INTRODUCTION

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Coprinus cinereus (Schaeff. ex Fr.) S. F. Gray is one of two model organisms commonly used to study developmental processes in the homobasidiomycetous fungi. Unlike the other model organism, the bracket fungus Schizophyllum commune, C. cinereus is a typical mushroom (246, 335, 338, 517). Although it is of limited edible value (10), studies of C. cinereus may be used to understand the development of numerous edible basidiomycetes which fail to grow and/or produce fruiting bodies in the laboratory and are not readily accessible to genetic approaches (84, 255a). C. cinereus was introduced early as an object for studies of development (26, 39, 46, 50, 53, 58), mainly because of its relatively short life cycle, which can be completed in the laboratory within 2 weeks (343). The natural substrate of the species is horse dung (50, 58), but it also grows and fruits well on various artificial media (73, 292, 406, 508a). Over the decades, the fungus has been isolated under various names including cinereus, delicatulus, fimentarius, lagopus, macrorhizus f. microsporus, and radiatus. The names lagopus and radiatus were also attributed to other Coprinus species, making it difficult to analyze the older literature and to be certain which fungus was dealt with. There is thus no guarantee that all articles listed in this paper were indeed on C. cinereus. The Coprinus lagopus of Borriss and Madelin, for example, has sometimes been interpreted to be Coprinus radiatus (344), but because of its fundamental character, the work is included here. At present, all isolates interfertile with C. lagopus sensu Buller are classified under the species name cinereus (222, 226, 344, 382), but another renaming is expected in the future. Traditionally, members of the genus Coprinus (commonly known as ink caps) were defined as saprophytic mushrooms whose gills and frequently the entire cap autodigest at maturity, giving rise to an inky black fluid that drips to the ground (10). Recent phylogenetic studies based on large-subunit rDNA sequences, however, indicate that the traditional genus Coprinus is polyphyletic. Unfortunately, the type species, Coprinus comatus, nests within the lepiotoid fungi while most other Coprinus species including C. cinereus are placed close to Psathyrella (177, 178, 192).

C. cinereus is a heterothallic basidiomycete. Two main types of mycelia are distinguished in its life cycle, the infertile monokaryon and the fertile dikaryon (71, 176). C. cinereus has a wide-ranging developmental potential in both the monokaryotic and the dikaryotic mycelial stages (Fig. 1). This potential ranges from the fruiting body (carpophore, basidiocarp, basidiome), with the meiotic basidiospores localized within the gills (314, 338) (see below), to mitotic aerial spores (oidia) and mitotic submerged spores (chlamydospores) to the multicellular structures of sclerotia, mycelial cords, pseudorhizas and rockeries (56, 60, 171, 261, 371, 385) (see below). There is considerable flexibility in the shapes formed and in when and where these different organs develop (89, 91, 171, 463; U. Kües, J. D. Granado, Y. Liu, E. Polak, and M. Aebi, unpublished data; E. Polak, M. Aebi, and U. Kües, submitted for publication). Fungal differentiation is clearly not a rigid process but is tolerant to imprecision in fungal morphogenesis, probably enabling the fungus to react to adverse conditions. Fungal morphogenesis is organized in an array of developmental pathways, or subroutines (335), which are genetically and physiologically distinct. These subroutines run in parallel or in sequence, and the order followed determines the final outcome (338-340). Numerous molecular, genetic and cytological techniques are now available (9, 41, 71, 101, 125, 126, 145, 267a, 283, 343, 366, 386, 388, 390, 391, 430, 446, 508b, 537, 539, 540, 543) to unravel these routes and we are beginning to gain insight into the fascinating regulatory mechanisms of development in C. cinereus.

View larger version (22K): [in this window] [in a new window]   FIG. 1.   Life cycle of C. cinereus.

GROWTH AND STRUCTURE OF THE MYCELIUM

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Monokaryons and Dikaryons

Monokaryons arise from germination of the haploid binucleate basidiospores or, alternatively, from germination of the uninucleate haploid oidia. During germination of the basidiospores, a single germ tube displaces the spore pore cap. Similarly, a single germ tube is formed by the oidium. Nuclei within the germ tubes divide, and septa may be formed between the daughter nuclei; alternatively, the germ tubes may stay aseptate through several cycles of nuclear division. Multinucleate side branches emerge, while septum formation occurs in the older portions of the germtubes to yield cell segments with one or only a few nuclei. The tip cells of the main hyphae and those of side branches remain multinucleate during further growth (26, 169). The literature contains reports of the older monokaryotic mycelium having only one nucleus per hyphal cell (452). However, nuclear staining of aerial mycelium of various strains revealed that in most cases up to half of all hyphal cell segments contain not just one but two and sometimes even three nuclei (385; E. Polak, personal communication). Thus, the primary mycelium of C. cinereus does not fulfill the strictest definition of a monokaryon, i.e., a specific homokaryotic mycelium with just one nucleus per cell (4, 121). Nevertheless, we prefer to continue calling the primary mycelium a monokaryon, since this term is traditionally used in C. cinereus (70, 75, 266, 343, 391), and thus follow the less stringent definition given by Hawksworth et al. (164) of a mycelium as having genetically identical haploid nuclei. For the geneticists, the term "monokaryon" nicely targets the most important difference from the dikaryotic mycelium, which typically contains two genetically distinct haploid nuclei in its cells (26, 175a, 187).

Cultures of monokaryons differ greatly in growth rate and also in their morphological appearance, primarily due to variations in the amount and structure of the aerial mycelium produced (Polak et al., submitted). The distance between branches varies between strains but branches usually form in a relatively wide angle of about 70 to 75°. Hyphae of monokaryons are usually thin, with a diameter of about 3 µm (58, 226; Polak et al., submitted). They are generally characterized by simple septa (26, 58, 475) with a dolipore, as is typical for the basidiomycetous fungi (132). The dolipore is barrel shaped due to the swollen edge of the cross-wall around the pore. Parenthosomes, dome-shaped double membranes thought to be a modified part of the endoplasmic reticulum (ER), cover the dolipore swellings and the pore channel to form the septal pore cap. Perforations within the membranes allow cytoplasmic continuity through the parenthosome barrier and between adjacent cellular compartments, giving the hyphae a coenocytic character. Passage of smaller organelles such as mitochondria is possible; however, migration of larger organelles such as nuclei is blocked (299, 348, 437).

