Biomechanics of Rhizomorph Development in Armillaria Mellea

Home , Armillaria, Coprinus sterquilinus

MIAMI UNIVERSITY The Graduate School

Certificate for Approving the Dissertation

We hereby approve the Dissertation

Levi Yafetto

Candidate for the Degree:

Doctor of Philosophy

______Director Dr. Nicholas P. Money

______Reader Dr. Diana J. Davis

______Reader Dr. John Z. Kiss

______Reader Dr. Nancy Smith-Huerta

______Graduate School Representative Dr. Richard T. Taylor

ABSTRACT

BIOMECHANICS OF RHIZOMORPH DEVELOPMENT IN ARMILLARIA MELLEA

Levi Yafetto

Fungal rhizomorphs are complex, multicellular, root-like organs formed through the aggregation, interlacing, and adhesion of millions of tip-growing hyphae. There has been very little research on the invasive mechanism utilized by rhizomorphs to penetrate compacted soils and woody substrates. Initial studies with Meruliporia incrassata, a wood-decay fungus that decomposes wooden components of buildings with an annual value of destruction estimated in millions of dollars, was aimed at inducing rhizomorphs in vitro. This attempt was not very successful, as only mycelial cords were produced. The pathogen Armillaria mellea was therefore chosen because it readily forms rhizomorphs in culture and serves as an excellent model for developmental studies. This dissertation presents findings from experiments designed to study (i) comparative features of rhizomorph anatomy in M. incrassata and A. mellea that support its invasive behavior; (ii) the adaptive growth response of rhizomorphs subjected to mechanical stress; (iii) the biochemical basis of turgor generation within rhizomorphs, and (iv) novel measurements of the forces exerted by growing rhizomorphs. Anatomical studies of rhizomorphs of A. mellea cultured in potato dextrose agar (PDA) revealed zones of hyphal tissues namely, an outer layer of peripheral hyphae, radial hyphae, longitudinal hyphae, and a central cavity. A. mellea rhizomorphs were observed to have faster growth than mycelia in PDA. We determined that increasing concentration of agar stimulated the production of more rhizomorphs, with those in media having higher concentration of agar extending faster with tapered tips. Turgor generation within A. mellea rhizomorphs was shown to be partially due to the accumulation of osmolytes. Erythritol and mannitol were identified using Gas Chromatography/Mass Spectrometry (GC/MS) and quantitatively determined to be the most dominant osmolytes that contribute to turgor generation. Osmometric studies revealed that substantial portion of turgor generated was used to exert pressure at the tip of the rhizomorphs

during invasive growth. The varying amounts of force that these A. mellea rhizomorph tips exert were measured, using a sensitive strain gauge. Our experiments provide the first clear picture of the mechanical processes that allow rhizomorphs to function as migratory, exploratory and invasive organs in low-moisture and nutrient-poor environments that present substantial obstacles to fungal colonization.

BIOMECHANICS OF RHIZOMORPH DEVELOPMENT IN ARMILLARIA MELLEA

A DISSERTATION

Submitted to the faculty of

Miami University in partial

fulfillment of the requirements

for the degree of

Doctor of Philosophy

Department of Botany

Levi Yafetto

Miami University

Oxford, Ohio

2008

Dissertation Director: Dr. Nicholas P. Money

Table of Contents Page

CERTIFICATE FOR APPROVING THE DISSERTATION ABSTRACT TITLE PAGE …………………………………………………………………………... i TABLE OF CONTENTS ……………………………………………………………… ii LIST OF TABLES ……………………………………………………………………... iv LIST OF FIGURES ……………………………………………………………………. vi DEDICATION ………………………………………………………………………….. x ACKNOWLEDGEMENTS ……………………………………………………………. xi

Chapter 1: INTRODUCTION AND LITERATURE REVIEW ……………………. 1 Wood decay basidiomycetes and the ecology of mycelial cords and rhizomorphs ……... 1 Dry rot in buildings: Meruliporia incrassata and Serpula lacrymans …………………... 2 Armillaria mellea: Model for the study of rhizomorphs ………………………………… 3 Mycelial cord and rhizomorph structure ………………………………………………… 5 Translocation in mycelial cords and rhizomorphs ………………………………………. 7 Specific experimental aims …………………………………………………………….. 8 References ……………………………………………………………………………….. 9

Chapter 2: Ultrastructural studies of Meruliporia incrassata and Armillaria mellea using scanning electron microscopy ……………………………………………………. 18 Abstract ………………………………………………………………………………….. 18 Introduction …………………………………………………………………………….... 19 Materials and Methods ………………………………………………………………...... 21 Results ……………………………………………………………………………………. 25 Discussion ………………………………………………………………………………... 27 References ………………………………………………………………………………… 31

Chapter 3: In vitro studies of rhizomorph extension in Armillaria mellea…………….. 43

Abstract …………………………………………………………………………………… 43 Introduction ……………………………………………………………………………..... 44 Materials and Methods ………………………………………………………………...... 45 Results …………………………………………………………………………………….. 48 Discussion ……………………………………………………………………………….... 50 References ………………………………………………………………………………… 54

Chapter 4: Osmolyte accumulation in Armillaria mellea rhizomorphs: Its role in turgor pressure generation ……………….……………………………………………………… 63 Abstract …………………………………………………………………………………… 63 Introduction ……………………………………………………………………………..... 64 Materials and Methods ………………………………………………………………...... 68 Results …………………………………………………………………………………….. 75 Discussion ……………………………………………………………………………….... 76 References ………………………………………………………………………………… 82

Chapter 5: The biomechanics of invasive growth in Armillaria mellea rhizomorphs … 108 Abstract …………………………………………………………………………………… 108 Introduction ……………………………………………………………………………..... 109 Materials and Methods ………………………………………………………………...... 111 Results …………………………………………………………………………………….. 115 Discussion ……………………………………………………………………………….... 117 References ………………………………………………………………………………… 122

Chapter 6: General discussion, Conclusions and Future Studies ……………………… 131 Research background ……………………………………………………………………... 131 General discussion ……………………………………………………………………….. 134 Conclusions……………………………………………………………………………...... 140 Future studies …………………………………………………………………………...... 142 References …………………………………………………………………………………. 144

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List of Tables Page

Table 3.1 Summary of rhizomorph length measurements in Armillaria mellea 62 in PD medium solidified with different concentrations of agar

Table 4.1 Wet and dry weight of Armillaria mellea rhizomorphs before and after 97 cellular extraction

Table 4.2 Mean concentrations of glycerol, erythritol and mannitol determined in 98 Armillaria mellea rhizomorphs

Table 4.3 Data of ion intensity from characteristic 103 ion from different 99 concentrations of glycerol standard solutions

Table 4.4 Data of ion intensity from characteristic 217 ion from different 100 concentrations of erythritol standard solutions

Table 4.5 Data of ion intensity from characteristic 139 ion from different 101 concentrations of mannitol standard solutions

Table 4.6 Data of ion intensity from characteristic 103 ion of glycerol in 102 rhizomorph samples cultured in PD broth with corresponding x values obtained from glycerol standard curve

Table 4.7 Data of ion intensity from characteristic 103 ion of glycerol in 103 rhizomorph samples cultured on cellophane with corresponding x values obtained from glycerol standard curve

Table 4.8 Data of ion intensity from characteristic 217 ion of erythritol in 104 rhizomorph samples cultured in PD broth with corresponding x values obtained from erythritol standard curve

Table 4.9 Data of ion intensity from characteristic 217 ion of erythritol in 105 rhizomorph samples cultured on cellophane with corresponding x values obtained from erythritol standard curve

Table 4.10 Data of ion intensity from characteristic 139 ion of mannitol in 106 rhizomorph samples cultured in PD broth with corresponding x values obtained from mannitol standard curve

Table 4.11 Data of ion intensity from characteristic 139 ion of mannitol in 107 rhizomorph samples cultured on cellophane with corresponding x values obtained from mannitol standard curve

Table 5.1 Individual measurements of applied forces and pressures from 129 Armillaria mellea rhizomorphs

Table 5.2 Mean values of applied force and pressure by rhizomorphs pressing 130 against strain gauge

List of Figures Page

Fig 1.1 The development of fungal mycelium from a spore 15

Fig 1.2 Formation of mycelial strands in Phymatotrichum omnivorum 16

Fig 1.3 (Top) Apical region of rhizomorph of Armillaria mellea 17 (Below) Apical region of rhizomorphs of Armillaria mellea growing in agar medium

Fig 2.1 Apical region of rhizomorphs of Armillaria mellea growing 33 in agar medium

Fig 2.2 Field-collected rhizomorphs of Meruliporia incrassata from an 34 infested home in California

Fig 2.3 Scanning electron micrograph of field-collected Meruliporia 35 incrassata rhizomorph showing constituent hyphae lying parallel to one another

Fig 2.4 (A) Scanning electron micrograph showing longitudinal hyphae 36 of Meruliporia incrassata rhizomorph (arrow). (B) Longitudinal hyphae are tightly packed, protecting the inner layers of hyphae in the middle portion of the organ. (C) Middle portion of loosely packed hyphae, m

Fig 2.5 (A) A vessel hypha of Meruliporia incrassata with thin wall and 37 wide diameter lumen (arrows) in close association with tightly packed regular hyphae. (B) Vessel hyphae embedded in and surrounded by hyphae and soil particles. (C) Vessel hyphae with remnants of degenerated internal wall components

Fig 2.6 Armillaria mellea rhizomorphs (r) growing in potato dextrose 38 broth media, with mycelium (m) on medium surface

Fig 2.7 Scanning electron micrograph of rhizomorph tip of Armillaria 39 mellea grown on cellophane covering potato dextrose agar

Fig 2.8 Scanning electron micrograph of transverse sections of 40 freeze-fractured rhizomorph of Armillaria mellea. (A) Complete hyphal arrangement in rhizomorph. (B) A close up micrograph of hyphal arrangement

Fig 2.9 Scanning electron micrographs of Armillaria mellea rhizomorph. 41 (A) Vessel hypha (v) in close association with tightly packed hyphae. (B) Wide lumen vessel hyphae

Fig 2.10 Scanning electron micrograph of Armillaria mellea rhizomorph 42

Fig 3.1 Relationship between agar concentration and medium gel 57 strength expressed in units of force per unit area (MPa or µN µm-2)

Fig 3.2 Comparative growth of mycelium and rhizomorphs in Armillaria 58 mellea cultured in PDA and incubated in the dark at room temperature over a period of 13 d

Fig 3.3 Effect of increasing gel strength on the production of Armillaria 59 mellea rhizomorphs, after four weeks of incubation

Fig 3.4 Length of rhizomorphs produced in increasing concentrations of 60 agar in PDB in 4-week old cultures of Armillaria mellea

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Fig 3.5 Effect of increasing gel strength on the extension of Armillaria 61 mellea rhizomorphs

Fig 4.1 A chromatogram of derivatized100 mM glycerol standard at a 88 retention time of 16.25 min, showing the corresponding spectrum (insert) with characteristic ion fragment of 103.

Fig 4.2 A log/log standard curve of glycerol showing a regression line 89 with concentrations ranging between 10 µM to 10 mM

Fig 4.3 A chromatogram of derivatized 100 mM erythritol standard at a 90 retention time of 19.25 min, showing the corresponding spectrum (insert) with characteristic ion fragment of 217

Fig 4.4 A log/log standard curve of erythritol showing a regression line 91 with concentrations ranging between 100 µM to 100 mM

Fig 4.5 A chromatogram of derivatized 100 mM mannitol standard at a 92 retention time of 23.80 min, showing the corresponding spectrum (insert) with characteristic ion fragment of 139

Fig 4.6 A log/log standard curve of mannitol showing a regression line 93 with concentrations ranging between 100 µM to 100 mM

Fig 4.7 GC/MS chromatogram of Armillaria mellea rhizomorph cultured on 94 cellophane covering potato dextrose agar, showing peaks of erythritol (*) and mannitol (x)

Fig 4.8 GC/MS chromatogram of Armillaria mellea rhizomorph cultured in 95 PD broth, showing peaks of erythritol (*) and mannitol (x)

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Fig 4.9 Concentrations of osmolytes in Armillaria mellea rhizomorphs 96 cultured on cellophane covering PD agar and in PD broth

Fig 5.1 Calibration curve for rigid-lever strain gauge used in the 125 rhizomorph force experiments

Fig 5.2 Measurement of force exerted by Armillaria mellea rhizomorphs 126 using a rigid-lever strain gauge

Fig 5.3 A representative chart recording the force applied by an 127 Armillaria mellea rhizomorph over a period of 30 min

Fig 5.4 A representative chart recording the force applied by an Armillaria 128 mellea rhizomorph over a period of about 30 min after the addition of 25 mM polyethylene glycol-6000

DEDICATION

This dissertation is dedicated to my father, Clemence Tennyson Agbele, and my mother, Gladys Ivy Dartey, for denying themselves of everything of their lives, and sacrificing them all towards the education of me and my siblings (Joana, Fidel, and Golda), so that we will have the opportunities that they never had. They will forever have a special place in our hearts.

ACKNOWLEDGMENTS

I am sincerely indebted to a number of individuals for their sacrifices and significant roles they played in my studies, resulting in the successful completion of my doctoral degree. First, I want to thank God for being my all in all – His Provision, Protection and Unconditional love – and for bringing me this far. I would like to thank my academic mentor and advisor, Dr. Nicholas P. Money, for his extraordinary dedicated guidance and support during the course of my studies and research, making sure he always brought out the best in me. I am greatly indebted to him for his ingenious and insightful suggestions during experimental designs. His patience, constructive criticisms, and willingness to listen and share ideas during our numerous discussions are qualities that make him one of the best mentors I have ever known. Ours is an exceptional mentor-student relationship which is based on mutual respect and great friendship. He really believed in my abilities, and never hid his desire to see me fulfill my dreams and succeed. His enthusiasm and interest in my research, coupled with his desire that I become a good, thinking scientist leaves me in awe of him. Without him, this work wouldn’t have been a success. For most part, I will forever remember Dr. Money for these words whenever things get tougher than expected: “Keep pushing.” What else can I say than “THANK YOU” for his kind heart, and for giving me a second chance. I am also greatly indebted to members of my doctoral committee: Dr. Diana J. Davis, Dr. John Z. Kiss, Dr. Nancy Smith-Huerta and Dr. Richard T. Taylor, whose excellent guidance and constructive criticisms ensured I was on track with my studies and research. I also express my indebtedness to them for reading this dissertation to make it better. I personally want to thank Dr. Richard T. Taylor for teaching me the use of the GC/MS technique, and for sharing the instrument with me, as he made sure it was always in a perfect condition for my work. I am most grateful for his patience and understanding during these periods of my study. My gratitude also goes to Dr. Diana J. Davis, who always took time off her busy schedule at the College of Mount St. Joseph, Cincinnati, to be with me in the laboratory to analyze GC/MS data, and to offer very useful suggestions. Without them, this work would not have been completed.

My sincere thanks goes to the Department of Botany, Miami University for providing funds through the Academic Challenge Grant to enable me acquire all the necessary laboratory materials for these studies. Two persons whose input in my research cannot be overlooked are Dr. Richard E. Edelmann and Mr. Matt Duley of the Electron Microscopy Facility of the Miami University, who painstakingly ensured I got the best from the facility, and for providing valuable suggestions to capture and edit some of the best images for this work. I thank them for being very patient with me, and being there all the time to answer my questions. Special thanks to my colleague, Mr. Aaron Kennedy for his friendship and his help with molecular identification of field-collected rhizomorphs. I acknowledge Luiz De La Cruz of the Luis De La Cruz Wood Preservation Services, Pasadena, CA, for providing field-collected rhizomorphs of M. incrassata for this work, and to the Forest Products Laboratory of the United States Department of Agriculture, Madison, Wisconsin, for providing M. incrassata strain FP-150521 for in vitro studies. I am grateful to Dr. Dana Richter, School of Forest Resources and Environmental Sciences, Michigan Technological University, Michigan for providing A. mellea strain DR-140 for this work. Finally, I want to take this unique opportunity to thank my beautiful, loving wife, Rodlyn Yafetto, for her understanding, patience and care when I have to leave her home alone to spend long hours in the laboratory and to write this dissertation. I am greatly indebted to you for your love and support in times of my shortcomings. I am very fortunate to have you as my best friend and wife. I love you!

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CHAPTER 1

INTRODUCTION AND LITERATURE REVIEW

1.1 Wood decay basidiomycetes and the ecology of mycelial cords and rhizomorphs

Fungi play vital ecological roles as decomposers of woody debris. Most species rot fallen timber in forest ecosystems, and cause plant diseases, but a few of them attack wood in buildings. Many of these wood-decay fungi are basidiomycetes (Schmidt, 2006). For example, fungi of the genera Armillaria, Auricularia, Coniophora, Fomitopsis, Meripilus, Meruliporia, Polyporus, and Serpula are well-known wood decomposers. In order to decompose their substrates (straw, foliage, and wood debris), wood-decay fungi employ many strategies in the heterogeneous environments in which they thrive (Lindahl and Olsson, 2004; Schwarze, 2007). The most common among these is the secretion of enzymes by fungal hyphae to break down complex molecules (cellulose, lignin, pectin, and chitin) in the substrates into smaller molecules (sugars, organic acids, and amino acids), after which the nutrients are absorbed and translocated from the substrate (the source) to parts of the hypha or mycelium where they are needed most (the sink) for growth (Lindahl and Olsson, 2004; Money, 2007). Dowson et al. (1989) observed that Phanerochaete velutina and Hypholoma fasciculare (both wood decay-fungi) extend their mycelia into soil as they deplete nutrients from already colonized wood substrates. Mycelial cords and rhizomorphs are used as exploratory and migratory organs that enhance the survival of basidiomycetes. Mycelial cords and rhizomorphs of wood-decay and pathogenic fungi serve as important sources of inoculum, enabling fungi to make contact with newly colonized substrate. In other words, mycelial cords and rhizomorphs become the inoculum potential, defined by Garrett (1956) as “the energy of growth of a fungus available for colonization of a substrate at the surface of the substrate to be colonized”. Although both are constructed from hyphae and function in similar fashion, they are structurally different. As mentioned above, hyphae secrete enzymes during growth to degrade nutrients in the substrates they colonize. Subsequent absorption of these nutrients sustains vegetative growth, and allows an increase in hyphal biomass, which is achieved through tip extension and branching. This radiating hyphal biomass is what is collectively called a mycelium (Fig 1.1; Carlile, 1994). As

fungi use this absorptive mechanism to acquire their nutrients, with time, these nutrients are depleted from the substrate, leaving the fungi no other option than to search for other supportive substrates rich in nutrients for growth and survival. This search is accomplished through mycelia, mycelial cords, and rhizomorphs (Hartig, 1874; Falck, 1912). Mycelial cords are less complex in structure than rhizomorphs. They are linear aggregate of hyphae formed behind an advancing mycelial margin in which the hyphae are loosely separated (Fig 1.2). Because of this arrangement, the appearance of mycelial cords characterized by a fan-like mycelial mat. A rhizomorph is a complex multicellular root-like organ formed through aggregation, interlacing, and adhesion of scores of hyphae. The rhizomorph has a waterproof surface, and a melanized rind that encloses a central open cavity, which is believed to serve as a channel for the conduction of oxygen under anoxic conditions (Fig 1.3).

1.2 Dry rot in buildings: Meruliporia incrassata and Serpula lacrymans

The rhizomorph-forming fungus Meruliporia incrassata causes “dry rot” of wood components in buildings. Its damage is sometimes referred to as brown rot. Since its description from pine as Merulius incrassatus Berk. & Curt. in South Carolina, M. incrassata (Berk & Curtis.) Murr. (Family Coniophoraceae) has been recognized as a wood-decay fungus in the North America (Berkley and Curtis, 1849). It has a wide distribution in the United States and has been reported occasionally in Canada (Verall, 1968; Palmer and Eslyn, 1980). It has been reported mainly in the southeastern United States. Its occurrence in the southern British Columbia, Canada is an interesting observation, since this region is very moist. There has been no report of the occurrence of M. incrassata in Arizona or New Mexico, an observation attributed to the dry climate of the desert southwest. Its distribution pattern suggests that it grows best in warm climates (Burdsall, 1991). M. incrassata causes a great deal of damage to homes in California. During attacks on homes, M. incrassata develops rhizomorphs from its depleted food base mostly from outside the home (tree stumps, landscaping lawns with wood chips, etc.). Under warm and very moist conditions, the rhizomorph is capable of translocating nutrients and water to its tip until it encounters a wooden component of the building. It then begins to colonize the wood swiftly by digesting and extracting cellulose, leaving behind the brown lignin component of the wood. The

description of M. incrassata as a dry rot fungus can be regarded as erroneous because wood decomposition is impossible without water. This fungus destroys dry wood, or wood that was dry, by transferring water through its rhizomorphs to the sites of active growth and decomposition (Verall, 1968). Possible sources of invasion of homes in California may include the following. First, when lands are cleared of trees for construction of homes, stumps from trees are sometimes left in the soil. An array of fungi colonizes these stumps until the nutrients are depleted, when the rhizomorph-forming species develop rhizomorphs in search of new sources of food. Second, topsoil and wood mulch used in landscaping may often provide a combination of optimum conditions (moisture and poor ventilation) for wood-decay fungi to develop rhizomorphs. This normally results from the continuous supply of water by sprinklers under low nutrient conditions. As long as the fungus has access to water and can channel it to the tips of its rhizomorphs, it survives until it locates a nutrient source in the building. Third, untreated wood used to construct buildings is another potential source of M. incrassata growth and rhizomorph development. Under all these conditions, the destructive abilities of the rhizomorphs manifest themselves when they contact wooden components of buildings (Money, 2004). Meanwhile, a different fungal species, Serpula lacrymans, is known to be responsible for the same kinds of destruction in Europe (especially in Britain), and other parts of the world, including Japan and Australia (Money, 2004). For example, S. lacrymans, together with other fungi, caused enormous historical damage to wood in ships. Rotting of timber was responsible for sinking the ship Royal George in 1782 and necessitated rebuilding of Queen Charlotte in 1810 (Money, 2004). Both M. incrassata and S. lacrymans produce brown basidiomes, with brown, thick-walled, dextrinoid basidiospores; both cause brown rot of their wood substrates; and both produce water- and nutrient-conducting rhizomorphs or mycelial cords (Burdsall, 1991; Moore, 1994).

1.3 Armillaria mellea: Model for the study of rhizomorphs

Armillaria (Fr.: Fr) Staude (Agaricales, Tricholomataceae) is a cosmopolitan genus of pathogenic basidiomycetes that contains about 40 species (Volk and Burdsall, 1995; Pegler,

2000) known for causing root rot and butt rot diseases. The term “honey fungus” has been applied to many species of Armillaria. It refers to the yellowish caps and sweet taste of their mushrooms (Yoshida et al. 1984). Armillaria has a wide range of hosts including deciduous and coniferous trees, shrubs, and vines (Smith et al. 1992), and it is considered as one of the most devastating disease-causing plant pathogens; it kills and decomposes plants in forests and plantations in both temperate and tropical regions of the world (Garraway et al. 1991). Of all species of the genus Armillaria, Armillaria mellea (Vahl: Fr) Kummer is the most pathogenic, and the best studied. Other species, including A. luteobubalina and A. ostoyea, have also been reported to be pathogenic (Fox, 2000). A. mellea, like other Armillaria species has the characteristic of producing reddish- to black-colored rhizomorphs, which have unique internal structure (Townsend, 1954). Interestingly, the mycelium and rhizomorphs of some Armillaria species are bioluminescent and have been studied over a century (Murrill, 1915; Buller, 1924; Harvey, 1952; Mihail and Bruhn, 2007). Surprisingly, their fruiting bodies are not luminescent (Buller, 1924; Harvey, 1952; Wassink, 1978). As a root pathogen, Armillaria is quite difficult to detect in nature, since it occurs everywhere in the soil, and its presence is known only when fruiting bodies are produced at the base of a tree trunk. This makes its control unfeasible (Schmidt, 2006). In 1992, for example, Smith et al. (1992) identified mycelium of A. bulbosa in Michigan using molecular genetics techniques. It was reported to have occupied an area of 15 hectares, weighed about 10,000 kg and was more than 1,500 years old, making it one of the largest and oldest living organisms known. In most cases, Armillaria is a saprobe, colonizing dead tree stumps in a field, but it becomes parasitic, either as a necrotroph or biotroph when it infects the roots of a healthy host using its networks of rhizomorphs (Day, 1927; Thomas, 1934; Patton and Riker, 1959; Watling et al. 1991). A rhizomorph adheres firmly to root tips of healthy plants after the mucilaginous material on its tip dries, sends out mycelia and continues to grow and spread beneath the bark of the tree producing mycelial fan. Competition from neighboring trees, insect attack and the prevalence of adverse climatic factors further affects the health of the infected host. In such a compromised condition, the host is killed outright if it fails to successfully defend itself and the pathogen manages to reach the xylem to interfere with flow of sap in the cambium (Wahlström

and Johansen, 1992; Woodward, 1992; Schmidt, 2006). Thus rhizomorphs are used by Armillaria as a means to propagate itself to effect infection over long distances (Hartig, 1874; Garrett, 1970). Initial research towards my doctoral degree was focused on mycelial cord and rhizomorph development in Meruliporia incrassata to understand the functions of these systems in relation to their structure and potential as organs of migration, translocation, and destruction. This interest was borne out by a number of facts. First, M. incrassata is a very destructive wood decay fungus that damages buildings, with costs running into several millions of dollars annually. Second, not much is understood about the mechanisms involved in the migration of mycelial cords and rhizomorphs from one location to another. Lastly, virtually no studies had been done on the biomechanics of mycelial cord and rhizomorph development, justifying the urgency in further studying rhizomorph-forming species that are of immense economic importance in real estate, forest ecology and plant pathology. Interestingly, in vitro cultures of M. incrassata produced only mycelial cords and not rhizomorphs, making the study of rhizomorph mechanics impossible. Since biomechanical studies form the core of this research, I shifted focus to the plant pathogenic fungus, Armillaria mellea, because it readily produces rhizomorphs in vitro, making it an excellent model for developmental studies in rhizomorph systems.

