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Abiotic and Biotic Factors: Effecting the Growth of Keratinophilic Fungi

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Manish Mathur and Neha Mathur

Submitted: October 8th, 2021Reviewed: February 15th, 2022Published: April 19th, 2022

DOI: 10.5772/intechopen.103716

Fungal Reproduction and GrowthEdited by Sadia Sultan

From the Edited Volume

Fungal Reproduction and Growth [Working Title]

Dr. Sadia Sultan and Dr. Gurmeet Kaur Surindar Singh

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Fungi portray an important role in decomposition of keratin, as their activity is tough to measure. According to an estimation, a quantity of cellulose is synthesized by primary producers over photosynthesis and then reinstated to the atmosphere as carbon dioxide and through the activity of fungi, which decompose the complex and inflexible polymer. Without this activity, the world would soon be submerged by plant residues, and this would probably exclude most living organisms from their natural habitat. This chapter deals with several abiotic and biotic factors, which effect the growth of keratinophilic fungus and the substrates, which can serve as potential growth promoters for them.


  • keratinophilic fungi
  • growth factors
  • fungus substrate

1. Introduction

The degradation is associated to the aptitude of the organism to develop on a substrate. The capacity of the growth depends upon the enzymes secreted by the microbe and conditions such as PH, protein content in the substrate, temperature, moisture, etc. The native feather keratin was degraded by species of Aphanoascus fulvescensand Chrysosporium articulatum, which are separated from soil. Recognition of strain was done by phenotypical qualities and nucleotide sequencing of protein. The ultimate deficit of substrate in comparison to the strain was recorded for Aphanoascus fulvescensand deficit of substrate for the Aphanoascus fulvescens (71.08%). Keratin is high in protein and abundant in nitrogen and sulfur [1, 2, 3]. Keratins have high mechanical and chemical resistance. Numerous disulfide bonds (S–S) are liable for the resistance. Except a few months, only keratinolytic microorganisms can grow and degrade native keratin including some bacteria of the genera Bacillus, Vibrio, Serratia([4], actinomycetes [5]). The genus Bacillus[6], representative geophilic dermatophytes are associated with the fungi known as Chrysosporia, name taken from the genus Chrysosporium[7]. The development of fungi is backed by a humus, neutral or a little alkaline pH reaction, fruitfulness in CaCO3 [8]. The Chrysosporimbelongs to this group, specialized in the growth of keratin, e.g., feathers, hairs [9]. Although it is considered safe in certain stages but can be converted into pathogenic form under certain conditions. Kushwaha [9] reported that genus Chrysosporiumis effective in disorienting keratin; however, keratin degradation varies in the species. Some microbes grow on keratinous materials including bacteria, biodegradation has been exhibited by Bacillus, particularly Bacillus licheniformis[10] and Bacillus subtilis[11], and Chryseobacterium. Actinomycetes from Streptomycesgenus produce keratinases [12, 13]. The most common keratinolytic fungi are Aspergillus, Penicillium[14], Fusarium[15].


2. Growth factors in keratinophilic fungi

To-Ka-Va hair baiting technique [16] was used in the experiments. Several workers projected that saprophytic phase of dermatophytic fungi in the soil are global [17, 18]. Dispersal of pathogenic fungi [17, 19, 20, 21, 22] specifies soil turns as pool meant for primary contamination may be for some pathogenic fungi. Potentially it is pathogenic to wild animals and acts as the resource of secondary infection to man and pets [23, 24, 25, 26]. Besides soil, birds’ nests and feathers [27, 28, 29, 30, 31, 32, 33, 34, 35], hairs [25, 36], water [37], plant debris [38], and dung [39] are the different ecological places for growth of capable dermatophytic fungi.

Fungi produce enzymes potent of degrading keratinized forms known as keratinophyles. Pathogenic strains find a suitable host in favorable environmental and physiological conditions and produce the symptoms known as “ringworm diseases” by growing on human dermis.

2.1 Climatic factors

2.1.1 Temperature

Keratinophilic fungi are mesophilic while a few have grown at 37°C such as C. tropicum, Chrysosporium keratinophilum, C. queenslandicum, etc. Nevertheless, geophilic dermatophytes grow best at 25–30°C. In 1970, Pugh and Evans mentioned span of 25–27°C in the keratinophilic fungi, which do not grow more than at 40°C.

The broadly recognized view about mesospheric in nature has been disquieted by the results of Battelli et al. [40], about the existence of Microsporuzri pypseiim, Trichophyton ajelloi, T. terrestrein alpine mountain. Pugh and Allsopp [41] stated a recurring existence of Chrysosporium pannorumand Mortierellaspp. with exceptional incidence of Trichophyton terrestreand Chrysosporiumsp. Some varieties are thermotolerant, and recently, it was shown that they can adapt to adverse temperatures for survival [41].

2.1.2 Light

It was known that the UV light is fungicidal, which creates about a reserve of spore sprouting leading to a hypha. Excellence of illumination on spore germination has entirely been analyzed by Buchnicek [42].

First shown by Berde [43], the growth reticence of dermatophytes is via means of lamp. Fungicidal effect in Candida albicansdiffered with the strength of light sensitizer and light intensity as was observed by Dickey [44]. There is no effect on illumination of spore suspension of T. mentagrophyteswithout photosensitization [44]. The red or blue light when applied separately prevents the growth greater than visible light. The quantity of inhibitions by both color lights utilized individually is more than the reserve by their mixture.

2.1.3 Seasonal variations

Season is quite anticipated with respect to dermatophytic flora of the soil, which has climatic differences resulting in change of temperature. Keratinomyces ajelloi, Trichophyton Terrestre, and Microsporum gypseumwere found in soil samples during April and in August.

