11

2. REVIEW OF LITERATURE

Marine Actinomycetes

Actinomycetes are the most economically and biotechnologically valuable

prokaryotes and are responsible for the production of half of the discovered

secondary metabolites. The actinomycetes are active components of marine

microbial communities and form a stable, persistent population in various marine

ecosystems. They are gram positive organisms with a wide range of habitat like

terrestrial, aquatic and extreme climatic conditions. They have high G+C (>55 per

cent) content in their DNA and they are originally considered as an intermediate

group between bacteria and fungi. The novel compounds with distinctive biological

activities have been isolated from these marine actinomycetes indicating that marine

actinomycetes are an important source for the discovery of novel secondary

metabolites. The discovery of numerous new marine actinomycete taxa, their

metabolic activity in their natural environments and their ability to form stable

populations in different habitats clearly illustrate that indigenous marine

actinomycetes indeed exist in the oceans. It is reported that marine actinomycetes

have novel characteristics from those of terrestrial counterparts and therefore, might

produce specific types of bioactive compounds. They are free living, saprophytic

bacteria, and a major source for production of antibiotics. Microbes are the most

interesting source of enzymes due to their broad biochemical diversity and

bioengineering potentiality (Bull et al., 2000; Stach et al., 2003; Magarvey et al.,

2004; Gesheva et al., 2005; Jensen et al., 2005; Fiedler et al., 2005). The demand for

enzymes from marine actinomycetes increased day by day because of its high

stability than enzymes derived from other sources like terrestrial microbes, plants

and animals. Marine microorganisms have recently appeared as rich sources for the

isolation of industrial enzymes. Actinomycetes isolated from the samples collected at

the marine environments such as the deep sea floor, marine invertebrates and marine

snow, all represent unique ecosystems that cannot be found anywhere else in the

world (Lam, 2006).

12

The distribution of actinomycetes in the sea is largely unexplored and the

presence of indigenous marine actinomycetes in the oceans remains elusive. As

marine microorganisms, particularly actinomycetes, have evolved the greatest

genomic and metabolic diversity, efforts should be directed towards exploring

marine actinomycetes as a source for the discovery of novel secondary metabolites.

Most of these actinomycetes are very different in terms of their 16S rRNA sequences

from their terrestrial counterparts and the ability to cultivate these novel

actinomycetes will provide a new source for the discovery of secondary metabolites

(Maldonado et al., 2005). Several ecologically significant properties of

actinomycetes were reported, which made the screening source expanded into

uncommon environments. Actinomycetes comprise 10.0 per cent of the total bacteria

colonizing marine aggregates (Valli et al., 2012). It is a boon in marine

bioprospecting for the exploration and exploitation of the rich biological and

chemical diversity found in marine organisms that inhabit the oceans. It is excited

that new groups of actinomycetes from unexplored or under exploited habitats be

persued as sources of novel bioactive secondary metabolites (Donia and Hamann,

2003).

Indian coastal area is the major source of actinomycetes and the isolation

from these unique ecosystems is a prolific source for the discovery of novel

secondary metabolites (Gulve and Deshmukh, 2012). Although isolation strategies

directed towards new marine-derived actinomycetes have been lacking, some

progress has recently been made in this area. Recent investigations using enrichment

techniques, new selection methods have led to the isolation of novel actinomycetes

from sediment samples (Magarvey et al., 2004; Mincer et al., 2005). Improved

recovery yields of marine actinomycetes from sponges using nutrient supplements

and enzymes have been reported (Olson et al., 2000; Webster et al., 2001). There is a

tremendous potential for the isolation of novel secondary metabolites from marine

actinomycetes. Balagurunathan and Subramanian, (2001) isolated the Streptomyces

from marine sediment samples. Sivakumar, (2001) isolated actinomycetes from the

samples collected from Pitchavaram mangrove, India. Few reports from the East

Coast of India, suggests that soil is a major source of Actinomycetes (Sivakumar

13

et al., 2005; Vijayakumar et al., 2007; Dhanasekaran et al., 2008; Vijayakumar et al.,

2009).

Among actinomycetes, Streptomyces are the dominant and the important

producers of novel bioactive compounds. Streptomyces are the most widely studied

and well known genus of the actinomycete family resembling fungi in their structure

but are intermediate between bacteria and fungi. Streptomyces is the largest genus of

Actinobacteria and belongs to the family Streptomycetaceae (Kämpfer, 2006). Over

500 species of Streptomyces bacteria have been described (Euzéby, 2008). As with

the other Actinobacteria, Streptomyces are gram-positive, and have genomes with

high GC content. The gene sequence analysis of 16S rRNA has been applied to

Streptomyces taxonomy to investigate relationships at the genus and species level.

Found predominantly in soil and decaying vegetation, most Streptomyces produce

spores and are noted for their distinct earthy odor that results from production of a

volatile metabolite, Geosmin. Streptomyces are characterised by a complex

secondary metabolism (Madigan and Martinko, 2005). The reports confirmed that

they produced over two-thirds of the clinically useful antibiotics of natural origin

(Kieser et al., 2000). Streptomyces are the filamentous bacteria which produce well

developed vegetative hyphae (between 0.5 - 2.0 µm in dm) with branches. Their

branching, filamentous arrangement of cells form a network called a mycelium that

aids in scavenging organic compounds from their substrates. The filament may be

long or short, depending on the species. Aerial myceliums are forms by them, much

smaller than those of fungi and many species produce asexual spores called conidia

(Mythili and Das, 2011). Their ability to produce spores contributes their survival

over long periods of drought, frost, hydrostatic pressure and anaerobic conditions

produced by water saturation. These species are widely distributed in the soils,

especially in those which are dry and not too acidic and which are rich in the organic

matter. They needed inorganic nitrogen sources for their growth but not require

vitamins and plenty of surfaces to support their mycelial growth in soil (Katsifas

et al., 2000).

14

Solid State and Submerged Fermentation

Fermentation is the technique of biological conversion of complex substrates

into simple compounds by various microorganisms such as bacteria and fungi. Both

bacteria and fungi yield an invaluable array of enzymes when fermented on

appropriate substrates. Both solid state and submerged fermentation are used for

enzymes production. Solid state fermentation is the growth of micro organisms under

controlled conditions using solid substrate with less moisture content for the

production of desired products of interest. It has advantages over submerged

fermentation such as high volumetric productivity, low cost of equipment involved,

better yield of product, lesser waste generation and lesser time consuming processes

(Robinson, 2001). Enzyme production using agro industrial substrates is one of the

most important advantages of solid state fermentation (Bhargav et al, 2008). In this

fermentation technique, the substrates are utilized uniformly and can be used for long

fermentation periods and this technique supports controlled release of nutrients. It is

best suited for fermentation techniques involving fungi and other microbes that

require less moisture content. The low availability of water reduces the possibilities

of contamination by bacteria and yeast. It exhibited low energy, easy downstream

processing, cheaper, less time consning, high recovery yield for the production of

industrial enzymes (Singh et al, 2010). The product obtained by this fermentation

showed different desired properties. Submerged fermentation utilizes free flowing

liquid substrates and is utilized quite rapidly; hence need to be constantly replaced

supplemented with nutrients. This fermentation technique is best suited for

microorganisms that require high moisture content. It is primarily used in the

extraction of secondary metabolites that need to be used in liquid form. Higher water

activity becomes the major cause of contamination in submerged fermentation and

makes downstream process difficult and very expensive. High quantity of liquid

waste is produced, causes difficulties in dumping. The cost of enzyme production by

submerged fermentation is higher than solid state fermentation (Mienda et al., 2011;

Subramaniyam and Vimala, 2012).

