in vitro antioxidant and antidiabetic activities of biomodified lignin from acacia nilotica wood...

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International Journal of Biological Macromolecules 75 (2015) 81–89

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International Journal of Biological Macromolecules

j ournal homepage: www.elsevier .com/ locate / i jb iomac

In vitro antioxidant and antidiabetic activities of biomodified ligninfrom Acacia nilotica wood

Anand Barapatre, Keshaw Ram Aadil, Bhupendra Nath Tiwary, Harit Jha∗

Department of Biotechnology, GuruGhasidas Vishwavidyalaya (A Central University), Bilaspur, Chhattisgarh 495009, India

a r t i c l e i n f o

Article history:

Received 5 June 2014

Received in revised form26 November 2014

Accepted 8 January 2015Available online 16 January 2015

Keywords:

Acacia lignin

Biotransformation

Antioxidant

Antidiabetic-Amylase

In vitro glucose movement

a b s t r a c t

The antioxidant and antidiabetic activity of biomodified alkali lignin extracted from a deciduous plant Acacia nilotica, was evaluated in vitro. The extracted alkali lignin was subjected to microbial biotransfor-mation by ligninolytic fungus Aspergillus flavus and Emericella nidulans. These modifications were done

under varying concentration of carbon to nitrogen sources. The structural feature of the lignin sampleswere compared by FTIR, functional group analysis and 13C solid state NMR. All lignin samples were tested

for antioxidantefficiency,reducing power and H2O2 scavenging power. Modifications in alllignin samplesshowed correlation with their antioxidant scavenging activity and reducing power. Antidiabetic prop-

erties were evaluated in terms of in vitro glucose movement inhibition and -amylase inhibition assay.Modified samples exhibited increased glucose binding efficiency as demonstrated by the decreased glu-

cose diffusion (55.5–76.3%) and 1.16–1.18-fold enhanced -amylase inhibition in comparison to theircontrol samples. The results obtained demonstrate that the structure and functional modifications in

lignin significantly affects its bioefficacy in term of antioxidant and antidiabetic activities.

© 2015 Elsevier B.V. All rights reserved.

1. Introduction

Plant phenolics and polyphenols have been increasingly entic-

ing the attraction due to their beneficial therapeutic valuesincluding antioxidant, antimicrobial, anti-inflammatory, cardio-protective, anticancerous, chemo-preventive and neuro-protectiveproperties. Polyphenols have been considered, a health food

supplement and are claimed to possess health promoting ordisease-preventing properties [1]. Lignin, the second most abun-dant natural macromolecule (polyphenolic in nature and 10–35%ofdry wt.of lignocellulosicbiomass), is a natural polymerized prod-

uct of optically active p-hydroxycinnamyl alcohol monomers andrelated monolignols (p-coumaryl, coniferyl,and sinapyl) formed byoxidative reactions. This polymer is the result of various inter unitlinkages in the monomer and monolignols (e.g. -O-4, -5, -,biphenyl (5-5), 4-O-5) [2]. The precise chemical structure of ligninis not known because of its complex polymeric nature and due tomany random coupling. Alkali lignin is currently the largest pro-duced among all lignin classes and a less valuable co-product of

biofuel and paper pulp industries, which is separated from fibersby a chemical pulping (mainly soda and sulphite) process [3].

∗ Corresponding author. Tel.: +91 9826630805.

E-mail address: [email protected](H. Jha).

Acacia nilotica, locally known as “Babul”, is a multipurposedeciduous tree of Mimosaseae family predominantly found in cen-tral India. This plant contains a variety of bioactive components

such as ellagic acid, isoquercitin, leucocyanadin, kaempferol-7-diglucoside, derivatives of (+)-catechin, apigenin derivatives etc.[4]. Traditionally in the central region of India leaves, pod, barkand root of A. nilotica is used for the treatment of various dis-

eases related to oral, bone and skin, like cold, bronchitis, diarrhoea,dysentery, biliousness, bleeding piles and Leucoderma [5].

