ...... 145
5.3.1. DGGE...... 145
5.3.2. T-RFLP...... 150
5.3.3. Cloning ...... 155
4.4. Discussion...... 168 v
CHAPTER 6: General Discussion...... 176
References ...... 197
Appendix I...... 249
vi
ABBREVIATIONS
4-MU 4-methylumbelliferyl
ARDRA amplified ribosomal DNA restriction analysis bp base pairs
BSA bovine serum albumin
C celsius ca approximately cm centimetre d day dATP deoxyadenosine triphosphate
DCA detrended correspondence analysis dCTP deoxycytidine triphosphate dGTP deoxyguanosine triphosphate
DGGE denaturing gradient gel electrophoresis diam. diameter
DNA deoxyribonucleic acid
DSE dark septate endophytes
DSF demateaceous surface fungi dTTP deoxythymidine triphosphate
E east
EDTA ethylenediaminetetraacetic acid
EP Epacris pulchella
FC forest Calluna vulgaris transect vii
FV forest Vaccinium myrtillus transect g gram
GC guanosine/cytidine h hour
ITS internal transcribed spacer
LiP lignin peroxidase
LSD least significant difference
LSU large subunit m metres
M molar
MC moorland Calluna vulgaris transect
MEA malt extract agar
MES morpholino-ethanesulfonic acid mg milligram min minute ml millilitres mm millimetres
MMN modified Melin-Norkrans
MnP manganese peroxidase nm nanometres
No. number
NSW New South Wales
PCR polymerase chain reaction viii
PDA potato dextrose agar pkat picokatals pmol picomoles rDNA ribosomal deoxyribonucleic acid
RFLP restriction fragment length polymorphism
RL Rhododendron lochiae
RPM revolutions per minute
S south sec second
TAE tris-acetate ethylenediaminetetraacetic acid
TC transition zone Calluna vulgaris transect
TMCF tropical montane cloud forest
T-RF terminal restriction fragment
T-RFLP terminal restriction fragment length polymorphism
UK United Kingdom
UV ultra violet
V volts w/v weight per volume
μg microgram
μl microliter
µm micrometer
µM micromolar
ix
ABSTRACT
Ericoid mycorrhizal fungi form mycorrhizal associations with Ericaceae plants and are regarded as essential to the ecological fitness of the plants in extremely nutrient-poor soils worldwide. The culturable fungal assemblages associated with hair roots of Epacris pulchella and Rhododendron lochiae (Ericaceae) from different forest habitats in eastern
Australia were investigated using rDNA internal transcribed spacer (ITS) restriction fragment length polymorphisms (RFLPs) and sequence analysis, and the abilities of the fungi to form ericoid mycorrhizas were tested. ITS sequences from cultured isolate assemblages were compared with fungi identified in DNA extracted directly from the same root systems by cloning and denaturing gradient gel electrophoresis (DGGE). A range of ericoid mycorrhizal and non-mycorrhizal fungi was identified from both host species using each approach, with the ericoid mycorrhizal fungi found to be taxonomically similar to those associated with Ericaceae in temperate habitats worldwide.
Although most of the abundant RFLP types identified in the cultured fungal assemblage were also the most abundant RFLP-types in the cloned assemblages and the most intense bands in DGGE profiles, each approach identified several unique fungi. This suggests that a combination of culturing and culture-independent approaches may be more efficacious than one method used individually.
Ericoid mycorrhizal endophytes and other root-associated fungi cultured from the E. pulchella, along with a Hymenoscyphus ericae isolate, were tested for their abilities to produce extracellular chitinolytic activities during growth in axenic culture. All fungi produced activities against at least one model chitin substrate. Specific activities for all x ericoid mycorrhizal fungi, including H. ericae, were of the same order of magnitude, suggesting that their chitinolytic potential is broadly similar. Chitinase activities were only produced by an ericoid mycorrhizal fungus when chitin was included in the medium, however no activity was produced if glucose was also present in the medium.
Ericaceae frequently represent the major understorey component of temperate and boreal forest habitats that are often adjacent to moorland heaths. While the structure of the
Ericaceae root associated fungal communities in these moorlands is relatively well studied, currently very little is known about these fungal communities in these adjacent forest understorey hosts. The community structure of Ericaceae root associated fungi was compared between adjacent moorland, transition zone and forest habitats in North-
East Scotland using DGGE, T-RFLP and cloning. Multivariate analyses identified clear differences in fungal endophyte community structure and composition between habitats.
Thus the communities in the open moorland and transition zone differed from each other, but the greatest differences were between these two and the community from the forest dwelling Ericaceae. DNA sequencing of selected clones revealed that typical mycorrhizal symbionts of Ericaceae, including fungi in the H. ericae complex, were common in Ericaceae roots across all habitats. This was also the case for a diverse array of Sebacinaceae fungi. Common ectomycorrhizal basidiomycetes were present in most samples from the forest but, were generally absent elsewhere. The functional significance of members of the H. ericae complex, Sebacinaceae and the ectomycorrhizal basidiomycetes are discussed particularly in regard to the possibility of symbiont sharing between Ericaceae and ectomycorrhizal hosts. 1
CHAPTER 1
General Introduction
1.1 The plant family Ericaceae
The plant family Ericaceae is currently one of 24 – 26 families assigned to the order
Ericales. This family division within the Ericales is based on recent phylogenetic studies that expanded the number of families from nine to this current figure (Anderberg et al.,
2002; Anderberg and Zhang, 2002; Bremer et al., 1998; Bremer et al., 2002). In the former nine family classification system, Empetraceae, Epacridaceae and Ericaceae were all assigned family status. Epacridaceae have an almost exclusively southern hemisphere distribution and the family was formerly regarded as the southern hemisphere Ericaceae equivalent (Cronquist, 1981). Cladistic analyses of morphological, phytochemical and embryological characters, along with phylogenies based on DNA sequence comparisons of a variety of chloroplastic and nuclear loci, increasingly suggest that Epacridaceae and
Empetraceae are derived lineages within the family Ericaceae (Andenberg 1993; Judd and Kron 1993; Kron and Chase 1993; Kron et al. 1996; Crayn and Quinn 2000; Bremer et al., 2002).
Considering this, recent taxonomic treatments regard the Ericaceae as a monophyletic family which comprises the eight sub-families Enkianthoideae, Monotropoideae,
Arbutoideae, Ericoideae, Cassiopoideae, Harrimanelloideae, Styphelloideae and
Vaccinioideae (Kron et al., 2002). Members of the former family Empetraceae are 2 morphologically distinct from all other Ericaceae, having reduced perianth parts and a propensity for wind pollination (Stevens, 1971), however, phylogenetic analyses placed this group as a tribe within the Ericoideae sub-family (Kron et al., 2002). Members of the former family Epacridaceae, by contrast, are morphologically similar to the other
Ericaceae with no consistent morphological characters separating the two families. For example, the dehiscence of pollen from epacrid anthers occurs via slits, while in the
Ericaceae dehiscence occurs primarily via pores. However, in some Ericaceae species dehiscence occurs via slit-like pores (Kron and Chase, 1993). This group represents the likely monophyletic sub-family Styphelloideae which is sister to the Vaccinioideae sub- family in the current Ericaceae taxonomy (Kron et al., 2002).
Ericaceae exist in a geographically widespread range of plant communities that includes
Magellanic tundra, boreal forests, tropical montane cloud forests, dry sclerophyll forests, sphagnum wetlands, mor humus heathlands and dry sand plains (Luteyn, 2002; Perotto et al., 2002). This habitat diversity has led to the Ericaceae evolving a multitude of life forms, however the greatest portion of species exist as shrubs in the tropical areas of the world (Luteyn, 2002). In terms of plant community structure, Ericaceae may represent a spectrum between the dominant vegetation form, as in the mor humus heathlands of the northern hemisphere, or be present only as rare epiphytes, which is often the case in tropical montane cloud forests (Luteyn, 2002). Despite the geographic and climatic diversity in Ericaceae distribution, the habitats they occupy have in common soils that are characteristically extremely low in nutrient, particularly nitrogen, status (Read, 1991) and have various combinations of extreme edaphic conditions including low pH, high metal 3 availability, poor drainage, free drainage, high levels of organic matter and high or low temperatures (Cairney and Meharg, 2003)
1.2 Fungal endophytes
The term ‘endophyte’ has been employed for all organisms that inhabit plants and mycologists use the term ‘fungal endophyte’ for fungi that inhabit plants without causing visible disease symptoms. The term refers only to fungi at the moment of detection, without regard for the future status of the interaction. These fungal endophytes represent a continuum of fungi with respect to physiological status, infection mode, colonisation pattern, secondary metabolism, life-history strategy, and developmental and evolutionary stages, but also with respect to the fungal and host taxa involved in the symbioses (Schulz and Boyle, 2005). The asymptomatic endophyte-host interaction seems to involve two actively antagonistic partners. However, a balanced antagonistic endophyte-host interaction does not exclude the possibility that the endophyte may play a beneficial role by increasing the fitness of the host, for example by inducing defence metabolites that may be active against pathogens and predators, by secreting phytohormones, by increasing the metabolic activity of the plant host (Schulz and Boyle, 2005) and (or) by supplying the host with nutrients from the rhizosphere (Smith and Read, 1997).
This notwithstanding, endophytes that, under certain conditions, interact mutualistically with their hosts may become pathogenic, for example when the host is stressed and the balance of the antagonism is tilted ‘in favour’ of the fungus (eg. Jumpponen 2001).
Balanced antagonistic interactions are dynamic and dependent upon the status of host and 4 endophyte and also on environmental factors. Furthermore, genetic variability between the individuals of a population, both host and fungal endophyte, means that endophytic interactions are flexible, having the ability to form a continuum of associations between more highly specialized mutualisms such as mycorrhizal fungi and highly specialized exploitations in the case of pathogens (Schulz and Boyle, 2005).
1.3 Ericoid mycorrhizas in Ericaceae
Although investigations carried out to date have revealed much about the diversity of fungal endophytes in the roots of Ericaceae, they have been restricted to relatively few
Ericaceae taxa from relatively few habitat types in primarily temperate to boreal areas
(Cairney and Ashford, 2002). The majority of Ericaceae root endophytes isolated to date are considered to be ericoid mycorrhizal fungi. Ericoid mycorrhizas are, by their etymology, mycorrhizas that occur exclusively with plants of the family Ericaceae.
Ericoid mycorrhizal symbiosis appears only to occur in the epidermal cells of their delicate hair roots which are involved in nutrient acquisition. The hair roots consist of three cell layers: a single layer of enlarged epidermal cells, an endodermis and a monarch stele (Cairney and Ashford, 2002). Chemicals, possibly polysaccarides, in the mucilage produced by host roots and fungal hyphae are thought to initiate infection (Bonfante-
Fasolo and Gianinazzi-Pearson, 1982) which leads to penetration of the plant cell wall via controlled expression of fungal cellulolytic enzymes (Duddridge and Read, 1982;
Bonfante-Fasolo and Gianinazzi-Pearson, 1982). After penetration of the cell wall, hyphae increase in thickness and begin to coil. McLean et al. (1998) noted that certain taxa produced dense coils which completely filled the epidermal cells, while other taxa 5 produced ‘loose’ coiling during infection, suggesting that the amount of coiling may be dependent on the fungal taxa. Coils are enveloped by the plant plasma membrane and it is believed that this position in the apoplast provides an increased surface area for exchange of material between host and symbiont (Cairney and Ashford, 2002).
The widespread occurrence of Ericaceae in edaphically harsh habitats is primarily due to the abilities of ericoid mycorrhizal fungi to minimise these environmental stresses.
Ericoid mycorrhizal fungi are, for example, thought to increase host plant resistance to toxic metals by detoxifying these potentially harmful compounds which are common in soils inhabited by many Ericaceae taxa (Cairney and Meharg, 2003). In addition to enhancing resistance to toxic metals, ericoid mycorrhizal fungi allow their hosts to acquire nutrients from organic sources via production of a range of hydrolytic and oxidative enzymes which give them access to a range of nitrogen and phosphorus sources in the soil that would be otherwise unavailable (Cairney and Meharg, 2003). Studies regarding functional aspects of ericoid mycorrhizas of Ericaceae have been primarily conducted using the fungal endophyte Hymenoscyphus ericae (Read) Korf and Kernan in the context of the northern hemisphere mor-humus heathland habitat. Although it has recently been proposed that this taxon is transferred to a new genus as Rhizoscyphus ericae (D.J. Read) W.Y. Zhuang & Korf (Zhang and Zhuang, 2004), this has not yet been universally adopted, thus H. ericae is retained throughout this thesis. H. ericae has been shown to produce a range of enzymes such as proteases and chitinases that give host plants direct access to organic sources of nitrogen (Kerley and Read, 1997). The same fungus also produces polyphenol oxidases which reduce inhibitory protein binding and 6 facilitate fungal access to tannin-bound proteins that frequently occur in the mor-humus habitats, as well as a suite of cell wall degrading enzymes that allow the endophytes to utilise complexed nutrient sources, such as proteins, which are associated with or internal to the plant cell wall (Bending and Read, 1996a; Cairney and Burke, 1998). In contrast to
H. ericae, very little is known in terms of the functional abilities of other ericoid mycorrhizal fungi from different host taxa or different habitats. Although multiple studies suggest that ericoid mycorrhizal fungi isolated from southern hemisphere
Ericaceae are capable of utilising organic nitrogen sources (eg. Midgley et al., 2004a), aside from one report of β-1-4 endoxylanase production by fungi isolated from Woollsia pungens Cav. (Cairney et al., 1996), there have been no investigations of hydrolytic or oxidative enzyme production by these fungi.
Ericoid mycorrhizal fungi appear to be represented by a limited number of ascomycete taxa (see Chapter 2). Many of those which are frequently isolated from Ericaceae roots in the northern hemisphere belong to the expanding group of Helotiales fungi considered to represent the H. ericae aggregate (Vrålstad et al., 2002b). A growing number of sterile unidentified fungi also with an affinity to the Helotiales, but taxonomically distinct from the H. ericae aggregate, have been isolated from both northern and southern hemisphere hosts. Myxotricaceae fungi, particularly members of the Oidiodendron genus, are also currently considered common mycorrhizal endophytes of temperate northern hemisphere
Ericaceae. Aside from forming ericoid mycorrhizal coils, the mutualistic nature of the association formed by fungi from these groups with Ericaceae hosts has been demonstrated. Thus, carbon has been shown to move from the host to the fungi, while 7 essential nutrients are known to pass reciprocally from the fungi to the host resulting in positive growth responses (Smith and Read, 1997). While ericoid mycorrhizal fungi are widely accepted as mutualistic, their roles within root systems under differing environmental conditions are less clear. Arbuscular mycorrhizal fungi have been variously cited as forming a continuum of positive, neutral and negative associations with their hosts (Jones and Smith, 2004) and it is likely that a similar range of relationships exists between the host and the various ericoid mycorrhizal taxa and other root-associated fungi from the Ericaceae (see below). In plant roots, fungi that are not normally considered to be mycorrhizal can also have positive effects and form 'mycorrhiza-like' associations or improve overall host fitness by other means. Conversely, known mycorrhizal fungi can potentially act in a negative fashion in some circumstances, for example, under conditions of fertilizer application (eg. Kiernan, 1983; Smith and Smith,
1996).
1.4 Other fungal root endophytes of Ericaceae
In many habitats in both the northern and southern hemispheres, a number of dark septate endophyte (DSE) taxa are commonly found to be associated with the roots of Ericaceae and other plant taxa (eg. Stoyke and Currah, 1991; Midgley, 2003; Vrålstad et al., 2002b;
Williams et al., 2004). Phialocephala fortinii Wang and Wilcox has been found as the dominant root endophyte of members of the Asteraceae, Caryophyllaceae, Ericaceae,
Poaceae, Primulaceae, and Rosaceae at an alpine field site in North America (Stoyke and
Currah, 1991; Stoyke et al., 1992). Infection by P. fortinii has been shown to improve plant growth in terms of total biomass in Pinus contorta Dougl. ex Loud. (Jumpponen 8 and Trappe, 1998b) and aid in the suppression of some fungal plant pathogens in
Solanum melongena L. (Narisawa et al., 2002). P. fortinii produces microsclerotia within the epidermal cells of Ericaceae rather than hyphal coils as seen in ericoid mycorrhizal fungi (Jumpponen and Trappe, 1998a). Another commonly isolated DSE is Cadophora finlandia Wang and Wilcox (= Phialophora finlandia Wang and Wilcox), which is associated with a broad range of plants including Ericaceae taxa (Vrålstad et al., 2000) and is likely to represent a true ericoid mycorrhizal fungus. Monreal et al. (1999) reported that, in ‘limited’ preliminary trials, C. finlandia formed typical ericoid mycorrhizal structures in roots of Gaultheria shallon Pursh, however this does not appear to have been confirmed. C. finlandia has Helotiales affinities and is therefore likely to be related to the large assortment of unidentified ericoid mycorrhizal fungi (LoBuglio et al.,
1999). Recently, phylogenetic analysis by Vrålstad et al. (2002b) has suggested that C. finlandia forms a part of the H. ericae aggregate. The endophytic status of other DSE isolated from Ericaceae roots remains unclear. At least some of these fungi have, however, been shown to increase host plant biomass and improve phosphorus uptake in non-Ericaceae plants, suggesting that a mutualistic relationship between the fungus and the plant is possible (Haselwandter and Read, 1982; Jumpponen et al., 1998).
A wide taxonomic range of ascomycete families other than those confirmed to be ericoid mycorrhizal fungi or DSE has been isolated from Ericaceae roots. Acremonium strictum
W. Gams (Hypocreales) forms ericoid mycorrhiza-like structures with G. shallon and was also shown to improve host productivity when grown with a variety of organic nitrogen sources (Xiao and Berch, 1995). Although A. strictum has not been reported in any other 9 ericoid mycorrhizal plant species, it is a common endophyte of a large number of grasses and sedges. Grasses infected with A. strictum are reported to have increased resistance to nematode infection, along with increased drought tolerance (Belesky et al., 1987; West et al., 1987) and allelopathy (Quigley et al., 1990). Allen et al. (2003) cultured multiple isolates from G. shallon that were closely related to the Capronia genus
(Herpotrichiellaceae) and subsequent gnotobiotic reinoculation experiments clearly showed that these isolates colonise host roots forming coil-like structures. Bergero et al.
(2000) also isolated Capronia-like fungi from Italian Ericaceae and showed that at least one isolate formed typical hyphal coils when reinoculated into the host. Midgley (2003) isolated Capronia-like fungi from the roots of W. pungens, however their physiological potential as mutualistic symbionts was not tested in any of the aforementioned investigations.
Xylariales-like isolates have also been obtained from Ericaceae roots, and a single culture from G. shallon formed ericoid mycorrhiza-like coils when reinoculated into the host
(Berch et al., 2002), however no studies to ascertain the nature of the association were carried out. Xylariaceae are common endophytes of multiple plant species, particularly in tropical regions where Ericaceae species are common. Studies of endophytic Xylaria from non-Ericaceae hosts have shown that they actively produce antibiotic secondary metabolites in axenic culture (Brunner and Petrini, 1992), and these may also be produced when the fungi inhabit living plant tissues. There has, however, been no research on how these important compounds may affect endophyte : host associations.
10
Multiple cultured isolates from a variety of habitats and Ericaceae taxa have been shown to be related to Cryptosporiopsis sp. (eg. Berch et al., 2002). Members of this genus are considered to be non-mycorrhizal root colonizers but have been shown to improve growth of non-Ericaceae hosts (Schulz and Boyle, 2005) which suggests a potential to improve Ericaceae host fitness. Colonisation of the roots of the Chinese cabbage
(Brassica campestris Ibaraki) by the endophyte Heteroconium chaetospira (Grove) M.B.
Ellis, almost completely suppressed disease caused by Plasmodiophora brassicae Ibaraki
(Narisawa et al. 1998, Usuki et al. 2002). This endophytic fungus has been isolated from soils inhabited by Ericaceae species in Canada and was shown to form ericoid mycorrhiza–like coils that neither promoted nor reduced host plant growth when inoculated into Rhododendron obtusum G. Don (Ericaceae) roots (Usuki and Narisawa,
2005). These findings suggest that this fungal endophyte could possibly increase
Ericaceae fitness by means other than enhanced nutrient uptake. Other ascomycete fungi commonly cultured from Ericaceae roots include isolates with affinities to Dermataceae,
Sordariales, Dothioraceae, Rhytismatales and Hyaloscyphaceae. None of these fungi have, however, been shown to form structures resembling typical ericoid mycorrhizal coils when used in gnotobiotic reinoculation experiments, thus the nature of their associations is unclear (eg. Monreal et al., 1999; Allen et al., 2003; Midgley et al., 2002,
2004c; Williams et al., 2004). Allen et al. (2003) suggested that these fungi are probably saprobic, parasitic or simply opportunistic with non-symbiotic colonisation.
Ericaceae from the sub-family Monotropoideae are considered to be myco-heterotrophic plants and form mycorrhizal associations that are distinct from those of other Ericaceae. 11
At this stage it seems highly unlikely that mycorrhizas formed by the heterotrophic
Ericaceae are mutualistic. The fungi involved in these associations are those traditionally thought of as ectomycorrhizal basidiomycetes such as Russula, Hydnellum and
Tricholoma spp. . The fungi form sheath-like structures around Monotropoideae roots which connect via a common hyphal link with ectomycorrhizal hosts. This hyphal link allows the Monotropoideae to obtain carbon from the ectomycorrhizal hosts without providing any benefit in return (Bidartondo, 2005).
Arctostaphylos, Arctous and Arbutus genera from the Arbutoideae sub-family form arbutoid mycorrhizal associations which are also distinct from other mycorrhizal associations in the Ericaceae. The fungi that form these associations are also typical ectomycorrhizal basidiomycetes and form a fungal sheath over the Arbutoideae roots
(Smith and Read, 1997). A major difference between arbutoid and ectomycorrhizal associations is that the hyphae of the former actually penetrate the outer cortical cells, and fill them with coils, making them more similar to ect-endomycorrhizal associations
(Smith and Read, 1997). Arbutoid mycorrhizal associations are considered mutualistic whereby there is a reciprocal transfer of nutrients to the host and carbon to the fungal endophyte (Smith and Read, 1997).
To date, most fungal root endophytes cultured from Ericaceae other than members of the
Monotropoideae and Arbutoideae sub-families have been identified as ascomycetes.
However, the unculturable basidiomycete, Clavaria argillacea Fr. has often been implicated as a possible mycorrhizal endophyte, due to the production of fruiting bodies 12 around Ericaceae plants in nature. Furthermore, the prescence of dolipore septa in root epidermal cells of Rhododendron around which Clavaria sp. were found, along with limited trials which suggested reciprocal transfer of carbon and phosphorus between C. argillacea and two ericaceous plants (Seviour et al., 1973; Bonfante-Fasolo, 1980;
Englander & Hull, 1980; Peterson et al., 1980) emphasise the possibility of this mycorrhizal relationship. Allen et al. (1989) observed dolipore septa in hyphal coils within the epidermal cells of Dracophyllum secundum (Poiret) R. Br roots, suggesting that at least some basidiomycetes may be involved in ericoid mycorrhizal symbiosis in southern hemisphere Ericaceae.
1.5 Community biology of Ericaceae endophytes
When Ericaceae host plants are collected from the field, a diverse array of endophyte taxa is isolated from the host root system, however, generally one taxon is isolated more frequently than other taxa (eg. Perotto et al., 1996; Xiao and Berch, 1996; Chambers et al., 2000; Sharples et al., 2000; Liu et al., 1998). By combining careful mapping of isolate origin and molecular analyses of isolate genetic diversity, Midgley et al. (2002,
2004c) were able to investigate fungal endophyte distribution within individual W. pungens and Leucopogon parviflorus (Andr.) Lindl. root systems. A single endophyte genotype was widely distributed within the root system of a single plant, indicating that individual Australian Ericaceae root systems, although inhabited by multiple taxa, are spatially dominated by a single fungal genotype. It has been shown that there is intra- and interspecific variation of nitrogen utilisation by Ericaceae endophytes, with various taxa and genotypes showing preferences for different amino acids or proteins (Cairney et 13 al., 2000; Whittaker and Cairney, 2001, Midgley et al., 2004a; 2004b). The availability of various inorganic and organic nitrogen fractions in Ericaceae habitat soils is known to vary according to edaphic factors such as water availability and fire (eg. Schmidt and
Stewart, 1997). Therefore, both interspecific and intraspecific diversity in nitrogen source utilisation by mycorrhizal endophytes of Ericaceae (those both spatially dominant and less common) may be functionally important in maintaining host nitrogen supply from diverse, complex and sometimes scarce pools, whereby different taxa and genotypes may be able to utilise different nitrogen sources (Whittaker and Cairney, 2001). Only two recent studies have investigated the effect of fungal species richness on host plant productivity, both of which focused on ectomycorrhizal associations. These found that increased species richness had varying affects on seedling productivity (Jonsson et al.,
2001) and increased phosphorus uptake, shoot nitrogen concentration and root biomass of the host plants (Baxter & Dighton, 2001). These studies suggest that the ability of an ectomycorrhizal host plant to form associations with a mixed population of fungi composed of multiple taxa may be beneficial to the host plant as different taxa may be able to access different nutrients and this is likely to also be the case for ericoid mycorrhizal associations.
Liu et al. (1998) isolated endophytes from four W. pungens plants at an eastern
Australian sclerophyll forest field site and found three of the plants to be dominated by a single taxon out of a total of six isolated at the site. It was shown that most of the isolates of this taxon within the host root systems were of a single genotype, suggesting that
Australian Ericaceae root systems at a single field site may be extensively colonised by 14 individual mycelia (Liu et al., 1998). In northern hemisphere habitats it appears that either H. ericae or Oidiodendron maius Barron may be dominant in individual hosts as well as at particular sites. For example, Calluna vulgaris L. Hull roots were colonised by multiple endophytes, but mostly by H. ericae at a site in south-west England, while O. maius was the dominant endophyte isolated from the same species at a site in northern
Italy (Perrotto et al., 1996; Sharples et al., 2000). These observations in both the southern and northern hemisphere suggest that, at particular sites representing a range of habitats, the endophyte community, although diverse, is dominated by a single ericoid mycorrhizal taxon and possibly even a single genotype of that taxon.
Although the community structure of fungi associated with Ericaceae roots has been well studied at the scale of an individual habitat, very little is known regarding if and/or how the community structure changes across adjacent but different habitats that are occupied by Ericaceae hosts. These situations occur in the southern hemisphere, for example in eastern and south-western Australia where Ericaceae taxa represent one of the dominant vegetation types in dry heathlands which are adjacent to sclerophyll forest habitats where
Ericaceae are also a common component of the understorey. In the northern hemisphere this scenario is even more common. Ericaceae frequently represent the major understorey component of temperate and boreal broadleaf and conifer forest habitats that are often adjacent to moorland heaths where Ericaceae represent the dominant vegetation type (Read, 1991). While the structure of the Ericaceae root-associated fungal communities in these moorland heaths is relatively well studied, currently very little is known about communities of these fungi in adjacent forest understorey Ericaceae. 15
Recent evidence suggests at least some members of the H. ericae aggregate have the ability to also form ectomycorrhizal associations, and it has been suggested that single isolates may be able to form both types of associations (Monreal et al., 1999). This seems increasingly likely based on the findings of Villarreal-Ruiz et al. (2004) who demonstrated that a single isolate from the H. ericae aggregate, simultaneously in time and from a continuous mycelium, developed typical ericoid mycorrhizal coils in epidermal cells of Vaccinium myrtillus L. and an ectomycorrhizal Hartig net and mantle with Pinus sylvestris L.. Based on these findings the authors suggested that some members of the H. ericae aggregate may have evolved as strict ericoid mycorrhizal endophytes in moorland heaths, but other, taxonomically distinct and primarily ectomycorrhizal members of the aggregate, form ericoid mycorrhizal associations with understorey Ericaceae. This idea implies that there may be a change in the Ericaceae root associated fungal communities across adjacent moorland heath and forest habitats.
Furthermore it raises some interesting questions regarding possible hyphal links between
Ericaceae and ectomycorrhizal hosts in both the forest habitats and bordering areas of expanding forest.
1.6 Analysis of Ericaceae fungal root endophytes using directly extracted DNA
Molecular community profiling techniques that utilise nucleic acids extracted directly from environmental samples have led to considerable advances in microbial ecology.
Techniques such as random cloning, denaturing gradient gel electrophoresis (DGGE), terminal restriction fragment length polymorphism (T-RFLP) analysis or amplified 16 ribosomal DNA restriction analysis (ARDRA) are now finding favour with fungal ecologists. These techniques have been important in confirming the limitations of culturing fungi as a means of assessing community diversity in a variety of substrates
(Anderson and Cairney, 2004).
Bergero et al. (2000) observed unidentified basidiomycete hyphae within epidermal cells of hair roots of Erica arborea L. The authors suggested that certain taxa might be overlooked as endophytes of Ericaceae since, as has been reported for other environmental samples (eg. Bridge and Spooner, 2001), commonly employed culture media are unsuitable for growing a variety of micro-organisms including fungi. In order to overcome this potential culture bias, Allen et al. (2003) compared partial rDNA internal transcribed spacer (ITS) sequences from endophytes cultured from roots of G. shallon (Ericaceae) with sequences cloned from DNA extracted directly from the same root systems. More than 50% of the cloned sequences from directly extracted DNA were similar to the basidiomycete genus Sebacina, however most cultured isolates were identified as ascomycetes with no Sebacina-like sequences identified in the cultured assemblage. Despite the molecular investigation of Allen et al. (2003), along with others who have observed basidiomycete fungi within ericaceous hair roots using morphological investigations (eg. Allen et al., 1989; Bergero et al., 2000), the possible inability to culture basidiomycete and other fungi from roots limits our understanding of the functional roles (possibly mutualistic) these fungi may play as Ericaceae endophytes.
17
1.7 Research Aims
It seems increasingly likely that culturing alone does not represent the actual fungal community present in Ericaceae roots. In order to investigate the effects of fungal endophyte (including ericoid mycorrhizal) diversity on Ericaceae host plant fitness and functioning, it is important to have information regarding the actual composition and structure of the endophyte community rather than that which is based solely on the culturable endophytes. Furthermore it is important to study a variety of Ericaceae taxa from multiple isolated and adjacent habitat types to make broad assessments regarding
Ericaceae root endophyte diversity and the nature of their associations. The research aims of the current study were therefore:
(i) to compare culturing with molecular profiling techniques based on direct DNA extraction for evaluating the diversity and structure of the root endophyte communities of two Australian Ericaceae species, Epacris pulchella Cav. from the understorey of dry sclerophyll forest habitats and Rhododendron lochiae F. Muell from a tropical montane cloud forest.
(ii) to investigate the physiology of the dominant ericoid mycorrhizal and root associated endophytes from E. pulchella with respect to their abilities to access an organic nitrogen source.
(iii) to investigate if and how the structure of Ericaceae root associated fungal communities change across adjacent moorland and forest habitats. 18
CHAPTER 2
Assemblages of ericoid mycorrhizal and other root-associated fungi from Epacris pulchella (Ericaceae) as determined by culturing and direct DNA extraction from roots
2.1 Introduction
2.1.1 Diversity of ericoid mycorrhizal fungi
Numerous ericoid mycorrhizal fungal endophytes have been isolated, however only a few have been identified, most of which are ascomycetes or their anamorphs (Cairney and
Ashford, 2002). Hymenoscyphus ericae is a common mycorrhizal endophyte of temperate northern hemisphere Ericaceae taxa such as Calluna, Vaccinium and Gaultheria (Read,
1974; Berch et al., 2002; Sharples et al., 2000; Vrålstad et al., 2000, 2002b). A number of unidentified endophytes which are distinct from, but have close affinity to, H. ericae have been isolated from the roots of a variety of Ericaceae and non-Ericaceae taxa. Based on phylogenetic analyses of sequences it has been suggested that these isolates, many of which form ericoid mycorrhizas, comprise part of a H. ericae aggregate (Vrålstad et al.,
2000, 2002b; Berch et al., 2002; Cairney and Ashford, 2002; Haug et al., 2004). There is no direct evidence suggesting that H. ericae is a mycorrhizal endophyte of Australian
Ericaceae taxa, however, several endophytes that may be closely related to members of the H. ericae aggregate have been isolated from Australian epacrids. These isolates, which have >90 % ITS sequence similarity to H. ericae, were isolated from Epacris impressa Labill., Woollsia pungens and Epacris microphylla R. Br. in eastern Australian habitats (McLean et al., 1998; Midgley et al. 2002; Williams et al., 2003). Furthermore, 19 the isolates from E. impressa and W. pungens have been shown to form ericoid mycorrhizas in laboratory inoculation experiments (McLean et al., 1998; Midgley et al.
2002). A rhizoid endophyte of the leafy liverwort Cephaloziella exiliflora Taylor with
97.9 % ITS sequence identity with H. ericae has also been isolated from an eastern
Australian habitat where epacrids are common members of the plant community
(Chambers et al., 1999).
It has been known for some time that Myxotricaceae Oidiodendron species are also common mycorrhizal endophytes of temperate northern hemisphere Ericaceae. Several
Oidiodendron species (O. cerealis (Thum.) Barron, O. citrinum Barron, O. flavum
Szilvinyi, O. maius, O. rhodogenum Robak and O. tenuissimum (Peck.) Hughes) have been shown to form ericoid mycorrhizas in gnotobiotic culture (Dalpé, 1986, 1989, 1991;
Douglas et al., 1989; Perotto et al., 1995; Smith and Read, 1997; Stoyke and Currah,
1991; Xiao and Berch, 1992, 1995). O. maius appears to be a dominant symbiont of many field collected Ericaceae examined to date (Perrotto et al., 1996; Hambleton and Currah,
1997). The number of mycorrhizal Oidiodendron species is uncertain due to misidentification of culture strains and field isolated endophytes, along with the fact that species boundaries within the genus are unclear (Hambleton et al., 1998; Lacourt et al.,
2001). The only evidence suggesting that Oidiodendron species may form mycorrhizal associations with Australian Ericaceae was the isolation of an endophyte having > 90%
ITS sequence similarity to O. maius from W. pungens growing in an eastern Australian
Sclerophyl forest (Chambers et al., 2000).
20
A diverse morphological array of slow-growing sterile endophytes has also been isolated from both northern and southern hemisphere Ericaceae. Many of these endophytes have been reinoculated into hair roots of epacrids (Read, 1996; McLean et al., 1998; Anthony et al., 2000; Midgley et al., 2004c) and other Ericaceae (Reed, 1996; Liu et al., 1998) and formed typical ericoid mycorrhizal structures, which suggests a mycorrhiza-forming ability. The precise taxonomic status of most of these endophytes remains uncertain, however ITS sequence comparisons are beginning to shed some light on the matter.
Phylogenetic analyses of numerous ITS sequences of both confirmed and putative ericoid mycorrhizal endophytes from a variety of Ericaceae taxa, along with a number of ascomycetes have been carried out (Monreal et al., 1999; Sharples et al., 2000; Berch et al., 2002; Vrålstad et al., 2002b). The analyses initially separated the fungi into two distinct clades; one being part of a Hymenoscyphus-like group, the other a poorly defined
Helotiales-like group (Monreal et al., 1999; Sharples et al., 2000). However, more recent analyses have indicated a greater diversity amongst these sterile endophytes and Berch et al. (2002) placed them into seven distinct groups. Four of these probably represent
Helotiales (including the H. ericae aggregate), Myxotricaceae (including Oidiodendron species), Chaetothyriales and a polyphyletic group with representatives from different orders. These groups appear to consist of geographically widespread taxa that are isolated from multiple habitats and Ericaceae species.
When Ericaceae host plants are collected from the field, a diverse array of endophyte taxa is isolated from their roots, however, generally one taxon is isolated more frequently than others. This suggests that, at particular sites, the endophyte community is dominated by a 21 single taxon (Perotto et al., 1996; Xiao and Berch, 1996; Chambers et al., 2000; Sharples et al., 2000; Liu et al., 1998). In some northern hemisphere habitats it appears that either
H. ericae or O. maius may be dominant at a particular site. For example, Calluna vulgaris roots were colonised by multiple endophytes, but mostly by H. ericae at a site in south-west England, while O. maius was the dominant endophyte isolated from the same species at a site in northern Italy (Perrotto et al., 1996; Sharples et al., 2000).
Liu et al. (1998) isolated endophytes from four W. pungens plants at an eastern
Australian field site and found three of the plants to be dominated by a single taxon out of a total of six isolated at the site. It was shown that most of the isolates of this taxon within the root system of a single plant were of a single genotype, suggesting that
Australian Ericaceae root systems may be extensively colonised by individual mycelia
(Liu et al., 1998). By combining careful mapping of isolate origin with molecular analyses of isolate genetic diversity, Midgley et al. (2002, 2004c) were able to investigate fungal endophyte distribution within individual W. pungens and Leucopogon parviflorus root systems. A single genotype was widely distributed within the root system of a single plant, indicating individual Australian Ericaceae root systems are spatially dominated by a single fungal genotype.
2.1.2 Identification of cultured endophytes
Initially, many cultured fungal endophytes from the roots of Ericaceae were identified based on morphological traits (Hambleton et al., 1998a), however many of the sterile fungal cultures obtained are morphologically similar, rendering species identification 22 problematic (Hambleton et al., 1998a). A number of studies have attempted to characterise ericoid mycorrhizal fungi into morphotypes based on gross culture characteristics such as colour, texture and production of exudates (eg. Hutton et al., 1994;
McLean et al., 1999). These characters are, however, largely dependent on the media in which the fungi are maintained and length of time in culture. Characters such as colour and texture can vary considerably between different cultures of genetically identical isolates when grown on media that differ in a single chemical constituent such as phosphorus (Reed, 1989; Steinke et al., 1996; Liu et al., 1998 and Midgley, 2003).
Due to these difficulties, molecular methods have become widely used for investigations of cultured ericoid mycorrhizal fungal isolates. The commonly-adopted method employs restriction fragment length polymorphism (RFLP) analyses of the internal transcribed spacer (ITS) region, and has given rise to most of what we currently know about the identity of ericoid mycorrhizal fungal endophytes. The ITS region is located between the large and small subunit ribosomal rRNA genes and contains two non-coding spacer regions and the 5.8S rRNA gene. In Helotiales ascomycetes the entire ITS region (ITS1-
5.8S-ITS2) is generally between 500 and 650 bp in length (eg. McLean et al., 1998;
Chambers et al., 2000; Sharples et al., 2000). RFLP analysis of the ITS region (ITS-
RFLP) has become a widely used method of distinguishing putative species and has been extensively utilised in studies of ericoid mycorrhizal fungal communities (eg. Monreal et al., 1999; Sharples et al., 2000; Vrålstad et al., 2002b; Midgley et al., 2002, 2004c). ITS-
RFLP analysis involves PCR amplification of the ITS region, followed by single restriction enzyme digests. Two to four restriction enzyme digests are typically sufficient 23 to distinguish most putative species (Horton and Bruns, 2001). ITS fragments representing putative taxa, as designated by RFLP analysis, are subsequently sequenced and compared to known sequences in the GenBank/EMBL/DDBJ databases in order to assign meaningful identities to the fungal endophytes.
2.1.3 Synthesis of ericoid mycorrhizas in gnotobiotic culture
Demonstrating the reciprocal transfer of nutrients between host and endophyte and positive growth responses by both are considered essential by some to assign true mycorrhizal status to a fungal endophyte (Smith and Read, 1997). This notwithstanding, the formation of typical ericoid mycorrhizal coils in the epidermal cells of Ericaceae host hair roots under gnotobiotic conditions is regarded as reasonable confirmation of the mycorrhizal status of an endophyte. Northern hemisphere Ericaceae, including various members of the genus Vaccinium, C. vulgaris and G. shallon have routinely been used in inoculation experiments to confirm the ericoid mycorrhizal status of fungal endophytes
(eg. Pearson and Read, 1973; Sharples et al., 2000; Xiao and Berch, 1995). These plants are ideal candidates for such experiments as they germinate easily and grow readily under axenic culture conditions. Unlike northern hemisphere Ericaceae, seeds of many
Australian Ericaceae are difficult or slow to germinate and some species produce seedlings that are difficult to maintain in axenic culture (Williams, 1986; Reed, 1989;
McLean et al., 1998; Anthony et al., 2000). Some success in growing cuttings of E. impressa has, however, been achieved using non-sterile soil which was likely to contain ericoid mycorrhizal fungal inoculum and other microorganisms (McLean et al., 1994). 24
Clonal micropropagation has also been achieved for Leucopogon objectus Benth. and E. impressa in tissue culture (Bunn et al., 1989; Anthony et al., 2000; Lawrie et al., 2001).
In order to overcome the difficulty of growing Australian Ericaceae, the mycorrhizal status of endophytes has often been tested using northern hemisphere Ericaceae as hosts
(Reed, 1989; Liu et al., 1998). Formation of ericoid mycorrhizal coils in northern hemisphere Ericaceae by endophytes derived from Australian Ericaceae allows them to be designated as mycorrhizal. However, failure to form typical ericoid mycorrhizal structures does not necessarily preclude those fungi from being mycorrhizal with
Australian Ericaceae in the field and factors such as host-fungus specificity, carbon- nitrogen ratios and production of phenolic compounds by the host plants have been routinely cited as reasons for failed mycorrhizal synthesis experiments involving northern hemisphere hosts (Liu et al., 1998; McLean et al., 1998). To overcome this complication it is therefore advisable to inoculate the fungi onto host plants that are as closely related as possible to the species from which it was originally obtained.
A number of protocols have been trialled in order to improve the germination, maintenance and ericoid mycorrhiza formation of both northern and southern hemisphere
Ericaceae. Improved germination of seed of some Australian Ericaceae species has been observed after treatment with smoke or aqueous smoke derivatives, soaking and/or heat shock (Dixon et al., 1995; Keith, 1997; DJ Midgley, pers. com.). Liu (1998) observed that formation of hair roots of Dracophylum secundum (Poiret) R. Br. was severely restricted when grown on homogeneous agar based media and this was also the case for 25
W. pungens (DS Bougoure, unpublished data). To overcome the problems of homogenous media, the use of sterile soil or a mixture of peat and vermiculite placed on top of water agar plates have been used and shown to increase hair root development and ericoid mycorrhiza formation (Pearson and Read, 1973; Starett et al., 2001a, 2001b). The authors suggested however, that soil and peat are difficult to chemically characterise and vary between sites, making replication difficult. Duclos and Fortin (1983) demonstrated increased mycorrhizal formation and maintenance in agar media supplemented with activated charcoal and glucose, however it has been suggested that excessive carbon can cause mycorrhiza formation between host and symbiont that would not normally occur in the field (Smith & Read, 1997). Other authors have advocated the use of modified
Melin-Norkrans (MMN) medium with reduced nitrogen, phosphorus and carbon (Xiao and Berch, 1995; Vrålstad et al., 2002a) in order to increase and validate mycorrhiza formation in northern hemisphere hosts. Despite the acknowledged difficulties in growing Australian Ericaceae, several studies using agar based systems have achieved infection of these hosts with endophytes originating from Australia in addition to the northern hemisphere (Reed, 1989; McLean et al., 1998).
Recently, some success has been achieved in maintaining seedlings of, and demonstrating ericoid mycorrhiza formation with, W. pungens. Palmer and Ashford (2004) demonstrated that seeds of this species germinated relatively easily on moist filter paper and in non-sterile soil in which plants were successfully maintained until maturation.
Furthermore, Midgley (2003) maintained the same species on a sphagnum-supplemented agar medium which was more heterogenous in nature than other agar based media that 26 have been used for inoculation experiments. Using this system Midgley (2003) was able to demonstrate ericoid mycorrhiza formation by endophytes that were isolated from W. pungens. Based on these observations it seems that W. pungens may not be as difficult to maintain in culture as other Australian Ericaceae and as such may represent a useful
‘model’ plant to use for inoculation experiments for fungal endophytes obtained from
Australian.
2.1.4 Are unculturable ericoid mycorrhizal fungal endophytes present in Ericaceae roots?
Most of what is known regarding diversity of ericoid mycorrhizal fungi, and for that matter all fungi in the environment, is based on data derived using culture–based techniques, the limitations of which have been highlighted elsewhere (e.g. Zak and
Visser, 1996; Bridge and Spooner, 2001). Based on information obtained using these techniques, most cultured ericoid mycorrhizal fungi appear to be ascomycetes, however, several reports suggest that basidiomycetes may also form ericoid mycorrhizal associations. Observations of fruiting bodies of the basidiomycete Clavaria argillacea around Ericaceae plants in nature has implicated this fungus as a possible ericoid mycorrhizal fungus (Seviour et al., 1973; Bonfante-Fasolo, 1980; Englander and Hull,
1980; Peterson et al., 1980). However, difficulties in culturing Clavaria spp. mean that no examination of its mycorrhizal potential have been undertaken under gnotobiotic culture conditions. Roots from a Rhododendron sp., around which Clavaria sp. fruiting bodies were found, have been studied, and hyphae with dolipore septa were found in epidermal cells, suggesting that a basidiomycete may form an ericoid mycorrhizal 27 association with this host (Bonfante-Fasolo, 1980; Peterson et al., 1980). Englander and
Hull (1980) also demonstrated reciprocal transfer of radioisotopes of carbon and phosphorus between C. argillacea and two ericaceous plants (Rhododendron sp. and
Pieris japonica D. Don.), which supported this suggestion. These experiments were carried out in non-sterile pot culture so it remains unclear whether Clavaria spp. were obtaining carbon from the host via a saprotrophic, necrotrophic or mycorrhizal relationship.
In a recent study it was suggested that the traditional plating-out technique employed for isolation of ericoid mycorrhizal fungal endophytes may be selective for certain taxa, and that many unculturable endophytes may have been overlooked in studies conducted to date (Bergero et al. 2000). These authors made this suggestion based on microscopic observations that showed unidentified basidiomycete hyphae within epidermal cells of hair roots of Erica arborea, and the fact that no basidiomycetes were identified when endophytes were cultured from roots of the same plants. This suggestion is in contrast with the general opinion that ericoid mycorrhizal fungi have a good saprophytic ability and are able to grow on common culture media (Leake and Read, 1991). The observation of Bergero et al. (2000) suggests that common culture media may be unable to replicate the in planta conditions that the fungi inhabit and a culturing bias may occur.
Most reports of basidiomycetes as potential symbionts in Ericaceae roots are restricted to northern hemisphere Ericaceae. Allen et al. (1989), however, observed dolipore septa in hyphal coils within the epidermal cells of a root of the Australian Ericaceae, D. 28 secundum, suggesting that at least some basidiomycetes may form ericoid mycorrhizal associations in the southern hemisphere. Despite these observations, the inability to culture basidiomycetes from roots has limited our understanding of the functional roles these fungi may play in ericoid mycorrhizas.
2.1.5 Identification of fungal root endophytes using DNA extracted directly from environmental samples
Based on the observations of unculturable basidiomycetes in Ericaceae roots, it is feasible that ericoid mycorrhizal fungal endophytes that can be cultured from their hair roots represent only a portion of the mycorrhizal community. Molecular analysis of nucleic acids extracted directly from Ericaceae hair roots will prove invaluable for identification of potentially unculturable ericoid mycorrhizal fungal endophytes in planta. Considerable advances have been made in understanding bacterial ecology in recent years as a result of widespread adoption of community profiling techniques that utilise nucleic acids extracted directly from environmental samples (reviewed Ranjard et al., 2000). Similar techniques are now finding favour with fungal ecologists, and are generally based on
PCR amplification with fungus-specific primers, coupled with a separation method such as random cloning, denaturing gradient gel electrophoresis (DGGE), terminal restriction fragment length polymorphism (T-RFLP) analysis or amplified ribosomal DNA restriction analysis (ARDRA) (reviewed Anderson and Cairney, 2004).
The direct DNA extraction methods have been important in confirming the limitations of culturing fungi as a means of assessing diversity in complex substrates such as soil (eg. 29
Viaud et al., 2000; Hunt et al., 2004), along with demonstrating changes in soil fungal communities associated with edaphic conditions, environmental stresses or vegetation type (van Elsas et al., 2000; Lowell and Klein, 2001; Chen and Cairney, 2002; Klamer et al., 2002; Anderson et al. 2003; Brodie et al. 2003; Jumpponen, 2003; Edel-Hermann et al., 2004). The methods have also been variously applied to investigate fungal diversity in plant roots (eg. Kowalchuk et al., 1997; Vandenkoornhuyse et al., 2002a), or a particular root-associated fungal group, such as arbuscular mycorrhizal fungi (eg.
Vandenkoornhuyse et al., 2002b; Johnson et al., 2003).
Random cloning from nucleic acids extracted directly from environmental samples is useful in terms of displaying the taxonomic diversity of a community and has successfully been used for assessing changes in soil fungal diversity (Chen and Cairney,
2002; Anderson et al. 2003; Jumpponen, 2003). However, compared to molecular techniques that produce community fingerprints such as DGGE, it may be less suited as a technique simply because, when analysing large numbers of samples it becomes too expensive, laborious and time consuming to sequence enough clones to obtain a true representation of the taxonomic diversity present (Muyzer et al. 1993). This can, in part, be overcome by using restriction enzyme analysis of cloned PCR products to minimise sequencing and/or constructing species abundance curves to ascertain sufficient numbers of clones to be sequenced in order to have fully sampled the diversity within a sample
(Anderson and Cairney, 2004).
30
In DGGE, the separation of different sequences is achieved by electrophoresis and so the laborious screening necessitated by the redundancy of clones is eliminated. DGGE as a technique for studying fungal diversity may be superior to cloning as it is overall less time consuming and expensive, and provides an immediate display of the members of a fungal community in both a qualitative (band numbers) and semiquantitative (band intensities) way (Muyzer et al., 1993).
In DGGE, PCR-amplified DNA fragments of the same length, but with different sequences, can be separated. Separation occurs due to the decreased electrophoretic mobility of partially melted (denatured) doubled stranded DNA molecules in a polyacrylamide gel containing a linear gradient of denaturants (urea and formamide).
