2. REVIEW OF LITERATURE - Shodhgangashodhganga.inflibnet.ac.in/bitstream/10603/9644/9/09_chapter 2.pdf · The market research report "Vision for Indian Poultry Industry: Current Scenario

Exploration of Keratinolytic Actinobacteria for the Bioconversion of Poultry Feather…. Mr. SUBHASISH SAHA 8

2. REVIEW OF LITERATURE

2.1. World population

The world population is estimated to number 7.029 billion on March 2012 by

the United States Census Bureau (USCB) and it is approximately assessed that apart

from India and China, very low percentage of the global population abstains from meat,

poultry, fish and avoiding all animal products. A large study of vegetarians

(Vegetarianism in America, published by Vegetarian Times) during 2008 showed that

3.2 % of U.S. adults (7.3 million people) follow a vegetarian-based diet. About 0.5 %

(1 million) of those are vegans, who consume no animal products whatsoever. In

addition, 10 % of U.S. adults (22.8 million people) say they follow a semi-vegetarian

diet, which includes occasional consumption of fish. In other developed country the

vegetarian populations are as follows: United Kingdom 6%, Italy 10% and Germany

9%. According to the 2006 Hindu-CNN-IBN State of the Nation Survey, 31% of

Indians are vegetarians, while another 9% consumes eggs.

2.2. Global demand of dietary animal protein

Global demand for dietary animal protein is rapidly increasing, largely due to

increased prosperity and urban population growth in developing and transition

economies. Because of favourable nutrient conversion efficiency relative to beef and

pork, global poultry production is projected to double by the year 2030 to meet this

demand. The present distribution of poultry production, vast majority of the global

demand for poultry products will be in the form of chicken meat. Production to meet

the regional demand for duck (83%) and goose (93%) will remain centred in Asia,

whereas turkey is highly famous in North America and Europe (92%). Using global

meat demand during the years 1997–1999 as a base, it is estimated that by 2030,

demand will increase by 45 %, 57 % and 106 % for bovine, pork and poultry meat,

respectively (Harlan, 2007).

2.3. Global trade in chicken products

While produced across the globe, 13 % of chicken products consumed globally

are currently traded across national boundaries. The United States of America and

Brazil combined represent 76 % of global exports in 2005 and they are expected to be

Review of Literature


the future big exporters. This trade in chicken products is expected to increase due to

the higher demand in developing economies, many of which lack adequate resources

and conditions needed for cost-effective poultry production. Additionally, the relatively

high production costs in many developed nations will provide market opportunities for

more competitive poultry-production regions. As tariffs decline, countries with

abundant grain production, such as Brazil, are positioned to expand production further,

as they offer a favourable value proposition to global customers (Harlan, 2007).

2.4. Chicken production in Asia

Asian countries are leading the world economic recovery out of the recession

providing a positive backdrop to the key drivers to boost the demand for chicken –

population growth, rising disposable incomes, urbanisation and improved price

competition against competitors. Although population growth is slowing, the global

total continues to head towards 9.2 billion in 2050. Faster economic growth in the

developing countries is lowering poverty rates enabling more people to buy meat.

Worldwide, chicken meat production represents around 86 % of poultry meat

output, however, in Asia this figure is a little lower (Fig. 1). But, in China it dips as low

as 72 % because of the production of large quantities of duck and goose meat, the

combined output of which is around five million tonnes a year.

China is easily the leading producing nation in the region accounting for around

44 % of total output. However, this country's share has actually declined since 2000 -

when it stood at 48 % as other countries in the region have expanded production more

rapidly. For example, while China's output looks to have risen by almost 39 % during

the current decade to an estimated 12.6 million tonnes this year, the corresponding

figures for India, the region's second largest producer, reveals a much more dramatic

expansion of 145 % with production currently around 2.7 million tonnes. Apart from

India, other Asian countries with sizeable annual chicken meat output, recording faster

industry growth than China are Myanmar, Pakistan, Iran, Indonesia, the Yemen,

Bangladesh, Turkey, Israel and the Republic of Korea (Table 2).



Fig. 1. Chicken meat production in Asia and world (Global poultry trends, June, 2010)

2.5. Indian poultry industry

India is the fifth largest producer of eggs and ninth largest producer of poultry

meat in the world, producing over 34 billion eggs and about 600,000 tons of poultry

meat. In the overall market for poultry products, India was positioned 17 in world

poultry production and analysts estimate that the poultry sector in India has been

growing at a much faster rate, along with other industries such as BPO and securities

market. Over the past decade the poultry industry in India has contributed

approximately US $ 229 million, to the Gross National Product (GNP).

Several breakthroughs in poultry science and technology have led to the

development of genetically superior breeds capable of higher production, even under

adverse climatic conditions that offer opportunities for overseas entrepreneurs to

expand export and import of poultry products on a large scale.

The average per capita poultry meat consumption is also estimated to increase

from 0.69 to 1.28 kilograms, during the 2000-2004. Overall, analysts studied that the

total egg consumption is estimated to increase from 34 billion in 2000 and to 106

billion in 2020, while poultry meat consumption is predicted to increase from 687

million kilograms to 1,674 million kilograms.



Table 2. Chicken meat production rankings in Asia

(Tonns, upto year 2010)

China 12,550.0 India 2,650.0 Iran Islamic Rep. 1,600.0 Indonesia 1,467.1 Japan 1,265.0 Thailand 1,240.0 Turkey 1,155.0 Malaysia 987.0 Myanmar 800.0 Philippines 785.5 Pakistan 640.0 Saudi Arabia 590.0 Korea Rep. 536.8 Israel 475.0 Viet Nam 380.0 Syrian Arab Rep. 182.0 Bangladesh 171.0 Lebanon 142.0 Jordan 139.2 Yemen 128.7 Sri Lanka 105.0 Iraq 102.0 Singapore 84.1 Occupied Palestinian Territory 71.5 Kazakhstan 66.6 Azerbaijan 57.0 Kuwait 44.1 United Arab Emirates 37.8 Korea DPR 31.0 Cyprus 30.0 Uzbekistan 25.9 Afghanistan 21.6 Brunei Darussalam 20.1 Cambodia 19.3 Lao PDR 19.2 Nepal 17.3 Turkmenistan 12.9 Armenia 8.0 Kyrgyzstan 7.4 Georgia 7.0 Oman 5.9 Bahrain 5.3 Qatar 5.1 Timor – Leste 2.0 Tajikistan 1.1 Bhutan 0.3 Mongolia 0.3



The market research report "Vision for Indian Poultry Industry: Current

Scenario and Future Prospects" predicts a relatively strong growth for the egg and

poultry meat industry, in both the urban and rural areas, in the next two decades. It has

been found that egg consumption has grown at a much faster pace, than the

consumption of poultry meat. With the continual rise in income, it is estimated to

nearly triple by 2020 (Fig. 2). The report also examines the consumption pattern of egg

and poultry meat for 2010 by taking into account urbanization and differences in

consumption patterns across various income groups, both in urban and rural areas

(Fig. 3). The report deals in detail with the market structure, as well as highlights the

production, consumption, import/export statistics etc., of the Indian poultry market,

including broilers and processed poultry (Samarendu and Rajendran, 2003).

Fig. 2. Average per capita poultry meat consumption in India: Rural vs. Urban

(Samarendu and Rajendran, 2003)

Fig. 3. Average per capita egg consumption in India: Rural vs. Urban

(Samarendu and Rajendran, 2003)

2.6. Feather waste

Depending on the popularity of chicken, worldwide 24 billion chickens are

killed annually to fulfil a huge demand of food habit for the non-vegetarian population

and around 8.5 billion tonnes of poultry feather are produced. According to a recent

report in leading news paper India's contribution alone is 350 million tonnes. The

poultry feathers are dumped, used for land filling, incinerated or buried, which involves

problems in storage, handling, emissions control and ash disposal. Discarded feather

also causes various human ailments including chlorosis, mycoplasmosis and fowl

cholera (Agrahari and Wadhwa, 2010).



Feathers represent 5-7% of the body weight of the domestic fowl. Although

they are of insulator, locomotory and conformational (structural) importance to the

birds, possible biological uses outside the body of the birds appear sub-optimally

harnessed, while it seems probable that poultry feather constitutes the most abundant

keratinous material in nature. Poultry feather accumulates as a waste after processing

the chickens for human consumption; thus the waste carries potent polluting

implications especially with burgeoning global poultry production (Onifade et al.,

1998). Feathers are produced in large amounts as a waste by-product of poultry

processing plant (Riffel and Brandelli, 2006).