1.4 Mycelial cord and rhizomorph structure

Fungal hyphae aggregate to serve as the “building blocks” of more complex structures in fungi, including fruiting bodies, sclerotia and rhizomorphs. Tube-shaped hyphal aggregates, particularly those formed by basidiomycetes, have been named variously as “strands,” “cords,” “rhizomorphs,” or “syrrotia” (Garrett, 1970; Watkinson, 1971; Thompson, 1984). Until recently, these names had been used interchangeably in literature by authors. To solve this confusing situation, Rayner et al. (1985) suggested the adoption of the terms “cords” and “rhizomorphs,” to denote apically diffuse mycelia aggregates and apically dominant growing tips, respectively. This was readily accepted and had been used since. In this study, we refer to hyphal aggregates of M. incrassata as mycelial cords because they possess apically diffused margins, whereas

5 hyphal aggregates of A. mellea are called rhizomorphs because they have apically dominant growing tips. Rhizomorphs have a similar appearance to the roots of higher plants and both function as organs of nutrient and water absorption and translocation. Rhizomorphs can also pass air along their lengths to prevent suffocation of the fungus while they explore the anoxic interior of substrates for nutrients (Money, 2004). Rhizomorphs have a highly organized apical growing tip with extreme apical dominance. This apical region contains a compact growing point of tightly packed cells, which is protected by a cap of intertwined hyphae in a mucilaginous matrix. Because of these root-like attributes, rhizomorphs were once thought to have meristems responsible for organized tip growth similar to the growth mechanism of roots (Garrett, 1963, 1970). It is now known that rhizomorphs do not grow as a result of meristematic activity, but rather by extension of the organized apical hyphae (Rayner et al. 1985). According to Moore (1994), the medullary zone behind the tip contains vessel hyphae composed of swollen, vacuolated and often multinucleate cells surrounded by copious air- or mucilage-filled spaces. However, other authors have reported wide lumen vessel hyphae in the medullary region of field-collected rhizomorphs which lack cytoplasmic content, and have suggested that long distance translocation of nutrients and water is their major role (Eamus et al. 1985; Jennings, 1987: Cairney et al. 1988). Constituent hyphae of mycelial cords that are relatively loosely aggregated develop as a result of the adhesion and growth of young branches of hyphae over an older leading hypha, which defines the building block for the development of the mycelial cord (Moore, 1994). Further, the inclusion of other lateral hyphae these constituent hyphae may encounter results in increasing the size of the cord (Nuss et al. 1991). Unlike rhizomorphs, mycelial cord formation is viewed as a secondary process that takes place behind the growing hyphal apices distal to the colony margin (Garrett, 1981), and it involves increasing aggregation of hyphae within the ageing mycelium. During this process, anastomoses between the hyphae on the surface of the cords occur, fusing them into a bundle. Three common hyphal types have been identified that constitute mycelial cords and rhizomorphs: “tendril” hyphae (or narrow hyphal branches) from the older regions of the main hyphae that interlace around the other hyphae (Nuss et al. 1991); central “vessel hyphae” wide in

diameter and thin walled; and thick-walled, but narrow “fiber” hyphae of old cords that are formed, running longitudinally through the mature cord to provide strength.

1.5 Translocation in mycelial cords and rhizomorphs

Mature cords and rhizomorphs have the ability to translocate nutrients and water towards their tips when fungi migrate from one food source to the other, especially over nutrient-poor surfaces (Jennings et al. 1974; Anderson and Ullrich, 1982; Eamus et al. 1985; Granlund et al. 1985; Cairney, 1992). During this period, the hyphae serve as a repository of absorbed nutrients, and grow as a cohesive unit. However, when these hyphae encounter a substrate with more nutrients cohesive growth is lost and the hyphae spread out with a characteristic invasive pattern of growth to completely colonize the new substrate (Moore, 1994). Until they come into contact with their new sources of food, they are regarded as migratory organs that develop and advance from one food base over nutrient-poor substrates such as bricks, stones, tiles, plastic sheets (Arora, 1986; Money, 2004). Brownlee and Jennings (1981) studied the content of soluble carbohydrates and their translocation in S. lacrymans colonies. Their analysis of sugars and polyols revealed diverse compounds in different mycelial regions (margin, submarginal and mid-region) and in the cords, including glycerol, mannitol, arabitol, trehalose, glucose, fructose, and sucrose. Trehalose and arabitol were reported to be the dominant carbohydrates in the mycelium. Trehalose was reported as the major carbohydrate in cords (over 70% of total soluble fraction) with arabitol occurring in high levels in marginal regions. 14Carbon labeling of the carbohydrates showed significant incorporation into trehalose, implicating trehalose as the main form by which carbohydrate is translocated in cords in S. lacrymans. Granlund et al. (1985) studied the translocation of solutes along rhizomorphs of A. mellea. Observations indicated that solutes (glycerol, mannitol, arabitol, trehalose, glucose, fructose and sucrose) were translocated bidirectionally along A. mellea rhizomorphs, and at similar velocity in both directions. Erythritol and glycerol were reported to be the major soluble carbohydrates, with maltose, sucrose and trehalose forming a significant part of the total amount of carbohydrates present. Along the length of the rhizomorph, glycerol was found to have a high

7 specific activity at the source where 14C glucose had been fed, suggesting that glycerol could be the major metabolic product of the assimilated glucose. However, away from the source, mannitol was detected at the highest specific activity, suggesting that mannitol could be the compound that was translocated, although this assertion remained unproven at the time. The authors, however, did not rule out the possibility that other compounds were translocated.

1.6 Specific experimental aims

Rhizomorphs have been reported to grow long distances over non-woody substrates such as concrete, rocks, metals, insulation materials (Arora, 1986: Money, 1994). Rishbeth (1968) reported that A. mellea rhizomorphs are capable of growing several meters through the soil. This phenomenon has also been reported in S. lacrymans, where cords formed could be several meters long over such nutrient-poor substrates (Butler, 1957, 1958; Watkinson, 1971; Jennings and Watkinson, 1982; Jennings, 1991). In the case of rhizomorphs, which are characterized by organized tip growth, not much is known about mechanisms of migration from one substrate to another, especially when the tip has to push through compact, heterogeneous soil medium. In this research, an attempt has been made to understand the stimulatory effects of substrate compactness on rhizomorph development and their success as organs for absorption, translocation and infection. To this end, specific objectives of these studies which have culminated in this dissertation include the following: 1. To examine in detail features of rhizomorph anatomy that support their migratory, translocatory and invasive behavior. 2. To study the effects of increasing medium gel strength on the production, growth and extension rates of A. mellea rhizomorphs in vitro. 3. To examine osmolyte accumulation in A. mellea and its role in the generation of tugor pressure within rhizomorphs. 4. To estimate, for the first time, how much force may be generated by these rhizomorphs as they migrate through compact soil and penetrate their hard woody substrates.

Findings from these experiments have contributed to a broader understanding of the functions of rhizomorphs in relation to their structure and development.

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Fig 1.1 – The development of fungal mycelium from a spore. (a) A germ tube germinating from a fungal spore (black dot); (b) Vegetative hyphae develops from germ tube and branches; (c) Further growth and branching in hyphae increases the biomass of the fungus producing (d) a highly-branched mycelium. Adapted from www.anbg.gov.au/fungi/mycelium

Fig 1.2 – Formation of mycelial strands in Phymatotrichum omnivorum, (top) Small hyphae beginning to grow over the surface of the wide, central, leading hypha; (middle) central hypha surrounded by loose network of small hyph ae; (below) deposition of the second hyphal layer. Adapted from Garrett (1970), after Rogers and Watkins ( 1938).

Fig 1.3 – (Top) Apical region of rhizomorph of Armillaria mellea. Adapted modified from Garrett (1970), after Hartig (1873). (Below) Apical region of rhizomorph of Armillaria mellea growing in agar medium.

CHAPTER 2

ULTRASTRUCTURAL STUDIES OF MERULIPORIA INCRASSATA AND ARMILLARIA MELLEA USING SCANNING ELECTRON MICROSCOPY

ABSTRACT

Rhizomorph structure of Meruliporia incrassata and Armillaria mellea was analyzed in detail using scanning electron microscopy. M. incrassata, as a wood decay basidiomycete, has the potential to destroy wooden components of buildings, using rhizomorphs that have the ability to migrate from one location to another in search of suitable substrates. Ironically, very little is understood about the structure of M. incrassata rhizomorphs and their role in water and nutrient translocation as it explores its environment. By learning in details the arrangement of these hyphal tissues in A. mellea rhizomorphs, we aimed to understand the structure of giant rhizomorphs of the dry rot fungus, M. incrassata, and how they operate when they destroy buildings. Calculations showed that rhizomorphs of field-collected M. incrassata and cultured A. mellea were constructed from millions of constituent hyphae. Scanning electron micrographs of field-collected rhizomorphs of M. incrassata showed less differentiation and organization of constituent hyphae than in A. mellea. We identified four types of constituent hyphae in A. mellea rhizomorphs with unique arrangements, namely, peripheral hyphae, radial hyphae, longitudinal hyphae (which is comprised of an outer thick-walled, narrow lumen hyphae, and thin-walled, wide lumen inner vessel hyphae), and a central cavity. We determined that the two types of longitudinal hyphal tissues are common in the rhizomorph structure of both M. incrassata and A. mellea. We also determined that M. incrassata rhizomorph does not have a conspicuous central cavity, whereas A. mellea is characterized with a central cavity. We concluded that the different hyphal arrangements and organization within rhizomorphs underscore the importance in their migratory, translocatory and invasive abilities as tip-growing organs.

1. INTRODUCTION

Wood-decay fungi have various strategies for spreading in the heterogeneous environments they inhabit. These include the use of mycelia, mycelial cords, rhizomorphs, and spores of fruiting bodies. Fungi direct much of their energy into the mass production of mycelium when nutrients in their substrate are being depleted, and especially when the mycelial front encounters new substrate (Lindahl and Olsson, 2004). Most basidiomycetous fungi use fruiting bodies to produce vast numbers of spores that are released from the gill surface, fall under gravitational pull, and are then blown away by wind currents. When these spores encounter favorable environmental factors, they germinate to produce mycelia that begin to explore the surrounding environment for nutrients. In other fungi, especially root-infecting and wood-decay fungi, mycelial cords and rhizomorphs have long been known as important agents by which these fungi spread (Hartig, 1874; Falck, 1912). It was only recently that much attention was paid to studying the structure and development of cords and rhizomorphs in an attempt to understand how they function. Townsend (1954) studied rhizomorph structure of sixteen basidiomycete fungi including Armillaria mellea (Fr.) Quel., Marasmius androsaceus Fr., Serpula lacrymans Fr., Phallus impudicus Pers. Although the rhizomorph structure of some of these fungi was described, Townsend (1954) acknowledged that other fungi did not form rhizomorphs in vitro, but results obtained provided some insights into the influence natural factors may have on their production in soil or decaying wood. Unlike M. incrassata, some studies on biochemical, physiological and anatomical aspects of Serpula lacrymans and A. mellea have been undertaken in the past couple of decades (Brownlee and Jennings, 1981a, b; Hornung and Jennings, 1981; Jennings and Watkinson, 1982; Granlund et al. 1984, 1985; Eamus, et al. 1985). Burdsall (1991) reviewed the distribution, significance and occurrence of M. incrassata in the United States. Hornung and Jennings (1981) studied the development of surface mycelium of S. lacrymans with emphasis on different stages of growth. Hyphal types observed within S. lacrymans mycelia were categorized as: undifferentiated hyphae common to all stages of mycelial development; main hyphae of faster growth rate that have the tendency to form hyphal aggregates; tendril hyphae from which hyphal aggregates originate; fiber hyphae which are often coenocytic, but thick-walled, and vessel hyphae which are often thin-walled with a wide lumen.

Jennings and Watkinson (1982) compared initial stages of mycelial cord development of S. lacrymans in culture with the structure of mature cords from infected wood. Mycelia cords have been reported to be formed by a secondary process, unlike rhizomorphs (Garrett, 1981). The extent to which hyphae are organized and differentiated varies between species (Butler 1966). Mycelial cords from S. lacrymans (Wulf ex Fr.) Schroet have earlier been reported to be made of “vessels” (wide but empty) and “fiber” hyphae (narrow but thick-walled) running longitudinally through the hyphal aggregate (Hartig and van Tubeuf, 1902; Falck, 1912). In another study, Granlund et al. (1984) performed scanning electron microscopy (SEM) studies on A. mellea rhizomorphs to investigate the internal and external features of these structures in relation to their reputed ability to translocate water and other solutes over long distances. The internal structure revealed the arrangement of tissues of the A. mellea rhizomorph: the different tissue zones observed were (i) peripheral hyphae that formed a layer approximately between 20 and 30 µm thick, (ii) cortex, (iii) medulla and (iv) the central space, usually occupied by a cottony pith of fine loosely-woven hyphae. During extensive preliminary studies in our laboratory, we were unable to generate rhizomorphs of M. incrassata using a variety of culture methods. Thin mycelial strands developed in older cultures, but nothing resembling the giant organs involved in building decay formed in vitro. For this reason, we refocused our experiments on field-collected rhizomorphs of M. incrassata and cultured rhizomorphs of A. mellea. Although Granlund et al. (1984) had already described the internal and external structure of A. mellea rhizomorphs using SEM, we saw the need to redo this work for two reasons; (i) so that we will have a standardized method of study that will enable us to compare the structure of field-collected M. incrassata rhizomorphs with that of A. mellea, and (ii) to confirm or disprove what they had already reported on A. mellea rhizomorph structure. We reasoned that information on the structure and function of rhizomorphs of A. mellea was likely to be applicable to the invasive organs of other basidiomycetes, including M. incrassata. In other words, we chose A. mellea as our experimental model for understanding rhizomorph biology. The main objective of the study was to analyze rhizomorph structure in detail, using electron microscopy, focusing on the arrangement of hyphal types that form them. Calculations indicated that rhizomorphs are constructed from millions of constituent hyphae. Comparison between M. incrassata organs and those of A. mellea was an interesting aspect of this work.

Scanning electron micrographs of field-collected rhizomorphs of M. incrassata showed less differentiation and organization of constituent hyphae. They also lacked organized tip-growth that is found in A. mellea rhizomorphs. A. mellea showed well organized rhizomorphs characterized by a complex arrangement of hyphal types and polarized tip-growth. M. incrassata and A. mellea both had thick-walled, narrow lumen, tightly packed hyphae arranged longitudinally, and thin-walled, wide lumen, loosely packed vessel hyphae in the central portion. Both were also characterized with a central cavity. Based on this, M. incrassata seem to be anatomically similar to mycelial cords of S. lacrymans as described by Hornung and Jennings (1981).

2. MATERIAL AND METHODS

2.1 Organisms and culture conditions

Meruliporia incrassata strain FP-150521 was obtained from the culture collection of the Forest Products Laboratory, Forest Service, USDA, Madison, WI, USA. This was cultured on 2% (w/v) malt marmite agar (20g malt extract, 1g marmite in 1000 mL distilled water, solidified with 2% (w/v) agar), since it was the only medium that could produce mycelial cords (not rhizomorphs) in the fungus in glass storage dishes. Fresh field-collected rhizomorphs of M. incrassata were obtained from residences in Pasadena, CA, by Luis De La Cruz Wood Preservation Services, Pasadena, CA, USA and shipped to our laboratory. They were thoroughly cleaned with distilled water and subsequently treated for ultrastructural studies, using scanning electron microscopy. Samples that were not used immediately were stored at -21 C and -70 C: samples stored at -21 C were chemically fixed for SEM studies, whereas those stored at -70 C were used for molecular identification and confirmation of field-collected M. incrassata. To confirm the identity of M. incrassata, molecular identification of both field-collected rhizomorphs acquired from California and cultured mycelial cords of strain FP-150521 was carried out. ITS regions of these fungi were PCR amplified using the primer combination ITS F and ITS 4b (Gardes and Bruns, 1993), cycle sequenced using Big Dye® (Version 3.1, Applied Biosystems, USA), and sequenced on ABI 3730x1 Capillary DNA Sequencer (Applied

Biosystems, USA). The resulting ITS sequences were identified with BLAST searches in NCBI’s GenBank, where a number of ITS sequences of dry rot fungi have been deposited. The ITS of field-collected rhizomorphs and cultured mycelial cords of strain FP-150521 were deposited into GenBank with Accession numbers EU429518 and EU429519, respectively. Additionally, a voucher specimen of the field-collected rhizomorphs of M. incrassata has been deposited at the Willard Sherman Turrell Herbarium of the Miami University, Oxford, Ohio, USA. Armillaria mellea strain DR-140 used in all experiments was obtained from Dr. Dana Richter (School of Forest Resources and Environmental Sciences, Michigan Technological University, USA). This strain has a unique characteristic: it readily forms rhizomorphs when cultured on potato dextrose agar (PDA; Difco, Becton, Dickinson & Company, MD, USA) and in potato dextrose broth (PDB; Difco, Becton, Dickinson & Company, MD, USA). A. mellea strain was maintained on PDA slants, but cultured on PDA in Petri dish covered with cellophane, or in round-bottomed borosilicate glass tubes (Kimax, USA) containing PDB. Both cultures were incubated in the dark throughout the experiments.

2.2 Media preparation

A quantity of 19.5 g of PDA mixture was dissolved in 500 mL distilled water in 1000 mL Erlenmeyer flask, autoclaved at 1.1 kg/cm2 at 121 C for 15 min and poured in disposable Petri dishes prior to inoculation. Potato dextrose broth was prepared by dissolving a quantity of 2.4 g in 100 mL distilled water, after which 10 mL of aliquots were dispensed into round-bottomed borosilicate glass tubes before autoclaving at 1.1 kg/cm2 at 121 C for 15 min.

2.3 Armillaria mellea rhizomorph observation under light microscope

Potato dextrose agar media without sterile cellophane covering the surface were inoculated with A. mellea inoculum blocks, and incubated in the dark at room temperature until rhizomorph began developing and extending. Extending tips of A. mellea rhizomorphs were

22 observed with an inverted microscope (OLYMPUS IX70-S8F, Olympus Optical Co. Ltd., Japan). The tip of an extending rhizomorph was brought into view and focused at a magnification of 10X. This was viewed on a computer monitor with a high performance digital 12 bit CCD video camera systems (PixelFly camera; Cooke Corporation, Romulus, MI) connected to the microscope. Sequences of still images of the extending rhizomorph tip were captured with CamWare software (Version 2.10, pco.imaging, Germany), and saved in 16 bit TIFF files. Calibration was done with stage micrometer at 10X magnification, where 1 unit of the stage micrometer was equal to 10 μm.

2.4 Rhizomorph culture for scanning electron microscopy

PDA in Petri dishes was covered with sterile 9 cm diameter circles of cellophane (Innovia Films, GA, USA) and inoculated with A. mellea mycelium from stock plates. The culture plates were incubated in the dark at room temperature until matured rhizomorphs were carefully excised and immediately placed in fixative. After aliquots of PDB in round-bottomed borosilicate glass tubes had been autoclaved and cooled, they were inoculated with A. mellea inoculum blocks and incubated in the dark at room temperature until rhizomorphs formed. Mature rhizomorphs in cultures were carefully removed from the tubes with forceps, rinsed with sterile distilled water, and immediately placed in fixative.

2.5 Fixation of field-collected Meruliporia incrassata rhizomorphs

Fresh field-collected rhizomorphs were thoroughly cleaned with distilled water and sectioned (longitudinal and transverse sections), after which they were treated by fixation and dehydration for analysis. The sectioned rhizomorph samples (approximately 6 mm x 6 mm on each side) were put in 2.5% (v/v) glutaraldehyde, 1% formaldehyde (v/v) in 0.05 M sodium cacodylate buffer (pH 7.2) overnight. The samples were then rinsed with 0.05 M sodium cacodylate buffer (pH 7.2) four times at intervals of 30 min. They were then put in 1% (w/v)

osmium tetroxide for 6 hrs at room temperature, and thereafter rinsed with distilled water four times at intervals of 15 min. The samples thus treated were then serially dehydrated with different percentages of ethanol at intervals of 15 min, and with 100% ethanol for 30 min. After dehydration, these samples were critically point dried, gold plated to approximately 20 nm after 90 sec, and observed for SEM imaging, using a JOEL JSM-840A scanning electron microscope (operating voltage, 15 kV; working distance, 39). Scanning electron micrographs were obtained showing details of the rhizomorph structure.

2.6 Fixation of Armillaria mellea rhizomorphs

Fresh rhizomorphs of A. mellea cultured in PDB and aerial rhizomorphs on cellophane covering PDA were excised and immediately transferred into 2.5% (v/v) glutaraldehyde, 1% (v/v) formaldehyde in 0.05 M (v/w) sodium cacodylate buffer (pH 7.2) for approximately two hours at room temperature. The samples were then rinsed with 0.05 M sodium cacodylate buffer (pH 7.2) three times at room temperature at intervals of 10 min. The samples thus treated were then serially dehydrated with different percentages of ethanol at intervals of 15 min, and with 100% ethanol for 30 min. During the first stage of dehydration with 100% ethanol, the samples were plunged into liquid nitrogen to freeze, after which a clean pre-cooled razor blade was used to press the samples on a fracturing plate to fracture the samples. The fractured samples were then dehydrated in 100% ethanol two more times, after which they were critically point dried, gold plated and studied for SEM imaging, using a JOEL JSM-840A scanning electron microscope (operating voltage, 15 kV; working distance, 39). Scanning electron micrographs were obtained showing details of the rhizomorph structure.

3. RESULTS

3.1 Structure of Armillaria mellea observed under light microscope

Extending rhizomorph tips of A. mellea in PDA observed under light microscopes were captured (Fig 2.1). Images showed an outer clear region that is composed entirely of peripheral hyphae forming the outline of the rhizomorph. This region is characterized by outliers of individual hyphae that extend randomly into the agar media. Within the cortical region is an inner dark region of central cavity surrounded by longitudinal hyphae.

3.2 Structure of field-collected Meruliporia incrassata rhizomorphs using scanning electron microscopy

Field-collected M. incrassata rhizomorphs (Fig 2.2) were promptly sectioned, fixed, critically point-dried, and gold-plated for observation. Remarkable scanning electron micrographs were obtained, revealing detailed structure of M. incrassata rhizomorphs. It was shown that M. incrassata rhizomorphs were constructed from millions of hyphae (Fig 2.3). For example, if a rhizomorph of diameter 1 cm (0.01 m) is considered (Fig 2.2), its cross-sectional area (πr2) will be 7.9 x 10-5 mm2. Assuming the diameter of each hypha in the rhizomorph in Fig 2.3 is 3 µm (3 x 10-6 m), it implies that the cross sectional area of each hyphae is 7.1 x 10-12 m2. We therefore deduced that about 11.2 million hyphae (7.9 x 10-5 mm2 ÷ 7.1 x 10-12 m2) will aggregate to form the rhizomorph under consideration. We deduced further that there would be a degeneration of about 1.12 million hyphae, representing 10% of total number of hyphae, to form a central cavity within the rhizomorph. Longitudinal sections of M. incrassata rhizomorphs revealed the differential arrangement of hyphae of the organ: hyphae on the exterior were thick-walled, tightly packed and longitudinally arranged (Figs 2.4 A and B), while those in the interior were loosely packed with no evidence of a conspicuous hollow central cavity (Fig 2.4 C). Interestingly, the presence of longitudinal vessel hyphae is well pronounced in M. incrassata. It is believed that through vessel hyphae, nutrients, osmolytes, water and, in some

cases, oxygen are translocated to the growing tip of the rhizomorphs (Fig 2.5). As delicate as they are, vessel hyphae are surrounded by narrow, thick-walled, longitudinal hyphae filled with cellular contents (Fig 2.8), protecting the former from desiccation and adverse weather conditions.