The melting of DNA fragments occurs in melting domains: stretches of base pairs with identical melting temperature. Once a certain point in the denaturing gradient of the gel is reached, a transition from helical to partially melted molecule occurs and migration of the molecule halts. Sequence variation within the melting domains causes melting temperatures to differ and molecules with different sequences will stop migrating at different positions in the gel (Muyzer and Smalla, 1998). By using DGGE, 50% of sequence variants can be detected in DNA fragments up to 500 base pairs, however, this can be increased to 100% by attaching a GC-clamp to the fragments. The clamp acts as a high melting domain to prevent the two strands of DNA from completely disassociating
(Muyzer and Smalla, 1998). It was determined that a bacterial species representing as little as 1% of the population in a soil sample could be detected using DGGE (Muyzer et al., 1993). Furthermore, individual bands in the gel can be excised and sequenced to 31 identify community members and determine phylogenetic relationships (Muyzer and
Smalla, 1998).
Since the first use of DGGE in microbial ecology (Muyzer et al., 1993), most studies using the technique have been concerned with investigating bacterial communities from bulk environmental samples (eg. Felske et al., 1996; Murray et al., 1996; Heuer and
Smaller, 1997). An increasing number of studies using DGGE are now being carried out to investigate community structure of eukaryotic microorganisms, for example, Anderson et al. (2003) investigated changes in soil fungal communities across a morland : forest gradient. However, only two studies have compared the use of DGGE and culturing for assessing fungal communities. DGGE analysis of DNA coding for SSU sequences was used to study fungal infections of Ammophila arenaria L. roots (Kowalchuk et al., 1997).
The level of fungal species richness per root segment, as determined by the number of
DGGE bands, was found to be consistent with that observed in attempts to culture root- infecting fungi on agar plates. However, the sequences of many DGGE recovered samples were not the same as those from the cultured fungi, indicating the presence of some unculturable fungal taxa inhabiting the roots (Kowalchuk et al., 1997).
Vanio and Hantula (2000) compared profiles of fungal species inhabiting Norway spruce stumps using culturing of pure mycelial samples from the wood or DGGE analysis of total fungal DNA extracted directly from wood samples. When analysed using the two techniques, adjacent wood samples displayed different fungal species profiles. This indicated that DGGE could detect fungal taxa that could not be cultured, and 32 demonstrated that the culture media used did not allow the growth of certain fungal species. A limited number of the culturable isolates could not be detected using DGGE and the authors suggested these cultures may have originated from spores or very sparse hyphae in the wood samples and so were not represented in the root samples for direct extraction. The diversity of fungal taxa detectable by DGGE analysis may therefore depend on the relative abundance of each species in a mixed population (Vanio and
Hantula, 2000). These studies indicate that DGGE can produce a different view of fungal diversity in a community to that obtained from culturing. The method can thus be used to complement conventional fungal isolation techniques to obtain a more accurate understanding of diverse communities.
To date, only two investigations have assessed community diversity of ericoid mycorrhizal fungi with techniques that utilise nucleic acids extracted directly from the hair roots of Ericaceae hosts, both of which used random cloning. Berch et al. (2002), when examining the roots of the North American Ericaceae species G. shallon, noted that although field collected hair roots appeared to be heavily colonised, only 1 –10% of root samples yielded culturable isolates. In order to investigate this further, the authors compared partial rDNA large subunit (LSU) sequences from endophytes cultured from roots of G. shallon with sequences cloned from DNA extracted directly from the same root systems. They found that 65% of the root segments examined contained sequences of a basidiomycete that was not found in the cultured assemblage (Berch et al., 2002).
Allen et al. (2003) carried out similar comparisons using partial ITS sequences from
DNA extracted from G. shallon roots and cultured endophytes. Most sequences for 33 cultured endophytes were attributed to ascomycetes, however, while similar ascomycete sequences were cloned from directly extracted DNA, > 50% of the cloned sequences were similar to the basidiomycete genus Sebacina. The absence of Sebacina-like sequences in DNA from the cultured endophyte assemblage led Allen et al. (2003) to conclude that unculturable Sebacina-like basidiomycete endophytes were present in the
G. shallon roots and represented a significant component of the root endophyte assemblage, however the role of these possibly mycorrhizal basidiomycetes remains to be determined.
The aims of the work described in this chapter were thus to (i) test whether the observations of Allen et al. (2003) also apply to endophytes of other Ericaceae taxa from different habitats by obtaining ITS sequences from cultured endophyte assemblages from hair roots of Epacris pulchella (Ericaceae), a common understorey shrub in south-eastern
Australian sclerophyll forests, and compare these with sequences cloned from DNA extracted directly from the root systems, (ii) compare partial ITS sequence profiles from the cultured endophyte assemblages and directly extracted DNA using DGGE, (iii) use the combined data from culturing, cloning and DGGE to examine the taxonomic richness and relative abundance of fungi associated with the roots of E. pulchella and compare combined sequence data to those from fungi associated with the roots of Ericaceae and other plant taxa and (iv) explore the ability of the cultured fungi to form ericoid mycorrhizal or other root associated structures with axenic Ericaceae seedlings.
34
2.2 Experimental Procedures
2.2.1 Collection of plants and isolation of fungal cultures
One 40 - 50 cm juvenile E. pulchella plant (EPA) was collected from a field site at
Lover’s Jump Creek, NSW, Australia (S 33º 35’ E 150º 05’) in March 2002 and two more 40 - 50 cm juvenile plants (EP1 and EP2), growing 20 m apart, were collected from a field site at Ridgecrop, NSW, Australia (S 33º 42’ E 149º 59’) in October 2002. Both sites were relatively dry, had soil derived from Hawksbury sandstone and supported an open dry sclerophyll forest plant community. EPA was investigated using only traditional culturing of root associated endophytes, while EP1 and EP2 were used to directly compare traditional culturing of endophytes with techniques that utilised direct extraction of fungal DNA from the plant roots. Upon collection, each plant (along with the adhering soil) was transferred to the laboratory, where soil was removed by careful rinsing under gently running tap water for 15 min. Any remaining soil or debris was gently removed with forceps. The hair root systems of EP1 and EP2 were divided into seven and six sections (of roughly equal size) respectively while the hair root system of EPA was left undivided as a single group. All root sections were surface sterilised in a 100%
-1 commercial bleach solution (Zixo, 4.5% available chlorine) containing 100 μl l of
Tween 20™ (Amersham Biosciences) for 30 sec, followed by 30 sec in a 70% ethanol solution and three one minute rinses in sterile Milli-Q® water. The sterilised root sections were cut into pieces (ca 5 mm in length). Fungal endophytes were cultured from the roots of EPA, whereas roots from EP1 and EP2 were used to culture endophytes and for direct
DNA extraction. All root pieces from EPA and alternating pieces from the EP1 and EP2 root sections were placed in Petri dishes (9.0 cm diam.) containing 2% malt extract agar 35
(MEA) with streptomycin (15 mg l-1, Sigma), gentamycin (15 mg l-1, Sigma) and tetracycline (12 mg l-1, Sigma) (Pearson and Read, 1973). Eighty root pieces were thus excised and plated from EPA, 70 pieces from EP1 and 65 pieces from EP2. The alternating root pieces from EP1 and EP2 root sections that were not used for endophyte isolation were pooled for subsequent direct DNA extraction.
Petri dishes were incubated in the dark at 23°C and observed daily for emerging hyphae.
Root pieces from which fast growing, rapidly sporulating fungi emerged were discarded, while slower growing fungi were subcultured onto 2% potato dextrose agar (PDA)
(Oxoid). Cultures were maintained in the dark at 21°C and subcultured every 12-16 weeks.
2.2.2 Testing for ericoid mycorrhiza formation
Due to difficulties in obtaining E. pulchella seed, W. pungens was selected as a host plant to use for mycorrhiza formation trials as its seed was readily available, easy to germinate and maintain, and, like E. pulchella, is a member of the Ericaceae sub-family
Styphelioideae. W. Pungens seeds were surface sterilised using a 20% solution of a
-1 commercially available bleach (Zixo, 4.5% available chlorine) containing 100 μl l
Tween 20™ (Amersham Biosciences) for 1 min, followed by two subsequent washes in sterile Milli-Q water. Sterilised seed was then soaked overnight (ca 8 h) in sterile Milli-Q water prior to being germinated on moist sterile filter paper. All seeds were maintained at
22°C during germination (16 h light: 8 h dark cycle). Sterile W. pungens seedlings were transferred to sterile 70 ml specimen containers, each of which contained ca 20 g of an 36 autoclaved growth medium consisting of 95% (dry weight) coarse sand and 5% peat moss. Three ml of sterile Milli-Q water and 0.5 ml of a nutrient solution containing 15.0
-2 -2 -3 µM KH2PO4; 17.0 µM MgSO4; 0.8 µM H3BO3; 0.4 µM MnCl2-4H2O; 0.3 µM ZnSO4-
-4 -5 H20; 0.8 µM CuSO4 and 0.8 µM (NH4)6MO7O24-4H20 was added to each container prior to seedling transfer. The medium composition and nutrient additions were based on the published analysis of the physical and chemical composition of soil from a similar sclerophyll forest site (King and Buckney, 2002).
After five weeks growth at 22°C (16 h light: 8 h dark cycle), root zones of seedlings were inoculated singly with 500 µl of a mycelial slurry of each cultured RFLP-type. To produce the slurry, mycelia were harvested from liquid modified Melin Norkrans (MMN) medium (Marx and Bryan, 1975) after three weeks growth, washed and macerated in sterile Milli-Q water. After 12 weeks hair roots were excised and rinsed to remove excess medium and were cleared and stained following a method modified from McLean and
Lawrie (1996). Root pieces were thus cleared in 10% KOH for 30 min at 90˚C and stained in 1.0% Trypan blue (Sigma) at 90˚C for 90 min. Roots were then destained for
25 min in a 1:1:1 mixture of lactic acid, glycerol and deionised water at room temperature
(Hutton et al., 1994). Mycorrhizal infection was assessed using a Leica DM RBE light microscope and AxioVision 3.0 image capture software.
2.2.3 DNA extraction
DNA was extracted from cultures obtained from all plants using the FastDNA® Kit (Bio
101) following the manufacturer’s instructions. Approximately 200 mg of mycelium was 37 removed from a PDA plate and placed into 2.0 ml screw-cap tubes which included
Lysing Matrix 3 and 1.0 ml of fungal lysing solution (CLS-Y, Bio 101). The tubes were then placed into the FastPrep® Instrument (Bio 101) and homogenised at speed 5.0 m sec-1 for 45 sec. The supernatant was separated by centrifugation at 12,300 RPM for 10 min. Subsequently, 600 μl of Binding Matrix (Bio 101) was added and the tubes were incubated at room temperature for five min, the matrix pelleted by centrifugation and washed in SEWS-M solution (Bio 101). DNA was eluted from the matrix in 80 μl Milli-
Q® water. DNA was extracted from pooled root pieces (from each root section) in the same manner with the following exceptions. The lysing solution used contained 0.4 ml of fungal lysing solution (CLS-Y, Bio 101), 0.4 ml of plant lysing solution (CLS-VF, Bio
101) and 0.2 ml of protein precipitation solution (PPS, Bio 101). The tubes were homogenised at speed 5.0 m sec-1 for 90 sec in the FastPrep® Instrument (Bio 101).
2.2.4 ITS amplification, cloning, RFLP analysis and sequence analysis
The ITS regions of the cultured fungi were amplified in 50 μl reaction volumes, as described by Midgley et al. (2002) using 25 pmol of each of the primers ITS1 and ITS4
(White et al., 1990). Amplifications were performed in a PCR Express thermocycler
(Hybaid) with 35 cycles of 95oC for one min, 50oC for one min and 72oC for one min, followed by a final extension at 72οC for 10 min (Midgley et al., 2002). The ITS regions of fungal endophytes colonising the pooled root pieces from each root section were amplified using the same reaction mixture with the exception of 25 pmol of the ITS1F
(fungal specific) primer in place of the ITS1 primer (Gardes and Bruns, 1993).
Amplifications were performed with the first 13 cycles at 95oC for 35 sec, 55oC for 55 38 sec and 72oC for 45 sec. Cycles 14 – 26 and 27 – 35 used the same parameters except that the extension steps were lengthened to 120 and 180 sec respectively for the next two sets of cycles. The 35 cycles were followed by a final extension at 72οC for 10 min
(Gardes and Bruns, 1993). Reactions were performed in duplicate and a negative control containing no DNA was included in every reaction run. Amplification products were electrophoresed in 2.0 % (w/v) agarose gels, stained with ethidium bromide and visualised under UV light.
ITS products (ca 1.0 μg) from cultured endophytes were digested using five units of the restriction endonucleases Hae III, Hinf I and Rsa I (Promega) for four h at 37 °C. Each digestion was performed in a volume of 15 μl containing five units of restriction endonuclease, 1.0 μl 10X reaction buffer (Promega), 1.0 μg BSA, ca 1.0 μg of amplified
ITS product and the remaining volume in sterile Milli-Q® water. The restriction fragments were separated by electrophoresis on a 3.0% agarose gel and stained and viewed as described above.
For sequencing, ITS-PCR products were cloned with the pGEM-T Easy vector system
(Promega). Representative isolates of each RFLP-type, as determined by different banding patterns, were sequenced using the ABI Big-Dye reaction kit in an ABI 373-A automated fluorescent DNA sequencer (Applied Biosystems, Foster City, CA, USA).
Sequencing reactions were performed with the T7 primer (Promega). ITS sequences were analysed using the FASTA 3.0 program (Pearson and Lipman, 1988) to find the closest sequence matches in the GenBank/EMBL/DDBJ databases. Selected ITS sequences were 39 aligned using the PILEUP program within the EGCG extentions to the Wisconsin
Package, Version 8.1.0 (Rice, 1996) and alignments were optimised first using the
ClustalW option and subsequently manually by visual inspection. Neighbour-joining and parsimony analyses of the patristic distance matrix (1000 bootstraps re-sampling replicates) were then conducted using PAUP (v. 4.01b9) to infer relationships between the fungal endophytes in the present study and with sequences available for other fungi in the GenBank and EMBL nucleotide databases.
ITS-PCR amplification products obtained using direct DNA extraction from hair root samples were cloned with the pGEM-T Easy vector system (Promega), and 102 and 98 clones were randomly selected from the EP1 and EP2 clone libraries respectively.
Transformed ITS products from each individual clone were re-amplified as above using the ITS1 and ITS4 primer pair, and RFLP and sequence analyses performed for each cloned product as described above.
2.2.5 ITS amplification for DGGE analysis
In preparation for DGGE, PCR was conducted using DNA directly extracted from pooled root pieces from each root section obtained from EP1 and EP2, and for each cultured fungus. The reaction mixtures were as described above, however 25 pmol of each of the primers ITS1F-GC and ITS2 were used. The ITS1F-GC primer had a 40 base GC-clamp attached to the 5' end to generate fragments suitable for DGGE analysis. Amplifications were performed on a PTC-100TM programmable thermo-cycler (MJ Research) using the following conditions: initial denaturation cycle at 94ºC for 85 sec, 12 cycles at 95ºC for
35 sec, 50ºC for 55 sec and 72ºC for 45 sec, 13 cycles at 95ºC for 35 sec, 50ºC for 55 sec 40 and 72ºC for 2 min, 9 cycles at 95ºC for 35 sec, 50ºC for 55 sec and 72ºC for 3 min, with a final extension cycle at 72ºC for 10 min (Gardes and Bruns, 1993). Negative controls were included in each PCR run and ITS amplified products were visualised as described above.
2.2.6 DGGE analysis
ITS products generated from pooled root pieces from each root section and combinations of products from cultured endophytes from EP1 and EP2 were separated by DGGE using the DCodeTM Universal Mutation Detection System (BioRad) following the manufacturer's instructions. Polyacrylamide gels (7.5% acrylamide/bisacrylamide
(37.5:5) solution (Amersco)) were prepared with a 25% (40% formamide and 7.0 M urea
(Sigma-Aldrich)) to 50% vertical denaturing gradient using a Gradient Delivery (475) system (BioRad). ITS-PCR products (10 µl) were loaded onto the gels and electrophoresis was performed in 1X TAE buffer (40 mM tris, 20 mM glacial acetic acid,
1.0 mM EDTA) at 75 V for eight h at a constant temperature of 60ºC. Gels were stained by gently shaking for 20 min in 100 ml 1X TAE containing a 1:10000 dilution of SYBR green (Molecular Probes) followed by a further 20 min in 1 x TAE and visualised under
UV light. Digitalised DGGE images were analysed using the Phoretix TM 1D Advanced software (Version 5.2) which performs band matching analyses based upon the prescence/absence and position of bands (Nonlinear Dynamics). Bands on the gels which were present in the pooled root sample direct DNA extraction lanes but absent in the cultured endophyte lanes were excised and reamplified as above with the ITS1 and ITS2 41 primer pair (Gardes and Bruns, 1993). Subsequent products were cloned, sequenced and analysed as described above.
2.3 Results
2.3.1 ITS-RFLP analysis of cultured fungal assemblages and clones from directly extracted DNA
Seventy slow growing, mycelial isolates were cultured from the 80 hair root pieces excised from EPA (Table 2.1). A total of 124 slow growing isolates was cultured from the 135 hair root pieces excised from EP1 and EP2, with 70 and 54 isolates obtained from each respectively (Table 2.1). Ninety-eight and 102 clones were selected for analysis from the clone assemblages generated from DNA extracted from pooled root pieces from each root section for EP1 and EP2 respectively. ITS-PCR amplification produced a single
PCR product (ca 500-700 bp) from each cultured isolate or clone. ITS-RFLP analysis grouped the cultured isolates from EPA, EP1 and EP2 as 11, 15 and 18 RFLP-types respectively, and those from the clone assemblages as 22 and 25 RFLP-types respectively from EP1 and EP2 (Table 2.2 and Table 2.3). Eight RFLP-types were common between the cultured assemblages from both EP1 and EP2, while seven and 10 RFLP-types were unique to EP1 and EP2 respectively, giving a total of 25 cultured RFLP types (Table 2.3 and Figure 2.2).
42
Table 2.1 The number and percentage of root pieces sampled for each of the three Epacris pulchella root systems from the Lover’s Jump Creek and Ridgecrop field site, from which slow growing fungal endophytes were obtained.
Root System Number of hair Number of Percentage (%) of hair root root pieces isolates pieces yielding slow examined obtained growing fungal isolates
E. pulchella (EPA) 80 70 87.5 E. pulchella (EP1) 70 70 100 E. pulchella (EP2) 65 54 83.1
Nine RFLP-types were common between the two cloned assemblages while 13 and 16
were unique to EP1 and EP2 respectively, giving a total of 38 cloned RFLP-types (Table
2.3 and Figure 2.2). Combined data for cultured isolate and clone assemblages from EP1
identified a total of 28 RFLP-types, nine of which were identified from both assemblages,
with six and 13 RFLP-types unique to the cultured isolate and clone assemblages
respectively (Table 2.3 and Figure 2.2). For EP2, 10 RFLP-types were present in both the
cultured isolate and clone assemblages, with seven and 14 RFLP-types unique to each
assemblage respectively (Table 2.3 and Figure 2.2).
The pattern of relative abundance of cultured isolate RFLP-types for EPA was
characterised by a few common and a few less frequently isolated RFLP-types (Figure
2.1). The majority of cultured isolates (ca 77%) from this plant were identified as EP
RFLP-types 1, 50, 52 and 56, with the remaining RFLP-types each comprising a
relatively small proportion of the assemblage (Figure 2.1). Patterns of relative abundance
of cultured isolate RFLP-types and those for the clones were broadly similar for both EP1
and EP2 root systems, with each assemblage characterised by a few common and a 43 greater number of less frequently isolated RFLP-types (Figure 2.2). The majority of cultured isolates (ca 85%) and clones (ca 75%) from EP1 were identified as EP RFLP- types 1, 2, 3, 12 and 13, with the remaining RFLP-types each comprising a relatively small proportion of the assemblages (Figure 2.2). For EP2, the majority of cultured isolates (ca 66%) were EP RFLP-types 2, 3, 15 and 17, with the remaining RFLP-types representing ca 35% of the isolate assemblage. EP RFLP-types 2, 3, 7 and 15 represented
50% of the cloned assemblage from this plant (Figure 2.2). The RFLP-types that were observed only as clones represented ca 11% and ca 17% of the assemblages from EP1 and EP2 respectively (Figure 2.2).
44
35
30
25
20
15
10 Percentage (%) of culture assemblage
5
0
52 50 56 1 57 54 53 49 51 55 58 EP RFLP-type
Figure 2.1 Presence of each EP RFLP-type as a percentage of total cultured isolates obtained from the roots of Epacris pulchella A (EPA), demonstrating that the fungal culture assemblage for this plant was characterised by a few common types and several less frequently isolated RFLP-types. Numbers for EP RFLP-types correspond to those in
Table 2.3. 35 45
30 a
25
20
15
10
5
0 1 13 2 12 3 35 14 21 34 4 15 22 23 26 36 37 38 5 28 39 6 16 24 29 30 32 40 41
EP RFLP-type 30
25 b
20
Percentage (%) of culture/clone assemblages 15
10
5
0 15 3 2 19 23 1 35 7 8 14 26 17 27 9 18 20 42 43 44 45 46 38 48 10 11 25 31 33 37 39 40 47 EP RFLP-type
Figure 2.2 Presence of each EP RFLP-type as a percentage of total cultured isolates (□) and/or clones (■) obtained from roots of two Epacris pulchella plants EP1 (a) and EP2 (b), demonstrating that the fungal culture and clone assemblages for each plant were characterised by a few common types and a larger number of less frequently isolated/cloned RFLP-types. Numbers for EP RFLP-types correspond to those in Table 2.3. 46
Table 2.2 Restriction fragment band sizes obtained by digestion with three restriction
endonucleases (HaeIII, HinfI and RsaI) for the rDNA ITS region amplified from representative
RFLP-types of fungal endophytes isolated from within the root systems of three Epacris pulchella
plants (EPA, EP1 and EP2).
EP RFLP HaeIII † HinfI † RsaI † type 1 300, 78, 60, 48, 28, 8 266, 172, 76, 8 358, 164 49 308, 137, 78 269, 170, 76, 8 359, 164 2 294, 137, 78, 14 266, 170, 79, 8 359, 164 3 384, 62, 48, 29 270, 245, 8 358, 165 4 384, 110, 29 270, 245, 8 358, 165 5 300, 78, 61, 48, 28, 8 269, 172, 74, 8 241, 164, 118 6 276, 78, 61, 48, 28, 24, 8 277, 172, 74 359, 164 7 386, 110, 29 269, 248, 8 358, 167 8 294, 140, 78, 14 266, 90, 81, 81, 8 362, 164 9 294, 138, 78, 14 266, 90, 81, 79, 8 360, 164 10 297, 140, 78, 14 269, 174, 78, 8 365, 164 11 301, 110, 79, 28 244, 149, 103, 14, 8 357, 161 50 275, 79, 79, 62 248, 191, 48, 8 358, 137 12 275, 79, 79, 62 245, 191, 51, 8 358, 137 51 269, 105, 77, 28, 15, 10, 8, 4 240, 140, 128, 8 424, 92 13 270, 77, 74, 31, 28, 19, 10, 8 240, 140, 129, 8 425, 92 14 270, 77, 63, 42, 28, 19, 10, 8 243, 137, 129, 8 425, 92 15 270, 77, 63, 42, 28, 19, 10, 8 240, 140, 129, 8 425, 92 16 235, 133, 78, 14, 11, 8, 4 239, 236, 8 260, 131, 92 52 368, 75, 61, 8, 6 248, 244, 18, 8 367, 96, 55 53 453, 63 256, 252, 8 364, 152 17 369, 76, 62, 8, 6 256, 246, 19 368, 97, 56 18 368, 76, 62, 8, 6 248, 245, 19, 8 261, 162, 97 54 464, 72 254, 159, 115, 8 387, 149 19 289, 139, 75, 6 257, 244, 8 358, 151 20 465, 72 254, 160, 115, 8 388, 149 21 281, 154, 77 258, 246, 8 357, 103, 52 22 280, 157, 77 259, 247, 8 360, 154 23 284, 137, 79, 17 243, 163, 103, 8 356, 161 55 338, 156, 18 234, 146, 124, 8 uncut 24 296, 93, 79, 63, 31, 25 296, 231, 52, 8 462, 99, 26
† Numbers refer to fragment sizes in bp. 47
Table 2.2 cont. Restriction fragment band sizes obtained by digestion with three restriction endonucleases (HaeIII, HinfI and RsaI) for the rDNA ITS region amplified from representative
RFLP-types of fungal endophytes isolated from within the root systems of three Epacris pulchella plants (EPA, EP1 and EP2).
EP RFLP HaeIII † HinfI † RsaI † type 25 290, 145, 72, 18 252, 222, 43, 8 uncut 26 285, 82, 78, 65, 7 257, 252, 8 uncut 27 268, 123, 52, 22, 16, 10, 9 247, 171, 74, 8 uncut 56 270, 77, 63, 43, 28, 19, 10, 8 241, 140, 129, 8 426, 92 28 uncut 303, 247, 48, 8 uncut 29 261, 99, 69, 58, 54, 15, 6 271, 97, 92, 69, 25, 8 uncut 30 uncut 247, 222, 126 551, 44 31 416, 149 273, 192, 92, 8 uncut 32 431, 52, 22, 8 195, 141, 109, 60, 8 410, 57, 46 33 246, 126, 79, 50 389, 112 462, 39 34 284, 74, 68, 54, 28, 22, 14, 5 277, 264, 8 492, 57 57 270, 77, 63, 43, 28, 19, 10, 8 241, 140, 129, 8 426, 92 58 314, 86, 66 199, 156, 65, 38, 8 uncut 35 484, 164 330, 310, 8 622, 26 36 460, 143 255, 227, 91, 22, 8 uncut 37 480, 133 179, 162, 146, 118, 8 uncut 38 uncut 334, 317, 8 641, 18 39 260, 205, 109, 88 349, 305, 8 uncut 40 uncut 322, 218, 102, 8 469, 181 41 uncut 321, 218, 102, 8 uncut 42 470, 142, 12 311, 112, 107, 86, 8 361, 263 43 302, 126, 102, 47 268, 227, 74, 8 uncut 44 uncut 321, 218, 102, 8 uncut 45 465, 94, 41, 29 185, 148, 118, 103, 67, 8 uncut 46 uncut 334, 194, 114 uncut 47 uncut 332, 186, 117, 8 uncut 48 354, 168 320, 117, 85 391, 131
† Numbers refer to fragment sizes in bp.
48
Table 2.3 Putative taxonomic affinities of EP RFLP-types present in cultured isolate assemblages from root systems of three Epacris pulchella plants (EPA, EP1 and EP2) as inferred from FASTA searches of ITS sequences in the GenBank/EMBL/DDBJ databases. EP RFLP-types highlighted in bold are regarded on the basis of inoculation experiments as being ericoid mycorrhizal endophytes.
EP GenBank Closest FASTA match FASTA No. of isolates in each host No. of clones in RFLP Accession expected each host type No. value EPA EP1 EP2 EP1 EP2 1 AY627804 ericoid mycorrhizal endophyte 7.1e-144 7 21 2 33 1 49 AY627805 ericoid mycorrhizal endophyte 2.2e-137 1 0 0 0 0 2 AY627806 ericoid mycorrhizal endophyte 5.2e-137 0 14 5 8 4 3 AY627807 ericoid mycorrhizal endophyte 9.2e-145 0 4 9 9 24 4 AY627808 ericoid mycorrhizal endophyte 1.3e-126 0 1 0 0 0 5 AY627781 ericoid mycorrhizal endophyte 2.0e-140 0 0 0 2 0 6 AY627782 ericoid mycorrhizal endophyte 4.9e-141 0 0 0 1 0 7 AY627809 ericoid mycorrhizal endophyte 1.7e-146 0 0 1 0 4 8 AY627810 ericoid mycorrhizal endophyte 7.4e-132 0 0 1 0 2 9 AY627811 ericoid mycorrhizal endophyte 8.2e-131 0 0 1 0 0 10 AY627783 ericoid mycorrhizal endophyte 1.4e-133 0 0 0 0 1 11 AY627784 ericoid mycorrhizal endophyte 1.5e-120 0 0 0 0 1 50 AY627812 Helotiales 9.1e-113 17 0 0 0 0 12 AY627813 Helotiales 7.6e-118 0 5 0 8 0 51 AY627814 Oidiodendron 1.0e-128 1 0 0 0 0 13 AY627815 Oidiodendron 6.5e-130 0 16 0 18 0 14 AY627816 Oidiodendron 2.7e-125 0 1 1 1 1 15 AY627817 Oidiodendron 9.7e-112 0 1 16 0 19 16 AY627785 Myxotrichaceae 6.2e-100 0 0 0 1 0 52 AY627818 Dermataceae 6.7e-114 22 0 0 0 0 53 AY627819 Dermataceae 5.0e-121 3 0 0 0 0 17 AY627820 Dermataceae 2.1e-123 0 0 6 0 0 18 AY627821 Dermataceae 5.8e-126 0 0 1 0 0 54 AY627822 ericaceae root endophyte 2.9e-96 4 0 0 0 0 19 AY627823 ericaceae root endophyte 1.3e-105 0 0 2 0 3 20 AY627824 ericaceae root endophyte 1.8e-91 0 0 1 0 0 21 AY627825 ericaceae root endophyte 1.1e-117 0 1 0 1 0 22 AY627826 ericaceae root endophyte 8.5e-124 0 1 0 0 0 23 AY627827 Hyaloscyphaceae 6.0e-123 0 1 2 0 2 55 AY627828 Sordariomycete 7.8e-96 1 0 0 0 0 24 AY627786 Sordariomycete 1.9e-116 0 0 0 1 0 25 AY627787 Sordariomycete 6.0e-91 0 0 0 0 1 26 AY627829 Ascomycota 1.2e-115 0 1 1 0 1 27 AY627830 Cladosporium 4.5e-122 0 0 2 0 0 56 AY627831 Hypocreales 1.3e-123 8 0 0 0 0 28 AY627788 Hypocreales 5.5e-145 0 0 0 2 0 29 AY627789 Hypocreales 3.2e-94 0 0 0 1 0 30 AY627790 Hypocreales 9.2e-78 0 0 0 1 0 31 AY627791 Hypocreales 6.9e-148 0 0 0 0 1 32 AY627792 Chaetothyriomycete 4.8e-71 0 0 0 1 0 33 AY627793 Ascomycota 4.0e-50 0 0 0 0 1 34 AY627832 Trichocomaceae 2.7e-123 0 1 0 1 0
49
Table 2.3 cont. Putative taxonomic affinities of EP RFLP-types present in cultured isolate assemblages from root systems of three Epacris pulchella plants (EPA, EP1 and EP2) as inferred from FASTA searches of ITS sequences in the GenBank/EMBL/DDBJ databases. EP RFLP- types highlighted in bold are regarded on the basis of inoculation experiments as being ericoid mycorrhizal endophytes.
EP GenBank Closest FASTA match FASTA No. of isolates in each host No. of clones in RFLP Accession expected each host type No. value EPA EP1 EP2 EP1 EP2 57 AY627833 Agaricales 4.8e-71 5 0 0 0 0 58 AY627834 Tremales 6.1e-77 1 0 0 0 0 35 AY627835 Tricholomataceae 4.8e-153 0 1 1 2 9 36 AY627836 Basidiomycota 7.7e-34 0 1 0 0 0 37 AY627794 Homobasidiomycota 6.4e-121 0 0 0 4 1 38 AY627795 Homobasidiomycota 6.1e-66 0 0 0 3 3 39 AY627796 Homobasidiomycota 1.1e-107 0 0 0 2 1 40 AY627797 Homobasidiomycota 1.5e-83 0 0 0 1 1 41 AY627798 Homobasidiomycota 2.7e-82 0 0 0 1 0 42 AY627837 Basidiomycota 4.6e-70 0 0 1 0 0 43 AY627838 Basidiomycota 5.2e-34 0 0 1 0 0 44 AY627799 Homobasidiomycota 2.7e-82 0 0 0 0 6 45 AY627800 Homobasidiomycota 2.3e-76 0 0 0 0 5 46 AY627801 Basidiomycota 1.3e-54 0 0 0 0 3 47 AY627802 Basidiomycota 2.0e-55 0 0 0 0 1 48 AY627803 Glomus 2.3e-83 0 0 0 0 2
2.3.2 Sequence analysis of ITS RFLP-types
One cultured isolate or clone representing each RFLP-type was selected for ITS
sequencing and comparison with available sequences in the GenBank nucleotide
database. Putative taxonomic affinities were assigned conservatively to RFLP-types
based on FASTA expected values and identities of the closest several sequence matches
obtained from the FASTA searches. Thus, for example, EP RFLP-types 51, 13 and 14
each had multiple matches (with very low expected values) with sequences for
Oidiodendron spp. and have been putatively designated as this genus. In contrast, for EP
RFLP-type 26, the closest several FASTA matches were for various ascomycetes, and it 50 has been putatively designated as Ascomycota (Table 2.3). On this basis EP RFLP-types
1–11 and 49 were designated as putative ericoid mycorrhizal fungal endophytes which belong to a group of currently unidentified Helotiales. EP RFLP-types 12 and 50 along with EP RFLP-types 13-15 and 51 were designated as unidentified Helotiales ascomycetes and Oidiodendron species respectively, groups from which some isolates have been shown to form ericoid mycorrhizas. Most of the remaining isolated fungi and/or clones were designated as a taxonomically diverse group of ascomycetes, some basidiomycetes or, for EP RFLP-type 48, in the clone assemblage from EP2, as a putative
Glomerales fungus. The closest several FASTA matches for three RFLP-types were contradictory and expected values were relatively high, thus no putative taxonomic affinity could be assigned to these (Table 2.3).
2.3.3 Ericoid mycorrhiza formation
Fourteen of the RFLP-types in the cultured isolate assemblage were found to form typical ericoid mycorrhizal coils in epidermal cells of W. pungens and were thus regarded as ericoid mycorrhizal endophytes (Table 2.3 and Figure 2.3). All mycorrhizal RFLP-types colonised many epidermal cells, within which they generally produced dense hyphal coils. When hyphal penetration could be observed, the walls of infected cells were generally only penetrated once, and no intercellular hyphae were observed to pass between adjacent cells in the plant roots (Figure 2.3). The cultured RFLP-types that formed coils were those putatively identified as Oidiodendron sp., Helotiales ascomycetes or those designated as unidentified ericoid mycorrhizal endophytes (Table
2.3). 51
a b
c d
e f
Figure 2.3 Hair roots of Woollsia pungens seedlings showing ericoid mycorrhizal coils in the epidermal cells after twelve weeks in gnotobiotic culture with fungal endophyte RFLP-types cultured from Epacris pulchella hair roots. a = EP RFLP-type 1, b = EP RFLP-type 49, c = EP RFLP-type 2, d = EP RFLP-type 3, e = EP RFLP-type 4 and f = EP RFLP-type 7. Bars represent 25 µm. 52
g h
i j
k l
Figure 2.3 cont. Hair roots of Woollsia pungens seedlings showing ericoid mycorrhizal coils in the epidermal cells after twelve weeks in gnotobiotic culture with fungal endophyte RFLP-types cultured from Epacris pulchella hair roots. g = EP RFLP-type 8, h = EP RFLP-type 9, i = EP RFLP-type 50, j = EP RFLP-type 12, k = EP RFLP-type 51 and l = EP RFLP-type 13. Bars represent 25 µm. 53
m n
Figure 2.3 cont. Hair roots of Woollsia pungens seedlings showing ericoid mycorrhizal coils in the epidermal cells after twelve weeks in gnotobiotic culture with fungal endophyte RFLP-types cultured from Epacris pulchella hair roots. m = EP RFLP-type 14 and n = EP RFLP-type 15. Bars represent 25 µm.
2.3.4 Phylogenetic analysis of ITS RFLP-types
Oidiodendron species, particularly O. maius, are regarded as common ericoid mycorrhizal endophytes of many northern hemisphere Ericaceae taxa, however, to date the only evidence to suggest that Oidiodendron species may form ericoid mycorrhizas with Australian Ericaceae was the isolation from W. pungens of a single endophyte having sequence similarity to O. maius (Chambers et al., 2000). Interestingly, the identities of several of the closest FASTA sequence matches to the mycorrhizal EP
RFLP-types 51, 13, 14 and 15 were O. maius. Furthermore, EP RFLP-types 13 and 15 represented one of the most frequently isolated RFLP-types from EP1 and EP2 respectively. On this basis, the taxonomic positions of EP RFLP-types 51, 13, 14 and 15 were investigated further using neighbour-joining and parsimony analyses. Sequences used in the analyses also included EP RFLP 16, which had several closest matches to
Oidiodendron sp., RL RFLP 31 from a fungal endophyte isolated from Rhododendron lochiae (Chapter 4), which had several closest matches to O. maius, and also formed 54 typical ericoid mycorrhizal coils during inoculation trials. The analysis also included selected ITS sequences representing Oidiodendron species from the
GenBank/EMBL/DDBJ databases.
The neighbour-joining analysis of the Oidiodendron sequences (Figure 2.4) placed the sequences into four groups. Within the strongly supported (100% bootstrap support) group I, all sequences for O. maius and O. citrinum along with RL RFLP 31 clustered in a strongly supported (100%) sub-group while EP RFLP 51 separated on a terminal branch. Also within group I, EP RFLP-types 13–15 clustered within a strongly supported
(97%) sub-group along with the sequence for the O. maius–like endophyte previously isolated from W. pungens. The poorly supported (53%) group II and strongly supported
(100%) group IV contained a variety of Oidiodendron sequences. EP RFLP 16 clustered separately with O. rhodogenum in the moderately supported (72%) group III. The parsimony analysis produced similar results (Figure 2.5) whereby the strongly supported
(100%) group I comprised all sequences for O. maius and O. citrinum along with RL
RFLP 31 in a strongly supported (100%) sub-group. EP RFLP 51 clustered with EP
RFLP-types 13–15 along with the sequence for the O. maius–like endophyte previously isolated from W. pungens within a strongly supported (97%) sub-group. The strongly supported (100%) group II, moderately supported (61%) group IV and strongly supported
(100%) group V contained a variety of Oidiodendron species sequences. Rather than grouping with O. rhodogenum, as was the case in the neighbour-joining analysis, EP
RFLP 16 clustered separately with O. cerialis and Oidiodendron echinulatum Barron in the strongly supported (100%) group III. 55
EP RFLP 13 (AY627815) 61 EP RFLP 15 (AY627817) 97 Ericaceae root endophyte (AF072294) EP RFLP 14 (AY627816) 81 Oidiodendron citrinum (AF062790) 57 Oidiodendron citrinum (AF307762) Oidiodendron citrinum (AF307761) 54 Oidiodendron maius (AF062798) Group I 63 Oidiodendron maius (AF062799) Oidiodendron maius (AF307766) Oidiodendron maius (AF062800) 100 Oidiodendron maius (AF062801) Oidiodendron maius (AF307767) 100 Oidiodendron maius (AF307768) Oidiodendron maius (AF307769) Oidiodendron maius (AF307770) 98 Oidiodendron maius (AF307771) Oidiodendron maius (AF307772) RL RFLP 31 (AY699655) EP RFLP 51 (AY627814) Oidiodendron pilicola (AF062787) Oidiodendron cerealis (AF062788) Oidiodendron echinulatum (AF062791) 84 Oidiodendron flavum (AF062792) Oidiodendron flavum (AF307763) 63 Oidiodendron griseum (AF062797) 71 88 Oidiodendron griseum (AF307764) 100 Oidiodendron tenuissimum (AF307774) Oidiodendron griseum (AF062793) 56 Oidiodendron griseum (AF062795) 84 91Oidiodendron griseum (AF062796) Group II Oidiodendron griseum (AF062794) 53 Oidiodendron setiferum (AF062805) 85 Oidiodendron tenuissimum (AF062807) 90 100 96 Oidiodendron tenuissimum (AF307773) 75 Oidiodendron tenuissimum (AF062808) Oidiodendron griseum (AF307765) 100 Oidiodendron truncatum (AF062809) Oidiodendron truncatum (AF307775) Oidiodendron periconioides (AF062802) Group III 72 Oidiodendron rhodogenum (AF062803) EP RFLP 16 (AY627785) Group IV 100 Oidiodendron chlamydosporicum (AF062789) Oidiodendron scytaloides (AF062804) Pseudogymnoascus roseus (AF062819) Geomyces pannorum (AF307760) 0.01 substitutions/site
Figure 2.4 Neighbour-joining tree showing the relationships between the ITS sequence for EP RFLP-types 13 – 16 and 51 from Epacris pulchella root systems, RL RFLP-type 31 from Rhododendron lochiae (Chapter 4) and selected sequences for Oidiodendron spp. from the GenBank/EMBL/DDBJ databases. Numerical values indicate bootstrap percentiles from 1000 replicates. 56
EP RFLP 13 (AY627815) 67 EP RFLP 14 (AY627816) 100 EP RFLP 15 (AY627817) 59 Ericaceae root endophyte (AF072294) EP RFLP 51 (AY627814) Oidiodendron citrinum (AF062790) 100 Oidiodendron citrinum (AF307761) 51 Oidiodendron citrinum (AF307762) Group I 100 Oidiodendron maius (AF062800) 67 83 Oidiodendron maius (AF307771) Oidiodendron maius (AF307772) RL RFLP 31 (AY699655) Oidiodendron maius (AF062798) 97 Oidiodendron maius (AF062799) 56 100 Oidiodendron maius (AF307766) Oidiodendron maius (AF062801) Oidiodendron maius (AF307767) Oidiodendron maius (AF307768) Oidiodendron maius (AF307769) Oidiodendron maius (AF307770) Oidiodendron periconioides (AF062802) Group II 100 Oidiodendron pilicola (AF062787) Oidiodendron rhodogenum (AF062803) 67 Oidiodendron cerealis (AF062788) Group III 100 EP RFLP 16 (AY627785) 100 Oidiodendron echinulatum (AF062791) 100 Oidiodendron flavum (AF062792) Oidiodendron flavum (AF307763) 63 Oidiodendron griseum (AF062793) Oidiodendron griseum (AF062794) 100 100 Oidiodendron griseum (AF062795) Oidiodendron griseum (AF062796) Oidiodendron griseum (AF062797) Group IV 100 Oidiodendron griseum (AF307764) Oidiodendron tenuissimum (AF307774) 61 Oidiodendron setiferum (AF062805) 60 Oidiodendron tenuissimum (AF062807) Oidiodendron tenuissimum (AF062808) Oidiodendron tenuissimum (AF307773) 100 Oidiodendron truncatum (AF062809) Oidiodendron truncatum (AF307775) Oidiodendron griseum (AF307765) 100 Oidiodendron chlamydosporicum (AF062789) Group V Oidiodendron scytaloides (AF062804) Pseudogymnoascus roseus (AF062819) Geomyces pannorum (AF307760) 5 changes
Figure 2.5 Parsimony tree showing the relationships between the ITS sequence for EP RFLP- types 13 – 16 and 51 from Epacris pulchella root systems, RL RFLP-type 31 from Rhododendron lochiae (Chapter 4) and selected sequences for Oidiodendron spp. from the GenBank/EMBL/DDBJ databases. Numerical values indicate bootstrap percentiles from 1000 replicates. 57
2.3.5 DGGE analysis
PCR amplification of the ITS1 regions in DNA from pooled root pieces from each root section and representative RFLP-types of the fungi isolated from EP1 and EP2 yielded products of the expected size (200-300 bp) (data not shown). The DGGE banding patterns for DNA from pooled root pieces from each root section from EP1 comprised a total of 22 bands, 10 of which aligned with bands from the cultured fungi from that plant
(Figure 2.6). Similarly, 12 of the 23 bands in the DGGE profiles from EP2 aligned with bands for cultured fungi (Figure 2.6). Several bands from DNA extracted directly from root pieces that were considered to represent the same bands from the cultured assemblages were sequenced to confirm that these represented the same sequence. These sequences were found to be identical, validating the assumption that bands migrating to the same position represented the same DNA sequence (data not shown). For both plants, the most intense bands observed in DNA extracted directly from pooled root pieces aligned with bands that represented the most frequently isolated RFLP-types in the culture assemblages. For each plant, only two bands (representing EP RFLP-types 6 and
11 in EP1, and EP RFLP-types 8 and 18 in EP2) were observed in DGGE profiles for cultured fungi, but not in DNA extracted directly from pooled root pieces (Figure 2.6).
All bands in DGGE profiles of DNA from pooled root pieces that did not align with those for cultured fungi were excised for PCR amplification and sequencing. Despite repeated attempts, DNA from six faint bands (four from EP1 and two from EP2) could not be amplified, however ITS1 sequences were obtained for the remaining 17 bands. Of the 58
eight bands sequenced from EP1, three were designated (using the criteria described
above) as putative ericoid mycorrhizal endophytes, two as Glomerales taxa and two as
ascomycetes, while one was unidentified (Table 2.4). For EP2, two were designated as
putative ericoid endophytes, one as an ascomycete and six as probable basidiomycetes
(Table 2.4).
DNA extracted from roots Cultured isolates DNA extracted from roots Cultured isolates 1 2 3 4 5 6 7 1 2 3 1 2 3 4 5 6 1 2 3 4
* 9 *
7 6 * 7 * 2
3 * 1 * 4 * 2 1 5 5 3 * 6
8 4 8 a b
Figure 2.6 DGGE profiles of ITS1 sequences amplified from DNA extracted directly from roots or cultured fungal isolates from Epacris pulchella plants EP1 (a) and EP2 (b). Each lane for DNA extracted from roots represents DNA from pooled root pieces from a root section, while each cultured isolate lane represents mixed ITS1 PCR products from several cultured isolates. Numbered bands are unique bands in profiles for DNA extracted from roots that were sequenced (numbers correspond to those in Table 2.3). Bands that were present in cultured isolate profiles, but absent from those for DNA extracted from roots are marked with an asterisk. 59
Table 2.4 Sequence information from excised DGGE bands for EP1 and EP2. The closest FASTA match represents the most accurate taxonomic match between our band sequences and those in the NCBI database. The FASTA expected value represents the number of sequence matches expected by random chance.
Band GenBank Closest FASTA match FASTA Band present in Number Accession No. expected host sample value EP1 EP2 1 AY669139 Helotiales 7.8e-42 + 2 AY669140 ericoid mycorrhizal endophyte 4.4e-55 + 3 AY669137 ericoid mycorrhizal endophyte 2.1e-58 + 4 AY669142 ericoid mycorrhizal endophyte 8.2e-57 + 5 AY669138 Glomerales 3.7e-31 + 6 AY669144 Glomerales 9.0e-25 + 7 AY669143 Ascomycota 1.4e-57 + 8 AY669141 Basidiomycota 2.0e-15 + 9 AY669128 ericoid mycorrhizal endophyte 7.0e-53 + 10 AY669133 Oidiodendron 2.0e-49 + 11 AY669129 Helotiales 7.0e-29 + 12 AY669132 Basidiomycota 2.0e-55 + 13 AY669130 Basidiomycota 4.5e-29 + 14 AY669136 Urediniomycetes 5.7e-17 + 15 AY669131 Basidiomycota 1.6e-19 + 16 AY669134 Basidiomycota 3.6e-17 + 17 AY669135 Basidiomycota 4.2e-20 +
2.4 Discussion
ITS-RFLP analysis indicated that the cultured isolate assemblage from EPA comprised
mostly unique and far fewer (ca 30-40%) RFLP-types (= putative taxa) than E. pulchella
plants EP1 and EP2. This may reflect differences in edaphic conditions between the two
field sites that the plants were collected from. EPA was collected in March (Autumn),
while EP1 and EP2 were collected in October (Spring). Seasonal differences affect both
the length and number of hair roots present and the level of ericoid mycorrhizal
colonisation for W. pungens, from eastern Australia (Kemp et al., 2003). Thus, as an 60 alternative, seasonal differences of collection could possibly result in the variations in fungal diversity associated with E. pulchella roots seen in the present study.
The cultured isolate assemblages from EP1 and EP2 comprised similar numbers of
RFLP-types (= putative taxa), although some RFLP-types were unique to each plant.
Clone assemblages from EP1 and EP2 also comprised similar numbers of RFLP-types with some unique to each plant, however for each plant the cloned assemblage contained seven more RFLP-types than the cultured isolate assemblage. In this respect, data from the present study are consistent with other investigations that have reported greater taxonomic richness in assemblages of fungi in soil or similar environments when determined by direct DNA extraction compared to cultured isolates (eg. Borneman and
Hartin, 2000; Viaud et al., 2000; Hunt et al., 2004).
When data for the two E. pulchella plants, EP1 and EP2, were combined, 15 RFLP-types were common between the cultured isolate and clone assemblages, with 10 observed exclusively in the cultured isolate assemblage and 23 only in the clone assemblage. Those
RFLP-types that were represented by the largest numbers of cultured isolates (EP RFLP- types 1, 2, 3, 12, 13 and 15) were also the most abundant in the clone assemblages from at least one plant. These, along with most other RFLP-types that were common between the cultured isolate and clone assemblages, were putatively identified as probable ascomycete ericoid mycorrhizal endophytes, suggesting that both methods provided broadly similar information regarding the most abundant ericoid mycorrhizal endophytes of E. pulchella. Of particular note amongst these were EP RFLP-types 13 and 15, both of which formed ericoid mycorrhizas with W. pungens, and which ITS sequence data 61 suggested are likely to represent Oidiodendron species. This genus has been reported widely as a mycorrhizal endophyte of northern hemisphere Ericaceae (Rice and Currah,
2001), but only a single Oidiodendron-like isolate has previously been obtained from roots of an Australian Ericaceae taxon (W. pungens) (Chambers et al., 2000). Data from the present investigation suggest that Oidiodendron-like fungi commonly occupy roots of
E. pulchella at the Ridgecrop field site and are likely to be important mycorrhizal endophytes. A single Oidiodendron-like isolate (EP RFLP-type 51) was obtained from the roots of EPA at the Lover’s Jump Creek site. Both neighbour-joining and parsimony analyses placed these Oidiodendron-like isolates from E. pulchella in a group which included O. maius and O. citrinum sequences, however they fell within a separate well- supported sister group which indicates that the isolates from this study probably represent a different Oidiodendron species. Ascertaining whether these ericoid mycorrhizal
Oidiodendron-like isolates have a preference for this plant host or different edaphic conditions at the field sites in the present and previous studies will require further investigation. RL RFLP-type 31 from R. lochiae, which formed ericoid mycorrhizal structures in hair roots of Vaccinium. macrocarpon Ait. (see Chapter 4), was grouped strongly with sequences for O. maius and O. citrinum isolates. These two taxa are regarded as conspecific (Hambleton et al., 1998), and it appears that EP RFLP type 31 is also conspecific with these taxa.