15

Lipase from bacteria and fungi

Lipolytic bacteria were isolated from oil contaminated soils and grown on

tributyrin media containing 1.0 per cent (w/v) olive oil and different media

parameters were optimized for maximal enzyme production. Lipase production was

maximum by using palm oil as carbon source, peptone as nitrogen source with pH

7.0 and temperature 37°C (Sirisha et al., 2010). B. subtilis showed maximum enzyme

production at 48 hrs with pH 8.0. Among carbon sources fructose and glycerol

showed maximum enzyme activity and yeast extract showed highest activity among

nitrogen sources (Hasan and Hameed, 2001). Sharma et al, (2009) studied about the

lipase production using Arthrobacter sp. BGCC 490. Iftikar et al, (2011) isolated 15

strains from oily products and screened for the production of lipases by solid state

fermentation. Among all the strains examined, Bacillus sp. MBLB 3 gave maximum

production of lipases. Different agricultural by products such as wheat bran, rice

husk, almond meal, cotton seed meal, soybean meal, sunflower meal and mustard

meal were used as substrates. Maximum extracellular lipase activity was observed

when almond meal and tween 80 of 0.5 per cent was used. Senthil Kumar and Selva

Kumar, (2008) isolated Bacillus sp. from slaughter house soil that showed maximum

lipase activity at 40ºC, pH 8.0 at 72 hrs incubation. A bacterial strain isolated from

an oil contaminated soil identified as Staphylococcus Lp 12 showed optimum

production of lipase at pH 8.0 and temperature 45°C. Maximum lipase production

was observed at 48 hrs of growth using peptone of 1.0 per cent and starch of 1.5

per cent was used as carbon and nitrogen sources respectively. Among the natural

oils, olive oil was able to induce more lipase rather than other oils like groundnut,

coconut and castor oils (Pogaku et al., 2010).

Imandi et al, (2010) investigated the production of lipase using Yarrowia

lipolytica NCIM 3589 from seed oil cake by solid state fermentation. The maximum

lipase activity of 26.42 U /gds was observed with the substrate of seed oil cake in

four days of fermentation. Khoramnia et al, (2011) evaluated the lipase production

ability of a newly isolated Acinetobacter sp. using both submerged and solid state

fermentations. The results showed that more enzyme production was observed using

solid state fermentation of coconut oil cake. The optimum temperature for the

16

enzyme activity was 45ºC where 63 per cent of its activity remained at 70

◦C after

2 hrs showed the feature of thermotolerant lipase which was active after 24 hrs in a

broad range of pH of 6.0 to 11.0. According to Veerapagu et al, (2013) the lipase

production was maximum at pH 7.0, temperature 37ºC and incubation time 48 hrs by

the lipase producing bacteria BLP 2 isolated from oil spilled soil from vegetable oil

processing factories The bacterial isolate was characterized and identified using

morphological, biochemical characteristics and 16S rRNA gene sequencing as

Pseudomonas gessardii. Increased enzymatic production was obtained by the

supplementation of 1.0 per cent peptone by P. gessardii. Mohan et al, (2008) isolated

Bacillus strains (B1 to B5) from the soil sample of coconut oil industry and the

strains were identified by morphological and biochemical characters. The lipase

production was measured with varying pH of 4.0 to 9.0 and temperature of 27, 37

and 47ºC. The maximum lipase production was recorded at pH 7.0 and temperature

37ºC at 24 hrs of the culture period by Bacillus strain B5. Among the substrates

tested, coconut oil at a concentration of 0.5 per cent was found to enhance the lipase

production in Bacillus strain B5. Amara and Saleem, (2009) used 2 strains of

P. aeruginosa for lipase production and both strains were able to grow at 37°C.

Rajesh et al., (2010) studied about the lipolytic activity of T. reesei by zone

of clearance on tributyrin agar medium. Lipase production was carried out using

modified mineral salt solution supplemented with edible oils as substrate in shake

flask at 30°C. The maximum lipase production was obtained after 96 hrs of

submerged fermentation in a media containing 5.0 per cent olive oil as a substrate at

initial pH of 5.0 and incubated in an orbital shaker at 150 rpm with 30°C. The

extracellular lipase present in the culture media was partially purified by ammonium

sulphate precipitation method, characterized for its pH and temperature optima. The

optimum pH and temperature on lipase activity were found to be pH 5.0 and 50°C

respectively. Gulati et al, (2005) extracted a low temperature, alkaline, and detergent

compatible lipase from Fusarium globulosum using an inexpensive source, neem oil.

Maximum lipase production was achieved in 96 hrs and the enzyme exhibited

remarkable stability in the presence of commercial detergents, bleaching agents, and

proteases. Rehman et al, (2011) used different agricultural wastes including canola

17

oilseed cake, sesame oilseed cake, linseed oil cake, cotton oilseed cake, rice bran and

wheat bran were used as substrates for their potential to produce lipase under solid

state fermentation conditions by Penicillium notatum. The maximum lipase activity

was observed using canola oil seed cake at 96 hrs of incubation period with pH 5.0,

incubation temperature of 30°C, 60.0 per cent moisture content and 3.0 per cent of

olive oil content. Enrichment with maltose as carbon source enhanced the lipase

production while nitrogen supplementation did not affect the lipase production

significantly. The optimization of conditions led to two-fold enhancement of lipase

activity as compared to the initial enzyme activity. Arthe et al, (2010) investigated

about the Lipase production by Trichoderma reesei and. According to Rifaat et al,

(2010) Fusarium oxysporum was the best lipase producer in submerged

fermentation. Serdar et al, (2011) used Trichoderma harzianum sp. for extracellular

lipase production.

Lipase from actinomycetes

Lipolytic activity was exhibited by the strains 3, 10, 28 from 56 actinomycete

strains isolated from marine sediments of south Indian coastal region of India

(Selvam et al., 2011). Aly et al, (2012) isolated a group of 20 bacterial isolates from

oil contaminated soil for lipase production using tributyrin and Tween 80 agar

media. Among the evaluated bacteria, the isolate LP10 was the most active isolate in

lipase production and was identified as S. exfoliates LP10. The molecular weight of

the purified enzyme was 60.0 kDa, determined using gel electrophoresis.

Gunalakshmi et al, (2008) isolated S. grieseochromogenes from shrimp pond which

showed highest lipase activity at pH 10.0, 55ºC with 4.0 per cent sodium chloride.

The enzyme activity was maximum using mannitol and L phenelylanine as carbon

and nitrogen source respectively. Sadati et al, (2013) studied about the lipase activity

of Microbacterium. The screening was performed based on clear zone formation on

tributyrin agar plates. It showed maximum enzyme activity at pH 6 and temperature

30°C. Jain et al, (2004) observed the maximum lipase activity from Streptomyces

strain SAP 1089. Cardenas et al, (2001) noted the maximum lipase activity using

S. halstedii. Dahiya et al, (2006) investigated about the lipase production from

B. megaterium AKG1 by submerged fermentation. Vishnupriya et al, (2010) studied

18

about S. griseus strain which showed maximum lipase activity at 24 and 48 hrs of

incubation period and the enzyme activity was maximum by using sunflower oil as

substrate. The lipase production using S. rimosus increased constantly when the

cultures entered stationary phase (Vujaklija et al., 2003).

Lipase production by Solid state fermentation

India is one of the world’s leading oilseeds producers. Oil cakes are by

products obtained after oil extraction from the seeds, rich in fibre, protein contents

and reported to be good substrate for enzyme production using microbes. Studies

using them for the production of industrial enzymes have shown promising results.

