The taxonomic placement of Gloeotinia is not clearly established. Wilson et al. (1954) placed G. temulenta within the family Sclerotiniaceae, based on its occurrence as a plant pathogen, presence of spermatia and macroconidia, and formation of a fleshy cupulate apothecium from a stroma. Although G. temulenta shares many features of the Sclerotiniaceae, it develops only an interwoven mycelium within the infected seed and does not form the true sclerotium that is characteristic of the Sclerotiniaceae. Ellis (1956) described Gloeotinia as structurally similar to Symphyosirinia, a member of the family Leotiaceae. Similar views were stated by Baral (1994) who considered Gloeotinia and Symphyosirinia related and members of the Leotiaceae, subfamily Hymenoscyphoideae. In 1997, Holst-Jensen et al. (1997) provided data from DNA analysis that Gloeotinia was distinct from other fungi within the Sclerotiniaceae. These studies support the concept that Gloeotina should be considered a member of the Leotiaceae, subfamily Hymenoscyphoideae.
Schumacher (1979) reported that a specimen described on Bromus erectus by Quelet (1883) as Phialea granigena was conspecific with G. temulenta and therefore represented an older name of the fungus. Alderman (1997) recognized G. temulenta and G. granigena as separate species, based on host range and morphological differences. Bromus erectus is not believed to be a host for G. temulenta (Hardison 1962, Alderman 1997). Little is known about G. granigena. Additional studies concerning species identity and their associated host range in the genus Gloeotinia are needed. Unfortunately, specimens of Gloeotinia from outside areas of commercial seed production are very rare in nature.
Two other species of Gloeotinia from Germany have been described: G. aschersoniana (P.C. Hennings and T. Ploettner) H.O. Baral on Carex and G. juncorum (J. Velenovsky) H.O. Baral on Juncus (Baral and Krieglsteiner 1985). Nothing is known of the life history of these species.
Stroma. Infection of the grass caryopsis results in the mummification of the caryopsis, creating a substratal stroma (Spooner 1987, Williams and Spooner 1991). Hyphae, 3-4 µm wide, ramify throughout the pericarp, teste, and endosperm and are not differentiated into rind and medullary parenchyma (Gray 1942, Wilson et al. 1945). A true sclerotium does not develop, although the infected seed functions similarly to a sclerotium as a means of survival through the winter.
Sporodochia. In late winter or early spring, pinkish, pulvinate, gelatinous sporodochia form either on the surface of the pales or between the pales and caryopsis (Neill and Hyde 1939, Gray 1942, Calvert and Muskett 1945, Griffiths 1959b). They are 0.4-1 × 0.5-1.5 mm in size (Prillieux 1897, Neill and Hyde 1939, Gray 1942, Calvert and Muskett 1945). Sporodochia consist of a core of closely septate, branching hyphae (Neill and Hyde 1939, Griffiths 1959b) with the terminal cells of each branch bearing 1-4 microconidiophores (Gray 1942, Griffiths 1959b).
Microconidiophores and microconidia (spermatia). Microconidiophores are 2-5 µm in diameter and 5-9 µm long, septate, guttulate, hyaline, and penicillate (branched 2 or 3 times) (Neill and Hyde 1939, Gray 1942, Griffiths 1959b). Microconidia are first formed by a constriction below the apex of the microconidiophore. The rest bud off in succession inside a tube formed by the terminal portion of the microconidiophore (Prillieux and Delacroix 1892b; Neill and Hyde 1939; Gray 1942; Wilson et al. 1945, 1954; Griffiths 1959b).
Microconidia are unicellular, uninucleate, ovoid, guttulate or biguttulate, hyaline, 1.8-3.0 × 2.3-6.0 µm (Gray 1942, Calvert and Muskett 1945, Griffiths 1959b). In microconidial germination, a terminal germ tube forms; or if a transverse septum forms, a terminal or lateral germ tube will be produced (Griffiths 1959b).
Macroconidiophores and macroconidia. Macroconidiophores are short barrel-shaped cells, 2-3 µm wide and 5-15 µm long, that arise laterally on the hyphae (Neill and Hyde 1939, Griffiths 1959b). Macroconidia are budded from the apex of the macroconidiophores (Griffiths 1959b) (figure 1) and arrange in clusters perpendicular to the hypha (Calvert and Muskett 1945, Wilson et al. 1945). Up to 30 macroconidia develop per conidiophore (Wilson et al. 1945).
Macroconidia are smooth, unicellular, uninucleate, hyaline, cylindrical to slightly cresentric with rounded ends, and usually biguttulate (figure 2) (Gray 1942; Calvert and Muskett 1945; Wilson et al. 1945, 1954; Spooner 1987). They are 2.5-6 × 11-21 µm in size. The vegetative nucleus is 3-5 × 2 µm and the nucleolus may be as large as 2 µm (Griffiths 1959b).
On the surface of the caryopsis, macroconidia are embedded in a pinkish, slimy mass (Spooner 1987) that dries to form a hard reddish-brown crust (Calvert and Muskett 1945, Hyde 1945) (figure 3, figure 4, figure 5). When germinating, macroconidia swell and produce one or two germ tubes (Griffiths 1959b).
Apothecia. Apothecia are small, fleshy, and cup-shaped. One to 7 (usually 1 to 3) apothecia emerge from each infected seed (Prillieux 1897; Gray 1942; Calvert and Muskett 1945; Wilson et al. 1945, 1954) (figure 6). The stipe is smooth, velutinous under magnification, externally white or gray, internally pinkish brown, enlarging upward (Neill and Hyde 1939), and longitudinally furrowed (Spooner 1987). The stipe varies from 1 to 10 µm in length and from 0.2 to 0.5 µm in diameter (Prillieux and Delacroix 1892b, Rehm 1900, Gray 1942, Calvert and Muskett 1945) and is composed of hyaline, parallel hyphae, 4-6 µm in diameter, occasionally intertwining and seldom branched (Gray 1942, Calvert and Muskett 1945).
