List of Publications

Papers Published

1. Deepak Pant, U.G. Reddy and Alok Adholeya. 2006. Cultivation of oyster

mushrooms on wheat straw and bagasse substrate amended with distillery effluent.

World Journal of Microbiology and Biotechnology 22: 267-275 (IF-

0.634).

2. Deepak Pant and Alok Adholeya. 2006. Isolation and screening of potential fungi

for decolourization of distillery wastewaters. In: Modern Multidisciplinary Applied

Microbiology: Exploiting Microbes and Their Interactions. Mendez-Vilas, Antonio

(Ed.). Pp. 95–102. Wiley-VCH, Weinheim.

3. Deepak Pant and Alok Adholeya. 2007. Biological approaches for treatment of

distillery wastewater: A review. Bioresource Technology 98: 2321-2334 (IF-

2.180)

4. Deepak Pant and Alok Adholeya. 2007. Enhanced production of ligninolytic

enzymes and decolorization of molasses distillery wastewater by fungi under solid

state fermentation. Biodegradation 18: 647-659 (IF-1.714)

5. Deepak Pant and Alok Adholeya. 2007. Identification, ligninolytic enzyme activity

and decolorization potential of two fungi isolated from a distillery effluent

contaminated site. Water, Air and Soil Pollution 183: 165-176 (IF-1.258)

6. Deepak Pant and Alok Adholeya. 2008. Nitrogen removal from biomethanated

spentwash using hydroponic treatment followed by fungal decolorization. (Revised

manuscript submitted in Environmental Engineering Science).

7. Deepak Pant and Alok Adholeya. 2008. Enzyme concentration by ultrafiltration

after fungal growth on solid-state fermentation. (Manuscript prepared for Journal

of Basic Microbiology).

Abstracts Published

1. Microbial diversity of industrially contaminated soil and water: community structure

and role in bioremediation. In: International conference on microbial diversity:

Current perspectives and potential applications (Microbial Diversity 2005),

9

List of publications

TERI University-Ph.D. Thesis, 2007

224

University of Delhi, South Campus, New Delhi during 16-18 April’ 2005. Abstract on

page 47.

2. Nitrogen removal from distillery wastewaters using plant based treatment system.

In: International workshop on Nitrogen in Environment, Industry and Agriculture,

Indian National Science Academy, New Delhi, India. 16-17 March’ 2006. Page 68-71.

3. Reclamation of industrial wasteland and development of green cover using

mycorrhizae: A demonstration at two industrial sites in India. In: 5th International

Conference on Mycorrhizae "Mycorrhiza for Science and Society" (ICOM5). 23-27

July'2006. Granada, Spain. Abstract page 229. Awarded as best poster.

4. Phytomicrobial management of black liquor from an agro-residue based paper mill

attaining zero discharge. Poster presented at 8th Specialized Conference on Small

Water and Wastewater Systems (SWWS) and the 2nd Specialized Conference on

Decentralised Water and Wastewater International Network (DEWSIN),

Coimbatore, India. February 06 – 08, 2008. Abstract page 103.

Cultivation of oyster mushrooms on wheat straw and bagasse substrate amended

with distillery effluent

Deepak Pant1, U. Gangi Reddy2 and Alok Adholeya1,2,*1Centre of Bioresources and Biotechnology, TERI School of Advanced Studies, DS Block, India Habitat Centre, LodhiRoad, 110003, New Delhi, India2Biotechnology and Management of Bioresources Division, The Energy and Resources Institute, DS Block, IndiaHabitat Centre, Lodhi Road, 110003, New Delhi, India*Author for correspondence: Tel.: +91-11-24682100, 24682111, Fax: +91-11-24682144, 24682145,E-mail: aloka@teri.res.in

Received 27 May 2005; accepted 24 July 2005

Keywords: Biological efficiency, distillery effluent, dry matter loss, Pleurotus spp., substrate amendment, yield

Summary

Molasses-based distilleries produce large quantities of dark coloured effluent, which is a major cause of environ-mental pollution. An experiment was conducted to investigate the efficacy of distillery effluent amendment for ediblemushroom production. Three species of oyster mushroom, namely Pleurotus florida Eger (EM 1303), Pleurotuspulmonarius (Fries) Quelet (EM 1302) and Pleurotus sajor-caju (Fries) Singer (EM 1304) were grown on wheat straw(variety UP 2338) and bagasse amended with post-anaerobic distillery effluent, a high organic load wastewater withhigh biochemical oxygen demand and chemical oxygen demand. Three different levels of effluent treatment wereapplied to bagasse and wheat straw. Wheat straw was found to be the preferred substrate and showed better resultsthan bagasse in all treatments with respect to yield, biological efficiency (BE) and dry matter loss. P. florida (EM1303) and P. pulmonarius (EM 1302) gave significantly enhanced yield with increasing levels of effluent, with BEreaching highest at 238.6% for P. florida (EM 1303). Using bagasse as a substrate, P. sajor-caju (EM 1304) andP. pulmonarius (EM 1302) exhibited a decreasing trend as compared to control. However, the effect of effluentconcentrations did not influence yield and BE significantly in case of bagasse. The dry matter loss of the substratevaried from 9.4% to 53.4% in wheat straw and 17.5% to 45.2% in bagasse respectively.

Introduction

Distilleries are among the most polluting industries astheir effluents, if discharged into water bodies, defile thenatural ecosystem. The wastewater from distilleries usingmolasses as the main substrate for fermentation is char-acterized by very high organic pollutant load. The bio-chemical oxygen demand (BOD) and chemical oxygendemand (COD) typically range between 35,000–50,000and 100,000–150,000 mg/l, respectively (Nandy et al.2002). An averagemolasses-based distillery generates 15 lof spent wash/l of alcohol produced. This effluent ischaracterized by its colour, high temperature, low pH,high ash content and high percentage of dissolved organicand inorganic matter (Beltran et al. 1999). Presently,there are 319 distilleries in India producing 3.25�109 l ofalcohol and generating 40.4�1010 l of wastewater annu-ally (Uppal 2004). This molasses spent wash (MSW) is apotential water pollutant in two ways. First, the highlycoloured nature of MSW can block out sunlight fromrivers and streams, thereby reducing oxygenation of thewater by photosynthesis and hence becomes detrimentalto aquatic life. Secondly, the MSW has a high pollution

load which would result in eutrophication of contami-natedwater courses (FitzGibbon et al. 1998). In addition,if disposed of on land, the spent wash functions as a soilpollutant with an ability to inhibit seed germination, re-duce soil alkalinity, cause soil manganese deficiency anddamage agricultural crops (Kannabiran & Pragasam1993; Ramana et al. 2002). However, organic wastescontained in distillery effluent are valuable source of plantnutrients especially N, P, K and organic substrates ifproperly utilized (Pathak et al. 1999).A number of fungi can be used to detoxify contami-

nated environments in the process of bioremediation.The white rot fungi (e.g. Phanerochaete chrysosporium)and brown rot fungi (e.g. Gloephyllum species) are themost widely used (Stamets 2000). Among the white rotfungi, oyster mushrooms of genus Pleurotus are wellknown for conversion of substrate mass into mush-rooms. Moreover, they are also the easiest and leastexpensive commercial mushroom to grow (Banik &Nandi 2004). This fungus is industrially producedfor human consumption, for the bioconversion ofagricultural and industrial lignocellulose waste and as asource of enzymes and other chemicals for industrial

World Journal of Microbiology & Biotechnology (2006) 22:267–275 � Springer 2005DOI 10.1007/s11274-005-9031-2

and medical applications. Pleurotus can be grown on awide range of substrates, agricultural sub-products suchas rice straw, wheat straw, sawdust, corn cobs (Poppe &Hofte 1995) and industrial wastes. Among these, pas-teurized wheat straw is the most commonly used sub-strate (Philippoussis et al. 2001). In the course ofdecomposing dry straw, nearly 50% of the mass is lib-erated as gaseous carbon dioxide, 20% is lost as residualwater, 20% remains as ‘spent’ compost and 10% isconverted into dry mushrooms (Stamets 2000). Theeconomic value of mushroom cultivation is the ability ofthese fungi in transforming inedible agricultural waste toedible biomass which is generally accepted as food ofhigh quality, flavour and nutritive value (Quimio 1978).Wheat production generates a considerable amount of

straw every year. This is an enormous underutilizedenergy resource, but of great potential as feed forruminants, and as raw material for the cultivation ofedible mushrooms (Zadrazil et al. 1996). According toAksu et al. (1996), production of Pleurotus spp., mostly-based on cereal straw, is second among oyster mush-rooms after Agaricus bisporus, with a share of 24.2% ofworld production. Wheat straw degradation is achievedby the influence of enzymes from P. ostreatus, particu-larly of cell-wall components, cellulose and lignin (Ad-amovic et al. 1998). Sugarcane bagasse is one of thelargest cellulosic agro-industrial by products, a fibrousresidue of cane stalk left over after the crushing andextraction of the juice from sugarcane. Use of bagasse isadvantageous due to its low ash content compared torice straw and wheat straw. Rice straw and wheat strawhave 17.5% and 11.0% ash contents respectively forusage in bioconversion processes using microbial cul-tures (Pandey et al. 2000).Very few studies have used industrial wastewaters as a

substrate amendment in the practice of mushroom cul-tivation. Wastewaters from a rice spirit distillery havebeen used as a substrate in solid-state fermentation ofGanoderma lucidum and also for culturing the yeastSaccharomyces cerevisiae (Yang & Tung 1996; Yanget al. 2003). Despite the rich organic content of distillerywaste, its use for mushroom cultivation has not beeninvestigated. Thus, the present work was carried outwith an objective of using agricultural wastes such aswheat straw and bagasse amended with distillery effluentas a substrate for mushroom cultivation, and to identifyappropriate mushroom strains that are well suited togrow on distillery effluent amended substrates. Theinformation thus obtained would be relevant for futureapplications using distillery effluent as substrateamendment for healthy mushroom production.

Methods

Distillery effluent

The post-anaerobically digested effluent was providedby Associated Alcohols and Breweries Limited (a cane

molasses-based distillery), Barwaha, Madhya Pradesh,Central India. Physico-chemical characterization of thiseffluent was carried out using standard methods (APHA1995). Reducing sugars were estimated using the dini-trosalicylate (DNS) method (Miller 1959) and totalsugars using the method described by Somogyi (1952).The effluent was found to contain potassium, sodiumand nitrogen in high concentrations, and in addition,manganese, magnesium, and other trace elements. Thedata is presented in Table 1.

Fungal strains

Three strains of Pleurotus, namely P. florida (EM 1303),P. pulmonarius (EM 1302) and P. sajor-caju (EM 1304),were procured from the Centre for Mycorrhizal CultureCollection (CMCC), TERI, New Delhi, India. Thesewere maintained and subcultured on Potato DextroseAgar (PDA) media (Hi Media, India) plates at 25 �C.

Spawn preparation

Spawn preparation was carried out according to themethod described by Stamets (2000). Three fungal discsof 7 mm diameter were cut from the edge of an activelygrowing colony and placed equidistantly from eachother in petri dishes (100�15 mm) containing PDAmedium. When the mycelial mat covered the entiresurface of the petri dish, a liquid culture medium wasprepared with the following composition: 40 g barleymalt sugar, 4 g hardwood sawdust, 2 g yeast, 1 g cal-cium sulphate and 1000 ml water. After mixing andsubdividing 750 ml of the broth into three 1500 mlErlenmeyer flasks, the flasks were sterilized in an auto-clave for 1 h at 121 �C.The mycelium from each petri dish was sectioned into

discs of 7 mm diameter with a heat-sterilized cork borerand aseptically transferred into the three 1500-mlErlenmeyer flasks. Each flask was incubated at roomtemperature at 150 rev/min for 48 h or till the mycelium

Table 1. Physico-chemical characteristics of distillery effluent (used in

the experiments).

Parameter Anaerobically treated

effluent (released in field)

Electrical conductivity (mS/cm) 33.16

PH 8.20

BOD5 (ppm) 5000

COD (ppm) 25,000

Total Kjeldahl Nitrogen (%) 3.50

Sodium (ppm) 500

Potassium (ppm) 2500

Manganese (ppm) 259.44

Magnesium (ppm) 98.00

Zinc (ppm) 272.97

Copper (ppm) 395.51

Total dissolved solids (ppm) 21,256

Total sugar (%) 2.80

Reducing sugar (%) 0.23

268 D. Pant et al.

was rendered in liquid form. This was done to achievevigorous regrowth. After 2–4 days of incubation in thenutrient-enriched broth, each flask hosted thousands ofstellar-shaped colonies of mycelium. This suspensionwas used to inoculate sterilised wheat grain filled glassjars at the rate of 30 ml/2-l jar.Wheat grain (1.5 kg) soaked in water (400 ml) was

precooked by autoclaving at 121 �C for 30 min. Oncooling, 50 g mixture of lime (CaCO3) and gypsum(CaSO4) in a ratio of 3:2 (w/w) was thoroughly mixedwith the moist wheat grain. The grains were then kept ina glass jar and tightly closed with a lid. This was againautoclaved for 15 min at 121 �C and 15 lb pressure. Thesterile wheat grain mixture was aseptically inoculatedwith 50 ml liquid inoculum suspension. The jars wereincubated in the dark at room temperature till the fungalmycelium covered the entire substrate.

Substrate preparation and inoculation

Wheat straw of cultivar UP 2338 was procured from acommercial supplier, Trace Scientific, New Delhi, Indiaand bagasse from Simbhaoli Sugar Mills Limited,Ghaziabad, Uttar Pradesh, India. The water-holdingcapacity (maximum liquid retention) was estimated as3.5 l water/kg in case of wheat straw and 3.0 l water/kgin case of bagasse. The chopped wheat straw (4 to 8 cm)and bagasse (3 to 5 cm) were soaked for 24 h in tap water(control). For the different treatments, distillery effluentwas added as 15, 30 and 50% (by volume) in tap water.All treatments were done in triplicate. After removingexcess liquid by draining manually, the substrate waspacked in pre-sterilized polypropylene bags (height20 cm, diameter 15 cm). Each bag was filled with 1.5 kgof substrate, sterilized in the autoclave for 1 h at 121 �C.The straw was left to cool overnight at 25 �C and inoc-ulated aseptically with spawns of P. florida (EM 1303), P.pulmonarius (EM 1302) and P. sajor-caju (EM 1304) atan amount of 2% (w/w) of substrate fresh weight.

Mushroom growth and harvesting

The mushrooms were grown in a 2.75�2.70�2.60 m3

environmental chamber where temperature (24±2 �C)and relative humidity (85±90%) were precisely con-trolled. The CO2 level was maintained by periodic aer-ation through the chamber. The inoculated bags wereplaced in the chamber with top of each bag sealed untilthe substrate in the bags was fully colonized. When thebags had been heavily colonized by the fungal mycelium,equal numbers of holes were punched in the substratebags for drainage and aeration. Two flushes of mush-rooms were harvested from each bag. At each harvest-ing, mushroom fruit bodies in each bag were pickedmanually by using sterile scalpel to collect wholemushroom clusters. Later these were cut into individualmushrooms and weighed for estimating the net yield.The substrate in the bag was also weighed before andafter the experiment for assessing the substrate drymatter loss. The experiment was repeated twice.

Statistical analysis

The biological efficiency was defined as the grams offresh mushrooms produced per 100 g of dry straw(substrate) used (Velazquez-Cedeno et al. 2002). Meanof three replications was considered as the final readingfor analysis. Data were analysed using Analysis of Var-iance (ANOVA) using Costat software (Cohort, Berke-ley, California). Significant means were compared by theDuncan’s Multiple Range Test (DMRT) at P£ 0.05 andstandard error of mean values (SEM) was also calcu-lated. All statistical analysis was conducted using theJandel Sigma Plot Version 9.0 (SAS Institute Inc. 2004).

Results

The mushroom yield, biological efficiency and substratedry matter losses after mushroom growth on wheatstraw and bagasse substrates, amended with differentconcentrations of distillery effluent are represented inFigures 1 and 2 respectively. The concentrations ofeffluent used were considered at 15, 30 and 50% (v/v)since further higher concentrations imposed problems ofsterility due to contamination from green moulds. Also,the relative yields obtained at higher concentrationswere comparatively much lower for satisfying the aim ofachieving maximum yield of mushrooms at a selectedconcentration of distillery effluent.

Wheat straw amended with distillery effluent

The observations revealed variable difference among thetreatments and exhibited statistically significant differ-ence in yield and biological efficiency in all threemushroom species with respect to the different effluentconcentrations. The representative experimental units ofP. sajor-caju (EM 1304) and P. pulmonarius (EM 1302)growing on wheat straw amended with distillery effluent-based substrate are shown in Figure 3. Maximum yieldwas observed in both P. florida (EM 1303) and P. pul-monarius (EM 1302) when substrate was amended with50% effluent, but in P. sajor-caju (EM 1304) yield wasfound to be significantly higher over the control at 15%treatment only, after which the yield declined on furtherincreasing the effluent concentration. Biological effi-ciency (BE) was found to be highest (238.6%) inP. florida (EM 1303) at 50% effluent amended treat-ment. There was no significant difference observed inyield and BE in case of P. pulmonarius (EM 1302) at 15and 30% effluent amended treatments. However, bothwere significantly higher at 50% effluent treatment.Substrate dry matter loss was significant in all the

treatments as well as control for P. sajor-caju (EM1304). Non-significant differences were observed in thecontrol, 15 and 30% effluent amended treatments forP. pulmonarius (EM 1302). However, here the drymatter loss was significantly higher at 50% effluenttreatment. Dry matter loss was observed significantly

Mushroom growth using distillery effluent 269

Control 15% 30% 50%

Mus

hroo

m y

ield

(g)

0

500

1000

1500

2000

2500

3000

Pleurotus florida

Pleurotus pulmonariusPleurotus sajor-caju

b

c

b

a

bc c

a

b

a

c

d

Control 15% 30% 50%

Bio

logi

cal E

ffici

ency

0

50

100

150

200

250

300

Pleurotus florida

Pleurotus pulmonariusPleurotus sajor-caju

b

c

b

a

bc c

a

b

a

cd

Treatments

Control 15% 30% 50%

Sub

stra

te D

ry M

atte

r Lo

ss(%

)

0

10

20

30

40

50

60

Pleurotus florida

Pleurotus pulmonariusPleurotus sajor-caju

b

b

aa

bb b

a

b

a

c

d

Figure 1. Comparison of mushroom yield, biological efficiency and substrate dry matter loss of three Pleurotus sp. grown on wheat straw as

substrate over different amendments of distillery effluent. Letters above the histogram bars represents Analysis of Variance (ANOVA). Bars with

different letters indicate values with significant difference. Bars represent means of three replicates at P £ 0.05.

270 D. Pant et al.

Control 15% 30% 50%

Bio

logi

cal E

ffici

ency

0

10

20

30

40

50

60

Pleurotus florida Pleurotus pulmonariusPleurotus sajor-caju

Control 15% 30% 50%

Mus

hroo

m Y

ield

(g)

0

200

400

600

800

1000

Pleurotus floridaPleurotus pulmonariusPleurotus sajor-cajua

bc d

a ab bc

a

bc

d

a

bc

d

a aab

b

a

c b

d

Treatments

Control 15% 30% 50%

Sub

stra

te D

ry M

atte

r Lo

ss(%

)

0

10

20

30

40

50

Pleurotus florida

Pleurotus pulmonariusPleurotus sajor-caju

a

b

c

d

bb

a

ca

bb

b

Figure 2. Comparison of mushroom yield, biological efficiency and substrate dry matter loss of three Pleurotus sp. grown on bagasse as substrate

over different amendments of distillery effluent. Letters above the histogram bars represents Analysis of Variance (ANOVA). Bars with different

letters indicate values with significant difference. Bars represent means of three replicates at P £ 0.05.

Mushroom growth using distillery effluent 271

higher at 30% and 50% treatment with respect to con-trol in case of P. florida (EM 1303). Regression analysisbetween mushroom yield (Y) and substrate dry matterloss (X) revealed a significant positive relationship for P.pulmonarius (EM 1302) (P = 0.0005; Y = 10.86X +475.83; �r2 = 0.686) and a significant negative correla-tion in case of P. sajor-caju (EM 1304) (P = 0.0005;Y = 41.74X – 223.88; �r2 = 0.894).

Bagasse amended with distillery effluent

Significant differences were observed in yield, BE anddry matter loss in all three species. The representativeexperimental units of P. florida (EM 1303) and P.pulmonarius (EM 1302) growing on bagasse amendedwith distillery effluent-based substrate are shown in

Figure 4. In case of P. florida (EM 1303), there wassignificant decline in yield, BE and substrate dry matterloss with increase in effluent concentrations. P. sajor-caju (EM 1304) and P. pulmonarius (EM 1302) showeda decreasing trend in yield and BE as compared to thecontrol. However, the different effluent concentrationsdid not influence the yield or the BE. Dry matter losswas non significant between control and 15% effluenttreatment for P. florida (EM 1303) and P. pulmonarius(EM 1302), but was observed significantly reduced at30% effluent amended treatment. Regression analysisbetween mushroom yield (Y) and substrate dry matterloss (X) revealed a significant positive relationship forP. pulmonarius (EM 1302) (P = 0.001; Y = 3.275X +395.38; �r2= 0.578). Here also, as in case of wheatstraw, a significant negative relationship was observed

Figure 3. (a) Pleurotus sajor-caju growing on hanging wheat straw substrate amended with distillery effluent. (b) Wheat straw amended with

distillery effluent supporting extensive Pleurotus pulmonarius production.

272 D. Pant et al.

in case of P. sajor-caju (EM 1304) as observed byregression analysis (P = 0.001; Y = 34.63X – 119.52;�r2= 0.596).

Discussion

The oyster mushroom, Pleurotus spp. is one of the mostcommercially important and successfully cultivatedspeciality mushrooms and is considered to be a delicacy(Chang 1996). In the present investigation, use of dis-tillery effluent as an amendment for mushroom pro-duction with three Pleurotus spp. was demonstrated.Response of each mushroom species varied significantly

with different amendments of effluent on wheat straw.However, this was not the case in bagasse, indicatingthat the fungus was not dependent on external source, asbagasse already contains a large quantity of residualsugar, readily available as nutrient source for the fun-gus. This cultivation of Pleurotus spp. on various ligni-nocellulosic materials has been extensively investigatedand well documented in several reports (Bisaria et al.1987; Madan et al. 1987). Zadrazil (1980) showed thatP. sajor-caju has a very high saprophytic colonizingability and can degrade wheat straw efficiently. Croan(2000) reported the use of wood waste to cultivatewood-inhabiting ligninolytic white-rot basidiomycetesof genus Pleurotus.

Figure 4. (a) Image showing extensive production of Pleurotus florida growing on bagasse amended with distillery effluent. (b) Bagasse substrate

covered with extensive mycelial network showing initiation of primordia of Pleurotus pulmonarius.

Mushroom growth using distillery effluent 273

In our study, the dry matter loss was observed to be inagreement with the mushroom yield and biologicalefficiency. The higher mushroom yield and biologicalefficiency corresponded to the higher dry matter loss.The dry matter loss of the substrate ranged from 9.4%[at 50% concentration for P. sajor-caju (EM 1304)] to53.4% [at 50% concentration for P. florida (EM 1303)]on wheat straw. Similarly, in case of bagasse, thereduction varied from 17.5% [at 30% concentration forP. sajor-caju (EM 1304)] to 45.2% [at 30% concentra-tion for P. florida (EM 1303)]. These results agree withfindings of Zhang et al. (2002) where rice and wheatstraw were used as substrate without nutrient supple-mentation for cultivation of P. sajor-caju. In the presentinvestigation, the biological efficiency was found in therange 33.1–238.6% for wheat straw and 33.9–46.9% forbagasse. Because most fresh mushrooms containapproximately 90% water (Zhang et al. 2002), it ispossible for the BE to be above 100%. This suggests thatthe amendment of substrate with distillery effluent couldbe beneficial as a nutrient supplement as well as pro-viding moisture to the growing system. Thus, mushroomcultivation proves to be a highly economical method fordisposing of the agricultural residues, such as wheatstraw and bagasse, along with providing by-product inthe form of manure for field application. In this regard,Kakkar & Dhanda (1998) compared wheat and paddystraws for the growth of Pleurotus and obtained a muchhigher yield of fruiting bodies with wheat straw thanwith paddy straw. As a low cost substrate, bioconver-sion of spent larval medium of Corcyra cephalonica forcultivation of P. sajor-caju was investigated by Muthu-krishnan et al. (2000).The present work is the first report to the best of our

knowledge where post-anaerobically digested distilleryeffluent on being used as a substrate amendment withwheat straw gave enhanced oyster mushroom produc-tion and thus may be used as a suitable substrateamendment for mushroom cultivation. Previously, olivemill wastewater (OMWW) has been used by Kalmis &Sargin (2004), who reported that OMWW at 25 and50% concentrations was observed ideal for cultivationof P. sajor-caju and P. cornucopiae. This is importantfrom the point of view of resource recovery, which is notpossible in other treatment methods. We conclude thaton the basis of higher yields, BE along with increasedsubstrate dry matter loss, that distillery effluent appearsas a better option when used at lower concentrations assubstrate amendment on wheat straw for oyster mush-room cultivation.

Acknowledgements

Authors wish to thank Dr. R. K. Pachauri, Director-General, TERI, New Delhi, India for offering theinfrastructure for carrying out the present investigation.Financial assistance from University Grants Commis-

sion, New Delhi in form of Junior Research Fellowshipto the first author is duly acknowledged.

References

Adamovic, M., Grubic, G., Milenkovic, I., Jovanovic, R., Protic, R.,

Sretenovic, L. & Stoicevic, Lj. 1998 The biodegradation of wheat

straw by Pleurotus ostreatus mushrooms and its use in cattle

feeding. Animal Feed Science and Technology 71, 357–362.