Oxidative damages creates by free radicals, play a substantialrole in the evolution of human diseases. Toxicity of free radicals

contributes to proteins and DNA damage, inflammation, tissueinjury and subsequent cellular apoptosis, which finally leads tocancer, emphysema, cirrhosis, arteriosclerosis and arthritis [6].Oxidative stress is created in the body due to a disruption in the

equilibrium between the production of reactive oxygen/nitrogenspecies (ROS/RNS)and the removal via the antioxidantdefence sys-tem [7]. Antioxidants can interfere with the oxidative processesby reacting with free radicals, chelating catalytic metals and also

by acting as oxygen scavengers thus helping the human body toreduce oxidative damage. From various epidemiological studies,it is proved that polyphenolic compounds possess an excellentantioxidant properties [6,8]. Previous studies also reported that

the polyphenols found in Acacia sp. plants having good antioxi-dant power [4,5,9,10]. As a complex phenolic polymer, lignin alsopossess a respectable medicinal properties [2,3,11].

http://dx.doi.org/10.1016/j.ijbiomac.2015.01.012

0141-8130/© 2015 Elsevier B.V. All rights reserved.

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82 A. Barapatre et al. / International Journal of Biological Macromolecules 75 (2015) 81–89

Over the last century changes in human behavior and life style

have resulted in a dramatic increase in the incidence of diabetesworld over. Presently, it is estimated that more than 220 mil-lion suffer from diabetes in which 90% is from type 2 diabetesType 2 diabetes is the results of ineffectiveness of insulin and

the primary cause of complications linked to cardiovascular dis-ease, renal failure, blindness, neurological complications, and soon [12]. Currently available conventional therapies for the treat-ment of diabetes include insulin and oral antidiabetic agents such

as sulfonylurea, biguanides, and alpha-glucosidase inhibitors. Tra-ditionally, many activecompoundsof plantorigin including several

Acacia sp. plants, have been also employed in the treatment of diabetes, mostly the secondary metabolites including alkaloids,

flavonoids, phenolics, steroids, carbohydrates, glycopeptides, ter-penoids etc. [13]. The inhibition of enzyme like -amylase as wellas the delay in glucose absorption to be an important strategy inthe management of blood glucose level in type 2 diabetic [14].

Based on the fact that structural heterogeneity will affect thechemical properties of lignin, we report the bioefficacy of biolog-ically modified lignin in terms of the antioxidant, antiradical andhydrogen peroxidase scavenging property and also evaluate the

effectiveness ofmodified ligninas aninhibitorof -amylase activityand in vitro glucose movement.

2. Materials and methods

2.1. Chemical and reagents

Gallic acid, d-glucose, -amylase (EC 3.2.1.1), 1,1-diphenyl-2-picrylhydrazyl (DPPH), neocuproine(Nc), catalase frombovine liver(966Umg−1) were purchased from Sigma–Aldrich Inc. (Mumbai,

India). All other chemicals and reagents used were of high purityanalytical grade and purchased from Merck Pvt. Ltd. (India). Ultra-pure water (Elix, Merck Milipore, India) was used throughout theexperiment. Wood dust (18 mesh size) of A. nilotica hardwood was

procured locally from the saw mill of Bilaspur, Chhattisgarh, India.

2.2. Extraction and characterization of alkali lignin

Alkali extraction of lignin was achieved by treatment of wooddust with an aqueous solution of NaOH (1.2%, w/v) in a 1 L glassflask for 1 h at 120 ◦C, using a solid/liquid ratio of 1:10 (g/mL).The solution was filtered through Whatman filter paper No. 4 to

remove wood dust. The filtrate (black liquor) was concentrated byslow heating at 60◦C in an oven and acidified up to pH 5.5 with6 M HCl. Thewater-soluble hemicellulosic fraction was removed byprecipitation, after adding two volumes of 95% ethanol (v/v). The

precipitated hemicellulosic were removed by gravity filtration. Theremaining filtrate was concentrated to 20–30mL, and the pH wasadjusted to 1.5–2.0 with 6 M HCl. The alkali lignin was precipitatedand sediments by centrifugation at 10,000 rpm (Remi R-24, India)

for 10min. The pellet was dried and stored at room temperaturefor further study [15].

2.3. Biodegradation and characterization of alkali lignin

2.3.1. Microorganism (fungus)

The two potent ligninolytic fungus A. flavus and E. nidulanswere used for the biotransformation of alkali lignin. The strainswere isolated from soil samples collected from Guru Ghasidas

Vishwavidyalaya campus, Bilaspur (C.G.) and near the effluentdischarge site of the Orient paper mill situated in Amalai (M.P.),India. Two potentially ligninolytic strains were characterized andidentified based on morphological characterization and partial

gene sequencing of Internal Transcribed Spacer (ITS) regions as

A. flavus (F10, NCBI accession no. KC911631.1) and E. nidulans

(APF4, NCBI accession no. KC911632.1) respectively. The strains

were maintained on malt agar slants for further use.

