66 The genus Chrysosporium, its physiology and biotechnological potential Rajendra Kumar Singh Kushwaha Department of Botany, Christ Church College, Kanpur, India Summary Key words The genus Chrysosporium is reviewed including its 11 species without intercalary conidia, 29 species with intercalary conidia, three species with uncertain position and four undescribed species with their report of isolation, physiology, antagonistic, keratinolytic and pathogenic potentials. Research revealed considerable biotechnological potential for recycling keratinous waste in soil and secretion of enzymes and antimicrobials. Its taxonomic relation with dermatophytes and their relatives is of immense importance. Germplasm collection of Chrysosporium and its teleomorphic connections is aplaused. Chrysosporium, Biotechnological Potential, Review The genus Chrysosporium was introduced by Corda [1]. Saccordo [2] placed it in synonymy with Sporotrichum. Later this genus was reviewed [3-5]. Twenty two species of Chrysosporium were recognised [5]. Since then several species have been added to this genus. Large number of workers have isolated Chrysosporium species from many different habitats around the world. Until recently, interest was restricted only to reports of its occurrence but as research into this fungus has gained momentum it has been evaluated as a potential fungus. Recently the research carried out on Chrysosporium has increased markedly and information has begun to appear. While isolating Chrysosporium by hair baiting and other methods its potential to degrade keratin was particularly emphasized. In view of this, its activity in soil and water sediments of polluted and fresh water sites is also receiving attention. Perhaps some species of Chrysosporium may be utilized for recycling of keratinous waste in soil and as water pollution indicators which certainly pave the way towards a congenial environment. Secretion of some of their metabolites, particularly enzymes and antimicrobials, is gaining the attention of pharmaceutical industries. The resemblance of Chrysosporium to dermatophytes, and their pathogenic potential, is directly related to health of human beings and animals. In addition to the keratinous substrates, this genus is now being found associated with other non keratinous substrate too. This potential, and its similarities to Myceliophthora, Emmonsia, Zymonema, Geomyces, Corresponding address: Dr. Rajendra K.S. Kushwaha Department of Botany, Christ Church College, Kanpur 208 001, India Tel: +91 512 637 318; Fax +91 512 311 627 ©2000 Revista Iberoamericana de Micología Apdo. 699, E-48080 Bilbao (Spain) Trichosporiella, Malbranchea, Ovadendron, Botryotrichum, Sepedonium, Mycogone, Sporotrichum and others and associated teleomorphs, is of immense diagnostic importance. Long term studies [6-11] on the biology of Chrysosporium, and other scattered reports, revealed that its wide distribution is due to its antagonistic potential and ability to produce enzymes and other extracellular metabolites [12-14]. Looking into all the above potential of Chrysosporium, it was intended to review all possible available work on this genus, which has not been reviewed before, in spite of the fact that the amount of new information has grown enormously in recent times. Forty - seven species of Chrysosporium are listed here along with their report of isolation including some salient features. Species with intercalary conidia White colony 1.Chrysosporium anamorph of Rollandina vriesii Apinis Trans. Brit. Mycol. Soc. 55:501, 1970. 25-30 mm on PYE in 14 days, terminal and lateral conidia smooth and thin walled, 1 celled, 3-6x2-3 µm, wide scar, 1-2 µm, keratinolytic. 2.Chrysosporium anamorph of Arthroderma curreyi Berk Outl. Bri. Fungol. 357, 1860. 30-50 mm on SGA in 14 days, terminal and lateral conidia mostly sessile, smooth or slightly echinulate, thin, 1 celled, rarely 2 celled, 3-6x2-3 µm, wide scar, 1-2 µm, keratinolytic [15-19]. 3.Chrysosporium anamorph of Arthroderma cuniculi Dawson Sabouraudia 2:187, 1963. 35-60 mm on SGA in 14 days, terminal and lateral conidia sessile, smooth, thin walled, 1 celled or 2-3 celled, mostly 3-8x2-3 µm, scar 1-2 µm, keratinolytic [16,17,20]. 4.Chrysosporium anamorph of Pectinotrichum llanense Varsavsky & Orr Mycopath. Mycol. Appl. 43:231, 1971. 5-15 mm on Czapeck agar in 14 days, terminal and lateral smooth and thin walled, 1 celled, 4-6.5x2-3 µm, scar 1-1.5 µm, keratinolytic [21]. Chrysosporium and its potential Kushwaha RKS 5.C. synchronum van Oorschot Stud. Mycol. 20:42, 1980. 80 mm on PYE in 7 days, terminal and lateral conidia thin walled, smooth or echinulate, 1 celled, 7.5-11x4-5.5 µm, scar 0.5-1 µm, not keratinolytic. Other than white 6.C. sulfureum (Fiedl.) van Oorschot & Samson Stud. Mycol.20:28, 1980. 10-15 mm on PYE in 14 days, pale creamy yellow, terminal and lateral mostly sessile, 2–8 per conidiogenous, smooth and thin becoming thick walled, some times echinulate, 1 celled, 3-8x3-6 µm, wide scar, 1.4 µm, not keratinolytic, shows preference for fatty or calcium rich material [22-24]. 7.C. georgii (Varsavsky & Ajello) van Oorschot Stud. Mycol.20:31, 1980. 10-30 mm on 2% malt agar in 14 days, white or pale buff or pink, terminal and lateral smooth and thin walled, 1 celled, rarely 2-3 celled, 3-8x2-3 µm, wide scar, 0.5-4.5 µm, keratinolytic [25,26]. 8.C. lucknowense Garg Mycopath. Mycol. Appl. 30: 224, 1966. 55 mm on SGA in 14 days, cream, terminal and lateral 1-4 conidia on one hyphal cell in close proximity, thin, 1 celled, 2.5-11x1.5-6 µ m, keratinolytic [22,23,26,27]. 9.C. filiforme Sigler, Carmichael & Whitney Mycotaxon 14:261, 1982. 40 mm in 21 days on PYE, white to buff, terminal or lateral conidia 0-1 or rarely 2 septate, sessile or on short pedicell, smooth, filiform upto 40 µm long, not keratinolytic. 10. C. mephiticum Sigler Can. J. Bot. 64: 1212, 1986. 50-62 mm on PYE and CER in 25 days, creamy white, terminal and lateral smooth, mostly sessile, in close proximity, 2.5-3.5x2.5-3 µm, scar 1-1.5, keratinolytic, strong and pungent odour. 11.C. gourii Jain, Deshmukh & Agrawal Mycoses 36:77, 1993. 74 mm on SGA in 18 days, white to cream to yellowish brown, terminal and lateral, smooth or rough, thin walled, 1 celled, 2.5-6x2-4 µm, keratinolytic. Species with intercalary conidia White colony 12.C. queenslandicum Apinis & Rees Trans. Brit. Mycol. Soc. 67:524,1976. 55-65 mm on PYE in 14 days, white, intercalary conidia several, smooth and slightly thick, 1 celled, 3.59.5x3.6 µm, broad basal scar, keratinolytic [17,21-23,25-27]. 13.Chrysosporium anamorph of Gymnoascus demonbreunii Ajello & Cheng Mycologia 59:692,1976. 25-33 mm in 14 days on hay infusion agar, white, intercalary conidia most abundant, smooth and thin, 5-11x4.5-6 µm, conidia always terminal, never lateral, smooth, thin walled, 1 celled, 6-9x5-6 µm wide basal scar, 1.5-2.5 µm, keratinolytic [21]. 14.C. carmichaelii van Oorschot Stud. Mycol. 20:15,1980. 15-30 mm in 14 days, white, intercalary less abundant, smooth, thin, 3-6x1.5-3 µm, terminal and lateral smooth or rarely rough, 1 or rarely 2 celled, 3-6x3-3.5 µm, not keratinolytic [17,18,22,23,27,28]. 15.C. tropicum Carmichael Can. J. Bot. 40: 1170, 1962. 50-60 mm in 40 days, white, intercalary not com- 67 mon, smooth and slightly thick, 3-4x6-10.5 µm, terminal and lateral smooth or slightly thick, 1 or rarely two celled, 3.5-7.5x3-4.5 µm, basal scar 1.5-2 µm, keratinolytic [1618,21-23,25-33,36-53]. 16.C. xerophilum Pitt Trans. Brit. Mycol. Soc. 49: 468, 1966. 85 mm on cherry deccoction agar in 7 days, white, intercalary conidia not always abundant, sometimes in chains of 2-6, smooth and thin or thick walled, 7.5-12x34.5 µm, terminal and lateral, smooth, thin or thick walled, 4-13x3-10 µm, wide basal scar, 1.5 µm, weak keratinolytic, osmophilic [21]. 17.C. pannicola (Corda) van Oorschot and Stalpers Stud. Mycol. 20:43,1980. 20-38 mm on PYE in 14 days, white, intercalary less abundant, smooth or echinulate, thick walled, 4-8x2-3 µm, keratinolytic [15,17,18,20,22,23,26,27,34,54,55]. 18.C. indicum (Randhawa & Sandhu) Garg Sabouraudia 4:262,1966. 40-50 mm on PYE in 14 days, white, intercalary conidia less abundant, smooth or slightly echinulate, thin walled, 6-12x2-3.5 µ m, keratinolytic [15-23,25-32,40, 44,45,48,54, 56]. 19.C. sinense Liang Acta Mycologica Sinica 10:50,1991. 30 mm in 20 days at 18°C on SDA, intercalary conidia abundant, 1.2-3.7x3.7-10 µm, terminal and lateral, 2.4-3.8x4.0-5.7 µm, development of synnemata, isolated from endosclerotium of Cordyceps sinensis [Berk.] Sacc. 20.C. geophilum Kushwaha & Shrivastava Curr. Sci. 58:970,1989. 60-70 mm on SGA, white, reverse pale creamy brown, intercalary conidia less, rough or smooth walled, 2-4 µm, lateral conidia sessile or on short protrusions, initially echinulate becoming smooth on transfers, thick, 1 or rarely 2 celled, 4-20x2-4 µm, keratinolytic. 21.C. botryoides Skou Mycotaxon 43:237,1992. White colony, ramose on MGYA, conidia 1 celled, thick walled, occur so close together that they look like bunches of grapes, globose to pyriform, up to 10.2 µm, intercalary conidia sparsely present, rounded in agar, single or a few in chains, not keratinolytic, osmophilic. 