Poultry industry is continuously producing increasing amount of poultry meat

and noticeable quantities of organic residues such as feather, bone meal, blood, offal

and so on. Chicken feathers, making up about 5% of the body weight of poultry, are a

considerable waste product of the poultry industry being produced about 4 million tons

per year world-wide. Disposal of waste feathers is a major concern for poultry industry

and accumulation of this huge volume of the waste feathers results in environmental

pollution and protein wastage (Salminen and Rintala, 2002; Salminen et al., 2003).

2.7. Specific characteristic of chicken feather and keratin protein

Feather consists of different parts (Fig. 4a) and nearly pure keratin protein

(Moran et al., 1966). Keratins are the most abundant proteins in epithelial cells of

vertebrates and represent the major constituents of skin and its appendages such as nail,

hair, feather and wool (Fig. 4b). Keratins are grouped into hard keratins (feather, hair,

hoof and nail) and soft keratins (skin and callus) according to sulphur content.

They are a major class of structural proteins that are highly resistant to

biological degradation. Common enzymes, which break down protein, such as trypsin,

do not affect keratin. Keratins are insoluble in water. Keratins, like other proteins, are

made of a long string of various amino acids, which fold into a final three-dimensional

form. Two types of KRTs, α-KRTs and β-KRTs, consist of tightly packed protein

chains in α-helices and β-sheets, respectively (Parry and North, 1998).



Fig. 4 a. Different parts of feather Fig. 4 b. Intermolecular hydrogen

bonding in keratin

The composition and molecular configurations of keratin constituent amino

acids warrant structural rigidity (Fig. 5 a, b, c, d). At least 30 different keratin

polypeptides are known falling into 2 evolutionary families designated type I and type

II. Within each polypeptide chain, the helical rod domain of about 310 amino acids is

flanked by a shorter nonhelical head and tail domains, which are thought to have a

flexible conformation (Cohlberg, 1993).

Furthermore, Keratin filament structures are stabilized by their high degree of

cross-linking of disulfide bonds, hydrophobic interactions and hydrogen bonds. Due to

their extremely rigid structures, KRTs are insoluble and hard to degrade (Esawy, 2007).

Chicken feathers are composed of over 90% of keratin protein, small amounts of lipids

and water. Feathers keratin consists of high quantities of small and essential amino acid

residues (Pencho, 1990; Salminen and Rintala, 2002). Keratin is also the main protein

components of hair, wool, nails, horn and hoofs. Animal hair, hoofs, horns and wool

contain α-keratin and bird’s feather contains β-keratin. The polypeptides in α-keratin

are closely associated pairs of α helices, whereas β-keratin has high proportion of

β pleated sheets. “This conformation confers an axial distance between adjacent

residues of 0.35 nm in β-sheets, compared to 0.15 nm in α-helices. The β sheets have a



far more extended conformation than the α helices” (Asquith, 1977; Morris et al., 1992;

Savitha et al., 2007). Keratins are insoluble macromolecule comprises a tight packing

of supercoiled long polypeptide chains with a molecular weight of approximately 10

kDa. High degree of cross linked cystine disulfide bonds between contiguous chains in

keratinous material imparts high stability and resistance to degradation (Schmidt and

Barone, 2004; Coward-Kelly et al., 2006; Tamilmani et al., 2008; Weidele, 2009).

Hence, a keratinous material is a tough, fibrous matrix being mechanically firm,

chemically unreactive, water insoluble and protease-resistant (Savitha et al., 2007).

Such a molecular structure makes feathers poorly degradable under anaerobic digestion

condition (Salminen and Rintala, 2002; Weidele, 2009).

Fig. 5. The composition and molecular configurations of keratin

(a) Sub-domain structure of epidermal keratin chains showing the basic short end regions E1 and E2, the variable glycine/serine-rich regions V1,V2 and the homologous regions H1 and H2 (Steinert, 1993).

(b) Sub-domain structure of hard α-keratin chains showing the basic (NB) and acidic (NA) regions of the N-terminal domain. The C-terminal domain of the type I chains is characterized by a repeated proline–cysteine–X motif. The C-terminal domain of type II chains contains a periodic distribution of hydrophobic residues (Parry and North, 1998).

(c) Model structure of keratin coiled-coil dimer, 45 nm in length. The hydrophobic amino acids of the two α-helices are meshed together in a regular interlocking pattern (Cohlberg, 1993).

(d) Organization of keratin micro fibrils, showing the globular head and tail domains (in black). The terminal domains can interact with segments in the rod domain and with other C domain in an antiparallel neighbouring molecule (Parry and North, 1998).



2.8. Composition of feather keratin

The average young feather consists of the following chemical substances:

moisture (6.72%); protein (81.46%); fat (11.36%); fiber (0.31%). Nitrogen is the most

abundant element present in the feather. In protein, almost all different amino acids are

present in feather such as taurine 0.01%, hydroxyproline 0.23%, aspartic acid 5.33%,

threonine 3.70%, serine 7.88%, glutamic acid 8.13%, proline 8.14%, lanthionine

1.65%, glycine 6.25%, alanine 3.57%, cysteine 4.99%, valine 6.28%, methionine

0.57%, isoleucine 3.79%, leucine 6.59%, tyrosine 2.33%, phenylalanine 3.97%,

hydroxylysine 0.01%, histidine 0.61%, ornithine 0.30%, lysine 1.79%, arginine 5.68%

and tryptophan 0.47%. Other elements found in feathers are sulphur (2.57%), chlorine

(0.53%), phosphorus, in the form of pentoxide (0.34%), silicon, in the form of silicic

acid (0.22%) and calcium, as oxide (0.10%). A crystalline sulphuric amino acid called

cystine can be extracted from feathers. One kind of feathers may differ slightly from

another in its chemical compositions. For example the fat content of the feathers of

geese and ducks is greater than that of the feathers of hens and turkeys.

2.9. Utilization of feather waste

Currently a minor quantity of waste feathers is used in other industrial

applications such as clothing, insulation and bedding (Poopathi and Abidha, 2007),

producing biodegradable polymers (Schmidt and Barone, 2004) and enzymes (Casarin

et al., 2008) and also as a medium for culturing microbes.

Anaerobic digestion is an environmentally and economically promising process

to recover feather waste and other solid organic wastes to valuable materials such as

biogas and fertilizers (Salminen and Rintala, 2002). However, slaughterhouse wastes

are in general considered as difficult substrates for anaerobic digestion because of their

high protein and lipid content leading in production of some by-products such as

unionised ammonia, floating scum and accumulated log chain fatty acids (LCFA)

during anaerobic degradation, which are toxic and inhibitory to anaerobic

microorganisms in high concentrations (Seisle, 2008). Such practical difficulties have

limited and hindered the successful efforts on anaerobic digestion of feathers and other

solid poultry slaughterhouse wastes (Salminen and Rintala, 2002).



A higher quantity of pretreated feather is utilized to produce a digestible dietary

protein feedstuff for poultry and livestock (Papadopoulos et al., 1985; Onifade et al.,

1998; Coward-Kelly et al., 2006; Tamilmani et al., 2008; Weidele, 2009).

Understandably, poultry feather locks up a great deal of potentially useful protein and

amino acids that could be beneficially harnessed as animal feedstuff. This makes

recycling of feather a subject of interest among animal nutritionists, because of its

potential as a cheap and alternative protein feedstuff. However, limitations to feather

utilization arise from its poor digestibility and low biological value and the deficiencies

of nutritionally essential amino acids such as methionine, lysine, histidine and

tryptophan (Baker et al., 1981; Papadopoulos et al., 1985; Dalev, 1994).

2.10. Conventional methods for feather degradation

Nevertheless, the conventional method of producing a more readily digestible

feather meal employed in hydrothermal degradation. But according to Papadopoulos

(1985), Latshaw et al., (1994) and Wang and Parsons (1997), hydrothermal treatment

achieves limited and varying nutritional improvement; sustains losses of essential

amino acids such as lysine, methionine and tryptophan and causes the formation of

non-nutritive amino acids such as lysinoalanine, lanthionine, etc (Table 3). Other side

chemical treatment of feather leads toxic end product which is equally useless. Beside

that considering the thermo energetic cost of conventional processing of feather against

the backdrop of its limited nutritional improvement, investigation into alternative

technology with prospects for nutritional enhancement, environmental friendliness or

compatibility, bioresources optimization and cost effectiveness seems justifiable.