2.3.2. Basal culture medium

Biodegradation of lignin was performed under different carbon

to nitrogen ratio i.e. low and high in Basal Salt Medium (BSM).The BSM contained gL −1 of KH2PO4, 0.2 g ; MgSO4·7H2O, 0.05g;CaC12, 0.01g supplemented with a 1 mL mineral solution.The min-eralsolutioncontained (in gL −1) nitrilotriacetate,1.5; MgSO

4·7H

2O,

3.0; MnSO4·H2O, 0.5; NaCl, 1.0; FeSO4.7H2O, 0.1; CoSO4, 0.1;CaC12, 0.082; ZnSO4, 0.1; CuSO4·5H2O, 0.01; AlK(SO4)2, 0.01;H3BO3, 0.010; NaMoO4, 0.01. The high carbon low nitrogen (HCLN)medium contained56 mMd-glucose asa carbonsource and2.4 mM

nitrogen (0.6 mM NH4NO3 and 0.6 mM l-asparagine) whereas thelow carbon high nitrogen (HNLC) medium contained 8.8 mM d-glucose was and 24mM nitrogen (NH4NO3 and l-asparagine, 6 mMeach).The pH of BSMwas maintained at 5.6–5.8for biomodification

of alkali lignin [16].

2.3.3. Biomodification and characterization of alkali lignin

Two hundred mililiter of HCLN and HNLC BSM medium main-

tained at pH 5.6–5.8 were transferred to 500mL erlenmeyer flasks.The autoclaved media were aseptically inoculated with 3 bores

(6mm diameter) of 7 days old culture of the strains F10 and APF4andincubatedat 28 ◦C for21 days under staticcondition. Thefungal

mat was separated fromthe mediumby filtration through sterilizedWhatmanfilterpaper No 4. Themodified ligninwas recoveredfromfiltrate by the same method as described in Section 2.2. Uninocu-lated media were used as negative control.

Biotransformed alkali lignin was characterized by FTIR. Asmall fraction of the sample was ground properly with equalamount of KBr. The FTIR spectrum was obtained in the range of 400–4000cm−1 using FTIR spectrophotometer Affinity A1, (Shi-

madzu, Japan).A high-resolution 1D solid-state 13CNMR spectra were obtained

with the Cross-Polarization Magic-Angle Spinning (CPMAS)technique on a Bruker spectrometer (Sophisticated Analytical

Instrument Facility (SAIF), Indian Institute of Science, Bangalore,India) operating at 100.525 MHz frequency for the 13C carbonnuclei. A total of300 mg solid sample was used for the NMR spectraat 294 K.The 2000–6000 scanwas performed to obtain a 1D spectra

with a 29.10 ms acquisition time and 5 s relaxation delay.

2.4. Total polyphenol content (TPC)

Total polyphenol content (TPC) was determined by reactionwith Folin-Denis reagent [17]. One mL of each lignin sample(50g/mL) was added to 0.5 mL of Folin-Denis reagent. After 30s,1 mL of 20%(w/v) sodiumcarbonate wasadded andthe volumewas

madeup to 5mL with distilled water. The mixture was allowed tostand at room temperature for 10min. The absorbance of result-

ing blue complex was measured at 765 nm against blank usingUV–visible double beam spectrophotometer (Shimatzu UV-1800,

Japan). A calibration curve of gallic acid was prepared, and pheno-liccontentswere determinedfrom thelinear regression equation of this curve. The results were expressed asg gallic acid equivalents(GAE) per milligram of dry material.

2.5. Functional group analysis

2.5.1. Phenolic hydroxyl groups by ultraviolet-spectroscopy

The content of hydroxyl phenolic units in lignin fractions wasdetermined by UV spectroscopy as described Aadil et al. [9]. Thismethod is based on the difference in absorption of lignin samples

atpH 6 (495mL of 0.2N potassium dihydrogen phosphate solution

mixed with 113 mL of 0.1 N NaOH and diluted to 2 L with distilled


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A. Barapatre et al. / International Journal of Biological Macromolecules 75 (2015) 81–89 83

water) and at alkaline buffer solution pH 12 (0.1N of boric acid in

0.1 N NaOH solution). The difference in spectra was obtained bytaking the absorbance of the alkaline solution relative to that of the neutralized solution in the range of 200–400 nm. The pheno-lic hydroxyl group content of lignin samples was calculated using

amax.

