1.shodhganga.inflibnet.ac.in/bitstream/10603/36513/15/15_publications.pdf1. Participated in the...

1. S. Vijayalaxmi, K. A. Anu Appaiah, S. K. Jayalakshmi, V. H. Mulimani & K. Sreeramulu. Production of Bioethanol from Fermented Sugars of Sugarcane Bagasse Produced by Lignocellulolytic Enzymes of Exiguobacterium sp. VSG-1. Applied Biochemistry and Biotechnology (2013) 171: 246–260.

2. S. Vijayalaxmi, S. K. Jayalakshmi, & K. Sreeramulu. Polyphenols from different agricultural residues: extraction, identification and their antioxidant properties. Journal of Food Science and Technology DOI 10.1007/s13197-014-1295-9

3. S. Vijayalaxmi, K.A. Anu Appaiah, S. K. Jayalakshmi, & K. Sreeramulu. Bioethanol production from the pentoses and hexoses of the agro industrial wastes hydrolyzed by the hydrolytic enzymes secreted by Exiguobacterium sp. VSG-1. Renewable Energy (Under review)

4. S. Vijayalaxmi, S. K. Jayalakshmi, & K. Sreeramulu. Ethanol fermentation from sugars of sugarcane bagasse hydrolyzates obtained by enzymatic saccharification of Micrococcus sp. VSG-5. Bioprocess and Biosystems Engineering (Under review)

5. S. Vijayalaxmi, S. K. Jayalakshmi, & K. Sreeramulu. Saccharification of rice straw and wheat straw pretreated with steam explosion by the enzymes of Micrococcus sp. VSG-5 for the production of bio-ethanol. FEMS Microbiology Letters (Under review)

6. Vijayalaxmi. S, Jayalakshmi. S.K, Sreeramulu. K. Fermentation of bio-ethanol from rice bran and wheat bran through hydrothermal pre-treatment and enzymatic hydrolysis by Micrococcus sp. VSG-5 (Communicated)

7. Vijayalaxmi. S, Jayalakshmi. S.K, Sreeramulu. Ethanol production by Candida tropicalis MTCC 230 using corn, groundnut and coffee husks through steam explosion and enzymatic hydrolysis (Communicated)

8. S. Vijayalaxmi, S. K. Jayalakshmi, & K. Sreeramulu. Oligosaccharides production from sugarcane bagasse treated with alkali and enzymes of Exiguobacterium sp. VSG-1 (Under preparation)

9. S. Vijayalaxmi, S. K. Jayalakshmi, & K. Sreeramulu. Production, purification and characterization of alkaliphilic, halotolerent and thermostable cellulase by Exiguobacterium sp. VSG-1 (Under preparation)

10. S. Vijayalaxmi, S. K. Jayalakshmi, & K. Sreeramulu. Production, purification and characterization of alkaliphilic, halotolerent and thermostable mannanase by Exiguobacterium sp. VSG-1 (Under preparation)

Publications other than thesis 1. S. Vijayalaxmi, P. Prakash, S. K. Jayalakshmi, V. H. Mulimani & K. Sreeramulu.

Production of Extremely Alkaliphilic, Halotolerent, Detergent, and Thermostable Mannanase by the Free and Immobilized Cells of Bacillus halodurans PPKS-2. Purification and Characterization. Applied Biochemistry and Biotechnology (2013) 171: 382–395.

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Poster Presented /

Conferences attended

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1. Participated in the National Workshop on “Challenging Techniques in

Phytochemical, Analytical and Pharmocological Evaluation of Herbal Products”

organized by Luqman College of Pharmacy, Gulbarga during 11-12th January,

2008.

2. Participated in the “2nd National Conference on Biotechnology for Industrial and

Rural Development (NCBIRD 2008)” organized by the Dept. of Biotechnology,

Gulbarga University, Gulbarga and Karnataka Rastriya Education society, Bidar,

Karnataka during 17-19th January, 2008.

3. Presented the poster in “International Conference on Current Trends in Chemistry

and Biochemistry (ICCTCB-2009)” organised by Central College Campus,

Bangalore University, Bangalore during 18-19th December, 2009.

4. Participated in the 79th Annual Meeting of The Society of Biological Chemists

(India) on “Regulation of Biochemical and Cellular Processes in Diverse

Systems” organised by Indian Institute of Science, Bangalore, India during 13-

15th December, 2010.

5. Presented the poster in “7th Kannada Vijnana Sammelana, Karnataka Science

Congress” organized by Swadeshi Vijnana Andolana, Karnataka, Bengaluru

during 15-17th September, 2011.

6. Participated in One Day Seminar on “Food Security and Natural Resource

Management” sponsored by UGC-New Delhi, organised by Science and

Technology, Gulbarga University, Gulbarga on 8th October, 2012.

7. Presented the poster in 82nd Annual Meeting of The Society of Biological

Chemists (India) and International Conference on “Genomes: Mechanism and

Function” organized by School of Life Sciences, University of Hyderabad, India

during 2-5th December, 2013.

8. Presented the poster in “Two Days KSTA Regional Conference on Science and

Technology for Development” organized by School of Earth sciences, Central

University of Karnataka funded by Karnataka Science and Technology Academy,

Government of Karnataka during 30-31st January, 2014.

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Production of Bioethanol from Fermented Sugarsof Sugarcane Bagasse Produced by LignocellulolyticEnzymes of Exiguobacterium sp. VSG-1

S. Vijayalaxmi & K. A. Anu Appaiah &

S. K. Jayalakshmi & V. H. Mulimani & K. Sreeramulu

Received: 3 March 2013 /Accepted: 23 June 2013 /Published online: 6 July 2013# Springer Science+Business Media New York 2013

Abstract Exiguobacterium sp. VSG-1 was isolated from the soil sample and characterized forthe production of lignocellulolytic enzymes. Production of these enzymes by the strain VSG-1was carried out using steam-exploded sugarcane bagasse (SCB) and found to secrete cellulase,pectinase, mannanase, xylanase, and tannase. The growth and enzyme production were foundto be optimum at pH 9.0 and 37 °C. Upon steam explosion of SCB, the cellulose increased by42 %, whereas hemicelluloses and lignin decreased by 40 and 62 %, respectively. Enzymatichydrolysis of steam-exploded SCB yielded 640 g/l of total sugars. Fermentation of sugarsproduced from pretreated SCB was carried out by using Saccharomyces cerevisiae at pH 5.0and 30 °C. The alcohol produced was calculated and found to be 62.24 g/l corresponding to78 % of the theoretical yield of ethanol. Hence, the strain VSG-1 has an industrial importancefor the production of fermentable sugars for biofuels.

Keywords Exiguobacterium sp. VSG-1 . Lignocellulolytic enzymes . Sugarcane bagasse(SCB) . Enzymatic hydrolysis . Bioethanol

Introduction

Extensive research has been carried on the conversion of agro waste cellulosic materials tobioethanol in the last two decades [1–7]. The demand for ethanol usage as a chemicals

Appl Biochem Biotechnol (2013) 171:246–260DOI 10.1007/s12010-013-0366-0

S. Vijayalaxmi : V. H. Mulimani :K. Sreeramulu (*)Department of Biochemistry, Gulbarga University, Gulbarga 585106, Karnataka, Indiae-mail: [email protected]

K. A. Anu AppaiahDepartment of Food Microbiology, Central Food Technological Research Institute,Mysore 570020, Karnataka, India

S. K. JayalakshmiAgricultural Research Station, University of Agricultural Sciences-Raichur,Gulbarga 585103, Karnataka, India

Author's personal copy

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feedstock or as an octane enhancer or petrol additive in the market is increasing day by day.Lignocellulosics that show potential for ethanol production include agricultural residues(i.e., corn stover, wheat straw, and rice straw), agricultural by-products (i.e., corn fiber, ricehull, sugarcane bagasse), energy crops (i.e., switch grass, sweet sorghum, high fiber sugar-cane, miscanthus) [8]. The major components of lignocellulosics are cellulose (polymers ofhexose sugars, 35–50 %), hemicelluloses (polymers of pentose sugars, 20–35 %), and lignin(polyphenols, 10–25 %) [8, 9]. The large-scale use of lignocelluloses for the production ofbiofuels or other value-added products depends on the breakdown of cellulose, hemicellu-loses, and lignin into their main components. Pretreatment of lignocelluloses is an importantstep for an efficient use of biomass for the production of fermentable sugars. Removal oflignin and hemicelluloses, reduction of cellulose crystallinity, and increase of porosity inpretreatment process can significantly improve the hydrolysis [10] and avoid the formationof inhibitors. Several pretreatment methods have been in use for the hydrolysis of lignocel-lulosics. The advantages of steam explosion pretreatment include lower energy requiredcompared to mechanical + combination and no recycling of environmental costs. Theconventional methods required 70 % more energy than steam explosion to achievethe same size reduction [11]. Steam explosion is recognized as one of the most cost-effective pretreatment processes for hardwoods and agricultural residues, but it is lesseffective for softwoods [12]. Alkaline [13] and acid [14] hydrolysis methods havebeen used to degrade lignocelluloses. Weak acids tend to remove lignin but result in poorhydrolysis of cellulose whereas strong acid treatment occur under relatively extremecorrosive conditions at high temperature and pH which necessitate the use of expensiveequipment.