2.11. Biodegradation of feather waste

Biotechnological approaches involving microorganisms and their enzymes

appear a conceptually appropriate processing technology. However, there is no

compendious literature on the prospects for industrial applications of keratinolytic

microorganisms, especially with emphasis on their production of keratinases, properties

of keratinases, mechanism and limitations of keratinolysis.



Table 3. Conventional methods for feather degradation looses essential amino acid during feather processing:

Protein and amino acid concentrations (g kg-l) of unprocessed and processed feather meal (Onifade et al., 1998)

S. No. Protein and amino acids

Latshaw et al., (1994) unprocessed

Latshaw et al., (1994) processed at 207 kPa for 24 min

Wang and Parsons (1997) processed at 160°C for 15 min

1. Protein 922.0 866.0 880.0 2. Alanine 28.8 37.7 39.6 3. Glycine 51.8 50.7 68.7 4. Isoleucine 39.4 41.3 42.3 5. Leucine 56.9 68.8 70.9 6. Valine 53.0 44.0 59.6 7. Phenylalanine 34.6 40.1 42.1 8. Arginine 67.6 62.5 61.0 9. Histidine 2.3 8.6 5.7

10. Lysine 15.4 22.6 18.8 11. Aspartic acid 41.8 55.9 55.2 12. Glutamic acid 82.2 72.3 97.2 13. Serine 87.3 72.1 100.0 14. Threonine 34.5 36.5 40.2 15. Proline 73.9 74.8 88.4 16. Cystine 65.8 48.7 42.9 17. Methionine 7.1 6.3 6.5



Therefore, we reviewed recent information on microbial keratinolysis in order to

stimulate the application of the biotechnology in feather processing as animal feedstuff.

The upgrading of feather nutritional value should yield a high-protein feedstuff that

may greatly spare the use of soyabean and fish meal in livestock diets. Furthermore,

bioconversion of feather will predictably benefit the poultry industry, man and the

environment.

2.12. Microbial proteolytic system

Proteases are essential constituents of all forms of life on earth, including

prokaryotes, fungi, plants and animals. They can be cultured in large quantities in a

relatively short time by established methods of fermentation and they also produce an

abundant, regular supply of the desired product. Microorganisms account for a two-

third share of commercial protease production in the world (Kumar and Takagi, 1999).

Microbial proteases are classified into various groups, dependent on whether they are

active under acidic, neutral or alkaline conditions and on the characteristics of the

active site group of the enzyme, i.e. metallo, aspartic, cysteine, sulphydryl, serine-type

(Kalisz, 1988; Rao et al., 1998). Alkaline proteases are defined as those proteases

which are active in a neutral to alkaline pH range. They either have a serine center

(serine protease) or are of metallo-type (metalloprotease) and the alkaline serine

proteases are the most important group of enzymes exploited commercially (Gupta et

al., 2002).

2.13. Sources of microbial keratinases

The amino acids composition of feather is highly variable (Wang and Parsons,

1997). Also, the total essential amino acids, especially, methionine, lysine and histidine

concentrations decreased with age in broiler chickens, while the total non-essential

amino acids as a percentage of total essential amino acids, increased as the birds aged

(Stillborn et al., 1997). Keratinase from different microbial source has been listed in

Table 4. The nutritional inferiority of native feather protein derives from the

composition and molecular configurations of constituent amino acids that are,

basically, to ensure structural rigidity for the role of feathers. The same reason explains

why native keratin is insoluble and undegradable by most proteolytic enzymes.



Feathers mechanical stability and resistance to proteolytic digestion are consequences

of the tight packing of the protein chain in the α-helix (α-keratin) or β-sheet (β-keratin)

into a supercoiled polypeptide chain. There is a high degree of cross linking of the

polypeptide chain caused by extensive formation of disulfide bonds. The high content

of cystein facilitates the formation of cystine bridges. Hydrogen bonding among the

polypeptides and the hydrophobic interaction and stabilization of the super coil further

confer strength and proteolytic resistance on keratin.

Keratinases are very widespread in the microbial world and they can be

identified from microorganisms of the three domains: Eucarya, Bacteria and Archaea.

These microorganisms have been isolated from the most distinct locations, from

Antarctic soils to hot springs, including aerobic and anaerobic environments. Therefore,

microbial keratinases present a great diversity in their biochemical and biophysical

properties.

2.14. Fungal keratinases

In natural environments, keratinolytic fungi are involved in recycling the

carbon, nitrogen and sulphur of the keratins. Their presence and distribution seem to

depend on keratin availability, especially where humans and animals exert strong

selective pressure on the environment (Filipello, 2000). A number of reports focused on

the keratinolytic potential of dermathophytic fungi such as Trichophyton and

Microsporum (Asahi et al., 1985; Qin et al., 1992; Filipello, 2000; Moallaei et al.,

2006), mainly due to their medical and veterinary implications. Although some studies

on the biotechnological potential of such genera are available (Anbu et al., 2008), little

commercial interest was attracted by this group because of their potential pathogenicity

(Gradisar et al., 2000; Blyskal, 2009). Among nondermatophytic fungi, keratinases

showing attractive biochemical properties were reported to be produced by Aspergillus

(Santos et al., 1996; Farag and Hassan, 2004), Trichoderma (Cao et al., 2008),

Doratomyces (Gradisar et al., 2000), Myrothecium (Moreira et al., 2009), Paecilomyces

(Gradisar et al., 2005), Scopulariopsis (Anbu et al., 2005) and also Acremonium,

Alternaria, Beauveria, Curvularia and Penicillium (Marcondes et al., 2008). Besides

the biotechnological interest, these investigations may help in understanding the role of



fungi in the degradation of complex keratinous substrates in the nature (Marcondes et

al., 2008).

2.15. Bacterial keratinases

Several keratinases have been isolated from a diversity of bacteria. Bacillus spp.

appears as the prominent keratinase producer. Diverse strains of Bacillus licheniformis

and Bacillus subtilis were described as keratinolytic (Lin et al., 1999; Suh and Lee,

2001; Manczinger et al., 2003; Balaji et al., 2008; Cai et al., 2008; Zhang et al., 2009a),

but other species such as Bacillus pumilus and Bacillus cereus also produce keratinases

(Kim et al., 2001; Werlang and Brandelli, 2005; Kumar et al., 2008; Ghosh et al.,

2008). Furthermore, B. licheniformis (Lin et al., 1992) is the source of Versazyme, the

first thermo-resistant commercial keratinase developed by Shih and co-workers at

Bioresource International, Inc. in the year 2000. Some thermophilic and alkaliphilic

strains of Bacillus have also been described to show keratin-degrading activity, such as

Bacillus halodurans AH-101 (Takami et al., 1992 and 1999), Bacillis pseudofirmus

AL-89 (Gessesse et al., 2003) and B. pseudofirmus FA30-01 (Kojima et al., 2006).

2.16. Actinobacterial keratinases

Keratinase producers have been also described among actinobacteria, mainly

from the Streptomyces genus. These microorganisms, isolated from several different

soil sites, were associated with the hydrolysis of a wide range of keratinous substrates

like hair, wool and feathers. For example, two highly keratinolytic actinobacterial

strains, Streptomyces flavis 2BG (mesophilic) and Microbispora aerata IMBAS-11A

(thermophilic) were isolated from Antarctic soil (Gousterova et al., 2005). The

thermophilic species Streptomyces gulbarguensis (Syed et al., 2009), Streptomyces

thermoviolaceus (Chitte et al., 1999) and Streptomyces thermonitrificans (Mohamedin,

1999) have also been isolated from soils. Besides these thermophilic strains, some

mesophilic Streptomyces have also been characterized like Streptomyces pactum DSM

40530 (Bockle et al., 1995), Streptomyces graminofaciens (Szabo et al., 2000) and

Streptomyces albidoflavus K1-02 (Bressollier et al., 1999).