Fourteen RFLP-types from the combined cultured assemblage formed typical ericoid mycorrhizal structures in gnotobiotic culture experiments, and these RFLP-types encompassed a substantial proportion of the total isolate assemblage from each plant, 62 particularly EP1 (ca 93%) and EP2 (ca 72%). All cultured RFLP-types that ITS sequence comparisons suggested to be putative ericoid mycorrhizal endophytes successfully formed ericoid mycorrhizas during the inoculation trials. Furthermore, cloned RFLP-types that were designated as putative ericoid mycorrhizal endophytes represented ca 80% (EP1) and ca 61% (EP2) of RFLP-types in the clone assemblages, indicating that cloning and culturing give similar indication of the ericoid mycorrhizal fungal community in at least this host species
The percentage of cultured endophytes from EP1 and EP2 that were mycorrhizal is comparable to that seen for two other epacrids sampled in the Sydney region, W. pungens and L. parviflorus (Midgley et al., 2004c). The lower percentage of mycorrhizal isolates obtained from EPA may be explained by variations in edaphic conditions between sites or seasonality of sampling as discussed above. The isolates shown to be mycorrhizal in this study, that sequence comparison suggested to belong to a group of currently unidentified
Helotiales, all appear to be closely related to fungal taxa that have been previously shown to form ericoid mycorrhizas with Australian Ericaceae. However, this is the first report of
Oidiodendron spp. being frequently observed members of the ericoid mycorrhizal fungal community in the roots of an Australian Ericaceae species.
Although some Australian Ericaceae are difficult to germinate and maintain in axenic culture, the peat/sand medium utilised in the current study was successfully utilised to maintain W. pungens seedlings in gnotobiotic conditions and to test for mycorrhizal ability of a number of endophyte taxa. Furthermore, the medium, which replicates to 63 some degree the soils from the Sydney region often inhabited by Australian Ericaceae species, is likely to provide a more ecologically realistic culture environment, in terms of assessing endophyte mycorrhizal status, than homogeneous agar- based media that have been previously employed. Such agar-based media have previously resulted in restricted hair root formation in Australian Ericaceae when used for inoculation trials (Liu et al.,
1998), whereas the heterogenous peat/sand medium used in this study resulted in healthy hair root formation and growth. These results suggest that the mycorrhizal status of endophytes from other Australian Ericaceae that have more demanding culture requirements could be tested by inoculation onto W. pungens in the 'model' microcosm that was employed in this study.
Several of the RFLP-types that were unique to either the cultured isolate or clone assemblages from E. pulchella were identified as putative ascomycete ericoid mycorrhizal endophytes. While only a single isolate of one basidiomycete RFLP-type (a putative Tricholomataceae taxon) was obtained from each of EP1 and EP2, multiple clones of this RFLP-type, along with several other putative basidiomycete RFLP-types
(totalling < 23% of the cloned assemblages) were observed. In a similar investigation of fungi associated with roots of G. shallon (Ericaceae) on Vancouver Island, Canada, Allen et al. (2003) found that, although only 5% of cultured isolates were basidiomycetes, 59% of sequences cloned from DNA extracted directly from the same root systems had strong sequence identity to the basidiomycete genus Sebacina. In contrast to the present study, there was thus a clear difference in the most commonly encountered taxa in the cultured isolate and clone assemblages. Some Sebacina taxa are known to form mycorrhizal associations with other plant taxa (Selosse et al., 2002), thus it is possible that the 64
Sebacina-like fungi represent unculturable ericoid mycorrhizal endophytes of G. shallon.
This suggestion is supported by the fact that, despite containing apparent mycorrhizal infection when viewed under the microscope, 74% of root pieces yielded no culturable fungi (Allen et al., 2003). This contrasts strongly with the ca 10% of root pieces that failed to yield culturable fungi in the present study. It is difficult to predict the functional status of the putative basidiomycetes identified in DNA cloned from E. pulchella roots.
While basidiomycete hyphae have occasionally been observed in epidermal cells of the hair roots of Australian Ericaceae (Allen et al., 1989), there is no evidence that these form ericoid mycorrhizal associations (Cairney and Ashford, 2002). On current evidence, these basidiomycetes seem more likely to be saprotrophs, and their absence from the cultured isolate assemblage may simply reflect an inability to grow on the medium used or their presence in or on roots as inactive propagules.
Approximately half of the DGGE bands in DNA extracted directly from E. pulchella roots matched those from cultured RFLP-types. Significantly, the most intense bands in the directly extracted DNA profiles matched those of the most commonly isolated RFLP- types (ericoid mycorrhizal endophytes). Since band intensity in DGGE gels can provide a rough estimate of relative abundance (Prosser, 2002), the DGGE data confirmed that these fungi were an important component of the E. pulchella root fungal assemblage.
Sequences obtained for DGGE bands that were observed only in the directly extracted
DNA profiles indicated that several of these probably also represented ericoid mycorrhizal endophytes, while several others were putatively identified as 65 basidiomycetes. The DGGE data were thus broadly supportive of those obtained by cloning from DNA extracted directly from E. pulchella roots.
As was the case with the cloned assemblage, several RFLP-types from the cultured isolate assemblage were not observed in DGGE profiles of the DNA extracted from E. pulchella roots. With the exception of EP RFLP-type 17, of which six isolates were cultured from EP2, only one or two isolates were obtained for these RFLP-types.
Similarly, other than six putative basidiomycete RFLP-types for which three to six clones were obtained, cloned RFLP-types that were absent from the cultured isolate assemblage were present as only one or two clones. Many of the DGGE bands that were absent from profiles from cultured isolates were relatively faint, suggesting that the relative abundance of the taxa they represent was probably low.
Sequences for two clones from EP2 and two DGGE bands from EP1 were identified as putative Glomerales taxa. These fungi form arbuscular mycorrhizas with a range of plant taxa and, while the presence of Glomerales-like hyphae in roots of Ericaceae has occasionally been reported (McGee, 1986; McLean and Lawrie, 1996; Reed, 1996), this appears likely to represent opportunistic, non-symbiotic colonisation rather than a mycorrhizal association (Leake and Read, 1991). Sequences for three cloned RFLP-types and one DGGE band from the DNA extracted from E. pulchella roots had low identity with database sequences. Assessment of these sequences in terms of the location of the nucleotide matches with closest matches from FASTA searches indicates that these probably do not represent chimeric sequences and are more likely to represent unknown 66 fungal groups. Sequences that have low similarity to database fungal sequences, and that may represent hitherto unknown fungal groups, have been obtained from DNA extracted directly from roots of other plant taxa (Vandenkoornhuyse et al. 2002a).
The potential bias associated with culture-based assessments of fungal diversity in environmental samples is widely accepted (Zak and Visser, 1996; Bridge and Spooner,
2001). There is, however, also a possibility that PCR-based molecular methods may introduce a degree of bias when amplifying samples inhabited by a taxonomically diverse community, as in the case of Ericaceae hair roots. PCR relies on target sequences binding to an excess of primers, however, competition may occur with samples containing mixed DNA. PCR may thus tend to preferentially amplify DNA from the most prevalent species (Bridge and Spooner 2001). In terms of the present study, this
PCR bias may have led to a reduction in detection of taxonomic diversity, based on the number of cloned RFLP-types or DGGE bands, relative to the actual diversity of fungal endophytes in E. pulchella roots.
The work described in this chapter combined both isolation and direct DNA extraction methods to investigate fungi associated with roots of E. pulchella. The data suggest that, for this taxon at the Ridgecrop field site, culturing and direct DNA extraction, coupled with either cloning or DGGE, were equally useful in identifying the most abundant endophyte taxa. On the basis of ITS sequence similarities and inoculation experiments, these endophyte taxa appear to be ericoid mycorrhizal fungi, implying that previous studies that have relied upon culture-based assessment of ericoid mycorrhizal endophytes 67 of Australian Ericaceae (eg. Chambers et al., 2000; Midgley et al., 2002; 2004c) have probably identified the major components of the endophyte assemblages. Each method, however, also identified a unique suite of less abundant fungi and, while more extensive sampling with a single method may have revealed more taxa, this emphasises the efficacy of using multiple approaches in investigations of this nature.
68
CHAPTER 3
Chitinolytic activities of ericoid mycorrhizal and other root-associated fungi from
Epacris pulchella
3.1 Introduction
3.1.1 Edaphic stresses in habitats of Ericaceae and their associated ericoid mycorrhizal fungi
Plants in the family Ericaceae generally inhabit heathland or forest habitats where soils are acidic and extremely nutrient deficient (Read, 1996; Cairney and Meharg, 2003). The northern hemisphere mor-humus heathlands have soil consisting primarily of decomposing organic material, and are characterised by high quantities of phenolic compounds, low pH and frequently anaerobic conditions (Smith and Read, 1997). These factors, often accompanied by relatively cool temperatures, hinder ammonification and moreover nitrification which results in soils being characteristically low in available inorganic nitrogen (Abuarghub and Read, 1988).
Australian heathlands generally occur on dry sandy soils with, in contrast, only a small
(5-10%) organic fraction and a large (ca 85%) component of silica (King and Buckney,
2002). Many Australian Ericaceae are common in acidic heathland and forest soils derived from nutrientpoor Hawkesbury sandstone (Read, 1991; Hutton et al., 1994;
McLean and Lawrie, 1996; Midgley et al., 2004a). Rates of nitrogen mineralisation in such soils are relatively low. Indeed rates of nitrogen immobilisation frequently equal or 69 exceed rates of mineralisation and the soils are regarded as nitrogen deficient (Adams and
Attiwill, 1986a, 1986b; King and Buckney, 2002). The low levels of nitrogen mineralisation in most heathland ecosystems worldwide has resulted in soil environments in which most of the nitrogen is present in various organic forms such as free amino acids, proteins or protein-phenol complexes (Abuarghub and Read, 1988; King and
Buckney, 2002). For example, in an Australian wet heathland (wallum) soil the two most abundant sources of nitrogen were found to be soluble protein and amino acids (Schmidt and Stewart, 1997). Other than information on wallum heathlands, there does not appear to be detailed information regarding the availability, persistence and composition of these organic nitrogen sources within other Australian heathland or forest soils. It is widely believed that the abundance and success of ericaceous plants worldwide in nutrient poor environments is primarily due to benefits, in terms of access to organic nitrogen and phosphorus stores, conferred to the host plant by mycorrhizal fungi (Read, 1996; Perotto et al., 2002).
3.1.2 Abilities of ericoid mycorrhizal fungi to utilise organic nitrogen sources
The mutualistic nature of the ericoid mycorrhizal symbiosis of Ericaceae was established by demonstrating carbon movement from a host to a root endophyte (Stribley and Read,
1974) and the demonstration that there was an endophyte-mediated enhancement of plant nitrogen and phosphorus nutrition via utilisation of organic sources of the nutrients in the growing media (Stribley and Read, 1980; Bajwa and Read, 1985; Myers and Leake,
1996; Xiao and Berch, 1999). The mycorrhizal nature of fungal endophytes in Australian
Ericaceae hair roots is yet to be confirmed by physiological investigations. To date, 70 mycorrhizal ability has been assumed on the basis that isolated endophytes can form typical ericoid mycorrhiza-like coils in Australian Ericaceae hair root cells when grown in gnotobiotic culture as well as being able to utilise organic nitrogen sources in axenic culture (Liu et al., 1998; McLean et al., 1998; Anthony et al., 2000; Midgley et al.,
2004a). Schmidt and Stewart (1999) showed that 15N-labelled glycine was incorporated by, presumably mycorrhizal, Australian Ericaceae, indicating a possible ability to use amino acids as a nitrogen source. Further investigation considered uptake of organic 15N- labelled plant debris by the same Australian Ericaceae, however due to similar levels of nitrogen accumulation by mycorrhizal and non-mycorrhizal plants it was suggested that mineralisation of the nitrogen may have occurred in the soil prior to acquisition (Bell and
Pate, 1996). As a result it could not be inferred from the data that ericoid mycorrhizal endophytes assist Australian Ericaceae host plants in the uptake of organic nitrogen sources.
Hymenoscyphus ericae, from the Northern hemisphere, has been shown in a number studies to utilise a wide range of nitrogen sources for growth (Stribley and Read, 1980;
+ Bajwa and Read, 1986; Chen et al., 1999). Inorganic nitrogen, in the forms of NH4 or
- NO3 , are both readily assimilated by the fungus (Bajwa and Read, 1986). In addition, H. ericae can also produce significant biomass on a range of neutral, acidic and basic amino acids and simple proteins (Stribley and Read, 1980; Bajwa and Read, 1986; Chen et al.,
1999). Similarly, a small number of other Helotiales-like ericoid mycorrhizal fungi from the northern hemisphere have also been shown to utilise a range of inorganic and organic nitrogen sources (Xiao and Berch, 1999). 71
The ability of endophytes isolated from the roots of the Australian Ericaceae species
Woollsia pungens to utilise inorganic and organic nitrogen sources in axenic culture has also been investigated (Chen et al., 1999; Whittaker and Cairney, 2001; Midgley et al.,
2004a). While the endophytes were found to utilise inorganic nitrogen sources, they also showed significant growth on acidic, basic and neutral amino acids, along with the protein bovine serum albumin (BSA), as sole nitrogen sources. For most W. pungens endophyte isolates, biomass yields were generally similar to a H. ericae isolate from the northern hemisphere, suggesting that endophytes from Australian Ericaceae hosts and those from other Ericaceae hosts have similar abilities to utilise organic forms of nitrogen
(Chen et al., 1999; Whittaker and Cairney, 2001).
The use of a soluble protein (BSA) as a source of nitrogen by ericoid mycorrhizal fungi implies the production of extracellular proteases, and H. ericae is known to produce carboxyl proteinase activity during growth in axenic culture (Leake and Read, 1989a;
Bajwa and Read, 1985). In axenic culture experiments this proteinase hydrolyses soluble proteins, releasing amino acids which are subsequently absorbed into the mycelium. H. ericae proteinase activities are optimal at pH 2.0 - 3.0, and it is widely assumed that this is an adaptation of the enzyme to the low pH of mor-humic soils of the northern hemisphere (Leake and Read, 1990a). Growth of ericoid mycorrhizal fungi from
Australian Ericaceae on media containing simple proteins such as BSA as a sole nitrogen source suggests that they also produce proteolytic enzymes (Chen et al., 1999; Midgley et al., 2004a). 72
Ericaceae are often associated with soil that contains high concentrations of phenolic compounds. These phenolic chemicals, such as phenolic acids, polyphenols including tannins, quinones, humic and fulvic acid, are derived partially from ericaceous leaf litter that is rich in phenolic compounds, and also from the partial degradation of lignin
(Bending and Read, 1996a; Rovira and Vallego, 2002). Polyphenols in soil are known to inhibit decomposition of cellular debris (Leake and Read, 1989b). The role of polyphenols in inhibition of decomposition is presumed to reflect the chemical abilities of these compounds to bind amino acids, peptides, proteins and other macromolecules, such as chitin or nucleic acids, into insoluble complexes (Leake and Read, 1989b; Rovira and
Vallego, 2002; Bending and Read, 1996a, 1996b; Bending and Read, 1997). H. ericae is known to produce polyphenol oxidase activities (Bending and Read, 1996a). When H. ericae is grown on soluble tannic acid, dark pigmented compounds are released and production of these, presumably phenolic, compounds has been correlated with removal of tannic acid from the growth media and presence of catechol oxidase activity (Bending and Read, 1996b). The enzyme is thought to reduce inhibitory protein binding and facilitate fungal access to tannin-bound proteins that frequently occur in the mor-humus habitats of H. ericae (Bending and Read, 1996b).
3.1.3 Abilities of ericoid mycorrhizal fungi to produce cell wall-degrading enzymes
In the mor humus habitat, many of the organic nitrogen and phosphorus substrates that are readily utilised by mycorrhizal endophytes are either chemically or physically complexed with moribund plant cell wall material (Cairney and Burke, 1998). Virtually all that is known about plant cell wall degrading enzymes in ericoid mycorrhizal 73 endophytes is based on investigations of H. ericae isolates. It has become clear that H. ericae produces a suite of cell wall-degrading enzymes that have previously been regarded as the domain of saprophytic fungi (Cairney and Burke, 1997). The enzymes probably contribute to the endophyte-host infection process as well as allowing the endophyte to utilise the complexed nutrient sources, such as proteins, which are associated with or internal to the plant cell wall. Production of these enzymes implies a direct role of the fungus in decomposition and nutrient cycling processes in mor-humus soils (Cairney and Burke, 1998). Burke and Cairney (1997) suggest that production of these enzymes may also allow H. ericae to supplement its carbon budget during symbiosis by accessing carbon from organic sources in the soil. H. ericae is considered to have some degree of saprophytic ability, as in vitro growth was demonstrated on cellulose-based substances such as filter paper (Varma and Bonfante, 1994). The extent of its saprophytic nature in its native mor-humus soil remains unclear, however growth on sterilised Calluna root material indicates that saprotrophic growth and persistence within dead plant material may be possible (Varma and Bonfante, 1994).
Cellulose and hemicelluloses in the plant cell wall are primarily degraded by hydrolytic cellulase and hemicellulase complexes, while lignin degradation is mediated by various ligninases (Kuhad et al., 1997). The cellulase enzyme complex is composed of three enzymes, namely: cellulase, cellobiohydrolase and β-D glucosidase. The activity of each of the three has been confirmed for H. ericae (Burke and Cairney, 1997; Varma and
Bonfante, 1994) and overall the activities were in the same order as those reported for the 74 known saprobe, Trichoderma reesei Simmons (Kolbe and Kubecik, 1990), indicating that the cellulolytic potential for H. ericae is considerable (Burke and Cairney, 1997).
H. ericae has been shown to produce a range of hemicellulolytic enzymes of which many are active against primary cell wall polymers. The enzymes include a xylanase complex, a mannanase complex and an enzyme acting against arabingalactan (Burke and Cairney,
1997). Each complex has an endo- and exo-acting enzyme acting synergistically to give complete hydrolysis of the target polymers (Coughlan et al., 1993). Perotto et al. (1997) also showed H. ericae also produced at least one pH-regulated polygalacturonase isozyme with pectinolytic abilities.
14 Radiorespirometric evidence, showing respiration of CO2 from lignin during growth in axenic culture, suggests that H. ericae may possess a degree of ligninolytic ability
(Haselwandter et al., 1990). In white rot fungi, lignin degradation happens via enzymatic combustion, whereby an enzyme produces an intermediate, which in turn causes oxidation within the polymer and initiates the combustion. The enzymes involved are lignin peroxidase (LiP), manganese peroxidase (MnP) and laccase. Lignin peroxidase cleaves the linkage between the lignin monomers facilitating complete lignin breakdown, while the other enzymes involved oxidise phenolic residues (Hatakka, 1994). In white rot fungi, LiP and MnP both use H2O2 to produce reactive intermediates to oxidise aromatic nuclei in lignin polymers (Gold et al., 1989). To date there are no reports of H. ericae producing extracellular LiP or MnP activity. Burke and Cairney (1998) suggest that H. ericae may induce lignin degradation via the action of another oxidising intermediate, the 75 hydroxyl or Fenton radical which is also produced during lignin degradation by brown rot fungi (Backa et al., 1992). It was suggested that lignin degradation by H. ericae may proceed in a way similar to the brown rot process (see Cairney and Burke, 1998).
Due to limited litter production, along with both fire and decomposer activity, the relatively dry habitats occupied by many Australian Ericaceae have relatively low organic matter content compared to mor-humus heathlands in the northern hemisphere (Straker,
1996). To date, there has been only one investigation of the enzymatic cell wall degrading abilities of endophytes obtained from Australian Ericaceae. In this report it was demonstrated that the endophytes produced extracellular β-1-4-endoxylanase activity
(Cairney et al., 1996). Midgley et al. (2004b) showed that ericoid mycorrhizal fungi from an Australian Ericaceae host were able to produce measurable biomass on a range of cell wall related carbohydrates as sole sources of carbon, which suggests that these fungi probably posses a suite of enzymes capable of degrading plant cell walls.
However, it has been suggested that mycorrhizal endophytes of Australian Ericaceae may be less well adapted to dealing with nitrogen sources complexed with moribund plant cell wall material (Cairney and Ashford, 2002). This may reflect the fact that soils from the
Australian Ericaceae habitats contain less organic material than their northern hemisphere counterparts.
76
3.1.4 Abilities of ericoid mycorrhizal fungi to produce chitinase enzymes
Aside from plant material, a significant proportion of soil biomass is present in the form of living and dead fungal mycelium (eg. Zvyagintsev, 1994). Fungal mycelium represents a major pool of soil nitrogen, much of which is deposited in cell walls as chitin (a polymer of β–1,4 linked N-acetylglucosamine units), and can represent up to around 30% of total soil nitrogen in some northern hemisphere heathlands (Kerley and Read, 1997).
The ericoid mycorrhizal endophytes H. ericae and Oidiodendron griseum can utilise purified chitin as a sole nitrogen source via absorption of the products of chitinolysis
(Leake and Read, 1990a; Mitchell et al., 1992; Kerley and Read 1995). Significantly, some of the nitrogen derived from purified chitin is transferred to the Ericaceae host plant
(Kerley and Read, 1995). Moreover, the likely ecological significance of these chitinolytic capabilities is emphasised by the observation that H. ericae can derive nitrogen from fungal necromass and effect its transfer to Ericaceae hosts (Kerley and
Read, 1997). Most current knowledge of chitin utilisation by ericoid mycorrhizal fungal endophytes, however, relates to H. ericae and the chitinolytic potential of other endophyte taxa remains unexplored.
Enzymatic hydrolysis of chitin to N-acetylglucosamine requires the action of both endo- and exo-acting activities (Patil et al., 2000). Thus the endo-acting chitinases [EC 3.2.1.14
(= 1,4-β-poly-N-acetylglucosaminidase)] randomly cleave 1,4-β-linkages in chitin macromolecules to release oligomers of N-acetylglucosamine of varying length. Exo- acting chitobiosidases (=N,N'-diacetylchitobiohydrolase) hydrolyse chitin and oligomers of N-acetylglucosamine from the non-reducing ends to release chitobiose, while exo- 77 acting β-N-acetylhexosaminidases (EC 3.2.1.52) cleave chitin and oligomers of N- acetylglucosamine from the non-reducing end to release N-acetylglucosamine monomers
(Patil et al., 2000; Kubicek et al., 2001). A number of ectomycorrhizal basidiomycetes produce both endo- and exo-acting chitinolytic activities (Hodge et al., 1995), and genes encoding β-N-acetylhexosaminidases are known to be widespread in ectomycorrhizal basidiomycetes (Lindahl and Taylor, 2004).
Although H. ericae has been shown to produce extracellular acid chitinase activity
(Mitchell et al., 1992; Kerley and Read, 1997), further characterisation of this activity has not been conducted. The aim of the research described in this chapter was to investigate the production of extracellular endo- and exo-acting chitinolytic activities by ericoid mycorrhizal endophytes from the Australian Ericaceae host Epacris pulchella, along with non-mycorrhizal root-associated fungi from the same host.
3.2 Experimental procedures
3.2.1 Fungal isolates and growth conditions
The fungi used in this work were nine isolates obtained from the hair roots of the E. pulchella (Ericaceae) plants sampled in Chapter 2, along with an isolate of H. ericae
(Read 101) which was included for comparison (Table 3.1). The identities of the isolates from E. pulchella (five ericoid mycorrhizal endophytes and four non-mycorrhizal root- associated fungi) have been inferred by internal transcribed spacer (ITS) rDNA sequence comparisons (Table 3.1) and their ability to form typical ericoid mycorrhizal structures in roots of the Australian Ericaceae W. pungens tested in gnotobiotic culture (see Chapter 78
2). Cultures were routinely maintained on 2% potato dextrose agar (PDA) (Oxoid) and were transferred to modified Melin-Norkrans (MMN) agar medium (Marx and Bryan,
1975) for 28 d prior to establishment of the experiments.
For investigation of chitinase activities, all isolates were grown in a basal liquid medium
-1 -1 to which 0.5 g l purified crystalline chitin and 0.25 g l (NH4)2HPO4 were added. The basal medium was based on MMN liquid medium (Marx and Bryan, 1975) and contained
-1 (l ): KH2PO4, 0.30 g; MgSO4.7H2O, 0.14 g; CaCl2, 50 mg; NaCl, 25 mg; ZnSO4, 3.0 mg; ferric EDTA, 12.5 mg; thiamine, 0.13 mg. To determine the effects of different media constituents on chitinase production, EP RFLP 1 was grown in basal medium containing
-1 -1 -1 (a) 0.5 g l purified crystalline chitin and 0.25 g l (NH4)2HPO4, (b) 0.5 g l purified
-1 -1 -1 crystalline chitin, (c) 0.25 g l (NH4)2HPO4, (d) 10 g l D-glucose or (e) 0.5 g l purified
-1 -1 crystalline chitin and 0.25 g l (NH4)2HPO4, and 10 g l D-glucose. All chemicals were from Sigma, and liquid media were adjusted to pH 4.0 with HCl prior to autoclaving.
Fifteen ml of liquid medium was dispensed into 9.0 cm diam., Petri dishes, which were inoculated with single (5.0 mm diam.) plugs of fungus excised from the growing edge of cultures on MMN agar medium. Petri dishes were maintained in the dark at 21oC and three replicate dishes were harvested for determination of chitinase activities for each isolate, 10, 20 and 30 d after inoculation. For determination of chitinase production by EP
RFLP 1, three replicate Petri dishes of the fungus in each medium were also harvested 10,
20 and 30 d after inoculation. Three uninoculated Petri dishes containing each medium were also harvested at each time point. At harvesting the contents of each Petri dish was 79
filtered through a 0.45 µm Millipore filter (Millipore Corporation) using gentle suction to
separate the culture solution from fungal mycelium and unused chitin.
Table 3.1 Details of isolates used for determination of chitinolytic activities.
Isolate Identity (ITS sequence GenBank Ericoid Culture collection accession) mycorrhizal status accession† EP RFLP 1 ericoid mycorrhizal endophyte (AY627804)1 yes BRIP 46109 a EP RFLP 2 ericoid mycorrhizal endophyte (AY627806)1 yes BRIP 46105 a EP RFLP 50 Helotiales (AY627812)1 yes BRIP 46112 a EP RFLP 13 Oidiodendron (AY627815)1 yes BRIP 46113 a EP RFLP 15 Oidiodendron (AY627817)1 yes BRIP 46114 a EP RFLP 52 Dermataceae (AY627818)1 no BRIP 46115 a EP RFLP 56 Hypocreales (AY627831)1 no BRIP 46116 a EP RFLP 57 Agaricales (AY627833)1 no BRIP 46117 a EP RFLP 58 Tremales (AY627835)1 no BRIP 46118 a Read 101 Hymenoscyphus ericae yes BRIP 46119 a 1see Chapter 2, †accessions in Plant Pathology Herbarium (BRIP), Queensland Department of Primary Industries.
3.2.2 Assay for extracellular chitinase activities
The chitinase assay was based on the method described by Hodge et al. (1995) using 4-
methylumbelliferyl (4-MU) glycosides of N-acetylglucosamine oligosaccharides (4MU-
(GlcNAc)1-3) (monomer, dimer and trimer) (Sigma) as fluorogenic substrates. Each
substrate was initially dissolved in 250 µl of dimethyl sulfoxide and stock solutions
subsequently prepared at a final concentration of 0.8 mM in distilled water and stored at –
20oC. Forty-five µl aliquots of culture filtrate were added to 1.5 ml microcentrfuge tubes.
One hundred µl of 0.1 M MES (2-(N-morpholino)-ethanesulfonic acid) buffer, pH 4.0,
along with 5.0 µl of monomer, dimer or trimer were added to each tube to initiate the
reaction. Controls containing buffer, substrate and 45 µl of autoclaved culture filtrate 80 were included for each isolate : substrate combination. The assay tubes, including controls, were incubated at 37 oC and after 1 h the reaction was terminated by adding 100
µl 1.0 M NaOH. Samples were transferred to UVettes® (Eppendorf) and the fluorescence produced by methylumbelliferyl released via chitinolytic activity determined immediately
(excitation 355 nm and emission 460 nm) using a Turner® QuantechTM digital filter fluorometer (Barnstead International). Fluorescence was within the linear range of a 0.0-
1.5 µg ml-1 methylumbelliferyl standard curve, from which enzyme activities in pkat (mol s-1) were calculated. The total crude protein concentration (mg ml-1) of each culture filtrate was determined using the method of Bradford (1976). One ml of culture filtrate was thus added to 1.0 ml of Bradford reagent (Sigma) in a 5.0 ml centrifuge tube, and incubated at room temperature for 45 min. Absorbance was then measured at 595 nm using a BioPhotometer (Eppendorf). Specific activities of chitinolytic enzymes were expressed in terms of total culture filtrate protein at each harvest (pkat mg-1 total protein).
Following checking for normality, data were analysed using one-way ANOVA, and significant differences determined by Fisher’s LSD using the Minitab® program v. 12.23.
3.3 Results
Activities against the 4-MU N-acetylglucosamine monomer (exo-acting activity), dimer
(exo-acting activity) and trimer (endo-acting activity) were detected in culture filtrates of most isolates (Figure 3.1). The exceptions were two non-mycorrhizal isolates (EP RFLP
52 and EP RFLP 57), for which activity against only the monomer was detected (Figure
3.1). For those isolates that produced measurable activity against the three substrates, activity against the monomer was greater at all time points, however in most cases the 81 differences were not statistically significant (P = 0.05). For some isolates (eg. EP RFLP
50 and 15) chitinolytic activities appeared to have peaked during the 30 d duration of the experiment, while, for others (eg. EP RFLP 1 and 2), highest specific activities were recorded after 30 d growth (Figure 3.1). Highest specific activities against the monomer ranged from 233 pkat mg-1 total protein (EP RFLP 2) to 25 pkat mg-1 total protein (EP
RFLP 52), however all ericoid mycorrhizal isolates, including H. ericae had highest specific activities against this substrate in the range 90-233 pkat mg-1 total protein (Figure
3.1). No activity was detected in the autoclaved culture filtrate controls, confirming the enzymatic nature of the activity.
No activity against any of the three substrates was detected for EP RFLP 1 when glucose was included in the growth medium (media d and e), even when purified crystalline chitin was also present (Figure 3.2). Highest specific activities against the three substrates were measured following growth in the media containing purified crystalline chitin (medium b) or purified crystalline chitin and (NH4)2HPO4 (medium a) (Figure 3.2). Only trace activities against each of the substrates were measured after 20 or 30 d growth when
(NH4)2HPO4, but neither purified crystalline chitin nor glucose, was present in the medium (Figure 3.2).
250 250 82 EP RFLP 1 EP RFLP 2 200 200
150 150
100 100
50 50
0 0 0 10 20 30 0 10 20 30
250 250 EP RFLP 50 EP RFLP 13 200 200
150 150
100 100
50 50
) 0 0 0 10 20 30 0 10 20 30 rotein
p 250 250 total EP RFLP 15
-1 EP RFLP 52 200 200 g 150 150 kat m
(p 100 100
y 50 50 0 0 0 10 20 30 0 10 20 30 me activit y 250 250 Enz EP RFLP 56 EP RFLP 57 200 200
150 150
100 100
50 50
0 0 0 10 20 30 0 10 20 30
250 250 EP RFLP 58 200 200 150 150 100 100 50 50
0 0 0 10 20 30 0 10 20 30 Time (days) Figure 3.1 Extracellular chitinolytic activities (pkat mg-1 total extracellular protein) against 4-MU N-acetylglucosamine monomer (●), dimer (▼) and trimer (■) in culture filtrates of selected ericoid mycorrhizal fungi and root-associated fungi from Epacris pulchella. Each point is the mean of three replicates and bars give LSD (P = 0.05) for each harvest.
250 Monomer 83 200
150
100
50
0 0 10 20 30
250
200 Dimer
total protein) total protein) -1 150
100
50
mg (pkat Enzyme activity 0 0 10 20 30 250
200 Trimer
150
100
50
0 0 10 20 30
Time (days) Figure 3.2 Extracellular chitinolytic activities (pkat mg-1 total extracellular protein) 4-MU N-acetylglucosamine monomer, dimer and trimer in culture filtrates of the ericoid mycorrhizal endophyte EP RFLP 1 following growth in basal medium containing chitin and (NH4)2HPO4 (▲), chitin (♦), (NH4)2HPO4 (■), glucose (●) or chitin, (NH4)2HPO4 and glucose (▼). Each point is the mean of three replicates and bars give LSD (P = 0.05) for each harvest. 84
3.4 Discussion
H. ericae has been shown previously to utilise chitin as a sole nitrogen and carbon source and to produce extracellular chitinolytic activity (Leake and Read, 1990a; Mitchell et al,. 1992; Kerley and Read, 1995; 1997). The data presented in this chapter are consistent with these observations and further demonstrate the chitinolytic abilities of five ericoid mycorrhizal, along with four non- mycorrhizal root-associated fungi from E. pulchella. Moreover, the data indicate that the ericoid mycorrhizal fungi, including H. ericae, produce both endo- and exo-acting chitinolytic activities.
Activity against the monomer is indicative of exo-acting β-N-acetylhexosaminidase activity, while release of fluorescence from the dimer and trimer reflect exo-acting chitobiosidase and endo-acting chitinase activities respectively (Tronso and Harman, 1993; Hodge et al., 1995).
Only β-N-acetylhexosaminidase activity was detected for a non-mycorrhizal ascomycete (EP
RFLP 52) and basidiomycete (EP RFLP 43), however activity against the three substrates was confirmed for all of the ericoid mycorrhizal fungi along with a non-mycorrhizal ascomycete (EP
RFLP 56) and basidiomycete (EP RFLP 58). Although only statistically significant at certain time points for some isolates, specific activities for β-N-acetylhexosaminidase were generally higher than for the other two activities, for all isolates tested. This is consistent with the data of
Hodge et al. (1995) who identified this activity as the predominant extracellular chitinolytic activity in ectomycorrhizal and non-mycorrhizal basidiomycetes. Highest specific activities for
β-N-acetylhexosaminidase recorded for the five ericoid mycorrhizal fungi from E. pulchella and for H. ericae were of the same order of magnitude, being in the range ca 90-233 pkat mg-1 total protein, suggesting that their chitinolytic potential may be broadly similar.
85
Highest chitinase activities were produced by the ericoid mycorrhizal isolate EP RFLP 1 in the presence of chitin, however the presence of glucose, in addition to chitin, repressed chitinase activities. Similar observations have been made previously for a range of ectomycorrhizal and non-mycorrhizal fungi (eg. Hodge et al., 1995; de la Cruz et al., 1993; El-Katatny et al., 2000) and for expression of some chitinase genes (eg. Dana et al., 2001). In the case of the ericoid mycorrhizal endophyte H. ericae, Kerley and Read (1997) found that extracellular chitinase production in a medium containing a fungal cell wall fraction as the sole nitrogen source, and half as much glucose as used in the current work, was negligible until 30 d growth. Such an observation is consistent with initial glucose repression of chitinase production followed by induction of activity in the presence of the cell wall material upon glucose depletion. In contrast,
Mitchell et al. (1992) found that extracellular chitinase activity was produced by a different isolate of H. ericae in both the presence and absence of chitin. Although the presence of chitin may induce expression of certain fungal chitinase genes (eg. Dana et al., 2001) expression of other chitinase genes may be triggered by carbon starvation rather than by the presence of the substrate or its breakdown products (eg. Mach et al., 1999). Extracellular chitinase activity in the absence of chitin was observed for nitrogen-sufficient cultures of H. ericae when a small amount of ‘starter’ glucose was present in the growth medium (Mitchell et al., 1992). Since this would have been utilised during the initial growth phase, it is likely that cultures would have become rapidly carbon-starved and this might induce chitinase production. The equivalent treatment in the present study (medium c), comprising basal medium + (NH4)2HPO4, contained no carbon source, and trace activities against each substrate were detected after 20 and 30 d growth, suggesting that carbon starvation induces low level expression of chitinase activities in EP RFLP
1. 86
In addition to demonstrating the chitinolytic potential of five ericoid mycorrhizal endophytes from E. pulchella, the data presented in this chapter also demonstrate that non-mycorrhizal fungi associated with roots of the same host produce chitinase activities. Although two non- mycorrhizal isolates produced activity against only the monomer (exo-acting β-N- acetylhexosaminidase activity), two of the non-mycorrhizal isolates produced activities that were active against all three chitin analogue substrates. A range of non-mycorrhizal fungi is associated with Ericaceae hair roots (see Chapter 2) and these may compete with ericoid mycorrhizal endophytes for nutrients sequestered in substrates such as fungal necromass. The extent to which this occurs in situ remains unclear, however it has been suggested that carbon supplied by the plant host would render the mycorrhizal endophytes more competitive than the relatively carbon- starved non-mycorrhizal fungi (Kerley and Read, 1995). Such an advantage would require that the supply of carbon from the host did not repress chitinase production. Data from the present study indicate that availability of a simple carbon source such as glucose can inhibit chitinase production in axenic culture. H. ericae has, however, been shown to supply its host with nitrogen from purified chitin and fungal wall material (Kerley and Read, 1995; 1997), providing strong evidence that chitinase activities are produced by the fungus during symbiosis. Further work will clearly be required to fully understand carbon source and host plant effects on chitinase production by ericoid mycorrhizal fungi.
87
CHAPTER 4
Fungi associated with hair roots of Rhododendron lochiae in an Australian tropical
montane cloud forest
4.1 Introduction
4.1.1 Habitat diversity of Ericaceae and associated ericoid mycorrhizal fungi
The ericoid mycorrhiza has in the past been regarded as the most specific of mycorrhizas because it is restricted to hosts in the family Ericaceae and the participation of a small group of ascomycete fungi as mycobionts in the association (Harley and Smith, 1983).
However, observations over the past 20 years have revealed that the taxonomic diversity of the fungal partners involved in the association is not as restricted as once thought (see
Chapter 2). Considering this diversity amongst fungal symbionts it is reasonable to suggest that different fungal groups have been, and perhaps still are able, to evolve these associations under common selective environmental pressures (Straker, 1996).
Plants that form ericoid mycorrhizas thrive in habitats where soils are subject to considerable climatic and edaphic stresses. These soils have characteristically extremely low nutrient status and display various levels of low pH, high metal availability, poor drainage, free drainage and high or low temperatures (Cairney an Meharg, 2003). The success of Ericaceae in these stressful habitats is thought to lie largely in the abilities of their ericoid mycorrhizal fungal endophytes to enhance plant fitness by reducing the effects of these stresses upon them (Cairney an Meharg, 2003). It has, for example, been 88 known for some time that certain ericoid mycorrhizal fungi produce a broad range of enzymes that are important in mobilising nutrients from organic complexes in the soil that would otherwise be unavailable to the host plants (see Chapter 3).
Understanding the benefits bestowed upon Ericaceae hosts by their fungal partners has driven much of the recent investigation and discovery regarding the diversity and function of these symbiotic associations. The habitats from which ericoid mycorrhizal associations of a limited number of Ericaceae taxa have been investigated include northern hemisphere mor humus heathlands, where the mycorrhiza forming-plants can occur as more or less pure stands. Ericoid mycorrhizas from mediterranean woodlands and boreal forests, where Ericaceae taxa represent a major component of the understorey, drier sandy heathlands and sclerophyl forests along with alpine heathlands of Australia have also been investigated (Cairney and Ashford, 2002). All of these habitats have in common some degree of climatic and edaphic stresses that are characteristic of the areas inhabited by Ericaceae.
As a family, the Ericaceae are diverse and geographically widespread, occurring in boreal, temperate and tropical regions of all continents except Antarctica. The mosaic of habitats that Ericaceae inhabit has led to evolution of varied lifeforms that include terrestrial, lithophytic and epiphytic herbs, shrubs, trees, lianas and mat forming cushion plants. The greatest diversity of Ericaceae genera and species lie in the montane cloud forests of the neotropics (tropical Central and South America) and Southeast Asia, including Papua New Guinea and northern Australia. Worldwide, there are 89 approximately 125 genera and 4500 species of Ericaceae with the five largest genera being Rhododendron, Erica, Vaccinium, Cavendishia and Gaultheria (Luteyn, 2002).
Although investigations carried out to date have significantly increased what is known about ericoid mycorrhizas, they have been restricted to relatively few Ericaceae taxa from relatively few habitat types in primarily temperate to boreal areas. Investigations of
Ericaceae from other habitats, particularly tropical montane cloud forests (TMCF), where the majority of ericaceous taxa occur, are likely to add considerably to what is known about the diversity and function of ericoid mycorrhizas.
TMCF comprise forest ecosystems of distinctive floristic and structural form. They typically occur in an altitudinal zone between 1000 m and 3000 m where the cooler atmospheric environment is characterised by persistent, frequent or seasonal cloud cover at the vegetation level causing "canopy wetting". The total precipitation in such forests is significantly enhanced by canopy interception and can be between 5 and 100 cm annually
(Hamilton et al., 1995; Bruijnzeel and Veneklaas, 1998). In comparison with lowland tropical forests, TMCF vegetation is characteristically reduced in stature with an increased stem density. Canopy trees usually exhibit gnarled trunks and branches, dense compact crowns and sclerophyllous leaves (Hamilton et al., 1995). TMCF are also characterised by having a high proportion of biomass as epiphytes and a corresponding reduction in vine climbers. As the epiphyte and canopy biomass dies and decomposes, it forms mats of organic material on tree branches which provide an environment for roots of host trees as well as epiphytic plants (Hamilton et al., 1995; Tanner et al., 1998). Soils are wet and frequently waterlogged, and are highly organic in the form of mor humus due 90 to relatively slower rates of microbial decomposition and mineralisation. Consequently, these soils are of low pH, contain high levels of phenolics and nutrient availability, particularly nitrogen, is limited (Tanner et al., 1998). Biodiversity in terms of plant species can be relatively high compared to lowland rainforest and endemism is often very common (Hamilton et al., 1995). The location, identification and extent of the various
TMCF are not well known as no detailed mappings have been produced. However, it is known that they occur globally in areas of the neotropics, central Africa, some Caribbean and Pacific Islands and Southeast Asia including Papua New Guinea and northern
Australia (Hamilton et al., 1995).
4.1.2 Ericoid mycorrhizas in tropical montane cloud forest
It is well established that temperate and boreal terrestrial Ericaceae form beneficial ericoid mycorrhizal associations (Smith and Read, 1997). However, there have been very few morphological studies and no molecular studies of ericoid mycorrhizal diversity in tropical Ericaceae. Nevertheless, based on the striking similarities between climatic and edaphic conditions in mor humus heathlands and TMCF, and observations presented in the following studies, it seems likely that symbiotic ericoid mycorrhizal associations are found in tropical Ericaceae. Bermudes and Benzig (1989) and Lesica and Antibus (1990) sampled roots of a variety of epiphytic Ericaceae (Cavendishia sp., Sphyrospermum sp.,
Macleania sp, and Psammisia sp.) from TMCF in Ecuador and Costa Rica respectively.
The authors observed high levels of fungal endophyte colonization, along with formation of coils similar to those that colonise their terrestrial Ericaceae counterparts. Bermudes and Benzig (1989) also noted that the same root samples were covered by an ECM-like 91 hyphal mantle that penetrated between epidermal root cells. They suggested that this structure may reflect a novel group of Ericaceae-associated mycorrhiza, the demateaceous surface fungi (DSF). Similar structures have been observed in temperate
Ericaceae species from the USA (Wurzburger and Bledsoe, 2001) and Canada (Xiao and
Berch, 1996). However none of the authors who observed these structures has been able to demonstrate the benefit of this putatative mycorrhiza type to the plant host. Rains et al. (2003) also observed ericoid mycorrhiza–like structures, along with structures similar to the DSF, in both epiphytic and terrestrial Ericaceae roots from a Costa Rican TMCF.
Kottke et al. (2004) observed that the roots of the non-ericaceous species, Graffenrieda emarginata (Ruiz and Pav.) Triana, from an Ecuadorian cloud forest were regularly colonised by fungi that formed a superficial Hartig net. Using sequence data, the authors identified two fungi that formed this association; one of which was related to the ascomycete Hymenoscyphus ericae, and the other to the Sebacinaceae basidiomycetes.
Fungi from both of these groups are known to form both ecto- and ericoid mycorrhizal associations (Weiss and Oberwinkler, 2001). Further investigations by Haug et al. (2004) identified 37 rDNA ITS sequences from fungi associated with roots of G. emarginata.
Twenty of these sequences had closest matches with sequences representing ascomycetes.
Phylogenetic studies grouped ten of these sequences in a distinct clade of the H. ericae aggregate that was most closely related to a clade from Vrålstad et al. (2002b), which contained sequences from ericoid mycorrhizal fungi, including the type species H. ericae.
Of the remaining 10 ascomycetes, three sequences were most similar to those from fungi commonly isolated from Ericaceae species from Canada (Monreal et al., 1998; Allen et 92 al., 2003). The presence of these fungi, particularly those similar to the common ericoid mycorrhizal fungi in the H. ericae aggregate suggests that if Ericaceae species inhabit the same TMCF field site, which is probable considering the diversity of Ericaceae found in the neotropics, they are likely to form ericoid mycorrhizal associations.
4.1.3 TMCF in Australia
Several areas in northeast Australia's Wet Tropics World Heritage Area are classified as
TMCF (McJannet and Reddell, 2003). These areas are at an altitude above 1000 m and have an annual average rainfall of about 20 cm. This rainfall is seasonal with mean precipitation in the wettest and driest quarters of about 10 cm (January to March) and 1.5 cm (August to October) respectively. Temperature ranges from an average minimum of
8ºC in the coolest period (June to August) to an average maximum of 29ºC in the warmest period (December to February) (Talbot et al., 2003). As is the case for other
TMCF throughout the world, Ericaceae taxa occur in cloud forests of northern Australia.
At least two ericaceous genera, Rhododendron and Agapetes, are found in these areas.
Both genera are members of the sub-family ericoideae, along with Calluna and Erica species (Kron et al., 2002). The main centers of diversity for the genus Rhododendron are the mountains of sub-continental and South-east Asia, Caucasus, the European Alps and temperate North America (Luteyn, 2002). The sole Australian species of Rhododendron described to date, R. lochiae, occurs in TMCF regions in North Queensland, and belongs in the largely South-east Asian R. sect. Vireya ser. Javinica Sleum (Sleumer, 1960). R. lochiae, which is endemic to the area, has been recorded as growing as a lithophytic, epiphytic or terrestrial shrub to three meters, in forest, in moss-covered boulder outcrops 93 or in windswept mossy thickets in cracks of bare rock exposures at altitudes between
900-1350m in the TMCF (Craven and Withers, 1996). The genus Agapetes comprises about 80 species but Agapetes meiniana F. Muell. is its sole Australian member. A. meiniana is a climber that occupies similar TMCF habitats to R. lochiae in north
Queensland (Craven and Withers, 1996).
Fungi that form ericoid mycorrhizal associations with Australian Ericaceae taxa that have been examined thus far are broadly similar to those observed in the northern hemisphere.
These belong to two broad groups; one with affinity to Helotiales the other with
Myxotrichaceae (See Chapter 2). To date, only members of sub-family Styphelioideae have been investigated and these were sampled from habitats that are relatively dry and lacking in organic matter compared to the TMCF of Northeast Australia. Furthermore, few detailed investigations of fungal diversity have been carried out at any level in
TMCF of north Queensland and so information obtained could prove useful for both understanding fungal biodiversity in a unique habitat as well as conservation of R. lochiae, which is considered a rare species.
The aims of the work described in this chapter were thus to (i) examine the taxonomic richness and relative abundance of fungi associated with the roots of R. lochiae using both culturing and direct extraction of fungal DNA from roots, (ii) compare these data to those from fungi associated with the roots of Ericaceae from temperate habitats and other plant taxa from tropical habitats, and (iii) explore the ability of the fungi to form ericoid mycorrhizal or other root associated structures with axenic Ericaceae seedlings. 94
4.2 Experimental Procedures
4.2.1 Collection of plants and isolation of fungal cultures
Three R. lochiae plants (RL1, RL2 and RL3) including relatively undisturbed rootballs were collected from a field site at Mount Lewis Forest Reserve, Queensland, Australia (S
16º 30’ E 145º 16’) in December 2003. The field site is classified as tropical montane cloud forest. The R. lochiae plants grew at an altitude of 1150 m and had a lithophytic/terrestrial habit, growing on moss-covered boulder outcrops along with
Agathis emergents, Orchidaceae and Leptospermum in a layer of granite based soil, which appeared to have a high organic matter content. RL1 was collected 50 m from RL2 and RL3, which were growing within 2 m of each other in a position more exposed to wind and sunlight. Upon collection, each plant (along with the adhering soil) was transferred to the laboratory within 48 h, where soil was removed by careful rinsing under gently running tap water for 15 min. Any remaining soil or debris was gently removed with forceps. The hair root system of each plant was divided into three sections of roughly equal size. These root sections were surface sterilised as detailed for Epacris pulchella (Chapter 2). The sterilised root sections were cut into pieces (ca 5 mm in length). Alternating pieces from each root section were then either pooled for direct DNA extraction and DGGE analysis or cultured as detailed for E. pulchella (Chapter 2) on
2.0% MEA. In total, 120 root pieces were excised and plated from each of the three plants. Petri dishes were incubated and emerging fungi were sub-cultured and maintained on PDA in the same manner as described in Chapter 2.
95
4.2.2 DNA extraction
DNA was extracted from cultures and pooled root pieces from each root section using the
FastDNA® Kit (Bio 101) and FastPrep® Instrument (Bio 101) as described in Chapter 2.