Both bacteria and fungi can produce cellulases for the hydrolysis of lignocellulosicmaterials. These microorganisms can be aerobic or anaerobc, mesophilic, or thermophilic.Bacteria belong to Clostridium, Cellulomanas, Bacillus, Thermomonospora, Ruminococcucs,Bacteroides, Erwinia, Acetovibrio, Microbispora, and Streptomyces can produce cellulases[15]. Although many cellulolytic bacteria particularly the cellulolytic anaerobes such asClostridium thermocellum and Bacteroides cellulosolvens produce cellulase with high specificactivity but they do not produce high enzyme titers [7]. Several fungi have been reported toproduce cellulases [16]. A non-exhaustive list of cellulolytic microorganisms of aerobic andanaerobic forms isolated from various habitats has been reported [17]. Fungi and yeasts havefrequently been applied in the development of industrial enzymes. However, bacteria haveseveral advantages over fungi in the production of hydrolytic enzymes, in terms of and enzymestiter and the time; for example, many strains have short generation times and can be easilycultured, making the use of bacteria in the biofuel industry more amiable. Additionally, bacteriaalso have increased resilience to environmental stresses due to their biochemical versatility (i.e.,temperature variations, salinity, oxygen limitation, and change in pH) [18]. Researchers havetypically focused on one group of enzymes during isolation, such as cellulases, hemicellulases,or lignases. For example, white rot fungi are among the greatest microorganisms which candegrade lignin and the most well studied [19]. However, anaerobic bacterium Clostridiumthermocellum and aerobic fungi Trichoderma reesei are among some of the greatest cellulase-producing microorganisms [20]. Nonetheless, none of these microorganisms are efficient atcellulolytic, hemicellulolytic, and ligninolytic activities simultaneously, rendering the opportu-nity for discovery of better lignocellulase-producing isolates. Thus, greater importance is beinggiven for the discovery and characterization of new microorganisms that are able to degradecomplex plant biomass more efficiently into fermentable sugars. An organism utilizes for thispurpose would have to express a mixture of several enzymes including cellobiohydrolases,hemicellulases, and pectinases. Cellulolytic and hemicellulolytic multienzyme complexes have

Appl Biochem Biotechnol (2013) 171:246–260 247


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also been reported in Bacillus circulans, Bacillus megaterium [21, 22], and Paenibacilluscurdlanolyticus [23].

Conventionally, the production of enzymes is very expensive and raw material translatesinto 40–60 % of the production cost [24]. Starch-based substrates such as maize, wheat, oats,cassava, potato, and rice were the potential sources for the ethanol fermentation by microbialprocesses [25]. In USA and Brazil, fuel ethanol is produced by fermentation of corn glucoseand or sucrose [26, 27], but any country with a significant agronomical-based economy canuse current technology for fuel ethanol fermentation. However, diverting food crops andtheir produce for ethanol production is a cause of concern for food and nutrition indeveloping and under developing countries. Lignocellulose materials which represent themost abundant alternative and cost-effective source of biomass can be converted into fuelethanol.

In this perspective, this study was aimed at the utilization of agro waste, sugarcanebagasse, as growth substrates for the production of cellulolytic, hemicellulolytic, andpectinase enzymes. Among the agricultural residues, sugarcane bagasse (SCB) is a substrateof high potential for biotechnological processes, which comprises 40–42 % cellulose, 24–28 %hemicelluloses, and 10–12 % lignin.

In this study, we isolated a new Bacillus sp. Exiguobacterium sp. VSG-1 able to hydrolyzeSCB more efficiently to fermentable sugars. There have been no reports on the production oflignolytic enzymes by Exiguobacterium sp. using sugarcane bagasse as a substrate. Further, thepresent study will investigate the saccharification of steam-exposed SCB by the cell-free extractof Exiguobacterium sp. VSG-1 followed by ethanol fermentation of glucose with Saccharo-myces cerevisiae.

Materials and Methods

Chemicals

Carboxymethylcellulose (medium viscosity, 400–800 cP), cellulose powder (Sigma cellCellulose, Type 20; particle size 20 μm), oat spelt xylan, and locust bean gum were purchasedfrom Sigma Chemical Company (St. Louis, MO, USA). Pectin, tannic acid, starch, and caseinwere purchased from Himedia Chemicals, Mumbai, India. All other reagents were of analyticalgrade.

Isolation and Screening of Microorganism

Soil samples were collected around grain mills of Gulbarga, India, and suspended in sterilesaline. Aliquots were inoculated on nutrient agar plates of pH 7.0. The plates were incubatedat 37 °C for 48 h. Isolated colonies were then purified through a serial streaking method onnutrient agar. From these plates, isolated colonies were taken and repeatedly streaked onnutrient agar to obtain pure cultures. This isolated culture was screened for their ability toproduce enzymes like protease, amylase, xylanase, pectinase, cellulase, and tannase. Thepure bacterial cultures were subsequently transferred into nutrient broth.

Microscopic and Biochemical Characterization

Cell morphology, motility, Gram reaction, and the presence of spores and capsule werestudied using standard protocols. Isolated colonies were tested for catalase, oxidase,

248 Appl Biochem Biotechnol (2013) 171:246–260


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utilization of sugars, growth at low and high pH, citrate utilization, indole and ureaseproduction, MR, VP, nitrate reduction, starch hydrolysis, and gelatin liquefaction. Furthertests, viz., growth at pH 5.0–10.0, temperature ambient from 30 to 50 °C, and salinitytolerance from 1 to 16 % sodium chloride, were carried out. Results were analyzed as perBergey’s Manual of Systematic Bacteriology [28].

16S rRNA-Based Identification

The partial sequence of the amplified DNA was determined by Ocimum Biosolutions,Hyderabad, India, and deposited in GenBank under the accession number JQ312121(http://www.ncbi.nlm.nih.gov/nuccore/JQ312121). Related sequences were obtained fromthe GenBank database (National Center for Biotechnology Information) using BLAST. Aphylogenetic tree was constructed by the neighbor-joining method using the MEGA 5.0software.

Sugar Cane Bagasse Pretreatment by the Steam Explosion Process

SCB was collected from local market, India. The SCB was ground and sieved until the SCBparticles were able to pass through a 60 mesh (0.3 mm) sieve and only these particles wereused for the pretreatment experiments. This material was washed with water until to neutralpH and dried at 50±5 °C to attain 10 % moisture content (untreated material) and steam-exploded separately at certain pressures for 10 to 15 min using autoclave under the followingconditions: 150 and 160 °C for 10 min at 1:10 w/v solid/liquid ratio with 6 rpm agitation. Thepretreated SCB was washed with water several times for the removal of residual quantities ofhemicellulosic hydrolysate and then the mass yield measured.

Enzyme Production

Bacterium was grown in a medium containing (in gram per liter): raw SCB, 10.0; peptone,5.0; NaCl, 5.0; K2HPO4, 2.0; MgSO4, 1.0; and yeast extract, 0.5. After incubation at 37 °C,150 rpm for 48 to 72 h, the contents were centrifuged at 8,000 rpm for 10 min and the cell-freeextract was used as an enzyme source.

Enzyme Assay

The cellulase, mannanase, xylanase, and pectinase activities were carried out according todinitrosalicylic acid method [29]. The reaction mixture consisted of 100 μl of enzymesolution, 400 μl of 1 % (w/v) corresponding substrate, and 500 μl of 50 mM phosphatebuffer of pH 9.0. The mixture was incubated at 50 °C for 10 min. The enzyme activity wasdetermined by measuring the release of reducing sugars. One unit (U) of activity was definedas the amount of enzyme producing 1 μmol/ml/min of reducing sugar under the standard assayconditions. The protein concentration was measured according Lowry’s method [30] usingbovine serum albumin as a standard.