Mohamedin, (1999) investigated on protease producing thermophilic

Streptomyces strain grown on chicken feather as a substrate from the soil of Mansoura

city, Egypt. Thermophilic actinobacteria was isolated from soil enriched with

hydrolysed wool waste (Gousterova et al., 2003). Streptomyces sp. S7 isolated from

slaughterhouse waste samples from Hyderabad, India by Radhika et al., (2008). A

keratinolytic Streptomyces sp. was isolated from poultry processing plant waste water

from Bazil (Tapia and Simoes, 2008). A novel keratin degrading actinobacterium

Actinomadura keratinilytica sp. isolated from bovine manure compost in China (Puhl

et al., 2009). A new Streptomyces sp. IF 5 was isolated from the feather dumped soil at

Thanjavur, Tamil Nadu, India (Ramakrishnan et al., 2011).

2.17. Actinobacteria

Actinobacteria are filamentous, branching bacteria with fungal type

morphology. They are part of the microbial flora of most natural substrates. Numerous

methods have been advocated to facilitate the isolation of actinobacteria and to separate

them from their relatives. It is not difficult to isolate actinobacteria from an intimate

mixture with fungi, since the physiological properties of these two groups of

microorganisms are different. For example, strictly antifungal antibiotics, which do not

affect the growth of actinobacteria, can be used successfully. It is more difficult to

separate actinobacteria from true bacteria. Nevertheless, some selective media have

been suggested and also various means for increasing the actinobacterial population of

the soil samples before the plating (Moustafa and Hubert, 1962).

2.17.1. Pre-treatment and preparation of soil suspensions

Calcium carbonate mixed with air dried soil enhances the explorations of

actinobacterial selective isolation (Tsao et al., 1960). Likewise sodium propionate was

added in a 0.4% (w/v) concentration to the arginine glycerol salt (AGS) medium before

sterilization of soil samples for actinobacterial isolation (Crook et al., 1950). Heat

treatment and phenol treatment were also applied in several studies to reduce the

unwanted microbial population while isolating actinobacteria. Soil suspensions for

actinobacteria isolation were prepared by serially diluting 1 g soil sample and



vigorously shaking in 10 ml of sterile distilled water for 30 min on a shaker. Serial

dilutions were used upto 10-6 dilution (Rehacek, 1959).

2.17.2. Isolation of soil actinobacteria

During the several years, research community has been using different medium

for the isolation of soil actinobacteria. Some of the popular medium for actinobacterial

isolation used were starch casein agar (SCA), arginine glycerol salt (AGS), chitin

medium (Lingappa and Lockwood, 1962); modified Benedict’s medium (Porter et al.,

1960); soybean meal glucose medium (Tsao et al., 1960); Bennett’s agar, complete

medium, Gauze’s agar medium (Rehacek, 1959); egg albumin medium, glucose-

asparagine medium and glycerol asparagine agar II (Waksman, 1961).

Keratinolytic actinobacterial isolates were also isolated from various soil

samples. Chitte et al., (1999) isolated keratinolytic feather-degrading thermophilic

Streptomyces thermoviolaceus SD8 from Lonar lake soil, a meteoritic crater situated in

a tectonic zone of Western Maharashtra using glucose yeast extract peptone agar.

Samples of agricultural soil collected from Mansoura city (Egypt) were

enriched with feather pieces and incubated at 50°C. Microbial growth that became

established on the pieces of feather within 2 weeks was scraped off and isolated on

starch casein plates. Plates were incubated at 50°C for 4 days and single colonies were

picked and transferred to a separate plate (Mohamedin, 1999).

Actinobacterial isolates with antimicrobial activity were obtained from saltpan

regions of Cuddalore and Parangipettai, Tamil Nadu, India by Dhanasekaran et al.,

(2005). In other subsequent studies during 2009, they investigated about 189

Streptomyces isolates from eight different soils of Cuddalore, Tamil Nadu, India and

among them, they found only 78 isolates were morphologically distinct (Dhanasekaran

et al., 2009a).



An aerobic mesophilic saprophytic actinobacterium was isolated from a

mediterranean sea (20-35ºC) nearby sea shore, Aexandria. Isolation and enrichment of

the keratin degrading microorganism was performed at 30ºC and pH 8.5 in a medium

containing KCl, MgSO4.7H2O, NH4Cl and 1g native chicken feather in 250 Erlenmeyer

flasks containing 50 ml sea water (Esawy, 2007).

Thermophilic actinobacteria was isolated from soil enriched with hydrolysed

wool waste using peptone, maize extract, starch, NaCl and CaCO3 containing medium

(Gousterova et al., 2003). Alkaliphilic keratinolytic enzyme producing Nocardiopsis sp.

TOA-1 isolated on skim milk and yeast extract medium in Japan by Mitsuiki et al.,

(2004). A new Streptomyces sp. IF 5 with tremendous keratinase activity was isolated

from the feather dumped soil Thanjavur, Tamil Nadu, India and same strain found

enable to degrade the chicken feathers very effectively in 60 h (Ramakrishnan et al.,

2011).

2.17.3. Screening of keratinolytic actinobacteria

A qualitative screening for the proteolytic activity of the actinobacterial isolates

was indicated by growth and clear zones appearance on casein agar media

(Mohamedin, 1999). Milk agar medium was used for the primary screening of

keratinolytic actinobacteria (Riffel and Brandelli, 2006). Medium supplemented with

cut sheep skin and wool wastes were used as a sole source of carbon and nitrogen for

screening the keratinolytic thermoactinobacteria by Gousterova et al., (2005).

In recent past Ningthoujam et al., (2009) reported on screening of extracellular

protease production from moderately halophilic alkalithermotolerant Nocardiopsis

prasina HA-4 from biotopes in Manipur, India was done on skim milk agar and

enzyme assay was performed by photographic film clearing, agar cup plate assay (Hsu

and Lockwood, 1975) and Hagihara-Anson method (Dingle et al., 1953).



2.17.4. Characterization of actinobacteria

2.17.4.1. Actinobacterial taxonomy

Taxonomy is the science of the classification of living organisms. There are

three important reasons for classifying organisms. Firstly, classification is a form of

database or information retrieval system containing a large amount of information

about an organism. Secondly, classification is important because organisms must be

categorized into groups before identification systems can be created for new isolates.

Thirdly, classification systems may provide an insight into the origins and evolutionary

pathways of organisms (Priest and Austin, 1995). A number of different methods have

been used to classify actinobacteria. These include morphological, biochemical and

genomic methods. The taxonomy of actinobacteria is, however, still evolving and the

taxonomic status of many taxa is currently being re-evaluated (Chiba et al., 1999).

Actinobacterial identification and characterization generally carried out based on

colony morphology, microscopic examination, biochemical features and molecular

conserved gene analysis.

2.17.4.2. Morphological and physiological methods

Actinobacteria have a wide range of morphologies, many of which can be used

in classification. Actinobacterial taxonomy was traditionally based on morphology and

some of the characteristics most considered included the size, shape and colour of

colonies on specific media. Gram’s stain, acid fastness, odor and pigment production

are also used when classifying using morphology. Other morphological features that

are taxonomically important include the colour, morphology and surface arrangement

of conidiospores (Shirling and Gottlieb, 1966). These techniques are more accurate on

samples that have been freshly isolated. Physiological attributes such as nutritional

requirements, fermentation products and growth conditions (oxygen, temperature and

inhibitory products) are also important when classifying actinobacteria (Ciantar et al.,

2005). Waksman (1940) classified actinobacteria based on the mycelium as (a)

mycelium rudimentary or absent, (b) true mycelium produced and (c) vegetative

mycelium normally remains undivided.



Waksman (1961) characterized 15 series based on the following properties viz.,

morphological structure notably structure of sporophores and spores, colour of aerial

mycelium on synthetic medium, formation of soluble brown pigment on protein media,

spore surface and other characteristics like colour of substrate growth, formation of

soluble non-melanoid pigments and rate of proteolysis or production of specific

antibiotics.

Actinobacterial isolates used to identify according to Bergey’s Manual of

Determinative Bacteriology (1974) and keys proposed by Shirling and Gottlieb (1972).

The cultural, morphological and physiological characteristics of the actinobacteria were

studied and well described by Shirling and Gottlieb (1966). Also electron microscopic

characterization of actinobacteria used to give an exact idea about the shape, size,

spores morphology, spores arrangement of different species of actinobacteria.

Macroscopically the actinobacterial isolates were differentiated by their colony

characters, e.g. size, shape, colour, consistency etc. For the microscopy, the isolates

were grown by cover slip culture method (Kawato and Sinobu, 1979). They were then

observed for their mycelial structure, conidiospores and arthrospore arrangements on

the mycelia under microscope. The observed morphology of the isolates was compared

with the actinobacterial morphology provided in Bergey’s Manual for the presumptive

identification of the isolates (Gurung et al., 2009).