Mixed substrate fermentation has been more advantageous for such applications. Oil

cakes are available throughout the year and cheapest source for enzymes production

(Ramachandran et al., 2007). Babassu oil cake was used for growth and lipase

production in SSF by a Brazilian strain of P. restrictum (Gutarra et al., 2005).

Emtiazi et al, (2003) studied extracellular lipase production by Pseudomonas strain

X using cotton seed cake. Maximum production of lipase was obtained on oil cake in

50 hrs. Another bacterial strain, Bacillus mycoides was identified as lipase producer

on coconut oil cake. The growth of the organism and lipase production was

maximum after 72 hrs of incubation under shaking. Olive oil and beef extract were

best carbon and nitrogen sources. Na+ induced more lipase than K

+ and Mg

2+

(Thomas et al., 2003). Production of lipases by Penicillium simplicissimum was

studied in SSF using soybean oil cake as substrate (Di Luccio et al., 2004).

According to Bapiraju et al, (2004) the maximum lipase production was obtained

using gingelly oil cake as substrate by Rhizopus sp. BTS-24. The optimal lipase

production was observed under the pH 5.0, incubation time 72 hrs, incubation

temperature 28°C and agitation speed of 100 rpm. Dharmendra and Parihar, (2012)

reported about the lipase production using linseed oilcake as substrate by SSF.

Singh et al, (2010) studied about B. subtilis for lipase production using solid

state fermentation. The maximum extracellular lipase activity was observed using

ground nut oil cake with 70 per cent initial moisture content at pH 8.0 after 48 hrs of

fermentation. Chaturvedi et al, (2010) investigated about B. subtilis for lipase

production using tributyrin agar medium. The maximum extracellular lipase activity

19

was observed using ground nut oil cake as a substrate with initial moisture content of

70.0 per cent and at pH 8.0. Imandi et al, (2010) observed the production of extra

cellular lipase by solid state fermentation using Yarrowia lipolytica NCIM 3589.

The maximum lipase activity was observed with the substrate of seed oil cake in four

days of fermentation. Babu and Rao, (2007) studied about the lipase production

using Yarrowia lipolytica NCIM 3589 with various mixed substrates and observed

that the lipase activity was high using mixed sugar cane bagasse and wheat bran.

Balaji and Ebenezer, (2008) reported about the maximum lipase production from

pongamia pinnate cake using Colletotrichum gloeosporoides in solid state

fermentation. The carbon source xylose, nitrogen source peptone and magnesium

sulphate increased the lipase activity for C. gloseosporoides. Parihar, (2012) studied

about the lipase production from linseed and mustard oil cake using P. aeruginosa

and observed that linseed oil cake with olive oil showed highest lipase activity at

30ºC, 50 per cent moisture content and 50 hrs incubation period.

Bacterial and fungal proteases

Mohamed et al, (2012) observed maximum protease activity when B. subtilis

was grown on corn steep liquor (2.0 %) followed by casein hydrolysate (2 %) and 12

per cent cane sugar molasses as a carbon source at pH 10.0 and 37ºC. The enzyme

exhibited an optimum pH of around 10.0 and maintained its stability over a broad pH

range from 5.0 to 12.0. Its optimum temperature was around 37°C and exhibited a

stability of up to 50ºC. Dutta and Ray, (2009) studied about the characterization of

alkaline thermostable crude lipase from Bacillus cereus. Padmapriya and Williams,

(2012) selected B. subtilis for protease production based on highest zone formation.

The protease enzyme was highly active and stable from pH 6.0 to 9.0 with an

optimum at pH 7.0. and its optimum temperature was 37°C and it proved stable up to

30 - 60°C. The best carbon and nitrogen sources were starch and whey protein for

maximum protease production. Protease was partially purified by ammonium

sulphate precipitation and dialysis and the molecular weight of purified enzyme was

characterized by SDS-PAGE and observed as 50.0 kDa.

Safey and Raouf, (2004) observed the following optimal conditions for

protease production by B. subtilis. It showed highest activity using 0.5 per cent of

20

substrate concentrations at 30 hrs incubation period with pH 7.0 and temperature

40ºC. The isolate exhibited maximum activity using lactose as carbon source,

ammonium sulphate as nitrogen source for protease production. The protease

enzyme was purified by ammonium sulfate precipitation and Sephadex G 200

filtration method. A protease producing moderately halo-alkali thermotolerant

Bacillus strain SH1 was isolated from salt spring sediment in Manipur, India. Corn

starch of 1.0 percent and peptone of 0.2 per cent were found as optimal carbon and

nitrogen sources for protease production. The enzyme was active over a wide range

of temperatures and pH with optima at 50º C and pH 8.0 and the production was

increased with the supplementation of by ferrous sulphate (Ningthoujam and

Kshetri, 2010). Prathamesh and Valasange, (2012) studied about the lipase and

protease production using Bacillus sp. isolated from compost soil. Dodia et al,

(2006) investigated the production of alkaline protease from halophilic and

alkaliphilic bacteria isolated from coastal Gujarat.

Patre and Dawande, (2010) examined Aspergillus niger for protease

production. The enzyme activity was found to be maximum using soyabean

(3.06 mg/l). Protease obtained using solid state fermentation was partly purified by

ammonium sulphate precipitation. According to Sahib, (2009) A. niger showed better

lipase production using 1.0 per cent glucose as carbon source, 1.5 per cent casein as

nitrogen source, 1.0 per cent sodium chloride at pH 10.0 and 40ºC for 5 days

incubation period. Oliveira et al, (2010) isolated 2 strains (Rhizobia sp. strain R-986

and Bradyrhizobia sp. strain R-993) from soils of the central Amazonian plain

region. The enzyme was active at pH 9.0 - 11.0 and temperature of 35 - 55oC.

Protease activities in the crude extracts were enhanced in the presence of 5.0 mM of

metal ions, such as Na+, Ca2

+, Mg2

+ and Mn2

+. Benazir et al, (2011) investigated the

potentiality of the fungi A. niger to produce the enzyme protease. Mishra et al,

(2008) studied the effect of different carbon sources on the proteolytic activity of

A. awamori MTCC 548. High protease production was observed using glucose with

1.0 per cent and pea nut meal with 2.0 per cent as an effective nitrogen source.

21

Protease from actinomycetes

S. pulvereceus MTCC 8374 was used for extra cellular alkaline protease

production by submerged fermentation and the maximum enzyme production was

obtained during early stationary phase with optimum pH, inoculum and temperature

of 9.0, 3.0 per cent and 33ºC respectively. Among carbon sources 0.3 per cent starch

gave a maximum production followed by maltose, xylose and fructose. High yield of

protease production was reported with 1.0 percent casein followed by soybean meal,

yeast extract and malt extract. Further, it was optimized with 0.5, 1.0 and 1.5

per cent of sodium chloride among which 1.0 per cent of sodium chloride resulted in

maximum protease production level of protease (Jayasree et al., 2009). Sayed et al,

2012 reported about the protease production from S. pseudogrisiolus NRC 15. Shafei

et al, (2010) studied about S. albidoflavus for protease production and observed

optimum enzyme production with pH of 10.0, inoculum size of 10.0 per cent.

Glucose and sodium hydroxide treated bagasse pith proved to be an efficient carbon

source for alkaline protease production and showed the highest enzyme activity.