Aksu, S., Isik, S.E. & Erkal, S. 1996 Development of cultivated

mushroom production in Turkey and general characteristics of

mushroom establishment. In Fifth National Congress of Edible

Mushrooms, November 5–7, 1996, Turkey, pp. 1–13.

APHA 1995 Standard methods for the examination of water and

wastewater. 19th edn. American Public Health Association,

Washington, DC. ISBN 0875532233.

Banik, S. & Nandi, R. 2004 Effect of supplementation of rice straw

with biogas residual slurry manure on the yield, protein and

mineral contents of oyster mushroom. Industrial Crops and Prod-

ucts 20, 311–319.

Beltran, F.J., Garcia-Araya, J.F. & Alvarez, P.M. 1999 Wine distillery

wastewater degradation. 1. Oxidative treatment using ozone and

its effect on the wastewater biodegradability. Journal of Agricul-

tural and Food Chemistry 47, 3911–3918.

Bisaria, R., Madan, M. & Bisaria, V.S. 1987 Biological efficiency and

nutritive value of Pleurotus sajor-caju cultivated on different agro-

wastes. Biological Wastes 19, 239–255.

Chang, S.T. 1996 Mushroom research and development equality and

mutual benefit. In Mushroom Biology and Mushroom Products,

Royse D.J (ed.) pp. 1–10, State College: the Pennsylvania State

University. ISBN 1-883956-01-3.

Croan, S. 2000 Conversion of wood waste into value-added products

into edible and medicinal Pleurotus (F.) P. Karst. Species (Agari-

cales sl., Basidiomycetes). International Journal of Medicinal

Mushrooms 2, 73–80.

FitzGibbon, F., Singh, D., McMullan, G. & Marchant, R. 1998 The

effect of phenolic acids and molasses spent wash concentration on

distillery wastewater remediation by fungi. Process Biochemistry

33, 799–803.

Kakkar, V.K. & Dhanda, S. 1998 Comparative evaluation of wheat

and paddy straws for mushroom production and feeding residual

straws to ruminants. Bioresource Technology 66, 175–177.

Kalmis, E. & Sargin, S. 2004 Cultivation of two Pleurotus species on

wheat straw substrates. International Biodeterioration and Biodeg-

radation 53, 43–47.

Kannabiran, B. & Pragasam, A. 1993 Effect of distillery effluent on

seed germination, seedling growth and pigment content of Vigna

mungo (L.) Hepper (CVT9). Geobios 20, 108–112.

Madan, M., Vasudevan, P. & Sharma, S. 1987 Cultivation of Pleurotus

sajor-caju on different wastes. Biological Wastes 22, 241–250.

Miller, G.L. 1959 Use of dinitrosalicylic acid reagent for determination

of reducing sugar. Analytical Chemistry 31, 426–428.

Muthukrishnan, N., Venugopal, M.S. & Janarthanan, R. 2000 Recy-

cling spent larval food of Corcyra cephalonica Stainton for pre-

paring spawn and sporophore of Pleurotus sajor-caju (Fr.) Singer.

World Journal of Microbiology and Biotechnology 16, 265–270.

Nandy, T., Shastry, S. & Kaul, S.N. 2002 Wastewater management in

a cane molasses distillery involving bioresource recovery. Journal

of Environmental Management 65, 25–38.

Pandey, A., Soccol, C.R., Nigam, P. & Soccol, V.T. 2000 Biotechno-

logical potential of agro-industrial residues. I: Sugarcane Bagasse.

Bioresource Technology 74, 69–80.

Pathak, H., Joshi, H.C., Chaudhary, A., Kalra, N. & Dwivedi, M.K.

1999 Soil amendment with distillery effluent for wheat and rice

cultivation. Water, Air, Soil Pollution 113, 133–140.

Philippoussis, A., Zervakis, G. & Diamantopoulou, P. 2001 Biocon-

version of agricultural lignocellulosic wastes through the cultiva-

tion of the edible mushrooms Agrocybe aegerita, Volvariella

274 D. Pant et al.

volvacea and Pleurotus spp. World Journal of Microbiology and

Biotechnology 17, 191–200.

Poppe, J.A. & Hofte, M. 1995 Twenty wastes for twenty cultivated

mushrooms. In Proceedings of the 14th International Congress on

the Science and Cultivation of Edible fungi, Oxford, pp. 171–179.

Rotterdam, Balkema, ISBN 9054105712 (v.1) and 9054105720

(v.2).

Quimio, J.H. 1978 Introducing Pleurotus flabellatus for your dinner

table. Mushroom Journal 68, 282–283.

Ramana, S., Biswas, A.K. & Singh, A.B. 2002 Effect of distillery

effluent on some physiological aspects of maize. Bioresource

Technology 84, 295–297.

Stamets, P. 2000. Growing Gourmet and Medicinal Mushrooms, 3rd

edition, pp. 201–325, Berkley, California: Ten Speed Press, ISBN

1580081754.

Somogyi, M. 1952 Notes on sugar determination. Journal of Biological

Chemistry 195, 19–23.

Uppal, J. 2004 Water Utilization and Effluent Treatment in the Indian

Alcohol Industry – An Overview. In Liquid Assets, Proceedings of

Indo-EU workshop on Promoting Efficient Water Use in Agro-based

Industries, New Delhi 15–16 January 2004, pp. 13–19. TERI Press,

New Delhi, India.

Velazquez-Cedeno, M.A., Mata, G. & Savoie, J.M. 2002

Waste-reducing cultivation of Pleurotus ostreatus and Pleurotus

pulmonarius on coffee pulp: changes in the production of some

lignocellulolytic enzymes. World Journal of Microbiology and

Biotechnology 18, 201–207.

Yang, F.C. & Tung, H.L. 1996 Reuse of stillage from rice spirit for the

culture of the yeast Saccharomyces cerevisiae. Process Biochemistry

31, 617–620.

Yang, F.C., Hsieh, C. & Chen, H.M. 2003 Use of stillage grain from a

rice-spirit distillery in the solid-state fermentation of Ganoderma

lucidum. Process Biochemistry 39, 21–26.

Zadrazil, F. 1980 Influence of ammonia nitrate and organic supple-

ments on the yield of Pleurotus sajor-caju. European Journal of

Applied Microbiology and Biotechnology 9, 31–35.

Zadrazil, F., Kamara, D.N., Isikhuemhen, O.S., Schuhardt, F. &

Lachowsku, G. 1996 Bioconversion of lignocellulose into ruminant

feed with white rot fungi- review of work done at the FAL

Braunschweig. Journal of Applied Animal Research 10, 105–124.

Zhang, R., Xiujin, L. & Fadel, J.G. 2002 Oyster mushroom cultivation

with rice and wheat straw. Bioresource Technology 82, 277–284.

Mushroom growth using distillery effluent 275

This article was originally published in a journal published byElsevier, and the attached copy is provided by Elsevier for the

author’s benefit and for the benefit of the author’s institution, fornon-commercial research and educational use including without

limitation use in instruction at your institution, sending it to specificcolleagues that you know, and providing a copy to your institution’s

administrator.

All other uses, reproduction and distribution, including withoutlimitation commercial reprints, selling or licensing copies or access,

or posting on open internet sites, your personal or institution’swebsite or repository, are prohibited. For exceptions, permission

may be sought for such use through Elsevier’s permissions site at:

http://www.elsevier.com/locate/permissionusematerial

Autho

r's

pers

onal

co

py

Review

Biological approaches for treatment of distillery wastewater: A review

Deepak Pant a, Alok Adholeya a,b,*

a Centre of Bioresources and Biotechnology, TERI University, DS Block, India Habitat Centre, Lodhi Road, New Delhi 110 003, Indiab Biotechnology and Management of Bioresources Division, The Energy and Resources Institute, DS Block, India Habitat Centre,

Lodhi Road, New Delhi 110 003, India

Received 12 June 2006; received in revised form 13 September 2006; accepted 25 September 2006Available online 7 November 2006

Abstract

Effluent originating from distilleries known as spent wash leads to extensive soil and water pollution. Elimination of pollutants andcolour from distillery effluent is becoming increasingly important from environmental and aesthetic point of view. Stillage, fermenter andcondenser cooling water and fermenter wastewater are the primary polluting streams of a typical distillery. Due to the large volumes ofeffluent and presence of certain recalcitrant compounds, the treatment of this stream is rather challenging by conventional methods.Therefore, to supplement the existing treatments, a number of studies encompassing physico-chemical and biological treatments havebeen conducted. This review presents an account of the problem and the description of colour causing components in distillery waste-water and a detailed review of existing biological approaches. Further, the studies dealing with pure cultures such as bacterial, fungal,algal and plant based systems have also been incorporated. Also, the roles of microbial enzymes in the decolourization process have beendiscussed to develop a better understanding of the phenomenon.� 2006 Elsevier Ltd. All rights reserved.

Keywords: Colour removal; Distillery effluent; Enzymes; Melanoidin; Microorganisms; Wastewater treatment

1. Introduction

Production of ethanol from agricultural materials foruse as an alternative fuel has been attracting worldwideinterest because of the increasing demand for limitednon-renewable energy resources and variability of oil andnatural gas prices. In India this demand is projected togo up because of a law for mixing 5% ethanol with petroland further raising this amount to 10% (The gazette ofIndia, 2002). Besides this, the other common usages of eth-anol are in the form of industrial solvent and beverages. Inthe year 1999, there were 285 distilleries in India producing2.7 · 109 L of alcohol and generating 4 · 1010 L of waste-water each year (Joshi, 1999). This number has gone up

to 319, producing 3.25 · 109 L of alcohol and generating40.4 · 1010 L of wastewater annually (Uppal, 2004). Overthe years as the sizes and number of distilleries have grown,bigger conventional aerobic-treatment plants have beenbuilt to deal with the constantly increasing effluent vol-umes. Space and money to construct these installationsare the biggest hindrances for such investments (Fumiet al., 1995).

In India, there are a number of large-scale distilleriesintegrated with sugar mills. The waste products fromsugar mill comprise bagasse (residue from the sugarcanecrushing), pressmud (mud and dirt residue from juiceclarification) and molasses (final residue from sugarcrystallization section). Bagasse is used in paper manufac-turing and as fuel in boilers; molasses as raw material indistillery for alcohol production while pressmud has nodirect industrial application (Nandy et al., 2002). The efflu-ents from molasses based distilleries contain large amountsof dark brown coloured molasses spent wash (MSW). Inthe distillation process, ethanol ranges from 5% to 12%

0960-8524/$ - see front matter � 2006 Elsevier Ltd. All rights reserved.

doi:10.1016/j.biortech.2006.09.027

* Corresponding author. Address: Biotechnology and Management ofBioresources Division, The Energy and Resources Institute, DS Block,India Habitat Centre, Lodhi Road, New Delhi 110 003, India. Tel.: +91 1124682100/24682111; fax: +91 11 24682144/24682145.

E-mail address: aloka@teri.res.in (A. Adholeya).

Bioresource Technology 98 (2007) 2321–2334

Autho

r's

pers

onal

co

py

by volume, hence it follows that the amount of waste variesfrom 88% to 95% by volume of the alcohol distilled. Anaverage molasses based distillery generates 15 L of spentwash L�1 of alcohol produced (Beltran et al., 2001).MSW is one of the most difficult waste products to disposebecause of low pH, high temperature, dark brown colour,high ash content and high percentage of dissolved organicand inorganic matter (Beltran et al., 1999). The biochemi-cal oxygen demand (BOD) and chemical oxygen demand(COD), the index of its polluting character, typically rangebetween 35,000–50,000 and 100,000–150,000 mg L�1,respectively (Nandy et al., 2002).

Worldwide environment regulatory authorities are set-ting strict norms for discharge of wastewaters from indus-tries. In India for instance, distillery industry had been toldto achieve zero discharge of spentwash by December 2005according to the charter of Central Pollution ControlBoard, the apex pollution control authority (CPCB,2003). It further says that till 100% utilization of spentwashis achieved, controlled and restricted discharge of treatedeffluent from lined lagoons during rainy season will beallowed by SPCBs/CPCB in such a way that the perceptiblecolouring of river water bodies does not occur. Physical,chemical and biological treatment approaches have beenemployed for the treatment of distillery wastewater. Thisreview focuses mainly on lab and field scale biologicalapproaches. The overall objective of the work is to presenta literature review on the state of the art in this field andaddress the issues requiring further research.

2. Pollution and toxicity profile of distillery effluent

The production and characteristics of spentwash arehighly variable and dependent on feedstocks and variousaspects of the ethanol production process. Wash waterused to clean the fermenters, cooling water blow down,and boiler water blow down further contributes to itsvariability (Duarte et al., 1997). In a distillery, sources ofwastewater are stillage, fermenter and condenser coolingwater and fermenter wastewater. The liquid residues duringthe industrial phase of the production of alcohol are:liquor, sugar cane washing water, water from the condens-ers and from the cleaning of the equipment, apart fromother residual water. This extract is extremely pollutingas it contains approximately 5% organic material and fertil-izers such as potassium, phosphorus and nitrogen. Theamount of water used in this process is large, generatinga high level of liquid residues (Borrero et al., 2003).

The MSW is a potential water pollutant in two ways.First, the highly coloured nature of MSW can block outsunlight from rivers and streams, thus reducing oxygena-tion of the water by photosynthesis and hence becomes det-rimental to aquatic life. Secondly, it has a high pollutionload which would result in eutrophication of contaminatedwater courses (FitzGibbon et al., 1998). Due to the pres-ence of putriciable organics like skatole, indole and othersulphur compounds, the MSW that is disposed in canals

or rivers produces obnoxious smell (Mahimaraja andBolan, 2004). Undiluted effluent has toxic effect on fishesand other aquatic organisms. The estimated LC50 for dis-tillery spent wash was found to be 0.5% using a bio-toxicitystudy on fresh water fish Cyprinus carpio var. communis

(Mahimaraja and Bolan, 2004). Impacts of distillery efflu-ent on carbohydrate metabolism of freshwater fish, C. car-

pio were studied recently by Ramakritinan et al. (2005).The respiratory process in C. carpio under distillery effluentstress was affected resulting in a shift towards anaerobiosisat organ level during sublethal intoxication.

Spent wash also leads to significant levels of soil pollu-tion and acidification in the cases of inappropriate land dis-charge. It is reported to inhibit seed germination, reducesoil alkalinity, cause soil manganese deficiency and damageagricultural crops (Kannabiran and Pragasam, 1993; Agra-wal and Pandey, 1994). However, effect of distillery effluenton seed germination is governed by its concentration and iscrop-specific. In a study by Ramana et al. (2002) the germi-nation percent in five crops decreased with increase in con-centration of the effluent. The germination was inhibited inall the five crops studied with concentration exceeding 50%.At the same time, organic wastes contained in distilleryeffluent are valuable source of plant nutrients especiallyN, P, K and organic substrates if properly utilized (Pathaket al., 1999). For instance, distillery effluent in combinationwith bioamendments such as farm yard manure, rice huskand Brassica residues was used to improve the propertiesof sodic soil (Kaushik et al., 2005). The use of fungi forbioconversion of distillery waste into microbial biomassor some useful metabolites has been recently reviewed byFriedrich (2004). The end products of bioconversion arefungal biomass, ethanol, enzymes etc. and substantiallypurified and decolourized effluents. Recently enhanced pro-duction of oyster mushrooms (Pleurotus sp.) using distill-ery effluent as a substrate amendment have been reported(Pant et al., 2006).

3. Colorants in distillery wastewaters

The molasses wastewater from alcoholic fermentationhas a large amount of a brown pigment. The colour ishardly degraded by the conventional treatments and caneven be increased during anaerobic treatments, due torepolymerization of compounds. Phenolics (tannic andhumic acids) from the feedstock, melanoidins from Mail-lard reaction of sugars (carbohydrates) with proteins(amino groups), caramels from overheated sugars, andfurfurals from acid hydrolysis mainly contribute to the col-our of the effluent (Kort, 1979). During heat treatment, theMaillard reaction (non enzymatic reaction) takes placeaccompanied by formation of a class of compounds knownas Maillard products. The reaction proceeds effectively at>50 �C and it is favored at pH 4–7 (Morales and Jimnez-Perez, 2001). Melanoidins are one of the final products ofthe Maillard reaction. They are complex compounds withtheir structures not fully understood.

2322 D. Pant, A. Adholeya / Bioresource Technology 98 (2007) 2321–2334

Autho

r's

pers

onal

co

py

Melanoidin is one of the biopolymers that is hardlydecomposed by microorganisms and is widely distributedin nature. Melanoidins have antioxidant properties, whichrender them toxic to aquatic micro and macroorganisms(Kitts et al., 1993). However, melanoidins present in Span-ish sweet wines were studied by Rivero-Perez et al. (2002)and they reported that high molecular weight brown pig-ments (mostly hypothesized to be melanoidins) isolatedby dialysis were correlated with the colour of the winesbut not to their antioxidant activity. Melanoidins, orrelated formation products can occur in different processesof beverage manufacture, such as heat concentrated juicesand musts, beers or wines (Kroh, 1994). From studies using13C and 15N CP-NMR spectrometry, Hayase et al. (1986)confirmed the presence of olefinic linkages and conjugatedenamines which were suggested to be important for thestructure of the chromophores in melanoidin. The reduc-tion in intensity of absorption for the ozonated sampleindicates a cleavage of the –C@C– on ozonation.

For melanoidins formed from carbohydrates and aminoacids, a new model of a basic melanoidin skeleton mainlybuilt up from amino-branched sugar degradation productswas suggested by Cammerer et al. (2002). They indicatedthat oligo- and polysaccharides reacted in the Maillardreaction preferentially as complete molecules at the reduc-ing end under water-free reaction conditions. Anotherapproach to estimate the chemical structure of melanoidinswas suggested by Kato and Hayase (2002). A blue pigment(Blue-M1, C27H31N4O13) was isolated from the reactionmixture of D-xylose and glycine in 60% ethanol stored at26.5 �C for 48 h (or 2 �C for 96 h) under nitrogen, whosechemical character was comparable to that of a nondialyz-able melanoidin preparation obtained from the reactionmixture of D-xylose and butylamine neutralized with aceticacid in methanol incubated at 50 �C for 7 days. Recently,the empirical formula of melanoidin has been suggestedas C17�18H26�27O10N. The molecular weight distributionis between 5000 and 40,000. It consists of acidic, polymericand highly dispersed colloids, which are negatively chargeddue to the dissociation of carboxylic acids and phenolicgroups (Manisankar et al., 2004).

4. Spent wash treatment

Biological treatments have been recognized as effectivemethods of treatment for highly polluted industrial waste-waters. Both anaerobic and aerobic systems are commonlyused to treat the wastewaters from agro-industrial plantsincluding distilleries as well. In the recent years, increasingattention is also being directed towards utilizing microbialactivity (pure bacteria and fungi) for the decolourizationand mineralization of spent wash. There are several reportsciting the potential of microorganisms for use in this pro-cess. Moreover, the biologically treated effluent could beused safely and effectively to increase the soil productivity.This section is discussed in detail as anaerobic and aerobictreatments.

4.1. Anaerobic treatment

Anaerobic digestion is widely accepted as the first treat-ment step in distilleries. Wilkie et al. (2000) have reviewedthe role of anaerobic digestion in stillage (spent wash)treatment. Anaerobic digestion can convert a significantportion (>50%) of the COD to biogas, which may be usedas an inplant fuel, and also saves the energy that would berequired for aeration using aerobic treatment. At present,the anaerobic biological treatment of distillery effluents iswidely applied as an effective step in removing 90% of theCOD in the effluent stream (Wolmarans and de Villiers,2002). During this stage, 80–90% BOD removal takes placeand biochemical energy recovered is 85–90% as biogas.

A list of common types of anaerobic reactors used fordistillery effluent treatment is given in Table 1. Akunnaand Clark (2000) studied the performance of a granularbed anaerobic baffled reactor (GRABR) in the treatmentof a whisky distillery wastewater having COD and BODconcentrations of 16,600–58,000 and 8900–30,000 mg L�1,respectively. The removal of total BOD and COD fromthe wastewater were 80–92% and 90–96%, respectively witha HRT of 4 days and at a loading rate of 2.37 kg CODm�3 day�1.

The highest BOD removal is possible in open lagoonwhereas highest biomethane produced is in upflow anaero-bic sludge blanket (UASB) type bioreactor. The UASBsystem has become the most widely applied reactortechnology for high rate anaerobic treatment of industrialeffluents. Its relative high treatment capacity compared toother systems permits the use of compact and economicwastewater treatment plants. Compared to aerobic system,it has slow growth rate, mainly associated with methano-genic bacteria. Therefore, it requires a long retention time,and also only a small portion of the degradable organicwaste is being synthesized to new cells. Full-scale thermo-philic (50–55 �C) anaerobic digestion of wastewater froman alcohol distillery was reported by Vlissidis and Zoubou-lis (1993). More than 60% removal of COD was achievedwith 76% of biogas comprising of methane thus makingit a valuable fuel.

Goodwin and Stuart (1994) studied two identical UASBreactors operated in parallel as duplicates for 327 days forthe treatment of malt whisky pot ale and achieved CODreductions of up to 90% for influent concentrations of3526–52126 mg L�1. When the OLRs of 15 kg m�3 dayand above were used, the COD removal efficiency droppedto less than 20%, in one of the duplicate reactors. A meso-philic two-stage system consisting of an anaerobic filter(AF) and an UASB reactor was found suitable for anaero-bic digestion of distillery waste, enabling better conditionsfor the methanogenic phase (Blonskaja et al., 2003). Theoptimum conditions for the stable work of reactor are:for the acidogenic stage, organic loading of 2–4 kgCOD m�3 day�1 at pH 6.0 and for the methanogenic stage,organic loading of 1–2 kg COD m�3 day�1 at pH 7.6.An advanced version of UASB system was reported by

D. Pant, A. Adholeya / Bioresource Technology 98 (2007) 2321–2334 2323

Autho

r's

pers

onal

co

py

Driessen and Yspeert (1999), wherein they used an internalcirculation (IC) reactor characterized by biogas separationin two stages within a reactor with a high weight/diameterratio and the gas driven internal effluent circulation. Thissystem could handle high upflow liquid and gas velocitiesmaking possible treatment of low strength effluents at shorthydraulic retention times as well as treating high strengtheffluents such as from brewery at very high volumetricloading rates up to 35 kg COD m�3. Effect of addition ofmacronutrients and micronutrients in the distillery effluentwas investigated on the performance of simulated UASBsystem by Sharma and Singh (2001). Calcium and phos-phate were found to be detrimental to treatment efficiency.Uzal et al. (2003) investigated the biochemical methanepotential (BMP) of malt whisky distillery wastewater bothwith and without basal medium to observe the effect ofnutrient supplementation. When a COD concentration of20,920 mg L�1 was maintained in the influent to the firststage, the effluent quality of the first stage began to deteri-orate. A significant colour change was observed from blackto brownish black and then to brown, and this was thoughtto be due to reduced metabolic activity owing to the toxiceffect of the wastewater on granular biomass, thus increas-ing oxidation-reduction potential. It was concluded thattwo stage UASB reactor configuration is an efficient systemfor malt whisky wastewater treatment until up to33,866 mg L�1 influent COD concentration. For the over-all sequential system (anaerobic/aerobic) treatment CODand BOD removal efficiencies were 99.5% and 98.1%,respectively, for the treatment of malt whisky wastewater.In aerobic phase, the effluent of anaerobic bioreactor isexposed to atmospheric oxygen in a tank with homogeniz-ers for proper mixing of the effluents. BOD is reduced to200 and effluent diluted with wastewater from bottlingand washer sections and disposed of after clarification(Ramendra and Awasthi, 1992). The stabilized sludgeserves as a soil conditioner and plant nutrient.

Cost economics of biogas production by anaerobictreatment was calculated by Ciftci and Ozturk (1995). They

suggested that for every $100 spent for the operation offull-scale anaerobic – aerobic treatment plants in a fermen-tation industry in Turkey producing baker’s yeast fromsugarbeet molasses, the biogas recovery is worth $300.Garcia-Calderon et al. (1998) reported the application ofthe down-flow fluidization technology for the anaerobicdigestion of red wine distillery wastewater. The systemachieved 85% TOC removal, at an organic loading rateof 4.5 kg TOC m3 day�1. Perlite was found to be a goodcarrier for the anaerobic digestion as it allowed a highbiomass hold-up, with minimum particle wash out, becauseof its density.

Immobilization of bacteria in biofilm and on bioflocs isa crucial step in anaerobic degradation because of advanta-ges such as higher activities, higher COD removal percentat short hydraulic retention times and better tolerance todisturbances such as toxic and organic shock loadings.At the same time there are certain disadvantages as wellbecause in addition to some readily biodegradable matter,vinasses contain compounds like phenols, which are toxicto bacteria and inhibit the digestion. Also, due to seasonalnature of many of these industries and the absence ofmicroorganisms in vinasses capable of carrying out anaer-obic digestion, long incubation periods are required forthe start-up stage. Besides, other operational problems inanaerobic digestion such as low growth rate of anaerobicbacteria and the loss of biomass in systems with highhydraulic rates frequently does not achieve a satisfactorypurification of vinasses (Beltran et al., 1999). The for-mation of H2S in anaerobic reactors is the result of thereduction of oxidized sulphur compounds. Methanogenicbacteria can tolerate sulphide concentration up to1000 mg L�1 total sulphide. A complete loss of methaneproduction occurred at 200 mg L�1 of un-ionized H2S dur-ing digestion of flocculent sludge. Anaerobic contact pro-cess incorporating an ultrafiltration (UF) unit was usedto treat distillery wastewater characterized by high andlow carbon to nitrogen concentrations. This treatment sys-tem showed methane yield of up to 0.6 m3 kg�1 VS and

Table 1Anaerobic methods employed for distillery effluent treatment

Reactor type Organic loading rate (OLR)(kg COD m�3 day�1)

CODremoval(%)

BODremoval(%)

Retentiontime(days)

Reference

Downflow fixed-film reactor – 60–73 85–97 – Bories and Ranyal, 1988Granular bed anaerobic baffled

reactor (GRABR)2.37 90–96 80–92 4 Akunna and Clark, 2000

Hybrid anaerobic baffled reactor 20 70 – – Boopathy and Tilche, 1991Upflow anaerobic sludge blanket

(UASB) reactor28 39–67 80 – Harada et al., 1996

Istanbul UASB reactor 6–11 90 – – Akarsubasi et al., 2006Tekirdag UASB reactor 2.5–8.5 60–80 – –Diphasic fixed-film reactor with

granular activated carbon (GAC)as support media

21.3 67.1 – 4 Goyal et al., 1996

Anaerobic contact filter 19,000 mg L�1

(influent COD concentration)73–98 – 4 Vijayaraghavan and

Ramanujam, 2000

2324 D. Pant, A. Adholeya / Bioresource Technology 98 (2007) 2321–2334

Autho

r's

pers

onal

co

py

removed up to 80% of the volatile acids (Kitamura et al.,1996). Two-phase anaerobic digestion of alcohol stillageproved to be superior to the single-phase process in termsof substrate loading rate and methane yield, without affect-ing the treatment performance (Yeoh, 1997). While main-taining BOD and COD reduction of 85% and 65%respectively, the two-phase achieved methane yield threetimes that of single-phase system.