3.3 Structure of Armillaria mellea rhizomorphs using scanning electron microscopy

Submerged rhizomorphs of A. mellea cultured in PDB in glass tubes (Fig 2.6), and aerial rhizomorphs on cellophane covering PDA were harvested. These were prepared for microscopic studies as described under methods above. Scanning electron micrographs were obtained. A. mellea rhizomorphs, constructed from millions of hyphae have a well organized growing tip covered with a combined layer of protective cap and mucilaginous sheath (just like roots of higher plants) that, presumably, protect them from abrasion as they push their way through soil and invade their substrates, and from desiccation (Fig 2.7). Not only was the rhizomorph tip covered with mucilaginous sheath, but the entire rhizomorph length. Transverse sections of freeze-fractured A. mellea rhizomorphs cultured in PDB showed a complex organ with distinct hyphal organization, arrangement and differentiation that reflect its translocatory function. These are described from the outside to the inside as follows: (i) an outer layer of peripheral hyphae, (ii) an inner cortical layer of radial hyphae, (iii) a medulla region, consisting of two layers of longitudinal hyphae – an outer layer of thick-walled, narrow lumen, tightly packed hyphae, and in inner layer of thin-walled, wide lumen, loosely packed hyphae usually called the vessel hyphae – and (iv) a central cavity, formed via a possible degeneration of vessel hyphae (Fig 2.8): it is thought that water, carrying dissolved ions and diverse compounds, is transported in the vessel hyphae. We chose the terms “radial” and “longitudinal” to easily identify the different hyphal types within the rhizomorphs, based on their arrangement, to take care of the usual confusion that is normally associated with their description. The arrangement of hyphae in A. mellea rhizomorph contrasted with M. incrassata: (i) radially arranged hyphae and peripheral hyphae that form the outer cortex in A. mellea were absent in M. incrassata, and (ii) the hollow central cavity prominent in A. mellea was absent in

M. incrassata. In M. incrassata rhizomorphs, the central portion consisted of loosely-packed hyphae with no conspicuous central cavity (Fig 2.4). As in M. incrassata, thin-walled vessel hyphae were prominent also in A. mellea (Figs 2.5 and 2.9) and were, in many cases, closely associated with an outer protective fortress of longitudinal hyphae in the medulla region formed around them (Fig 2.10).

4. DISCUSSION

The extent to which hyphae are organized and differentiated varies between species (Butler 1966). Data available on the different types of hyphae formed during mycelial cord and rhizomorph development in S. lacrymans and A. mellea indicate that sizes of constituent hyphae of these structures vary. For example, Granlund et al. (1984) reported that the diameter of the hyphae constituting the various tissues in A. mellea increased towards the center of the rhizomorph. They recorded 2.2 µm, 2.3 µm, and 13.9 µm as the mean diameter of peripheral hyphae, cortex hyphae and medulla hyphae of the rhizomorph, respectively. In addition, the distance between septa of hyphae increased towards the center. While a distance of 20 µm between septa was recorded in the cortex, 70 µm was recorded in the medulla. These data support observations from our micrographs, which indicate that indeed, the diameter of hyphal lumen of the various hyphal types increases from the periphery to the vessel hyphae in the central region of the rhizomorph (Fig 2.8). The arrangement of hyphae in this fashion is of importance to the translocatory needs of rhizomorph-forming fungi, since an aseptate lumen is required for mass translocation of nutrients and water within the vessel hyphae towards the rhizomorph tip, with less expenditure of energy in the form of the application of pressure. This may also suggest why the vessel hyphae are thin- walled to allow easy passage of water and osmolytes across the wall surface. Eamus et al. (1985) provided evidence that supports the view that long-distance translocation in mycelial cord and rhizomorphs occurred predominantly by the movement of solution along the vessel hyphae. We inferred from this that unlike thin-walled vessel hyphae, thick-walled medulla longitudinal hyphae render them with the required strength to withstand internal turgor pressure created by their cytoplasmic contents, causing them to move towards rhizomorph tip. We believe that this is

27 one of the main sources for rhizomorph tip extension that we have observed in vitro. We have provided data and discussed the role of internal turgor pressure and force exertion by rhizomorphs tips in Chapters 4 and 5. From the micrographs, vessel hyphae in both M. incrassata and A. mellea showed the widest lumen, usually with no visible plate-like “cellular partition” and no cytoplasmic contents in sight (Figs 2.5 and 2.9). Additionally, these vessel hyphae appeared wrinkled, especially in A. mellea rhizomorphs (Fig 2.9). In the medulla longitudinal hyphae that surround the vessel hyphae, cellular partitions were quite common (Fig 2.10). The certainty of these cellular partitions as septa could not be readily ascertained. However, these partitions could also be cellular contents within the hyphae that were at various stages of dehydration and crystallization at the time the rhizomorphs samples were processed for SEM. In some of the medulla hyphae, where these cellular partitions were absent, cell content was almost always present (Fig 2.8). Jennings and Watkinson (1982) compared initial stages of mycelial cord development of S. lacrymans in culture with the structure of mature strands from infected wood, and reported that cultures with mycelial cords stained with 1% (v/w) Nile Blue at early stages of development had three types of differentiated hyphae: 1) approximately 10 µm diameter wide, empty aseptate hyphae that stained evenly blue; 2) non-stained, septate hyphae with visible cytoplasmic contents and 3) narrow and septate unbranched tendril hyphae with dark-stained dense cytoplasm of about 2 µm diameter. Meanwhile, a comparative transmission electron microscopy (TEM) and SEM study of mature mycelial cords (obtained from an actively growing colony in a house) by Jennings and Watkinson (1982) revealed three distinguishable regions: an outer surface covered by colorless thin-walled hyphae growing separately from one another; a zone lying beneath the outer surface characterized with by a few hyphae that were cracked with holes, and a central portion characterized with longitudinal channels. TEM work indicated that hyphae constituting the outer surface were thick-walled with narrow lumen, and were found only in mature cords. These hyphae were characterized by three-layers of walls. In other words, the outer hyphae had layers of walls that were deposited inside the original wall. In the central part, hyphae with dense cytoplasmic contents were found. The hyphal walls were much thinner compared to those on the outer surface, but these walls were also deposited in three layers. In our study, three-layered hyphal wall types were not found in any of the hyphal types that constituted M. incrassata or A. mellea rhizomorphs. However, it was interesting that the general description of the mycelial cord

of S. lacrymans precisely fits the structure of M. incrassata rhizomorphs examined in our study. For example, M. incrassata showed an outer layer of longitudinally arranged thick-walled narrow lumen hyphae (Fig 2.4) that surrounded a group of loosely-packed central hyphae, some of which were identified as vessel hyphae with thin walls and wide lumen. One remarkable observation was that the transition of hyphal types from the periphery to the central part of M. incrassata rhizomorph was not as distinct as compared to A. mellea rhizomorph, where there was a distinct organization of hyphal types from the periphery to the central part. Moreover, no hyphal type was identified to be characterized by cracked holes in M. incrassata or A. mellea as observed by Jennings and Watkinson (1982) in S. lacrymans. From our micrographs and previous data available, it seems that a decrease in the thickness of hyphal walls and increase in distance between septa pertaining to the different hyphal types from the outside of the rhizomorph to the central region plays a very significant and concerted role in the functions of mycelial cords and rhizomorphs. Micrographs of M. incrassata and A. mellea showed that both have tightly packed, thick- walled, longitudinally arranged fiber hyphae surrounding the inner region characterized by thin- walled, wide lumen vessel hyphae. However, there were some structural differences between their rhizomorphs. Whereas an outer peripheral hyphae and a central cavity were absent in M. incrassata rhizomorph, they were conspicuously present in A. mellea (Figs 2.4 and 2.8). Interestingly, the occurrence of the radial hyphae forming the cortex of the rhizomorph was observed in mature regions of A. mellea and not the tip (Figs 2.7 and 2.8). This supports the description of in A. mellea rhizomorphs by Granlund et al. (1984). In M. incrassata, however, there were only tightly packed, longitudinally arranged hyphae with no peripheral hyphal covering. Their absence in M. incrassata rhizomorphs may be attributed to the following: (i) they may have been lost in the process of rhizomorph removal from the infested home; (ii) rhizomorphs used in the study may have been immature; (iii) migration through compact soil may have deprived rhizomorphs of peripheral hyphae, or (iv) they are simply not formed at all. According to Granlund et al. (1984), peripheral hyphae could serve as infective hyphae such that they could be used as a means by which A. mellea makes contact and invades new plant hosts. They argued that their absence in the rhizomorph tip, which is constantly supplied with nutrients and water to support its active tip growth, may suggest that peripheral hyphae may also serve absorptive roles for the rhizomorphs. As the tip extends, the peripheral hyphae, which

provides a large surface area in the mature region, may have the potential to laterally absorb nutrient and water from the immediate surroundings of the rhizomorph into its interior. The dissolved nutrients and water are thus transported into the inner medulla region for translocation to the tip. It suffices to say that with such intake of nutrients and water, and the accumulation of osmolytes, internal turgor pressure is created within the rhizomorph. Some of this turgor pressure is eventually used as the driving force for the growing tip to migrate and penetrate through hard, compact substrates. In an attempt to understand the structure and function of hyphal differentiation in M. incrassata rhizomorph, our study with mainly A. mellea and data from studies with S. lacrymans has provided more information on the detailed features of rhizomorph anatomy that support their migratory, translocatory and invasive behavior. It is evident from our study that M. incrassata rhizomorphs morphologically appear more similar to S. lacrymans than to A. mellea, which is more complex. We concluded from our study that, first, two hyphal types are prominent in the structure and function of rhizomorphs in both M. incrassata and A. mellea: the thick-walled, narrow lumen, longitudinally arranged hyphae, and inner thin-walled, wide lumen vessel hyphae. Secondly, whereas M. incrassata rhizomorph does not have a conspicuous central cavity, A. mellea is characterized with a central cavity believed to be formed as a result of the degeneration in vessel hyphae, a view supported by Hartig and van Tubeuf (1902) and Falck (1912). Third, A. mellea rhizomorphs indeed have well organized tips. As a result, these tip-growing organs have the potentials that make them an essential part of rhizomorph-producing fungi’s successful explorations and survival in their environments. Not only did we confirm findings by Granlund et al. (1984) on the structure of A. mellea rhizomorphs, but we were also able to provide better micrographs, and revised the terminology used to identify the different hyphal types, based on their arrangement, to take care of the usual confusion that is normally associated with them. We suggest that this study be expanded to investigate if M. incrassata has organized tip growth, which to our knowledge, has never been reported in literature. Moreover, investigating the role of mechanics in rhizomorph construction is one area that needs urgent attention to provide insight into factors that are involved in the initiation and sustenance of rhizomorph development and extension.

REFERENCES

Brownlee C, Jennings DH, 1981a. Further observations on tear or drop formation by mycelium of Serpula lacrymans. Transactions of the British Mycological Society 77: 33-40.

Brownlee C, Jennings DH, 1981b. The content of soluble carbohydrates and their translocation in mycelium of Serpula lacrymans. Transactions of the British Mycological Society 77: 615-619.

Burdsall HH, 1991. Meruliporia (Poria) incrassata: Occurrence and significance in the United States as a dry rot fungus. In: Jennings DH, Bravery AF (eds), Serpula lacrymans: Fundamental Biology and Control Strategies. John Wiley & Sons, Chichester, pp. 189- 191.

Butler GM, 1966. Vegetative structures in the fungi. In: Ainsworth GC, Sussman AS (eds), The Fungi. Academic Press, London, U.K.

Eamus D, Thompson W, Cairney JWG, Jennings DH, 1985. Internal structure and hydraulic conductivity of basidiomycete translocating organs. Journal of Experimental Biology 36: 1110-1116.

Falck R, 1912. Die Merulius faule des Bauholzes. Hausschwammforschung 6: 1-405.

Gardes M, Bruns TD, 1993. ITS primers with enhanced specificity for basidiomycetes: application to the identification of mycorrhizae and rusts. Molecular Ecology 2:113-118.

Garrett SD, 1981. Soil and Soil Fertility: An Introduction to Soil Mycology, second edn. Pergamon Press, Oxford.

Granland HI, Jennings DH, Veltkamp K, 1984. Scanning electron microscope studies of rhizomorphs of Armillaria mellea. Nova Hedwigia 39: 85-100.

Granlund HI, Jennings DH, Thompson W, 1985. Translocation of solutes along rhizomorphs of Armillaria mellea. Transactions of the British Mycological Society 84: 111-119.

Hartig R, 1874. Wichtige Krankheiten der Waldbäume. Beiträge zur mycologie und Phytopathologie für Botaniker und Forstmänner. Berlin: Springer. 127 p. [Important Diseases of Forest Trees. Contributions to Mycology and Phytopathology for Botanists and Foresters. Phytopathological Classics No. 12; 1975. St. Paul, MN: American Phytopathological Society.]

Hartig R, van Tubeuf CF, 1902. Der echte Hausschwamm und andere das Bauholz zerstörende Pilze. Berlin: Springer-Verlag.

Hornung U, Jennings DH, 1981. Light and electron microscopical observations of surface mycelium of Serpula lacrymans: Stages of growth and hyphal nomenclature. Nova Hedwigia 34: 101-126.

Jennings L, Watkinson SC, 1982. Structure and development of mycelial strands in Serpula lacrymans. Transactions of the British Mycological Society 78: 465-474.

Lindahl BD, Olsson S, 2004. Fungal translocation – creating and responding to environmental heterogeneity. Mycologist 18: 79-88.

Schmidt O, Moreth U, 2002, Data bank of rDNA-ITS sequences from building rot fungi for their identification. Wood Science and Technology 36: 429-433.

Townsend BB, 1954. Morphology and development of fungal rhizomorphs. Transactions of the British Mycological Society 37: 222-232.

Fig 2.1 – Apical region of rhizomorph a of Armillaria mellea growing in agar medium, showing outliers of individual hyphae extending randomly into PDA media. The dark inner region is composed of longitudinal hyphae.

Fig 2.2 – Field-collected rhizomorphs of Meruliporia incrassata from an infested home in California. Note the root-like, woody appearance. Specimen provided courtesy of Luis De La Cruz Wood Preservation Services, Pasadena, CA, USA.

Fig 2.3 – Scanning electron micrograph of field-collected Meruliporia incrassata rhizomorph showing constituent hyphae lying parallel to one another. Scale bar = 10 µm

Fig 2.4 – (A) Scanning electron micrograph showing longitudinal hyphae of Meruliporia incrassata rhizomorph (arrow). Scale bar = 10 µm. (B) Longitudinal hyphae are tightly packed, protecting the inner layers of hyphae in the middle portion of the organ. Scale bar = 10 µm. (C) Middle portion of loosely packed hyphae, m. Scale bar = 100 µm.

Fig 2.5 – (A) A vessel hypha of Meruliporia incrassata with thin wall and wide diameter lumen (arrows) in close association with tightly packed regular hyphae (*). Scale bar = 10 µm. (B) Vessel hyphae embedded in and surrounded by hyphae and soil particles. Scale bar = 10 µm. (C) Vessel hyphae with remnants of degenerated internal wall components. Scale bar = 10 µm.

Fig 2.6 – Armillaria mellea rhizomorphs (r) growing in potato dextrose broth media, with mycelium (m) on medium surface. Scale bar = 20 mm.

Fig 2.7 – Scanning electron micrograph of rhizomorph tip of Armillaria mellea grown on cellophane covering potato dextrose agar. A protective cup of mucilage (arrow) is localized to the tip, and a mucilaginous sheath (mu) covers the entire length of the rhizomorphs. Scale bar = 100 µm.

Fig 2.8 – Scanning electron micrograph of transverse sections of freeze-fractured rhizomorph of Armillaria mellea. (A) Complete hyphal arrangement in rhizomorph. (B) A close up micrograph of hyphal arrangement From the outer layer to the inner zones are; a cortical layer of loosely-packed, radially arranged, thin hyphae (r), covered with peripheral hyphae (p); an inner medulla of tightly-packed, longitudinally arranged, thick- walled hyphae with narrow lumens (m) with cellular contents (arrows); longitudinally arranged thin-walled vessel hyphae with wide lumens (v), and a central cavity that may be formed by degeneration of vessel hyphae (c). Scale bar = 100 µm (A); Scale bar = 10 µm (B).

Fig 2.9 – Scanning electron micrographs of Armillaria mellea rhizomorph. (A) Vessel hypha (v) in close association with tightly packed longitudinal hyphae (Arrow I). Some of the vessel hyphae have wrinkled wall (*), while others have collapsing wall (Arrow J). Scale bar = 10 µm. (B) Wide lumen vessel hyphae (arrows) in close association with small lumen, thick-walled longitudinal hyphae (m) that form a rind around the vessel hyphae. Scale bar = 10 µm. (C) Wrinkled, collapsed vessel hyphae (v), protruding from freeze-fractured surface. Scale bar = 10 µm.

Fig 2.10 – Scanning electron micrograph of Armillaria mellea rhizomorph. A group of longitudinal hyphae that that surround inner thin-walled vessel hyphae. Hyphae are characterized by thick walls (Arrow J) and putative septa (arrows I). Scale bar = 10 μm.

CHAPTER 3

IN VITRO STUDIES OF RHIZOMORPH EXTENSION IN ARMILLARIA MELLEA

ABSTRACT

Mycelial and rhizomorph growth rates of Armillaria mellea were determined in vitro. Studies were conducted on the stimulatory effects of different agar concentrations on the production and extension of rhizomorphs. In vitro growth rate studies of mycelia and rhizomorphs of A. mellea were conducted with potato dextrose (PD) medium augmented with increasing agar concentrations to produce a range of medium gel strength or hardness. Results indicate that rhizomorphs of A. mellea grow faster than mycelia in PD medium, extending beyond the colony radius of the mycelia. Increasing agar concentrations stimulated the production of lengthier rhizomorphs. Still images obtained with video microscopy also showed that rhizomorphs in PD 7% agar medium produced rhizomorphs that extended faster than those in normal PD 2% agar medium. Additionally, rhizomorphs in PD 2% agar showed broad apices and multiple individual hyphae that extended from the surface of the organ, unlike rhizomorphs penetrating PD 7% agar, which showed more tapered apices, with a zone of compression in the agar evident around the apices. We concluded therefore that (i) A. mellea rhizomorphs extended faster than mycelia from which they were generated, and (ii) the rate of rhizomorph extension was faster in media with higher concentrations of agar.

1. INTRODUCTION

Growth and extension of mycelia is of immense importance to the spread and survival of fungi. In search of nutrients in the environment, fungi produce mycelia from the source of their inoculum, and spread as the nutrient base in the colonized substrate is depleted. Fungal rhizomorphs and mycelial cords, on the other hand, are specially constructed structures produced by mostly wood-decay and forest-inhabiting pathogenic fungi. They (i) translocate resources (nutrients, water and oxygen), (ii) allow fungi to migrate from one substrate to another usually over non-nutrient substrates, and (iii) enhance inoculum potential (Garraway et al. 1991). Thus, rhizomorphs are important in the spread of basidiomycete fungi including Armillaria mellea, Marasmius brevipes, Meruliporia incrassata, and Serpula lacrymans. Armillaria generally has a slow growth rate in soil (Rishbeth, 1968), and it is known to cause diseases in many trees by first making contact with healthy tissues through rhizomorphs or mycelium (Redfern 1975; Rykowski, 1984). Hartig (1874) and Nechleba (1915) reported the gravity of Armillaria disease in hardwood plantations that had been replaced by conifers. These reports generated interesting debates on the superiority of substrate quality in disease development between hardwood and conifer species. Contrary to the assertion that hardwood species served as better substrates than conifers, Ono (1965, 1970) reported the occurrence of severe disease in coniferous stump substrates, with other reports showing that conifer stumps are effective inoculum sources for infections of surrounding trees (Filip 1979; Roth et al. 1980). Additionally, others variously reported the production of rhizomorphs among hardwood and conifer stumps (Redfern 1975; Rykowski, 1984; Benjamin and Newhook, 1984; Pearce and Malajczuk, 1990). Factors that affect the growth of rhizomorphs have been studied, and it has been determined that growth of rhizomorphs varies among species with environmental conditions playing immense role (Morrison, 1976, 1982; Rishbeth, 1982). It has been demonstrated that pH, aeration, temperature, moisture, organic matter, light, etc. have an effect on rhizomorph growth and development. Of these, moisture level and substrate temperature are the most extensively studied. For example, Pearce and Malajczuk (1990) reported that rhizomorphs of A. luteobubalina grew at faster rate on a substrate with a water potential of –0.6 MPa, with no growth at –0.001 MPa. As observed by Garrett (1944), Armillaria root diseases are common in

44 wet soil conditions. Cruickshank et al. (1997) found that saprotrophic Armillaria species are suppressed under dry conditions. Although other factors have been studied, we were mindful of our observation from preliminary experiments that rhizomorphs extended much faster than mycelial margins in agar medium. Based on this, the objectives of this study were to determine the growth rate of both Armillaria mycelium and rhizomorphs in vitro; to study the stimulatory effects of different agar concentrations on the production of rhizomorphs, and to compare the rate of rhizomorph extension in different agar concentrations. In vitro growth rate studies of mycelia and rhizomorphs of A. mellea were conducted with potato dextrose medium solidified with different concentrations of agar (PD 2 - 8% agar). Results indicate that rhizomorphs of A. mellea grow faster than mycelia in PDA medium; although developing much later after mycelial growth has been initiated, the rhizomorphs extend beyond the colony radius of the mycelia within a few days. Additionally, PD5 - 8% agar produced rhizomorphs that extended much faster than those in PD 2% agar. Interestingly, increasing agar concentrations increased the extension rate of individual rhizomorphs and stimulated the production of lengthier rhizomorphs.

2. MATERIALS AND METHODS

2.1 Organism and culture conditions

Armillaria mellea strain DR-140 used in all experiments was obtained from Dr. Dana Richter (School of Forest Resources and Environmental Sciences, Michigan Technological University, USA). This strain has a unique characteristic: it readily forms rhizomorphs when cultured on PDA, and in PDB (Difco, Becton, Dickinson & Company, MD, USA). A. mellea strain was maintained on PDA slants, but cultured on PDA in Petri dish in the dark during throughout the experiments.

2.2 Media preparation and gel strength determination

A quantity of 19.5 g of PDA mixture was dissolved in 1 L distilled water in 1000 mL Ehrlenmeyer flask, autoclaved at 1.1 kg/cm2 at 121 C for 15 min and poured in disposable Petri dishes prior to inoculation. Separate quantities of 0.5 g and 2 g of agar, and 2.4 g dried PDB medium were dissolved in 100 mL distilled water in a 500 mL Erlenmeyer flask to prepare PD 0.5% and PD 2% agar media, respectively, after which they were autoclaved at 1.1 kg/cm2 at 121 C for 15 min and poured in disposable Petri dishes prior to inoculation. In the case of PD 5% (w/v) agar medium, 5 g of agar and 2.4 g PDB was dissolved in 100 mL distilled water in a 500 mL Erlenmeyer flask. The medium was covered with a metal cap and heated on a magnetic stirring hot plate (Barnstead/Thermolyne Cimarec® 2, Model # SP46925, Iowa, USA) until the agar was melted and well mixed, after which it was immediately poured in Petri dishes to solidify, prior to inoculation. The same method was used to prepare PD 7% agar and PD 8% agar media, using 7 g and 8 g of agar, respectively. Medium gel strength was determined as described by Brush et al. (1999) (Fig 3.1).

2.3 Mycelial growth and rhizomorph extension rate in PDA

Aliquots of 20 mL molten sterile PDA medium were poured into 9 cm diameter sterile Petri dishes and allowed to solidify. The dishes were then inverted and two diameters at right angles to one another were drawn at the bottom of the Petri dishes with an ultra fine point permanent marker. The dishes were turned over and inoculum blocks of A. mellea were carefully placed at the intersections of the two diameters under sterile conditions, with the culture-bearing surface directly in contact with the fresh medium. The Petri dishes were incubated in the dark at room temperature, and monitored daily for mycelial growth and rhizomorph initiation and extension under the microscope. The growth rate of mycelium was measured every 24 h after an observation of initial mycelial growth by marking the extending periphery of the mycelium along the two diameters drawn on the base of the Petri dish; the actual rate of extension of the mycelium was measured

46 with the eyepiece micrometer (ocular graticule) at 0.7x/2x/10x magnification under a dissecting microscope (Olympus SZH10-ILLD, Olympus Optical Co. Ltd. Japan). The extension rate of randomly selected rhizomorphs initiated close to the inoculum block was also measured by the same method, but since the rhizomorphs extended in haphazard fashion, the measurements were taken along their path in the medium.

2.4 Armillaria mellea rhizomorph growth in media with different concentrations of agar

Armillaria mellea was cultured in Petri dishes containing PDA with different agar concentrations (0.5%, 2.0%, 5.0% and 8.0%; all in w/v) with four replicates per treatment. Four- week old cultures incubated in the dark at room temperature were used to determine the extent of rhizomorph growth. Rhizomorphs of A. mellea cultures were measured by first scanning the bottom of the Petri dishes using an Epson 4870 PHOTO scanner. Using Image-Pro Plus 4.5 imaging software (Media Cybernetics, MA, USA), the lengths of individual rhizomorphs were measured. All individual measurements in millimeters were automatically generated in Microsoft Excel 2003 for retrieval and analyses. The mean total length of the replicates for each treatment of the different agar concentrations was computed.