Casein in presence of yeast extract was considered as the best nitrogen source for

maximum protease prouction. The alkaline protease enzyme exhibited an optimum

activity at pH 9.0 and was stable over a pH range from 6.0 to 11.0. Debananda et al,

(2009) studied about the production of extracellular proteases using halophilic alkali

thermotolerant actinomycetes. Jani et al, (2012) reported about a protease producing

actinomycete Saccharomonospora viridis SJ 21 which was isolated from hot water

spring of Gujarat and exhibited growth in broad temperature range of 35 to 60ºC and

pH range of 7.0 to 10.0. The optimum conditions observed for protease production

were 55ºC and pH 9.5, with 3.0 per cent inoculum in the medium containing metal

ions like Mg2+

and Ca2+

. The best carbon and organic nitrogen sources for the

organism were observed to be glycerol and aspargine, respectively, while the most

effective inorganic nitrogen source was observed to be sodium nitrate. Among the

raw sources used, the maximum protease production was found with wheat bran of

0.5 per cent. Thumar and Singh, (2007) isolated an alkaliphilic and salt tolerant

actinomycete, S. clavuligerus strain Mit1, from Mithapur, the Western coast of India.

The organism was gram-positive, having filamentous, long thread like structure and

22

exhibited the optimum level of alkaline protease (130 U/ml) during the early

stationary phase at pH 9.0. The strain could produce protease at optimum sodium

chloride concentration of 5.0 per cent (w/v) and sucrose and gelatin were the best

carbon and nitrogen sources respectively. Ningthoujam et al, (2009) studied about

the protease production using Nocardiopsis prasina HA 4 which was isolated from

Manipur and screened on skim milk agar. The organism grew well at temperatures of

20 to 42°C, pH 7.0 to 10.0 and in presence of 2.0 to 10.0 per cent of sodium

chloride. Sucrose (5 %, w/v) and peptone (0.2 %, w/v) were found to be the best

carbon and nitrogen sources for protease production. The optimum temperature for

the enzyme was 55°C and it had two pH optima at 7.0 and 10.0. Interestingly Fe2+

was found to have stimulatory effect and Ca2+

, Mg2+

partially inhibited enzyme

activity whereas Hg2+

completely inhibited the enzyme activity.

Vijayaraghavan et al, (2012) studied about the protease production using

halotolerant Actinobacterium sp. for solid state fermentation while apple pomace

supported maximum protease production. The optimum conditions required for

enzyme production were a fermentation period of 72 hrs, 10.0 per cent of sodium

chloride, pH 7.0 and 10.0 per cent of inoculum. The enzyme exhibited activity to a

range of pH (7.0 - 9.0) and temperature (30 - 45°C), with optima at 8.0 and 40°C,

respectively. Ca2+

ion was required for its activity and stability and the enzyme was

widely active at the range of sodium chloride concentration of 5.0 to 15.0 per cent.

According to Rifaat et al, (2006) a neutral protease was detected in the culture

medium of S. microflavus isolated from some Egyptian soils and the enzyme was

purified by precipitation with ammonium sulphate and gel filtration on Sephadex G-

75. The optimal pH and temperature for catalytic activity of protease were pH 7.0

and 40ºC respectively. Calcium and manganese stimulated protease activity while

Ag+ inhibited the enzyme activity. Vonothini et al, (2008) isolated 28 strains of

actinomycetes from the sediment samples of an estuarine shrimp pond located along

the South East Coast of India. Among them Streptomyces roseiscleroticus showed

higher protease activity. The enzymatic activity was maximum at pH 7.0,

temperature 40ºC and 3.0 per cent of sodium chloride concentration, carbon

compound such as sucrose and L-glutamine as amino acids.

23

Guravaiah et al, (2010) collected 300 marine samples from different locations

of Bay of Bengal from Pulicat lake to KanyaKumari. Among them 208 isolates of

marine actinomycetes were isolated using differential media such as starch casein

agar medium. Alkaliphilic actinomycetes isolated from water samples were studied

for the production of protease activity. Strain MA1-1 was selected as a good alkaline

protease producer and starch was found to be effective for the growth and enzyme

production with highest specific activity. The optimum pH and temperature of the

partially purified protease were determined as pH 9.0 and 50°C, but high activity was

also observed at pH 8.0 - 13.0 at 35° - 50°C. Fulzele et al, (2011) isolated 9 different

cultures from marine samples of the Indian Ocean which grew optimally at 30ºC

temperature and pH 7.0 - 8.0. The isolate MBRI 7 exhibited extracellular protease

activity on skim milk agar plates and identified as Marinobacter based on 16S rRNA

sequencing. The optimum temperature and pH for enzyme activity were found to be

40ºC and 7.0 respectively. The crude enzyme was stable at temperature range of 30

to 80ºC and pH 5.0 - 9.0 and the enzyme remained 50.0 per cent active at pH 9.0

after 1 hr. Awad et al, (2012) used S. pseudogrisiolus NRC-15 isolated from

Egyptian soil sample for and maximum protease production was obtained in the

medium supplemented with 1.0 per cent glucose, 1.0 per cent yeast extract, 6.0

per cent sodium chloride and 100 μmol/l of tween 20, initial pH 9.0 at 50 ºC for 96

hrs.

Protease production by Solid State Fermentation

The extra-cellular alkaline protease was produced by B. horikoshii which

showed maximum enzyme activity using soybean oil cake (1.5 %) and casein (1.0

%) at pH 9.0 and 34°C over 16 to 18 hrs incubation periods. The enzyme had an

optimum pH of around 9.0 and maintained its stability over a broad pH of 5.5 and

12.0 (Joo et al., 2002). Protease production from soyabean oil cake using

Penicillium sp. in solid state fermentation was reported by Germano et al, (2003).

Joo et al, (2003) observed the production of alkaline protease by B. clausii and

maximum activity was observed using soybean cake and the enzyme had an

optimum pH of around 11.0 and optimum temperature of 60°C. The suitability of

several oil cakes such as coconut oil cake, palm kernel cake, sesame oil cake, olive

24

oil cake were evaluated for neutral protease production using A. oryzae along with

several agro industrial residues such as wheat bran, rice husk, rice bran, spent

brewing grain (Sandhya et al., 2005). Coconut oil cake in combination with wheat

bran was used for production of neutral metallo protease by A. oryzae NRRL 2217

(Sumantha et al., 2005). Rauf et al, (2010) reported about the protease production

from Rhizopus oligosporus using sunflower meal as a substrate. The maximum

enzyme activity was observed with inoculum size of 1.0 per cent, substrate

concentration of 20.0 g with pH 3.0 and temperature 35oC. Sumantha et al, (2006)

investigated Rhizopus microsporus NRRL 3671 for protease enzyme production by

rice bran as a substrate with 44.44 per cent moisture content, temperature of 30°C for

72 hrs. Addition of casein resulted in a marginal increase in protease yield and the

enzyme was found to be a metalloprotease, being activated by Mn2+

, with maximal

activity at a temperature of 60°C and pH 7.0. The enzyme was partially purified by

precipitation, dialysis and 3 fold increase in the enzyme purity was achieved.

Divakar et al, (2006) optimized the process parameters for alkaline protease

production using solid state fermentation by Thermoactinomyces thalpophilus

PEE 14.