A number of environmental factors affect the activity ofwastewater microbial populations and the rate of biochem-ical reactions. Of particular importance are temperature,pH, nutrients, and inhibiting toxic compounds. Due toacidic nature of vinasses, pH is one of the most relevantfactors affecting the microbiological activity in the biolog-ical process. The pH of wastewater increases from 4.0 to7.5 after anaerobic digestion due to the oxidation oforganic acids to CO2 and the reaction between the CO2

and basic compounds to form carbonates and bicarbonates(Beltran et al., 1999). Because of its high organic load, thedistillery wastewater is often diluted with tap water to sim-ulate the concentration of a typical industrial effluent enter-ing a wastewater treatment plant. However, spent washeven after anaerobic treatment does not meet the stringenteffluent standards laid down by CPCB, India, in terms ofvery high levels of BOD, COD, solids etc. (Asthanaet al., 2001). Also, the secondary spent wash produced bythe anaerobically digested primary molasses spent wash(DMSW) effluent is darker in colour, needing huge vol-umes of water to dilute it and is currently used in irrigationwater causing gradual soil darkening. The effluent there-fore is released after diluting with fresh water which is avery dear commodity to industries. Besides, sometimes fail-ure of the anaerobic digestion threatens the criteria for dis-charge limit. To overcome this problem either a largeamount of water is used to dilute the wastewater prior toanaerobic digestion or chemical coagulants are added.These actions require expansion of the anaerobic digestervolume, large amounts of water for dilution, and addi-tional costs for coagulants (Kim et al., 1997). Hence aero-bic treatment is necessary for anaerobically treated finaleffluent.

4.2. Aerobic treatment

4.2.1. Bacterial treatment

Microbial treatments employing pure bacterial culturehave been reported frequently in past and recent years. Adetailed list of bacteria tried by different researchers fordecolourization of distillery effluent is given in Table 2.Kumar and Viswanathan (1991) isolated bacterial strainsfrom sewage and acclimatized on increasing concentrationsof distillery waste. These strains were able to reduce CODby 80% in 4–5 days without any aeration. The major prod-ucts left after treatment were biomass, carbon dioxide andvolatile acids. Petruccioli et al. (2000) used an air bubblecolumn reactor with activated sludge carrying self adaptedmicrobial population in both free and immobilized on

polyurethane particles for treating aerobic winery wastewa-ter. The highest COD removal rate was with free activatedsludge in the bubble column reactor. The most prominentbacterial species isolated from the reactor liquid belongedto Pseudomonas while Bacillus was isolated mostly fromcolonized carriers. Pseudomonas fluorescens, decolourizedmelanoidin wastewater (MWW) up to 76% under non-ster-ile conditions and up to 90% in sterile samples (Dahiyaet al., 2001a). The difference in decolourization might bedue to the fact that melanoidin stability varies with pHand temperature and at higher temperature during sterili-zation melanoidin-pigments decompose to low molecularweight compounds (Ohmomo et al., 1988b). The effect ofimmobilization on the decolourization of a melanoidinsolution may be explained by the fact that Lactobacillus hil-

gardii requires a small amount of oxygen for the decolou-rization and immobilization within Ca-alginate gel leadsto suitably limited aeration, supplying a small amount ofoxygen continuously (Ohmomo et al., 1988a).

Acetogenic bacteria are capable of oxidative decomposi-tion of melanoidins. Cibis et al. (2002) achieved biode-gradation of potato slops (distillation residue) by a mixedpopulation of bacteria under thermophilic conditions upto 60 �C. A COD removal of 77% was achieved undernon-optimal conditions. Marine cyanobacteria such asOscillatoria boryna have also been reported to degrade mel-anoidin due to production of H2O2, hydroxyl, perhydroxyland active oxygen radicals, resulting in the decolourizationof the effluent (Kalavathi et al., 2001). 96%, 81% and 26%decolorisation of distillery effluent through bioflocculationby Oscillatoria sp., Lyngbya sp. and Synechocystis sp.respectively was reported by Patel et al. (2001).

Distillery spent wash, despite carrying high organicload contains little readily available carbon. Isolationof bacterial strains capable of degrading recalcitrant com-pounds of anaerobically digested spent wash from soil ofeffluent discharge site was reported by Ghosh et al.(2004). These were Pseudomonas, Enterobacter, Steno-trophomonas, Aeromonas, Acinetobacter and Klebsiella allof which could carry out degradation of some componentof spent wash. Maximum 44% COD reduction wasachieved using these bacterial strains either singly or col-lectively. Sirianuntapiboon et al. (2004) used an acetogenicbacterium to obtain a decolourization yield of 76.4%under optimal nutrient conditions. However, this valuewas only 7.3%, by using anaerobic pond. Also, it requiredsugar, especially glucose and fructose for decolourizationof MWWs. The decolourization activity might be due toa sugar oxidase.

4.2.2. Fungal treatment

In recent years, several basidiomycetes and ascomycetestype fungi have been used in the decolourization of naturaland synthetic melanoidins in connection with colour reduc-tion of wastewaters from distilleries. The aim of fungal treat-ment is to purify the effluent by consumption of organicsubstances, thus, reducing its COD and BOD, and at the

D. Pant, A. Adholeya / Bioresource Technology 98 (2007) 2321–2334 2325

Autho

r's

pers

onal

co

py

same time to obtain some valuable product, such as fungalbiomass for protein-rich animal feed or some specific fungalmetabolite. Filamentous fungi have lower sensitivity to vari-ations in temperature, pH, nutrients and aeration and havelower nucleic acid content in the biomass (Knapp et al.,2001). Several fungi have been investigated for their abilityto decolourise melanoidins and MSW (Table 3).

Coriolus sp. no. 20, in class basidiomycetes was the firststrain for the application of its ability to remove melanoi-dins from MWW (Watanabe et al., 1982). This isolatedid not show any decolourization activity when molassespigment was used as carbon source but it showed the activ-ity when sorbose or glucose was added. In 1985, Ohmomoet al. used Coriolus versicolour Ps4a for MWW decolouri-zation and obtained 80% decolourization in darknessunder optimum conditions. Later, Ohmomo et al. (1988b)

used autoclaved mycelium of Aspergillus oryzae Y-2-32that adsorbed lower weight fractions of melanoidin anddegree of adsorption was influenced by the kind of sugarsused for cultivation. The wine distilleries produce large vol-umes of wastewaters having phenolic compounds, whichgive a high inhibitory and anti-bacterial activity to thiswastewater, thus slowing down the anaerobic digestionprocess. Partial elimination of these phenolics compoundswas obtained by using Geotrichum candidum (Borja et al.,1993). Rhizoctonia sp. D-90 decolourized molasses mela-noidin medium and a synthetic melanoidin medium by87.5% and 84.5% respectively, under experimental growthconditions. Electron microscopy revealed that the myceliaabsorbed melanoidin pigment, which was in the form ofelectron dense material in the cytoplasm. However, mela-noidin could be eluted from the mycelia by washing in a

Table 2Bacteria employed for the decolourization of distillery effluent

S. no. Name Comments Colour Removal (%) Reference

1 Xanthomonas fragariae All the three strains needed glucose as carbon source andNH4Cl as nitrogen source. The decolourization efficiency offree cells was better than immobilized cells

76 Jain et al., 2002

2 Bacillus megaterium 763 Bacillus cereus 824 Bacillus smithii Decolourization occurred at 55 �C in 20 days under anaerobic

conditions in presence of peptone or yeast extract assupplemental nutrient. Strain could not use MWW as solecarbon source

35.5 Kambe et al., 1999

5 Lactobacillus hilgardii Immobilized cells of the heterofermentative lactic acidbacterium decolourized 40% of the melanoidins solution within4 days aerobically

40 Ohmomo et al., 1988a

6 Acetobacter acetii The organism required sugar especially, glucose and fructosefor decolourization of MWWs

76.4 Sirianuntapiboon et al.,2004

7 Pseudomonas

fluorescens

This decolourization was obtained with cellulose carrier coatedwith collagen. Reuse of decolourized cells reduced thedecolourization efficiency

94 Dahiya et al., 2001a

8 Pseudomonas putida The organism needed glucose as a carbon source, to producehydrogen peroxide which reduced the colour

60 Ghosh et al., 2002

9 Acinetobacter sp. All these organisms were isolated from an air bubble columnreactor treating winery wastewater after 6 months ofoperation. Most isolates from the colonized carriers belongedto species of the genus Bacillus

Not checked in thisstudy

Petruccioli et al., 2000

10 Aeromonas sp.11 Alcaligens faecalis

12 Bacillus sp.13 Flavobacterium sp.14 F. meningosepticum

15 Pseudomonas sp.16 P. paucimobilis

17 P. vescicularis

18 Sphingobacterium

multivorum

19 Bacillus thuringiensis Addition of 1% glucose as a supplementary carbon source wasnecessary

22 Kumar and Chandra,2006

20 Bacillus brevis 27.421 Bacillus sp. 27.422 Pseudomonas

aeruginosa

The three strains were part of a consortium which decolourizedthe anaerobically digested spent wash in presence of basal saltsand glucose

67 Mohana et al., 2007

23 Stenotrophomonas

maltophila

24 Proteus mirabilis

2326 D. Pant, A. Adholeya / Bioresource Technology 98 (2007) 2321–2334

Autho

r's

pers

onal

co

py

Table 3Fungi employed for the decolourization of Distillery effluent

S.no.

Name Comments ColourRemoval (%)

Reference

1 Phanerochaete

chrysosporium

Both the fungi required a readily available carbon source formelanoidin decolourization while N source had no effect. Maximumdecolourization was observed in 6.25% (v/v) spent wash

53.5 Kumar et al., 1998

2 Coriolus versicolor 71.53 Trametes versicolor COD and N–NH4 removal observed in presence of sucrose and

KH2PO4 as nutrient source82 Benito et al., 1997

4 Geotrichum

candidum

Fungus immobilized on polyurethane foam showed stabledecolourization of molasses in repeated-batch cultivation

80 Kim and Shoda, 1999

5 Coriolus hirsutus A large amount of glucose was required for colour removal butaddition of peptone reduced the decolourizing ability of the fungus

80 Miyata et al., 2000

6 Penicillium sp. All fungi produced decolourization from first day of incubation,with maximum being shown byP. decumbens at fourth day with areduction of 70% of the phenolic content of the wastewater

30 Jimnez et al., 2003

7 Penicillium

decumbens

41

8 Penicillium

lignorum

28

9 Aspergillus niger 2510 Aspergillus niger

UM2Decolourization was more by immobilized fungus and it was able todecolourize up to 50% of initial effluent concentrations

80 Patil et al., 2003

11 Aspergillus

fumigatus G-2-6Thermophilic strain tried for molasses wastewater decolourizationbut colouring compounds hardly degraded

56 Ohmomo et al., 1987

12 Mycelia sterilia Organism required glucose for the decolourizing activity 93 Sirianuntapiboon et al., 198813 Aspergillus niger Maximum colour removal was obtained when MgSO4, KH2PO4,

NH4NO3 and a carbon source was added to wastewater69 Miranda et al., 1996

14 Flavodon flavus MSW was decolourized using a marine basidiomycete fungus. It alsoremoved 68% benzo(a)pyrene, a PAH found in MSW

80 Raghukumar and Rivonkar,2001; Raghukumar et al., 2004

15 Rhizoctonia sp. D-90

Mechanism of decolourization of melanoidin involved absorption ofthe melanoidin pigment by the cells as a macromolecule and itsintracellular accumulation in the cytoplasm and around the cellmembrane as a melanoidin complex, which was then graduallydecolourized by intracellular enzymes

90 Sirianuntapiboon et al., 1995

16 Coriolus versicolor

Ps4aTwo types of enzymes, sugar-dependent and sugar-independent,were found to be responsible for melanoidin decolourizing activity

80 Ohmomo et al., 1985

17 Aspergillus oryzae

Y-2-32The thermophilic strain adsorbed lower molecular weight fractionsof melanoidin and required sugars for growth

75 Ohmomo et al., 1988b

18 Phanerochaete

chrysosporium

JAG-40

This organism decolourized synthetic and natural melanoidins whenthe medium was supplemented with glucose and peptone

80 Dahiya et al., 2001b

19 Coriolus hirsutus

IFO4917Melanoidins present in heat treatment liquor were subjected tosequencing batch decolourization by the immobilized fungal cells

45 Fujita et al., 2000

20 Aspergillus niveus The fungus could use sugarcane bagasse as carbon source andrequired other nutrients for decolourization

56 Angayarkanni et al., 2003

21 Trametes sp. I-62 No colour observed associated with either fungal mycelium orpolysaccharides secreted by the fungus and therefore colour removalwas attributed to fungal degradation and not to a simple physicalbinding

73 Gonzalez et al., 2000

22 Aspergillus niger All these organisms were isolated from an air bubble column reactortreating winery wastewater after 6 months of operation

Not checkedin this study

Petruccioli et al., 2000

23 Candia sp.24 C. lambica

25 C. lypolitica

26 Fusarium sp.27 Penicillium sp.28 P. roquefortii

29 Saccharomyces

cerevisiae

30 Trichoderma

koningii

31 Coriolus sp. no. 20 First strain for the application of its ability to remove melanoidinsfrom MWW, showed decolourization activity in 0.5% melanoidinwhen sorbose or glucose was added as carbon source

80 Watanabe et al., 1982

(continued on next page)

D. Pant, A. Adholeya / Bioresource Technology 98 (2007) 2321–2334 2327

Autho

r's

pers

onal

co

py

solution of NaOH and the relative amount of melanoidineluted from the mycelia increased with increase in the con-centration of NaOH (Sirianuntapiboon et al., 1995). Asper-

gillus awamori var. kawachi has been used for productionof single cell protein from Japanese distillery (Shochu)wastewater after aerobic cultivation (Kida et al., 1995).The supernatant after cultivation could be anaerobicallytreated, at a high TOC loading rate, by the addition ofNi2+ and Co2+. Also, NHþ4 , accumulated in the anaerobi-cally treated wastewater, was efficiently removed by utiliza-tion of residual volatile fatty acids (VFA) as electrondonors during biological denitrification and nitrification,and the residual organic matter could be removed simulta-neously. Colour elimination from MSW using Aspergillus

niger was studied by Miranda et al. (1996). Under optimalnutrient concentration 83% of the total colour removedwas eliminated biologically and 17% by adsorption onthe mycelium. Ohmomo et al. (1988a) concluded that

microbial decolourization of melanoidins is due to twodecomposition mechanisms; in the first the smaller molecu-lar weight melanoidins are attacked and in the second thelarger molecular weight melanoidins are attacked.

Under nutrient limiting conditions, fungal cells gener-ally cannot remain active during a long-term cultivation.Therefore, the continuous-culture method is not practicaland the semi-batch or repeated-batch method can be analternative for long-term cultivation. The immobilizationof the fungus on a solid support is an appropriate meansfor controlling the thickness of the biofilm. The immobili-zation of the fungus offers advantages such as short reten-tion time, easy recovery of the cells and increased activity.Furthermore, in the presence of the foam matrix, pelletsize is restricted by the size and the physical propertiesof the foam (Kim and Shoda, 1999). Miyata et al. (2000)suggested an inhibitory effect of organic nitrogen on mel-anoidin decolourization by fungus Coriolus hirsutus. At

Table 3 (continued)

S.no.

Name Comments ColourRemoval (%)

Reference

32 Williopsis saturnus

strain CBS 5761Yeast isolates from a rotating biological contactor (RBC) treatingwinery wastewater. Only 43% COD removal could be achieved

Not checkedin this study

Malandra et al., 2003

33 Pichia

membranaefaciens

strain IGC 500334 Candia intermedia

JCM 160735 Eremothecium

gossyphi

36 Saccharomyces

cerevisiae strain J237 Hanseniaspora

uvarum

38 Coriolus versicolor

sp no. 2010% diluted spent wash was used with glucose @ 2% added as carbonsource

34.5 Chopra et al., 2004

39 Phanerochaete

chrysosporium

Sugar refinery effluent was treated in a RBC using polyurethane foamand scouring web as support

55 Guimaraes et al. (2005)

40 Pycnoporus

coccineus

Immobilized mycelia removed 50% more colour than free mycelia 60 Chairattanamanokorn et al.,2005

41 Coriolus versicolor Cotton stalks were added as additional carbon source which stimulatedthe decolourization activity of all fungi in 30% vinasses

63 Kahraman and Yesilada,2003

42 Phanerochaete

chrysosporium

37

43 Funalia trogii 5744 Pleurotus

pulmonarius

43

45 Aspergillus-UB2 This was with diluted wastewater with optimum values ofsupplemented materials

75 Shayegan et al. (2004)

46 MarineBasidiomyceteNIOCC # 2a

Experiment was carried out at 10% diluted spent wash 100 D’souza et al. (2006)

47 Phanerochaete

chrysosporium

Molasses medium decolourization was checked in stationary andsubmerged cultivation conditions

Thakkar et al., 2006

NCIM 1073 0NCIM 1106 82NCIM 1197 76

48 Citeromyces sp.

WR-43-6Organism required glucose, Sodium nitrate and KH2PO4 for maximaldecolourization

68.91 Sirianuntapiboon et al., 2003

49 Hansenula fabianii The flocculant strains could reduce 28.5% TOC from wastewaterwithout dilution

Not checkedin this study

Moriya et al., 1990

50 Hansenula anomala

2328 D. Pant, A. Adholeya / Bioresource Technology 98 (2007) 2321–2334

Autho

r's

pers

onal

co

py

the same time glucose was also required for enhancingdecolourization as the peroxidases require H2O2, whichis generated by glucose oxidation, to decolourize melanoi-din. Chopra et al. (2004) reported that absence of addi-tional nitrogen could not inhibit activity of fungus C.

versicolour sp no. 20 considerably, as significant decolouri-zation and COD reduction occurred even in the absence ofit.

Colour removal from distillery effluent using a marinefungus, Flavodon flavus has been reported by Raghukumarand Rivonkar (2001). This fungus was more effective indecolourizing raw MSW than was the molasses wastewatercollected either after anaerobic treatment or after aerobictreatment. The oxygen demand of the fungus was quitehigh. P. chrysosporium JAG-40 decolourized syntheticand natural melanoidins present in spentwash up to 80%(Dahiya et al., 2001b). The larger molecular weight frac-tions of melanoidin were decolourized rapidly, while thesmall molecular weight fractions remained in solutionand were metabolized slowly. Also, the decolourizationwas less in sterilized spentwash than in non-sterile solution.This observation is completely opposite of the one whenPseudomonas fluorescens was used by same authors(Dahiya et al., 2001a). Kahraman and Yesilada (2003)reported molasses decolourization in semi solid state(SSS) cultivation by fungi C. versicolour, Funalia trogii,P. chrysosporium and Pleurotus pulmonarius with cottonstalks being used as additional source of carbon. C. versi-

colour decolourized 48% of 30% diluted vinasse withoutany additional carbon source which increased to 71% onaddition of cotton stalks. Aspergillus niveus, a litter degrad-ing fungi was used by Angayarkanni et al. (2003) for thetreatment of distillery effluent using paddy straw, sugar-cane bagasse, molasses and sucrose as carbon source forgrowth of fungus in the effluent. Sugarcane bagasse at1% (w/v) concentration resulted in maximum removal ofcolour (37%) and COD (91.68%). The decrease in colourremoval in this study might be due to the fact that the efflu-ent taken for study was alkaline (pH 9.0) and the melanoi-dins responsible for colour were more soluble in thealkaline pH. In the acidic pH, the melanoidins might beprecipitated and removed easily. Shayegan et al. (2005)used an Aspergillus species isolated from the soil fordecolourization of anaerobically digested (UASB) andaerobically treated distillery wastewater. With dilutedwastewater at optimum values of supplemented materials75% decolourization was achieved which reduced to 40%on using undiluted wastewater. It was suggested that deco-lourization by fungi takes place due to the destruction ofcoloured molecules and partially because of sorption phe-nomena. A longer aeration period causes the adsorbed col-our molecules to be released as a result of endogenousrespiration and cell death, hence reducing decolourizationefficiency.

Yeast Citeromyces was used for treating MWW andhigh and stable removal efficiencies in both colour intensityand organic matter were obtained. However, the semi-pilot

and pilot-scale experiments are to be tested for checkingthe stability of Citeromyces sp. (Sirianuntapiboon et al.,2003). Malandra et al. (2003) studied the microorganismsassociated with a rotating biological contactor (RBC)treating winery wastewater. One of the yeast isolates wasable to reduce the COD of synthetic wastewater by 95%and 46% within 24 h under aerated and non-aerated condi-tions, respectively. Moriya et al. (1990) used two flocculantstrains of yeast, Hansenula fabianii and Hansenula anomala

for treatment of wastewater from beet molasses-spirits pro-duction and achieved 25.9% and 28.5% removal of TOCrespectively from wastewater without dilution. Dilutionof wastewater was not favourable for practical treatmentof wastewater due to the longer treatment time and higherenergy cost.

4.2.3. Mixed consortium treatment

During last two decades, several attempts have beenmade to investigate the possibility of using cell immobili-zation in the technology of aerobic wastewater treatment(Fedrici, 1993; Sumino et al., 1985). Early experimentswere restricted to the use of selected pure cultures im-mobilized on solid supports for the degradation of specifictoxic compounds (Anselmo et al., 1985; Livernoche et al.,1983). Later, immobilized consortia of two or moreselected strains were employed (Kowalska et al., 1998;Zache and Rehm, 1989) but of late activated sludge hasbeen immobilized on different carriers and used for waste-water treatment (Shah et al., 1998). Jet loop reactors(JLR), the efficiency of which has already been shown inboth chemical and biological processes have also beenevaluated for aerobic treatment of winery wastewater. AJLR of 15 dm3 working volume was used for the aerobictreatment of winery wastewater (Petruccioli et al., 2002).COD removal efficiency higher than 90% was achievedwith an organic load of the final effluents that rangedbetween 0.11 and 0.3 kg COD m�3. Most isolates belongto the genus Pseudomonas and the yeast Saccharomyces

cerevisiae. Later, Eusibio et al. (2004) reported the opera-tion of a JLR for more than one year treating winerywastewater collected in different seasons and achieved anaverage COD removal efficiency of 80%. JLR have higheroxygen transfer rates at lower energy costs. They alsoobserved Bacillus apart from Pseudomonas and the yeastSaccharomyces cerevisiae. Adikane et al. (2006) studieddecolourization of molasses spent wash in absence ofany additional carbon or nitrogen source using soil asinoculum. A decolourization of 69% was obtained using10% (w/v) soil and 12.5% (v/v) MSW after 7 daysincubation.

4.2.4. Phytoremediation approach

Algal growth potential bioassay is a standard assay todetermine the potential of water bodies, natural watersand wastewaters, to support or inhibit the microalgaegrowth. Algae growth potential was determined in distill-ery wastewater pretreated by anaerobic processes and by

D. Pant, A. Adholeya / Bioresource Technology 98 (2007) 2321–2334 2329

Autho

r's

pers

onal

co

py

a combined anaerobic-aerobic system. The biologicallytreated distillery wastewater provided satisfactory condi-tions for microalgae growth (Travieso et al., 1999). Billoreet al. (2001) used Phragmites kharka in a constructed wet-land for treatment of wastewater from the same industryand obtained 36% removal of total Kjeldahl nitrogen and48% removal of total suspended solids (TSS). Enhanceddecolourization was achieved by phytoremediation ofdistillery effluent by a macrophyte, Spirodela polyrrhiza

(L.) Schliden pretreated with Bacillus thuringiensis (Kumarand Chandra, 2004). Recently, macrophyte Potamogeton

pectinatus was used for bioaccumulating heavy metals fromdistillery effluent (Singh et al., 2005). Increasing concentra-tion of the effluent greatly reduced the biomass of the plantwith maximum accumulation of Fe being recorded inplants growing in 100% effluent. In another study Trivedyand Nakate (2000) employed Typha latipholia for distilleryeffluent treatment in a constructed wetland. The systemresulted in 78% and 47% reduction in COD and BODrespectively in a period of 10 days. Using a combined treat-ment with Lemna minuscula and Chlorella vulgaris Valder-rama et al. (2002) reported 52% colour removal fromdistillery effluent. The microalgal treatment removed nutri-ents and organic matter from wastewater and producedoxygen for other organisms. The macrophyte removedorganic matter and eliminated the microalgae form treatedwastewater. However, despite the potential of aquatic mac-rophytes in cleaning wastewaters the use of these plants indesigning a low cost treatment system is still at experimen-tal stage and is considered to be a potentially importantarea of environmental management.