Dikaryons arise upon fusion of monokaryons of different mating types (Fig. 2), either by hypha-hypha fusion or by fusion of hyphae with germinating or resting oidia (26, 75, 223). Cellular fusion (anastomosis) may occur between two hyphal tips (tip-to-tip fusion), between a hyphal tip and a lateral hyphal wall (tip-to-side fusion), between a hyphal tip and a lateral swelling of a hypha (tip-to-peg fusion), or between lateral swellings of two neighboring hyphae (peg-to-peg fusion) (59, 410, 440). Upon fusion, nuclei enter the mycelium of the opposite mating type and, with dissolution of the hyphal septa (69, 75, 132), migrate through the hyphae until they reach a hyphal tip cell (26, 58) (Fig. 2). In C. cinereus, the speed of nuclear migration is between 1 and 3 mm h¹, which is up to 20 times faster than that of the hyphal tip growth (58, 236, 460; for a compilation of data for various basidiomycetes, see reference 64). Buller (58) found that migration takes place mainly through the outer part of the colony, and he assumed that migration involved the regular division of the invading nucleus. In Schizophyllum commune, invading nuclei apparently do not densely colonize the established host mycelium and there are areas through which the migrant nuclei obviously pass but in which they cannot be detected (407, 441). Niederpruem (361) monitored nuclear movements over short distances in living hyphae in compatible matings of S. commune and did not observe any mitosis. The observations in S. commune suggest that there may be no or little nuclear division during nuclear migration in basidiomycetes (361, 407, 441). Other authors, however, dispute this conclusion. At least directly after hyphal fusion, at the initial stage of nuclear exchange and migration, there are sometimes nonsynchronous (265, 409) and often also synchronous mitotic divisions of the two nuclei while they are together in the fusion compartment (410, 471). In addition, synchronous nuclear divisions in "migration" hyphae have been visualized by indirect immunofluorescence microscopy (410). The age of the hyphae strongly influences the rate of nuclear migration (425) and might also have an effect on the mitotic activity of the nuclei in the fusing hyphae (M. Raudaskoski, personal communication). Whatever the situation in C. cinereus, once a migrating nucleus reaches a hyphal tip cell, the two types of haploid nuclei pair at average distances of 15 to 20 µm (175a, 211) and the paired nuclei divide synchronously (26, 58, 459) (Fig. 2). Simultaneously, specialized clamp cells (or hook cells) are formed at the position where a new septum will appear (26, 59, 459) (Fig. 2).

View larger version (31K): [in this window] [in a new window]   FIG. 2.   Dikaryon formation following peg-to-peg fusion of hyphae of two compatible monokaryons. This model has been composed from observations by Buller (59), Iwasa et al. (187), and Raudaskoski (410). For a discussion of A and B mating-type control in the process of hyphal fusion and dikaryon formation, see references 79, 172, 410, and the text.

During the mitotic prophase, the nucleus localized closer to the tip will enter the developing clamp cell, and in the anaphase, it divides with a short spindle. The second nucleus stays in the hyphal cell beneath the clamp cell and divides with a longer spindle. Due to the different lengths of the spindles, the foremost of the dividing nuclei in the hyphal cell will pass the member of the other pair of dividing nuclei that is leaving the clamp cell. Septa are generated between the dividing nuclei, enclosing one nucleus into the clamp cell, one nucleus of the other type in the newly formed subapical cell, and a nucleus of each type in the newly arising hyphal tip cell. After septum formation, the backward-looped clamp cell fuses with a peg induced at the subapical cell. This fusion releases the enclosed nucleus from the clamp cell (59, 187, 464, 465) (Fig. 2). As a result of this coordinated mechanism of clamp cell formation and synchronized nuclear division, the order of nuclei reverses between the apical and subapical cell with each cellular division (187). The mechanism ensures that every cellular segment within a dikaryon contains one of each of the two distinct nuclei (26, 59, 187) (Fig. 2). However, occasionally, more than two nuclei are found within cells of a C. cinereus dikaryon (175a), and other species are known where the homokaryon also possesses clamp cells (225) or where a dikaryon is perfectly formed in the absence of any clamp cells (224, 459). Consistent with earlier data obtained with S. commune (427), recent analysis of mutant strains indicates that the actin cytoskeleton and microtubules with ₁- and ₁-tubulins play a role both in nuclear migration for dikaryosis and in subsequent nuclear positioning and movements within the established dikaryon (199, 209, 211, 410, 464, 465).

As in the monokaryon, the septa of the dikaryon have a dolipore, irrespectively of being formed between two hyphal segments or between a hyphal cell and a clamp cell (132). Biochemically, the septa of monokaryons and dikaryons are probably somewhat distinct, as suggested by in vitro enzymic dissolution of S. commune septa (69, 75, 519). The more resistant nature of dikaryotic septa had been suggested to be a cause of blockage of nuclear migration through septa within the dikaryon (69, 80). The literature on the cell wall composition of the monokaryotic and the dikaryotic mycelium in C. cinereus is unfortunately poor (42, 43, 195, 296, 298, 432). The innovative work of Marchant (298) and Bottom and Siehr (42, 43) indicates that major differences in chitin content and sugar compositions exist between the cell walls of the two different mycelial stages. In addition, Ásgeirsdóttir et al. (15) recently isolated monokaryon-specific hydrophobins. These are small, hydrophobic, cysteine-rich secreted proteins that assemble at the hyphal wall when hyphae emerge from a submerged culture into the air (116, 231, 521, 522). Visually, there are further differences between monokaryons and dikaryons. Dikaryons of C. cinereus tend to grow faster, with a denser, more protuberant and conspicious aerial mycelium, the hyphae are about 7 µm in diameter, and branches arise with a relatively acute angle of 10 to 45° (58, 266). Due to the vigorous, rapidly propagating character of dikaryotic mycelium, outgrowth of a dikaryon from a cross of compatible monokaryons is easily detected. In the ideal case, there are four positions where outgrowth will be observed. Two dikaryotic sectors will arise at the junction of the growing monokaryotic colonies, where their hyphae met and fuse. Two other dikaryotic mycelia are generated at the outward sides of the two monokaryons after migration of nuclei through the opposite mycelium (58, 158). Since only the nuclei, not the mitochondria, migrate, these latter dikaryons are distinguished from each other by their mitochondrial content. In contrast, the dikaryotic sectors arising from the junction zones of the monokaryotic colonies are expected to be mitochondrial mosaics due to mixed hyphal populations with distinct mitochondrial DNA populations. Usually, such mixed dikaryotic mycelia rapidly segregate for one mitochondrial type. Although mitochondrial inheritance in C. cinereus is basically uniparental, recombined mitochondrial DNA is occasionally found in places of hyphal anastomosis (23, 75, 117, 118, 306; for a recent review of mitochondrial inheritance and recombination, see reference 420).