Lazim et al, (2009) used Streptomyces sp. CN902 for protease production

under solid state fermentation. The combination of wheat bran (WB) with chopped

date stones (CDS) (5:5) proved to be an efficient mixture for protease production as

it gave the highest enzyme activity when compared to individual WB or CDS as

substrates with 60 per cent moisture content and incubated at 45°C. Rathakrishnan

and Nagarajan, (2011) studied about the protease production from B. cereus using

agro residue red gram husk with maximum enzyme production of 262 U/g of

substrate. Maximum protease production was observed at a growth of 48 hrs, pH of

9.0, 50 per cent substrate moisture, temperature of 40ºC and 7.5 g of substrate

concentration. Enrichment with fructose as carbon source and nitrogen

supplementation with beef extract also enhanced the enzyme yield.

Rajmalwar and Dabholkar, (2009) reported about the six oil seed cakes as

substrates for protease production by solid state fermentation using Aspergillus sp.

isolated from leather. The agro industrial by product (oilseed cakes) namely soybean,

25

groundnut, sesame, linseed, mustard and cotton were evaluated as inducer for

production of protease using solid state fermentation and the enzyme synthesis was

maximum when soybean oil seed cakes was used as substrate followed by sesame

oilseed cake. Paranthaman et al, (2009) carried out a comparative study of the

production of protease using different varieties of rice brokens (PONNI, IR-20,

CR-1009, ADT-36 and ADT-66) from rice mill wastes as substrates in solid-state

fermentation by A. niger. Among them rice broken PONNI produced the highest

activity while ADT-66 produced lowest protease under solid state fermentation

conditions. The optimized conditions for producing maximum yield of protease were

incubation at 35°C, 96 hrs, pH 7.0 and the protease could be commercially used in

detergents and leather industry.

According to Ravi kumar et al, (2012) the optimum conditions for maximum

protease production under solid state fermentation by Pleurotus sajor-caju was

found to be on the fourth day of incubation, pH 7.0, temperature 30ºC, 3.0 per cent

corn as substituted carbon source, 0.8 per cent ammonium nitrate as nitrogen source

concentration. Metal ions (Ca2+

and Na 2+

) caused moderate reduction in enzyme

activity. Protease was partially purified by ammonium sulphate precipitation (70 %

saturation). It showed 2.72 fold with recovery of 53.0 per cent and the purified

enzyme preparation gave a single protein band in SDS-PAGE analysis indicating a

molecular weight of 48.0 kDa which exhibited its optimal activity at 60ºC and at pH

8.0. Kuberan et al, (2010) used halophilic strain of Bacillus sp. for protease

production under solid state fermentation. Among the different Agro-industrial waste

products like various oil cakes, wheat bran, rice bran, sugarcane bagasse, moong

dhal husk, cotton seed and green gram husk screened as substrates, green gram husk

followed by moong dhal husk supported maximum protease production. Among the

tested nitrogen compounds 0.5 per cent (w/w) urea followed by tryptone was served

as best nitrogen sources and 0.5 per cent (w/w) lactose followed by fructose was

served as the best carbon sources for protease production by Bacillus sp. The

protease production was optimum at 5.0 per cent (w/v) sodium chloride. The highest

protease activity was observed at pH 8.0 and 35ºC for 48 hrs incubation period.

26

Applications

Dairy industry effluent treatment

Industrialization has become a major concern due to its deteriorating activity

on environment. Most of the industries discharge their untreated effluent into the

nearby ecosystem results in harmful effects to both terrestrial and aquatic ecosystem.

Dairy industry is one of the important industries in India that cause water pollution.

Dairy effluent has high organic loads as milk is its basic constituent with high levels

of chemical oxygen demand, biological oxygen demand, oil and grease and nitrogen

and phosphorous content. Biotreatment by microbes is an efficient methodology for

the removal of toxicants from the effluent. Silambarasan et al, (2012) used

Pithophora sp. for the treatment of dairy industry effluent. The various physico

chemical characteristics of the effluent were analyzed before and after treatment.

From the results it was observed that except nitrate all the other nutrients level was

decreased during the treatment. Mongkolthanaruk and Dharmsthiti, (2002) studied

about a mixed bacterial culture comprising P. aeruginosa LP602, Bacillus sp. B304

and Acinetobacter calcoaceticus LP009 for use in treatment of lipid-rich wastewater.

One litre of wastewater was treated with a mixed inoculum in 20.0 ml of basal salt

solution. The BOD value and lipid content were reduced within 12 days.

Bioremediation of high fat and oil based waste water from palm oil, dairy,

soap industries were treated by selective lipase producing bacteria like B. subtilis,

B.licheniformis, B.amyloliquefaciens, P. aeruginosa, Serratia marscesens. Among 6

isolates P. aeruginosa showed maximum reduction of BOD (111 mg/l) in palm oil

effluent, 82.0 mg/l in dairy effluent, 145 mg/l in soap effluent (Prasad and

Manjunath, 2011). Bhumibhamon et al, (2002) carried out biotreatment of high fat

and oil wastewater by selected lipase producing bacteria (Acinetobacter sp. KUL8,

Bacillus sp. KUL39 and Pseudomonas sp. KLB1). The treatment was done using

single and mixed culture and in the first experiment, wastewater of bakery industry

was treated with bacterial isolates. Results showed that fat, oil and COD decreased

remarkably with treatments. With single culture, the removal of fat, oil and COD

were 73.0 to 88.0 per cent and 81.0 to 99.0 per cent and was observed that the

isolates KUL8 and KUL39 showed better activities. In the second experiment,

27

wastewater of palm oil and bakery industries were treated. The isolates KUL8 and

KUL39 showed better degradation activities which removed fat and oil by 87.7, 80.6

per cent in palm oil wastewater and 70.0, 64.0 per cent in bakery wastes respectively.

The decreasing of COD was found to be 90.0 to 96.0 per cent when mixed culture of

KUL8, KUL39 and KLB1 were applied for both kinds of wastewater. Vida et al,

(2007) isolated 10 micro flora from the dairy factory effluent, Tehran and screened

for their ability to reduce the organic matter content and COD of the effluents at

30°C and pH 11.0. Highest COD reduction was obtained by two isolates, BP3 and

BP4 with 70.7 and 69.5 per cent respectively.

Dairy effluent in germination studies

Seed germination, seedling growth and certain biochemical parameters in

paddy variety ADT-38 were studied by Dhanam, (2009) and observed that 100

per cent concentration of effluent caused inhibitory effect on paddy. According to

Gaikar et al, (2010) the germination and seedling growth of Soybeans was carried

out using dairy effluent and was observed that increase in effluent concentration

there was a corresponding decrease in per cent germination, but seedling growth

gradually increased up to 50.0 per cent of effluent concentration. The utilization of

100 per cent effluent completely inhibit for both seed germination and seedling

growth of maize variety Nithyashri. At lower dilutions and in treated effluent

samples, maize showed favorable effects on seed germination, seedling growth, dry

matter production and biochemical parameters. Among them 75.0 and 100 per cent

concentrations of raw effluent caused inhibitory effects and the study suggested that

the effluent can be used safely for irrigation of maize, only after proper treatment and

dilutions (Manu et al., 2012).