2.1.4 Soil pH

Effects of pH on microbial life are explored extensively. Bohme and Ziegler [45] observed about soil pH as verified by Pugh [32]. Ziegler [46] observed the degradation of keratin over a wide range. Findings uncovered the optimum pH for amylase 7–2, alkyl phosphates 8–7, lipids 6–8, proteinase 5.4–6.9. Ectoenzymes are shown dormant at pH below 4.5, and the enzymatic catabolism of keratinophyles occurs at pH 6.9. Ziegler [46] further confirmed that the most luxuriant growth and highest frequency of keratinophilic fungi were seen in soils with pH 6.9.

2.1.5 Carbon

Decomposition of carbon can be done by keratinophilic fungi but there is not enough information regarding carbonate impacting the delivery in soils. Chmel et al. [47] described favored incidence of M. qypseumin carbonate field soil, an alliance of K. ajelloiwith soils and a predilection of T. terrestrein alluvial soils. It was studied that a plenty of keratinophilic fungi in carbonate field and chernozemic soils have greater humus substance in comparison to gray podzolic soils. Keratinophyles can decompose extremely compound carbon complexes in many culture media, a simple carbon source can be used and not influence their qualitative spreading but the findings on their quantitative dispersal may disclose fascinating truths.

2.1.6 Nitrogen

Keratinolytic fungi can decompose keratin, which is a rich source of nitrogen. Rate of keratinolysis has been reported by Ziegler [46] at pH 6–9 supporting the earlier finding.

Growth of Microsporum gypseumis fine revealed by the experiments utilizing optically dissimilar forms of cystine, quite large amounts in the scleroprotein [48]. Carbon and nitrogen application was perceived when surplus sulfur was secreted in the medium and became oxidized. Oxidation of sulfur in the extracellular medium makes use of L-cystine, which was showing a slow form. The way of consumption of L- and D-L-cystine was the same. The nitrogen-containing substances of keratin differ from one another and in opportunity would impact colonization of the last.

2.1.7 Sulfur

Kunert [49, 50] calculated that the inorganic and organic sulfur resources for the development of the fungus M. gypseum., sulfite, disulfide, peroxodisulfate dithionate, and sodium sulfate were the best hints of inorganic sulfur. Inorganic sources sulfide produced a primary inhibition of the fungal growth supportive to a previous report that mineral water comprising H2S is repressing for the growth of dermatophytes. Amino acids such as cystine, cysteine, glutathione, S-sulfocysteine, lanthioneine, taurine, and serine-sulfate are the leading organic sulfur sources of the fungus.

2.1.8 Moisture content

An enzymatic reaction takes place in aqueous solution in the cell cytoplasm and is vital to life. The systems of germination, growth, and reproduction are energetic associated to the substrate moisture content. Pugh and Evans described a better percentage of spore growing in Arthroderma uncinatumand Ctenomyces serratusat 90–100% RH. It was also stated that an plenty of keratinophyles in birds’ nest with moisture usually showed a greater occurrence of keratinophilic fungi in bird nests with 15–20% water content [51].

2.1.9 Humus

High humus content in soils invites M. gypseumand T. terrestrein soils as revealed by Chmel et al.[47]. Whereas distribution of K. ajelloiand C. keratinophilumis regardless of soil humus. Soils made up of fragmented lava with little organic matter in the Galapagos Islands revealed by Ajello and Padhye [52] reported the occurrence of A. quadrifidum, C. indicum, C. keratinophilum, C. tropicum,and Ctenomyces serratus. A higher number of keratinophilic fungi were described in soils with exceptional humus value. In all levels of soil profile extreme in the region with high humus content, there exist M. gypseum and T. georgii,while T. oanbreuseqhemiiand its flawless state A. gertleriwere primarily found in soils with low humus value.

2.1.10 Fatty acids and oils

In 1899, Clarke antifungal estates of fatty acids were known to scientific world. The anti-dermatophytic properties were discussed by Rigler and Greathouse in 1940, which was later validated by Das and Banerjee [53].

Hajini et al. [54] studied unsaturated fatty acids, hair oils, and various natural fatty acids for their anti-dermatophytic properties. Reticence of growth of T. rubrumand associated dermatophytes was at 0.1% strength of mustard oil, oleic, linoleic. Linolenic, and aracliidionic acids while coconut oil, castor oil, till oil, Bryl cream, vaseline hair tonic, palmitic acid, and stearic acid did not prevent the growth even at 10% strength.

2.1.11 Salts

In coastal soils, various keratinophilic fungi can survive. A. curreiioccur although the rate of availability of keratin must be small. Varieties of hares, rabbits, and birds exhibited A. curreyiand C tenomices serratus. Padhye et al.[55] isolated Clirysosporium tropicumand Microsporumgypseum from long immersed marine soils in Bombay. C. indicumand Cteiiomyces sermonsand inaccessible to somewhat irregularly. The fungi remain to be revealed, for no such fungi have been found from coastal Mediterranean soils, which have salinity. A repressive sodium chloride impact on the growth has been stated of dermatophytes. Growth was reserved by NaCl in Microsporum, Epidermophyton, Trichophyton,etc.

2.2 Biotic factors

Biotic component influences the occurrence of keratinophilic fungi as the main causal factor in spread and persistence of ringworm diseases produced by dermatophytes up to a great extent. Biotic component influences the occurrence of fungi in birds and animals.

2.2.1 Birds

Pugh [56] displayed the existence of keratinophilic fungi on the experimental birds although validated them on birds in Australia. An association exists in the keratinophilic fungi and birds of correct order, e.g., A. curreyiand Turdus[31]; C. serratusand representatives of Galliforme, particularly partridges and chickens. Regularly conveyed from most of the bird forms [56], in Chrysosporiumspp. Commonly found species are C. k eratinophilum, Keratinomyces ajelloi,and T. terreste. There was significant habitation for the cleistrocarpic stage of C. serraius.

