Septal Pore Caps in Basidiomycetes Composition and Ultrastructure

Septal Pore Caps in Basidiomycetes Composition and Ultrastructure

Septumporie-kappen in Basidiomyceten Samenstelling en Ultrastructuur

(met een samenvatting in het Nederlands)

Proefschrift

ter verkrijging van de graad van doctor aan de Universiteit Utrecht op gezag van de rector magnificus, prof.dr. J.C. Stoof, ingevolge het besluit van het college voor promoties in het openbaar te verdedigen op maandag 17 december 2007 des middags te 16.15 uur

door

Kenneth Gregory Anthony van Driel

geboren op 31 oktober 1975 te Terneuzen Promotoren: Prof. dr. A.J. Verkleij Prof. dr. H.A.B. Wösten

Co-promotoren: Dr. T. Boekhout Dr. W.H. Müller voor mijn ouders Cover design by Danny Nooren. Scanning electron micrographs of septal pore caps of Rhizoctonia solani made by Wally Müller.

Printed at Ponsen & Looijen b.v., Wageningen, The Netherlands.

ISBN 978-90-6464-191-6 CONTENTS

Chapter 1 General Introduction 9

Chapter 2 Septal Pore Complex Morphology in the Agaricomycotina 27 (Basidiomycota) with Emphasis on the Cantharellales and Hymenochaetales

Chapter 3 Laser Microdissection of Fungal Septa as Visualized by 63 Scanning Electron Microscopy

Chapter 4 Enrichment of Perforate Septal Pore Caps from the 79 Basidiomycetous Fungus Rhizoctonia solani by Combined Use of French Press, Isopycnic Centrifugation, and Triton X-100

Chapter 5 SPC18, a Novel Septal Pore Cap Protein of Rhizoctonia 95 solani Residing in Septal Pore Caps and Pore-plugs

Chapter 6 Summary and General Discussion 113

Samenvatting 123

Nawoord 129

List of Publications 131

Curriculum vitae 133

Chapter 1

General Introduction

Kenneth G.A. van Driel*, Arend F. van Peer*, Wally H. Müller, Teun Boekhout & Han A.B. Wösten

*Both authors equally contributed to this work Chapter 1

Filamentous Fungi Filamentous fungi grow by means of hyphae that extend at their apices while branching subapically. This mode of growth together with hyphal fusion (anastomosis) results in an interconnected network of hyphae called a mycelium. Hyphae of the lower fungi, i.e. the Glomeromycota, Zygomycota, and Chytridiomycota (Schüßler et al., 2001; Bauer et al., 2006), are sparsely, if at all, septated (Barr, 2001; Benny et al., 2001). In contrast, the hyphae of the filamentousAscomycota and Basidiomycota are regularly septated. Their septa contain pores of about 50 to 500 nm, which are covered with a continuous plasma membrane and allow streaming of cytoplasm including large organelles like mitochondria (Bracker & Butler, 1963, 1964; Gull, 1978; Van Driel et al., 2007). The continuity of the cytoplasm discriminates cells of filamentous fungi from those of plants and animals. In these latter two kingdoms there are also intercellular cytoplasmic connections but they are much smaller. Gap junctions in animals have pores of about 1.6 to 3.0 nm in diameter and allow streaming of inorganic ions and small water-soluble organic molecules (Veenstra, 1996; Perkins et al., 1997). Plasmodesmata in plants are small microchannels of 1.5 to 2.0 nm, restricting also in this case the intercellular flow to small molecules (reviewed by Ghoshroy et al., 1997). It should be noted that the channels in plasmodesmata are dynamic and can be closed or increased in diameter to 5 to 9 nm.

In this chapter the ultrastructure, the composition and the function of fungal septa and their associated organelles (i.e. the Woronin bodies and the septal pore caps) are described. The mechanisms underlying the formation of the septa and the associated organelles are also briefly discussed. This is followed by a description of Rhizoctonia solani (Basidiomycota) as a model system to study septal pore caps in Basidiomycota. At the end of this chapter the aim of this Thesis is described followed by a brief summary of each of the chapters.

The Septum and Woronin Bodies in Filamentous Ascomycota The ascomycetous septum consists of a cross-wall with a central pore (Figure 1A). The septum is tapered towards the pore and is usually referred to as “simple” septum. The diameter of the septal pore varies between 50 to 500 nm allowing passage of mitochondria, nuclei and other organelles (Shatkin & Tatum, 1959; Moore & McAlear, 1962; Gull, 1978). Although the septa are extensions of the lateral cell wall, their chemical compositions differ (Gull, 1978). The septum plate is build up by chitin microfibrils and β-glucans, but α-glucan, which is a component of the lateral wall, is not found (Griffin, 1994). Older septa may be covered with an amorphous protein layer (Gull, 1978).

The ascomycetous septum is associated with Woronin bodies (Figure 1A) that are also found at the hyphal tip (Momany et al., 2002). Woronin first observed Woronin bodies in 1864 by light microscopy (Woronin, 1864; Buller, 1933). They have a spherical

10 General introduction

Figure 1 – Transmission electron micrographs of several septum types that are found in the fungal kingdom. A) Septal pore of Aspergillus nidulans (Ascomycota). The septum is tapered towards the pore, and is also referred as “simple” septum. Woronin bodies are associated at the pore (image is adopted from Momany et al., 2002). B) Septal pore of Sporidiobolus ruineniae (Pucciniomycotina). Septum is tapered towards the pore, but without Woronin bodies (Boekhout et al., 1992). C) Dolipore septum of Itersonilia perplexans (Agaricomycotina) without septal pore caps (SPC). Membrane bands are located between the swollen pore that is covered by endoplasmic reticulum (Boekhout, 1991). D) Dolipore septum (DP) of Rhizoctonia solani (Agaricomycotina) associated with perforate SPCs. A nucleus (Nu) is blocked passage by the pores in the septal pore caps (Müller et al., unpublished). Bars represent 250 nm in A, 500nm in B and D, and 200 nm in C.

or hexagonal shape with a diameter of 150 to 500 nm (Markham & Collinge, 1987). Woronin bodies rapidly plug septal pores when hyphae are damaged to prevent loss of cell content (Trinci & Collinge, 1974). HEX-1 is the main protein of these organelles in Neurospora crassa (Jedd & Chua, 2000). It forms the crystalline core of the Woronin body (Yuan et al., 2003), which is surrounded by a single membrane (Markham & Collinge, 1987). Phosphorylation of HEX-1 is important for multimerization of the protein and proper formation of the Woronin bodies (Juvvadi et al., 2007). The HEX-1 protein has a C-terminal peroxisome targeting signal (PTS1) and therefore Woronin bodies have been suggested to be peroxisomes (Jedd & Chua, 2000; Tenney et al., 2000). HEX-1 homologues have been found in several other filamentous Ascomycota, like Aspergillus nidulans and Magnaporthe grisea (Jedd & Chua, 2000; Soundarajan et al., 2004) but not in ascomycetous yeasts nor in Basidiomycota.

Ultrastructure of the Septum and the Septal Pore Cap in Basidiomycota Within the Basidiomycota, the three major groups Pucciniomycotina (Urediniomycetes; Swann & Taylor, 1995), Ustilaginomycotina (Ustilaginomycetes; Swann & Taylor, 1995), and Agaricomycotina (Hymenomycetes; Swann & Taylor, 1995) are distinguished (Bauer et al., 2006; James, et al., 2006; Hibbett et al., 2007). These groups are amongst others characterized by the ultrastructure of their septum. The Pucciniomycotina contain the rust fungi, which have septa with a pore morphology as found in the filamentous

11 Chapter 1

Ascomycota (Oberwinkler & Bandoni, 1982; Swann et al., 2001; Bauer et al., 2006), though without Woronin bodies (Figure 1B). The Ustilaginomycotina that include the smut fungi also possess a septum with a septal pore that is similar to that found in the hyphae of Ascomycota, but may have a slightly swollen rim around the pore. These septal pores may also be associated with membrane caps or membrane bands (Bauer et al., 1997, 2001). The Agaricomycotina (i.e. Tremellomycetes, Dacrymycetes, and Agaricomycetes) includes jelly fungi and mushroom-forming fungi. They have a barrel-shaped swelling around the pore, the dolipore (Figure 1C, D), which generally is associated with a septal pore cap (SPC) (Figure 1D) (Bracker & Butler, 1963). The SPC is also known as Verschlußband (Girbardt, 1958) or parenthesome (Moore & McAlear, 1962). The dolipore channel is about 70 to 500 nm in diameter (Bracker & Butler, 1964 ; Setliff et al., 1972; Patton & Marchant, 1978), but SPCs that cover the dolipore restrict the passage of large organelles, such as nuclei (Figure 1D).

Like in the Ascomycota, the composition of the basidiomycetous septum is different from that of the lateral cell wall. The lateral cell wall consists of chitin, β-1,3/β-1,6- glucan and α-1,3-glucan. The septal plate contains chitin and β-1,3/β-1,6-glucan but no α-1,3-glucan, whereas the septal swelling contains α-1,3-glucan, β-1,3-glucan, and β- 1,6-glucan (Janszen & Wessels, 1970; Müller et al., 1998a, 2000a). The dolipore swelling contains more β-1,6-glucan than the septal plate (Müller et al., 1998a, 2000a). Staining of the polysaccharides according to Thiéry (1967) showed that filaments of the inner electron dense septal layer radiate into the swelling of the dolipore and form a distinct rim visible in median and traverse sections. This rim intertwines as a loose network of stained fibrous material into the non-stained peripheral part of the septal swelling (Bracker & Butler, 1963; Van der Valk & Wessels, 1976).

At the dolipore septum, several SPC-types can be distinguished, which can be used as a phylogenetic marker (e.g. McLaughlin et al., 1995; Fell et al., 2001; Hibbett & Thorn, 2001; Wells & Bandoni, 2001; Lutzoni et al., 2004). The vesicular (tubular or saccular) SPC-type is found in members of the Tremellomycetes. This SPC-type consists of a group of vesicles or tubules arranged in a hemisphere surrounding the dolipore, e.g. in Trichosporon sporotrichoides (Figure 2A). The imperforate SPC-type is found in the Dacrymycetes and Agaricomycetes (Wells & Bandoni, 2001) and consists of a slightly flattened closed membranous structure, e.g. in Epulorhiza anaticula (Figure 2B). This SPC-type may have an inward growth with reduced thickness or a minute pore in the centre of the membrane (Müller et al., 2000b). These imperforate SPCs are about 270 to 800 nm in width (Patton & Marchant, 1978; Müller et al., 1998b, 2000a). Next to imperforate SPCs, perforate SPCs are found in the Agaricomycetes (Hibbett & Thorn, 2001; Wells & Bandoni, 2001). The perforate SPC-type can have many small perforations like in Schizophyllum commune (Figure 2C) or have few large perforations like in R. solani (Figure 2D). The diameter of

12 General introduction

Figure 2 – Scanning electron micrographs of the main SPC-types found in the Agaricomycotina. A) The vesicular-tubular SPC in Trichosporon sporotrichoides (Tremellales). B) The imperforate SPC in Epulorhiza anaticula (Agaricomycetes). C) The perforate SPC with small perforations in Schizophyllum commune (Agaricomycetes). D) The perforate SPC with few large holes in Rhizoctonia solani (Agaricomycetes). Bars represent 250 nm in A and C, 500 nm in B, and 1000 nm in D. Images are adopted from Müller et al., 1998a (A and C) and Müller et al., 1998b (B and D). the SPCs of S. commune is 450 to 600 nm with perforations of about 100 nm (Müller et al., 1994; 1998a). The SPCs of R. solani that have a diameter of 1600 to 2000 nm contain only 3 to 5 perforations with a diameter of 800 nm (Müller et al., 2000a). Pores are larger at the apex than at the base, as found in S. commune (Müller et al., 1994, 1999).

The SPC is a layered structure that has an inner and outer membrane confining a matrix (Girbardt, 1958, 1961; Moore & McAlear, 1962; Bracker & Butler, 1963; Marchant & Wessels, 1973; Müller et al., 1998a, 2000a). The different SPC-types are connected at their base to endoplasmic reticulum (ER) (Girbardt, 1961; Moore, 1975; Müller et al., 1998a, 2000b). This is especially clear in young hyphae. In older cells, the ER has degenerated and the connection is lost. Next to this basal connection, ER is often observed overlaying imperforate SPCs as reported in Cyclomyces fuscus, Coltricia perennis (Müller et al., 2000b), Sebacina vermifera (Currah & Sherburne, 1992; Müller et al., 1998b), Basidiodendron cinereum, Tremellodendropsis tuberosa, and Botryobasidium vagum (Wells, 1994; Langer, 1994). The outer cap observed in the case of perforate SPCs in agaricoid

13 Chapter 1 fungi like Agaricus bisporus (Thielke, 1972; Craig et al., 1977), Coprinus cinereus (Van der Valk & Marchant, 1978), and Agrocybe praecox (Gull, 1976) may be similar to the ER plates that are associated with the imperforate SPCs. The presence of the outer cap in these perforate SPCs may depend on the developmental stage of the hyphal cells as it occurred at dolipore septa in subhymenial cells (Gull, 1976; Craig et al., 1977).

As the base of SPCs is continuous with the ER, SPCs are regarded as subdomains of this organelle (Wilsenach & Kessel, 1965; Müller et al., 1998a, 2000a). Fluorescent markers that stain ER, like ER-tracker, DIOC-6, and Brefeldin A conjugated to BODIPY also highlight the SPC region (Chapter 5). However, differences are observed with a zinc- iodine osmium tetroxide (ZIO) staining that marks calcium-affinity sites (Gilloteaux & Naud, 1979; Müller et al., 1998a, 1999, 2000a). The SPCs of T. sporotrichoides were as densely stained as the ER. In contrast, perforate SPCs of S. commune and R. solani showed staining only of the SPC inner and outer membranes (Müller et al., 1998a, 1999, 2000a). This suggests that there are no structural or functional differences between ER and the vesicular-tubuar SPC-type in T. sporotrichoides (Müller et al., 1998a), while this may be the case in perforate SPCs of S. commune and R. solani.

Formation of Septa Formation of septa in filamentous fungi takes place in germinating spores (Jersild et al., 1967; Niederpruem & Jersild, 1972), within hyphal compartments (Niederpruem & Jersild, 1968; Volz & Niederpruem, 1968; Niederpruem, 1971), after branching (Jersild et al., 1967), nuclear translocation (Giesy & Day, 1965), and clamp formation (Jersild et al., 1967; Niederpruem, 1971) and during basidiospore formation (Wells, 1965). Formation of septa has been studied in quite some detail in the yeasts Saccharomyces cerevisiae and Schizosaccharomyces pombe (Forsburg & Nurse, 1991; Morris & Enos, 1992; Sipiczki, 2007). It should be noted that these yeasts display complete cytokinesis whereas filamentous fungi remain coenocytic (Harris et al., 1994). Yet, the yeast models, together with recent data on Aspergillus sp. provide a strong framework to describe septum formation in the Ascomycota (Doonan, 1992; Harris, 2001; Walther & Wendland, 2003). Little is known about the mechanism in the Basidiomycota but it may be similar to that in the Ascomycota.

Analysis of temperature sensitive A. nidulans mutants that are defective in septum synthesis showed that sites for septum formation are established during growth (Morris, 1975; Trinci & Morris, 1979). Harris and Momany (2004) proposed three models for septum localization in filamentous fungi. Firstly, by cortical landmarks that gather at specific sites of the cell cortex that also occurs in yeast. Secondly, by activation of specific cortical receptors (located in the cell cortex) at the future septum site, and thirdly, by stochastic signal fluctuation (Harris & Momany, 2004). The strict organization

14 General introduction and regular distribution of septa makes the third model the least probable. In yeast, cortical landmarks direct the formation of a medial ring. Early in mitosis the medial ring is placed overlying the nucleus marking the start site for septum synthesis. It is still unclear whether the cortical landmarks that direct the site for septum synthesis are present at their site prior to mitosis or that they are formed during mitosis, e.g. by the mitotic machinery (reviewed by Chang et al., 1996; Chang & Nurse, 1996). The septum in filamentous fungi also develops at the site that has previously been occupied by the dividing nuclei (Jersild et al., 1967; Trinci, 1978), in particularly at the site occupied during the metaphase (Bourett & McLaughlin, 1986). This correlation between nuclear division and septum formation is also observed in the Basidiomycota (Bourett & McLaughlin, 1986).

The yeast medial ring is a very complex structure that includes actin, type II myosin, profilin, tropomyosin, a myosin light chain-like protein, a formin, an SH3 domain containing protein and the product of the rng2 gene (for references see Gould & Simanis, 1997). Synthesis of the septum is initiated at the end of the anaphase, which starts at the cortex and is directed inwards to the centre of the cell (Johnson et al., 1973). The medial ring constricts the cell membrane (Girbardt, 1979; Marks & Hyams, 1985; Jochová et al., 1991; Fankhauser et al., 1995; McCollum et al., 1995; Chang et al., 1997; Kitayama et al., 1997) and the developing septum is surrounded by F-actin patches (Marks & Hyams, 1985). The polarization of such patches adjacent to the ring is obligate for septum formation (Gould & Simanis, 1997). The leading edge of the developing septum also contains a ring of F-actin (Jochová et al., 1991).

In filamentous fungi the plasma membrane is also furrowed at the site of the emerging septum (reviewed by Wendland & Walther, 2006). The septal belt (Patton & Marchant, 1978; Girbardt, 1979; Hoch & Howard, 1981; Roberson, 1992) pulls the plasmamembrane together with radial and parallel filaments (Orlovich & Ashford, 1994). This belt has been shown to contain actin microfilaments (Harris et al., 1994; and references herein) and might be functionally equivalent to the medial ring in yeast. This mechanical contraction is also observed in the contractile actin ring of animal cells (Jochová et al., 1991) suggesting a highly conserved mechanism. More detailed information on the septal band of filamentous fungi has been obtained in Aspergillus sp. After SepA (a formin) localizes to the septation site, it co-assembles with AspB (a septin) and actin to form the septal band. At this point the daughter nuclei have undergone mitotic exit. The AspB ring then splits into two rings, that flank the actin and SepA rings. Then the SepA ring and the actin ring constrict and septum deposition takes place (reviewed by Harris, 2001).

15 Chapter 1

Formation of the Dolipore Septum and Septal Pore Caps The dolipore is formed late during synthesis of the basidiomycetous septum. After the septum has been expanded to its near final size, lamellae of the septum branch off at the pore rim and extend into the matrix of the forming dolipore (Bracker & Butler, 1963). The swelling is then enlarged. Growth of the swelling takes place at both sides of the septum and in the middle of the dolipore channel (Moore, 1975). Little is known about the formation of SPCs. Based on electron microscopy, Moore proposed a model for the synthesis of these structures (Moore, 1975). In this model ER sheaths would align in a parallel orientation and aggregate into an irregularly shaped structure. This structure would develop into two SPCs that stay attached during their growth process. After separation, they would be positioned at the dolipore septum (Moore, 1975; Orlovich & Ashford, 1994). Positioning of the SPCs at the septum supposedly takes place at the final stage of the septum formation process. Alternatively, SPCs may originate from a microfilament-based matrix situated at the developing dolipore instead of ER (Patton & Marchant, 1978). This matrix may contain phospholipids and proteins. Only at the completion of SPC formation, laying down of the ER along the septum takes place where after the ER is attached to the SPC (Patton & Marchant, 1978). It should be noted that studies done so far were based on chemical fixation that could have induced artificial side effects. The morphologically highly reliable cryo-preparation techniques such as high-pressure freezing combined with freeze-substitution (Müller et al., 2000a) could reveal additional details on dolipore and SPC formation.

Functions of the Fungal Septum and Septum-Associated Structures The role of the septa in the fungal mycelium seems diverse. These cross walls may mechanically strengthen hyphae, but this is probably of secondary importance (Gull, 1978). Another role of the septa is compartmentalization of the hyphae into cellular compartments. As in regular cell division, distribution of cell constituents upon septation is not random, at least in the Basidiomycota with complex hyphae. In the Ascomycota, the maintenance of distribution of organelles over compartments seems not controlled. Nuclei, mitochondria and other particles often traverse the septum (Shatkin & Tatum, 1959; Moore & McAlear, 1962; Hunsley & Gooday, 1974). In contrast, in the higher Basidiomycota (i.e. Agaricomycotina) the dolipore septum with the SPC is thought to prevent migration of nuclei and other organelles, though mitochondria can be observed traversing the dolipore septum (Bracker & Butler, 1963, 1964; Van Driel et al., 2007).

In general, the dolipore septa have to be reorganized to simple septa to allow organelle redistribution (Bracker & Butler, 1964; Giesy & Day, 1965; Wessels & Marchant, 1974; Todd & Aylmore, 1985). This occurs for instance during the nuclear exchange that follows dikaryotization (Jersild et al., 1967; Koltin & Flexer, 1969; Niederpruem & Wessels, 1969).

16 General introduction

Degradation of a septum starts at the dolipore that initially loses its membrane integrity (Giesy & Day, 1965; Marchant & Wessels, 1973; Mayfield, 1973). The dolipore swelling is progressively degraded until it is completely removed. Septal degradation has been reported to be associated with an invasion of vesicles and multi-vesicular bodies into the septum region (Marchant & Wessels, 1973; Mayfield, 1973) that are thought to contain degrading enzymes (Marchant & Wessels, 1974). The degradation process is directed from the septal pore to the hyphal wall. Degradation continues until a septum- base is left. The fate of the SPC during this process is obscure. It is removed from the septum region but its place in the degradation sequence is not known. Some reports state that the SPC is degraded even before the initial breakdown of the dolipore (Giesy & Day, 1965). Others show that the SPC can be found floating freely in the cytoplasm (Jersild et al., 1967; Marchant & Wessels, 1974).

Apart from functioning as a sieve for organelles, septa can be completely closed sealing the compartment. Closure of the septal pore is observed during heterokaryon incompatibility (HI) and during differentiation of mycelium (Morris, 1975; Gull, 1978; Glass & Kaneko, 2003). In the event of the incompatible fusion of two hyphae the four bordering septa of the fusion cell are plugged. The fusion cell is then lysed by an apoptosis-like process whereas the adjacent compartments remain intact (Glass & Kaneko, 2003). The function of septal closure in development is indicated from the fact that mutations in septum formation often hamper development of fungi (Morris, 1975; Gull, 1978). Another main function for closure of septal pores is to prevent lysis of the mycelium as a response to hyphal damage (Markham, 1994). Septal pores in ascomycetous hyphae are plugged by Woronin bodies (see above). Emergency plugging of the septal pores in hyphae of the Basidiomycota follows a different mechanism. Aylmore et al. (1984) suggested that septal sealing in the Basidiomycota is a two-stage process. First, an electron-dense pore plug is formed. This plug of unknown composition is formed in an instant and restricts the dolipore entrance from each side of the disrupted compartment (Moore & McAlear, 1962; Bracker & Butler, 1963; Koltin & Flexer, 1969; Casselton, 1971; Moore & Marchant, 1972; Setliff et al., 1972; Craig et al., 1977). Then the plug is extended into the dolipore channel further sealing the opening between two neighboring hyphal compartments, but usually leaving the inside of the plug translucent (Aylmore et al., 1984; Müller et al., 2000a). The plug material can be degraded by trypsin and chymotrypsin, showing that it is composed, at least partially, of protein (Flegler et al., 1976). Polysaccharide staining of the plug material did not give consistent results as staining according to Thiéry (1967) did not stain the plug material (Flegler et al., 1976) whereas alkaline bismuth polysaccharide staining did (Shinji et al., 1975, 1976; Müller et al., 1998a). In addition, β-1,6-glucan was not detected in the plug material of S. commune, whereas β-1,6-glucan was present in plugs of T. sporotrichoides (Müller et al., 1998a).

17 Chapter 1

Initiation of the immediate response is also observed in adjacent septa of the damaged compartment, but plugging here is aborted and eventually reversed (Aylmore et al., 1984). In young hyphal cells plugging is also thought to be reversible, while in mature cells the plug is more permanent (Bracker, 1967). Both the dolipore and SPCs have been proposed to participate in the plugging process. The dolipore could function as an acceptor for plugging material (Bracker & Butler, 1963; Setliff et al., 1972) whereas the SPC has been suggested to act as a repository for plugging material (Moore, 1985; Markham, 1994; Müller et al. 1998a, 2000a). The localization of the SPC in close vicinity of the dolipore would support this view as it enables a quick delivery of plugging material to the pore. Moreover, filamentous structures have been observed between the plug and the SPC in R. solani (Müller et al., 2000a; Van Driel et al., 2007), Pisolithus arhizus (Orlovich & Ashford, 1994), and S. commune (Müller et al., 1998a). They could be involved in transport of the plugging material from the SPC to the pore, function as an anchor to keep the plug in its position, or function as a scaffold for plug formation. It should be noted, however, that Basidiomycota, which do not have SPCs at their dolipore (i.e. Pucciniomycotina, Ustilaginomycotina) also plug the septal pore (Aylmore et al., 1984).

Rhizoctonia solani as a Model for Studies on the Septal Pore Cap The plant pathogen Rhizoctonia solani Kühn (1858) affects economically important crops worldwide, like rice (Johanson et al., 1998), potato (Ceresini et al., 2002a, b), soy bean (Naito et al., 1993; Fenille et al., 2002), tobacco (Ceresini et al., 2002a), and sugar beet (O’Sullivan and Kavanagh, 1991). Rhizoctonia solani is a soil-borne plant pathogen and its sclerotia (Tu & Kimbrough, 1975), aggregations of thick-walled melanized cells, are the primary survival structures and source of infection. Rhizoctonia solani is associated with the teleomorph Thanatephorus cucumeris (Frank) Donk (Talbot, 1970; Tu & Kimbrough, 1978) and is considered to be a species complex, Rhizoctonia sensu lato (s.l.) (Adams, 1988; Farr et al., 1989; Sneh et al., 1991). Next to many plant pathogens, most orchid mycorrhizal fungi have been assigned to the Rhizoctonia s.l. complex (Sneh et al., 1991; Andersen & Rasmussen, 1996; Shan et al., 2002). The strains from the Rhizoctonia species complex are multinucleate or binucleate and are subdivided into several anastomosis groups that group closely related isolates capable to form hyphal anastomosis reactions (hyphal fusion) (Parmeter et al., 1969; Ogoshi, 1987; Cubeta & Vilgalys, 1997). The teleomorphs that are connected with the Rhizoctonia s.l. complex are amongst others Ceratobasidium, Thanatephorus, Uthatobasidium, Tulasnella, Sebacina and Waitea (Kataria & Hoffmann, 1988; Stalpers & Andersen, 1996).