Tannery industry effluent treatment

Tannery is one of the problematic industries in India which generate high

quantity of wastewater with high TDS (Total Dissolved Solids) and toxicity due to

chromium hence discharge of tannery effluent is a very serious issue. Many

conventional treatment processes has been carried out in these industry before the

discharge of effluent. One of the most common treatment process applied is

biological treatment process by activated sludge process and upflow anaerobic

28

sludge blanket process which is cost oriented. Therefore the biotreatment using

microbes are used for tannery effluent as a low cost ecosafe methodology. Hozzein

et al, (2012) selected 10 actinomycete isolates which belong to the genera of

Nocardia, Streptomyces, Rhodococcus, Gordonia and Nocardiopsis. The results

showed that most of the selected actinomycetes were effective in the biological

treatment of the wastewater and have the ability to decrease the values of BOD,

COD, TSS and also to reduce the concentrations of the tested heavy metals (Cu, Fe,

Mn, Pb and Zn) markedly. The Streptomyces strain C11 was found to be the most

efficient organism in respect to biological treatment of the wastewater and removal

of the heavy metals from the raw wastewater. Sivaprakasam et al, (2008)

demonstrated about the biological treatment using salt tolerant microbes. The salt

tolerant bacteria isolated from marine and tannery effluent samples were identified as

P. aeruginosa, B. flexus, Exiguobacterium homiense and Staphylococcus aureus. Salt

tolerant bacterial mixed consortia showed appreciable biodegradation at all saline

concentrations (2.0, 4.0, 6.0, 8.0 and 10.0 % w/v) with 80.0 per cent COD reduction

in particular at 8.0 per cent salinity level. Midha and Dey, (2008) studied about the

sulfide removal of tannery effluent using microbes. Ingole et al, (2012) isolated

Pseudomonas sp. from tannery effluent and its degradation efficiency was analyzed

by comparing the changes in the physicochemical analysis of the tannery effluent

before and after bacterial treatment. The results revealed that the toxic effects were

mostly reduced after treatment, making the treated effluent suitable to be discharged

into the environment. Subramani and HariBalaji, (2012) studied about the 2 bacterial

spp. (Bacillus sps and Pseudomonas aeruginosa) and one fungi (Aspergillus niger)

for biodegradation study. A. niger was effective in degradation followed by

P. aeruginosa and Bacillus sp. The physio chemical parameters analysis showed a

gradual decreased value in microbially treated sample when compared to the

untreated sample.

Chromium removal from tannery effluent

Chromium is commonly used for leather processing in tannery industries and

Cr (VI) is a potent carcinogen to humans and animals as it enters cells via surface

transport system and gets reduced to Cr (III) that inducing genotoxicity (Matsumoto

29

et al., 2006). Thus, Cr loaded effluent used for irrigation disrupts several

physiological and cytological processes in cells (Chidambaram et al., 2009). It also

results in reduced root growth, biomass, seed germination and early seedling

development, chlorosis, photosynthetic impairment and finally leading to plant death

(Irfan and Akinici, 2010). Biological processes, such as biosorption using microbial

cells have been examined for their Cr (VI) removal capabilities (Cheung and Gu,

2003). Cr (VI) reduction by bacterial consortia has been extensively studied and

most studies reported that micro-organisms are capable of reducing Cr (VI) at low

concentrations, usually within the range from 0.1 to 0.5 mM (Chardin et al., 2002).

Similarly the bioaccumulation characterization of zinc and cadmium was studied by

using S. zinciresistens, a novel actinomycete (Lin et al., 2012). Some micro-

organisms lose viability and Cr (VI) reducing capacity in the presence of high

concentrations of Cr (VI), which further complicates the biological treatment of Cr

(VI) polluted wastewater (Pattanapipitpaisal et al. 2001). McLean and Beveridge,

(2001) used Pseudomonas sp. for reduction of chromate. Barrera et al, (2008)

studied about a fungal strain Trichoderma inhamatum capable of removing

hexavalent chromium. The results indicated that T. inhamatum fungal strain may

have potential applications in bioremediation of Cr (VI) contaminated wastewater.

Germination studies using tannery effluent

The phytotoxic and genotoxic effects of tannery effluent and chromium (Cr)

were investigated in Allium cepa which was exposed to various concentrations of

tannery effluent and Cr. In all treated plants, a significant (p>0.05) reduction in root

length was observed. At highest effluent and Cr concentration both the plants

showed pyknosis condition after 168 hrs (Gupta et al., 2012). Mishra et al, (2012)

studied about the toxicity study of tannery effluent using Allium cepa. Mythili and

Karthikeyan, (2011) used the selected isolates Bacillus, Pseudomonas and

Micrococcus for the bioremediation of tannery effluent. The study of seed

germination (Black gram and Sunflower) was carried out at different concentrations

of treated and untreated effluent using soil sowing method and observed that the

shoot length of the seedlings were increased for treated effluent comparing to

untreated effluent. The results revealed that effluent treated by microorganisms had

30

no negative impact on the seed germination and can be effectively used for irrigation

purpose.

Biogas production

The agricultural wastes that are released every day create serious

environmental problems. They are widely available, renewable and virtually free,

hence they can be an important resource and can be converted into animal feed,

compost and biogas. However, many of the agricultural wastes are still largely under

utilized and burned in the field results in serious environmental problems. Water

pollution by solid waste disposal is also becoming a severe problem. Renewable

resources for energy production come more and more into public focus because of

problems caused by the predictable shortage of fossil fuels in the next decades. The

ever increasing energy demands force us to look for a replacement of the

nonrenewable resources of energy by other (renewable) resources. Besides

generating revenue from the energy produced, waste to energy schemes offer an

alternative and environmentally acceptable means of waste disposal. Additionally,

the schemes also provide a valuable by product with a good quality, agricultural

fertilizer that is nearly odourless. With the concern over future energy shortages and

increasing costs of conventional fuels and electricity derived from them, there is

increasing interest in using anaerobic digestion as a source of renewable energy

while providing acceptable waste management (Sabiiti, 2011). In rural areas of

developing countries various cellulosic biomass (cattle dung, agricultural residues,

etc.) are available in plenty which have a very good potential to cater to the energy

demand, especially in the domestic sector (Kashyap et al., 2003). Anaerobic

technologies offer an attractive manner of use of biomass resources for the purpose

of partial satisfying of the energy needs of our society (Yadvika et al., 2004).

A comparative study on the effect of different pre-treatment methods on the

biogas yield from water hyacinth was carried out in a batch digester for a 60 days

retention period. Biomethanation was carried out in mesophilic temperature range of

30 to 37°C. The results of the study showed the highest cumulative biogas yield was

from water hyacinth combined with poultry waste and the biogas yield of the fresh

water hyacinth was negligible. The overall results showed that blending water

31

hyacinth with poultry waste and primary sludge had significant improvement on the

biogas yield, and treating water hyacinth with sodium hydroxide increased the biogas

yield slightly. It also indicated that water hyacinth was a very good biogas producer

and the yield can be improved by drying and combining it with poultry waste and

primary sludge (Patil et al., 2011). Anaerobic codigestion of Jatropha deoiled cake

and orange peel waste for biogas production was studied by Elaiyaraju and Partha,

(2012). The experimental data showed a maximum gas gas production at 1:2 ratio of

Jatropha deoiled cake with orange peel waste obtained for a period of 17 days. The

biogas production was measured by liquid displacement system on daily basis and

the digested slurry can be used as a fertilizer for agricultural purpose.