% phenolic hydroxyl =amax × 17

41

2.5.2. Carboxyl groups determination by aqueous titration

method

An amount of25 mg of recovered lignin samples wassuspendedin 25mL of an alkaline 0.1 N NaOH solution and stirring for 3 h. The

pHwasadjustedto12with0.1NNaOH.Followedbypotentiometrictitration with 0.1 N HCl, as described Aadil et al. [9].

2.6. Total reducing power assay

Reducing the power of allsamples wasdetermined according tothemethod of Oyaizu[18]. Briefly, different concentrations of lignin

samples were prepared in sodium phosphate buffer (200 mM, pH6.6) and 2.5 mL of each sample was separately mixed with 2.5 mL

of 1% (w/v) potassium ferricyanide. The mixture was incubated at50 ◦C for 20min. A mixture containing all the reaction reagents

except the test material serves as the control. The reaction wasstopped by adding 2.5 mL of 10% (w/v) TCA and the mixture wascentrifuged at 1750rpm for 10min. The upper layer (2mL) wasmixedwith1mL of0.1%(w/v)offerricchlorideandmadeupto8mL

with deionized water. The perl’s prussian blue color formed due toreduction in Fe3+ was measured at A700. The EC50 of extracts werecalculated from the graph of A700 versus extracts concentration.

2.7. DPPH free radical scavenging activity

The antiradical activity of lignin samples was measured basedon their reaction with stable free radical DPPH* and subsequent

reduction in max of DPPH* [19]. In this reaction 1.5mL sample,(100g/mL, prepared in 50mM phosphate buffer, pH 7.5) wasallowed to react with 1.5mL of 100M methanolic solution of DPPH* for 30min in darkness at room temperature. The decrease in

absorbance was measured at A515 . DPPH*radical scavenging capac-ity was calculated using the following equation

DPPH ∗ scavenging (%) =

( A0 − A1)

A0× 100

where A0 and A1 are absorbance of DPPH* radical at 515 nm in theabsence and presence of the samples.

2.8. Hydrogen peroxide scavenging (HPS) assay

HPS capacity of lignin samples was estimated by cupric reduc-ing antioxidant capacity method according to Ozyurek et al. [20].In this method, hydrogen peroxide incubation solution and scav-enger solutions were prepared. The hydrogen peroxide incubationsolution (used as a reference) contained 0.7mL of phosphate buffer

(0.2M, pH7.4), 0.4 mL of1mM H2O2,0.4mL of 0.1 mM CuCl2.2H2O,whereas scavenger solutions (I and II) were prepared in twotest tubes containing 0.2 mL of test samples, 0.5 mL of phosphatebuffer (0.2M, pH 7.4), 0.4 mL of 1 mM H2O2 and 0.4 mL of 0.1 mM

CuCl2.2H2O (identicalup to this step). Themixtures were incubatedfor 30min at 37 ◦C in water bath. After incubation 0.4 mL of H2Owas added to the reference and scavenger solution-I and 0.4 mL of catalase solution (268U mL −1) in scavenger solution-II. The mix-

ture wasvortexesfor 30 s. From theaboveincubated mixtures,1 mL

solution was mixed with 1 mL of Nc (0.0075M, freshly prepared

in ethanol), 1 mL of 0.1mM CuCl2·2H2O and 2mL of ammonium

acetate buffer (1M, pH 7). After 30min, the absorbance of the finalsolution was taken at 450 nm against the reagent blank. The HPSactivity (%) of samples was calculated using the following formula:

HPS(%) =[ A0 − { A1 − A2}]

A0× 100

where A0 is the absorbance of reference hydrogen peroxide incuba-

tion solution, A1 and A2 are the absorbance of scavenger solution-Iand -II, respectively.