Following dikaryon formation, there is no habitual restriction to hyphal fusion with other monokaryotic or dikaryotic strains, but, in contrast to monokaryons, dikaryons do not accept invading nuclei (237, 456, 457, 460). However, they are capable of contributing fertilizing nuclei to a haploid monokaryon, resulting in a new dikaryon (57, 58, 69); this reaction has been termed the Buller phenomenon (400). "Legitimate di-mon matings" (for "dikaryon-monokaryon matings") are those where both kinds of nuclei in the dikaryon are of compatible mating type to the nuclei of the monokaryon. In these cases, heterokaryotization of the monokaryon occurs readily. Either one or both types of the nuclei from the dikaryon migrate into the mycelium of the recipient monokaryotic partner, giving rise to one or two new dikaryotic mycelia. Occasionally, both nuclei from the dikaryon replace the original resident nucleus of the monokaryon (237, 238, 304, 401, 457). In di-mon matings, as in matings of compatible monokaryons, mitochondria are not exchanged but mitochondrial mosaics may arise by a mixed hyphal population (304, 306). In hemicompatible matings, where one nuclear type of the dikaryon is incompatible with the nuclei of the monokaryotic partner, only the compatible nuclei might enter the monokaryon or, alternatively, both kinds of nuclei invade the monokaryon and replace the resident nucleus (304). In illegitimate matings, where all nuclei are incompatible due to differences at either one of the two mating-type loci (see below), heterokaryotization of the monokaryon may occur but only after a delay during which compatible nuclei are generated by somatic recombination (58, 401). Exchange of nuclei between different dikaryons is also possible despite the blockage of nuclear migration, but only in areas where hyphae of both strains intermingle. New dikaryons are thought to form either by outgrowth of fused cells after a fresh sorting of the original pairs of nuclei or by dikaryotization of monokaryotic hyphae that occasionally are observed in the growing front of a dikaryon. Somatic recombination between nuclei has never been observed in dikaryon-dikaryon combinations (456), probably due to the low frequency with which nuclei of the different strains are found together within the same cell.

One of the most remarkable aspects of the life cycle of most homobasidiomycetes is the temporal and spatial separation of cellular fusion from nuclear fusion. Karyogamy normally occurs in the fruiting bodies within the specialized cells, the basidia. It is directly followed by meiosis, thereby restricting the diplophase to a single nuclear generation (283, 469) (see below). The extended dikaryotic growth phase, together with the unusual mating system developed in Coprinus and other basidiomycetes (see below), has special consequences for the understanding of the individual fungus. The free fusion between any mono- and dikaryotic mycelia and the variety of donating, accepting, and expelling of nuclei allows the formation of genetic mosaic colonies composed of genetically different sectors. However, somatic incompatibilities (mycelial antagonisms) lead to processes that rapidly separate newly formed and original dikaryons, despite their history of hyphal fusions and a common nucleus, ensuring that different dikaryons can represent discrete individuals. The vegetative self-nonself recognition of dikaryons is associated with pigmented zones, sparse hyphae or, less commonly, tight hyphal knitting, and an increased production of sclerotia at the interface of colonies. In most cases, the nuclear genomes but occasionally also the mitochondria seem to determine somatic incompatibility. Usually, mitochondrial differences are tolerated and a mitochondrial mosaic behaves as one individual (304; for a further discussion of somatic incompatibility and the problem of fungal individualism, see references 122, 227, 412, and 511).

Dikaryons are only one type of heterokaryon found in C. cinereus. Dikaryon formation is governed by the two independent mating type loci, A and B. For a mating to be successful, monokaryons must be different at both loci (for recent reviews, see references 79, 172, and 245) (see below). However, monokaryons can fuse independently of whether different mating-type specificities are present (438, 440). If monokaryons differ in the B but not in the A mating-type loci, a so-called common A heterokaryon is formed that is not distinct in hyphal morphology from the monokaryon, although such strains usually grow less vigorously and more slowly than their component monokaryons. If strains differ within the A but not the B loci, a common B heterokaryon results which grows as vigorously as a dikaryon and which is characterized by formation of unfused clamp cells at the hyphal septa. Common B heterokaryons may form fruiting bodies but not as readily as does the dikaryon. In contrast, fruiting body formation in common A heterokaryons is the exception (399, 460, 459; for further review, see references 75, 76, and 407). Usually, common A heterokaryons are not stable and can be recognized and kept as such only under strong selection pressure (forced heterokaryon), e.g., by selection for complementation of auxotrophies. Similarly, unstable clampless common AB heterokaryons can be isolated by a forced mating of monokaryotic strains of the same mating type but with different auxotrophies. By contrast, common B heterokaryons are normally quite stable. Like dikaryons, common B heterokaryons have been reported to act as donors but not as acceptors of nuclei upon fusion with suitable monokaryons. In contrast, common A and common AB heterokaryons will usually act both as donors and as acceptors of nuclei, probably also because they frequently produce hyphae with only one type of nucleus (455, 459, 460).

Normally, it is the dikaryon that produces the fruiting bodies (see below). However, under specific conditions, monokaryons may also form fruiting bodies, e.g., under total nutritional depletion (159, 503, 508a). Certain mutations outside of the mating-type loci (su-A [103]), fis^c [324, 373, 480, 491], pcc1, and CopD5 [356, 357]) also lead to formation of fruiting bodies, karyogamy, and meiosis on monokaryons. These mutations are all accompanied by the formation of unfused clamp cells at hyphal septa (356, 357), and the su-A mutation is known to block nuclear acceptance (103). A specific class of mutants comprises homokaryons with defects in the mating-type loci that overcome the self-incompatibility of monokaryotic strains. Such strains, commonly called Amut Bmut homokaryons, are fully self-compatible and will initiate fruiting-body development under certain environmental conditions (452; see below). Depending on the general genetic constitution of the strain, fruiting-body initials and primordia will mature and karyogamy, meiosis, and the production of four basidiospores will occur within the gills of the fruiting body (165, 452) (Fig. 3). Amut Bmut homokaryons resemble dikaryons in their vigorous growth and hyphal morphology. They have fused clamp cells at the hyphal septa, but their distribution is usually not as regular as in dikaryons, and septa with unfused or no clamps are frequently been detected (452; U. Kües, unpublished data). The nuclei in aerial cells of Amut Bmut homokaryons are distributed in similar patterns to those of related monokaryons (385), but in submerged mycelium a fairly strict binucleate distribution has been reported (452). Although fully self-compatible, Amut Bmut homokaryons might mate with any strain in the junction zones of a cross, irrespective of which wild-type (self or nonself) or mutant mating types are present in the mating partner. However, heterokaryons formed with strains of related mating types appear to be short-lived and unstable. Nuclear acceptance of Amut Bmut homokaryons might be restricted but donation by nuclei into monokaryons is possible (269; Kües, unpublished).

View larger version (154K): [in this window] [in a new window]   FIG. 3.   Basidia with mature basidiospores of homokaryon AmutBmut (A43mut B43mut). Courtesy of J. D. Granado.

To generate Amut Bmut homokaryons, independent mutations were selected in the A and B mating-type loci and combined into the same nucleus by crosses (165, 452). Amut homokaryons with only a mutated A locus mimic common B heterokaryons and are characterized by mycelia with unfused ("false") clamp cells at some but usually not all septa. Amut homokaryons may have one, two or occasionally even three nuclei per cell (103, 165, 452) at frequencies comparable to what we have found in monokaryons (384, 385). Amut homokaryons are hemicompatible and form stable dikaryons with any monokaryotic strain as long it has a different B specificity (103, 165, 452). In mating, Amut homokaryons may act as both donors and acceptors of nuclei (Kües, unpublished). Bmut homokaryons are also hemicompatible. They mate bilaterally and accept and donate nuclei in crosses with monokaryons as long as these differ in the A mating-type locus. Bmut homokaryons have simple septa like normal monokaryons (165, 452). In contrast to the analogous Bmut homokaryons of S. commune (243, 361), the dolipore septa are not disrupted in the C. cinereus Bmut homokaryons and the nuclear distribution within the cells is even over the entire hyphal length as in monokaryons (165, 452).