5. Role of enzymes in effluent decolourization

Although the enzymatic system related with decolouriza-tion of melanoidins is yet to be completely understood, itseems greatly connected with fungal ligninolytic mech-anisms. The white-rot fungi have a complex enzymaticsystem which is extracellular and non-specific, and undernutrient-limiting conditions is capable of degrading ligno-lytic compounds, melanoidins, and polyaromatic com-pounds that cannot be degraded by other microorganisms(Benito et al., 1997). A large number of enzymes from avariety of different plants and microorganisms have beenreported to play an important role in an array of wastetreatment applications.

Several studies regarding degradation of melanoidins,humic acids and related compounds using basidiomyceteshave also suggested a participation of at least one laccaseenzyme in fungi belonging to Trametes (Coriolus) genus.The role of enzymes other than laccase or peroxidases inthe decolourization of melanoidins by Trametes (Coriolus)strain was reported during the 1980s. Several reportsclaimed that intracellular sugar-oxidase- type enzymes (sor-bose-oxidase or glucose-oxidase) had melanoidin-decolou-rizing activities. It was suggested that melanoidins weredecolourized by the active oxygen (O2; H2O2) produced

by the reaction with sugar oxidases (Watanabe et al.,1982). Decolourization by microbial methods includes theenzymatic breakdown of melanoidin and flocculation bymicrobially secreted substances. Ohmomo et al. (1985) usedC. versicolour Ps4a, which decolourized molasses wastewa-ter 80% in darkness under optimum conditions. Decolouri-zation activity involved two types of intracellular enzymes,sugar-dependent and sugar-independent. One of theseenzymes required no sugar and oxygen for appearance ofthe activity and could decolourize MWW up to 20% indarkness and 11–17% of synthetic melanoidins. Thus, theparticipation of these H2O2 producing enzymes as a partof the complex enzymatic system for melanoidin degrada-tion by fungi should be taken into account while designingany treatment strategy. One of the more complete enzy-matic studies regarding melanoidin decolourization wasreported by Miyata et al. (1998). Colour removal of syn-thetic melanoidin by C. hirsutus involved the participationof peroxidases (MnP and MIP) and the extracellular H2O2

produced by glucose-oxidase, without disregard of a partialparticipation of fungal laccase. Mansur et al. (1997)obtained a maximum decolourization of around 60% onday 8 after inoculating with fungus Trametes sp. I-62. Hereeffluent was added at a final concentration of 20% (v/v)after 5 days of fungal growth, the time at which high levelsof laccase activity were detected in the extracellularmycelium.

The white-rot basidiomycete T. versicolour is an activedegrader of humic acids as well as of melanoidins. A mel-anoidin mineralizing 47 kDa extracellular protein corre-sponding to the major mineralizing enzyme system fromT. versicolour was isolated by Dehorter and Blondeau(1993). This Mn2+ dependent enzyme system required oxy-gen and was described to be as peroxidase. Uniform, smalland spongy pellets of the fungus T. versicolour were used asinoculum for colour removal using different nutrients suchas ammonium nitrate, manganese phosphate, magnesiumsulphate and potassium phosphate and also sucrose as car-bon source (Benito et al., 1997). Maximum colour removalof 82% and 36% removal of N–NHþ4 was obtained on usinglow sucrose concentration and KH2PO4 as the only nutri-ent. Some studies have identified the lignin degradationrelated enzymes participating in the melanoidin decolouri-zation. Intracellular H2O2 producing sugar oxidases havebeen isolated from Coriolus strains. Also, C. hirsustus havebeen reported to produce enzymes that catalyze melanoidindecolourization directly without additions of sugar and O2.Miyata et al. (1998) used C. hirsutus pellets to decolourize amelanoidin-containing medium. It was elucidated thatextracellular H2O2 and two extracellular peroxidases, amanganese-independent peroxidase (MIP) and manganeseperoxidase (MnP) were involved in decolourizationactivity.

Lee et al. (2000) investigated the dye-decolourizing per-oxidase by cultivating Geotrichum candidum Dec1 usingmolasses as a carbon source. Components in the molas-ses medium stimulated the production of decolourizing

2330 D. Pant, A. Adholeya / Bioresource Technology 98 (2007) 2321–2334

Autho

r's

pers

onal

co

py

peroxidase but inhibited the decolourizing activity of thepurified enzyme. It was found that the inhibitory effect ofmolasses can be eliminated at dilution ratios of more than25. Recently D’souza et al. (2006) reported 100% decolou-rization of 10% spent wash by a marine fungal isolatewhose laccase production increased several folds in thepresence of phenolic and non-phenolic inducers.

A combined treatment technique consisting of enzymecatalyzed in situ transformation of pollutants followed byaerobic biological oxidation was investigated by Sangaveand Pandit (2006a) for the treatment of alcohol distilleryspent wash. It was suggested that enzymatic pretreatmentof the distillery effluent leads to in situ formation of thehydrolysis products, which have different physical proper-ties and are easier to assimilate than the parent pollutantmolecules by the microorganisms, leading to faster initialrates of aerobic oxidation even at lower biomass levels.In another study, Sangave and Pandit (2006b) used irradi-ation and ultrasound combined with the use of an enzymeas pretreatment technique for treatment of distillery waste-water. The combination of the ultrasound and enzymeyielded the best COD removal efficiencies as compared tothe processes when they were used as stand-alone treatmenttechniques. Enzymatic decolourization of molasses med-ium has also been tried using P. chrysosporium (Thakkaret al., 2006). Under stationary cultivation conditions, noneof the strains could decolourize molasses nor produceenzymes lignin peroxidase, manganese peroxidase and lac-case. All of them could produce lignin peroxidase and man-ganese peroxidase when cultivated in flat bottom glassbottles under stationary cultivation conditions.

6. Conclusions

For industries using large quantities of water such asdistilleries, it is essential to treat and reuse their wastewater.However, most of the times, the discharge standardsapplied to most agro-industries, including distilleries areoften too stringent and below the levels that can beachieved with appropriate biological treatment technolo-gies. It has been observed and often reported that the useof an individual process alone may not treat the wastewatercompletely. A combination of these processes is necessaryto achieve the desirable goal. An anaerobic, or chemicalcoagulation/oxidation pretreatment followed by aerobicbiological oxidation is a common technique used for decol-orizing wastewater. But as discussed above, these processesare not efficient enough to treat these large volumes of col-oured wastewaters. A combination of different treatmentprocesses including a decolourization step could result inan effective bioremediation of the molasses wastewaters.

In general, microbial decolourization is an environ-ment-friendly and cost-competitive alternative to chemicaldecomposition process. However, the problem still persistsbecause several organisms that have been shown to degrademelanoidin are not best suited for treating MSW. This is

because they deplete oxygen in the effluent and further,higher fungi are not easily adopted for aquatic habitats.The investigations so far can be seen as an initial steptoward solving the problem. Moreover, most of thesemicrobial decolourization studies required effluent dilutionfor optimal activity. While using microorganisms, use ofmedia supplement pose extra burden on overall effluenttreatment process.

The use of cellulose carriers for microbial treatmentoffers an alternative method for cell immobilization.Gel entrapment has been conventional process of cellimmobilization. However, in processes like wastewatertreatment where large volumes are involved, entrapmentin gel beads is not as practical and economic when usedon an industrial scale. Further, the emerging treatmentmethods like enzymatic treatment have technologicaladvantages and yet are in its infancy, requiring economicalconsiderations in order to apply it on the plant scale. Cap-ital and operating costs of the available physicochemicaland biological treatment processes of distillery wastestream are inevitably high thus making these processes lesslucrative to the industry. Nevertheless, the feasibility ofapplication of the process to full-scale would need furtherresearch in this continuous culture set-up, in order to min-imize the added nutrients and extend the biomass activityfor a longer period. An understanding of complete profileof the effluent and the structures of coloring compoundswould also be helpful in achieving the appropriate treat-ment solutions.

Acknowledgements

Authors wish to thank Dr. R. K. Pachauri, Director-General, TERI, New Delhi, India for support in research.Financial assistance from University Grants Commission,New Delhi in form of Senior Research Fellowship to thefirst author is duly acknowledged.

References

Adikane, H.V., Dange, M.N., Selvakumari, K., 2006. Optimization ofanaerobically digested distillery molasses spent wash decolourizationusing soil as inoculum in the absence of additional carbon and nitrogensource. Biores. Technol. 97, 2131–2135.

Agrawal, C.S., Pandey, G.S., 1994. Soil pollution by spentwash discharge:Depletion of manganese (II) and impairment of its oxidation. J.Environ. Biol. 15, 49–53.

Akarsubasi, A.T., Ince, O., Oz, N.A., Kirdar, B., Ince, B.K., 2006.Evaluation of performance, acetoclastic methanogenic activity andarchaeal composition of full-scale UASB reactors treating alcoholdistillery wastewaters. Process Biochem. 41, 28–35.

Akunna, J.C., Clark, M., 2000. Performance of a granular-bed anaerobicbaffled reactor (GRABBR) treating whisky distillery wastewater.Biores. Technol. 74, 257–261.

Angayarkanni, J., Palaniswamy, M., Swaminathan, K., 2003. Biotreat-ment of distillery effluent using Aspergillus niveus. Bull. Environ.Contam. Toxicol. 70, 268–277.

Anselmo, A.M., Mateus, M., Cabral, J.M.S., Novais, J.M., 1985.Degradation of phenol by immobilized cells of Fusarium flocciferum.Biotechnol. Lett. 7, 889–894.

D. Pant, A. Adholeya / Bioresource Technology 98 (2007) 2321–2334 2331

Autho

r's

pers

onal

co

py

Asthana, A., Mishra, S.K., Chandra, R., Guru, R., 2001. Treatment ofcolour and biochemical oxygen demand of anaerobically treateddistillery effluent by aerobic bacterial strains. Ind. J. Environ. Prot.21, 1070–1072.

Beltran, F.J., Garcia-Araya, J.F., Alvarez, P.M., 1999. Wine distillerywastewater degradation 2. Improvement of aerobic biodegradation bymeans of an integrated chemical (ozone)-biological treatment. J. Agric.Food Chem. 47, 3919–3924.

Beltran, F.J., Alvarez, P.M., Rodriguez, E.M., Garcia-Araya, J.F., Rivas,J., 2001. Treatment of high strength distillery wastewater (cherrystillage) by integrated aerobic biological oxidation and ozonation.Biotechnol. Prog. 17, 462–467.

Benito, G.G., Miranda, M.P., Santos, D.R., 1997. Decolourization ofwastewater from an alcoholic fermentation process with Trametes

versicolor. Biores. Technol. 61, 33–37.Billore, S.K., Singh, N., Ram, H.K., Sharma, J.K., Singh, V.P., Nelson,

R.M., Dass, P., 2001. Treatment of a molasses based distillery effluentin a constructed wetland in central India. Water Sci. Tech. 44, 441–448.

Blonskaja, V., Menert, A., Vilu, R., 2003. Use of two-stage anaerobictreatment for distillery waste. Adv. Environ. Res. 7, 671–678.

Boopathy, R., Tilche, A., 1991. Anaerobic digestion of high strengthmolasses wastewater using hybrid anaerobic baffled reactor. WaterRes. 25, 785–790.

Bories, A., Ranyal, J., 1988. Anaerobic digestion of high-strength distillerywastewater (cane molasses stillage) in a fixed-film reactor. Biol.Wastes. 23, 251–267.

Borja, R., Martin, A., Luque, M., Duran, M.M., 1993. Enhancement ofthe anaerobic digestion of wine distillery wastewater by the removal ofphenolics inhibitors. Biores. Technol. 45, 99–104.

Borrero, M.A.V., Pereira, J.T.V., Miranda, E.E., 2003. An environmentalmanagement method for sugar cane alcohol production in Brazil.Biomass Bioenergy. 25, 287–299.

Cammerer, B., Jalyschkov, V., Kroh, L.W., 2002. Carbohydrate structuresas part of the melanoidins skeleton. Int. Congress Series 1245, 269–273.

Chairattanamanokorn, P., Imai, T., Kondo, R., Sekine, M., Higuchi, T.,Ukita, M., 2005. Decolourization of alcohol distillery wastewater bythermotolerant white rot fungi. Appl. Biochem. Microbiol. 41, 583–588.

Chopra, P., Singh, D., Verma, V., Puniya, A.K., 2004. Bioremediation ofmelanoidins containing digested spent wash from cane-molassesdistillery with white rot fungus, Coriolus versicolour. Ind. J. Microbiol.44, 197–200.

Cibis, E., Kent, C.A., Krzywonos, M., Garncarek, Z., Garncarek, B.,Miskiewicz, T., 2002. Biodegradation of potato slops from a ruraldistillery by thermophilic aerobic bacteria. Biores. Technol. 85, 57–61.

Ciftci, T., Ozturk, I., 1995. Nine years of full-scale anaerobic-aerobictreatment experiences with fermentation industry effluents. Water Sci.Technol. 32, 131–139.

CPCB, 2003. Charter on corporate responsibility for environmentalprotection. Workshop organized at Mumbai by MPCB on 03.01.2003.<http://cpcb.nic.in//Charter/charter5.htm>.

Dahiya, J., Singh, D., Nigam, P., 2001a. Decolourisation of molasseswastewater by cells of Pseudomonas fluorescens immobilized on porouscellulose carrier. Biores. Technol. 78, 111–114.

Dahiya, J., Singh, D., Nigam, P., 2001b. Decolourisation of synthetic andspentwash melanoidins using the white-rot fungus Phanerochaete

chrysosporium JAG-40. Biores. Technol. 78, 95–98.Dehorter, B., Blondeau, R., 1993. Isolation of an extracellular Mn-

dependent enzyme mineralizing melanoidins from the white rot fungusTrametes versicolour. FEMS Microbiol. Lett. 109, 117–122.

Driessen, W., Yspeert, P., 1999. Anaerobic treatment of low, medium andhigh strength effluent in agro-industry. Water Sci. Technol. 40, 221–228.

D’souza, D.T., Tiwari, R., Sah, A.K., Raghukumar, C., 2006. Enhancedproduction of Laccase by a marine fungus during treatment of

coloured effluents and synthetic dyes. Enzyme Microb. Technol. 38,504–511.

Duarte, E., Martins, M., Carvalho, E., Costa, S., Spranger, I., 1997. Anintegrated approach for overcoming the environmental impacts ofwineries wastewaters a Portuguese case study. In: Proceedings ofInternational Symposium of the Vine and wine, 07–10 October,1997. Yangling, China, pp. 1–5.

Eusibio, A., Petruccioli, M., Lageiro, M., Fedrici, F., Duarte, J.C., 2004.Microbial characterization of activated sludge in jet-loop bioreactorstreating winery wastewaters. J. Ind. Microbiol. Biotechnol. 31, 29–34.

Fedrici, F., 1993. Potential applications of viable, immobilized fungal cellsystems. World J. Microbiol. Biotechnol. 9, 495–502.

FitzGibbon, F., Singh, D., McMullan, G., Marchant, R., 1998. The effectof phenolics acids and molasses spent wash concentration on distillerywastewater remediation by fungi. Process Biochem. 33, 799–803.

Friedrich, J., 2004. Bioconversion of distillery waste. In: Arora, D.K.(Ed.), Fungal biotechnology in agriculture, food and environmentalapplications. Marcel Dekker Inc., New York, pp. 431–442.

Fujita, M., Era, A., Ike, M., Soda, S., Miyata, N., Hirao, T., 2000.Decolourization of heat-treatment liquor of waste sludge usingpolyurethane foam-immobilized white rot fungus equipped with anultramembrane filtration unit. J. Biosci. Bioeng. 90, 387–394.

Fumi, M.D., Parodi, G., Parodi, E., Silva, A., 1995. Optimization oflong-term activated-sludge treatment of winery wastewater. Biores.Technol. 52, 45–51.

Garcia-Calderon, D., Buffiere, P., Moletta, R., Elmaleh, S., 1998.Anaerobic digestion of wine distillery wastewater in down-flowfludized bed. Water Res. 32, 3593–3600.

Ghosh, M., Ganguli, A., Tripathi, A.K., 2002. Treatment of anaerobicallydigested distillery spentwash in a two-stage bioreactor using Pseudo-

monas putida and Aeromonas sp. Process Biochem. 7, 857–862.Ghosh, M., Verma, S.C., Mengoni, A., Tripathi, A.K., 2004. Enrichment

and identification of bacteria capable of reducing chemical oxygendemand of anaerobically treated molasses spent wash. J. Appl.Microbiol. 96, 1278–1286.

Gonzalez, T., Terron, M.C., Yague, S., Zapico, E., Galletti, G.C.,Gonzalez, A.E., 2000. Pyrolysis/gas chromatography/ mass spectro-metry monitoring of fungal biotreated distillery wastewater usingTrametes sp. I 62 (CECT 20197). Rapid Commun. Mass Spectrom. 14,1417–1424.

Goodwin, J.A.S., Stuart, J.B., 1994. Anaerobic digestion of malt whiskydistillery pot ale using upflow anaerobic sludge blanket reactor. Biores.Technol. 49, 75–81.

Goyal, S.K., Seth, R., Handa, B.K., 1996. Diphasic fixed-film biometh-anation of distillery spentwash. Biores. Technol. 56, 239–244.

Guimaraes, C., Porto, P., Oliveira, R., Mota, M., 2005. Continuousdecolourization of a sugar refinery wastewater in a modified rotatingbiological contactor with Phanerochaete chrysosporium immobilized onpolyurethane foam discs. Process Biochem. 40, 535–540.

Harada, H., Uemura, S., Chen, A.C., Jayadevan, J., 1996. Anaerobictreatment of a recalcitrant distillery wastewater by a thermophilicUASB reactor. Biores. Technol. 55, 215–221.

Hayase, F., Kim, S., Kato, H., 1986. Analyses of the chemical structuresof melanoidins by 13C and 15N CP-MAS NMR spectrophotometry.Agric. Biol. Chem. 50, 1951–1957.

Jain, N., Minocha, A.K., Verma, C.L., 2002. Degradation of predigesteddistillery effluent by isolated bacterial strains. Ind. J. Exp. Bot. 40, 101–105.

Jimnez, A.M., Borja, R., Martin, A., 2003. Aerobic-anaerobic biodegra-dation o beet molasses alcoholic fermentation wastewater. ProcessBiochem. 38, 1275–1284.

Joshi, H.C., 1999. Bio-energy potential of distillery effluents. Bio. EnergyNews. 3, 10–15.

Kahraman, S., Yesilada, O., 2003. Decolourization and bioremediation ofmolasses wastewater by white-rot fungi in a semi-solid state condition.Folia Microbiol. 48, 525–528.

Kalavathi, D.F., Uma, L., Subramanian, G., 2001. Degradation andmetabolization of the pigment-melanoidin in distillery effluent by the

2332 D. Pant, A. Adholeya / Bioresource Technology 98 (2007) 2321–2334

Autho

r's

pers

onal

co

py

marine cyanobacterium Oscillatoria boryana BDU 92181. EnzymeMicrob. Technol. 29, 246–251.

Kambe, T.N., Shimomura, M., Nomura, N., Chanpornpong, T., Naka-hara, T., 1999. Decolourization of molasses wastewater by Bacillus sp.under thermophilic and anaerobic conditions. J. Biosci. Bioeng. 87,119–121.

Kannabiran, B., Pragasam, A., 1993. Effect of distillery effluent on seedgermination, seedling growth and pigment content of Vigna mungo (L.)Hepper (CVT9). Geobios. 20, 108–112.

Kato, H., Hayase, F., 2002. An approach to estimate the chemicalstructure of melanoidins. Int. Congress Series 1245, 3–7.

Kaushik, A., Nisha, R., Jagjeeta, K., Kaushik, C.P., 2005. Impact of longand short term irrigation of a sodic soil with distillery effluent incombination with bioamendments. Biores. Technol. 96, 1860–1866.

Kida, K., Morimiura, S., Abe, N., Sonoda, Y., 1995. Biological treatmentof Shochu distillery wastewater. Process Biochem. 30, 125–132.

Kim, S.J., Shoda, M., 1999. Batch decolourization of molasses bysuspended and immobilizes fungus of Geotrichum candidum Dec 1.J. Biosci. Bioeng. 88, 586–589.

Kim, J.S., Kim, B.G., Lee, C.H., Kim, S.W., Jee, H.S., Koh, J.H., Fane,A.G., 1997. Development of clean technology in alcohol fermentationindustry. J. Cleaner Prod. 5, 263–267.

Kitamura, Y., Maekawa, T., Tagawa, A., Hayashi, H., Farrell-Poe, K.L.,1996. Treatment of strong organic, nitrogenous wastewater by ananaerobic contact process incorporating ultrafiltration. Appl. Eng.Agric. ASAE 12, 709–714.

Kitts, D.D., Wu, C.H., Stich, H.F., Powrie, W.D., 1993. Effects ofglucose-glycine maillard reaction products on bacterial and mamma-lian cells mutagenesis. J. Agric. Food Chem. 41, 2353–2358.

Knapp, J.S., Vantoch-Wood, E.J., Zhang, F., 2001. Use of wood-rottingfungi for the decolourisation of dyes and industrial effluents. In: Gadd,G.M. (Ed.), Fungi in Bioremediation. British Mycological Society.Cambridge University Press, p. 242.

Kort, M.J., 1979. Colour in the sugar industry. In: de Birch, G.G., Parker,K.J. (Eds.), Science and Technology. Applied Science, London, pp.97–130.

Kowalska, M., Bodzek, M., Bohdziewicz, J., 1998. Biodegradation ofphenols and cyanides with immobilized microorganisms. ProcessBiochem. 33, 189–197.

Kroh, L.W., 1994. Caramelisation in food and beverages. Food Chem.51, 373–379.

Kumar, P., Chandra, R., 2004. Detoxification of distillery effluent throughBacillus thuringiensis (MTCC 4714) enhanced phytoremediationpotential of Spirodela polyrrhiza (L.) Schliden. Bull. Environ. Contam.Toxicol. 73, 903–910.

Kumar, P., Chandra, R., 2006. Decolurisation and detoxification ofsynthetic molasses melanoidins by individual and mixed cultures ofBacillus spp. Biores. Technol. 97, 2096–2102.

Kumar, S., Viswanathan, L., 1991. Production of biomass, carbondioxide, volatile acids, and their interrelationship with decrease inchemical oxygen demand, during distillery waste treatment by bacterialstrains. Enzyme Microb. Technol. 13, 179–186.

Kumar, V., Wati, L., Nigam, P., Banat, I.M., Yadav, B.S., Singh, D.,Marchant, R., 1998. Decolourization and biodegradation of anaero-bically digested sugarcane molasses spent wash effluent from biometh-anation plants by white-rot fungi. Process Biochem. 33, 83–88.

Lee, T.H., Aoki, H., Sugano, Y., Shoda, M., 2000. Effect of molasses onthe production and activity of dye-decolourizing peroxidase fromGeotrichum candidum Dec 1. J. Biosci. Bioeng. 89, 545–549.

Livernoche, D., Jurasek, L., Desrochers, M., Dorica, J., 1983. Removal ofcolour from kraft mill wastewaters with cultures of white-rot fungi andimmobilized mycelium of Coriolus versicolour. Biotechnol. Bioeng. 25,2055–2065.

Mahimaraja, S., Bolan, N.S., 2004. Problems and prospects of agriculturaluse of distillery spentwash in India. SuperSoil 2004. 3rd AustralianNew Zealand Soils Conference. 5-9 December’2004. University ofSydney, Australia.

Malandra, L., Wolfaardt, G., Zietsman, A., Viljoen-Bloom, M., 2003.Microbiology of a biological contactor for winery wastewater treat-ment. Water Res. 37, 4125–4134.

Manisankar, P., Rani, C., Vishwanathan, S., 2004. Effect of halides in theelectrochemical treatment of distillery effluent. Chemosphere 57,961–966.

Mansur, M., Suarez, T., Fernandez-Larrea, J., Brizuela, M.A., Gonzalez,A.E., 1997. Identification of a Laccase gene family in the new lignin-degrading basidiomycete CECT 20197. Appl. Environ. Microbiol. 63,2637–2646.

Miranda, P.M., Benito, G.G., Cristobal, N.S., Nieto, C.H., 1996. Colourelimination from molasses wastewater by Aspergillus niger. Biores.Technol. 57, 229–235.

Miyata, N., Iwahori, K., Fujita, M., 1998. Manganese-independent and –dependent decolourization of melanoidin by extracellular hydrogenperoxide and peroxidases from Coriolus hirsutus pellets. J. Ferment.Bioeng. 85, 550–553.

Miyata, N., Mori, T., Iwahori, K., Fujita, M., 2000. Microbial decolouri-zation of melanoidin-containing wastewaters: combined use of activatedsludge and the fungus Coriolus hirsutus. J. Biosci. Bioeng. 89, 145–150.

Mohana, S., Desai, C., Madamwar, D., 2007. Biodegradation anddecolourization of anaerobically treated distillery spent wash by anovel bacterial consortium. Biores. Technol. 98, 333–339.

Morales, F., Jimnez-Perez, S., 2001. Free radical scavenging capacity ofMaillard reaction products as related to colour and fluorescence.Food Chem. 72, 119–125.

Moriya, K., Iefuji, H., Shimoi, H., Sato, S., Tadenuma, M., 1990.Treatment of distillery wastewater discharged from beet molasses-spirits production using yeast. J. Ferment. Bioeng. 69, 138–140.

Nandy, T., Shastry, S., Kaul, S.N., 2002. Wastewater management in acane molasses distillery involving bioresource recovery. J. Environ.Manag. 65, 25–38.

Ohmomo, S., Aoshima, I., Tozawa, Y., Sakurada, N., Ueda, K., 1985.Purification and some properties of melanoidin decolourizing enzymes,P-3 and P-4, from mycelia of Coriolus vericolour Ps4a. Agric. Biol.Chem. 49, 2047–2053.