2.2.2 Animals

The role of wild animals as carrier of these diseases has been documented earlier in literature.

Occurrence of Tinea capitiswas in newborns who obtained disease from stray kittens. Incidence of scalp abrasions in an 8-year-old boy and a 3-year-old Yorkshire was seen as well as subsequent isolation of Microctenopoma nanumfrom soil. Soil acts as a group for dermatophytes [57].


3. Fungal growth on feathers

Keratin makes feather obstinate to common proteases such as trypsin, pepsin, papain, slowing its degradation process. Each bird has up to 125 gm of feather and approx. 400 million chickens processed universally the daily accumulation of which reaches 5 million tons [58, 59]. According to Lin et al.[60], the waste disposal is a global issue foremost to pollution of both air and water resources. Keratinase-treated feather is considered as a viable source of dietary protein in food and feed, as it has high nutritive value. Keratinases are potential market as proteases. Microorganisms are described to produce keratinase as Doratomyces microsporus, Alternaria radicina, Trichurus spiralis, Aspergillus sp., Rhizomucor sp., Absidia sp,etc., and actinomycetes as Streptomycespactum, S. alvs, Streptomyces thermoviolaceus, Streptomyces fradiae, Thermoactinomyces candidusetc.), and as bacterial species (Fervidobacterium islandicum, Pseudomonas aeruginosa, Microbacterium sp., and Bacillusincluding Bacilluslicheni formisand B. pumilus) [61]. Feather is insoluble fibrous protein and highly resistant [62] to enzymatic digestion [63]. Though, fungi often colonize on various keratinous substrates, degrade them and enhance the minerals in soil [64].

Hydrolytic enzymes are synthesized by filamentous fungi. Various species are used to produce industrially important enzymes as distinct proteases, carbohydrates, and lipases. It is the key enzymes in fungal incursion of skin found in dermatophytes as Trichophyton[65]. Candidaalso contributes to skin infections. Enzymes are found in Streptomyces[66] and Bacillusspp. [60]. Novel and Nickerson [67] examined bacteria, actinomycetes, and fungi for keratinolytic activity and found Streptomycesas most active in the decomposition of sheep wool. Keratin hydrolysis was the most active in Verticillium tenuipes, Trichophyton equinum,and T. mentagrophytesin peacock feathers. T. mentagrophytes, T. verrucosum,and Keratinomyces ajelloidegraded hair, whereas only T. gallinaedegraded chicken feathers.


4. Fungal growth on hair

Keratin, the fibrous protein, is a codified part of hair, wool, and related structures, which differ from other proteins in their high cystine content.

Five keratinophilic fungi, i.e., Chrysosporium indicum, Geotrichum candidum, Gymnoascoideus petalosporus, Scopulariopsis brevicaulis,and Talaromyces trachyspermus,which grow on human hair in stationary culture, have been examined. Hair was studied on criteria of cysteine, cystine, inorganic sulfate, thiosulfate, total protein, keratinase, and change in alkalinity. Gymnoascoideus petalosporusshowed degradation to remaining isolates when grown on human scalp hair as the sole resource of nutrients in vitro.

Kunert [68, 69] described a release of cystine, cysteine, and sulfate in the culture filtrate of Microsporum gypseumgrowing on hair. Ruffin et al. [70] discovered the S-sulfo cysteine in culture fluid and established the role of sulfitolysis during keratin degradation by Keratinomyces ajelloi.Stahl et al. [71], Chesters and Mathison [72], and Ziegler & Bohme [73] could not detect cysteine in filtrates of the dermatophytes studied. Weary et al. [63] recorded the production of 21 pg./ml and 38 pg./ml of cysteine by two strains of Trichophyton rubrum.


5. Fungal growth on leather

Samples collected from different museums of feather and leather objects and deposited dusts were studied for the isolation of keratinopliilic and non-keratinophilic fungi. Throughout the study, five species of Chrysosporium, four of Aspergillus, one of Penicillium,and two each of Acremoniumand Fusariumwere isolated.

Deterioration of objects of cultural value, in the conservation of cultural heritage, is a real problem. Microbial activity on museum objects, especially skin or leather, starts with a surface infection that eventually invades the full thickness of the skin. The most common molds belonging to the genera Aspergillusand Penicilliumemerge as black and green surface discoloration. There are, however, other molds, although of rare incident compared with the above, causes degradation of skin objects [74, 75, 76].

From different museums of Northern India, 24 keratinophilic and non-keratinophilic fungi represented by five genera and 15 species were isolated. Strains of C. keraiinophilum,three of C. tropicum,two each of C. evolceanuiand C. indicum,and one Chrysosporiumsp. were found in museums. Some non-keratinophilic fungi, i.e., three isolates of Aspergillus flavus,three of A. niger,one each of A. sulphureusand A. luchuensis,two species of Penicillium, i.e., P. ciirinum, P. chrysogenum,and two species of both of Fusariumand Acremoniumwere found and called as non-keratinophilic [77]. These were involved in the degradation of feather and leather objects.

Keratinophilic and non-keratinophilic fungi arise in regions where they can find dissimilar types of keratinaceous substrates. The existence of the fungi was connected with numbers of people inhabiting [47]. English [78] found 31 saprophytic fungi capable of inhabiting keratin along with non-keratinophilic fungi on keratinic substrate witnessed by Nigam and Kushwaha [77]. Aspergillusand Penicilliumcolonized keratinous substrates, and pathogenic behavior was revealed by Kishimoto and Baker [79]. The capacity of these fungi to colonize keratinous substrates was confirmed by Carmichael [80]. Some saprophytes are involved in microbial deterioration of leather [81, 82].