4.2.3 ITS amplification, RFLP analysis and sequence analysis
The ITS region from all cultured fungi was amplified in a 50 μl reaction volume, as described by Midgley et al. (2004a) using 25 pmol of each of the primers ITS1 and ITS4
(White et al., 1990). Amplifications were performed in a PCR Express thermocycler
(Hybaid) using the cycling parameters outlined in Chapter 2 for the ITS region of the cultured fungi. Reactions were performed in duplicate and a negative control containing no DNA was included in every reaction run. Amplification products were electrophoresed in 2.0 % (w/v) agarose gels, stained with ethidium bromide and visualised under UV light. ITS products (ca 1.0 μg) from cultures were digested using the restriction endonucleases Hae III, Hinf I and Rsa I (Promega) as described in Chapter
2. The restriction fragments were separated by electrophoresis on a 3.0% agarose gel and stained and viewed as described above.
For sequencing, ITS-PCR products were cloned with the pGEM-T Easy vector system
(Promega). Representative isolates of each RFLP-type, as determined by different banding patterns, were sequenced using the ABI Big-Dye reaction kit in an ABI 373-an automated fluorescent DNA sequencer (Applied Biosystems, Foster City, CA, USA).
Sequencing reactions were performed with the T7 primer (Promega). ITS sequences were analysed using the FASTA 3.0 program (Pearson and Lipman, 1988) to find the closest 96 sequence matches in the GenBank/EMBL/DDBJ databases. Selected ITS sequences were aligned using the PILEUP program [within the EGCG extentions to the Wisconsin
Package, Version 8.1.0 (Rice, 1996)], alignments were optimised first using the ClustalW option and subsequently manually by visual inspection. Neighbour-joining and parsimony analyses of the patristic distance matrix (1000 bootstraps re-sampling replicates) were then conducted using PAUP (v. 4.01b9) to infer relationships between the fungal endophytes in the present study and with sequences available for other fungi in the
GenBank and EMBL nucleotide databases.
4.2.4 Testing for ericoid mycorrhiza formation
Due to difficulties in obtaining R. lochiae seed, or any seed from a species in subfamily ericoideae, Vaccinium macrocarpon (subfamily vaccinoideae) was selected as a host plant to use for mycorrhiza formation trials. V. macrocarpon seeds were extracted from commercially available frozen fruits and sterilised, germinated and axenically cultured as per Woollsia pungens in Chapter 2. After three weeks growth, root zones of seedlings were singly inoculated with a mycelial slurry of each cultured RFLP-type from the R. lochiae plants as described for W. pungens seedlings in Chapter 2. After 12 weeks, hair roots were excised and rinsed to remove excess medium and were cleared, stained and assessed for mycorrhizal infection following the methods used in Chapter 2.
4.2.5 ITS amplification for DGGE analysis
In preparation for DGGE, PCR was carried out using DNA directly extracted from pooled root pieces from each root section obtained from RL1, RL2, RL3 and each of the 97 cultured fungi. The reaction mixtures and amplification conditions were as described in
Chapter 2, using 25 pmol of each of the primers ITS1F-GC and ITS2. Negative controls were included in each PCR run and ITS amplified products were visualised as described above.
4.2.6 DGGE analysis
ITS products generated from pooled root pieces from each root section, along with combinations of products from cultured endophytes, were separated by DGGE using the
DCodeTM Universal Mutation Detection System (BioRad). This was performed following the method outlined in Chapter 2, except that a 27.5% to 50% vertical denaturing gradient was employed and 1.0 µl ITS-PCR product was loaded into each lane. Gels were stained with SYBR green (Molecular Probes) following the methods detailed in Chapter 2 and visualised under UV light. Digitalised DGGE images were analysed using the PhoretixTM
1D Advanced software (Version 5.2) (Nonlinear Dynamics). Bands on the gels which were present in the pooled root sample direct DNA extraction lanes but absent in the cultured endophyte lanes were excised and reamplified as above with the ITS1 and ITS2 primer pair (Gardes and Bruns, 1993). Reamplified products were cloned, sequenced and analysed as described above.
4.3 Results
4.3.1 ITS-RFLP analysis of cultured fungal assemblages
A total of 331 slow growing, mycelial isolates was cultured from the 360 hair root pieces excised from the three R. lochiae plants, with 126, 98 and 108 isolates obtained from
RL1, RL2 and RL3 respectively (Table 4.1). ITS-PCR amplification produced a single 98
PCR product (ca 500-650 bp) from each isolate. ITS-RFLP analysis grouped the cultured isolates from RL1, RL2 and RL3 as 38, 22 and 25 RFLP-types respectively (Tables 4.2 and Table 4.3). Nine RFLP-types were common between the cultured assemblages from all three plants, while 23, seven and 10 RFLP-types were unique to RL1, RL2 and RL3 respectively. Three RFLP-types each were common between the RL1 and RL2; RL1 and
RL3; and RL2 and RL3 assemblages respectively, giving a total of 58 cultured RFLP types for the three plants (Table 4.2 and Figure 4.1).
Table 4.1 The number and percentage of root pieces sampled for each of the three
Rhododendron lochiae plants from the Mt Lewis Forest Reserve field site, from which slow growing fungal endophytes were obtained.
Root System Number of hair Number of Percentage (%) of hair root root pieces isolates pieces yielding slow growing examined obtained fungal isolates R. lochiae 1 (RL1) 120 126 100.0 R. lochiae 2 (RL2) 120 97 84.9 R. lochiae 3 (RL3) 120 108 90.0
99
Patterns of relative abundance of cultured isolate RFLP-types were broadly similar for all three root systems, with each assemblage characterised by a few common and a greater number of less frequently isolated RFLP-types. However, the assemblage from RL1 consisted of more RFLP-types than those from the other two plants, many of which were represented by only a single isolate (Fig. 4.1). Approximately 55% of the cultured isolates from RL1 were identified as RL RFLP-types 1, 3, 14 and 21, with the remaining
RFLP-types each comprising a relatively small proportion each of the assemblage. From
RL2, approximately 60% of the cultured isolates were identified as RL RFLP-types 1, 5, and 14 while RL RFLP-types 4, 9, 14 and 23 represented approximately 60% of the isolate assemblage from RL3 (Fig. 4.1).
4.3.2 Sequence analysis of ITS RFLP-types
One cultured isolate representing each RFLP-type was selected for ITS sequencing and comparison with available sequences in the GenBank database. Putative taxonomic affinities were assigned conservatively to RFLP-types based on FASTA expected values and identities of the closest several sequence matches obtained from the FASTA searches
(see Chapter 2). Thus, for example, RL RFLP-type 19 had multiple close matches (low expected values) with sequences for Hymenoscyphus spp. and has been putatively designated as this genus. In contrast, for RL RFLP-type 45, the closest several FASTA matches were with various ascomycetes, and it has been putatively designated as
Ascomycota (Table 4.3). On this basis, ca 45% (RL1), ca 40% (RL2) and ca 50% (RL3) of the cultured RFLP-types were designated as putative ericoid mycorrhizal fungal endophytes (Figure 4.1 and Table 4.3). The majority of these isolates (RL RFLP-types 1 18
16 RL1 100 14
12
10 8
6
4 2
0 1 14 3 21 18 35 2 6 10 11 13 24 26 29 31 41 4 5 7 12 15 16 19 20 23 25 27 33 34 36 37 38 39 40 41 43 45 46
25
RL2 20
15
assemblage per plant 10
5
0 14 1 5 47 23 4 24 42 43 49 2 32 44 48 50 8 9 12 21 28 31 51
30 RL3 25 Percentage (%) of culture 20
15
10
5
0 9 14 23 4 24 8 2 3 18 26 47 21 30 43 1 17 22 44 52 53 54 55 56 57 58
RFLP-type Figure 4.1 Presence of each RFLP-type as a percentage of total cultured isolates obtained from the three Rhododendron lochiae root systems examined (RL1, RL2 and RL3), demonstrating that the fungal culture assemblage for each plant were characterised by a few common types and a larger number of less frequently isolated RFLP-types. Numbers for RFLP-types correspond to those in Table 4.3.
101
Table 4.2 Restriction fragment band sizes obtained by digestion with three restriction
endonucleases (HaeIII, HinfI and RsaI) for the rDNA ITS region amplified from representative
RFLP-types of fungal endophytes isolated from within the root systems of three Rhododendron
lochiae plants (RL1, RL2 and RL3).
RL RFLP HaeIII † HinfI † RsaI † type 1 282, 77, 61, 48, 29, 14, 12 268, 247, 8 360, 163 2 444, 78 269, 245, 8 358, 164 3 294, 141, 78, 14 269, 171, 79, 8 363, 164 4 327, 78, 61, 48, 8 268, 172, 74, 8 359, 163 5 306, 138, 77 268, 245, 8 358, 163 6 288, 138, 77, 19 269, 245, 8 358, 164 7 287, 78, 75, 62, 21 269, 246, 8 359, 164 8 331, 78, 62, 48, 8 270, 174, 75, 8 362, 165 9 294, 140, 77, 14 268, 171, 78, 8 362, 163 10 429, 73, 23, 15 276, 134, 122, 8 uncut 11 405, 74, 35, 15, 14 172, 133, 108, 73, 49, 8 uncut 12 256, 120, 72, 55, 36, 23 296, 258, 8 447, 115 13 293, 121, 72, 64 282, 136, 124, 8 uncut 14 429, 73, 23, 15 276, 134, 122, 8 uncut 15 243, 134, 94, 68 276, 255, 8 uncut 16 412, 163 309, 132, 126, 8 385, 140, 50 17 446, 154 332, 180, 80, 8 410, 191 18 275,79, 79, 62 245, 191, 51, 8 358, 137 19 277, 140, 83, 17 246, 246, 17, 8 359, 158 20 439,114 302, 165, 78, 8 287, 200, 66 21 299, 79, 75, 62 243, 163, 101, 8 356, 159 22 265, 79, 75, 62, 34 243, 163, 101, 8 356, 159 23 321, 76, 61, 48, 8 245, 242, 19, 8 262, 156, 96 24 370, 76, 61, 8, 6 248, 246, 19, 8 369, 96, 56 25 Uncut 265, 170, 51, 26, 8 361, 159 26 270, 87, 76, 62, 14, 12 248, 246, 19, 8 262, 162, 52, 45 27 Uncut 171, 154, 107, 79, 8 409, 110 28 287, 138, 93, 6 274, 242, 8 259, 169, 96 29 295, 142, 71 253, 167, 80, 8 360, 148 30 465, 72 254, 160, 115, 8 388, 149 31 263, 106, 77, 28, 15, 10, 8, 7, 4 241, 140, 129, 8 425, 93
† Numbers refer to fragment sizes in bp. 102
Table 4.2 cont. Restriction fragment band sizes obtained by digestion with three restriction endonucleases (HaeIII, HinfI and RsaI) for the rDNA ITS region amplified from representative
RFLP-types of fungal endophytes isolated from within the root systems of three Rhododendron lochiae plants (RL1, RL2 and RL3).
RL RFLP HaeIII † HinfI † RsaI † type 32 216, 103, 87, 54, 19, 17 351, 137, 8 uncut 33 436, 69 262, 235, 8 uncut 34 343, 176 237, 221, 26, 18, 9, 8 uncut 35 343, 176 237, 221, 26, 18, 9, 8 361, 158 36 220, 111, 75, 61, 18, 11 243, 113, 92, 18, 13, 9, 8 uncut 37 395, 102 247, 242, 8 397, 100 38 353, 177 247, 222, 26, 18, 9, 8 uncut 39 343, 188 237, 221, 38, 18, 9, 8 uncut 40 345, 74, 68, 31 261, 249 uncut 41 268, 170, 86, 44, 5 285, 280, 8 469, 104 42 438, 78, 31, 10, 5 181, 169, 113, 91, 8 uncut 43 401, 87, 17 250, 247, 8 uncut 44 287, 138, 93, 6 274, 242, 8 259, 169, 96 45 271, 147, 57, 25, 18, 18 254, 159, 115, 8 227, 176, 133 46 Uncut 220, 121, 87, 8 uncut 47 377, 85, 58 261, 251, 8 469, 51 48 270, 83, 78, 64, 19 256, 250, 8 497, 17 49 389, 127, 29 260, 146, 131, 8 uncut 50 403, 87, 17 252, 247, 8 uncut 51 372, 85, 64 261, 252, 8 uncut 52 298, 147, 73, 18 274, 254, 8 227, 175, 134 53 403, 87, 17 250, 249, 8 uncut 54 356, 108, 45, 13, 12 271, 140, 115, 8 377, 138, 19 55 278, 94, 86, 69, 20 262, 251, 26, 8 uncut 56 348, 82, 63, 19 256, 125, 92, 31, 8 386, 126 57 Uncut 217, 120, 87, 8 330, 102 58 306, 271, 128, 51 256, 178, 173, 141, 8 293, 190, 167, 106
† Numbers refer to fragment sizes in bp.
103
Table 4.3 Putative taxonomic affinities of RFLP-types present in cultured isolate assemblages from root systems of three Rhododendron lochiae root plants (RL1, RL2 and RL3) as inferred from FASTA searches of ITS sequences in the GenBank/EMBL/DDBJ databases. RFLP-types highlighted in bold are regarded on the basis of inoculation experiments as being ericoid mycorrhizal endophytes. RL RFLP Accession Putative taxonomic affinity FASTA No. of isolates in each host type code expected value RL1 RL2 RL3 1 AY699664 ericoid mycorrhizal endophyte 1.0e-129 21 19 1 2 AY699665 ericoid mycorrhizal endophyte 6.2e-121 4 2 3 3 AY699666 ericoid mycorrhizal endophyte 1.9e-133 14 0 3 4 AY699667 ericoid mycorrhizal endophyte 1.5e-138 1 3 7 5 AY699668 ericoid mycorrhizal endophyte 1.5e-123 1 13 0 6 AY699670 ericoid mycorrhizal endophyte 6.5e-119 3 0 0 7 AY699681 ericoid mycorrhizal endophyte 2.4e-125 1 0 0 8 AY699683 ericoid mycorrhizal endophyte 9.0e-138 0 1 4 9 AY699684 ericoid mycorrhizal endophyte 3.4e-133 0 1 32 10 AY699645 Xylariaceae 8.5e-110 3 0 0 11 AY699646 Xylariaceae 3.0e-110 2 0 0 12 AY699647 Xylariaceae 3.5e-84 1 1 0 13 AY699648 Xylariaceae 9.4e-89 2 0 0 14 AY699649 Xylariaceae 2.4e-109 20 24 16 15 AY699660 Xylariaceae 2.1e-82 1 0 0 16 AY699675 Xylariaceae 1.8e-105 1 0 0 17 AY699685 Xylariaceae 3.1e-76 0 0 1 18 AY699659 Helotiales 2.5e-115 6 0 3 19 AY699657 Hymenoscyphus 1.6e-126 1 0 0 20 AY699674 Helotiales 8.4e-110 1 0 0 21 AY699650 Hyaloscyphaceae 5.9e-134 14 1 2 22 AY699686 Hyaloscyphaceae 2.1e-128 0 0 1 23 AY699651 Dermataceae 7.5e-125 1 4 12 24 AY699652 Dermataceae 2.4e-131 2 3 5 25 AY699656 Dermataceae 1.4e-115 1 0 0 26 AY699662 Dermataceae 8.4e-133 2 0 3 27 AY699672 Dermataceae 1.0e-111 1 0 0 28 AY699687 Phialocephala 6.9e-130 0 1 0 29 AY699661 Ericaceae root endophyte 2.5e-117 2 0 0 30 AY699688 Ericaceae root endophyte 1.4e-91 0 0 2 31 AY699655 Oidiodendron 1.4e-137 2 1 0 32 AY699689 Myxotrichaceae 2.2e-47 0 2 0 33 AY699653 Hypocreales 8.1e-126 1 0 0 34 AY699658 Chaetosphaeriales 1.6e-126 1 0 0 35 AY699663 Chaetosphaeriales 1.0e-129 5 0 0 36 AY699673 Chaetosphaeriales 2.2e-92 1 0 0 37 AY699676 Chaetosphaeriales 1.0e-83 1 0 0 38 AY699679 Chaetosphaeriales 2.0e-91 1 0 0 39 AY699682 Chaetosphaeriales 7.3e-115 1 0 0 40 AY699677 Cladosporium 5.2e-123 1 0 0 41 AY699680 Sordariomycete 2.7e-78 1 0 0 42 AY699690 Trichocomaceae 1.9e-141 0 3 0 43 AY699654 Ascomycota 3.5e-66 1 3 2 44 AY699669 Ascomycota 3.3e-83 2 2 1 104
Table 4.3 cont. Putative taxonomic affinities of RFLP-types present in cultured isolate assemblages from root systems of three Rhododendron lochiae root plants (RL1, RL2 and RL3) as inferred from FASTA searches of ITS sequences in the GenBank/EMBL/DDBJ databases. RFLP-types highlighted in bold are regarded on the basis of inoculation experiments as being ericoid mycorrhizal endophytes. RL RFLP Accession Putative taxonomic affinity FASTA No. of isolates in each host type code expected value RL1 RL2 RL3 45 AY699671 Ascomycota 8.5e-124 1 0 0 46 AY699678 Ascomycota 5.3e-44 1 0 0 47 AY699691 Ascomycota 1.3e-93 0 6 3 48 AY699692 Ascomycota 3.3e-82 0 2 0 49 AY699693 Ascomycota 5.4e-136 0 3 0 50 AY699694 Ascomycota 2.7e-63 0 2 0 51 AY699695 Ascomycota 2.2e-93 0 1 0 52 AY699696 Ascomycota 2.3e-126 0 0 1 53 AY699697 Ascomycota 1.1e-63 0 0 1 54 AY699698 Ascomycota 6.4e-86 0 0 1 55 AY699699 Ascomycota 1.3e-72 0 0 1 56 AY699700 Ascomycota 2.8e-72 0 0 1 57 AY699701 Ascomycota 9.6e-43 0 0 1 58 AY699702 Basidiomycota 7.6e-28 0 0 1
to 9) had closest ITS sequence affinity to known ericoid mycorrhizal fungi from other northern or southern hemisphere Ericaceae. RL RFLP-types 18 and 19 showed affinity to
H. ericae and known members of the H. ericae aggregate sensu Vrålstad et al. (2002b), while RL RFLP-type 31 had a majority of closest ITS sequence matches to
Oiodiodendron maius and an unidentified fungus, tentatively identified as an
Oidiodendron sp. previously isolated from W. pungens. RL RFLP-type 28 had a majority of closest sequence matches with low expected values to a known DSE, Phialocephela fortinii, and is probably conspecific with this taxon. RL RFLP-types 10-17 had multiple matches with sequences for Xylariaceae, and in the roots of each plant represented a significant portion of the isolate assemblage (ca 23% (RL1), ca 25% (RL2) and ca 16%
(RL3)) (Figure 4.1 and Table 4.3). Other RFLP-types that comprised significant portions 105
of the isolate assemblages were RL RFLP-type 21 (ca 11% of RL1) and RL RFLP-type
23 (ca 11% of RL3), which were putatively designated as Hyaloscyphaceae and
Dermataceae. The remaining isolates were designated as a range of probable ascomycetes or, for RL RFLP-type 58 from RL3, as a putative basidiomycete fungus
(Figure 4.1 and Table 4.3).
4.3.3 Ericoid mycorrhiza formation
Fourteen of the RFLP types in the cultured isolate assemblage were found to form typical ericoid mycorrhizal coils in epidermal cells of V. macrocarpon and were thus regarded as ericoid mycorrhizal endophytes (Figure 4.2). On the basis of ITS sequence comparisons these were identified as Hymenoscyphus sp., Oidiodendron sp., or as belonging to a group of currently unidentified Helotiales ascomycetes of which members are known to form ericoid mycorrhizal associations with Ericaceae plants in temperate environments (Table
4.3). Sequences for selected RFLP-types that were designated as ericoid mycorrhizal fungal endophytes from the three R. lochiae plants (RL RFLP-types 1 to 9) and the three
E. pulchella plants from Chapter 2 (EP RFLP-types 1 to 11 and 49) were further investigated using both neighbour-joining and parsimony analyses. These sequences were supplemented by selected ITS sequences from ericoid mycorrhizal endophytes and their close relatives from the GenBank/EMBL/DDBJ databases. The sequences were chosen based upon the identities of the closest several sequence matches obtained from the
FASTA searches for each of the RFLP-types. These phylogenetic analyses were carried out primarily to determine if any patterns were evident in relation to aspects such as geographical location, host range and habitat type. 106
a b
c d
e f
Figure 4.2 Hair roots of Vaccinium macrocarpon seedlings showing ericoid mycorrhizal coils in the epidermal cells after twelve weeks in gnotobiotic culture with fungal endophyte RFLP-types cultured from Rhododendron lochiae hair roots. a = RL RFLP- type 3, b = RL RFLP-type 4, c = RL RFLP-type 5, d = RL RFLP-type 6, e = RL RFLP- type 7 and f = RL RFLP-type 8. Bars represent 25 µm.
107
g h
i j
k l
Figure 4.2 cont. Hair roots of Vaccinium macrocarpon seedlings showing ericoid mycorrhizal coils in the epidermal cells after twelve weeks in gnotobiotic culture with fungal endophyte RFLP-types cultured from Rhododendron lochiae hair roots. g = RL RFLP-type 9, h = RL RFLP-type 18, i = RL RFLP-type 19, j = RL RFLP-type 29, k = RL RFLP-type 30 and l = RL RFLP-type 31. Bars represent 25 µm.
108
m n
Figure 4.2 cont. Hair roots of Vaccinium macrocarpon seedlings showing ericoid mycorrhizal coils in the epidermal cells after twelve weeks in gnotobiotic culture with fungal endophyte RFLP-types cultured from Rhododendron lochiae hair roots. m = RL RFLP-type 1 and n = RL RFLP-type 2. Bars represent 25 µm.
4.3.4 Phylogenetic analysis of ITS RFLP-types
In the neighbour-joining analysis, (Figure 4.3), most sequences fell within one large group which was separated into seven smaller groups. The exceptions were EP RFLP 11 and an Ericaceae root associated endophyte from Canada which were separated from all other sequences in the moderately supported group VIII (60% bootstrap support). The main large group, (group I), which was only weakly supported (52%), contained RL
RFLP 4, EP RFLP-types 1, 5 and 6, along with sequences from endophytes associated with roots of Australian Ericaceae and Australian ericoid mycorrhizal endophytes. Group
II, which was also weakly supported (50%), contained RL RFLP-types 2, 3, 5, 6, 7, 9, EP
RFLP types 49, 2, 8, 9 and 10, along with sequences from Australian ericoid mycorrhizal 109
endophytes, endophytes associated with the roots of Australian Ericaceae and one
Canadian Ericaceae root associated endophyte. EP RFLP types 3, 4 and 7 clustered in the relatively weakly supported (58%) group III, along with sequences from an endophyte associated with the roots of Australian Ericaceae and three Australian ericoid mycorrhizal endophytes.
Group IV, which contained RL RFLP 1, and group V, which contained RL RFLP 8, were both strongly supported (85% and 90% respectively) and contained sequences from endophytes associated with the roots of Canadian Ericaceae and Canadian ericoid mycorrhizal endophytes. All sequences for mycorrhizal and non-mycorrhizal endophytes associated with the roots of Ericaceae from the UK fell within the well supported groups
VI and VII. There were only a few minor differences in the phylogram produced by the parsimony analysis (Figure 4.4), but with respect to isolates from the present study, RL
RFLP 12 was not grouped separately to the large main group and sequences from the weakly supported group II in the neighbour-joining analysis were split into two separate strongly supported (100%) groups (group IIa and IIb). Sequences for RL RFLP-types 2,
5, 6 and 7 were placed individually outside the main groups.
H. ericae is regarded as a common ericoid mycorrhizal endophyte of many northern hemisphere Ericaceae taxa, however, to date there has been no direct evidence to suggest
H. ericae forms ericoid mycorrhizas in Australian Ericaceae taxa. Since the identities of
110
54 EP RFLP 1 (AY627804) 61 ERM Australia (AF097312) ERM Australia (AF072302) 59 ERM Australia (AF441196) ERM Australia (AF072290) 98 ERM Australia (AF072296) 82 ERM Australia (AF072301) Group I 56 ERM Australia (AF072292) ERM Australia (AF072297) ERE Australia (AY268213) 79 98 EP RFLP 6 (AY627782) EP RFLP 5 (AY627781) 52 80 ERM Australia (AF099091) ERM Australia (AF099979) RL RFLP 4 (AY699667) ERM Australia (AF099090) EP RFLP 49 (AY627805) 99 EP RFLP 2 (AY627806) 54 ERM Australia (AF072298) 73 ERE Australia (AY268186) 51 74 EP RFLP 8 (AY627810) 82 EP RFLP 9 (AY627811) 85 EP RFLP 10(AY627783) Group II 66 100 RL RFLP 9 (AY699684) 98 RL RFLP 3 (AY699666) 50 ERE Canada (AF149076) RL RFLP 7 (AY699681) 89 RL RFLP 2 (AY699665) 97 RL RFLP 5 (AY699668) RL RFLP 6 (AY699670) 70 EP RFLP 4 (AY627808) 56 ERM Australia (AF072295) 60 99 EP RFLP 3 (AY627807) 97 EP RFLP 7 (AY627809) Group III 58 ERM Australia (AF072293) ERM Australia (AF072303) ERE Australia (AY268188) 79 ERE Canada (AF149073) 96 ERE Canada (AF300738) 51 ERE Canada (AF149075) Group IV 85 RL RFLP 1 (AY699664) ERM Canada (AF252839) 100 90 ERM Canada (AF149077) RL RFLP 8 (AY699683) Group V 80 ERM UK (AF252838) ERE UK (AF252850) Group VI 80 ERM UK (AF252844) 90 ERM UK (AF252845) ERM UK (AF252846) 100 ERE UK (AF252848) Group VII ERE UK (AF252847) ERM Canada (AF300737) ERE Australia (AY268217) 60 ERE Canada (AF300739) EP RFLP 11 (AY627784) Group VIII Hymenoscyphus ericae (AF069440) Hymenoscyphus ericae (AF069505) 0.005 substitutions/site
Figure 4.3 Neighbour-joining tree showing the relationships between ITS sequences for selected RFLP-types designated as ericoid mycorrhizal fungal endophytes from the thr ee
Rhododendron lochiae plants, the three Epacris pulchella plants from Chapter 2 as well as related taxa from the GenBank/EMBL/DDBJ databases. Numerical values indicate bootstrap percentiles from 1000 replicates. 111
EP RFLP 1 (AY627804) ERM Australia (AF072290) 100 ERM Australia (AF072296) ERM Australia (AF072301) ERM Australia (AF072292) 100 ERM Australia (AF072297) ERM Australia (AF072302) 100 ERM Australia (AF097312) Group I ERM Australia (AF441196) 100 ERE Australia (AY268213) 100 EP RFLP 6 (AY627782) EP RFLP 5 (AY627781) ERM Australia (AF099091) ERM Australia (AF099979) RL RFLP 4 (AY699667) EP RFLP 49 (AY627805) 100 EP RFLP 2 (AY627806) ERM Australia (AF072298) Group IIa 100 ERE Australia (AY268186) EP RFLP 4 (AY627808) EP RFLP 7 (AY627809) 100 100 ERM Australia (AF072293) Group III ERM Australia (AF072303) EP RFLP 3 (AY627807) ERM Australia (AF072295) EP RFLP 8 (AY627810) EP RFLP 9 (AY627811) 100 EP RFLP 10 (AY627783) 100 RL RFLP 3 (AY699666) Group IIb RL RFLP 9 (AY699684) ERE Canada (AF149076) 100 ERM Australia (AF099090) ERE Canada (AF149073) 64 ERE Canada (AF149075) Group IV 100 ERE Canada (AF300738) RL RFLP 1 (AY699664) 100 ERM Canada (AF149077) RL RFLP 8 (AY699683) Group V 100 ERM UK (AF252838) ERE UK (AF252850) Group VI ERM Canada (AF252839) 72 ERM UK (AF252844) ERM UK (AF252845) 100 ERM UK (AF252846) Group VII ERE UK (AF252847) 74 ERE UK (AF252848) ERM Canada (AF300737) ERE Canada (AF300739) ERE Australia (AY268188) ERE Australia (AY268217) EP RFLP 11 (AY627784) RL RFLP 2 (AY699665) RL RFLP 5 (AY699668) RL RFLP 6 (AY699670) RL RFLP 7 (AY699681) Hymenoscyphus ericae (AF069440) Hymenoscyphus ericae (AF069505) 5 changes
Figure 4.4 Parsimony tree showing the relationships between ITS sequences for selected
RFLP-types designated as ericoid mycorrhizal fungal endophytes from the three Rhododendron lochiae plants, the three Epacris pulchella plants from Chapter 2 as well as related taxa from the GenBank/EMBL/DDBJ databases. Numerical values indicate bootstrap percentiles from
1000 replicates. 112 the closest several sequence matches to RL RFLP 19 obtained from the FASTA search were members of the H. ericae aggregate with close affinity to the type species, the taxonomic position of this RFLP-type was investigated using neighbor-joining and parsimony analyses. Sequences included in the analyses were from other fungal endophytes from the R. lochiae plants (RL RFLP18) and the three E. pulchella plants from Chapter 2 (EP RFLP-types 50 and 12) that had FASTA sequence matches to members of the H. ericae aggregate, along with selected ITS sequences representing members of the H. ericae aggregate and their close relatives from the
GenBank/EMBL/DDBJ databases. The neighbour-joining analysis (Figure 4.5) clustered all sequences except for Hyaloscypha aureliella (Nyl.) Huhtinea in one large weakly supported (53%) group. Within this group, sequences for EP RFLP-types 50, 12 and RL
RFLP 18 clustered in the strongly supported (100%) group I along with those from endophytes isolated from roots of other Australian Ericaceae, two of which have been shown to be ericoid mycorrhizal. The remaining sequences clustered into four other groups with moderate to high support (Groups II – V). The well supported (94%) group
III divided into two well supported sub-groups. One of these, which had 93% support, included sequences for the type species H. ericae and H. ericae-like endophytes, while the other, also with strong support (91%), contained RL RFLP 19 along with sequences for uncultured fungal clones from ectomycorrhizal roots of a non-ericaceous host in a tropical montane cloud forest (TMCF). The parsimony analysis (Figure 4.6) produced a phylogram with similar topology and levels of support to that of the neighbour-joining tree and with respect to the sequences from this study, the same sequences clustered within groups I and III, however group III was only moderately supported with 65%. 113
EP RFLP 12 (AY627813) 72 92 EP RFLP 50 (AY627812)
100 RL RFLP 18 (AY699659) Epacrid root endophyte E1-9 (AF099978) 99 Epacrid root endophyte MG110 (AF441192) 89 Epacrid root endophyte (AY268190) Phialophora finlandia (AF011327) 98 52 Hymenoscyphus sp. G11 (HSP292203) Helotiales sp. (HSP308341) 84 Helotiales sp. (HSP308340) Hymenoscyphus sp. 21 (AF069439) Hymenoscyphus ericae 101 (AF069505) 60 Hymenoscyphus ericae (HER319078) 67 Hymenoscyphus ericae (HER308337) 93 ericoid endophyte (cf.H. ericae) (HER430104) Hymenoscyphus ericae UBCtra274 (AF149069) 53 93 Hymenoscyphus ericae BH (AF069440) 57 84 Hymenoscyphus ericae (HER319077)
94 Hymenoscyphus ericae UBCtra241 (AF149068) 99 TMCF endophyte clone K9-15 (AY394684) 91 TMCF endophyte clone K11-14 (AY394686) 91 TMCF endophyte clone K10-16 (AY394687) 74 TMCF endophyte clone K8-24 (AY394685) RL RFLP 19 (AY699657) 100 Epacris microphylla root endophyte 13 (AY268197) 100 ericoid endophyte (cf.H. ericae) (HER430113)
67 Hymenoscyphus sp. G1(HSP292199) Hymenoscyphus sp. G2 (HSP292200)
82 64 Salal mycorrhizal fungus UBCtra43 (AF149082) 100 Salal mycorrhizal fungus UBCtra323 (AF149083) Salal mycorrhizal fungus UBCtra69 (AF149084) 86 66 Salal mycorrhizal fungus UBCtra56 (AF149085) Ericoid endophyte sp. GU32 (AF252837) Phialocephala fortinii (AY033087) Hyaloscypha aureliella (HAU57495) Sclerotinia borealis (AF067644) Sclerotinia sclerotiorum (M96382) 0.01 substitutions/site
Figure 4.5 Neighbour-joining tree showing the relationships between ITS sequences for selected RFLP-types from the cultured isolate assemblage from Rhododendron lochiae root systems and the three Epacris pulchella plants from Chapter 2 designated as probable members of the Hymenoscyphus ericae aggregate as well as selected sequences for H. ericae and related taxa from the GenBank/EMBL/DDBJ databases. Numerical values indicate bootstrap percentiles from 1000 replicates. 114
62 EP RFLP 14 (AY627813) 96 EP RFLP 50 (AY627812)
100 RL RFLP 18 (AY699659) Epacrid root endophyte E1-9 (AF099978) Group I 91 Epacrid root endophyte MG110 (AF441192) 92 Epacrid root endophyte (AY268190) Phialophora finlandia (AF011327)
74 61Hymenoscyphus sp. G11 (HSP292203) Group II Helotiales sp. (HSP308341) Helotiales sp. (HSP308340) Hymenoscyphus sp. 21 (AF069439)
95 Hymenoscyphus ericae BH (AF069440) Hymenoscyphus ericae (HER319077) Hymenoscyphus ericae 101 (AF069505) 92 54 Hymenoscyphus ericae (HER319078) Hymenoscyphus ericae UBCtra241 (AF149068) Group III Hymenoscyphus ericae UBCtra274 (AF149069) Hymenoscyphus ericae (HER308337) 54 65 100 62 ericoid endophyte (cf.H. ericae) (HER430104)
85 TMCF endophyte clone K9-15 (AY394684) 81 TMCF endophyte clone K11-14 (AY394686) TMCF endophyte clone K8-24 (AY394685) 100 100 TMCF endophyte clone K10-16 (AY394687) RL RFLP 19 (AY699657) Phialocephala fortinii (AY033087) 100 Epacris microphylla root endophyte 13 (AY268197) ericoid endophyte (cf.H. ericae) (HER430113)
66 Hymenoscyphus sp. G1(HSP292199) Hymenoscyphus sp. G2 (HSP292200) Group IV Salal mycorrhizal fungus UBCtra43 (AF149082) 79 100 Salal mycorrhizal fungus UBCtra323 (AF149083) Salal mycorrhizal fungus UBCtra69 (AF149084) Group V 85 77 Salal mycorrhizal fungus UBCtra56 (AF149085) Ericoid endophyte sp. GU32 (AF252837) Hyaloscypha aureliella (HAU57495) Sclerotinia borealis (AF067644) Sclerotinia sclerotiorum (M96382) 10 changes
Figure 4.6 Parsimony tree showing the relationships between ITS sequences for selected RFLP-types from the cultured isolate assemblage from Rhododendron lochiae root systems and the three Epacris pulchella plants from Chapter 2 designated as probable members of the Hymenoscyphus ericae aggregate as well as selected sequences for H. ericae and related taxa from the GenBank/EMBL/DDBJ databases. Numerical values indicate bootstrap percentiles from 1000 replicates.
115
The taxonomic affinities of frequently isolated RFLP-types with FASTA matches with
Xylariaceae (RL RFLP-types 10 to 16) were investigated using neighbor-joining and parsimony analyses. These analyses were conducted on sets of sequences obtained from the GenBank/EMBL/DDBJ databases. In the neighbour-joining analysis of the
Xylariaceae sequences (Figure 4.7), all sequences fell within one large well supported
(100%) group. Within this large group, the well supported (85%) group I clustered all but one of the Xylaria sequences, along with all the Rosellinia sequences except Rosellinia pepo Pat.. RL RFLP-types 10, 11 and 14 formed a strongly supported (100%) sub-group within group I which was sister to three other Xylaria containing sub-groups, while RL
RFLP 16 was located on a separate terminal branch. RL RFLP-types 12 and 13 clustered in the moderately supported (67%) group II along with R. pepo, while the Daldinia and
Hypoxylon sequences formed the well supported (87%) group III. RL RFLP 15 was located separately on a terminal branch. The parsimony analysis of Xylariaceae sequences (Figure 4.8) separated the sequences into two groups that excluded R. pepo.
The large moderately supported (75%) group I contained sequences from the equivalent neighbour-joining analysis groups I and III. Within group I, Daldinia and Hypoxylon sequences clustered as a moderately supported (72%) sub-group along with RL RFLP 15 and X. cubensis while, as was the case in the neighbour-joining analysis, RL RFLP-types
10,11 and 14 formed a strongly supported (99%) sub-group, that was sister to three other
Xylaria containing sub-groups. RL RFLP 16 was located on a separate terminal branch.
RL RFLP-types 12 and 13 clustered separately as the strongly supported (100%) group II in the parsimony phylogram.
116
Rosellinia necatrix (AB017657) 100 Rosellinia necatrix (AB017658) 77 Rosellinia arcuata (AB017660) Rosellinia quercina (AB017661) Xylaria acuta (AF163026) 73 91 Xylaria longipes (AF163038) 76 Xylaria longipes (AF163039) 85 90 Xylaria castorea (AF163030) 91 Xylaria enteroleuca (AF163033) Xylaria fioriana (AF163034) Xylaria apiculata (AF163027) Group I 97 100 Xylaria arbuscula (AF163028) Xylaria arbuscula (AF163029) 54 78 Xylaria hypoxylon (AF163035) 62 100 Xylaria mali (AF163040) Xylaria cornu-damae (AF163031)
93 Xylaria hypoxylon (AF163036) 88 Xylaria hypoxylon (AF163037) 100 Xylaria polymorpha (AF163041) 100 Xylaria polymorpha (AF163042) 100 RL RFLP 10 (AY699645) 100 RL RFLP 14 (AY699649) RL RFLP 11 (AY699646) RL RFLP 16 (AY699675) Rosellinia pepo (AB017659) Group II 67 100 RL RFLP 12 (AY699647) RL RFLP 13 (AY699648) 85 Daldinia concentrica (AF163021) Group III 100 Daldinia sp. (AF163023) 87 HypoxylonDaldinia vernicosa sp. (AF163024) (AF163022)
Xylaria cubensis (AF163032) RL RFLP 15 (AY699660) Cordyceps militaris (AF163020) Nectria cinnabarina (AF163025) 0.01 substitutions/site Figure 4.7 Neighbour-joining tree showing the relationships between ITS sequences for selected RFLP-types from the cultured isolate assemblage from Rhododendron lochiae root systems and selected Xylariaceae sequences from the GenBank/EMBL/DDBJ databases.
Numerical values indicate bootstrap percentiles from 1000 replicates.
117
Rosellinia necatrix (AB017657) 100 Rosellinia necatrix (AB017658) 72 Rosellinia arcuata (AB017660) Rosellinia quercina (AB017661) Daldinia concentrica (AF163021) 97 100 Daldinia sp. (AF163023) 69 Daldinia vernicosa (AF163022) Hypoxylon sp. (AF163024) 72 Xylaria cubensis (AF163032) RL RFLP 15 (AY699660) Xylaria acuta (AF163026) 60 Xylaria longipes (AF163038) 75 77 Xylaria longipes (AF163039) Xylaria castorea (AF163030) 77 Xylaria enteroleuca (AF163033) Group I 75 Xylaria fioriana (AF163034) Xylaria apiculata (AF163027) Xylaria arbuscula (AF163028) 82 100 87 Xylaria arbuscula (AF163029) 100 Xylaria hypoxylon (AF163035) Xylaria mali (AF163040) Xylaria cornu-damae (AF163031)
70 97 Xylaria hypoxylon (AF163036)
97 Xylaria hypoxylon (AF163037)
100 Xylaria polymorpha (AF163041) Xylaria polymorpha (AF163042)
100 RL RFLP 10 (AY699645) 100 99 RL RFLP 14 (AY699649) RL RFLP 11 (AY699646) RL RFLP 16 (AY699675) Rosellinia pepo (AB017659)
100 RL RFLP 12 (AY699647) RL RFLP 13 (AY699648) Group II Cordyceps militaris (AF163020) Nectria cinnabarina (AF163025) 10 changes
Figure 4.8 Parsimony tree showing the relationships between ITS sequences for selected
RFLP-types from the cultured isolate assemblage from Rhododendron lochiae root systems and selected Xylariaceae sequences from the GenBank/EMBL/DDBJ databases. Numerical values indicate bootstrap percentiles from 1000 replicates.
118
4.3.5 DGGE analysis PCR amplification of the ITS1 regions in DNA from pooled root pieces from each root section and representative RFLP-types of the fungi isolated from RL1, RL2 and RL3 yielded products of the expected size (150-300 bp) (data not shown). Despite repeated attempts, amplification of DNA from RL RFLP-types 15, 16, 17, 40 and 58 was unsuccessful. The DGGE banding patterns for DNA from pooled root pieces from each root section from RL1 comprised a total of 32 bands, 25 of which aligned with bands from the cultured fungi from that plant. Similarly, 16 of the 24 bands in the DGGE profiles from RL2, and 18 of the 30 bands in the DGGE profiles from RL3, aligned with bands for cultured fungi (Figure 4.9). Several bands from DNA extracted directly from pooled root pieces that were considered as matches with bands from the cultured assemblages were sequenced to confirm that they represented the same sequence. These sequences were found to be identical, which validated the assumption that bands migrating to the same position represented the same DNA sequence (data not shown).
For all plants, many of the most intense bands observed in DNA extracted directly from pooled root pieces aligned with bands that represented the most frequently isolated
RFLP-types in the cultured isolate assemblages. There were, however, a few notable exceptions. RL RFLP-type 14, a Xylariaceae, represented a significant proportion of isolates in each RL assemblage, but did not match with any band observed in DNA extracted directly from pooled root pieces. Furthermore, bands 1 and 6 from RL2, along with bands 1 and 2 from RL3 (Figure 4.9), which were present in DNA extracted directly from pooled root pieces and did not match with any bands representing isolates in the cultured fungi lanes were intense bands relative to many others in the direct DNA extraction lanes. For each plant, multiple bands (representing RL RFLP-types 7, 10 - 14, 119
34, 43, 44 and 46 in RL1, RL RFLP-types 12, 14, 43, 44, 49 and 50 in RL2 and RL
RFLP-types 14, 43, 44 and 53 - 56 in RL3) were observed in DGGE profiles for cultured fungi, but not in DNA extracted directly from pooled root pieces (Figure 4.9).
All 27 bands from DGGE profiles of DNA from pooled root pieces that were not observed in the cultured endophyte lanes were excised for PCR amplification and sequencing. Of the seven bands sequenced from RL1, four were designated (using the criteria described above) as putative ericoid mycorrhizal endophytes, one as the DSE P. fortinii, one as an ascomycete and one as a probable basidiomycete (Table 4.4). Of the eight bands sequenced from RL2, three were designated as putative ericoid mycorrhizal endophytes, one as the DSE P. fortinii, one as a Trichocomaceae, one as a Rhodotorula sp. and two as ascomycetes (Table 4.4). Of the 12 bands sequenced from RL3, four were designated as putative ericoid mycorrhizal endophytes, one as a Rhodotorula sp., one as a
Mycosphaerellaceae, five as ascomycetes, while one was a probable basidiomycete
(Table 4.4).
DNA extracted from Cultured isolates DNA extracted from Cultured isolates roots roots 120
1231 23456 78 123 12345 6 7 * * 6 5 * 6 *
* * * * * 3 * * 1 * * 1 4 2 2 3 * 8 5 4
7 * * a b
DNA extracted from Cultured isolates roots
123 12 345
6
9
5
3 4 * 1
2 * * * 8 * 10
12 11 c 7
Figure 4.9 DGGE profiles of ITS1 sequences amplified from DNA extracted directly from roots or cultured fungal isolates from Rhododendron lochiea plants RL1 (a), RL2 (b) and RL3 (c). Each lane for DNA extracted from roots represents DNA from pooled root pieces from a root section, while each cultured isolate lane represents mixed ITS1 PCR products from several cultured isolates. Numbered bands are unique bands in profiles for DNA extracted from roots that were sequenced (numbers correspond to those in Table 4.4). Bands that were present in cultured isolate profiles, but absent from those for DNA extracted from roots are marked with an asterisk. 121
Table 4.4 Putative taxonomic affinities of ITS sequences from selected DGGE bands amplified from DNA extracted directly from root systems of three Rhododendron lochiae root plants (RL1, RL2 and RL3) as inferred from FASTA searches of ITS sequences in the GenBank/EMBL/DDBJ databases. RFLP-types highlighted in bold are regarded on the basis of FASTA matches to be probable ericoid mycorrhizal endophytes.
DGGE Accession Putative taxonomic affinity FASTA Band code expected value RL1 (1) AY823033 ericoid mycorrhizal endophyte 1.8e-51 RL1 (2) AY823034 ericoid mycorrhizal endophyte 1.1e-54 RL1 (3) AY823035 Helotiales 3.2e-43 RL1 (4) AY823036 Hymenoscyphus ericae 2.2e-50 RL1 (5) AY823037 Phialocephala fortinii 7.2e-56 RL1 (6) AY823038 Ascomycota 5.9e-23 RL1 (7) AY823039 Basidiomycota 1.4e-12 RL2 (1) AY823040 ericoid mycorrhizal endophyte 6.8e-55 RL2 (2) AY823041 ericoid mycorrhizal endophyte 2.1e-51 RL2 (3) AY823042 Helotiales 3.9e-43 RL2 (4) AY823043 Phialocephala fortinii 3.6e-55 RL2 (5) AY823044 Trichocomaceae 2.8e-50 RL2 (6) AY823045 Rhodotorula sp. 4.0e-27 RL2 (7) AY823046 Ascomycota 7.9e-24 RL2 (8) AY823047 Ascomycota 4.1e-25 RL3 (1) AY823048 ericoid mycorrhizal endophyte 6.9e-55 RL3 (2) AY823049 ericoid mycorrhizal endophyte 1.2e-51 RL3 (3) AY823050 ericoid mycorrhizal endophyte 1.4e-51 RL3 (4) AY823051 ericoid mycorrhizal endophyte 2.9e-50 RL3 (5) AY823052 Rhodotorula sp. 9.3e-26 RL3 (6) AY823053 Mycosphaerellaceae 8.6e-28 RL3 (7) AY823054 Ascomycota 6.4e-27 RL3 (8) AY823055 Ascomycota 7.8e-28 RL3 (9) AY823056 Ascomycota 8.0e-25 RL3 (10) AY823057 Ascomycota 4.9e-15 RL3 (11) AY823058 Ascomycota 6.2e-07 RL3 (12) AY823059 Basidiomycota 4.8e-14
122
4.4 Discussion
Although the presence of ericoid mycorrhizal fungal infection in roots of tropical
Ericaceae taxa has been reported previously (Bermudes and Benzig, 1989; Lesica and
Antibus, 1990; Rains et al., 2003), the work described in this chapter represents the first attempt to isolate and infer the taxonomic affinities of the endophytes. These data indicate that root systems of R. lochiae in an Australian tropical montane cloud forest harbour diverse assemblages of ericoid mycorrhizal endophytes and other root associated fungi. While some fungal taxa were common to the root systems of the three R. lochiae plants investigated, others were present only in a single plant, and the relative abundance of fungal taxa varied between individual plants.
ITS RFLP analysis indicated that the cultured isolate assemblage from RL1 comprised a greater number of RFLP-types (= putative taxa) than either of the other R. lochiae plants.
Furthermore, over half of the RFLP-types from RL1 were unique to that plant. The cultured isolate assemblages from RL2 and RL3 comprised similar numbers of RFLP- types (= putative taxa), and about a third of the RFLP-types from each plant were unique.
RL1 was collected from a sheltered understorey site while RL2 and RL3 were collected approximately 50 m away on a more exposed rock ledge site. It is reasonable to suggest that subtle differences in edaphic conditions between collection sites may have contributed to unique endophytes being obtained from each plant. A greater number of
RFLP-types were cultured from the R. lochiae plants than the E. pulchella plants investigated in Chapter 2. However, levels of recovery of fungal isolates from root pieces were similar for the two Ericaceae hosts, but fewer root pieces were sampled from 123 the E. pulchella plants. Thus, if more had been taken, similar numbers of RFLP-types may have been observed for each plant taxon. Alternatively, it may be the case that the edaphic conditions at the TMCF Mount Lewis Forest Reserve site are more conducive than the sclerophyl forests sites of Eastern Australia in terms of enhancing the diversity of fungi associated with the roots of ericaceous hosts. Further investigation of both the edaphic conditions at each of the field sites, along with the root morphology and anatomy of R. lochiae and E. pulchella, is required to resolve the issue of variation in fungal endophyte isolation from these hosts.
The R. lochiae plants were colonised by a few dominant RFLP-types and a greater number of less frequently isolated types, which is consistent with results for the relative abundance of root endophytes for the E. pulchella plants in Chapter 2, along with other investigations of Australian and northern hemisphere Ericaceae taxa (Perotto et al., 1996;
Xiao and Berch 1996; Chambers et al., 2000; Sharples et al., 2000; Liu et al., 1998;
Midgley et al., 2002; 2004c). Many of the RL RFLP-types that were represented by the largest numbers of cultured isolates (RL RFLP-types 1, 2, 3, 4, 5 and 9) were identified as ericoid mycorrhizal endophytes.
Of the 58 RFLP types isolated from the three R. lochiae plants, 14 were found to form typical ericoid mycorrhizal structures in roots of V. macrocarpon under gnotobiotic conditions, and are thus regarded as representing ericoid mycorrhizal endophytes. V. macrocarpon was used to test for mycorrhiza-forming ability because seeds of R. lochiae were not available. Ericoid mycorrhizal fungi are thought to show little, if any, 124 specificity for different Ericaceae hosts (Cairney and Ashford, 2002). The observation that all of the RFLP types from R. lochiae that ITS sequence comparisons suggested were ericoid mycorrhizal endophytes formed ericoid mycorrhiza with V. macrocarpon provides support for this idea. On this basis, ca 41–51% of the cultured isolate assemblage from each R. lochiae plant were ericoid mycorrhizal endophytes. The majority of the ericoid mycorrhizal endophytes had strong ITS sequence similarity to unidentified ericoid mycorrhizal endophytes that have been isolated from roots of
Ericaecae in temperate habitats of Australia, North America and Europe (eg. Sharples et al., 2000; Berch et al., 2002; Bougoure and Cairney, 2005). Although currently unnamed, these endophytes appear to belong to several groups of Helotiales ascomycetes (Berch et al., 2002; Cairney and Ashford, 2002), and these observations indicate that they are also important ericoid mycorrhizal endophytes of tropical Ericaceae. The habitats from which these Helotiales-like fungi have been obtained, although having in common some degree of edaphic stresses such as limited nutrient availability, show distinct differences, such as soil moisture levels, soil organic matter content and temperature. These fungi, like H. ericae, are likely to posses a broad suite of enzymes that ensure a degree of benefit to the host plants, although this is yet to be determined.