2.9. Antidiabetic assay by in vitro glucose movement

To evaluate the effects of biotransformed lignin on glucosemovement an in vitromodel system was used according to Büyük-balci and Nehir [21] with slight modification. The dialysis tube(6cm×14.3 mm) (HiMedia, Mumbai, India; pore size 2.4 nm) was

filled with a total volume of 6 mL test sample mixtures containing1 mg/mL biotransformed lignin and 1.65 mM d-glucose (preparedin 0.15M NaCl) in theratio of 2:1 (v/v). The dialysis tube was sealedat both ends and placed in a flask containing 45mL 0.15M NaCl.

Dialysis experiment was performedon an orbital shakerwaterbath

(100rpm) at 37◦C for 3h to induce the movement of glucose intothe external solution. Concentration of glucose outside the dialy-sis tubing was measured by DNS (dinitrosaliacylic acid) reagent.

The control experiment was conducted in the absence of the testsample.

2.10. In vitro ˛-amylase inhibition assay

The -amylase inhibitory activity was determined by themethod of Quesille-Villalobos et al. [22]. A total of 500L of dif-ferent concentrations of each sample (0.1, 0.5, 1 and 5 mg/mL) and500L of 0.02M sodium phosphate buffer (pH 6.9 with 0.006MNaCl) containing-amylase (0.5mg/mL) was incubated for 10minat 25 ◦C. After preincubation, 500L of 1% (w/v) starch solution in

0.02M sodium phosphate buffer (pH 6.9 with 0.006 M NaCl) wasadded to each of the pre-incubated tubes. The reaction mixtures

were then incubated at 25◦C for 10 m in and stopped with 1mL of DNS reagent. The test tubes were further incubated in a boilingwaterbathfor10min and cooledto roomtemperature. The reactionmixture was dilutedwith10 mL distilled water and the absorbance

was measured at 540nm. The absorbance of blank samples (bufferinstead of enzyme solution) and a control (buffer in place of thesample extract) was also recorded forcomparison.The final activityof -amylase was calculated by subtracting the final A540 of samplewith its corresponding A540 blank.

2.11. Statistical analysis

Unless otherwise stated experiments were performed in tripli-cate and statistical analysis was done in term of mean± standarddeviation (SD). Significance levels were calculated using Graph PadPrism 5.0 by one way analysis of variance (ANOVA) followed by

Tukey’s multiple comparison test.

3. Results and discussion

The biotransformation and biodegradation of lignin occur

in a multistep process involving Lignin Peroxidase (LiP),Manganese–dependent Peroxidase (MnP), Laccase, VersatilePeroxidases (VPs) and dioxygenases enzymes, working in asso-ciation with small molecules and radicals [23]. The production

of these enzymes is associated with nutrient stress conditions.

White rot basidiomycetes and brown rot ligninolytic fungi display


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a broad diversity in response to carbon and nitrogen source and

their C/Nratio. In most of theligninolyticfungi LiP, MnPand laccaseproduction is primarily regulated by the nitrogen concentration[24,25]. Biodegradation of lignin starts with depolymerizingactivity of LiP and MnP followed by demethylation activity of

laccase. It is reported that the side chain and aromatic rings of lignin model compounds and synthetic lignin (DHPs) were cleaved

via aryl cation radical and phenoxy radical intermediates which ismediated by LiP/H

2O

2 and laccase/O

2/mediator system. Whereas

MnP catalyze demethylation, C-C cleavage, alkyl-aryl cleavageand C oxidation of phenolic syringyl type -1 and -O-4 ligninstructures [26].

In present study biomodified lignin was obtained by biomodifi-

cation of lignin using two ligninolytic fungi F10 and APF4. Duringmicrobial transformation of lignin by F10 and APF4, the onset of MnP and LiP activity in HCLN medium starts from 3rd day and con-tinuously increase up to the 21st day, whereas in HNLC medium it

starts from 3rd in F10 and 12th day and APF4. The activity of bothenzymes is minimal in HNLC as compared to HCLN medium. On theother hand, thelaccase activity washigh in HCLN medium,whereasin HNLC mediumit wasvery low(unpublished data). After biomod-

ification recovered biomodified lignin samples were evaluated fortheirTPC, functionalgroup analysis, reducing power, hydrogen per-

oxide scavenging capacity, DPPH free-radical scavenging capacity,-amylase and in vitro glucose movement inhibition activity.