Vivekanandan and Kamaraj, (2011a) investigated about the rice chaff with

co-digestion of cow dung for biogas production. The combination of 50.0 per cent of

boiled rice chaff with 50.0 per cent weight of cow dung showed a cumulative biogas

production of 161.5 ml for the retention time of 60 days. The combination of 75.0

per cent of boiled rice chaff and 25.0 per cent of cow dung showed the biogas

production of 140.5 ml; 50.0 per cent of raw rice chaff with 50.0 per cent of cow

dung showed no significant gas production due to high percent of lignin in raw rice

chaffs. Vivekanandan and Kamaraj, (2011b) studied the anaerobic digestion of rice

crop residues rice chaff, rice straw and rice husk as cosubstrate with cow dung. Each

waste was mixed in different proportion: Rice chaff / cow dung; Rice straw/ cow

dung and; Rice husk / cow dung. Plastic digester was used and the digestion of

residues was undertaken by batch-type anaerobic process, the digester operated at a

temperature (25 - 35oC). The low performance of cow dung / rice husk was due to

the high lignin content which contained unfavourable non lignin-carbon to nitrogen

ratio (70:1). Kapadiya et al, (2010) reported about the biogas production potential

using powdered tobacco (Nicotiana tabacum) stem and with cow dung. The digested

biogas slurry of running biogas plant was added in all the digesters (10.0 %). Biogas

production was measured by water displacement method and the results revealed that

addition of tobacco stem waste significantly increased the daily average rate of

biogas production than control. Sridevi and Ramanujam, (2012) reported about the

biogas production using mixture of vegetable wastes like carrot, beans and brinjal.

32

Iyagba et al. (2009) investigated the co-digestion of cow dung with rice husk

for biogas production. The study was carried out at 26 to 29°C for a period of 52

days and the biogas produced was collected by water displacement method. Sample

A (50 % cow dung with 50 % rice husk) showed a cumulative biogas production of

161.5 ml at the end of the 38th

day of the experiment after which there was no further

production. The production from sample B (25 % cow dung and 75 % rice husk) was

not significant, while there was no production from sample C (100 % rice husk). The

use of oil cakes offers good alternative to traditional applications by their

exploitation in the production of environmentally friendly green bio fuel.

Satyanarayan et al, (2008) used mustard oil cake (Brassica compestries) as a

substrate for biogas production. The addition of 30.0 per cent mustard oil cake

showed a tremendous increase of 63.44 per cent of biogas production. Powdered

leaves of some plants and legumes have been found to stimulate biogas production

that lead to high localized substrate concentration and a more favourable

environment for growth of microbes. The additives also help to maintain favourable

conditions for rapid gas production in the reactor, such as pH, promotion of

acetogenesis and methanogenesis for the best yield, etc. Alkali treated (1%) plant

residues when used as a supplement to cattle dung resulted in almost twofold

increase in biogas and CH4 production (Yadvika et al., 2004). Sharma, (2002)

observed an increase of 40.0 to 80.0 per cent in biogas production on addition of 1.0

per cent onion storage waste to cattle dung.

Utility of microbes and enzymes for biogas production

In today’s energy demanding life style, need for exploring and exploiting

new sources of energy which are renewable as well as eco friendly is a must. To

protect the environment, waste management in an efficient way by microbes is now

encouraged. Nowadays the increasing attention has been paid for developing and

implementing this type of innovative technology for efficient waste management.

Anaerobic digestion is an attractive technology for the treatment of organic waste

(e.g., manure, food-processing wastes and green wastes). Masse et al, (2001)

reported the degradation by microbes prior to the biodigestion process results in the

easier absorption of nutrients for anaerobic microorganisms and enhanced for more

33

complete biodegradation. Zhong et al, (2011) used the microbial agents for the

pretreatment of corn straw at ambient temperature of 20°C to improve its

biodegradability and anaerobic biogas production. These treatment conditions

resulted in 33.07 per cent more total biogas yield, 75.57 per cent more methane

yield, and 34.6 per cent shorter technical digestion time compared with the untreated

sample. Biological pretreatment could be an effective method for improving

biodegradability and enhancing the highly efficient biological conversion of corn

straw into bioenergy.

Weiss et al, (2010) used Bacteroides sp. and Azospira oryzae immobilised on

zeolite to enhance he biogas production. In batch-culture experiments there was an

increase of methane by 53.0 per cent in microbially pretreated samples than the

control. Potivichayanon et al, (2011) studied the enhanced biogas production from

bakery waste by the addition of Pseudomonas aeruginosa. The grease waste from

bakery industry’s grease trap was initially degraded by P. aeruginosa. The

biodigesters of 3 numbers were set up by using 3 different substrates: non degraded

waste as substrate in first biodigester, degraded waste as substrate in second

biodigester, and degraded waste mixed with swine manure in ratio 1:1 as substrate in

third biodigester. The highest concentration of biogas was found in third biodigester

that was 44.33 per cent of methane and 63.71 per cent of carbon dioxide. The lower

concentration at 24.90 per cent of methane and 18.98 per cent of carbon dioxide was

exhibited in secondary biodigester whereas the lowest was found in non-degraded

waste biodigester. It was demonstrated that the biogas production was greatly

increased with the initial grease waste degradation by P. aeruginosa.

Fungal mediated degradation of chopped and moist paddy straw was

investigated using Trichoderma reesei MTCC 164 and Coriolus versicolor MTCC

138 by Phutela et al, (2011). Paddy straw pretreated with T. reesei MTCC 164

showed 20.8 per cent enhanced biogas production after 25 days treatment. The paddy

straw treated with C. versicolor MTCC 138 showed 26.2 per cent enhanced biogas

production. Tuesorn et al, (2013) assessed the potential of a lignocellulolytic

microbial consortium for enhancing biogas production from fibre rich swine manure.

It led to increase in biogas production under mesophilic and thermophilic conditions

34

and the greatest enhancement was observed at 37°C. Biological degradation of

organic waste is normally facilitated by enzymes extracted from microbes. The

addition of exogenous enzymes can improve the performance of anaerobic digestion

systems (Ramachandran et al., 2007). Sonakya et al, (2001) pretreated wheat grains

with cellulase, a-amylase and protease prior to anaerobic digestion and observed an

increase in methane production by 7.0 - 14.0 per cent. Romano et al, (2009) studied

about the effects of the addition of enzyme (cellulase, hemicellulase, and

b-glucosidase) to anaerobic digestion of tall wheat grass. Anaerobic digestion tests

were performed using batch reactors operated at 50°C. The enzyme products showed

positive effects on the solubilization of wheat grass and enhanced biogas production.

Crude oil degradation using microbes

The rate of biodegradation of crude oil was studied by Ekpo and Udofia,

(2008) using Micrococcus varians, B. subtilis and P. aeruginosa. The microbe

P. aeruginosa degraded 97.2 per cent of the oil introduced into the medium followed

by M. varians with 85.7 per cent degradation. The least was B. substilis with 72.3

per cent degradation of the oil. P. aeruginosa was found to have the highest rate of

degradation. Zhang et al, (2005) reported about the potential biodegradation of crude

oil using P. aeruginosa. Christova et al, (2004) analysed the degradation ability of

B. subtilis 22BN strain. Sathiskumar et al, (2008) reported about the degrading

ability of Bacillus sp. IOS1-7, Corynebacterium sp. BPS2-6, Pseudomonas sp.

HPS2-5, and Pseudomonas sp. BPS1-8. The mixed bacterial consortium showed

more growth and degradation than did individual strains. At 1.0 per cent crude oil

concentration, the mixed bacterial consortium degraded a maximum of 77.0 per cent

of the crude oil. This was followed by 69 .0 per cent by Pseudomonas sp. BPS1-8,

64.0 per cent by Bacillus sp. IOS1-7, 45 per cent by Pseudomonas sp. HPS2-5, and

41.0 per cent by Corynebacterium sp. BPS2-6. The percentage of degradation by the

mixed bacterial consortium decreased from 77.0 to 45.0 per cent as the concentration

of crude oil was increased from 1.0 to 12.0 per cent. The temperature of 35ºC and pH

7.0 were found to be optimum for maximum degradation. Aparna et al, (2011)

investigated about the hydrocarbon degrading Pseudomonas sp. for efficient oil

degradation. Liang et al, (2005) studied the biodegradation of crude oil using

35

rhamnolipids by P. aeruginosa. Christova et al, (2004) used B. subtilis for the

enhanced degradation of hydrocarbon.