The SPC ultrastructure of R. solani has been studied by transmission electron microscopy (Bracker & Butler, 1963, 1964; Setliff et al., 1972; Müller et al., 2000a), scanning electron microscopy (Lisker et al., 1975; Müller et al., 1998b, 2000a) and automated electron tomography (Müller et al., 2000a). The Rhizoctonia s.l. complex contains strains with

18 General introduction dolipore septa associated with imperforate SPCs (Tulasnella, Sebacina), perforate SPCs with small perforations (Ceratobasidium, Waitea) and perforate SPCs with large perforations (Thanatephorus, R. solani) (Andersen, 1996; Müller et al., 1998b). As a model to study SPCs, we choose the multinucleate strain R. solani (CBS 346.84), which belongs to AG-group 10 as was determined by phylogenetic analyses of ITS sequences of Rhizoctonia species (Nakatani, 2006). This strain has broad hyphae with dolipore septa associated with large SPCs (Müller et al., 2000a). Furthermore, the extensively studied SPC ultrastructure of R. solani (see above) supports our research with many ultrastructural data.

Aim and Outline of this Thesis In this Thesis the SPCs from Rhizoctonia solani were studied (Chapter 3 - 5). Furthermore, as SPCs are important taxonomic characters, the known SPC ultrastructure of several basidiomycetous fungi was related with the present fungal classification Chapter( 2). In addition, the SPC ultrastructure of Rickenella fibula and Cantharellus formosus were examined, as their SPC ultrastructure was unclear. In Chapter 3 and Chapter 4 two isolation methods are described that can be used to study the structural components of the septal region in R. solani. The first method (Chapter 3) uses the PALM microbeam system to isolate septal regions from sectioned hyphae by laser microdissection. The second method (Chapter 4) uses a combination of French press, isopycnic centrifugation, and Triton X-100 treatment to enrich SPCs. In Chapter 5, the SPC protein SPC18 was identified that localized to SPCs and pore-plug material. The results are summarized and discussed in Chapter 6.

19 Chapter 1 REFERENCES

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Chapter 2

Septal Pore Complex Morphology in the Agaricomycotina (Basidiomycota) with Emphasis on the Cantharellales and Hymenochaetales

Kenneth G.A. van Driel, Bruno M. Humbel, Arie J. Verkleij, Joost Stalpers, Wally Müller & Teun Boekhout Chapter 2

ABSTRACT

The ultrastructure of septa and septum-associated septal pore caps are important taxonomic markers in the Agaricomycotina (Basidiomycota, Fungi). The septal pore caps covering the typical basidiomycetous dolipore septum are distinguished into three main morphotypes: vesicular, imperforate, and perforate. Until recently, the septal pore cap-type reflected the higher-order relationships within the Agaricomycotina. However, the new classification of Fungi resulted in many changes including addition of new orders. Therefore, the septal pore cap ultrastructure of more than 350 species as reported in literature was related to this new classification. In addition, the septal pore cap ultrastructure of Rickenella fibula and Cantharellus formosus was examined by transmission electron microscopy. Both fungi were shown to have dolipore septa associated with perforate septal pore caps. These results combined with data from the literature show that the septal pore cap type within orders of the Agaricomycotina is generally monomorphic, except for the Cantharellales and Hymenochaetales.

INTRODUCTION

Morphology of for example fruiting bodies (e.g. Fries, 1874; Patouillard, 1900; Fennel, 1973; Müller & Von Arx, 1973; Jülich, 1981; Berbee & Taylor, 1992), basidia (e.g. Martin, 1957; Donk, 1958; Talbot, 1973), spindle pole bodies (SPB) (e.g. McLaughlin et al., 1995; Celio et al., 2006), and septa (e.g. Moore, 1980, 1985, 1996; Khan & Kimbrough, 1982; Oberwinkler & Bandoni, 1982; Kimbrough, 1994; Wells, 1994; McLaughlin et al., 1995; Bauer et al., 1997; Müller et al., 2000b; Hibbett & Thorn, 2001) as well as physiological and biochemical characteristics (Bartnicki-Garcia, 1968; Van der Walt & Yarrow, 1984; Prillinger et al., 1993; Kurtzman & Fell, 1998; Boekhout & Guého, 2002) have strongly contributed to fungal systematics. The structural and biochemical database for fungi (Celio et al., 2006) aims to capture several of these characters in a comprehensive manner. Next to these morphological and physiological characteristics, sequence data from ribosomal DNA (i.e. nSSU and nLSU rDNA), mitochondrial DNA and protein coding genes (e.g. EF1, RPB1, RPB2) have been instrumental in fungal systematics (e.g. Swann & Taylor, 1993, 1995; Liu, 1999, 2006; Fell et al. 2000; Schüßler et al., 2001; Lutzoni et al., 2004; Tanabe et al., 2004). More recently, complete fungal genomes were used in phylogeny (phylogenomics) and revealed consistency with the molecular studies done so far (Fitzpatrick et al., 2006; Kuramae et al., 2006). Collaborations between fungal systematics (AFTOL/Deep Hyphae) have increased the resolution of the fungal tree of life that resulted in an upgraded classification of the Fungi (James et al., 2006; Hibbett et al., 2007).

28 Septal pore complex morphology in the Agaricomycotina

Since the last overview of septal ultrastructure in relation with fungal phylogeny (Fell et al., 2001; Hibbett & Thorn, 2001; Wells & Bandoni, 2001) many new orders have been proposed in the Agaricomycotina (equivalent to Hymenomycetes; Swann & Taylor, 1995) (Larsson et al., 2004; Binder et al., 2005; Hosaka et al., 2006; Hibbett et al., 2007) and the fundamental distinction between Heterobasidiomycetes and Homobasidiomycetes has disappeared. At present the Agaricomycotina contains three main clades, namely the Tremellomycetes, the Dacrymycetes, and the Agaricomycetes and 21 orders are recognized (Hibbett, 2006; Hibbett et al., 2007). In general, members of the Agaricomycotina have a dolipore septum that is flared towards the pore and may be associated with septal pore caps (SPCs) (Girbardt, 1958; Moore & McAlear, 1962; Bracker & Butler, 1963; Müller et al., 1998a, 2000b). These SPCs are distinguished into three main morphotypes: the vesicular (tubular, saccular), the imperforate (continuous) and the perforate SPC-type.

The ultrastructure of the septum and septum-associated subcellular structures reflected the higher-order relationships within the Agaricomycotina, and until recently, the orders herein contained only one SPC-type, either vesicular, imperforate, or perforate (e.g. Wells, 1994; Müller et al., 1998b, 2000b; Fell et al. 2001; Hibbett & Thorn, 2001; Wells & Bandoni, 2001). However, the basic changes inferred by molecular data necessitated a reconsideration of the septal ultrastructure in relation with the new classification. Furthermore, the orders Cantharellales and Hymenochaetales both were considered having only imperforate SPCs (Hibbett & Thorn, 2001), but at present these orders probably include also members with perforate SPCs (Larsson et al., 2006; Moncalvo et al., 2006). Into the Cantharellales the Ceratobasidiales were placed, to which, Thanatephorus, Uthatobasidium and Ceratobasidium belong that all have perforate SPCs (Bracker & Butler, 1963; Lisker et al., 1975; Tu et al., 1977; Langer, 1994; Andersen, 1996; Müller et al., 1998b, 2000a; Moncalvo et al., 2006). Moreover, the position of Cantharellus itself is unclear, as it has been reported to contain perforate SPCs (Keller, 1997) as well as imperforate SPCs (Hibbett & Thorn, 2001; Larsson et al., 2004; Moncalvo et al., 2006). Hyphoderma praetermissum with perforate SPCs (Langer & Oberwinkler, 1993; Keller, 1997) is now classified in the Hymenochaetales (Larsson et al., 2004, 2006). Finally, the omphalinoid fungi that previously were classified in the Agaricales (Singer, 1986) revealed tobe polyphyletic and a biotrophic group, including Rickenella fibula (Bull.) Raitelhuber (1973), was placed in the Hymenochaetales (Moncalvo et al., 2002; Redhead et al., 2002; Larsson et al., 2004, 2006).

Here, SPC ultrastructural data from the literature was related with the recently proposed classification of the Agaricomycotina. Moreover, the SPC ultrastructure of Cantharellus formosus and R. fibula was examined by transmission electron microscopy. It is concluded that the SPC-type within the orders of the Agaricomycotina is generally monomorphic, except for the Cantharellales and Hymenochaetales.

29 Chapter 2

MATERIALS & METHODS

Strain, Media, and Culture Conditions Rickenella fibula (CBS 116393) was grown on X-agar medium (110 ml cherry extract, 600 ml pepton-glucose-saccharose, 600 ml oatmeal extract, 480 ml water, and 25 g agar; Gams et al., 1998) at room temperature. After 5 weeks a colony with a diameter of about 1 cm was used for chemical fixation and high pressure freezing. Cantharellus formosus was obtained from a commercial source. The identity of both isolates was checked by sequence analyses of the internal transcribed spacers (ITS) 1 and 2, and the D1/D2 region of the nuclear large subunit (nLSU) ribosomal DNA using standard primers, PCR and sequence conditions (White et al., 1990; Hopple & Vilgalys, 1999).

Chemical Fixation Peripheral parts of the R. fibula colony of about 34 mm, and approximately 1 mm tissue blocks from the stipe and the cap of C. formosus were cut. The mycelium was chemically fixed in freshly prepared ice-cold 1% (w/v) aqueous potassium permanganate for20 min on ice. After rinsing with ice-cold distilled water, the mycelium was dehydrated in a series of 70%, 80%, 90%, 95% and 100% (v/v) ethanol on ice. Subsequently, the ethanol was replaced by 1,2-propylene oxide (Merck KGaA, Darmstadt, Germany) (25%, 50%, 75%, and 100%) and the fungal cells were infiltrated (25%, 50%, 75%, 100%) and embedded in Spurr’s resin (Spurr, 1969), which was polymerized at 65ºC for 2 days.

High-pressure Freezing and Freeze-substitution From the periphery of the R. fibula colony, pieces of about 3 mm in diameter were cut and sandwiched between aluminum planchettes (Engineering Office M. Wohlwend GMbH, Sennwald, Switzerland), which were filled with 1-hexadecene (Müller & Moor, 1984; Studer et al., 1995) and subsequently high-pressure frozen with a Leica EM HPF (Leica Microsystems, Vienna, Austria) according to the supplier’s manual. After freezing the sandwich, it was put into liquid nitrogen and the two aluminum planchettes were separated. The excess of 1-hexadecene was removed by gently scratching the surface of the hyphae with a fine needle in liquid nitrogen (Müller et al., 2002). The fungal cells with the supporting planchette were transferred in liquid nitrogen to a CS auto freeze- substitution apparatus (Reichert-Jung, Vienna, Austria). In the substitution chamber the frozen fungal cells were rapidly put into the freeze-substitution fluid, containing

1% OsO4, 3% glutaraldehyde (EM grade, Polysciences Inc, Warrington, PA, USA), and 0.3% uranylacetate (Merck) in anhydrous methanol (Merck) (modified from Müller et al., 1980). Fungal cells were freeze-substituted for 4.5 days at -85ºC, after which the temperature was gradually raised (3ºC per hr) to 0ºC. Vials containing the freeze- substituted fungal cells were put on ice. After 1 hr the fungal cells were rinsed with anhydrous methanol, followed by anhydrous acetone. After rinsing, they were infiltrated

30 Septal pore complex morphology in the Agaricomycotina and embedded in Spurr’s resin, and polymerized as described above.

Transmission Electron Microscopy Sections of 90 nm and 300 nm were post-contrasted with 4% (w/v) aqueous uranylacetate (Merck) and 0.4% (w/v) aqueous lead citrate (Merck) (Venable & Coggeshall, 1965) and viewed in a TECNAI 10 transmission electron microscope (FEI Company, Eindhoven, The Netherlands) at an acceleration voltage of 100 kV.

RESULTS & DISCUSSION

Septal Pore Cap Ultrastructure of Rickenella fibula and Cantharellus formosus Rickenella fibula is a small gilled mushroom commonly found between moss (Bas et al., 1995) and strongly suspected to be biotrophic (Redhead, 1981; Kost, 1984). It was previously classified in the family Tricholomatacea within the order Agaricales (Singer, 1986). Sections of chemically fixed R. fibula hyphal cells revealed a dolipore septum associated with perforate septal pore caps (SPCs) (Figure 1A), which corresponds with previous observations in R. aulacomniophila (= R. fibula; Kost, 1984). SPCs had a width of about 300 to 400 nm, a height of about 180 nm, and small perforations of about 50 to 60 nm in diameter. The SPCs of R. fibula were comparable to those observed in Oxyporus latemarginatus (cited as Poria latemarginata; Setliff et al., 1972). The base of the SPC was connected with endoplasmic reticulum (ER) (Figure 1A), supporting previous views that the SPC is a subdomain of the ER (Girbardt, 1961; Bracker & Butler, 1963; Müller et al., 1995, 1998a; Chapter 5.). Sections of high-pressure frozen (HPF) and freeze-substituted hyphal cells of R. fibula confirmed the presence of perforate SPCs at the dolipore septum (Figure 1B). In these hyphal cells the SPC had a width of about 320 to 400 nm at its base, a height of about 200 nm, and perforations of about 50 to 60 nm. In some cells perforations of about 80 nm were found. Cryo-fixation by HPF confirmed the results obtained by chemical fixation, but gave a more detailed view of the SPC membranes and plug morphology. The SPC existed of an inner and an outer membrane enclosing the SPC matrix with an electron-dense layer in the centre (result not shown). Filamentous structures connected the inside of the SPC with the pore-occluding material as was reported previously in Schizophyllum commune (Müller et al., 1998a) and Rhizoctonia solani (Müller et al., 2000a; Van Driel et al., 2007).

Sections of chemically fixed mycelium of Cantharellus formosus revealed a dolipore septum associated with perforate SPCs (Figure 2). Tissue from both stipe and hymenophore were analyzed. Stipe tissue revealed few dolipore septa and SPCs were often degenerated, while tissue from the hymenophore gave intact SPCs. Sections

31 Chapter 2 showed that the SPCs were about 630 to 810 nm in diameter with perforations of about 100 to 200 nm (Figure 2). SPCs of C. formosus were comparable to SPCs observed in Ceratobasidium cornigerum (Müller et al., 1998b). ER membrane covering the SPC and forming an outercap region was observed (Figure 2) as previously reported in other fungi (Thielke, 1972; Gull, 1976; Craig et al., 1977; Van der Valk & Marchant, 1978; Desole, 1982).

Figure 1 – Transmission electron micrographs of the dolipore-septal pore cap (SPC) complex in Rickenella fibula after chemical fixation (A) and after high-pressure freezing and freeze substitution (B). The dolipore (DP) septum is covered with perforate SPCs. The SPCs in Figure B are near median cut and tangentially cut, the latter showing the surface view. Bars represent 200 nm.

Figure 2 – Transmission electron micrographs of the dolipore-septal pore cap (SPC) complex in chemically fixed hyphae of Cantharellus formosus. The dolipore (DP) is covered with SPCs. Arrows indicate the membrane that forms an outer cap region above the SPC, which may be endoplasmic reticulum. Figure B is a magnification of Figure A. Bars represent 250 nm.

32 Septal pore complex morphology in the Agaricomycotina

Septal Pore Cap Morphology in the Agaricomycotina According to the current classification, the Agaricomycotina contains three classes (Tremellomycetes, Dacrymycetes, and Agaricomycetes) and 21 orders (Hibbett, 2006; Hibbett et al., 2007). The SPC ultrastructure of more than 350 species has been published (Appendix, page 42). Table 1 shows a summary of the Appendix by giving the SPC-type per order. The current use of species names was checked in Mycobank (www.mycobank. org; Crous et al., 2004). In the Tremellomycetes, the SPC is absent (Cystofilobasidiales) or has the vesicular morphology (Filobasidiales, Trichosporonales, Tremellales) (Table 1). The Dacrymycetes (Dacrymycetales) contains only species with imperforate SPCs (Table 1). The previously recognized clades that now belong to the Agaricomycetes contained either the imperforate SPC-type (Tulasnellales, Auriculariales, Hymenochaetoid, and Cantharelloid clade) or the perforate SPC-type (Polyporoid, Euagarics, Bolete, Thelephoroid, and Russuloid clade) (Hibbett & Thorn, 2001; Wells & Bandoni, 2001), with the exception of the gomphoid-phalloid clade that contained both perforate and imperforate SPCs

Class Subclass Order SPC-type

Tremellomycetes Cystofilobasidiales absent Tremellales absent or vesicular Trichosporonales absent or vesicular Filobasidiales absent or vesicular

Dacrymycetes Dacrymycetales imperforate

Agaricomycetes Sebacinales imperforate Cantharellales perforate and imperforate Auriculariales imperforate Phallomycetidae Geastrales imperforate * Phallomycetidae Hysterangiales unknown Phallomycetidae Phallales perforate ** Phallomycetidae Gomphales imperforate * Trechisporales imperforate * Hymenochaetales imperforate and perforate Thelephorales perforate Polyporales perforate Gloeophylalles perforate ** Corticiales perforate Russulales perforate Agaricomycetidae Agaricales perforate Agaricomycetidae Boletales perforate Agaricomycetidae Atheliales perforate

Table 1 – SPC-type per order level in the Agaricomycotina (summary of the Appendix). The SPC-type in Hysterangiales is unknown as no SPC ultrastructure has been published. * SPC-type determined in one species. ** SPC-type determined in two species.

33 Chapter 2

(Hibbett & Thorn, 2001). However, the SPC-type of the latter clade was unclear, as only few taxa were included. Present classification combined with SPC morphology data shows that the orders in the Agaricomycetes have in general only one SPC-type. The imperforate SPC-type is found in the Geastrales, Gomphales, Trechisporales, Auriculariales, and Sebacinales (Table 1). The perforate SPC-type is found in the Agaricales, Atheliales, Boletales, Phallales, Corticiales, Gloeophyllales, Polyporales, Russulales, and Thelephorales (Table 1). However, both perforate and imperforate SPCs are found in the Cantharellales and Hymenochaetales (Table 1). The SPC-type for members of the Hysterangiales has not been determined yet. Furthermore, the SPC-type in the Trechisporales, Geastrales, and Gomphales was examined only in one species, whereas the SPC-type in Gloeophyllales and Phallales was examined in two species. For these orders, more data on the SPC ultrastructure are required to allow reliable statements concerning their SPC-type. An overview of the SPC-type in relation with the current tree topology of the Agaricomycotina (Hibbett, 2006) is given in Figure 3.

The descriptions of the SPC-type of Typhula uncialis, Bolbitius vitellinus, Plicatura nivea, Basidiodendron rimulentum, Phanerochaete sordida, Tremella encephala, Trechispora subsphaerospora, Hydnocristella himantia (Keller, 1997), Auricularia polytricha, A. mesenterica (Patton & Marchant, 1978), and Coltricia perennis (Moore, 1980) were not included in this study as either the images were of suboptimal quality and could be interpreted differently, or the material was misidentified. Furthermore, few irregularities on the SPC-type were found in the Agaricales (i.e. Lepista glaucocana, Mycena galopus, and Radulomyces confluens), the Russulales (i.e. Scytinostromella olivaceoalba), and the Tremellales (Ditangifibulae dikaryotae) suggesting that the SPC-type in these orders is not monomorphic (Appendix). However, as misidentifications were made in the past, these anomalies should be confirmed or supported by genetic data (e.g. ITS or nLSU sequence data) and high-quality images of the dolipore-SPC complex, for example, obtained after high-pressure freezing and freeze-substitution. However, a recent study of the SPC ultrastructure in two species of Mycena, showed perforate SPCs in M. hiemalis, while M. galopus has imperforate SPCs (Rexer & Stepanova, 2004). A reversal from perforate to imperforate SPC-type could have taken place in this genus, which would suggest that perforate SPCs might not be morphologically stable. Nevertheless, this is the only reported anomaly within a genus so far. In addition, the authors suggested that Mycena is heterogeneous (Rexer & Stepanova, 2004).

Septal Pore Cap Morphology in the Hymenochaetales The Hymenochaetales order has six clades: the Oxyporus, Rickenella, Kneiffiella, Hyphodontia, Coltricia, and Hymenochaetaceae clades (Larsson et al., 2006). The SPC ultrastructure is known for many of its members. Imperforate SPCs have been found in Inonotus, Hymenochaete, Hydnochaete, Phellinus, Onnia, Asterodon, Schizopora, Hyphodontia, Coltriciella, Coltricia, and Trichaptum (Appendix). Perforate SPCs were found in the Rickenella clade, i.e.

34 Septal pore complex morphology in the Agaricomycotina

perforate SPC imperforate SPC vesicular SPC endoplasmic reticulum

Figure 3 – Schematic phylogenetic diagram of the Agaricomycotina adopted from Hibbett (2006). In the Tremellomycetes septal pore caps (SPCs) are absent (Cystofilobasidiales) or have the vesicular morphology (Filobasidiales, Tremellales). In the Dacrymycetes (Dacrymycetales) dolipore septa are associated with imperforate SPCs. In the Agaricomycetes dolipore septa are covered either with imperforate SPCs (Auriculariales, Sebacinales, Gomphales, Trechisporales, and Geastrales) or perforate SPCs (Phallales, Corticiales, Gloeophyllales, Polyporales, Thelephorales, Russulales, Boletales, Atheliales, and Agaricales). Both imperforate and perforate SPCs occur in the Cantharellales and Hymenochaetales. The SPC-type in the Hysterangiales is unknown as no SPC ultrastructure was published. The ER-like strands covering the dolipore in the Cystofilobasidiales seem ancestral to the vesiculate and imperforate SPC-type. It appears that the perforate SPC-type has arisen several times in the Agaricomycetes. Eventually, the perforate SPC was lost in the Cantharellales and Hymenochaetales (grey boxes) and reversed to the imperforate SPC-type.

35 Chapter 2

R. fibula (Figure 1) and Hyphoderma praetermissum (Langer & Oberwinkler, 1993; Keller, 1997). Furthermore, the perforate SPC-type occurs in the Oxyporus clade as Oxyporus latemarginatus (cited as Poria latemarginata) has dolipore septa associated with perforate SPCs (Setliffet al., 1972). Thus the basal clades, viz. the Rickenella and the Oxyporus clade in the Hymenochaetales have perforate SPCs, whereas all the other clades have imperforate SPCs. This suggests that after the perforate SPC-type appeared in the Rickenella clade and the Oxyporus clade, it was subsequently lost and reversed into the imperforate type in the other clades.

Septal Pore Cap Morphology in the Cantharellales The Cantharellales order consists of four clades: a core cantharelloid clade (including Cantharellus, Craterellus, Hydnum, Sistotrema, Clavulina, Multiclavula, and Membranomyces), the Botryobasidium clade, the Ceratobasidiales clade (including Ceratobasidium, Thanatephorus, and Uthatobasidium) and the Tulasnella clade (Moncalvo et al., 2006). The literature on the septal pore morphology in Cantharellus is confusing. Cantharellus cinereus was reported having dolipore septa associated with perforate SPCs (Keller, 1997). On the other hand, others interpreted Cantharellus having imperforate SPCs based on this publication (Hibbett & Thorn, 2001; Larsson et al., 2004; Moncalvo et al., 2006). Our examination of the SPC of C. formosus showed dolipore septa covered with perforate SPCs (Figure 2) and confirmed Keller’s interpretation (Keller, 1997). Next to Cantharellus, Sistotrema brinkmannii also has dolipore septa with perforate SPCs (Dong et al., 1981; Langer, 1994), and thus, members of the core cantharelloid clade have perforate SPCs. The Botryobasidium clade, which is sister to the core cantharelloid clade, has been studied extensively with respect to its SPC ultrastructure (Appendix). It has dolipore septa with imperforate SPCs. Interestingly, the Ceratobasidiales, which is the sister group of the core cantharelloid clade and the Botryobasidium clade, all do have perforate SPCs (Appendix). Finally, members of the Tulasnella clade have dolipore septa that are covered with imperforate SPCs (Appendix). The exact position of Tulasnella remained unclear, but it may be in basal position within the Cantharellales (Moncalvo et al., 2006). After the perforate SPC-type appeared in the Cantharellales it disappeared in the Botryobasidium clade and reversed to the imperforate SPC-type, which is schematically drawn in Figure 4.