3.1. Characterization of modified lignin

3.1.1. FTIR of lignin samples

TheFTIR spectrum ofthe control andbiomodified liningsamplesis presented in Fig. 1a and b. All modified lignin samples (B, C, E andF) showed a broad absorption band at 3410–3460cm−1, attributedto the O–H groups stretching in phenolic and aliphatic structures

and oscillation of the hydroxyl group. The relative intensity of thisband stretching was more in the treated sample (both HCLN F10and HCLN APF4) as compared to control sample, which indicatedthe phenolic ring modification by addition of a hydroxyl group.

Whereas in modified HNLC samples (HNLC F10 and HNLC APF4),the intensity of O–H groups stretching was almost same in com-parison to modified HCLN samples. A strong bands at 2847cm−1

arising from C–H stretching in the aliphatic methylene group was

less intense in treated sample (B, C, E and F) as compared to theircontrol samples (A and D). Bands centered on 2938 and 2842cm−1,predominantly arising from C–H stretching in aromatic methoxylgroups and in methyl and methylene groups of side chains. The

intensity of both peaks is reduced in treated samples as comparedto control samples. The increase in the intensity of –OH group anddecrease in C–H stretching also an outcome of laccase activity assuggested in the literature [27,28].

All samples displayed weak bands in the carbonyl/carboxylregion 1705–1720 cm−1, assigned for unconjugated ketone or

unconjugated carbonyl stretching. The intensity of aromatic skele-ton vibrations at 1600, 1515 and 1426cm−1 characteristic of the

aromatic ring in alkaliligninshowed significant decrease in treatedsamples as compared to controls,indicating thecleavageof thearo-maticring structure. The C–H deformationcombined with aromatic

ring vibration at 1462cm−1 was observed in the untreated con-trol samples, however, these peaks were absent in treated samples.These results suggested that the structural and functional groupsof alkali lignin were altered by both the fungal strains [29].

3.1.2. 13C NMR

A comparison of the 1 3C NMR spectra of an untreated controllignin samples with biodegraded lignin sampleunder two different

nutritionalconditionsis presented in Fig.2. InallthreeNMR spectra

of lignin, there was an absence of signals between 90 and 102ppm

which indicates that the samples were free of carbohydrate

contamination. The signals for unconjugated carboxylic acids–COOH (178.0–167.5 ppm) were high in HNLC sample in compar-ison to control while low in HCLN sample. The relative increase inthis signal could be attributed to the formation of aldehydes, acids

and aroxiacetic structures probably because of the oxidation of theside-chains. These results suggest oxidative attack on the lignin bythe microbial enzymatic system of fungus [30].

The signal for the aliphatic (171–168.5ppm) and phenolic

(168.5–167ppm) hydroxyl groups were observed in the ligninsamples treated with fungus (HCLN and HNLC) whereas they areabsent in control lignin. The three aromatic region signals for pro-tonated aromatic speciallyunsubstituted aromatic carbons orthoor

para to the substituted carbon (125–103ppm), the condensed aro-matic mainly C-substituted aromatic carbon (141–125ppm), andthe oxygenated aromatic mainly O-substituted aromatic carbon of guaiacol (160–141ppm) were higher in control lignin as compared

to treated lignin [31].Theregion between162 to 103ppm belongs to aromatic carbon,

a syringal units produce strong signal at 153–151 ppm (C-3 ester-ified) whereas the guaicyl unit signals at 119 ppm (C-6), 115 ppm

(C-5) and 111ppm (C-2) were also observed in control and treatedsamples. The changes in the aromatic-C region was observed,

notably the decrease in the syringyl and guaiacyl amount (signalsat 153ppm and 148 ppm respectively) after the fungal degradation

of the alkali-lignin. The relative decrease in syringyl units reflectsan easier accessibility of the microorganisms to the less condensedsyringyl units of the polymer [30,31].

The signals at 90–57.5 ppm display aliphatic C–O bonds and

, , carbons on the lignin side chain. In which C, C, andC in -O-4 can be identified in the regions 79.0–67.0, 90.0–78.0,and 61.5–57.5 ppm, respectively [31]. A significant decrease wasobserved in C in -O-4 (61.8ppm) both treated samples as com-pare to control, while the other two signals for C and C in-O-4 increased in HNLC sample as compared to control. Somenew signal intensity (30.5, 28.98 and 26.08) were also observedin the 46–10 p pm spectral region, mainly in the case of HNLC

lignin sample, suggesting accumulation of saturated alkyl struc-tures (Aliphatic CH2, 35.9ppm) in the biomodified lignin. One of the signal at 35.9 corresponding to C in arylpropanol unit wasdecreased in HCLN sample in comparison to control while it was

unaffected in HNLC lignin sample [30].