Keratinophilic fungi were commonly found on birds and animals and also isolated in the Antarctic [74]. Isolation of Chrysosporiumspp. is supported by other work, and it was frequently isolated [55, 83, 84, 85, 86, 87, 88]. The presence of Chrysosporiumand related keratinophilic fungi was reported from museum objects, their surroundings, or their deposited dusts.

In 2006, several valued leather objects were found during archeological excavation of Ghalee-Kooh-i Ghaen (historic stronghold from the Seljuk period, 11th–13th centuries) in the South Khorasan province of Iran. When examined after 5 years, there were red stains on the fragments of a shoe with poor strength and powdery surface like red rot decay. Since red rot is more common in manmade leathers from the mid-nineteenth century as clarified by the structural features and degradation factors responsible for red stains on the shoe.

As leather production is an ancient industrial activity [89], historical and archeological leathers represent an important part of any society’s cultural materials. Ghalee-Kooh-i Ghaen isa fortified castle from the Seljuk period (11th–13th centuries) located 3.4 km from Ghaen city in the South Khora-san province of Iran [90]. It was destroyed in 1066 A.D. by an intensive earthquake in the Ghaen area [91]. Later, Hossein Ghaeni rebuilt the castle in the late eleventh century to use as headquarters in the southern part of Khorasan known as Ghohestan [92]. The castle was registered as no. 4803 in the list of Iranian national monuments due to its historical and archeological value in 2002.

Several leather bottles, shoes, a fur, and some pieces of leather were found in the archeological investigation. Studies of the leather bottles indicated goat skin treated with lime depilation, vegetable tanning, and animal fats as lubricant for leather making. After excavation, the leathers were stored in inappropriate conditions at the base of the cultural heritage organization of South Khorasan province without any preventive measures. During the time of storage, there was no information about the environmental condition of the leathers within their archeological context in 2011 one of the excavated shoes was examined. Red rot is more common in manufactured leathers from the mid-nineteenthcentury, but the decomposition pattern may be found in more ancient leathers as well [93]. Therefore, this leather shoe was studied to better understand the deterioration mechanism prior to any interceptive activities.

Previous investigations on the structural features and degradation of leather and parchment artifacts have revealed important information, which is of value to our study [94, 95]. Sulfuric acid is regarded as main deterioration factor of red rot [96], possibly originating from environmental pollution or the materials used in the leather-making process [97]. Additionally, the production of acid from vegetable tannins, especially condensed and disintegration of the collagen-tannin complex, may result in red rot [96]. Many methods are available for identifying the decay, such as assessment of esthetic properties, pH and evaluation of the physical and mechanical characteristics of the object [96, 97, 98]. Red rot probably occurs in leathers with pH under 2.8. However, the degree of degradation cannot be determined just by pH measurement [98]. Moreover, biodeterioration is another important factor altering the artistic and useful properties of leather [99]. Fungi species attach to materials such as paper, textile, wood, paints, leather, etc., and produce characteristic signs that can be used for identification [100]. Leather composed of collagen, i.e., fibrous protein, provides a source for the evolution of proteolytic fungi. Some species of Aspergillus and Penicilliumhave been identified as the most common species related to leather and parchment molds [101]. There are also other rare molds responsible for biodeterioration of proteinous materials. Ebrahimi et al. [100] observed the activity of Aspergillusspp., Penicilliumspp., Chrysosporiumspp., Madurellaspp., Trichophytonspp., and Zygomycotaon the leather artifacts in Shahrekord Museum, Iran. Abdel-Maksoud [102] identified Penicillium spp., Aspergillusspp., and Fusariumsp. as the most profuse fungi found on a leather book binding of a Quranic manuscript from the nineteenth century. Nigam et al. [103] attributed degradation of leather and feather objects in Indian museums to keratinophilic and non-keratinophilic fungi.


6. Fungal growth on wool

Common method of isolating dermatophytes of so-called “keratinophilic” fungi is by baiting with hair [104, 105]. Little is known of the relationship of these non-dermatophytic fungi to the decomposition and utilization of keratinous substrates, although previous studies [78] have indicated that role in the breakdown of keratinous tissue may be significant.

Fungal succession on woolen baits was studied and was found that the initial colonizers on woolen baits are non-keratinophilic fungi, while the late colonizers are keratinophilic fungi comprising six phases during fungal succession. The successional tendencies obtained during degradation of wool in samples collected from plain and hilly areas, apart from for the prevailing colonization in the last phase, composed of chrysosporium tropicumfor the plain, but Miorosporumgypseum and M. fulvumfor the hilly area.

Various workers [68, 69, 75, 76, 106, 107, 108, 109] studied degradation of keratinous material, but none of them reported the involvement of other soil microbes in the decomposition of keratin. On succession of fungi on keratinous material [104], little but adequate information on fungal succession on woolen baits is still lacking.

The archeological textiles have a unique position in archaeology, textiles probably contain much archeological information, even more than ceramics. During the Bronze Age (1700–500 BCE), the trade of wool textiles was complex, widespread, significant, and possibly important as the metals [110]. Foundation of economic and political development in the late Middle Ages (c. CE 1100–1500) [111] was Trans-European trade of these materials.