Trends in the Evolution of Septal Pore Cap Morphology in the Agaricomycotina As the position of certain orders is uncertain, the fungal phylogeny is not final yet (Hibbett et al., 2007) and future phylogenetic studies may involve changes in the current tree topology. Furthermore, the SPC ultrastructure in certain orders (Geastrales, Gloeophyllales, Gomphales, Phallales, and Trechisporales) has been studied only in few species, and thus, these studies should be extended to get a better-supported SPC-type in these orders. Therefore, we cannot be conclusive on the SPC morphology evolution in

36 Septal pore complex morphology in the Agaricomycotina

Figure 4 – Simplified phylogenetic diagram of the Cantharellales showing the four main clades according to Moncalvo et al. (2006). The core cantharelloid clade and Ceratobasidiales both have dolipore septa associated with perforate septal pore caps (SPCs), whereas the Botryobasidium and Tulasnella clades both have imperforate SPCs. Probably the perforate SPC-type has been lost in the Botryobasidium clade and reversed to the imperforate SPC-type. the Agaricomycotina. Still, certain trends can be inferred from the SPC morphology data combined with the current classification. As the basal lineage in the Agaricomycotina has dolipore septa without SPCs (Cystofilobasidiales) but covered with ER-like strands (e.g. Itersonilia perplexans; Boekhout, 1991), we assume this might be ancestral to both the vesicular and imperforate SPC-types. Evidence showing that the vesicular SPC- type resembles the ER membrane when stained with zinc-iodine (Müller et al., 1995, 1998a) may support this view of a close relation between ER and the vesicular SPC-type. Eventually, the imperforate has given rise to the perforate SPC-type in the Agaricomycetes, which might have reversed to the imperforate SPC-type (Figure 3). This view differs from the one stated by Moore (1996), who suggested a SPC phylogeny that would progress from imperforate to perforate to vesicular forms. The Cantharellales and Hymenochaetales both have imperforate and perforate SPCs. After perforate SPCs have appeared, they subsequently were lost and reversed to the imperforate SPC-type. However, the presented phylogeny of the Cantharellales is probably not final, as, for example, the position of the Tulasnella clade is still not clear (Moncalvo et al., 2006). Moreover the classification of most orders in the Agaricomycetes is still considered uncertain (incertae sedis; Hibbett et al., 2007). Future phylogenetic studies together with ultrastructural studies of the septal pore complex morphology may shed a more definitive light on SPC morphology evolution.

37 Chapter 2 REFERENCES

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41 Chapter 2 , 1979; Van der , 1979; Van ., 1975 et al. et al 1998b (reference herein) ., 1976; Patton & Marchant, 1978a 1977; Patton & Marchant, 1978a; ., 1972 et al et al., et al., et al Author Craig Thielke, 1972 Manocha, 1965 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, Gull, 1976 Flegler Müller 1997 Keller, Berliner & Duff, 1965; Moore, 1965 1997 Keller, 1997 Keller, 1997 Keller, McLaughlin, 1974; Moore & Marchant, 1978 Valk 1965; Waters Giesy & Day, Desole, 1982 Ellis 1985 Oberwinkler, 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, SPC-type perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate Order Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales cited as Agaricus essettei Agrocybe aegerita Coprinus cinereus Coprinus radiatus Coprinus stercorarius Coprinus lagopus Species Agaricus bisporus Agaricus campestris Agaricus silvicola Agaricus xanthoderma Agrocybe arvalis Agrocybe cylindracea Agrocybe dura Agrocybe praecox Amanita muscaria Amanita rubescens Amanita strobiliformis Armillaria mellea Calocybe chrysenteron Clitocybe martiorum Clitocybula lacerata Coprinopsis cinerea Coprinopsis radiata Coprinopsis stercorea Coprinus comatus Cortinarius odorifer Cortinarius orellanus Cortinarius trivialis Cortinarius xanthophyllus Crepidotus amygdalosporus Appendix Chapter 2 - Septal pore cap type in the Agaricomycotina Coprinopsis lagopus

42 Appendix ., 1989 , 1965 ., 1976 et al et al. et al Author Boekhout Patrignani & Pellegrini, 1986 Foerster Besson & Froment, 1968 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, Beneke, 1963 Rexer & Stepanova, 2004 (reference herein) 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, Flegler 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, Rexer & Stepanova, 2004 Rexer & Stepanova, 2004 1997 Keller, Khan & Kimbrough, 1979 SPC-type perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate imperforate perforate perforate perforate perforate perforate perforate perforate perforate perforate imperforate perforate perforate perforate Order Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales cited as Collybia velutipes Galera paludosa Collybia peronata Calvatia gigantea Limacella glioderma . glioderma var Species Disporotrichum dimorphosporum Fistulina hepatica Flammulina velutipes Galerina paludosa Gymnopilus sapineus Gymnopus peronatus Hygrophorus karstenii Laccaria amethystina Lachnella alboviolascens Langermannia gigantea Lentinula edodes Lepiota grangei Lepista glaucocana Lepista luscina Limacella delicata perlatum Lycoperdon favrei Lyophyllum ulmarium Lyophyllum Macrocystidia cucumis Melanoleuca subalpina Melanoleuca subpulverulenta Melanoleuca verrucipes Mucronella calva Mycena galopus Mycena hiemalis Mycena pseudocorticola Nematoloma puiggarii Appendix Chapter 2 - continued

43 Chapter 2

et al., 1976 1976 1967; Marchant & Wessels, 1973, 1967; Marchant & Wessels, et al., et al., et al., Author Patton & Marchant, 1978a Patrignani & Pellegrini, 1986 Lingle, 1989 1997 Keller, 1997 Keller, 1978 Wells, Moore, 1977; Moore & Patton, 1975 1997 Keller, & Kimbrough, 1978 Tu Flegler 1997 Keller, 1997 Keller, Clemencon, 2004 Jersild 1974; Moore & Patton, 1975; Müller 1994, 1995, 1998a, 1999, 2000c; Patton & der Marchant, 1978a; Raudaskoski, 1972; Van 1965 & Marchant, 1978; Wells, Valk 1997 Keller, 1997 Keller, Thielke, 1972 1997 Keller, 1997 Keller, 1997 Keller, Flegler 1997 Keller, 1997 Keller, SPC-type perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate imperforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate Order Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales Agaricales cited as Clitocybe olearia Rhodocybe mundula anthracophilum, Lyophyllum spaerosporum Lyophyllum boudieri Lyophyllum coracinum Lyophyllum Appendix Chapter 2 - continued Species Nidularia confluens Omphalotus olearius Panellus stipticus Phaeolepiota aurea Phaeomarasmius erinaceus Pholiota terrestris Pleurotus cystidiosus Pluteus salicinus Psilocybe cubensis Psilocybe mexicana Radulomyces confluens Resupinatus applicatus Rhodocybe popinalis Schizophyllum commune Strobilurus esculentus Stropharia aeruginosa Stropharia rugosoannulata anthracophila Tephrocybe boudieri Tephrocybe coracina Tephrocybe bombycina Volvariella Xeromphalina cornui Xerula caussei

44 Appendix ., 1977 et al Author Tu 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, 1994 Wells, Lü & McLaughlin, 1991; Moore, 1978b; & Kimbrough, 1978; 1985; Tu Oberwinkler, & Bandoni, 2001 1994; Wells Wells, 1994 1980; Wells, McLAughlin, 1997 Keller, 1994 Wells, Khan & Kimbrough, 1980 1985 Oberwinkler, 1985 Oberwinkler, 1985 Oberwinkler, 1994 Wells, 1997; Moore, 1978b; Patton & Keller, 1994 Marchant, 1978a; Wells, 1964 Wells, 1985 Oberwinkler, 1997 Keller, Patton & Marchant, 1978a 1994; Williams & Andersen, 1996; Wells, 1989 Thilo, 1997 Keller, SPC-type perforate perforate perforate perforate perforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate Order Atheliales Atheliales Atheliales Atheliales Atheliales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales cited as Sclerotium rolfsii Fibulomyces mutabilis Piloderma croceum Elmerina caryae Hirneola auricula-judae, auricula Auricularia Sebacina calcea . sp . . sp sp . sp Appendix Chapter 2 - continued Species Athelia rolfsii Athelopsis glaucina Cristinia helvetica Leptosporomyces mutabilis Piloderma bicolor Aporpium caryae auricula-judae Auricularia fuscosuccinea Auricularia mesenterica Auricularia Basidiodendron cinereum Basidiodendron eyrei Basidiodendron Ductifera Eichleriella Exidia candida Exidia glandulosa Exidia nucleata Exidia Exidia thurentiana Exidia truncata Exidiopsis calcea Exidiopsis effusa

45 Chapter 2 ., et al 1974 et al., Author 1985 Oberwinkler, Khan & Kimbrough, 1980 1989 Andersen, 1996; Williams & Thilo, Kirschner & Chen, 2004 1985 Oberwinkler, 1994 Wells, 1994 Wells, 1994 1997; Moore, 1996; Wells, Keller, 1994 Wells, 1985 Oberwinkler, 1997 Keller, Patrignani & Pellegrini, 1986 1997 Keller, Patrignani & Pellegrini, 1986 Patton & Marchant, 1978a Becket 1997 Keller, Langvad, 1971 1997 Keller, Orlovich & Ashford, 1994; Shepherd 1993 1997 Keller, Hofmann, 1989 1994 Wells, 1994 Langer, SPC-type imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate imperforate imperforate Order Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Auriculariales Boletales Boletales Boletales Boletales Boletales Boletales Boletales Boletales Boletales Boletales Cantharellales Cantharellales cited as Sebacina umbrina Heterochaetella dubia Guepinia rufa Pulveroboletus gentilis Boletus rubinellus Coniophora cerebella Pisolithus tinctorius tuberosa Tremellodendropsis . . sp . sp sp Appendix Chapter 2 - continued Species Exidiopsis Exidiopsis sublivida Exidiopsis umbrina Helicomyxa everhartioides Myxarium Patouillardina cinerea Protodontia oligacantha Pseudohydnum gelatinosum Stypella dubia Stypella Stypella vermiformis helvelloides Tremiscus gentilis Aureoboletus Boletus cramesinus Boletus edulis Chalciporus rubinellus Coniophora fusispora Coniophora puteana Leucogyrophana mollusca Pisolithus arhizus Serpula lacrymans Xerocomus chrysenteron Aphelaria tuberosa Botryobasidium candicans

46 Appendix ., 1977; et al Figure 2 ., 1998b, 2000c; ., 1998b, et al Chapter 2

., 2004 et al., et al Author 1994 Langer, 1994 Langer, 1994 Langer, 1994 Langer, 1994 Langer, 1994 Langer, 1994 Langer, 1997 Keller, 1994 Langer, 1985 1994; Oberwinkler, Langer, 1994 1997; Langer, Keller, 1994 Langer, 1997 Keller, Driel Van Andersen, 1996 2001 & Oberwinkler, Weiss Andersen, 1996; Currah & Sherburne, 1992; 1997; Müller Keller, Patton & Marchant, 1978a; Tu & Bandoni, 2001 1994; Wells Wells, Andersen, 1996; Currah & Sherburne, 1992 1997 Keller, Weiss Andersen, 1996 Andersen, 1996 1985 1997 (perforate); Oberwinkler, Keller, (imperforate) SPC-type imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate perforate perforate perforate imperforate perforate perforate perforate perforate perforate perforate perforate / imperforate Order Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales cited as Botryobasidium botryosum Botryobasidium isabellinum Ceratobasidium ramicola Rhizoctonia cerealis Rhizoctonia fragariae . sp Appendix Chapter 2 - continued Species Botryobasidium conspersum Botryobasidium curtisii Botryobasidium grandisporum Botryobasidium lacinisporum Botryobasidium laeve Botryobasidium longisporum Botryobasidium obtusisporum Botryobasidium pruinatum Botryobasidium simile Botryobasidium subcoronatum Botryobasidium vagum Botryohypochnus isabellinus Cantharellus cinereus Cantharellus formosus Ceratobasidium anceps Ceratobasidium calosporum Ceratobasidium cornigerum Ceratobasidium obscurum Ceratobasidium pseudocornigerum Ceratobasidium Ceratorhiza cerealis Ceratorhiza fragariae Clavulicium macounii

47 Chapter 2 1998b, 2000a; 1998b, , 1998b, 2000c; Tu 2000c; Tu , 1998b, et al., et al. ., 1998b, 2000c ., 1998b, ., 1975; Müller ., 1972 ., 1981; Langer, 1994 ., 1981; Langer, et al et al et al et al ., 1977 Author Andersen, 1996; Currah & Sherburne, 1992; Müller 1994 (reference herein) Wells, 1994 (reference herein) Wells, Currah & Sherburne, 1992 1994 (reference herein) Wells, 1985 Oberwinkler, Andersen, 1996 Andersen, 1996 Andersen, 1996 Andersen, 1996 Andersen, 1996; Currah & Sherburne, 1992 1963, 1964; Andersen, 1996; Bracker & Butler, Lisker Setliff Dong Currah & Sherburne, 1992 1994 (references herein) Wells, 1963; Andersen, 1996; Bracker & Butler, 1994; Müller Langer, et al Andersen, 1996; Currah & Sherburne, 1992 1994 Langer, 1994 Langer, 1994 Wells, Andersen, 1996 SPC-type imperforate imperforate imperforate perforate imperforate imperforate perforate perforate perforate perforate imperforate perforate perforate perforate imperforate perforate perforate perforate perforate imperforate imperforate Order Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales cited as Rhizoctonia anaticula Heteroacanthella variabile Epulorhiza repens Rhizoctonia dichotoma Aquathanatephorus pendulus, Thanatephorus praticola, Thanathephorus sasakii, Pellicularia filamentosa . sp Species Epulorhiza anaticula Heteroacanthella acanthophysa Heteroacanthella variabilis Moniliopsis anomala Monosporonella termitophila Paullicorticium pearsonii Rhizoctonia endophytica Rhizoctonia oryzae Rhizoctonia praticola Rhizoctonia ramicola Rhizoctonia repens Rhizoctonia solani Sistotrema Stilbotulasnella conidiophora Thanatephorus cucumeris Thanatephorus pennatus biapiculata Tofispora repetospora Tofispora araneosa Tulasnella calospora Tulasnella Appendix Chapter 2 - continued Sistotrema brinkmannii

48 Appendix ., et al ., 1977 ., et al et ., 1998b; Tu ., 1998b; Tu 2001 ., 2004; Wells & ., 2004; Wells et al et al., et al 2003 ., 1992 (inflated non-perf. septum) 2004 et al., , 1993 , 1993 et al et al., et al. et al. Suh Guého Author Moore, 1978b Andersen, 1996 1994; Weiss Langer, Bandoni, 2001 1992 Keller & Job, 1992 Keller & Job, Tu 1994; Langer, 1992; Job, & Keller 1985 Oberwinkler, 1997 Keller, 1997 Keller, Hoch & Howard, 1981 Patton & Marchant, 1978a 1994 Wells, 1997 Keller, 1997 Keller, Diederich Andersen, 1996 Andersen, 1996; Müller 1977 1993 Suh & Sugiyama, Weiss Suh Boekhout, 1991; Fell SPC-type imperforate imperforate imperforate imperforate imperforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate absent absent absent absent absent absent Order Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Cantharellales Corticiales Corticiales Corticiales Corticiales Corticiales Corticiales Corticiales Corticiales Corticiales ? Corticiales ? Cystofilobasidiales Cystofilobasidiales Cystofilobasidiales Cystofilobasidiales Cystofilobasidiales Cystofilobasidiales cited as Laeticorticium lundellii Laetocorticium roseum Corticium fuciforme Galzinia culmigena Leucosporidium lari-marini . sp . sp Species fuscoviolacea Tulasnella irregularis Tulasnella Tulasnella violacea Tulasnella violea Tulasnella Uthatobasidium fusisporum Uthatobasidium Corticium boreoroseum Corticium roseum Laetisaria arvalis Laetisaria fuciformis Limonomyces culmigenus Lindtneria flava Lindtneria trachyspora Marchandiomyces corallinus Rhizoctonia zeae circinata Waitea Cystofilobasidium capitatum Cystofilobasidium ferigula Cystofilobasidium infirmo- miniatum Itersonilia perplexans Mrakia frigida pullulans Trichosporon Appendix Chapter 2 - continued

49 Chapter 2 1976; Keller & Job, 1992; Moore, 1976; Keller & Job, , 1997; Deml & Oberwinkler, 1981; , 1997; Deml & Oberwinkler, 2004. et al., et al. et al. Author & Kimbrough, 1978; Wells, Keller 1992; Tu 1994 Keller 1992; Patton & Marchant, 1978a 1994 (reference herein) Wells, 1994 Wells, & Kimbrough, 1978 Tu 1994 Wells, 1994 (reference herein) Wells, Moore, 1965 Flegler 1978b; Mossebo & Amougou, 2001; Wells, 1994. 1994 (reference herein) Wells, 1992 Keller & Job, Bauer Weiss Rij, 1972 Moore & Kreger-Van Rij, 1972 (SPC absent, Moore & Kreger-Van 1994 (sacculate) ER-vesicles); Wells, 1994 (sacculate, poorly defined; Wells, reference herein) Hibbett & Thorn, 2001 SPC-type imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate absent vesicular absent or vesicular absent or vesicular imperforate Order Dacrymycetales Dacrymycetales Dacrymycetales Dacrymycetales Dacrymycetales Dacrymycetales Dacrymycetales Dacrymycetales Dacrymycetales Dacrymycetales Dacrymycetales Entorrhizomycetes Filobasidiales Filobasidiales Filobasidiales Geastrales minor cited as Guepiniopsis chrysocoma Dacrymyces deliquescens var. Dacrymyces deliquescens Femsjonia peziziformis Leucosporidium capsuligenum sp. Species Calocera cornea Calocera viscosa Cerinomyces aculeatus Cerinomyces altaicus Dacrymyces abietinus Dacrymyces chrysocomus Dacrymyces dendrocalami Dacrymyces minor Dacrymyces stillatus Ditiola haasii Ditiola peziziformis Entorrhiza casparyana Filobasidium capsuligenum Filobasidium floriforme Filobasidium uniguttulatum Geastrum Appendix Chapter 2 - continued

50 Appendix , 2000 ., 2000b ., 2000b ., 2000b ., 2000b ., 2000b et al. et al et al et al et al et al Author 1966 Hyde & Walkinshaw, 1997 Keller, Patrignani & Pellegrini, 1986 Müller Müller Müller Müller Müller 1985 Oberwinkler, 1993 1997; Langer & Oberwinkler, Keller, 1993 Langer & Oberwinkler, 1997 Keller, Greslebin 1997 Keller, 1993 Langer & Oberwinkler, 1997 Keller, 1993 1997; Langer & Oberwinkler, Keller, 1993 Langer & Oberwinkler, 1997 Keller, & Huang, 1997 Wu 1993 Langer & Oberwinkler, 1993 Langer & Oberwinkler, 1997 Keller, 1993 1997; Langer & Oberwinkler, Keller, & Huang, 1997 Wu SPC-type perforate perforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate perforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate Order Gloeophyllales Gloeophyllales Gomphales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales cited as Lenzites sepiaria Lentinus lepideus Clavaria ignicolor Cyclomyces fuscus Basidioradulum radula Hyphodontia verruculosa Hyphoderma sambuci Species Gloeophyllum sepiarium Neolentinus suffrutescens Ramaria ignicolor Asterodon ferruginosum Coltricia perennis Coltriciella dependens Hydnochaete japonica Hymenochaete cyclolamellata Hymenochaete rubiginosa Hyphoderma praetermissum Hyphodontia alutaria Hyphodontia arguta Hyphodontia australis Hyphodontia barba-jovis Hyphodontia cineracea Hyphodontia crustosa Hyphodontia floccosa Hyphodontia gossypina Hyphodontia hastate Hyphodontia mollis Hyphodontia pallidula Hyphodontia radula Hyphodontia rimosissima Hyphodontia sambuci Hyphodontia subglobosa Appendix Chapter 2 - continued

51 Chapter 2 ., et al Figure 1 Chapter 2 , ., 2000b; Setliff et al. et al ., 2005 et al ., 2000b ., 1972 et al et al Patton & Marchant, 1978a 1997 Keller, 1997 Keller, Moore, 1980 Author Moore, 1980 Moore, 1980 Müller Moore, 1980 Moore, 1980; Müller 1993 Langer & Oberwinkler, Moore, 1985 & McKeen, 1978 Traquair Rexer & Stepanova, 2004 (reference herein) Patton & Marchant, 1978a 1997; Moore & Marchant, 1972; Keller, 1975, 1985; Patton & Marchant, 1978a, b 1997 Keller, Bianchinotti 1972 Setliff Shukla, 1975 Moore, 1980 1997 Keller, 1997 Keller, Driel Kost, 1984; Van perforate perforate perforate perforate SPC-type imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate perforate perforate perforate perforate perforate perforate imperforate imperforate imperforate imperforate perforate Polyporales Polyporales Polyporales Polyporales Order Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Phallales Phallales Polyporales Polyporales Polyporales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Hymenochaetales Cerocorticium sulfureoisabellinum cited as Onnia leporina Polyporus tomentosus Poria latemarginata Fomes igniarius Phellinus chrysoloma Rickenella aulacomniophila Hirschioporus abietinus Hirschioporus pargamenus Polyporis biennis Rhizochaete americana Hydnum septentrionale . igniarius var Conohypha terricola Flavophlebia sulfureoisabellinum Fomes fomentarius Appendix Chapter 2 - continued Species Inonotus hispidus Inonotus leporinus Inonotus weirii Onnia circinata Onnia tomentosa Oxyporus latemarginatus Phellinus igniarius Phellinus torulosus Phellinus tuberculosus Porodaedalea chrysoloma Rickenella fibula Schizopora paradoxa abietinum Trichaptum biforme Trichaptum Clathrus cancellatus Phallus impudicus Abortiporus biennis Bulbillomyces farinosus Ceraceomyces americanus Climacodon septentrionalis

52 Appendix ., 1993 et al ., 2005 ., 2005 ., 2005 ., 2005; Tsuneda ., 2005 ., 1989 et al et al et al et al et al ., 1976 et al et al Bianchinotti Bianchinotti 1997 Keller, Bianchinotti Flegler Wilsenach & Kessel, 1965 1997 Keller, Author 1997 Keller, 1997 Keller, 1993 1997; Langer & Oberwinkler, Keller, 1997 Keller, 1993 Langer & Oberwinkler, 1997 Keller, 1993 Langer & Oberwinkler, 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, 1962 Moore & McAlear, 1997 Keller, 1997 Keller, 1997 Keller, Moore, 1980 Boekhout Bianchinotti Bianchinotti perforate perforate perforate perforate perforate perforate perforate SPC-type perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Order Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales cited as Hypochnicium polonense Hypochnicium sphaerosporum Sporotrichum pruinosum, Chrysosporium xerophilum Rhizochaete filamentosa Rhizochaete radicata Favolus alveolaris Phlebia radiata Polyporus rugulosus Polyporus squamosus Appendix Chapter 2 - continued Species Ganoderma lucidum Grifola frondosa Gyrophanopsis polonensis Hyphoderma mutatum Hyphoderma setigerum Hyphoderma subdefinitum Hypochnicium bombycinum Hypochnicium eichleri Hypochnicium lundellii Hypochnicium punctulatum Meruliopsis taxicola Merulius tremellosus Mycoacia fuscoatra Mycoacia uda Osteina obducta Phaeolus schweinitzii Phanerochaete chrysosporium Phanerochaete filamentosa Phanerochaete radicata Phanerochaete velutina Phlebia ochraceofulva Phlebia rufa Polyporus alveolaris

53 Chapter 2 ., 2005 ., 1989 et al ., 1984; Girbardt, 1958,1961 ., 1976 ., 2000b et al et al et al et al Wilsenach & Kessel, 1965 1997 Keller, Patrignani & Pellegrini, 1986 Boekhout 1997 Keller, Aylmore 1997 Keller, 1997 Keller, 1997 Keller, Müller 1997 Keller, 1997 Keller, 1997 Keller, 1997 Keller, Flegler 1997 Keller, 1997 Keller, Besson & Fremont, 1964 1997 Keller, Hanlin, 1978 Patrignani & Pellegrini, 1986 1997 Keller, Author Moore, 1980 Bianchinotti perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate perforate imperforate perforate perforate perforate SPC-type perforate perforate Polyporales Polyporales Polyporales Polyporales Polyporales Polyporales Russulales Russulales Russulales Russulales Russulales Russulales Russulales Russulales Russulales Russulales Russulales Russulales Russulales Russulales Russulales Russulales Order Polyporales Polyporales Albatrellus pes-caprae Steccherinum robustius Coriolus versicolor, Polystictus versicolor Megalocystidium lactescens citrinus Vesiculomyces Confertobasidium olivaceoalbum cited as Poria monticola Scutiger oregonensis Sparassis crispa Sporotrichum aurantiacum Steccherinum bourdotii versicolor Trametes Albatrellus ovinus Albatrellus subrubescens Aleurodiscus aurantius Asterostroma medium vulgare Auriscalpium Gloeocystidiellum lactescens Gloeocystidiellum porosum Gloiothele citrina Hericium coralloides Laxitextum bicolor Peniophora laeta Scytinostroma duriusculum Scytinostromella olivaceoalba Spiniger meineckellus Stereum hirsutum Zelleromyces stephensii Appendix Chapter 2 - continued Species Polyporus tuberaster Rhizochaete brunnea Rhodonia placenta

54 Appendix 1998 ., 1998b; Williams & Thilo, 1989 ., 1998b; Williams & Thilo, et al et al., Author 1997 Keller, 1982 & Oberwinkler, Wells Khan & Kimbrough, 1980 (perforate); 1988 (imperforate; reference Berbee & Wells, 1994 (imperforate; reference herein); Wells, herein) Verma 1997 Keller, 1989 Andersen, 1996; Williams & Thilo, 1997 Keller, Khan & Kimbrough, 1980 Currah & Sherburne, 1992; Oberwinkler, 1989 1985; Williams & Thilo, Müller & Khan & Kimbrough, 1980; Wells 1982 Oberwinkler, 1982 & Oberwinkler, Wells 1982 & Oberwinkler, Wells 1985 Oberwinkler, 1997 Keller, 1997 Keller, 1997 Keller, Patrignani & Pellegrini, 1986 1994 1997; Langer, Keller, SPC-type imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate imperforate perforate perforate perforate perforate perforate Order Sebacinales Sebacinales Sebacinales Sebacinales Sebacinales Sebacinales Sebacinales Sebacinales Sebacinales Sebacinales Sebacinales Sebacinales Sebacinales Sebacinales Thelephorales Thelephorales Thelephorales Thelephorales Thelephorales cited as Exidiopsis fugacissima, Sebacina fugacissima Exidiopsis grisea, Exidiopsis plumbescens Sebacina vermifera, Exidiopsis vermifera Thelephora palmata . sp . sp Appendix Chapter 2 - continued Species Craterocolla cerasi Efibulobasidium rolleyi Microsebacina fugacissima Piriformospora indica Sebacina epigaea Sebacina grisea Sebacina helvelloides Sebacina incrustans Sebacina Serendipita vermifera candidum Tremellodendron australiensis Tremelloscypha gelatinosa Tremelloscypha Tremelloscypha Bankera violascens Hydnellum concrescens Sarcodon versipellis Thelephora anthocephala Thelephora terrestris