2.5 Video microscopy of rhizomorph extension in PD 2% and PD 7% agar

The observation that, increasing concentrations of agar stimulated faster growth and development of rhizomorph prompted the idea to observe and compare the movement of rhizomorphs in PD 2% agar and PD 7% agar medium. Petri dishes of A. mellea were observed with an inverted microscope (OLYMPUS IX70-S8F, Olympus Optical Co. Ltd., Japan). The tip of an extending rhizomorph was brought into view and focused at a magnification of 10x. This was also viewed on a computer monitor with a high performance digital 12 bit CCD video camera system (PixelFly camera; Cooke Corporation, Romulus, MI) connected to the microscope. Sequences of still images of the extending rhizomorph tip were captured at 10 min intervals with CamWare software (Version 2.10, pco.imaging, Germany), and saved in 16 bit

TIFF files. Measurements were done with eyepiece micrometer at 100x magnification, where 1 eyepiece unit was equal to 10 µm. Images were processed using Corel Photo-Paint® x3 (Version 13.0.0.739, Corel Corporation, USA)

3. RESULTS

3.1 Mycelial growth and rhizomorph extension rate in PDA

Initiation of mycelial growth in A. mellea occurred in most cultures after 2 d of incubation. Markings on the base of the Petri dishes made measurement easy; the colony radius of the growing mycelia was measured until the extending rhizomorphs on the periphery of the mycelia interfered with any meaningful measurement of the colony radius. Measurements were terminated whenever there was such interference. Additionally, rhizomorph measurement was terminated whenever the organs touched the sides of the Petri dishes and extended out of the media to become aerial rhizomorphs (at which point the rhizomorphs turn brown and stop extending). Generally, rhizomorph initiation and development in most cultures occurred between 6 and 8 d, after which extension increased rapidly to outpace the extending hyphae (Fig 3.2). It was observed that colony radius continued to increase, but daily extension rate was remarkably reduced, especially as more rhizomorphs were initiated in culture, and those that had already formed continued to extend in length. The observation was confirmed by the data obtained, where rhizomorph extension rate was four fold that of hyphal growth at the end of day 11 (Fig 3.2). This may suggest an incremental channeling and investment of resources by the fungus towards mass production of hyphae for the subsequent initiation and production of rhizomorphs, as the nutrients in the media are depleted, and the fungus searches for other potential sources of food.

3.2 Armillaria mellea rhizomorph growth in media with different concentrations of agar

The observation that rhizomorphs extended faster than the hyphal mycelium in PDA prompted analysis of the effect of increasing agar concentration (and gel strength) on the extension rate and total length of rhizomorphs produced under such conditions. Increasing the agar concentration in the media provided conditions that mimicked those of soil conditions, and decaying wood substrates encountered by A. mellea in its natural environments. Potato dextrose medium augmented with 0.5%, 5%, and 8% agar (w/v) were used as the other treatments for this study. The lengths of all rhizomorphs in the 4-week old cultures were measured in all the replicates for each treatment, after which the average total lengths were calculated. It was observed that uniform mycelial growth occurred in both PD 0.5% agar and PD 2% agar media until initiation of rhizomorph development, which occurred between 6 and 8 d, after incubation. Also, the mycelial mat was more pronounced in these two media than in PD 5% agar and PD 8% agar media. In some cases, no growth or very little growth of mycelium was observed in PD 5% agar and PD 8% agar media before initiation of rhizomorphs. Once initiated, rhizomorph extended in all media in a fashion similar to that observed in the previous experiment (Fig 3.2) with the exception of PD 0.5% agar media, where there was lower extension rate of the rhizomorphs. Measurement of rhizomorph length produced in the various treatments revealed that PD 0.5% agar media produced the least average total length of rhizomorphs (651 ± 22 mm; n = 6) compared to that produced in PD 8% agar media (3166 ± 237 mm; n = 6) (Table 3.1; Figs 3.3, 3.4). The results obtained indicate that increasing agar concentration indeed resulted in increased number of rhizomorphs produced, although there was a slight decrease in the mean total length of rhizomorphs produced in PD 5% agar media, which may be due to variability in the conditions of the culture (Fig 3.4). Overall, the results suggest that increasing gel strength in media may stimulate faster growth, extension and greater production of rhizomorph in A. mellea.

3.3 Video microscopy of rhizomorph extension in PD 2% and 7% agar

Since it had been established that rhizomorphs extended faster than mycelia and, most importantly, increasing concentration of agar impacted the production of rhizomorphs in A.

49 mellea, this experiment was designed to study the extension of individual rhizomorphs in PD 2% agar and PD 7% agar media. Still images of extending rhizomorphs in these two growth media were captured at 10 min interval, using CamWare software, (see Materials and Methods), after which their respective rates of extension were compared. The comparison shows that rhizomorphs in PD 7% agar extended faster and farther than rhizomorphs in PD 2% agar after 60 min of measurement (Fig 3.5). This observation confirms our data that rhizomorphs are indeed stimulated to grow and extend faster (thus, producing lengthier rhizomorphs) by media augmented with higher agar concentrations (Figs 3.3 and 3.4). One significant observation made from the still images is that, whereas the rhizomorphs in PD 2% agar had broad apices and multiple individual hyphae extending from the surface of the organ, rhizomorphs penetrating PD 7% agar were more compact, showing extreme tapering of their apices, with compression of agar evident. Because the tip is compressed by a more solid media, few individual hyphae are observed to extend from the main body of the rhizomorph into the PD 7% agar media, unlike in PD 2% agar media.

4. DISCUSSION

The development of rhizomorphs in PD 0.5% agar (w/v), our medium with lowest concentration of agar, and other PD agar media (2%, 5%, 7% and 8%) used in the experiments may suggests the innate ability of A. mellea to produce rhizomorphs as a characteristic vital to the survival of the fungus in its environment. Results obtained in the first experiment were used as a guide in the design of subsequent experiments. Since rhizomorphs are used to translocate resources such as nutrients, water and oxygen, to migrate from one substrate to another, and to enhance inoculum potential (Garraway et al. 1991), they allow A. mellea to explore and colonize other woody substrates, both dead and alive, in the immediate vicinity or farther away from the depleted substrates they colonize. From in vitro experiments conducted with A. mellea, both mycelial growth, and rhizomorph formation and extension occurred in all the media used, although there was a reduction in mycelial mat production in media with increasing concentrations of agar (data not shown). This seems to imply that the easier it is for A. mellea to colonize and utilize nutrients

available in a soft substrate, the more mycelia it produces to spread from one food source to another, as opposed to the massive rhizomorph production. On the other hand, the production of more rhizomorphs in high concentrations of agar media suggest that the degree of hardness of a woody substrate may be a critical factor in the determination of (i) the suitability of a substrate as an effective nutrient source, (ii) the ability of Armillaria to make a “choice” between production of just mycelium alone, or rhizomorphs as it attempts to colonize its substrate, and (iii) the tendency to produce more rhizomorphs to explore the immediate surroundings for other suitable sources of nutrients. As shown in Fig 3.2, mycelial growth had been initiated before the formation of rhizomorphs, and yet the rhizomorphs, once produced, extended faster and radially outpaced the mycelia colony over time. This seems to suggest the ability of A. mellea to channel resources for the massive production of mycelia from hyphae to form rhizomorphs once nutrients in the substrate are being depleted. Data from the experiments also revealed that rhizomorph length increased with increasing concentration of agar (Fig 3.4). For example, in PD 8% agar (w/v), the average total length of rhizomorphs produced in four 90 mm Petri dishes was recorded to be a staggering 3166 mm (over 3 meters in 4 weeks), compared to 651 mm in PD 0.5% agar (w/v) (Table 3.1). This supports earlier reports that hardwood species inoculated with Armillaria isolates produced lengthier and greater dry weight of rhizomorphs than coniferous species (Morrison, 1972). Rishbeth (1972) also showed that English oak was a superior substrate for Armillaria than pine, since the former produced more rhizomorphs, both in number and in biomass. That lengthier rhizomorphs are produced in hardwoods than in pines supports our data that show rhizomorphs in higher concentrations of agar extending faster and further than those in lower concentrations of agar (Fig 3.5). This suffices to say that the faster rhizomorphs extend, the lengthier they get, which subsequently produces a larger rhizomorph biomass. Careful observation of extending rhizomorphs in PD 2% agar (w/v) and PD 7% agar (w/v), with respective gel strength of 0.01 MPa and 0.08 MPa, indicate that rhizomorphs in the latter were more compact, with tapering apices than those in the former (Fig 3.5). The tapering apices of rhizomorphs in PD 7% agar may be attributed to the more solid nature of the medium; the harder the medium, the higher its resistance to penetration to the rhizomorph tip. This ultimately compresses the tip to an extent where it rather splits open the medium with its tapering tip, by acting like a wedge, thus increasing its acceleration in the media. Similar observations

have been reported in fungal hyphae where, during extension, the hyphal tip was characterized by a tapering apex, whereas non-extending hyphal tip appear broad (Harold et al. 1996). The fact that rhizomorphs are able to penetrate media with gel strength of 0.08 MPa underscores their uniqueness in migration through compact substrates such as soil and wood, and their importance in plant pathogen and wood decomposers. For example, Findlay (1951) reported a 9-meter growth in A. mellea rhizomorphs along a brick-lined drainage tunnel in a period of 7 years. Similarly, Shaw and Roth (1976) estimated that A. mellea expanded in a ponderosa pine field at a rate of 2 m per year, an observation that has been attributed to the development of tip-extending rhizomorphs. Experiments have shown that rhizomorph production may be higher in other wood types than in hardwood (Redfern 1975; Rykowski, 1984; Benjamin and Newhook, 1984; Pearce and Malajczuk, 1990). However, inadequate data to completely generalize the superiority of one wood type over the other makes these in vitro experiments useful. This is because data from the studies suggest that substrate hardness may not be the only overriding determining factor in the production of rhizomorphs in Armillaria, since Armillaria may be also influenced by other factors (for example, depleted of nutrient sources, low oxygen, moisture content, the amount of mycelia as inoculum source, etc.) to initiate the production of rhizomorphs. These notwithstanding, we suggest that the level of compactness of soil in itself may be a stimulatory factor on rhizomorph production and extension. In other words, the more compact a soil, the more lengthy rhizomorphs would be produced. We base this on our data which provides evidence that (i) PD 8% agar media produce lengthier rhizomorphs than media augmented with less agar concentration, (Fig 3.3) and (ii) rhizomorphs in PD 7% agar extended faster than those in PD 2% agar (Fig 3.5). We predict that this situation may be true for more compact soils, where rhizomorphs are stimulated to extend faster and spread more rapidly in the soil environment: the probability of the rhizomorphs to encounter roots of hosts for infection, and hence disease incidence, is higher. Thus, soil compactness greatly determines the success rate, either directly or indirectly, of A. mellea. From our experiments, we concluded that (i) A. mellea rhizomorphs extended faster than mycelia from which they were generated, and (ii) the rate of rhizomorph extension was faster in media with higher concentrations of agar.

The extent to which A. mellea rhizomorphs are successful is dependent on the interplay between several environmental factors, with substrate hardness serving as an important stimulatory factor which cannot be overlooked. This is evident from our comparative gel strength experiments. Based on the fact that A. mellea produces rhizomorphs irrespective of prevailing environmental conditions, we suggest that a complete molecular study be conducted to identify genes that control the initiation, growth and development of rhizomorphs. These can be extended to M. incrassata for comparative purposes, and to learn more about the role of rhizomorphs in building decay.

REFERENCES

Benjamin M, Newhook FJ, 1984. The relative susceptibility of various Eucalyptus spp. and Pinus radiata to Armillaria grown in different food bases. Proceedings of the 6th International Conference on Root and Butt Rots of Forest Trees, Melbourne, Australia: 140-147.

Brush L, Money NP, 1999. Invasive hyphal growth in Wangiella dermatitidis is induced by stab inoculation and shows dependence upon melanin biosynthesis. Fungal Genetics and Biology 28: 190-200.

Filip GM, 1979. Root disease in Douglas-fir plantations is associated with infected stumps. Plant Disease Reporter 63: 580-583.

Findlay WPK, 1951. The development of Armillaria mellea rhizomorphs in a water tunnel.

Transactions of the British Mycological Society 34: 146.

Garraway MO, Hüttermann A, Wargo PM, 1991. Ontology and physiology. In: Shaw GC, Kile GA (eds), Armillaria Root Disease. United States Department of Agriculture-Forest Service Agriculture Handbook No. 691. pp. 21-46.

Garrett SD, 1944. Root disease fungi. In: Chronica Botanica Co. Waltham, Massachusetts.

Harold RL, Money NP, Harold FM, 1996. Growth and morphogenesis in Saprolegnia ferax: Is turgor required? Protoplasma 191: 105-114.

Hartig R, 1874. Wichtige Krankheiten der Waldbäume. Beiträge zur mycologie und Phytopathologie für Botaniker und Forstmänner. Berlin: Springer.127 p. [ImportantDiseases of Forest Trees. Contributions to Mycology and Phytopathology for Botanists and Foresters. Phytopathological Classics No. 12; 1975. St. Paul, MN: American Phytopathological Society.]

Morrison DJ, 1972. Studies on the biology of Armillaria mellea. Ph.D. Thesis, University of Cambridge.

Morrison DJ, 1976.Vertical distribution of Armillaria mellea rhizomorphs in soil. Transactions of the British Mycological Society 66: 393-399.

Morrison DJ, 1982. Effects of soil organic matter on rhizomorph growth by Armillaria mellea. Transactions of the British Mycological Society 78: 201-207.

Nechleba A, 1915. Der Hallimasch: studien beobechtungen und hypothesen. [The honey agaric.] Fortwissenschaftliches Centralblatt 59: 384-392.

Ono K, 1965. Armillaria root rot in plantations of Hokkaido. Effects of topography and soil conditions on its occurrence. Meguro: Bulletin of the Government Forest Experiment Station. 179: 1-6. In Japanese. Review of Applied Mycology. 45: 635

Ono K, 1970. Effect of soil conditions on the occurrence of Armillaria root rot of Japanese larch. Meguro: Bulletin of the Government Forest Experiment Station. 229: 123-219. In Japanese. Review of Plant Pathology 50: 2001

Pearce MH, Malajczuk N, 1990. Factors affecting the growth of Armillaria luteobubalina rhizomorphs in soil. Mycological Research 94: 32-37.

Redfern DB, 1975. The influence of food base on rhizomorph growth and pathogenicity of Armillaria mellea isolates. In: Bruehl GW (ed), Biology and Control of Soil-Borne Plant Pathogens. St. Paul, MN: American Phytopathological Society: pp. 69-73.

Rishbeth J, 1972. The production of rhizomorphs by Armillaria mellea from stumps. European Journal of Forest Pathology 2: 193-205.

Rishbeth J, 1968. The growth rate of Armillaria mellea. Transactions of the British Mycological Society 51: 575-586.

Rishbeth J, 1982. Species of Armillaria in southern England. Plant Pathology 31: 9-17.

Roth LF, Rolf L, Cooley S, 1980. Identifying infected ponderosa pine stumps to reduce costs of controlling Armillaria root rot. Journal of Forestry 78: 145-151

Rykowski K, 1984. Niektóre troficzne uwarunkowania patogeniczności Armillaria mellea (Vahl) Quél. w uprawach sosnowych. [Some trophic factors in the pathogenicity of Armillaria mellea in Scotch pine plantations.] Prace Instytutu Badaweczego Leśnictwa 640: 1-140. In Polish.

Shaw CG, Roth LF, 1976. Persistence and distribution of a clone of Armillaria mellea in a ponderosa pine forest. Phytopathology 66: 1210-113.

0.12

) -2

m 0.10 μ

μ 0.08

0.06

0.04

0.02

Strength (MPa or 0.00

02468

Agar concentration (g/100 mL)

Fig 3.1 – Relationship between agar concentration and medium gel strength expressed in units of force per unit area (MPa or µN µm-2). Each data point is mean ± S.E. (n = 5).

80 70 60 50 Mycelium radius 40 Rhizomorph length 30 Length (mm) Length 20

10 0 12345678910111213 Time (d)

Fig 3.2 – Comparative growth of mycelium and rhizomorphs in Armillaria mellea cultured in PDA and incubated in the dark at room temperature over a period of 13 d ± S.E (n = 6). Rhizomorph initiation was delayed, but extension rate was faster than that of colony radius of mycelia (i.e., hyphal length), a situation that may be responsible for reducing mycelial expansion

0.5% agar

8% agar

Fig 3.3 - Effect of increasing gel strength on the production of Armillaria mellea rhizomorphs, after 4 weeks of incubation. Scale bar = 20 mm.

4000

3500

3000

2500

2000

1500 Length (mm) Length 1000

500 0

0.5% 2.0% 5.0% 8.0% Agar concentration

Fig 3.4 – Length of rhizomorphs produced in increasing concentrations of agar in PDB in 4-week old cultures of Armillaria mellea ± SE (n = 4).

0 min

10 min

20 min

30 min

40 min

50 min

60 min

2% agar 7% agar (gel strength 0.01 MPa) (gel strength 0.08 MPa)

Fig 3.5 – Effect of increasing gel strength on the extension of Armillaria mellea rhizomorphs. Rhizomorphs in 2% (w/v) agar show broad apices and multiple individual hyphae extend from the surface of the organ. Rhizomorphs penetrating 7% agar are more compact, show extreme tapering of their apices, and compression of agar is evident. Scale bar = 200 µm.

Table 3.1 – Summary of rhizomorph length measurements in Armillaria mellea in PD medium solidified with different concentrations of agar. aMean rhizomorph lengths significantly different from each other. bMean rhizomorph lengths not significantly different from each other. Data is shown as mean ± S.E. (n = 4).

Agar concentration (%) Mean length (mm)

0.5 651 ± 22a

2.0 1114 ± 62a,b

5.0 977 ± 221a,b

8.0 3166 ± 237a

CHAPTER 4

OSMOLYTE ACCUMULATION IN ARMILLARIA MELLEA RHIZOMORPHS: ITS ROLE IN TURGOR PRESSURE GENERATION

ABSTRACT

Accumulation of osmolytes creates differential osmotic pressure within fungal cells and their environment to drive water influx into the cells for the generation of tugor pressure. The generated turgor pressure contributes to cell expansion and pathogen infection of host in the yeast and filamentous fungi. Osmolyte accumulation and composition, together with their possible role in the generation of turgor pressure and tip growth in Armillaria mellea rhizomorphs was determined using Gas Chromatography/Mass Spectroscopy (GC/MS) and osmometry. A. mellea cultured on cellophane covering potato dextrose (PD) agar and in PD broth accumulated glycerol, erythritol and mannitol in its rhizomorphs. Chromatograms indicate that erythritol and mannitol were the most dominant osmolytes present in A. mellea rhizomorphs. Erythritol and mannitol occurred with equal concentrations in rhizomorphs cultured on cellophane covering PD agar, while erythritol was the most concentrated osmolyte in rhizomorphs cultured in PD broth. Generally, glycerol concentration was far less than erythritol and mannitol under both conditions. The total concentration of these osmolytes were determined to be 43 mM and 39 mM in rhizomorphs cultured on cellophane covering PD agar and those cultured in PD broth, respectively. Determination of osmolality of cellular extracts from rhizomorphs cultured in PD broth indicated that rhizomorphs generated a total osmotic pressure of 0.90 MPa, out of which an osmotic pressure of 0.09 MPa, constituting 10% of the total osmotic pressure is contributed by glycerol, erythritol and mannitol in rhizomorphs cultured in PD broth. In rhizomorphs cultured on cellophane, an osmotic pressure of 0.10 MPa contributed by glycerol, erythritol and mannitol constituted 11% of the total osmotic pressure. We conclude, therefore, that A. mellea accumulates osmolytes within its rhizomorphs and uses the accumulated osmolytes to create osmotic pressure to facilitate water influx, with resultant generation of turgor pressure within the rhizomorph, some of which is used to exert force for invasive growth.

1. INTRODUCTION

Mature mycelial cords and rhizomorphs have the ability to translocate nutrients and water towards their tips (Jennings et al., 1974). The hyphae are a likely repository of nutrients when the mycelial cord is migrating to explore new sources of nutrient over poor nutrient surfaces and, under such a condition, the integrity of the mycelial cord is maintained. However, the stimulus to cohesive growth is lost within the mycelial cord and the mycelia colonize the new substrate, upon encountering an external source of nutrient greater than its own supply (Moore, 1994). Until they come into contact with their new sources of food, they are described as migratory organs that develop and advance from one food-base over nutrient-poor substrates such as, bricks, stones, tiles, plastic sheets etc. to another (Arora, 1986; Money, 2004). In Serpula lacrymans for example, mycelial cords formed could be several meters long over such poor nutrient substrates (Butler, 1957, 1958; Watkinson, 1971; Jennings and Watkinson, 1982; Jennings, 1991). Mycelial cord formation is mostly associated with ageing mycelium that has virtually exhausted nutrient from its substrate. In an event of nutrient exhaustion, existing hyphae, acting as the main nutrient stores support the growth and existence of other new hyphae that grow alongside them. As long as such condition prevails, the integrity of a mycelial cord thus formed is maintained between these hyphae (the nutrient-supplier and the nutrient- dependent) until potential sources of food are encountered and explored. The tips of rhizomorphs can grow at great distances from the colony’s source of water and nutrients. Many questions, however, remain unanswered about how the tip growth is facilitated, and the exact mechanisms involved in mycelial cords and rhizomorphs migration and function. Some of the questions include the following: (i) does water move through vessel hyphae in some manner comparable to the movement of water through vessels in plant organs; (ii) does drying of the rhizomorph tip create the necessary gradient in water potential for water movement that is analogous to plant transmission; (iii) are certain osmolytes concentrated in the tips of rhizomorphs; (iv) if so, are these involved in driving osmotic flow of water along the organ; and, (v) do osmotically-active molecules moved through rhizomorphs also serve as energy source for the growing tip? The accumulation of polyols is linked with tolerance to drought and/or salinity in many species, including microorganisms, marine algae, higher plants and animals (Bohnert et al.

1995). Generally, microorganisms from most of the major groups are represented in environments that have reduced water activity. Filamentous fungi and yeasts have developed mechanisms and strategies of growth in environments containing high sugar concentration by accumulating solutes within their cells (Hocking, 1993). According to Hocking (1993) and Dijksterhuis and de Vries (2006), the accumulation of these solutes by fungi does not interfere with the functions of proteins and other biomolecules, when they are present in high amounts in the cell. For this reason, they are called “compatible” solutes. The term was introduced by Brown and Simpson (1972) to loosely define such solutes as those that, at high concentration, allow an enzyme to function effectively. In literature, they are sometimes referred to as osmolytes or polyols. In this chapter, the terms “osmolytes” and “polyols” are used interchangeably. Since it was established in 1945 that osmophilic yeasts produce polyhydric alcohols from sugar fermentation, there have been extensive studies on the production and accumulation of polyols in fungi. For example, Nikerson and Carroll (1945) identified glycerol in Zygosaccharomyces acidifaciens (= Zygosaccharomyces bailii), after which Spencer and Sallans (1956) found that polyol production occurred in 79 strains of yeast they studied. They reported that faster-growing strains produced mainly glycerol and D-arabitol, whereas the slower-growing strains produced mainly glycerol and erythritol. Osmotic adjustment and osmoprotection seem to be the two most important functions of polyols that are difficult to separate mechanistically; thus, during osmotic adjustments, the polyols act as osmolytes, aiding the retention of water in the cytoplasm and allowing sodium sequestration to the vacuole or apoplast, whereas in osmoprotection, protection of cellular structures might be accomplished through interactions of such osmolytes. Lewis and Smith (1967) speculated that polyols may function (i) as carbohydrate reserves; (ii) as translocated compounds; (iii) in helping to generate the appropriate internal solute potential; and (iv) in coenzyme regulation. Osmolyte biosynthesis, specifically, mannitol has been widely studied in plants, fungi and even animals. In fungi, the mannitol biosynthetic pathway was proposed by Hult and Gatenbeck (1978), and had since been variously reported to play crucial roles in their survival, especially in plant, animal and human pathogenesis, drought and stress tolerance, sporulation and spore discharge (Webster et al. 1995; Chaturvedi et al. 1996; Jennings et al. 2002; Ruijter et al. 2003; Solomon et al. 2006).