Effect of Temperature, pH, and NaCl Concentrations on the Growth and LignocellulolyticEnzymes Production by Exiguobacterium sp. VSG-1

This was carried out by growing the organism at different temperatures (20–60 °C), differentinitial pH values using 50 mM adequate buffers (4–6, acetate buffer; 7–8, phosphate buffer;



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http://www.ncbi.nlm.nih.gov/nuccore/JQ312121

and 9–12, glycine–NaOH buffer), and NaCl concentrations, 0–16 % (w/v). The enzymesactivity and biomass were measured at optimum growth (48 h).

Enzymatic Hydrolysis of Steam-Exploded Materials

Different concentrations (2 to 50 mg) of the cell-free extract of Exiguobacterium sp. VSG-1grown at 48 h, was added to 500 ml Erlenmeyer flask containing carbonate buffer (50 mM,pH 9.0) and 10 g of steam-exploded or unexploded SCB so that the slurry concentrationbecame 10 % w/v. The flasks were gently mixed to make the slurry uniform. Enzymatichydrolysis experiments were carried out at 40 °C under the static conditions. Liquid lossfrom evaporation of the buffer solution was prevented by tightly sealing the flask. Hydro-lysis was terminated by boiling at 100 °C for 5 min at the end of stipulated time intervals,filtered, and the filtrate was collected. The enzymatic hydrolyzation was calculated aspercentage of reducing sugars released.

Estimation of Total Sugars, Reducing Sugars, and Polyphenols

The filtrate was assayed for total sugars by sulfuric acid method [31], reducing sugars bydinitrosalicylic acid method [29], and inhibitory compounds like polyphenols [32].

Estimation of Sugars by HPLC

Sample slurry was centrifuged (8,000 rpm, 4 °C, 10 min) and filtered. The glucose, xylose,and arabinose concentrations were quantified by high-performance liquid chromatography(HPLC) on a micro Bond pack Amino Carbohydrate column (4.1×300 mm). Samples (20 μl)were injected and eluted with acetonitrile–water (70:30 ratio) at a flow rate of 1 ml/min. Thehydrolyzed products were detected using a refractive index detector.

Culture Conditions of S. cerevisiae

S. cerevisiae (MTCC S-170) is a kind gift of Dr. Anu Appaiah, CFTRI, Mysore. The yeastculture was maintained in YPD agar media containing (in gram per liter): glucose, 20.0;peptone 20.0; yeast extract 10.0; and agar, 20.0 at pH 5.5, temperature at 30 °C.

Inoculum Preparation and Fermentation

Inoculum (25 ml) was prepared in 100 ml Erlenmeyer flask with the following mediacomposition (in gram per liter): glucose, 20.0; peptone 20.0; and yeast extract 10.0 at pH5.0 and incubated on a rotary shaker for 24 h (150 rpm) at 30 °C. After 24 h, the cells wererecovered by centrifugation. The fermentation medium (100 ml) was prepared in 250 mlErlenmeyer flask containing hydrolyzed SCB (about 30 g/l of glucose) with the supplemen-tation of 0.2 % of yeast extract or fish protein as a nitrogen source. The pH of the mediumwas adjusted to 5.0 using 1.0 M acetic acid by inoculating with 10 % of 24 h inoculum andincubated at 37 °C in a rotary shaker at 150 rpm for 24 h.

Growth Versus Ethanol Production of S. cerevisiae

Glucose fermentation was carried out to characterize the time-dependent changes of cellgrowth, sugar consumption, and ethanol production by the yeast. Thus, the culture was



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incubated under aerobic conditions at an agitation speed of 150 rpm. Erlenmeyer flasks(100 ml) plugged with cotton were used for a working volume of 25 ml containinghydrolyzed SCB (30 g/l), yeast extract (10 g/l), or fish protein (10 g/l) at pH 5.0. The flaskswere incubated at 30 °C on a rotary shaker at 150 rpm for 72 h.

Effect of Initial pH on Ethanol Production by S. cerevisiae

The effect of the initial pH of the fermentation medium was studied by using pH values of4.0, 4.5, 5.0, 5.5, and 6.0. The pH was adjusted with 0.1 M acetic acid. The overnight YPDculture was inoculated into 25 ml of medium containing 10 g/l yeast extract and 30 g/l of thedesired sugar described above, and incubated at 30 °C, 150 rpm for 48 h. Precultured cellswere inoculated as described above.

Estimation of Alcohol

Estimation of alcohol was done according to [33]. In brief, standards and sample (2 ml) weretaken in a distillation flask. Volume was made up to 50 ml with distilled water and distilled at50–60 °C. Fifteen milliliters of distillate was collected in a clean conical flask and 25 mlchromic acid solution was added. Volume was made up to 50 ml with distilled water. Conicalflasks were incubated at 50 °C on water bath for 30 min, then the solutions were brought toroom temperature and optical density was read at 600 nm. The ethanol yield was calculatedby modified formula proposed by Gunasekharan and Kamini [34].

Estimation of Alcohol by Gas Chromatography

The alcohols present in the samples were determined using gas chromatography ShimadzuGC-6A. A Porapak Q column with a temperature of 180 °C was used with nitrogen as acarrier gas along with a flow rate of 40 ml/min. The injection and departure temperaturewere 220 and 230 °C. Injected into the column were 0.2 μl of alcohol standards and sampledistillates. Peaks were identified by comparing the retention time of the standard alcohols.

Statistical Analysis

The results are the means of three independent experiments. The data were statisticallyevaluated using parametric statistic program, version 1.01 (Lundon software, Inc., ChagrinFalls, OH, USA).

Results

Identification of the Bacterial Isolate

Strain VSG-1 is a gram-positive bacteria, non-sporulating rods, 0.5–1.06×2–10 mm, occur-ring singly, in pairs, or in short chains, and motile by means of peritrichous flagella. It ismoderately thermophilic (growth between 30 and 50 °C, no growth above 50 °C, optimumat 37 °C) and halotolerant (growth in the presence of 12 % NaCl, optimum 1 % NaCl) andpH range for growth is 5.0–10.0 with optimum at pH 9.0. The isolate produced yellowishorange, smooth, glossy, and opaque colonies on nutrient agar plate. It was positive formethyl red, citrate utilization, arginine utilization, catalase and oxidase reactions, but



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negative for VP reaction, nitrate reduction, indole utilization, and H2S production. Casein,gelatin, and starch were hydrolyzed. It utilized glucose, fructose, sucrose, maltose, mannitol,and trehalose as sole carbon sources. Strain VSG-1 was identified by sequence analysis ofthe amplified 1,029-bp segment of its 16S rRNA gene. The strains VSG-1 belonged to thegenus Exiguobacterium, order Bacillales, family Bacillaceae and showed 99 % similarity tothe members of Exiguobacterium (Fig. 1).

Enzyme Production and Assay

Lignocellulolytic enzymes production was observed in the fermentation broth as soon as thebacterium entered the exponential phase (18 h) and reached maximum in the stationaryphase (48 h). The optimum culture conditions for growth and enzymes production were 48–60 h of incubation after which remained more or less stable until 72 h and then decreased withincrease of incubation time (Fig. 2). Cellulase, pectinase, mannanase, and xylanase activitieswere recorded as 38.4, 48.2, 26.6, and 22.8 U/ml, respectively, at 48 h of incubation.

Effect of pH, Temperature, and NaCl Concentrations on the Growth of Exiguobacteriumsp. VSG-1

The highest growth was observed in alkaline pH (7–10) with an optimum at 9.0. There waslow growth at pH 7, whereas high growth was noticed at pH 8–10. Maximum growth wasobserved in the temperature range of 30–50 °C with optimum at 37 °C. The Exiguobacteriumsp. strain VSG-1 was able to grow in a broad-range NaCl concentration (1–16 %) withoptimum at 1 % NaCl. This clearly indicates the halotolerant nature of the strain VSG-1 (datanot shown).

Fig. 1 Neighbor-joining phylogenetic dendrogram based on 16S rRNA gene sequence data indicating theposition of strain VSG-1 among members of the genus Exiguobacterium. Accession numbers of 16S rRNAgene sequences of reference organisms are indicated. Bootstrap values from 1,000 replications are shown atbranching points; only values above 60 are shown. Bar, 0.01 substitutions per 100 nt



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Pretreatment of SCB

The composition of the raw material used in this work is shown in Table 1. The raw SCBshowed a high carbohydrate content (about 42.6 % of cellulose). This table also shows theproportional increase of the cellulose by 40 %, whereas hemicellulose and lignin contentdecreased by 50 and 60 %, respectively, in the pretreated SCB (150 °C; 160 °C for 10 min)compared the raw SCB, due to solubilization of the hemicellulosic fractions.