55 Chapter 2 , 1995 (cupulate) 1991 (cupulate) ., 1991 (cupulate) et al. 1995 ., 1981 et al., et al et al et al., Kwon-Chung & Popkin, 1976 1994 (reference herein) Wells, 1994 (reference herein) Wells, Moore, 1978a (ampulliform vesicles) 1994 (references herein) Wells, 1994 (reference herein) Wells, 1994 (references herein) Wells, Moore, 1978b 1994 (reference herein) Wells, Moore, 1978b (reference herein) Author 1997 Keller, Calonge, 1969 1997 Keller, 1997 Calonge, 1969; Keller, 1997 Keller, 1997 Keller, 1997 Keller, Boekhout Boekhout Rhodes Adams 1994 (reference herein) Wells, Kwon-Chung SPC-type perforate perforate perforate perforate perforate perforate imperforate vesicular vesicular absent reticulate vesicular vesicular absent absent vesicular vesicular absent absent absent vesicular vesicular vesicular Order Thelephorales Thelephorales Thelephorales Thelephorales Thelephorales Thelephorales Trechisporales Tremellales Tremellales Tremellales Tremellales Tremellales Tremellales Tremellales Tremellales Tremellales Tremellales Tremellales Tremellales Tremellales Tremellales Tremellales Tremellales cited as fibrosa, Tomentella bombycina Tomentella Carcinomyces effibulatus Christiansenia pallida . laurentii var Species crinalis Tomentella fuscoferruginosa Tomentella pilosa Tomentella fibrosa Tomentellina echinospora Tomentellopsis submollis Tomentellopsis Subulicystidium longisporum Bullera variabilis Bulleromyces albus Cryptococcus laurentii Ditangifibulae dikaryotae Fibulobasidium inconspicuum Filobasidiella depauperata Filobasidiella neoformans Phragmoxenidium mycophilum Rhynchogastrema coronatum Sirobasidium magnum Syzygospora alba Syzygospora effibulata Syzygospora pallida brasiliensis Tremella foliacea Tremella fuciformis Tremella Appendix Chapter 2 - continued

56 Appendix ., 2000). al

et ., 1995, 1998a, (Fell ., 2001 ., 2001 et al al

et al et Trichosporonales ., 1992 (tubular/vesicular) ., 1992 (non-perforate septum) ., 1992 (tubular) ., 1992 ., 1992; Fell ., 1992; Fell ., 1992 ., 1992 ., 1992; Müller ., 2004 (sacculate) et al et al et al et al et al et al et al et al et al et al Author 1985 1988; Oberwinkler, Berbee & Wells, (sacculate) 1994 Moore, 1978b; Wells, 1994 (reference herein) Wells, Weiss 1988 (references herein); Berbee & Wells, 1994 (reference herein) Wells, 1988 (references herein); Berbee & Wells, 1994 (reference herein) Wells, Guého Guého Guého Guého Guého Guého Guého Guého Guého globular) 2000c (tubular, Moore, 1986 ., 2007). Current use of names was verified in Mycobank (www. al

et SPC-type vesicular vesicular vesicular vesicular vesicular vesicular vesicular absent vesicular vesicular absent vesicular vesicular absent vesicular vesicular . (2007) plus addition of the order al

et in Corticiales is still uncertain. (Hibbett Order Tremellales Tremellales Tremellales Tremellales Tremellales Tremellales Trichosporonales Trichosporonales Trichosporonales Trichosporonales Trichosporonales Trichosporonales Trichosporonales Trichosporonales Trichosporonales Wallemiomycetes Rhizoctoina zeae Agaricomycotina and . Orders according to Hibbett Waitea circinata Waitea Agaricomycotina are two unplaced classes in the cited as uliginosa Tetragoniomyces ., 2004). The placement of al

et Wallemiomycetes and . sp Species globospora Tremella mesenterica Tremella rhytidhysterii Tremella Tremella uliginosa Tremella papilionaceus Trimorphomyces asahii Trichosporon brassicae Trichosporon coremiiforme Trichosporon cutaneum Trichosporon inkin Trichosporon laibachii Trichosporon moniliiforme Trichosporon mucoides Trichosporon sporotrichoides Trichosporon sebi Wallemia Entorrhizomycetes mycobank.org; Crous Appendix Chapter 2 - continued Table 1 – Septal pore cap type in the

57 Chapter 2 REFERENCES APPENDIX

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62 Chapter 3

Laser Microdissection of Fungal Septa as Visualized by Scanning Electron Microscopy

Kenneth G.A. van Driel, Teun Boekhout, Han A.B. Wösten, Arie J. Verkleij & Wally H. Müller

Published in Fungal Genetics and Biology 44, 466 – 473 (2007) Chapter 3

ABSTRACT

Laser microdissection has been proven a successful technique to isolate single cells or groups of cells from animal and plant tissue. Here, we demonstrate that laser microdissection is suitable to isolate subcellular parts of fungal hyphae. Dolipore septa of Rhizoctonia solani containing septal pore caps were cut by laser microdissection from sections of mycelium and collected by laser pressure catapulting. Subsequently, microdissected septa were visualized using wheat germ agglutinin labeling in combination with scanning electron microscopy. The use of laser microdissection on fungal cells opens new ways to study subcellular fungal structures and the biochemical composition of hyphal cells.

INTRODUCTION

Laser microdissection has become a standard technique to select single cells or specific cell populations from animal or human tissue to study them separately from their heterogeneous environment. Recently, laser microdissection has also been used in plant research (Day et al., 2005; Nelson et al., 2006). Several laser microdissection techniques have been developed (Day et al., 2005; Pinzani et al., 2006), of which three systems are mainly used. Firstly, the PixCell system (Arcturus Bioscience, Inc., Mountain View, CA, USA, www.arctur.com) that uses a near infrared laser to capture the cells of interest by fusing them to a thermoplastic film (Emmert-Buck et al., 1996). Secondly, the AS LMD laser microscope (Leica Microsystems AG, Wetzlar, Germany, www.leica-microsystems. com), which is a non-contact method that uses an UV laser of 337 nm wavelength to dissect cells, which then fall by gravity into a tube cap placed underneath the section (De Souza et al., 2004). Thirdly, the PALM MicroBeam system (P.A.L.M. Microlaser Technologies AG, Bernried, Germany, www.palm-mikrolaser.com) that uses a pulsed UV-A laser of 337 nm wavelength to microdissect the cells of interest. This is combined with non-contact transfer by laser pressure catapulting (LPC), by which the force of a focused laser beam is used to overcome the gravity and to transfer the dissected tissue sample upwards into the cap of a sample tube placed above the section (Schütze & Lahr, 1999; Westphal et al., 2002).

Once the cells of interest are dissected and collected, their protein and genetic profile can be analyzed by several down-stream applications. DNA and RNA extracted from the laser microdissected tissue can be subjected for instance to PCR (Wong et al., 2000), RT- PCR (Fend et al., 1999; Mojsilovic-Petrovic et al., 2004; Shimamura et al., 2004; Wang et al., 2004), real-time quantitative RT-PCR (Wong et al., 2000), or gene expression studies (Scheidl et al., 2002; Burbach et al., 2003; Fuller et al. 2003; Mohr et al., 2004). RNA isolated

64 Laser microdissection of fungal septa from laser microdissected samples is of high quality and low copy number mRNA can be isolated (Mikulowska-Mennis et al., 2002). Proteins from laser microdissected tissue can be analyzed for example by 2D-gel electrophoresis (Banks et al., 1999; Ornstein, 2000; Craven & Banks, 2001; Craven et al., 2002; Moulédos et al., 2003; De Souza et al., 2004), peptide mass fingerprinting (Craven et al., 2002), protein arrays (Grubb et al., 2003), immunoassays (Simone et al., 2000), and LC-MS/MS (Schad et al., 2005a). Compared to traditional isolation techniques, laser microdissection has no gross effects on protein profiles, antigenicity, and mass spectrometric profiles (Banks et al., 1999). Next to genetic and protein profiles it was demonstrated that cell-type specific metabolite profiles could be generated from laser microdissected plant tissue (Schadet al., 2005b).

Laser microdissection has also been applied in several studies in which bacterial, viral, and fungal infections were involved that were recognized either by morphological changes of the tissue (Ryan et al., 2002; Yazdi et al., 2004), or by visualization by fluorescencein situ hybridization (FISH) (Klitgaard et al., 2005) or by using a fluorescent fungal cell wall stain (Xue et al., 2005). In addition, laser microdissection can be applied for isolation of subcellular structures. For instance, individual chloroplasts could be isolated from plant cells for single-organelle analysis (Meimberg et al., 2003), and metaphase chromosomes were isolated from plant and animal cells (Schermelleh et al. 1999; Hobza et al., 2004).

In the current study, we used the PALM MicroBeam system to isolate fungal septa from hyphae of the filamentous basidiomcyetous fungusRhizoctonia solani, which has dolipore septa with perforate septal pore caps (SPCs) (Bracker & Butler, 1963; Müller et al., 1998, 2000). Though many ultrastructural details of the SPC and the dolipore septum of R. solani are available (Bracker & Butler, 1963; Lisker et al., 1975; Müller et al., 1998, 2000), nothing is known about the composition of the dolipore-SPC complex. Therefore, an appropriate isolation technique of the dolipore-SPC complex is needed to understand its biochemical composition and hence its role in basidiomycetous fungi. Here, laser microdissection is used for the first time to isolate fungal structures, and therefore we focus on the visualization of the isolated and collected septa to confirm the isolation procedure. The microdissected and laser catapulted septa were visualized by scanning electron microscopy using WGA-gold-Ag labeling.

MATERIALS & METHODS

Strain and Growth Conditions Rhizoctonia solani (CBS 346.84) was grown on malt extract agar (Oxoid, Hampshire, UK) for 4 days at 25oC between two perforated polycarbonate (PC) membranes with a

65 Chapter 3 diameter of 47 mm and a 0.6 μm pore size (Poretics, Osmonics Inc., Minnetonka, MN, USA).

Preparation of Mycelium for Laser Microdissection in Buffer The upper PC membrane was removed from the sandwiched culture and the mycelium was fixed with a picric acid and formaldehyde containing fixative (PA/FA fixative: 1.5 parts saturated picric acid (PA), 2.5 parts 8% (w/v) formaldehyde (FA), 5 parts 200 mM sodium citrate buffer, pH 6.2, and 1 part distilled water) (Mülleret al., 2002). After 30 min, the mycelium was transferred to fresh PA/FA fixative and incubated for another 30 min. Then, the sandwich culture was washed and stored in 1% formaldehyde in phosphate buffered saline (PBS) (137.0 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, 1.5 mM KH2PO4) at 4oC. Before use, cultures were washed with PBS. Parts of about 10 x 5 mm were cut from the periphery of the colony, placed on a microscope glass, and kept in buffer to prevent desiccation of the mycelium.

Cryo-sectioning and Lowicryl HM20 Embedded Sections Sandwiched cultures were fixed in PA/FA fixative as described above and embedded in 12% (w/v) gelatin in PBS. Cubes of about 1 mm3 were cut, infused with 2.3 M sucrose, mounted on a cutting pin and frozen for cryo-sectioning according to Tokuyasu (1973). Cryo-sections of about 150 nm and 1 μm thick were cut with a glass knife using a cryo- ultramicrotome (Leica Microsystems AG, Wetzlar, Germany). The cryo-sections were picked-up with 2.3 M sucrose using a Perfect Loop (Electron Microscopy Sciences (EMS), Hatfield, PA, USA) and transferred to a formvar film-coated 1 × 2 mm single slot nickel grid (L2 × 1, Veco grid, EMS). The grids with cryo-sections were floated on droplets of PBS, with the section-side down, to remove the sucrose. Subsequently, the sections were stained with 0.25% (w/v) toluidine blue, and 0.5% (v/v) acetic acid in distilled water for 10 min to enhance the visibility of the hyphal cells and the identification of the septa. Sections were washed three times with distilled water and embedded in a thin film of methylcellulose (Müller et al., 2002) to prevent drying of the cryo-sections. The grids with cryo-sections were placed on a microscope glass slide.

As an alternative to cryo-sectioning, R. solani mycelium was low-temperature embedded in lowicryl HM20. Parts of the sandwiched cultures were first cryo-fixed by high-pressure freezing (HPF) using a Leica EM HPF (Leica). The mycelium was subsequently freeze- substituted in 0.3% (w/v) uranyl acetate and 0.01% (v/v) glutaraldehyde in anhydrous methanol, and infiltrated and low-temperature embedded in lowicryl HM20 as described by Müller et al. (2002). After 48 hr of polymerization at –40oC and 24 hr of polymerization at room temperature under UV light, 300 nm thick sections were cut with a diamond knife (Diatome, Hatfield, PA, USA) using an ULTRACUT E ultramicrotome (Leica) and labeled as described below.

66 Laser microdissection of fungal septa

WGA-gold-Ag Labeling Sections were labeled and washed while floating and were transferred with a Perfect Loop between droplets. HM20 sections were blocked for 30 min with 3% (w/v) BSA-c (Aurion, Wageningen, The Netherlands) in TBS buffer (10 mM Tris-HCl, 100 mM NaCl, pH 7.4). Sections were subsequently incubated for 60 min with wheat germ agglutinin (WGA) conjugated to 10 nm colloidal gold (Sigma Aldrich, St. Louis, MO, USA), 1:50 diluted in 0.1% (w/v) BSA-c. Sections were washed with 0.1% (w/v) BSA-c in TBS followed by three washes with PBS and three washes with distilled water for 5 min each. Gold particles were silver enhanced (Aurion) during 30 min at room temperature, and washed subsequently with distilled water. WGA-gold-Ag labeled HM20 sections were picked-up with a single hole nickel grid with an aperture of 2 mm (GA2000-Ni, EMS) and placed up side down on a PEN membrane covered microscope slide (Figure 1). Excess of fluid was removed by placing a small piece of Whatman paper against the rim of the grid.

Figure 1 – Schematic representation of the preparation of labeled sections of Rhizoctonia solani hyphae embedded in HM20 placed on a PEN membrane. Sections were labeled with WGA-gold and silver enhanced (A). A single hole grid was placed on the droplet in such a way that the grid enclosed the section (B). By picking up the grid, the section was taken from the droplet (C). The grid was subsequently placed up side down on a PEN membrane-covered microscope slide (D) and excess of fluid was removed with a piece of Whatman paper (E, F).

67 Chapter 3

Figure 2 – Schematic representation of laser microdissection and laser pressure catapulting of septa of Rhizoctonia solani from WGA-gold-Ag labeled HM20 sections. A section was placed with a single hole grid on PEN-membrane (A) and microdissected with the laser (B). A formvar film-coated single slot grid (L2x1) was put on top of the single hole grid that enclosed the HM20 section and the microdissected region was catapulted upwards by LPC (C). During the transfer (D) the microdissected samples turned over and were collected at the formvar film-coated grid (E), which was analyzed by electron microscopy (F).

Laser Microdissection and Laser Pressure Catapulting Laser microdissection and laser pressure catapulting were performed with the PALM MicroBeam system (P.A.L.M. Microlaser Technologies AG) equipped with an Axiovert 200 Zeiss inverted microscope (Carl Zeiss AG, Oberkochen, Germany) and a 3CCD color camera (HV-D30, Hitachi Kokusai Electric Inc., Tokyo, Japan). We selected fungal septa on the computer screen with the PALM RoboSoftware (v2.2). Around the septum a circle was drawn along which the laser followed to cut the septum. The laser power and laser focus were adjusted until the cut was at its narrowest during microdissection. The following settings have been used with a 40× objective: PA/FA fixed mycelium was microdissected using a laser UV energy setting of 76 and a focus setting of 51 in the PALM Robosoftware. Cryo-sections were microdissected using UV energy settings between 73 – 75, and UV focus settings of 51 – 52 in the PALM Robosoftware. The microdissected material was catapulted into the cap of a sample tube placed above the section. Lowicryl

68 Laser microdissection of fungal septa

HM20 sections on a PEN membrane-covered microscope glass slide were cut using a laser UV energy setting of 67 and a focus setting of 50 in the P.A.L.M. software. The diameter of the dissected specimen was between 14.7 and 19.5 μm. The microdissected material was catapulted with an UV energy setting of about 77 in the PALM Robosoftware and collected on a formvar film-coated single slot grid (L2 × 1, Veco grid, EMS). A schematic representation of the laser microdissection and laser catapulting procedure of lowicryl HM20 sections is shown in Figure 2.

Electron Microscopy Sections of HPF-fixed, freeze-substituted, and HM20 embedded R. solani hyphae were labeled with WGA-gold-Ag and viewed with a TECNAI 10 (FEI Company, Eindhoven, The Netherlands) transmission electron microscope at an acceleration voltage of 100 kV prior to laser microdissection. Microdissected and laser pressure catapulted HM20 embedded material was examined with a light microscope or an XL30 scanning field emission gun microscope (FEI Company). At low magnification, an acceleration voltage of 5.0 kV, spot size of 3.0, work distance of 5.0 mm and the SE detector were used (Figure 7A, B). At high magnification, an acceleration voltage of 20.0 kV and the TLD detector were used with a spot size of 4.0 and a work distance of 5.1 mm (Figure 7C) or a spot size of 3.0 and a work distance of 4.7 mm were used (Figure 7D, E).

RESULTS

Laser Microdissection of Fungal Septa from Mycelium in Buffer Picric acid and formaldehyde fixed mycelium of Rhizoctonia solani was placed on a microscope slide with adherent buffer. We did not use air-dried mycelium or critical point dried mycelium, because the septa could not be recognized anymore (result not shown). The septa were selected on the computer screen (Figure 3A), microdissected with the laser (Figure 3B, C), and released into the surrounding buffer (Figure 3C). However, the released septa could not be collected in the cap of a sample tube by laser pressure catapulting (LPC), as was shown by light microscopy (data not shown). In addition, the LPC shot, used to transfer the microdissected samples upwards, affected a larger proportion of the mycelium than only the dissected septum, resulting in a dislocation of the surrounding hyphae (result not shown). Though the hyphae were successfully cut by the laser, the microdissected septa could not be collected from fluid by LPC.

69 Chapter 3

A B C Figure 3 – Light microscopic images of picric acid and formaldehyde fixed Rhizoctonia solani hyphae in buffer before (A) and after (B, C) laser microdissection at either side of a septum indicated with an asterisk. Bar represents 10 µm.

Laser Microdissection and Laser Pressure Catapulting of Fungal Septa from Cryo- sections Transmission electron microscopy showed that dolipore septa with associated perforate septal pore caps were well preserved in picric acid and formaldehyde fixed R. solani mycelium (Figure 4). Preceding laser microdissection, the cryo-sections were stained with toluidine blue to enhance contrast of the hyphal cells and the septa (Figure 5A). The septa were laser microdissected (Figure 5B), and transferred upwards in the cap of a sample tube by LPC, thereby leaving small holes in the cryo-section (Figure 5C). However, the collected material was wrinkled, therefore hampering examination by electron microscopy (data not shown). Thus, cryo-sections could be successfully microdissected and fungal septa could be collected by LPC, but the material cannot be examined by electron microscopy.

Laser Microdissection and Laser Pressure Catapulting of Fungal Septa from Lowicryl HM20 Embedded Mycelium Sections of HPF-fixed, freeze-substituted and lowicryl HM20 embedded R. solani mycelium were labeled with WGA-gold, and subsequently silver enhanced (Figure 1). Cell walls, septa and septal pore caps were decorated with gold-Ag particles as visualized by transmission electron microscopy (TEM) (Figure 6). The WGA-gold-Ag labeled sections were placed on a PEN-membrane covered microscope slide with the labeled side upwards (Figure 2) and septal regions were selected on the computer screen. After laser microdissection, a formvar film-coated single slot grid was placed on top of the single hole grid that enclosed the HM20 section (Figure 2). Examination of the grids in the scanning electron microscope showed a thickness of about 0.045 mm each, thus resulting in a total transfer distance of about 0.09 mm between the sample and the formvar film. The microdissected material was transferred upwards by LPC, and collected at the formvar film as was observed by light microscopy (inset of Figure 7A). The microdissected stack consisting of PEN membrane and the microdissected septum

70 Laser microdissection of fungal septa

A B

Figure 4 – Transmission electron micrograph of a 150 nm cryo-section of picric acid and formaldehyde fixed Rhizoctonia solani hyphae. A) Overview of longitudinally sectioned hyphae. The box indicates the septum (S) with the dolipore (DP) and associated septal pore caps (SPC). B) Higher magnification of a part of the septum as shown in (A). Perforate SPCs cover the dolipore septum at both sides. Electron-dense plugging material (*) is present at the orifice of the dolipore. Bar represents 10 µm (A) and 1 µm (B).

A B C Figure 5 – Light microscopic images of 1 µm thick cryo-sections of Rhizoctonia solani mycelium stained with toluidine blue before (A) and after (B) laser microdissection. The septum of interest (*) was microdissected along a predrawn circle (B) and subsequently laser pressure catapulted (C). Bar represents 10 µm.

Figure 6 – Transmission electron micrograph of part of a HPF-fixed, freeze-substituted and HM20 embedded Rhizoctonia solani hypha that was labeled with WGA-gold, and subsequently silver- enhanced. Cell walls (CW), septum (S) and septal pore caps (SPC) are decorated with WGA-gold-Ag. Bar represents 2 µm.

71 Chapter 3

A B

D

C E

Figure 7 – Scanning electron micrographs of laser microdissected septa of Rhizoctonia solani that were WGA- gold-Ag labeled and collected by laser pressure catapulting (LPC) to a formvar film. A) Low magnification of a microdissected stack that consisted of PEN membrane and part of a HM20 embedded hypha with a septum. Inset shows light microscopic visualization of microdissected septa collected on a formvar film by LPC. B) Low magnification of a laser microdissected stack, in which the LPC shot introduced a hole (*) at the site where the LPC shot hit the microdissected stack to transfer it upwards. C) High magnification of a microdissected septum showing the gold-Ag particles (arrows) that decorated the cell walls, septum and septal pore caps and a hole that was introduced by an LPC shot (*). D) Higher magnification of a part of the septum as shown in (C) with WGA-gold-Ag labeling of septum and both septal pore caps. E) Higher magnification of a part of the septum showing WGA-gold-Ag labeling of cell wall and septum. Bar represents 5 µm (A, B), 10 µm (inset A), 2 µm (C), 0.5 µm (D), and 1µm (E). CW = cell wall, S = septum, SPC = septal pore cap, P = PEN-membrane.

72 Laser microdissection of fungal septa in HM20, appeared electron impermeable in the TEM at 100kV (result not shown), but could be examined by scanning electron microscopy (Figure 7). Furthermore, at the site where the LPC shot hit the microdissected stack to transfer it upwards (Figure 7B, C), a hole was visible occasionally (Figure 7B, C). Examination at high magnification showed WGA-gold-Ag labeling of the cell walls, septa, and septal pore caps at the surface of the microdissected HM20 section (Figure 7C – E).

DISCUSSION

Laser microdissection is a powerful tool to obtain pure samples of specific cells or subcellular structures and is more and more routinely used in animal and plant studies. Surprisingly, laser microdissection has not been used to study fungi or fungal subcellular structures. Although fungal cells are much smaller compared to animal or plant cells, they can be viewed with a light microscope, which is a prerequisite for laser microdissection. We report here the use of laser microdissection to cut fungal hyphae. Dolipore septa with associated septal pore caps were laser microdissected and collected by laser pressure catapulting (LPC) from hyphae of Rhizoctonia solani as was shown by scanning electron microscopy. The presented isolation method allows future biochemical analysis of the dolipore septum and septal pore caps, necessary for understanding its function in the fungal cell.

Fixation of the R. solani mycelium is necessary to prevent leakage of the cellular content from the hyphae during microdissection. Ethanol and formaldehyde fixations are often used in laser microdissection studies (Ahram et al., 2003). Precipitative fixatives (e.g. ethanol, acetone) are preferred above cross-linking fixatives (e.g. formaldehyde, glutaraldehyde), because the cross-linking reduces the amount of extractable DNA, RNA, and proteins (Day et al., 2005). However, ethanol fixation resulted in poor morphology of the septal region, and therefore a 2% formaldehyde-containing fixative was used instead. Low percentage formaldehyde fixations form reversible cross-linkings, and future DNA, RNA and protein extractions of the microdissected hyphae will be possible (Baschong et al., 1983; Finke et al., 1993; Pinzani et al., 2006). Future studies have to determine the most optimal fixative that gives both a good histological detail of the tissue and good recovery of the biomolecules of interest (Nelson et al., 2006).

Though we were able to microdissect the septal area from fixed R. solani hyphae in buffer, we could not collect these selected parts by LPC. Probably, the LPC energy was not powerful enough to overcome the water surface tension of the buffer to catapult the septum out of the buffer. Furthermore, the LPC energy was transferred to a larger area than the selected point of catapulting and changed the position of the sample. To

73 Chapter 3 collect the microdissected material in buffer, optical tweezers should be used (reviewed by Grier, 2003), for example the PALM MicroTweezers (P.A.L.M. Microlaser Technologies AG). When a laser microdissection microscope is not equipped with optical tweezers, sectioned tissue is needed to overcome the collection difficulties. For this reason, we used cryo-ultramicrotomy or a combination of high pressure freezing, freeze substitution and low temperature embedding.