7. Conclusions

The keratinophilic fungi can cause superficial mycosis both in humans and animals. They include a variety of taxonomic groups of filamentous fungi, one of them being the dermatophytes fungi. The keratinophilic fungi can produce a specific enzyme named keratinase that is responsible for keratin degradation. Keratinases can be serine proteases or metalloproteases [86]. The keratinophilic fungi could use keratin from keratinized materials (superficial layers of the skin, hair shaft and nails in humans and claws, horns, wool in animals) as the unique source of carbon and nitrogen. Keratin is found predominantly in feathers, hair, nails, horns, hooves, furs, claws, bird beaks, skin and consists of two types of keratin: α (alpha) and β (beta)-keratin; α keratin (soft) is usually found in hair, wool, horns, nails, claws, and hooves, whereas β keratin (harder) is found in bird feathers, beaks, and claws. Understanding of abiotic and biotic factors and role of different substrates for the growth of these fungi can help researchers to study their growth pattern and conduct their studies for better management of these fungi.


Conflict of interest

The authors declare no conflict of interest.


  1. 1.Korniłłowicz-Kowalska T, Bohacz J. Biodegradation of keratin waste: Theory and practical aspects. Waste Management. 2011;31:1689-1701. DOI: 10.1016/j.wasman.2011.03.024
  2. 2.Kunert J. Physiology of keratinophilic fungi. In: Kushwaha RKS, Guarro J, editors. Biology of Dermatophytes and Other Keratinophilic Fungi. 1st ed. Bilbao: Revista Iberoamericana de Micología; 2000. pp. 77-85
  3. 3.Onifade AA, Al-Sane NA, Al-Musallam AA, Al-Zarban S. A review: Potentials for applications of keratin-degrading microorganisms and their enzymes for nutritional improvement of feathers and other keratins as livestock feed resources. Bioresource Technology. 1998;66:1-11. DOI: 10.1016/S0960-8524(98)00033-9
  4. 4.Kim JM, Lim WJ, Suh HJ. Feather-degrading Bacillus species from poultry waste. Process Biochemistry. 2001;37:287-291. DOI: 10.1016/ S0032-9592(01)00206-0
  5. 5.Vasileva-Tonkova E, Gousterova A, Neshev G. Ecologically safe method for improved feather wastes biodegradation. International Biodeterioration & Biodegradation. 2009;63:1008-1012. DOI: 10.1016/j.ibiod.2009.07.003
  6. 6.Farag AM, Hassan MA. Purification, characterization and immobilization of a keratinase from Aspergillus oryzae. Enzyme and Microbial Technology. 2004;34:85-93. DOI: 10.1016/j.enzmictec.2003.09.002
  7. 7.Simpanya MF. Dermatophytes: Their taxonomy, ecology and pathogenicity. In: Kushwaha RKS, Guarro J, editors. Biology of Dermatophytes and Other Keratinophilic Fungi. 1st ed. Bilbao: Revista Iberoamericana de Micología; 2000. pp. 1-12
  8. 8.Garg AP, Gandotra S, Mukerji KG, Pugh GJF. Ecology of keratinophilic fungi. Proceedings / Indian Academy of Sciences. 1985;94:149-163. DOI: 10.1007/BF03053134
  9. 9.Kushwaha RKS. The genus Chrysosporium, its physiology and biotechnological potential. In: Kushwaha RKS, Guarro J, editors. Biology of Dermatophytes and Other Keratinophilic Fungi. 1st ed. Bilbao: Revista Iberoamericana de Micología; 2000. pp. 66-76
  10. 10.Ni H, Chen Q , Chen F, Ml F, Dong YC, Cai HN. Improved keratinase production for feather degradation by Bacillus licheniformis ZJUEL31410 in submerged cultivation. African Journal of Biotechnology. 2011;10:7236-7244
  11. 11.Gröhs Ferrareze PA, Folmer Correa AP, Brandelli A. Purification and characterization of a keratinolytic protease produced by probiotic Bacillus subtilis. Biocatalysis and Agricultural Biotechnology. 2016;7:102-109. DOI: 10.1016/j.bcab.2016.05.00
  12. 12.Lopes FC, Dedavid e Silva LA, Daroit DJ, Velho RV, Pereira JQ , et al. Production of proteolytic enzymes by a keratin degradingAspergillus niger. Enzyme Research. 2011;2011:487093. DOI: 10.4061/2011/487093
  13. 13.Syed DG, Lee JC, Li WJ, Kim CJ, Agasar D. Production, characterization and application of keratinase from Streptomyces gulbargensis. Bioresource Technology. 2009;100:1868-1871. DOI: 10.1016/j.biortech.2009.08.026
  14. 14.Kannahi M, Ancy RJ. Keratin degradation and enzyme producing ability of Aspergillus flavus and Fusarium solani from soil. Journal of Chemical and Pharmaceutical Research. 2012;4:3245-3248. DOI: 10.1139/w06-067
  15. 15.Sowjanya NC, Chary CM. Degradation of few avean feathers by Microsporum gypseum. Journal of Phytology. 2012;4:21-23
  16. 16.Vanbreuseghem R. Technique biologique pour 1’ isolement des dermatophytes du sol. Annales de la Societe belge de medecine tropicale. 1952;32:173-178
  17. 17.Ajello L. Natural history of the dermatophytes and related fungi. Mycopathologia et Mycologia Applicata. 1974;53:93-110
  18. 18.Hejtmanek M, Hejtmankova N, Kunert J. On the occurrence of geophilic dermatophytes in Asia. Ceska Mykol. 1973;27:139-161