Although the diplophase is normally restricted to the basidia in the life cycle of C. cinereus, nuclear fusion occasionally occurs within the vegetative mycelium of heterokaryons. Diploid mycelia can be selected from common A and common AB heterokaryons by plating and germinating uninucleate oidia at frequencies ranging from approximately 10³ to 10⁴ (68, 187, 355). There are no reports in the literature on the rescue of diploids from common B heterokaryons or from dikaryons, but indirect evidence for the infrequent existence of diploid nuclei comes from the observations of somatic recombinations (98, 237, 401, 457, 458).

Diploid oidia of common A and of common AB heterokaryons have increased cell size, with a single enlarged nucleus that is twice the size of a haploid nucleus. Germinated hyphae are significantly broader than those of the monokaryotic parents (68, 187, 355, 364). Diploid common A and common AB mycelia readily mate with monokaryons and with other diploid common A strains, provided that these strains contain other A mating-type genes. However, common A diploids serve only as nuclear donors in matings, whereas common AB diploids both donate and accept nuclei (77, 80, 98, 364). In agreement with their different behavior in nuclear acceptance, septal dissolution was not detected in common A diploids, in contrast to common A heterokaryons, indicating that the cytological difference between the two nuclear conditions has an impact on the mating behavior of the mycelium (364). Possibly due to uneven nuclear distribution within colonies of common A heterokaryons (78, 455), common A diploids and common A heterokaryons also differ in complementation efficiency of metabolic functions whereas common A diploids and ordinary dikaryons usually behave similarly (78, 105, 367, 472). However, bringing genes together in a single nucleus in common A diploids or separating them into two haploid nuclei within dikaryotic cells can also cause changes in expression, as observed with certain resistance genes (267, 365).

Diploid mycelia are often fairly stableonly 0.5% of oidia produced by such mycelia were found to be haploid (68, 78)but treatment with the fungicide griseofulvin can induce haploidization by a gradual loss of chromosomes (364). Most interestingly, diploid-haploid and diploid-diploid pairings produce dikaryons in which the diploid nuclear components are unstable in the vegetative growth phase, indicating that the dikaryotic situation is preferred over the diploid. Loss of chromosomes in such dikaryons is also progressive as aneuploid and haploid nuclei are recovered from these colonies (68, 78, 397). Finally, it is interesting that common A diploids are not fully compatible even if they are distinct in their A mating-type loci. In dikaryons formed by such common A diploids, the produced clamp cells at first fail to fuse, but progressive haploidization, eliminating one of the two B chromosomes from the diploid nuclei, eventually results in a compatible situation and clamp cell fusion (78).

ASEXUAL SPORE FORMATION

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Oidiophores and Oidia

Monokaryons of C. cinereus have for a long time been known to constitutively produce abundant uninucleate asexual spores on specialized aerial structures (oidiophores). These spores are rod shaped, and because of this shape, they are called oidia (26, 47). Until recently, dikaryons were believed not to form these mitotic spores (48, 70, 234), but their production has been observed after light induction (232, 452) (Fig. 4). Likewise, formation of oidia on Amut and Amut Bmut homokaryons is under light control, whereas Bmut homokaryons and common A heterokaryons behave like monokaryons and constitutively produce numerous oidia (232, 385, 452). Common B heterokaryons are also reported to produce abundant oidia in the light (459); due to the presence of different A mating-type genes within these strains, repression of oidiation in the dark is expected (232, 261) (see below) but was not tested. Common A and common AB diploids also give rise to oidia, and spore production is presumably constitutive (68, 78, 364). Extensive use of oidia has been made in single-cell purification of cultures, in generation of uninucleate protoplasts for transformation, and in providing isolated uninucleate cells in classical and modern mutagenesis techniques (30, 31, 68, 145, 366, 386, 388). Regardless of their importance in genetic studies, attention was only recently drawn to the cytological process of formation of oidia (385; Polak et al., submitted). Under certain genetic conditions, oidiation is light induced, and light acts at the level of oidiophore formation, which has enabled the whole process of sporulation to be monitored over time in microslide cultures (385).

View larger version (160K): [in this window] [in a new window]   FIG. 4.   Young oidiophore in the aerial mycelium of dikaryon FA2222 (A5 B6) × 306 (A43 B43). Note the clamp cells at the septa of the hyphae and the absence of clamp cells on the oidiophore. Courtesy of E. Polak.

Oidiophores are formed predominantly in the aerial mycelium (47, 385), although in most strains small numbers of oidiophores can regularly be detected in the submerged mycelium of agar cultures (Polak et al., submitted). Not every cell of the aerial mycelium becomes an oidiophore foot cell, i.e., the hyphal cell that will give rise to the oidiophore. Unlike the oidiophore foot cells, undifferentiated aerial cells vary greatly in length, indicating that the former are physiologically predetermined (Polak et al., submitted). Development starts with the protrusion of a young oidiophore from its foot cell (385) (Fig. 5). Subsequently, a nucleus in the foot cell divides, one daughter nucleus migrates into the bulging young oidiophore, and a septum is formed which separates the first oidiophore stemcell from its foot cell. The stem cell elongates and may divide once or twice, and side stems may branch from the cells of the main stalk. From the tip of stem cells of both main and side stalks, short, oblong oidial hyphae bud off one after the other. Successive nuclear divisions within the stem cells ensure that each oidial hypha is provided with a nucleus before it is separated from the stem cell by septation. Within an oidial hypha, a further nuclear division occurs followed by formation of a septum that separates the oidial hypha into two equally sized cells. Eventually, these cells are released by schizolysis of the septa between the oidial hypha and between the oidial hypha and its stem cell. A few hundred free spores are successively collected in a sticky liquid droplet secreted at the tips of the oidiophores (385) (Fig. 5). Since their mode of generation involves budding and septum schizolysis, oidia are classified as arthroconidia (228, 385). Mature haploid spores average about 2 by 4 to 6 µm in size, and mature diploid oidia measure about 3 by 7 µm (68, 364, 385; Polak et al., submitted). Oidia are enclosed by mucilage and possess a double-layered primary cell wall with hair-like structures, except at former sites of cell attachments, where a single-layer secondary cell wall is present (168, 169, 385). These secondary walls are later involved in the germination of the spores (168). Oidia are distinguished as "wet" spores due to their strong hydrophilic character (15, 222). Consistent with this, proteins with the characteristic properties of hydrophobins are not detected on oidia (15), unlike the "dry" hydrophobic spores of other fungal organisms, which are coated with hydrophobins (116, 231, 518, 521, 522). Oidia of C. cinereus are not wind-borne spores. Instead, they stick to flies and other insects attracted by the horse dung substrate and are thereby distributed to new substrates (47). Independently of mating types, oidia attract hyphae of coloniesa process called oidial homingand act as spermatia in matings with compatible C. cinereus cells (223) (see above). Most interestingly, they also attract and fuse to hyphae of other related coprophilous fungi. Upon fusion, somatic incompatibility reactions are initiated, leading to the death of the foreign hypha. Oidia thereby act as a killing agent of opponents competing for the same limited substrate (223).