Effect of Enzyme Concentration on the Hydrolysis of Pretreated SCB

Significant increase in the production of reducing sugars was observed as the concentrationof enzyme increases and reached enzyme loading at 40 mg protein/g steam-exploded SCB.Thus, the enzymes are saturated at 40 mg protein/g steam-exploded SCB under the exper-imental conditions (Fig. 3).

Estimation of Total Sugars, Reducing Sugars, and Polyphenols

The estimation of total sugars, reducing sugars, and polyphenols were calculated from thehydrolyzate of steam-exploded SCB and found to be 640, 45, and 4.6 mg/ml, respectively,

Table 1 Chemical composition of SCB before and after pretreatments with steam explosion

Components Pretreatment conditions of SCB

Raw SCB 150 °C/10 min 160 °C/10 min

Cellulose 42.6±0.1* 57.3±0.2* 58.8±0.1*

Hemicelluloses 24.7±0.2* 14.8±0.1* 13.5±0.2*

Lignin 22.3±0.1* 23.8±0.2* 23.4±0.1*

Ash 1.5±0.3* 2.2±0.1* 2.8±0.1*

Extractives 5.6±0.1* 1.2±0.2* 1.1±0.2*

Total 96.7±0.1* 98.1±0.1* 98.5±0.1*

± standard deviation

*P<0.05 (two paired sample Student’s t test)

0

0.5

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1.5

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0

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30

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0 8 16 24 32 40 48 56 64

Bio

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ivit

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/ml)

Incubation time (h)

Cellulase Pectinase Mannanase Xylanase Growth

Fig. 2 Effect of incubation time onthe growth and lignocellulolyticenzyme production byExiguobacterium sp. VSG-1 atpH 9.0 and temperature 37 °C



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whereas unexploded SCB hydrolyzate showed low levels of sugars and high levels ofpolyphenols (Table 2).

Effect of pH and Temperature on the Growth and Alcohol Fermentation by S. cerevisiae

Maximum growth and alcohol production by S. cerevisiae S-170 were observed in acidic pH(5.0–6.0) with an optimum at 5.5 and the temperature range of 25–35 °C with optimum at30 °C during the alcohol fermentation (data not shown).

HPLC Analysis of Sugars

The products obtained from enzymatic hydrolysis of pretreated SCB were analyzed andfound to be as glucose, xylose, and arabinose (Fig. 4).

Fermentation of Alcohol

Alcohol production was started in the fermentation broth at 48 h and reached maximum at 72 h.The optimum cultural conditions for the production of alcohol were up to 72 h, which will remainmore or less up to 96 h and then decreased with increase of incubation time (Fig. 5). The estimation

0

20

40

60

80

100

0 2 4 6 8 10 12

Glu

cose

con

cent

rati

on (

g/l)

Reaction time (h)

2 mg/g 5 mg/g 10 mg/g

20 mg/g 30 mg/g 40 mg/g

Fig. 3 Changes in glucose concentrations during enzymatic hydrolysis of 10 % w/v SCB at different loadingsof extracellular enzymes by Exiguobacterium sp. VSG-1 under static condition. Enzyme loading is inmilligram protein/gram steam-exploded SCB

Table 2 Estimation of total sugars, reducing sugars, and polyphenols after pretreatment of steam explosion(hydrothermal) and enzymatic hydrolysis of SCB

Untreated SCB Steam-exploded SCB Enzymatic hydrolyzed SCB

Total sugars (mg/ml) 17.5±0.1* 67±0.2* 640±0.1*

Reducing sugars (mg/ml) 1.5±0.2* 8.2±0.1* 45±0.2*

Polyphenols (mg/ml) 7.4±0.1* 6.2±0.2* 4.6±0.1*





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of total sugars, reducing sugars, and polyphenols was calculated from the hydrolyzate of steam-exploded SCB before and after the fermentation and the results were presented in Table 3.

Estimation and Detection of Alcohol

The alcohol percentage was estimated and found to be 7.8 or 7.2 % when the fermentationwas carried out with the fish protein or yeast extract, respectively, as a nitrogen source(Table 4). The alcohol obtained from yeast fermentation was analyzed by gas chromatogra-phy and identified as ethyl alcohol (Fig. 6).

Discussion

Temperature is a major environmental condition that affects microbial physiology andgrowth. Every bacterial sp. has its own optimum temperature and cultural conditions for

Minutes3 4 5 6 7 8 9 10

0.000

0.002

0.004

4.3

67

4.7

25

5.4

00

4.3-glucose4.7-xylose5.4-galactose

Fig. 4 HPLC profile of sugars present in hydrolysed SCB before fermentation

0

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0 24 48 72 96

Alc

ohol

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cose

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%)

Fermentation days (h)

Glucose concentration (%)

Alcohol content (%v/v)

Fig. 5 Changes in sugar concen-tration and alcohol contents ofSCB sugar syrup over thefermentation period



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survival. However, Exiguobacterium sp. have been isolated from and molecularly detectedin a wide range of habitats. The unique feature of this bacterial sp. is to grow under extremeenvironmental conditions with the temperature ranging from −12 to 55 °C with minimumnutrients. The Exiguobacterium genus comprises psychrotrophic, mesophilic, and moderatethermophilic species and strain with pronounced morphological diversity (ovoid, rods,double rods, and chains) depending on the species, strain, and environmental conditions[35]. Until now, studies of Exiguobacterium sp. mainly focused on characteristics ofresistance to extreme conditions such as high/low temperature, alkaline environment, andhigh concentrations of salts [36]. The biotechnological applications of these strains, espe-cially in the biodegradation of lignocellulosic materials have not been explored.

The characterization of cellulolytic and hemicellulolytic bacteria have been given muchattention as readily available abundance of lignocellulosics carbon source, which can bedegraded to fermentable sugars for the production of bioethanol [37]. Therefore, evaluatingthese activities in our isolate is pertinent to finding an efficient lignocellulosic bacterium. Asa result, it was important for us to distinguish those strains which can degrade amorphous andcrystalline cellulose by cellulase in addition, to xylanase activity. Hence, we could isolate astrain producing high titer of lignocellulosic enzymes. Enzymatic activities of lignocellulolyticsuggest their prominent role during the breakdown of SCB. Several bacterial strains have beenisolated and screened for the lignocellulolytic enzymes [38].

Pretreatment is necessary for lignocellulosics to achieve a highly efficient enzymatichydrolysis and fermentation. Composition of untreated and steam explosion pretreated SCBis summarized in Table 1. Considerable amounts of hemicelluloses (50.20 %) and lignin(65.02 %) were removed during the process. However, the cellulose content is increased bymore than 40 % in the treated SCB. Kim and Lee [39] reported 53–79 % delignification with75–97% removal of hemicelluloses and 4–11% removal of cellulose from corn stover by a twostage hot water and ammonia recycle percolation process at high temperature (170–210 °C).The main objective of the steam explosion pretreatment is the delignification (65.02 %) SCB,which further increases the area and porosity of hemicelluloses and cellulose and its utilizationfor production of value-added materials.

Table 3 Calculation of total sugars, reducing sugars and polyphenols before and after fermentation of alcohol

Before fermentation After fermentation

Total sugars(mg/ml) 640.6±0.1* 58.8±0.1*

Reducing sugars (mg/ml) 45.2±0.1* 8.5±0.1*

Polyphenols (mg/ml) 3.8±0.1* 0.5±0.1*



Table 4 Ethanol yields for untreated and pretreated SCB

g/l Theoretical yield (%)

Untreated SCB 12.3±0.1* 21±0.1*

Pretreated SCB 62.24±0.1* 78±0.1*





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Enzymatic hydrolysis of cellulosic substances was carried out by cellulase enzymes,which are highly specific [40]. The products of the hydrolysis are usually reducing sugarsincluding glucose. Utility cost of enzymatic hydrolysis is low compared to acid or alkalinehydrolysis because enzyme hydrolysis is usually conducted at mild conditions and does nothave any corrosion problem and environmental problems [7]. Untreated and treated SCBwere hydrolyzed for 48 h using cell-free extract of Exiguobacterium sp. VSG-1. The highpercentage of digestibility in the treated material can be attributed to lignin removal. Thecrude supernatant preparation had a wide range of activities on insoluble substrates. Thepredominant was cellulase, but mannanase and pectinase were also present. Since thexylanase activity was low, it is advantageous for hydrolysis process as the supernatantcontains low level of pentoses which are not desirable for the fermentation of alcohol. Thestructures of multienzyme complexes (MEC) have not been elucidated in detail, but many ofthem contain predominantly cellulase activity found in the cellulosomes. In this study, it wasfound that the MECs were able to hydrolyze insoluble cellulose. The ability to bindinsoluble substrates has been considered important due to the fact that degradation ofinsoluble substrates was inextricably linked to the enzymes/complex’s ability to bind andthus remain in close proximity to the substrate while it is hydrolyzed. Further, binding tocrystalline cellulose has been a feature of the cellulosome as the scaffolding protein of thecellulosome which contains a CBM3a domain which is able to bind crystalline substrates[19]. Cellulosomes have been found to bind only very weakly to insoluble xylan [41]. Use ofmixture of cellulases and other enzymes in the hydrolysis of cellulosic materials have beenextensively studied [42–44]. A cellulose conversion yield of 90 % was achieved in theenzymatic saccharification of 8 % alkali-treated sugarcane bagasse when a mixture ofcellulases from Aspergillus ustus and Trichoderma viride [45]. A nearly complete sacchar-ification of steam explosion pretreated Eucalyptus viminalis chips was obtained using acellulose mixture of commercial celluclast and novozyme preparations [46]. Use of com-mercial enzymes for saccharification increases the cost of the process.