  19. 19.Canizares O. Dermatology in India. Archives of Dermatology. 1976;112:93-97
  20. 20.Palsson G. Gcophilie dermatophytes in the soil in Sweden. Studies on their occurrence and pathogenic properties. Acta Veterinaria Scandinavica. 1968;25:89
  21. 21.Philpot CM. Geographical distribution of dermatophyte: A review. Epidemiology & Infection. 1978;80:301-314
  22. 22.Vanbreuseghem R, De Vroey C. Geographic distribution of dermatophytes. International Journal of Dermatology. Apr-Jun 1970;9(2):102-109. DOI: 10.1111/j.1365-4362.1970.tb04587.x. PMID: 5426624
  23. 23.Ajello L. The dermatophyteM. gypseumas a saprophyte and parasite. The Journal of Investigative Dermatology. 1953;21:157-171
  24. 24.Ajello L. Occurrence of Histoplasma capsulatum and other human pathogenic molds in Panamanian soils. The American Journal of Tropical Medicine and Hygiene. 1954;3:897-904
  25. 25.Alteras I, Feuerman EI, Grunwald M, Shvili D. Tinea capitis due to Mlicrosporum canis in infants. Mycopathologia. 1984;86:89-92
  26. 26.George LK. Animal Ringworm in Public Health. Publication No.727. Washington DC: United States Government Printing Office; 1960. p. 57
  27. 27.English MP, Morris P.Trichophyton mentagrophytesvar.erinaceiin hedgehog nests. Sabouraudia. 1969;7:118-121
  28. 28.Hubalek Z. Keratinophilic fungi or wild birds. Mykosen. 1972;15:207-211
  29. 29.Hubalek Z. Occurrence of keratinophilic fungi in nests of tree sparrow (f'esser montozirs) in relation to the substrate moisture. Ceska Mykol. 1976a;30:106-109
  30. 30.Hubalek Z. Interspecific a&nity among keratinolytic fungi associated with birds. Folia Parasital. 1976b;23:267-272
  31. 31.Pugh GJF. Dispersal ofArihroderma curreyiby birds, and its role in the soil. Sabowaudia. 1964;3:275-278
  32. 32.Pugh GJF. Associations between birds, nests. their pH and keratinophilic fungi. Sabouraudia. 1966b;5:49-53
  33. 33.Pugh GJF. Fungi on birds in India. Journal of the Indian Botanical Society. 1966a;4S:296-303
  34. 34.Pugh GJF. The contamination of birds of feathers by fungi. Ibis. 1972;14:172-177
  35. 35.Sur B, Ghosh GR. Keratinophilic fungi from Orissa, India. I. Isolation from soils. Sabouraudia. 1980;18:269-274
  36. 36.Alteras I, Nesterov V, Ciolofan I. The occurrence of dermatophytes in wild animals from Romania. Sabouraudia. 1966;4:215-218
  37. 37.Siniordova M, Hejtmanek M. Orraiatophytes and other keratinopbytic fungi in surface and waste waters. Mykosen. 1970;13:467-471
  38. 38.Balbaiioff VA, Usunov P. Organic waste matter of plant origin—Natural source ot’ primitive dermatophytes. I. Communication. Mycopathologia et Mycologia Applicata. 1967;33:43-48
  39. 39.Caretta G, Frate D. Funghi cheratinofilie dell’ isola di Montecristo: isolamenti da sualo. escrementi e pelo di arlimali e piume di uecelli. Chi Psi. Rot. Unit. Lah Criitogam Parlas. 1976;6:203-208
  40. 40.Battelli G, Bianchedi M, Frigo W, Amorati P, Mantovani AC, Pagliani A. Survey of keratinophilic fungi in alpine marmat (Marmata marmata) burrow soil and in adjoining soils. Sabouraudia. 1978;16:83-86
  41. 41.Pugh GJF, Allsopp D. Microfungi on Signy Island, South Orkney Islands. Br. infarct. Sum. Bull. 1982;57:55-67
  42. 42.Buchnicek J. Ober den einfluss der langwelligen UV-strahlung auf da’s wachstums von dermatophytes III. Symp. int. Dermetol. Bratislava Abs. 1966:10-16
  43. 43.Berde K. Verhalten von Fadenpilzkolonieo unter der Wirkung des Lichter. Archiv für Dermatologie und Syphilis. 1929;158:35-50
  44. 44.Dickey RF. Investigative studies in fungicidal powers of photodynamic action. Journal of Investigative Dermatology. 1961;39:7-19
  45. 45.Bohme H, Ziegler H. Verbreitung und Keratinophilie von Anixiopsis siercoraria (Hansen) Hansen. Archiv für Klinische und Experimentelle Dermatologie. 1965;223:422-428
  46. 46.Ziegler H. Ektoenzyme der dermatemyzeten. II. Miit. Arch. Klin. Exp. Dermatol. 1966;226:282-299
  47. 47.Chmel L, Hasilikova A, Hrasko J, Vlacilikova A. The influence of some ecological factors on keratinophilic fun•pi in soil. Sabouraudia. 1972;10:26-34
  48. 48.Kunert J. Utilization of L- and D-cystine by the fungus Microsporum gypseum. Folia Microbiologica. 1982;27:390-394
  49. 49.Kunert J. Inorganic sulphur sources for the growth of the dermatophytes Microsporum gypseum. Folia Microbiologica. 1981a;26:196-200
  50. 50.Kunert J. Organic sulphur sources for the growth of the dermatophyte Microsporum gypseum. Folia Microbiologica. 1981b;26:201-206
  51. 51.Hubalek Z, Rush-Munro FM. A dermatophyte from birds: Microsporum ripariae sp. nov. Sabauraudia. 1973;11:287-292
  52. 52.Ajello L, Padhye A. Ktratinophilic fungi of the Galapagos Islands. Mykosen. 1974;17:239-243
  53. 53.Das SK, Banerjee AB. Effect of undecanoic acid on phospholipid metabolism in Trichophyton rubnim. Sabouraudia. 1982;20:267-272
