Place the items into the appropriate category characterizing fungal mitosis or animal mitosis.

“Deuteromycetes,” the Fungi Imperfecti

The deuteromycetes, commonly called molds, are “second-class” fungi that have no known sexual state in their life cycle, and thus reproduce only by producing spores via mitosis, This asexual state is also called the anamorph state. Despite its “-mycetes” ending the deuteromycetes are not a class; rather this term is now used as a common name for this dumping-ground group of fungi. About 90% of these have affinities to the Ascomycota. Most food spoilage (Figures 9 and 10) and fungal human diseases are caused by members of this group. They are also known as the fungi imperfecti, because of their “imperfect” lack of sex. When the “perfect state” of one of these organisms is discovered, as happens every year, the fungus is more properly classified with the teleomorph name, although in practicality, like for people, it is difficult to remember to switch to their “married” name. Notice that this group is not classified as one of the phyla. It is just a loose assemblage of organisms that we have not been sure where to place accurately in the taxonomic order. Although we can accurately place most of these fungi into phyla using molecular biology techniques such as PCR and DNA sequencing, mycologists maintain this group and its genera for convenience and tradition (St-Germain and Summerbell, 1996).

Defining Features of the ‘Class’

The deuteromycetes is an artificial grouping in that the phylogenetic relationships among taxa are mostly unknown or not apparent. They are the mitotic states of meiotic groups such as the basidiomycetes and especially the ascomycetes, or have evolved from them. A very small number of taxa have been correlated with meiotic states but the majority have not. Thus there is a residual body of taxa which cannot easily be incorporated into the classifications for meiotic fungi. This situation will become less problematic as the results of molecular characterization and their application to fungal systematics become more widespread. For the moment, however, there is a serious lack of information about DNA-based typification, and apart from classifications for ascomycetes and basidiomycetes there is still no taxonomic system to cope with these other fungi. Over the last 200 years, separate classifications have been developed for mitotic fungi and until recently these have arisen independently from classifications for ascomycetes and basidiomycetes. Several names, both formal (nomenclatural) and informal (colloquial), have been used in the past for groups of mitotic fungi. These include Deuteromycotina, Deuteromycetes, Fungi Imperfecti, asexual fungi, conidial fungi, and anamorphic fungi. The most recent suggestion, accepted in the Dictionary of the Fungi (8th edn), is ‘mitosporic fungi.’ Colloquial names such as this and others have no nomenclatural standing. Neither these nor the formal names that have been proposed are in any way equivalent to the names used for taxonomic categories accepted in basidiomycete and ascomycete systematics that are governed by the International Code of Botanical Nomenclature. Although many class, subclass, order, suborder, and family names have been used in the group in the past none of these is currently accepted, and if they are used at all it is in an informal manner. Even the use of generic and specific names (which are allowed by the Code) must be with qualification, for they are also not equivalent to those employed in ascomycetes and basidiomycetes. Sometimes they are referred to as form genera and form species. Despite this there is still a need for a framework on which to hang the information used in identification of mitosporic fungi. Until the 1950s, taxa were differentiated primarily by the nature of the fruiting structures and conidial morphology. However, since that time the systematics of the group has largely depended on aspects of the conidiogenous processes exhibited by the fungi involved.

Discussions concerning the classification of, or information frameworks for, mitosporic fungi ignore the ultimate aim to do away with the group and incorporate its members into the classifications for ascomycetes and basidiomycetes. The use of DNA technology is the key to accomplishing this.

The group is characterized by the absence of teleomorphic (meiotic) states. It is heterogeneous, i.e., polyphyletic. Reproduction is commonly by spores (conidia) produced mitotically (asexually) from conidiogenous cells which are sometimes free (as in yeasts) or more commonly formed on separate supporting hyphae (conidiophores) and/or cells which may be produced in or on organized fruiting structures (conidiomata). Taxa are separated by differences in conidiogenous events and the structures involved, conidiomatal form and development, conidial morphology, colony characteristics, and the presence and nature of vegetative structures.