View larger version (14K): [in this window] [in a new window]   FIG. 5.   Model of the development of oidiophores and oidia in C. cinereus.

Analysis of various C. cinereus strains showed that oidiophore formation is not a fixed cytological process. Oidiophores may have simple stems or may be branched, and the stem cells may not elongate or may totally be absent (26; Polak et al., submitted), indicating that the different steps in oidiophore development, as defined in Fig. 5, are independent of each other and that some may be omitted without terminating development. To denote the main structural differences, four main types of oidiophores have been defined. Type 1 oidiophores have unbranched elongated stems and produce oidial hyphae at only their tips. Type 2 oidiophores form oidial hyphae at the tip of a main stem and also at the tip of side branches; oidial hyphae may also bud directly from the uppermost stem cell (type 2A) and from stem cells beneath (type 2B). In contrast to these more advanced structures, type 3 oidiophores do not elongate their stem cells. Type 4 oidiophores are those lacking a stem cell, where oidial hyphae bud off either singly (type 4A) or in bundles (type 4B) from cells of the aerial hyphae. These various types of oidiophores occur within the same aerial mycelium of all strains so far analyzed, although at strain-specific frequencies (Polak et al., submitted).

Chlamydospores are large, thick-walled mitospores of variable forms and with condensed cytoplasm. They are found in brown patches present on the mycelial mating of older cultures of dikaryons (6, 71, 121, 266) and, occasionally, on certain monokaryons (26, 261) (Fig. 6). The generation mode of this type of spore in C. cinereus is not well documented. In general, chlamydospores may arise endogenously in cells of the vegetative hyphae following compression of the cytoplasm (chlamydospores in the strictest sense) or by transfer of the compressed cytoplasm from a hyphal cell into a bud (blastocyst) (95). Diagrams found in some publications (6, 121, 266), combined by occasional observations in our laboratory (U. Kües and E. Polak, unpublished data), suggest that both generation modes exist in C. cinereus.

View larger version (136K): [in this window] [in a new window]   FIG. 6.   Young chlamydospores in the submerged mycelium of monokaryon 218 (A3 B1).

Despite their poor cytological description, chlamydospores have been used to collect component monokaryons from dikaryotic mycelia. Chlamydospores from dikaryons generally contain one nucleus of each parental type. Dikaryotic chlamydospores germinate with either one germ tube or two germ tubes, one at each end. When two germ tubes are present, each will receive one nuclear type. Monokaryotic hyphal cells can be isolated by microsurgery using a sharp scalpel before the germ tubes newly dikaryotize each other (121, 266). However, the fact that oidia production has now been observed in dikaryotic cultures after light illumination simplifies the process of monokaryotic culture isolation, but there is normally an uneven recovery of the two nuclear types (175a, 383).

FRUITING-BODY DEVELOPMENT

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Developmental Course of Fruiting

Fruiting-body formation is the most complex developmental process in the life cycle of C. cinereus. Fruiting-body development involves a dramatic change of the growth pattern from a relatively regular three-dimensional loose mesh of free, undifferentiated hyphae to a compact multihyphal structure composed of many different cell types that associate with each other in distinct hypha-hypha interactions (335, 337, 338). Fruiting-body formation in Coprinus is a rapid process. From the first sign of fruiting (formation of initials from hyphal knots) to maturation (autolysis of the cap of the mature fruiting body), it takes about 4 to 5 days (263, 277, 344, 353, 450) (Fig. 7). To initiate and continue fruiting-body development, several environmental signals are called for, of which dark and light periods are the most important (see below). Indeed, fruiting-body development is perfectly synchronized to the light-dark rhythm fixed by the normal day-night cycle (21, 40, 184, 203, 277, 350, 351, 353, 476) (Fig. 7). Hyphal knots are formed in the dark (261, 330), and light then induces the formation of globose fruiting-body initials. Within the initials, cellular differentiation begins, but a further light signal is needed to develop primordia with distinctive cellular tissues (21, 184, 203, 277, 350, 351), otherwise the initials will grow into elongated structures with underdeveloped caps (see below), variously called pseudorhizas (32, 56, 60) dark stipes (353, 476) or etiolated stipes (40, 120). Following another dark period, light is needed to complete meiosis within the basidia, while stipe elongation and cap maturation take place in parallel. Meiosis will be finished by midnight, and basidiospore formation and cap opening occur in the early-morning hours. Within a few more hours, the cap will autolyze and the black basidiospores will be released within a brown liquid (283, 286, 353) (for further details of light regulation, see below).

View larger version (18K): [in this window] [in a new window]   FIG. 7.   Fruiting-body development of homokaryon AmutBmut (A43mut B43mut) in a 12-h-dark/12-h-light regime (light intensity, 50 to 75 µE m2 s1; light source: Osram L36W) at 25°C and 90% humidity (Boulianne et al., submitted; Kües, Granado, et al., unpublished). Dark periods are indicated by black boxes, and light periods are indicated by white boxes. "Light" arrows indicate time points where light positively influences a developmental process: the first signal causes the induction of initial formation, the following two induce gill development in the cap and suppress the formation of etiolated stipes, which will appear when, instead, cultures are transferred in constant dark. Further on, light will induce premeiotic DNA replication and another signal influences meiosis. For details on karyogamy and meiosis, see the text. Note that changing the light intensities and/or the length of light incubation time might change the length of the intervals for the different developmental steps, especially the course of initial and primordium formation and the length of meiosis (U. Kües and J. D. Granado, unpublished data; B. C. Lu, personal communication).

Fruiting-body development in C. cinereus is monocentric: a hyphal knot (also referred to as a hyphal tuft [335] or nodulus [93, 94]) forms as a single independent organ from which a fruiting-body initial may arise (302, 303). In the literature, there is no sharp distinction between a hyphal knot and an initial, and the difference between the initial and the primordium is not easy to understand due to the different uses of these terms by different authors (see, e.g., references 93, 94, 303, 335, 338, and 344). Hyphal knots as defined by Reijnders (413, 415) occur in various tissues of basidiomycetous fruiting bodies and their primordia (e.g., in the stem, veil, and cap). Reijnders (413, 415) defines hyphal knots as aggregates of hyphae that form small communities consisting of an induction hypha with the surrounding hyphae brought under its influence (for further discussion, see reference 338).