Ohmomo, S., Kaneko, Y., Sirianuntapiboon, S., Somchai, P., Att-hasumpunna, P., Nakamura, I., 1987. Decolourization of molasseswastewater by a thermophilic strain Aspergillus fumigatus G-2-6.Agric. Biol. Chem. 51, 3339–3346.

Ohmomo, S., Daengsabha, W., Yoshikawa, H., Yui, M., Nozaki, K.,Nakajima, T., Nakamura, I., 1988a. Screening of anaerobic bacteriawith the ability to decolourize molasses melanoidin. Agric. Biol.Chem. 57, 2429–2435.

Ohmomo, S., Kainuma, M., Kamimura, K., Sirianuntapiboon, S., Oshima,I., Atthasumpunna, P., 1988b. Adsorption of melanoidin to the myceliaof Aspergillus oryzae Y-2-32. Agric. Biol. Chem. 52, 381–386.

Pant, D., Reddy, U.G., Adholeya, A., 2006. Cultivation of oystermushrooms on wheat straw and bagasse substrate amended withdistillery effluent. World J. Microbiol. Biotechnol. 22, 267–275.

Patel, A., Pawar, P., Mishra, S., Tewari, A., 2001. Exploitation of marinecyanobacteria for removal of colour from distillery effluent. Ind. J.Environ. Prot. 21, 1118–1121.

Pathak, H., Joshi, H.C., Chaudhary, A., Kalra, N., Dwivedi, M.K., 1999.Soil amendment with distillery effluent for wheat and rice cultivation.Water Air Soil Poll. 113, 133–140.

Patil, P.U., Kapadnis, B.P., Dhamankar, V.S., 2003. Decolourisation ofsynthetic melanoidin and biogas effluent by immobilized fungal isolateof Aspergillus niger UM2. AIDA News Letter., 53–57.

Petruccioli, M., Duarte, J.C., Fedrerici, F., 2000. High rate aerobictreatment of winery wastewater using bioreactors with free andimmobilized activated sludge. J. Biosci. Bioeng. 90, 381–386.

Petruccioli, M., Duarte, J.C., Eusibio, A., Federici, F., 2002. Aerobictreatment of winery wastewater using a jet-loop activated sludgereactor. Process Biochem. 37, 821–829.

Raghukumar, C., Rivonkar, G., 2001. Decolourization of molasses spentwash by the white-rot fungus Flavodon flavus, isolated from a marinehabitat. Appl. Microbiol. Biotechnol. 55, 510–514.

D. Pant, A. Adholeya / Bioresource Technology 98 (2007) 2321–2334 2333

Autho

r's

pers

onal

co

py

Raghukumar, C., Mohandass, C., Kamat, S., Shailaja, M.S., 2004.Simultaneous detoxification and decolourization of molasses spentwash by the immobilized white-rot fungus Flavodon flavus isolatedfrom a marine habitat. Enzyme Microb. Technol. 35, 197–202.

Ramakritinan, C.M., Kumaraguru, A.K., Balasubramanian, M.P., 2005.Impact of distillery effluent on carbohydrate metabolism of freshwaterfish, Cyprinus carpio. Ecotoxicology 14, 693–707.

Ramana, S., Biswas, A.K., Singh, A.B., 2002. Effect of distillery effluenton some physiological aspects of maize. Biores. Technol. 84, 295–297.

Ramendra, Awasthi, M., 1992. Anaerobic and aerobic fermentation - aproven biotechnology for distillery effluent treatment. Ind. J. Environ.Prot. 12, 835–838.

Rivero-Perez, M.D., Perez-Magarino, S., Jose, M.L.G., 2002. Role ofmelanoidins in sweet wines. Anal. Chim. Acta. 458, 169–175.

Sangave, P.C., Pandit, A.B., 2006a. Enhancement in biodegradability ofdistillery wastewater using enzymatic pretreatment. J. Environ.Manag. 78, 77–85.

Sangave, P.C., Pandit, A.B., 2006b. Ultrasound and enzyme assistedbiodegradation of distillery wastewater. J. Environ. Manag. 80, 36–46.

Shah, S.S., Desai, J.D., Ramakrishna, C., Bhatt, N.M., 1998. Aerobicbiotreatment of wastewater from dimethyl terephthalate plant usingbiomass support particles. J. Ferment. Bioeng. 86, 215–219.

Sharma, J., Singh, R., 2001. Effect of nutrient supplementation onanaerobic sludge development and activity for treating distilleryeffluent. Biores. Technol. 79, 203–206.

Shayegan, J., Pazouki, M., Afshari, A., 2005. Continuous decolourizationof anaerobically digested distillery wastewater. Process Biochem. 40,1323–1329.

Singh, N.K., Pandey, G.C., Rai, U.N., Tripathi, R.D., Singh, H.B.,Gupta, D.K., 2005. Metal accumulation and ecophysiological effects ofdistillery effluent on Potamogeton pectinatus L. Bull. Environ. Contam.Toxicol. 74, 857–863.

Sirianuntapiboon, S., Sihanonth, P., Somchai, P., Atthasampunna, P.,Hayashida, S., 1995. An adsorption mechanism for melanoidindecolourization by Rhizoctonia sp. Biosci. Biotechnol. Biochem. 59,1185–1189.

Sirianuntapiboon, S., Somchai, P., Ohmomo, S., Atthasampunna, P.,1988. Screening of filamentous fungi having the ability to decolourizemolasses pigments. Agric. Biol. Chem. 52, 387–392.

Sirianuntapiboon, S., Zohsalam, P., Ohmomo, S., 2003. Decolourizationof molasses wastewater by Citeromyces sp. WR-43-6. Process Biochem.39, 917–924.

Sirianuntapiboon, S., Phothilangka, P., Ohmomo, S., 2004. Decolouriza-tion of molasses wastewater by a strain no. BP103 of acetogenicbacteria. Biores. Technol. 92, 31–39.

Sumino, T., Kon, M., Mori, N., Nakajima, K., 1985. Development ofwastewater treatment techniques by immobilized microorganisms.J. Water Waste 27, 1024–1029.

Thakkar, A.P., Dhamankar, V.S., Kapadnis, B.P., 2006. Biocatalyticdecolourisation of molasses by Phanerochaete chrysosporium. Biores.Technol. 97, 1377–1381.

The gazette of India: extraordinary [part i- sec. I], Ministry of Petroleumand Natural Gas Resolution, New Delhi, 3rd September, 2002 no.P-45018/28/2000-C.C.

Travieso, L., Benitz, F., Dupeyron, R., 1999. Algae growth potentialmeasurement in distillery wastes. Bull. Environ. Contam. Toxicol. 62,483–489.

Trivedy, R.K., Nakate, S.S., 2000. Treatment of diluted distillery waste byconstructed wetlands. Ind. J. Environ. Prot. 20, 749–753.

Uppal, J., 2004. Water utilization and effluent treatment in the indianalcohol industry – an overview. In: Liquid Assets, Proceedings of Indo-EU workshop on Promoting Efficient Water Use in Agro-basedIndustries. TERI Press, New Delhi, India, pp. 13–19.

Uzal, N., Gokacay, C.F., Demirer, G.N., 2003. Sequential anaerobic/aerobic) biological treatment of malt whisky wastewater. ProcessBiochem. 39, 279–286.

Valderrama, L.T., Del Campo, C.M., Rodriguez, C.M., Bashan, L.E.,Bashan, Y., 2002. Treatment of recalcitrant wastewater from ethanoland citric acid using the microalga Chlorella vulgaris and themacrophyte Lemna minuscula. Water Res. 36, 4185–4192.

Vijayaraghavan, K., Ramanujam, T.K., 2000. Performance of anaerobiccontact filters in series for treating distillery spentwash. Bioproc. Eng.22, 109–114.

Vlissidis, A., Zouboulis, A.I., 1993. Thermophilic anaerobic digestion ofalcohol distillery wastewaters. Biores. Technol. 43, 131–140.

Watanabe, Y., Sugi, R., Tanaka, Y., Hayashida, S., 1982. Enzymaticdecolourization of melanoidin by Coriolus sp. Agric. Biol. Chem. 46(20), 1623–1630.

Wilkie, A.C., Riedesel, K.J., Ownes, J.M., 2000. Stillage characterizationand anaerobic treatment of ethanol stillage from conventional andcellulosic feedstocks. Biomass Bioenergy. 19, 63–102.

Wolmarans, B., de Villiers, G.H., 2002. Start-up of a UASB effluenttreatment plant on distillery wastewater. Water SA. 28, 63–68.

Yeoh, B.G., 1997. Two-phase anaerobic treatment of cane-molassesalcohol stillage. Water Sci. Tech. 36, 441–448.

Zache, G., Rehm, H.J., 1989. Degradation of phenol by a coimmobilizedentrapped mixed culture. Appl. Microbiol. Biotechnol. 30, 426–432.

2334 D. Pant, A. Adholeya / Bioresource Technology 98 (2007) 2321–2334

Identification, Ligninolytic Enzyme Activityand Decolorization Potential of Two Fungi Isolatedfrom a Distillery Effluent Contaminated Site

Deepak Pant & Alok Adholeya

Received: 16 October 2006 /Accepted: 9 February 2007 / Published online: 7 March 2007# Springer Science + Business Media B.V. 2007

Abstract Two fungal strains producing ligninolyticenzymes and having the potential to decolorizedistillery effluent were isolated from the soil of adistillery effluent contaminated site. DNA was isolat-ed from the pure cultures of these fungi andpolymerase chain reaction (PCR) amplification oftheir internal transcribed spacer (ITS) region ofnuclear ribosomal DNA was carried out. Further, theDNAwas sequenced and the comparison of generatedsequence with database led to their identification asPenicillium pinophilum TERI DB1 and Alternariagaisen TERI DB6 respectively. These two isolatesalong with one isolate of Pleurotus florida EM 1303were assessed for their ligninolytic enzyme activity inculture filtrate as well as after solid state fermentationon two substrates wheat straw and corncob powder.Ergosterol was measured to assess the growth of fungion solid media. Both P. pinophilum TERI DB1 and A.

gaisen TERI DB6 were found to produce laccase,manganese-dependent peroxidases (MnP) and ligninperoxidases (LiP). The immobilized fungal biomasswas then used for decolorization of the post bio-methanated wastewater from the distillery. Reductionin color up to the magnitude of 86, 50 and 47% wasobserved with P. florida, P. pinophilum and A. gaisenrespectively.

Keywords Fungi . ITS . PCR . Ligninolytic enzymes .

Distillery effluent . Decolorization

1 Introduction

In India, cane-molasses based distilleries are includedamong the 17 most polluting industries by the CentralPollution Control Board (CPCB). In 2004, the totalnumber of distilleries had gone up to 319. On anaverage, collectively, these distilleries are reported toproduce 3.25 × 109 l of alcohol and 40.4 × 1010 l ofwastewater annually (Uppal 2004). Due to the largevolumes of effluent generated, their management isparticularly difficult for the industries. The untreatedeffluent is characterized by dark color, high temper-ature, low pH, high ash content and high percentageof dissolved organic and inorganic matter (Beltranet al. 1999). The conventionally treated distilleryeffluent still contains chemical oxygen demand(COD) around 25–30,000 mg l−1 and is dark brownin color. The disposal of this partially treated effluent

Water Air Soil Pollut (2007) 183:165–176DOI 10.1007/s11270-007-9366-4

D. Pant :A. AdholeyaCentre of Bioresources and Biotechnology,TERI University,DS Block, India Habitat Centre, Lodhi Road,New Delhi 110003, India

A. Adholeya (*)Biotechnology and Management of Bioresources Division,The Energy and Resources Institute,DS Block, India Habitat Centre, Lodhi Road,New Delhi 110003, Indiae-mail: aloka@teri.res.in

on vast stretches of land for solar drying is a commonpractice among distilleries (CPCB 2003). The soils ofthese contaminated sites are ideal habitats for micro-organisms adapted to such environments and aretherefore explored and exploited for bioremediationpurpose. A number of filamentous fungi have beenisolated from such sites and utilized for bioremedia-tion practices (Miranda et al. 1996; Raghukumar andRivonkar 2001).

Filamentous fungal isolates are usually identifiedby microscopic examination of characteristic morpho-logic structures after growth on appropriate media.Identification on this basis becomes extremely diffi-cult if the isolate fails to form the diagnosticallyappropriate structures (Brandt et al. 1998). In theabsence of any distinct morphological feature of thesefungi when grown on different types of media,molecular identification becomes the sole criteria.The PCR has become a preferred method for theidentification of fungal cultures from environmentalsamples. Numerous molecular techniques such asquantitative polymerase chain reaction (qPCR), re-striction fragment length polymorphism (RFLP) anal-ysis, random amplified polymorphic DNA (RAPD)analysis, and image analysis have been developed,which are capable of identification and quantificationof fungal species (Dean et al. 2004; Dean et al. 2005).

In recent years, the ligninolytic system of fungiwith respect to their enzymatic potential for thebioremediation of persistent pollutants has beenextensively studied (Pointing 2001). These enzymesexhibit different characteristics depending on thespecies, strains and culture conditions (Saparrat et al.2002). LiP, MnP and laccase are the three majorlignin-degrading enzymes with great potential inindustrial applications (D’souza et al. 2006). Themajor industrial application of these enzymes is foundin degradation of polymeric dyes from the textileindustry and paper whitening. In the present study, P.florida EM 1303, which has been earlier reported bythe authors as being highly efficient in growing ondistillery effluent amended substrate was used forcomparison as a positive control. In the study thisisolate gave significantly enhanced mushroom yieldwith increasing levels of effluent, with biologicalefficiency (BE) as high as 238.6% (Pant et al. 2006).Prior to this, Kahraman and Yesilada (2003) havereported using Pleurotus pulmonarius 43% decolor-

ization of 30% vinasses when cotton stalks wereadded as additional carbon source which stimulatedthe decolourization activity.

The objective of this study was to identify twofungal cultures isolated from distillery-contaminatedsite showing decolorization potential for distilleryeffluent by sequencing. Further the enzymatic activityof these isolates was investigated. An attempt wasmade to increase their enzymatic yield through solid-state fermentation (SSF) using two agro-residues,wheat straw and corn cob as a substrate.

2 Materials and Methods

2.1 Isolation of Soil Fungi

Fungal cultures employed in the study were isolatedfrom effluent contaminated soil of Associated Alco-hols and Breweries Limited (a cane molasses-baseddistillery) located at Barwaha, Madhya Pradesh,Central India. A total of 21 isolates were subjectedto screening on a poly R-478 dye medium fordecolorization activity and six potential isolates werechosen for further study (Pant and Adholeya 2006).Four of these six were identified morphologically butthe two isolates could not be identified on the basis ofmorphological features were identified by sequencing.P. florida EM 1303 was procured from the Centre forMycorrhizal Culture Collection (CMCC), TERI, NewDelhi, India. All the isolates were grown on potatodextrose broth (PDB) (HiMedia, India) by transferringfour discs of 7 mm cut out from the zone of anactively growing colony on a potato dextrose agar(PDA) plate in a 250 ml Erlenmeyer flask with 75 mlPDB. These were incubated for 1 week or till themycelium covered the whole surface of the medium.Fungi on SSF were produced according to the methoddescribed earlier by authors (Pant and Adholeya2006). The fungi were grown in glass bottles onwheat straw and corn cob powder procured locally(40 g based upon oven-dry weight). One-third of thebottle from top was left as airspace. The moisturecontent for each substrate was adjusted to 75% (v/w)by soaking with four different moistening agents viz.water, 1% solution of molasses, PDB and anaerobi-cally digested distillery effluent. This substrate wassteam sterilized at 121°C for 15 min and inoculated

166 Water Air Soil Pollut (2007) 183:165–176

with 3 ml of mycelium suspension. The cultures wereincubated at 25°C.

2.2 Morphologic Characterization

Micro- and macroscopic observations were performedon Petri dishes with PDA and malt extract agar (MEA),respectively. The optimum temperature and pH ofisolates were also determined for both the isolates.

2.3 DNA Isolation

DNA was extracted according to a slightly modifiedmethod of Hamelin et al. (1996). The fungalmycelium growing on PDB was harvested and anymedia attached to it was removed by washing withsterile water. This was dried at −20°C for 30 min andthen ground under liquid N2 using a sterile pestle andmortar. 15 ml of extraction buffer (1.4 M NaCl, 0.1 MTris-HCl (pH 8), 0.02 M EDTA, 200 μl β-2-mercaptoethanol, 2% cetyl-trimethyl-ammonium bro-mide) was added to 100 mg of this groundedmycelium. The mixture was incubated at 65°C for1 h and extracted by adding 10 ml of phenol:choloroform:isoamyl alcohol (25:24:1). This wasgently shaken and centrifuged at 15,000 rpm. Super-natant was transferred to a new tube and the aqueousphase was precipitated with cold isopropanol byincubating for 1 h at −20°C, and then centrifugedfor 15 min at 15,000 rpm. The pellets were washedwith 70% ethanol, air dried, resuspended in 50 μl ofTE buffer (10 mM Tris-HCl (pH 8), 1 mM EDTA)and stored at 4°C.

2.4 PCR and Sequencing

Universal primers ITS1 (5′-TCCGTAGGTGAAC-CTGCGG) and ITS4 (5′-TCCTCCGCTTATTGA-TATGC) (White et al. 1990) were procured fromBangalore Genie, India. DNA amplifications wereperformed in a model PTC 200 thermal cycler (MJResearch). PCR was performed in a 25 μl reactionmixture with final concentration (per reaction) of 1 ×PCR core buffer, 2.5 mM MgCl2, 0.2 mM (each)dNTPs, 10 pmol of each primer, 1 U of AmpliTaq and50 ng of template. First denaturation was carried outfor 3 min at 95°C. Initial amplifications wereperformed as 5 cycles of 95°C for 30 s (denaturation),

52°C for 30 s (annealing) and 72°C for 1.5 min(extension). Further amplification was performed as25 cycles of 95°C for 30 s (denaturation), 51°C for30 s (annealing) and 72°C for 1.5 min (extension),followed by a 10 min final extension of 72°C.Negative controls were carried out without templateto ensure there was no contamination. PCR productswere resolved by electrophoresis in 2% low meltingpoint agarose, and visualized by ethidium bromidestaining. Sequencing of the PCR products was carriedout by LabIndia (Gurgaon, India) on an automatedmulti-capillary DNA sequencer, ABI Prism 3130xlGenetic analyzer (Applied Biosystems, Foster city,CA, USA) using the Big Dye Terminator v.3.1 ReadyReaction Cycle Sequencing Kit (Applied Biosys-tems). Sequences were then matched with thosealready known using the BLAST search option atNCBI Genbank (http://www.ncbi.nlm.nih.gov).

2.5 Measurement of Ergosterol

Ergosterol content in different substrates after SSFwas measured by the method described by Martinet al. (1990). Fifty milligrams of fungal myceliumcovered SSF substrate was ground in a microfugetube using a plastic pestle. 1 ml of absolute ethanoladded and the tube was shaken for 30 s and set in icefor 1 h. It was then centrifuged for 5 min at14,000 rpm. The supernatant was collected and pelletwas resuspended in 1 ml of absolute ethanol andtreated once again as mentioned above. The twosupernatants were pooled, filtered using 0.22 μmnitrocellulose filters (Milipore) and the filtrate ana-lyzed for ergosterol on a HPLC (Agilent 1100 series,Agilent technologies, Deutschland). Only wheat strawand corn cob powder moistened with different agentswithout fungal mycelium growth were used ascontrols. Ergosterol was detected, separated andquantified using C-18 column, 150 × 4.6 mm (SSWakosil, HG, SGE). The samples were eluted with97:3 methanol/water (v/v) with a flow rate of 0.5 mlmin−1 and monitored at 282 nm using variablewavelength UV detector. Fifty microliters of filtratewas injected into the HPLC system. Peak surface areawas measured and compared to the data obtained withstandard of known ergosterol concentration, whichwas injected before and after each series of sample

Water Air Soil Pollut (2007) 183:165–176 167

using ergosterol procured from Sigma chemicals(79% pure).

2.6 Enzyme Extraction from SSF Substrates

This was carried out according to the methoddescribed by Makkar et al. (2001) along with slightmodification. Briefly, the mycelium growing on thesubstrates was taken along with the substrate after 10days and extracted with 1:2.5 (w/v) 50 mM triethanol-amine-maleic buffer, pH 6.0, with continuous stirring.Another set was extracted using 1:2.5 (w/v) water.The extraction step was repeated for a total of fivetimes. Extracts thus obtained were pooled and filteredthrough a glass filter. Resulting filtrate was centri-fuged at 10,000 rpm for 25 min. Coloring materialsmainly consisting of polyphenolic compounds (For-rester et al. 1990) were removed by addition of cross-linked form polyvinylpyrrolidone (PVP) at a finalconcentration of 7.7% (w/v) (Loomis 1969). Theextracts thus obtained were filtered through a glassfilter and processed for the second PVP treatmentfollowed by centrifugation (10,000 rpm, 25 min) toremove fine PVP particles. The treatment withpolyvinylpyrrolidone (PVP) removed the interferen-ces in the enzymatic assays caused either due to thephenolics or polymeric aromatic compounds (Feniceet al. 2003). The decolorized extracts were filteredthrough Millipore filters (pore size, 0.22 μm) andfinally centrifuged at 16,000 rpm for 60 min. Sampleswere processed at 4°C in all steps.

2.7 Enzyme Assays

Lignin Peroxide (LiP), (EC 1.11.1.14) activity wasdetermined by monitoring the oxidation of Veratrylalcohol to veratraldehyde at 37°C as indicated by anincrease in A310 (Tien and Kirk 1988). The reactionmixture (2.5 ml) contained 500 μl enzyme extract,500 μl H2O2 (2 mmol L−1), 500 μl Veratryl alcoholsolution (10 mmol L−1) and 1.0 ml sodium tartaratebuffer pH 3.0 (10 mmol L−1). One unit of enzymeactivity is defined as the amount of enzyme oxidizing1 μmol of substrate per minute. Manganese Peroxi-dase (MnP), (EC 1.11.1.13) activity was measuredwith phenol red as the substrate at A610 (Kuwaharaet al. 1984). Reaction mixture contained 500 μlenzyme extract, 100 μl phenol red solution (1.0 gL−1), 100 μl sodium lactate pH 4.5 (250 mmol l−1),

200 μl bovine serum albumin solution (0.5%), 50 μlmanganese sulphate (2 mmol l−1) and 50 μl H2O2

(2 mmol l−1) in sodium succinate buffer pH 4.5(20 mmol l−1). Activity is expressed as increase inA610 per minute per mililitre. One unit of enzyme

Substrate

0

200

400

600

Water Molasses Effluent PDB

Erg

os

tero

l (

g g

-1)

0

100

200

300

400

500

600

0

100

200

300

400

500

600

Wheat Straw Corn cob

a

b

c

a

c abc

bc

a

bcbc

ab

b

c

a

b

d d d d

bb b b

c

a

b b

LSD (p≤ 0.01)= 207.26

LSD (p≤ 0.01)= 79.48

LSD (p≤ 0.01)= 39.33

Fig. 1 Ergosterol in SSF samples. a P. pinophilum TERI DB1,b A. gaisen TERI DB6 and cP. florida EM 1303. Letters abovethe histogram bars represents Analysis of Variance (ANOVA).Bars with different letters are significantly different at P ≤ 0.01

168 Water Air Soil Pollut (2007) 183:165–176

activity is defined as the amount of enzyme oxidising1 μmol of substrate per minute. Laccase (EC1.10.3.2) activity is determined by the oxidation of2, 2′-azino-bis (3-ethylthiazoline-6-sulfonate), i.e.,ABTS at 37°C (Buswell and Odier 1987). Thereaction mixture (total volume 1 ml) contained600 μl enzyme extract, 300 μl sodium acetate bufferpH 5.0 (0.1 M) and 100 μl ABTS solution (1 mM).Oxidation was followed via the increase in absor-bance at 420 nm. One unit of enzyme activity isdefined as the amount of enzyme oxidizing 1 mmol ofABTS per minute.

2.8 Effluent Decolorization

Effluent used in the experiment was collected fromthe lagoon of the industry where it was kept for solardrying. The different SSF substrates covered withfungal mat were evenly homogenized under sterileconditions and equal amounts (5 g of immobilizedsubstrate) were used to inoculate 100 ml of distilleryeffluent (50% v/v) in a 500 ml Erlenmeyer flask asdescribed earlier by authors (Pant and Adholeya2006). Distillery effluent used for decolorizationstudies had undergone hydroponic treatment to reducehigh nitrogen content (data not shown). Controls withno fungal inoculations were also used to account fordecolorization due to natural microbial action. Incu-bation was carried out without agitation for 28 days at25°C. The flasks were prepared in triplicates for allthe treatments. The absorbance of effluent wasmeasured at 475 nm. Decolorization was calculatedaccording to the formula:

Decolorization (%) = [(initial absorbance j finalabsorbance)/initial absorbance] � 100 (Itoh 2005). Inorder to check the decolorization due to adsorption onsubstrates, controls were taken where effluent wasinoculated only with wheat straw and corncob without

any fungus. In order to rule out effluent decolorizationdue to adsorption, same amount of heat killed fungalbiomass was used for inoculation.

2.9 Statistical Analysis

In all the experiments described in this studytriplicates were set up for each parameter tested. Themeans of three replicate values for all data weresubjected to DMRT (Duncan's Multiple Range Test)at P ≤ 0.01 in a one-way ANOVA (Analysis ofVariance) using the Costat programme.

3 Results

3.1 Morphologic Characterization

Morphologically, isolate TERI DB1 was characterizedby slow growing colonies, reaching 37 ± 0.85 mm indiameter after 4 days on PDA at 25°C; aerial myceliaat first white and later turning black and reverseblack. The optimum pH for this isolate was 8.5. Theisolate TERI DB6 was characterized by slow growingcolonies, reaching 32 ± 0.76 mm in diameter after 4days on PDA at 25°C; aerial mycelia at first white andlater turning dull white and reverse black. Theoptimum pH for this isolate was found to be 10.