Brownlee and Jennings (1981) had earlier studied the content of soluble carbohydrates and their translocation in S. lacrymans. Analysis of sugars and polyols revealed various carbohydrates in different mycelial regions (margin, submarginal and mid-region) and in the mycelial cords. Notable amongst them were glycerol, mannitol, arabitol, trehalose, glucose, fructose and sucrose. Trehalose and arabitol were reported to be the most dominant in the mycelium. However, this could not be said of the mycelial cords. Trehalose was reported as the major carbohydrate in mycelial cords, making up over 70% of total soluble fraction, with arabitol occurring in high levels in marginal regions. 14Carbon labeling of the carbohydrates showed significant incorporation into trehalose, implicating trehalose as the main form by which carbohydrate is translocated in cords in S. lacrymans. Coggins et al. (1980) suggested that the flow of water along the hyphae of mycelial cord in S. lacrymans was driven by internal hydrostatic pressure generated by uptake of solutes causing uptake of water into hyphae by osmosis. Later, Jennings and Watkinson (1982) supported this idea by suggesting that hydrostatic pressure in hyphae could lead to the flow of water along the spaces in a mycelial cord. They explained that the structure of mycelial cord of the S. lacrymans is such that as the volume of water inside the living hyphae within the mycelial cord increases by osmosis, the resulting inward pressure created by the surrounding firm layer of rind and other wall materials will generate a hydrostatic pressure inside the mycelial cord that will only result in flow of water out of the living hyphae, and along the central cord spaces towards the tip. It is not clear, however, if the accumulation and translocation of osmolytes may be responsible for the generation of hydrostatic pressure and subsequent flow of water in rhizomorphs. Granlund et al. (1985) studied translocation of solutes along rhizomorphs of A. mellea. They were able to demonstrate that polyols and sugar were translocated along rhizomorphs of A. mellea. These polyols and other sugars were extracted and identified using gas-liquid chromatography (GLC), based on a method earlier described by Holligan and Drew (1971) and Holligan and Jennings (1972). Carbohydrates were extracted from rhizomorphs with water and ethanol, after which they were dried and derivatized using trimethylchlorosilane (TMCS). This method converted the sugars into trimethylsilyl esthers (TMS) that were then analyzed with the GCL. They reported that erythritol and glycerol were the major soluble carbohydrates found within the rhizomorphs of A. mellea, with mannitol, maltose, sucrose and trehalose forming a

significant part of the total amount of carbohydrates present and translocated. Although Granlund et al. (1985) had identified some major osmolytes in A. mellea rhizomorphs, we also repeated this work for the following reasons: 1. Whereas the studies of Granlund et al. (1985) focused on osmolyte and sugar translocation in A. mellea rhizomorphs and did not quantify the concentration of these osmolytes, our studies were conducted not only to identify and confirm the identity of these osmolytes, but to understand the role these osmolytes play in the generation of osmotic and turgor pressure in A. mellea rhizomorphs by quantifying their concentrations and overall contributions to turgor generation. 2. We adopted the most reliable, highly optimized and widely used alditol acetate derivatization method, coupled with GC/MS for the analyses of our samples. This is because the conversion rate of sugars to their alditol acetate is high, unlike in TMS derivatization of samples used by Granlund et al. (1985), where there is a high possibility of anomer formation and ring isomerization within samples that require measurement of as many as four derivatives of each sugar (Holligan and Drew, 1971; Brunton et al. 2007). Moreover, alditol acetate derivatization gives more simplified chromatograms of samples that readily produce mass spectra of peaks, making identification of compounds easier. With the above reasons we believe that data provided in our studies on osmolyte composition of A. mellea rhizomorphs are more accurate, reliable and up-to-date than that of Granlund et al. (1985) The main objective of this study is to examine the accumulation and composition of osmolyte in A. mellea and their role in the generation of turgor pressure within rhizomorphs. This was achieved in two ways: (i) by the use of GC/MS techniques to identify and quantify osmolytes present in the rhizomorphs, and (ii) by osmolality measurement to determine the level of osmotic pressure that these osmolytes generate within rhizomorphs. Results obtained from experiments indicate that the osmolytes erythritol, mannitol and glycerol were common in A. mellea rhizomorphs cultured on cellophane in PDA (aerial rhizomorphs), and in PD broth (submerged rhizomorphs). Although other osmolyte peaks were present in the chromatograms, they were not easily identified, because the fingerprints of their spectra closely resemble that of mannitol. Pinitol was readily identified, but could not be

quantified. In A. mellea, glycerol was present in low concentration, whereas erythritol and mannitol were the dominant osmolytes. The total concentrations of the three osmolytes identified were determined to be higher in A. mellea rhizomorphs cultured on cellophane covering PD agar than those in PD broth. Osmolality determination indicates that there is a differential between rhizomorph extracts and aliquots of PD broth in which they were cultured, implying this differential result in water influx that eventually results in the generation of turgor pressure within these rhizomorphs. Osmotic pressure estimated in rhizomorph sap indicates that glycerol, erythritol and mannitol present in A. mellea rhizomorphs cultured in PD broth accounted for 10% of the total osmotic pressure (0.90 MPa) compared to 11% of that in rhizomorphs cultured on cellophane.

2. MATERIALS AND METHODS

2.1 Organism and culture conditions

Armillaria mellea strain DR-140 used in these experiments was obtained from Dr. Dana Ritcher (School of Forest Resources and Environmental Sciences, Michigan Technological University, USA). A. mellea strain was maintained on PDA slants and cultured on PDA in Petri dishes covered with cellophane or in round-bottomed borosilicate glass tubes (Kimax, USA) containing PD broth. Both cultures were incubated in the dark during the duration of experiments. All stock cultures were subcultured at regular intervals and stored at 4 C to ensure their viability for subsequent experiment.

2.2 Media preparation

A quantity of 39 g of PD agar mixture was dissolved in 1 L distilled water, autoclaved at 1.1 kg/cm2 at 121 C for 15 min and poured in disposable Petri dishes prior to inoculation. Potato dextrose broth was prepared by dissolving a quantity of 24 g in 1 L distilled water, after which

10 mL of aliquots were dispensed into round-bottomed borosilicate glass tubes before autoclaving at 1.1 kg/cm2 at 121 C for 15 min.

2.3 Armillaria mellea rhizomorph culture for GC/MS

Solidified potato dextrose agar media in Petri dishes was covered with sterile circular cellophane, after which they were inoculated with A. mellea inoculum blocks. The culture plates were incubated in the dark at room temperature until matured rhizomorphs were carefully excised and immediately prepared for cellular extraction. After 10 mL aliquots of PD broth in round-bottom Borosilicate glass tubes had been autoclaved and cooled, they were inoculated with A. mellea inoculum blocks and incubated in the dark at room temperature until rhizomorphs were ready for use after 4-weeks. Matured rhizomorphs in cultures were carefully removed from the tubes with forceps, briefly rinsed with sterile distilled water, blotted with Kimwipes to dry, and immediately prepared for cellular extraction.

2.4 Sample extraction from Armillaria mellea

Well-developed rhizomorphs of A. mellea were carefully selected for cellular extraction to determine the composition, identity and concentration of organic compounds (osmolytes) therein using GC/MS, and to measure osmolality using Wescor® 5520C vapor pressure deficit osmometer (Wescor, Inc., Utah, USA). Sterile Eppendorf tubes were assigned to rhizomorphs. These tubes were weighed and their weight noted. Rhizomorphs were removed from the culture and placed in their appropriately labeled tubes and the tubes were weighed again. The wet weight of the fungal specimens was then computed (Table 4.1). A volume of 150 µL of distilled water was added to each tube and the tubes were placed in a water bath (Aloe Scientific, MO, USA) at 80 C for about 3 min to deactivate enzymes. Cultured rhizomorphs of A. mellea were crushed manually with a pointed plastic rod, since they were too tough to be disrupted with a sonicator. After crushing and disrupting samples, they were centrifuged (Micromax centrifuge, IEC,

Massachusetts, USA) at 10000 rpm for 5 min, after which the supernatant, hereafter referred to as the extracts, were transferred with pipettes (Gilson, France) into fresh tubes. The solid fungal samples that remain in the tubes were used to determine the dry weight of the fungi after extraction.

2.5 Dry weight determination of samples

Pellets of wet rhizomorph samples were individually collected on 55 millimeter-diameter Whatman’s No. 1 filter paper (Whatman International Ltd. Maidstone, England) that had been previously dried and weighed. The filter papers with the harvested pellets were placed in glass Petri dishes, covered, and dried in an electrically heated oven (Fisher Scientific) at 80 C for 24 hours, after which they were removed and then allowed to cool. Each filter paper with its harvested pellet was weighed with an electronic balance (model AG 135, Mettler Toledo, Switzerland) and the dry weight of the pellet calculated, accordingly (Table 4.1).

2.6 Derivatization of samples for Gas Chromatography/Mass Spectrometry

Cellular contents of rhizomorphs and mycelial cords were extracted with 150 µL of sterile distilled water and stored in Eppendorf tubes at -20 C until ready for use. For chemical derivatization, frozen extracts were thawed and transferred to glass vials. The samples were dried with heat under vacuum using a speed vac concentrator (model RH40-12, Savant Industries, Inc., NY, USA) at high speed for approximately 5 to 6 hrs. After drying, 300 µL of acetic anhydride was added to each sample, and the vials were tightly capped and placed in a drying oven for 15 to 19 hrs at 100 C. The glass vials were then removed and placed in an ice bath to cool. After adding 750 µL of distilled water to each sample, the vials were tightly capped, taped horizontally on a platform shaker (model G-560, Scientific Industries, Inc., Bohemia, NY, USA) and shaken for 1 hr. Then 1 mL of chloroform was added to each sample and vortexed for approximately 1 min. The upper aqueous phase was removed and discarded, using a 200 µL pipette, after which 0.8 mL cold refrigerated ammonium hydroxide was added to the solvent phase, and mixed briefly. After this, the lower portion of the sample was carefully transferred into newly labeled

Eppendorf tubes. These derivatized samples, thus obtained, were dried without heat under vacuum using a speed vac concentrator at high speed for 18 to 30 min, then re-dissolved in 10 µL chloroform before analysis by GC/MS. Unless these derivatized samples were immediately used, they were tightly capped and stored at room temperature.

2.7 Alditol acetate standards for fungal osmolyte and concentration determination

Erythritol, glycerol, and mannitol standards were prepared to calibrate the GC/MS instrument. Standard curves obtained were used to determine the concentration of osmolytes later identified from derivatized fungal samples (Figs 4.2, 4.4, and 4.6). Different concentrations of erythritol standards were prepared by dissolving 0.122 g of powdered erythritol in 1 mL sterile distilled water to make 1 M erythritol stock solution, after which aliquots of 100 µL, 10 µL, and 1 µL, constituting 100 mM, 10 mM, 1 mM of the 1 M stock solution, were drawn and placed in appropriately labeled glass vials. To prepare 100 µM, 10 µM and 1 µM standard solutions, 1 µL of 1 M erythritol stock solution was diluted to 1 mL to make 1 mM erythritol stock solution, after which aliquots of 100 µL, 10 µL and 1 µL were drawn and placed in appropriately labeled glass vials. Assuming all the standard solutions were diluted to 1 mL, they were dried and derivatized (see 2.6 of Materials and methods). After derivatization, the dried sample was dissolved in 1 mL chloroform. The same procedure was used to prepare mannitol standards: the standards were prepared from a 1 M mannitol stock solution prepared by dissolving 0.182 g in 1 mL of sterile distilled water. A volume of 1 µL of each concentration of erythritol and mannitol solutions was analyzed with GC/MS in triplicates. In the case of glycerol, 188.4 µL of liquid glycerol triacetate (GTA) was dissolved in 1 mL of chloroform to make a concentration of 1 M. Additionally, 1 mM stock solution was prepared from 1 M GTA stock solution. Without derivatizing, serial dilutions were prepared, after which 1 µL of each concentration was directly analyzed with GC/MS in triplicates. Blank runs of chloroform between different concentrations of the various osmolytes were carried out to clean out the column to ensure that concentrations of the solutions were not compromised, and to obtain clean chromatograms.

2.8 Determination of osmolyte identity in rhizomorph samples using Gas Chromatography/Mass Spectrometry

Derivatized fungal samples were re-dissolved in 10 µL of chloroform prior to analyses using a CP-3800 GC and a Saturn 2000 MS (Varian). One microliter of the sample was injected into the GC column using a Hamilton syringe (Reno, NV). Using the front injector, set to a temperature of 250 C, and a split ratio of 1:100, the samples were separated on Varian silica- coated CP-SIL 8 CB low-bleed capillary column with dimension 30m x 0.35 i.d. The temperature of the GC column was held at 160 C for 3 min, and then ramped at 12.5 C/min to 265 C. Each sample was run in triplicates. Blank runs of chloroform between samples were carried out to clean out the column to prevent cross contamination of samples, and to obtain clean chromatograms. Osmolytes were identified by comparison of retention times of major peaks from fungal samples to the retention time of peaks from GC/MS of alditol acetate standards, and their mass spectra obtained from the NIST/EPA/NIH Mass Spectral Library (NIST, Gaithersburg, MD).

2.9 Calculation of osmolyte concentration in rhizomorph samples

The concentrations of glycerol, erythritol and mannitol in derivatized rhizomorph samples A. mellea were calculated using x values determined from osmolyte standard curves (Figs. 4.2, 4.4, and 4.6; Tables 4.3, 4.4, 4.5) and log ion intensities of characteristic ion fragments specific for osmolytes (103 for glycerol, 217 for erythritol, and 139 for mannitol) from derivatized rhizomorphs samples (Tables 4.6, 4.7, 4.8, 4.9, 4.10 and 4.11).

To calculate the concentration of osmolytes in rhizomorph samples, the following mathematical expression was used:

conc. of osmolyte in sample = 10x x total vol. of standard x fraction of standard injected x total sample vol.

vol. of extracted rhizomorph sample

where, x = log of the concentration of the standard with the same ion count as the sample, from − by the standard curve using x = , (y is the log ion intensity of characteristic ion m of osmolyte under consideration, b is the y intercept of standard curve, and m is the slope of standard curve). The sample ion count was the mean of n = 3.

Total Volume of standard = 0.001 L,

Fraction of standard injected = 1/1000 µL of standard solution,

Total sample volume = 10 µL, and

Volume of extracted rhizomorph sample is represented by the difference between wet weight of rhizomorph sample (before extraction) and dry weight of rhizomorph sample (after extraction) (Table 4.1). The mass of the extract was converted to volume using a density of 1g/ml.

Below is an example of procedure used to determine the concentration of erythritol in rhizomorph cultured in PD broth:

The volume of the standard solutions were I mL and 1 µL was injected to produce the standard curve. A volume of 1 µL of the sample, which was resuspended in 10 µL of chloroform, was injected. The original volume of the rhizomorph extract, determined as wet weight – dry weight = 186 µL, which is 0.000186 L. Thus, the concentration of erythritol in the rhizomorph from equation 1 is,

()10− 39.0 1000/1001.0 10 μμ LxLxLxM

000186.0 L = 0.022 M = 22.0 mM

The above procedure was used to determine the concentrations of all other osmolytes identified in derivatized rhizomorph samples.

2.10 Osmolality measurement and osmotic pressure estimation in Armillaria mellea

Inoculum blocks of Armillaria mellea about 5 mm square were excised and used to inoculate 10 mL of sterile PD broth in round-bottomed borosilicate glass tubes (Kimax, USA). These were incubated at room temperature for two weeks. Rhizomorphs produced by A. mellea cultures were used for cellular extraction and osmolality measurement. Unless these extracts were immediately used, they were stored in tightly-capped glass vials at -20 C. Ten microliters of the extracted samples were pippeted onto 6 mm diameter filter paper sample discs already placed in a sample chamber of the Wescor® 5520C vapor pressure deficit osmometer (Wescor Inc., Logan, Utah, USA) to determine the osmolality (mol/kg) of the samples. Aliquots of the PD broth used in culturing the rhizomorphs were retained and used for osmolality measurement. All samples were run in triplicates. Between sample runs, the osmometer thermocouple was cleaned with distilled water and calibrated with NaCl standards. The mean value of osmolality registered for PD broth was subtracted from the mean value of osmolality for extracted sample to compute the actual value of osmolality for the samples. Using the van’t Hoff’s equation, the osmolality measurements were converted to osmotic pressure. Thus, Osmotic pressure (MPa) = RTc, where RT = 2.454 kg MPa mol-1 at 22 C, and c = osmolality in mol kg-1. From Davis et al. (2000), the osmotic pressures (MPa) of glycerol, erythritol and mannitol identified within A. mellea rhizomorphs were calculated using the expression, osmotic pressure (MPa) = ax + bx2, where x is the molarity of osmolytes from rhizomorph sample (Table 4.2), and a and b are curve parameters provided by Davis et al. (2000).

3. RESULTS

3.1 Determination of osmolyte identity and concentration in Armillaria mellea rhizomorph extracts using Gas Chromatography/Mass Spectrometry

Derivatized extracts of A. mellea rhizomorphs cultured on cellophane and in PD broth were analyzed using GC/MS. In A. mellea rhizomorphs cultured on cellophane and in PD broth, three common osmolytes (glycerol, erythritol and mannitol) were identified by comparing their mass spectra to those obtained from osmolyte standards and from NIST/EPA/NIH Mass Spectral Library (Figs. 4.7 and 4.8). Additionally, the retention times of the standard osmolytes were crucial in ascertaining the authenticity of the peaks of glycerol, erythritol and mannitol derived from the chromatograms of the fungal samples. Glycerol, erythritol and mannitol standards had retention times of 16.25 min, 19.25 min and 23.80 min, respectively (Figs 4.1, 4.3 and 4.5). Retention time for peaks of glycerol, erythritol and mannitol detected in the rhizomorph samples differed by a range of ± 0.20 min. Erythritol and mannitol were the most dominant osmolytes detected. They were represented by prominently large peaks in the chromatogram, although erythritol peaks in both samples were larger than mannitol peaks. Glycerol, on the other hand, although present in both samples, was characterized by remarkably small peaks (Figs 4.7 and 4.8). Calculations of osmolyte concentrations within the rhizomorphs indicate that erythritol and mannitol were the most concentrated osmolyte present in rhizomorphs cultured on cellophane covering PD agar (Fig 4.9; Table 4.2): erythritol concentration was determined to be 21.3 ± 4.0 mM; mannitol concentration was determined to be 21.0 ± 5.0 mM. The situation was surprisingly different in rhizomorphs cultured in PD broth: a concentration of 30.3 ± 5.0 mM erythritol was determined, compared to a reduced mannitol concentration of 8.2 ± 2.0 mM. In general, the total identified osmolyte concentrations of 43 mM and 39 mM were determined in rhizomorphs cultured on cellophane and rhizomorphs cultured in PD broth, respectively. One peak in the gas chromatogram was identified as pinitol, a cyclohexitol found in many living systems (Duquesnoy et al. 2008). We suspect that the unidentified peaks in the chromatograms may represent sorbitol and arabitol, since they are high molecular weight compounds and are closer to the mannitol peak than to erythritol peak (Figs 4.7 and 4.8).

3.2 Osmolality measurement and osmotic pressure estimation in Armillaria mellea

Osmolality of extracted samples of A. mellea rhizomorphs cultured in PD broth was determined. Aliquots of the PD broth used in culturing the rhizomorphs were retained and used to determine their osmolality. The mean value of osmolality registered for PD broth (53 ± 1 mol kg-1; n = 15) was subtracted from the mean value of osmolality of extracted samples (361 ± 25 mol kg-1; n = 15) to compute the actual value of osmolality for the samples, which was 308 mol kg-1. Using the van’t Hoff’s equation, the osmotic pressure of rhizomorph sap was calculated to be 0.90 MPa, predicting a maximum osmotic pressure of 0.77 MPa for rhizomorphs growing in PD broth, whose osmotic pressure was 0.13 MPa. This suggest that differential in osmotic pressure induces water influx to generate turgor pressure within the rhizomorphs. Respective osmotic pressures of glycerol, erythritol and mannitol in rhizomorphs cultured on cellophane covering PD agar generated osmotic pressures of 0.0005 MPa, 0.047 MPa and 0.051 MPa, whereas those cultured in PD broth generated osmotic pressures of 0.0002 MPa, 0.069 MPa and 0.020 MPa, respectively. Thus, the total osmotic pressure of identified osmolytes estimated from rhizomorph cultured on cellophane covering PD agar was 0.10 MPa, which accounted for 11% of the overall osmotic pressure (0.90 MPa), compared to 10% (0.09 MPa) of the overall osmotic pressure estimated in rhizomorphs in PD broth. These estimations were based on the overall osmotic pressure of the rhizomorph extract from PD broth (0.90 MPa).

4. DISCUSSION

Osmolyte accumulation and function in fungi and other organisms are well documented in literature (Bohnert et al. 1995; Hocking, 1993; Dijksterhuis and de Vries 2006). In general, the strategy of accumulating a mixture of osmolytes is a common occurrence in many fungi (Davis et al. 2000). Reports from studies with fungal spore discharge, mycelia and mycelial cords, and appressoria have highlighted the possible functions of these osmolytes on the growth, dispersal, and survival strategies of fungi in their environments, and their interactions with other fungi, especially in pathogen-host settings (Butler, 1957, 1958; Watkinson, 1971; Jennings and Watkinson, 1982; Jennings, 1991; Webster et al. 1995; Chaturvedi et al. 1996; Ruijter et al.

2003; Jennings et al. 2002; Trail et al. 2002, 2005; Fischer et al. 2004; Solomon et al. 2006). For example, mannitol is one of the most common osmolytes that have been implicated to play various roles in fungi and plants, many of which include osmoregulation, serving as a storage or translocated carbohydrate, serving as a source of reducing power, regulating coenzymes, regulating cytoplasmic pH, acting as a sink or source for protons, etc, (Lewis and Smith, 1967; Jennings, 1984). Nonetheless, the actual physiological roles of osmolytes metabolized and accumulated in fungi are unclear (Clark et al. 2003). In fungal rhizomorphs, not much is understood about the role of such osmolytes, especially in the generation of turgor pressure, and extension mechanism of rhizomorph tip growth. In our study, our main objective was to examine the composition of osmolytes in A. mellea and their possible role in the generation of turgor pressure within rhizomorphs and on tip growth. We hypothesized that the accumulation of osmolytes in rhizomorph will generate sufficient turgor pressure to contribute to rhizomorph tip extension. In our experiments, we extracted cellular contents of A. mellea rhizomorphs cultured in PD broth. The osmotic pressure of the cellular contents, which constitute the identified osmolytes, and other unidentified sugars and inorganic ions, was determined to evaluate its contribution towards the provision of turgor pressure during growth. Erythritol, mannitol and glycerol were identified, using CG/MS as the common osmolytes present in the rhizomorphs of A. mellea cultured on cellophane covering PD agar and in PD broth. In both cases, erythritol and mannitol were the most dominant osmolytes identified. Glycerol, although present, was in very low levels in our A. mellea samples as compared to erythritol and mannitol (Figs 4.7, 4.8 and 4.9; Table 4.2). This was a surprising outcome considering the fact that glycerol has been reported to generally play an important role in creating osmotic potential that facilitates uptake of water in fungi (Kelly and Budd, 1991; Dixon et al. 1999). For instance, de Jong et al. (1997) reported that the accumulation of glycerol in the appressoria of the rice blast fungus Magnaporthe grisea provided the needed osmotic force for the penetration of leaf epidermis. The reason for the low levels of glycerol in the rhizomorphs is unclear, especially when Granlund et al. (1985) had reported that glycerol and erythritol were the major osmolytes present in A. mellea rhizomorphs they had worked with, but did not estimate the concentrations of the osmolytes they identified. Although they used GLC to identify their osmolytes, they failed to provide chromatograms and spectra of these osmolytes, for reasons we