Krasilnikov (1960) considered pigment production as a constant specific

property, although the nature of the pigments varied with the composition of the

medium. The specificity of sugar, amino acid combination for the optimal brewing of

tyrosine might be used as criteria in the classification, which is similar to and

comparable with that of utilization of nitrogen compound (Kuster and Locci, 1963).

Hydrolysis of starch, casein and liquefaction of gelatin were also considered as

important characters of actinobacteria (Gordon and Smith, 1955; Waksman, 1961;

Gottlieb, 1961).



Subsequently Pridham and Gottlieb (1948) tested 27 cultures of Streptomyces,

Nocardia and Micromonospora for their ability to utilize 33 different carbon sources in

a chemically defined medium. The actinobacteria prefers yeast extract, peptone, urea,

asparagine, ammonium sulphate and sodium nitrate in the order of preference for their

growth (Pridham and Gottlieb, 1948; Okami, 1952).

Acid production from lactose, maltose, xylose and mannose was used as one of

the criteria for differentiating among the different species of actinobacteria. The

utilization of acetate, malate, propionate, pyruvate and succinate were considered as

specific characteristics of Streptomyces (Gordon and Mihm, 1959). Waksman (1959),

Jones and Bradley (1964) found that only a few actinobacteria used cellulose as source

of carbon and few Nocardia utilized phenol. Slack et al., (1971) studied the

morphological, biochemical and serological characters of 64 strains of actinobacteria

which found to be acid producer from various carbon sources.

Dhanasekaran et al., (2009b) reported about detailed morphological,

physiological, chemotaxonomic characterization of actinobacteria isolated from Vellar

Estuary, Annagkoil, Tamil Nadu, India. In 2009, Gurung et al., characterized

antimicrobial actinobacteria isolated from soil samples of Kalapatthar, Mount Everest

region based on various biochemical tests such as catalase, oxidase, citrate utilization,

nitrate reduction, starch hydrolysis, tween 20 hydrolysis, urea hydrolysis, gelatin

hydrolysis, esculin hydrolysis, acid production from sugar and the physiological test

included motility, NaCl resistance and temperature tolerance.

2.17.4.3. Molecular characterization

Identification of actinobacteria using microscopic techniques alone was not

enough to ensure certainty. Biochemical methods would be the best method to identify

actinobacteria to their species level. But this test consumes a lot of time and chemicals.

With the advancement of technology in molecular study, primers had been developed

by researchers to target specifically the 16S rRNA sequence of the actinobacteria and

that made identification of actinobacteria to genus level was made possible in a fast and

accurate manner (Schwieger and Tebbe, 1998; Wang et al., 1999; Jeffrey, 2008).



The comparison of the DNA nucleotide sequences of two strains provides a

rapid and accurate method of establishing relatedness. Techniques for carrying out the

comparisons include DNA-DNA hybridization and PCR based gene analysis. The

analysis of RNA for taxonomic purposes focuses on three different molecules of

ribosomal RNA, 5S (~120 nucleotides), 16S (~1600 nucleotides) and 23S (~3000

nucleotides). These molecules are important indicators of relatedness of organisms

because the rRNAs are essential elements in protein synthesis and are therefore present

in all living organisms (Priest and Austin, 1995). Other factors that make these

molecules ideal for the analysis of evolutionary relationships are that i) the lateral

transfer of rRNAs between different organisms is extremely rare ii) the longer rRNAs

(16S, 18S and 23S) contain regions of highly conserved, moderately variable and

highly variable sequences. The conserved regions are essential as they provide primer

directed sites for PCR as well as convenient hybridization targets for the cloning of

rRNA genes (Letowski et al., 2004; Gentry et al., 2006).

16S rRNA is a major component of the small (30S) ribosomal subunit. It is

important for subunit association and translational accuracy. The 16S rRNA gene,

consisting of 1542 bases, is highly conserved among microorganisms and is therefore

an excellent tool for studying phylogenetic relationships (Sacchi et al., 2002). PCR-

based methods are considered to be a rapid and accurate way of identifying bacteria

(Cook and Meyers, 2003). In sequence based techniques, primers to the extremities of

the gene are used to amplify the DNA. The amplified DNA can either be sequenced

directly or cloned into a phage or plasmid vector prior to sequencing. After the

sequences have been generated they are compared by aligning the corresponding

nucleotide sites. These type of simple comparisons of sequence positions will provide

an estimate of how closely related the organisms are (Priest and Austin, 1995).

Analysis of the 16S rRNA gene offers a time saving alternative to the classical methods

of identification summarised above (Alfaresi and Elkosh, 2006).

16S rRNA sequencing is very valuable in clinical settings such as for the

accurate identification of Nocardia species. Identification of Nocardia isolates to the

species level is very important for the estimation of pathogenicity, virulence and in



predicting how susceptible a strain will be to antimicrobial agents (Roth et al., 2003).

Other medically important actinobacteria that can be identified using 16S rRNA

sequencing include Actinomadura, Gordonia, Rhodococcus, Saccharomonospora,

Saccharopolyspora, Streptomyces and Tsukamurella (Cook and Meyers, 2003).

16S rRNA gene analysis has been used to reclassify actinobacteria species that

were incorrectly classified using classical identification methods. An example is the

reclassification of the actinobacteria strain ATCC 39727 which produces the

glycopeptides antibiotic A40926. This actinobacteria was originally classified on the

basis of morphology and cell wall composition into the genus Actinomadura. However,

phylogenetic analysis revealed that the strain ATCC 39727 belongs to the genus

Nonomuraea (Monciardin and Sosio, 2004).

Although any one approach used to assess diversity cannot claim to be more

efficient than another, 16S rRNA gene sequence analysis allows for the assessment of a

broader range of diversity than that obtained by physiological studies (Brambilla et al.,

2001). The 16S rRNA gene can also be analysed by a number of non-sequence based

methods which include amplified rDNA restriction analysis (ARDRA), restriction

fragment length polymorphisms (RFLP), random amplified polymorphic DNA analysis

(RAPD), amplified fragment length polymorphisms (AFLP) and rep-DNA (Gurtler and

Mayall, 2001).

Streptomyces coelicolour A3 and Streptomyces lividans 66 were examined for

their morphological and cultural characteristics as well as DNA-DNA relatedness in

comparison with other Streptomyces coelicolour and Streptomyces lividans in order to

clarify their taxonomic status (Hatano et al., 1994). Song et al., (2001) reported the

phylogenetic diversity of thermophilic actinobacteria and Thermoactinomyces sp.,

isolated from mushroom composts in Korea, based on 16S rRNA gene sequence

analysis. The cultivation and independent population analysis of bacterial endophytes

in three potato varieties based on eubacterial and actinobacterial specific PCR of 16S

rRNA gene have been reported (Sessitsch et al., 2002). Phylogenetic analysis of

Nocardiopsis quinghaiensis sp. nov., isolated from saline soil in China was reported by

Chen et al., (2008).



Suihko et al., (2009) studied about 122 bacterial isolates from water damaged

building material and among them they identified actinobacteria or

thermoactinobacteria were present in 48% of the samples based on 16S rRNA gene

analysis. The dominant genus was Streptomyces (58% of isolates), followed by

Thermoactinomyces (23%), Laceyella (14%), Nocardiopsis (3%), Pseudonocardia

(1%) and Saccharomonospora (1%). Dhanasekaran et al., (2012b) was reported the

phylogenetic analysis and predicted the structure of 16S rDNA gene for soil

Streptomyces sp. Similarly Ramakrishnan et al., (2011) identified a feather degrading

keratinolytic Streptomyces sp. IF5 based on 16S rRNA gene analysis.