22.C. globiferum Skou Mycotaxon 43:237,1992. White, dense with slightly ramose margin on MGYA, conidia 1 celled, thick walled, globose to pyriform, upto 11.1 µm, intercalary conidia abundant, rounded in agar, not keratinolytic, osmophilic. 23.C. hispanicum Skou Mycotaxon 43:237,1992. White, dense on MGYA, conidia 1 celled, thick walled, globose to pyriform, upto 13.4 µm, in agar only terminal conidia, intercalary conidia very sparcely in agar, not keratinolytic, osmophilic. 24.C. holmii Skou Mycotaxon 43:237,1992. White, intercalary conidia turncate never in pronounced amount, not in agar, terminal globose conidia, pyriform up to 12.8 µm, not keratinolytic, osmophilic. 25.C. medium Skou Mycotaxon 43:237,1992. White, flat on MGYA, conidia 1 celled, globose to pyriform, upto 9.8 µm, intercalary abundant in agar, not keratinolytic, osmophilic. 26.C. minor Skou Mycotaxon 43:237,1992. White, flat, conidia 1 celled, globose to pyriform, up to 8.4 µm, intercalary conidia single and rounded in agar, not keratinolytic, osmophilic. 27.C. pyriformis Skou Mycotaxon 43:237,1992. White, dense, flat on MGYA, conidia 1 celled, thick walled, globose to pyriform up to 11.0 µm, intercalary conidia in agar more or less rounded, not keratinolytic, osmophilic. 68 Other than white 28.C. pseudomerdarium van Oorschot Stud. Mycol. 20:14,1980. 7-20 mm on PYE in 14 days, white, locally pale yellow, intercalary conidia intially smooth and thin walled or becoming echinulate and / or thick walled, 3-6x3-6 µm, terminal and lateral in chains up to 4, initially smooth and thin becoming echinulate, 1 celled, 2-6.5x1.5-5 µm, weak keratinolytic [25,26]. 29.C. merdarium (Link ex Grev.) Carmichael Can. J. Bot.40:1160,1962. 30-35 mm on PYE in 14 days, white becoming bright yellow, pink or green, intercalary conidia few, 512x3-6 µm, terminal and lateral 1 celled, 4-10x3-6 µm, not keratinolytic. G. uncinatum do not develop conidia on PYE while these are abundant on hay infusion agar [15,16,20-24,31,36]. 30.C. keratinophilum D. Frey ex Carmichael Can. J. Bot. 40:1157,1962. 40-50 mm on PYE in 7 days, white to cream, sulphor yellow, intercalary conidia less abundant or rare, smooth or echinate, thick walled, 6-25x3.5-7 µm, terminal and lateral smooth or echinate, thick, 1 celled, 3.5-22x3.511 µ m, keratinolytic [15,17,20-23,25,26,30,32,34-38, 40,41,43-45,47,54-66]. 31.C. inops Carmichael Can. J. Bot. 40:1156,1962. 0.5-10 mm on PYE in 14 days, cream, conidia terminal or intercalary, smooth and thick walled, 1 celled, 6.5-12x5-9 µm, not keratinolytic. 32.Chrysosporium anamorph of Renispora flavissima Sigler et al Mycotaxon 10:133, 1979. 30-40 mm on PYE in 14 days, pale yellow, buff centre, intercalary conidia rare, initially smooth, thin or thick walled, verrucose, 1 celled, 6-8x5-8 µm, keratinolytic. 33.C. lobatum Scharapov Nov. Syst. niz. Rast. 15:144, 1978. 30-35 mm on PYE in 14 days, white becoming pale green or pale grey, intercalary conidia rare, smooth, thin walled becoming echinulate, thick walled, 1 celled, 34x2-3 µm, terminal and lateral, smooth and thin walled becoming reddish brown to dark brown, echinulate, thick walled 1 celled, 2-4x1.5-3.5 µm, scar 0.5-1.5 µm, keratinolytic. 34.C. vespertilium Guarro, Vidal & de Vroey Mycotaxon 59:189,1996. 34-45 mm on PYE in 14 days, yellow, intercalary conidia rare, smooth and thin walled, 1-3 celled, 5-20x2-5 µm, scar 2.5 µm, keratinolytic, coiled sterile hyphae. 35.C. pilosum Gene, Guarro & Ulfig Mycotaxon 50:107,1994. Restricted, 0.4-0.7 mm on PYE, raised, yellowish white to mustared yellow or light brown, reverse brownish or dark brown, intercalary conidia smooth, thin becoming thick walled and verrucose, 1 celled, 3.5-5.5x3-4 µm, terminal and lateral smooth and thick walled, becoming coarsly verrucose, 1 celled, 4-6x3.5-5.5 µm, scar 1.5-2 µm, poor keratinolytic, broad, simple, thick walled, brownish sterile hyphae present. 36.C. europae Sigler, Guarro & Punsola Can. J. Bot. 64:1212,1986. 50-60 mm on PYE in 35 days, vinaceous buff or brown diffusing pigment, intercalary 4.5 µm, terminal and lateral 8.5x2.5-3.5 µm, keratinolytic. 37.C. zonatum Al - Musallam & Tan Persoonia 14:69,1989. 55 mm on PYE in 14 days, white to buff, intercalary conidia abundant, 5-12x2-4 µm, terminal and lateral thick walled and verrucose at maturity, 1-2 celled, 5-7x35 µm, scar 2-2.5 µm, keratinolytic and cellulolytic. 38.C. vallenarense van Oorschot & Piontelli Persoonia 12:487,1985. Colonies restricted on YpSs at 25°C, white becoming sulphor yellow, conidia terminal, often developing sympodially, rarely intercalary, tuberculate, 3.5-5.5x5-7 µm. Isolated from keratinous substrates. Conidia resembling those of Chrysosporium anamorph of Renispora flavissima. 39.C. siglerae Cano & Guarro Mycotaxon 51:75,1994. 10-15 mm on PYE in 21 days at 28°C, pale yellow, conidia mostly lateral, 1 celled, smooth to slightly verrucose, 5-30x2-3.5 µm, 2 celled, 10-16x2-3 µm, intercalary in series of two or more, 10-15x2-2.5 µm, keratinolytic. 40.C. farinicola (Burnside) Skou Friesia 11:70.1975. 27-45 mm in 14 days on honey agar, initially white or becoming pale greenish yellow or pale brown, intercalary conidia abundant, smooth, thick walled, 3-12x5-12 µm, terminal and lateral smooth, thick walled, 1 celled, 6-15x4.5-9 µm, wide basal scar, 1.5-5 µm, not keratinolytic, osmophilic [22,23]. Species with uncertain position 41.C. parvum [or Emmonsia?] 42.C. crescens [or Emmonsia?] Placed under Emmonsia Cif & Montemartini on the basis of blastic conidia and thick walled chlamydospore- like celles [5,67]. 43.C. racemosus Sharma, Bhattacharjee & Bhadauria Indian Phytopath. 46:404,1993. 1-1.2 cms in 7 days at 28±1°C on PDA and Czapecks dox agar, initially white, later changes to cream, green, dark brown to blackish brown. Conidiophore branched with thick, numerous scars [hilum]. Acropleurogenous and pedicellate aleurospores aggregate to form clusters at intercalry position of hyphae. The described species without camera leucida drawings or photographs does not seems to be Chrysosporium because conidia do not have scars and are round. The cultures were not available. Species in press 44.C. undulatum Vidal, Ulfig & Guarro 45.C. fluviale Vidal, Ulfig & Guarro 46.C. submersum Vidal, Ulfig & Guarro 47.C. minutisporosum Vidal, Ulfig & Guarro Genus Chrysosporium have following teleomorphs also Gymnoascus arxii, G. uncinatus, Nannizziopsis spp., Amauroascopsis perforatus, Aphanoascus durus, Aph. clathratus, Aph. fulvescens, Aph. hispanicus, Aph. reticulisporus, Aph. saturnoideus, Aph. terreus, Aph. verrucosus, Arthroderma multifidum, Arth. tuberculatum, Arth. croccatum, Arth. silverae, Ctenomyces serratus, Ajellomyces dermatitidis, Aj. capsulatus, Apinisia graminicola, Api. queenslandica, Neoxenophila foetida, Renispora flavissima, Amauroascus albicans, Am. aureus, Am. volatilis-patellis, Orromyces spiralis, Phaneochaeta chrysosporoidea, Pseudarachmiotus orissi, Bettsia alvei, Bettsia species, Cordyceps species. Chrysosporium and its potential Kushwaha RKS 69 Temperature Mycelial growth of C. tropicum There are many reports of the ability of Chrysosporium to grow at sub zero and above 37°C. C. keratinophilum, C. tropicum and C. queenslandicum grows at 37°C. C. asperatum, C. pannorum and C. indicum grows in antarctic soil [68,69] and C. evolceanui in alpine soil [70]. Garg et al [71] found 21-50 and 25-30°C optimum for C. pannicola and C. keratinophilum. The growth of C. tropicum was rapid at 27°C and 37°C showed minimum rate of growth [72]. Maximum germination of conidia of C. tropicum took place at 32°C within 24 hours. Low temperature also supported conidial germination [72]. Slight growth of C. pannorum at -6°C [73] and good growth at -5°C and at 5°C was reported [74-75]. Rapid growth of this fungus occurred at 15°C [76], maximum growth was at 25°C [74,75] and its rate of growth reduced at 30-37°C [75,77,78]. Growth of C. tropicum was measured on 12 agar media as follows: oat meal> YpSs> glucose asparagine> potato dextrose> Czapecks dox> SAA> malt extract> tryptone agar> Czapecks dox +yeast extract> peptone dextrose> Sabouraud dextrose> Sabouraud dextrose+ yeast extract. Maximum sporulation was recorded on Sabouraud dextrose agar supplemented with yeast extract followed by tryptone agar [89]. This fungus grows in varying concentrations of glucose while 6% glucose favoured maximum growth. A study of carbon metabolism in C. tropicum was made at 5-15 days of incubation. The rate of assimilation of glucose, sucrose, mannose, maltose and lactose by C. tropicum was studied chromatographically [90]. Growth and sporulation of 15 species of Chrysosporium including 6 strains of C. tropicum were different on 7 media. Six strains could be categorised in 3 groups based on their growth characteristics [91]. Glucose supported maximum mycelial growth of C. tropicum and fructose was assimilated more slowly than mannose among the monosaccharides. Utilization of sucrose, maltose and polysaccharide was average. Mannose and starch showed an increase in growth rate up to 10 days of incubation and then gradually decreased. Mannose and maltose were utilized very rapidly by this fungus and exhausted within 5 days. Glucose, fructose, sucrose, lactose and starch were not completely utilized [90]. C. tropicum synthesized galactose, glucose and fructose [90]. C. tropicum exhibited carbon heterotrophy, as was reported in other soil saprophytes, and