Biological Aspects

Growth of imperfect fungus Aspergillus depends upon having both cell wall-bound and extracellular proteolytic enzymes to satisfy requirements for nutrition, because the fungi feed entirely by absorption, not by photosynthesis or ingestion. Aspergillopepsins are extracellular proteinases secreted by fungal mycelia. The enzymes are of practical importance for fungal nutritional in an acidic environment. Aspergillopepsin I from A. saitoi shows two forms of activity at acidic pH, pepsin-like catalytic function and trypsinogen-activating activity like enteropeptidase (Chapter 586) [6,38].

The major extracellular protease from A. niger, which is responsible for 80–85% of the total activity, is aspergillopepsin A, a protein of approximately 43 kDa, the activity of which is inhibited by pepstatin [51]. The gene for aspergillopepsin A (pepA) is located on chromosome 1 of A. niger.

Hydrolysis of structural proteins in the lung by extracellular proteinases secreted by A. fumigatus is thought to play a significant role in invasive aspergillosis. The fungus secretes not only elastolytic aspartic proteinase (aspergillopepsin F) but also an elastinolytic serine proteinase, oryzin (Chapter 713) and a metalloproteinase (Chapter 286) [12]. Aspergillopepsin F can catalyze hydrolysis of the major structural proteins of basement membrane, elastin, collagen and laminin. Immunogold electron microscopy showed that the aspartic proteinase was secreted by A. fumigatus invading neutrogenic mouse lung and its secretion was directed towards the germ tubes of penetrating hyphae [12]. Deletion or disruption of the gene for aspergillopepsin I has no effect on the virulence of A. fumigatus [52], although such treatment reduces extracellular proteolytic activity 85% or more in A. niger [53].

Characteristics of the Genus and Relevant Species

Mitosporic Fungi

The ‘mitosporic fungi’ (asexual, anamorphic, imperfect, conidial, deuteromycete fungi) are an artificial group without a formal nomenclature above the generic level, comprising the mitotic states (anamorphs) of the meiotic ascomycetes and basidiomycetes (teleomorphs) and mitotic fungi that have not been correlated with any meiotic states. They are characterized by the formation of conidia as a result of presumed mitosis. Separation of genera is primarily by mode of conidiogenesis and growth of the conidiogenous cell, with morphology of conidiomata, conidia, and conidiophores as subsidiary criteria.

Acremonium – see Emericellopsis and Nectria, but many species of polyphyletic ascomycete affinity have no known teleomorph.

Alternaria – see Clathrospora and Leptosphaeria, but many species of undoubted ascomycete affinity have no known teleomorph.

Aspergillus – see Emericella, Eurotium and Neoasartorya, but many species of ascomycete affinity have no known teleomorph.

Aureobasidium – colonies covered by slimy, yellow, cream, pink, brown or black masses of spores. Aerial mycelium scanty, immersed mycelium often dark brown. Conidiogenous cells undifferentiated, procumbent, intercalary or on short lateral branches. Conidia produced synchronously on multiple loci in dense groups on short scars or denticles, hyaline, smooth, with a truncate base. Distribution worldwide, saprobic, from soil, leaf surfaces, cereal seed, on flour, tomato, pecan nuts, fruit, fruit drinks.

Basipetospora – see Applications of Monascus-Fermented Products.

Botrytis – see Botryotinia, but many species have not been linked to teleomorphs.

Brettanomyces – see Dekkera. There are a number of species not linked to teleomorphs.

Candida – see Debaryomyces; Issatchenkia; Kluyveromyces; Pichia; Saccharomyces; Torulaspora and Yarrowia, but many species of polyphyletic ascomycete affinity have not been linked with teleomorphs.

Chrysonilia – see Neurospora.

Cladosporium – see Mycosphaerella, but many species have not been linked to teleomorphs.

Epicoccum – colonies fluffy, yellow, orange, red, brown, or green. Conidiophores formed in black sporodochial conidiomata, closely branched, compact and dense. Conidiogenous cells pale brown, smooth or verrucose, integrated, terminal, determinate, cylindrical. Conidia solitary, dry, subspherical to piriform, dark golden-brown, often with a pale, protuberant basal cell, muriform, rough, opaque. Distribution worldwide, from soil, cereal seed, beans, mouldy paper, textiles.