Phylogenetic analyses of sequences for this group of Helotiales-like fungi confirmed that there may be limited specificity in terms of geographic location, habitat type and host range. Many of the groups identified in both neighbour-joining and parsimony analyses included sequences for fungi from varied geographic locations, habitat types and host plant taxa. For example, sequences for fungi isolated from R. lochiae in TMCF and E. 125 pulchella from a sclerophyll forest in this study, along with fungi isolated from other
Australian Ericaceae and Gaultheria shallon from temperate woodland in Canada, grouped together. However, sequences for fungi isolated from Calluna vulgaris inhabiting mor humus heathlands in the United Kingdom grouped separately to all other sequences in both analyses, suggesting these fungal isolates may be specific to the mor humus heathlands of the UK and/or C. vulgaris as a host. Based upon the observations of the present and other studies, it seems reasonable to suggest that this group of ericoid mycorrhizal Helotiales-like fungi is as common, and likely to be just as beneficial to their hosts, as members of the H. ericae aggregate or Oidiodendron spp.
H. ericae and related taxa of the H. ericae aggregate are common ericoid mycorrhizal and ectomycorrhizal partners of Northern Hemisphere plants (Vrålstad et al., 2002;
Villarreal-Ruiz et al., 2004). Although isolates with strong ITS sequence identity to H. ericae sensu stricto have been obtained from liverwort rhizoids in Australia and
Antarctica (Chambers et al., 1999), Hymenoscyphus-like isolates that have previously been isolated from Australian Ericaceae have been grouped separately to the H. ericae type isolate and related genotypes in phylogenetic analyses (Williams et al., 2004). This was confirmed in the phylogenetic analyses conducted during the present study, however,
RFLP type 19 from R. lochiae grouped strongly with the H. ericae type isolate and related genotypes. This establishes that such isolates, although uncommon in the cultured isolate assemblages from R. lochiae, are, in fact, ericoid mycorrhizal partners of
Australian Ericaceae. Moreover, RL RFLP type 19 formed a well-supported monophyletic terminal group with sequences cloned from ectomycorrhizal roots of G. 126 emarginata from a tropical cloud forest in Ecuador. These fungi have been suggested to represent a distinct ectomycorrhiza- forming taxon within the H. ericae aggregate (Haug et al., 2004). These data not only suggest that the taxon may be widespread in tropical cloud forests, but also suggest that it contains ericoid mycorrhiza-forming as well as ectomycorrhizal isolates. These data also provide support for the suggestion that ericoid- and ecto- mycorrhizal plants may share some common symbiotic partners in the H. ericae aggregate (Vrålstad et al., 2000) (See Chapter 5).
A significant proportion (16–23%) of the cultured isolate assemblage from each of the R. lochiae root systems was identified as Xylariaceae taxa. Although many putative
Sordariales isolates have been obtained from roots of other Ericaceae taxa in temperate habitats in Australia and elsewhere, putative Xylariaceae isolates have only rarely been reported (Berch et al., 2002; Williams et al., 2004). Xylariaceae are, however, commonly found as endophytes of plant parts, including roots, in tropical habitats (eg.
Whalley, 1996; Bayman et al., 1997), and the observations from the present study suggest that taxa within this family are also common endophytes of at least one tropical Ericaceae species in Australia. A single Xylariales-like isolate obtained from G. shallon was shown to form typical ericoid mycorrhizal coils when reinoculated into the host (Berch et al.,
2002), however in the present study none of the Xylariales-like isolates formed any structures resembling ericoid mycorrhizal coils in V. macrocarpon roots. This, however, does not necessarily preclude the possibility that these isolates might represent ericoid mycorrhizal fungi, and inoculation experiments using Rhododendron or a more closely 127 related host may be more appropriate for assigning mycorrhizal status to this group of fungi in the future.
Allen et al. (2003) found that partial rDNA internal transcribed spacer (ITS) sequences from endophytes cultured from roots of G. shallon (Ericaceae) were markedly different to sequences cloned from DNA extracted directly from the same root systems. Specifically,
>50% of the cloned sequences represented Sebacina-like basidiomycetes, a group that was absent from the cultured isolate assemblage. In contrast, the results from Chapter 2 indicated that cultured isolate assemblages from roots of the Australian Ericaceae taxon
E. pulchella were broadly similar to those predicted from cloned sequences and DGGE profiles from directly extracted DNA. In common with the results presented in Chapter 2, several bands were found to be present in DGGE profiles of DNA extracted from R. lochiae root pieces that were not present in profiles for cultured isolates. Sequence comparisons indicated that these arose from putative ericoid mycorrhizal fungi, along with other ascomycetes and basidiomycetes. While it is possible that some of these may represent unculturable root-associated fungi, it is equally likely that their absence from the cultured isolate assemblage reflects their relatively low abundance on the community.
Several cultured isolate RFLP types from E. pulchella in Chapter 2 were not present in the directly extracted DNA profiles, however, these were restricted to those that were represented by only one or a few isolates in the cultured isolate assemblages. Similarly, several RFLP types from the cultured isolate assemblage from R. lochiae in the present study were not observed in DGGE profiles of DNA extracted from root pieces. In all but 128 one case these RFLP types were represented by only one or a few isolates in the cultured isolate assemblages, and their absence from the DGGE profiles of DNA extracted from root pieces presumably reflects the relative scarcity of their DNA in the total DNA pool.
The exception was RFLP type 14, a Xylariaceae taxon that constituted c. 15–25% of the cultured isolate assemblage from each R. lochiae root system. The absence of this RFLP type from the DGGE profiles of DNA extracted from root pieces may indicate that, although widespread in the root systems, only a relatively small amount of mycelium was present. This is supported by the fact that ericoid mycorrhizal endophytes form dense hyphal coils in multiple cortical cells (Cairney and Ashford, 2002) and, as such, might be expected to comprise a greater proportion of the overall fungal biomass in R. lochiae roots than non-mycorrhizal fungi such as RFLP type 14. In addition, this RFLP type was observed to grow considerably faster in culture than most of the other fungi isolated (data not shown), and it is possible that its rapid growth, and the concomitant suppression of other fungi during isolation may have resulted in an overestimation of its importance in the root-dwelling fungal assemblage. This phenomenon has been observed by other authors (eg. Hambleton and Currah, 1997) and highlights one potential limitation of the cultured isolate approach to investigating communities of root-dwelling fungi.
The data from the present investigation suggest that the fungi that form ericoid mycorrhizal associations with R. lochiae in an Australian tropical montane cloud forest habitat belong to similar taxonomic groupings to those already identified as ericoid mycorrhizal partners of Ericaceae in temperate zones of Australia and other parts of the world. Whereas results from Chapter 2 concluded that culture based methods and culture- 129 independent DNA methods were similarly efficacious in their abilities to identify the most abundant members of root-associated fungal communities, the data in this chapter, along with those of Allen et al. (2003), indicate that this is not always true. Given that both culture-based and culture-independent methods introduce potential bias (Bridge and
Spooner, 2001; Anderson and Cairney, 2004), a combination of both types of approach should probably be adopted in future investigations of this nature.
130
CHAPTER 5
Assemblages of ericoid mycorrhizal and other root-associated fungi from Ericaceae
across a moorland : forest gradient in North-East Scotland
5.1 Introduction
In the past, ericoid mycorrhizal and ectomycorrhizal symbioses were generally thought to involve distinct groups of fungi. Ectomycorrhizal relationships were regarded as predominately involving a broad taxonomic range of basidiomycetes, while ericoid mycorrhizal fungal partners have been identified as ascomycetes, including members of the Hymenoscyphus ericae aggregate, Oidiodendron spp. and other unidentified
Helotiales–like fungi (Smith and Read, 1997). Recent advances in molecular techniques have revealed that belowground communities of ectomycorrhizal fungi are far more complex than previously thought (Horton and Bruns, 2001). Furthermore, these techniques have resulted in a significant increase in the number and variety of fungal taxa that are known to form mycorrhizal symbioses. One of the most significant advances from the molecular approaches has been the realization that multiple ascomycete taxa are ectomycorrhizal fungal symbionts (Horton and Bruns, 2001).
Piceirhiza bicolorata is a common ectomycorrhizal morphotype identified on various host species from boreal and temperate forests of the northern hemisphere (Vrålstad et al., 2000). Analysis of ITS sequence data have indicated that the fungi that form this ectomycorrhizal morphotype are related to the ericoid mycorrhizal fungus H. ericae 131 blurring the established distinction between ectomycorrhizal and ericoid mycorrhizal fungi (Vrålstad et al., 2000). Indeed, many fungi that have been isolated from both ericoid mycorrhizal and ectomycorrhizal associations worldwide are now regarded as comprising part of the H. ericae aggregate (eg. Vrålstad et al., 2000; 2002b; Allen et al.,
2003; Rosling et al., 2003 and Haug et al., 2004). Vrålstad et al. (2002b) conducted a phylogenetic study of rDNA ITS sequences from the H. ericae aggregate and found that they clustered within four well-supported clades. Clades I and III contained predominantly, but not exclusively, fungi isolated from or detected in the roots of
Ericaceae in Europe and North America, and included the type isolate H. ericae and its putative anamorph Scytalidium vaccini (Read) Korf and Kernan (Pearson and Read,
1973; Read, 1974; Egger and Sigler, 1993). In contrast, the majority of fungi that fell within clades II and IV had been detected or isolated from roots of ectomycorrhizal hosts in Scandinavia (Wang and Wilcox, 1985; Vrålstad et al., 2002b; Rosling et al., 2003).
Recently Hambleton and Sigler (2005) proposed species names corresponding to three of the H. ericae aggregate clades identified by Vrålstad et al. (2002b). Thus, Hambleton and Sigler (2005) proposed that members of clade I represent Meliniomyces variabilis
Hambleton and Sigler sp. nov., clade II represent Meliniomyces vraolstadiae Hambleton and Sigler sp. nov. and several clade IV isolates represent Meliniomyces bicolor
Hambleton and Sigler sp. nov. Since this nomenclature has only recently been proposed and has not yet gained broad acceptance, the clade numbers used by Vrålstad et al.
(2002b) have been retained here.
Vrålstad et al. (2000) suggested that ericoid mycorrhizal and ectomycorrhizal plants may share some common symbiotic partners in the H. ericae aggregate. This idea is supported 132 by the fact that a strain of Cadophora finlandia, (= Phialophora finlandia), a member of clade IV, was shown to form ectomycorrhizal associations with woody hosts (Wilcox and
Wang, 1987). Monreal et al. (1999) subsequently indicated that the same strain formed typical ericoid mycorrhiza with Gaultheria shallon in ‘limited trials'. Furthermore, genetically identical isolates from the H. ericae aggregate have been found in both ectomycorrhizal and ericoid mycorrhizal roots in two instances. A single genotype from clade I was found to associate with the ectomycorrhizal host Quercus robur L. and the ericoid mycorrhizal host Calluna vulgaris, while a genotype from clade IV was isolated from the ectomycorrhizal host Salix herbaceae L. along with the ericoid mycorrhizal host
Vaccinium vitis-idaea L. (Vrålstad et al., 2002b). Moreover, in Chapter 4, RL RFLP-type
19, was shown on the basis of ITS sequence analysis to be closely related to sequences cloned from ectomycorrhizal roots of Graffenrieda emarginata from a tropical cloud forest in Ecuador and also formed typical ericoid mycorrhizal coils when used in colonisation trials. The fungi from G. emarginata roots have been suggested to represent a distinct ectomycorrhiza-forming taxon within the H. ericae aggregate (Haug et al.,
2004), however the data in Chapter 4 suggest that the taxon also contains ericoid mycorrhiza-forming isolates. These observations, along with the results of Bergero et al.
(2000), who found that ectomycorrhizal roots of Quercus ilex L. harboured fungi capable of forming typical ericoid mycorrhizas with Erica arborea, provide additional support for the suggestion that ericoid mycorrhizal and ectomycorrhizal plants may share some common symbiotic partners in the H. ericae aggregate (Vrålstad et al., 2000).
In order to test this hypothesis, Vrålstad et al. (2002a) performed inoculation trials between selected isolates of the H. ericae aggregate and potential ectomycorrhizal and 133 ericoid mycorrhizal plant hosts. Ectomycorrhiza formation was generally restricted to isolates of ectomycorrhizal host origin which grouped in clade IV along with C. finlandia. None of the ectomycorrhiza-forming isolates produced any structures that resembled ericoid mycorrhizas when inoculated onto V. vitis-idaea. In contrast, isolates from clade III formed typical ericoid mycorrhizal structures when inoculated onto V. vitis-idaea hosts, but were unable to form ectomycorrhizal associations (Vrålstad et al.,
2002a). The data confirmed that both the ericoid and the ectomycorrhizal condition have evolved within the H. ericae aggregate, but provided no support for the suggestion that a single genotype could form both ectomycorrhizas and ericoid mycorrhizas.
More recently, Villarreal-Ruiz et al. (2004) demonstrated that a single C. finlandia–like clade IV isolate from a P. bicolorata ectomycorrhizal root tip produced typical ericoid mycorrhizal coils in epidermal cells of V. myrtillus and an ectomycorrhizal Hartig net and mantle with Pinus sylvestris. Furthermore, fungal colonisation induced a substantial growth response in V. myrtillus roots, and no sign of pathogenic reactions were observed in either of the host plants. These data represent substantial evidence that a single isolate from the H. ericae aggregate can form both ectomycorrhizal and ericoid mycorrhizal associations. Based on these observations, Villarreal-Ruiz et al. (2004) suggested that isolates that fall within clade III of the H. ericae aggregate may have evolved as obligate ericoid mycorrhizal endophytes in moorlands, but those members of clade IV (including
C. finlandia) are primarily ectomycorrhizal fungi. They further suggested that clade IV isolates may have the potential to be ericoid mycorrhizal symbionts in woodlands where plant communities comprise both Ericaceae and ectomycorrhizal host plants. 134
Despite the fact that Ericaceae are a common component of the understorey of many temperate and boreal conifer and broadleaf forest habitats worldwide, investigations of ericoid mycorrhizal fungal diversity in these regions have largely focused on heathlands where they represent the dominant taxa. Conversely, the equivalent work on ectomycorrhizal fungi has focused largely on forest habitats. Temperate and boreal forest and heathland habitats are characterised by substantial accumulation of organic matter due to low temperatures and recalcitrant leaf litter that is rich in polyphenolic compounds
(Lindahl et al., 2002). Consequently, nutrient and carbon cycling depend heavily on the activities of saprotrophic and mycorrhizal fungi (Lindahl et al., 2002; Read and Perez-
Moreno, 2003). Recent studies have revealed that the diversity and likely functional impact of ectomycorrhizal fungi in these forests is large (eg. Taylor et al., 2000; Horton and Bruns, 2001). However, despite the abundance of Ericaceae hosts as understorey vegetation, relatively little is known about the fungi involved in these ericoid mycorrhizal relationships of forest dwelling Ericaceae. In the only study to have considered this so far, a variety of ericoid mycorrhizal fungal genotypes were isolated from forest Ericaceae hosts and, although these fell largely within clades I and III of the H. ericae aggregate, some isolates formed part of clade IV (Vrålstad et al., 2002b).
The C. finlandia-like isolate that was shown by Villarreal-Ruiz et al. (2004) to form both ectomycorrhizal and ericoid mycorrhizal associations was isolated from a P. sylvestris root tip from a Caledonian forest in north east Scotland. In such a habitat, where light might be a limiting factor for understorey vegetation, the possibility that ectomycorrhizal trees and Ericaceae shrubs may be interconnected via a common mycelium is of interest.
As demonstrated to occur between ectomycorrhizal plants in the field (Simard et al., 135
1997), such a fungal link might provide a conduit for transfer of carbon between the coniferous overstorey and Ericaceae understorey (Vrålstad, 2004).
Caledonian forests that are dominated by P. sylvestris, are an extremely rare, indigenous habitat in the UK, with remnants distributed sparsely throughout the central and north- eastern Grampians, along with the northern and western Highlands of Scotland. These boreal forests represent a precious natural resource of high biodiversity value and consequently current UK policy is to encourage their expansion (Anderson et al., 2003).
While expansion of these forests is being encouraged, little is known regarding how below-ground microbial diversity that is associated with bordering ecosystems might affect the expansion process, however it is likely to aid the natural regeneration process.
In Scotland, much of the Caledonian forest expansion occurs on highly organic soils that support C. vulgaris-dominated moorland plant communities (Anderson et al., 2003).
Anderson et al. (2003) investigated soil fungal community structure and diversity using
ITS-DGGE profiles along a gradient from moorland into an expanding P. sylvestris forest in north east Scotland. Whereas the extremes of the transects represented mature P. sylvestris forest with an Ericaceae understorey and moorland vegetation, the intermediate region was characterised by the presence of colonizing P. sylvestris saplings, and represented a zone of transition between the two habitat types. The authors hypothesised that a gradual change in the soil fungal community might occur in the transition zone as a result of colonisation by the P. sylvestris saplings in conjunction with their associated ectomycorrhizal fungi. However, the data indicated that the presence of the saplings in the transition zone had little effect on soil fungal community structure (Anderson et al., 136
2003). The ITS-DGGE profiles revealed a sharp boundary between the mature forest and transition zone, the latter having similar profiles to the established moorland.
Anderson et al. (2003) suggested that changes in the ericaceous vegetation structure, along with ecological host specificity for symbionts between the forest and moorland ends of the transects, may play a significant role in influencing the soil fungal community. Supporting this idea is the suggestion of Villarreal-Ruiz et al. (2004) that members of clade IV of the H. ericae aggregate are primarily ectomycorrhizal fungi, and may have the potential to be ericoid mycorrhizal symbionts with only forest understorey
Ericaceae hosts, while members of clade III may have evolved as obligate moorland symbionts. If this were the case, then it is possible that at least some of the variation observed between DGGE profiles reflected the presence of clade IV ectomycorrhizal fungi and clade III members of the H. ericae aggregate in the samples from the forest and moorland ends of the transects respectively.
The aim of the work described in this chapter was to investigate and compare the community structure of fungi associated with roots of C. vulgaris and V. myrtillus in mature Caledonian forest, along with C. vulgaris from a bordering transition zone with developing P. sylvestris saplings and an open moorland. This investigation was carried out in order to test the hypotheses that (i) isolates from clade IV of the H. ericae aggregate are present as mycorrhizal symbionts in Ericaceae hosts and (ii) that these isolates are more prevalent as ericoid mycorrhizal symbionts in understorey hosts rather than in transitional zone and moorland hosts. DGGE and T-RFLP were used to compare fungal community profiles between habitat type and host species. Cloning was also 137 carried out to further investigate the fungal community profiles from a taxonomic perspective with particular emphasis placed on sequences representing members of the H. ericae aggregate.
5.2 Experimental procedures
5.2.1 Field site description and sampling
The field site for this work was at Abernethy forest, Cairngorm, Scotland (National grid reference NJ027122) and sampling was carried out in March 2005. Three 80 m transects that were roughly parallel to each other and approximately 50 m apart were established at the field site. The transects ran in an approximately south-west – north-east direction.
One transect was located well within the boundary of a mature P. sylvestris forest, while another fell within an intermediate zone of moorland where there was sapling colonisation by P. sylvestris that, at the time of sampling, was 10–20 years old. The third transect was located solely within an area of open moorland. Each transect was divided into 10 sections with 11 points in total, 8 m apart starting from point 1 at the north-east end of the transect. Vegetation at the moorland and intermediate zone transects was dominated by C. vulgaris, while the forest transect was dominated by a sparse coverage of V. myrtillus and C. vulgaris. At each transect point a C. vulgaris plant, including the root system, was removed with a spade. Along the forest transect, V. myrtillus plants were also similarly removed at each point, giving two sampling sets from this transect
(FC and FV) and one each from the other two (TC and MC). In cases where no plant was located at the sampling point, which was frequently the case in the forest transect, the 138 closest plant to the sampling point was collected and in each case was no more than 2 m from the sampling point. Samples were transported to the laboratory and stored at 4oC overnight. From each plant collected, a portion of hair root material was carefully sorted and washed to remove adhering soil and organic matter. Hair roots were surface sterilised
-1 in a 100% commercial bleach solution (4.5% available chlorine) containing 100 μl l of
Tween 20™ (Amersham Biosciences) for 30 sec, followed by 30 sec in a 70% ethanol solution and three one minute rinses in sterile Milli-Q® water before being frozen at -
20oC .
5.2.2 DNA extraction
DNA was extracted from hair root material from each of the sampled plants using liquid nitrogen grinding and the DNeasy mini® extraction kit (Qiagen) following the manufacturer’s instructions. Approximately 200 mg of hair root material was ground using a micropestle in a 1.5 ml microcentrifuge tube with liquid nitrogen and the ground powder was immediately placed into 2.0 ml screw-cap tubes which included Lysing
Matrix E (Qbiogene) and 1.0 ml of Lysing Buffer AP1. The tubes were then placed into the FastPrep® Instrument (Bio 101) and homogenised at speed 5.0 m s-1 for 45 sec then incubated at 65oC for 10 min. To each tube, 140 µl of buffer AP2 was added and tubes were incubated for five min on ice. The lysate was applied to QIAshredder Mini Spin
Columns placed in 2.0 ml collection tubes and centrifuged for 2 min at 14000 rpm. The flow-through fraction was transferred to a 1.5 ml Eppendorf tube without disturbing the cell-debris pellet and 1.5 volumes of Buffer AP3 was added to the cleared lysate and mixed by pipetting. This mixture, including any precipitate which may have formed, was 139 applied to DNeasy Mini Spin Columns sitting in 2.0 ml collection tubes and this mixture was centrifuged for one min at 8000 rpm and flow-through was discarded. DNeasy Mini
Spin Columns were placed in new 2.0 ml collection tubes and 500 µl of buffer AW was added to the DNeasy Mini Spin Columns and centrifuged for one min at 8000 rpm with flow-through discarded. This step was repeated once. DNeasy Mini Spin Columns were transferred to 1.5 ml microcentrifuge tubes and 100 µl of Buffer AE was pipetted directly onto the DNeasy membrane. This mixture was incubated for 5 min at room temperature and then centrifuged for 1 min at 8000 rpm to elute extracted DNA.
5.2.3 ITS amplification
A first round PCR was carried out to amplify the rDNA ITS region from fungal endophytes colonising the sampled hair roots. Amplifications were carried out in 50 μl reaction volumes containing approximately 50 ng of template DNA; 20 pmol of each of the primers ITS1F and ITS4 (White et al., 1990); 2.0 mM MgCl2; 250 µM of each of dATP, dCTP,dGTP and dTTP; 5.0 µl 10x reaction buffer; 1.0 µl bovine serum albumin
(BSA) and 0.75 µl of High Fidelity Taq polymerase (Roche). Amplifications were performed on a Dyad DNA engine thermal cycler (MJ Research) with an initial 5 min stage at 95οC followed by 30 cycles of 95oC for 30 sec, 55oC for 30 sec and 72oC for 1 min, followed by a final extension at 72οC for 10 min. Amplification products were electrophoresed in 1.5 % (w/v) agarose gels, stained with ethidium bromide and visualised under UV light. These products were then used as templates (0.5 µl) in a nested PCR to generate products for DGGE analysis. DGGE products were amplified using 20 pmol of each of the primers ITS1F and ITS2 (Gardes and Bruns, 1993) with a 140
40 bp GC clamp attatched to the 5’ end of the ITS1F primer to stabilise the melting behaviour of the DNA fragments. PCR and cycling conditions were as described above except that 0.5 µl of BIOTAQ™ polymerase (Bioline) was used in each reaction and
BSA was omitted from the reactions. To generate products for T-RFLP analysis, the rDNA ITS region from fungal endophytes colonising the sampled hair roots was amplified using the same reaction mixture as for the nested PCR described above except that 20 pmol of each of the primers ITS1F and ITS4 (White et al., 1990) were used. The primers were labelled at their 5’ ends with fluorescent dyes HEX and 6-FAM, respectively. The cycling parameters were as described above.
5.2.4 DGGE analysis
DGGE analyses were carried out using the DCode universal mutation detection system
(Bio-Rad). Polyacrylamide gels (8% Acrylogel 2.6 solution (BDH)) were prepared with a 20% (8% (v/v) formamide and 1.4 M urea) to 60% (24% (v/v) formamide and 4.2 M urea) vertical denaturing gradient using a gradient former (Fisher Scientific) and a peristaltic pump with a flow rate of 5 ml min-1. Gels were poured onto the hydrophilic side of GelBond PAG film (BioWhittaker) to facilitate handling during the staining process. ITS-PCR products (1.0 µl) and a combination DNA marker (0.5µl Hyperladder
V (Bioline) and 0.5 µl Phix Hinf1 (Promega)) were loaded onto the gels and electrophoresis was performed in 1x TAE buffer (40 mM tris, 20 mM glacial acetic acid,
1.0 mM EDTA) at 75 V for 16 h at a constant temperature of 60ºC.
141
Silver staining was employed for visualising the bands. The gels were gently shaken for
2 h in a fixing solution containing 10% v/v ethanol and 0.5% v/v glacial acetic acid. After removal of the fixing solution, gels were gently shaken in the staining solution (0.1% w/v
AgNO3) for 20 min. Following a brief rinse in tap water, gels were shaken in developing solution (1.5% w/v NaOH, 0.4% v/v formaldehyde, 0.01% w/v NaBH4) until bands became just visible. After another brief tap water rinse, gels were shaken in a final fixing solution (0.75% w/v Na2CO3) for 10 min before being scanned. Digitalised images of
DGGE gels were analysed using GelCompare II software to compare community profiles of C. vulgaris endophytes between transects as well as community profiles of C. vulgaris and V. myrtillus endophytes from transect A. Band matching, using GelCompare II, was performed with 7% minimum profiling and cluster analysis using the Jaccard algorithm with 1% optimisation and 1% position tolerance. Using the same parameters a composite data set was used to produce a binary table based on only the presence or absence and not intensity of bands in each sample lane. This binary information was used for further data analysis.
5.2.5 T-RFLP analysis
Prior to T-RFLP analysis the fluorescently labelled ITS products were purified using the magnetic bead based ChargeSwitch® purification kit (Invitrogen) as per manufacturer’s instructions. Two restriction endonuclease enzymes (Hinf1 and Taq1 (Promega)) were used, and digestions were performed in 96 well plates with each 10 µl reaction containing
0.5 µl restriction enzyme, 1.0 µl buffer, 0.1 µl BSA and 1.0 µl PCR product. The Hinf1 digests were incubated for 4 h at 37°C while the Taq1 digests were incubated for 4 h at 142
65°C. Mixtures containing 2.0 µl of each of the digest products, 0.5 µl of GeneScan 500-
ROX size standard (Applied Biosystems) and 12 µl Hi-Di Formamide (Applied
Biosystems) were denatured at 95°C for 5 min then cooled on ice before being applied to an ABI 3130xl sequencer. Digested amplicons were separated along a 50 cm column using the POP4 polymer (Applied Biosystems). Terminal restriction fragment (T-RF) lengths were determined by Genescan® software (Applied Biosystems) and peaks below
50 base pairs and above 500 base pairs were ignored. Based on the Genescan® output a binary table was constructed including each sample point from all four sampling transects using both restriction enzymes and both fluorescently labelled T-RFs to indicate only the presence or absence of T-RFs between 50 and 500 base pairs and not their peak heights
(fluorescence intensity). This binary information was used for further data analysis.
5.2.6 Cloning and sequence analysis
ITS-PCR products generated with the unlabelled ITS1F and ITS4 primer pair were cloned with the pGEM-T Easy vector system (Promega). In order to make the number of clones more manageable, the ITS products generated from each sample point were pooled prior to cloning. Sampling points 1-5 and 6-11 from each of the four sample sets (MC,
TC, FC and FV) were thus pooled together respectively, resulting in two samples for each set and a total of eight samples. The pooled PCR products (blunt ended due to using High
Fidelity Taq polymerase) were then A-tailed and purified with the ChargeSwitch® purification kit (Invitrogen) to make them suitable for ligation.
143
Ninety-six clones were randomly selected from clone libraries representing each of the two pooled samples from each transect. Transformed ITS products from each individual clone were re-amplified as described above using the M13 forward and reverse primer pair and RFLP screened using Taq1 in order to group clones with the same ITS sequence insert. Restriction fragments were run on a 3% agarose gel, stained with ethidium bromide and visualised under UV light. The resulting restriction fragment patterns were compared using GelCompare II software, with the aid of the Hyperladder IV DNA marker
(Bioline). PCR products that represented distinct RFLP patterns were then sequenced using the Big Dye Terminator Cycle Sequencing Kit v3.1 (Applied Biosystems) and an
ABI 3130xl sequencer (Applied Biosystems). The sequences obtained were analysed using Sequencher software (Version 3.0, Gene Codes Corp.) and closest matches in the
EMBL/GenBank/DDBJ databases were identified using the FASTA program (version
3.4t21) (Pearson & Lipman, 1988). Selected ITS sequences were aligned using the
BioEdit program (Version 5.0.9. (Hall, 1999)), firstly using the ClustalW option and subsequently manually by visual inspection. In order to infer taxonomic relationships for sequences representing both H. ericae and Sebacinaceae-like fungal endophytes in the present study along with related sequences in the GenBank, EMBL and DDBJ nucleotide databases, neighbour-joining analysis of the patristic distance matrix (1000 bootstraps re- sampling replicates) was conducted using PAUP (v. 4.01b9).
5.2.7 Data analysis
All multivariate data analyses were carried out using PC-ORD 4.0 for Windows (McCune and Mefford, 1999). A single ordination plot based on the DGGE binary information 144 from each transect was constructed using detrended correspondence analysis (DCA). All samples for each transect were run on separate DGGE gels, thus it is probable that this may account for at least some of any separation observed between samples in the ordination plot. The possibility of such bias was addressed by constructing a separate ordination plot using only binary data from standard lanes to investigate the similarities between banding positions for standard lanes from each gel. Due to the possibility of such bias no further multivariate analyses were carried out on the DGGE data.
The T-RFLP binary information was summed to give the total number of T-RFs for each enzyme and each fluorescent label at each sample point. Data from the total T-RF table were used in a single factor ANOVA to determine if there was a significant difference in the number of T-RFs from each transect. Furthermore, the table of total T-RFs for each sample point was used to construct a frequency distribution histogram which indicated the T-RFs to be used in additional analyses. An initial DCA of the T-RFLP data was carried out using all 675 T-RFs, however the ordination was unstable. Subsequent ordination plots were constructed, firstly, down weighting the rare species (in proportion to their frequency where F is < Fmax/5) and, secondly, using only those 238 T-RFs which had a frequency of five or more. Ordinations of separate T-RF binary data sets, for example ITS 1F-HEX Hinf data only, were also carried out.
Two-way indicator species analysis (TWINSPAN) based on the 238 T-RFs that had a frequency of five or more was carried out in order to disclose the sample composition of the main groupings and the T-RFs that are characteristic of each group. Further indicator 145 species analysis was carried out using 1000 Montecarlo simulations to calculate the probability that the distribution of T-RFs and subsequent sample assignment to each group could have occurred by chance.
5.3 Results
5.3.1 DGGE
PCR amplification of the rDNA ITS1 regions of fungal endophytes from pooled samples from each of the four sample transects yielded products of the expected size (150-300 bp)
(data not shown). Visual comparison of ITS1 rDNA band profiles obtained from the four
DGGE gels indicated that there were distinct and, in terms of the number of bands, diverse communities associated with the different transects (Figure 5.1). The moorland and transition zone sample transects appeared distinct from the forest sample transects, with fewer bands in the upper portion of the gels and relatively more in the bottom portions compared to the forest transect (Figure 5.1). The GelCompare II bandmatching software revealed there were 95 different bands across all transects with 82-88 bands observed in each transect. Many bands were observed in multiple samples often from more than one transect, while only four bands were unique to a single transect.
The DCA of DGGE binary data resulted in a plot with two axes which explained 21.27 % of the variation between samples (Figure 5.2). The DCA separated the samples into four distinct clusters that represented each of the four sample transects (MC, TC, FC and FV).
The two forest clusters were separated from the moorland and transitional zone clusters along the x axis, which explained most variation, indicating that the forest samples were 146 more similar to each other than the other two sample sets. It must be noted however, that all samples for each transect were run on separate DGGE gels and it is possible that this may account for at least some of the separation observed in the ordination plot. The possibility of such bias was addressed by constructing a separate ordination plot using only binary data generated from the DNA marker lanes of each gel. Sample positions on that plot (Figure 5.3) indicated there were differences between standard lanes on each gel, particularly those from the moorland and transition zone gels. It thus seems likely that at least some of the separation seen in the sample DCA plot (Figure 5.2) was due to running
PCR products from each transect on separate gels.
147
MC TC S 1 2 3 S 4 5 6 S 7 8 9 S 10 11 S S 1 2 3 S 4 5 6 S 7 8 9 S 10 11 S
FC FV
S 1 2 3 S 4 5 6 S 7 8 9 S 10 11 S S 1 2 3 S 4 5 6 S 7 8 9 S 10 11 S
Figure 5.1 DGGE analysis of ITS1 sequences amplified from Ericaceae roots collected from four sampling transects (MC = moorland Calluna vulgaris, TC = transition zone C. vulgaris, FC = forest C. vulgaris and FV = forest Vaccinium myrtillus) at Abernethy forest. Lane numbers correspond to sampling points along each transect and lanes marked S correspond to standard DNA marker lanes.
148
FC3
FC5
T10 FC4 T8 M3 T1
T4 T1 FC1 FC6 FC10 T7 T6 M9 FV1 T3 T2 FV2 T9 FC8 FC9 FC2 FC7 T5 FV3 M1 FV9 FC1 M2 en value: 0.0629
g FV8 FV1 M8 M7 M5
FV4 Axis 2 Ei M1 FV11 FV5 M1 M4 FV7 M6 FV6
Axis 1 Eigen value: 0.1498
Figure 5.2 Detrended Correspondence Analysis ordination of samples of Calluna vulgaris from moorland (M), transition zone (T) and forest (FC) transects and Vaccinium myrtillus from the forest (FV) transect at Abernethy forest based on 95 DGGE bands representing the ITS1 region of fungal rDNA extracted from surface-sterilised hair roots.
Numbers correspond to the sample point along each transect.
149
SFV
SFC
SFV SFV SFV
SFC
ST
SM SM
Axis 2 SFC
ST ST SM
SFC SM SM
SFC
ST SFV
ST
Axis 1
Figure 5.3 Detrended Correspondence Analysis ordination of standard lanes based on
DNA marker bands from each of four DGGE gels (SM = moorland Calluna vulgaris gel,
ST = transition zone C. vulgaris gel, FC = forest C. vulgaris gel and FV = forest
Vaccinium myrtillus gel).
150
5.3.2 T-RFLP
T-RFs were obtained from all samples using both restriction enzymes with the exception of TC1, TC11 and FV1 from which only Taq1 T-RFs were generated. These three samples were thus omitted from any further analyses. The frequencies of individual T-
RFs were calculated for each enzyme and each fluorescent label at each sample point
(Table 5.1). Using the Taq1 restriction enzyme, a total of 153 FAM and 191 HEX T-RFs were observed respectively from all sampling transects with 67-126 observed for each transect (Table 5.1). The numbers of T-RFs obtained using Hinf1 were similar, with a total of 151 FAM and 190 HEX T-RFs observed respectively from all transects and 57-
132 identified for each transect (Table 5.1). Although there appeared to be fewer T-RFs in the transition zone samples regardless of the enzyme used (Table 5.1), a single factor
ANOVA indicated there was no significant difference in the number of T-RFs observed between any of the sampling transects. Combined data revealed a total of 675 T-RFs across all sampling transects. These T-RFs were plotted on a frequency distribution histogram (Figure 5.4) which showed that 238 T-RFs were observed five or more times, while the remaining 437 T-RFs were observed less than five times.
151
Table 5.1 Abundance of T-RFs generated from fungal rDNA ITS regions obtained from Ericaceae roots from each of four sam pling transects (MC = moorland Calluna vulgaris, TC = transition zone C. vulgaris, FC = forest C. vulgaris and FV = forest Vaccinium myrtillus) at Abernethy forest using Taq1 and Hinf1 restriction enzymes.
Taq1 Hinf1 Sample HEX FAM HEX FAM Totals MC1 15 12 47 33 107 MC2 9 13 6 5 33 MC3 24 20 52 33 129 MC4 11 11 16 8 46 MC5 50 32 47 27 156 MC6 23 19 25 25 92 MC7 15 9 12 6 42 MC8 33 26 37 21 117 MC9 25 17 37 32 111 MC10 20 19 31 26 96 MC11 18 18 15 13 64 Total different MC T-RFs 100 74 132 103 TC1# 20 19 3 2 44 TC2 20 15 25 14 74 TC3 27 21 27 18 93 TC4 10 6 31 29 76 TC5 20 20 30 24 94 TC6 13 8 8 8 37 TC7 8 2 6 3 19 TC8 7 3 4 2 16 TC9 19 12 11 7 49 TC10 30 32 27 19 108 TC11# 21 15 6 3 45 Total different TC T-RFs 72 67 79 57 FC1 53 32 22 13 120 FC2 29 27 30 24 110 FC3 50 29 22 15 116 FC4 33 24 20 9 86 FC5 18 13 17 10 58 FC6 17 19 25 24 85 FC7 36 27 22 29 114 FC8 23 12 23 16 74 FC9 26 20 36 27 109 FC10 27 20 36 27 110 FC11 48 27 32 17 124 Total different FC T-RFs 120 88 104 83 FV1# 51 29 0 0 80 FV2 31 22 22 20 95 FV3 47 28 28 21 124 FV4 32 20 27 17 96 FV5 48 29 39 29 145 FV6 25 18 18 15 76 FV7 16 14 22 20 72 FV8 19 14 19 17 69 FV9 8 6 5 2 21 FV10 22 18 18 13 71 FV11 44 30 34 28 136 Total different FV T-RFs 126 93 87 64 71 Total different T-RFs 191 153 190 151 675
# Samples that were omitted from Total different T-RF counts and multivariate analysis.
152
Frequency distribution of TRFs
45
40
35
30
25
20
Frequency 15
10
5
0 1 41 81 121 161201 24 1 281 32 1 361 401 441 481 52 1 56 1 601 641 681 T-RF num ber
Figure 5.4 Frequency distribution histogram of the total 675 T-RFs (combing HEX and
FAM T-RFs obtained using both Taq1 and Hinf1 restrict ion enzy me s) from fungal rDNA
ITS regions obtained from Ericaceae roots at Abernethy forest.
DCA of all 675 T-RFs re sulted in an unstable ordination plot due to the software having difficulty recognising the main axi s of var iatio n, how ever a DCA that down-weighted the
437 T-RFs with a frequency of less than five displayed reasonable separation between the moorland/transition zone and forest samples (data not shown). The best separation between samples was observed using DCA of only the 238 T-RFs with a frequency of 153 five or greater. Samples were thus separated into four distinct clusters that represented each of the four sampling transects (Figure 5.5). The two forest transects were separated from the moorland and transitional zone transects along the axis 1, which explained most variation (28.17 %), indicating that the forest samples were more similar to each other than the other two sample sets. There was however, some overlap between samp le clusters, particularly the forest clusters, on the T-RF plot (Figure 5.5). DCA of separat e
T-RF binary data sets, for example HEX Hinf1 data only, all grouped the samp les similarly to the combined DCA plot (data not shown).
TWINSPAN classification of the 238 T-RFs with a frequency of five or more initially identified two groups, one of which contained all forest samples, while the other contained all moorland and transition zone samples. A second level of classification divided the combined forest group into two further sub-groups, one containing predominately forest Calluna samples and the other containing mainly forest Vacciniu m samples (Table 5.2). Similarly, the moorland/transition zone group was further divided into two sub-groups, one containing mainly moorland samples, and the other mainly transition zone samples (Table 5.2). The TWINSPAN analysis identified 66, 26, 75, and
25 T-RFs that were characteristic of each group (MC, TC, FC and FV) respectively and indicator species analysis using 1000 Montecarlo simulations confirmed that these T-RFs were reliable (P< 0.05) indicators of the four predetermined sampling transects (MC, TC,
FC and FV) respectively (data not shown).
154
T2
FC3
T5
FC5 FC1 T8 T3 FC4 T7
FC11 T4 T9 T6 M FC2 FC9
FV4 FC8 M10 en value: 0.1272 g FV5 FV6 M7 M5 FC10 T10 M2 Axis 2 Ei FV2 FV3 FV11 M3 FV8 M8
FC7 FV10 M1
FV7 FC6 FV9
M6 M9
M11
Axis 1 Eigen value: 0.2817
Figure 5.5 Detrended Correspondence Analysis ordination of samples of 31 Calluna vulgaris from moorland (M), transition zone (T) and forest (FC) transects and 10
Vaccinium myrtillus from the forest (FV) transect at Abernethy forest based on 675 terminal restriction fragments generated from the ITS region of fungal rDNA extracted from surface-sterilised hair roots.. Numbers correspond to the sample point along each transect.
155
Table 5.2 TWINSPAN classification of sample points from transects (MC, TC, FC and
FV) at Abernethy forest based on 238 T-RFs with a frequency of 5 or more. Initial classification identified two groups (1 and 2). Secondary classification further divided these two groups giving four groups: 1a, 1b, 2a and 2b which contained samples from predominately the MC, TC, FC and FV sampling transects respectively.
Group Transect sample points
1a (MC) MC1, MC3, MC4, MC5, MC6, MC7, MC8, MC9, MC10, MC11, TC10
1b (TC) TC2, TC3, TC4, TC5, TC6, TC7, TC8, TC9, MC2
2a (FC) FC1, FC3, FC4, FC5, FC8, FC9, FC11, FV3, FV5, FV11
2b (FV) FV2, FV4, FV6, FV7, FV8, FV9, FV10, FC2, FC6, FC7, FC10
5.3.3 Cloning
Ninety-six clones were randomly selected for analysis from each of the eight clone assemblages that represented the four sampling transects. Of the total 768 clones, thirty- three did not contain an ITS insert, leaving 735 in total. GelCompare II analysis of Taq1 156
ITS-RFLP patterns obtained from clones grouped them into 148 RFLP-types (data not shown). One clone representing each RFLP-type was selected for ITS sequencing. Six
RFLP-types could not be sequenced even after repeated attempts, however all of these
RFLP-types were represented by only single clones. The two closest sequence matches for each RFLP-type resulting from FASTA searches of sequences from the GenBank database are presented in Table 5.3. Twenty-six, 20, 22 and 23 RFLP-types were unique to the moorland, transition zone, forest Calluna and forest Vaccinium sampling transects respectively. Of the total of 729 clones representing the 148 RFLP-types, 50% were of sixteen different RFLP-types while the remaining clones comprised 132 RFLP-types
(Table 5.3).
RFLP-types 1-21 , that comprised ca 28% of the clones across all transects had closest matches with sequences for members of the H. ericae aggregate or taxonomically similar
Helotiales fungi ( Table 5.3). One of the closest sequence matches to RFLP-type 12, that representing seve n clones, was the isolate that was shown by Villarreal-Ruiz et al. (2004) to simultaneously form ericoid mycorrhizal and ectomycorrhizal structures. The closest sequence matches for RFLP-types 22-34 (which represented clones from all transects, but mainly the forest Calluna transect) were Capronia-like sequences for cultured fungi, some of which are mycorrhizal (Allen et al., 2003), or clones from other Ericaceae species.
RFLP-types with closest matches to sequences of Sebacinaceae basidiomycetes were also common, with 32 different types identified (RFLP-types 72-103) (Table 5.3). Of these, 157
15 RFLP-types were found only in the moorland and/or transition zone transects while eight were found only in the forest transects. The closest sequence matches to RFLP- types 104-124, representing ca 11% of the clone population, were ectomycorrhizal fungi from the families Atheliaceae, Thelephoraceae, Corticiaceae, Clavulinaceae, Russulaceae and Tricholomataceae (Table 5.3). Seven of these RFLP-types (representing twelve clones) were from the moorland/transition zone transects, while the remaining 16 RFLP types (representing 65 clones) were from the forest transects (Table 5.3).
The closest sequence matches to the remaining RFLP-types were with sequences from a range of ascomycetes, basidiomycetes and zygomycetes, howeverRFLP-types 145-148 had matches with multiple fungal phyla, making identification at any level impossible.
Of the RFLP-types that matched ascomycete taxa, several had closest matches with sequences representing fungi frequently obtained from the roots of other Ericaceae species (Table 5.3). Of these, RFLP-types 35-38 had matches with sequences for fungi that have been shown to form ericoid mycorrhizal associations), while several others,
RFLP-types 39-43, matched sequences representing dark septate endophytes, which are common colonisers of the roots of Ericaceae and other plant taxa worldwide (Jumpponen,
2001).