The maximum yield of total reducing sugars (640 g/l) was obtained when 10 g of SCBwas mixed with 40 ml of crude enzyme extract. There was not much significant increase ofreducing sugars yield with increasing concentration of crude enzyme extract. The yield ofglucose (395 g/kg) and xylan (135 g/kg) achieved in this study were higher than those sugars

Fig. 6 Gas chromatogram of alcohol from fermented SCB. Oven temperature, 180 °C; injector and detectortemperatures, 220 and 230 °C, respectively



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of sugarcane bagasse (225 g glucose/kg dry and 62 g xylose/kg biomass) [47] and sorghum(glucose, 400–470 g/kg; xylose, 130–170 g/kg dry sorghum) [48] pretreated with ammonia–water.

Several reports have been published on the production of ethanol fermentation bybacteria, yeast, and fungi [49]. The most commonly used yeast, S. cerevisiae, producedethanol as high as 18 % of the fermentation broth and is the preferred one for most of theethanol fermentation. This yeast can grow both on simple sugars, such as glucose, and on thedisaccharide, sucrose. Saccharomyces is also generally recognized as safe as a food additivefor human consumption and is therefore ideal for producing alcoholic beverages and forleavening bread. Ethanol concentration reached its highest peak at 72 h (Figs. 5 and 6) andno further increase was observed at 96 h (data not shown). At the end of the fermentationprocess, ethanol concentrations reached 642 g/kg of pretreated SCB. This yield is higherthan those reported using dilute ammonia-treated sorghum, 250 g/kg [48], and sulfuric acidpretreated sorghum, 141 g/kg [50]. Mamma [51] reported 115 g/kg dry sorghum ethanolfrom sorghum fibers using mixed culture of Fusarium oxysporum and S. cerevisiae. Sugar-cane bagasse and sorghum are grass plants and have the similar composition of lignocellu-losic materials. Shaibani et al. [52] have reported that the only 50 % ethanol was producedby simultaneous saccharification and fermentation of sugarcane bagasse with crude enzymesolutions of Trichoderma longibrachiatum and S. cerevisiae yeast.

Production of fermentable sugars from the cheap agro waste lignocellulosic materials is acrucial step for the success of alcohol and beverage industries. It is further constrained by costlyinputs, process operation, and time. Overall, on comparison with the other strains of bacteriaand pretreatmentmethods, the present study demonstrates that the strain ofExiguobacterium sp.VSG-1 is the most promising candidate, producing high levels of fermentable of sugars fromsteam-exploded SCB. The overall time taken to produce 64.2 g/l of ethanol from pretreatedSCB is 5 days.

The newly isolated strain, Exiguobacterium sp. VSG-1, is an excellent source forlignocellulolytic enzymes for the production of fermentable sugars from SCB. This strainmay have great potential for developing bacterial consortiums in the near future to enhancethe decomposition of lignocellulosic biomass and helps to overcome costly hurdled beingfaced in the industrial production of biofuels. Further, we achieved significant removal oflignin from SCB, which resulted in higher yield of fermentable sugars for the production ofethanol.

Acknowledgments This research was supported by research grants to KS from DST and UGC-SAP, theGovernment of India, New Delhi. VS thank UGC-SAP for providing JRF. We are grateful to Dr. Anu Appaiahfor providing facilities to carry out the experiments in CFTRI.

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ORIGINAL ARTICLE

Polyphenols from different agricultural residues: extraction,identification and their antioxidant properties

S. Vijayalaxmi & S. K. Jayalakshmi & K. Sreeramulu

Revised: 18 January 2014 /Accepted: 10 February 2014# Association of Food Scientists & Technologists (India) 2014

Abstract Agricultural residues like sugarcane bagasse (SCB),corn husk (CH), peanut husk (PNH), coffee cherry husk(CCH), rice bran (RB) and wheat bran (WB) are low-valuebyproducts of agriculture. They have been shown to containsignificant levels of phenolic compounds with demonstratedantioxidant properties. In this study, the effects of two types ofsolvent extraction methods: solid–liquid extraction (SLE) andhot water extraction on the recovery of phenolic compoundsfrom agricultural residues were investigated to optimize theextraction conditions based on total phenolic content (TPC),total tannin content (TTC) and total flavonoids content (TFC).Methanol (50 %) was found to be the most efficient solvent forthe extraction of phenolics with higher DPPH, nitric oxideradical scavenging and reducing power activity, followed byethanol and water. The phenolic compounds of methanolicextracts (50 %) were determined by reverse phase high perfor-mance liquid chromatography; in addition gallic acid became

the major phenolic acid present in all the agricultural residueswhereas ferulic acid, epicatechin, catechin, quercitin andkampferol present in lesser amounts. The present investigationsuggested that agricultural residues are potent antioxidants. Theoverall results of this research demonstrated the potential ofagricultural residues to be an abundant source of natural anti-oxidants suitable for further development into dietary supple-ments and various food additives.

Keywords Agriculture residues . Polyphenols . Tannins .

Flavonoids . Antioxidant activity

Introduction

Polyphenols and flavonoids are widely distributed in plantswith significant applications in the health and food industries(Sarikaya and Ladisch 1999; Ventura et al. 2008). Such com-pounds include a variety of phenolic acids (hydroxybenzoicacids and hydroxycinnamic acids), flavonoids (flavonols, fla-vones, flavonones, isoflavones and anthocyanidins), stilbenesand lignans (Hooper and Cassidy 2006). These phytochemi-cals were known to reduce many chronic diseases such ascardiovascular diseases, heart diseases, diabetes, obesity andcertain cancer and improve endothelial functions and reduceblood pressure (Liu 2007; Yawadio et al. 2007). Among thesecompounds, a strong correlation between antioxidant activityand the total phenolic content in the plants has been observed,suggesting that phenolic compounds could be the major con-tributor of their antioxidant capacity (Li et al. 2008).

Phenolic compounds in large quantities are known to bepresent in edible and non edible plants (Gharras 2009), veg-etables, fruits, herbs and other plant materials which are rich inphenolics are increasingly being used for the extraction ofphenolics at the industrial level. Using valuable horti cropswhich are human food resources, for phenolic production or

Highlights • Agricultural residues like sugarcane bagasse (SCB), cornhusk (CH), peanut husk (PNH), coffee cherry husk (CCH), rice bran (RB)and wheat bran (WB) are used for the extraction of polyphenols.• The effects of two types of solvent extraction methods: solid–liquidextraction (SLE) and hot water extraction on the recovery of phenoliccompounds from agricultural residues were investigated to optimize theextraction conditions based on total phenolic content (TPC), total tannincontent (TTC) and total flavonoids content (TFC).• The types of polyphenols from different agricultural residues wereanalyzed by HPLC.• The antioxidant activities like DPPH, NO and FRAP were also studied.

S. Vijayalaxmi :K. Sreeramulu (*)Department of Biochemistry, Gulbarga University,Gulbarga 585106, Karnataka, Indiae-mail: [email protected]

S. Vijayalaxmie-mail: [email protected]

S. K. JayalakshmiAgricultural Research Station, University of AgriculturalSciences-Raichur, Gulbarga 585103, Karnataka, India

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extraction raisesmajor ethical and moral issues. This would bea very costly affair for the developing and under developingcountries. Expanding phenolics production could divert valu-able fruits and vegetables needed to feed the people. Thisfurther would increases malnutrition in the developing andunder developing countries.

Alternatively, for the production or extraction of bioactivephenolic compounds, agro industrial residues could beexploited. Large quantities from these materials, includingseeds, peels, husks among others are generated every year inthe form of wastes and are poorly harvested or left to decay onthe land. Increased attention is being given for these materialsas abundantly available and cheap renewable feedstocks forthe production of value added compounds.