  54. 54.Hajini GH, Khandari KC, Mohapatra LN, Bhutani LK. Effmt of hair oil and fatty acids on growth of dermatophytes and their in uitro penetration of human scalp half. Sabowaudia. 1970;8:174-176
  55. 55.Padhye AA, Mishra SP, Thirumalachar MJ. Occurrence of soil inhabiting dermatophytes and other keratinophilic fungi from soil in Poona. Hindustan Antibiotic Bulletin. 1966;9:90-93
  56. 56.Pugh GJF. Cellulolytic and keratinophilic fungi recorded on birds. Sabouraudia. 1965;4:85-91
  57. 57.Ajello L, Varsavsky E, Satgin G, Mazzoni A, Mantovani A. Survey of soils for human pathogenic fungi from the Emilia-Romagna region of Italy. Mycopathologia et Mycologia Applicata. 1965;26:65-71
  58. 58.AcdaMN. Waste chicken feather as reinforcement in cement- bonded composites. Philippine Journal of Science. 2010;139:161-166
  59. 59.Han M, Luo W, Gu Q , Yu X. Isolation and characterization of a keratinolytic protease from a feather-degrading bacterium Pseudomonas aeruginosa C11. African Journal of Microbiology Research. 2012;6:2211-2222
  60. 60.Lin X, Lee CG, Casale ES, Shih JCH. Purification and characterization of a keratinase from a feather-degrading Bacillus licheniformis strain. Applied and Environmental Microbiology. 1992;58:3271-3275
  61. 61.Suneetha V, Lakshmi VV. Optimisation of parameters for fermentative production of keratinase by feather degrading microorganisms. World Journal of Microbiology and Biotechnology. 2005;7:106-115
  62. 62.Page RM. Observations on keratin digestion by Microsporumgypseum. Mycopathologia. 1950;42:591-602
  63. 63.Weary PE, Canby CM, Cowley EP. Keratinolytic activity of Microsporumcanis and Microsporumgypseum. Journal of Investigative Dermatology. 1965;44:300-310
  64. 64.Kunert J. Biochemical mechanism of keratin degradation by theactinomyceteStreptomyces fradiaeand the fungusMicrosporum gypseum: A comparison. Journal of Basic Microbiology. 1989;29:597-604
  65. 65.Yu RJ, Harmon SR, Blank F. Isolation and purification of an extracellular keratinase of Trichophyton mentagrophytes. Journal of Bacteriology. Oct 1968;96(4):1435-1436
  66. 66.Böckle B, Galunsky B, Müller R. Characterization of a keratinolytic serine proteinase from Streptomyces pactum DSM 40530. Applied and Environmental Microbiology. Oct 1995;61(10):3705-3710. DOI: 10.1128/aem.61.10.3705-3710.1995. PMID: 7487006; PMCID: PMC167669
  67. 67.Noval JJ, Nickerson WJ. Decomposition of native keratin by Streptomyces fradiae. Journal of Bacteriology. 1959;77:251-263
  68. 68.Kunert J. Keratin decomposition by dermatophytes, I: Sulfate production as a possible way of substrate decom- position. Zeitschrift für Allgemeine Mikrobiologie. 1973a;13(6):489-498
  69. 69.Kunert J. Keratin decomposition by dermatophytes. Zeitschrift für Allgemeine Mikrobiologie. 1973b;13:189-198
  70. 70.Ruffin P, Andrieu S, Biserte G, Biguet J. Sulfitolysis in keratinolysis: Biochemical proof. Sabouraudia. 1976;14:181-184
  71. 71.Stahl WH, MsQue B, Mandels GR, Siu GH. Studies on microbiological degradation of wool, I: Sulfur metabolism. Archives of Biochemistry. 1949;20:422-432
  72. 72.Chesters CGE, Mathison GE. The decomposition of wool keratin by Keratinomyces ajelloi. Sabouraudia. 1963;2:225-237
  73. 73.Ziegler H, Bohme H. Untersuchungen iiber den Haarabbau durch Dermatomyzeten. Dermat Wschr. 1963;148:429-454 Original not seen. Cited by Kunert, Z Allg Microbiol. 1973;13(6):489-498
  74. 74.Allsopp D, Seal KI. Introduction to Biodeterioration. London: Edward Arnold; 1986. pp. 1-136
  75. 75.Nigam N, Kushwaha RKS. Biodegradation of keratinous substrates. Biodeierior. of Cultural Propert y, Japan. 1992a:180-185
  76. 76.Nigam N, Kushwaha RKS. Biodegradation of wool by Chrysosporium keratinophilum acting singly or in combination with other fungi. Transactions of the Mycological Society of Japan. 1992b;33:481-486
  77. 77.Nigam N, Kushwaha RKS. Occurrence of non-keratinophilic fungi on keratin. Transactions of the Mycological Society Japan. 1989;30:l-8
  78. 78.English MP. The saprophytic growth of nonkeratinophilic fungi on keratinized substrate and a comparison with keratinophilic fungi. Transactions of the British Mycological Society. 1965;48:219-235
  79. 79.Kishimoto RA, Baker GE. Pathogenic and potentially pathogenic fungi isolated from beech sands and selected soil of Oahu, Hawaii. Mycologia. 1969;61:537-548
  80. 80.Carmichael JW. Chrysosporium and some other aleurosporic hyphomycetes. Canadian Journal of Botany. 1962;40:l137-l173
  81. 81.Orlita A. Occurrence of fungi on book leather binding from the baroque period. International Biodeterioration Bulletin. 1977;13:45-47
  82. 82.Kowalik R. Microbiodeterioration of library materials. Part-2 Micro biodecomposition of basic library materials. Restaurator. 1980;4:135-219
  83. 83.Deshmukh SK. Isolation of dermatophytes and other keratinophilic fungi from soil of Mussoorie (India). Mycoses. 1985;28:98-101