77 do not know. Additionally, their claim that glycerol and erythritol were the major osmolytes present was contradicted by the data presented in their report, which show that it was rather erythritol and mannitol that were the major osmolytes translocated. This ambiguity in their report may only be attributed to ineffectiveness of their derivatization methods that possibly affected the reliability of their data (Holligan and Drew, 1971; Brunton et al. 2007). This is why we are confident that our data are more accurate and reliable: this is evident from chromatograms and spectra of our identified osmolytes obtained from GC/MS analyses of A. mellea rhizomorphs. In our studies, we determined that erythritol and mannitol are the two most dominant osmolytes present in A. mellea rhizomorphs (Figs 4.7, 4.8 and 4.9; Table 2). Also, we estimated their concentrations, which totaled 43 mM and 39 mM within rhizomorphs cultured on cellophane and in PD broth, respectively (Table 4.2). We estimated, using osmometric techniques that the osmotic pressure of rhizomorph sap was 0.9 MPa; this predicts a maximum osmotic pressure of 0.77 MPa for rhizomorphs growing in PD broth with an osmotic pressure of 0.13 MPa. From Davis et al. (2000), we calculated that glycerol, erythritol and mannitol in rhizomorphs cultured on cellophane covering PD agar generated osmotic pressures of 0.0005 MPa, 0.047 MPa and 0.051 MPa, respectively. The total osmotic pressure of these identified osmolytes was 0.10 MPa, which accounted for 11% of the overall osmotic pressure (0.90 MPa) estimated from rhizomorph sap. This implies that the remaining osmotic pressure of 0.80 MPa generated within the rhizomorph may be accounted for by other unidentified osmolytes, sugars, and inorganic ions present in the rhizomorph sap extracted. Compared to rhizomorphs cultured in PD broth, osmotic pressures calculated for glycerol, erythritol and mannitol were 0.0002 MPa, 0.069 MPa and 0.020 MPa, respectively, totaling an osmotic pressure of 0.09 MPa, and accounting for 10% of the overall osmotic pressure estimated. These estimations correlate with the estimated concentrations of the individual isolated osmolytes within the rhizomorphs (Table 4.2). Based on our data from GC/MS (Figs 4.7 and 4.8), and calculated concentrations of individual osmolytes identified (Fig 4.9; Table 4.2), it is obvious that erythritol and mannitol are the two main osmolytes among the osmolyte mixture that contribute largely to creating the osmotic pressure within the rhizomorph. We inferred from our data that the presence of osmolytes, together with other sugars and inorganic ions in the rhizomorphs, create differences in osmotic pressure between the cellular content of the A. mellea rhizomorphs and the media in which they were cultured. This difference in the osmotic pressures induces water influx into the

rhizomorph, which results in the generation of turgor pressure. (Money, 1994; 1997, Davis et al. 2000). This was indeed the case in our osmolality measurements of extracts of rhizomorphs cultured in PD broth and the aliquots in which they were cultured. The difference between the osmolality of the rhizomorph extract (361 mol kg-1; 0.90 MPa) and the aliquots (53 mol kg-1; 0.13 MPa) accounts for the actual osmolality of 308 mol kg-1 (0.77 MPa) that is responsible for the generation of turgor pressure from water influx. The mechanism underlying water influx in the mycelial cords of S. lacrymans was described by Coggins et al. (1980), and later supported by Jennings and Watkinson (1982). Eamus and Jennings (1984) also demonstrated that long distance transport in various wood decay fungi was due to turgor gradients. This notwithstanding, it may also be possible that the presence of different osmolytes within rhizomorphs may play other functions which include; (i) being stored as carbohydrate reserves; (ii) being used as translocated compounds; (iii) being used in the generation of appropriate internal solute potential; and (iv) involvement in coenzyme regulation as speculated by Lewis and Smith (1967). The rhizomorphs of A. mellea were cultured under two extreme environments conditions – rhizomorphs cultured on cellophane covering PD agar were confronted with a drier environment in sharp contrast to rhizomorphs cultured in PD broth, where the rhizomorphs were completely submerged in liquid. In rhizomorphs cultured on cellophane covering PD agar, the concentrations of both osmolytes were comparatively high, but were not significantly different from each other (Fig 4.9; Table 4.2). Additionally, mannitol concentration in these rhizomorphs was higher than in rhizomorphs in PD broth. Interestingly, erythritol concentration estimated in rhizomorphs cultured in PD broth was far higher than mannitol. Since rhizomorphs that are cultured on cellophane covering PD agar encountered drier environments, we suspect that the higher concentration of mannitol, together with that of erythritol, may possibly serve an additional role by facilitating water uptake from the atmosphere to provide the cells with the needed osmotic balance and turgor pressure (Brown, 1976). This may ensure that the rhizomorphs generate and maintain sufficient internal osmotic and turgor pressure to help them extend at the tips and migrate across the cellophane surface from one end to another. Surprisingly, the total osmotic pressures generated by these identified osmolytes in rhizomorphs cultured under both conditions do not differ significantly from each other. In a comparative experiment with mycelial cords of M. incrassata cultured on cellophane covering malt marmite

agar, GC/MS chromatograms showed high peaks of erythritol and mannitol (data not shown), an observation that may be interpreted to suggest that erythritol and mannitol may be the most dominant osmolytes in fungi that thrive in dry environment. According to Dijksterhuis and de Vries (2006), erythritol and glycerol are linked to osmotic stress and growth at lower water activity. Niederpreum and Hunt (1967) reported that glycerol, mannitol, and arabitol as the main osmolytes present in Schizophyllum commune, a wood decomposing fungus. Additionally, Davis et al. (2000) reported that comparisons between the osmotic effects of polyols with three, four, five and six carbon atoms at identical concentrations showed a statistically significant increase in osmotic pressure with increasing molecular weight, with the most profound effect at high osmotic concentration. This may suggest that, erythritol, a four carbon molecule, is required in high concentrations in preference to glycerol, a three carbon molecule, to contribute to the overall osmotic pressure within the rhizomorphs. These support our data that much as it might not be the dominant osmolyte in other fungi, erythritol may be a preferred osmolyte of extraordinary importance to A. mellea rhizomorphs than glycerol, especially in those cultured in PD broth. Also, it there is a possibility that rhizomorphs in PD broth naturally favor the accumulate erythritol than mannitol, hence its higher concentration (Fig 4.9; Table 4.2). Information available in literature suggests that attempts have been made in the past to provide insights into erythritol metabolism, catabolism, and function in bacteria, yeast and other filamentous fungi (Hajny et al. 1964; Braun and Niederpruem, 1969; Sperry and Robertson, 1975; Kim et al. 1997; Ookura et al. 2005; Slotnick and Dougherty, 1972). In one recent study, Kogej et al. (2007) reported that erythritol played a major role in the osmotic adaptation of the halophilic fungus Hortaea werneckii. Based on data obtained from our studies, we conclude, therefore, that (i) A. mellea accumulates osmolytes within its rhizomorphs; (ii) erythritol, unlike glycerol in other fungi, and mannitol are the two most dominant osmolytes in A. mellea rhizomorphs; (iii) identified osmolytes within A. mellea rhizomorphs contribute up to 11% of the total osmotic pressure generated within rhizomorphs; and (iv) there is an established differential in osmotic pressure that drives water influx to generate turgor pressure within A. mellea rhizomorphs. We suggest that further work focuses on the identification of other osmolytes that were not easily identified in this work to complete the list of osmolyte mixture that is accumulated in A. mellea. Another area of interest is to investigate the metabolic pathways that result in the

80 accumulation of the dominant osmolytes in the rhizomorphs, and their actual physiological roles in the rhizomorphs, considering the two extremes of environment that these rhizomorphs were cultured.

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Fig 4.1 – A chromatogram of derivatized 100 mM glycerol standard at a retention time of 16.25 min, showing the corresponding mass spectrum (insert) with characteristic ion fragment of 103.

4.5

4.0

3.5

3.0

2.5 y intercept = 5.44

Log intensity ofLog Ion intensity 103 slope = 0.73

2.0 2 r = 0.99

1.5

-5.5 -5.0 -4.5 -4.0 -3.5 -3.0 -2.5 -2.0 -1.5

Log concentration (M)

Fig 4.2 – A log/log standard curve of glycerol showing a regression line with concentrations ranging between 10 µM to 10 mM.

Fig 4.3 – A chromatogram of derivatized 100 mM erythritol standard at a retention time of 19.25 min, showing the corresponding mass spectrum (insert) with characteristic ion fragment of 217.

4.5

4.0

3.5

3.0

2.5 y intercept = 4.98

2.0 slope = 0.83

Log intensity of Ion 217 r2 = 0.98 1.5

1.0 -4.5 -4.0 -3.5 -3.0 -2.5 -2.0 -1.5 -1.0 -0.5

Log concentration (M)

Fig 4.4 – A log/log standard curve of erythritol showing a regression line with concentrations ranging between 100 µM to 100 mM.

Fig 4.5 – A chromatogram of derivatized 100 mM mannitol standard at a retention time of 23.80 min, showing the corresponding mass spectrum (insert) with characteristic ion fragment of 139.

5.0

4.5

4.0

3.5

3.0 y intercept = 5.26 2.5 slope = 0.80

Log intensity of Ion 139 Log intensity 2.0 r2 = 0.95

1.5 -4.5 -4.0 -3.5 -3.0 -2.5 -2.0 -1.5 -1.0 -0.5

Log concentration (M)

Fig 4.6 – A log/log standard curve of mannitol showing a regression line with concentrations ranging between 100 µM to 100 mM.

* Erythritol mass spectrum

Mannitol mass spectrum

= erythritol peak * x x = mannitol peak

Fig 4.7 – GC/MS chromatogram of Armillaria mellea rhizomorph cultured on cellophane covering potato dextrose agar, showing peaks of erythritol (*) and mannitol (x).

Erythritol mass spectrum

Mannitol mass spectrum

= erythritol peak * x = mannitol peak

Fig 4.8 – GC/MS chromatogram of Armillaria mellea rhizomorph cultured in PD broth, showing peaks of erythritol (*) and mannitol (x).

40 Glycerol Erythritol Mannitol

10 Osmolyte concentration (mM) Osmolyte

0 Rhizomorph Rhizomorph on cellophane in PD broth

Fig 4.9 – Concentrations of osmolytes in Armillaria mellea rhizomorphs cultured on cellophane covering PD agar and in PD broth. Means ± S.E. for samples grown on cellophane versus PD broth are not significantly different for glycerol, but are different for erythritol and mannitol. The osmolyte concentration of samples was the mean of n = 9.

Table 4.1 – Wet and dry weight of Armillaria mellea rhizomorphs before and after cellular extraction. The difference between wet and dry weight represents mass of cellular extract. *The mass of the extract was converted to volume using a density of 1g/ml, and used to calculate concentration of osmolytes in rhizomorphs.

Sample extraction from Armillaria mellea rhizomorphs

Wet weight (mg) Dry weight (mg) *Mass extracted (mg)

Sample 1 90.55 4.44 86.11

Rhizomorphs cultured on Sample 2 70.61 8.40 62.21 cellophane

Sample 3 72.64 8.10 64.54

Sample 1 190.37 3.90 186.47 Rhizomorphs

cultured in PD broth Sample 2 302.14 22.99 279.15

Sample 3 315.47 30.71 284.76

Table 4.2 – Mean concentrations of erythritol, mannitol and glycerol determined in Armillaria mellea rhizomorphs. Means for samples grown on cellophane versus PD broth are not significantly different for glycerol and erythritol, but are different for mannitol. The osmolyte concentration of samples was the mean of n = 9.

Mean concentration of osmolytes ± S.E. (mM)

Armillaria mellea rhizomorphs Armillaria mellea rhizomorphs Osmolyte grown on cellophane covering PD agar grown in PD broth (n = 9) (n = 9)

Glycerol 0.2 ± 0.1 0.1 ± 0.03

Erythritol 21.3 ± 4.0 30.3 ± 5.0

Mannitol 21.0 ± 5.0 8.2 ± 2.0

Total Osmolyte Concentration 42.5 38.6

Table 4.3 – Data of ion intensity from characteristic 103 ion from different concentrations of glycerol standard solutions. *Parameters used to plot log/log glycerol standard curve.

Ion intensity of characteristic 103 ion of glycerol standard

Ion intensity of Concentration *Log concentration characteristic ion 103 Mean ion intensity *Log ion intensity

92 10 µM -5 77 77.00 1.89

62 227

100 µM -4 239 251.33 2.40 288

1910 1 mM -3 2020 1970.67 3.30

1982 8469

10 mM -2 11781 10165.00 4.01 10245

Table 4.4 – Data of ion intensity from characteristic 217 ion from different concentrations of erythritol standard solutions. *Parameters used to plot log/log erythritol standard curve.

Ion intensity of characteristic 217 ion of erythritol standard

Ion intensity of Concentration *Log Concentration characteristic ion 217 Mean ion intensity *Log ion intensity

27 100 µM -4 39 34.33 1.54

37 388

1 mM -3 427 485.33 2.69 641

1744 10 mM -2 2209 2023.67 3.31

2118 10991

100 mM -1 13290 12133.33 4.10 12119

Table 4.5 – Data of ion intensity from characteristic 139 ion from different concentrations of mannitol standard solutions. *Parameters used to plot log/log mannitol standard curve.

Ion intensity of characteristic 139 ion of mannitol standard

Ion intensity of Concentration *Log Concentration characteristic ion 139 Mean ion intensity *Log ion intensity

100 µM -4 187 156.67 2.19

197

501

1 mM -3 1126 763.33 2.88

663

20651

10 mM -2 19122 21480.10 3.33

24670

47507

100 mM -1 52109 50040.67 4.69

50506

Table 4.6 – Data of ion intensity from characteristic 103 ion of glycerol in rhizomorph samples with corresponding x values obtained from glycerol standard. *Parameters used to calculate glycerol concentration in Armillaria mellea rhizomorphs cultured in PD broth.

Ion intensity of characteristic 103 ion of glycerol with corresponding x value from glycerol standard curve obtained from Armillaria mellea samples

Ion intensity of

characteristic ion 103 Mean ion intensity *Log ion intensity *x value

5755 Sample 1 6005 4764.70 3.68 -2.426

2534 3953

Sample 2 4262 3637.33 3.56 -2.591 2697

1949 Sample 3 3662 3201.33 3.51 -2.660

3993

Table 4.7 – Data of ion intensity from characteristic 103 ion of glycerol in rhizomorph samples with corresponding x values obtained from glycerol standard. *Parameters used to calculate glycerol concentration in Armillaria mellea rhizomorphs cultured on cellophane covering PD agar.

Ion intensity of characteristic 103 ion of glycerol with corresponding x value from glycerol standard curve obtained from Armillaria mellea samples

Ion intensity of

characteristic ion 103 Mean ion intensity *Log ion intensity *x value

842 Sample 1 1128 1036.70 3.02 -3.335

1140 2101

Sample 2 3414 2360.33 3.37 -2.853 1566

2388 Sample 3 3835 2587.67 3.41 -2.811

1540

Table 4.8 – Data of ion intensity from characteristic 217 ion of erythritol in rhizomorph samples with corresponding x values obtained from erythritol standard. *Parameters used to calculate erythritol concentration in Armillaria mellea rhizomorphs cultured in PD broth.

Log ion intensity of characteristic 217 ion of erythritol obtained from Armillaria mellea samples with corresponding x value from erythritol standard curve

Ion intensity of

characteristic ion 217 Mean ion intensity *Log ion intensity *x value

49786 Sample 1 62806 46227.70 4.66 -0.390

26091 64601

Sample 2 102195 81699.67 4.91 -0.080 78303

85206 Sample 3 90789 98735.33 5.00 0.024

120211

Table 4.9 – Data of ion intensity from characteristic 217 ion of erythritol in rhizomorph samples with corresponding x values obtained from erythritol standard. *Parameters used to calculate erythritol concentration in Armillaria mellea rhizomorphs cultured on cellophane covering PD agar.

Log ion intensity of characteristic 217 ion of erythritol obtained from Armillaria mellea samples with corresponding x value from erythritol standard curve

Ion intensity of

characteristic ion 217 Mean ion intensity *Log ion intensity *x value

22735 Sample 1 13371 16744.00 4.22 -0.920

14126 22458

Sample 2 23774 21759.67 4.34 -0.770 19047

19553 Sample 3 29059 19343.00 4.29 -0.830

9417

Table 4.10 – Data of ion intensity from characteristic 139 ion of mannitol in rhizomorph samples with corresponding x values obtained from mannitol standard. *Parameters used to calculate mannitol concentration in Armillaria mellea rhizomorphs cultured in PD broth.

Log ion intensity of characteristic 139 ion of mannitol obtained from Armillaria mellea samples with corresponding x value from mannitol standard curve

Ion intensity of

characteristic ion 139 Mean ion intensity *Log ion intensity *x value

42989 Sample 1 44969 40636.70 4.61 -0.818

33952 38533

Sample 2 38659 37855.00 4.58 -0.516 36373

36326 Sample 3 48418 41967.67 4.62 -0.805

41159

Table 4.11 – Data of ion intensity from characteristic 139 ion of mannitol in rhizomorph samples with corresponding x values obtained from mannitol standard. *Parameters used to calculate mannitol concentration in Armillaria mellea rhizomorphs cultured on cellophane covering PD agar.

Ion intensity of characteristic 139 ion of mannitol with corresponding x value from mannitol standard curve obtained from Armillaria mellea samples

Ion intensity of characteristic ion 139 Mean ion intensity *Log ion intensity *x value 30540 Sample 1 26301 29250.00 4.47 -0.994 30909 39813 Sample 2 47739 41713.67 4.62 -0.805 37589 45172 Sample 3 46016 42851.00 4.63 -0.792 37365 CHAPTER 5

THE BIOMECHANICS OF INVASIVE GROWTH IN ARMILLARIA MELLEA RHIZOMORPHS

ABSTRACT

The accumulation of osmolytes creates a differential osmotic pressure between fungal cells and their environment that drives water influx into the cells and generates turgor pressure. The exertion of forces from turgor pressure created within fungal hyphae and other infection structures, such as appressoria, is well documented. Oomycete hyphae have also been studied and various measurements of forces using strain gauge have been reported in the literature. This chapter reports the first measurements of the forces exerted by extending rhizomorphs. Forces exerted by rhizomorphs of Armillaria mellea cultured in potato dextrose agar were measured using a strain gauge. Rhizomorph tips exerted forces that ranged between 1.44 x 10-3 and 5.84 x 10-3 N., with applied pressures ranging between 0.04 MPa and 0.30 MPa. Calculations indicated that a mean force of 2.67 x 10-3 ± 3.20 x 10-4 N, corresponding to a mean applied pressure of 0.13 ± 0.02 (MPa), was exerted by rhizomorph tips that made contact with the strain gauge. This mean applied pressure (0.13 ± 0.02 MPa), constitutes 17% of the total turgor pressure (0.77 MPa) generated within rhizomorphs. The addition of 25 mM polyethylene glycol-6000 to a rhizomorph tip already exerting force on a beam of strain gauge resulted in initial increase in force, after which there was a steady decrease. We conclude from this study that rhizomorphs exert substantial levels of pressure, which are derived from turgor pressure and are sufficiently powerful to penetrate the solid materials that they invade.

108 1. INTRODUCTION

Rhizomorphs are produced in abundance when some fungi are cultured on potato dextrose agar and other media. Some of the rhizomorphs are aerial, projecting a few millimeters into the air; others are invasive, radiating from the central inoculum and penetrating the surrounding agar. The exertion of force by rhizomorphs is evident from the compression and splitting of agar media in advance of their apices. Cultured rhizomorphs extend well beyond the fringe of hyphae that expand from the inoculum, demonstrating that they extend more swiftly than the “unbundled” vegetative hyphae. Many observations suggest that in vitro growth rates reflect the fast extension of rhizomorphs in nature. Disease centers of A. mellea in stands of ponderosa pine have been estimated to expand at a rate of up to 2 m per year, which is an astonishing 5.4 mm per day (Shaw and Roth, 1976). Most of this spread was probably achieved by tip-extending rhizomorphs in compact soils and solid debris through force exertion at their tips. Sustained fast extension of rhizomorphs was proven by Findlay (1951), who showed that rhizomorphs of A. mellea had grown more than 9 meters along a brick-lined drainage tunnel in 7 years, corresponding to a sustained average growth rate of 3.6 mm per day. It is common knowledge that rhizomorphs of wood-decomposing and plant pathogenic fungi have remarkable invasive abilities. However, to our knowledge, no study has been conducted on the source of the strength for rhizomorphs that enable them to push through compacted soils. The closest attempt made was by Coggins et al. (1980) who suggested that the flow of water along the hyphae of mycelial cords in Serpula lacrymans was driven by internal hydrostatic pressure generated by uptake of solutes causing uptake of water into hyphae by osmosis. Jennings and Watkinson (1982) supported this idea and further suggested that hydrostatic pressure in hyphae could lead to the flow of water along the spaces within a mycelial cord. They explained that the structure of mycelial cord of the S. lacrymans is such that the volume of water inside the living hyphae within the cord increases by osmosis. The resulting inward pressure created by the surrounding firm layer of rind and other wall materials will, as the volume of water increases, generate a hydrostatic pressure inside the mycelial cord that will only result in flow of water out of the living hyphae and along the central cord spaces towards the tip. Interestingly, they did not calculate the amount of force that is created at the tip of the mycelial cords resulting from the internal hydrostatic pressure that was generated. This may be due to the

109 fact that the idea of measuring forces may not have occurred to them. There is also a possibility that instruments capable of making force measurements were unavailable at the time. Basidiomycetes have been reported to crack open bark of decaying timber, asphalts, and concrete paving through forces during fruiting body expansion (Gooday 1985; Niksic, 2004). The strength of the fruiting bodies originates from hydrostatic pressure. Many decades ago, Buller (1931) estimated that hyphae within an elongating stipe exerted a pressure of 0.07 MPa to raise a weight of more than 200 g. Money and Ravishankar (2005) have measured that a pressure of 0.059 MPa is associated with stipe elongation in Coprinopsis cinerea fruit bodies. Over the past decade, studies on fungal hyphal growth have generally addressed the biochemical and physiological processes involved in cell wall synthesis and expansion (Jackson and Heath, 1993; Lopez-Franco et al. 1994; Shapiro and Mullins, 2002; Horio and Oakley, 2005). Although an enormous literature exists on hyphal growth, the elongation process has been very controversial. For example, Lopez-Franco et al. (1994) reported that the hyphal elongation was not steady, but fluctuated continuously with alternating periods of fast and slow growth at more or less regular intervals. While this issue is still unsolved, recent studies on fungal hyphal elongation have focused on the mechanics of tip extension with emphasis on the exertion of force by hyphal tips as they penetrate their substrates (Ravishankar et al. 2001; MacDonald et al. 2002). Many other studies have led to a general acceptance that the expansion of hyphae at the tip is mainly controlled by forces produced by turgor, enabling fungi to penetrate all kinds of solid substrates (Strange, 1993; Wessel, 1993; Griffin, 1994; Koch, 1994; Money, 1995; Ravishankar, et al. 2001; MacDonald et al. 2002; Money and Ravishankar, 2005). Some fungal infection of plants has been largely attributed to the role of turgor pressure created in infection apparatus such as appressorium. It was shown that turgor pressure created in the appressoria in Magnaporthe grisea generates the driving force for leaf penetration (Howard et al. 1991). Experiments have reported turgor as high as 8.0 MPa in M. grisea (Howard and Ferrari, 1989; Howard et al. 1991; Money and Howard, 1996). Money (2007) explicitly explains the processes that are involved in generation of hyphal turgor pressure, as a source of force needed by hyphal tips for invasive growth. Bastmeyer et al. (2002) summarize it this way: “The driving force of hyphal invasion growth is turgor pressure.” Although data from earlier studies provided insight into processes that are involved in hyphal tip growth, there has been little comparable work on rhizomorph development and

110 mechanics. By understanding how hyphae bundle during the formation of these organs, and how they extend in a cooperative fashion, it is possible to elaborate upon the findings made so far on tip growth at the hyphal level. This will allow us to understand how rhizomorphs are able to spread through forest soils. The rhizomorph system of A. mellea strain DR-140 represents an excellent experimental model for this research because (i) they develop quickly and plentifully in standard culture media, (ii) they are composed of extending bundled hyphae, and (iii) they are reasonably larger than hyphae, making it suitable for easy handling, manipulation, and observation during force measurements. Rhizomorphs of other species are also composed of lots of hyphae, but because they do not form in vitro (at least as easily), they don’t lend themselves to these experiments. The main objective of this study was to estimate how much force may be generated from turgor pressure by rhizomorphs as they migrate through compact soil and penetrate woody substrates. Force measurements from rhizomorph tips indicate that fungal rhizomorphs exert pressures ranging between 0.04 and 0.30 MPa. Overall, the mean force measured was 2.67 x 10-3 ± 3.20 x 10-4 N, corresponding to an overall mean applied pressure of 0.13 ± 0.02 MPa, which is 17% of the total turgor pressure calculated from osmometric measurements in chapter 4. Polyethylene glycol-6000 added to rhizomorph tips already exerting forces on a beam of strain gauge caused an initial steady increase in voltage output, which was sustained for only a period of time, after which there was a dramatic decrease in voltage output. An interesting observation from chart recording of output voltage suggests that rhizomorphs may be extending in a pulsed fashion. We recommend further investigation into this observation.

2. MATERIALS AND METHODS

2.1 Organism and culture conditions

Armillaria mellea strain DR-140 used in these experiments was generously supplied by Dr. Dana Richter (School of Forest Resources and Environmental Sciences, Michigan Technological University, USA). A. mellea strain DR-140 was maintained on PDA slants. For

111 experiments, this strain was cultured on PDA in Petri dishes in the dark at room temperature until rhizomorphs emerged.

2.2 Media preparation

A quantity of 39 g of PDA mixture (Difco, Becton, Dickinson & Company, MD, USA) was dissolved in 1 L distilled water, autoclaved at 1.1 kg/cm2 at 121 C for 15 min. A quantity of 50 mL sterile PDA was poured in disposable Petri dishes prior to inoculation to provide the depth needed to access the rhizomorph tips with the strain gauge.

2.3 Rhizomorph culture for force measurements

Solidified PDA with greater than normal depth between, 8 mm and 10 mm was inoculated with inoculum blocks of A. mellea mycelium from stock plates. The cultures were incubated in the dark at room temperature until rhizomorphs emerged. Submerged rhizomorphs at a depth of approximately 6 mm within the PDA were selected for force measurements. This was to ensure easy access to these rhizomorphs when they extended from a well cut from the agar in advance of their extending tips.