meets its requirements from various sources. pH requirements A. uncinatum and A. curreyi are acidophilic and C. tropicum, C. keratinophilum, and A. quadrifidum were found to be alkalophilic [79]. C. pannorum grew well at sea water salinity [77] but 20% sodium chloride inhibited its growth when used in Czapecks medium [80]. A. curreyi survives in mud and sand dunes of coastal soils [81] and C. tropicum and C. indicum in marine soils [82]. For C. tropicum 7 pH was optimum but it grows at 3 and 7 pH also [72]. Garg et al. [71] gave a range of soil pH in relation to the distribution of A. fulvescens, C. keratinophilum, C. pannorum, C. tropicum, C. asperatum, C. evolceanui and C. serratus. Moisture contents Occurrence of A. curreyi was reported in nests with 11.81% and 19.81% water content while C. keratinophilum was isolated from nests with higher moisture content showing a hygrophilic nature [83]. The hygrophilic nature of A. fulvescens and C. keratinophilum was also confirmed [84]. C. pannorum survived in habitats of little or no biotic influence [68]. A high percentage of spore germination in A. uncinatum and Ctenomyces serratus at 90-100 RH was recorded [85]. Minimum aW for growth at 25°C on NaCl of C. pannorum, C. xerophilum and C. fastidium was 0.92, 0.71 and 0.69 respectively. The growth rate of C. fastidium in medium containing glycerol was lower than with glucose and fructose. Pugh and Evans [85] reported higher percentages of spore germination in A. uncinatum and C. serratus at 90-100 % RH. Humus Distribution of Chrysosporium was not affected by humus [86] and this fungus along with C. indicum and C. tropicum occurred in soil made up of disintegrated lava with low organic matter [74]. Nigam and Kushwaha [23] also reported C. carmichaelii, C. evolceanui, C. indicum, C. keratinophilum, C. merdarium, C. pannicola, C. queenslandicum and C. tropicum in house dust, which is very low in organic matter. Katiyar and Kushwaha [88] reported C. keratinophilum, C. tropicum and Chrysosporium spp. with 100 % hair perforation ability from sand of Mediterranean sea which was very poor in organic matter. Nitrogen nutrition and metabolism Analysis of filtrates of C. tropicum at 4 days revealed arginine, asparagine, aspartic acid, hydroxyl proline and threonine, while at 12 days cystine, proline and serine were also detected. Mycelial extract revealed the presence of arginine, Y amino butyric acid, asparagine, cystine, histidine, hydroxy proline and serine.The asparagine utilization by C. tropicum was rapid. Suitability of ammonium nitrate and some other nitrogen sources for the growth of C. tropicum was studied by replacing asparagine. The rate of utilization of asparagine and synthesis of cell bound and cell free aminoacids was studied [92]. C. tropicum grew fairly well in nitrogen provided in the form of nitrate but nitrate from ammonium source supported completely less mycelial growth. Cystine supported maximum mycelial yield. Peptone was found to be the best source for the fungus while tyrosine seems to be poor in this respect. Alanine, arginine, aspartic acid, cystine, glycine, glutamic acid, histidine, leucine, methionine, phenyl alanine, proline, serine, tyrosine, aspartic acid+glutamic acid+arginine and alanine+ asparagine+ histidine+ phenyl alanine+ proline+ cystine and sodium nitrate, ammonium sulphate and peptone were also all tested for the growth of C. tropicum. Vitamin requirements A mixture of biotin, cyanocobalmin, pyridoxin, riboflabin was used with the omission of one vitamin each time and the mycelial weight of C. tropicum was determined. It grew in the medium provided with all the five vitamins whereas it showed a sudden decrease when biotin 70 and cyanocobalmine were omitted individually from the combination, pointing towards their deficiency [93]. Spore germination Germ tubes of conidia of C. tropicum attained an average length in plain agar in 24 hours while in SDA mycelial clumps were developed in 24 hours. One percent glucose and 0.01% peptone induced 100% germination [94]. Maximum germination of conidia of C. tropicum took place at 32 ºC within 24 hours. A low temperature of 12 ºC also supported conidial germination [94]. The keratinized and non-keratinized propagules of C. tropicum and C. keratinophilum showed differences when germinated on different substrates, but were similar in their ability to tolerate temperature exposure by two methods [95]. Glucose was found to be a good carbon source for germination. Fructose, mannose, sucrose, maltose, lactose and starch did not favour germination [94]. It was shown that in C. tropicum with an increase up to 0.2% in the concentration of nitrogen content of sodium nitrate, the percentage of spore germination was also increased upto 60% within 24 hours. Twelve per cent germination was recorded when 0.01% nitrogen from nitrate source was added to the medium. Nitrogen given in the form of ammonium sulphate exhibited an opposite effect. In this case the maximum germination was recorded at 0.01% nitrogen. In organic nitrogen, asparagine could induce germination up to 52%. An equimolar mixture of asparagine, aspartic acid and glutamic acid supported 58% spore germination of C. tropicum [94]. Antibiotic response to growth and spore germination Dermostatin showed the highest inhibition of C. tropicum at its lowest concentration of 200 µ g/ml while aureofungin showed maximum inhibition at 1000 µ g/ml when supplemented with SDA. MICs of aureofungin and dermostatin was 600 and 200 µ g/ml. Maximum inhibition was caused by griseofulvin at 1000 µg/ml [96]. Complete inhibition of spore germination of C. tropicum was caused by aureofungin at a concentration of 500 µg/ml. Aureofungin was found to be effective even at its lower dose as it also inhibits the length of the germ tube. Dermostatin induced the swelling of the spores before the emergence of the germ tube [96]. The culture filtrates of M. fulvum, M. gypseum and T. mentagrophytes inhibited conidial germination of C. tropicum [72]. Biomass of C. tropicum was reduced by prednisolone at its lowest concentration [97]. Sensitivity to plant extracts, volatile substances, soaps, detergents, oils and biocides Plant extracts of garlic, ginger, neem, ocimum, onion, yellow oliander and a mixture of all these inhibited growth of C. tropicum [72]. Mycostatic fumes of some volatile substances such as formic acid, acetic acid, ethyl, methyl, isopropile and butyl alcohols, diethyl ether and chloroform were able to reduce the growth of C. tropicum in liquid culture to more than half [72]. Biomass loss was noticed of some keratinophilic fungi including C. tropicum by ethyl alcohol, isopropyle alcohol, chloroform, carbon tetra chloride, carbon disulphide, acetone and benzene [98]. Opportunistic fungal strains have been found to be implicated in mycotic diseases of man and animals. Influence of some homeopathic drugs was studied to inhibit the hair invasion activity of A. terreus, C. keratinophilum and C. tropicum mechanically and ezymatically. Mezerium, petroleum, ustilago and sepia caused 100% inhibition of hair perforation and no protein could be released in the culture filtrates at their higher doses [99]. Similarily some hair dyes: black diamond, black rose, mehndi and amla were used to inhibit hair perforation. Complete inhibition of hair perforation was at 100 µg/ml for black diamond and black rose, and the other two were comletely inhibitory at 1000 µg/ml for these fungi. Five Indian soaps: kesh nikhar, godrej shikakai, swastik shikakai, vipro shikakai were inhibitory for hair perforation at 1000 µ g/ml; shampoos: Optima, Organics, Pantene, Sunsilk, detergents; Areil, Rin, Surf Excel, Wheel; and agrochemicals: bavistin, thiram, neem, ocimum, urea, rock phosphate, NPK and zinc sulphate were also completely inhibitory for hair perforation and protein release by the above three fungi at their higher doses [100]. Six plant extracts of Lawsonia inermis, Eclipta alba, Nyctanthus arbortristis, Datura stramonium and a mixture of all the extracts were used for testing antifungal activity of C. tropicum [101]. Essential oils of Mentha arvensis, Trachyspermum ammi, Cymbopogan nardus and Eucalyptus citriodora also caused more than 77% inhibition of C. tropicum [102]. Chrysosporium species were inhibited by himax and tree burb [103]. Mycostatic effect of tea, coffee, proteinex, coconut oil, linseed oil and vegetable oil was studied on C. tropicum. Coffee and tea extracts were inhibitory for growth when these were used with Sabouraud dextrose agar while proteinex showed negligible inhibition, mustard and linseed oil were found to be the most inhibitory [104]. C. pannorum was reported to be resistant for higher concentration of organic mercurial preparation [105]. This fungus was also used in the removal of phosphate from sewage [106]. C. keratinophilum was resistant to cadmium concentrations as high as 560 ppm [107]. Keratinolytic potential, enzymes and secondary metabolites C. pannicola, C. keratinophilum and C. tropicum took as little as 5 days to colonize human hair [108]. C. carmichaelii, C. evolceanui and C. indicum were found to be late colonizers of hair. A perforating group of C. keratinophilum, C. pannicola, C. queenslandicum and C. tropicum and a non - perforating group of