Antioxidants are any substance that delay or inhibits oxi-dative damage to a target molecule. Antioxidants prevent celland tissue damage as they act as scavenger. Antioxidants canterminate or retard the oxidation process by scavenging freeradicals. Overproduction of the free radicals can be responsi-ble for tissue injury. Anti-oxidants are substances capable tomop up free radicals and prevent them from causing celldamage. Free radicals are responsible for causing a widenumber of health problems which, include cancer, aging, heartdiseases, gastric problems etc.

Antioxidant capacity is widely used as a parameter formedicinal bioactive components. Various methods are cur-rently used to assess the antioxidant activity of plant phenoliccompounds. ABTS or DPPH radical scavenging methods arecommon spectrophotometric procedures for determining theantioxidant capacities of components (Gulçin et al. 2010).

In the present study, waste feedstocks such as sugarcanebagasse (SCB), corn husk (CH), peanut husk (PNH), coffeecherry husk (CCH), rice bran (RB) and wheat bran (WB) werescreened for the extraction of different bioactive phenolics andcharacterized.

Materials and methods

Materials

Agricultural residues such as sugarcane bagasse (SCB), cornhusk (CH), peanut husk (PNH), rice bran (RB) and wheat bran(WB)were obtained from a local agriculture farm at Gulbarga,Karnataka, India. Coffee cherry husks (CCH) were obtainedby grain mill, CFTRI, Mysore.

Chemicals

Methanol and alcohol were purchased from Merck(Darmstadt, Germany). All the chemicals and reference com-pounds were purchased from Sigma Aldrich (Steinheim,Germany).

Sample pretreatment

Agricultural residues were ground between 10 and 30mesh (0.5 mm and 2.0 mm) were collected and defattedusing n-hexane (ratio of solid/liquid 1/10, w/w) at roomtemperature and then the solvent was evaporated.

Extraction and concentration

The extraction of phenolic compounds was carried out usingsolvents at different polarity: 50 %, 70 % and 100 % ofmethanol; 50 %, 70 % and 100 % of ethanol and water. Thepowdered samples were added to a solvent 1:10 mixed welland kept at room temperature, for 3 days under constantstirring. The mixture was centrifuged at 6,000 rpm for15 min and the supernatant was filtered through a filter paper.Then, the solvent was evaporated in a rotavapor. The extrac-tion yield was expressed as dry matter percentage.

Total phenols content (TPC)

Total polyphenols analysis was performed by the colorimetricmethod, as described by Vazquez et al. (2008), with somemodifications. The sample was re-dissolved in the extractionmedium. To 100 μl of sample, 500 μl of Folin-Ciocalteureactive, and 400 μl of 7.5 % aqueous solution of Na2CO3

were added. The mixture was kept for 30 min in the dark atroom temperature. The absorbance was read at 720 nm using aUV/Vis spectrophotometer (UV-6450; Jenway, UK). Gallicacid (5–50 mM) was used for constructing the standard curve,and the results were expressed as g of gallic acid equivalents(GAE)/100 g of extract.

Total tannins content (TTC)

Total tannins analysis were performed by the colorimetricmethod, as described by Price and Butler (1977). To 100 μlof sample, 1 ml of 1 % potassium ferricyanide and 8 % ofaqueous solution of ferric chloride was added. The mixturewas kept for 5 min at room temperature. The absorbance wasread at 720 nm using a UV/Vis spectrophotometer (UV-6450;Jenway, UK). Tannic acid (5–50 mM) was used for construct-ing the standard curve, and the results were expressed as g oftannic acid equivalents (TAE)/100 g of extract.

Total flavonoids content (TFC)

Total flavonoid content was determined by a colorimetricmethod (Bao et al. 2005). 0.5 ml extracts were added to15 ml volumetric flasks containing 2 ml ddH2O and mixedwith 5 % of 0.15 ml NaNO2. After reacting for 5 min, 0.15 ml10 % AlCl3.6H2O solution was added. After another 5 min,1 ml 1 M NaOH was added. The reaction solution was well

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mixed, kept for 15 min and the absorbance was determined at415 nm. Quantification was done using the quercitin as stan-dard and the results were expressed as g of quercitin equiva-lents (QE)/100 g of extract.

HPLC determination of individual polyphenols

The concentration of individual polyphenols was determinedby HPLC Shimadzu LC-10Awith a UV detector recording at280 nmwas used to detect the phenolic compounds. A reversephase C-18 column (15 cm) was used with a flow rate of 0.8/min of a solvent system containing water, methanol, aceticacid were in the ratio of 80:18:2. The standard mixtures ofpolyphenols were prepared in the concentration of 1 mg/ml.The standard and sample of 0.02 ml were injected into thecolumn. Peaks were identified by the retention time of thecommercial standard phenolic compounds.

Antioxidant assays

DPPH (1, 1-diphenyl-2-picrylhydrazyl) radical scavengingassay

The electron donation ability of the obtained methanol ex-tracts of agricultural residues was measured by bleaching ofthe purple colored solution of DDPH (1, 1-diphenyl-2-picrylhydrazyl) radical according to the method of Sun et al.(1988). Methanolic extracts (2 ml, 10–1,000 μg/ml) wereadded to 0.5 ml of 0.2 mM/L DDPH. After incubation periodof 30 min at room temperature, the absorbance was measuredagainst a blank at 517 nm using UV/Vis spectrophotometer(UV-6450; Jenway, UK) versus ethanol as a blank. The anti-oxidant activity was calculated by the following ratio: (blank-sample/blank) × 100, where blank is the absorption of theDPPH solution and sample is the absorption of the DPPHsolution after the addition of the sample.

Nitric oxide (NO) radical scavenging assay

The assay was carried out according to Isfahlan et al. (2010).The reaction mixture contained 10 mM SNP (Sodium nitro-prusside), phosphate-buffered saline (pH 7.4), and the variousconcentrations of the samples. After incubation for 150 min at25 °C, 1 ml of sulfanilic acid (0.33 % in 20 % glacial aceticacid) was added to 0.3 ml of the incubated solution, and themixture allowed to stand for 5 min. NED (naphthyl ethylenediamine dihydrochloride, 0.5 ml, 0.1 % w/v) was added, andthe mixture was incubated for 30 min at 25 °C. The pinkchromophore, generated during diazotization of nitrite ionswith sulfanilic acid and subsequent coupling with NED, wasmeasured from the absorbance at 540 nm, using an appropri-ate blank.

IC50 values were calculated from the plotted graph ofscavenging activity against the concentrations of the samples.IC50 is defined as the total antioxidant necessary to decreasethe initial DPPH and NO radical by 50%. IC50 was calculatedfor all the extracts based on the percentage of DPPH radicalsscavenged. Ascorbic acid was used as the reference com-pound (positive control) with concentrations 50 to500 μg/ml for both the above spectroscopic methods.

FRAP (ferric ion reducing antioxidant potential) assay

The antioxidant activity of samples was determined by theFRAP assay, according to Barriera et al. (2008). Variousconcentrations of extracts (100–500 μg/ml) were mixed with2.5 ml of 200 mM sodium phosphate buffer (pH, 6.6) and2.5 ml of 1 % potassium ferricyanide. The mixture wasincubated at 50 °C for 20 min then 2.5 ml of 1 % TCA(w/w) was added. This was followed by the addition of 5 mlof distilled water and 1 ml of 0.1 % of ferric chloride. Theabsorbance was recorded at 700 nm after 5 min. The antiox-idant activity was calculated from the calibration curve ofascorbic acid (0.1–1 mM). Results were expressed as μmolof ascorbic acid equivalents (AAE)/mg of extract.