2.4 Force measurements from Armillaria mellea rhizomorphs

Forces exerted by A. mellea rhizomorphs were measured using the strain gauge method described for hyphae by Ravishankar et al. (2001) and MacDonald et al. (2002). The strain gauge was calibrated with standard weights (Troemner, Philadelphia, PA) ranging from 100 mg to 500 mg, providing forces of 0.98 to 4.90 millinewtons (mN; force = mass x gravitational acceleration; Fig 5.1). Force measurements were made from rhizomorphs that extended from agar medium into an open through that was made by removing a slab of agar (5 x 10 x 10 mm, width x length x

112 depth) in advance of the growing tips. Cultures were placed on the stage of an inverted microscope (OLYMPUS IX70-S8F, Olympus Optical Co. Ltd., Japan), for observation during the force measurements. The tip of a rhizomorph that has extended into the trough, after about an hour, was brought into view and focused at a magnification of 2X. Force measurements were made with a strain gauge (FORT 10 load cell force transducer; World Precision Instruments, Sarasota, FL) by maneuvering it into the trough under the control of a micromanipulator such that the rhizomorph tip pushed against the beam of the instrument as it extended from the cut shelf of agar (Fig 5.2). Because the rhizomorph tip was extending horizontally from the agar into the trough, the strain gauge was carefully tilted such that it was perpendicularly aligned to the extending rhizomorph (Fig 5.2). Additionally, about 50 µL of sterile distilled water was transferred into the trough to ensure that the immediate surroundings of the extending rhizomorph were sufficiently moistened to prevent dehydration of the rhizomorph tip. This was carefully done to ensure that the extending rhizomorph and the strain gauge did not come into contact with the sterile distilled water, as this would interfere with data acquisition. The strain gauge output was adjusted to read zero, using the bridge amplifier, after which the free end of the strain gauge (adapted with a 7 x 7 mm chip cut from a plastic cover slip glued over the hole at the end of the aluminum beam) was carefully manipulated to contact the tip of the extending rhizomorph. After the initial contact, the free end was moved slightly away from the rhizomorph tip until the strain gauge read zero. Force exerted by the extending rhizomorph was then monitored as the tip pressed against the beam of the strain gauge. After output voltage (mV) of the strain gauge had peaked and remained constant or declined, the free end of the strain gauge in contact with the rhizomorph tip was repositioned to separate the rhizomorph tip from the strain gauge. A successful recording was made when the strain gauge output immediately dropped to zero at the baseline, providing a measure of the maximum force applied by the extending rhizomorph (Fig 5.3). A standard chart recorder (mode BD12E; Kipp and Zonen, Delft) was used for data acquisition. The diameter of the rhizomorph tip that made contact with the strain gauge was noted and later used in calculating pressure exerted.

113 2.5 Effect of polyethylene glycol-6000 on force exertion by Armillaria mellea rhizomorphs

Polyethylene glycol-6000 (PEG-6000) (Fluka Chemical Corp., New York) solution was prepared by dissolving 150 g in 1 L distilled water (25 mM; 0.3 MPa), based on Money (1989). During force measurement, about 10 μL of PEG-6000 solution was carefully added to the rhizomorph tip when it was still in contact with strain gauge and pushing against it. Force exerted by the extending rhizomorph was then monitored. After output voltage (mV) of the strain gauge had declined and remained constant, the free end of the strain gauge in contact with the rhizomorph tip was repositioned to separate the rhizomorph tip from the strain gauge. A successful recording was made when the strain gauge output immediately dropped to zero at the baseline, providing a measure of the maximum force applied by the extending rhizomorph (Fig 5.3). Data acquired on a standard chart recorder was analyzed, and the diameter of the rhizomorph tip that made contact with the strain gauge was noted and later used in calculating pressure exerted.

2.6 Calculation of pressure exerted by the rhizomorph tip

Measured forces were divided by the cross-sectional area of rhizomorph tip that made contact with the stain gauge at its thinnest point. This provided a measure of the pressure exerted by the rhizomorph as it pushed against the strain gauge. A set of calculations is provided as an example:

Consider a plateau output voltage recorded from the strain gauge during contact with rhizomorph of 61 mV

Voltage is directly converted to force with reference to a calibration curve (Fig 5.1). Thus, force = voltage (mV) x 4 x 10-5 = 61 mV x 4 x 10-5 = 2.44 x 10-3 N or 2.44 mN

114 The area of rhizomorph cross section in contact with the strain gauge, πr2, (100 µm diameter) = 8.0 x 10-9 m2

Pressure = force ÷ area = 2.44 x 10-3 N ÷ 8.0 x 10-9 m2 = 0.30 x 106 Pa = 0.30 MPa = 3 atmospheres.

This procedure was used to determine pressure from measured forces (Tables 5.1 and 5.2).

3. RESULTS

3.1 Force measurements from Armillaria mellea rhizomorphs

Rhizomorph tips pushed against the strain gauge which registered the applied force. The maximum output voltage (mV) from the strain gauge (Fig 5.3), was used to calculate the forces exerted by rhizomorphs as described in the Materials and Methods. Calculations showed that forces exerted by the rhizomorphs ranged between 1.44 x 10-3 and 5.84 x 10-3 N, and applied pressures ranging between 0.04 and 0.30 MPa: the rhizomorphs that exerted these forces varied in their tip area that made direct contact with the strain gauge as they pressed against it. For example, the highest force of 5.84 x 10-3 recorded by the strain gauge was exerted by a rhizomorph tip with a contact area of 4.9 x 10-8 m2. An average force of 2.67 x 10-3 ± 3.20 x 10-4 N with a corresponding mean applied pressure of 0.13 ± 0.02 MPa (n = 16) was recorded. When compared with our estimate of the turgor pressure generated by osmolyte accumulation within rhizomorphs (0.76 MPa), discussed in Chapter 4, this mean applied pressure corresponds to 17% of the turgor pressure. The pressures exerted by the rhizomorphs varied from 0.04 MPa to 0.30 MPa, corresponding to 5% and 39% of the mean pressure of the rhizomorph (P = 0.77 MPa; Chapter 4).

115 As the rhizomorph tips pushed against the strain gauge, they flattened slightly, producing a larger contact area than anticipated from the morphology of the usually acute rhizomorph tips. The diameter of these flattened contact areas was used to calculate the applied pressures (Table 5.1; see Section 2.6 of Material and Methods). Another notable observation made during recording of force exerted by the rhizomorph tip seemed to suggest a pulsatile extension of rhizomorphs (Fig. 5.3). Although each rhizomorph exhibited a steady overall increase in output voltage until it became constant, chart recordings showed dramatic variations in force. We recommend that more studies in this aspect of rhizomorph extension are conducted to ascertain the veracity of this observation. This is important because pulsatile extension patterns in fungal hyphae remain highly controversial. Lopez-Franco et al. (1994) have reported that the rate of hyphal elongation was not steady but fluctuated continuously, with alternating periods of fast and slow growth at more or less regular intervals.

3.2 Effect of polyethylene glycol-6000 on force exertion by Armillaria mellea rhizomorphs

Upon exposure to 25 mM PEG-6000, the force exerted by extending rhizomorphs was boosted for some minutes, and then showed a substantial decrease (Fig 5.4). Assuming the force exerted by the rhizomorph tip is derived from the turgor pressure within the constituent hyphae of the organ, the initial rise in applied force must have resulted from an increase in rhizomorph turgor, and/or from increased cell wall loosening. The reason for this effect is unclear, but is probably related to the fact that only the tip of the rhizomorph was exposed to the PEG-6000 solution in these experiments, with the majority of the organ buried within the agar medium. The immediate disturbance of the differential in water potential between the rhizomorph tip and surrounding fluid would tend to induce water loss from the tip; this might include water flow toward the rhizomorph apex, causing the measure increase in force. We do not have direct evidence for this, but this interpretation is at least consistent with the observed behavior of the rhizomorphs. Subsequent diffusion of PEG-6000 through out the culture would eventually dehydrate the entire organ, causing the sustained loss in force registered by the strain gauge. Irrespective of the explanation for the counterintuitive initial increase in applied force in response to hyperosmotic

116 shock, the extreme sensitivity of the force exerted by the rhizomorphs to changes in medium osmolality is consistent with the presumed osmotic origin of the force-generating mechanism of rhizomorphs.

4. DISCUSSION

Observations of rhizomorph extension in agar media are fascinating. What makes it intriguing is that the “movement” of the rhizomorphs is appreciable to the naked eye, unlike that in tiny fungal hyphae where one could only view extension and penetration with the aid of a microscope. We were tempted to ask how much force the rhizomorphs exerted as they extend through the solid medium, especially when the medium is augmented with differing concentrations of agar. More thrilling is the thought of the rhizomorphs migrating in compact soils, where the amount of force needed to migrate over long distances could be challenging due to the heterogeneous nature of the soil environment. Based on data presented in Chapters 3 and 4, we conceived the idea to measure forces that rhizomorphs could exert as they extended through agar, and what role tugor pressure might play in the exertion of these forces. In spite of its importance as a means by which A. mellea propagates itself to effect infection in healthy plants over long distances, the mechanics of rhizomorph extension are poorly studied and understood. To our knowledge, the measurement of force exerted by elongating and invasive rhizomorphs of A. mellea in this study is the first of its kind. Previous studies on the mechanics of fungal growth have focused on biomechanics of hyphal and fruiting body extension, and on infection mechanisms of appressoria in fungal pathogens of plants (Gooday, 1985; Howard and Ferrari, 1989; Howard et al. 1991; Jackson and Heath, 1993; Lopez-Franco et al. 1994; Money and Howard, 1996; Brush and Money, 1999; Ravishankar et al., 2001; Shapiro and Mullins, 2002; Niksic et al. 2004; Horio and Oakley, 2005). We believe that the A. mellea rhizomorph system represents a unique experimental model for extending research on hyphal growth to understanding the biomechanics of organ development in fungi. In order words, the main contribution of this work is to provide data and insights to fill the existing knowledge gap between understanding hyphal extension and complex structural morphogenesis in fungi. Our

117 studies also demonstrate the remarkable improvements that have been made in the area of instrumentation to measure forces exerted by hyphal tips, and now rhizomorphs, since Buller’s classic experiments with Coprinus sterquilinus. This subject of interest has been given a detailed review by Bastmeyer et al. (2002). Buller’s remarkable demonstration in 1931 that a basidiocarp of C. sterquilinus could raise a weight of 200 g obviously shows how powerful tugor pressure generated by fungal hyphae could be. As recently as 2005, Money and Ravishankar reported that elongating stipes of C. cinerea exerted a pressure of 0.059 MPa as they pressed against a strain gauge. Despite the differences in their methods to determine the amount of pressure generated by elongating fruiting bodies, the data reported by Buller (1931), and Money and Ravishankar (2005) are strikingly similar. Many recent studies that have focused on oomycete and fungal hyphae have also reported measurements of turgor pressure and the consequent force that they generate (Money, 1990; Money, 1995; Ravishankar, et al. 2001; MacDonald et al. 2002; Money and Ravishankar, 2005). Additionally, force measurements from hyphae using strain gauges have produced interesting data. For example, Johns et al. (1999) made total force measurement of 107 µN that was exerted by hyphae of the oomycete water mold Achlya bisexualis. MacDonald et al. (2002) also measured a force of approximately 2 µN exerted by hyphae of two oomycete pathogens, P. graminicola and P. insidiosum. The use of A. mellea rhizomorph in our study adds a new dimension to the analysis of force exerted by fungi during penetration of their substrates. As stated earlier, the novelty of this study is the nature of the experimental system used. Money (2001) details the difficulty in working with slow-growing fungal hyphae that are as small as 5 µm or less in diameter, although he also stated that hyphae of Achlya grow swiftly and can exceed 30 µm in diameter. We believe that rhizomorphs of A. mellea represent an excellent complementary experimental model for studying tip growth in fungi. The average rhizomorph tip diameter alone of rhizomorphs used in force measurements in these experiments was 172 ± 13 µm (Table 5.2). Because instrumentation had already been perfected for measuring forces exerted by hyphal tips, which are far smaller than rhizomorphs, data acquisition from rhizomorphs using the same basic approach was straightforward (Fig 5.2). Experiments indicated that forces exerted by individual rhizomorphs (n = 16) ranged from 1.44 x 10-3 and 5.84 x 10-3 N. Corresponding pressures are 0.08 MPa and 0.12 MPa,

118 respectively, based on the contact area of the rhizomorph tip with the strain gauge (Table 5.1). The maximum applied pressure of 0.31 MPa recorded by the strain gauge was exerted by a rhizomorph tip that made a contact area of 8.0 x 10-9 m2 with the strain gauge (this maximum pressure, which is equivalent to 3 atmospheres, is more than the pressure in automobile tire). Force data were derived from the maximum voltage output (mV) recorded from the strain gauge after output peaked and remained constant. (Fig 5.3; see Material and Methods). When all data was collated, it became clear that an average force of 2.67 x 10-3 ± 3.20 x 10-4 N was exerted by the rhizomorphs, with a corresponding average pressure of 0.13 ± 0.02 MPa (n = 16). This suggests that the average pressure generated by the rhizomorph tip represents only 17% of the average total turgor pressure within the rhizomorphs (0.77 MPa). It is presumed that this pressure contributes to the overall driving force for extension and penetration. Buller’s estimated pressure of 0.07 MPa by an elongating stipe in 1931, together with a pressure of 0.059 MPa reported by Money and Ravishankar (2005) show that A. mellea rhizomorphs in vitro exert higher pressure than fruiting bodies. This may also be true in their natural soil environments, a scenario which may be attributed to the narrow rhizomorph tip. The narrow rhizomorph tip of A. mellea as shown in PD 7% agar in Fig. 3.5 of Chapter 3, suggests that rhizomorphs may serve as a wedge for penetrating compact soil, as compared to a broad cap of a young fruit body trying to push its way through the soil upwards. If this is the case in their natural environment, it then suffices to say that rhizomorphs are well adapted for extending through heterogeneous surroundings. For example, Findlay (1951) reported a 9-meter growth in A. mellea rhizomorphs along a brick-lined drainage tunnel in a period of 7 years. Similarly, Shaw and Roth (1976) estimated that A. mellea expanded in a ponderosa pine field at a rate of 2 m per year, an observation that has been attributed to the development of tip-extending rhizomorphs. Strain gauge experiments we have conducted have undoubtedly demonstrated that rhizomorph tips exert significant invasive pressure derived from turgor pressure. This is evident from our data in Chapter 4 showing an osmotic pressure of 0.90 MPa within rhizomorphs, of which 0.77 MPa constitutes the tugor pressure. Applied pressures calculated from measured forces provided in this chapter are consistent with the view that part of the total tugor pressure that is generated in rhizomorphs is used for rhizomorph extension and invasive growth. In one case, for example, a single rhizomorph exerted a total force of 1.44 x 10-3 N over a flattened contact area of 1.8 x 10-8 m2 (diameter 150 µm). This corresponds to an applied pressure of 0.08

119 MPa, which is equivalent to approximately 11% of rhizomorph turgor pressure (Table 5.1). Interestingly, this pressure is sufficient for mechanical penetration of medium solidified with up to 5% agar (which has a gel strength of 0.05 MPa), but the fact that rhizomorphs plough through 8% agar (gel strength 0.09 MPa; Chapter 3) shows that these organs are be capable of exerting higher pressures (Tables 5.1 and 5.2), which is confirmed by an applied pressure of 0.13 MPa we are reporting. Previous studies have shown that individual hyphae of a variety of fungi and oomycete water molds apply pressures ranging from 0.04-0.09 MPa (Money, 2007). The match between rhizomorph and hyphal pressure suggests that rhizomorphs behave as a relatively simple sum of their parts. In this respect, rhizomorphs are surprisingly similar to the elongating stipe of basidiomes (Money and Ravishankar, 2005). We conclude from the study that rhizomorphs exert substantial levels of turgor pressure, of which a variable proportion, up to 39%, is used to create the needed pressure at their tips to penetrate and invade substrates.

4.1 FUTURE STUDIES

This study is a crucial beginning in understanding the biomechanics of rhizomorphs in that it raises a series of questions that needs to be critically addressed in the future. To this end, future studies should seek to address the following questions: • How is rhizomorph development initiated from constituent hyphae after they aggregate, and do mechanics play any role in hyphal interlacing; do patterns of cell division lay the foundation for hyphal differentiation as seen in a mature rhizomorph?

• Is hyphal extension within the interior of the rhizomorph the driving force behind overall rhizomorph extension? Are differential forces created within hyphal cells responsible for bending and branching? Where are the zones of extension within the rhizomorph?

• What mathematical models will best explain elongation and branching mechanics, and how do these compare with those of other systems, such as hypha and root?

Findings from experiments designed to answer these important questions will be very significant as they will be the first attempt to elucidate rhizomorph morphogenesis, and the mechanics involved in its invasive abilities. Understanding the initial stages of development will be very helpful in appreciating rhizomorph invasive growth in fungal phytopathogens and wood

120 decay basidiomycetes, since these are of economic importance in real estate, forest ecology and plant pathology. As these questions are addressed, it will also be an important step towards documenting for the first time, the role of stress and strain, coupled with concerted hyphal extension, on the elongation and branching of rhizomorphs. It is also envisioned that this will begin a new interest in the biomechanical studies, not only of rhizomorph development, but of fungal fruiting bodies, appressoria, and other fungal structures.

121 REFERENCES

Bastmeyer M, Deising HB, Bechinger C, 2002. Force exertion in fungal infection. Annual Review of Biophysics and Biomolecular Structures 31: 321-341.

Brush L, Money NP, 1999. Invasive hyphal growth in Wangiella dermatitidis in induced by stab inoculation and shows dependence upon melanin biosynthesis. Fungal Genetics and Biology 28: 190-200.

Buller AHR, 1931. Researches on Fungi. Volume 4. Longmans, Green & Co., London.

Coggins CR, Jennings DH, Clarke RW, 1980. Tear or drop formation by mycelium of Serpula lacrymans. Transactions of the British Mycological Society 75: 63-7.

Findlay WPK, 1951. The development of Armillaria mellea rhizomorphs in a water tunnel.

Transactions of the British Mycological Society 34: 146.

Gooday GM, 1985. Elongation of the stipe of Coprinus cinereus. In: Moore D, Casselton LA, Wood DA, Frankland JC (eds), Developmental Biology of the Higher Fungi. Cambridge University Press, Cambridge, UK, pp 311–331.

Griffin DH, 1994. Fungal Physiology. 2nd edn. Wiley-Liss, New York.

Horio T, Oakley BR, 2005. The role of microtubules in rapid hyphal tip growth of Aspergillus nidulans. Molecular Biology of the Cell 16: 918–926.

Howard JR, Ferrari MA, 1989. Role of melanin in appressorium function. Experimental Mycology 13: 403-418.

Howard RJ, Ferrari MA, Roach DH, Money NP, 1991. Penetration of hard substrates by a fungus employing enormous turgor pressure. Proceedings of the National Academy of Sciences USA 88: 11281-11284.

122 Jackson SL, Heath IB, 1993. Roles of calcium ions in hyphal tip growth. Microbiology and Molecular Biology Review 57: 367-382.

Jennings L, Watkinson SC 1982. Structure and development of mycelial strands in Serpula lacrymans. Transactions of the British Mycological Society 78: 465-474.

Johns S, Davis CM, Money NP, 1999. Pulses in turgor pressure and water potential: resolving the mechanics of hyphal growth. Microbial Research 154: 225-231.

Koch AL, 1994. The problem of hyphal growth in Streptomycetes and fungi. Journal of Theoretical Biology 171: 137-150.

Lopez-Franco R, Bartnicki-Garcia S, Bracker CE, 1994. Pulsed growth of fungal hyphal tips. Proceedings of the National Academy of Sciences, USA 91: 12228-12232.

MacDonald E, Millward L, Ravishankar JP, Money NP, 2002. Biomechanical interaction between hyphae of two Pythium species (Oomycota) and host tissues. Fungal Genetics and Biology 37: 245–249.

Money NP, 1989. Osmotic pressure of aqueous polyethylene glycols: relationship between molecular weight and vapor pressure deficit. Plant Physiology 91: 766-769.

Money NP, 1995. Turgor pressure and the mechanics of fungal penetration. Canadian Journal of Botany 73: S96-S102.

Money NP, 2001. Biomechanics of invasive hyphal growth. In: Howard RJ, Gow NAR (eds), The Mycota, Volume 8, Biology of the Fungal Cell, first edn. Springer-Verlag Berlin Heidelberg, pp. 3-17.

123 Money NP, 2007. Biomechanics of invasive hyphal growth. In: Howard RJ, Gow NAR (eds), The Mycota, Volume 8, Biology of the Fungal Cell, second edn. Springer-Verlag Berlin Heidelberg, pp. 237-249.

Money NP, Howard RJ, 1996. Confirmation of a link between fungal pigmentation, turgor pressure, and pathogenicity using a new method of turgor measurement. Fungal Genetics and Biology 20: 217-227.

Money NP, Ravishankar JP, 2005. Biomechanics of stipe elongation in the basidiomycete Coprinopsis cinerea. Mycological Research 109: 628–635.

Niksic M, Hadzic I, Glisic M, 2004. Is Phallus impudicus a mycological giant? Mycologist 18: 21–22.

Ravishankar JP, Davis CM, Davis DJ, MacDonald E, Makselan SD, Millward L, Money NP, 2001. Mechanics of solid tissue invasion by the mammalian pathogen Pythium insidiosum. Fungal Genetics and Biology 34: 167-175.

Shapiro A, Mullins JT, 2002. Hyphal tip growth in Achlya bisexualis. I. Distribution of 1,3-β- glucans in elongating and non-elongating regions of the wall. Mycologia 94: 267–272.

Shaw CG, Roth LF, 1976. Persistence and distribution of a clone of Armillaria mellea in a ponderosa pine forest. Phytopathology 66: 1210-113.

Strange RN, 1993. Plant disease control: regulation of cell wall mechanical properties. Annual Review of Plant Physiology 35: 585-657.

Wessels JGH, 1993. Wall growth, protein excretion and morphogenesis in fungi. Transley Review 45. New Phytologist 123: 397

124

140

120

100

60 40

Strain gauge output (mV) 0

0 100 200 300 400 500

Applied mass (mg)

Fig 5.1 – Calibration curve for rigid-lever strain gauge used in the rhizomorph force experiments (FORT 10 load cell force transducer; WPI, Sarasota, FL). Each data point represents duplicate measurements.

125

Fig 5.2 – Measurement of force exerted by Armillaria mellea rhizomorphs using a rigid-lever strain gauge. Rhizomorph tip (on left) pressing against beam of strain gauge (on right), after extension into a well created by the removal of a block of agar in its path.

126

Fig 5.3 – A representative chart recording the force applied by an Armillaria mellea rhizomorph over a period of 30 min. The recording shows the maximum level of force exertion of the rhizomorph tip on the right. At end of the recording, the strain gauge separated from the rhizomorph tip, resulting in the immediate drop in output voltage to the baseline. A peak force of 2.52 mN was exerted by this rhizomorph with a cross sectional area of 3.1 x 10-8 m2, accounting for a pressure of 0.08 MPa (= 0.8 atmospheres).

Fig 5.4 – A representative chart recording the force applied by an Armillaria mellea rhizomorph over a period of about 30 min after the addition of 25 mM polyethylene glycol-6000. The recording shows the maximum level of force exertion of the rhizomorph (asterix), after which there was a decrease in force exerted. A peak force of 3.0 mN was exerted by this rhizomorph with a cross sectional area of 3.1 x 10-8 m2, accounting for a pressure of 0.10 MPa (= 1 atmosphere).

Table 5.1 – Individual measurements of applied forces and pressures from Armillaria mellea rhizomorphs (n = 16).

Maximum Rhizomorph tip Rhizomorph output Applied Force diameter contact area Applied Pressure (mV) (N) (µm) (m2) (MPa)

38 1.50 x 10-3 150 1.8 x 10-8 0.08

60 2.40 x 10-3 200 3.1 x 10-8 0.08

80 3.20 x 10-3 200 3.1 x 10-8 0.10

48 1.92 x 10-3 100 8.0 x 10-9 0.20

80 3.20 x 10-3 150 1.8 x 10-8 0.20

52 2.10 x 10-3 150 1.8 x 10-8 0.12

51 2.04 x 10-3 150 1.8 x 10-8 0.11

68 2.72 x 10-3 150 1.8 x 10-8 0.15

68 2.72 x 10-3 200 3.1 x 10-8 0.09

61 2.44 x 10-3 100 8.0 x 10-9 0.30

42 1.70 x 10-3 100 8.0 x 10-9 0.21

146 5.84 x 10-3 250 4.9 x 10-8 0.12

136 5.44 x 10-3 250 4.9 x 10-8 0.11

36 1.44 x 10-3 150 1.8 x 10-8 0.08

50 2.00 x 10-3 200 3.1 x 10-8 0.06

50 2.00 x 10-3 250 4.9 x 10-8 0.04

129

Table 5.2 – Mean ± S.E. values of applied force and pressure by rhizomorphs pressing against strain gauge (n = 16).