C. carmichaelii, C. evolceanui and C. indicum were recognised. C. indicum, C. keratinophilum and 2 Chrysosporium spp. isolated from sand of a Mediterranean beach were able to perforate human hair in 18 days and completely digested it [88]. Five strains of C. tropicum caused 100 % hair perforation and 12 strains decolourised the hair, out of 12 C. indicum 5 caused 100% perforation and all 11 decolourised hair, of 10 C. queenslandicum 6 perforated hair and all decolourised hair, of 5 C. keratinophilum 3 perforated hair and all decolourised hair, of 3 C. pannicola all perforated hair and decolourised hair. All these strains when grown on dermatophyte test medium develop zones ranging from 3-17 mm [109]. Fifteen strains of C. tropicum took 7-40 days for colonization and perforation of human hair in soil [110]. In an another study C. carmichaelii, C. evolceanui, C. farinicola, C. indicum, C. keratinophilum, C. lucknowense, C. pannicola, C. queenslandicum and C. tropicum Chrysosporium and its potential Kushwaha RKS colonized human hair in vitro in 3-9 days [111]. Among the eight species of Chrysosporium, C. keratinophilum was able to perforate and degrade buffalo, cow, dog, goat, horse and human hairs rapidly. Infected hair showed undulation, lifting and disruption of cuticle, narrow and broad perforating organs, projection of medulla and decolouration of hair as induced by this fungus [111]. English [112] observed cuticle lifting, erosion of cortex and perforating organs in hair penetrated by C. keratinophilum. Seven types of perforators developed by A. terreus, C. keratinophilum and C. tropicum were also observed [88,113-116]. The manner in which Chrysosporium species attacked hair was intermediate between dermatophytes and keratinophilic fungi [117]. C. keratinophilum, C. tropicum, C. indicum and A. fulvescens were reported as keratin colonizers [118132]. Most active keratinolysis was shown by A. terrreus, A. fulvescens, C. pannicola, C. queenslandicum, C. xerophilum, C. tropicum, C. keratinophilum, Chrysosporium anamorph of A. cuniculi and Chrysosporium anamorph of A. curreyi [133]. The penetration of hair in vitro by C. tropicum was similar to dermatophytes showing the ability to infect [134]. The ability of A. fulvescens and A. verrucosus to penetrate hair, was found to be different from A. keratinophilus [135]. Determination of this, and keratinolytic ability was demonstrated by a new method by inoculating the fungus directly on hair tied in a tube [136]. A. terreus, C. serratus, C. tropicum and C. keratinophilum isolated from museum soil were found to deteriorate feathers and produce 98, 44, 96 and 76 ku/ml keratinase and release 680, 789, 830 and 650 µg/ml net protein [137]. C. crassitunicatum and C. tropicum released 690 and 788 µg/ml protein and produced 88 and 33 ku/ml keratinase from hen feathers [138]. C. crassitunicatum, C. tropicum and C. indicum produced 130.2, 77.2 and 75 ku/ml keratinase when pig hairs were used [139] and role of C. carmichaelii, C. evolceanui, C. indicum and C. tropicum in keratin degradation [140] and characterization of extracellular proteolytic enzyme of C. tropicum and its role in keratin degradation was also monitored [141]. C. tropicum degraded buffalo horn, woman hair and wool [142]. Amylase production by C. tropicum was reported [143]. C. merdarium, C. keratinophilum, C. indicum, C. crassitunicatum, Chrysosporium anamorph of Pectinotrichum llanense and Chrysosporium anamorph of A. cunicuili were found to degrade chicken feathers [144]. Liquification of gelatin is a criterion for identification of the filamentous phase of Chrysosporium, Blastomyces and Histoplasma [145]. C. indicum is able to digest gelatin [146]. C. carmichaelii, C. evolceanui, C. indicum, C. keratinophilum and C. merdarium, two strains of C. queenslandicum, four strains of C. tropicum also liquified gelatin [147]. C. tropicum, C. zonatum and Chrysosporium anamorph of A. curreyi utilizes standard lipids and fatty acids [cholesterol, palmitic and linolytic acids] and evidence is available for the uptake and degradation of cholesterol by C. keratinophilum. Nineteen enzymes were produced by 390 strains of Chrysosporium [148]. C. tropicum- amylase, urease, pectinase, keratinase, esterase lipase, leucine aryl amidase, cystine arylamidase, alpha galactosidase, alpha glucosidase, beta glucosidase, N acetyl glucosaminidase, alpha mannosidase. C. merdarium- amylase, keratinase, urease, esterase lipase, leucine arylamidase, alpha galactosidase, beta galactosidase, alpha glucosidase, N acetyl glucosaminidase. C. indicum- amylase, cellulase, pectinase, keratinase, leucine arylamidase, cystine arylamidase, alpha galactosidase, beta glucosidase, N acetyl-glucosaminidase, alpha mannosidase, C. keratinophilum- amylase, cellu- 71 lase, urease, pectinase, keratinase, esterase, esterase lipase, lipase, leucine arylamidase, chymotrypsin, alpha galactosidase, beta glucuronidase, alpha glucosidase, beta glucosidase, N acetyl glucosaminidase, alpha fucosidase. C. queenslandicum- amylase, urease, pectinase, keratinase, esterase, lipase, leucine arylamidase, cystine aryla midase, alpha galactosidase, alpha glucosidase, N acetyl glucosaminidase. Chrysosporium anamorph of A. curreyiamylase, keratinase, esterase lipase, lipase, leucine arylamidase, cystine arylamidase, trypsin, chymotrypsin, alpha galactosidase, beta glucuronidase, alpha glucosidase, N acetyl glucosaminidase. C. carmichaelii- amylase, urease, keratinase, esterase lipase, lipase, leucine arylamidase, cystine arylamidase, alpha galactosidase, alpha glucosidase, beta glucosidase, N acetyl glucosaminidase. C. georgii- amylase, urease, pectinase, keratinase, esterase lipase, lipase, leucine arylamidase, cystine arylamidase, alpha galactosidase, beta glucosidase, N acetyl glucosaminidase. Production of anthroquinines [questin and questinol] and asteric acid by C. merdarium was reported [149]. Secondary metabolites produced by keratinophilic fungi were discussed [140]. Elastase was produced by C. evolceanui and C. indicum [150]. Phospholipids and acetone soluble lipids were detected in cold mycelial extracts of C. tropicum [151]. C. indicum produces L-Arginyl- D allothreonil- L-phenylalanine wich is antifungal [152]. C. pannorum produces cryscandin, antibacterial and anticandida and pinnorin-3-hydroxy-3-methylglutaryl co enzyme, a reductase inhibitor [153,154]. Antagonistic potential In vitro studies revealed that C. tropicum was inhibited by staling substances of C. evolceanui, C. indicum, C. lucknowense, Aspergillus flavus, A. niger, Chaetomium globosum, Cladospora species, three species of Penicillium, T. vanbreuseghemii and M. gypseum. The ability of C. tropicum to interact with 12 keratinophilic and saprophytic fungi was evaluated in dual cultures. It was overgrown by A. niger and C. globosum but other fungi showed inhibition when in opposition to it. Eight fungi were able to cause hyphal inhibition of C. tropicum at a distance. Frequent curling, penetration, granulation, lysis and chlamydospore formation in C. tropicum were observed during hyphal interference. C. tropicum penetrated the mycelium of A. niger [12]. Antagonistic potential of C. tropicum against C. indicum, one Penicillium species and M. gypseum is indicative of production of some inhibitory substances [12]. The growth of C. indicum was promoted by nine keratinophilic fungi [155]. With C. tropicum it exhibited maximum positive antagonistic potentiality while with A. benhamiae - strain it showed least promotion in growth. With both of these fungi C. indicum produced intermingling colonies, the score being 1 and no zone of inhibition. C. queenslandicum inhibited this to the maximum extent thus exhibiting a maximum negative antagonistic nature. The inhibiting species developed a zone of inhibition exceeding 2 mm. This intermingled freely with 10 fungal partners while with 3 isolates it produced a zone of inhibition less than 2 mm, the score being 2 as described for soil fungi [156]. C. queenslandicum showed the greatest capacity to interact with as many as 14 fungi, which it exhibited a promoting nature. Maximum antagonistic potential of C. queenslandicum to promote was observed against Chrysosporium species with mutual inhibition and it was assigned a score of 2. This was more inhibited when grown against Chrysosporium, an ana- 72 morph of R. vriesii, with the score being 3, Malbranchea species could intermingle freely with it. Its higher index of dominance showed its high cometitive nature. The study of dual culture interaction showed that out of 17 isolates interacted, two C. queenslandicum showed a similar antagonistic potential exhibiting a maximum antagonistic nature against other fungi. The overall sequence of antagonistic potential can be summarized as C. queenslandicum 264 and 265> Malbranchea sp.