Fusarium – see Nectria and Gibberella, but many species have no known teleomorph.

Geotrichum – see Galactomyces, but many species of polyphyletic ascomycete affinity have not been linked to teleomorphs.

Moniliella – colonies acidophilic, restricted, smooth, velvety or cerebriform, cream then pale olivaceous or black-brown. Cells often budding to produce a pseudomycelium. Conidiophores undifferentiated, hyaline, smooth, repent. Conidia formed in acropetal chains from individual (conidiogenous) cells of the mycelium, hyaline, smooth, aseptate, ellipsoid. Thallic conidia also formed by fragmentation of hyphae, becoming thick-walled and brown. From Europe and the United States, occurring in pickles and vinegar, fruit juices, syrups, and sauces.

Paecilomyces – see Byssochlamys and Thermoascus, but many species of ascomycete affinity have no known teleomorph.

Penicillium – see Talaromyces and Eupenicillium, but many species of ascomycete affinity have no known teleomorph.

Phialophora – colonies slow-growing, olivaceous black, sometimes pink or brown. Conidiophores erect, hyaline or pale brown, branched or reduced to simple hyphae. Conidiogenous cells clustered or single, phialidic, lageniform or cylindrical, with a distinct darker collarette. Conidia formed in slimy heads or in chains, aseptate, globose to ellipsoid or curved, mostly hyaline, smooth. See Coniochaeta, Mollisia, Pyrenopeziza, with Phialophora-like anamorphs, also linked with Geaumannomyces, but several species with no known teleomorph. Worldwide in distribution but most common on decaying wood, wood pulp, secondarily soil-borne, from water, fermented corn dough, foodstuffs, butter, wheat.

Phoma – colonies comparatively fast-growing, gray, olivaceous, brown, fluffy. Conidiomata pycnidial, black-brown, ostiolate, sometimes setose. Conidiophores absent. Conidiogenous cells ampulliform to doliiform, hyaline, smooth, phialidic. Conidia hyaline, smooth, aseptate or sometimes septate, ellipsoid, ovate, cylindrical. Dark-brown unicellular or multicellular chlamydospores sometimes formed. Some teleomorphs in Pleosporaceae (Pleospora), but most species with no known teleomorphs. Distribution worldwide, from soil, butter, rice grain, cement, litter, paint, wool, and paper; also produces mycotoxins.

Rhodotorula – colonies pink, with carotenoid pigment soluble in organic solvents, mycelium and/or pseudomycelium formed, cells usually small and narrow. Conidia spherical, ovate or clavate, with a narrow or rather broad base, budding. Sometimes assimilates nitrate, but fermentation absent. Teliospores absent, but basidium-like structures in some species indicate basidiomycete affinity with Rhodosporidium (Sporidiobolaceae). From wood, involved in spoilage of dairy products, fresh fruit, vegetables and seafoods, especially refrigerated foods.

Scopulariopsis – see Microascus, but many species of ascomycete affinity have no known teleomorph.

Stachybotrys – colonies black to black-green, powdery. Conidiophores erect, separate, simple or branched, septate, becoming brown and rough at the apex. Conidiogenous cells grouped at the conidiophore apex, phialidic, obovate, ellipsoid, clavate or broadly fusiform, becoming olivaceous, with a small locus and no collarette. Conidia in large, slimy black heads, ellipsoid, reniform or subglobose, hyaline, gray, green, dark brown or black, sometimes striate, coarsely rough or warted, aseptate. Distribution worldwide, from soil, paper, cereal seed, textiles. Trichothecene mycotoxins produced such as satratoxin but its toxicity is unknown.

Trichoderma – see Hypocrea, but many species of ascomycete affinity have no known teleomorph.

Trichosporon – colonies slow-growing, white to cream, butyrous, smooth or wrinkled. Mycelium repent, hyaline. Conidiophores absent. Conidia of two types: (1) thallic, formed by fragmentation of the mycelium, cylindrical to ellipsoid; (2) blastic, formed in clusters near the ends of the thallic conidia or by budding of the lateral branches of the mycelium, subglobose, with a narrow distinct scar. Distribution worldwide, from humans and animals, saprobic in soil, fresh and sea water, plant material, fermented corn dough.