3.2 DNA Isolation and Sequencing

Genomic DNA was successfully isolated from themycelia of both the fungal isolates. These genomicDNAs were subjected to PCR-amplification of theITS region of the rDNA using ITS1 and ITS4 primers.The amplified PCR products turned out to beapproximately 600 bp in length. The partial ITSsequences determined in this study have been depos-ited in the GenBank database with the assigned

Table 1 Screening results for ligninolytic enzyme production

Isolate Laccase MnP LiP

P. pinohilum TERI DB1 0.285 ± 0.051b 0.437 ± 0.019a 7.231 ± 0.248b

A. gaisen TERI DB6 0.285 ± 0.024b 0.123 ± 0.019b 14.597 ± 0.667a

P. florida Eger EM 1303 0.806 ± 0.112a ND ND

ND - Not detected.

Mean ± standard error. Means within a column with different superscript letters are significantly different at p ≤ 0.01 according toDuncan’s Multiple Range Test.

Water Air Soil Pollut (2007) 183:165–176 169

0.0

0.1

0.2

0.3

0.4

Water extraction Buffer extraction

La

cc

as

e p

rod

uc

tio

n (

U m

l-1)

0.0

0.1

0.2

0.3

0.4

Substrate

0.0

0.1

0.2

0.3

0.4

Water Molasses Effluent PDB Water Molasses Effluent PDB

Wheat straw Corn cob

bcd

bcd

a

abcabc

cdd

bcd

ab

ab

a

d

ab

abc d d

fgg

a

efg

bcd

bc

def

efg fg

defg

a

fg

cde

defg

efg

ab

abcd

abc

bcd

abc

efgefg

g

abc

cde

ab

efg

a

fg

def

bcd

ab

a

b

c

LSD (p≤ 0.01) = 0.09

LSD (p≤ 0.01)= 0.08

LSD (p≤ 0.01)= 0.09

Fig. 2 Laccase productionby fungal isolates in SSFwith different substrates.a P. pinophilum TERI DB1,b A. gaisen TERI DB6 andc P. florida EM1303.Lettersabove the histogram barsrepresents Analysis of Vari-ance (ANOVA). Bars withdifferent letters are signifi-cantly different at P ≤ 0.01

170 Water Air Soil Pollut (2007) 183:165–176

accession numbers: 18S ribosomal RNA gene, partialsequence; ITS 1, 5.8S ribosomal RNA gene, and ITS2, complete sequence; and 28S ribosomal RNA gene,partial sequence P. pinophilum strain TERI DB1partial sequence, no. DQ778959 and 18S ribosomalRNA gene, partial sequence; ITS 1, 5.8S ribosomalRNA gene, and ITS 2, complete sequence; and 28Sribosomal RNA gene, partial sequence, A. gaisenTERI DB6, no. DQ778960.

3.3 Ergosterol Estimation

Ergosterol, the component of fungal cell wall wasused as a measure of fungal biomass on various

substrates. The growth of each isolate varied accord-ing to the substrate and the moistening agent used. Nopeak of ergosterol was obtained in control samples. InP. pinophilum TERI DB1, maximum ergosterol wasdetected when grown on wheat straw with water,which was not significantly different from its growthon corncob with water (Fig. 1a). The least ergosterolproduction was observed when P. pinophilum TERIDB1 was grown on wheat straw with molasses asmoistening agent. In A. gaisen TERI DB6, maximumergosterol production was observed when grown onwheat straw with effluent as moistening agent, whichwas significantly higher than all the other treatments(Fig. 1b). For P. florida EM 1303 significantly high

Mn

P p

rod

uct

ion

(U

ml-1

)

0

1

2

3

4

Water extraction

Buffer extraction

0

1

2

3

4

Substrate

Water Molasses Effluent PDB Water Molasses Effluent PDB

Wheat straw Corn cob

e

ee

de

e

e

e

e

ab

e

a

e

bc

e

cd

e

cde

de

a

bc

cde

bccd

bc

cde

e

cd

b

bc

de

cdbc

LSD (p≤0.01)= 0.69

LSD (p≤0.01)= 0.53

a

b

Fig. 3 MnP production byfungal isolates in SSF withdifferent substrates.a P. pinophilum TERI DB1,b A. gaisen TERI DB6.Letters above the histogrambars represents Analysis ofVariance (ANOVA). Barswith different letters aresignificantly different atP ≤ 0.01

Water Air Soil Pollut (2007) 183:165–176 171

growth was recorded when grown on corncob withmolasses. The minimum growth was observed whenthe same was grown on corncob with water (Fig. 1c).

3.4 Enzyme Activity

For initial screening, ligninolytic enzyme activity waschecked in culture filtrates. Both P. pinophilum TERIDB1 and A. gaisen TERI DB6 were positive for allthe three enzymes tested viz., laccase, MnP and LiP,while only laccase activity was detected in P. floridaEM 1303 (Table 1). In SSF mode, enzyme productionby fungi again varied according to the substrate used.Both water and a buffer were used to check if there isa difference in the yield of the enzyme extracted. In

general, extraction with water led to better recovery ofenzymes. For laccase, maximum activity for P.pinophilum TERI DB1 was detected when grown oncorncob with molasses as moistening agent andextracted with water (Fig. 2a). Similar was the casewith A. gaisen TERI DB6, when maximum activitywas detected in same substrate (Fig. 2b). However,for P. florida EM 1303, maximum laccase activitywas observed when extracted with buffer (Fig. 2c). Incase of MnP, maximum activity for P. pinophilumTERI DB1 was observed when grown on corncobwith molasses and extracted with water, which wassignificantly higher than all other treatments irrespec-tive of the extractant used (Fig. 3a). For A. gaisenTERI DB6, maximum MnP was observed when

0

5

10

30

35

40Water extraction Buffer extraction

LiP

Pro

du

cti

on

(U

ml-1

)

0

5

10

30

35

40

Substrate

Water Molasses Effluent PDB Water Molasses Effluent PDB

Wheat straw Corn cob

bb b b

a

b

b

bb

b

b b

b

b

b

b

cd

bcd

cdd

a

bc

cdd

bcd

cd

bcdbcd

bcdbcd

b b

a

b

LSD (p≤ 0.01)= 7.58

LSD (p≤ 0.01) = 4.61

Fig. 4 LiP production byfungal isolates in SSF withdifferent substrates.a P. pinophilum TERI DB1,b A. gaisen TERI DB6.Letters above the histogrambars represents Analysis ofVariance (ANOVA). Barswith different letters aresignificantly different atP ≤ 0.01

172 Water Air Soil Pollut (2007) 183:165–176

grown on wheat straw with molasses and extractedwith water (Fig. 3b). In case of LiP, significantly highactivity was found when P. pinophilum TERI DB1was grown on wheat straw with effluent and extractedwith water (Fig. 4a). Due to extraordinarily highactivity detected in the above treatment, there was nosignificant difference in rest of the treatments. Similarwas the case with A. gaisen TERI DB6 where againhighest activity was observed when grown on wheatstraw with effluent followed by water extraction(Fig. 4b).

3.5 Effluent Decolorization

When this immobilized fungal biomass was appliedfor decolorization of post anaerobically treated dis-tillery wastewater, 86% decolorization was achievedusing P. florida EM 1303 immobilized on corncobpowder (Fig. 5). This was followed by 50 and 47%reduction in color obtained for P. pinophilum TERIDB1 and A. gaisen TERI DB6, respectively. Therewas no color removal due to adsorption as nodecolorization of effluent was observed in wheat

straw and corncob without the fungal inoculum.Besides, in the heat-killed immobilized fungus treat-ment, no decolorization was noticed throughout theexperiment.

4 Discussion

The internal transcribed spacer (ITS) regions offungal rDNA have been successfully used for speciesidentification. The fungal strains isolated during thisresearch were described as P. pinophilum and A.gaisen based on the genetic (ITS region) character-ization. The ITS-1/4 primers, which lie on either sideof the ITS1, 5.8S rDNA and cover regions of both thenuclear 18S rDNA and the nuclear 28S rDNA, werethe primers of choice because they are considered toamplify DNA sequences from a wide range of fungi(Velegraki et al. 1999). The ITS region is present at avery high copy number in the genome of fungi, aspart of tandemly repeated nuclear rDNA whichcoupled with PCR amplification produces a highlysensitive assay (Jasalvich et al. 2000). Earlier, wooddecaying fungi were identified by hybridization ofimmobilized sequence specific oligonucleotide probeswith PCR-amplified fungal rDNA ITS (Oh et al.2003). By sequencing the ITS and large subunitrDNA, three new Penicillium species were described(Peterson et al. 2004). Using a similar approach, anew fungal strain of Bjerkandera audusta wasidentified very recently (Kornillowicz-Kowalskaet al. 2006) and used in decolorization of daunomycinwastes.

Ergosterol is a signature compound of fungal cellwall. Very recently ergosterol was used as a biomark-er to assess the characteristics and abundance of thefungal biomass in atmospheric aerosols (Lau et al.2006). Therefore, by estimating ergosterol production,fungal growth was assessed on different substrates.This gave an idea on the extent of growth of fungalbiomass on the substrates as amended by differentmoistening agents. It was expected that by usingwheat straw and corn cob powder as a carbon source,the rate of effluent decolorization by the fungi couldbe increased because both these substrates containsligno-cellulose, providing conditions that induce theproduction of ligninolytic enzymes. The use of wheatstraw has also been reported for growth of a numberof white-rot fungi for bioremediation of polycyclic

Fungal Isolates

Pleuro

tus f

lorid

a Ege

r EM

1303

Penici

llium

pino

philu

m TERID

B1

Altern

aria

gaise

n TERI D

B6

% D

eco

lori

zati

on

0

20

40

60

80

100

a

bb

Fig. 5 Effluent decolorization by fungal isolates. Letters abovethe histogram bars represents Analysis of Variance (ANOVA).Bars with different letters indicate means with significantdifference at P ≤ 0.01

Water Air Soil Pollut (2007) 183:165–176 173

aromatic hydrocarbon (PAH) contaminated soils(Matsubara et al. 2006). Likewise, Barley grains wereused as a bulking agent for fungi during bio-remediation of creosote contaminated soil (Ataganaet al. 2006). Moreover, in recent years, solid-statefermentation (SSF) has gained ground as a mean ofenhancing enzyme production in fungi. Compared tosubmerged fermentation, SSF is simpler, less capitalintensive, has superior productivity, reduced energyrequirement, simpler fermentation media, uses lesswater and produce lower wastewater, has easiercontrol of bacterial contamination and requires lowcost for downstream processing (Anto et al. 2006;Pandey 1994).

In this study, the production of three ligninolyticenzymes viz., laccase, MnP and LiP by P. pinophilumand A. gaisen and their subsequent use for decolor-ization of distillery effluent is reported. An enhancedproduction of enzymes was also observed after solid-state fermentation. In earlier studies, the molassesdecolorization in solid-state cultivation has only beenreported once by white rot fungi (Kahraman andYesilada 2003) where Coriolus versicolor, Funaliatrogii, Phanerochaete chrysosporium and P. pulmo-narius with cotton stalks were used as additionalcarbon source. In the present study also, agriculturalresidue like wheat straw and corn cob were used as amean for supporting fungal growth, thereby acting asa carbon source.

Traditionally, it has been mainly white rot fungiwhich are used for decolorization of distillery waste-waters (Dahiya et al. 2001; Kumar et al. 1998).Therefore, several screening works about ligninolyticenzymes of white-rot basidiomycetes have beencarried out. The correlation between decolorizingand ligninolytic abilities of white-rot fungi has beencommented upon by several authors (Banat et al.1996; Revankar and Lele 2006; Zhang et al. 2006).Essentially, the main color-causing compound indistillery effluent is melanoidin, which has a chemicalstructure quite similar to humic acid, another recalci-trant compound found in soils and lignin (Plavšićet al. 2006). The ligninolytic activity of white rotsis thought to be responsible for degradation ofmelanoidin.

However, other fungi, representatives of differenttaxonomic and ecophysiological groups, are able todegrade lignocellulosic substrates and produce ligni-

nolytic enzymes (Saparrat et al. 2002). Recently, theproduction of xylanase and protease by P. janthinel-lum from different agricultural wastes has beenreported (Oliveira et al. 2006). MnP Activity wasreported in Alternaria alternata with a possible role inhumic acid degradation by Řezáčová et al. (2006).Penicillium has also been reported to produceligninolytic enzymes (Sack and Gunther 1993) thoughits degrading mechanism is considered to be differentfrom that of the white-rot fungi (Hy et al. 2005). Incase of beet molasses alcoholic fermentation waste-water 41% decolorization by Penicillium decumbenshas been reported (Jimnez et al. 2003). UsingFlavodon flavus, a white-rot basidiomycete fungusisolated from a marine habitat 73% decolorization of10% diluted molasses spent wash (MSW) wasachieved in 7 days (Raghukumar et al. 2004). In astudy similar to the present one, a novel Penicilliumisolate was reported by Zheng et al. (1999) whichcould aerobically decolorize polymeric dyes. Since, ithas been reported that fungal ligninolytic enzymesplay a major role in humic acid and melanoidindecolorization (Blondeau 1989), the degradation ofmelanoidin and hence in its subsequent decolorizationin the present study, can also be attributed to thesame.

5 Conclusion

To conclude, the present work led to the identificationof two novel fungal isolates (P. pinophilum and A.gaisen), which have not been reported earlier fordistillery effluent decolorization. Both the isolatesexhibited ligninolytic enzyme activity production,which was enhanced by solid-state fermentation, thusproviding a scope for their future application in otherbioremediation purposes. Further investigations willentail the characterization of these enzymes and scale-up the process of high biomass recovery in order todevelop a consortium for the treatment at pilot level.

Acknowledgements Authors wish to thank Dr. R. K.Pachauri, Director-General, TERI, and Chancellor, TERIUniversity, New Delhi, India for offering the infrastructure forcarrying out the present investigation. Financial assistance fromUniversity Grants Commission (UGC), New Delhi in the formof Senior Research Fellowship to the first author is dulyacknowledged.

174 Water Air Soil Pollut (2007) 183:165–176

References

Anto, H., Trivedi, U. C., & Patel, K. C. (2006). Glucoamylaseproduction by solid-state fermentation using rice flakemanufacturing waste products as substrate. BioresourceTechnology, 97, 1161–1166.

Atagana, H. I., Haynes, R. J., & Wallis, F. M. (2006). Fungalbioremediation of creosote-contaminated soil: A laborato-ry scale bioremediation study suing indigenous soil fungi.Water, Air and Soil Pollution, 172, 201–219.

Banat, I. M., Nigam, P., Singh, D., & Marchant, R. (1996).Microbial decolorization of textile-dye containing efflu-ents: A review. Bioresource Technology, 58, 217–227.

Beltran, F. J., Garcia-Araya, J. F., & Alvarez, P. M. (1999).Wine distillery wastewater degradation 2. Improvement ofaerobic biodegradation by means of an integrated chemical(ozone)-biological treatment. Journal of Agricultural andFood Chemistry, 47, 3919–3924.

Blondeau, R. (1989). Biodegradation of natural and synthetichumic acids by the white rot fungus Phanerochaetechrysosporium. Applied and Environmental Microbiology,55, 1282–1285.

Brandt, M. E., Padhye, A. A., Mayer, L. W., & Holloway, B. P.(1998). Utility of random amplified polymerase DNAPCR and TaqMan automated detection in molecularidentification of Aspergillus fumigatus. Journal of ClinicalMicrobiology, 36, 2057–2062.

Buswell, J. A., & Odier, E. (1987). Lignin biodegradation. CRCCritical Reviews in Biotechnology, 6, 1–60.

CPCB. (2003). Charter on Corporate Responsibility forEnvironmental Protection. Workshop organized at Mum-bai by MPCB on 03.01.2003 http://cpcb.nic.in//Charter/charter5.htm.

Dahiya, J., Singh, D., & Nigam, P. (2001). Decolorisation ofsynthetic and spentwash melanoidins using the white-rotfungus Phanerochaete chrysosporium JAG-49. Biore-source Technology, 78, 95–98.

Dean, T. R., Betancourt, D., & Menetrez, M. Y. (2004). A rapidDNA extraction method for PCR identification of fungalindoor air contaminants. Journal of MicrobiologicalMethods, 56, 431–434.

Dean, T. R., Roop, B., Betancourt, D., & Menetrez, M. Y.(2005). A simple multiplex polymerase chain reactionassay for the identification of four environmentallyrelevant fungal contaminants. Journal of MicrobiologicalMethods, 6, 9–16.

D’souza, D. T., Tiwari, R., Sah, A. K., & Raghukumar, C.(2006). Enhanced production of Laccase by a marinefungus during treatment of colored effluents and syntheticdyes. Enzyme and Microbial Technology, 38, 504–511.

Fenice, M., Sermanni, G. G., Federici, F., & D’Annibale, A.(2003). Submerged and solid-state production of laccaseand Mn-peroxidase by Panus tigrinus on olive millwastewater-based media. Journal of Biotechnology, 100,77–85.

Forrester, I. T., Grabski, A. C., Mishra, C., Kelly, B. D.,Strickland, W. N., & Leatham, G. F., et al. (1990).Characteristics and N-terminal amino acid sequence of amanganese peroxidase purified from Lentinus edodes

cultures grown on a commercial wood substrate. AppliedMicrobiology and Biotechnology, 33, 359–365.

Hamelin, R. C., Berube, P., Gignac, M., & Bourassa, M.(1996). Identification of root rot fungi in nursery seedlingsby nested multiplex PCR. Applied Microbiology andBiotechnology, 62, 4026–4031.

Hy, Y., Zeng, G. M., Huang, G. H., Huang, D. L., & Chen, Y. N.(2005). Lignin degradation by Penicillium simplicissimum.Huanjing Kexue, 26, 167–171.

Itoh, K. (2005). Decolorization and degradation of methyleneblue by Arthrobacter globiformis. Bulletin of Environmen-tal Contamination and Toxicology, 75, 1131–1136.

Jasalvich, C. A., Ostrofsky, A., & Jellison, J. (2000). Detectionand identification of decay fungi in spruce wood byrestriction fragment length polymorphism analysis ofamplified genes encoding rRNA. Applied and Environ-mental Microbiology, 66, 4725–4734.

Jimnez, A. M., Borja, R., & Martin, A. (2003). Aerobic-anaerobic biodegradation of beet molasses alcoholicfermentation wastewater. Process Biochemistry, 38,1275–1284.

Kahraman, S., & Yesilada, O. (2003). Decolorization andbioremediation of molasses wastewater by white-rot fungiin a semi-solid state condition. Folia Microbiologica, 48,525–528.

Kornillowicz-Kowalska, T. K., Wrzosek, M., Ginalska, G.,Iglik, H., & Bancerz, R. (2006). Identification andapplication of a new fungal strain Bjerkandera audustaR59 in decolorization of daunomycin wastes. Enzyme andMicrobial Technology, 38, 583–590.

Kumar, V., Wati, L., Nigam, P., Banat, I. M., Yadav, B. S., &Singh, D., et al. (1998). Decolorization and biodegradationof anaerobically digested sugarcane molasses spent washeffluent from biomethanation plants by white-rot fungi.Process Biochemistry, 33, 83–88.

Kuwahara, M., Glenn, J. K., Morgan, M. A., & Gold, M. H.(1984). Separation and characterization of two extracellu-lar H2O2-dependent oxidases from ligninolytic cultures ofPhanerochaete chrysosporium. FEBS Letters, 169, 247–250.

Lau, A. P. S., Alex, K. Y., Lee, A. K. Y., Chan, C. K., & Fang, M.(2006). Ergosterol as a biomarker for the quantification ofthe fungal biomass in atmospheric aerosols. AtmosphericEnvironment, 40, 249–259.

Loomis, W. D. (1969). Removal of phenolic compounds duringthe isolation of plant enzymes. Methods in Enzymology,13, 555–563.

Makkar, R. S., Tsuneda, A., Tokuyasu, K., & Mori, Y. (2001).Lentinula edodes produces a multicomponent proteincomplex containing manganese (II)-dependent peroxidase,laccase and L-glucosidase. FEMS Microbiology Letters,200, 175–179.

Martin, F., Delaruelle, C., & Hilbert, J. L. (1990). An improvedergosterol assay to estimate fungal biomass in ectomycor-rhizas. Mycological Research, 94, 1059–1064.

Matsubara, M., Lynch, J. M., & De Leij, F. A. A. M. (2006).A simple screening procedure for selecting fungi withpotential for use in the bioremediation of contaminatedland. Enzyme and Microbial Technology, 39, 1365–1372.

Water Air Soil Pollut (2007) 183:165–176 175

Miranda, P. M., Benito, G. G., Cristobal, N. S., & Nieto, C. H.(1996). Color elimination from molasses wastewater byAspergillus niger. Bioresource Technology, 57, 229–235.

Oh, S., Kamden, D. P., Keathley, D. E., & Han, K. H. (2003).Detection and species identification of wood-decayingfungi by hybridization of immobilized sequence specificoligonucleotide probes with PCR-amplified fungal ribo-somal DNA internal transcribed spacers. Holzforschung,57, 346–352.

Oliveira, L. A., Porto, A. L. F., & Tambourgi, E. B. (2006).Production of xylanase and protease by Penicilliumjanthinellum CRC 87M-115 from different agriculturalwastes. Bioresource Technology, 97, 862–867.

Pandey, A. (1994). Solid-state fermentation (pp. 12–17). NewYork: Wiley.

Pant, D., & Adholeya, A. (2006). Isolation and screening ofpotential fungi for decolorization of distillery wastewaters.In A. Mendez-Vilas (Ed.), Modern multidisciplinaryapplied microbiology exploiting microbes and their inter-actions (pp. 95–102). Weinheim: Wiley.

Pant, D., Reddy, U. G., & Adholeya, A. (2006). Cultivation ofoyster mushrooms on wheat straw and bagasse substrateamended with distillery effluent. World Journal of Micro-biology & Biotechnology, 22, 267–275.

Peterson, S. W., Bayer, E. M., & Wicklow, D. T. (2004).Penicillium thiersii, Penicillium angular and Penicilliumdecaturense, new species isolated from wood-decay fungiin North America and their phylogenetic placement frommultilocus DNA sequence analysis. Mycologia, 96, 1280–1293.

Plavšić, M., Ćosović, B., & Lee, C. (2006). Copper complexingproperties of melanoidins and marine humic material.Science of the Total Environment, 366, 310–319.

Pointing, S. B. (2001). Feasibility of bioremediation by white-rot fungi. Applied Microbiology and Biotechnology, 57,20–33.

Raghukumar, C., & Rivonkar, G. (2001). Decolorization ofmolasses spent wash by the white-rot fungus Flavodonflavus, isolated from a marine habitat. Applied Microbiol-ogy and Biotechnology, 55, 510–514.

Raghukumar, C., Mohandass, C., Kamat, S., & Shailaja, M. S.(2004). Simultaneous detoxification and decolorization ofmolasses spent wash by the immobilized white-rot fungus

Flavadon flavus isolated from a marine habitat. Enzymeand Microbial Technology, 35, 197–202.

Revankar, M. S., & Lele, S. S. (2006). Synthetic dye decolor-ization by white rot fungus, Ganoderma sp. WR-1.Bioresource Technology, 98, 775–780.

Řezáčová, V., Hršelová, H., Gryndlerová, H., Mikšík, I., &Gryndler, M. (2006). Modifications of degradation-resis-tant soil organic matter by soil saprobic microfungi. SoilBiology & Biochemistry, 38, 2292-2299.

Sack, U., & Gunther, T. (1993). Metabolism of PAH by fungiand correlation with extracellular enzymatic activities.Journal of Basic Microbiology, 33, 269–77.

Saparrat, M. C. N., Martinez, M. J., Cabello, M. N., &Arambarri, A. M. (2002). Screening for ligninolyticenzymes in autochthonous fungal stains from Argentinaisolated from different substrata. Revista Iberoamericanade Micología, 19, 181–185.

Tien, M., & Kirk, T. K. (1988). Lignin peroxidase ofPhanerochaete chrysosporium. Methods in Enzymology,6, 238–249.

Uppal, J. (2004). Water utilization and effluent treatment in theindian alcohol industry – An overview. In Liquid assets,proceedings of Indo-EU workshop on Promoting EfficientWater Use in Agro-based Industries, New Delhi 15–16January 2004 (pp. 13–19). New Delhi, India: TERI Press.

Velegraki, A., Kambouris, M. E., Skiniotis, G., Savala, M.,Mitroussia-Ziouva, A., & Legakis, N. J. (1999). Identifi-cation of medically significant fungal genera by polymer-ase chain reaction followed by restriction enzyme analysis.FEMS Immunology and Medical Microbiology, 23, 303–312.

White, T. J., Bruns, T., Lee, S., & Taylor, J. (1990).Amplification and direct sequencing of fungal ribosomalRNA genes for phylogenetics. In M. Innis (Ed.) PCRprotocols: A guide to methods and applications (pp. 315–322). San Diego: Academic.

Zhang, M., Wu, F., Wei, Z., Xiao, Y., & Gong, W. (2006).Characterization and decolorization ability of a laccasefrom Panus rudis. Enzyme and Microbial Technology, 39,92–97.

Zheng, Z., Levin, R. E., Pinkham, J. L., & Shetty, K. (1999).Decolorization of polymeric dyes by a novel Penicilliumisolate. Process Biochemistry, 34, 31–37.