> Chrysosporium sp. 234, Chrysosporium anamorph of R. vriesii 208> Chrysosporium sp. 215, C. indicum, C. indicum 201, 221, C. tropicum 263, 449> C. indicum 238> Chrysosporium sp. 442> Chrysosporium sp. 267> A. benhamiae +> A. benhamiae -> A. ciferrii [155]. The antifungal potential of C. carmichaelii, C. queenslandicum and six strains of C. tropicum was studied against Acrophialophora fusispora, Alternaria alternata, A. tenvissima, Aspergillus flavus, A. niger, Aureobasidium sp., Botrytis sp., Cladosporium sp., Cunninghamella sp., Fusarium sp., Harposporium sp., Mucor sp., Penicillium citrinum, Rhizopus nigricans, Torula sp., white sterile mycelium and three unidentified fungi. One strain of C. tropicum allowed minimum number of fungi to grow. Staling substances of other Chrysosporium spp. also caused inhibition of soil fungi. Metabolites secreted by C. evolceanui, C. pannicola, C. queenslandicum and one strain of C. tropicum showed strong antifungal activity against A. niger [157]. Thirty Chrysosporium spp. caused inhibition of the radial extension of M. gypseum, ranging from 20-100%. Twenty one showed more than 50% inhibition of M. gypseum, 15 caused more than 50% inhibition of A. niger, three strains completely inhibited A. niger, six caused more than 50 % inhibition of P. citrinum while four completely inhibited this fungus. Neither of the C. evolceanui were inhibitory. Out of eight strains of C. indicum three inhibited M. gypseum. C. keratinophilum was less than 50% inhibitory. C. lobatum inhibited M. gypseum. One strain of C. tropicum was 100% inhibitory for M. gypseum and another for P. citrinum while other showed less than 50% inhibition of these fungi [155]. Interaction experiments on Chrysosporium anamorph of Arthroderma were also carried out [158]. The volatiles emnated from three strains of C. tropicum were inhibitory for T. mentagrophytes. Inhibition of T. rubrum was caused by volatiles of C. indicum, C. lobatum, and two strains of C. tropicum. The effect of fungal staling substances of eight Chrysosporium spp. on soil mycoflora was studied [157] and it was found that among four strains of C. tropicum one allowed a minimum number of fungi to grow. Staling substances of other Chrysosporium spp. also caused inhibition of soil fungi. Metabolites secreted by C. evolceanui, C. pannicola, C. queenslandicum showed strong antifungal activity against A. niger [157]. The effect of staling products of keratinophilic and non-keratinophilic fungi on the growth and spore germination of C. tropicum was studied [159]. The inhibition /promotion in dual cultures depends on many factors such as staling products of interacting colonies, pH change, nutrient media depletion or alteration of nutritional ingredients besides hyphal interference. Colony interaction in a number of cases is represented by mutual inhibition in growth of both fungal partners. The relative difference reveals the measure of susceptibility and antagonistic ability of the fungus. The development and subsequent formation of chlamydospores, lysis, coiling, deformation, granulation and swelling in dual cultures in most cases are evidence of their successful competition [14]. Combined effect of Chrysosporium with other keratinophilic fungi on hair and feather decomposition Combination of C. keratinophilum with C. queenslandicum, C. tropicum and M. gypseum enhanced feather decomposition. The synergistic action of C. keratinophilum and M. gypseum was most effective as it caused highest protein release in the course of feather decomposition in vitro. The course of wool degradation by C. keratinophilum acting singly and in combination with C. carmichaelii, C. tropicum and M. gypseum was studied by measuring protein released in the culture medium and weight loss of wool up to four weeks. The combined effect of these fungi on the continual breakdown of wool by C. keratinophilum and the effect of the addition of C. keratinophilum on the continual degradation of wool by C. carmichaelii, C. tropicum and M. gypseum were also studied. The synergistic action of C. keratinophilum and M. gypseum on wool was found to be more effective. The biodegrading potential of C. keratinophilum can be made more effective by M. gypseum and C. carmichaelii, if the latter fungi join halfway through keratinolysis. C. keratinophilum did not act as a follower fungus in wool degradation [167]. A series of experiments were performed to see if keratinolytic C. carmichaelii and C. tropicum could be effective on the amount of wool decomposed by C. keratinophilum. It would be expected to occur because some of these might be utilising protein released as it breaks down. In doing so, it should relieve competitive inhibition of keratinase action and also relieve repression of keratinase synthesis [167]. C. indicum, C. keratinophilum, C. pannicola, C. queenslandicum and C. tropicum were also used for their capacity to degrade horn, hair and wool [142]. C. europae decomposed feather and released 457.33 µ g/ml protein along with another 47 Chrysosporium strains tested. NaCl, KCl and Ca Cl2 inhibited protein release from feathers when C. keratinophilum was used. This fungus was also used for feather decomposition during solid state fermentation [169]. Fungal colonization of hair in contact with soil C. tropicum appeared in 6 days on fresh feathers in peptone and water amended soil while it took more time in the presence of glucose to colonize defatted and sterilized feathers. In the presence of water 16-20 days were taken to colonize 100% fresh and defatted feathers. More than 25 days were required for achieving 100% colonization in the presence of glucose. Sterilized human hair was not completely colonized in peptone and glucose amended soil up to 25 days. Cow, goat and horse hairs were colonized completely in 20 days while fresh horse hair required more than 25 days in glucose amended soil [111]. Five strains of A. keratinophilus perforated hair in soil and caused 48-54% hair perforation in 30 days [100]. A. terreus perforated 63-100% hair. C. articulatum and C. carmichaelii colonized hair in 12 and 18 days but could not perforate them. Six C. indicum colonized hair in 4-15 days and 5 strains perforated hair. Five strains each of C. keratinophilum, C. pannicola and C. tropicum colonized 100% hair in 3-5 days. C. queenslandicum, C. zonatum and C. xerophilum also colonized hair through soil but later did not perforate hair. Chrysosporium and its potential Kushwaha RKS Decomposition of hair and feather in soil The hair from humans and animals and feather from birds which come to the soil either as dropped off or dead are affected by microbial decomposition. In the past few decades some studies on the decomposition of keratin in submerged cultures appeared. Biodegradation of keratin by using Chrysosporium and other related fungi in submerged culture is reviewed [160] and scattered reports are available in literature [137,138,142,161-168,170-172]. The process of keratin decomposition has also been found to be very fast in soil and it plays a very important role in energy transformation and nutrient cycling in soil. Decomposition of keratin in soil has received very little attention [167,174,175]. Several species of Chrysosporium were found to degrade hair through soil by a new method [166]. Combinations of C. indicum and C. keratinophilum, C. pannicola and C. keratinophilum, C. queenslandicum and C. keratinophilum, C. tropicum and C. keratinophilum, M. gypseum and C. keratinophilum were employed for finding out weight loss of hair and feather in soil The midway addition of the C. keratinophilum in C. queenslandicum experimental set showed 100% weight loss of feathers in 3 weeks. It is evident from overall observations made during experiments that, in general, the fungi acting in combination are found to be more effective in keratin decomposition than the individual action of various fungi. The midway addition of C. keratinophilum was the most effective in C. tropicum amended soil while midway additon of C. queenslandicum was more successful when initially inoculated with C. keratinophilum. Feather and wool degradation by C. keratinophilum was studied in natural garden and sterilized garden soil, where it was presumed that C. keratinophilum utilises keratin without any competition from microorganisms for up to 3 weeks, and later a slight decline was noted. This showed that C. keratinophilum can efficiently cause keratin breakdown while competing with other fungi in soil [167]. The effect of C. tropicum on soil which had received decomposed wool products was studied during seed germination and seedling growth of Brassica campestris Linn, Zea mays Linn and Phaseolus Roxb. and it was concluded that this fungus could be used for the slow release of nitrogen fertilizer in soil [176]. Pathogenicity Chrysosporium sp. was cultured from a tissue biopsy of the nasal mucosa which was found in brain, lungs and left kidney as well as nasal and sinus regions [177]. Chrysosporium sp. was isolated from 2 biopsy specimens of a 24 - year - old man [178] and three other male patients [179]. Pathogenecity of Chrysosporium spp. and C. parvum var crescens was also confirmed [180-182]. Chrysosporium infection in a bone marrow transplant 73 recipient was noted as Chrysosporium which caused an invasive infection in an 18 - year - old woman where infection began as a facial swelling and which extended into the central nervous system [183]. Studies were carried out on the epidemiological, immunological, biochemical and physiological properties of Chrysosporium sp. and C. keratinophilum along with some dermatophytes [184]. The pathogenic role of C. keratinophilum, C. asperatum, C. georgii, C. tropicum, C. pannorum, Chrysosporium state of A. curreyi, Chrysosporium