158
Table 5.3 Two closest sequence matches obtained from FASTA searches between cloned RFLP-types and sequences from the GenBank/EMBL/DDBJ databases. The number of clones representing each RFLP-type found in each of the four sampling transects (MC = moorland Calluna vulgaris, TC = transition zone C. vulgaris, FC = forest C. vulgaris and FV = forest Vaccinium myrtillus) at Abernethy forest is included. Number of clones in RFLP Accession Closest 2 FASTA matches Identity% Overlap e score each transect type code MC TC FC FV 1 DQ309226 Hymenoscyphus ericae isolate pkc 29 (AY394907) 98.195 554 2.3 e-67 0 1 0 0 Hymenoscyphus sp. UBCM8 (AF081435) 96.570 554 3.9 e-65 2 DQ309143 Hymenoscyphus ericae isolate pkc 29 (AY394907) 99.273 688 1.3 e-89 1 0 0 0 Hymenoscyphus ericae isolate 21 (AF069439) 99.110 337 1.9 e-40 3 DQ309150 Hymenoscyphus ericae strain UBCtra 141 (AF149067) 98.355 547 2.4 e-62 9 13 12 8 Hymenoscyphus ericae isolate pkc 29 (AY394907) 98.311 533 7.7 e-61 4 DQ309163 Hymenoscyphus ericae strain UBCtra 141 (AF149067) 91.985 549 6.5 e-53 2 0 0 0 Hymenoscyphus ericae isolate pkc 29 (AY394907) 92.150 535 6.6 e-52 5 DQ309185 Hymenoscyphus sp. UBCM8 (AF081435) 96.618 680 5.9 e-81 0 0 1 0 Hymenoscyphus ericae isolate pkc 29 (AY394907) 94.767 707 4.7 e-80 6 DQ3092 13 Hymenoscyphus ericae strain UBCtra 141 (AF149067) 94.402 393 5.2 e-41 1 0 0 0 Hymenoscyphus ericae isolate pkc 29 (AY394907) 94.667 375 1.1 e-39 7 DQ309218 Hymenoscyphus ericae strain UBCtra 141 (AF149067) 92.350 549 1.3 e-53 7 14 18 14 Hymenoscyphus ericae isolate pkc 29 (AY394907) 92.523 535 1.3 e-52 8 DQ309197 Hymenoscyphus ericae isolate pkc 29 (AY394907) 99.088 658 7.5 e-81 2 1 3 0 Hymenoscyphus sp. UBCM8 (AF081435) 96.840 633 1.1 e-73 9 DQ309108 cf. Hymenoscyphus sp. GU30 (AF252836) 95.388 412 4.4 e-50 0 0 0 1 Hymenoscyphus ericae isolate 21 (AF069439) 95.355 409 3.9 e-50 10 DQ309155 cf. Hymenoscyphus sp. he7-23-5 (AY112936) 88.694 513 8.0 e-47 2 0 0 0 Salal root associated fungus UBCtra 180 (AF149071) 88.153 498 1.0 e-44 11 DQ309154 Salal root associated fungus UBCtra 264 (AF149070) 98.221 506 2.7 e-56 1 0 0 0 Phialophora finlandia (AJ534704) 94.600 537 4.0 e-55 12 DQ309223 Salal root associated fungus UBCtra 264 (AF149070) 98.617 506 1.1 e-56 4 3 0 0 Ectomycorrhizal isolate LVR4069 (AY579413) 94.054 555 1.4 e-55 13 DQ309198 Uncultured ectomycorrhiza (Phialophora) 4097 (AY634154) 88.038 627 5.4 e-49 0 3 0 0 Phialophora verrucosa (PV31846) 84.834 633 1.5 e-48 14 DQ309215 Ectomycorrhizal root tip 165-sepB Ny2.B1-27.3 (AF481385) 87.715 407 6.7 e-34 2 0 0 0 Phialophora finlandia (AJ534704) 87.469 407 7.4 e-34 15 DQ309164 Uncultured mycorrhizal fungus clone c12.1 (AY394903) 89.083 458 3.3 e-35 12 1 0 0 Uncultured fungus isolate RFLP 66 (AF461627) 84.073 496 1.9 e-30 16 DQ309130 Salal root associated fungus UBCtra 264 (AF149070) 100.000 505 6.7 e-62 0 0 1 0 Axenic ericoid root isolate cf. H. ericae (AJ430114) 100.000 475 7.8 e-58 17 DQ309172 Uncultured mycorrhizal fungus clone bw 2.3 (AY394897) 90.783 575 9.8 e-54 0 5 2 1 Mycorrhizal fungal sp. pkc24 (AY394893) 91.334 577 1.3 e-53 18 DQ309217 Mycorrhizal fungal sp. pkc24 (AY394893) 96.187 577 9.5 e-60 20 10 8 10 Un cultured mycorrhizal fungus clone 2.13 (AY394918) 96.014 577 1.6 e-59 19 DQ309128 My corrhizal fungal sp. pkc12 (AY394886) 99.542 437 2.2e-58 2 3 3 1 Mycorrhizal fungal sp. pkc22 (AY394890) 99.542 437 2.2e-58 20 DQ3091 40 Mycorrhizal sp. UBCS246 (AF081443) 92.941 510 2.3e-49 0 2 1 0 Salal mycorrhizal fungus UBCtra 179 (AF149077) 92.843 517 2.0 e-49 21 DQ3091 57 Mycorrhizal fungal sp. pkc11 (AY394892) 84.429 578 3.2e-40 1 0 0 0 Ericoid mycorrhizal sp. D02 (AF072302) 87.277 448 2.5 e-35 22 DQ3091 94 Mycorrhizal fungal sp. pkc12 (AY394886) 99.470 566 1.7 e-66 0 0 2 1 Mycorrhizal fungal sp. pkc2 (AY394890) 99.470 566 1.8 e-66 23 DQ3091 46 Mycorrhizal ascomycete of Rhododendron type 3 (AB089663) 87.405 524 1.6 e-47 1 0 0 0 Leaf litter ascomycete strain its 233 (AF502761) 86.591 440 2.7 e-38 24 DQ3092 01 Mycorrhizal ascomycete of Rhododendron type 3 (AB089661) 84.817 382 2.9 e-28 4 0 0 3 Ericoid mycorrhizal sp. Sm2 (AY046398 ) 84.921 378 3.1 e-27 25 DQ3091 29 Capronia villosa (AF050261) 79.204 553 6.8e-31 0 0 2 0 Woollsia root associated fungus IX (AY230779) 78.276 557 2.0 e-30 159
Table 5.3 cont. Two closest sequence matches obtained from FASTA searches between cloned RFLP-types and sequences from the GenBank/EMBL/DDBJ databases. The number of clones representing each RFLP-type found in each of the four sampling transects (MC = moorland Calluna vulgaris, TC = transition zone C. vulgaris, FC = forest C. vulgaris and FV = forest Vaccinium myrtillus) at Abernethy forest is included. Number of clones in RFLP Accession Closest 2 FASTA matches Identity% Overlap e score each transect type code MC TC FC FV 26 DQ309133 Woollsia root associated fungus IX (AY230779) 90.018 551 1.4 e-52 0 0 1 0 Capronia villosa (AF050261) 84.517 549 1.1 e-42 27 DQ309103 Woollsia root associated fungus IX (AY230779) 89.091 550 1.3 e-47 0 0 0 1 Capronia sp. UBCTRA1522.6 (AF284126) 97.784 361 9.8 e-38 28 DQ309106 Woollsia root associated fungus IX (AY230779) 89.111 551 1.3 e-47 0 0 5 3 Capronia villosa (AF050261) 83.942 548 2.3 e-39 29 DQ309144 Woollsia root associated fungus IX (AY230779) 75.763 557 1.7 e-28 1 0 0 0 Capronia villosa (AF050261) 76.036 555 9.6 e-27 30 DQ309151 Woollsia root associated fu (AY230779) 89.895 475 2.7 e-47 1 0 0 0 Fungal sp. TRN242 (AY843097) 88.627 466 1.0 e-44 31 DQ309170 Capronia sp. UBCTRA 1322.11 (AF284128) 90.358 363 8.6 e-31 0 1 0 0 Capronia sp. UBCTRA 1.01 (AF284127) 90.110 364 2.4 e-30 32 DQ309244 Capronia sp. UBCTRA1522.6 (AF284126) 98.338 361 2.1 e-41 0 1 0 2 Capronia sp. UBCTRA 1322.11 (AF284128) 98.338 361 2.1 e-41 33 DQ309188 Capronia sp. UBCTRA1522.6 5 (AF284126) 98.061 361 3.7 e-38 0 0 5 0 Capronia sp. UBCTRA 1322.11 (AF284128) 98.061 361 3.7 e-38 34 DQ309192 Woollsia root associated fungus IX (AY230779) 89.111 551 1.4 e-47 0 0 2 0 Fungal sp. TRN242 (AY843097) 86.863 510 1.9 e-41 35 DQ309235 Woollsia root associated fungus IX (AY230779) 89.655 551 8.5 e-50 0 1 1 1 Capronia villosa (AF050261) 84.799 546 4.3 e-41 36 DQ309127 Woollsia root associated fungus IX (AY230779) 79.710 552 5.8 e-34 0 0 1 0 Uncultured fungus (AJ582965) 80.289 553 9.8 e-34 37 DQ309126 Cladophialophora sp. TRN488 (AY843173) 82.979 470 2.1e-31 0 0 2 0 Capronia villosa (AF050261) 81.239 549 1.5e-31 38 DQ309101 Cf. Phialocephala fortinii (AY078132) 93.878 392 1.6 e-39 0 0 0 1 Cf. Phialocephala fortinii (AY078151) 93.590 390 5.2 e-39 39 DQ309109 Cf. Phialocephala fortinii (AY078132) 97.243 544 9.7 e-62 0 0 0 5 Phialocephala fortinii CBS 109302 (AY078136) 97.238 543 5.6 e-62 40 DQ309113 Cf. Phialocephala fortinii (AY078133) 95.115 655 5.5 e-75 0 0 0 3 Phialocephala fortinii 92-109 (AY078135) 95.090 611 3.9 e-70 41 DQ309112 Cf. Phialocephala fortinii (AY078133) 88.336 583 7.4 e-54 0 0 0 2 Cf. Phialocephala fortinii (AY078132) 89.927 546 1.2 e-52 42 DQ309152 Phialophora sp. P3901 (AF083199) 88.770 561 9.8 e-48 4 7 0 0 Phialocephala virens strain CBS 452.92 (AF486132) 88.869 566 5.9 e-47 43 DQ309122 Hymenoscyphus monotropae ATCC52305 (AF169309) 91.440 514 4.8 e-49 0 0 0 1 Uncultured fungus isolate RFLP 98 (AF461659) 86.438 553 3.8 e-46 44 DQ309238 Salal root associated fungus UBCTRA 1542.10 (AF284136) 86.525 423 1.6 e-40 2 1 0 0 Salal root associated fungus UBCtra 153 (AF149078) 90.909 319 1.7 e-34 45 DQ309240 Salal root associated fungus UBCtra 153 (AF149078) 95.984 498 2.4 e-52 0 4 0 1 Epacris microphylla root associated fungus 16 (AY268200) 94.882 508 2.5 e-52 46 DQ309241 Solenopezia solenia (U57991) SSU57991) 73.633 512 4.4 e-26 2 5 0 0 Epacris microphylla root associated fungus 15 (AY268199) 75.506 494 3.9 e-25 47 DQ309191 Solenopezia solenia (U57991) 73.633 512 4.4 e-26 3 2 3 0 Epacris microphylla root associated fungus 15 (AY268199) 75.506 494 3.9 e-25 48 DQ309236 Epacris microphylla root associated fungus 16 (AY268200) 95.385 325 4.0 e-32 0 1 0 0 Salal root associated fungus UBCtra 153 (AF149078) 96.238 319 5.8 e-32 49 DQ309228 Uncultured fungus isolate RFLP 34 (AF461595) 76.825 548 5.2 e-33 0 1 0 0 Salal root associated fungus UBCtra 264 (AF149070) 98.239 284 1.1 e-32 50 DQ309134 Scleropezicula alnicola strain CBS 200.46 (AF141168) 90.519 559 2.9 e-52 0 0 1 0 Scleropezicula alnicola strain CBS 474.97 (AF141169) 89.502 562 1.6 e-50 160
Table 5.3 cont. Two closest sequence matches obtained from FASTA searches between cloned RFLP-types and sequences from the GenBank/EMBL/DDBJ databases. The number of clones representing each RFLP-type found in each of the four sampling transects (MC = moorland Calluna vulgaris, TC = transition zone C. vulgaris, FC = forest C. vulgaris and FV = forest Vaccinium myrtillus) at Abernethy forest is included. Number of clones in RFLP Accession Closest 2 FASTA matches Identity% Overlap e score each transect type code MC TC FC FV 51 DQ309139 Scleropezicula alnicola strain CBS 200.46 (AF141168) 90.000 560 5.7 e-51 0 0 1 0 Scleropezicula alnicola strain CBS 474.97 (AF141169) 88.988 563 3.1 e-49 52 DQ309118 Scleropezicula alnicola strain CBS 200.46 (AF141168) 90.647 556 6.9 e-51 0 0 0 1 Scleropezicula alnicola strain CBS 474.97 (AF141169) 89.982 559 1.2 e-49 53 DQ309162 Pezicula sporulosa strain CBS 224.96 (AF141172) 89.587 557 3.0 e-51 1 0 0 0 Cryptosporiopsis ericacea isolate UBCtra 1204 (AY442321) 89.209 556 4.0 e-51 54 DQ309147 Phialocephala virens strain CBS 452.92 (AF486132) 90.036 562 7.3 e-51 1 0 0 0 Cryptosporiopsis ericacea isolate UBCtra 1204 (AY442321) 88.669 556 1.7 e-50 55 DQ309136 Uncultured fungus clone D38 (AF504874) 80.275 436 3.4 e-10 0 1 1 1 Uncultured fungus clone D7 (AF504875) 83.776 339 1.2 e-9 56 DQ309153 Uncultured ascomycete clone K11-10 (AY394668) 94.255 470 2.2 e-63 2 2 0 0 Uncultured Mycosphaerella clone p4-7 (AF207671) 82.944 428 7.1 e-38 57 DQ309176 Ascomycete sp. Olrim543 (AY805603) 85.714 483 8.2 e-36 0 1 1 0 Leaf litter ascomycete strain its 249 (AF502775) 82.090 402 9.4 e-27 58 DQ309190 Leaf litter ascomycete strain its 233 (AF502761) 78.657 417 1.2 e-21 0 0 1 0 Ascomycete sp. Olrim401 (AY781244) 81.989 372 1.3 e-21 59 DQ309131 Uncultured fungus clone D3 (AF504848) 97.161 634 8.0 e-74 0 0 1 0 Uncultured fungus clone D5 (AF504849) 94.228 641 4.5 e-69 60 DQ309121 Uncultured fungus clone D3 (AF504848) 98.210 447 5.3 e-49 0 0 0 1 Verticillium sp. olrim438 (AY805596) 98.353 425 2.4 e-46 61 DQ309239 Neofabraea malicorticis (AF141189) 78.198 555 1.3 e-31 0 2 0 0 Neofabraea alba (AF141190) 78.261 552 1.9 e-31 62 DQ309243 Uncultured ascomycete clone ot1c3 (AY273315) 96.994 499 3.2 e-51 0 3 0 0 Neofabraea malicorticis (AF141189) 90.942 552 1.7 e-50 63 DQ309137 Cladosporium fulvum strain ATCC 44960 (AF393701) 85.504 407 4.1 e-30 0 0 1 0 Mycosphaerella pini (AF013227) 85.856 403 1.1 e-29 64 DQ309148 Lunulospora curvula (AY729938) 90.871 482 7.6 e-45 1 0 0 0 Alatospora acuminata strain ccm-F12186 (AY204588) 87.434 565 6.3 e-44 65 DQ309206 Pseudocyphellaria homoeophy (AF351145) 73.383 541 3.5 e-26 8 7 0 0 Pseudocyphellaria rufovires (AF351142) 73.198 541 5.7 e-26 66 DQ309119 Kluyveromyces lactis strain NRRL Y-1140 (CR382124) 59.184 441 2.2 e-1 0 0 0 1 Erysiphe pulchra (AY870864) 63.387 437 3.3 e-1 67 DQ309200 Bisporella citrina (AF335454) 91.328 542 2.6 e-49 4 1 1 2 Calycina herbarum isolate 1549 (AY348594) 95.816 478 3.0 e-48 68 DQ309166 Taphrina caerulescens strain CBS 351.35 (AF492081) 86.631 187 5.6 e-15 2 0 0 0 Taphrina aff. virginica 046 (AY188378) 85.789 190 4.5 e-15 69 DQ309179 Penicillium thomii strain FRR 2077 (AY373934) 99.642 558 3.8 e-63 0 4 3 3 Penicillium spinulosum strain FRR 1750 (AY373933) 99.640 556 3.0 e-63 70 DQ309184 Penicillium lividum strain FRR 1228 (AY373922) 99.822 562 1.4 e-63 0 0 1 0 Uncultured fungus isolate RFLP 68 (AF461629) 97.059 578 1.3 e-61 71 DQ309227 Penicillium thomii (AF034460) 91.189 454 7.9 e-41 0 9 0 0 Eupenicillium lapidosum strain NRRL 718 (AF033409) 91.189 454 7.9 e-41 72 DQ309141 Salal root associated fungus UBCTRA 1322.6 (AF284137) 94.363 408 2.1 e-42 0 0 0 9 Salal root associated fungus UBCTRA 1041.2 (AF284135) 93.674 411 2.6 e-41 73 DQ309105 Salal root associated fungus UBCTRA 1322.6 (AF284137) 93.199 397 1.2 e-41 0 1 0 1 Salal root associated fungus UBCTRA 1041.2 (AF284135) 92.714 398 1.1 e-41 74 DQ309110 Salal root associated fungus UBCTRA 1041.2 (AF284135) 92.910 409 3.1 e-40 0 0 0 2 Salal root associated fungus UBCTRA 1322.6 (AF284137) 92.892 408 1.0 e-39 75 DQ309111 Salal root associated fungus UBCTRA 1542.10 (AF284136) 96.635 416 5.0 e-45 0 0 0 2 Salal root associated fungus UBCTRA 1041.2 (AF284135) 92.048 415 7.2 e-40 161
Table 5.3 cont. Two closest sequence matches obtained from FASTA searches between cloned RFLP-types and sequences from the GenBank/EMBL/DDBJ databases. The number of clones representing each RFLP-type found in each of the four sampling transects (MC = moorland Calluna vulgaris, TC = transition zone C. vulgaris, FC = forest C. vulgaris and FV = forest Vaccinium myrtillus) at Abernethy forest is included. Number of clones in RFLP Accession Closest 2 FASTA matches Identity% Overlap e score each transect type code MC TC FC FV 76 DQ309160 Salal root associated fungus UBCTRA 1322.6 (AF284137) 92.875 407 5.8 e-50 1 0 0 0 Salal root associated fungus UBCTRA 1041.2 (AF284135) 92.176 409 2.1 e-49 77 DQ309161 Salal root associated fungus UBCTRA 1322.6 (AF284137) 93.382 408 2.8 e-41 1 0 0 0 Salal root associated fungus UBCTRA 1041.2 (AF284135) 92.944 411 3.4 e-41 78 DQ309178 Salal root associated fungus UBCTRA 1322.6 (AF284137) 92.067 416 4.2 e-12 0 0 1 1 Salal root associated fungus UBCTRA 1041.2 (AF284135) 89.835 423 9.6 e-12 79 DQ309180 Salal root associated fungus UBCTRA 1322.6 (AF284137) 98.515 404 4.7 e-47 0 0 7 11 Salal root associated fungus UBCTRA 1041.2 (AF284135) 97.783 406 1.4 e-46 80 DQ309158 Salal root associated fungus UBCTRA 1322.6 (AF284137) 94.118 408 3.2 e-42 5 8 0 0 Salal root associated fungus UBCTRA 1542.10 (AF284136) 92.010 413 1.9 e-41 81 DQ309221 Salal root associated fungus UBCTRA 1542.10 (AF284136) 87.112 419 3.5 e-34 1 0 0 0 Salal root associated fungus UBCTRA 1322.6 (AF284137) 87.470 415 4.0 e-34 82 DQ309224 Salal root associated fungus UBCTRA 1322.6 (AF284137) 94.321 405 2.0 e-41 2 2 0 0 Salal root associated fungus UBCTRA 1542.10 (AF284136) 92.457 411 2.7 e-40 83 DQ309230 Salal root associated fungus UBCTRA 1322.6 (AF284137) 93.627 408 2.5 e-41 0 2 0 0 Salal root associated fungus UBCTRA 1041.2 (AF284135) 93.171 410 4.1 e-41 84 DQ309231 Salal root associated fungus UBCTRA 1322.6 (AF284137) 93.137 408 3.7 e-47 0 1 0 0 Salal root associated fungus UBCTRA 1041.2 (AF284135) 92.476 412 4.4 e-46 85 DQ309233 Salal root associated fungus UBCTRA 1322.6 (AF284137) 90.580 414 1.6 e-37 0 2 0 0 Salal root associated fungus UBCTRA 1041.2 (AF284135) 90.315 413 3.8 e-37 86 DQ309142 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 81.832 677 4.2 e-40 2 0 0 0 Salal root associated fungus UBCTRA 1542.10 (AF284136) 86.747 415 6.0 e-35 87 DQ309149 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 82.422 677 8.7 e-40 3 2 1 0 Salal root associated fungus UBCTRA 1322.6 (AF284137) 87.775 409 3.9 e-35 88 DQ309159 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 82.743 678 1.9 e-39 4 3 0 0 Salal root associated fungus UBCTRA 1542.10 (AF284136) 87.740 416 5.0 e-35 89 DQ309165 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 79.911 672 1.9 e-36 2 0 0 0 Salal root associated fungus UBCTRA 1542.10 (AF284136) 87.470 415 3.8 e-35 90 DQ309174 Salal root associated fungus UBCTRA 1322.6 (AF284137) 96.774 403 5.1 e-44 0 0 5 3 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 81.166 669 2.6 e-43 91 DQ309187 Salal root associated fungus UBCTRA 1322.6 (AF284137) 97.022 403 1.7 e-44 0 0 6 5 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 81.614 669 2.9 e-44 92 DQ309199 Salal root associated fungus UBCTRA 1322.6 (AF284137) 92.611 406 7.0 e-40 1 2 0 0 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 80.882 680 1.9 e-36 93 DQ309196 Salal root associated fungus UBCTRA 1322.6 (AF284137) 92.308 403 1.1 e-39 1 0 0 0 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 80.769 676 3.1 e-36 94 DQ309202 Salal root associated fungus UBCTRA 1542.10 (AF284136) 87.648 421 2.9 e-35 1 0 0 0 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 81.688 628 8.3 e-32 95 DQ309204 Salal root associated fungus UBCTRA 1322.6 (AF284137) 92.647 408 3.2 e-40 1 1 1 0 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 80.294 680 2.7 e-36 96 DQ309205 Salal root associated fungus UBCTRA 1322.6 (AF284137) 94.258 418 1.9 e-41 2 0 0 0 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 81.910 691 1.3 e-36 97 DQ309208 Salal root associated fungus UBCTRA 1322.6 (AF284137) 94.626 428 1.7 e-48 2 1 1 0 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 81.212 660 3.7 e-36 98 DQ309210 Salal root associated fungus UBCTRA 1322.6 (AF284137) 93.250 400 1.9 e-41 10 7 5 3 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 80.938 682 3.0 e-33 99 DQ309211 Salal root associated fungus UBCTRA 1322.6 (AF284137) 92.632 380 2.8 e-40 9 3 1 9 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 79.908 652 3.3 e-40 100 DQ309214 Uncultured orchid mycorrhizal fungus (AJ549120) 87.937 572 6.4 e-51 3 6 5 12 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 81.442 652 1.6 e-45 162
Table 5.3 cont. Two closest sequence matches obtained from FASTA searches between cloned RFLP-types and sequences from the GenBank/EMBL/DDBJ databases. The number of clones representing each RFLP-type found in each of the four sampling transects (MC = moorland Calluna vulgaris, TC = transition zone C. vulgaris, FC = forest C. vulgaris and FV = forest Vaccinium myrtillus) at Abernethy forest is included. Number of clones in RFLP Accession Closest 2 FASTA matches Identity% Overlap e score each transect type code MC TC FC FV 101 DQ309219 Uncultured orchid mycorrhizal fungus (AJ549120) 87.741 571 7.7 e-50 2 5 6 2 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 82.300 661 1.0 e-46 102 DQ309225 Uncultured orchid mycorrhizal fungus (AJ549120) 83.649 581 3.5 e-44 0 2 2 0 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 82.265 671 5.6 e-41 103 DQ309100 Uncultured orchid mycorrhizal fungus (AJ549120) 87.916 571 1.9 e-50 0 0 0 4 Uncultured mycorrhiza (Sebacinaceae) 4078 (AY634132) 82.451 661 2.6 e-47 104 DQ309124 Tomentellopsis submollis (AY641459) 97.028 673 1.7 e-78 0 0 1 0 Tomentellopsis submollis (AJ438983) 97.019 671 3.5 e-78 105 DQ309171 Ectomycorrhizal root tip 203 Ny3.O-32.2 (AF476974) 91.935 434 4.1 e-47 0 1 0 0 Tomentellopsis sp. Pink/L2 (AJ410766) 91.935 434 6.0 e-47 106 DQ309175 Thelephoralean ectomycorrhiza AT97LC10b3 (AF274778) 97.204 608 7.2 e-43 0 0 8 3 Ectomycorrhizal root tip 116 Ny1.B1-1.1 (AF476967) 94.818 521 5.5 e-35 107 DQ309117 Thelephoralean ectomycorrhiza AT97LC10b3 (AF274778) 99.332 599 2.2 e-72 0 0 0 1 Pseudotomentella tristis JS20643(O) (AF274772) 99.314 583 2.4 e-71 108 DQ309183 Pseudotomentella tristis JS20643(O) (AF274772) 99.829 584 6.4 e-71 0 0 4 1 Thelephoralean ectomycorrhiza LC 169 (AF274777) 99.827 579 1.1 e-70 109 DQ309132 mycorrhizal basidiomycote isolate T01 (AB089818) 99.136 579 2.4 e-67 0 0 11 10 Amphinema byssoides (AY219839) 83.806 599 1.7 e-43 110 DQ309107 mycorrhizal basidiomycote isolate T01 (AB089818) 96.381 525 7.9 e-64 0 0 3 4 Amphinema byssoides (AY219839) 83.148 540 1.8 e-43 111 DQ309173 Russula decolorans (AF418637) 98.836 687 8.3 e-82 0 4 0 0 Russula decolorans 1-502IC5 (AY061670) 97.163 705 2.3 e-81 112 DQ309138 Sistotrema sernanderi isolate olrim 216 (AY805624) 91.873 566 4.3 e-56 0 0 2 0 Sistotrema sernanderi strain KHL8576 (AF506476) 93.785 354 5.0 e-35 113 DQ309120 Sistotrema coronilla strain NH 7598 (AF506475) 93.566 373 1.1 e-36 0 0 0 2 Uncultured ectomycorrhiza (Cantharellales) (AF440673) 78.369 601 1.1 e-36 114 DQ309156 Uncultured ectomycorrhizal basidiomycete (AY641458) 74.214 636 2.4 e-34 1 0 0 0 Uncultured cf. Tricholoma sp. Clone d485.8 (AY254876) 79.925 533 1.3 e-32 115 DQ309114 Uncultured mycorrhizal fungus clone bw2.12 (AY394895) 100.000 636 1.3 e-78 0 0 0 3 Uncultured cf. Piloderma (AY097053) 99.687 638 2.0 e-77 116 DQ309186 Uncultured mycorrhizal fungus clone bw2.12 (AY394895) 99.283 558 3.2 e-84 0 0 1 2 cf. Piloderma reticulatum 212 Ny3.E1-4.1 (AF481387) 99.283 558 5.0 e-84 117 DQ309102 Clavulinaceae sp. F46 (CSP534708) 78.338 674 3.1 e-32 0 0 0 1 Uncultured ectomycorrhizal fungus TAM 209 (AY310836) 76.789 629 5.1 e-32 118 DQ309232 Gymnopus dryophilus isolate FB 11015 (AF505787) 99.023 614 9.0 e-90 0 1 0 0 Gymnopus bicolor AWW116 (AY263423) 97.900 619 5.4 e-89 119 DQ309123 Uncultured mycorrhizal fungus clone bw 2.17 (AY394904) 86.593 455 3.2 e-30 0 1 0 1 Uncultured fungus isolate RFLP 40 (AF461601) 84.917 484 3.8 e-30 120 DQ309209 Uncultured mycorrhizal fungus clone bw 2.17 (AY394904) 88.027 451 3 e-31 2 0 0 2 Uncultured fungus isolate RFLP 40 (AF461601) 85.239 481 1.4 e-30 121 DQ309145 Fibulorhizoctonia carotae strain ATCC 10866 (U85789) 96.749 646 2.9 e-73 1 1 0 0 Athelia arachnoidea (U85791) 96.749 646 2.9 e-73 122 DQ309189 Fibulorhizoctonia centrifuga (U85790) 83.282 652 2.7 e-49 0 0 3 0 Athelia epiphylla (U85794) 83.282 652 2.7 e-49 123 DQ309181 Uncultured ectomycorrhiza 4074 (AY634127) 92.239 670 2.1 e-66 0 0 1 0 Uncultured ectomycorrhiza 6139 (AY634119) 92.227 669 2.3 e-66 124 DQ309098 Cenococcum geophilum isolate c12.19 (AY394919) 94.840 407 2.7 e-16 0 0 0 1 Cenococcum geophilum isolate c11.21 (AY394913) 94.595 407 3.8 e-16 125 DQ309216 Uncultured Termitomyces DNA AB081118) 63.123 602 6.3 e-12 4 0 0 0 Cryptococcus victoriae strain S762 (AY301025) 69.755 367 7.3 e-12 163
Table 5.3 cont. Two closest sequence matches obtained from FASTA searches between cloned RFLP-types and sequences from the GenBank/EMBL/DDBJ databases. The number of clones representing each RFLP-type found in each of the four sampling transects (MC = moorland Calluna vulgaris, TC = transition zone C. vulgaris, FC = forest C. vulgaris and FV = forest Vaccinium myrtillus) at Abernethy forest is included. Number of clones in RFLP Accession Closest 2 FASTA matches Identity% Overlap e score each transect type code MC TC FC FV 126 DQ309116 Ectomycorrhizal root tip (Cenococcum) 1 Ny1.O-1.1 (AF476965) 78.443 668 3.5 e-39 0 0 0 1 Mycena sp. F14061 (AF335445) 82.852 554 3.6 e-39 127 DQ309097 Nolanea conferenda (AF538624) 99.691 648 6.5 e-82 0 0 14 12 Mycena galopus isolate olrim 541 (AY805614) 98.322 596 1.2 e-71 128 DQ309099 Mycena sp. Olrim480 (AY805615) 96.675 421 7.4 e-50 0 0 0 1 Unidentified basidiomycota (AF241332) 85.178 533 3.1 e-47 129 DQ309104 Unidentified basidiomycota (AF241332) 96.508 630 8.0 e-73 0 0 0 2 Mycena sp. Olrim480 (AY805615) 99.111 450 4.9 e-53 130 DQ309237 Mycena sp. F14061 (AF335445) 85.179 560 7.5 e-49 1 1 0 0 Nolanea conferenda (AF538624) 88.920 361 6.0 e-34 131 DQ309167 Unidentified basidiomycota (AF241317) 92.081 442 1.3 e-47 0 1 0 0 Mycena galopus isolate olrim 541 (AY805614) 86.719 512 1.6 e-47 132 DQ309169 Nolanea conferenda (AF538624) 99.339 605 5.5 e-83 0 2 0 0 Mycena galopus isolate olrim 541 (AY805614) 98.407 565 8.5 e-75 133 DQ309193 Mycena sp. F14061 (AF335445) 95.433 416 5.1 e-44 0 0 1 0 Uncultured cf. Mycena sp. Clone d485.19 (AY254873) 94.417 412 1.4 e-42 134 DQ309195 Nolanea sericea strain CBS2 (AF357021) 90.970 598 6.3 e-57 1 0 0 0 Entoloma nitidum (AF335449) 84.375 672 5.4 e-52 135 DQ309203 Nolanea conferenda (AF538624) 99.520 625 3.7 e-85 4 0 1 4 Mycena galopus isolate olrim 541 (AY805614) 98.464 586 1.6 e-76 136 DQ309207 Unidentified basidiomycota (AF241317) 89.322 487 2.0 e-54 5 2 0 0 Mycena aff. murina F14062 (AF335444) 89.004 482 5.3 e-54 137 DQ309222 Mycena sp. F14061 (AF335445) 86.260 655 3.2 e-80 0 2 0 0 Unidentified basidiomycota (AF241317) 89.117 487 2.4 e-65 138 DQ309229 Mycena aff. murina F14062 (AF335444) 89.984 609 7.7 e-55 0 3 2 0 Unidentified basidiomycota (AF241317) 91.466 539 1.2 e-51 139 DQ309168 Uncultured basidiomycete clone K8-15 (AY394671) 85.486 627 1.6 e-50 0 1 0 0 Uncultured basidiomycete clone K7-13 (AY394672) 85.327 627 2.8 e-50 140 DQ309115 Salal root associated fungus UBCTRA 1322.6 (AF284137) 98.515 404 2.1 e-46 0 0 0 1 Salal root associated fungus UBCTRA 1041.2 (AF284135) 97.783 406 6.0 e-46 141 DQ309125 Uncultured fungus clone D20 (AF504850) 98.472 589 7.3 e-70 0 0 5 0 Zygomycete sp. Olrim313 (AY781213) 99.632 544 1.5 e-65 142 DQ309177 Zygomycete sp. Olrim456 (AY805544) 99.632 543 5.0 e-63 0 0 1 0 Zygomycete sp. olrim287 (AY781212) 99.807 519 3.7 e-60 143 DQ309182 Mortierella cf. hyalina (AY157495) 95.859 652 2.5 e-72 0 0 1 0 Zygomycete sp. GFI 1 (AJ608979) 95.433 635 3.5 e-69 144 DQ309234 Mortierella sp. Finse 15-07-00 (ASP541799) 89.385 537 9.9 e-42 0 1 0 0 Zygomycete sp. olrim922 (AY787740) 88.330 497 1.0 e-41 145 DQ309135 Sporopachydermia sp. 91-110 (AF202921) 79.870 154 2.3 e-0 0 0 1 0 Pleurotus dryinus isolate 7 (AY450343) 78.481 158 2.2 e-0 146 DQ309242 Uncultured glomeraceous AM fungus (AY819659) 85.000 500 6.6 e-40 0 1 0 0 Uncultured cf. Tricholoma sp. (AY254876) 85.143 525 2.3 e-33 147 DQ309212 Uncultured glomeraceous AM fungus (AY819659) 84.958 472 2.9 e-41 1 0 0 0 Uncultured cf. Tricholoma sp. (AY254876) 85.282 496 2.3 e-37 148 DQ309220 Uncultured glomeraceous AM fungus (AY819659) 85.800 500 5.7 e-40 3 0 0 0 Uncultured cf. Tricholoma sp. (AY254876) 86.286 525 1.8 e-33
164
The taxonomic positions of selected RFLP-types that had closest sequence matches to members of the H. ericae aggregate or close Helotiales relatives were further investigated using neighbour-joining a nalysis. This analysis (Figure 5.6) groupe d most o f the sequences into one well supported cluster (81% bootst rap su ppo rt) wh ich com p ris ed sequences, including several from the present study, which are lik ely t o b e mem be rs o f the H. ericae aggregate. This cluster comprised seven g roups, wh ile Gr oups I-I V (all with m oderate to high support) i ncluding sequences that correspond to those in clades I-
IV res pectively from Vrålstad et al. (2002b). In terms of sequences fro m the pr es en t study, Group I contained RFLP- types 17 and 18, while Gr oup II I, w hich in cluded mai nly ericoi d mycorrhizal isolate sequences, contained RFLP- types 1 a nd 3 . No seq ue n ces from the present study clustered in Group II or Group I V, wh ich co nta in ed ectomycorrhizal sequences and the sequence for the iso late th at Villarreal-Ruiz et al.
(2004) demonstrated could simultaneously form ericoid and ectomyco rrhizal stru c tur es .
Group V (100% support) contained sequences from an ericoid mycorrhizal fungus isolate d from Rhododendron lochiae (Chapter 4) along w ith se que nces fr om unc ul tu red funga l clones from ectomycorrhizal roots of a non-Ericace ae hos t G. emar ginata. RFLP- types 11, 12 and 14 clustered in G roup VI (67% support) with se que nce s repre se nti ng fungi obtained from a variety of Ericaceae hosts that were orig inal ly p lac ed in cl ad e IV by Vr ålstad et al. (2002b). Group VII (67% support) contained onl y se qu ences fr om the presen t study, including those from RFLP-types 2 and 4-8. The remaining se que n ces from the present study grouped in two clusters that were separa te t o th e m ain gro u p a nd probably represent Helotiales fungi that are distantly related to the other groups (Fig ure 165
5.6). One of these clusters (96% support) contained RFLP-types 15 and 16, while the cluster (55% support) contained RFLP-types 10, 13, 19, 20, 21, 114 and 118.
The taxonomic positions of selected RFLP-types that had closest sequence matches to members or putative members of the Sebacinaceae were further investigated using neighbour-joining analysis. This analysis (Figure 5.7) grouped most of the sequences into two groups. Group I contained only sequences from the GenBank, EMBL and
DDBJ databases that represented multiple unidentified and several known Sebacinaceae fungi, many of which form orchid mycorrhizas and/or ectomycorrhizas. Group II contained the RFLP types from the current study, along with sequences representing
Sebacina calcea (Pers.) Bres., Sebacina grisea (Pers.) Bres. and an unidentified
Sebacinacea. Within this group the sequences for S. calcea and S. grisea formed a separate well supported (100%) subgroup IIa, while the remaining sequences clustered in the well supported (98%) subgroup IIb.
Figure 5.6 Neighbour-joining tree showing the relationships between ITS sequences for selected fungal clone RFLP-types from the moorland Calluna vulgaris, transition zone C. vulgaris, forest C. vulgaris and forest Vaccinium myrtillus transects at Abernethy forest that had FASTA matches with Hymenoscyphus ericae or related sequences, along with selected sequences for H. ericae and related taxa from the GenBank/EMBL/DDBJ databases. The host species is indicated for sequences included from the databases. Groups are represented by Roman numerals while numerical values indicate the bootstrap percentiles from 1000 possible replicates. = confirmed ectomycorrhizal isolate. = confirmed ericoid mycorrhizal isolate. = confirmed ecto/ericoid mycorrhizal isolate. Other sequences are either non-mycorrhizal or have not been tested for their mycorrhizal status. 166
Pinus sylvestris AJ292201 RFLP-type 18 Gaulteria shallon AF149083 G. shallon AF149082 Group I Populus tremula AJ308338 100 G. shallon AF149085 G. shallon AF149084 RFLP-type 17 93 Betula pubescens AJ292199 B. pubescens AJ292200 Group II Calluna vulgaris AF069505 Hymenoscyphus ericae C. vulgaris AJ319078 H. ericae C.vulgaris AJ308337 P. sylvestris AJ430168 P. sylvestris AJ430159 V. angustifolium AJ319077 Scytalidium vaccinii RFLP-type 3 Group III 70 RFLP-type 1 Cephaloziella exiliflora AF069439 H. ericae 21 Empetrum nigrum AJ430104 G. shallon AF149069 G. shallon AF149068 56 C. exiliflora AF069440 H.ericae BH Graffenrieda emarginata AY394684 G. emarginata AY394686 G. emarginata AY394685 Group V 100 G. emarginata AY394687 Rhododendron lochiae AY699657 P.sylvestris AF011327 Cadophora finlandia Conifer AF476977 56 Picea abies AJ292202 81 P. sylvestris AJ430160 60 V. vitis idaea AJ430119 Conifer AF481386 P. abies AJ308340 Group IV Conifer AF476973 62 P. sylvestris AJ430174 65 B. pubescens AJ430144 Quercus robur AJ292203 P. tremula AJ308341 B. pubescens AJ430136 P. sylvestris AY579413 # Deschampsia flexuosa AJ430122 RFLP-type 14 V. myrtillus AJ430113 V. myrtillus AJ430114 RFLP-type 11 Group VI RFLP-type 12 67 100 V. vitis idaea AJ430120 Epacris microphylla AY268197 RFLP-type 5 RFLP-type 2 67 RFLP-type 8 RFLP-type 6 Group VII RFLP-type 4 RFLP-type 7 P. sylvestris AJ430176 C. vulgaris AF252837 RFLP-type 16 96 RFLP-type 15 RFLP-type 9 RFLP-type 114 RFLP-type 13 Helotiales RFLP-type 19 RFLP-type 118 55 RFLP-type 20 RFLP-type 21 RFLP-type 10 Phialocephala fortinii AY3949212 P. fortinii AY033087 0.01 substitutions/site 167
100 Hexalectris spicata AF202728 Sebacina vermifera Caladenia carnea AY643802 Neottia nidus-avis AF465191 Sebacina sp. AJ966750 Sebacina helvelloides Eucalyptus sp. AY093438 Sebacinaceae ectomycorrhiza AY296254 Sebacinaceae sp. SEB534910 Sebacinaceae sp. Neottia nidus-avis AF440664 Sebacina sp. AF465185 Sebacinaceae ectomycorrhiza AJ893265 Sebacinaceae ectomycorrhiza AY143340 Sebacina incrustans AJ879694 Sebacinaceae ectomycorrhiza Neottia nidus-avis AF490395 Sebacina incrustans AJ966753 Sebacina incrustans AF490394 Tremelloscypha gelatinosa AF509964 Sebacinaceae ectomycorrhiza Neottia nidus-avis AF490393 Sebacina epigaea 75 50 AY940652 Sebacinaceae ectomycorrhiza AY940650 Sebacinaceae ectomycorrhiza Group I I Neottia nidus-avis AF440663 Sebacina sp. AY296259 Sebacinaceae sp. SEB534907 Sebacinaceae sp. AY634133 Sebacinaceae sp. AY534208 Sebacinaceae ectomycorrhiza Quercus ilex AY825521 Sebacinaceae ectomycorrhiza AJ549198 Sebacinaceae sp. AY452678 Sebacinaceae sp. Epipactis microphylla AY286193 Sebacinaceae sp. Neottia nidus-avis AF440647 Sebacina sp. 100 AF384862 Tremellodendron pallidum AJ966754 Sebacina epigaea Hexalectris spicata AY243533 84 Hexalectris revoluta AY243532 100 85 AJ966755 Sebacina incrustans Neottia nidus-avis AF490397 Sebacina epigaea AF384860 Efibulobasidium albescens Neottia nidus-avis AF490396 Sebacina allantoidea 100 SCA427408 Sebacina calcea Group IIa SGR427410 Sebacina grisea RFLP 72 RFLP 80 RFLP 96 RFLP 97 RFLP 82 RFLP 76 RFLP 83 RFLP 78 80 RFLP 90 RFLP 91 RFLP 79 71 RFLP 75 RFLP 73 RFLP 74 RFLP 93 83 RFLP 99 Group IIb RFLP 92 RFLP 95 RFLP 81 RFLP 94 RFLP 86 58 RFLP 88 82 RFLP 89 RFLP 87 RFLP 85 98 RFLP 98 99 AY634132 Sebacinaceae sp. RFLP 77 RFLP 84 RFLP 101 89 RFLP 103 RFLP 100 RFLP 102 100 AY880930 Russula pectinatoides AB211275 Russula sororia AY245542 Russula californiensis 0.01 substitutions/site
Figure 5.7 Neighbour-joining tree showing the relationships between ITS sequences for selected RFLP-types from the clone assemblages from Calluna vulgaris and Vaccinium myrtillus root systems at Abernethy forest and selected Sebacinaceae sequences from the GenBank/EMBL/DDBJ databases. Numerical values indicate bootstrap percentiles from 1000 replicates. 168
5.4 Discussion
Based upon the numbers of DGGE bands, fungal species richness in communities associated with roots of Calluna across the moorland : forest gradient and the Vaccinium in the forest transect was similar. DCA of DGGE data clearly grouped the samples according to their transect of origin, suggesting that differences in fungal endophyte community structure and composition exist between the four sampling transects. The endophyte community structure in the moorland and transition zone differed from each other, but the greatest differences were between these two and samples from the C. vulgaris and V. myrtillus forest transects. A separate ordination plot of only data from
DNA marker lanes, however suggested that at least some of the separation observed in the sample ordination plot was a consequence of running samples from individual transects on separate gels. This problem could have been negated by running the samples from each transect randomly across the lanes of the four DGGE gels.
Based on the number of T-RFs identified, T-RFLP analysis detected a greater level of fungal endophyte species richness than DGGE gels. This was not unexpected since T-
RFLP analyses have previously been shown to distinguish a greater level of species richness than DGGE from the same environmental samples (eg. Moeseneder et al., 1999).
The difference in species richness observed using both techniques might partially be explained by pseudo-T-RFs that can occur due to either restriction enzyme malfunction at sub-optimal conditions leading to cutting at additional secondary sites or the presence of single-stranded amplicons both leading to an overestimation of T-RFs (Moeseneder et al.,
1999; Egert and Friedrich, 2003). Given that restriction enzyme digests were carried out 169 according to manufacturer’s instructions, this seems unlikely to be the case. It seems more likely that fluorescent detection of T-RFLP is simply more sensitive than the silver staining of DGGE in terms of detecting rare sequence types (Moeseneder et al., 1999).
When the data obtained with the two techniques were compared for individual sampling transects (data not shown), the level of species richness detected using T-RFLP was similar to or greater than that detected using DGGE for all but the transition zone transect. In the latter transect, the omission of two samples from the analysis (due to unsuccessful digestion) may have reduced the total number of T-RFs identified, and this might explain the decreased species richness. Despite this, ANOVA indicated there were no significant differences in the levels of species richness detected from each transect using T-RFLP, which confirmed the observation made on the basis of DGGE results.
DCA of T-RFLP data indicated that fungal endophyte community structure and composition varied between the four sampling transects. As was suggested by the DGGE results, the endophyte community structure in the moorland and transition zone differed from each other, however, the greatest differences were between these two and samples from the C. vulgaris and V. myrtillus forest transects. TWINSPAN and indicator species analysis of T-RF data confirmed the DCA results whereby the samples were clustered into four groups that consisted primarily of moorland, transition zone, forest C. vulgaris and forest V. myrtillus samples respectively. Notwithstanding the potential confounding effects of running gels from individual transects separately, the same general pattern of sample separation according to their transect and host origin was obtained by DCA of both DGGE and T-RFLP data. Not only does this suggest that the fungal endophyte 170 communities in the roots of C. vulgaris vary between the moorland, transition zone and forest sampling transects, it also indicates that the fungal endophyte communities in the roots of forest dwelling V. myrtillus and C. vulgaris are different.
These results support the findings of Anderson et al. (2003) who used DGGE to investigate soil fungal community structure and diversity along a moorland: forest gradient at the same sampling location. They identified a distinct profile boundary between the transition zone and forest samples, and suggested that the presence of P. sylvestris saplings in the transition zone had little effect on soil fungal community structure. Previous vegetation surveys at the Abernethy forest site identified V. myrtillus as the dominant Ericaceae shrub at the forest end of each transect, while C. vulgaris was dominant at the moorland end, although it extended into the forest (Chapman et al.,
2003). Anderson et al. (2003) suggested that this difference in Ericaceae vegetation structure between the forest and moorland, particularly in terms of potential specificity for symbionts between host species, may play a significant role in changing the respective soil fungal communities. Data from the present study suggest that fungal communities from roots of forest dwelling C. vulgaris and V. myrtillus were more similar to each other than to those from C. vulgaris roots in the transition zone or moorland. It therefore seems likely that the changing Ericaceae vegetation and potential host specificity of symbionts are not entirely responsible for changes observed between fungal communities from soil or Ericaceae roots in the moorland/transition zone and forest.
This notwithstanding, the fact that the endophyte communities from forest dwelling C. 171 vulgaris and V. myrtillus show some difference suggests that there may be a level of host specificity for symbionts.
Cloning confirmed the distinction observed between endophyte communities from each transect, with 20 - 26 unique RFLP-types being observed in clone assemblages from each sampling transect. DNA sequencing of the clone RFLP-types revealed that typical mycorrhizal symbionts of Ericaceae, particularly fungi related to members of the H. ericae aggregate and other closely related Helotiales ascomycetes, were common in C. vulgaris roots throughout the moorland : forest gradient and V. myrtillus roots in the forest. The H. ericae aggregate sequences used in the neighbor-joining analysis from the present study were separated into seven groups, which suggests that the H. ericae aggregate may be more complex than the four clades suggested by Vrålstad et al.
(2002b). The analysis placed two sequences from the current study in each of two groups along with sequences from clades I and III from Vrålstad et al. (2002b). RFLP-type 3, which was frequently identified in the clone assemblages from all transects, grouped with clade III sequences that represent known and putative ericoid mycorrhizal fungi. This
RFLP-type may therefore represent an ericoid mycorrhizal isolate that is common in both
C. vulgaris and V. myrtillus across the entire moorland : forest gradient. Interestingly, no sequences from the current study clustered with ectomycorrhizal sequences from clade IV of the H. ericae aggregate. Several sequences from the current study were placed in a
‘new’ group with sequences from isolates that were obtained solely from Ericaceae hosts that had previously been grouped in clade IV along with ectomycorrhizal sequences
(Vrålstad et al., 2002b). This supports the notion that isolates from ectomycorrhizal hosts 172 in the H. ericae aggregate are taxonomically distinct from those obtained from Ericaceae hosts (Vrålstad et al., 2002a). However, until isolates representing all the sequences from the aggregate are tested for their colonising ability with both ectomycorrhizal and ericoid mycorrhizal hosts, the true extent of their mycorrhizal status cannot be confirmed.
Villarreal-Ruiz et al. (2004) demonstrated that a single isolate from clade IV simultaneously developed ericoid mycorrhizal coils in epidermal cells of V. myrtillus and an ectomycorrhizal Hartig net and mantle with P. sylvestris, supporting the suggestion that ericoid mycorrhizal and ectomycorrhizal plants may share some common symbiotic partners from the H. ericae aggregate in forests where plant communities comprise both
Ericaceae and ectomycorrhizal host plants Vrålstad et al. (2000). This is not supported by data from the present study, however, since no sequences grouped with ectomycorrhizal sequences. The most common H. ericae aggregate sequences from the present study were obtained from the clone assemblages from each of the transects. Together with the fact that these sequences were obtained from both C. vulgaris and V. myrtillus, suggests that they show little specificity and may not have evolved to preferentially colonise either forest or moorland dwelling Ericaceae hosts at the Abernethy site.
Multiple sequences obtained from C. vulgaris and V. myrtillus hosts in the present study grouped separately to the those sequences in the main group of the neighbour-joining tree and therefore probably represent closely related Helotiales fungi rather than true members of the H. ericae aggregate. Helotiales fungi that are closely related to, but distinct from, the H. ericae aggregate have been frequently isolated from a variety of Ericaceae hosts 173 and locations (see Chapter 2 and 4) and their presence in the current study provides further evidence of their widespread distribution and limited host specificity. Many of these Helotiales isolates have been shown to form ericoid mycorrhizal structures with
Ericaceae hosts in inoculation trials (see Chapter 2 and 4), suggesting that the unidentified Helotiales sequences from the current study may represent important ericoid mycorrhizal fungi across the moorland : forest gradient.
A diverse array of Sebacinaceae-like sequences were obtained from V. myrtillus and C. vulgaris across all sampling transects. The rDNA ITS region in this group of fungi is extremely variable and, therefore, most phylogenetic studies of this group of fungi use the rDNA LSU gene to infer taxonomic relationships (Weiss et al., 2004). As a result of this, a limited range of sequences were available to use in the Sebacinaceae phylogenetic analysis in the current study. Furthermore, based on the results of this analysis, no identities could be ascertained for the sequences included from the current study, other than that they probably represent Sebacinaceae taxa. The fact that Sebacinaceae have been proposed as possible ericoid mycorrhizal symbionts in the Ericaceae species
Gaultheria shallon (Allen et al., 2003) suggests that these fungi may represent ericoid mycorrhizal symbionts of both hosts across all transects in the current study.
Sebacinaceae fungi also form ectomycorrhizal associations (Weiss et al., 2004) and therefore those sequences from the transition zone may represent fungi that are an important source of ectomycorrhizal inoculum for developing P. sylvestris saplings in the areas of forest expansion. 174
Sebacinaceae can simultaneously form orchid and ectomycorrhizal associations that are linked by a common hyphal network (Weiss et al., 2004). Thus, it is possibile that ericoid mycorrhizal Sebacinaceae isolates could simultaneously form ectomycorrhizal associations with P. sylvestris saplings. If this were to be the case, carbon transfer from dominant Ericaceae hosts to establishing ectomycorrhizal hosts via a common mycelial network in the transitional habitats, might be important in context of expansion of P. sylvestris forests. Further studies are clearly required to ascertain the mycorrhizal status of the Sebacinaceae taxa identified in the present study and their potential interactions with both ericoid and ectomycorrhizal hosts.
Multiple sequences that matched common basidiomycete ectomycorrhizal fungi, particularly from Atheliaceae and Thelephoraceae, were present in clone assemblages mainly from the forest understorey transects. Similarly, Anderson et al. (2003) found that many bands in DGGE profiles of soil fungal communities that were unique to the forest end of their transects at Abernethy forest represented ectomycorrhizal basidiomycetes. The Atheilaiceae and Thelephoraceae basidiomycets have previously been regarded exclusively to form ectomycorrhizal associations with forest trees (Smith and Read, 1997). Ectomycorrhizal fungi are also known to form mycelial networks that interconnect individual host trees (Simard et al., 1997) and the data presented here suggest that understorey Ericaceae could possibly be integrated into these networks. Such putative symbiont sharing could be functionally significant in carbon and nutrient cycling in forest ecosystems where light might be the limiting factor for understorey Ericaceae vegetation. Ectomycorrhizal fungi of forest trees are known to have been ‘captured’ by 175 understorey green orchids and by members of the Ericaceae in subfamily
Monotropoideae in fully or partial mycoheterotrophic relationships (Bidartonndo, 2005).
Monotropoideae are considered a derived lineage which lack the hair roots characteristic of many Ericaceae (Kron et al., 2002). The data from the present study represents the first indication that common ectomycorrhizal basidiomycetes associate with hair roots of fully autotrophic woody perennials, such as C. vulgaris, and V. myrtillus in the subfamilies
Ericoideae and Vaccinoideae respectively. The potential for evolution of mycoheterotrophy may thus extend to these lineages within Ericaceae and, if so, would establish a new paradigm as to how such associations may have evolved in this important and diverse plant family. It is acknowledged, however, that the ectomycorrhizal basidiomycetes may not exist as endophytes of the forest dwelling V. myrtillus and C. vulgaris perse, and their presence might simply reflect some sort of facultative or opportunistic infection of roots. Further work will be required to elucidate the nature of these associations.