Work in our laboratory focuses on fruiting-body initiation. This has made it necessary to define the initial hyphal knot in fruiting-body formation of C. cinereus more specifically as areas of intense localized branching of undifferentiated hyphae within the vegetative mycelium (R. P. Boulianne, Y. Liu, M. Aebi, B. C. Lu, and U. Kües, submitted for publication). (Fig. 8, left). Mature hyphal knots are about 0.2 mm in size (201, 302, 344; Y. Liu, personal communcation). Microscopic studies revealed that the simplest hyphal knot arises from just a single hypha that will intensely ramify with short branches, which in turn give rise to further generations of branches with restricted tip growth (46; Liu, personal communication). In the more common case, hyphal knots originate from more than one generative hypha. Branches of neighboring aerial hyphae will grow toward and alongside each other and possibly merge by anastomosis through the lateral hyphal walls to form an intricate and easily distinguishable lattice. Such locally restricted interlacings will serve as the active center for the intense production of short hyphal branches. Hyphal cells within the resulting bunches of branches are often of a globose or inflated morphology (201, 302; Liu, personal communication). The process as described so far occurs in the dark and is repressed by illumination with blue light (261, 330). However, continuation of development toward fruiting-body formation is light dependent (277, 350) (Fig. 8, middle), indicating a major change in regulation and the beginning of a new, distinct developmental phase. This new phase includes a switch from a (purely) ramifying to an aggregating mode of hyphal growth and results in the formation of a round, aggregated body of tightly interwoven hyphae (95, 335, 338; Boulianne et al., submitted). For this reason, we restrict the definition of a hyphal knot to the locally restricted bunch of hyphal branches which is formed in the dark and which does not necessarily involve hyphal aggregation (Boulianne et al., submitted). These dark-formed structures are not fruiting-body specific but also serve as precursors in the dark-dependent development of sclerotia (330) (Fig. 8, right) (see below). Thus, the formation of globose hyphal aggregates from hyphal knots in the light is the first fruiting-body-specific stage (330, 338), and these structures are referred to as the fruiting-body initials (see also references 263, 344, and 450). This distinction between hyphal knots (i.e., the primary nodules of Clémençon [93, 94]) and initials (in their early appearance, the secondary nodules of Clémençon [93, 94]) has recently been confirmed by using molecular markers. Two different galectins, Cgl1 and Cgl2, belonging to a -galactoside sugar binding subgroup of the lectin family, had been isolated from Coprinus mushrooms of a wild-type dikaryon and an Amut Bmut homokaryon (44, 96; Boulianne et al., submitted). Expression of Cgl2 in the Amut Bmut homokaryon has been shown to correlate with hyphal knot formation and to be repressed by light, whereas Cgl1 expression starts with the development of initials and is light-induced (Boulianne et al., submitted). Mutants have served before to elucidate fungal development (95), and we generated a collection of mutants from an Amut Bmut homokaryon by restriction enzyme-mediated DNA integration (REMI) and UV mutagenesis (145; Kües, Granado, et al., unpublished). Among these mutants, we identified a number of strains with specific defects in fruiting-body initiation, having a block either before or directly after hyphal knot formation (knt and prm, respectively). Most interestingly, we found one mutant that formed hyphal knots but produced neither Cgl1 nor Cgl2, some knotless strains that produced only Cgl2, and mutants that formed knots but no initials and produced either only Cgl2, or, in rare cases, both galectins at a low level. These classes of mutants correlate well with the distinction of hyphal knot and initial and a two-phase regulation of development (Boulianne et al., submitted; Liu, unpublished). Galectins are thought to be involved in hypha-hypha aggregation (44, 96), and the results obtained with the mutants indicate further that galectins are not essential for hyphal knot formation (Y. Liu, M. Aebi, and U. Kües, unpublished data), consistent with the notation that aggregation is not essential for hyphal knot formation.

View larger version (59K): [in this window] [in a new window]   FIG. 8.   Hyphal knots in the vegetative mycelium of monokaryon 218 (A3 B1) that were formed in the aerial mycelium the dark after transformation of the strain with the A43 mating-type gene a1-2 (left). When the cultures are transferred to a 12-h-light/12-h-dark regime at 25°C, primordia develop (middle); when they remain in the dark, sclerotia form from hyphal knots (right) (261).

In the hyphal knot, there is a noticeable reduction in the chitin content with respect to that in the vegetative hyphae (201). However, the globose hyphal aggregate of the initial, the "primordial bud" (277), is the first structure that shows clear histological differentiation. It is enclosed within a coat of large, vacuolated, and mainly outwardly directed hyphae covered with various amounts of amorphous material that protect a compact core of prosenchymal tissue. In the young initial (about 0.2 mm in diameter), this core of heavily branched short cells is only loosely grouped, but in older initials (up to 0.5 mm in diameter), the cells are tightly twisted and fixed together (303, 500). The densely packed cells in the core are rich in glycogen, and between the cells, a large amount of mucilaginous material is present which is probably involved in aggregation. Anastomosis and presumably exchange of cytoplasm occur frequently between adjacent cells. Septa between core cells are usually surrounded at one or both sides by a parenthosome and an additional hemispherical cap also composed of ER membranes. In contrast, cellular fusions do not occur between the large hyphae of the outer layer and the septa are dolipores with normal parenthosomes (500). The current lack of suitable molecular markers makes it difficult to give any information about the possible mechanisms involved in defining the further morphogenesis of the structures which eventually lead to distinctly differentiated tissues within the core area. Since the fruiting-body embryo is initiated within a hyphal agglomeration and is protected by outer layers of large cells (veil cells), which rupture later in development, fruiting-body formation in C. cinereus is classified as hemiangiocarp (95, 335, 512) (Fig. 9). Most important is the change in polarity: the lower and the upper parts of the core of the initial give rise to the fruiting-body stipe and cap, respectively (303, 362). It is not clear when the change from undirected hyphal growth to a defined polarity is determined, and the responsible physiological determinants are also unknown. As a step forward in elucidating this problem, a promising method of vital staining with Janus green has recently been developed to identify initials easily and to study the physiological processes involved in fruiting-body initiation (430).

View larger version (46K): [in this window] [in a new window]   FIG. 9.   Schematic presentation of a slightly tangential section of a primordium. The main tissue organization is indicated. Further details are described by Reijnders (413, 414).

Following the definitions of Moore and coworkers (263, 344, 450), up to a maximal size of 2 mm we will call the developing globular, later more oval aggregation the initial. Staining of fixed material indicated that the first cytological differentiation within the aggregate becomes obvious at a size of about 0.4 to 0.7 mm. Differentiation starts visibly in the upper third of the aggregate, the prepileus region. This ventral area is still prosenchymal and is the base for the upper peripheral cells (pileus trama) and, underneath, the ring of cells in which polysaccharides (glycogen) accumulate. Cells in the middle part of the initial line up to give rise to the stipe tissue, whereas cells in the lower third are randomly orientated to form the basal plectenchyma, representing a tissue of tightly packed hyphal cells not belonging to the stipe. A distinct mushroom shape emerges at a size of 0.8 to 1.0 mm, when hyphal tissue grows from the subapical region in an outward direction to give the annular boundaries of the pileus (303, 335, 344, 362, 414).