state of A. multifidum, Chrysosporium state of A. tuberculatum is uncertain but their ability to remain viable for several weeks in skin and peritonial tissue indicates that they could become pathogenic in certain circumstances [181,182]. Antigenic activity within the genus Chrysosporium was demonstrated [185]. A. keratinophilus, A. fulvescens, A. reticulisporus and A. verrucosus produced nodular lesions when inoculated intraperitonially [186]. The isolation of C. merdarium from nail, C. pseudomerdarium from lung of rodent, C. carmichaelii from human skin and sputum, C. queenslandicum from snake, Chrysosporium anamorph of Gymnoascus demonbreunii from human, Chrysosporium anamorph of Pectinotrichum llanense and C. inops from human skin, C. sulfureum from bones, Chrysosporium anamorph of Rollandina vriesii from skin and lung of lizard, C. lobatum from scrappings of human and skin crest and C. pannicola from dog [5] are all also indicative of their potential for pathogenecity. Future Prospects It is felt that there has been no attempt to determine the geographical distribution of Chrysosporium. Large areas of the world have yet to be sampled for its teleomorphic connections and germplasm collection. Molecular characterization of Chrysosporium species, its teleomorphs and related genera may be able to provide help in solving diagnostic problems but this criterion should be cosidered as the second step because visualization of characteristics has always had the upper hand. More information on the pathogenic and saprophytic survival, spread and nature of the life cycle of Chrysosporium is needed. An approach in using combinations of wild as well as genetically engineered strains of a single species or different species of Chrysosporium for enhancing biodegrading potential merits study. Detailed ecological and physiological studies followed by suitable selection and development of specific strains may lead to a commercially viable use of fast - growing non - pathogenic strains of this fungus. Most of the work cited here is financed by Department of Science and Technology and University Grants Commission, New Delhi, India. 74 References 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. Sturm J. Deutschlands Flora 3 1833; Nurnberg. Saccardo PA. Sylloge Fungorum 1901; 15. Carmichael JW. Chrysosporium and some other aleuriosporic hyphomycetes. Can J Bot 1962; 40:1137-1173. Dominik T. Chrysosporium Corda. Zesz nauk wyzszkroln szczec 1967;24:37-66. Oorschot CAN van. A revision of Chrysosporium and allied genera. Stud Mycol 1980;20:1-89. Kushwaha RKS. Distribution of dermatophytes and keratinophilic fungi in soils of India. ICMR Project Report 1985;13pp. Kushwaha RKS. Biology of Chrysosporium and allied fungi with special reference to their morphology and taxonomy. CSIR Project Report 1987;44pp. Kushwaha RKS. Biology of keratinophilic fungi pathogenic to man and animals. DST Project Report 1994;116pp. Nigam N, Kushwaha RKS. Distribution of Chrysosporium in Indian soil. IV Internat Mycol Cong 1990;3:144. Nigam N, Kushwaha RKS. Ecology of soil inhabiting keratinophilic fungi with special reference to Chrysosporium. In: Rai B, Arora DK, Dubey NK, Sharma PD (Eds) Fungal Ecology and Biotechnology Rastogi Publications. India, 1993;173-182. Kushwaha RKS. Isolation and conservation of germplasm with special reference to Onygenaceae and their keratin degrading ability. DST Project Report 1999;68pp. Nigam N, Kushwaha RKS. Hyphal interference among Chrysosporium tropicum, keratinophilic and saprophytic fungi. Trans Jap Mycol Soc 1990;31:399-404. Kushwaha RKS. Human and animal hair invasion by keratinophilic fungi, dermatophytes and opportunistic keratin colonizing fungi. UGC Project Report 1997;101pp. Nigam N. Antagonistic properties of Chrysosporium Corda. DST [YS] Project Report 1993;22pp. Ulfig K. Keratinophilous fungi in the sediments of surface water bottoms. Acta Mycol 1987;23:3-11. Chabasse D. Taxonomic study of keratinophilic fungi isolated from soil and some mammals in France. Mycopathologia 1988;101:133-140. Al-Musallan AA. Distribution of keratinophilic fungi in animal folds in Kuwait. Mycopathologia 1990;112:65-67. Dixit AK, Kushwaha RKS. Keratinophilic fungi from Andaman Islands, India. Ind J Microbiol 1990;30:349-350. Ulfig K, Terakowski M, Plaza G, Kosarewicz O. Keratinophilic fungi in sewage sludge. Mycopathologia 1996;136:41-46. Piontelli E, Alicia M, Toro SM, Dunny Casanova Z. Diversity, dominance and succession of fungal communities in sandy soil [A beach of V. Region – Chile] on keratinic substrates. Bolet Micologico 1984;2:73-89. Singh CJ, Singh BG, Singh BS. Keratinophilic fungi of Ghana birds sanctuary Bharatpur, Rajasthan. Ad Plant Sci 1994;7:280-291. Nigam N, Kushwaha RKS. Some new reports on keratinophilic fungi. Curr Sci 1989;58:1374. Nigam N, Kushwaha RKS. Occurrence of keratinophilic fungi with special reference to Chrysosporium species in India. Sydowia 1990;42:200-208. Abdel-Gawad KM. Fungi on claws of buffalo and cow in Egypt. J Basic Microbiol 1989;29:323-328. Eshmm All AF. The occurrence of keratinophilic fungi in sewage sludge from Egypt. J Basic Microbiol 1990;30:73-79. Abdel Hafez AII, EL-Sharouny HMM. Keratinophilic and saprophytic fungi isolated from students nail in Egypt. J Basic Microbiol 1990;30:13-20. Dixit AK, Kushwaha RKS. Occurrence of keratinophilic fungi on Indian birds. Folia Microbiol 1991;36:383-386. 28. Ghawana CVK, Shrivastava JN, Kushwaha RKS. Some observations on fungal succession during decomposition of wool. Mycoscience 1997;38;79-81. 29. Caretta G, Piontelli E. Isolation of keratinophilic fungi from soil in Pavia Italy. Sabouraudia 1975;13:33-37. 30. Gugnani HC, Randhawa HS, Shrivastava JB. Isolation of dermatophytes and other keratinophilic fungi from apparently healthy skin coats of domestic animals. Ind J Med Res 1971;59:16991702. 31. Caretta G, Mancianti F, Ajello L. Dermatophytes and keratinophilic fungi in cats and dogs. Mycoses 1989;32:620626. 32. Bagy MMK. Saprophytic and keratinophilic fungi isolated from desert and cultivated soils continously exposed to cement dust particles in Egypt. Zentralblatt fur Microbiologie 1992;147:418-426. 33. Nigam N, Dhawan S, Nair MV. Deterioration of feather and leather objects of some Indian museums by keratinophilic and non keratinophilic fungi. Internat Biodeterio Biodegra 1994;33:145-152. 34. Ulfig K. Badania Wstepne nad wystepowaniem dermatofilow i innychgrzybow keratinofilnych w osadachdennych rzek i zbiornikow. Acta Mycologica 1983;19:331-340. 35. Soo-Hoo TS. Isolation of keratinophilic fungi from soil in Malaysia. Mycopathologia 1991;113:115-158. 36. Irena M, Dominik T. Further contribution to the knowledge of keratinolytic and keratinophilic fungi in immediate surroundings of cattle. Ekologia Polska-Seria A 1969;17:1-30. 37. Mangiarotti AM, Caretta G. Keratinophilic fungi isolated from small pool. Mycopathologia 1984;85:9-11. 38. Abdel-Hafez AII, Moharram AM, Abdel Gawad KM. Survey of keratinophilic and saprobic fungi in the cloven-hooves and horns of goat and sheep from Egypt. J Basic Microbiol 1990;30:13-20. 39. Oyeka CA. Isolation of dermatophytes and related keratinophilic fungi from mammals in Anambra State, Nigeria. Bull Animal Health Prod Africa. 1989; 37:143-146. 40. Ajello L, Kuttin ES, Burner AN, Kaplan W, Padhye AA. Occurrence of H. capsulatum Darling with a review of the current status of histoplasmosis in the middle East. Am J Trop Med Hyg 1977;26:140147. 41. Gugnani HC, Wattal BL, Sandhu RS. Dermatophytes and other keratinophilic fungi recovered from small mammals in India. Mykosen 1975;18:529-538. 42. Fonseca OJ de M. Keratinophilic soil fungi of Manaus. Acta Amazonica 1976;6:63-65. 43. Takahashi Y. Mycotic diseases of skin in Northern Japan. Jap J Dermatol 1971;81:187-213. 44. Piontelli E, Caretta G. Ecological considerations of some geomycetes isolated on keratinic substrates in mountainous localities in Chilean Andes. Rivista Patol Veg 1974;10:261-314. 45. Volz PA, Hsu YC, Lin CH, Ho SM, Chen ZC. The keratinophilic fungi of Taiwan. Taiwania 1975;20:23-31. 46. Bakerspigel A. The keratinophilic fungi of Ontario Canada. Mycopath Mycol Appl 1974;53:1-11. 47. Sharapov VM. Species fungorum ceratinophilorum novae et in URSS primum in ventae. Nov Syst niz Rast 1974;11:266271. 48. Kushwaha RKS, Agrawal SC. Some keratinophilic fungi and related dermatophytes from soil. Ind Natl Sci Acad 1976;42:102-110. 49. Garg AK. Isolation of dermatophytes and other keratinophilic fungi from soil in India. Sabouraudia 1966;4:259-264. 50. Padhye AA, Mishra SP, Thirumalachar MJ. Occurrence of soil inhabiting dermatophytes and other keratinophilic fungi from soil in Poona. Hind Ant Bull 1966;9:90-93. 51. Randhawa HS, Sandhu RS. A survey of soil inhabiting dermatophytes and related keratinophilic fungi of India. Sabouraudia 1965;4:71-79. 52. Jain M, Shukla PK, Shrivastava OP. Keratinophilic fungi and dermatophytes in Lucknow soils and their global distribution. Mykosen 1985;28:98-101. 53. Kushwaha RKS. Four atypical strains of Chrysosporium tropicum. Mycol Soc America News Letter 1982;33:37. 54. Ulfig K. Grzy by keratynofilne W osadach Dennych Zbiornika Zaporowego. Przeczyce Archiwum Ochrony Srodowiska 1989;1-2:67-71. 