Apothecia emerge from the caryopsis and elongate (figure 7). The disk of the apothecium is at first closed (Gray 1942) but opens to cup-shaped and with age becomes saucer-shaped and then flat (Gray 1942, Calvert and Muskett 1945, Spooner 1987) (figure 8, figure 9). The disc diameter ranges from 1.0 to 7.0 µm (Prillieux and Delacroix 1892b, Rehm 1900, Neill and Hyde 1939, Gray 1942, Calvert and Muskett 1945). The disk color changes from light pinkish brown to deep brown (Calvert and Muskett 1945), orange brown (Spooner 1987), or pale pinkish cinnamon, darkening to cinnamon when old (Neill and Hyde 1939, Gray 1942). The margin is smooth and entire (Neill and Hyde 1939, Gray 1942, Calvert and Muskett 1945, Spooner 1987) and is radially wrinkled around the stipe apex (Spooner 1987).
Hymenium. The hymenium is 100-140 µm deep (Williams and Spooner 1991). The subhymenium consists of intricately intertwined and coiled hyphae 2.5-3 µm in diameter. The subhymenium blends into the medullary excipulum, a 22-27 µm deep layer composed of fine, densely intertwining hyphae 2-5 µm broad (Neill and Hyde 1939, Gray 1942, Williams and Spooner 1991). The outermost layer (the ectal excipulum) is 35-40 µm thick and is composed of parallel to somewhat interwoven hyphae 3.5-4.5 µm in diameter (Williams and Spooner 1991) (figure 10).
Asci. The asci are cylindrical and clavate, with 8 spores obliquely placed in a single row (uniseriate) in the upper two-thirds of the ascus (Neill and Hyde 1939, Gray 1942, Calvert and Muskett 1945, Spooner 1987) (figure 11). Ascus size is variable but falls within the range of 66-120 µm long × 3-8 µm wide. The ascus base tapers to about 2-5 µm (Spooner 1987, Williams and Spooner 1991). The apical cap is 1-3 µm thick (Alderman 1997), and the apical plug does not stain blue with iodine (Prillieux and Delacroix 1892b, Neill and Hyde 1939, Gray 1942, Calvert and Muskett 1945, Wilson et al. 1954, Spooner 1987).
Ascospores. Ascospores are hyaline, smooth, elliptical, fusoid to broadly fusoid, and usually biguttulate (Neill and Hyde 1939, Gray 1942, Calvert and Muskett 1945). One side is often flattened, or curved, continuous, or rarely developing a central septum (Spooner 1987, Williams and Spooner 1991). Ascospore size is variable, 7-14 × 2.5-4.5 µm. Germinating ascospores swell to about 10 × 5 µm (Neill and Hyde 1939) (figure 12). The first germ tube is terminal, followed by a second that is frequently lateral in position and usually constricted at the point of origin. They normally develop a central septum and two polar hyphae, but often lack a septum and have a single polar or lateral hypha (Neill and Hyde 1939, Calvert and Muskett 1945)
Paraphyses. Paraphyses are fusiform, hyaline, nonseptate (Neill and Hyde 1939, Gray 1942, Calvert and Muskett 1945) or sparsely septate (Spooner 1987) and 1.5-4 µm wide (Neill and Hyde 1939, Gray 1942). Spooner (1987) described the paraphyses as enlarging at the apex to 2.5-3.0 µm, but others (Neill and Hyde 1939, Gray 1942, Calvert and Muskett 1945) reported that the apex was not swollen. Paraphyses are as long as or slightly longer than the asci (figure 13).
On a nutrient medium such as potato dextrose agar, G. temulenta grows slowly and produces a partly submerged, branching, hyaline, septate mycelium (Neill and Hyde 1939, Calvert and Muskett 1945). Sporulation and slime production occur after 7 days (Calvert and Muskett 1945, Wilson et al. 1945, Hair 1952) and in culture appears reddish brown (Neill and Hyde 1939) or chocolate brown (Wilson et al. 1945). The addition of 1-percent peptone to PDA or malt agar increases spore mucilage production (Calvert and Muskett 1945). However, some cultures are predominantly mycelial while others are conidial (Wilson et al. 1945).
In culture, macroconidia are produced from short conidiophores formed at intervals perpendicular to the hypha (Calvert and Muskett 1945, Wilson et al. 1945). Conidia from culture may be larger (Wilson et al. 1945) or appear less regular than those from seed (Calvert and Muskett 1945). Growth is slow at 5 °C, optimal at about 20 °C, less at 27 °C, and restricted at 30 °C (Neill and Hyde 1939, Alderman 1992). Radial growth slows with decreasing water potential through -9.0 to -1.0 MPa (Alderman 1992).
Sporodochia develop in culture at 5 °C to room temperature after about 1-3 months (Calvert and Muskett 1945). Growth characteristics on various media were described by Neill and Hyde (1939) and Calvert and Muskett (1945).
Calvert and Muskett (1945) collected other discomycetes associated with ryegrass and detritus that are similar to G. temulenta but differ in morphology in culture and do not produce spores. Unfortunately, neither species identification nor technical descriptions of these other fungi were recorded.
Neill and Hyde (1939) found a fungus on Lolium that is similar to G. temulenta. They defined it as Lolium fungus number 2. Unfortunately, the taxonomic description and species identity of this fungus was not established either.
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Original posting: October 2001.