Cryo-sectioning according to Tokuyasu (1973) is a technique compatible with mild fixation without other denaturing steps involved, like dehydration with organic solvents or embedding in resins. This makes cryo-sectioned tissue useful for future extraction of DNA, RNA and proteins. Although laser microdissection of the cryo- sections of R. solani hyphae was successful, we could not verify whether fungal septa were collected after laser pressure catapulting. During its transport to the collection cap by the LPC shot, the microdissected material became wrinkled, hence, impossible to examine by light or electron microscopy. To examine the microdissected septa by electron microscopy, to confirm the isolation procedure, we used sections of lowicryl HM20 embedded R. solani mycelium that were labeled with WGA-gold-Ag, a lectin that binds the N-acetylglucosamine residues in cell walls and septa (Benhamou et al., 1993). Furthermore, the sections were placed on a PEN-membrane covered slide to give them extra support during the transfer. After laser microdissection and transfer of the septa by LPC to a formvar film-coated single slot grid (Figure 2), we observed by scanning electron microscopy that most microdissected septa did not transfer vertically, but instead, turned over during the transfer (Figure 2). WGA-gold-Ag labeling of the cell walls, septum and septal pore caps verified microdissection and transfer by laser pressure catapulting of fungal septa.

Here, we show the potential of laser microdissection as an isolation tool in fungal cell biological research. Specific hyphae or hyphal cells could be isolated from a mycelium in order to analyze their biomolecules. Alternatively, parts of hyphae such as tips or subapical regions or organelles such as individual nuclei can be isolated and studied individually.

ACKNOWLEDGMENTS

We express our thanks to Dr. Elly Hol (Netherlands Institute for Brain Research) for technical advice and using the PALM Microbeam system (Figures 3 and 5). We would also like to thank DSM Food Specialties (Delft, The Netherlands) and SenterNovem (The Netherlands) for using their PALM Microbeam system (Figure 7).

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Chapter 4

Enrichment of Perforate Septal Pore Caps from the Basidiomycetous Fungus Rhizoctonia solani by Combined Use of French Press, Isopycnic Centrifugation, and Triton X-100

Kenneth G.A. van Driel, Arend F. van Peer, Han A.B. Wösten, Arie J. Verkleij, Teun Boekhout & Wally H. Müller

Published in Journal of Microbiological Methods doi:10.1016/j.mimet.2007.09.013 Chapter 4

ABSTRACT

Septal pore caps occur in many filamentous Basidiomycota fungi. They are located at both sides of the dolipore septum and are at their base connected to the endoplasmic reticulum. The septal pore cap ultrastructure has been described extensively, but its composition and function are not yet known. To enable biochemical and functional analyses, we here describe an enrichment method for perforate septal pore caps from Rhizoctonia solani. Our method is based on the combined use of French press, isopycnic centrifugation using a discontinuous sucrose gradient, and a treatment with Triton X-100. Enrichment was monitored by the use of scanning electron microscopy and transmission electron microscopy. Using the same isolation method, smaller septal pore caps were isolated from two other basidiomycetous fungi as well. Furthermore, we showed that pore-occluding material co-purified with the septal pore caps. This observation supports the hypothesis that septal pore caps play a key role in the plugging process of the septal pores in filamentousBasidiomycota fungi.

INTRODUCTION

The septal pore cap (SPC) or parenthesome is a membranous structure located at both sides of the dolipore septum in many filamentousBasidiomycota fungi (Moore & McAlear, 1962; Bracker & Butler, 1963). In 1958, Girbardt described for the first time the SPC ultrastructure at the septum of Trametes versicolor by the use of transmission electron microscopy (cited as Polystictus versicolor; Girbardt, 1958). Since then, many studies on the SPC ultrastructure have followed (e.g. Bracker & Butler, 1963; Setliffet al., 1972; Khan & Kimbrough, 1982; Patrignani & Pellegrini, 1986; Wells, 1994; Moore, 1996; Keller, 1997; Müller et al., 1998b, 2000b). These studies have shown that the ultrastructure of SPCs is diverse and can be divided into the following types: the vesicular (saccular, cupulate or tubular) type, the imperforate (or continuous) type, and the perforate type (McLaughlin et al., 1995). The absence or presence of a SPC, the SPC-type, and septal morphology are important markers that reflect the main phylogenetic groups in the Basidiomycota (Khan & Kimbrough, 1982; Wells, 1994; McLaughlin et al., 1995; Moore, 1996; Müller et al., 2000b, Weiss et al., 2004; Chapter 2).

Although the ultrastructure of SPCs has been studied extensively, its precise function is still a matter of speculation. As the base of SPCs is continuous with the endoplasmic reticulum (ER), the SPC has been proposed to be a subdomain of the ER (Girbardt, 1961; Bracker & Butler, 1963; Müller et al., 1995a). However, differences in calcium-affinity sites between the ER and SPCs have been observed by the use of zinc-iodin-osmium tetroxide staining (Müller et al., 1995a, 1998a). Possibly, SPCs act as a repository for

80 Enrichment of septal pore caps from Rhizoctonia solani certain proteins that are produced and processed in the ER, and that are translocated from the SPC to the septal pore when pore sealing is needed in cases of stress or hyphal damage (Müller et al., 1998a). Several other functions of SPCs have been suggested: they may act as a sieve to discriminate between organelles that pass the pore (Wilsenach & Kessel, 1965), they may guide cytoplasmic streams to the pore (Orlovich & Ashford, 1994), or they may function in protoplasmic streaming, by which SPCs protect the dolipore for accidental closing or diminishing the pore by organelles that could hit the sides of the swelling in case SPCs are absent (Bracker & Butler, 1964). Furthermore, in Schizophyllum commune, Rhizoctonia solani, and Pisolithus tinctorius filamentous structures were observed that connect the inside of the SPC with the pore-occluding material (Orlovich & Ashford, 1994; Müller et al., 1999, 2000a). These observations suggest that SPCs play a key role in plug formation after hyphal damage or stress and consequently, are of importance in intercellular communication within the hyphae (Thielke, 1972; Aylmore et al., 1984; Markham, 1994; Müller et al., 1999, 2000a).

The many ultrastructural studies of SPCs have led to a number of hypotheses on SPC functioning. Biochemical analysis of SPCs may help to understand its function in the hyphal cells of the Basidiomycota. However, an isolation procedure of SPCs necessary to biochemically study these organelles has never been published. We here describe for the first time a method to enrich SPCs from the basidiomycetous fungus R. solani. Using the same procedure, we could also isolate SPCs from two other species belonging to the Rhizoctonia sensu lato (s.l.) complex (Müller et al., 1998b), namely Thanatephorus cucumeris and Ceratobasidium cornigerum. Furthermore, we showed that a structural complex consisting of SPCs attached by filaments to pore-occluding material could be isolated. The presented method will also allow the future isolation and biochemical analysis of proteins of perforate SPCs from different basidiomycetous fungi.

MATERIALS & METHODS

Organisms, Media and Culture Conditions Rhizoctonia solani (CBS 346.84), Thanatephorus cucumeris (CBS 700.82), and Ceratobasidium cornigerum (CBS 132.82) were grown on malt extract agar (Oxoid, Hampshire, UK) at 25oC for 4 days. A Sorvall omni-mixer (Kendro Laboratory Products GmbH, Langenselbold, Germany) was used for 10 sec at speed 4 to homogenize the culture in 100 ml complete medium (20.0 g glucose, 2.0 g peptone L37 (Oxoid), 2.0 g yeast extract (Difco, Detroit,

MI, USA), 0.5 g MgSO4.7H2O, 0.46 g KH2PO4, 1.0 g K2HPO4 per liter) containing 100 mg/ l Penicillin-G (Yamanouchi Pharma, Leiderdorp, The Netherlands) and 100,000 units/ l Streptomycin (Radiumfarma-Fissiopharma, Naples, Italy). After growing for 48 hr at 175 rpm and 25oC, the culture was again homogenized in an omni-mixer for 10 sec at

81 Chapter 4 speed 4. Thereafter, 10 ml of this homogenate was used to inoculate 100 ml complete medium and growth was allowed for 3 days at 175 rpm and 25oC.

Subcellular Fractionation by Isopycnic Centrifugation The mycelium from three-days-old submerged cultures was harvested by centrifugation at 2000 rpm and 4oC for 3 min in a Mistral 400 centrifuge (MSE Scientific Instruments, West Sussex, UK). Subsequently, the mycelium was washed twice in HEPES/KAc buffer (20 mM HEPES, pH 6.8, 50 mM potassium acetate (KAc), 200 mM D-sorbitol, 1 mM EDTA) (Rieder & Emr, 2000) and resuspended in half a volume of HEPES/KAc buffer supplemented with 1/200 volume of protease inhibitor cocktail (Sigma-Aldrich, St. Louis, MO, USA). The mycelium was disrupted by two passages of 500 PSI (equivalent to 3447 kPa) through a French press (American Instrument Company, Silver Spring, MD, USA), resulting into homogeneous fungal cell extracts.

For isopycnic centrifugation, 12 ml of fungal cell extracts was layered on top of a discontinuous gradient consisting of 8 ml 70% (w/v), 8 ml 50% (w/v), and 8 ml 30% (w/v) sucrose in HEPES/KAc buffer in a 38.5 ml polyallomer thinwall ultracentrifuge tube (Herolab GmbH, Wiesloch, Germany). After centrifugation at 85,000 × g (21,500 rpm) and 4oC for 90 min in a Centrikon T-2180 ultracentrifuge (Kontron Instruments, Watford, UK), fractions were collected from underneath using a pasteur pipette with a 180 degrees bent tip. The fractions were diluted with HEPES/KAc buffer containing 2% (w/v) Triton X-100 (GE Healthcare, Uppsala, Sweden) to a final sucrose concentration of 10% (w/v). After one hour on ice, fractions were centrifuged at 5,000 × g (5,300 rpm) and 4oC for 1 hr. The resulting pellet was resuspended in 1 ml HEPES/KAc buffer and further processed for electron microscopy as described below. To analyze the supernatant (remaining after the 5,000 × g centrifugation step) for the presence of SPCs, the supernatant was centrifuged at 100,000 × g (23,500 rpm) and 4oC for another 45 min, and subsequently the resulting pellet was resuspended in 1 ml HEPES/KAc buffer and processed for electron microscopy. A flow chart of the SPC-enrichment procedure is presented in Figure 1.

Transmission Electron Microscopy The obtained fraction samples were fixed chemically or by high-pressure freezing. For the chemical fixation, samples were fixed in 1.5% glutaraldehyde (Agar Scientific Ltd, Essex, UK) buffered with 20 mM HEPES, pH 6.8, overnight at o4 C. Samples were washed twice in PBS and post-fixed in 1% (w/v) aqueous osmium tetroxide (EMS, Hatfield, PA, USA) for one hour at room temperature. Subsequently, the samples were washed twice in distilled water, followed by gradual dehydration in a series of ascending concentrations of acetone, namely 60% (v/v), 70% (v/v), 80% (v/v), 90% (v/v), and three times of 100% acetone containing 1% (v/v) acidified 2,2-dimethoxypropane (DMP) for 30 min each.

82 Enrichment of septal pore caps from Rhizoctonia solani

Figure 1 – Flow diagram of the septal pore cap-enrichment procedure.

The samples were gradually infiltrated in Spurr’s resin (Spurr, 1969) using a series of 25% (v/v), 50% (v/v), 75% (v/v), 90% (v/v), 95% (v/v) Spurr’s resin in 100% acetone containing 1% (v/v) acidified DMP and twice 100% Spurr’s resin respectively for 1 hr each. Finally, the samples were embedded in freshly prepared 100% Spurr’s resin and polymerized at 65oC for 48 hr in BEEM capsules (EMS).

83 Chapter 4

Mycelium of R. solani was subjected to high-pressure freezing (HPF) and freeze- substitution as described by Müller et al. (1998a). To high-pressure freeze the SPC- enriched fraction samples, an aluminum lecithin-coated planchette (100 µm-deep well) used for HPF (Engineering Office M. Wohlwend GMbH, Sennwald, Switzerland) was dipped into the SPC-enriched pellet and we drew material from the bottom of the tube. A second lecithin-coated planchette (300 µm-deep well) was placed with the flat side as a lid on top of the first planchette, thus subjecting 100 µm thick SPC-enriched pellet material to HPF by the use of a Leica EM HPF (Leica Microsystems, Vienna, Switzerland). After separating the planchettes in liquid nitrogen, the samples were freeze-substituted in a mixture of 1% (w/v) osmium tetroxide, 3% (v/v) glutaraldehyde, and 0.3% (w/v) uranyl acetate in anhydrous methanol at -85oC for 2 days. After rinsing with methanol, the samples were gradually infiltrated using a series of 25% (v/v), 50% (v/v), 75% (v/v) lowicryl HM20 (EMS) for 2 hr each, and three times 100% lowicryl HM20 for 1 day each. Finally, the samples were low-temperature embedded in lowicryl HM20, and were polymerized after 48 hours at -35oC and 24 hr at room temperature under UV light.

Sections of about 90 nm and 350 nm were cut with a diamond knife (Diatome, Hatfield, PA, USA) using an ULTRACUT E ultramicrotome (Leica Microsystems, Vienna, Austria). The sections were picked up with formvar film-coated, carbon-stabilized copper grids (hexagonal 150 mesh Veco grids, EMS). Sections were contrasted with 4% (w/v) aqueous uranyl acetate (Merck) for 10 min and 0.4% (w/v) aqueous lead citrate (Merck) for 2 min (Venable & Coggeshall, 1965). The sections were viewed with a TECNAI 10 (FEI Company, Eindhoven, The Netherlands) transmission electron microscope at an acceleration voltage of 100 kV.

Scanning Electron Microscopy A small aliquot of the R. solani SPC-enriched fraction was placed on a formvar film-coated, carbon-stabilized grid and incubated for 10 min at room temperature. Thereafter, excess of fluid was removed carefully by touching the edge of the grid with a piece of Whatman paper, and subsequently the grid was air-dried overnight. Then, it was incubated 2 min with 4% (w/v) aqueous uranyl acetate, and subsequently washed thoroughly in distilled water. The grid was dried carefully with a piece of Whatman paper and further air-dried. Finally, the grids were mounted on a stub, coated with 5 nm Pt/Pd by using a Cressington sputter coater 208HR (Cressington Scientific Instruments, Watford, UK), and viewed in a XL30 scanning electron microscope (FEI Company) at an acceleration voltage of 15 kV and a working distance of about 8 mm.

84 Enrichment of septal pore caps from Rhizoctonia solani

Figure 2 – Transmission electron micrograph of 350 nm thick sections of the dolipore septum in a high-pressure frozen, freeze-substituted and lowicryl HM20 embedded hypha of Rhizoctonia solani showing septal pore caps (SPCs) covering the dolipore (DP). The dolipore channel may be open, allowing passage of mitochondria (A) or the dolipore channel can be plugged with electron-dense occluding material (B). Filaments (arrows) are present between SPCs and the plugging material. O = occluding material. Bars represent 500 nm.

RESULTS

Rhizoctonia solani has dolipore septa with perforate SPCs of about 1.6 to 2.0 µm in diameter as was observed in high-pressure frozen (HPF) and freeze-substituted mycelium examined by transmission electron microscopy (TEM) (Figure 2). The dolipore channel may be open allowing passage of small organelles, i.e. mitochondria (Figure 2A), or is plugged with electron-dense pore-occluding material (Figure 2B). After two passages of 500 PSI (equivalent to 3447 kPa) through a French press, most of the harvested R. solani mycelium was broken as was examined by light microscopy (Figure 1) and TEM (result not shown). The resulting cell homogenate was subjected to isopycnic centrifugation in a discontinuous 30-50-70% sucrose gradient. Cell walls were pelleted after centrifugation, while SPCs were mainly found on top of the 70% sucrose layer (Figure 3A). Furthermore, many membrane vesicles were observed, and likely are formed as a result of the French press treatment to cell organelles and plasma- and vacuolar membranes. After treatment of this SPC-containing fraction with 2% Triton X-100, followed by centrifugation at 5000 × g, the amount of membrane vesicles was reduced and SPCs were further enriched (Figure 3B). Few vesicles, but no SPCs were observed in the supernatants (result not shown).

Scanning electron microscopy showed that isolated SPCs were flattened and had a diameter of approximately 1.5 µm and two to four perforations of about 400 to 600 nm

85 Chapter 4

86 Enrichment of septal pore caps from Rhizoctonia solani

(Figure 4). Furthermore, the base of the SPC was continuous with membranes, which may be part of endoplasmic reticulum (ER) (Figure 4, inset). TEM showed that thin- sectioned chemically fixed isolated SPCs maintained their characteristic dome-shaped structure after the enrichment procedure (Figure 5) and could be co-isolated with occluding material (Figure 5B-D). Additional structures that were necessary to retain the SPC dome-shape were not observed in SPCs isolated without plugging material (Figure 5A). In the isolated complexes of SPC and occluding material, putative filaments occurred in between the SPC and the occluding material, which were visualized as a grey zone in chemical fixed SPC-enriched samples (Figure 5B). An electron-translucent zone was visible between the putative filaments and the inside of the SPC (Figure 5B). To examine the ultrastructure of the isolated SPCs in more detail, SPC-enriched fractions from R. solani were fixed by HPF, followed by freeze-substitution. Examinations of these samples showed that isolated SPCs were connected to the occluding material by

Figure 4 – Scanning electron micrographs of the septal pore cap (SPC) enriched fraction of Rhizoctonia solani. SPCs are encircled. The inset shows a higher magnification of an isolated SPC with putative endoplasmic reticulum attached at the base (arrows). Bars represent 1 µm.

Ô Figure 3 – Transmission electron micrographs of the subcellular fraction of Rhizoctonia solani isolated from above the 70% sucrose layer before (A) and after (B) treatment of the fraction with 2% Triton X-100 followed by centrifugation at 5,000 × g for 1 hour. Cell organelles other than septal pore caps (SPCs) (encircled) could not be identified. SPCs were co-purified with plugging material (*). Arrows indicate possible isolated plug material. Bars represent 1µm.

87 Chapter 4

Figure 5 – Transmission electron micrographs of chemical fixed (A and B) or high-pressure frozen (HPF) and freeze-substituted (C and D) septal pore cap (SPC) enriched fractions of Rhizoctonia solani. A) Transverse section of an isolated SPC showing 3 perforations. B) Median section of an isolated SPC that is connected to electron-dense occluding material (o) via a putative filamentous network (black arrows). An electron- translucent zone is visible between the inside of the SPC and the filaments (white arrows). HPF-fixed and freeze-substituted SPC-enriched samples (C and D) show clearly the filamentous structures (arrows) that connect the electron-dense pore-occluding material (o) to the inside of the SPCs. Plug morphology varied from a loose structure (C), to a compact and dense structure (D). The plug with a loose structure (C) was connected to a more extensive filamentous network than the compact and dense structured plug (D). An amorphous layer is visible at the inside of the SPC (white arrows). Bars represent 200 nm. a filamentous network (Figure 5C, D). Furthermore, HPF-fixed and freeze-substituted SPC-enriched samples gave a detailed view of the plug morphology that varied from loosely structured (Figure 5C) to compact and densely structured (Figure 5D). Between the inside of the SPC and the extensive filamentous network an amorphous layer was visible (Figure 5C) that was less visible in the isolated SPC in Figure 5D.

To investigate whether our method could be applied for the isolation of SPCs from other basidiomycetous species, we subjected Thanatephorus cucumeris and Ceratobasidium cornigerum to the combined use of French press and isopycnic centrifugation. SPCs of T. cucumeris and C. cornigerum are smaller than those of R. solani and are about 1200 nm and

88 Enrichment of septal pore caps from Rhizoctonia solani

850 nm in diameter respectively. Cell homogenates were successfully generated and for both, SPCs could be isolated after isopycnic centrifugation on top of the 70% sucrose layer (result not shown). However, the SPC-enriched fraction of C. cornigerum contained also mycelial fragments and remnants of broken cell walls, whereas T. cucumeris cell walls and mycelial fragments passed the 70% sucrose layer and were pelleted at the bottom of the ultracentrifuge tube, like in R. solani (result not shown). Moreover, C. cornigerum SPCs were also found in the 50% sucrose layer (result not shown). SPC fractions from C. cornigerum and T. cucumeris contained complexes of SPCs co-isolated with electron-dense occluding material as observed in R. solani SPC-enriched fractions (result not shown).

DISCUSSION

A wealth of information is available on the ultrastructure of SPCs in Basidiomycota, but about the function one can only speculate at present. The ultrastructure of the SPC in Rhizoctonia solani has been extensively studied by transmission electron microscopy (TEM) (Bracker & Butler, 1963, 1964; Setliff et al., 1972), scanning electron microscopy (SEM) (Lisker et al., 1975; Müller et al., 1998b) and automated electron tomography (Müller et al., 2000a). However, SPCs have never been subjected to biochemical analysis of the proteins residing in the SPCs. Prior to analysis of these SPC-related proteins, a method is needed to enrich SPCs. Ideally, this method must also be applicable to other basidiomycetous fungi.

Fungal organelles can be isolated using several approaches. For example, Penicillium chrysogenum microbodies were isolated by protoplasting followed by isopycnic centrifugation (Müller et al., 1995b). Protoplasting does not affect the structural integrity of SPCs (Müller et al., 1998a), however, the yield of SPC-containing protoplasts is low and release of SPCs from protoplasts was unsuccessful (E. Boon & W.H Müller, unpublished results). Woronin bodies from Neurospora crassa were enriched after the cells were frozen in liquid nitrogen and ground to a fine powder that was separated on a sucrose cushion (Jedd & Chua, 2000). This study led to the findings that Woronin bodies are pre-formed peroxisomes that consist of HEX-1 protein and are necessary for septal pore sealing in filamentous Ascomycota (Jedd & Chua, 2000). More recently, laser microdissection by the use of a PALM laserbeam system resulted into isolated septal regions of R. solani (Van Driel et al., 2007). Though fungal septa including SPCs were successfully isolated, laser microdissection is a very laborious technique that is not available in every laboratory. Furthermore, the isolation of proteins from sectioned fungal septa has not been optimized yet. Compared to described isolation methods, the success of our SPC isolation method is based on the combination of French press, isopycnic centrifugation, and a treatment with Triton X-100 that resulted into a high

89 Chapter 4 yield of isolated SPCs (Figure 3B).

Cell homogenates of R. solani, T. cucumeris, and C. cornigerum were prepared by passage of the mycelium through a French press. We experienced that the smaller hyphae of C. cornigerum were more resistant to the pressure of the French press than the broader hyphae of R. solani and T. cucumeris, resulting in more mycelial fragments present in the fractions. Therefore, to generate cell homogenates from other fungi one may need additional passages or higher PSI values to break all the cells. By combining the French press with isopycnic centrifugation we could isolate the different sized SPCs as visualized by TEM. However, we cannot rule out that by the inclusion of the French press step in our method, parts of the SPCs will be ripped into smaller components that are not recognized in the SPC-enriched fraction by TEM. Still, when isolating these SPC- components together with the enriched intact SPCs, this will result into a high yield of SPC components enhancing future biochemical studies.

To enrich the SPCs, we included a treatment with Triton X-100 detergent to solubilize the many membrane structures and vesicles that developed after French press treatment and isopycnic centrifugation. Triton X-100 is a non-ionic detergent often used to solubilize lipid membranes to isolate membrane proteins or detergent-resistant membrane domains like lipid rafts (reviewed by London & Brown, 2000). Interestingly, this Triton-treatment did not solubilize the SPC membranes, the filamentous network or the pore-occluding material as was only seen in the HPF-fixed and freeze-substituted SPC-enriched samples. Therefore, the membranes of the perforate SPCs of R. solani may not exclusively build-up by lipids that are solubilized by detergent, but they also may consist other components like sphingolipids or proteins that keep the SPC detergent- resistant. Wheat germ agglutinin labeling showed that N-acetyl glucosamine residues are present in SPCs (Benhamou et al., 1993; Van Driel et al., 2007) that may indicate the presence of glycoproteins in the SPC matrix, in the SPC-membranes or in both. Alternatively, a fibrous layer on top of the SPC (Figures 5C and 5D) that was also observed in other studies (C.E. Bracker, pers. comm.; Müller et al., 2000a) and the amorphous layer at the inside of the SPC (Figure 5C) that was also observed in S. commune (Müller et al., 1998a) may prevent SPC membrane solubilization by Triton X-100.

Next to analysis of the fractions by TEM, SEM was used as a fast and easy-to-use method to analyze the quality of the SPCs and the degree of enrichment. SEM analysis showed that SPCs in the R. solani SPC-enriched fraction were slightly reduced in size and the characteristic dome-shaped morphology was flattened compared to earlier observations (Müller et al., 1998b). Both observations may be a result of the air-drying procedure in the preparation for SEM analysis. In contrast, TEM analysis showed isolated SPCs with the characteristic dome-shaped morphology. In addition, isolated SPCs that were not

90 Enrichment of septal pore caps from Rhizoctonia solani associated with plugging material did not show any additional structures that were necessary to retain the dome-shaped structure of SPCs as was proposed by Orlovich and Ashford (1994). This was also demonstrated in free-lying SPCs in Schizophyllum commune protoplasts (Müller et al., 1998a), which showed the same morphology as in intact hyphal cells. These observations, together with the fact that the SPC structure was not broken down or solubilized after the enrichment procedure that includes passages through a French press and a Triton X-100 treatment, we conclude that SPCs are resistant to high pressures and non-ionic surfactants.