55. Ulfig K. Occurrence of keratinophilic fungi in soil of homestead attached gardens polluted by industrial emissions. Rocz IV. PZH 1991;62:445-449. 56. Ulfig K. Keratinophilic fungi in the environment. Wiadomosci Botaniczne 1990;34:510. 57. Chabasse D, Simmon B. Keratinophilic fungi isolated from pathways of parks and gardens in Angers District. Bull de la Soc Française de Mycologie Médicale. 1986;15:407-411. 58. Sundaram BM. Fungal flora of rice field soils. Proc Ind Acad Sci 1977;85:90-99. 59. Bracalenti BJC, de Alvarej DP, Colella MG. Ecology of dermatophytes I Correlation between dermatophytes and keratinophilic soil fungi of Rosario. Sabouraudia 1975;13:255-262. 60. Feurerman E, Alters I, Honig E, Lehrer N. The isolation of keratinophilic fungi from soil in Israel:A preliminary report. Mycopathologia 1975;56:41-46. 61. Schonborn C, Winden F. Investigations on the occurrence of fungi in air and dust of hospital wards. Mykosen 1973;16: 385-391. 62. Hejtmanek M, Hejmankova N, Kunert J. On the occurrence of geophilic dermatophytes in Asia. Ceska Mycologie 1973;27:159-161. 63. Visset MF, Vermeil C. Contribution to the knowledge of keratinophilic fungi of west France. V. Scanning electron microscopic study. Mycopath Mycol Appl 1973;49: 89-100. 64. Sarpakes E, Belegrake A, Maseloukinte O. Soil dermatophytes and atmospheric mycoflora of Bolos. Acta Microbiologica Hellenica 1985;30:73-84. 65. Youssef YA, El-Din AAK, Hussanein SN. Occurrence of keratinolytic fungi and related dermatophytes in soils of Cairo Egypt. Zentralblatt fur Microbiologie 1992;147:80-85. 66. Ulfig K, Korcz M. Isolation of keratinolytic fungi from coal mine dump. Mycopathologia 1995;129:83-86. 67. Arx JA von. Further observations on Sporotrichum and some similar fungi. Persoonia 1973;7:127-130. 68. Pugh GJF, Allsopp D. Microfungi in Signy Island, South Orkney Islands. Br Antarct Surv Bull 1982;57:55-67. 69. Caretta G, Piontelli E. Microsporum magellanicum and Cunninghamella antarctica, new species isolated from Australic and Antarctic soil of Chile. Sabouraudia1977;15:1-10. 70. Meinhof W, Grabowski A. Geophilic dermatophytes and other keratinophilic fungi in soil samples of alpine region. Hautarzt 1972;23:359-362. 71. Garg AP, Gandotra S, Mukerji KG, Pugh GJF. Ecology of keratinophilic fungi. Proc Ind Acad Sci 1985;94:149-163. 72. Kushwaha RKS. Studies on keratinophilic fungi from soil. Ph.D.Thesis. Univ Sagar India, 1976. 73. Brooks FT, Hansford CG. Mould growth upon cold stored meat. Trans Br Mycol Soc 1923;8:113-142. Chrysosporium and its potential Kushwaha RKS 74. Haines RB. The influence of temperature on the rate of growth of Sporotrichum carnis from –10°C to +30°C. J. Exp Biol 1931;4:379-388. 75. Vartiovaara V. Studies on metabolism of soil fungi. Acta Agral Fenn 1935;32:1-112. 76. Dickinson CH, Kent JW. Critical analysis of fungi in two sand dune soils. Trans Br Mycol Soc 1972;58:269-280. 77. Poole NJ, Price PC. The occurrence of Chrysosporium pannorum in soils receiving incremental cellulose. Soil Biol Biochem 1971;3:161-166. 78. Taylor JJ. Further clarification of Sporotrichum species. Mycologia 1970;62:797-825. 79. Hubalek Z. Fungi associated with free living birds in Czechoslovakia and Yugoslavia. Acta Sci Natl Brno 1974;8: 1-62. 80. Grimes M, Kennelly UCE, Cummiins HA. A study of fungi found in butter. Sci Proc R Dublin Soc NS 1930;19:549-569. 81. Pugh GJF, Mathison GE. Studies on fungi in coastal soils III An ecological survey of keratinophilic fungi. Trans Br Mycol Soc 1962;45:567-572. 82. Padhye AA, Pawar VH, Sukapure RS, Thirumalachar MJ. Keratinophilic fungi from marine soils of Bombay, India. Hind Antibiot Bull 1967;10:138-141. 83. Hubalek Z. Intraspecific affinity among keratinophilic fungi associated with birds. Folia Parasitologica 1976:23;267-272. 84. Hubalek Z, Balat F. Seasonal distribution of keratinophilic fungi in nests of tree sparrow [Passer montanus L.]. Zbl Bakt II 1976;131:179-197. 85. Pugh GJF, Evans MD. Keratinophilic fungi associated with birds. I Fungi isolated from feathers, nests and soils. Trans Br Mycol Soc 1970;54:233-240. 86. Chmel L, Hasilikova A, Hrasko J, Vlacilikova A. The influence of some ecological factors on keratinophilic fungi in soil. Sabouraudia 1972;10:26-34. 87. Ajello L, Padhye AA. Keratinophilic fungi of the Galapagos Island. Mykosen 1974;17:239-243. 88. Katiyar S, Kushwaha RKS. Human hair invasion by keratinophilic fungi isolated from Mediterranean Sea beach. Natl Acad Sci Letters 1997;20:71-74. 89. Kushwaha RKS, Agrawal SC. Physiological studies of some common Indian keratinophilic fungi. Proc Natl Acad Sci 1981; 51:105-106. 90. Kushwaha RKS, Agrawal SC. Carbon metabolism in some soil inhabiting keratinophilic fungi. Trans Jap Mycol Soc 1977;18:47-56. 91. Nigam N, Kushwaha RKS. Growth variations in fifteen Chrysosporium species. Ind J Microbiol 1988;28:209-210. 92. Kushwaha RKS. Nitrogen nutrition and free and bound aminoacids in three dermatophytes and Chrysosporium tropicum. Proc Ind Natl Sci Acad 1984;50:218-222. 93. Kushwaha RKS. Vitamin requirements of somesoil inhabing keratinophilic fungi. Bull Bot Soc Univ Sagar 1974;21:71-72. 94. Kushwaha RKS. Spore germination in some keratinophilic fungi and dermatophytic molds. J Ind Bot Soc 1983;62: 176-180. 95. Nigam N, Kushwaha RKS. Thermotolerance of keratinized and non keratinized spores of Chrysosporium tropicum and C. keratinophilum. Ind J Plant Sci 1986;4:5-8. 96. Kushwaha RKS, Agrawal SC. Efficacy of some antibiotics against spore germination of keratinophilic fungi. Acta Bot Indica 1977;5:174-177. 97. Kushwaha RKS. In vitro effect of prednisolone on some keratinophilic fungi. Ind Drugs 1976;14:11-12. 98. Singh CJ. Biomass loss by some volatile compounds in some keratinolytlic fungi and dermatophytes. Geobios 1998;25:6970 99. Katiyar S, Kushwaha RKS. Influence of some homeopathic drugs on human hair penetration by keratinophilic fungi. Biomedical Letters 1998;57:153-158. 100. Katiyar S. The keratinomycetes from bottom sediments and their hair invasion ability. Ph.D. Thesis. Kanpur University, India, 1998. 101. Singh CJ, Singh BG. Antifungal activity of some plant extracts against dermatophytes and related keratinophilic fungi. Adv Plant Sci 1997;10:249-251. 102. Singh CJ, Singh BS, Singh BG. In vitro antifungal activity of essential oils of official plants against dermatophytes and allied keratinofers. Adv Plant Sci 1997;9:35-37. 103. Nooruddin M, Randhawa SS, Singh B, Gupta RP, Gill BS, Dhaliwal PS. In vitro studies on antibacterial and antifungal activities of Himax and Teeburb. J Res Punjab Agri Univ 1986;23:499-504. 104. Nigam N, Shrivastava JN, Kushwaha RKS. Mycostatic effect of some oils and beverages on keratinophilic fungi. Bull Bot Soc Univ Sagar 1987;33:17-20. 105. Williams J. Pugh GJF. Fungal biological flora: Chrysosporium pannorum [Link] Hughes. Internat Biodeterio Bull 1974;10:75-80. 106. Simard RE. Removal of phosphate from sewage. Mycopath Mycol Appl 1973; 51:223-232. 107. Pugh GJF, Garg AP, Smith SN. Inhibition of keratinophilic fungus Chrysosporium keratinophilum. In: Barrys Houghton DR (Ed.) Biodeterioration CAB IMI 1986;6: 142-147. 108. Bahuguna S, Kushwaha RKS. Hair perforation by keratinophilic fungi. Mycoses 1989;32:340-343. 109. Gupta M. Decomposition of hair and feathers by keratinophilic fungi. Ph.D. Thesis. Kanpur University, India, 1994. 110. Jahan F. Keratin decomposition by Chrysosporium tropicum. Ph.D. Thesis. Kanpur University, India, 1998. 111. Bahuguna S. Hair perforation leading to its degradation by keratinophilic fungi. Ph.D. Thesis. Kanpur University, India, 1993. 112. English MP. The saprophytic growth of keratinophilic fungi on keratin. Sabouraudia 1963;2:115-130. 113. Katiyar S, Kushwaha RKS. Human hair invasion by keratinophilic fungi isolated from house dust. Ind Sci Cong Assoc 1997;84:11. 114. Katiyar S, Kushwaha RKS. Hair invasion by keratinophilic fungi: A review. Microbial biotechnology for sustainable development and productivity. 1998: 161-162. 115. Katiyar S, Kushwaha RKS. Invasion of human hair by Chrysosporium of bottom sediments. Microbes in Sustainable Environment 1997;159. 116. Katiyar S, Kushwaha RKS. Demolition of human hair by keratinophilic fungi isolated from bottom sediments. Proc Ind Bot Soc 1997;20:13. 117. English MP. Destruction of hair by Chrysosporium keratinophilum. Trans Br Mycol Soc 1969;52:247-255. 118. Dominik T, Majchrowicz I. A trial for isolating keratinolytic and keratinophilic fungi from the soils of cementeries and forests of Szczecin. Ekol Pol 1964;13:415-447. 119. Majchrowicz I, Dominik T. Further contribution to the knowledge of keratinolytic and keratinophilic soil fungi of the region of Szczecin. Ekol Pol 1969;17:87-116. 120. Dominik T, Ihnatowicz A, Kopilow H, Mietkiewski R. Mycoflora of sand boxes in kindergardens in Szczecin. Ekol Pol 1973;21:901-923. 121. Rippon JW, Lee FC, Mc Millan S. Dermatophyte infection caused by Aphanoascus fulvescens. Arch Dermatol 1970;102:552-555. 122. Caretta G, Del Frate G. Funghi cheratinofili dell isola di Motecristo, isolamenti da suolo, escrementi e pelo di animali e piume di uecelli. Atti Ist Bot Lab Critt Pavia 1976;11:203-208. 123. Caretta G, Del Frate G, Piontelli E, Todaro F. Micoflora cheratinofila del pelo e dello sterco di mucca, del foraggio e del suolo di fattoria: Considerazioni sulla loro distribuzione. Riv Parassit 1976;37:333-361. 