Results and discussion

Extraction yield

The results of the extraction yields are presented in Table 1.Organic solvents such as methanol and ethanol with differentratio were used for the extraction of polyphenols and tanninsfrom the agricultural residues. The highest yields are usuallyachieved with methanol and ethanol and their mixtures withwater, although other solvents have been widely used in theextraction of polyphenols from plants, as ethyl acetate oracetone. Water and ethanol are the most widely used becauseof their low toxicity and high extraction yield, with the ad-vantage of modulating the polarity of the solvent by usingethanol/water mixtures at different ratios. The main drawbackof the aqueous extraction is the low yield in antioxidants withlow polarity or liposoluble antioxidants as, for example, thecarotenoids. Solubility of polyphenols depends mainly on thehydroxyl groups and the molecular size and the length ofhydrocarbon. For all used solvents, methanol (50 %) provedto be the most suitable solvent for the extraction of thepolyphenols as compared to the different concentrations ofethanol. The optimization procedure employed in this studyresulted in a similar recovery of phenolic compounds usingjust one extraction stage, thereby allowing for a reduction insolvent consumption and time required for extraction. Amongall agricultural residues, coffee cherry husk (CCH) showed thehigher extraction yields. The largest yield value for CCH was

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Table 1 Extraction yield, total polyphenols, tannins and flavonoids content of the agricultural residues

Agriculturalsamples

Extractionyield (%)

Total polyphenols(g GAE/100 g extract)

Total tannins(g TAE/100 g extract)

Total flavonoids(g RE/100 g extract)

Sugarcane bagasse (SCB)

Water 2.8±0.20 16.20±0.13 9.25±0.20 6.58±0.15

50 % CH3OH 6.2±0.18 52.45±0.20 38.18±0.18 10.20±0.21

70 % CH3OH 5.8±0.16 46.30±0.15 36.12±0.15 9.64±0.20

100 % CH3OH 4.6±0.14 42.68±0.16 32.33±0.11 9.23±0.18

50 % C2H5OH 4.5±0.25 40.15±0.18 31.65±0.22 8.16±0.11

70 % C2H5OH 3.8±0.27 36.33±0.25 28.48±0.14 8.35±0.16

100 % C2H5OH 3.6±0.30 33.70±0.23 24.85±0.31 7.13±0.12

Corn husk (CH)

Water 2.5±0.28 12.40±0.20 7.80±0.08 4.84±0.14

50 % CH3OH 6.0±0.22 48.50±0.24 38.12±0.12 8.23±0.18

70 % CH3OH 5.5±0.16 45.45±0.12 37.15±0.18 7.82±0.23

100 % CH3OH 4.4±0.12 41.35±0.15 35.76±0.13 6.80±0.12

50 % C2H5OH 4.0±0.18 42.65±0.13 33.65±0.22 8.72±0.11

70 % C2H5OH 3.8±0.24 38.70±0.20 29.78±0.25 7.67±0.15

100 % C2H5OH 3.6±0.25 35.80±0.22 29.33±0.15 7.35±0.14

Peanut husk (PNH)

Water 2.6±0.15 14.20±0.16 8.88±0.10 5.65±0.08

50 % CH3OH 6.3±0.18 62.58±0.20 48.28±0.18 9.26±0.12

70 % CH3OH 6.2±0.28 56.35±0.22 38.85±0.21 8.35±0.18

100 % CH3OH 5.8±0.30 50.48±0.24 41.92±0.16 7.68±0.15

50 % C2H5OH 5.6±0.15 48.72±0.18 39.56±0.13 8.62±0.11

70 % C2H5OH 4.5±0.12 45.68±0.12 37.73±0.18 6.87±0.18

100 % C2H5OH 4.3±0.20 42.35±0.15 33.45±0.10 7.73±0.16

Coffee cherry husk (CCH)

Water 3.2±0.23 22.10±0.18 14.28±0.16 7.03±0.08

50 % CH3OH 9.2±0.25 85.25±0.16 72.56±0.12 10.15±0.10

70 % CH3OH 10.3±0.12 81.50±0.15 70.63±0.20 9.23±0.18

100 % CH3OH 9.7±0.13 80.44±0.21 71.36±0.24 9.45±0.13

50 % C2H5OH 9.3±0.22 81.38±0.11 65.44±0.08 8.67±0.17

70 % C2H5OH 8.5±0.20 77.72±0.15 62.23±0.13 8.45±0.15

100 % C2H5OH 8.3±0.18 72.64±0.20 58.45±0.18 7.78±0.12

Rice bran (RB)

Water 3.6±0.30 15.80±0.22 8.75±0.12 5.76±0.14

50 % CH3OH 6.2±0.25 46.72±0.21 36.56±0.14 7.75±0.13

70 % CH3OH 6.0±0.17 44.35±0.20 32.88±0.16 7.75±0.16

100 % CH3OH 5.4±0.24 40.65±0.15 30.45±0.18 7.42±0.19

50 % C2H5OH 5.6±0.10 42.22±0.13 31.65±0.21 6.54±0.22

70 % C2H5OH 4.8±0.20 38.36±0.18 30.47±0.23 6.24±0.10

100 % C2H5OH 4.3±0.11 33.53±0.20 28.32±0.18 6.16±0.15

Wheat bran (WB)

Water 3.3±0.16 16.80±0.15 9.84±0.10 6.57±0.23

50 % CH3OH 6.4±0.21 53.32±0.12 45.57±0. 13 7.42±0.18

70 % CH3OH 5.9±0.14 48.43±0.15 42.56±0.15 6.75±0.13

100 % CH3OH 5.7±0.20 45.56±0.10 38.23±0.22 6.24±0.20

50 % C2H5OH 5.2±0.15 46.73±0.20 38.12±0.20 5.86±0.08

70 % C2H5OH 4.7±0.28 43.81±0.15 37.54±0.16 6.31±0.12

100 % C2H5OH 4.0±0.22 40.12±0.20 33.35±0.17 5.86±0.16

Each value is an average of triplicate determination; ± standard deviation

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10.3 % obtained with 70 % methanol, while the lowest resultwas 3.3 % obtained using water as solvent. The yields of cornhusk (CH) were 2.5 % and 6.0 % corresponding to theextraction with water and 50 % methanol respectively(Table 1).

Organic solvents are commonly used for the extraction ofpolyphenols from plant material. The most important factorthat determines the recovery of polyphenols from plant mate-rials is the solubility of the phenolic compounds in the solventused for the extraction process. Ethanol, methanol and acetoneand their aqueous mixtures are commonly used for the extrac-tion purposes. Zhou and Yu (2006) used 50 % acetone/wateras an extraction solvent for the extraction of polyphenols fromColorado grown vegetables. Absolute methanol was used byAbas et al. (2006) and Wijngaard et al. (2009) for the extrac-tion of phenolic antioxidants from leafy vegetables and Irishfruit and vegetable wastes, respectively, whereas Sreeramuluand Raghunath (2010) used 60 % methanol with 0.1 % HClfor the extraction of polyphenols from vegetables and tubers.

Total polyphenols and tannins and flavonoids contentof the extracts

The total phenols, tannins and flavonoids content were widelyhigher for CCH extracts, when compared with other agricul-tural residues (SCB, CH, PNH, RB and WB) extracts, for alltype solvent extraction. The values for CCH were 22.10;85.50 and 81.38 g GAE/100 g of extract, for extraction withwater, 50 % methanol and 50 % ethanol respectively. Thecontent of total tannins in CCH extracts found to be 14.28;72.56 and 62.44 g TAE/100 g of extract and the total flavo-noids content was found to be 7.03; 10.15 and 8.67 g TFC/100 g for extraction with water, 50 % methanol and 50 %ethanol respectively. Among the studied solvents methanolicextract (50 %) exhibited significantly higher total phenoliccontent for all the agriculture residues while among them,coffee cherry husk had the highest phenolic content. Thisimplies that the phenolic compounds in the agriculture resi-dues might be readily soluble in aqueous methanol. Theamount and quantity of phenolic molecules by extraction insolvents depends on the plant materials, the solvent used(Marinova and Yanishlieva 1997; Moure et al. 2000), as wellas the contact time of extraction (Delgado et al. 2010). Awikaet al. (2005) employed aqueous acetone for phenol and anti-oxidant activity on sorghum bran.

The extraction yield and the antioxidant activity of theextracts from plants highly depend on the solvent polarity,which determines both qualitatively and quantitatively of theextracted antioxidant compounds. The highest yields are usu-ally achieved with ethanol and methanol and their mixtureswith water.

Several authors (Lou et al. 2004; Yu et al. 2005, 2006;2007; Wang et al. 2007) have reported that peanut skins

contain phenolic compounds with demonstrated antioxidantproperties. Yu et al. (2006) observed three classes of com-pounds in peanut skin extracts including phenolic acids, fla-vonoids and stilbene (resveratrol). Some authors (Yu et al.2005, 2006; Nepote et al. 2002, 2005) have employed tradi-tional solid–liquid extraction techniques using different organ-ic solvents to extract antioxidants from peanut skins. Nepoteet al. (2005) investigated the effects of several parameters onthe extraction of phenolic compounds from peanut skins usingsolid–liquid extraction. In that study, optimum extraction con-ditions were solely based on the quantity of total phenoliccompounds extracted as determined by Folin-Ciocalteu re-agent, and no identification of phenolics were reported.Wang et al. (2007) extracted phenolics from peanut skins bymaceration of the skins with 50 % (v/v) aqueous ethanol atroom temperature and reported a total phenolics content of90 mg/g of extract. More research is needed to developalternative extraction procedures and to obtain a more detailedprofile of the phenolic composition of peanut skin extracts.