Trichothecium – colonies powdery, pink. Conidiophores erect, separate, simple, unbranched, septate near the base, rough, apical cell functioning conidiogenously. Conidia formed in retrogressively delimited basipetal chains, appearance zigzagged, hyaline, smooth, one-septate, ellipsoid or piriform, thick-walled, with an obliquely truncate scar. Distribution worldwide, from soil, water, decaying plant material, leaf litter, cereal seed, pecan nuts, stored apples, fruit juices, foodstuffs especially flour products; also a potent producer of trichothecene mycotoxins, but significance to human health is unknown.

Ulocladium – colonies black to olivaceous black. Conidiophores erect, separate, simple or branched, septate, smooth, straight, flexuous, often geniculate, geniculations associated with preformed loci (pores). Conidia dry, solitary or in short chains, obovoid to short ellipsoid, with several transverse and londitudinal or oblique eusepta, medium brown to olivaceous, smooth or verrucose, base conical, apex broadly rounded and becoming conidiogenous. Not uncommon, widely distributed, from soil, water, dung, paint, grasses, fibres, wood, paper, corn, seeds.

Verticillium – colonies cottony, white to pale yellow, sometimes becoming black due to resting mycelium. Conidiophores erect, separate, septate, smooth, hyaline, simple, unbranched or branched. Conidiogenous cells solitary or produced in verticillate divergent whorls, long lageniform to aculeate, hyaline, phialidic. Conidia form in droplets at the apices of conidiogenous cells, hyaline, aseptate, smooth, ellipsoid to cylindrical. Hyaline multicellular chlamydospores and microsclerotia sometimes formed. Distribution worldwide, commonly causing plant wilt diseases, from soil, paper, insects, seeds, bakers' yeast, potato tubers, commercially grown fungi; also forms mycotoxins.

Wallemia – colonies xerophilic, restricted, fan-like or stellate, powdery, orange brown to black brown. Conidiophores erect, separate, cylindrical, smooth, pale brown. Conidiogenous cells apical, long lageniform to cylindrical, finally verrucose, forming a phialidic aperture without collarette from which a short chain of four thallic conidia is formed. Conidia initially cuboid, later globose, pale brown, finely warted. Distributed worldwide, from dry foodstuffs such as jams, marzipan, dates, bread, cake, salted fish, bacon, salted beans, milk, fruit, soil, air, hay, textiles.

SUMMARY OF EXISTING KNOWLEDGE

Several groups of mitosporic fungi occur in freshwater, although the species present at particular sites vary with the types of habitat and substratum represented. The best-known and most studied group of mitosporic fungi is the “aquatic,” or “Ingoldian,” hyphomycete group, members of which are distinguished by their tetraradiate, branched, or sigmoid conidia that are released into and dispersed by water (Ingold 1975; Webster and Descals 1981; Bärlocher 1992). Although the first species was described in 1880 (Saccardo 1880), it was not until Ingold discovered a varied mycota on alder and willow leaves submerged in a stream (Ingold 1942) that the diversity of this distinctive group of fungi was recognized. About 300 species have been described thus far (Bärlocher 1992). Most aquatic hyphomycetes occur in lotic habitats on decaying deciduous leaves and woody debris of allocthonous origin. A few species, however, have been reported from lentic habitats (Suzuki and Nimura 1961), brackish waters (Jones and Oliver 1964; Müller-Haeckel and Marvanová 1979b), and terrestrial habitats (Park 1974; Webster 1977, 1981; Sridhar and Kaveriappa 1987; Iqbal et al. 1995).