The isolation of structural complexes consisting of SPCs connected to pore-occluding material by a filamentous network shows the close interaction of these two structures. Chemically fixed fractions showed an electron-translucent zone between the inside of the SPC and the filaments. We assume this zone is due to the preparation, because in HPF-fixed and freeze-substituted samples, an amorphous layer was visible instead, which agrees with previous observations (Müller et al., 1998a). This may be material from the SPC that contributes to the build-up of filaments or pore-occluding material. We, however, cannot rule out that during the isolation method proteins may be trapped into this zone due to the many filaments that are present (Figure 5C). When few filaments are present, these proteins then can be easily washed off (Figure 5D). The filamentous network between SPCs and occluding material agrees with previously reported observations of SPCs in intact hyphae after automated electron tomography (Müller et al., 2000a). The filamentous connections between SPCs and pore-occluding material and the isolation of these three structures as a structural complex suggest that SPCs may take part in the plugging process of dolipores and thereby fulfill a crucial role in maintaining cell homeostasis. Furthermore, the filamentous network showed dynamics in density and was found to be more extensive when the plug was loosely structured (Figure 5C) instead of compact (Figure 5D). In addition, those parts of the SPC that are connected with the filaments show a less electron dense SPC matrix than the remainder of the SPC matrix (Müller et al., 2000a). This indicates that the filaments between SPC and occluding material may be involved in the formation of pore-occluding material.

91 Chapter 4 REFERENCES

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Müller, W.H., Stalpers, J.A., Van Aelst, A.C., De Jong, M.D.M., Van der Krift, T.P. & Boekhout, T. (2000b) The taxonomic position of Asterodon, Asterostroma and Coltricia inferred from the septal pore cap ultrastructure. Mycol. Res. 104, 1485 – 1491. Orlovich, D.A. & Ashford, A.E. (1994) Structure and development of the dolipore septum in Pisolithus tinctorius. Protoplasma 178, 66 – 80. Patrignani, G. & Pellegrini, S. (1986) Fine structures of the fungal septa on varieties of basidiomycetes. Caryologia 39, 239 – 250. Rieder, S.E. & Emr, S.D. (2000) Isolation of subcellular fractions from the yeast Saccharomyces cerevisiae. Pp. 3.8.45. In Current protocols in cell biology. Bonifacino, J.S., Dasso, M., Lippincott- Schwartz, J., Harford, J.B. & K.M. Yamada, K.M. (eds.), John Wiley and Sons, New York, USA. Setliff, E.C., MacDonald, W.L. & Patton, R.F. (1972) Fine structure of the septal pore apparatus in Polyporus tomentosus, Poria latemarginata, and Rhizoctonia solani. Can. J. Bot. 50, 2559 – 2563. Spurr, A.R. (1969) A low viscosity resin embedding medium for electron microscopy. J. Ultrastruct. Res. 26, 31 – 43. Thielke, C. (1972) Die Dolipore der Basidiomyceten. Arch. Mikrobiol. 82, 31 – 37. Van Driel, K.G.A., Boekhout, T, Wösten, H.A.B., Verkleij, A.J. & Müller, W.H. (2007) Laser microdissection of fungal septa as visualized by scanning electron microscopy. Fungal Genet. Biol. 44, 466 – 473. Venable, J.H. & Coggeshall, R. (1965) A simplified lead citrate stain for use in electron microscopy. J. Cell Biol. 25, 407 – 408. Weiss, M., Bauer, R. & Begerow, D. (2004) Spotlights on heterobasidiomycetes. Pp. 7 – 48. In Frontiers in Basidiomycete Mycology. Agerer, M., Piepenbring & M., Blanz, P. (eds.), IHW-Verlag, Berlin, Germany. Wells, K. (1994) Jelly fungi, then and now. Mycologia 86, 18 – 48. Wilsenach, R. & Kessel, M. (1965) On the function and the structure of the septal pore of Polyporus rugulosus. J. Gen. Microbiol. 40, 397 – 400.

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Chapter 5

SPC18, a Novel Septal Pore Cap Protein of Rhizoctonia solani Residing in Septal Pore Caps and Pore-plugs

Kenneth G.A. van Driel, Arend F. van Peer, Jan Grijpstra, Han A.B. Wösten, Arie J. Verkleij, Wally H. Müller & Teun Boekhout Chapter 5

ABSTRACT

Fungal septa play an important role in the biology of filamentous fungi. In the higher Basidiomycota (i.e. Agaricomycotina), the dolipore septum is associated with septal pore caps. Although the ultrastructure of the septal pore caps has been studied extensively, neither their biochemical composition nor their function are known. Here, we report the identification of the N-glycosylated protein SPC18 that was identified in the septal pore cap-enriched fraction of Rhizoctonia solani. Based on its N-terminal sequence the SPC18 gene was isolated. It encodes a protein of 158 amino acid residues, which contains a hydrophobic signal peptide for targeting to the endoplasmic reticulum, an N-glycosylation motif, and a putative endoplasmic reticulum-retention signal. Immuno-localization showed that SPC18 is not only present in the septal pore caps but also resides in plug material. These data supports the hypothesis that septal pore caps are derived from endoplasmic reticulum and that septal pore caps are involved in dolipore plugging, and thus contribute to hyphal homeostasis in Basidiomycota.

INTRODUCTION

Compartmentation of fungal hyphae dates back to the origin of the Dikarya (i.e. Ascomycota and Basidiomycota), which occurred approximately 500 to 600 million years ago (Berbee & Taylor, 2001; James et al., 2006; Hibbett et al., 2007). Fungal septa and septum-associated structures play an important role in the biology of filamentous Ascomycota and Basidiomycota, as they separate the fungal hyphae into functional cellular compartments, and give the hyphal filaments rigidity. In addition, they are involved in mycelial differentiation (Gull, 1978). The regularly formed septa have a central pore through which protoplasm is continuous, and organelles like mitochondria can pass (Moore & McAlear, 1962; Moore, 1997). Consequently, mycelia act as functional coenocytic systems, and need some mechanism to restrict the effects of cellular damage within the system. Therefore, a damaged hyphal compartment needs to be isolated from the rest of the mycelial body to prevent cell lysis. In filamentous Ascomycota (Pezizomycotina), Woronin bodies that are closely associated with the septum, seal the septal pores upon damage (Woronin, 1864; Markham & Collinge, 1987; Jedd & Chua, 2000). In Neurospora crassa it was shown that Woronin bodies are peroxisomes and contain the HEX-1 protein (Jedd & Chua, 2000). Homologues of this protein have been found in other Pezizomycotina like Aspergillus nidulans, A. oryzae, Botrytis cinerea, Trichoderma reesei and Magnaporthe grisea (Jedd & Chua, 2000; Tenney et al., 2000; Curach et al., 2004; Soundarajan et al., 2004; Juvvadi et al., 2007), but not in ascomycetous yeasts (Saccharomycotina) nor in Basidiomycota (Jedd & Chua, 2000; Tenney et al., 2000).

96 SPC18, a novel septal pore cap protein

Within the three main lineages of Basidiomycota, i.e. Pucciniomycotina (= Urediniomycetes; Swann & Taylor, 1995), Ustilaginomycotina (= Ustilaginomycetes; Swann & Taylor, 1995), and Agaricomycotina (= Hymenomycetes; Swann & Taylor, 1995), the septal pore morphology clearly differ (Patrignani & Pellegrini, 1986; Wells, 1994; McLaughlin et al., 1995; Bauer et al., 2001, 2006; Hibbett & Thorn, 2001; Swann et al., 2001; James et al., 2006). Pucciniomycotina and Ustilaginomycotina have hyphae with a septum morphology similar to that found in hyphae of Ascomycota, though without Woronin bodies (Swann et al., 2001; Bauer et al., 2001, 2006). Agaricomycotina, on the other hand, are characterized by dolipore septa that are associated with a septal pore cap (SPC) (Girbardt, 1958; Moore & McAlear, 1962; Bracker & Butler, 1963; Müller et al., 1998b, 2000b; Hibbett & Thorn, 2001). The endoplasmic reticulum (ER) is connected at the base of the SPC, thus suggesting that the SPC is a subdomain of the ER (Girbardt, 1961; Bracker & Butler, 1963; Müller et al., 1998a). The ultrastructure of SPCs has been studied extensively and based on these observations, several functions of the SPC have been proposed. SPCs may act as a sieve to discriminate between organelles that may pass the septal pore (Wilsenach & Kessel, 1965), guide cytoplasmic streams towards the pore (Orlovich & Ashford, 1994), and would function in protoplasmic streaming to protect the dolipore region in such way that protoplasmic streaming is not disturbed by organelles (Bracker & Butler, 1964). Alternatively, they may be involved in dolipore plugging (Thielke, 1972; Aylmore et al., 1984; Markham, 1994; Müller et al., 1999, 2000a).

In this study, we describe for the first time an SPC component. The glycoprotein SPC18 was identified in the SPC-enriched fraction of the filamentous basidiomycetous fungus Rhizoctonia solani. Immuno-labeling localized SPC18 in the SPC and in the pore-plugging material. Data support the hypothesis that SPCs are derived from ER and are involved in dolipore-plugging in hyphae of higher Basidiomycota.

MATERIALS & METHODS

Preparation of a Septal Pore Cap-enriched Fraction SPCs were enriched from Rhizoctonia solani (CBS 346.84) by isopycnic centrifugation as described by Van Driel et al. (2007b). Rhizoctonia solani was grown for 3 days in complete medium (20.0 g glucose, 2.0 g peptone L37 (Oxoid, Hampshire, UK), 2.0 g yeast extract

(Difco, Detroit, MI, USA), 0.5 g MgSO4.7H2O, 0.46 g KH2PO4, 1.0 g K2HPO4 per liter) at 175 rpm and 25oC. Mycelium was harvested and subsequently disrupted by using a French press (American Instrument Company, Silver Spring, MD, USA). The cell extract was subjected to isopycnic centrifugation on a discontinuos density gradient of 30-50-70% (w/v) sucrose. The SPC fraction on top of the 70% sucrose layer was treated with 2% (w/v) Triton X-100 (GE Healthcare, Uppsala, Sweden) followed by centrifugation at 5000 × g

97 Chapter 5 for 1 hr. SPCs were enriched in the resulting pellet.

Fluorescence Microscopy Endoplasmic reticulum was stained with ER-tracker (Invitrogen, Breda, The Netherlands) or Brefeldin A conjugated to BODIPY 558/568 (BODIPY-BFA) (Invitrogen) according to the manufacturer’s instructions. Mycelial samples were mounted on a glass slide for light microscopy using a Zeiss Axioskop microscope (Carl Zeiss AG, Oberkochen, Germany). The filter set BP365, FT395, and LP397 was used for ER-tracker, whereas filter set BP510-560, FT580, and LP590 was used for BODIPY-BFA.

Protein Analysis Protein content was determined using the bicinchoninic acid (BCA) method as provided by the supplier (Pierce, Rockford, IL, USA) using bovine serum albumin as a standard. Protein samples were concentrated by addition of nine volumes of ice-cold acetone. After 30 min incubation on ice samples were centrifuged for 10 min at 14,000 rpm at 4oC. The pellet was air-dried and subsequently dissolved in sample buffer (Pierce) containing dithiothreitol (DTT) with a final concentration of 50 mM. SDS-PAGE was carried out by the method of Laemmli (1970) in a 10-20% gradient Tris-HCl gel (Bio-Rad Laboratories, Hercules, CA, USA) in 25 mM Tris, 192 mM glycine, and 0.1% (w/v) SDS, pH 8.3. A molecular weight marker (Bio-Rad) was used with band sizes 10, 15, 20, 25, 37, 50, 75, 100, 150, and 250 kDa. Proteins were stained with Coomassie R350 (GE Healthcare).

Deglycosylation of the protein with the enzymes EndoH (New England Biolabs, Ipswich, MA, USA), and Sialidase A and O-glycanase (Prozyme, San Leandro, CA, USA) were performed according to the manufacturer’s instructions.

For N-terminal sequencing, proteins were subjected to SDS-PAGE and electroblotted onto an Immobilon-P transfer membrane (Millipore, Billerica, MA, USA). After staining with Coomassie R350, the protein of interest was cut out and subjected to N-terminal sequencing using an Applied Biosystems 476A (Applied Biosystems, Foster City, CA, USA).

Isolation of the Full-length SPC18 cDNA and Sequence Analysis A three-days-old submerged culture of R. solani was snap-frozen in liquid nitrogen, and subsequently grinded to a fine powder in a mortar. Total RNA was isolated using TRIzol reagent (Invitrogen, Paisley, UK). A forward degenerate primer SPC18#02 (Table 1) was designed according to the N-terminus of mature SPC18 and adjusted for codon usage of R. solani. Codon usage was determined by analyzing the nucleotides of the coding sequences of the laccase, gpd, ABC transporter (partial), aromatic polypeptide, and ste3- like (partial) genes of R. solani (Coding sequence of Gly: GGN GGH, Ile: ATH ATY, Val:

98 SPC18, a novel septal pore cap protein

GTN GTB; N = A, T, C, or G; H = A, C, or T; Y = C or T; B = C, G, or T). cDNA was made using ImProm-II Reverse Transcriptase (Promega, Madison, WI, USA), recombinant RNAsin Ribonuclease Inhibitor (Promega), the degenerate primer SPC18#02 and the oligo(dT) primer (RT-PCR: 5 min at 25oC, 60 min at 42oC, and 15 min at 70oC; PCR amplification: 4 min at 94oC, followed by 35 cycles of 20 sec at 94oC, 20 sec at 47oC, and 2 min at 72oC, followed by 10 min at 72oC). To obtain the full SPC18 cDNA sequence, 5’RACE was performed with the 5’/3’ RACE kit (Roche, Basel, Switzerland) using SPC18_SP1, SPC18_ SP2, and SPC18_SP3 primers (Table 1) according to manufacturer’s instructions. The genomic sequence of SPC18 was obtained by PCR with SPC18-fw1 and SPC18-rv1 primers (Table 1) using genomic DNA as a template (5 min at 95oC, followed by 35 cycles of 1 min at 95oC, 1 min at 54oC, and 2 min at 72oC, followed by 7 min at 72oC). PCR products were cloned into the pGEM-T vector (Promega) and sequenced using BigDye v3.1 Chemistry (Applied Biosystems) and analyzed on an ABI 730XL DNA analyzer (Applied Biosystems) according to the manufacturer’s instructions. Hydropathy analysis of the protein was done according to Kyte and Doolittle (1982) with a window of 11 using ProtScale (Gasteiger et al., 2005). ProtParam (Gasteiger et al., 2005) was used to calculate molecular weights, SignalP 3.0 (Bendtsen et al., 2004) was used to predict signal sequence, and ScanProsite (De Castro et al., 2006) was used to predict N-glycosylation sites. Similarity searches were done with BLAST against the sequences in Genbank and Swissprot (http:// www.ncbi.nlm.nih.gov/BLAST), and fungal genomes of the Broad Institute (http://www. broad.mit.edu/annotation/fgi/), DOE Joint Genome Institute (http://www.jgi.doe.gov/) and the J. Craig Venter Institute (http://www.tigr.org/tdb/fungal/). DNA hybridization (Southern) was done according to Schuren et al. (1993). Genomic DNA of R. solani was digested with HindIII and hybridized with the probe that was derived by PCR with SPC18-fw1 and SPC18-rv1 primers (Table 1) using genomic DNA as a template.

Primer Sequence (5’ – 3’) SPC18#2 GCNATGGGHGAYGTBATYGT

Oligo(dT) TTAAT22(AGC) SPC18_SP1 GGTGGTCGGTGATAATGAGG SPC18_SP2 GTCGGAAGGCTTGTTGATGT SPC18_SP3 GAACAACGTAGCCCTGAGGA RSFWTotalSPC18 TCAGCATATGATCTTCCCCGTCGCC RSRVSPC18 GTTAAAGCTTAGAGTTCCTTGAAGTCCTTG SPC18-fw1 AGCCTCCTCAGGGCTACG SPC18-rv1 GGATATGCAAAAAGGCAAGC Table 1 – Primers used in this study.

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Overexpression of SPC18 in Escherichia coli The coding sequence of SPC18 was amplified by PCR with RSFWTotalSPC18 and RSRVSPC18 primers (Table 1) using cDNA as a template (5 min at 94oC, followed by 35 cycles of 30 sec at 94oC, 30 sec at 55oC, and 1 min at 72oC, followed by 7 min at 72oC). The cDNA fragment was cloned into pET11a (Promega). The recombinant SPC18 protein was expressed in Escherichia coli BL21 by IPTG induction. Inclusion bodies were isolated by sonication of the cells in ice-cold 10 mM Tris/HCl, 3 mM EDTA and 1X protease inhibitor (Roche), followed by centrifugation at 4000 rpm for 10 min. Finally, inclusion bodies were purified by centrifugation 75000 rpm for 1 hr using a Beckman TL-100 and were dissolved in 20 mM Tris/HCL, 100 mM glycine, and 8 M urea.

Preparation of Antibodies Polyclonal antibodies were raised in rabbits using recombinant SPC18 that had been cut out of a SDS-PAA gel (Eurogentec, Seraing, Belgium). IgGs were purified from the serum by using the Melon Gel IgG Purification System (Pierce) according the manufacturer’s instructions.

Immunoblot Analysis After SDS-PAGE, proteins were electroblotted on to an Immobilon-P transfer membrane. After blocking with 3% TopBlock (Fluka, Buchs, Switzerland), the membrane was incubated with anti-recSPC18 antibodies (1:500). This was followed by incubation with goat-anti-rabbit antibodies conjugated with alkaline phosphatase (1:15.000) (Sigma- Aldrich). These antibodies were detected using 5-bromo-4-chloro-3-indolyl phosphate and nitro blue tetrazolium as substrate (Harlow & Lane, 1988).

Chemical Fixation and Embedding in Spurr’s Resin Samples were fixed overnight at o4 C in 1.5% glutaraldehyde (Agar Scientific LTD., Essex, UK) buffered with 20 mM HEPES, pH 6.8. This was followed by postfixation in 1% (w/v) aqueous osmium tetroxide (EMS, Hatfield, PA, USA) for 1 hr at room temperature. Samples were gradually dehydrated in a series of ascending concentrations of acetone, followed by a gradual infiltration in Spurr’s resin (Spurr, 1969). The resin was polymerized at 65o C for 48 hr in BEEM capsules (EMS).

High-pressure Freezing, Freeze-substitution, and Lowicryl HM20 Embedding Mycelium was high-pressure frozen and freeze-substituted as described by Müller et al. (1998a). Sections of about 90 nm and 350 nm were contrasted with 4% (w/v) aqueous uranyl acetate (Merck) for 10 min and 0.4% (w/v) aqueous lead citrate (Merck) for two min (Venable & Coggeshall, 1965). For immuno-gold labeling, mycelium was freeze- substituted with 0.1% (w/v) uranyl acetate and 0.01% (w/v) osmium tetroxide in acetone. SPC-enriched fractions were high-pressure frozen as described by Van Driel et al. (2007b)

100 SPC18, a novel septal pore cap protein and freeze-substituted with 0.1% (w/v) uranyl acetate and 0.01% (w/v) osmium tetroxide in acetone. Thereafter, samples were low-temperature embedded in lowicryl HM20 (Müller et al., 1998a).

Immuno-gold Labeling Lowicryl HM20 sections were incubated overnight with anti-recSPC18 antibodies (1:60). The antigen-antibody complexes were visualized by incubation with secondary goat- anti-rabbit antibodies (1:10) conjugated with 10 nm gold particles (Aurion, Wageningen, The Netherlands) for 1 hr at room temperature. Following immunolabeling, sections were contrasted with 4% (w/v) aqueous uranylacetate for 2 min.

Transmission Electron Microscopy Sections were viewed in a TECNAI 10 (FEI Company, Hillsboro, OR, USA) transmission electron microscope at an acceleration voltage of 100 kV.

RESULTS

Septal Pore Caps of Rhizoctonia solani Perforate SPCs of about 1.6 to 2.0 µm in diameter cover the dolipore septum of Rhizoctonia solani as was observed in high-pressure frozen and freeze-substituted mycelium examined by transmission electron microscopy (TEM) (Figure 1A, B). The dolipore channel may be plugged with electron-dense occluding material (Figure 1A),

Figure 1 – A, B) Transmission electron micrographs of a high-pressure frozen, freeze-substituted and lowicryl HM20 embedded Rhizoctonia solani hyphae showing dolipore septa associated with perforate septal pore caps (SPCs). Endoplasmic reticulum (ER) (arrows) is connected with the SPC. The dolipore-channel is plugged with electron-dense occluding material (o) (A). A mitochondrion (mi) is passing the dolipore channel that is surrounded by a ring-like structure of electron-dense material (B). Fluorescence micrograph showing hyphae stained with ER-tracker (C) and BODIPY-BFA (D). In both cases fluorescence was localized at the SPCs. Bar represents 500 nm in A and 200 nm in B.

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Figure 2 – A) Transmission electron micrograph of chemically fixed septal pore cap (SPC) enriched fraction with isolated SPCs encircled. Bar represents 1 µm. B) SDS-PAGE analysis of the subcellular fractions of Rhizoctonia solani obtained after isopycnic centrifugation. Lane 1, fraction on top of 30% sucrose layer; lane 2, fraction on top of 50% sucrose layer; lane 3, fraction 50% sucrose layer; lane 4, fraction on top of 70% sucrose layer (SPC-containing fraction); lane 5, SPC-enriched fraction (SPC-containing fraction treated with 2% Triton X-100). The arrow indicates the position of SPC18 protein. thereby blocking protoplasmic streaming. When open, the SPC and dolipore allow passage of mitochondria (Figure 1B). Endoplasmic reticulum (ER) parallel to the septum is connected to the SPC base (Figure 1A). In fact, the ER-specific fluorochromes ER-tracker and BODIPY-BFA stained the SPCs (Figure 1C, D).

Enrichment of Septal Pore Caps SPCs were enriched by a two-step procedure (Van Driel et al., 2007b). First, R. solani extracts were subjected to isopycnic centrifugation using a discontinuous 30-50-70% sucrose gradient. Second, the subcellular fraction on top of the 70% sucrose layer was treated with 2% Triton X-100 followed by centrifugation at 5000 × g (Figure 2A). Fractions obtained during the SPC-enrichment procedure were analyzed by SDS-PAGE (Figure 2B). A protein with an apparent molecular weight of about 18 kDa was enriched in the SPC- enriched fraction (Figure 2B) and was named Septal Pore Cap 18 protein (SPC18).

Molecular Cloning and Sequence Analysis of SPC18 The N-terminal amino acid sequence of the SPC18 protein was determined as RIIAPPSVPRAMGDVIVLQP. A partial cDNA of SPC18 was generated by RT-PCR using the oligo(dT) primer and the degenerate oligonucleotide primer SPC18#02 that were designed on the basis of the sequence of residues 11 – 17 (AMGDVIV). We performed 5’ RACE PCR to obtain the complete cDNA sequence of 722 bp (Figure 3). The open reading frame (ORF) consisted of 477 bp encoding a protein of 158 amino acids with a predicted molecular weight of 17664 Da. Homologous genes or proteins were not found in the sequence databases or in any of the sequenced fungal genomes. The N-terminal sequence

102 SPC18, a novel septal pore cap protein

Figure 3 – Nucleotide sequence and deduced amino sequence of the SPC18 gene from Rhizoctonia solani. Numbers on the left mark the position of the nucleotide residues. Numbers on the right mark the position of amino residues (mature protein). Single underline represents amino acid residues determined by Edman degradation. Amino acid residues of the signal peptide are shaded, the amino acid residues that are underlined with ~~~ represent the N-glycosylation site, and the –FKEL N-terminus is doubly underlined. The position shown by the triangle (s) is the cleavage site of the putative signal peptide, the residues in bold represent the position of the degenerate primer, the positions showed by V (1 and 2) are the two intron sites and an asterisk (*) represents the stop codon. The intron sequences are shown separately and the typical fungal 5’ and 3’ intron splice sequences are underlined. The Genbank accession number is EF636668.

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Figure 4 – Kyte and Doolittle hydropathy plot (window 11) of the SPC18 amino acid sequence. Positive and negative values indicate hydrophobic regions and hydrophilic regions, respectively. The hydrophobic region that corresponds to the signal peptide (SP) is indicated with a bar.

Figure 5 – SDS-PAGE analysis of EndoH- treated septal pore cap (SPC) enriched fraction (panel A), and Western blot analysis of SPC18 protein as detected by anti-recSPC18 antibodies (panel B). Lane 1, SPC-enriched fraction; lane 2, EndoH-treated SPC-enriched fraction. The upper and lower arrow indicate the position of SPC18 before and after EndoH-treatment, respectively. The asterisk (*) indicates the EndoH protein.

of SPC18 obtained with Edman degradation was recovered in the cDNA sequence with exception of the first amino acid residue that was found to be a Thr (T) instead of an Arg (R). Comparison of the cDNA and genomic sequence revealed two short introns of 51 bp and 52 bp (Figure 3) with 5’ and 3’ splice sites typical for filamentous fungi (Ballance, 1991; Johansson & Nyman, 1996). Southern analysis showed one band after hybridization (result not shown) indicating the presence of one copy of SPC18 in the R. solani genome.

The deduced SPC18 protein sequence encompasses a putative N-terminal signal sequence with a predicted signal sequence cleavage site between positions 17 and 18 of the protein (position 0 of the mature protein; Figure 3) that corresponded with the determined N-terminus of mature SPC18. Figure 4 shows the hydropathy plot of SPC18 using the Kyte and Doolittle algorithm. The strongly hydrophobic N-terminal domain is predicted to be a signal peptide that targets SPC18 to the ER. The cleaved form of SPC18 has a calculated molecular mass of 15963 Da. The difference in apparent

104 SPC18, a novel septal pore cap protein

Figure 6 – Transmission electron micrographs of high-pressure frozen, freeze-substituted, and lowicryl HM20 embedded Rhizoctonia solani hyphae (A) and septal pore cap-enriched fractions (B) labeled with anti-recSPC18 antibodies. Antigen-antibody complexes were visualized with secondary goat-anti-rabbit antibodies conjugated with 10 nm gold. Gold label (arrows) was present at the SPC. Bars represents 200 nm.