2.18. Keratinases from other microbes and archaea

In addition to these Bacillus spp. and actinobacteria, keratinase production has

been associated to an increasing number of bacteria. Since keratin degradation is

facilitated at high temperatures, pH and thermostable hydrolases are employed in

various industrial processes, the thermophilic and alkaliphilic microorganisms are of

great interest. Fervidobacterium pennavorans (Friedrich and Antranikian, 1996),

Fervidobacterium islandicum (Nam et al., 2002), Meiothermus ruber H328 (Matsui et

al., 2009), Clostridium sporogenes (Ionata et al., 2008) and strains of

Thermoanaerobacter sp. (Riessen and Antranikian, 2001; Kublanov et al., 2009) were

isolated from extreme environments like hot springs, geothermal vents, solfataric muds

and volcanic areas. Some alkaliphilic strains such as Nesternkonia sp. (Gessesse et al.,

2003) and Nocardiopsis sp. TOA-1 (Mitsuiki et al., 2002) have been also characterized,

showing keratinase activity in strongly alkaline pH. Several feather degrading bacterial

strains have been isolated from soil, poultry wastes and other sources, were

characterized as mesophilic keratinase producers. These include some Gram positive,

such as Lysobacter NCIMB 9497 (Allpress et al., 2002), Kocuria rosea (Bernal et al.,

2006) and Microbacterium sp. kr10 (Thys et al., 2004) and a few Gram negative, such

as Vibrio sp. (Sangali and Brandelli, 2000b), Xanthomonas maltophilia (De Toni et al.,

2002), Stenotrophomonas sp. (Yamamura, et al., 2002; Cao et al., 2009),

Chryseobacterium sp. (Riffel and Brandelli, 2002; Wang et al., 2008) and Serratia sp.

(Khardenavis et al., 2009).



Table 4. Diversity of keratinolytic microorganisms and some biochemical properties of keratinases

Microorganism Catalytic type Molecular

mass (kDa) Optimal pH Optimal T (°C)

Reference

Bacillus cereus DCUW Serine 80 8.5 50 Ghosh et al., (2008)

Bacillus licheniformis FK14 Serine 35 8.5 60 Suntornsuk et al., (2005)

Bacillus licheniformis MSK103 Serine 26 9–10 60–70 Yoshioka et al., (2007)

Bacillus licheniformis RPk Serine 32 9.0 60 Fakhfakh et al., (2009)

Bacillus pumilis Serine 65 8.0 65 Kumar et al., (2008)

Bacillus subtilis KD-N2 Serine 30.5 8.5 55 Cai et al., (2008)

Bacillus subtilis MTCC (9102) Metallo 69 6 40 Balaji et al., (2008)

Bacillus subtilis RM-01 Serine 20.1 9 45 Rai et al., (2009)

Clostridium sporogenes – 28.7 8 55 Ionata et al., (2008)

Chryseobacterium sp. kr6 Metallo 64 8.5 50 Riffel et al., (2007)

Chryseobacterium indologenes TKU014

Metallo

Metallo

Metallo

P1: 56

P2: 40

P3: 40

P1: 10

P2: 7–8

P3: 8–9

P1: 30–50

P2: 40

P3: 40–50

Wang et al., (2008)

Fervidobacterium pennavorans Serine 130 10 80 Friedrich and Antranikian (1996)

Kocuria rosea Serine 240 10 40 Bernal et al., (2006)

Microbacterium sp. kr10 Metallo 42 7.5 50 Thys and Brandelli (2006)

Nesternkonia sp. AL-20 Serine 23 10 70 Gessesse et al., (2003)

Nocardiopsis sp. TOA-1 Serine 20 >12.5 60 Mitsuiki et al., (2004)

Stenotrophomonas maltophilia Serine 35.2 7.8 40 Cao et al., (2009)

Streptomyces sp. S7 Serine-metallo 44 11 45 Tatineni et al., (2008)



Streptomyces sp. strain 16

Serine

Serine

Serine

Serine

KI: 203.2

KII: 100.8

KIII: 31.8

KIV: 19.2

KI: 9

KII: 9

KIII: 9

KIV: 9

KI: 50

KII: 50

KIII: 50

KIV: 60

Xie et al., (2010)

Streptomyces albidoflavus Serine 18 6–9.5 40–70 Bressollier et al., (1999)

Streptomyces pactum Serine 30 7–10 40–75 Bockle et al., (1995)

Streptomyces gulbagensis DAS 131 – 46 9 45 Syed et al., (2009)

Streptomyces thermoviolaceus – 40 8 55 Chitte et al., (1999)

Streptomyces sp. _ _ 10 60 Jayalakshmi et al., (2011)

Nocardiopsis sp. Protease - - - Cavalcanti et al., (2005)

Thermoanaerobacter sp. 1004-09 Serine 150 9.3 60 Kublanov et al., (2009)

Aspergillus oryzae Metallo 60 8 50 Farag and Hassan (2004)

Doratomyces microsporum Serine 30–33 8–9 50 Gradisar et al., (2005)

Myrothecium verrucaria Serine 22 8.3 37 Moreira-Gasparin et al., (2009)

Paecilomyces marquandii Serine 33 8.0 60–65 Gradisar et al., (2005)

Scopulariopsis brevicaulis Serine 36–39 8.0 40 Anbu et al., (2005)

Trichoderma atrvoviride F6 Serine 21 8–9 50–60 Cao et al., (2008)

Trichophyton sp. HÁ-2 Serine 34 7.8 40 Anbu et al., (2008)

Trichophyton vanbreuseghemii Serine 37 8.0 – Moallaei et al., (2006)

Aspergillus flavipes - 60 7 - El-Ayouty et al., (2012)



Many archaea grow in environments usually lethal to most cells, including

extremes in temperature, pH, salt content and pressure. Thus, archaea are valuable

resource of proteases for fundamental microbiology and enzymology studies, also

possessing the potential for biotechnological applications. Archaea displaying

keratinolytic activities were recently revealed through the in situ enrichment of

thermophilic prokaryotes with hydrolytic activities in hot springs (68–87°C and pH

4.1–7.0) (Kublanov et al., 2009). One isolate, identified as 1507-2, grew on α-keratin at

70°C and pH 6.0, was found to be an archaeon of the Crenarchaeota phylum,

representing a cluster of the so-called unknown Desulfurococcales. In the same

investigation, a 220-kDa thermostable keratinase showing broad pH (6.0 to 10.0) and

temperature (30 to 80°C) ranges of activity, with an optimum at pH 7.0 and 66°C, was

found in the culture supernatant of strain 1523-1 growing on keratin (Kublanov et al.,

2009).

2.19. Physiology of keratinase production and keratinolysis

Microbial keratinases are predominantly extracellular when grown on

keratinous substrates; however, a few cell-bound (Friedrich and Antranikian, 1996;

Onifade et al., 1998; Rissen and Antranikian, 2001; Nam et al., 2002) and intracellular

keratinases have also been reported (El-Naghy et al., 1998; Onifade et al., 1998). The

intracellular fraction in most of these reports mainly contributes to disulfide reductases,

sulfite or thiosulfate that synergistically assists the extracellular keratinases to degrade

keratin by reducing the disulfide bonds of keratin. To be more explicit, it can be put

forth that there are two steps in keratinolysis: sulfitolysis or reduction in disulfide

bonds and proteolysis. It may be speculated that sulfitolysis requires either the presence

of live cells (Bockle and Muller, 1997; Ramnani et al., 2005); reductants like sodium

sulfite, DTT, mercaptoethanol, glutathione, cysteine and thioglycolate (Onifade et al.,

1998) or disulfide reductases (Yamamura et al., 2002; Ramnani et al., 2005), which act

in cooperation with keratinolytic proteases to bring about complete degradation of

keratin.

However, the order of these events and their exact nature are still debatable.

Keratinases are largely produced in a basal medium with a keratinous substrate. Most



of the organisms are capable of using keratin as the sole source of carbon and nitrogen

(Williams et al., 1990; El-Naghy et al., 1998; Lin et al., 1999; Szabo et al., 2000;

Gousterova et al., 2005). However, the type of exogenous keratin inducer may range

from whole chicken feather, feather powder, wool, horns, nails and stratum corneum to

hair. In most cases, keratin serves as the inducer; however, soy meal is also known to

induce enzyme production (Cheng et al., 1995; Gradisar et al., 2000). Most of the

reports available on keratinases group them as inducible enzymes; however, few

constitutive keratinases have also been reported (Gessesse et al., 2003; Manczinger et

al., 2003). It is important to mention that in most of the reports on constitutive

keratinases, the nature of the enzyme is based on their caseinolytic rather than

keratinolytic activity. Hence, it is proposed that keratinolytic activity is by and large

inducible. Further, simple sugars such as glucose have been reported to suppress the

synthesis of keratinase due to catabolite repression (Santos et al., 1996; Ignatova et al.,

1999; Mohamedin, 1999; Singh, 1999; Wang and Shih, 1999; Yamamura et al., 2002;

Bernal et al., 2003; Gessesse et al., 2003; Suntornsuk and Suntornsuk, 2003; Thys et

al., 2004), which is a well known phenomenon for microbial proteases (Gupta et al.,

2002). However, comparison of keratinolytic titter of various microorganisms is

difficult due to the variety of substrates and the definitions of keratinase units

employed.