The “aeroaquatic hyphomycetes,” whose conidia are modified in a variety of ways to trap air for flotation, comprise a second group of mitosporic fungi (Fisher 1979; Michaelides and Kendrick 1982; Webster and Descals 1981; Premdas and Kendrick 1991). Mycelia of aeroaquatic fungi are thought to grow on submerged decaying leaves, woody debris, dead emergent macrophytes, and a variety of other decaying plant parts. Unlike most aquatic hyphomycetes, which require submersion for sporulation, the aeroaquatic fungi sporulate when they are exposed to air. Aeroaquatic fungi can be found in all types of aquatic habitats, but they occur most commonly in lentic habitats with fluctuating water levels, such as small ponds, marshes, swamps, and ditches.

Dematiaceous, and to a lesser extent, hyaline hyphomycetes, nematode-trapping hyphomycetes, and coelomycetes all are encountered regularly on a wide variety of submerged plant substrata in both lentic and lotic habitats. Those fungi usually go undetected, however, unless a substratum on which they occur is incubated in a moist chamber after collection (Shearer 1972; Lamore and Goos 1978; Shearer and Crane 1986; Révay and Gönczöl 1990; Sivichai et al. 2000; Tsui et al. 2000). We will not discuss those fungi or the aeroaquatic fungi in detail. Generally, however, the methods used to collect, isolate, and culture them are very similar to those used for the aquatic ascomycetes. Therefore, we refer the reader to the previous section on Ascomycetes and the papers cited therein as starting points for dealing with these fungi. No comprehensive keys to the freshwater mitotic fungi exist. The best strategy for identifying them is to use a general reference to arrive at a suitable genus (Barron 1968; Ellis 1971, 1976; Carmichael et al. 1980; Sutton 1980; Barnett and Hunter 1998; Kiffer and Morelet 2000) and then to consult the CABI Bioscience Database of Fungal Names (Appendix III) to find the species described in the genus. Literature citations provided in that database can be used to obtain the original descriptions of species needed for identification.

In the remainder of this section, we deal only with the aquatic hyphomycetes. Descals (1997) described detailed methods for the collection, identification, isolation, and deposition of aquatic hyphomycetes. Basic, and in some instances, different techniques are described in the following sections.

13.3.2.2 Beauveria bassiana

B. bassiana, a Deuteromycete long known for its entomopathogenic properties, causes an insect disease known as white muscardine. The organism produces a number of cyclodepsipeptides such as beauvericin which may account for at least part of its toxicity to insects (Miller et al., 1983). Beauvericin may also have antimicrobial, cytotoxic, and apoptotic activity (Klaric and Pepeljnjak, 2005). B. bassiana has been used as a medicant in Japan for over a millenium (Ignoffo, 1973). Allergic responses have been reported in humans following inhalation of spore preparations, though repeated handling of cultures did not reveal adverse effects in another study (Ignoffo, 1973). A mouse study from China noted hypersensitivity-like pulmonary reactions in mice and rats after a single exposure to B. bassiana. However, the low room temperatures may have constituted a significant stress to the animals (Song et al., 1989, cited in Semalulu et al., 1992). Russian investigators determined the LD50 to be greater than 1.1 × 1010 and greater than 2.2 × 1010 fungal cells in albino rats exposed intragastrically and intraperitoneally, respectively, and greater than 4 × 1010 fungal cells in rabbits exposed intravenously (Mel’nikova and Murza, 1980). No significant toxicity or pathogenicity by the oral, dermal, or pulmonary routes were noted by the U.S. EPA in reviews of a series of acute studies on B. bassiana strain HF23 submitted to support its registration as a microbial pesticide, though there was mild eye irritation (U.S. EPA, 2006). Acute inhalation, hypersensitivity, and immune response studies were waived for this strain based on the evidence for clearance and the absence of toxicity in the other studies, as well as the low toxicity potential of the inert ingredients.