124. Caretta G, Del Frate G, Piontelli E, Todaro F. Distribuzione di funghi cheratinofili nel suolo del vulcano Etna [Sicilia]. Riv Parassit 1977;38:115-127. 75 125. De Vroey C, Rechacochea M. Keratinophilic fungi isolated from soil in northwestern Nepal. Fr Mycol Med 1977;6:207-210. 126. Mercantini R, Marsella R, Caprilli F. Isolation of keratinomycetes from soil of wild animal cages and enclosures in the zoo of the Parco Nazionale d’ Abruzzo, Italy. Sabouraudia 1978;16:285-289. 127. Muhammed SI, Lalji N. The distribution of geophilic dermatophytes in Kenyan soils. Mycopathologia 1978;63:123-128. 128. Van Glederen De Komaid A, Elias F. Presence of dermatophytes in the soil of school in the province of Tucuman, Argentina. Rev Latinoam Microbiol 1978;20:95-98. 129. Filipello Marchisio V, Luppi Mosca AM, Attivita cheratinolitica in vitro di miceti isolati dalle sabbie di un arenile in un parcogiochi. Allonia 1980;24:127-131. 130. Mc Aleer R. Investigation of keratinophilic fungi from soil in western Australia: a preliminary survey. Mycopathologia 1980;72:155-166. 131. Mercantini R, Marsella R, Caprilli F, Dovgiallo G. Isolation of dermatophytes and correlated species from the soil of public gardens and parks in Rome. Sabouraudia 1980;18:123-128. 132. Abdel – Fattah HM, Moubasher AH, Maghazy SM. Keratinophilic fungi in Egyptian soils. Mycopathologia 1982;79:49-54. 133. Filipello Marchisio V. Keratinolytic and keratinophilic fungi of children’s sand pits in the city of Turin. Mycopathologia 1986;94:163-172. 134. Fusconi A, Filipello Marchisio V. Ultrastructural aspects of demolition of human hair in vitro by Chrysosporium tropicum. Mycoses 1991;34:153-165. 135. Cano J, Guarro J, Figueras MJ. Study of the invasion of human hair in vitro by Aphanoascus spp. Mycoses1991;34: 145-152. 136. Agrawal SC, Kushwaha RKS. A method for the determination of keratinophilic molds. Curr Sci 1972;43:791-792. 137. Kushwaha RKS. Biodeterioration of feathers by keratinomycetes isolated from a museum of Spain. In : Aranayanak and Singhasiri (Eds.) Biodeterioration of Cultural Property. Bangkok, 1995: 438-442. 138. Kushwaha RKS. In vitro degradation of peacock feathers by some fungi. Mykosen 1983;26:324-326. 139. Kushwaha RKS, Nigam N. Keratinase production by some geophilic fungi. Natl Acad Sci Letters 1996;19:107-109. 140. Deshmukh SK, Agrawal SC. Biology of keratinophilic fungi and related dermatophytes. In : Varma A (Ed.) Microbes: For Health, Wealth and Sustainable Environment. MPH New Delhi, 1998: 253-272. 141. Singh CJ, Singh BG. Characterization of extracellular, proteolytic enzyme of Chrysosporium tropicum CF34 and its role in keratin degradation. Ind J Microbiol 1995;35:311-315. 142. Kushwaha RKS, Agrawal SC. Microbial degradation of keratin. Proc Natl Acad Sci 1981;51:181-183. 143. Kushwaha RKS, Agrawal SC. Amylolytic capability of some soil inhabiting keratinophilic molds. Proc Natl Acad Sci 1975;45:182-184. 144. Singh CJ, Singh BG, Singh BS. Biodegradation of keratin substrates in vitro by some keratinophilic fungi. Adv Plant Sci 1995;8:271-276. 145. Berliner MD. Gelatin hydrolysis for identification of the filamentous phase of Histoplasma, Blastomyces and Chrysosporium species. Sabouraudia 1967;5:274-277. 146. Rajak RC, Parwekar S, Malviya H, Hasija SK. Keratin degradation by fungi isolated from ground of gelatin factory in Jabalpur, India. Mycopathologia 1991;114:83-87. 147. Nigam N. Studies on Chrysosporium and allied fungi from soil. Ph.D. Thesis. Kanpur University, India, 1987. 76 148. Calvo RM, Calvo MA, Larrondo J. Enzymatic activities in Chrysosporium strains. Mycopathologia 1991;116:177179. 149. Haskins RH. Production of yellow pigment anthroquinones [questin and questinol] and asteric acid. Can J Microbiol 1971;17:1575-1579. 150. Hopsu-Havu VK, Sonck CE, Tunnela E. Production of elastase by pathogenic and non pathogenic fungi. Mykosen 1972;15:105-110. 151. Nigam N, Kushwaha RKS. Biomass and lipids of keratinophilic fungi. Acta Bot Indica 1987;15:135-137. 152. Koenig WA, Loeffler W, Meyr W, Uhmann R. L-Arginyl – D-allothreonil – L phenylalanine, ein Aminosaure – Antagonist aus dem Pilz, Keratinophyton terreum. Chem Bericht 1973;106:816. 153. Yamashita M, Tsurumi Y, Hosoda J, Komori T, Kosak M, Imanaka H. Cryscandin, a novel pepodyl nuclioside antibiotic. I Taxonomy, fermentation, isolation and characterization. J Antibiotics 1984;37:1279-1283. 154. Ogawa H, Hasumi K, Sakai K. Pannorin: A new 3 hydroxy 3 methyl glutaryl co enzyme, a reductase inhibiters. J Antibiotics 1991;44:762-767. 155. Kushwaha RKS. Interactions between keratinophilic fungi and dermatophytes. UGC Project Report 1993;33 pp. 156. Johnson LF, Curl EA. Methods for research on the ecology of soil borne plant pathogens. Minnesota USA: Burgess Publishing Co.1978. 157. Bahuguna S, Kushwaha RKS. Antifungal activity of some Chrysosporium species. Biomedical Letters 1992;47:99-103. 158. Saxena V, Kushwaha RKS. Interaction experiments on Chrysosporium anamorph of Arthroderma. Proc Ind Sci Cong 1992; 79:27. 159. Nigam N. Kushwaha RKS. Effect of staling products of keratinophilic and non keratinophilic fungi on the growth and spore germination of Chrysosporium tropicum. In Microbes and Sustainable Development 1997;62. 160. Kushwaha RKS. Biodegradation of keratin by soil inhabiting fungi. A Review. In: Varma A (Ed.) Microbes: For Wealth, Health and Sustainable Environment. MPH, New Delhi 1998:273-280. 161. Parihar P, Kushwaha RKS. Rapid degradation of human hair by Chrysosporium indicum isolated from soil of Thailand. Biotechnology New Trends and Prospects 1996;40. 162. Parihar P, Kushwaha RKS. Keratin decomposition in forest soil in successive manner. Environmental Biopollution 1997;25. 163. Parihar P, Kushwaha RKS. Keratin decomposition in soil. Recent Advances in Diagnosis and Management of Important Plant Diseases 1997;99. 164. Parihar P, Kushwaha RKS. Decomposition of feathers by some keratinophilic fungi. Proc Ind Bot Soc 1997; 20:13-14. 165. Nigam N, Kushwaha RKS. Decomposition of feathers and hairs by keratinophilic fungi. Ind J Microbiol 1989;29:241-244. 166. Nigam N, Kushwaha RKS. A method for the determination of hair penetration by keratinophilic fungi through soil. Ind J Microbiol 1989;29:359-360. 167. Nigam N, Kushwaha RKS. Biodegradation of wool by Chrysosporium keratinophilum acting singly and in combination with other fungi. Trans Jap Mycol Soc 1992;33: 481-486. 168. Nigam N, Kushwaha RKS. Biodeterioration of keratinous substrates. In: Toishi et al. (Eds.) Biodeterioration of Cultural Property Koyoto, Japan 1992; 180-185. 169. Parihar P. Exploitation of non pathogenic keratinophilic fungi for their potential to degrade keratin. Ph.D. Thesis. Kanpur University India, 1998. 170. Kunert J. Keratin decomposition by dermatophytes: evidence of sulphitolysis of the protein. Experentia 1972;28:10251026. 171. Kunert J. The digestion of human hair by the dermatophyte Microsporum gypseum in submerged culture. Mykosen 1972;15: 59-71. 172. Kunert J. Growth of keratinolytic and non keratinolytic fungi on human hairs. A physiological study. Acta Univ Olmouc Fac Med 1989;122:25-38. 173. Kunert J. Biochemical mechanism of keratin degradation by keratinolytic fungi. J Ind Bot Soc 1995;74A:89-98. 174. Wainwright M. A new method for determining the microbial degradation of keratin in soil. Experientia 1982;38:243-244. 175. Kundart AJ, Rowe FW. A study of hair degradation in agricultural soil. Biodeterioration Res 1989;2:91-98. 176. Shrivastava JN, Ghawana VK, Mathur M, Kumar A, Bhatnagar VP. Potential role of keratin decomposing fungi in soil fertility. Assoc Microbiologists India 1997;38:3. 177. Levy FE, Larson George E, Maisel RH. Invasive Chrysosporium infection of the nose and paranasal sinuses in an immunocmpromised host. OtolaryngologyHead and Neck Surgery 1991;104: 384-388. 178. Stillwell WT, Rubin BD, Axelord JL. Chrysosporium, a new causative agent in osteomylitis. A case Report. Cli Orthopad Related Res 1984;184:190-192. 179. Furtado M, dos des Thara LT, Maroja M, de Jose JI, Castrillon Al. Dermatophytoses na cidade de Manaus-AM. Anais Brasileiros de Dermatologia 1987; 62:195-196. 180. Chabasse D, Gentle L de, Bouchara JP. Pathogenecity of some Chrysosporium species isolated in France. Mycopathologia 1989;106:171-177. 181. Chabasse D, Gentle L de, Bouchara JP. Pathogenecity of some keratinophilic fungi belonging to the genus Chrysosporium. A preliminary study. Bull Soc Fr Mycol Med 1989;18:185-190. 182. Otcenasek M, Hubalek Z. Chrysosporium parvum var crescens – little known animal and human fungal pathogen. IV Internat Mycol Cong 1990;IIE 248. 183. Warwick A, Ferrieri P, Burke B, Blazar BR. Presumptive invasive Chrysosporium infection in bone marrow transplant recipient. Bonemarrow Transplantation 1991;8:319-322. 184. Balabanoff V, Danev P, Radev S. Comparative study of physiological, biochemical, epidemiological and immunological properties of dermatophytes and Candida albicans and their evolutionary aspects. Rev Iberoam Micol 1990;7: 27-30. 185. Tomsikova A, Novackova D. Contribution to antigenic activity within the genus Chrysosporium Corda. Dermat Monatsschrift 1976;162:135-141. 186. Cano J, Mayayo E, Guarro J. Experimental pathogenicity of Aphanoascus spp. Mycoses 1990;33: 41-45.