As far as physical parameters for production are concerned, they are species-

specific and thus vary with respect to the organism (Williams et al., 1990; Friedrich and

Antranikian, 1996; El-Naghy et al., 1998; Sangali and Brandelli, 2000b; Vidal et al.,

2000; Rissen and Antranikian, 2001; Rozs et al., 2001; Yamamura et al., 2002; Riffel et

al., 2003; Thys et al., 2004). It has been observed that alkaline pH from 6 to 9 supports

keratinase production and feather degradation in most microorganisms. Alkaline pH

possibly favours keratin degradation as higher pH modifies cystine residues to

lathionine (Friedrich and Antranikian, 1996), making it accessible for keratinase action.

Temperature for keratinase production ranges from 28 to 50°C for most bacteria,

actinobacteria and fungi to as high as 70°C for Thermoanaerobacter and

Fervidobacterium spp. (Friedrich and Antranikian, 1996; Rissen and Antranikian,



2001; Nam et al., 2002). Psychrotrophic production of keratinase has also been reported

for Stenotrophomonas sp. D1 (Yamamura et al., 2002).

Keratinase has been produced under submerged shaking conditions, except for a

few thermophilic bacteria (Friedrich and Antranikian, 1996; Nam et al., 2002; Rissen

and Antranikian, 2001) and fungi (Kaul and Sambali, 1999; Singh, 1999) where static

submerged fermentation has been reported. However, there are no reports available on

solid state fermentation for keratinase production. Therefore, since keratin is used as an

inducer, all fermentations leading to keratinase production are also accompanied by

subsequent degradation of keratin substrate. However, it is interesting to note that the

kinetics of keratinase production and that of keratin degradation do not overlap. Thus,

keratinolysis cannot serve as a marker for keratinase production and vice versa. This

can be exemplified from the literature where keratinase is mainly produced during the

late exponential or stationary phase of microbial growth (Williams et al., 1990; Cheng

et al., 1995; Sangali and Brandelli, 2000b; Vidal et al., 2000; Kim et al., 2001;

Ramnani and Gupta, 2004; Thys et al., 2004), whereas keratinase degradation takes

from 24 h (Ramnani and Gupta, 2004) to several days (Kaul and Sambali, 1999). This

is probably attributed to the complex mechanism of keratinolysis of these

microorganisms.

Other parameters accompanied during keratinolysis include increase in

alkalinity and thiol groups in the medium by most microorganisms. The higher

alkalinity is attributed to deamination reactions leading to the release of ammonium and

thus increase in pH (Dozie et al., 1994; Cheng et al., 1995; Friedrich et al., 1999;

Ignatova et al., 1999; Gradisar et al., 2000; De Toni et al., 2002; Riffel et al., 2003) and

consequent increase in keratinolysis (Riffel et al., 2003). Release of thiol groups is

largely due to reduction in disulfide bonds by enzymatic (disulfide reductases) or

chemical mechanisms (sulfite or thiosulfate) (Ignatova et al., 1999; Sangali and

Brandelli, 2000b; Yamamura et al., 2002; Ramnani et al., 2005).



2.20. Actinobacterial keratinolysis

Thermophilic actinobacteria produce many degradive enzymes (Ball and

McCarthy, 1989; Lin and Stutzenberger, 1995) and can play a major role in the

biodegradation of keratinous waste materials (Abdel Hafez et al., 1995). Proteolytic

and keratinolytic activity of protease producing thermophilic Streptomyces strain grown

on chicken feather as a substrate were demonstrated by Mohamedin (1999) and his

investigation revealed that proteolytic activity exhibited by Streptomyces sp. was

superior than keratinolytic activity because of protein substrate variation.

Biodegradation of keratin feather waste using actinobacteria was also studied by

Dhanasekaran (2012a) where he mentioned about the role of potential actinobacteria in

order to control feather pollution.

The novel mesophilic marine Streptomyces albus AZA strain was investigated

for its ability to produce constitutive and inducible extracellular keratinase by Esawy

(2007) and research outcome revealed that maximum keratinase production was

achieved with wheat flour among the different substrates used as the sole carbon and

nitrogen source for keratinase production.

Degradation of keratin and collagen containing wastes by newly isolated

thermoactinobacteria was reported by Gousterova et al., (2005). Keratinolytic and

proteolytic activity from the broth of a feather degrading thermophilic Streptomyces

thermoviolaceous strain SD8 suggested about potential enzymatic degradation of

various substrate such as fibrin, muscle, collagen, nail and hair (Chitte et al., 1999).

Keratin hydrolysis studies by alkaliphilic Nocardiopsis sp. TOA-1 was investigated by

Mitsuiki et al., (2002) and an enzyme called NAPase was identified responsible for

keratinolytic activity.

A Streptomyces sp. was isolated from poultry plant waste water, showed high

keratinolytic activity when cultured on feather meal medium (Tapia and Simoes, 2008).

Another alkaline keratinase producing Streptomyces sp. was reported from

slaughterhouse waste samples, Hyderabad, India by Radhika et al., (2008).



Electron microscopic study has been utilized for the detection of keratinolytic

feather degradation. Feather samples from inoculated broth of Streptomyces sp. S7

were examined with SEM for their degradation (Radhika et al., 2008). Similarly

Ramakrishnan et al., (2011) studied the scanning electron microscopic analysis of

feather degradation by Streptomyces sp. isolated from the feather dumped soil in

Thanjavur, Tamil Nadu India. More recently Tiwary and Gupta’s (2012) electron

microscopic study report revealed the effect of keratinase enzyme produced by Bacillus

licheniformis ER-15 on degradation of feather with respect to different time interval.

Physical parameters play a big role when it comes to any enzymes activity.

Effect of pH and temperature on enzyme activity is always considered as pioneer to

characterize the enzyme. Generally proteases and keratinase exhibits optimum activity

at alkaline range but there were reports suggested optimum enzyme activity had been

achieved in broad range of pH. Streptomyces sp. poultry processing plant waste water

in Brazil was exhibited optimum keratinolytic activity at 40°C and pH 8 and it stability

between 40 - 60°C (Tapia and Simoes, 2008). Keratinase activity against keratin azure

by Streptomyces sp. S7 showed optimum at 45°C and at highly alkaline pH 11

(Radhika et al., 2008). Streptomyces albidoflavus exhibited keratinolytic activity when

it was cultured on feather meal based medium and it showed stable enzyme activity

from 6-9.5 pH and temperature ranging from 40-70°C (Bressollier et al., 1999).

Similarly some other Streptomyces sp. such as S. pactum DSM40530, Streptomyces sp.

strain 16 showed optimum enzyme activity in the pH range of 7-10 and temperature

from 40-75°C (Bockle et al., 1995; Fuhong et al., 2010).

Mitsuiki et al., (2004) reported about a keratinolytic enzyme NAPase from

Nocardiopsis sp. TOA-1, which exhibited a much greater keratinolytic activity

compared with proteinase K and subtilisin, especially at a higher alkaline pH. The

maximal activity toward keratin was observed at a pH above 12.5 and at 60°C. A recent

report on protease producing Saccharomonospora viridis SJ-21, isolated from water

sample of water spring in Gujarat, showed optimum enzyme activity at 55ºC and pH

9.5 (Jani et al., 2012).



Keratinase enzymes have huge industrial importance and applications. Thus

effect of several chemicals, enzyme inhibitors, chelator, detergents etc. on keratinase

activity plays a significant role to classify the enzymes. Hence study on industrially

important microbial keratinase does not come to an end without investigating the effect

of various chemicals on their activity. NAPase a keratinolytic enzyme isolated from

Nocardiopsis sp. TOA-1, showed strong resistance to reducing agents such as

dithiothreitol, β-mercaptoethanol and sodium thioglycolate (Mitsuiki et al., 2004).

Bockle et al., (1995) reported that keratinolytic serine protease isolated from

Streptomyces pactum DSM 40530 showed a high level of stability with different

additives. In presence of SDS and thioglycolate, proteinase activity was reduced,

whereas DMSO and DTT showed slightly positive effect on proteinase activity.