Figure 7 – Transmission electron micrographs of high-pressure frozen, freeze-substituted, and lowicryl HM20 embedded septal pore cap (SPC) enriched fractions labeled with anti-recSPC18 antibodies. Antigen-antibody complexes were visualized with secondary goat-anti-rabbit antibodies conjugated with 10 nm gold. Gold label was preferentially localized at the base of the SPC (A, B). Figure B is a higher magnification of part of Figure A. Furthermore, label (indicated by white arrows) was preferentially localized at the periphery of the plug as shown in this detailed image of a plug (C). Bars represent 200 nm in A and C, and 100 nm in B.

105 Chapter 5 molecular weight as determined by SDS-PAGE is due to glycosylation of the protein. Residues 137-139 (N–I–S) of the protein represent a potential N-glycosylation site (residues 119 – 121 of the mature protein; Figure 3). Indeed, enzymatic deglycosylation of SPC18 by EndoH resulted in a reduction of the molecular weight from 18 to about 16 kDa (Figure 5A). A treatment with the enzymes Sialidase A and O-glycanase that cleave simple O-linked sugar residues did not reduce the molecular mass any further (result not shown). Finally, the C-terminal sequence of SPC18 was determined as –FKEL, which may represent an ER-retention signal.

Localization of SPC18 Polyclonal antibodies were raised against recombinant SPC18 (anti-recSPC18). Western blot analysis showed that the anti-recSPC18 IgG antibodies (1:500) recognized SPC18 before and after EndoH-treatment (Figure 5B). Immuno-gold labeling was performed to examine the localization of the SPC18 protein. Anti-recSPC18 antibodies localized SPC18 at the SPCs in sections of hyphae (Figure 6A) and in sections of the SPC-enriched fraction (Figure 6B). The label intensity at the SPCs was denser in sections of SPC-enriched fractions than in hyphal sections. Label preferentially localized at the basal parts of the SPCs (Figure 7A, B). Furthermore, label was also observed at the plug material, again preferentially localized at the periphery of the plug (Figure 7C).

DISCUSSION

The divergence of the Dikarya (Ascomycota and Basidiomycota) from the non-septate basal lineages of the Fungi (i.e. Glomeromycota, Chytridiomycota, and Zygomycota) coincided with the occurrence of regularly septate hyphae (Figure 8). It appears that following these evolutionary innovation two different strategies evolved to plug the septal pores during stress or hyphal damage. Filamentous Ascomycota seal their septal pores with peroxisomal Woronin bodies that are associated with the septum (Woronin, 1864; Markham & Collinge, 1987; Jedd & Chua, 2000). In contrast, septal pore caps (SPCs) seem to fulfill this role in higher Basidiomycota (Thielke, 1972; Aylmore et al., 1984; Markham, 1994; Müller et al., 1999, 2000a). Fifty years after the first transmission electron microscopic observation of SPCs in Trametes versicolor (cited as Polystictus versicolor; Girbardt, 1958), we enriched and biochemically analyzed SPCs from Rhizoctonia solani for the first time. Here we describe a novel SPC protein that was located in the SPCs and in the septal pore plugging material in hyphae of R. solani.

SPC18, a protein with an apparent Mw of 18 kDa, was identified in the SPC-enriched fraction of R. solani. This protein has a hydrophobic signal sequence and is N-glycosylated. Both features are characteristic for proteins that are imported into the endoplasmic

106 SPC18, a novel septal pore cap protein

Figure 8 – Schematic overview of the phylogeny of main lineages in the fungal kingdom (adopted from James et al., 2006). The “lower” fungi, i.e. the Chytridiomycota, Zygomycota, and Glomeromycota, are aseptate or irregular septate. Regular septate fungi occurred about 500-600 million years ago (Ma) (Berbee & Taylor, 1991). Approximately 500 Ma, the Ascomycota diverged from Basidomycota (Berbee & Taylor, 1991) and two different septum-associated structures that have a role in septal pore plugging, namely Woronin bodies (peroxisomes) and septal pore caps (ER-derived), developed in both lineages respectively. reticulum (ER) (Blobel & Dobberstein, 1975; Helenius & Aebi, 2001; Keenan et al., 2001). The presence of glycoproteins in the SPCs was suggested before by detection of N-acetyl glucosamine groups by wheat germ agglutinin (WGA) labeling of SPCs in R. solani (Nagata & Burger, 1974; Benhamou et al., 1993; Van Driel et al., 2007a). The putative ER- nature of SPC18 together with the staining of SPCs by fluorochromes that label ER (Cole et al., 2000) strongly supports that SPCs are specialized subdomains of ER as has been proposed previously (Wilsenach & Kessel, 1965; Bracker & Butler, 1963; Müller et al., 2000a). Furthermore, the C-terminal –FKEL of SPC18 has characteristics for ER retention signals, namely the Glutamic acid-Leucine (EL) residues as in the C-terminal sequence (H/K/A/D)DEL (Munro & Pelham, 1987; Jackson et al., 1990; Teasdale & Jackson, 1996; Kasuya et al., 1999). Future studies should establish whether the –FKEL sequence can act as a (temporarily) ER retention signal.

Immuno-gold labeling localized SPC18 both at the SPC and the plug material. The SPC-enriched fraction labeled more intense than the hyphal sections. This difference could be explained by the fact that the SPC-enriched fractions were treated with Triton X-100, which might increase the accessibility of protein epitopes to the antibodies. Furthermore, it should be noted that the plug material was not always labeled, and that labeling was mainly observed at the periphery of the plug. Also the base of the SPC, which is the part that is connected with ER, was found to be preferentially labeled. This difference in label intensity suggests that other unidentified proteins reside in the central parts of SPC and plug. In addition, SPC18 residing in SPCs and in the plug may

107 Chapter 5 exist in different conformations that are detected differentially by the antibodies.

We hypothesize that the SPC18 protein is targeted to ER where glycosylation takes place. As the ER is continuous with the base of the SPC, SPC18 may then be transported from the ER lumen to the SPC (Müller et al., 1998b). Furthermore, modifications of the N- glycan group on SPC18 may occur in the ER or possibly also in the SPC that define the destiny of the protein. The transport route of SPC18 from ER to SPC to the cytoplasm at the dolipore is different from the secretory pathway where proteins are destined for Golgi compartment, endosomes, lysosomes, the plasma membrane or the extracellular environment and resembles more the retrotranslocation route from ER to cytoplasm of misfolded proteins (Molinari, 2007) or an unknown mechanism for the export of small glycoproteins (Suzuki & Lennarz, 2000). The filaments between the inside of the SPC and the plugging material (Orlovich & Ashford, 1994; Müller et al., 1999, 2000a, b; Van Driel et al., 2007b) may transport or guide SPC18 from the SPC to the orifice of the dolipore. During this transport, the SPC matrix may become more electron-translucent at sites where filaments are attached at the inside of the SPC (Mülleret al., 1998a, 2000a). Furthermore, other post-translational modifications of SPC18 may be of importance and need further investigation. A recent study of Juvvadi et al. (2007) showed that PKC phosphorylation of HEX-1 is needed for multimerization of the protein and proper localization in Woronin bodies in Ascomycota. As ATPase activity was found at the plug site (Schramm, 1971), phosphorylation of SPC18 or other unidentified proteins that are part of the SPC-dolipore-plugging complex might also play a role in the dolipore- plugging process of filamentousBasidiomycota fungi.

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111

Chapter 6

Summary and General Discussion Chapter 6

Filamentous fungi, including Ascomycota and Basidiomycota, form mycelia that consist of a network of apical growing hyphae. These hyphae are separated into cellular compartments by septa that have pores of about 70 to 500 nm in diameter. The cytoplasm within the mycelium is thus continuous (coenocytic). The septum in the higher Basidiomycota (i.e. Agaricomycotina) is flared towards the pore, forming a barrel-shaped structure that is called the dolipore. These dolipore septa are generally associated with septal pore caps (SPCs). The ultrastructure of the SPC has been relatively well studied by the use of electron microscopy, but the composition and the function of these organelles is unknown. Several functions have been suggested based on the morphological studies. SPCs may be involved in dolipore plugging to prevent lysis of the mycelium after hyphal damage either by producing pore-plug material or by acting as a repository of the pore- plug material. SPCs may also function as a sieve, guide organelles to the dolipore, or prevent organelles to block the entrance of the dolipore.

The aim of this Thesis was to identify components of the SPC and to assess whether the SPC ultrastructure reflects the recently revised phylogeny of the Agaricomycotina. It is shown that two orders in the Agaricomycotina, i.e. the Hymenochaetales and Cantharellales, contain two SPC morphologies. Thus, SPCs within the orders of the Agaricomycotina are not monomorphic per se. In addition, two methods were developed to isolate and enrich SPCs from Rhizoctonia solani. This basidiomycetous fungus was chosen because of its well studied, large SPCs that have a diameter of about 1600 to 2000 nm and pores of about 600 to 800 nm (Müller et al., 1998b). With one of the isolation methods an SPC protein was identified for the first time, which is named SPC18.

Septal Pore Caps as Taxonomic Character In 1958, Girbardt produced the first transmission electron image of an SPC in Trametes versicolor (cited as Polystictus versicolor; Girbardt, 1958). Since then, many ultrastructural studies of fungal septa followed, as it appeared that septa and SPC morphology were useful in fungal taxonomy. While the basal lineages in the Basidiomycota (i.e. Pucciniomycotina, Ustilaginomycotina) have a septal morphology more similar to that found in the filamentous Ascomycota, the Agaricomycotina on the other hand have dolipore septa that may be associated with SPCs. These membranous structures are connected at the base with endoplasmic reticulum (ER), and therefore considered as subdomains of the ER. Three morphologically distinct SPC-types have been distinguished. First, the vesicular (tubular or cupulate) SPC-type that consists of a group of vesicles or tubules surrounded in a hemisphere around the dolipore. Second, the imperforate (continuous) SPC-type that consists of a closed membrane structure covering the dolipore. This imperforate SPC may have a decreased thickness in the center or a minute pore (Müller et al., 2000b). And third, the perforate SPC-type that consists of a dome-shaped structure, which may have many small perforations as observed in Schizophyllum commune (Müller et al., 1998a)

114 Summary and general discussion or few large holes as observed in R. solani (Müller et al., 2000a).

Until recently, SPC morphology followed fungal phylogeny, and the orders in the Agaricomycotina contained only one SPC-type, either vesicular, imperforate, or perforate (Hibbett & Thorn, 2001; Wells & Bandoni, 2001). However, phylogeny of the Agaricomycotina was recently updated (Hibbett, 2006; Hibbett et al., 2007). Therefore, it was addressed whether SPC-morphology still correlates with the classification based on molecular phylogenetic analyses (Chapter 2). The SPC morphology of more than 350 species as reported in the literature was considered in relation with this new classification. Within the Agaricomycotina three classes (Tremellomycetes, Dacrymycetes, and Agaricomycetes) are recognized. In the most basal group, the Tremellomycetes, SPCs are absent or may have the vesicular morphology. The imperforate SPC-type occurs in the Dacrymycetes, whereas both imperforate and perforate SPC-types were found in the Agaricomycetes.

Contrary to previous reports (Hibbett & Thorn, 2001; Larsson et al., 2004; Moncalvo et al., 2006), Cantharellus was shown to have perforate SPCs (Chapter 2). The order Cantharellales within the Agaricomycetes thus contains species with perforate SPCs (Cantharellus, Sistotrema, and Ceratobasidiales) and imperforate SPCs (Tulasnella and Botryobasidium). The order Hymenochaetales also contains species with both SPC-types. Perforate SPCs were found in Hyphoderma praetermissum (Langer & Oberwinkler, 1993; Keller, 1997), Oxyporus latemarginatus (cited as Poria latemarginata; Setliffet al., 1972) and R. fibula (Chapter 2), while all other clades have imperforate SPCs (Moore, 1980; Langer & Oberwinkler, 1993; Müller et al., 2000b; Larsson et al., 2006). Within evolution SPC morphology thus changed several times. It seems that the ER-like strands observed above the dolipore in the Cystofilobasidiales (e.g. Itersonilia perplexans; Boekhout, 1991) may be ancestral to both the vesicular and imperforate SPC-type. Eventually, the perforate SPC has arisen several times in the Agaricomycetes. After the latter type has appeared in the Cantharellales and Hymenochaetales, it subsequently reversed to the imperforate SPC- type.

Biochemical Analysis of SPCs from Rhizoctonia solani A procedure was developed to enrich SPCs from R. solani (Chapter 4). It was also successfully applied for the enrichment of SPCs from Ceratobasidium cornigerum and Thanatephorus cucumeris. These fungi also belong to the Cantharellales and are characterized by SPCs of about 700 to 900 nm and 1400 to 1500 nm in diameter respectively (Müller et al., 1998b). The enrichment procedure was based on the combined use of French press, isopycnic centrifugation using a discontinuous sucrose gradient, followed by a treatment with Triton X-100. From the fact that this detergent dissolves lipid membranes but leaves the SPCs intact, it is concluded that the main

115 Chapter 6 part of these organelles is composed of protein and/or detergent-resistant membranes. SDS-PAGE of the SPC-enriched fraction of R. solani revealed several abundant protein bands (Chapter 5) and WGA labeling showed the presence of N-acetyl glucosamin residues at the SPCs (Chapter 3). This may indicate the presence of glycoproteins. One of these proteins running at an apparent molecular weight of 18 kDa was enriched in this fraction (Chapter 5). The encoding gene called SPC18 was isolated on basis of the N-terminal sequence. The gene encodes a protein of 158 amino acid residues, which contains both a signal peptide to direct it to the ER, and an N-glycosylation motif. Enzymatic deglycosylation confirmed the N-glycosylation of SPC18. Immuno- localization confirmed that SPC18 is located in the SPCs, but it was also shown to reside in pore-plug material. SPC18 was not detected on the filaments between the SPC and the pore-plug. The glycoproteins may originate from ER connected to or covering the SPC. In Chapter 5 evidence is presented that SPCs are closely connected or even part of the ER, as SPCs became highly fluorescent when hyphae ofR. solani were incubated with the ER-specific stains ER-tracker and BODIPY-Brefeldin A Chapter( 5). The presence of lipid material, possibly ER, surrounding the SPCs of S. commune was also biochemically shown (A.F. van Peer & H.A.B. Wösten, unpublished).

The pore-plug material co-purified with the SPCs. It was tightly attached to the inside of the SPCs with a filamentous network Chapter( 4). These filaments were observed in previous studies (Orlovich & Ashford, 1994; Müller et al., 2000a) and are, like the SPCs, resistant against the detergent Triton X-100, thus indicating the presence of proteins in the filaments. The function of these filaments is unclear. It has been suggested that they keep the SPC in its dome-shaped structure (Orlovich & Ashford, 1994). However, isolated SPCs with no additional structures (Chapter 4) or free-lying SPCs in protoplasts (Müller et al., 1998a) do not lose their characteristic form. Furthermore, the filaments could guide organelles and cytoplasmic streams towards the entrance of the dolipore (Orlovich & Ashford, 1994). Finally, a role in pore plugging has been suggested. Pore- plug material that is stored in SPCs could be transported via the filaments to the dolipore (Müller et al., 1998a, 2000a). This may happen in a pulsate manner as a banded pattern of electron-dense material has been observed between the SPC and the plug (Müller et al., 1998a, 2000a). It was shown that a dense network of filaments is observed when the plug is still amorphous, while the number of filaments decreases when the plug becomes more condensed, probably as a result of maturation (Chapter 5). These results favor a role of the filaments in dolipore plugging.

Taken together, I propose that SPCs of R. solani consist of a proteinaceous core, which is covered with ER-membranes. SPC18 is one of the proteins residing in the SPC core. It may be transported from the ER that is connected to the SPC. The C-terminal –FKEL sequence of SPC18 resembles the K/HDEL sequence of ER-residing proteins, and may

116 Summary and general discussion thus act as a temporary ER-retention signal. The dolipore-plug may also be formed by the SPC/ER since SPC18 was also detected in this material. The transport route of SPC18 from ER to SPC to cytoplasm at the orifice of the dolipore is different from the secretory pathway. Moreover, it resembles the retrotranslocation route from ER to cytoplasm of misfolded proteins (Molinari, 2007) or a still unknown mechanism for the export of small glycoproteins (Suzuki & Lennarz, 2000). It is rather surprising that SPC18 was not found in the genomes of other basidiomycetous fungi, like Coprinopsis cinereus and Phanerochaete chrysosporium that also have dolipore septa associated with perforate SPCs. This suggests that SPC18 only occurs in R. solani and possibly its close relatives. Recently, the SPC14 protein was isolated from a purified SPC fraction of S. commune (A.F. van Peer & H.A.B. Wösten, unpublished). The encoding gene is highly conserved in genomes of members of the Basidiomycota with perforate SPCs but is absent in those that lack these organelles. In contrast to SPC18, SPC14 is predicted to be a cytosolic protein. Future research aims to identify the role of SPC14 in S. commune and possibly R. solani.

Laser Microdissection as a Tool for Comparative Analysis in Fungal Cell Biology Laser microdissection has been proven a successful technique to isolate single cells or groups of cells from animal and plant tissue (Emmert-Buck et al., 1996; Day et al., 2005; Nelson et al., 2006). Until now, this technique had not been used in fungal cell biology. In Chapter 3, the hyphal region around the dolipore septa of R. solani was cut by laser microdissection and collected by laser pressure catapulting. Scanning electron microscopy confirmed the isolation of the septa that were visualized using wheat germ agglutinin (WGA) labeling. The WGA labeling may have detected the N-linked sugar residues of SPC18 located in the SPCs (Chapter 5).

The use of laser microdissection on fungal cells opens new ways to study subcellular fungal structures and the biochemical composition of hyphal cells. For example, septal regions in apical and subapical hyphal compartments could be compared. Similarly, septal composition of different parts of the fruiting body (e.g. stipe, hymenophore, gills) or from hyphae of different age can be analyzed. These studies, together with the biochemical enrichment method and the identification of SPC proteins could shed light on the dynamics of the composition of the dolipore-SPC complex.

117 Chapter 6 REFERENCES

Boekhout, T. (1991) Systematics of the genus Itersonilia Derx: a comparative phenetic study. Mycol. Res. 95, 135 – 146. Bracker, C.E. & Butler, E.E. (1963) The ultrastructure and development of septa in hyphae of Rhizoctonia solani. Mycologia 55, 35 – 58. Day, R.C., Grossniklaus, U. & Macknight, R.C. (2005) Be more specific! Laser-assisted microdissection of plant cells. Trends Plant Sci. 10, 397 – 406. Emmert-Buck, M.R., Bonner, R.F., Smith, P.D., Chuaqui, R.F., Zhuang, Z., Goldstein, S.R., Weiss, R.A. & Liotta, L.A. (1996) Laser capture microdissection. Science 274, 998 – 1001. Girbardt, M. (1958) Über die Substruktur von Polystictus versicolor. Arch. Mikrobiol. 28, 255 – 269. Hibbett, D.S. (2006) A phylogenetic overview of the Agaricomycotina. Mycologia 98, 917 – 925. Hibbett, D.S. & Thorn, R.G. (2001) Basidiomycota: Homobasidiomycetes. Pp 121 – 168. In The Mycota VII, Systematics and evolution, Part B. McLaughlin, D.J., McLaughlin, E.G. & Lemke, P.A. (eds.), Springer-Verlag, Berlin, Germany. Hibbett, D.S., Binder, M., Bischoff, J.F., Blackwell, M., Cannon, P.F., Eriksson, O.E., Huhndorf, S., James, T., Kirk, P.M., Lucking, R., et al. (2007) A higher-level phylogenetic classification of the Fungi. Mycol. Res. 111, 509 – 547. Keller, J. (1997) Atlas des Basidiomycètes vus aux microscopes electroniques. Union des Sociétés Suisses de Mycologie, Neuchâtel, Suisse. Langer, E. & Oberwinkler, F. (1993) Corticioid basidiomycetes. I. Morphology and ultrastructure. Windahlia 20, 1 – 28. Larsson, K.H., Larsson, E. & Kõljalg, U. (2004) High phylogenetic diversity among corticoid homobasidiomycetes. Mycol. Res. 108, 983 – 1002. Larsson, K.H., Parmasto, E., Fischer, M., Langer, E., Nakasone, K.K. & Redhead, S.A. (2006) Hymenochaetales: a molecular phylogeny for the hymenochaetoid clade. Mycologia 98, 926 – 936. Molinari, M. (2007) N-glycan structure dictates extension of protein folding or onset of disposal. Nat. Chem. Biol. 3, 313 – 320. Moncalvo, J.M., Nilsson, R.H., Koster, B., Dunham, S.M., Bernauer, T., Matheny, P.B., Porter, T.M., Margaritescu, S., Weiss, M., Garnica, S., et al. (2006) The cantharelloid clade: dealing with incongruent gene trees and phylogenetic reconstruction methods. Mycologia 98, 937 – 948. Moore, R.T. (1980) Taxonomic significance of septal ultrastructure in the genus Onnia Karsten (Polyporineae/Hymenochaetaceae). Bot. Notiser 133, 169 – 175. Müller, W.H., Montijn, R.C., Humbel, B.M., Van Aelst, A.C., Boon, E.J.M.C., Van der Krift, T.P. & Boekhout, T. (1998a) Structural differences between two types of basidiomycete septal pore caps. Microbiology 144, 1721 – 1730. Müller, W.H., Stalpers, J.A., Van Aelst, A.C., Van der Krift, T.P. & Boekhout, T. (1998b) Field emission gun-scanning electron microscopy of septal pore caps of selected species in the Rhizoctonia s.l. complex. Mycologia 90, 170 – 179. Müller, W.H., Koster, A.J., Humbel, B.M., Ziese, U., Verkleij, A.J., Van Aelst, A.C., Van der Krift, T.P., Montijn, R. & Boekhout, T. (2000a) Automated electron tomography of the septal pore cap in Rhizoctonia solani. J. Struct. Biol. 131, 10 – 18. Müller, W.H., Stalpers, J.A., Van Aelst, A.C., De Jong, M.D.M., Van der Krift, T.P. & Boekhout, T. (2000b) The taxonomic position of Asterodon, Asterostroma and Coltricia inferred from the septal pore cap ultrastructure. Mycol. Res. 104, 1485 – 1491. Nelson, T., Tausta, S.L., Gandotra, N. & Liu, T. (2006) Laser microdissection of plant tissue: what you see is what you get. Annu. Rev. Plant Biol. 57, 181 – 201. Orlovich, D.A. & Ashford, A.E. (1994) Structure and development of the dolipore septum in Pisolithus tinctorius. Protoplasma 178, 66 – 80. Setliff, E.C., MacDonald, W.L. & Patton, R.F. (1972) Fine structure of the septal pore apparatus in Polyporus tomentosus, Poria latemarginata, and Rhizoctonia solani. Can. J. Bot. 50, 2559 – 2563.

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Suzuki, T. & Lennarz, W.J. (2000) In yeast the export of small glycopeptides from the endoplasmic reticulum into the cytosol is not affected by the structure of their oligosaccharide chains. Glycobiology 10, 51 – 58. Wells, K. & Bandoni, R.J. (2001) Heterobasidiomycetes. Pp 85 – 120. In The Mycota VII, Systematics and evolution, Part B. McLaughlin, D.J., McLaughlin, E.G. & Lemke, P.A. (eds.), Springer-Verlag, Berlin, Germany.

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List of Publications

Curriculum vitae

Samenvatting

Samenvatting

Filamenteuze schimmels (ascomycete en basidiomycete schimmels) vormen uitgebreide netwerken (mycelia) die bestaan uit schimmeldraden (hyfen). Deze hyfen worden door dwarswanden (septa) opgedeeld in compartimenten. De septa hebben een centrale opening (porie) met een diameter van tussen de 70 en 500 nm, waardoor de celinhoud (cytoplasma) van het mycelium continu is. Het septum in de hogere basidiomycete schimmels (deze groep heet de Agaricomycotina en bevat o.a. trilzwammen en paddestoelvormende schimmels) is rond de porie opgezwollen en vormt de doliporie. De doliporie wordt vaak vergezeld van septumporie-kappen. De morfologie van deze kappen is goed bestudeerd door middel van elektronenmicroscopie, maar de samenstelling en de functie is nog steeds onbekend. Aan de hand van de morfologie studies zijn verschillende functies verondersteld. Zo zouden de kappen een rol spelen in het afsluiten van de doliporie met plugmateriaal om verlies van celinhoud te voorkomen bij beschadiging van de hyfen. De kappen zouden dit doen door plugmateriaal te produceren of door plugmateriaal op te slaan. Daarnaast is gesuggereerd dat de septumporie-kappen als een zeef werken, dat ze celorganellen dirigeren naar de doliporie, of dat ze voorkomen dat organellen de ingang van de doliporie blokkeren.

Het doel van mijn proefschrift is het identificeren van componenten van de septumporie-kappen. Er zijn twee methoden ontwikkeld waarmee de septumporie- kappen van de schimmel Rhizoctonia solani geïsoleerd en verrijkt kunnen worden. Deze bodemschimmel veroorzaakt ziekten bij verschillende belangrijke plantgewassen zoals suikerbieten en aardappelen. De grote septumporie-kappen van deze schimmel zijn goed bestudeerd. De kappen hebben een diameter tussen de 1600 en 2000 nm en zijn geperforeerd met gaten die een diameter hebben tussen de 600 en 800 nm. Met één van de isolatiemethoden is voor het eerst een eiwit uit de kappen geïdentificeerd. Dit septumporie-kapeiwit heet SPC18. Daarnaast is onderzocht of de verschillende typen septumporie-kappen de recent herziene classificatie van schimmels volgen. In het proefschrift wordt aangetoond dat twee ordergroepen in de Agaricomycotina (de Hymenochaetales en Cantharellales) twee verschillende typen septumporie-kappen bevatten in plaats van één. Binnen de ordergroepen zijn de septumporie-kappen dus niet altijd uniform in morfologie.