176
CHAPTER 6
General Discussion
The data presented in this thesis demonstrated that root systems of the Australian
Ericaceae species Epacris pulchella and Rhododendron lochiae harbour diverse communities of both ericoid mycorrhizal and other root-associated fungi. This is consistent with other studies of Ericaceae root associated fungal communities in both the northern and southern hemispheres (eg. Perotto, 1996; Monreal et al., 1999; Midgley,
2003; Williams et al., 2004). A greater number of RFLP-types were cultured from the R. lochiae plants than the E. pulchella plants. Differences in edaphic conditions between habitats (tropical montane cloud forest and dry sclerophyll forests for R. lochiae and E. pulchella respectively) or differences in the ability of the two host taxa to sustain fungal endophytes were suggested as possible explanations for this. However, differences in the number of root samples taken for each host plant make it difficult to be sure without further investigation. As is the case for other northern hemisphere and Australian
Ericaceae taxa (Perotto et al., 1996; Xiao and Berch 1996; Chambers et al., 2000;
Sharples et al., 2000; Liu et al., 1998; Midgley et al., 2002, 2004c), the root systems of each of the E. pulchella and R. lochiae plants were colonised by a few dominant RFLP- types, mostly identified as ericoid mycorrhizal endophytes, and a greater number of less frequently isolated types. The exception to this was an RFLP-type that was identified, based on sequence matches and phylogenetic analyses, as a probable member of the
Xylaria genus which was frequently isolated from the roots of all three from R. lochiae 177 plants. Xylaria spp. are common tropical endophytes whose functions in terms of a host plant association remain largely unknown.
The common RFLP-types, along with the less common types obtained from E. pulchella, were putatively identified on the basis of ITS sequence matches as ericoid mycorrhizal endophytes and then confirmed as such by gnotobiotic inoculation experiments with
Woollsia pungens as a host. A total of fourteen RFLP-types from the three host plants formed typical ericoid mycorrhizal structures in gnotobiotic culture experiments, and these RFLP-types encompassed a substantial proportion of the total isolate assemblage from each plant. Similar to the endophytes from E. pulchella, most of those frequently isolated, along with some of the less common RFLP-types, from R. lochiae were putatively identified and subsequently confirmed as ericoid mycorrhizal fungi when grown in gnotobiotic culture with Vaccinium macrocarpon. Interestingly, none of the eight RFLP-types that were indentified as Xylariaceae, including the common RL RFLP- type 14, formed structures resembling those of ericoid mycorrhizas when grown in gnotobiotic culture with V. macrocarpon. However, this does not necessarily preclude the possibility that fungi representing these RFLP-types, or any others that did not form typical ericoid mycorrhizal structures, may form beneficial associations with other
Ericaceae hosts.
While many of the dominant RFLP-types from both host species were grouped with unidentified Helotiales-like fungi that are known to be common ericoid mycorrhizal fungi in the northern and southern hemisphere (eg. Xiao and Berch 1996; Chambers et al., 178
2000; Sharples et al., 2000; Liu et al., 1998; Midgley et al., 2002; 2004c), several RFLP- types were of particular interest. The frequently cultured EP RFLP-types 13 and 15 from
E. pulchela are likely to represent, based on sequence matches and phylogenetic analyses,
Oidiodendron sp. closely related to O. maius. Oidiodendron spp. are considered common ericoid mycorrhizal fungi in many northern hemisphere Ericaceae but only a single isolate had been previously identified in an Australian Ericaceae host (W. pungens)
(Chambers et al., 2000). Data from the present investigation suggest that Oidiodendron- like fungi commonly occupy roots of E. pulchella and are likely to be important mycorrhizal endophytes.
Sequence matches and phylogenetic analysis suggested that RL RFLP-type 19 from R. lochiae is part of the Hymenoscyphus ericae aggregate, and this is the first report of a member of this aggregate being cultured from an Ericaceae host in Australia. Members of the H. ericae aggregate are common ericoid mycorrhizal endophytes of Northern
Hemisphere hosts (Vrålstad et al., 2002), however until now no fungi from the aggregate have been observed in roots of Australian Ericaceae. Moreover, this RFLP type appears to be closely related to a group of ectomycorrhiza-like fungi from the H. ericae aggregate observed in the roots of Graffenrieda emarginata in a tropical cloud forest in Ecuador
(Haug et al., 2004). These data suggest that this taxon may be widespread in tropical cloud forests and that it contains both ericoid mycorrhiza-forming and ectomycorrhizal isolates.
179
Results from phylogenetic analyses of the RFLP-types (from both E. pulchella in the sclerophyll forest habitat and R. lochiae in the TMCF habitat) that were identified as members of a taxonomically unplaced group of Helotiales-like ericoid mycorrhizal fungi further support the suggestion that ericoid mycorrhizal fungi show little, if any, specificity for different Ericaceae hosts or habitats (Cairney and Ashford, 2002). The geographically widespread distribution of these Helotiales-like fungi, along with the discovery of a member of the H. ericae aggregate in R. lochiae and the confirmation of
Oidiodendron spp. in Australian Ericaceae hosts, highlights the similarities between northern and southern hemisphere ericoid mycorrhizal fungi. This provides further advocacy for a proposed monophyletic origin of plants that form ericoid mycorrhizas
(Cairney, 2000) and emphasises a probable radiation of ericoid mycorrhizas from a common ancestral symbiosis. The Ericaceae centre of diversity is the southern hemisphere and this is thought to reflect a Godwanan origin of taxa ancestral to the
Ericaceae, followed by subsequent spread northwards (Cullings, 1996).
The clone assemblages from E. pulchella contained several more RFLP-types than their corresponding cultured isolate assemblages, and this is consistent with other investigations that have reported greater taxonomic richness in fungal assemblages from other environments when determined by direct DNA extraction compared to cultured isolates (eg. Borneman and Hartin, 2000; Viaud et al., 2000; Hunt et al., 2004).
Although the combined cultured and clone assemblages each contained a suite of unique
RFLP-types, those cultured most frequently were also the most abundant clones. These
RFLP-types, along with most others that were common between the culture and clone 180 assemblages, were identified as ericoid mycorrhizal fungi. This, along with the fact that the most frequently observed clones had the same sequence matches to the common culture RFLP-types that formed mycorrhizas, suggested that cloning and culturing give a similar representation of the ericoid mycorrhizal fungal community in E. pulchella.
Several cloned RFLP-types that were identified as basidiomycetes, including a putative
Tricholomataceae taxon, represented a substantial component (< 23%) of the EP1 and
EP2 clone assemblages. In contrast, only a few putative basidiomycete RFLP-types, that represented a small proportion of the cultured assemblages, were identified from the three
E. pulchella plants. In a similar investigation, Allen et al. (2003), found that, although only 5% of cultured isolates from Gaultheria shallon were basidiomycetes, 59% of sequences cloned from DNA extracted directly from the same root systems had strong sequence identity to the basidiomycete genus Sebacina. Thus, in contrast to the present study, there was a clear difference in the most commonly encountered taxa in the cultured isolate and clone assemblages, emphasising that the results from one study cannot be extrapolated to another host and/or habitat.
Most of the bands in DGGE profiles of DNA extracted directly from E. pulchella and R. lochiae roots matched those from cultured RFLP-types. Many of the DGGE bands that were absent from profiles from cultured isolates were relatively faint, suggesting that the relative abundance of the taxa they represent was probably low. Of these, several probably also represented ericoid mycorrhizal endophytes, while several others were putatively identified as ascomycetes, basidiomycetes and Glomerales. It is difficult to 181 predict the functional status of the putative basidiomycetes and Glomerales identified from these DGGE bands and in the DNA cloned from E. pulchella roots. Although these could possibly represent unculturable mycorrhizal fungi, on current evidence the basidiomycetes, at least, seem likely to be saprotrophs. The absence of these fungi from the cultured isolate assemblage may simply reflect an inability to grow on the medium used, particularly in the case of the putative Glomerales which are widely regarded as unculturable (Smith and Read, 1997), or their presence in or on roots as inactive propagules.
Conversely, several RFLP-types from the cultured isolate assemblages were not observed in DGGE profiles of DNA extracted from E. pulchella and R. lochiae roots, however only one or two isolates were obtained for these RFLP-types. The notable exception to this was RL RFLP-type 14, a Xylariaceae taxon that constituted ca 15–25% of the cultured isolate assemblage from each R. lochiae root system and was absent from the DGGE profiles of DNA extracted from root pieces. The absence of this RFLP type may indicate that, although widespread in the root systems, only a small amount of mycelium was present relative to that present in the form of ericoid mycorrhizal coils. Furthermore, this
RFLP-type grew considerably faster in culture than most of the other fungi isolated and may have suppressed other fungi during isolation and resulted in an overestimation of its importance. The likely overestimation of this RFLP-type in the cultured assemblages highlights a potential limitation of using culturing to investigate communities of root- dwelling fungi. Significantly, the most intense bands in the directly extracted DNA profiles matched those of the most commonly cultured ericoid mycorrhizal RFLP-types 182 which confirmed that these fungi were an important component of the E. pulchella and R. lochiae root fungal assemblages.
The DGGE data for E. pulchella were thus broadly supportive of those obtained by cloning from DNA extracted directly from the same roots. Furthermore, results for E. pulchella indicated that the culture based method and DGGE were similarly efficacious in their abilities to identify the most abundant members of root-associated fungal communities from E. pulchella. However, the data for the R. lochiae suggest, as highlighted by Allen et al. (2003), that this is not always true. Overall, data generated from both culturing and direct DNA extraction, coupled with either cloning or DGGE, were equally useful in identifying the majority of taxa from the Ericaceae fungal root endophyte communities, which on the basis of sequence similarities and inoculation experiments appear to be ericoid mycorrhizal fungi. However, given the varying DGGE results from Chapters 2 and 4 of this thesis and the observation that each technique identified a unique suite of less abundant fungi, along with the fact that both culture- based and culture-independent methods introduce potential bias (Bridge and Spooner,
2001; Anderson and Cairney, 2004), a combination of approaches should probably be adopted in future investigations regarding the diversity of fungal root endophytes in
Ericaceae.
Plants in the family Ericaceae generally inhabit heathland or forest habitats where soils are extremely nutrient deficient (Read, 1996; Cairney and Meharg, 2003) and the dry sclerophyll forest habitats from which the E. pulchella plants in this study were collected 183 are no exception to this. It is widely believed that the abundance and success of
Ericaceae in these habitats is primarily due to benefits, in terms of access to particularly organic nitrogen stores, conferred to the host plant by ericoid mycorrhizal fungi (Read,
1996; Perotto et al., 2002). Aside from plant material, a significant pool of soil organic nitrogen is found in fungal mycelium as chitin in heathland habitats (Kerley and Read,
1997).
Chitin utilisation has been demonstrated for H. ericae (Kerley and Read, 1995), however the chitinolytic potential of other fungal endophyte taxa from Ericaceae roots was previously unexplored. The work in Chapter 3 demonstrated for the first time the chitinolytic abilities of fungal root endophytes from Ericaceae in the southern hemisphere, along with confirming this ability for H. ericae. Endo- and exo-acting chitinolytic activities were demonstrated for five ericoid mycorrhizal fungi, one non- mycorrhizal ascomycete and one non-mycorrhizal basidiomycete cultured from the three
E. pulchella plants. Only exo-chitinolytic activity was detected for a non-mycorrhizal ascomycete and basidiomycete from the three E. pulchella plants. Exo-chitinolytic activities were generally higher than endo-chitinolytic activities for all isolates tested, which is consistent with the data of Hodge et al. (1995) who found extracellular chitinolytic activity was predominant in ectomycorrhizal and non-mycorrhizal basidiomycetes. Highest chitinolytic activities recorded for the five ericoid mycorrhizal fungi from E. pulchella and for H. ericae were of the same order of magnitude, suggesting that their chitinolytic potential may be broadly similar.
184
Reports on the effects of glucose and a chitin source on fungal chitinolytic activities are contradictory. Some evidence suggests that expression of chitinolytic activity may not be inhibited by the presence of glucose (eg. Mach et al., 1999), and this supports the suggestion of Kerley and Read (1995) that carbon supplied by the plant host would render ericoid mycorrhizal endophytes more competitive than relatively carbon-starved non-mycorrhizal fungi in the roots of Ericaceae. Such an advantage would require that the supply of carbon from the host did not repress chitinase production. Suppression of chitinolytic activity in EP RFLP-type 1 in the presence of glucose does not appear to support this suggestion. However, there is evidence to suggest that ericoid mycorrhizal fungi produce chitinase activities during symbiosis (Kerley and Read, 1995; 1997).
Therefore further work is clearly required to fully understand carbon source and host plant effects on chitinase production by ericoid mycorrhizal fungi. This notwithstanding, results from the present study suggest that ericoid mycorrhizal fungi from E. pulchella are potentially capable of utilising chitin as a nitrogen source which could, in turn, be passed on to their hosts. While it is not possible to elucidate the precise contribution the chitinolytic activities displayed by the ericoid mycorrhizal fungi from this study may make in terms of Ericaceae host nitrogen acquisition, the results provide further support for the idea that ericoid mycorrhizal fungi from the southern hemisphere posses a similar enzymatic potential to their northern hemisphere counterparts. However, this clearly needs to be further examined for a variety of enzymes, such as proteases, with a range of fungal taxa from a variety of hosts and habitats.
185
The above highlights a limitation of using isolated fungi and model substrates, as has been the case for most studies regarding nutrient acquisition by ericoid mycorrhizal fungi, to infer the abilities of fungi for in situ nutrient mobilization. The key to understanding the role of ericoid mycorrhizal fungi in nature lies in determining the extent to which their nutrient mobilising potentials are expressed in the mycorrhizal condition and when colonised plants are exposed to substrates more similar to those likely to occur in their natural habitats. Studies of this nature have been carried out using Vaccinium plants colonised with H. ericae to assess their potential in acquiring nitrogen from both mycelial and plant-derived necromass (Kerley and Read, 1997; 1998). In both instances, colonisation by H. ericae conferred host plant access to the nutrient. Previously, work of this nature has not been possible with Australian Ericaceae due to difficulties with their propagation. The success achieved in using W. pungens as a host for ericoid mycorrhizal inoculation trials in the present study means that such work may now be possible.
Moreover, the heterogeneous medium used was more similar to conditions in nature compared to media adopted in other such studies, emphasizing that there is now an opportunity to conduct more meaningful in planta physiological experiments using this model system. Such experiments would be useful for initially determining if ericoid mycorrhizal fungi have the ability to enhance the fitness of Australian Ericaceae hosts from the subfamily Stypheloideae in terms of nutrient acquisition, as this is yet to be confirmed and is assumed based only on circuitous evidence. Furthermore, investigations could be undertaken regarding the nutrient mobilising potentials of these mycorrhizal fungi, specifically considering variations in the type and amount of the nutrient sources they are able to utilise and pass on to their hosts. Such investigations would potentially 186 reveal much about the evolution of ericoid mycorrhizas and their roles in structuring plant communities in the northern, and, particularly, the southern hemisphere.
The work described in Chapter 5 compared, using DGGE, T-RFLP and cloning, the community structure of fungi associated with roots of understorey Calluna vulgaris and
Vaccinium myrtillus in a Pinus sylvestris-dominated forest, along with C. vulgaris from a bordering transition zone that contained P. sylvestris saplings and an open moorland in
North-East Scotland. Multivariate analyses of both DGGE and T-RFLP data showed that the fungal endophyte communities in the roots of C. vulgaris vary between the moorland, transition zone and forest sampling transects and that the fungal endophyte communities in the roots of understorey V. myrtillus and C. vulgaris are different. Using both methods, the endophyte community structure in roots of C. vulgaris in the moorland and transition zone were found to differ from each other, however, the greatest differences were between these two and samples from the C. vulgaris and V. myrtillus forest transects. This suggests that differences in the community structure of Ericaceae vegetation across the moorland : forest gradient and potential differences in Ericaceae host specificity of symbionts were not primarily responsible for changes observed between fungal communities from Ericaceae roots in the moorland/transition zone and forest habitats. It thus seems reasonable to suggest that differences in edaphic and environmental conditions may produce different selection pressures on both Ericaceae hosts and their root-associated fungal endophytes across the moorland : forest gradient.
This is supported by the results of Chapman et al. (2001) who identified clear distinctions 187 in soil physical, chemical and biological properties between forest and transition zone/moorland habitats at the same sampling site.
Phylogenetic studies of H. ericae sequences, including those from the present study, revealed that the aggregate is more complex than the four clades suggested by Vrålstad et al. (2002b). While sequences from the present study grouped with known ericoid mycorrhizal isolates, interestingly and in contrast to recent suggestion (Villarreal-Ruiz et al., 2004), no sequences from this study grouped with ectomycorrhizal H. ericae sequences. This implies that the H. ericae aggregate isolates obtained from ectomycorrhizal hosts are not frequent colonisers of understorey Ericaceae hosts at the field site. These findings add support to an earlier suggestion that isolates from ectomycorrhizal hosts in the H. ericae aggregate are taxonomically distinct from those obtained from Ericaceae hosts (Vrålstad et al., 2002a). Furthermore, it suggests that the isolate Villarreal-Ruiz et al. (2004) identified as forming both ectomycorrhizal and ericoid mycorrhizal associations may be an exception rather than the rule. Equally, the possibility that fungal isolates associate differently under laboratory and natural field conditions with potential mycorrhizal hosts can not be discounted. It is important to emphasise, however, that until an H.ericae isolate is tested for its ability to form ectomycorrhizal and ericoid mycorrhizal associations with appropriate hosts in gnotobiotic culture, its taxonomic position within the aggregate is only likely to imply, but not indicate for certain, its true mycorrhizal potential.
188
Allen et al. (2003) identified unculturabale Sebacinaceae fungi as probable ericoid mycorrhizal fungi, and Sebacinaceae sequences were also obtained from V. myrtillus and
C. vulgaris across all sampling transects in the present study. The frequent observation of these fungi, particularly in the moorland transect, suggests that the multitude of studies which have investigated ericoid mycorrhizal diversity from the same hosts in similar northern hemisphere moorland habitats may have overlooked important ericoid mycorrhizal fungi by using culturing as the sole method of isolation. It would probably prove beneficial to use methods based on direct DNA extraction from host roots to investigate ericoid mycorrhizal diversity at sites other than the Abernethy forest in northern hemisphere moorland habitats to investigate the presence of previously uncultured fungi such as members of the Sebacinaceae. The existence of such fungi in
Ericaceae roots across a broader geographic range could change the current view of ericoid mycorrhizal fungal diversity particularly in terms of the perceived dominance of
H. ericae and O. maius in northern hemisphere habitats.
The ability of some Sebacinaceae to form ectomycorrhizal associations suggests that they may represent a possible source of ectomycorrhizal inoculum for P. sylvestris saplings in the transitional zone at the Abernethy forest site. Furthermore, the ability of these fungi to simultaneously form orchid and ectomycorrhizal associations linked by a common hyphal network (Weiss et al., 2004) raises the intriguing possibility that ericoid mycorrhizal Sebacinaceae isolates could simultaneously form ectomycorrhizal associations with P. sylvestris saplings. This potential association could possibly provide 189 nutritional benefits to developing ectomycorrhizal saplings in the transition zone habitat dominated by Ericaceae. Clearly further studies are required to ascertain the mycorrhizal status of the Sebacinaceae identified in the present study with the view of understanding the interactions between these fungi with both ericoid and ectomycorrhizal hosts.
Of particular interest were multiple sequences that were obtained predominately from the forest understorey sample transects that matched common basidiomycete ectomycorrhizal fungi. In trying to explain differences between DGGE profiles representing soil fungi across a moorland : forest gradient at the Abernethy forest site, Anderson et al. (2003) identified basidiomycete ectomycorrhizal fungi in the forest end as the likely source of variation between community profiles. Similarly, these fungi may be responsible for some of the variation observed between fungal community profiles from Ericaceae roots in the present study. Furthermore, the data presented in this thesis show that understorey
Ericaceae could possibly be integrated into mycelial networks that interconnect ectomycorrhizal host trees. The possibility of such symbiont sharing is intriguing and could be functionally significant in carbon and nutrient cycling in forest ecosystems where light might be the limiting factor for understorey Ericaceae vegetation. However, it is acknowledged that it is equally likely that these fungi may not form beneficial endophytic associations within the understorey Ericaceae roots, but rather may represent inactive propagules or opportunistic colonisers in or on the sampled roots. This notwithstanding, ectomycorrhizal fungi of forest trees are known to have been ‘captured’ by understorey members of the Ericaceae subfamily Monotropoideae in mycoheterotrophic relationships (Bidartonndo, 2005). The results from the current study 190 are the first indication that common ectomycorrhizal basidiomycetes may associate with roots of the Ericaceae, C. vulgaris, and V. myrtillus in the subfamilies Ericoideae and
Vaccinoideae respectively and thus the potential for evolution of mycoheterotrophy may extend to these lineages within Ericaceae.
Although speculation regarding the possible functions of the basidiomycete fungi, including the Sebacinaceae, identified in Ericaceae roots at the Abernethy forest site have been discussed, in the future it will be useful to attempt to culture these fungi. Many basidiomycetes are considered unculturable, however some reports suggest members of the Thelephoraceae, to which many basidiomycete sequences from the present study had an affinity with, along with some members of the Sebacinaceae can be maintained in pure culture (eg. Finlay et al., 1992; Weiss et al., 2004). This suggests that there is at least the possibility of culturing some of the basidiomycetes identified in the current study.
Obtaining pure cultures of these fungi would allow a multitude of novel investigations to be carried out which would expand what is known about ecosystem processes in
Ericaceae habitats. Such studies might consider, for example, their functional capabilities in terms of forming beneficial mycorrhizal associations with Ericaceae hosts and how this might allow Ericaceae hosts to interact with potential ectomycorrhizal host species via common hyphal networks.
Despite the fact that Ericaceae are a common component of the understorey of many forest habitats worldwide, investigations of ericoid mycorrhizal fungal diversity have largely focused on heathland habitats and the current study is the first to comprehensively 191 examine the community structure of Ericaceae root associated fungi in a forest habitat. In the current study, the fungal communities from Ericaceae roots in the forest, adjacent transition zone and moorland habitats appear to be dominated by the same few broad fungal groups, including members of the H. ericae aggregate, Helotiales-like fungi and
Sebacinaceae, however, many other less common taxa were also identified in the root systems from each habitat. This pattern, where a few taxa dominate the ericoid mycorrhizal fungal community is similar to that observed for other investigations of
Ericaceae root associated fungal communities including the work in Chapters 2 and 4.
Ectomycorrhizal fungal communities have been observed to have similar patterns of relative abundance. For example, Gehring et al. (1998) investigated ectomycorrhizal communities on mycorrhizal root tips of Pinus edulis Engelm. The study indicated that at some field sites colonisation of root tips by a single taxon accounted for nearly half of the root tips sampled at a single site. Similarly, Natarajan et al. (1992) observed that Amanita muscaria L. Fr. colonised a similar proportion of root tips from Pinus patula Schiede,
Schltdl. and Cham. seedlings in an Indian forestry plantation. Indeed, these patterns of relative abundance are common to other organisms including plant, animal and bacterial communities in a variety of ecosystems worldwide (Huhta, 2002; Labaune and Magnin,
2002).
Regardless of the dominance of a few fungal taxa in the communities of individual mycorrhizal host root systems or habitats, it is likely, based on a number of studies, that the diversity of fungi reflects, at both the inter- and intraspecific level, a significant degree of functional diversity in terms of nutrient utilization (eg. Anderson et al., 1999; 192
Whittaker and Cairney, 2001). There is increasing evidence to suggest that increased mycorrhizal fungal diversity may strongly affect host fitness, plant community structure and ecosystem processes. Baxter and Dighton (2001), for example, grew Betula populfolia Marshall seedlings with one to four ectomycorrhizal fungi and examined the biomass and phosphorus uptake in seedlings. The study demonstrated that greater ectomycorrhizal diversity was positively correlated with root biomass and increased phosphorus uptake in seedlings. Although to date no studies have examined the effects of ericoid mycorrhizal fungal diversity on host plant function, it is possible that colonisation by multiple species and/or multiple genotypes of single species might facilitate increased fitness of host plants. Having different taxa or genotypes which have differing abilities to access and utilise different niches within the soil may result in enhanced host fitness and overall functioning of the symbiosis. This may be vital for Australian Ericaceae in heathland and sclerophyll forest habitats as it is well documented that the relative quantities and composition of various organic nitrogen sources in soils of these habitats vary considerably with respect to biotic and abiotic factors such as fire, species composition and waterlogging (Adams and Attiwill, 1982; Erskine et al., 1996; Schmidt and Stewart, 1997). Given that this heterogenous mixture of nitrogen sources may vary in both chemical composition and temporal availability, intra- and interspecific diversity of ericoid mycorrhizal fungi may be functionally important to the success of Ericaceae in these Australian soils.
Functional diversity in communities of ericoid mycorrhizal fungi may also contribute to maintaining balanced ecosystem processes, especially nutrient cycling, at particularly a 193 habitat scale. For example, a taxonomically, and possibly functionally, diverse range of ericoid mycorrhizal fungi associated with understorey Ericaceae may be important in terms of nutrient cycling in forest habitats that are also occupied by ectomycorrhizal hosts, such as the pine forests at Abernethy or the sclerophyll forests in eastern Australia described in the current study. This may be particularly relevant considering that when comparisons are made between ecto- and ericoid mycorrhizal fungi, the abilities of the former group to access complexed organic nutrient sources are generally lower than those of ericoid mycorrhizal fungi. (eg. Bending and Read, 1996a). Thus, the diverse range of ericoid mycorrhizal fungi within the forest communities described in this thesis may be critical in maintaining a balance in nutrient cycling that might not be possible if ectomycorrhizal and saprotrophic fungi were solely responsible for such a role.
Furthermore, on a larger scale, nutrient cycling by ericoid mycorrhizal fungi resulting in the removal of nitrogen and phosphorus bound to organic complexes may increase the
C:N and C:P ratios of soil residues in their habitats (Read and Perez-Moreno, 2003). This might contribute to soil carbon retention which ultimately influences the carbon source- sink relationships upon which global climate systems depend (Read and Perez-Moreno,
2003).
Aside from influencing host fitness and ecosystem functioning, a broad spectrum of ericoid mycorrhizal fungi may be potentially important in terms of dictating the community structure of a habitat by altering the outcomes of competitive interactions between plant taxa. Several studies have demonstrated that the presence or absence of arbuscular mycorrhizal fungi can alter plant community structure (eg. O'Conner et al., 194
2002). Thus, the diverse nutrient acquiring abilities associated with a broad group of ericoid mycorrhizal fungi may aid their hosts in competitively excluding other plant groups less well equipped to deal with a range of bound organic nutrient sources resulting in habitats, such as moorlands, being dominated by Ericaceae taxa.
The presence of a taxonomically broad group of less common, and probably non- mycorrhizal, fungi in the Ericaceae roots sampled in the current study may reflect a diverse range of ecological roles ranging from mutualistic to parasitic. In plant roots fungi not normally considered to be mycorrhizal can have positive effects and form
'mycorrhiza-like' associations. For example, Aspergillus ustus (Bain.) growing within roots of Atriplex canescens (Pursh) Nutt has been shown to enhance the overall fitness of host plants (Barrow, 2002). Similarly, infection of plant hosts by Acremonium sp., have been variously purported to increase host fitness via enhanced nutrient acquisition and other means such as resistance to pathogenic fungi (Xiao and Berch, 1999; Narisawa et al., 2000). These examples suggest that non-mycorrhizal fungi could possibly form beneficial associations with Ericaceae hosts that are also likely to be important in terms of functioning of ecosystem process such as nutrient cycling. Conversely, some of the less common fungi from these Ericaceae root associated communities may compete with ericoid mycorrhizal endophytes for nutrients sequestered in organic substrates. The extent to which this occurs in situ remains unclear, however it has been suggested that carbon supplied by the Ericaceae host would render the mycorrhizal endophytes more competitive than the relatively carbon-starved non-mycorrhizal fungi (Kerley and Read,
1995). 195
In conclusion, the data presented in this thesis demonstrated that culturing and directly extracting DNA both reveal a range of ericoid mycorrhizal and non-mycorrhizal fungi in the root systems of E. pulchella and R. lochiae, with the ericoid mycorrhizal fungi found to be taxonomically similar to those associated with Ericaceae in temperate habitats worldwide. Each approach identified several unique fungi, suggesting that a combination of culturing and culture-independent approaches may be more efficacious than one method used individually for investigating Ericaceae root associated fungal diversity.
Several of the ericoid mycorrhizal endophytes and root-associated fungi from the E. pulchella sampled were shown to produce extracellular chitinolytic activities during growth in axenic culture indicating these fungi have the potential to supplement their own as well as their host’s nitrogen supply via naturally occurring sources chitin.
DGGE, T-RFLP and cloning identified clear differences in Ericaceae root associated fungal endophyte community structure across a moorland : forest gradient at the
Abernethy forest site. DNA sequencing of selected clones revealed that typical mycorrhizal symbionts of Ericaceae along with Sebacinaceae were common in Ericaceae roots throughout the field site. Common ectomycorrhizal basidiomycetes were also present in most samples from the forest understorey but, were generally absent elsewhere.
Investigating the functional significance of Australian ericoid mycorrhizal fungi in planta as well as evaluating the nature of the associations basidiomycetes form with Ericaceae roots represent potential areas of future research that might be pursued. Such research 196 could contribute significantly to further understanding the ericoid mycorrhizal association.
197
REFERENCES
Abuarghub, S.M. and Read, DJ. (1988) The biology of mycorrhizas in the Ericaceae. XI.
The distribution of nitrogen in the soil of a typical upland Callunetum with special reference to the ‘free’ amino acids. New Phytologist 108: 425-431.
Abuzinadah, R.A. and Read, D.J. (1988) Amino acids as nitrogen sources for ectomycorrhizal fungi: utilization of individual amino acids. Transactions of the British
Mycological Society 91: 473-479.
Adams, M.A. and Attiwill, P.M. (1982) Nitrogen mineralization and nitrate reduction in forests. Soil Biology and Biochemistry 14: 197-202.
Adams, M.A. and Attiwill, P.M. (1986a) Nutrient cycling and nitrogen mineralization in eucalypt forests of south-eastern Australia. I. Nutrient cycling and nitrogen turnover.
Plant and Soil 92: 319-339.
Adams, M.A. and Attiwill, P.M. (1986b) Nutrient cycling and nitrogen mineralization in eucalypt forests of south-eastern Australia. II. Indices of nitrogen mineralization. Plant and Soil 92: 341-362.
Aerts, R. (2002) The role of various types of mycorrhizal fungi in nutrient cycling and plant competition. In: Mycorrhizal Ecology (eds M.G.A. van der Heijden and I.R.
Sanders). Springer-Verlag Berlin., Heidelberg, Germany. 198
Ahlich, K. and Sieber, T.N. (1996) The profusion of dark septate endophytic fungi in non-ectomycorrhizal fine roots of forest trees and shrubs. New Phytologist 132: 259-270.
Allen, W. K., Allaway, W. G., Cox, G. C., and Valder, P. G. (1989) Ultrastructure of mycorrhizas of Dracophyllum secundum R. Br. (Ericales: Epacridaceae). Australian
Journal of Plant Physiology 16: 147-153.
Allen, T. R., Millar, T., Berch, S. M., and Berbee, M. L. (2003) Culturing and direct
DNA extraction find different fungi from the same ericoid mycorrhizal roots. New
Phytologist 160: 255-272.
Amann, R.I. (1995) Fluorescently labelled rRNA-targeted oligonucleotide probes in the study of microbial ecology. Molecular Ecology 4: 543-554.
Anderberg, A.A. (1993) Cladistic inter-relationships and major clades of the Ericales.
Plant Systematics and Evolution 184: 207-231.
Anderberg, A.A., Rydin, C. and Kallersjo, M. (2002) Phylogenetic relationships in the order Ericales SL: Analyses of molecular data from five genes from the plastid and mitochondrial genomes. American Journal of Botany 89: 677-687.
Anderberg, A.A. and Zhang, X.P. (2002) Phylogenetic relationships of Cyrillaceae and
Clethraceae (Ericales) with special emphasis on the genus Purdiaea Planch. Organisms
Diversity and Evolution 2: 127-137. 199
Anderson, I.C., and Cairney, J.W.G. (2004) Diversity and ecology of soil fungal communities: increased understanding through the application of molecular techniques.
Environmental Microbiology 6: 769-779.
Anderson, I. C., Campbell, C. D., and Prosser, J. I. (2003) Diversity of fungi in organic soils under a moorland - Scots pine (Pinus sylvestris L.) gradient. Environmental
Microbiology 5: 1121-1132.
Anthony, J. McLean, C.B. and Lawrie, A.C. (2000) In vitro propagation of Epacris impressa (Epacridaceae) and the effects of clonal material. Australian Journal of Botany
48: 215-221.
Ashford, A.E., Allaway, W.G. and Reed, M.L. (1996) A possible role for the thick-walled epidermal cells in the mycorrhizal hair roots of Lysinema ciliatum R. Br. and other
Epacridaceae. Annals of Botany 77: 375-381.
Ashford, A.E., Ryde, S. and Barrow, K.D. (1994) Demonstration of a short chain polyphosphate in Pisolithus tinctorius and the implications for phosphorus transport. New
Phytologist 126: 239-247.
Attiwill, P.M. and Leeper, G.W. (1987) Forest soils and nutrient cycles. Melbourne
University Press. Melbourne, Australia. 200
Backa, S., Gierer J., Reitberger, T. and Nilsson, T. (1992) Hydroxyl radical activity in brown-rot fungi studied by a new chemiluminescence method. Holzforschung 46: 61-67.
Bailey, M.J., Biely, P. and Poutanen, K. (1992) Interlaboratory testing of methods for assay of xylanase activity. Journal of Biotechnology 23: 257-270.
Bajwa, R., Abuarghub, S. and Read, D.J. (1985) The biology of mycorrhiza in the
Ericaceae. X. The utilisation of proteins and the production of proteolytic enzymes by the mycorrhizal endophyte and by mycorrhizal plants. New Phytologist 101: 469-486.
Bajwa, R. and Read, D.J. (1985) The biology of mycorrhiza in the Ericaceae. IX.
Peptides as nitrogen sources for the ericoid endophyte and for mycorrhizal and non- mycorrhizal plants. New Phytologist 101: 459-467.
Bajwa, R. and Read, D.J. (1986) Utilisation of mineral and amino N sources by the ericoid mycorrhizal endophyte Hymenoscyphus ericae and by the mycorrhizal and non- mycorrhizal seedlings of Vaccinium. Transactions of the British Mycological Society 87:
269-277.
Barrow, J.R. and Osuna, P. (2002) Phosphorus solubilization and uptake by dark septate fungi in fourwing saltbush, Atriplex canescens (Pursh) Nutt. Journal of Arid
Environments 51: 449-459. 201
Baxter, J.W. and Dighton, J. (2001) Ectomycorrhizal diversity alters growth and nutrient acquisition of grey birch (Betula populifolia) seedlings in host-symbiont culture conditions. New Phytologist 152: 139-149.
Bayman, P., Lebrón, L.L., Trembalay, R.L. and Lodge, D.J. (1997) Variation in endophytic fungi from roots and leaves of Lepanthes (Orchidaceae). New Phytologist
138: 143-149.
Belesky, D.P., Devine, O.J., Pallas, J.E.J., and Stringer, W.C. (1987) Photosynthetic activity of Tall Fescue as influenced by a fungal endophyte. Photosynthetica 21: 82-87.
Bell, T.L. and Pate, J.S. (1996) Nitrogen and phosphorus nutrition in mycorrhizal
Epacridaceae of south-west Australia. Annals of Botany 77: 389-397.
Bell, T.L., Pate, J.S. and Dixon, K.W. (1994) Response of mycorrhizal seedlings of SW
Australian sandplain Epacridaceae to added nitrogen and phosphorus. Journal of
Experimental Botany 45: 779-790.
Bending, G.D. and Read, D.J. (1996a) Effects of the soluble polyphenol tannic acid on the activities of ericoid and ectomycorrhizal fungi. Soil Biology and Biochemistry 28:
1595-1602.
202
Bending, G.D. and Read, D.J. (1996b) Nitrogen mobilization from protein-polyphenol complex by ericoid and ectomycorrhizal fungi. Soil Biology and Biochemistry 28: 1603-
1612.
Bending, G.D. and Read, D.J. (1997) Lignin and soluble phenolic degradation by ectomycorrhizal and ericoid mycorrhizal fungi. Mycological Research 101: 1348-1354.
Berch, S.M., Allen, T.R. and Berbee, M.L. (2002) Molecular detection, community structure and phylogeny of ericoid mycorrhizal fungi. Plant and Soil 244: 55-66.
Bergero, R., Perotto, S., Girlanda, M., Vidano, G. and Luppi, A.M. (2000) Ericoid mycorrhizal fungi are common root associates of a Mediterranean ectomycorrhizal plant
(Quercus ilex). Molecular Ecology 9: 1639-1649.
Bermudes, D. and Benzig, D.H. (1989) Fungi in neotropical epiphyte roots. BioSystems
23: 65-73.
Bidartondo, M.I. (2005) The evolutionary ecology of myco-hetertrophy. New Phytologist
167: 335-352.
Bindslev, L., Oliver, R.P. and Johansen, B. (2002) In situ PCR for detection and identification of fungal species. Mycological Research 106: 277-279.
203
Blaiseau, P.L., Kunz, C., Grison, R., Bertheau, Y. and Brygoo, Y. (1992) Cloning and expression of a chitinase gene from the hyperparasitic fungus Aphanocladium album.
Current Genetics 21: 61-66.
Bonfante-Fasolo, P. (1980) Occurence of a basidiomycete in living cells of mycorrhizal hair roots of Calluna vulgaris. Transactions of the British Mycological Society 75: 320-
325.
Bonfante-Fasolo, P., Berta, G. and Gianinazzi-Pearson, V. (1981) Ultrastructural aspects of endomycorrhiza in the Ericaceae. II. Host-endophyte relationships in Vaccinium myrtillus. New Phytologist 89: 219-244.
Bonfante-Fasolo, P. and Gianinazzi-Pearson, V. (1979) Ultrastructural aspects of endomycorrhiza in the Ericaceae I. Naturally infected hair roots of Calluna vulgaris L.
Hull. New Phytologist 83: 739-744.
Bonfante-Fasolo, P. and Gianinazzi-Pearson, V. (1982) Ultrastructural aspects of endomycorrhiza in the Ericaceae. III. Morphology of the dissociated symbionts and modifications occuring during their reassociation in axenic culture. New Phytologist 91:
691-704.
Borneman, J., and Hartin, R. J. (2000) PCR primers that amplify fungal rRNA genes from environmental samples. Applied Environmental Microbiology 66: 4356-4360. 204
Bougoure, D.S. and Cairney, J.W.G. (2005) Assemblages of ericoid mycorrhizal and other root associated fungi from Epacris pulchella (Ericaceae) as determined by culturing and direct DNA extraction from roots. Environmental Microbiology 7: 819-827.
Bradford, M.M. (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein using the principle of protein-dye binding. Analytical Biochemistry
72: 248-254.
Bradley, R., Burt, A.J. and Read, D.J. (1981) Mycorrhizal infection and resistance to heavy metal toxicity in Calluna vulgaris. Nature 292: 335-337.
Bradley, R., Burt, A.J. and Read, D.J. (1982) The biology of mycorrhiza in the Ericaceae.
VIII. The role of mycorrhizal infection in heavy metal resistance. New Phytologist 91:
197-201.
Bremer, B., Bremer, K., Heidari, N., Erixon, P., Olmstead, R.G., Anderberg, A.A.,
Kallersjo, M. and Barkhordarian, E. (2002) Phylogenetics of asterids based on 3 coding and 3 non-coding chloroplast DNA markers and the utility of non-coding DNA at higher taxonomic levels. Molecular Phylogenetics and Evolution 24: 274-301.
Bremer, K., Chase, M.W., Stevens, P.F., Anderberg, A.A., Backland, A., Bremer, B.,
Briggs, B.G., Endress, P.K., Fay, M.F., Goldblatt, P., Gustafsson, M.H.G., Hoot, S.B., 205
Judd, W.S., Kallersjo, M., Kellogg, E.A., Kron, K.A., Les, D.H., Morton, C.M., Nickrent,
D.L., Olmstead, R.G., Price, R.A., Quinn, C.J., Rodman, J.E., Rudall, P.J., Savolainen,
V., Soltis, D.E., Soltis, P.S., Sytsma, K.J. and Thulin, M. 1998. An ordinal classification for the families of flowering plants. Annals of the Missouri Botanical Garden 85: 531-
553.
Bridge, P. and Spooner, B. (2001) Soil fungi: diversity and detection. Plant and Soil 232:
147-154.
Briggs, C.L. and Ashford, A.E. (2001) Structure and composition of the thick wall in hair root epidermal cells of Woollsia pungens. New Phytologist 149: 219-232.
Brodie, E., Edwards, S., and Clipson, N. (2003) Soil fungal community structure in a temperate upland grassland soil. FEMS Microbiology Ecology 45: 105-114.
Bruijnzeel, L.A. and Veneklasas, E.J. (1998) Climatic conditions and tropical montane forest productivity: The fog has not lifted yet. Ecology 79: 3-9
Brunner, F. and Petrini, O. (1992) Taxonomy of some Xylaria species and xylariaceous endophytes by isozyme electrophoresis. Mycological Research 96: 723-733.
Bruns, T.D. (2001) ITS Reality. Inoculum-Mycologia supplement 52: 2-3.
206
Bunn, E., Dixon, K.W. and Langley, M.A. (1989) In vitro propgation of Leucopogon obtectus Benth. (Epacridaceae). Plant Cell, Tissue and Organ Culture 19: 77-84.
Burke, R.M. and Cairney, J.W.G. (1997a) Carbohydrolase production by the ericoid mycorrhizal fungus Hymenoscyphus ericae under solid state fermentation conditions.
Mycological Research 101: 1135-1139.
Burke, R.M. and Cairney, J.W.G. (1997b) Purification and characterisation of a -1,4- endoxylanase from the ericoid mycorrhizal fungus Hymenoscyphus ericae. New
Phytologist 135: 345-352.
Burke, R.M. and Cairney, J.W.G. (1998) Carbohydrate oxidases in ericoid and ectomycorrhizal fungi: a possible source of Fenton radicals during degradation of lignocellulose. New Phytologist 139: 637-645.
Cairney, J.W.G. (2000) Evolution of mycorrhiza systems. Naturwissenschaften 87: 467-
475.
Cairney, J.W.G. and Ashford, A.E. (2002) Biology of mycorrhizal associations of epacrids (Ericaceae). New Phytologist 154: 305-326.
207
Cairney, J.W.G. and Burke, R.M. (1998) Extracellular enzyme activities of the ericoid mycorrhizal endophyte Hymenoscyphus ericae (Read) Korf and Kernan: their likely roles in decomposition of dead plant tissue in soil. Plant and Soil 205: 181-192.
Cairney, J.W.G. and Burke, R.M. (1996a) Physiological heterogeneity within fungal mycelia: an important concept for a functional understanding of the ectomycorrhizal symbiosis. New Phytologist 134: 685-695.
Cairney, J.W.G. and Burke, R.M. (1996b). Plant cell-wall degrading enzymes in ericoid and ectomycorrhizal fungi. In: Mycorrhizas in integrated systems. (eds C. Azcon-Aguilar and J.M. Barea), pp. 218-221. European Commission, Brussels, Belgium.
Cairney, J.W.G., Burke, R.M., Webster, M.A. and Steinke, E. (1996) A β-1-4- endoxylanse from the ericoid mycorrhizal fungus Hymenoscyphus ericae and putative epacrid mycorrhizal endophytes. Proceedings of the Australian Society for Biochemistry and Molecular Biology 28, POS 74.
Cairney, J.W.G. and Meharg, A.A. (2003) Ericoid mycorrhiza: a partnership that expoloits harsh edaphic conditions. European Journal of Soil Science 54: 735-740.
Cairney, J.W.G., Sawyer, N.A., Sharples, J.M. and Meharg A.A. (2000) Intraspecific variation in nitrogen source utilisation by isolates of the ericoid mycorrhizal fungus 208
Hymenoscyphus ericae (Read) Korf and Kernan. Soil Biology and Biochemistry 32:
1319-1322.
Carpita, N.C. and Gibeaut, D.M. (1993) Structural models of primary cell walls in flowering plants: consistency of molecular structure with the physical properties of the walls during growth. Plant Journal 3: 1-30.
Chambers, S.M., Liu, G. and Cairney J.W.G. (2000) ITS rDNA sequence comparison of ericoid mycorrhizal endophytes from Woollsia pungens. Mycological Research 104: 168-
174.
Chambers, S.M., Williams, P.G., Seppelt, R.D. and Cairney J.W.G. (1999) Molecular identification of a Hymenoscyphus species from rhizoids of the leafy liverwort
Cephaloziella exiliflora (Tayl.) Steph. In Australia and Antarctica. Mycological Research
103: 286-288.
Chapman, S.J., Campbell, C.D. and Puri, G. (2003) Native woodland expansion: soil chemical and microbiological indicators of change. Soil Biology and Biochemistry 35:
753-764.
Chen, A., Chambers, S.M. and Cairney J.W.G. (1999) Utilisation of organic nitrogen and phosphorus sources by mycorrhizal endophytes of Woollsia pungens (Cav.) F. Muell.
(Epacridaceae). Mycorrhiza 8: 181-187. 209
Chen, D.M., and Cairney, J.W.G. (2002) Investigation of the influence of prescribed burning on ITS profiles of ectomycorrhizal and other soil fungi at three Australian sclerophyll forest sites. Mycological Research 106: 532-540.
Christensen, M. (1989) A view of fungal ecology. Mycologia 81: 1-19.
Colpaert, J.V., Van Laere, A. and Van Assche, J.A. (1996) Carbon and nitrogen allocation in ectomycorrhizal and non-mycorrhizal Pinus sylvestris L. seedlings. Tree
Physiology 16: 787-793.
Coughlan, M.P., Tuohy, M.G., Filho, E.X.F., Puls, J., Claeyssens, M., Vranska, M. and
Hughes, M.M. (1993) Enzymological aspects of microbial hemicellulases with emphasis on fungal systems. In: Hemicellulose and hemicellulases (eds. M.P. Coughlan and G.P.
Hazlewood). Portland Press, London, UK.
Couture, M., Fortin, J.A. and Dalpé, Y. (1983) Oidiodendron griseum Robak: an endophyte of ericoid mycorrhiza in Vaccinium spp. New Phytologist 95: 375-380.
Craven, L.A. and Withers, R.M. (1996) A second species of Rhododendron from
Australia. Edinburgh Journal of Botany 53: 27-37.
210
Crayn, D.M., Kron, K.A., Gadek, P.A. and Quinn, C.J. (1998) Phylogenetics and evolution of epacrids - a molecular analysis using the plastid gene rbcl with a reappraisal of the position of Lebetanthus. Australian Journal of Botany 46: 187-200.
Crayn, D.M. and Quinn, C.J. (2000) The evolution of the atpβ-rbcL intergenic spacer in the epacrids (Ericales) and its systematic and evolutionary implications. Molecular
Phylogenetics and Evolution 16: 238-252.
Cronquist, A. (1981) An integrated system of classification of flowering plants. Columbia
University Press, New York.
Cullings, K.W. (1996) Single phylogenetic origin of ericoid mycorrhizae within the
Ericaceae. Canadian Journal of Botany 74: 1896-1909.
Dalpé, Y. (1986) Axenic synthesis of ericoid mycorrhiza in Vaccinium angustifolium Ait. by Oidiodendron species. New Phytologist 103: 391-396.
Dalpé, Y. (1989) Ericoid mycorrhizal fungi in the Myxotrichaceae and Gymnoascaceae.
New Phytologist 113: 523-527.
Dalpé, Y. (1991) Statut endomycorrhizen du genre Oidiodendron. Canadian Journal of
Botany 69: 1712-1714.
211
Dana, M.M., Limón, M.C., Mejías, R., Mach, R.L., Benítez, T., Pintor-Toro, J.A. and
Kubicek, C.P. (2001) Regulation of chitinase 33 (chit33) gene expression in Trichoderma harzianum. Current Genetics 38: 335-342.
De la Cruz, J., Rey, M., Lora, J. M., Hidalgo-Gallego, A., Domínguez, F., Pintor-Toro, J.
A., Llobell, A. and Benítez, T. (1993) Carbon source control on β-glucanases, chitobiase and chitinase from Trichoderma harzianum. Archives of Microbiology 159: 316-322.
Douglas, G.C., Heslin, M.C. and Reid, C. (1989) Isolation of Oidiodendron maius from
Rhododendron and ultrastructural characterization of synthesised mycorrhizas. Canadian
Journal of Botany 67: 2206-2212.
Duckett, J.G. and Read, D.J. (1995) Ericoid mycorrhizas and rhizoid-ascomycete associations in liverworts share the same mycobiont: isolation of the partners and resynthesis of the associations in vitro. New Phytologist 129: 439-447.
Duclos, J.L. and Fortin, J.A. (1983) Effect of glucose and active charcoal on in vitro synthesis of ericoid mycorrhiza with Vaccinium spp. New Phytologist 94: 95-102.
Duddridge, J.A. and Read, D.J. (1982) Ultrastructural analysis of the development of mycorrhizas in Rhododendron ponticum. Canadian Journal of Botany 60: 2345-2356.
212
Edel-Hermann, V., Dreumont, C., Pérez-Piqueres, A., and Steinberg, C. (2004) Terminal restriction fragment length polymorphism analysis of ribosomal RNA genes to assess changes in fungal community structure in soils. FEMS Microbioogyl Ecology 47: 397-
404.
Egert, M. and Friedrich, M.W. (2003) Formation of pseudo-terminal restriction fragments, a PCR related bias affecting terminal restriction fragment length polymorphism analysis of microbial community structure. Applied Environmental
Microbiology 69: 2555-2562.