176 Water Air Soil Pollut (2007) 183:165–176

ORIGINAL PAPER

Enhanced production of ligninolytic enzymesand decolorization of molasses distillery wastewaterby fungi under solid state fermentation

Deepak Pant Æ Alok Adholeya

Received: 2 September 2006 / Accepted: 22 November 2006 / Published online: 20 December 2006� Springer Science+Business Media B.V. 2006

Abstract Selected isolates of fungi were grown

on wheat straw and corncob in the presence of

different moistening agents such as water, molas-

ses, potato dextrose broth and distillery effluent.

All the fungal isolates responded differently with

respect to growth and ligninolytic enzyme pro-

duction. Fungal growth on different substrates

was checked by calculating ergosterol content,

which varied widely within a single species when

grown on different substrates. The maximum

laccase production was obtained for Aspergillus

flavus TERI DB9 grown on wheat straw with

molasses. For manganese peroxidase, highest

production was in Aspergillus niger TERI DB20

grown on corncob with effluent. Among the two

isolates positive for lignin peroxidase, the highest

production was in Fusarium verticillioides ITCC

6140. This immobilized fungal biomass was then

used for decolorization of effluent from a cane

molasses based distillery. Maximum decoloriza-

tion (86.33%) was achieved in Pleurotus ostreatus

(Florida) Eger EM 1303 immobilized on corncob

with molasses in a period of 28 days.

Keywords Decolorization � Distillery effluent �Ergosterol � Fungal biomass � Ligninolytic

enzymes � Solid state fermentation

Introduction

White-rot fungi constitute a diverse ecophysio-

logical group comprising mostly of basidiomyce-

tous and litter-decomposing fungi. These fungi

exhibit extensive bioremediation activities that

are mainly based upon their capabilities to pro-

duce one or more extracellular lignin-modifying

enzymes (Wesenberg et al. 2003). Lignin perox-

idases (LiP), manganese-dependent peroxidas-

es (MnP) and laccase are the three major

lignin-degrading enzymes with great potential in

industrial applications (D’Souza et al. 2006).

Production of these enzymes from white-rot fungi

has been well documented. However, in recent

years, there are several reports of these lignino-

lytic enzymes being produced from other fungi

like Phylosticta, Aspergillus, Fusarium and Peni-

cillium (Sahoo and Gupta 2005; Shah et al. 2005;

Kumari et al. 2002). Recently, laccase, lignin

peroxidase, xylanase, endo-1,4-b-D-glucanase and

exo-1,4-b-D-glucanase production by Aspergillus

sp. on agricultural waste of banana under solid

D. Pant � A. AdholeyaCentre of Bioresources and Biotechnology, TERIUniversity, DS Block, India Habitat Centre, LodhiRoad, New Delhi 110003, India

A. Adholeya (&)Biotechnology and Management of BioresourcesDivision, TERI, DS Block, India Habitat Centre,Lodhi Road, New Delhi 110003, Indiae-mail: aloka@teri.res.in

123

Biodegradation (2007) 18:647–659

DOI 10.1007/s10532-006-9097-z

state fermentation (SSF) condition was reported

by Shah et al. (2005). Extracellular MnP produc-

tion under alkaline conditions has been reported

in Aspergillus terreus (Kanayama et al. 2002).

Laccase production in Fusarium proliferatum

cultures, using wheat bran as a natural lignin–

carbon source and benzyl alcohol as laccase

inducer has been reported by Fernaud et al.

(2006). Edible and medicinal mushroom Pleuro-

tus are well known for their ability to produce

extracellular ligninolytic enzymes: laccase (Lac),

two peroxidases: Mn dependent peroxidase

(MnP), versatile peroxidase (VP) and aryl-

alcohol oxidase (AAO) (Munoz et al. 1997).

Recently production of laccase and MnP by

Pleurotus eryngii, P. ostreatus and P. pulmonarius

both under conditions of submerged fermentation

(SF) and SSF was reported (Stajic et al. 2006).

Phanerochaete chrysosporium in submerged

mode has been used for decolorization of olive oil

mill wastewater (Kissi et al. 2001). Earlier, a

peroxidase from Geotrichum candidum has been

used for decolorization of dyes (Kim and Shoda

1999). In recent years, SSF has gained ground as a

mean of enhancing enzyme production in fungi.

Immobilization is considered as a natural state for

fungi, since in nature most fungi tend to attach

firmly on surfaces (Pandey et al. 2001). Thus,

artificially immobilized microorganisms tend to

produce extracellularly secondary metabolites.

Wheat straw is the most common among all

the various substrates that have been employed

for this purpose (Pickard et al. 1999; Aikat

and Bhattacharyya 2000; Valaskova and

Baldrian 2006). Recently sago hampas was used

(Vikineswary et al. 2006) for laccase production by

Pycnoporus sanguineus in SSF. Corncob has also

been used by several researchers as a SSF substrate

for enhanced enzyme production (Couto and

Ratto 1998; Cabaleiro et al. 2002; Oliveira et al.

2006).

These lignin-degrading fungal enzymes lack

substrate specificity thus making them capable of

degrading a wide range of xenobiotics including

industrial colored wastewaters such as dyes

and distilleries (Novotny et al. 2001; Pant and

Adholeya 2006a). Distilleries are among the

most polluting industries generating large amount

of wastewater known as molasses spent wash

(MSW). In India, there are 319 distilleries at

present, producing 3.25 · 109 l of alcohol and

generating 40.4 · 1010 l of wastewater annually

(Uppal 2004), thus making this problem particu-

larly acute. This dark brown colored effluent,

when discharged into water bodies, defiles the

natural ecosystem (FitzGibbon et al. 1998). The

conventionally treated effluent has dark brown

color, strong objectionable odor and contains

COD in the range of 25–30,000 mg l–1 thereby

making it imperative to explore new micro-

organisms and methods for its effective treatment.

The objective of this study was to investigate

the production of ligninolytic enzymes by fungi

isolated from distillery effluent and effluent con-

taminated soils. An attempt was made to enhance

the production of these ligninolytic enzymes by

using two agricultural residues wheat straw and

corncob powder as substrate. The effect of

different extractants viz. water and buffer in

enzyme recovery is evaluated. The subsequent

use of this immobilized fungal biomass in molas-

ses distillery wastewater decolorization has been

reported. Further, the role of different moistening

agents on fungal growth and enzyme yield is also

discussed.

Materials and methods

Microorganisms

The soil samples and effluent after primary

treatment was collected from effluent dumping

site of Associated Alcohols and Breweries Lim-

ited, Barwaha, Madhya Pradesh, a distillery in

central India. Sampling was done three times in a

year during different seasons in order to have

maximum diversity of microorganisms. Through-

out sampling, the soil pH was in between 8.5 and

9.5. The soil was serially diluted 10-fold in 0.85%

saline, and diluted sample (0.1 ml) was spread on

the potato dextrose agar (PDA) plate. The plates

were incubated at 25�C for 4 days. The microbial

colonies (fungi) that appeared on the PDA plates

were isolated and purified. These were character-

ized at Indian Type Culture Collection (ITCC) at

Indian Agricultural Research Institute (IARI),

New Delhi, India based on their morphological

648 Biodegradation (2007) 18:647–659

123

structures such as color, diameter of the mycelia

and microscopic observation of spore formation.

One isolate Pleurotus ostreatus (Florida) EM

1303 was procured from the Centre for Mycor-

rhizal Culture Collection (CMCC), TERI, New

Delhi, India. All these isolates were maintained

and subcultured on PDA media (Hi Media, India)

plates at 25�C.

Preparation of SSF substrates and culture

conditions

The water holding capacity was estimated to be

100 ml per 50 g of wheat straw and 150 ml per

50 g of corncob powder. The culture bottles were

filled with wheat straw and corncob such that

80% of their volume was left as headspace to

increase the oxygen transfer inside the lignocel-

lulose complex material. The moisture content for

each substrate was adjusted to 75% (v/w). Four

different types of moistening agents viz., water,

1% (v/v) molasses, 100% distillery effluent and

Potato Dextrose Broth (PDB, a commercial

media) were used to check their effect and

suitability for enhancing fungal growth and

subsequent enzyme production. The bottles were

subsequently sealed and autoclaved for 60 min at

121�C and cooled to room temperature prior to

inoculation. After sterilization, each flask was

surface inoculated with five discs containing

culture mycelium and agar, 6 mm in diameter,

from agar plate cultures (4–5 day old) of the fungi

on PDA medium. Three replications were pre-

pared for each treatment, and an uninoculated

flask served as control. All the flasks were

incubated at 25�C in the dark.

Enzyme extraction from SSF substrates

This was done according to an existing method

(Makkar et al. 2001) and its slight modifications.

Here, water and buffer were used for enzyme

extraction to check the difference in extracted

enzyme yield. Briefly, mycelium growing on

the substrates was taken along with the sub-

strate after 10 days and extracted with 1:2.5 (w/

v) 50 mM triethanolamine–maleic buffer, pH

6.0, with continuous stirring. Another set was

extracted using 1:2.5 (w/v) water. The extraction

step was repeated five times. Extracts obtained

were finally pooled and filtered through a glass

filter. Resulting filtrate was centrifuged at

10,000 rpm for 25 min. Coloring materials mainly

consisting of polyphenolic compounds in the

supernatant (Forrester et al. 1990) were removed

by addition of cross-linked form polyvinylpyrroli-

done (PVP) at a final concentration of 7.7% (w/v)

(Loomis 1969). The extracts thus obtained were

filtered through a glass filter and processed for the

second PVP treatment followed by centrifugation

(10,000 rpm, 25 min) to remove fine PVP parti-

cles. The treatment with PVP removed the

interferences in the enzymatic assays caused

either due to the phenolics or polymeric aromatic

compounds (Fenice et al. 2003). The decolorized

extracts were filtered through Millipore filters

(pore size, 0.22 lm) and finally centrifuged at

16,000 rpm for 60 min. Samples were processed at

4�C in all steps.

Measurement of ergosterol

Ergosterol, a component of fungal cell wall was

used as a measure of fungal biomass on various

substrates. Ergosterol content in different sub-

strates after SSF was measured by an earlier

described method (Martin et al. 1990). About

50 mg of fungal mycelium covered SSF substrate

was ground in a microfuge tube using a plastic

pestle. About 1 ml of absolute ethanol added and

the tube was shaken for 30 s, set in ice for 1 h and

then centrifuged for 5 min at 14,000 rpm. The

supernatant was collected and pellet was resus-

pended in 1 ml of absolute ethanol and treated

once again as mentioned above. The two super-

natants were pooled, filtered using 0.22 lm nitro-

cellulose filters (Millipore) and the filtrate

analyzed for ergosterol on a HPLC (Agilent

1100 series, Agilent technologies, Deutschland).

Ergosterol was detected, separated and quantified

using C-18 column, 150 · 4.6 mm (SS Wakosil,

HG, SGE). The samples were eluted with 97:3

methanol/water (v/v) with a flow rate of

0.5 ml min–1 and monitored at 282 nm using

variable wavelength UV detector. About 50 ll

of filtrate was injected into the HPLC system.

Peak surface area was measured and compared

to the data obtained with standard of known

Biodegradation (2007) 18:647–659 649

123

ergosterol concentration, which was injected

before and after each series of sample using

ergosterol procured from Sigma chemicals (79%

pure).

Enzyme activity determination

Lignin peroxide (LiP), (EC 1.11.1.14) activity was

determined by monitoring the oxidation of verat-

ryl alcohol to veratraldehyde at 37�C as indicated

by an increase in A310 (Tien and Krik 1988). The

reaction mixture (2.5 ml) contained 500 ll

enzyme extract, 500 ll H2O2 (2 mmol l–1),

500 ll veratryl alcohol solution (10 mmol l–1)

and 1.0 ml sodium tartrate buffer pH 3.0

(10 mmol l–1). One unit of enzyme activity is

defined as the amount of enzyme oxidizing

1 lmol of substrate per minute. Manganese

Peroxidase (MnP), (EC 1.11.1.13) activity was

measured with phenol red as the substrate at A610

(Kuwahara et al. 1984). Reaction mixture con-

tained 500 ll enzyme extract, 100 ll phenol red

solution (1.0 g l–1), 100 ll sodium lactate pH 4.5

(250 mmol l–1), 200 ll bovine serum albumin

solution (0.5%), 50 ll manganese sulfate

(2 mmol l–1) and 50 ll H2O2 (2 mmol l–1) in

sodium succinate buffer pH 4.5 (20 mmol l–1).

Activity is expressed as increase in A610 per

minute per milliliter. One unit of enzyme activity

is defined as the amount of enzyme oxidizing

1 lmol of substrate per minute. Laccase (EC

1.10.3.2) activity is determined by the oxidation of

2, 2¢-azino-bis (3-ethylthiazoline-6-sulfonate), i.e.,

ABTS at 37�C (Buswell and Odier 1987). The

reaction mixture (total volume 1 ml) contained

600 ll enzyme extract, 300 ll sodium acetate

buffer pH 5.0 (0.1 M) and 100 ll ABTS solution

(1 mM). Oxidation was followed via the increase

in absorbance at 420 nm. One unit of enzyme

activity is defined as the amount of enzyme

oxidizing 1 mmol of ABTS per minute.

Effluent decolorization

The different SSF substrates covered with fungal

mat were evenly homogenized under sterile

conditions and equal amounts (5 g of immobi-

lized substrate) were used to inoculate 100 ml of

distillery effluent in a 500 ml Erlenmeyer flask as

described previously (Pant and Adholeya 2006b).

Distillery effluent used for decolorization studies

had undergone hydroponic treatment to reduce

high nitrogen content (data not shown). In

this process effluent was treated using root

zone system of Phragmites kharka and Vetiveria

zizanoides. Physico-chemical characteristics of

this effluent are given in Table 1. Controls with

no fungal inoculations were also used to account

for decolorization due to natural microbial action.

In order to rule out effluent decolorization due to

adsorption, same amount of heat killed fungal

biomass was used for inoculation. Incubation was

carried out without agitation for 28 days at 25�C.

The flasks were prepared in triplicates for all the

treatments. The absorbance of effluent was mea-

sured at 475 nm. Decolorization was calculated

according the formula given (Itoh 2005): Decol-

orization (%) = [(initial absorbance–observed

absorbance)/initial absorbance] · 100. In order

to check the decolorization due to substrates

alone, non-sterile blank controls were taken to

mimic the actual effluent from the industries

where effluent was inoculated with wheat straw

and corncob without any fungal growth on them.

Table 1 Physico-chemical characteristics of distilleryeffluent

Parameter Anaerobically treatedeffluent (releasedin field)

Electrical conductivity (mS cm–1) 33.16pH 8.20BOD5 (ppm) 5,000COD (ppm) 25,000Total Kjeldahl Nitrogen (%) 3.50Sodium (ppm) 500Potassium (ppm) 2,500Manganese (ppm) 259.44Magnesium (ppm) 98.00Zinc (ppm) 272.97Copper (ppm) 395.51Total dissolved solids (ppm) 21,256Total sugar (%) 2.80Reducing sugar (%) 0.23

Pant et al. (2006)

650 Biodegradation (2007) 18:647–659

123

Statistical analysis

In all the experiments described in this study

triplicates were set up for each parameter tested.

Completely randomized design was used and sam-

pling was random. The means of three replicate

values for all data in the experiments obtained

were tested in a One-way analysis of variance

using the Costat software (CoHort, Berkeley).

Results and discussion

Identification of isolated fungi

The search for newer strains of fungi producing

ligninolytic enzymes and their use in decoloriza-

tion of industrial wastewaters especially from

distilleries is a continuous process. Apart from

white rot fungi, which have been exploited so far,

there are fungi from other groups as well which

can be used for this purpose. A number of fungi

were isolated from both the effluent as well as

soils contaminated with this effluent. Based on

their morphology, the isolates with bioremedia-

tion potential were identified as Aspergillus flavus

TERI DB9, Fusarium verticillioides ITCC 6140,

Aspergillus niger TERI DB18 and A. niger TERI

DB20.

Ergosterol based biomass estimation

Ergosterol is the membrane component of most

fungi and ergosterol levels are commonly used to

estimate fungal biomass on various substrates

(Charcosset and Chauvet 2001). For instance, in a

recent study, ergosterol of basidiomycete Gano-

derma lucidum was used to study its growth after its

solid state fermentation on cornmeal (Han et al.

2005). In our study, all isolates exhibited different

growth response on different SSF substrates as

evidenced by varying amount of ergosterol pro-

duced (Table 2). Even within the same substrate,

the effect of different moistening agents was quite

pronounced. In A. flavus TERI DB9, maximum

ergosterol was detected when grown on wheat

straw with PDB, which was significantly higher

than corncob with PDB. The minimum ergosterol

in case of A. flavus TERI DB9 was when grown on Ta

ble

2E

rgo

ste

rol

con

ten

t(l

gg

–1)

infu

ng

al

iso

late

sg

row

no

nw

he

at

stra

wa

nd

corn

cob

we

tte

dw

ith

dif

fere

nt

mo

iste

nin

ga

ge

nts

Iso

late

Wa

ter

Mo

lass

es

Effl

ue

nt

PD

B

Wh

ea

tst

raw

Co

rnco

bW

he

at

stra

wC

orn

cob

Wh

ea

tst

raw

Co

rnco

bW

he

at

stra

wC

orn

cob

A.

fla

vu

sT

ER

ID

B9

32

.86

c±

2.9

87

2.9

5b

±9

.97

40

.83

c±

2.6

58

2.4

7b

±5

.05

68

.40

b±

1.2

23

9.4

6c

±1

.45

11

4.8

1a

±3

.20

63

.46

b±

0.5

0L

SD

(0.0

1)

18

.18

F.

ver

tici

llio

ides

ITC

C6

14

07

6.5

3b

c±

3.9

41

41

.72

a±

3.3

76

0.9

3c

±1

.75

12

8.2

01

a±

25

.90

74

.22

bc

±5

.53

11

0.3

6ab

±7

.16

10

8.5

4ab

±4

.50

10

5.4

ab

c±

3.6

2L

SD

(0.0

1)

41

.76

A.

nig

erT

ER

ID

B1

85

6.8

5c

±2

.50

96

.77

b±

2.1

71

01

.87

b±

3.3

13

5.0

0d

±2

.13

23

2.8

3a

±4

.29

45

.77

cd

±3

.39

11

3.4

7b

±8

.33

63

.89

c±

4.1

1L

SD

(0.0

1)

17

.45

A.

nig

erT

ER

ID

B2

06

9.4

4f

±4

.57

73

.40

ef

±3

.32

85

.49

de

±2

.59

41

.61

g±

2.1

99

2.1

7cd

±1

.75

98

.82

c±

3.4

01

38

.27

a±

2.4

11

15

.61

b±

2.1

7L

SD

(0.0

1)

12

.11

P.

ost

rea

tus

(Flo

rid

a)

EM

13

03

22

2.6

4b

±2

7.6

28

4.4

3c

±1

1.4

51

96

.80

b±

21

.26

46

4.6

1a

±1

2.7

71

76

.34

b±

35

.37

23

5.5

1b

±0

.18

20

1.9

3b

±5

.42

23

8.5

3b

±1

3.1

2

LS

D(0

.01

)7

9.4

8

Me

an

±st

an

da

rde

rro

r.M

ea

ns

wit

hin

aro

wfo

llo

we

db

yth

esa

me

sup

ers

crip

tle

tte

ra

ren

ot

sig

nifi

can

tly

dif

fere

nt

acc

ord

ing

toD

un

can

’sM

ult

iple

Ra

ng

eT

est

(P£

0.0

1)

Biodegradation (2007) 18:647–659 651

123

wheat straw with water. However, this was not

significantly different from its growth on wheat

straw with molasses, and corncob with effluent.

The maximum ergosterol in F. verticillioides ITCC

6140 was observed when grown on corncob powder

with water. However, this was not significantly

different from the one when it was grown on

corncob with molasses or effluent or PDB and

wheat straw with PDB. The growth was signifi-

cantly less when grown on wheat straw with water,

molasses or effluent. For A. niger TERI DB18,

growth was maximum, when grown on wheat straw

with distillery effluent as moistening agent which

was significantly higher than all other treatments.

This was followed by growth on wheat straw with

PDB, wheat straw with molasses and corncob with

water, which did not differ significantly. The

growth was significantly less when it was grown

on wheat straw with water, corncob with effluent

and corncob with PDB. In A. niger TERI DB20,

growth in terms of ergosterol was highest on

growing it on wheat straw with PDB followed by

corncob on PDB and corncob on effluent. All other

treatments resulted in significantly reduced

growth. P. ostreatus (Florida) Eger EM 1303

recorded maximum growth on corncob with molas-

ses which was significantly higher than rest of the

treatments. There was no significant difference in

growth due to all the other treatments except for

corncob with water where growth was significantly

less.

SSF based enzyme production

For enzyme production, SSF mode was chosen in

this study since it has been reported that produc-

tion of ligninolytic enzymes is repressed by

agitation in submerged liquid culture (Galhaup

et al. 2002). Most of the studies so far have been

carried out in submerged liquid culture conditions

or solid cultures on agar plates, which do not

reflect the natural living conditions of these fungi

(Boer et al. 2004). Agriculture residues such as

wheat straw and corncob powder as a support

provides the fungus a similar environment to its

natural habitat and offers the possibility of re-

using an agricultural waste. Recently, enhanced

production of P. ostreatus (Florida) EM 1303 on

wheat straw amended with distillery effluent was

reported by Pant et al. (2006).

Among the five selected fungi, only two, A.

flavus TERI DB9 and P. ostreatus (Florida) EM

1303 were found as laccase producers (Table 3).

Like ergosterol, enzyme production also followed

the similar trend with same isolate producing

varying amount of enzyme when grown on

different SSF substrates (Table 4). Also, extrac-

tion with water and buffer influenced the amount

of enzyme detected. In A. flavus TERI DB9,

maximum laccase production was detected when

grown on wheat straw with effluent and extracted

with water, which was significantly higher from

the one, which was extracted with buffer. All

other treatments had no significant difference on

laccase production by A. flavus TERI DB9. With

P. ostreatus (Florida) EM 1303, maximum laccase

was detected when grown on corncob with

molasses and extracted with buffer, which was

non-significantly different from when this isolate

was grown on corncob with PDB; wheat straw

with molasses and extracted with buffer, wheat

straw with water and extracted with water and

buffer. In general, extraction with water gave

significantly reduced yield of enzyme for

P. ostreatus (Florida) EM 1303 when grown on

wheat straw with effluent and PDB, and corncob

with water and effluent. Earlier, Bucher et al.

(2004) reported the production of wood

decay enzymes including laccase from tropical

Table 3 Screening results for ligninolytic enzyme produc-tion (U ml–1)

Isolate Laccase MnP LiP

A. flavusTERI DB9

0.45a ± 0.01 0.51b ± 0.05 ND

F. verticillioidesITCC 6140

ND 0.34b ± 0.08 2.03a ± 0.35

A. nigerTERI DB18

ND 1.12a ± 0.14 ND

A. nigerTERI DB20

ND 0.81ab ± 0.76 0.89b ± 0.08

P. ostreatus(Florida)EM 1303

0.81a ± 0.11 ND ND

ND—Not detected. Mean ± standard error. Means withina column followed by the same superscript letter are notsignificantly different according to Duncan’s MultipleRange Test (P £ 0.01)

652 Biodegradation (2007) 18:647–659

123

freshwater fungi. Recently, laccase production by

two strains of Aspergillus has been reported

(Souza et al. 2005), which were further used for

treatment of delignification effluent from a nitro-

cellulose industry.

Four isolates viz., A. flavus TERI DB9,

F. verticillioides ITCC 6140, A. niger TERI

DB18 and A. niger TERI DB20 were found

positive for MnP (Table 5). In A. flavus TERI

DB9, maximum MnP was detected when grown

on corncob powder with molasses and extracted

with water, which was not significantly different

when extracted with buffer. Minimum yield was

observed when A. flavus TERI DB9 was grown

on wheat straw with water and extraction was

also with water. For F. verticillioides ITCC 6140,

maximum MnP was detected when grown on

wheat straw with water, and extraction with

water, which again was not significantly different

from its extraction with buffer. Growth on wheat

straw with effluent and extraction with water and

buffer also did not vary significantly when grown

on wheat straw with molasses. However, rest of

the treatments resulted in significantly reduced

MnP yield with minimum in case of growth on

corncob with molasses and extracted with buffer.

MnP in A. niger TERI DB18 was found highest

when grown on corncob with molasses and

extracted with buffer. The minimum was found

when A. niger TERI DB18 was grown on wheat

straw with water and extracted with buffer. Here,

in many treatments such as wheat straw with PDB

and corncob with water, non-significant differ-

ence was observed in yield irrespective of the

extractant used. Similar trend was observed when

grown on corncob with effluent and PDB. For A.

niger TERI DB20, maximum yield was found in

corncob with effluent, and extraction with water.

Again this was not significantly different when

extracted with buffer. Among other treatments,

there was no significant difference in MnP yield

when A. niger TERI DB20 was grown on wheat

straw with water and PDB, corncob with water,

molasses and PDB irrespective of the extractant.