Septumporie-kappen als Kenmerk voor Classificatie In 1958 fotografeerde Girbardt voor de eerste keer de septumporie-kap in de schimmel Trametes versicolor met behulp van een transmissie-elektronenmicroscoop. Vanaf toen volgden meerdere studies naar het septum in schimmels, vanwege de bruikbaarheid bij de classificatie van schimmels. De basale groepen in de basidiomycete schimmels

123 Samenvatting hebben een eenvoudig type septum dat vergelijkbaar is met het septumtype dat voorkomt in de ascomycete schimmels. De hogere basidiomycete schimmels (de Agaricomycotina) hebben echter een septum met een doliporie die vergezeld kan zijn van een septumporie-kap. Dit zijn membraanstructuren die aan hun basis verbonden zijn met het endoplasmatisch reticulum. Daarom worden de kappen ook wel als subdomeinen van dit organel gezien. Hoofdzakelijk worden drie verschillende typen septumporie-kappen onderscheiden. Ten eerste het vesiculaire kaptype dat bestaat uit een groep zakvormige of buisvormige structuren die in een hemisfeer rond de doliporie zijn gegroepeerd. Ten tweede het niet-geperforeerde kaptype. Dit is een gesloten membraanstructuur die over de doliporie ligt. Het midden van het membraan kan dunner zijn of zelfs een minuscule porie bevatten. Ten derde, het geperforeerde kaptype dat als een koepel over de doliporie ligt. Dit kaptype kan veel kleine gaten bevatten zoals in Schizophyllum commune (het waaiertje) of een aantal grote gaten zoals in R. solani.

Tot nu toe volgde het kaptype de classificatie van schimmels. De order-groepen in de Agaricomycotina bevatten slechts één kaptype, ofwel vesciulair, ofwel niet-geperforeerd, ofwel geperforeerd. Onlangs echter is de classificatie van het koninkrijk schimmels herzien. Daarom is hier bestudeerd of het septumporie-kaptype nog overeenkomt met de classificatie gebaseerd op moleculaire fylogenetische studies (Hoofdstuk 2). De kapmorfologie van meer dan 350 soorten uit de literatuur werd gerelateerd aan deze nieuwe classificatie. DeAgaricomycotina wordt onderverdeeld in drie klasse-groepen (dit zijn de Tremellomyceten, de Dacrymyceten en de Agaricomyceten). In de meest basale groep, de Tremellomyceten, zijn de kappen afwezig of hebben ze het vesiculaire kaptype. Het niet- geperforeerde kaptype wordt gevonden in de Dacrymyceten, terwijl in de Agaricomyceten zowel het niet-geperforeerde als het geperforeerde kaptype worden gevonden.

In tegenstelling tot eerdere publicaties is gebleken dat de schimmel Cantharellus (de Cantharel) geperforeerde septumporie-kappen heeft (Hoofdstuk 2). De ordergroep Cantharellales in de klassegroep Agaricomyceten bevat dus soorten met geperforeerde kappen (Cantharellus, Sistotrema en de Ceratobasidiales) en niet-geperforeerde kappen (Tulasnella en Botryobasidium). Ook de ordergroep Hymenochaetales bevat beide kaptypen. Geperforeerde kappen zijn gevonden in de schimmels Hyphoderma praetermissum, Oxyporus latemarginatus en Rickenella fibula (het oranjegeel trechtertje) (Hoofdstuk 2). De rest van de Hymenochaetales hebben niet-geperforeerde kappen. Gedurende de evolutie van de septumporie-kap is de morfologie dus een aantal keren veranderd. Het lijkt erop dat de strengen endoplasmatisch reticulum gelegen over de doliporie in bijvoorbeeld Itersonilia perplexans (behoort tot de basale order-groep Cystofilobasidiales) de voorouder zijn van zowel de vesiculaire als de niet-geperforeerde kappen. Het geperforeerde kaptype is daarna een aantal keren ontstaan in de Agaricomyceten, maar is ook teruggekeerd naar het niet-geperforeerde kaptype zoals in de Cantharellales en

124 Samenvatting

Hymenochaetales is gebeurd.

Biochemische Analyse van Septumporie-kappen van Rhizoctonia solani In het onderzoek is een methode ontwikkeld om de kappen van R. solani te verrijken (Hoofdstuk 4). Met deze methode werden ook kappen van Ceratobasidium cornigerum en Thanatephorus cucumeris verrijkt. Deze drie schimmels behoren allen tot de order- groep Cantharellales. Ceratobasidium cornigerum en T. cucumeris hebben geperforeerde septumporie-kappen met een diameter van respectievelijk 700 tot 900 nm en 1400 tot 1500 nm. De verrijkingsprocedure bestaat uit een combinatie van drie technieken. Zo wordt een French press gebruikt, om door middel van hoge druk de schimmels kapot te maken, waardoor de celinhoud vrij komt. De vrijgekomen celinhoud wordt vervolgens gescheiden op dichtheid (isopycnische centrifugatie). De fractie die de septumporie- kappen bevat wordt behandeld met het detergens Triton X-100, om lipide membranen op te lossen. Uit de waarneming dat de kappen intact blijven na behandeling met Triton X-100 kan worden geconcludeerd dat het hoofdonderdeel van de septumporie-kappen bestaat uit eiwitten en/of membranen die resistent zijn voor dit type detergens.

SDS-PAGE (een methode om eiwitten te scheiden) laat een eiwitpatroon van de met kappen verrijkte fractie zien (Hoofdstuk 5). Een positieve labeling van de kappen met het lectine WGA (Hoofdstuk 3) suggereert de aanwezigheid van geglycosyleerde eiwitten (eiwitten met een suikerketen aan het oppervlak). In de kappen-verrijkte fractie was ook een eiwit verrijkt met een geschat moleculair gewicht van 18 kDa (Hoofdstuk 5). Aan de hand van de sequentie van het N-uiteinde van het eiwit werd het coderende gen SPC18 geïsoleerd. Het gen codeert voor een eiwit van 158 aminozuren en bevat een signaalpeptide om het eiwit te dirigeren naar het endoplasmatisch reticulum. Daarnaast heeft SPC18 een N-glycosyleringsmotief (aan dit motief wordt een suikerketen gekoppeld). Deglycosylering met behulp van enzymen bevestigde de N-glycosylering van het SPC18 eiwit. Lokalisatie-studies met antilichamen bevestigen de aanwezigheid van SPC18 in de septumporie-kappen. SPC18 werd ook gevonden in het porie-plugmateriaal. De geglycosyleerde eiwitten kunnen hun oorsprong hebben in het endoplasmatisch reticulum dat verbonden is met of gelegen is over de kappen. De resultaten in Hoofdstuk 5 laten zien dat de septumporie-kap verbonden is met, of zelfs deel is van het endoplasmatisch reticulum. De kappen kleurden fluorescent nadat de hyfen van R. solani werden behandeld met specifieke kleuringen om het endoplasmatisch reticulum aan te tonen (Hoofdstuk 5).

Het porie-plugmateriaal werd tegelijkertijd met de septumporie-kappen geïsoleerd. Het was stevig verankerd aan de binnenkant van de kappen door middel van een netwerk van filamenten (Hoofdstuk 4). Deze filamenten zijn ook in eerdere studies waargenomen en zijn net als de kappen resistent voor het detergens Triton X-100.

125 Samenvatting

Waarschijnlijk zijn er dus ook in de filamenten eiwitten aanwezig. De functie van deze filamenten is onduidelijk. Voorheen is gesuggereerd dat ze de septumporie-kappen in hun karakteristieke vorm houden. Echter, geïsoleerde kappen zonder additionele structuren (Hoofdstuk 4) of vrij liggende kappen in protoplasten (schimmelcellen waarbij de celwand met enzymen is verwijderd) behouden hun koepelvorm. Daarnaast zouden de filamenten organellen en cytoplasma-stromen naar de opening vande doliporie kunnen leiden. Als laatste wordt een rol in het pluggen van het doliporie kanaal gesuggereerd. Het plugmateriaal dat is opgeslagen in de septumporie-kappen kan via deze filamenten naar de doliporie worden getransporteerd. Dit zou op een pulserende manier gebeuren aangezien een gebandeerd patroon is geobserveerd tussen de SPC en de plug. In Hoofdstuk 5 is aangetoond dat het netwerk een grotere dichtheid heeft aan filamenten als de plug amorf is terwijl de dichtheid aan filamenten afneemt naarmate de plug meer gecondenseerd wordt. Dit kan het gevolg van een maturatie-proces zijn. Deze resultaten maken de rol van de filamenten in het pluggen van de doliporie de favoriete mogelijke functie.

Concluderend bestaan de septumporie-kappen van R. solani uit een kern van eiwitmateriaal die omgeven is door membranen afkomstig van het endoplasmatisch reticulum. SPC18 is één van de eiwitten die voorkomen in de kern van de kappen. Het kan getransporteerd worden vanuit het endoplasmatisch reticulum dat verbonden is met de kappen. Het C-uiteinde van het SPC18 eiwit heeft de aminozuur sequentie FKEL. Dit lijkt op het H/KDEL C-uiteinde van eiwitten die verblijven in het endoplasmatisch reticulum. Het FKEL uiteinde kan dus als een tijdelijk signaal dienen om SPC18 te laten verblijven in het endoplasmatisch reticulum totdat het naar de kap getransporteerd kan worden. Het plugmateriaal voor het doliporie-kanaal kan ook gevormd worden in septumporie-kappen of het endoplasmatisch reticulum aangezien SPC18 ook gevonden is in het plugmateriaal. De transportroute van SPC18, van endoplasmatisch reticulum naar septumporie-kap en verder via het cytoplasma naar de opening van het doliporie-kanaal, verschilt van de gewoonlijke route voor secretoire eiwitten (deze eiwitten worden uitgescheiden door de schimmel). De transportroute lijkt meer op de retrotranslocatieroute van ER naar cytoplasma van niet goed gevouwen eiwitten of op een nog onbekende exportroute van kleine geglycosyleerde eiwitten. Het is verrassend dat SPC18 of een hieraan homoloog eiwit niet gevonden is in het genoom van andere basidiomycete schimmels, zoals Coprinopsis cinereus (de inktzwam) en Phanerochaete chrysosporium, die ook septa bezitten met een doliporie vergezeld van septumporie- kappen. Het lijkt er dus op dat het SPC18 eiwit alleen voorkomt in R. solani en misschien in aanverwante schimmels. Onlangs is het eiwit SPC14 geïsoleerd uit een septumporie- kap verrijkte fractie van S. commune. Het coderende gen hiervoor is geconserveerd in de genomen (die tot nu toe bekend zijn) van basidiomycete schimmels met geperforeerde kappen, maar is afwezig in de genomen van schimmels zonder kappen. In tegenstelling

126 Samenvatting tot SPC18 is SPC14 waarschijnlijk een eiwit dat in het cytosol voorkomt. Verder onderzoek moet de rol van SPC14 in S. commune en mogelijk in R. solani bevestigen.

Lasermicrodissectie als een Hulpmiddel voor Vergelijkende Analyses in de Biologie van de Schimmelcel Microdissectie met behulp van een laser is een beproefde methode om enkele cellen of groepjes cellen van dierlijk en plantaardig weefsel te isoleren. Deze techniek was nog niet gebruikt met schimmelweefsel. In Hoofdstuk 3 wordt de hyferegio rond het septum met de doliporie van R. solani uitgesneden met behulp van lasermicrodissectie en daarna opgevangen door het omhoog te schieten met een laserpuls. Met een scanning- elektronenmicroscoop is de isolatie van de septa bevestigd. De septa waren gelabeld met het lectine WGA, zodat ze zichtbaar waren in de scanning-elektronenmicroscoop. De labeling met het lectine WGA heeft vermoedelijk de suikergroepen van het SPC18 eiwit in de septumporie-kappen hebben aangetoond (Hoofdstuk 5).

Het gebruik van lasermicrodissectie op schimmelcellen opent nieuwe wegen om celstructuren van schimmels en de biochemische samenstelling van hyfe- compartimenten te bestuderen. Zo kunnen aan de hyfetop gelegen compartimenten vergeleken worden met verder van de top gelegen compartimenten. Daarnaast kan de samenstelling van septa van verschillende delen van het vruchtlichaam (zoals de steel, de hoed of de plaatjes van de paddestoel) of van hyfen van verschillende leeftijd worden geanalyseerd. Deze studies kunnen samen met de biochemische verrijkingsmethoden en de identificatie van kapeiwitten opheldering geven over de samenstelling vande doliporie en de septumporie-kappen.

127

Nawoord

Nawoord

Toen ik aan deze klus begon, had ik nooit gedacht dat ik mijn proefschrift hier in Bangkok zou afronden. Dankzij het internet en e-mail hoeft dit tegenwoordig geen probleem meer te zijn. Een bijkomend voordeel is dat je niet meer de handgeschreven commentaren op je manuscripten hoeft te ontcijferen. Ondanks dat alleen mijn naam op de voorkant staat gedrukt heb ik dit onderzoek natuurlijk niet in mijn eentje gedaan. Ik wil daarom een aantal mensen bedanken voor hun inzet, hulp en enthousiasme, maar ook voor de gezelligheid tijdens mijn promotie-onderzoek.

Allereerst wil ik de leden van de “SPC-club” bedanken. Arie, bedankt voor je inzet achter de schermen. Het was een genoegen om op jouw afdeling te mogen werken. Daarnaast wil ik Han bedanken. Je hebt me op het juiste moment het zetje gegeven waarbij de afronding van mijn proefschrift in een hogere versnelling raakte. Arend, ik heb wel eens het idee dat we de enigen op de wereld zijn die aan septal pore caps werken. We waren altijd weer benieuwd naar wat de ander had ‘ontdekt’. Ik hoop dat je nog veel nieuwe ontdekkingen doet en ik kijk uit naar jouw proefschrift en publicaties! Beste Wally. Ik heb veel van je kunnen leren. Heel erg bedankt hiervoor. Je was meer dan alleen een begeleider. Ik zal de goede samenwerking, de leuke e-mails en de gezellige lunches niet vergeten! En bedankt voor de mooie scanningfoto’s die gebruikt zijn in de omslag. Teun, ik heb bewondering voor je enorme enthousiasme en je onuitputtelijke ideeënbron. Je kon genieten van de behaalde resultaten en dat werkte behoorlijk inspirerend. Helaas zijn de “groene mannetjes” niet in het proefschrift beland (misschien voor een volgende promovendus). Naast deze vaste leden van de “SPC-club” zijn er ook nog de tijdelijke leden geweest. HaciAli, Annemieke en Linda, ik wil jullie bedanken voor jullie inzet en voor jullie hulp als stagiairs.

Naast de “SPC-club” hebben mijn collega’s in de werkgroep op het CBS het leven op het lab aangenaam gemaakt. Ferry, naast een heel harde werker ben je ook nog een gezellig lab-maatje! Verder was je nooit te beroerd om te helpen, zelfs toen ik hier in Bangkok zat! Daarnaast ben je een echte globetrotter en ik hoop dat je ongebreidelde reislust niet is verdampt nu je zelf promovendus bent. Aangezien Azië nog te ontdekken valt voor jou nodig ik je uit voor een heerlijke Thaise curry! Marjan, we konden bij elkaar terecht als er problemen waren, maar ook voor een gezellig babbeltje! Eiko, geen Portugese tekst voor jou, want ik denk dat je het Nederlands al onder de knie hebt. Ik kon altijd bij je terecht met vragen over bio-informatics of als ik andere vragen had. Daarnaast zal ik je Caipirinha en overheerlijke sushi niet vergeten! Bart, ik heb bewondering voor je optimisme in het leven! Gijs, ik zal je verjaardag nooit vergeten. Carlos, je hebt me overtuigd om een Macbook aan te schaffen.

129 Nawoord

Onze werkgroep was erg dynamisch vanwege de vele gastonderzoekers die op bezoek kwamen. Twee hiervan wil ik in het bijzonder noemen. Cara Andreia, trabalhamos juntos sobre la Rhizoctonia. Durante el tiempo que estuviste en el laboratorio fuiste su luz. Te deseo todo lo mejor. Querida Tere, tu visita al CBS fue breve, sin embargo, nos hicimos amigos rapidamente. Espero que todo te vaya muy bien, y que nos volvamos a encontrar pronto, ya sea en Cuba o en Holanda (quizas en la defesa de tu Tesis?). Ook wil ik alle andere CBS collega’s niet vergeten te bedanken voor hun bijdrage in het verschaffen van schimmelcultures, het leveren van chemicaliën, het bereiden van buffers en media, en het lezen van manuscripten. Zij maakten het werk een stuk aangenamer! In het bijzonder wil ik Jan bedanken. Voordat ik aan dit promotie-onderzoek begon heeft hij me op zeer enthousiaste wijze geïntroduceerd in het onderzoek aan schimmels.

Delen van het onderzoek zijn uitgevoerd op de afdeling Cellular Architecture and Dynamics (Universiteit Utrecht). Bruno, Elly, Chris, Hans en Pim, bedankt voor jullie hulp bij de elektronenmicroscopie. Zonder jullie had ik niet de mooie platen kunnen maken die her en der in het proefschrift staan. Mireille en Marc, bedankt voor het “lenen” van jullie goudbolletjes. De collega’s van de afdeling Microbiologie (schimmelsectie) ook bedankt voor jullie input bij de werkbesprekingen. Het was altijd weer verfrissend om te horen wat jullie van mijn resultaten vonden.

Naast mijn collega’s op het lab en elders wil ik ook mijn vrienden niet vergeten te bedanken. Ze hebben niet direct bijgedragen aan dit onderzoek door een pipet te hanteren of een manuscript te lezen, maar zonder hen waren het maar saaie jaren geweest! Ik heb ze nooit lastig gevallen met schimmels en septal pore caps (tenzij ze erom vroegen en perse een artikel van mij wilde lezen!). Toch wil ik jullie bedanken voor de vele spelletjes, kroegbezoeken, etentjes, films, etc. die we gedaan hebben ter afleiding van de dagelijkse bezigheden. En Danny, bedankt voor het ontwerpen van de omslag.

Mijn directe omgeving wil ik ook niet vergeten. Andy, bedankt dat ik je favoriete broer mag zijn. Cara Yolanda, espero que, algún día, pueda explicaros en español a ti, a Elisa y a ... lo que está escrito en este libro. Xenia, jouw positieve kijk op het leven heeft me veel geleerd. Jouw liefde voor mij (en andersom) heeft me naar Bangkok doen verhuizen en van beide heb ik geen spijt. Als laatste (maar niet als onbelangrijkste) wil ik mijn ouders bedanken. Zonder hen was ik niet zover gekomen. Pa en ma, bedankt!

130 List of publications

List of Publications

Publications in Peer Reviewed Journals Van Driel, K.G.A., Van Peer, A.F., Wösten, H.A.B, Verkleij, A.J., Boekhout, T. & Müller, W.H. (2007) Enrichment of perforate septal pore caps from the basidiomycetous fungus Rhizoctonia solani by combined use of French press, isopycnic centrifugation, and Triton X-100. J. Microbiol. Meth. doi:10.1016/j.mimet.2007.09.013 Van Driel, K.G.A., Boekhout, T., Wösten, H.A.B., Verkleij, A.J. & Müller, W.H. (2007) Laser microdissection of fungal septa as visualised by scanning electron microscopy. Fungal Genet. Biol. 44, 466 – 473. Dijksterhuis, J., Van Driel, K.G.A., Sanders, M., Molenaar, D., Houbraken, J., Samson, R.A. & Kets, E.P.W. (2002) Trehalose degradation and glucose efflux precede cell ejection during germination of heat-resistant ascospores of Talaromyces macrosporus. Arch. Microbiol. 178, 1 – 7. Te Biesebeke, R., Ruijter, G., Rohardjo, Y., Hoogschagen, M., Heerikhuizen, M., Levin, A., Van Driel, K.G.A., Dijksterhuis, J., Yang, Z., Van de Hondel, C.A.M.M., Rinzema, A. & Punt, P.J. (2002) Solid-state fermentations with Aspergillus oryzae: physiological and molecular aspects. FEMS Yeast Res. 2, 245 – 248.

Abstracts Van Driel, K.G.A., Van Peer, A.F., Wösten, H.A.B., Verkleij, A.J., Müller, W.H. & Boekhout, T. (2006) Ultrastructure and biochemical analyses of septal pore caps in Basidiomycetes. 8th International Mycological Congress IMC8, August 20 – 25, Cairns, Australia. (invited speaker) Van Peer, A.F., Van Driel, K.G.A., Boekhout, T, Müller, W.H. & Wösten, H.A.B. (2006) Structure and function of the septal pore cap (SPC) in Basidiomycetes. 8th European Conference on Fungal Genetics (ECFG8), Vienna, April 8 – 11, Vienna, Austria Van Driel, K.G.A., Van Peer, A.F., Wösten, H.A.B., Verkleij, A.J., Müller, W.H. & Boekhout, T. (2005) Isolation of septal pore caps from basidiomycetous fungi. 23rd Fungal Genetics Conference, March 15 – 20, Asilomar, CA, USA. (poster) Van Driel, K.G.A., Van Peer, A.F., Wösten, H.A.B., Verkleij, A.J., Müller, W.H. & Boekhout, T. (2005) Isolation of septal pore caps from basidiomycetous fungi. Wetenschappelijke Voorjaarsvergadering, Nederlandse Vereniging voor Microbiologie (NVMM), April 11 – 13, Arnhem, The Netherlands. (selected speaker) Van Driel, K.G.A., Van Peer, A.F., Wösten, H.A.B., Verkleij, A.J., Müller, W.H. & Boekhout, T. (2004) Characterization of the septal pore cap structure in basidiomycetous fungi. Eurofung - “The genomic era”, November 17 – 19, Wageningen, The Netherlands. (selected speaker)

131 List of publications

Van Driel, K.G.A., Van Peer, A.F., Wösten, H.A.B., Verkleij, A.J., Müller, W.H. & T. Boekhout. (2004) Characterization of the septal pore cap structure in basidiomycetous Fungi. IB Conference on Biomembranes, October 22, Utrecht, The Netherlands. (poster) Van Driel, K.G.A., Müller, W.H., Van Peer, A.F., Verkleij, A.J., Wösten, H.A.B. & Boekhout, T. (2004) Enrichment of isolated septal pore caps of the plant pathogen Rhizoctonia solani. 13th European Microscopy Congress (EMC 2004), August 22 – 27, Antwerpen, Belgium. (poster) Van Driel, K.G.A., Müller, W.H., Van Peer, A.F., Verkleij, A.J., Wösten, H.A.B. & Boekhout, T. (2004) Enrichment of isolated septal pore caps of the plant pathogen Rhizoctonia solani. 7th European Conference on Fungal Genetics (ECFG-7), April 17 – 21, Copenhagen, Denmark. (poster) Van Driel, K.G.A., Müller, W.H., Van Peer, A.F., Verkleij, A.J., Wösten, H.A.B. & Boekhout, T. (2004) Enrichment of isolated septal pore caps of the plant pathogen Rhizoctonia solani. Wetenschappelijke voorjaarsvergadering, Nederlandse Vereniging voor Microbiologie (NVMM). April 16 – 17, Arnhem, The Netherlands. (poster) Van Driel, K.G.A., Müller, W.H., Yigittop, H., Verkleij, A.J., Wösten, H.A.B. & Boekhout, T. (2003) Isolation and characterization of the septal pore cap structure in the plant pathogen Rhizoctonia solani. Wetenschappelijke voorjaarsvergadering, Nederlandse Vereniging voor Microbiologie (NVMM). April 15 – 16, Arnhem, The Netherlands. (poster) Müller, W.H., Humbel, B.M., Koster, B., Van der Krift, T.P., Verkleij, A.J., Van Aelst, A.C., Montijn, R.C., Stalpers, J., Van Driel, K.G.A. & Boekhout, T. (2002) The septal pore cap as a key structure in cell to cell transport. IMC7 International Mycology Conference. August 11 – 17, Oslo, Norway.

132 Curriculum vitae

Curriculum vitae

Kenneth van Driel was born on October 31, 1975 in Terneuzen, The Netherlands. He followed his high school education at the Zeldenrustcollege in Terneuzen and received his VWO diploma in 1994. The same year, he started to study Chemistry at the Catholic University of Nijmegen (KUN, since 2004 Radboud University Nijmegen) in Nijmegen. In 1995, he changes the subject for Biology. In his graduation year, scientific research was done at the departments of Molecular Biology (a molecular study towards the development of a vaccine against malaria) and Microbiology (aspects of the activation and germination of the ascospores of the heat-resistant fungus Talaromyces macrosporus). He graduated in August 2000 and obtained his MSc degree. In January 2001 he started working as a junior scientist at the Agrotechnological Research Institute (ATO), partner of the Wageningen Centre for Food Sciences (WCFS) in Wageningen and did research on physiological aspects of Aspergillus oryzae in solid-state fermentation. From May 2002 he worked as an ‘onderzoeker-in-opleiding’ (OIO) at the Centraalbureau voor Schimmelcultures (CBS), an institute of the Royal Netherlands Academy of Arts and Sciences (KNAW) in Utrecht. The research during this period was done under supervision of Dr. T. Boekhout and Dr. W.H. Müller and is described in this Thesis. In June 2007, he moved to Bangkok, Thailand.

133

The research presented in this Thesis was conducted at the Centraalbureau voor Schimmelcultures (CBS), an Institute of the Royal Netherlands Academy of Arts and Sciences (KNAW), Uppsalalaan 8, 3584 CT, Utrecht, The Netherlands.

Electron microscopy was performed at Cellular Architecture and Dynamics, Utrecht University, The Netherlands.

Financial support for the research was provided by the Odo van Vloten Stichting.