Egger, K.N. and Sigler, L. (1993) Relatedness of the ericoid endophytes Scytalidium vaccinii and Hymenoscyphus ericae inferred from analysis of ribosomal DNA. Mycologia
85: 219-230.
El-Katatny, M.H., Somitsch, W., Robra, K., El-Katatny, M.S. and Gübitz, G.M. (2000)
Production of chitinase and β-1,3-glucanase by Trichoderma harzianum for control of the phytopathogenic fungus Sclerotium rolfsii. Food Technology and Biotechnology 38: 173-
180.
van Elsas, J.D., Duarte, G. F., Keijzer-Wolters, A., and Smit, E. (2000) Analysis of the dynamics of fungal communities in soil via fungal-specific PCR of soil DNA followed by denaturing gradient gel electrophoresis. Journal of Microbiological Methods 43: 133-
151. 213
Englander, L. and Hull, R.J. (1980) Reciprocal transfer of nutrients between ericaceous plants and a Clavaria sp. New Phytologist 84: 661-667.
Erskine, P.D., Stewart, G.R., Schmidt, S., Turnbull, M.H., Unkovich, M. and Pate, J.S.
(1996) Water availability - a physiological constraint on nitrate utilization in plants of
Australian semi-arid mulga woodlands. Plant, Cell and Environment 19: 1149-1159.
Felske, A., Engelen, B., Nubel, U. and Backhaus, H. (1996) Direct ribosomal isolation from soil to extract bacterial rRNA for community analysis. Applied and Environmental
Microbiology 62: 4162-4167.
Finlay, R.D., Frostegård, Å. and Sonnerfeldt, A.M. (1992). Utilisation of organic and inorganic nitrogen sources by ectomycorrhizal fungi in pure culture and in symbiosis with Pinus contorta Dougal. Ex Loud. New Phytologist 120: 105-115.
Gardes, M. and Bruns, T.D. (1993) ITS primers with enhanced specificity for basidiomycetes: application to the identification of mycorrhiza and rusts. Molecular
Ecology 2: 113-118.
Gehring, C.A., Thiemer, T.C., Whitham, T.G., and Keim, P. (1998) Ectomycorrhizal fungal community structure of pinyon pine growing in two environmental extremes.
Ecology 79: 1562-1572.
214
Gianinazzi-Pearson, V., Bonfante-Fasolo, P. and Dexheimer, J. (1986) Ultrastructural studies of surface interactions during adhesion and infection by ericoid endomycorrhizal fungi. In: Recognition in Microbe-Plant Symbiotic and Pathogenic Interactions. (ed B.
Lugtenberg), 4: 273-282. Springer-Verlag., Berlin, Germany.
Gold, M.H., Wariishi, H. and Valli, K. (1989) Extracellular peroxidases involved in lignin degradation by the white rot basidiomycete Phanerochaete chrysosporium.
American Chemistry Society Symposium Serials 389: 127-140.
Goldstein, J.L. and Swain, T. (1965) The inhibition of enzymes by tannins.
Phytochemisty 4: 185-192.
Gooday, G.W. (1990) The ecology of chitin degradation. In Advances in Microbial
Ecology (ed. K.C. Marshall), 11: 387-430. Plenum Press: New York.
Gooday, G.W. (1993) Physiology and microbial degradation of chitin and chitosan. In:
Biochemistry of Microbial Degradation (ed. C. Ratledge), pp.279-312. Klewer,
Dordrecht.
Grunig, C.R., Sieber, T.N., Rogers, S.O. and Holdenrieder, O. (2002) Spatial distribution of dark septate endophytes in a confined forest plot. Mycological Research 106: 832-840.
215
Hambleton, S. and Currah, R.S. (1997) Fungal endophytes from the roots of alpine and boreal Ericaceae. Canadian Journal of Botany 75: 1570-1581.
Hambleton, S., Currah, R.S. and Egger, K.N. (1998a) Phylogenetic relationships of ascomycetous root endophytes of the Ericaceae inferred from 18S rDNA sequence analysis. In: Abstracts of the Second International Conference on Mycorrhiza (eds U.
Ahonen-Jonnarth, E. Dannell, P. Fransson, P. Kåren, B. Lindahl, I. Rangel and R.
Finlay), p. 78. Swedish University of Agricultural Sciences, Uppsala, Sweden.
Hambleton, S., Egger, K.N. and Currah, R.S. (1998b) The genus Oidiodendron - species delimitation and phylogenetic relationships based on nuclear ribosomal DNA analysis.
Mycologia 90: 854-868.
Hambleton, S. and Sigler, L. (2005) Meliniomyces, a new anamorph genus for root- associated fungi with phylogenetic affinities to Rhizoscyphus ericae (= Hymenoscyphus ericae), Leotiomycetes. Studies in Mycology 53: 1-27.
Hamilton, L.S., Juvik, J.O. and Scatena, F.N. (1995) The Puerto Rico tropical cloud forest symposium: Introduction. In: Tropical Montane Cloud Forests (eds. L.S.
Hamilton, J.O. Juvik and F.N. Scatena) pp.1-24. Springer-Verlag, New York.
Harley, J.L. (1969) The Biology of Mycorrhiza. 2nd edn. Leonard Hill, London.
216
Harley, J.L. and Smith, S.E. (1983) Mycorrhizal Symbiosis. Academic Press, London,
U.K.
Haselwandter, K., Bobleter, O. and Read D.J. (1990) Degradation of 14C-labelled lignin and dehydropolymer of coniferyl alcohol by ericoid and ectomycorrhizal fungi.
Microbiology 153: 352-354.
Hatakka, A. (1994) Lignin-modifying enzymes from selected white-rot fungi: production and role in lignin degradation. FEMS Microbiology Review 13: 125-135.
Haug, I., Lempe, J., Homeier, J., Weis, M., Setaro, S., Oberwinkler, F. and Kottke, I.
(2004) Graffenrieda emarginata (Melastomataceae) forms mycorrhizas with
Glomeromycota and with a member of the Hymenoscyphus ericae aggregate in the organic soil of a neotropical mountain rainforest. Canadian Journal of Botany 82: 340-
356.
Heuer, H. and Smalla, K. (1997) Application of denaturing gradient gel electrophoresis
(DGGE) and temperature gradient gel electrophoresis (TGGE) for studying soil microbial communities. In: Modern Soil Microbiology (eds. J.D. van Elsas, J.T. Trevors and
E.M.H. Wellington), pp 353-373. Marcel Dekker, New York, USA.
Hodge, A., Alexander, I.A. and Gooday, G.W. (1995) Chitinolytic enzymes of pathogenic and ectomycorrhizal fungi. Mycological Research 99: 935-941. 217
Hooper, D.U. and Vitousek, P.M. (1998) Effects of plant composition and diversity on nutrient cycling. Ecological Monographs 68: 121-149.
Horton, T.R. and Bruns, T.D. (2001) The molecular revolution in ectomycorrhizal ecology: peeking into the black-box. Molecular Ecology 10: 1855-1871.
Huhta, V. (2002) Soil macroarthropod communities in planted birch stands in comparison with natural forests in central Finland. Applied Soil Ecology 20: 199-209.
Hunt, J., Boddy, L., Randerson, P. F., and Rogers, H. J. (2004) An evaluation of 18S rDNA approaches for the study of fungal diversity in grassland soils. Microbial Ecology
47: 385-395.
Hutton, B.J., Dixon, K.W. and Sivasithamparam, K. (1994) Ericoid endophytes of
Western Australian heaths (Epacridaceae). New Phytologist 127: 557-566.
Jalal, M.A.F. and Read, D.J. (1983a) The organic acid composition of Calluna heathland with special reference to phyto- and fungi- toxicity. I. Isolation and identification of organic acids. Plant and Soil 70: 257-272.
Jalal, M.A.F. and Read, D.J. (1983b) The organic acid composition of Calluna heathland with special reference to phyto- and fungi- toxicity. II. Monthly quantitative 218 determination of the organic acid content of Calluna and spruce dominated soils. Plant and Soil 70: 273-286.
Johnson, D., Vandenkoornhuyse, P.J., Leake, J.R., Gilbert, L., Booth, R.E., Grime, J.P.,
Young, J.P.W. and Read, D.J. (2003) Plant communities affect arbuscular mycorrhizal fungal diversity and community composition in grassland microcosms. New Phytologist
161: 503-515.
Jones, M.D. and Smith, S.E. (2004) Exploring functional definitions of mycorrhizas: Are mycorrhizas always mutualisms? Canadian Journal of Botany 82: 1089-1109.
Jonsson, L.M., Nilsson, M.C., Wardle, D.A. and Zackrisson, O. (2001) Context dependent effects of ectomycorrhizal species richness on tree seedling productivity.
Oikos 93: 353-364.
Judd, W.S. and Kron, K.A. (1993) Circumscription of Ericaceae (Ericales) as determined by preliminary cladistic analyses based on morphological, anatomical, and embryological features. Brittonia 45: 99-114.
Jumpponen, A. (1999) Spatial distribution of discrete RAPD phenotypes of a root endophytic fungus, Phialocephala fortinii, at a primary succesional site on a glacier forefront. New Phytologist 141: 333-344.
219
Jumpponen, A. (2001) Dark septate endophytes - are they mycorrhizal? Mycorrhiza 11:
207-211.
Jumpponen, A. (2003) Soil fungal community assembly in a primary successional glacier forefront ecosystem as inferred from rDNA sequence analysis. New Phytologist 158: 569-
578.
Jumpponen, A., Mattson, K.G. and Trappe, J.M. (1998) Mycorrhizal functioning of
Phialocephala fortinii with Pinus contorta on glacier forefront soil - interactions with soil nitrogen and organic matter. Mycorrhiza 7: 261-265.
Jumpponen, A. and Trappe, J.M. (1998a) Dark septate endophytes - a review of facultative biotrophic root-colonizing fungi. New Phytologist 140: 295-310.
Jumpponen, A. and Trappe, J.M. (1998b) Performance of Pinus contorta inoculated with two strains of root endophytic fungus, Phialocephala fortinii: effects of synthesis system and glucose concentration. Canadian Journal of Botany 76: 1205-1213.
Kemp, E., Adam, P. and Ashford, A.E. (2003) Seasonal changes in hair roots and mycorrhizal colonization in Woollsia pungens (Cav.) F. Muell. (Epacridaceae). Plant and
Soil 250: 241-248.
220
Kerley, S.J. and Read, D.J. (1995) The biology of mycorrhiza in the Ericaceae. XVIII.
Chitin degradation by Hymenoscyphus ericae and transfer of chitin-nitrogen to the host plant. New Phytologist 131: 369-375.
Kerley, S.J. and Read, D.J. (1997) The biology of mycorrhiza in the Ericaceae. XIX.
Fungal mycelium as a nitrogen source for the ericoid mycorrhizal fungus Hymenoscyphus ericae and its host plants. New Phytologist 136: 691-701.
Kerley, S.J. and Read, D.J. (1998) The biology of mycorrhiza in the Ericaceae. XX. Plant and mycorrhizal necromass as nitrogenous substrates for the ericoid mycorrhizal fungus
Hymenscyphus ericae and its host. New Phytologist 139: 353-360.
Kiernan, J.M., Hendrix, J.W., and Maronek, D.M. (1983) Fertilizer-induced pathogenicity of mycorrhizal fungi to sweetgum seedlings. Soil Biology and Biochemistry
15: 257-262.
King, S.A. and Buckney, R.T. (2002) Invasion of exotic plants in nutrient-enriched urban bushland. Austral Ecology 27: 573-583.
Klamer, M., Roberts, M. S., Levine, L. H., Drake, B. G., and Garland, J. L. (2002)
Influence of elevated CO2 on the fungal community in a coastal scrub oak forest soil investigated with terminal restriction fragment length polymorphism analysis. Applied
Environmental Microbiology 68: 4370-4376. 221
Klironomos, J.N., McCune, J., Hart, M. and Neville, J. (2000) The influence of arbuscular mycorrhizae on the relationship between plant diversity and productivity.
Ecology Letters 3: 137-141.
Kolbe, J. and Kubicek, C.P. (1990) Quantification and identification of the main components of the Trichoderma cellulase complex with monoclonal antibodies using an enzyme-linked immunosorbent assay (ELISA). Applied Microbiology and Biotechnology
34: 26-30.
Kottke, I., Beck, A., Oberwinkler, F., Homeier, J. and Neill, D. (2004) Mycorrhizal state of trees in neotropical cloud forests. Journal of Tropical Ecology 20: 1-6.
Kowalchuk, G.A., Gerards, S. and Woldendorp, J.W. (1997) Detection and characterization of fungal infections of Ammophila arenaria (marram grass) roots by denaturing gradient gel electrophoresis of specifically amplified 18S rDNA. Applied and
Environmental Microbiology 63: 3858-3865.
Kron, K.A. (1996) Phylogenetic relationships of Empetraceae, Epacridaceae, Ericaceae,
Monotropaceae, and Pyrolaceae - evidence from nuclear ribosomal 18s sequence data.
Annals of Botany 77: 293-303.
222
Kron, K.A. and Chase, M.W. (1993) Systematics of the Ericaceae, Empetraceae,
Epacridaceae and related taxa based upon rbcl sequence data. Annals of the Missouri
Botanical Garden 80: 735-741.
Kron, K.A., Fuller, R., Crayn, D.M., Gadek, P.A. and Quinn, C.J. (1996) Molecular systematics of the vacciniods and epacrids using matK sequence data. American Journal of Botany 83 (Supplement): 170.
Kron, K.A., Fuller, R., Crayn, D.M., Gadek, P.A. and Quinn, C.J. (1999) Phylogenetic relationships of epacrids and vaccinioids (Ericaceae s. l.) based on matK sequence data.
Plant Systematics and Evolution 218: 55-65.
Kron, K.A., Powell, E.A. and Luteyn, J.L. (2002) Phylogenetic relationships within the blueberry tribe (Vaccinieae, Ericaceae) based on sequence data from matK and nuclear ribosomal ITS regions, with comments on the placement of Satyria. American Journal of
Botany 89: 327-336.
Kubicek, C. P., Mach, R. L., Peterbauer, C. K. and Lorito, M. (2001). Trichoderma: from genes to biocontrol. Journal of Plant Pathology 83 (Special issue): 11-23.
Kuhad, R.C., Singh, A. and Eriksson, K-El. (1997) Microorganisms and enzymes involved in the degradation of plant fibre cell walls. In: Biotechnology in the pulp and paper industry (ed. K-El. Eriksson), pp 45-125. Springer-Verlag, Berlin, Germany. 223
Labaune, C. and Magnin, F.D. (2002) Pastoral management vs. land abandonment in
Mediterranean uplands: impact on land snail communities. Global Ecology and
Biogeography 11: 237-245.
Lacourt, I., Girlanda, M., Perotto, S., Del Pero, M., Zuccon, D. and Luppi, A.M. (2001)
Nuclear ribosomal sequence analysis of Oidiodendron: towards a redefinition of ecologically relevant species. New Phytologist 149: 565-576.
Lawrie, A.C., McDonald, N. and McLean, C.B. (2001) Fungi isolated from ericoid mycorrhizal roots of Australian family Epacridaceae improve rooting of shoot cuttings.
In: Abstracts from the 3rd International Conference on Mycorrhizas, P140. The
University of Adelaide.
Leake, J.R. (2001) Is diversity of ectomycorrhizal fungi important for ecosystem function? New Phytologist 152: 1-8.
Leake, J.R. and Miles, W. (1996) Phosphodiesters as mycorrhizal P sources. I.
Phosphodiesterase production and the utilization of DNA as a phosphorus source by the ericoid mycorrhizal fungus Hymenoscyphus ericae. New Phytologist 132: 435-443.
Leake, J.R. and Read, D.J. (1989a) The biology of mycorrhiza in the Ericaceae. XIII.
Some characteristics of the extracellular proteinase activity of the ericoid endophyte
Hymenoscyphus ericae. New Phytologist 112: 69-72. 224
Leake, J.R. and Read, D.J. (1989b) The effects of phenolic compounds on nitrogen metabolism by ericoid mycorrhizal fungi. Agricultural Ecosystems and Enivonment. 29:
225-236.
Leake, J.R. and Read, D.J. (1990a) Proteinase activity in mycorrhizal fungi. I. The effect of extracellular pH on the production and activity of proteinase by ericoid endophytes from soils of contrasted pH. New Phytologist 115: 243-250.
Leake, J.R. and Read, D.J. (1990b) Proteinase activity in mycorrhizal fungi. II. The effects of mineral and organic nitrogen sources on induction of extracellular proteinase in
Hymenoscyphus ericae (Read) Korf and Kernan. New Phytologist 116: 123-128.
Leake, J.R. and Read, D.J. (1990c) Chitin as a nitrogen source for mycorrhizal fungi.
Mycological Research 94: 993-995.
Leake, J.R. and Read, D.J. (1991) Experiments with mycorrhiza. Methods in
Microbiology 23: 435-459.
Leake, J.R. and Read. D.J. (1997) Mycorrhizal fungi in terrestrial habitats. In: The
Mycota IV Environmental and Microbial Relationships (eds. D. Wicklow and B.
Söderström) pp 281-301. Springer –Verlag, Berlin, Germany.
225
Lee, K.E. and Correll, R.L. (1978) Litter fall and its relationship to nutrient cycling in a
South Australian dry sclerophyll forest. Australian Journal of Ecology 3: 243-252.
Lesica, P. and Antibus, R.K. (1990) The occurrence of mycorrhizae in vascular epiphytes of two Costa Rican rainforests. Biotropica 22: 250-258.
Lindahl, B.D., Taylor, A.F.S., and Finlay, R.D. (2002) Defining nutritional constraints on carbon cycling in boreal forests – towards a less “phytocentric” perspective. Plant and
Soil 242: 123-135.
Lindahl, B.D. and Taylor, A.F.S. (2004) Occurance of N-acetylhexosaminidase-encoding genes in ectomycorrhizal basidiomycetes. New Phytologist 164: 193-199.
Liu, G. (1998) Molecular diversity of mycorrhizal endophytes of Woollsia pungens.
(MSc Thesis) University of Western Sydney, Nepean. Sydney, Australia.
Liu, G., Chambers, S.M, Cairney, J.W.G. (1998) Molecular diversity of ericoid mycorrhizal endophytes isolated from Woollsia pungens. New Phytologist 140: 145-153.
LoBuglio, K.F., Berbee, M.L. and Taylor, J.W. (1999) Phylogenetic Origins of the
Asexual Mycorrhizal Symbiont Cenococcum geophilum Fr. and Other Mycorrhizal Fungi among the Ascomycetes. Molecular Phylogenetics and Evolution 6: 287-294.
226
Lowell, J.L., and Klein, D.A. (2001) Comparative single-stranded conformation polymorphism (SSCP) and microscopy-based analysis of nitrogen cultivation interactive effects on the fungal community of a semiarid steppe soil. FEMS Microbiology Ecology
36: 85-92.
Luteyn, J.L (2002) Diversity, adaptation and endemism in Neotropical Ericaceae:
Biogeographical patterns in the Vaccinieae. The Botanical Review 68: 55-87
Mach, R.L., Peterbauer, C.K., Payer, K., Jaksits, S., Woo, S.L., Zeilinger, S., Kullnig,
C.M., Lorito, M. and Kubicek, C.P. (1999) Expression of two major chitinase genes of
Trichoderma atroviride (T. harzianum P1) triggered by different regulatory signals.
Applied and Environmental Microbiology 65: 1858-1863.
Martino, E., Turnau, K., Girlanda, M., Bonfante, P. and Perotto, S. (2000) Ericoid mycorrhizal fungi from heavy metal polluted soils: their identification and growth in the presence of zinc ions. Mycological Research 104: 338-344.
Marx, D.H. and Bryan, W.C. (1975) Growth and ectomycorrhizal development of loblolly pine seedlings in fumigated soil infested with the fungal symbiont Pisolithus tinctorius. Forest Science 21: 245-254.
McCreath, K.J. and Gooday, G.W. (1992) A rapid and sensitive microassay for determination of chitinolytic activity. Journal of microbiological methods 14: 229-237. 227
McCune, B. and Mefford, M.J. (1999) Multivariate analysis of ecological data, Version
4.25, MjM Software, Gleneden Beach, Oregon, USA.
McJannet, D. and Reddell, P. (2002) Where earth meets sky: Cloud forests of the wet tropics. CSIRO, Canberra.
McLean, C.B. and Lawrie, A.C. (1996) Patterns of root colonization in epacridaceous plants collected from different sites. Annals of Botany 77: 405-411.
McLean, C.B., Lawrie, A.C. and Blaze, K.L. (1994) The effect of soil microflora on the survival of cuttings of Epacris impressa. Plant and Soil 166: 295-297.
McLean, C.B., Anthony, J., Collins, R.A., Steinke, E. and Lawrie, A.C. (1998) First synthesis of ericoid mycorrhizas in the Epacridaceae under axenic conditions. New
Phytologist 139: 589-593.
McLean, C.B., Cunnington, J.H. and Lawrie, A.C. (1999) Molecular diversity within and between ericoid endophytes from the Ericaceae and Epacridaceae. New Phytologist 144:
351-358.
Meharg, A.A. and Cairney, J.W.G. (2000) Co-evolution of mycorrhizal symbionts and their hosts to metal contaminated environments. Advances in Ecological Research 30: 69-
112. 228
Midgley, D.J. (2003) Diversity and distribution of mycorrhizal and root associated fungal endophytes of Leucopogon parviflorus and Woollsia pungens. (PhD Thesis) University of
Western Sydney. Sydney, Australia.
Midgley, D.J., Chambers, S.M. and Cairney, J.W.G. (2002) Spatial distribution of fungal endophyte genotypes in a Woollsia pungens (Ericaceae) root system. Australian Journal of Botany 50: 559-565.
Midgley, D.J., Chambers, S.M. and Cairney, J.W.G. (2004a) Inorganic and organic substrates as sources of nitrogen and phosphorus for multiple genotypes of two ericoid mycorrhizal fungal taxa from Woollsia pungens Cav. (Muell.) and Leucopogon parviflorus (Andr.) Lindl. (Ericaceae). Australian Journal of Botany 52: 63-71.
Midgley, D.J., Chambers, S.M. and Cairney, J.W.G. (2004b) Utilisation of carbon substrates by multiple genotypes of ericoid mycorrhizal fungal endophytes from eastern
Australian Ericaceae. Mycorrhiza 14: 245-251.
Midgley, D.M., Chambers, S.M., and Cairney, J.W.G. (2004c) Distribution of ericoid mycorrhizal endophytes and root-associated fungi in neighbouring Ericaceae plants in the field. Plant and Soil 259: 137-151.
229
Mitchell, D.T. and Read, D.J. (1981) Utilisation of inorganic and organic phosphates by the mycorrhizal endophytes of Vaccinium macrocarpon and Rhododendron ponticum.
Transactions of the British Mycological Society 76: 255-260.
Mitchell, D.T., Sweeney, M., and Kennedy, A. (1992) Chitin degradation by
Hymenoscyphus ericae and the influence of H. ericae on the growth of ectomycorrhizal fungi. In: Mycorrhizas in Ecosystems (eds. D.J. Read, D.H. Lewis, A.H. Fitter and I.J.
Alexander), pp 246-251. CAB International: Oxon, U.K.
Moeseneder, M.M., Arrieta, J.M., Muyzer, G., Winter, C. and Herndl, G.J. (1999)
Optimization of terminal restriction fragment length polymorphism analysis for complex marine bacterioplankton communities and comparison with denaturing gradient gel electrphoresis. Applied and Environmental Microbiology 65: 3518-3525.
Monreal, M., Berch, S.M. and Berbee, M. (1999) Molecular diversity of ericoid mycorrhizal fungi. Canadian Journal of Botany 77: 1580-1794.
Murray, A.E., Hollibaugh, J.T. and Orrego, C. (1996) Phylogenetic compositions of bacterioplankton from two California estuaries compared by denaturing gradient electrophoresis of 16S rDNA fragments. Applied and Environmental Microbiology 62:
2676-2680.
230
Muyzer, G., DeWaal, E.C. and Uitterlinden, G. (1993) Profiling of complex microbial populations by denaturing gradient gel eletrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Applied and Environmental
Microbiology 59: 695-700.
Muyzer, G. and Smalla, K. (1998) Application of denaturing gradient gel electrophoresis
(DGGE) and temperature gradient gel electrophoresis (TGGE) in microbial ecology.
Antonie van Leeuwenhoek 73: 127-141.
Myers, M.D. and Leake, J.R. (1996) Phosphodiesters as mycorrhizal P sources. II.
Ericoid mycorrhiza and the utilisation of nuclei as a phosphorus and nitrogen source by
Vaccinium macrocarpon. New Phytologist 132: 445-452.
Naeem, S., Hakansson, K., Lawton, J.H., Crawley, M.J. and Thompson, L.J. (1996)
Biodiversity and plant productivity in a model assemblage of plant species. Oikos 76:
259-264.
Narisawa, K., Tokumasu, S. and Hashiba, T. (1998) suppression of clubroot formation in
Chinese cabbage by the root endophytic fungus, Heteroconium chaetospira. Plant
Pathology 47: 206-210.
231
Narisawa, K., Oski, K.T., and Hashiba, T. (2000) Suppression of clubroot and
Verticillium yellows in Chinese cabbage in the field by the root endophytic fungus,
Heterconium chaetospira. Plant Pathology 49: 141-146.
Narisawa, K., Kawamata, H., Currah, R.S. and Hashiba, T. (2002) Suppression of
Verticillium wilt in eggplant by some fungal root endophytes. European Journal of Plant
Pathology 108: 103-109.
Natarajan, K., Mohan, V. and Ingleby, K. (1992) Correlation between basidiomata production and ectomycorrhizal formation in Pinus patula plantations. Soil Biology and
Biochemistry 24: 279-280.
O'Conner, P.J., Smith, S.E., and Smith, F.A. (2002) Arbuscular mycorrhizas influence plant diversity and community structure in a semiarid herbland. New Phytologist 154:
209-218.
Palmer, J. and Ashford, A.E. (2004) Woollsia pungens – Germination and seedling growth. Australian Plants 22: 243-252.
Pate, J.S., Stewart, G.R. and Unkovich, M. (1993) 15N natural abundance of plant and soil components of a Banksia woodland ecosystem in relation to nitrate utilisation, life form, mycorrhizal status and N2-fixing abilities of component species. Plant, Cell and
Environment. 16: 365-373. 232
Patil, R.S., Ghormade, V. and Deshpande, M.V. (2000) Chitinolytic enzymes: an expoloration. Enzyme and Microbial Technology 26: 473-483.
Pearson, V., and Read, D. J. (1973) The biology of mycorrhizas in the Ericaceae. I. The isolation of the endophyte and synthesis of mycorrhizas in aseptic cultures. New
Phytologist 72: 371-379.
Pearson, V. and Read, D.J. (1975) The physiology of the mycorrhizal endophyte of
Calluna vulgaris. Transactions of the British Mycological Society 64: 1-7.
Pearson, W.R., and Lipman, D.J. (1988) Improved tool for biological sequence analysis.
Proceedings of the National Academy of Sciences USA 85: 2444-2448.
Perotto, R., Bettini, V. and Bonfante, P. (1993) Evidence of two polygalaturonases produced by a mycorrhizal ericoid fungus during its saprotrophic growth. FEMS
Microbiology Letters 114: 85-92.
Perotto, S., Actis-Perino, E., Perugini, J. and Bonfante, P. (1996) Molecular diversity of fungi from ericoid mycorrhizal roots. Molecular Ecology 5: 123-131.
Perotto, S. and Bonfante, P. (1998) Genetic and Functional Diversity of Ericoid
Mycorrhizal Fungi. Symbiosis 25: 19-27.
233
Perotto, S., Coisson, J.D., Perugini, I., Cometti, V. and Bonfante, P. (1997) Production of pectin-degrading ezymes by ericoid mycorrhizal fungi. New Phytologist 135: 151-162.
Perotto, S., Girlanda, M. and Martino, E. (2002) Ericoid mycorrhizal fungi: some new perspectives on old acquaintances. Plant and Soil 244: 41-53.
Perotto, S., Perotto, R., Faccio, A., Schubert, A., Varma, A. and Bonfante, P. (1995)
Ericoid mycorrhizal fungi: cellular and molecular bases of their interactions with the host plant. Canadian Journal of Botany 73: S557-S568.
Peterson, T.A., Mueller, W.C. and Englander, L. (1980) Anatomy and ultrastructure of a
Rhododendron root-fungus association. Canadian Journal of Botany 58: 2421-2433.
Piercey, M.M., Thormann, M.N. and Currah, R.S. (2002) Saprobic characteristics of three fungal taxa from ericalean roots and their association with the roots of Rhododendron groenlandicum and Picea mariana in culture. Mycorrhiza 12: 175-180.
Powell, J.M., Crayn, D.M., Gadek, P.A., Quinn, C.J. and Morrison, D.A. (1996) A re- assessment of relationships within Epacridaceae. Annals of Botany 77: 305-315.
Prosser, J.I. (2002) Molecular and functional diversity in soil micro-organisms. Plant and
Soil 244: 9-17.
234
Quigley, P.E., Snell, F.J., Cunningham, P.J., and Frost, W. (1990). The effects of endophyte infected ryegrass on the establishment persistence and production of mixed pastures in Australia. In Proceedings of the International Symposium on Acremonium-
Grass Interactions., pp. 22. (Abstr.), Baton Rouge, La. USA.
Rains, K.C., Nadkarni, N.M. and Bledsoe, C.S. (2003) Epiphytic and terrestrial mycorrhizas in a lower montane Costa Rican cloud forest. Mycorrhiza 13: 257-264.
Ranjard, L., Poly, F., and Nazaret, S. (2000) Monitoring complex bacterial communities using culture-independent molecular techniques: application to soil environment.
Research in Microbiology 151: 167-177.
Read, D.J. (1974) Pezizella ericae sp. nov., the perfect state of a typical mycorrhizal endophyte of the Ericaceae. Transactions of the British Mycological Society 63: 381-419.
Read, D.J. (1983) The biology of mycorrhiza in the Ericales. Canadian Journal of Botany
61: 985-1004.
Read, D.J. (1991) Mycorrhizas in ecosystems. Experientia 46: 376-390.
Read, D.J. (1990) Mycorrhizas in ecosystems - Nature's response to the "law of the minimum" In: Frontiers in Mycology. Honorary and General Lectures from the Fourth 235
International Mycological Congress. (eds. D.L. Hawkesworth), pp 101-129. CABI
Publishing. Wallingford, UK.
Read, D.J. (1992). The mycorrhizal mycellium. In: Mycorrhizal Functioning (ed M.F.
Allen), pp 102-133. Chapman and Hall, London, UK.
Read, D.J. (1996) The structure and function of the ericoid mycorrhizal root. Annals of
Botany 77: 365-374.
Read, D.J., Leake, J.R. and Langdale, A.R. (1989) The nitrogen nutrition of mycorrhizal fungi and their host plants. In: Nitrogen, Phosphorus and Sulphur Utilisation by Fungi
(eds. L. Boddy, R. Marchant and D.J. Read), pp 181-204. Cambridge University Press,
Cambridge, U.K.
Read, D.J., Duckett, J.G., Francis, R., Ligrone, R. and Russell, A. (2000) Symbiotic fungal associations in 'lower' land plants. Philosophical Transactions of the Royal
Society, London 355: 815-831.
Read, D.J. and Perez-Moreno, J. (2003) Mycorrhizas and nutrient cycling in ecosystems - a journey towards relevance? New Phytologist 157: 475-492.
Read, D.J. and Stribley, D.P. (1973) Effect of mycorrhizal infection on nitrogen and phosphorus nutrition of ericaceous plants. Nature 244: 81. 236
Reed, M.L. (1987) Ericoid mycorrhiza of Epacridaceae in Australia. In: Mycorrhizae in the Next Decade. (eds. D.M. Sylvia, L.L. Hung and J.H. Graham), p. 335. Institute of
Food and Agricultural Sciences. University of Florida., Gainsville FL. USA.
Reed, M.L. (1989) Ericoid Mycorrhizas of Styphelieace: Intensity of Infection and
Nutrition of the Symbionts. Australian Journal of Plant Physiology 16: 155-160.
Reed, M.L. (1996) Diversity of mycorrhizal fungi in the roots of epacrids. In:
Gondwanan Heritage. Past, Present and Future of the Western Australian Biota. (eds
S.D. Hopper, J.A. Chappill, M.S. Harvey and A.S. George), pp 308-313. Surrey, Beatty and Sons, Sydney, Australia.
Rice, P. (1996) Program Manual for the EGCG Package. The Sanger Centre, Cambridge.
UK.
Rice, A.V. and Currah, R.S. (2001) Physiological and morphological variation in
Oidiodendron maius. Mycotaxon 79: 383-396.
Rice, A.V. and Currah, R.S. (2002) New perspectives on the niche and holomorph of the myxotrichoid hyphomycete, Oidiodendron maius. Mycological Research 106:1463-1467.
237
Rosling, A., Landeweert, R., Lindahl, B.D., Larson, K.H., Kuyper, T.W., Taylor, A.F.S., and Finlay, R.D. (2003) Vertical distribution of ectomycorrhizal fungal taxa in a podsol soil profile. New Phytologist 159: 775-783.
Rovira, P. and Vallejo, V.R. (2002) Labile and recalcitrant pools of carbon and nitrogen in organic matter decomposing at different depths in soil: an acid hydrolysis approach.
Geoderma 107: 109-141.
Saikkonen, K., Faeth, S.H., Helander, M. and Sullivan, T.J. (1998) Fungal Endophytes: A continuum of interactions with host plants. Annual Review of Ecology and Systematics
29: 319-343.
Schindler, T.M. (1998) The new view of the primary cell wall. Zeitschrift fur
Pflanzenernahrung und Bodenkunde 161: 499-508.
Schmidt, S. and Stewart, G.R. (1997) Waterlogging and fire impacts on nitrogen availability and utilisation in a subtropical wet heathland (wallum). Plant, Cell and
Environment 20: 1231-1241.
Schmidt, S. and Stewart, G.R. (1999) Glycine metabolism by plant roots and its occurrence in Australian plant communities. Australian Journal of Plant Physiology 26:
253-264.
238
Schulz, B. and Boyle, C. (2005) The endophytic continuum. Mycological Research 109:
661-686.
Selosse, M-A., Weiß, M., Jany, J-L. and Tillier, A. (2002) Communities and populations of sebacinoid basidiomycetes associated with the achlorophyllous orchid Neottia nidus- avis (L.) L.C.M. Rich. and neighboring tree ectomycorrhiza. Molecular Ecology 11:
1831-1844.
Seviour, R.J., Willing, R.R. and Chilvers, G.A. (1973) Basidiocarps assocated with ericoid mycorrhizas. New Phytologist 72: 381-385
Sharples, J.M., Cairney, J.W.G. (1997) Organic nitrogen utilisation by an unidentified mycobiont isolated from mycorrhizas of Pisonia grandis. Mychological Research 101:
315-318.
Sharples, J.M., Meharg, A.A., Chambers, S.M. and Cairney, J.W.G. (2000a) Evolution -
Symbiotic solution to arsenic contamination. Nature 404: 951-952
Sharples, J.M., Chambers, S.M., Meharg, A.A. and Cairney, J.W.G. (2000b) Genetic diversity of root-associated fungal endophytes from Calluna vulgaris at contrasting field sites. New Phytologist 148: 153-162.
239
Sharples, J.M., Meharg, A.A., Chambers, S.M. and Cairney, J.W.G. (2001) Arsenate resistance in the ericoid mycorrhizal fungus Hymenoscyphus ericae. New Phytologist
151: 265-270.
Simard, S.W., Perry, D.A., Jones, M.D., Myrold, D.D., Dural, D.M. and Molina, R.
(1997) Net transfer of carbon between ectomycorrhizal tree species in the field. Nature
388: 579-582.
StLeger, R.J., Cooper, R.M. and Charnley, A.K. (1986) Cuticle-degrading enzymes of entomopathogenic fungi:regulation and production of chitinolytic enzymes. Journal of
General Microbiology 132: 1509-1517.
Sleumer, H. (1960) Florae Malesianae precursors XXIII. The genus Rhododendron in
Malaysia. Reinwardtia 5: 45-231.
Smith, F.A. and Smith, S.E. (1996) Mutualism and Parasitism: Diversity in Function and
Structure in the "Arbuscular" (VA) Mycorrhizal Symbiosis. Advances in Botanical
Research. 22: 1-43.
Smith, S.E. and Read, D.J. (1997) Mycorrhizal Symbiosis. Academic Press., London, UK.
Söderström, B. and Bååth, E. (1978) Soil microfungi in three Swedish coniferous forests.
Holarctic Ecology 1: 62-72. 240
Sowden, F.J., Morita, H. and Levesque, M. (1978) Organic nitrogen distributions in selected peats and peat fractions. Canadian Journal of Soil Science 58: 237-249.
Specht, R.L. (1963) Dark island heath (Ninety-Mile Plain, South Australia.) VII. The effect of fertilisers on composition and growth, 1950-1960. Australian Journal of Botany
11: 67-94.
Specht, R.L. and Rayson, P. (1957) Dark island heath (Ninety-Mile Plain, South
Australia.) 1. Definition of the ecosystem. Australian Journal of Botany 5: 52-85.
Specht, R.L. and Rundel, P.W. (1990) Sclerophylly and foliar nutrient status of mediterranean-climate plant communities in southern Australia. Australian Journal of
Botany 38: 459-474.
Straker, C.J. (1996) Ericoid mycorrhiza: ecological and host specificity. Mycorrhiza 6:
215-225.
Starrett, M.C., Blazich, F.A., Shafer, S.R. and Grand, L.F. (2001a) In vitro colonization of micropropagated Pieris floribunda by ericoid mycorrhizae. I. Establishment of mycorrhizae on microshoots. Hortscience 36: 353-356.
241
Starrett, M.C., Blazich, F.A., Shafer, S.R. and Grand, L.F. (2001b) In vitro colonization of micropropagated Pieris floribunda by ericoid mycorrhizae. II. Effects on acclimatization and growth. Hortscience 36: 357-359.
Steinke, E., Williams, P.G. and Ashford, A.E. (1996) The structure and fungal associates of mycorrhizas in Leucopogon parviflorus (Andr.) Lindl. Annals of Botany 77: 413-419.
Stevens, P.F. (1971) A classification of Ericaceae: subfamilies and tribes. Botanical
Journal of the Linnean Society 64: 1-53.
Stewart, G.R., Pate, J.S. and Unkovich, M. (1993) Characteristics of inorganic nitrogen assimilation of plants in fire-prone Mediterranean type vegetation. Plant, Cell and
Environment 16: 351-363.
Stoyke, G. and Currah, R.S. (1991) Endophytic fungi from the mycorrhizae of alpine ericoid plants. Canadian Journal of Botany : 347-352.
Stoyke, G. and Currah, R.S. (1993) Resynthesis in pure culture of a common subalpine fungus root association using Phialocephala fortinii and Menziesia ferruginea
(Ericaceae). Arctic and Alpine Research 25: 189-193.
Stoyke, G., Egger, K.N. and Currah, R.S. (1992) Characterization of sterile endophytic fungi from mycorrhizae of subalpine plants. Canadian Journal of Botany 70: 2009-2016. 242
Straker, C.J., Gianinazzi-Pearson, V., Gianinazzi, S., Cleyet-Marel, J-C. and Bousquet,
N. (1989) Electrophoretic and immunological studies on acid phosphatase from a mycorrhizal fungus of Erica hispida. New Phytologist 111: 215-222.
Straker, C.J. (1996) Ericoid mycorrhiza:ecological and host specificity. Mycorrhiza 6:
215-225.
Straker, C.J. and Mitchell, D.T. (1986) The activity and characterization of acid phosphatases in endomycorrhizal fungi of the Ericaceae. New Phytologist 104: 243-256.
Stribley, D.P. and Read, D.J. (1974) The biology of mycorrhiza in the Ericaceae. III.
Movement of carbon-14 from host to fungus. New Phytologist 73: 731-741.
Stribley, D.P. and Read, D.J. (1976) The biology of mycorrhiza in the Ericaceae. VI. The effect of mycorrhizal infection and concentration of ammonium nitrogen on the growth of cranberry (Vaccinium macrocarpon Ait.) in sand culture. New Phytologist 77: 63-72.
Stribley, D.P. and Read, D.J. (1980) The biology of mycorrhiza in the Ericaceae. VII.
The relationship between mycorrhizal infection and capacity to utilise simple and complex organic nitrogen sources. New Phytologist 86: 365-371.
Swofford, D.L. (2002) PAUP*. Phylogenetic analysis using parsimony (*and other methods). Version 4. Sinauer Associates, Sunderland, Massachusetts. 243
Taaffe, G., Brown, E.A., Crayn, D.M., Gadek, P.A. and Quinn, C.J. (2001) Generic concepts in Styphelieae: resolving the limits of Leucopogon. Australian Journal of
Botany 49: 107-120.
Talbot, L.M., Turton, S.M. and Graham, A.W. (2003) Trampling resistance of tropical rainforest soils in the wet tropics of north east Australia. Journal of Environmental
Management 69: 63-69.
Tanner, E.J., Vitousek, P.M. and Cuevas, E. (1998) Experimental investigation of nutrient limitation of forest growth on wet tropical mountains. Ecolgy 79: 10-22
Tenkanen, M. (1998) Action of Trichoderma reesei and Aspergillus oryzae esterases in the deacetylation of hemicelluloses. Biotechnology and Applied Biochemistry 27: 19-24.
Thormann, M.N., Currah, R.S. and Bayley, S.E. (2002) The relative ability of fungi from
Sphagnum fuscum to decompose selected carbon substrates. Canadian Journal of
Microbiology 48: 204-211.
Tilman, D., Wedin, D. and Knops, J. (1996) Productivity and sustainability influenced by biodiversity in grassland ecosystems. Nature 379: 718-720.
244
Tronso, A. and Harman, G. E. (1993). Detection and quantification of N-acetyl-β-D- glucosaminidase, chitobiosidase, and endochitinase in solutions and on gels. Analytical
Biochemistry 208: 74-79.
Ulhoa, C.J. and Peberdy, J.F. (1992) Purification and some properties of the extracellular chitinase produced by Trichoderma hazianum. Enzyme and Microbial Technology 14:
236-240.
Usuki, F., Narisawa, K., Yonezawa, M., Kakishima, M. and Hashiba, T. (2002) An efficient inoculation method for colonisation of Chinese cabbage seedlings by the root endophytic fungus, Heteroconium chaetospira. Journal of General Plant Pathology 68:
326-332.
Usuki, F. and Narisawa, K. (2005) Formation of structures resembling ericoid mycorrhizas by the root endophytic fungus Heteroconium chaetospira within roots of
Rhododendron obtusum var. kaempferi. Mycorrhiza 15: 61-64.
Vainio, E. and Hantula, J. (2000) Direct analysis of wood-inhabiting fungi using denaturing gradient gel electrophoresis of amplified ribosomal DNA. Mycological
Research 104: 927-936.
Vandenkoornhuyse, P., Baldauf, S.L., Leyval, C., Straczek, J., and Young, J.P.W.
(2002a) Extensive fungal diversity in plant roots. Science 295: 2051. 245
Vandenkoornhuyse, P., Husband, R., Daniell, T.J., Watson, I.J., Duck, J.M., Fitter, A.H. and Young, J. P. W. (2002b) Arbuscular mycorrhizal community composition associated with two plant species in a grassland ecosystem. Molecular Ecology 11: 1555-1564.
Van der Heijden, M.G.A., Klironomos, J.N., Ursic, M., Moutoglis, P., Streitwolf-Engel,
R., Boller, T., Wiemken, A. and Sanders, I.R. (1998) Mycorrhizal fungal diversity determines plant biodiversity, ecosystem variability and productivity. Nature 396: 69-72.
Van der Heijden, M.G.A., Klironomos, J.N. and Ursic, M. (1999) “Sampling effect”, a problem in biodiversity manipulation? A reply to David A. Wardle. Oikos 87: 408-410.
Van Leerdam, D.M., Williams, P.A. and Cairney, J.W.G. (2001) Phosphate-solubilising abilities of ericoid mycorrhizal endophytes of Woollsia pungens (Epacridaceae).
Australian Journal of Botany 49: 75-80.
Varma, A. and Bonfante, P. (1994) Utilisation of cell-wall related carbohydrates by ericoid mycorrhizal endophytes. Symbiosis 16: 301-313.
Viaud, M., Pasquier, A., and Brygoo, Y. (2000) Diversity of soil fungi studied by PCR-
RFLP of ITS. Mycological Research 104: 1027-1032.
246
Villarreal-Ruiz, L. Anderson, I.C. and Alexander, I.J. (2004) Interaction between an isolate from the Hymenoscyphus ericae aggregate and roots of Pinus and Vaccinium. New
Phytologist 164: 183-192.
Vrålstad, T., Fossheim, T. and Schumacher, T. (2000) Piceirhiza bicolorata - the ectomycorrhizal expression of the Hymenoscyphus ericae aggregate? New Phytologist
145: 549-563.
Vrålstad, T., Schumacher, T. and Taylor, A.F.S. (2002a) Mycorrhizal synthesis between fungal strains of the Hymenoscyphus ericae aggregate and potential ectomycorrhizal and ericoid hosts. New Phytologist 153: 143-152.
Vrålstad, T., Myhre, E. and Schumacher, T. (2002b) Molecular diversity and phylogenetic affinities of symbiotic root-associated ascomycetes of the Helotiales in burnt and metal polluted habitats. New Phytologist 155: 131-148.
Vrålstad, T. (2004) Are ericoid and ectomycorrhizal fungi part of a common guild? New
Phytologist 164: 7-10.
Wang, C.J.K. and Wilcox, H.E. (1985) New species of ectendomycorrhizal and pseudomycorrhizal fungi: Phialophora finlandia, Chloridium pacisporum and
Phialocephala fortinii. Mycologia 77: 951-958.
247
Wardle, D.A. (1999) Is “sampling effect” a problem for experiments investigating biodiversity-ecosystem function relationships? Oikos 87: 403-407.
Weiss, M. and Oberwinkler, F. (2001) Phylogenetic relationships in Auriculariales and related groups – hypotheses derived from nuclear ribosomal DNA sequences.
Mycological Research 105: 403-415.
Weiss, M., Selosse, M-A., Rexer, K-H., Urban, A. and Oberwinkler, F. (2004)
Sebacinales: a hitherto overlooked cosm of heterobasidiomycetes with a broad mycorrhizal potential. Mycological Research 108: 1003-1010.
West, C.P., Izekor, E., Oosterhuis, D.M., and Robbins, R.T. (1987) Endophytic fungus effects on tall fescue tolerance of drought and nematodes. Arkansas Farm Research 37: 1.
Whalley, A.J.S. (1996) The xylariaceous way of life. Mycological Research 100: 897-922.
White, T.J., Bruns, T., Lee, S. and Taylor, J. (1990). Analysis of phylogenetic relationships by amplification and direct sequencing of ribosomal RNA genes. In: PCR
Protocols: A Guide to Methods and Applications (eds. M.A. Innis, D.H. Gefland, J.J.
Sninsky and T.J. White), pp 53-65. Academic Press, New York, USA.
Whittaker, S.P. and Cairney, J.W.G. (2001) Influence of amino acids on biomass production by ericoid mycorrhizal endophytes from Woollsia pungens (Epacridaceae).
Mycological Research 105: 105-111. 248
Williams, R. (1986) Research into propagation of Australian native plants. International
Plant Propagators Society 36: 183-187.
Williams, A.F., Chambers, S.M., Davies, P.W., McLean, C.B. and Cairney, J.W.G.
(2004) Molecular investigation of sterile root-associated fungi from Epacris microphylla
(Ericaceae) and other epacrids at alpine, subalpine and coastal heathland sites.
Australasian Mycologist 23:94-104.
Wurzburger, N., Bidartondo, M.I. and Bledsoe, C.S. (2001) Characterizationof Pinus ectomycorrhizas from mixed conifer and pygmy forests using morphotyping and molecular methods. Canadian Journal of Botany 79: 1211-1216.
Xiao, G. and Berch, S.M. (1992) Ericoid mycorrhizal fungi of Gaultheria shallon.
Canadian Journal of Botany 74: 337-346.
Xiao, G. and Berch, S.M. (1995) The ability of known ericoid mycorrhizal fungi to form mycorrhizae with Gaultheria shallon. Mycologia 87: 467-470.
Xiao, G. and Berch, S.M. (1996) Diversity and abundance of ericoid mycorrhizal fungi of
Gaultheria shallon on forest clearcuts. Canadian Journal of Botany 74: 337-346.
249
Xiao, G. and Berch, S.M. (1999) Organic nitrogen use by salal ericoid mycorrhizal fungi from northern Vancouver Island and impacts on growth in vitro of Gaultheria shallon.
Mycorrhiza 9: 145-149.
Zak, J.C., and Visser, S. (1996) An appraisal of soil fungal biodiversity: the crossroads between taxonomic and functional biodiversity. Biodiversity and Conservation 5: 169-
183.
Zhang, Y-H. and Zhuang, W-Y. (2004) Phylogenetic relationships of some members in the genus Hymenoscyphus (Ascomycetes, Helotiales). Nova Hedwigia 78: 475-484.
Zvyagintsev, D. G. (1994) Vertical distribution of microbial communities in soils. In:
Beyond the Biomass (eds. K. Ritz, J. Dighton and K. E. Giller), pp 29-37. John Wiley and
Sons, Chichester, UK.
250
APPENDIX I
Modified Melin Norkrans medium (MMN) (Marx & Bryan, 1975).
The agar medium consisted of (mg l-1):
(NH4)2HPO4 500
KH2PO4 300
MgSO4.7H2O 140
Glucose 10000
Agar 13000 CaCl2 50
NaCl 25
ZnSO4 3
Thiamine 0.13
The pH was then adjusted to 5 – 5.5 with 2M HCl before the addition of
Citric acid + Fe EDTA 12.5
The medium was then sterilised by autoclaving at 121oC for 15 min and cooled to approximately 55oC before the plates were poured.
Agar was excluded for liquid MMN medium.