Complexipes moniliformis C. Walker

Field-collected spores in PVLG
Spore from PGA culture in PVLG
Field-collected spore (SEM)

SPORES occur singly in the soil or form in agar cultures; develop terminally, rarely laterally from the tip of a single subtending hypha; spores from the field are greyish orange (5B6) to burn Sienna (7D8); globose to subglobose; (20-)75-125(-140) µm diam.

 

 

SUBCELLULAR STRUCTURE OF SPORES consists of one wall with two layers (swl1 and 2).

 

Field-collected spores in PVLG
Field-collected spores (SEM)
Layer 1 permanent, hyaline to greyish orange (5B6) to golden yellow (5B8), 1-2.5 µm thick, ornamented with warts, globular, finger- or cup-shaped processes, 1-5 µm high.
Layer 2 laminate, greyish orange (5B6) to burn Sienna (7D8), 3.5-10.0 µm thick.

Layers 1 and 2 do not stain in Melzer’s reagent.

Most juvenile spores with spore wall layer 1 only.

From field-collected spore

SUBTENDING HYPHA cylindrical, flared or constricted, straight or recurved, light yellow (4A4) to yolk yellow (4B8), ornamented with small warts or spines, divided into 3-8 septa, with constrictions at them; the distance between septa ranges from 5-22.5 µm. The subtending hypha is connected with hyaline to pale yellow (3A3), 2.5-8.8 µm wide hyphae, ornamented with hyaline to pale yellow (3A3) blisters.

 

 

Spores and hyphae from PGA agar culture in PVLG
On potato glucose agar medium, C. moniliformis formed slow-growing, caramel (6C6) to rust brown (6E8), low, downy cultures with an irregular margin. After 35 days, these cultures reached 60-70 mm diam and started to form spores. The hyphae from the agar medium were hyaline to pale yellow (3A3), verrucose, 5-7.5 µm diam, divided by septa; the distance between the septa ranged from 22.5 to 50 µm. The spores were hyaline to pale yellow (3A3), globose to subglobose, 30-70 µm diam, and had one 2-layered wall, 1-5 µm thick. This wall was ornamented with warts, spines, finger- or cup-shaped processes, up to 10 µm high. The subtending hypha was cylindrical, flared or constricted, hyaline to pale yellow (3A3), 10-20 µm wide at the spore base and 20-130 µm long, with 2-11 septa.

MYCORRHIZAE. Generally, fungi of the genus Complexipes are considered to occur in associations with ectendomycorrhizae (Laiho 1965; Mikola 1965; Thomas and Jackson 1982; Wilcox et al. 1974), although Wilcox et al. (1974) insisted that some of them produce ectendomycorrhizae with a given pine while others form ectomycorrhizae in the same pine. The species of pine is also important whether the association will be ecto- or ectendomycorrhizal.

The function of the fungi in mycorrhizal associations is poorly known. Pachlewski (1983) isolated a fungus called MrgX from ectendomycorrhizae of Pinus sylvestris L., which seems to be of the genus Complexipes. In his laboratory and field studies, the MrgX isolates were highly active mycorrhizal fungi forming a biotic barrier for ectomycorrhizal symbionts of pine. Danielson et al. (1984a-d) also classified fungi of Complexipes to the most effective mycorrhizal fungi.

DISTRIBUTION. The fungi of the genus Complexipes are widely distributed on coniferous and deciduous hosts in tree nurseries, sand dunes, burned sites, and disturbed areas in Canada (Danielson 1991; Danielson et al. 1984a-d), the United States (Laiho 1965; Wilcox et al. 1974, 1983; Yang and Wilcox 1984), the United Kingdom (Thomas et al. 1983), Finland (Mikola 1965), Poland (Blaszkowski 1989), Kenya (Ivory and Pearce 1991), and New Zealand and Australia (Mosse and Bowen 1968).

REFERENCES

Blaszkowski J. 1989. The occurrence and geographic distribution of E-strain ectendomycorrhizal fungi in Poland. Bull. Pol. Ac. Sci. Biol. Sci. 37, 19-31.

Danielson R. M. 1991. Temporal changes and effects of amendments on the occurrence of sheathing (ecto-) mycorrhizas of conifers growing in oil sands tailings and coal spoil. Agric. Ecosyst. Environ. 35, 261-281.

Danielson R. M., Griffiths C. L., Parkinson D. 1984a. Effects of fertilization on the growth and mycorrhizal development of container grown pine seedlings. Forest Sci. 30, 828-835.

Danielson R. M., Visser S., Parkinson D. 1984b. Microbial activity and mycorrhizal potential of four overburden types used in the reclamation of extracted oil sands. Can. J. Soil Sci. 63, 363-375.

Danielson R. M., Visser S., Parkinson D. 1984c. Production of ectomycorrhizae on container-grown jack pine seedlings. J. For. Res. 14, 33-36.

Danielson R. M., Zak J. C., Parkinson D. 1984d. Mycorrhizal inoculum in a peat deposit formed under a white spruce stand in Alberta. Can. J. Bot. 63, 2557-2560.

Ivory M. H., Pearce R. B. 1991. Wilcoxina mikolae newly identified as a mycorrhizal fungus on pines in Africa. Mycol. Res. 95, 250-253.

Laiho O. 1965. Further studies on the ectendotrophic mycorrhiza. Acta For. Fenn. 79, 1-34.

Mikola P. 1965. Studies on the ectendotrophic mycorrhiza of pine. Acta For. Fenn. 75, 1-56.

Mosse B., Bowen G. D. 1968. A key to the recognition of some Endogone spore types. Trans. Br. Mycol. Soc. 51, 469-483.

Pachlewski R. 1983. Grzyby symbiotyczne i mikoryzy sosny (Pinus silvestris L.). Prace IBL 615, 3-123.

Thomas G. W., Jackson R. M. 1982. Scanning electron microscopy of sheathing mycorrhizas of Sitka spruce. Trans. Br. Mycol. Soc. 79, 31-39.

Thomas G. W., Rogers D., Jackson R. M. 1983. Changes in the mycorrhizal status of Sitka spruce following outplanting. Plant Soil 71, 319-323.

Wilcox H. E., Ganmore-Neumann R., Wang C. J. K. 1974. Characteristics of two fungi producing ectendomycorrhizae in Pinus resinosa. Can. J. Bot. 52, 2279-2282.

Wilcox, H. E., Yang C. S., LoBuglio K. 1983. Responses of pine to E-strain ectendomycorrhizal fungi. Plant and Soil 71, 239-247.

Yang C. S., Wilcox H. E. 1984. An E-strain ectendomycorrhiza form by a new species, Tricharina mikolae. Mycologia 76, 675-684.