Flavonoids are major group of polyphenols, which posses abasic C15 phenyl-benzopyrone skeleton modified with differ-ing numbers and positions of substituents, including hydroxyl,methoxyl and glycosyl groups. The concentration of flavonoidderivatives in the studied agriculture residues is presented inTable 1. Methanol was found to be optimum solvent for theextraction of flavonoids from agriculture residues.

HPLC analysis

Phenolic compounds from the various agricultural residueswere detected by HPLC analysis. The chromatogram of all thesamples showed two major peaks (1 and 2) along with threeminor peaks (3 and 5) (Fig. 1). The peaks (1 to 5) wereidentified as gallic acid, ferulic acid, epicatechin, quercitinand kampferol by comparing the retention time of the authen-tic standards.

Antioxidant assays

DPPH radical scavenging activity

The DPPH (1, 1- diphenyl-2-picrylhydrazyl) radical scaveng-ing activity of agricultural residues is shown in (Fig. 2). Thisactivity was found to increase with increasing concentration of50 % methanolic extract. It is well known that the antioxidantactivity of plant extracts containing polyphenol components isdue to their capacity to donate hydrogen atoms or electronsand scavenge free radicals (Sreeramulu and Raghunath 2010).Among the samples SCB showed the least scavenging activity(IC50 = 26.3μg/ml) while PNH (IC50 = 22.6μg/ml), CH (IC50 =24.25 μg/ml), CCH (IC50 = 24.3 μg/ml) and WB (IC50 =25.1 μg/ml) showed a better result than the positive controlascorbic acid (IC50 = 38.0 μg/ml). The results are summarized

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in Table 2. These suggested that SCB has an efficient radicalscavenger, its activity being confined to the hydroxymethanolicfraction.

Agricultural residues have a great potential as source ofantioxidants, many of which are polyphenols. Solvent andprocess variables must be carefully chosen to optimize theirextraction. Antioxidant activity of every extract must be mea-sured by several methods: radical scavenging activities(DPPH), oxidation of lipids (TBARS), micellar systems,etc., because none of them is representative of all real system;for example it is possible to obtain extracts with high radicalscavenging activity but unable to protect an oil for oxidationbecause of its low miscibility.

The antioxidant activity of plant extracts can be in largepart attributed to the presence of polyphenolic compounds

located within the plant tissues. Polyphenols are attracting agreat deal of attention due to evidence suggesting that anincrease in their consumption in the diet may prevent cancer,strokes and neurological diseases. They are the most abundantantioxidants in our diets and it is estimated that we consumeabout 1 g of polyphenols per day (Scalbert and Williamson2000). Several thousands of natural polyphenols have beenidentified in plants and plant foods. Polyphenolic compoundsare present in high concentrations in a variety of fruits, vege-tables and beverages such as tea and wine. They are alsoabundant in agricultural byproducts such as peanut skins,hulls and roots, grape seeds and skins and in a number ofherbs and spices (rosemary, sage, thyme and oregano).Polyphenols are important to plant growth and developmentand provide a defense mechanism against infection and injury

Fig. 1 HPLC profile of hydromethanolic extract (50 %) of agricultureresidues, a sugarcane bagasse (SCB), b corn husk (CH), c peanut husk(PNH), d coffee cherry husk (CCH), e rice bran (RB) and f wheat bran

(WB). Peak 1: gallic acid; peak 2: ferulic acid; peak 3: epicatechin; peak4: quercitin and peak 5: kampferol

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(Karakaya and Tas 2001). Many polyphenolic compoundshave been found to have a much stronger antioxidant activitythan vitamins C and E and β-carotene within the same food(Chu et al. 2002).

Nitric oxide radical scavenging activity

Nitric oxide (NO) has an important role in various inflamma-tory processes. Its sustained production is toxic to tissues andcan contribute to the vascular collapse associated with septicshock. Chronic generation of nitric oxide is associated withcarcinomas and various inflammatory conditions includingjuvenile diabetes, multiple sclerosis, arthritis, and ulcerativecolitis (Tylor et al. 1997). Nitric oxide is generated from thedecomposition of SNP and measured. SNP in aqueous solu-tion at physiological pH spontaneously generates NO, whichinteracts with oxygen to produce nitrite ions that can bemeasured. All the extracts for both these methods analyzed

at the same range of concentration (50–500 μg/ml). A signif-icant decrease in the NO radical is due to the scavengingactivity of the extracts. Our results showed dose-dependentNO scavenging activity by all the test samples (Fig. 3). IC50

values (Table 2) were calculated from the graphs plottedscavenging activity versus the concentrations of samples.The IC50 values of SCB (27.0 μg/ml), CH (27.5 μg/ml),PNH (28.0 μg/ml), CCH (29.0 μg/ml), RB (30.0 μg/ml) andWB (31.0 μg/ml) for scavenging NO revealed that SCB andCH were more potent than the positive control, ascorbic acid(IC50 = 38.0μg/ml). in this case also, SCBwas the most activeas compare to other agricultural residues.

Table 2 Antioxidant activity (IC50 values) of the agricultural residuesextracted in 50 % methanol

Agricultural residues DPPH assay NO assay

Ascorbic acida 39.0±0.02 38.0±0.08

SCB 26.3±0.02 27.0±0.01

CH 24.25±0.05 27.5±0.04

PNH 22.6±0.02 28.0±0.07

CCH 24.3±0.01 29.0±0.01

RB 24.3±0.06 30.0±0.03

WB 25.1±0.03 31.0±0.05

Each value is an average of triplicate determination; ± standard deviationa Standard drug

0

20

40

60

80

100

0 50 100 150 200 250 300

% S

cave

ngin

g

Concentration (µg/ml)

Ascorbic acid SCB CH

GNH CCH RB

WB

Fig. 3 Nitric oxide scavenging activity of agricultural residues withhydromethanol extracts (50 %). Values are the average of triplicateexperiments and represented as mean ± SD

0

0.2

0.4

0.6

0.8

1

0 50 100 150 200

Abs

orba

nce

Concentration (µg/ml)

PNH SCB CHCCH RB WBAscorbic acid

Fig. 4 Ferrous ion reducing activity of agricultural residues withhydromethanol extracts (50 %). Values are the average of triplicateexperiments and represented as mean ± SD

0

20

40

60

80

100

0 100 200 300 400 500

% S

cave

ngin

g

Concentration ((µg/ml)

Ascorbic acid SCBCH GNHCCH RBWB

Fig. 2 DPPH radical scavenging activity of agricultural residues withhydromethanol extracts (50 %). Values are the average of triplicateexperiments and represented as mean ± SD

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Ferric ion reducing assay

The reducing power of a compound may serve as a significantindicator of its potential antioxidant activity (Meir et al. 1995).Hence, the Fe3+ reducing powers of agricultural residues aswell as its various fractions were investigated, and the resultscompared with that of the reference compound, Ascorbic acid.The reducing powers of the samples were found to increaseconcentration dependently (Fig. 4). The RP 0.5 AU wascalculated and depicted in Table 3. In this assay, a higherabsorbance of reaction mixture indicates higher reducingpower of the sample. The absorbances of assay mixturecontaining a fixed concentration (200 μg/ml) of SCB, CH,PNH, CCH, RB,WB and ascorbic acid were found to be 0.39,0.28, 0.44, 0.36,0.35, 0.33 and 0.47 respectively (Table 3).The order of the reducing powers of the samples wasCH>WB>RB>CCH>SCB>PNH>ascorbic acid. Thus, allsamples showed better reducing powers than ascorbic acid.

The antioxidant properties of phenolic compounds areassociated with their reducing power (Jayaprakasha et al.2001), which is associated with the presence of reductones(Duh 1998). The reducing power of agricultural residuesincreases significantly with phenol content (Fig. 4 & Table 3).

Conclusion

Agricultural residues have great potential and cost effectivesources of antioxidants, many of which are polyphenols. Toobtain the maximum yield of extraction, 50 % methanol isrecommended for phenolic compounds extraction from agri-cultural residues. The beneficial effects of polyphenols havebeen ascribed to their strong antioxidant activity that is, theirability to scavenge oxygen radicals and other reactive species.These features make phenols a potentially interesting materialfor the development of functional foods or possible therapyfor the prevention of some diseases.

Acknowledgments This research was supported by research grants toKS from DST and UGC-SAP, the Government of India, New Delhi. VSthanks UGC-SAP for providing JRF.

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CH 0.28±0.08

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CCH 0.36±0.03

RB 0.35±0.07

WB 0.33±0.02

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