Lipase and protease as detergent additive

The enzyme based detergents have better cleaning properties as compared to

synthetic detergents and active at low washing temperatures and environment

friendly also. The enzymes in the detergents do not lose their activity after removing

stain and the enzyme containing detergents also improve the fabric quality and

keeping color bright. The enzyme based detergents are used in small quantity as

compared to synthetic chemicals and work at very low and high temperature,

environment friendly and completely biodegradable. They have specific cleaning

action and can also be used at lower temperatures and produce effluents with lower

COD and noncorrosive nature (Dsouza and Mawson, 2005). Enzymes are used in

detergents to increase the cleaning ability of detergents and can be used for an

alternative to chlorine bleach for removing stains on cloth. Enzymes used in

detergents must be effective at low levels, compatible with various detergent

components and to be active at wide range of temperatures. The enzymes are used in

detergent preparations and it should have the ability to withstand the other

components of the detergents and also stable at high pH values. Enzymes can reduce

the environmental load of detergent products because they save energy by enabling a

lower wash temperature to be used (Vakhlu and Kour, 2006). Addition of lipase in

detergent make it biodegradable, leaves no harmful residues, have no negative

impact on sewage treatment processes and do not have risk for aquatic life (Joseph et

al., 2007). Detergent enzymes should be cost-effective and safer to use. Bacterial

lipases added to household detergents reduce or replaces synthetic detergents, which

have considerable environmental problems. Detergent lipases are especially selected

to meet the following requirements like a low substrate specificity, ability to

withstand relatively harsh washing conditions (pH 10.0 - 11.0 and 30 - 60º C) and

ability to withstand damaging surfactants and enzymes which are important

ingredients of many detergent formulations. Proteases and other enzymes used in

detergent formulations should have high activity and stability over a broad range of

36

pH and temperature and are effective at low levels of 0.4 to 0.8 per cent (Rahman

et al., 2006; Hasan et al., 2010).

A presoak formulation was developed using an alkaline lipase produced by

Trichosporon asahii MSR 54, used for removing oil stains at ambient temperature

(Kumar et al., 2009). Lipase isolated from a thermo alkalophilic Pseudomonas

species was used as additive to improve the degree of olive oil removal from cotton

fabric in the presence of surfactants. The application of lipase from Pseudomonas

species act as an additive which has improved oil removal by 36.0 per cent at 70°C

for 20 min (Khoo and Ibrahim, 2009). A detergent stable lipase was produced from

Burkholderia cepatia (Rathi et al., 2001). Suzuki, (2001) got patent for an alkaline

pseudomonas lipase which was active at low temperatures and improved the wash

performance. A. niger is a highly potent fungus used in the production of alkaline

protease. The enzyme was most compatible with commercial detergent Tide (Dubey

et al., 2010). Khoramnia et al, (2011) studied about the detergent tolerance potentials

of lipase from Acinetobacter sp. Iftikar et al, (2011) investigated about the lipase

from Bacillus sp. was used as a good additive in detergents.

The protease enzyme produced from A. fumigatus retain 100 per cent activity

with detergents like Wheel and Vim at 37ºC for 30 min indicated the possible

suitability of its exploitation in detergent industries (Wathore and Patil, 2008). The

most potent thermophilic bacterial isolate Shewanella putrefaciens EGKSA21

showed alkaline thermostable protease production at optimum condition of 50°C and

pH 9.0. The crude enzyme of this bacterial strain was used as a detergent additive

(Bayoumi and Bahobil, 2011). Shukla et al, (2009) reported the detergent activity of

Staphylococus sp. isolated from Uttarakhand, India which showed good stability and

compatibility with Surf Excel detergent. Saraswathy et al, (2008) studied about the

laundry applications of protease from Bacillus sp. KCT I and the compatibility

studies showed that the enzyme was able to retain 50.0 per cent of initial activity

after 1.5 hrs incubation in the detergent solution and the enzyme was found to be

effective for the removal of blood and egg yolk stains in cotton cloth. The wash

performance analysis of alkaline protease produced by soil bacterium Bacillus

sp.GOS 2 was studied by Selvakumar et al, (2008). The enzyme showed more

37

compatibility with Power detergent powder. Ahmed et al, (2011a) studied about the

detergent compatibility of purified protease from A. niger.

Role of lipase and protease in dehairing of hides

Soaking, dehairing of hides and bating are traditionally being carried out by

using different chemicals and results in serious pollution effects. Both alkaline stable

and acid active lipases can be used in skin and hide degreasing and lipases offer the

tanner two advantages over solvents or surfactants: fat dispersion and production of

waterproof and low-fogging leathers. If surfactants alone are used for sheepskins,

they are usually not as effective and may be harmful to the environment while lipase

enzymes can remove fats and grease from skins and hides, particularly those with a

moderate fat content. Both alkaline stable and acid active lipases can be used in skin

and hide degreasing, deliming and bating. Lipases hydrolyse triglyceride to glycerol

and free fatty acids. Acid active lipases can be used to treat skins that have been

stored in a pickled state and alkaline lipases are applied for soaking and liming with

protease for efficient results. Lipases with acidic nature be applied for pickled skin or

wool and fur (Hasan et al., 2006). The alkaline proteases induced the swelling of hair

roots and allow easy removal of hair and for bating process it enhanced desired

softness and tightness of leather in a short time. Alkaline proteases with keratinolytic

activity have been reported for remarkable dehairing properties (Mukherjee et al.,

2008).

Verma et al, (2011) showed the use of protease from Thermoactinomyces sp.

RM4 for dehairing goat hides. Bacillus cereus MCM B-326, isolated from buffalo

hide, produced an extracellular protease that exhibited molecular weights of 45.0

kDa and 36.0 kDa. The enzyme could be effectively used to remove hair from

buffalo hide indicating its potential in leather processing industry (Nilegaonkar et al.,

2007). Alkaline protease was produced in a lab scale fermentor using B. subtilis

IH 72 and was applied to the goat skin for the removal of hair. The best results with

the skin processing were obtained, when the skin was treated with crude enzyme in

combination with 7.0 per cent lime sulphide. The quality of pelt (color, grain, stretch,

scud) and physical properties of the finally prepared leather (tensile strength, tear

strength, bursting strength etc) were also improved with the use of proteolytic

38

enzymes produced by B. subtilis IH 72 (Muktar and Haq, 2008). The protease

produced from A. niger and P. aeruginosa were used for dehairing of goat skin

(Patre and Dawande, 2010). The protease enzyme produced from S. nogalator was

used for dehairing of goat skin (Mitra and Chakrabartty, 2005). Zambare et al,

(2007) studied about the protease enzyme produced from B. cereus MCM B-326 and

its application as dehairing agent in buffalo hide. In the control sample, hair

loosening was not observed, even by mechanical means such as plucking by forceps.

The enzyme treated hide showed visible dehairing activity after 12 hrs of incubation

and complete dehairing at 21 hrs. Arunachalam and Saritha, (2009) reported about

the use of protease from B. subtilis for dehairing process. The enzyme activity found

maximum at 45oC and at pH 11.0. An index of dehairing comparable to the use of

conventional sodium sulfide method was achieved in 7 hrs of its application on wet

goat skin. Enzyme had the potential for replacing sodium sulfide in the dehairing

process of the leather industry.