Earlier MnP production by A. terreus has been

reported (Kanayama et al. 2002). Besides this, the

expression of P. chrysosporium MnP in A. niger

has also been reported (Conesa et al. 2000). Low

levels of MnP in certain isolates of FusariumTa

ble

4L

acc

ase

pro

du

ctio

n(U

ml–

1)

by

fun

gal

iso

late

sin

SS

Fw

ith

dif

fere

nt

sub

stra

tes

Wh

eat

stra

wC

orn

cob

Iso

late

Wate

rM

ola

sses

Effl

uen

tP

DB

Wate

rM

ola

sses

Effl

uen

tP

DB

Wate

rB

uff

er

Wate

rB

uff

er

Wate

rB

uff

er

Wate

rB

uff

er

Wate

rB

uff

er

Wate

rB

uff

er

Wate

rB

uff

er

Wate

rB

uff

er

A.

flavu

sT

ER

ID

B9

0.0

97

ab

c

±0.0

13

0.0

68

bc

±0.0

13

0.1

94

ab

c

±0.0

10.0

98

ab

c

±0.0

06

0.3

47

a

±0.0

48

0.0

48

c

±0.0

03

0.0

95

ab

c

±0.0

18

0.1

09

ab

±0.1

08

0.2

47

ab

c

±0.0

13

0.2

41

ab

c

±0.0

13

0.1

64

ab

c

±0.0

06

0.5

63

ab

c

±0.4

25

0.2

76

ab

c

±0.0

06

0.1

54

ab

c

±0.0

09

0.0

79

bc

±0.0

15

0.2

31

ab

c

±0.0

23

LS

D (0.0

1)

0.2

26

P.

ost

reatu

s(F

lori

da)

EM

1303

0.2

38

ab

cd

±0.0

07

0.2

65

ab

c

±0.0

15

0.2

25

bcd

±0.0

14

0.2

55

ab

c

±0.0

51

0.1

1efg

±0.0

13

0.1

2efg

±0.0

23

0.0

57

g

±0.0

08

0.2

65

ab

c

±0.0

36

0.1

77

cd

e

±0.0

12

0.2

96

ab

±0.0

12

0.1

06

efg

±0.0

04

0.3

23

a

±0.0

19

0.0

73

fg

±0.0

06

0.1

51

def

±0.0

23

0.2

21

bcd

±0.0

10.2

88

ab

±0.0

32

LS

D (0.0

1)

0.0

83

Mean

±st

an

dard

err

or.

Mean

sw

ith

ina

row

foll

ow

ed

by

dif

fere

nt

sup

ers

crip

tle

tters

are

sign

ifica

ntl

yd

iffe

ren

tacc

ord

ing

toD

un

can

’sM

ult

iple

Ran

ge

Test

(P£

0.0

1)

Biodegradation (2007) 18:647–659 653

123

Ta

ble

5M

nP

pro

du

ctio

n(U

ml–

1)

by

fun

gal

iso

late

sin

SS

Fw

ith

dif

fere

nt

sub

stra

tes

Wh

eat

stra

wC

orn

cob

Iso

late

Wate

rM

ola

sses

Effl

uen

tP

DB

Wate

rM

ola

sses

Effl

uen

tP

DB

Wate

rB

uff

er

Wate

rB

uff

er

Wate

rB

uff

er

Wate

rB

uff

er

Wate

rB

uff

er

Wate

rB

uff

er

Wate

rB

uff

er

Wate

rB

uff

er

A.

flavu

sT

ER

ID

B9

0.2

54

d

± 0.0

42

0.9

27

cd

± 0.0

54

2.8

18

a

± 0.1

95

0.2

9b

c

± 0.0

47

0.6

5cd

e

± 0.0

72

0.4

19

bc

± 0.0

33

1.1

29

cd

± 0.1

3

0.3

21

bc

± 0.0

27

0.8

cd ± 0.0

54

0.4

48

cd

± 0.0

59

3.9

09

a

± 0.4

45

3.8

34

a

± 0.3

32

0.3

51

d

± 0.0

54

0.2

91

d

± 0.0

45

0.6

43

cd

± 0.0

33

0.5

68

cd

± 0.0

42

LS

D(0

.01)

0.6

35

F.

ver

tici

llio

ides

ITC

C6140

0.5

68

ef

± 0.0

4

0.8

45

def

± 0.0

66

2.3

92

a

± 0.1

37

2.2

2a

± 0.0

72

2.1

9a

± 0.3

69

1.9

36

ab

± 0.2

21

1.2

71

cd

± 0.0

61

1.1

21

cd

e

± 0.0

98

1.5

32

bc

± 0.0

61

1.2

26

cd

± 0.0

6

0.6

05

ef

± 0.1

49

0.5

01

f

± 0.0

95

0.8

07

def

± 0.1

04

0.7

17

def

± 0.0

45

0.8

59

def

± 0.0

95

0.8

45

def

± 0.0

4L

SD

(0.0

1)

0.5

21

A.

nig

erT

ER

ID

B18

0.2

84

ef

± 0.0

52

0.2

17

f

± 0.0

27

0.7

62

cd

± 0.1

27

0.7

17

cd

e

± 0.0

85

1.7

49

a

± 0.2

63

1.3

3b

± 0.1

66

0.3

89

def

± 0.0

49

0.3

51

def

± 0.0

54

0.4

33

def

± 0.0

45

0.4

04

def

± 0.0

47

1.0

16

bc

± 0.1

3

1.9

51

a

± 0.0

93

0.6

88

cd

e

+ 0.0

94

0.6

73

cd

e

± 0.0

47

0.7

1cd

e

± 0.0

42

0.6

73

cd

e

± 0.0

34

LS

D(0

.01)

0.4

01

A.

nig

erT

ER

ID

B20

0.4

19

e

± 0.0

33

0.4

04

e

± 0.0

47

3.6

77

ab

c

± 0.5

57

0.9

34

e

± 0.1

22

3.2

96

bc

± 0.0

59

2.6

88

cd

± 0.1

57

1.7

79

de

± 0.0

84

1.1

66

e

± 0.0

85

0.9

57

e

± 0.1

76

0.8

52

e

± 0.1

13

0.3

89

e

± 0.0

2

0.3

59

e

± 0.0

26

4.9

18

a

± 1.0

88

4.2

23

ab

± 0.3

71

0.9

57

e

± 0.2

51

0.8

74

e

± 0.2

5L

SD

(0.0

1)

1.3

22

Mean

±st

an

dard

err

or.

Mean

sw

ith

ina

row

foll

ow

ed

by

dif

fere

nt

sup

ers

crip

tle

tters

are

sign

ifica

ntl

yd

iffe

ren

tacc

ord

ing

toD

un

can

’sM

ult

iple

Ran

ge

Test

(P£

0.0

1)

654 Biodegradation (2007) 18:647–659

123

solani have also been observed (Saparrat et al.

2000).

Two isolates namely, F. verticillioides ITCC

6140 and A. niger TERI DB20 were positive for

LiP (Table 6). In F. verticillioides ITCC 6140,

growth on wheat straw with molasses and water

extraction recorded maximum LiP yield fol-

lowed by corncob with PDB and water extrac-

tion. Rest of the treatments gave significantly

reduced yield with minimum when grown on

corncob with water and extraction with buffer.

In case of A. niger TERI DB20, maximum LiP

production was found when grown on wheat

straw with effluent, and extraction with water,

which was significantly higher from the one

extracted with buffer. The minimum yield in

this case, was for growth on corncob powder

with effluent and buffer extraction. LiP has

been reported from Aspergillus sp., isolated

from a mangrove area whose best activity was

in coir pith substrate at 3% concentration

(Ahammed and Prema 2002). Recently LiP

production by Penicillium decumbens was re-

ported (Yang et al. 2005). Although the enzy-

matic system related with decolorization of

melanoidins is yet to be completely understood,

it seems greatly connected with fungal lignino-

lytic mechanisms. One of the enzymatic studies

regarding melanoidin decolorization was re-

ported by Miyata et al. (1998). Color removal

of synthetic melanoidin by C. hirsutus involved

the participation of peroxidases (MnP and MIP)

and the extracellular H2O2 produced by glu-

cose–oxidase, without disregard of a partial

participation of fungal laccase. These authors

used C. hirsutus pellets to decolorize a mela-

noidin containing medium. It was elucidated

that extracellular H2O2 and two extracellular

peroxidases, a manganese–independent peroxi-

dase (MIP) and MnP were involved in decolor-

ization activity.

Distillery effluent decolorization

Maximum decolorization of distillery effluent

achieved was 86.33% in case of P. ostreatus

(Florida) EM 1303 grown on corncob with

molasses after 28 days of incubation, followed

by 74.67% decolorization by A. flavus TERI DB9Ta

ble

6L

iPp

rod

uct

ion

(Um

l–1)

by

fun

ga

lis

ola

tes

inS

SF

wit

hd

iffe

ren

tsu

bst

rate

s

Wh

ea

tst

raw

Co

rnco

b

Iso

late

Wa

ter

Mo

lass

es

Effl

ue

nt

PD

BW

ate

rM

ola

sse

sE

fflu

en

tP

DB

Wa

ter

Bu

ffe

rW

ate

rB

uff

er

Wa

ter

Bu

ffe

rW

ate

rB

uff

er

Wa

ter

Bu

ffe

rW

ate

rB

uff

er

Wa

ter

Bu

ffe

rW

ate

rB

uff

er

F.

ver

tici

llio

ides

ITC

C6

14

09

.29

bc

± 0.6

8

8.9

6cd

± 0.7

3

34

.14

a

± 3.5

23

6.6

4cd

e

± 2.7

58

3.9

43

de

± 0.6

29

5.3

32

de

± 0.6

13

2.1

2d

e

± 0.0

6

4.9

6d

e

± 0.2

8

3.5

de

± 0.2

6

1.9

44

e

± 0.2

87

4.4

09

de

± 0.3

98

1.7

92

cd

e

± 0.0

10

6.6

31

de

± 1.2

69

2.7

42

de

± 0.3

17

17

.9b

± 0.9

4

2.4

91

de

± 0.1

16

LS

D(0

.01

)5

.68

5A

.n

iger

TE

RI

DB

20

2.3

9d

e

± 0.1

2

3.3

9cd

± 0.4

2

4.5

43

c

± 0.1

73

1.6

13

e

± 0.1

17

23

.86

a

± 0.3

27

9.4

44

b

± 0.9

87

3.7

2cd

± 0.1

1

1.3

7e

± 0.3

8

4.2

3c

± 0.1

1

3.2

24

cd

± 0.2

03

4.0

14

cd

± 0.4

76

4.8

03

c

± 0.2

76

2.5

09

de

± 0.0

59

1.5

5e

± 0.2

82

3.7

8cd

± 0.4

6

3.1

99

cd

± 0.2

59

LS

D(0

.01

)1

.43

3

Me

an

±st

an

da

rde

rro

r.M

ea

ns

wit

hin

aro

wfo

llo

we

db

yd

iffe

ren

tsu

pe

rscr

ipt

lett

ers

are

sig

nifi

can

tly

dif

fere

nt

acc

ord

ing

toD

un

can

’sM

ult

iple

Ra

ng

eT

est

(P£

0.0

1)

Biodegradation (2007) 18:647–659 655

123

grown on wheat straw with effluent, 68.33% by F.

verticillioides ITCC 6140 grown on wheat straw

with molasses, 64.67% by A. niger TERI DB18

grown on wheat straw with effluent and 54.67%

by A. niger TERI DB20 grown on wheat straw

with effluent (Fig. 1). Besides this, there was a

reduction in pH, COD and BOD of the effluent

by all the fungi (Fig. 2). Prior to this, Kahraman

and Yesilada (2003) reported molasses decolor-

ization in solid state cultivation by fungi Coriolus

versicolor, Funalia trogii, P. chrysosporium and

Pleurotus pulmonarius with cotton stalks being

used as additional source of carbon. C. versicolor

decolorized 48% of 30% diluted vinasse without

any additional carbon source which increased to

71% on addition of cotton stalks. Very recently

100% decolorization of 10% spent wash by a

marine fungal isolate was reported whose laccase

production was increased several folds in the

presence of phenolic and non-phenolic inducers

(D’souza et al. 2006). Miranda et al. (1996)

reported 69% color removal when MgSO4,

KH2PO4, NH4NO3 and a carbon source was

added to wastewater from alcohol fermentation

involving beet molasses.

Color removal by adsorption is ruled out in

this case, since no decolorization of effluent was

observed in wheat straw and corncob without the

fungal inoculum. In the heat-killed immobilized

fungus treatment, no decolorization was noticed

throughout the experiment. Using Flavodon

flavus, a white-rot basidiomycete fungus isolated

from a marine habitat 73% decolorization of 10%

diluted molasses spent wash (MSW) was achieved

in 7 days (Raghukumar et al. 2004). However, in

this case decolorization was achieved in 50%

diluted effluent. It was proposed that hydrogen

peroxide produced as a result of enzyme activity

might act as a bleaching agent. Also since the

production of H2O2 in several fungi depend on

substrates used and other culture conditions,

several workers have attempted to correlate the

production of ligninolytic enzymes in white-rot

fungi and the rates of decolorization. It has been

shown that it is possible to stimulate the yield of

laccase activity of T. versicolor by using several

Fungal isolate

Pleuro

tus o

strea

tus (

florid

a) E

M 1

303

Asper

gillus

flavu

s TERI D

B9

Fusar

ium ve

rticil

lioide

s ITCC 6

140

Asper

gillus

nige

r TERI D

B18

Asper

gillus

nige

r TERI D

B20

% D

ecol

oriz

atio

n

0

20

40

60

80

100

a

b

bcc

d

Fig. 1 Effluentdecolorization by fungalisolates. Letters above thehistogram bars representsanalysis of variance(ANOVA). Bars withdifferent letters indicatemeans with significantdifference. Bars representmean of three replicatesat P £ 0.01

656 Biodegradation (2007) 18:647–659

123

agricultural wastes (Lorenzo et al. 2002); however,

the decolorizing capacity of the extracellular

liquid was not found proportionately increased.

Conclusions

This is the first report of decolorization of highly

recalcitrant distillery effluent using indigenously

isolated fungi at a higher concentration (50%

v/v). The previous studies carried out for the

decolorization of distillery effluent can be linked

by the fact that almost all of them used effluent at

lower dilutions ranging from 6.5% (Kumar et al.

1998) to 20% (Gonzalez et al. 2000). To the best

of our knowledge, this is also the first report for

extracellular MnP and LiP activities of F. verti-

cillioides. The results of this study clearly show

the suitability of wheat straw and corncob powder

moistened with different liquids such as molasses

and distillery effluent as a support medium for

enhanced production of enzymes by fungi. These

agro-residues would be an effective supplement

to support the decolorization process because of

their low cost and easy availability in most

tropical and subtropical developing countries.

Besides, their usage is environmentally safe and

no extra carbon and/or nitrogen source in the

media is required, when this material is used for

immobilization process to fungus. After immobi-

lization, the fungi could be employed in a reactor

such as a rotating biological contractor (RBC).

This may be a practically economic and easily

employable technique for treating large volumes

of distillery wastewaters.

Acknowledgments Authors wish to thank Dr R. K.Pachauri, Director-General, TERI and Chancellor, TERIUniversity, New Delhi, India for offering the infrastructurefor carrying out the present investigation. Financialassistance from University Grants Commission, NewDelhi in the form of Senior Research Fellowship to thefirst author is duly acknowledged.

References

Ahammed S, Prema P (2002) Influence of media nutrientson synthesis of lignin peroxidase from Aspergillus sp.Appl Biochem Biotechnol 103:327–336

P. ostr

eatus

(flor

ida) E

M13

03

A. flav

us T

ERI DB9

F. ver

ticilli

oides

ITCC 61

40

A. nige

r TERI D

B18

A. nige

r TERI DB20

Per

cen

t re

du

ctio

n

0

20

40

60

80

pH COD BOD

Fungal Isolate

Fig. 2 Per cent change inpH, COD and BOD overcontrol by fungal isolates

Biodegradation (2007) 18:647–659 657

123

Aikat K, Bhattacharyya BC (2000) Protease extraction insolid state fermentation of wheat bran by a local strainof Rhizopus oryzae and growth studies by the soft geltechnique. Process Biochem 35:907–914

Boer CG, Obici L, deSouza CGM, Peralta RM (2004)Decolorization of synthetic dyes by solid state cul-tures of Lentinula (Lentinus) edodes producing man-ganese peroxidases as the main ligninolytic enzyme.Bioresour Technol 94:107–112

Bucher VVC, Pointing SB, Hyde KD, Reddy CA (2004)Production of wood decay enzymes, loss of mass, andlignin solubilization in wood by diverse tropicalfreshwater fungi. Microb Ecol 48:331–337

Buswell JA, Odier E (1987) Lignin biodegradation. CRCRev Biotechnol 6:1–60

Cabaleiro DR, Rodrıguez-Couto S, Sanroman A, LongoMA (2002) Comparison between the protease pro-duction ability of ligninolytic fungi cultivated in solidstate media. Process Biochem 37:1017–1023

Charcosset JY, Chauvet E (2001) Effect of cultureconditions on ergosterol as an indicator of biomassin the aquatic hyphomycetes. Appl Environ Microbiol67:2051–2055

Conesa ACA, Hondel VD, Punt PJ (2000) Studies on theproduction of fungal peroxidases in Aspergillus niger.Appl Environ Microbiol 66:3016–3023

Couto SR, Ratto M (1998) Effect of veratryl alcohol andmanganese (IV) oxide on ligninolytic activity in semisolid cultures of Phanerochaete chrysosporium. Bio-degradation 9:143–150

D’souza DT, Tiwari R, Sah AK, Raghukumar C (2006)Enhanced production of Laccase by a marine fungusduring treatment of colored effluents and syntheticdyes. Enzyme Microb Technol 38:504–511

Fenice M, Sermanni GG, Federici F, D’Annibale A (2003)Submerged and solid-state production of laccase andMn-peroxidase by Panus tigrinus on olive mill waste-water-based media. J Biotechnol 100:77–85

Fernaud JRH, A Marina A, Gonzalez K, Vazquez J,Falcon MA (2006) Production, partial characteriza-tion and mass spectrometric studies of the extracel-lular laccase activity from Fusarium proliferatum.Appl Microbiol Biotechnol. 70:212–221

FitzGibbon F, Singh D, McMullan G, Marchant R (1998)The effect of phenolics acids and molasses spent washconcentration on distillery wastewater remediation byfungi. Process Biochem 33:799–803

Forrester IT, Grabski AC, Mishra C, Kelly BD, StricklandWN, Leatham GF, Burgess RR (1990) Characteristicsand N-terminal amino acid sequence of a manganeseperoxidase purified from Lentinus edodes culturesgrown on a commercial wood substrate. Appl Micro-biol Biotechnol 33:359–365

Galhaup C, Wagner H, Hinterstoisser B, Haltricha D(2002) Increased production of laccase by the wood-degrading basidiomycete Trametes pubescens. En-zyme Microb Technol 30:529–536

Gonzalez T, Terron MC, Yague S, Zapico E, Galletti GC,Gonzalez AE (2000) Pyrolysis/gas chromatography/mass spectrometry monitoring of fungal biotreated

distillery wastewater using Trametes sp. I 62 (CECT20197). Rapid Commun Mass Spectrom 14:1417–1424

Han JR, An CH, Yuan JM (2005) Solid-state fermentationof cornmeal with the basidiomycete Ganodermalucidum for degrading starch and upgrading nutri-tional value. J Appl Microbiol 99:910–915

Itoh K (2005) Decolorization and degradation of methy-lene blue by Arthrobacter globiformis. Bull EnvironContam Toxicol 75:1131–1136

Kahraman S, Yesilada O (2003) Decolorization andbioremediation of molasses wastewater by white-rotfungi in a semi-solid state condition. Folia Microbiol48:525–528

Kanayama N, Suzuki T, Kawai K (2002) Purification andcharacterization of an alkaline manganese peroxidasefrom Aspergillus terreus LD-1. J Biosci Biochem93:405–410

Kim SJ, Shoda M (1999) Purification and characterizationof a novel peroxidase from Geotrichum candidumDec 1 involved in decolorization of dyes. ApplEnviron Microbiol 65:1029–1035

Kissi M, Mountadarb M, Assobhei O, Gargiulo E, Palm-ieri G, Giardina P, Sannia G (2001) Roles of twowhite-rot basidiomycete fungi in decolorisation anddetoxification of olive mill waste water. Appl Micro-biol Biotechnol 57:221–226

Kumar V, Wati L, Nigam P, Banat IM, Yadav BS, SinghD, Marchant R (1998) Decolorization and biodegra-dation of anaerobically digested sugarcane molassesspent wash effluent from biomethanation plants bywhite-rot fungi. Process Biochem 33:83–88

Kumari M, Yadav RS, Yadav KD (2002) Secretion ofligninperoxidase by Penicillium citrinum, Fusariumoxysporum and Aspergillus terreus. Ind J Exp Biol40:802–806

Kuwahara M, Glenn JK, Morgan MA, Gold MH (1984)Separation and Characterization of two extracellularH2O2-dependent oxidases from ligninolytic cultures ofPhanerochaete chrysosporium. FEBS Lett 169:247–250

Loomis WD (1969) Removal of phenolic compoundsduring the isolation of plant enzymes. MethodsEnzymol 13:555–563

Lorenzo MD, Moldes D, Rodriguez CS, Sanroman A(2002) Improving laccase production by employingdifferent lignocellulosic wastes in submerged culturesof Trametes versicolor. Bioresour Technol 82:109–113

Makkar RS, Tsuneda A, Tokuyasu K, Mori Y (2001)Lentinula edodes produces a multicomponent proteincomplex containing manganese (II)-dependent perox-idase, laccase and L-glucosidase. FEMS MicrobiolLett 200:175–179

Martin F, Delaruelle C, Hilbert JL (1990) An improvedergosterol assay to estimate fungal biomass in ecto-mycorrhizas. Mycol Res 94:1059–1064

Miranda PM, Benito GG, Cristobal NS, Nieto CH (1996)Color elimination from molasses wastewater byAspergillus niger. Bioresour Technol 57:229–235

Miyata N, Iwahori K, Fujita M (1998) Manganese-inde-pendent and -dependent decolourization of melanoi-din by extracellular hydrogen peroxide and

658 Biodegradation (2007) 18:647–659

123

peroxidases from Coriolus hirsutus pellets. J FermentBioeng 85:550–553

Munoz C, Guillen F, Martınez TA, Martınez JM (1997).Induction and characterization of laccase in theligninolytic fungus Pleurotus eryngii. Curr Microbiol34:1–5

Novotny C, Rawal B, Bhatt M, Patel M, Sasek V, MolitorisHP (2001). Capacity of Irpex lacteus and Pleurotusostreatus for decolorization of chemically differentdyes. J Biotechnol 89:113–122

Oliveira LA, Porto ALF, Tambourgi EB (2006)Production of xylanase and protease by Penicilliumjanthinellum CRC 87M-115 from different agricul-tural wastes. Bioresour Technol 97:862–867

Pandey A, Soccol CR, Rodriguez-Leon JA, Nigam P(2001) Solid-state fermentation in biotechnology.fundamentals and applications. History and develop-ment of solid-state fermentation. P-3. Asiatech Pub-lishers, New Delhi

Pant D, Adholeya A (2006a) Biological approaches fortreatment of distillery wastewater: a review, BioresourTechnol doi:10.1016/j.biortech.2006.09.027 (in press)

Pant D, Adholeya A (2006b) Isolation and screening ofpotential fungi for decolourization of distillery waste-waters. In: Mendez-Vilas A (eds.) Modern multidisci-plinary applied microbiology exploiting microbes andtheir interactions. Wiley-VCH, Weinheim, pp 95–102

Pant D, Reddy UG, Adholeya A (2006) Cultivation ofoyster mushrooms on wheat straw and bagasse sub-strate amended with distillery effluent. World JMicrobiol Biotechnol 22:267–275

Pickard MA, Vandertol H, Roman R, Duhalt V (1999)High production of ligninolytic enzymes from whiterot fungi in cereal bran liquid medium. Can JMicrobiol 45:627–631

Raghukumar C, Mohandass C, Kamat S, Shailaja MS(2004) Simultaneous detoxification and decolorizationof molasses spent wash by the immobilized white-rotfungus Flavadon flavus isolated from a marine habitat.Enzyme Microb Technol 35:197–202

Sahoo DK, Gupta R (2005) Evaluation of ligninolyticmicroorganisms for efficient decolorization of a smallpulp and paper mill effluent. Process Biochem40:1573–1578

Saparrat MCN, Martınez MJ, Tournier HA, Cabello MN,Arambarri AM (2000) Production of ligninolyticenzymes by Fusarium solani strains isolated fromdifferent substrata. World J Microbiol Biotechnol16:799–803

Shah MP, Reddy GV, Banerjee R, Babu PR, Kothari IL(2005) Microbial degradation of banana waste undersolid state bioprocessing using two lignocellulolyticfungi (Phylosticta spp. MPS-001 and Aspergillus spp.MPS-002). Process Biochem 40:445–451

Souza JVB, da Silva ES, da Silva FT, Paiva TCB (2005)Fungal treatment of a delignification effluent from anitrocellulose industry. Bioresour Technol 96:1936–1942

Stajic M, Persky L, Friesem D, Hadar Y, Wasser SP, NevoE, Vukojevic J (2006) Effect of different carbon andnitrogen sources on laccase and peroxidases produc-tion by selected Pleurotus species. Enzyme MicrobTechnol 38:65–73

Tien M, Kirk TK (1988) Lignin peroxidase of Phanero-chaete chrysosporium. Methods Enzymol 6:238–249

Uppal J (2004) Water utilization and effluent treatment inthe indian alcohol industry—an overview. In: Liquidassets, proceedings of Indo-EU workshop on promot-ing efficient water use in agro-based industries, TERI,New Delhi, 15–16 January 2004, pp 13–19. TERIPress, New Delhi, India

Valaskova V, Baldrian P (2006) Estimation of bound andfree fractions of lignocellulose-degrading enzymes ofwood-rotting fungi Pleurotus ostreatus, Trametes ver-sicolor and Piptoporus betulinus. Res Microbiol157:119–124

Vikineswary S, Abdullah N, Renuvathani M, Sekaran M,Pandey A, Jones EBG (2006) Productivity of laccasein solid substrate fermentation of selected agro-residues by Pycnoporus sanguineus. Bioresour Tech-nol 97:171–177

Wesenberg D, Kyriakides I, Agathos SN (2003) White-rotfungi and their enzymes for the treatment of industrialdye effluents. Biotechnol Adv 22:161–187

Yang JS, Yuan HL, Wang HX, Chen WX (2005) Purifi-cation and characterization of lignin peroxidases fromPenicillium decumbens P6. World J Microbiol Bio-technol 21:435–440

Biodegradation (2007) 18:647–659 659

123