B. bassiana has been implicated in at least two cases of keratomycosis, though both patients had long histories of antibiotic and corticosteroid use (Ishibashi et al., 1986). Separate clinical studies identified Beauveria subspecies colonizing the liver (Henke et al., 2002) or deep skin structures (Tucker et al., 2004) of immunocompromised patients under treatment for leukemia. Direct inoculation into rabbit corneas of B. bassiana isolated from a patient with keratitis resulted in inflammation, corneal ulcers, corneal haze, injection of the iris, and sparse-to-moderate fungal growth in the cornea, though the severity was less than that seen in parallel eyes treated with Candida albicans and tended to resolve with time (Ishibashi et al., 1986). Injection of B. bassiana into the quadriceps muscles of CD-1 mice led to focal muscle necrosis, edema, and inflammation, with the severity of the responses dependent on the number of organisms injected (Semalulu et al., 1992). Muscle regeneration was visible by 7 days. Viable spores capable of initiating colonies in artificial media were not detected after 3 days. While it is unlikely that this organism, which does not grow well at temperatures above 32°C, can infect or colonize humans under normal circumstances, it appears that exposure to Beauveria must be avoided when sick or immunocompromised individuals are present.

Scald

Scald, caused by the imperfect fungus Rhynchosporium secalis (Oudem.) J.J. Davis f. sp. hordei, is a major foliar disease of barley grown in areas of the world where the climate is cooler and more humid because the leaf remains wet for a longer period. Reports about the pathogenic variability of R. secalis are numerous (Brown, 1985; Mc Donald et al., 1999), and in some countries an increase in genetic variability of the fungus has been described (Tekauz, 1991). Numerous papers report pathogenic variability of scald on all five continents (Hansen and Magnus, 1973; Williams and Owen, 1973; Ali et al., 1976; Jackson and Webster, 1976; Metcalfe et al., 1977; Ceolini, 1980; Brown, 1985; Cromey, 1987; Jørgensen and Smedegaard-Petersen, 1995), which directly influences the estimates of the diversity of scald resistance genes in the regions of the world. It is clear that the virulence spectrum of scald populations within regions of barley cultivation is able to change in short periods of time (Tekauz, 1991) and correlations indicate that the variability of scald populations reflect the cultivar composition.

Several evaluations of barley for resistance to scald have been carried out (Fukuyama et al., 1998; Yitbarek et al., 1998), and many resistance genes have also been described (Table 8.2). However, reviews about scald and corresponding resistance genes (Bockelman et al., 1977; Beer, 1991) have been confused, on account of contradictory results about reaction patterns between lines possessing the same resistance. This is most probably due to the use of local pathotypes, a known problem for many foliar diseases in barley. Also, a set of standard differentials to distinguish pathotypes is necessary for the systematic analysis of resistance.

Table 8.2. Main resistance genes to scald.

Characteristics of the Genus and Species

The genus Botrytis belongs to the class Fungi Imperfecti (Deuteromycetes) which contains all those fungi that do not have a sexual (perfect) stage. The conidial stages of these fungi are similar to those of Ascomycetes. The genus Botrytis belongs to Moniliaceae, the largest form-family of the class Fungi Imperfecti. The family includes all imperfect fungi which produce conidia on unorganized hyaline conidiophores or directly on the somatic hyphae. Most species are saprobic, but many are well-known plant pathogens and others are human pathogens. In Botrytis, large oval or spherical conidia are produced at the tips of erect conidiophores which are simple or branched. The conidia are not in chains but in head-like formations and are attached singly on sterigmata. They may be hyaline or, in some species, brightly coloured. Many species are known to have teleomorphs in the genus Botryotinia. The organisms include many plant pathogenic species, e.g. B. allii, B. cinerea and B. fabae. The classification of Fungi Imperfecti is based on the secondary fruiting forms and other external traits and serves exclusively the practical aims of naming and identification.

Clinical Features

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Caused by three genera of imperfect fungi—Epidermophyton, Trichophyton, and Microsporum—that cause superficial infections involving keratinized tissues such as the cornified layer of epidermis, the hair, and the nails

Dermatophytosis involving different anatomic sites are named with site-specific terms such as tinea capitis (scalp), tinea barbae (beard area), tinea faciei (face), tinea corporis (trunk), tinea cruris (intertriginous areas), tinea pedis et manus (feet and hands), and tinea unguium (nails)

Typical lesions of superficial dermatophytosis present as sharply demarcated patches with an arcuate border

Tinea capitis and tinea barbae present as folliculitis; tinea unguium is characterized by yellow-gray discoloration of nails