Rhizomorph tip Rhizomorph Max. Output Applied Force diameter contact area Applied Pressure (mV) (N) (µm) (m2) (MPa)

Mean ± S.E. 67 ± 8 2.67 x 10-3 ± 3.20 x 10-4 172 ± 13 2.52 x 10-8 ± 3.60 x 10-9 0.13 ± 0.02

CHAPTER 6

GENERAL DISCUSSION, CONCLUSIONS AND FUTURE STUDIES

1. RESEARCH BACKGROUND

Fungi play vital ecological roles as decomposers of woody debris in the terrestrial ecosystem. Many of these wood-decay fungi are basidiomycetes (Schmidt, 2006). Fungi of the genera Armillaria, Auricularia, Coniophora, Fomitopsis, Meripilus, Meruliporia, Polyporus, and Serpula are examples of well-known wood decomposers. Fungi have various strategies to decompose their wood substrates in heterogeneous environments (Lindahl and Olsson, 2004; Schwarze, 2007). They secrete enzymes at the tips of their growing hyphae to break down complex molecules (cellulose, lignin, pectin, and chitin) in their substrates into smaller molecules (sugars, organic acids, and amino acids), after which the nutrients are absorbed and translocated from the substrate to parts of the hypha or mycelium where these nutrients are needed most for growth (Lindahl and Olsson, 2004; Money, 2007). Mycelia spread by actively growing fungi to other parts of the substrate as nutrients are depleted. In their quest for new substrates, these fungi form mycelial cords and rhizomorphs over long distances, which are used as exploratory and migratory organs that enhance their spread and survival (Hartig, 1874; Falck, 1912). Thus, mycelial cords and rhizomorphs of wood-decay and pathogenic fungi serve as important sources of inoculum, enabling fungi to make contact with newly colonized wood substrates both in soil and in buildings, and other wooden structures (Money, 2004). For example, the rhizomorph-forming fungus Meruliporia incrassata causes “dry rot” of wood components in buildings, and is commonly found causing damage to homes in California. During attacks on homes, M. incrassata develops rhizomorphs from its depleted food base mostly from outside the home (tree stumps, landscaping lawns with wood chips, etc.). Under warm and very moist conditions, the rhizomorph is capable of translocating nutrients and water to its tip until it encounters a wooden component of the building. It then begins to colonize the wood swiftly by digesting and extracting cellulose, leaving behind the brown lignin component of the wood. This fungus destroys dry wood, by transferring water through its rhizomorphs to the sites of active growth and decomposition (Verall, 1968). A different fungal

131 species, Serpula lacrymans, is known to be responsible for the same kinds of destruction in Europe, especially in Britain (Money, 2004). S. lacrymans, together with other fungi, was responsible for rotting of timber, and eventual sinking of the ship Royal George in 1782 (Money, 2004). Both M. incrassata and S. lacrymans cause brown rot of their wood substrates; and both produce water- and nutrient-conducting rhizomorphs or cords (Burdsall, 1991; Moore, 1994). Research interest in the structure and development of mycelial cord in S. lacrymans has generated enormous amount of information in literature (Falck, 1912; Hartig and van Tubeuf, 1902; Brownlee and Jennings 1981a; Hornung and Jennings 1981; Jennings and Watkinson 1982). Ironically, and to the best of our knowledge, virtually no studies have been done on rhizomorph development and its invasive growth in M. incrassata. We reasoned that studies on these aspects of rhizomorphs in M. incrassata would very insightful in understanding the biochemical and biomechanical processes that underlie their invasive growth and destruction. This was the main reason behind my decision to initiate research on M. incrassata rhizomorphs for my doctoral degree, in addition to the fact that M. incrassata is an economically important wood-decay basidiomycete that causes building decay, with costs running into several millions of dollars annually (Money, 2004). One thing became clear from in vitro preliminary studies with M. incrassata: unlike its behavior in the building environment, it does not produce rhizomorphs in vitro, but only mycelial cords! The difficulty to induce M. incrassata to generate rhizomorphs in vitro might be one of the main reasons why virtually no information is available in literature on its rhizomorph structure and development. I shifted focus to the plant pathogenic fungus, Armillaria mellea, because it readily produces rhizomorphs in vitro, making it an excellent model for developmental and biomechanical studies in rhizomorph systems. This research was designed as an attempt to understand how rhizomorphs work as organs for translocation, migration, invasion, and infection. To this end, specific objectives of the various studies were aimed to:

1. Examine in detail features of rhizomorph anatomy that support their migratory, translocatory and invasive behavior.

2. Study the effects of increasing medium gel strength on the production, growth and extension rates of A. mellea rhizomorphs in vitro.

132 3. Examine osmolyte accumulation in A. mellea and its role in the generation of turgor pressure within rhizomorphs.

4. Estimate, for the first time, how much force may be generated by these rhizomorphs as they migrate through compact soil and penetrate their hard woody substrates.

To summarize, we wanted to analyze the hyphal construction of rhizomorphs that endow them with their reputed role in translocating nutrients and water as they migrate from one location to another. Having accomplished this, understanding invasive growth in rhizomorphs within their heterogeneous soil environments was the next logical area we pursued. We sought to investigate what might stimulate rhizomorph growth and extension, such that they are successful with their invasive growth and migration over long distances, especially across non-nutrient substrates. Based on information in literature about the presence of osmolytes within cords of S. lacrymans and rhizomorphs of A. mellea, it was important to identify and quantify some of these osmolytes, in order to understand their roles in turgor generation as a source of pressure needed for invasive growth. This we did with the use of GC/MS and osmometric techniques. The next logical question was how much force do these rhizomorphs exert at their tips, and how much of the turgor generated was used to exert these forces. For the first time, we have been able to measure forces exerted by rhizomorphs using a strain gauge, a novel approach that broadens our knowledge in biomechanics of hyphal tip growth, on one hand, and organized tip growth, on the other hand, especially in structures as mushroom fruiting bodies and rhizomorphs. Findings and conclusions from these experiments have contributed to a broader understanding of the functions of rhizomorphs in relation to their structure and development, and are presented in this dissertation. In this chapter, I present a brief discussion of findings and their conclusions, and propose future studies that will expand upon our work.

133 2. GENERAL DISCUSSION

2.1 Ultrastructural studies of Meruliporia incrassata and Armillaria mellea using scanning electron microscopy

Fungi direct much of their energy into the mass production of mycelium when nutrients in their substrate are being depleted, and especially when the mycelial front encounters new substrate (Lindahl and Olsson, 2004). Through the mycelia, these fungi are able to spread from one location to the other. In other fungi, especially root-infecting and wood-decay fungi, mycelial cords and rhizomorphs have long been known as important agents by which these fungi spread (Hartig, 1874; Falck, 1912). It was only recently that more attempts were made to study and understand the structure and development of cords and rhizomorphs in relation to their functions as translocatory, migratory and invasive organs. Townsend (1954) studied rhizomorph structure of sixteen basidiomycete fungi including Armillaria mellea (Fr.) Quel., Marasmius androsaceus Fr., Serpula lacrymans Fr., Phallus impudicus Pers. In the last couple of decades, Brownlee and Jennings (1981a, b), Hornung and Jennings (1981), Jennings and Watkinson (1982), Granlund et al. (1984, 1985), and Eamus, et al. (1985) have quite extensively studied the biochemical, physiological and anatomical aspects of mycelial cords in Serpula lacrymans (some of these aspects of rhizomorphs in A. mellea were also studied). In the case of M. incrassata, Burdsall (1991) reviewed its distribution, significance and occurrence in the United States. Of the studies earlier conducted, that of Granlund et al. (1984) was of most interest to us, because it involved anatomical studies of A. mellea rhizomorphs. Granlund et al. (1984) carried out SEM studies on A. mellea rhizomorphs to investigate the internal and external features of these organs in relation to their reputed ability to translocate water and other solutes over long distances. The internal structure revealed the arrangement of tissues of the A. mellea rhizomorph: the different zones of hyphal tissues observed were (i) peripheral hyphae that formed a layer approximately between 20 and 30 µm thick, (ii) cortex, (iii) medulla and (iv) the central space, usually occupied by a cottony pith of fine loosely-woven hyphae. Our findings from SEM studies confirmed the same zones of hyphal tissues within strain DR-140 A. mellea rhizomorphs which we described from the outside to the inside as follows: (i)

134 an outer layer of peripheral hyphae, (ii) an inner cortical layer of radial hyphae, (iii) a medulla region, consisting of two layers of longitudinal hyphae; an outer layer of thick-walled, narrow lumen, tightly packed hyphae, and inner layer of thin-walled, wide lumen, loosely packed hyphae usually called the vessel hyphae, and (iv) a central cavity. Further, comparison between M. incrassata rhizomorphs and those of A. mellea was another interesting aspect of this study. Scanning electron micrographs of field-collected rhizomorphs of M. incrassata showed less differentiation and organization of constituent hyphae. A. mellea showed well organized rhizomorphs characterized by beautifully arranged hyphal types as described above. M. incrassata and A. mellea both had thick-walled, narrow lumen, tightly packed hyphae arranged longitudinally, and thin-walled, wide lumen, loosely packed vessel hyphae in the central portion. Also, M. incrassata rhizomorphs showed lack of were central cavity, whereas A. mellea were determined to be characterized with a central cavity. These results have undoubtedly provided some insight into the structure of cords and rhizomorph, which in turn helps our understanding of their functions as organs for the translocation and transport of nutrients and water along the length of mycelial cords and rhizomorphs towards their tips. We also agree strongly that the central cavity described from our SEM and those from previous studies (Granlund et al. 1984), is used as air-conducting channels. A close observation suggests that these central cavities may have been generated as a result of the collapse of vessel hyphae, supporting the views of Hartig and van Tubeuf (1902) and Falck (1912).

2.2 In vitro studies of rhizomorph extension in Armillaria mellea

Growth and extension of mycelia is of immense importance to the spread and survival of fungi. Fungal rhizomorphs are specially constructed root-like structures produced by mostly wood-decay and forest-inhabiting pathogenic fungi. Rhizomorphs (i) translocate resources (nutrients, water and oxygen), (ii) allow fungi to migrate from one substrate to another usually over non-nutrient substrates, and (iii) enhance their inoculum potential (Garraway et al. 1991). Armillaria generally has a slow growth rate in soil (Rishbeth, 1968). Factors that affect the growth of rhizomorphs have been studied, and it has been determined that growth of

135 rhizomorphs varies among species with environmental conditions playing immense role (Morrison, 1976, 1982; Rishbeth, 1982). pH, aeration, temperature, moisture, organic matter, and light, have been demonstrated to affect rhizomorph growth and development. Of these, moisture level and substrate temperature are the most extensively studied. For example, Pearce and Malajaczuk (1990) reported that rhizomorphs of A. luteobubalina grew at faster rate on a substrate with a water potential of –0.6 MPa, with no growth at –0.001 MPa. As observed by Garrett (1944), Armillaria root diseases are common in wet soil conditions. Cruickshank et al. (1997) found that saprotrophic Armillaria species are suppressed under dry conditions. We investigated the growth rate of both Armillaria mycelium and rhizomorphs in vitro, studying the stimulatory effects of different agar concentrations on the production of rhizomorphs. Results indicate that rhizomorphs of A. mellea grow faster than mycelia in PDA medium. This seems to suggest the ability of A. mellea to channel resources for the massive production of mycelia from hyphae to form rhizomorphs once nutrients in the substrate are being depleted. Additionally, PD 5-8% agar produced rhizomorphs that extended much faster than those in PD 2% A. Careful observation of extending rhizomorphs in PD 2% (w/v) agar and PD 7% (w/v) agar, with respective gel strengths of 0.01 MPa and 0.08 MPa, indicate that rhizomorphs in the latter were more compact, with more tapered apices than those in the former. This may suggest that, with their tapering apices, rhizomorph tips are able to ultimately function more or less as a wedge they invade they penetrate and invade their substrates. Similar observations have been reported in fungal hyphae where, during extension, the hyphal tip was characterized by a tapering apex, whereas non-extending hyphal tips appeared broad (Harold et al. 1996). The fact that rhizomorphs are able to penetrate media with gel strength of 0.08 MPa underscores their uniqueness in migrating through compact substrates such as soil and wood, and highlights their importance in plant pathogen and wood decomposers. Interestingly, increasing agar concentrations increased the extension rate of individual rhizomorphs and stimulated the production of a greater number of rhizomorphs. Data from the experiments also revealed that rhizomorph length increased with increasing concentration of agar. For example, in PD 8% agar, the average total length of rhizomorphs produced in four 90 mm Petri dishes was recorded to be a staggering 3166 ± 237 mm, compared to 651 ± 22 mm in PD 0.5% agar. This supports earlier reports that hardwood species inoculated with Armillaria isolates produced lengthier and greater dry weight of rhizomorphs than coniferous species

136 (Morrison, 1972). Rishbeth (1972) also showed that English oak was a superior substrate for Armillaria than pine, since the former produced more rhizomorphs, both in number and in biomass. That lengthier rhizomorphs are produced in hardwoods than in pines supports our data that show rhizomorphs in higher concentrations of agar extending faster than those in lower concentrations of agar. This suffices to say that the faster rhizomorphs extend, the lengthier they get, which subsequently produces a larger biomass.

2.3 Osmolyte accumulation in Armillaria mellea rhizomorphs: Its role in turgor pressure generation and tip-growth

Osmolyte biosynthesis, specifically mannitol production has been widely studied in plants, fungi and even animals. Brownlee and Jennings (1981b) had earlier studied the content of soluble carbohydrates and their translocation in S. lacrymans. Analysis of sugars and polyols revealed various carbohydrates in different mycelial regions (margin, submarginal and mid- region) and in the mycelial cords. Notable amongst them were glycerol, mannitol, arabitol, trehalose, glucose, fructose and sucrose. Granlund et al. (1985) studied translocation of solutes along rhizomorphs of A. mellea. Observations indicated that solutes (glycerol, mannitol, arabitol, trehalose, glucose, fructose and sucrose) were translocated bidirectionally along A. mellea rhizomorphs, and at similar velocity in both directions. Erythritol and glycerol were reported to be the major soluble carbohydrates found with maltose, sucrose and trehalose forming a significant part of the total amount of carbohydrates present. In fungal rhizomorphs not much is understood about the role of such osmolytes, specifically in the development and extension mechanism of rhizomorph tips. In our study, we examined the accumulation and composition of osmolyte in A. mellea and their role in the generation of turgor pressure within rhizomorphs by identifying and quantifying osmolytes present in the rhizomorphs, and by osmolality measurement. Results indicated that A. mellea cultured on cellophane covering PD agar and in PD broth accumulated glycerol, erythritol and mannitol in its rhizomorphs. Chromatograms and calculations showed that erythritol and mannitol were the most dominant osmolytes present in A. mellea rhizomorphs. Calculated concentrations of the osmolytes showed that mannitol was the most concentrated osmolyte

137 present in rhizomorphs cultured on cellophane covering PD agar compared to erythritol, although the opposite was true in rhizomorphs cultured in PD broth. Generally, glycerol concentration was far less than erythritol and mannitol under both conditions. This was a surprising outcome considering the fact that glycerol has been reported to generally play an important role in creating osmotic pressure that facilitate uptake of water in fungi (Kelly and Budd, 1991; Dixon et al. 1999). For instance, de Jong et al. (1997) reported that the accumulation of glycerol in the appressoria of the rice blast fungi Magnaporthe grisea provided the needed osmotic force for the penetration of leaf epidermis. The reason for the low levels of glycerol in the rhizomorphs is unclear, especially when Granlund et al. (1985) had reported that glycerol and erythritol were the major osmolytes present in A. mellea rhizomorphs they had worked with, but did not estimate the concentrations of the osmolytes they identified. Although they used GLC to identify their osmolytes, they failed to provide chromatograms and spectra of these osmolytes, for reasons we do not know. Additionally, their claim that glycerol and erythritol were the major osmolytes present was contradicted by the data presented in their report, which show that it was rather erythritol and mannitol that were the major osmolytes translocated. This ambiguity in their report may only be attributed to ineffectiveness of their derivatization methods that possibly affected the reliability of their data (Holligan and Drew, 1971; Brunton et al. 2007). This is why we are confident that our data are more accurate and reliable: we have been able to use quantitative GC/MS to estimate that actual concentrations of these identified osmolytes within these rhizomorphs. In addition, using osmometry, the osmotic pressure of rhizomorph sap was measured to be 0.90 MPa, predicting a maximum turgor pressure of 0.77 MPa for rhizomorphs growing in PD broth, whose osmotic pressure was 0.13 MPa, indicating that the differential in osmotic pressure induces water influx to generate turgor pressure within the rhizomorphs.

2.4 The biomechanics of rhizomorph invasive growth in Armillaria mellea rhizomorphs

138 surrounding agar. The exertion of force by rhizomorphs is evident from the compression and splitting of agar media in advance of their apices. Cultured rhizomorphs extend well beyond the fringe of hyphae that expand from the inoculum, demonstrating that they extend more swiftly than the “unbundled” vegetative hyphae. Many observations suggest that in vitro growth rates reflect the fast extension of rhizomorphs in nature. It is common knowledge that rhizomorphs of wood-decomposing and plant pathogenic fungi have remarkable invasive abilities. However, to our knowledge, no study has been conducted on the source of the strength for rhizomorphs that enable them to push through compacted soils. The closest attempt made was by Coggins et al. (1980) who suggested that the flow of water along the hyphae of mycelial cords in Serpula lacrymans was driven by internal hydrostatic pressure generated by uptake of solutes causing uptake of water into hyphae by osmosis. Jennings and Watkinson (1982) supported this idea and further suggested that hydrostatic pressure in hyphae could lead to the flow of water along the spaces within a mycelial cord. We have been able to identify and quantify three important osmolytes in A. mellea rhizomorphs, and have estimated the percentage of generated turgor that may be exerted at the tips of these rhizomorphs. And for the first time, we are reporting estimated forces that rhizomorphs exert in vitro. Basidiomycetes have been reported to crack open bark of decaying timber, asphalt, and concrete paving through forces during fruiting body expansion (Gooday 1985; Niksic, 2004). The strength of the fruiting bodies originates from hydrostatic pressure. Many decades ago, Buller (1931) estimated that hyphae within an elongating stipe exerted a pressure of 0.07 MPa to raise a weight of more than 200 g. Money and Ravishankar (2005) have measured that a pressure of 0.06 MPa is associated with to stipe elongation in Coprinopsis cinerea fruit bodies. Money (2007) explicitly explains the processes that are involved in generation of hyphal turgor pressure, as a source of force needed by hyphal tips for invasive growth. Bastmeyer et al. (2002) beautifully summarize the whole process this way: “The driving force of hyphal invasion growth is turgor pressure.” Although data from earlier studies provided insight into processes that are involved in hyphal tip growth, there has been little comparable work on rhizomorph development and mechanics. By understanding how hyphae bundle during the formation of these organs, and how they extend in a cooperative fashion, it is possible to elaborate upon the findings made so far on

139 tip growth at the hyphal level. Ultimately, this will allow us to understand how rhizomorphs are able to spread through forest soils. The rhizomorphs of A. mellea strain DR-140 represent an excellent experimental model for this research because (i) they develop quickly and plentifully in standard culture media, (ii) they are composed of extending bundled hyphae, and (iii) they are much larger than hyphae, making them suitable for easy handling, manipulation, and observation during force measurements. Rhizomorphs of other species are also composed of lots of hyphae, but because they do not form in vitro (at least as easily), they don’t lend themselves to these experiments. Strain gauge experiments we conducted have undoubtedly demonstrated that rhizomorph tips exert significant invasive pressure derived from turgor pressure. Applied pressures calculated from measured forces provided in this chapter are consistent with the view that part of the total turgor pressure that is generated in rhizomorphs is used for rhizomorph invasive growth. This is evident from our data in Chapter 4 showing a differential osmotic pressure of 0.77 MPa, which is equivalent in magnitude to the maximum turgor pressure of the rhizomorph.

3. CONCLUSIONS

We have studied rhizomorphs of A. mellea in vitro and present here a complete picture of how rhizomorphs work. It must also be emphasized that comparative work with field-collected rhizomorphs of M. incrassata helped us to understand, for the first time, some anatomical and biochemical aspects of this important wood-decay fungus. We anticipate that this will provide a platform upon which further investigations will be conducted. Experiments designed and conducted with A. mellea rhizomorphs have been very insightful. Enormous amounts of data have been generated as demonstrated in this dissertation. Based on these data, we conclude that:

1. M. incrassata is only able to produce mycelial cords in vitro, and not rhizomorphs. Moreover, field-collected rhizomorphs obtained from Luis De La Cruz in Pasadena, California are those of M. incrassata, as confirmed by comparative molecular studies with M. incrassata strain FP-150521.

140 2. A. mellea rhizomorphs are composed of peripheral hyphae, radial hyphae, two types longitudinal hyphae and a central cavity. The two types of longitudinal hyphae are prominent in the structure of rhizomorphs in both M. incrassata and A. mellea: thick- walled, narrow lumen hyphae and inner thin-walled, wide lumen vessel hyphae. M. incrassata rhizomorphs do not have a conspicuous central cavity, whereas A. mellea rhizomorphs have a central cavity. Well organized tip was prominent in A. mellea rhizomorphs.

3. A. mellea produce rhizomorphs, which extend faster than mycelia in culture. The extent to which rhizomorphs are successful is dependent on the interplay between several environmental factors, with substrate hardness serving as an important stimulatory factor. This is evident from our comparative gel strength experiments, which shows rhizomorphs in PD 7% agar produced lengthier rhizomorphs and extended faster than those in PD 2% agar.

4. A. mellea synthesizes and accumulates osmolytes within its rhizomorphs, notable among them mannitol, erythritol and glycerol. Mannitol and erythritol are the two most dominant osmolytes present in A. mellea rhizomorphs. Accumulated osmolytes within rhizomorphs generate turgor pressure by water influx and a substantial amount of this turgor pressure is used for force exertion at their tips during tip-growth and substrate invasion. Confirmation of this was derived from strain gauge experiment showing a steady drop in force exertion after the addition of 25 mM PEG-6000 to a rhizomorph tip.

5. A. mellea rhizomorphs exert substantial levels of pressure, with a maximum estimation of 0.30 MPa, which is more than the estimated pressure in an automobile tire. These estimated pressures are derived from turgor pressure.

141 4. FUTURE STUDIES

Some aspects of these studies provide us the opportunity to seek answers to other important questions that arose during the course of our investigations. We, therefore, suggest the following areas that need urgent attention in future studies, as we believe A. mellea rhizomorphs are excellent models for such studies:

1. Investigation should be conducted to find out if M. incrassata rhizomorphs possess organized tip growth as seen in A. mellea. This has never been reported in literature. We identified osmolytes (mannitol, erythritol and glycerol) within M. incrassata rhizomorphs and cords with the GC/MS technique. Although this was a preliminary study and was not reported in this dissertation, we suggest that biochemical studies on a full scale be conducted in M. incrassata to afford us the opportunity to quantitatively compare their concentrations and contributions to turgor generation with those in A. mellea. Also, we believe that a consistent effort should be made to develop nutrient medium that has the potential to induce the production of rhizomorphs in M. incrassata. Success with this will be a breakthrough of unprecedented importance in rhizomorph studies.

2. Since data on profiled gene sequence of A. mellea already exists, work to identify genes that control the growth and development of rhizomorphs should be initiated. This can be extended to M. incrassata for comparative purposes. We believe there is a huge potential in this area for understanding the molecular basis of these processes within these wood- decay and plant pathogenic fungi will help design effective control strategies in the future.

3. Further work should focus on the identification of other osmolytes that were not easily identified in this work to complete the list of the osmolyte mixture that is accumulated in A. mellea. Another area of interest to investigate is the metabolic pathways that result in the accumulation of the dominant osmolytes in the rhizomorphs, and their actual physiological roles in the rhizomorphs, considering the two extremes of environment in which our A. mellea rhizomorphs were cultured.

142

4. Finally, we propose that a set of experiments are designed as a crucial step to further understand the biomechanics of rhizomorphs extension, since we believe we have set the stage for more progress to be made in this area by reporting the first successful measurement of forces exerted by rhizomorphs. To this end, future studies should seek to address the following questions:

• How is rhizomorph development initiated from constituent hyphae after they aggregate, and do mechanics play any role in hyphal interlacing; do patterns of cell division lay the foundation for hyphal differentiation as seen in a mature rhizomorph?

• What mathematical models will best explain elongation and branching mechanics, and how do these compare with those of other systems, such as hyphae and roots?

Findings from experiments designed to answer these important questions will be very significant, since they will help to elucidate rhizomorph morphogenesis, and the mechanics involved in invasive growth. This will lead to designing effective control strategies against these phytopathogenic and wood decay basidiomycetes. In conclusion, we highly recommend a multidisciplinary approach in investigating these unsolved questions. This is important since there are obviously biological, molecular physical and mathematical components necessary for the proposed studies.

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