The whole complex is surrounded by a marginal veil of hyphae which originate from the top of the pileus (303, 344, 353, 362, 414, 422, 424). Veil cells are large, multinucleate, and septate, but they lack clamp connections (277). With an increasing size of 1.5 to 2 mm, a dense band develops which clearly defines the boundary of the hymenium and stipe (277, 333, 344, 362, 414, 422, 424). To suppress the formation of etiolated stipes and to continue hymenium development (referred to as day 0 by Lu [277]), light is needed at this stage (277). In the lateral part of the cap, between the trama of the cap (pileus) and the differentiated veil cells, a zone of cell division (meristemoid) of strictly parallel hyphae appears (413, 414). Glycogen deposits can be detected in the basal plectenchyma and in the hymenium (89, 197, 303, 344). The hymenium differentiates further into dome-shaped rudiments, which eventually will become the gills (lamella), which carry the basidia in their outer hymenial cell layer (89, 277, 333, 413, 414) (Fig. 9). Primary gill formation proceeds from the edge closest to the stipe toward the cap and from the margin to the apex of the cap (277, 333, 422, 424). Autodigestion of the cells in areas of the gill cavities has been proposed to be the origin of gill construction (277, 282), although the cell debris and the multivesicular and membraneous residual bodies seen in electron micrographs and taken as remnants of degenerating cells possibly represent fixation artifacts (335, 337, 338). Support for the occurrence of morphogenetic cell death as part of the programmed process of primordia development comes from studies with Agaricus bisporus and other mushrooms. In Agaricus, programmed cell death repeatedly occurs during establishment of the hymenial chamber within the primordia (339, 478, 479). It has been suggested that this cellular autolysis contributes to the formation of the primary extracellular matrix (ECM) (479).

Light is essential to enter the next stage of development (203, 277, 351) (Fig. 7), termed stage 1 primordia by Moore and coworkers (263, 344, 450), day 1 primordia by Lu (277), and stage 0 by Kamada et al. (203). Light induces hymenium differentiation and ultimately karyogamy (203, 277, 351) (see below), but the nuclear prefusion phase is long (taking about 24 h) (479). During this time, stage 1 primordia grow from about 2-3 mm to 7-9 mm in diameter. They are characterized by well-developed gills containing basidia of about 12 µm and cystidia (large sterile cells sized between 20 and 25 µm) at their surface (263, 344, 402, 450). Gills are essentially vertical plates arranged radially around the stipe. Primary gills are connected with their tramal tissue of their apex to the outer layers of the stipe; consequently, they have a hymenium at both their sides but not at their edge. With increasing primordium size, space for further gills becomes available within the cap. Secondary and lesser ranked gills are formed by bifurcation of the primary gills. Secondary gills grow radially outward, away from the stipe, by a replication fork-like movement of the gill organizer (a hypothetical formative element in the tissue at the extreme end of the gill cavity) into the trama of the primary gills. As the gill cavity moves outward, it expands tangentially, making room for a new gill organizer to appear, which initiates further gill splitting (89, 333, 414, 422, 424). When space becomes available, further gills may arise from prosenchymal tissue as folds of the cap. Gills of this kind appear as convoluted plates usually in the most apical regions of the cap (89, 422; for further reading on the process of gill formation, see references 95, 335, 337, and 338). Due to their mode of formation, secondary gills are not attached to the stipe. Therefore, their hymenium is continuous over the gill edge. At the edge of the secondary gills, there is an increased number of cystidia (333, 414, 422, 424). The hymenium develops as a layer of similar-sized palisade-shaped cells by branching from sister hyphae of the prosenchymal subhymenium. This hymenial layer of cells is occasionally interrupted by larger cystidia (90, 179, 344, 422-424). Hymenial cells have clamp cells, and the nuclei stay paired, indicating that the dikaryotic stage is preserved (277, 344). During further development of stage 1 primordia, the basidia change from a cylindrical to a club-shaped structure (402). Glycogen accumulates in the subhymenium and cystidia and in the basal plectenchyma of the stipe (263, 344, 450). The timing of this stage is difficult to determine, and this makes it particularly difficult to compare the observations on primordium development made by different researchers (21, 203, 277, 344, 351). Karyogamy is therefore used as a reference point (277). Karyogamy occurs toward the end of this phase and is considered, according to Lu's definition, to have begun when 5% of the basidia have undergone nuclear fusion (277, 402) (see below). At this point, by the beginning of the following day, tissue development in the cap is completed (277).

Stage 2 (day 3) primordia (Fig. 7) are 6 to 10 mm (occasionally up to 15 mm) tall. Meiosis starts within the basidia (see below), which are now 15 to 18 µm long. The cystidia enlarge up to 45 µm. Primary gills are still connected to the stipe, but the gills begin to separate. Initially, the veil of stage 2 primordia is still intact, but it becomes more and more free. In the cap, more glycogen accumulates, while the glycogen content in the basal plectenchyma decreases (263, 277, 344, 450). Galectin distribution has been analyzed in stage 2 primordia. Galectin antibodies localize preferentially to the veil cells, to the outer layer of narrow hyphae of the stipe, and to the apex of the primary gills connected to the stipe (Boulianne et al., submitted). Significantly, these different tissues (called in specialist terms lemmablem [the external tissue of the cap] and lipsanoblem [the external tissue of the stipe], respectively) are functionally related in protection of the inner primordium tissues, and at least the veil cells of the cap are dispensible at later stages and degenerate or are discarded (95, 155). To a lesser extent, galectins are also present in the gill trama and the inner tissues of the stipe. Galectins are found attached to cell walls and in the extracellular matrix and, within the cells, localized in vesicles and in the dolipores (Boulianne et al., submitted). Consistent with the immunolocalization studies, Northern blot analysis indicates high expression of galectin genes in stage 1 and stage 2 primordia, but at later meiotic stages (late pachytene/diplotene, metaphase I) at the transition to the next developmental stage (fruiting-body maturation) galectin expression is halted in both the cap and the stipe (85).

The further development of stage 2 primordia into mature basidiocarps takes another 24 h (263, 277, 344, 353, 394, 450). Maturation consists of at least four processes: stipe elongation, basidium maturation (including meiotic divisions and spore formation), pileus expansion, and autolysis of the pileus (see below).

The mature stipe is a thin-walled hollow cylinder with two types of hyphae, i.e., narrow and wide hyphae. It has been found that 23 to 54% of all hyphae are narrow. Most of these narrow hyphae are concentrated at the exterior of the stipe, but they are also found at the lumen and in between the wide, inflated, and highly vacuolated hyphae that make up most of the interior tissues (39, 156). The narrow hyphae resemble vegetative hyphae, since they clearly have clamp cells and form interconnections independent of the inflated hyphae (39). This network of narrow hyphae has been suggested to act in nutrient translocation from stipe to cap (188, 335, 338). Cytologically, the cell walls of the narrow hyphae at the different locations differ from each other and from those of the inflated hyphae, as indicated in various staining experiments (39, 139, 156) and by a different immunohistochemical response with antigalectin antibodies (Boulianne et al., submitted). On the whole, the composition of cell walls of the stipe seems very much like that of monokaryotic hyphae and is very different from that of vegetative dikaryotic hyphae (298). However, compared to the monokaryotic mycelium, the overall chitin content is increased in the stipe (298), consistent with the facts that chitin synthesis is especially active in stipe formation (106, 137, 140, 143) and that the chitin synthesis inhibitor polyoxin D suppresses stipe elongation (144). Chitin microfibrils, 7 to 25 nm in diameter, are indeed a major component of the walls of stipe cells. They are arranged as shallow helices transverse to the long axis of the cell. Two-thirds of these helices are left-handed, and one-third are right handed (140, 195, 210). In contrast, chitin microfibrils are randomly orientated in the vegetative mycelium; the shift to a parallel transverse arrangement can occur in hyphal knots of only 0.1 to 0.2 mm in diameter, although at this point it is still incomplete (201).