Fuhong et al., (2010) was reported that even one millimolar of PMSF completely

inhibited the keratinolytic activities of four keratinase enzymes KI, KII, KIII and KIV

isolated from Streptomyces sp. strain 16.

Similarly a keratinolytic Streptomyces sp. isolated from poultry plant waste

water exhibited slight inhibition of keratinolytic activity against CaCl2, ZnCl2 and

BaCl2, whereas keratinolytic activity was not affected by EDTA, DMSO and Tween 20

(Tapia and Simoes, 2008). Keratinolytic activity of Streptomyces sp. S7 increased

substantially in presence of Ca2+ and inhibited in presence of PMSF and EDTA

(Radhika et al., 2008). They also reported that stability of the keratinolytic enzyme

against detergents; surfactants and solvents make this keratinase extremely useful for

biotechnological process involving keratin hydrolysis or in the leather industry.

Jayalakshmi et al., (2011) has been conducted the studies on purification and

characterization of keratinase enzyme from Streptomyces sp. JRS 19. The extracellular

concentrated crude enzymes were precipitated using ammonium sulphate (80%) and

dialyzed to remove salt. SDS-PAGE was used to analyze the protein profiling.

Hydrolytic activity of keratinase enzyme was detected by Native-PAGE and

zymography study.



Fuhong et al., (2010) reported about purification and characterization of four

keratinases produced by Streptomyces sp. strain 16 in native human foot skin medium.

Four extracellular keratinases (designated KI, KII, KIII, and KIV) were detected by

zymogram analysis and later the molecular weights of these enzymes 25, 50, 34 and 19

kDa, respectively were determined by SDS–PAGE after the purification of enzymes by

sephacryl S-200 column and DEAE FF column.

Bressollier et al., (1999) reported on keratinolytic serine proteinase from

Streptomyces albidoflavus. They detected serine proteinase by zymogram analysis and

after purification through DEAE-cellulose chromatography and carboxymethyl accel

plus chromatography; low molecular weight (18 kDa) serine proteinase was detected by

SDS-PAGE. Similarly Radhika et al., (2008) purified and characterized the alkaline

keratinase from Streptomyces sp. S7. After purification of the same enzyme by

sephacryl S-100 column, they have detected the keratinolytic nature of the enzyme

through zymogram analysis and the SDS-PAGE revealed the presence of 44 kDa

purified keratinase enzyme.

MALDI-TOF analysis of culture supernatant containing peptides produced

during enzymatic hydrolysis of hair by B. subtilis AMR revealed fragments in a range

of 800–2600 Da (Mazotto et al., 2009). The MALDI-TOF analysis of HMY after

cultivation for 4 days revealed multiple peaks from 816 to 2080 m⁄z, indicating the

presence of peptides generated by hydrolysis, when compared to control. Analysis of

commercial hydrolysed keratin also showed peptides of low molecular weight 900–

1200 Da.

A new alkaline keratinase extracted from Bacillus sp. 50-3 was isolated and

purified. The purified keratinase was determined through SDS-PAGE and the

molecular weight of the enzyme was found at 27 kDa by the MALDI-TOF-MS (Zhang

et al., 2009b). Bacillus licheniformis N22 was exhibited keratinase activity and the

molecular weight of purified keratinase was measured as 28 KD by SDS-PAGE and

confirmed by MALDI-TOF MS. Optimum keratinase activity was obtained at pH 8.5

and 50°C. This strain produced a distinct MALDI-TOF MS spectrum which was

different from that of the reference strain B. licheniformis PWD-1. The keratinase has a



unique peptide spectrum and was able to significantly degrade melanised feather

(Okoroma, 2012).

2.21. Feather meal formulation and biotechnological applications

Biodegradation of feathers by microorganisms represents a method for

improving the utilization of feathers as a feed protein (Hussein and Swelim, 1989;

Williams et al., 1991) and amino acids as pure chemicals (Williams et al., 1990). Most

investigators agreed that microbial conversion of feather (keratin) represents a

biotechnology for improving the utilization of feather as a feed protein (Table 5).

Biodegradation of feather can be achieved by cultivation of keratin-degrading

microorganism(s) on feather and the subsequent elaboration of extracellular keratinase;

the use of culture filtrates containing the keratinase or crude enzyme alone without the

microorganism and the use of purified enzyme alone without the microorganism.

Table 5. Amino acids composition (g kg-1) of microbially treated and untreated

samples (Onifade et al., 1998)

Amino acids Processed feather Unprocessed feather

Glycine 98.0 162.0

Valine 8.0 20.0

Leucine 45.0 83.0

Isoleucine 21.0 43.0

Arginine 41.0 17.0

Lysine 41.0 18.0

Methionine 4.0 -

Cysteine 46.0 76.0

Threonine 55.0 8.0

Phenylalanine 28.0 43.0

Tyrosine 19.0 16.0

Histidine 4.0 3.0

Tryptophan - -

Asparagine 81.0 67.0

Serine 82.0 72.0

Glutamine 105.0 97.0

Proline 222.0 188.0

Alanine 104.0 84.0



Studies by Elmayergi and Smith (1971) appeared to us as the pioneering

attempt to assess the nutritional complementarity between amino acids of feather meal

and microbial biomass following the fermentation with Streptomyces fradiae. The

methionine content of the product was higher than the unfermented, though methionine,

lysine and tryptophan contents were still low in the fermented product. Elmayergi and

Smith (1971) showed that all the concentrations of the amino acids were increased

considerably after fermentation. However, the results of the feeding trial with chickens

indicated no significant difference in the nutritional value between the fermented and

unfermented feather meal. This was explained to be the result of product

unacceptability by the chickens. The continuation of the experiment with

supplementation of methionine up to requirement eventually caused a comparable

growth rate of broilers with those fed isolated soyabean.

In a similar study, Onifade et al., (1998) reported that lysine, methionine and

arginine contents of feather meal were higher in the microbially fermented feather than

in the intact feather. The authors conducted two feeding trials with rats and their

findings were that those which received feather hydrolysates did not lose weight but

those fed a protein-deficient diet recorded weight loss. Feather hydrolysate digestibility

and utilization were confirmed in the studies, though sub-deficiency of methionine

would seem to predicate the lower or lack of weight gain of rats fed feather meal. It

was concluded from their investigations that not only feather meal (keratin) could be

used as protein for animal food, but also the biomass of the enzyme producing strain as

well.

The application of microbial technology for feather processing holds the

following nutritional significance. First, culturing of the microorganisms and keratinase

activity may result in a modification of the structure of feather keratin. This may alter

its resistance to digestive enzymes of the consuming animals (Elmayergi and Smith,

1971; Benedek et al., 1985; Williams and Shih, 1989). Furthermore, there can be

nutritional enrichment of the feather meal from microbial protein biomass that may be

complementary or additive. Onifade et al., (1998) reported about higher amounts of

lysine, methionine and arginine in fermented than in unfermented feather, leading to



their conclusion that it is not only the feather keratin that can be used as the protein

source, but the microbial biomass as well. Earlier, Elmayergi and Smith (1971) had

reported a marginal increase in methionine and lysine contents of feather fermented by

a methionine-secreting mutant of Streptomyces fradiae (Table 6).

Table 6. Concentration (g / 100 g protein) of selected amino acids in fermented

and unfermented feather meal (Elmayergi and Smith, 1971)

Amino acid Feather

meal (FM) FM fermented by

parent strain of S. fradiae

FM fermented by mutant

strain of S. fradiae

Methionine 0.37 0.43 0.90

Tyrosine 0.15 0.38 0.74

Lysine 1.77 2.14 3.23

Histidine 0.15 0.21 0.73

Cystine 4.74 3.45 2.18

Thirdly, the production of amino acids, especially feed-grade lysine and others,

from microbial fermentation of feather is also possible (Mohammed EI-Akied, 1987;

Williams and Shih, 1989).

Since there are not much studies on keratinolytic potential of soil actinobacteria

isolated from feather waste soil samples from Tiruchirappalli and Namakkal, Tamil

Nadu, India in terms of feather waste management, we planned to explore the

keratinolytic actinobacteria from the above locations, which can come up as a solution

of feather waste management by converting feather waste into alternative poultry feed

supplement.

2. REVIEW OF LITERATURE - Shodhgangashodhganga.inflibnet.ac.in/bitstream/10603/9644/9/09_chapter 2.pdf · The market research report "Vision for Indian Poultry Industry: Current Scenario

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