  84. 84.Garg AK. Isolation of dermatophytes and other keratinophilic fungi from soil of India. Sabouraudia. 1966;4:259-264
  85. 85.Jain M, Shukla PK, Shrivastava OP. Keratinophilic fungi and dermatophytes in Lucknow soils and their global distribution. Mykosen. 1985;28:148-153
  86. 86.Kushwaha RKS, Agarwal SC. Some keratinophilic fungi and related dermatophytes from soil. Proceedings of the Indian National Science Academy. 1976;42:102-110
  87. 87.Nigam N, Kushwaha RKS. Occurrence of keratinophilic fungi with special reference to Chrysosporium species in soils of India. Sydowia. 1990;42:200-208
  88. 88.Randhawa HS, Sandhu RS. A survey of soil inhabiting dermato- phytes and keratinophilic fungi of I ndia. Sabourauâia. 1965;4:71-79
  89. 89.Gaidau C. Applicative Chemistry of Tanning Metallic Heterocomplexes. Bucharest, Romania: Bentham Science Publishers; 2013
  90. 90.Koochakzaei A, Ahmadi H, Mohammadi Achachluei M. Leather making of Seljuk period at Qohestan of Khorasan (Skin and tanning characterization of leather objects excavated at Ghalee-Kuh-i Ghaen historic site). Quarterly Bulletin of Greater Khorasan. 2012;3(7):34-44 [In Farsi]
  91. 91.Ibn al-Athir E. The Complete History. Vol. 6. Tehran: Asaatir; 1995 [In Farsi]
  92. 92.Rajabi N. History and geography of the Ghaen city. Tehran: Shahr-i-Ashoub; 2005 [In Farsi]
  93. 93.Cameron E, Spriggs J, Wills B. The conservation of archaeological leather. In: Kite M, Thomson R, editors. Conservation of Leather and Related Materials. London: Butterworth-Heinemann; 2006. pp. 244-263
  94. 94.Kennedy CJ, Hiller JC, Lammie D, Drakopoulos M, Vest M, Cooper M, et al. Microfocus x-ray diffraction of historical parchment reveals variations in structural features through parchment cross sections. Nano Letters. 2004;4(8):1378-1380
  95. 95.Maxwell CA, Wess TJ, Kennedy CJ. X-ray diffraction study into the effects of liming on the structure of collagen. Biomacromolecules. 2006;7(8):2321-2326
  96. 96.Creanga DM. Novel aspects in leather covers conservation manuscripts from Punta monastery. European Journal of Science and Theology. 2006;2(2):91-97
  97. 97.Lama A, Antunes A, Covington A, Fletcher Y, Guthrie-Strachan J. Red rot in historic leather. Leather International. 2011;213(4814):34-38
  98. 98.Thomson R. Testing leathers and related materials. In: Kite M, Thomson R, editors. Conservation of Leather and Related Materials. London: Butterworth-Heinemann; 2006. pp. 58-65
  99. 99.Orlita A. Microbial biodeterioration of leather and its control: A review. International Biodeterioration & Biodegradation. 2004;53(3):157-163
  100. 100.Ebrahimi A, Karimi S, Lotfalian S, Majidi F. Allergenic fungi in deteriorating historic objects of Shahrekord Museum, in Iran. Jundishapur Journal of Microbiology. 2011;4(4):261-265
  101. 101.Polacheck I, Salkin IF, Schenhav D, Ofer L, Maggen M, Haines JH. Damage to an ancient parchment document by Aspergillus. Mycopathologia. 1989;106(2):89-93
  102. 102.Abdel-Maksoud G. Analytical techniques used for the evaluation of a 19th century quranic manuscript conditions. Measurement. 2011;44(9):1606-1617
  103. 103.Nigam N, Dhawan S, Nair MV. Deterioration of feather and leather objects of some Indian museums by keratinophilic and non-keratinophilic fungi. International Biodeterioration & Biodegradation. 1994;33(2):145-152
  104. 104.Griffin DM. Funal colonization of sterile hair in contact with soil. Transactions of the British Mycological Society. 1960;43:583-559
  105. 105.Karling JS. Keratinophilic chytrids. I. Rhizophydrum keratinophilum n. sp. , a saprophyte isolated on human hair and its parasite, Phylctidium mycetophagum n. sp. American Journal of Botany. 1946;33:751-757
  106. 106.Kunert J. Keratin decomposition by dermatophytes. Zeitschrift für Allgemeine Mikrobiologie. 1976;16:97-105
  107. 107.Kushwaha RKS, Agrawal SC. Microbial degradation of keratin. Proceedings of the National Academy of Sciences, India. 1981;51:181-183
  108. 108.Safranek WW, Goos RD. Degradation of wool by saprophytic fungi. Canadian Journal of Microbiology. 1981;28:137-140
  109. 109.Wainright MA. A new method for determining the microbial degradation of keratin in soil. Experimentia. 1982;38:243-244
  110. 110.Frei KM, Mannering U, Berghe IV, Kristiansen K. Bronze Age wool: Provenance and dye investigations of Danish textiles. Antiquity. 2017;91(357):640-654
  111. 111.Von Holstein ICC, Walton Rogers P, Craig OE, Penkman KEH, Newton J, Collins MJ. Provenancing archaeological wool textiles from medieval northern Europe by light stable isotope analysis (δ13C, δ15N, δ2H). PLoS One. 2016;10(11):e0162330. DOI: 10.1371/journal.pone.0162330

Written By

Manish Mathur and Neha Mathur

Submitted: October 8th, 2021Reviewed: February 15th, 2022Published: April 19th, 2022