Stipe growth is mainly the result of manifold cell elongation rather than cell division (39, 99, 142, 196). However, in the very young primordium, stipe cells are cuboid or polyhedral, and proliferation still occurs in the apex meristematic region of the stipe (39, 155, 302). At the stage of karyogamy, many of the cells inflate and become cylindrical, and it is obvious that these enlarged cells lack clamp connections, in contrast to cells found in the central yet closed region of the stipe (39, 155, 156, 277). During stipe elongation, inflated hyphae inflate further but only vertically; the width of the stipe cells remains almost constant (155, 156, 335, 337, 338). However, the proportion of narrow hyphae declines as the stem grows from 45 to 70 mm, indicating that normal vertical stem extension involves both an increase in the cross-sectional area of inflated hyphae and recruitment of narrow hyphae into the inflated population (156, 335, 337, 338).

The enlarged stipe cells are usually multinucleate, and more nuclei are progressively produced by mitosis as the cells grow by vertical elongation (277, 449). Indeed, there is a linear correlation between the length of the stipe and the numbers of nuclei found within the elongated cells. There are between 2 and 8 nuclei in stipe cells of 2- to 4.5-mm primordia, about 16 at a primordial size of 5 to 7.5 mm, and more than 150 when the stipes are longer than 20 mm. In 29% of stipe cells, nuclei are present in uneven numbers and there is no indication of synchronized nuclear division (142, 449). The increase in the number of nuclei is indicated by the dramatic increase in DNA content 3 h before the onset of rapid stipe elongation (142, 202). In contrast, the RNA content increases continuously with stipe maturation and the protein content is constant (141, 202, 344). The young stipe contains large amounts of glycogen inclusions, whereas glucans, including glycogens, are almost absent in the fully expanded stipe, probably because they serve as a source of cell wall precursor material (32, 195, 197, 198). Apart from glycogen inclusions, insoluble protein inclusions were also detected which first increase but then decrease in size with progressive stipe development (32). This is most interesting, since, apart from the intercellular compartments and cell walls, the galectins have been localized to vesicles within stipe cells of stage 2 primordia and young elongating fruiting bodies (Boulianne et al., submitted).

Decapitation of the stipe before meiosis arrests stipe expansion, showing that the cap is needed for the early development of the stipe (39, 99, 138, 150, 195). Consistent with this, grafting experiments revealed that during meiosis, the cap produces a diffusible substance acting on basidiocarp maturation, including stipe elongation (200). Rapid stipe elongation correlates with the end of meiosis 8 h after the nuclei in the bulk of basidia have fused (155, 202). Although the cap still influences the absolute length a stipe can achieve, in the postmeiotic basidiomes the stipe and cap develop independently from each other. Also, the stipe does not need to be connected to the vegetative mycelium, indicating that stipe elongation is an autonomous endotrophic event requiring no exogenous nutrients, growth factors, or water (39, 99, 138, 142, 155). Elongation is not evenly distributed over the length of a stipe. The ultimate apex of the stipe does not extend much, although in general the cells in the apical half elongate most, in contrast to the cells of the basal half, which show reduced stretching, and the cells in the base, which show no stretching at all (39, 99, 156, 196, 198, 344). Stipe cells elongate in an intercalary fashion by diffuse extension in which the extension of the cell surface is not confined to the hyphal tip but occurs throughout the whole cell (139, 141, 156, 195). The stipe has to be formed quickly and as strongly and economically as possible. Therefore, cell walls have to be plastic but also rigid to withstand the high turgor pressure. Plasticity is given by the ordered arrangement of chitin helices, and rigidity is given by cross-linking between chitin and glucans (195, 196, 198). Throughout elongation, the chitin content stays constant and insertion of new chitin microfilaments occurs over the whole length of the cell in a uniform intercalary fashion, probably mediated by a constant unfastening and resealing of linkages between chitin and glucan (139, 141, 197, 210). Consistent with active cell wall hydrolysis playing a role in the mechanism of stipe elongation, stipes are rich in chitinases and glucanases (204, 205, 207).

An average stipe may elongate 80 mm in less than 12 h (195, 341, 449). It is very strong in its longitudinal axis and withstands pressures of 7 × 10⁴ N m², a fact impressively shown by the occasional documentations of fungal breaks through asphalt (58, 142, 327). Rapid stipe elongation requires high turgor pressures (142, 327), and throughout elongation the osmotic value stays almost constant at 0.45 to 0.5 M (99, 195). This high turgor must be actively maintained, probably by organic metabolites which have not yet been identified (39, 141, 142, 338). Trehalose and polyols have been excluded as the sole osmotic stabilizers, since the concentration of the former decreases during elongation whereas the latter are hardly present in stipes at all (141, 142, 338, 406). Amino nitrogen compounds and urea have also been implicated in osmostabilization, but their contents are low and decline with development (123, 142, 344). Regularly in fruiting-body development, shortly before rapid stipe elongation, clear or sometimes yellowish droplets are excreted at the region where the cap with the gills is attached to the stipe (241; Kües, unpublished). These droplets are rich in potassium oxalacetate, but it is not known whether this contributes to the high turgor pressure of the stipe (241). Cox and Niederpruem (99) reported that a brown gel was present within the lumen of young stipes but disappeared during elongation, and this has also been discussed as a source of an organic osmotic stabilizer (195). Since no sugars or other substances have been found to be of prime importance, cocktails of simple carbohydrates together with inorganic ions are anticipated to control the osmotic pressure in stipe elongation (338, 344).

Basidiospore formation. Basidia are the only C. cinereus cells that express developmental comittment. Probasidia can easily be arrested in development (86, 277, 285), but once meiosis is initiated, maturation of basidia is an autonomous, endotrophic process (86, 314) (Fig. 10). The basidium usually arises as the terminal cell of a hyphal branch and is separated from the subjacent cell by a dolipore septum with a well-formed septal cap at the side of the subjacent cell (312, 314). The cytoplasm in the probasidia is relatively simple with numerous free ribosomes, few vacuoles, mitochondria, and limited ER. At the time of nuclear fusion, glycogen is evident at the basidial base. An increase in the amount of rough ER is observed at the end of prophase I, and ER extends throughout the cell in the postmeiotic basidium. Usually, as basidiospores form, a vacuole at the base of the basidium gradually expands to fill the cell (169, 314).

View larger version (16K): [in this window] [in a new window]   FIG. 10.   Karyogamy, meiosis, and basidiospore formation.