3.2. Total phenolic content and functional group analysis

All treated samples exhibited significantly lower TPC as com-pared to their respective controls (Table 1). It was detected that inF10and APF4 modified lignin samples, thequantity of TPC was lowunder HCLN than HNLC condition.In lignindegradationwhenlignin

is exposed to peroxidases, it undergoes decomposition into lowermolecular weight fragments containing methoxyl groups. Laccase

demethylates these fragments and peroxidases further degradethem into smaller fragments. These smaller fragments reduce into

their respective phenols by MnP, LiP and VPs which undergoes ringcleavage and form keto acids. The keto acids enter through kreb’scycleand aremetabolizedby thefungus [32]. VariationsinTPCwere

observed possibly due to difference in the activity of ligninolyticenzymes under varying nutrient conditions. The results obtainedrevealed that the TPC of samples was influenced by lignin degra-dation under carbon and nitrogen surplus/stress conditions. Under

HCLN condition enzymatic activity was high due to which lignindepolymerized into smaller fragments and subsequently metabo-lized by the fungus. Previous results also supported our contentionthat the ligninolytic enzymes activities of different fungi were

low/diminished in nitrogen rich medium as compare to nitrogen

deficient medium [16,23].


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Fig. 2. 13C CPMAS NMR spectra of thelignin samples.(A) Control lignin; (B)HCLN lignin; (C)HNLC lignin.

3.3. Reducing power assay

A reducing agent contributes to antioxidant activity by donat-

ing its electron to free radicals, which result in neutralization of the reactivity of the radical, and the reduced species subsequently

acquire a proton from the solution. It was previously reportedthat the Acacia species contain a number of active phenol andpolyphenolic contents which were implied in the reducing reac-

tions [9,10,33,34]. Reducing power of all samples was expressedin the form of EC50 value (Table 1), which ranged from 405.41 to


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Table 1

Total polyphenol content, phenolic hydroxyl content (%), carboxyl group (%, w/w), reducing power EC50, H2O2 scavenging (%) and DPPH scavenging activity (%) of control

and biomodified lignin samples.

Samples Polyphenol

contentaPhenolic hydroxyl

group (%)

Carboxyl group

(% w/w)

Reducing power

EC50 (in g)bH2O2scavenging (%)c

DPPH

scavenging (%)c

HCLN Control 362.30±8.42 1.24±0.01 8.89 ± 0.16 462.96 ± 14.28 30.09±3.64 56.27±0.33

HCLN F10 235.78±12.14** 1.19±0.01** 9.83 ± 0.13** 405.41 ± 4.38*** 55.54±5.36*** 71.44±0.93 ***HCLN APF4 277.54±6.30*** 1.26±0.01** 8.24 ± 0.13** 535.71 ± 11.47*** 54.29±5.29*** 63.27±2.35 ***

HNLC Control 427.50±10.46 1.09±0.02 10.13 ± 0.13 441.18 ± 5.19 63.56±0.16 55.33±0.89

HNLC F10 320.64±21.14*** 1.70±0.03** 10.06 ± 0.13* 842.69 ± 28.41** 57.17±3.07*** 37.94±0.64*HNLC APF4 197.10±8.16** 1.62±0.02** 9.6 ± 0.12** 1056.34 ± 14.87** 56.07±2.63*** 42.07±1.56*

Allvalues aremean±SD (n=3).

Mean±SD, significantly different from their respective control at *** p< 0.001, ** p


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Table 2

Concentration of d-glucose(in g)outside thedialysis tubeafter3 h in vitroglucose

movement test.

Samples Concentration of

d-glucose (ing/mL)

Decrease of

movement (%)a

Blank (in absence of sample) 72.77±2.25 –

HCLN Control 71.19±8.47 NDHCLN F10 17.26±9.10* 76.3%*

HCLN APF4 32.39±15.39* 55.5%*

HNLC Control 67.43±1.96 NDHNLC F10 23.21±11.26* 68.1%*

HNLC APF4 32.22±9.49* 55.7%*

Allvalues aremean±SD (n=3).

ND – notdetectable.* Significantly different, p60% inhibition) inhibitor, whereas others inhibit

the glucose diffusion in the range of 6–48%. They also reported thatthe plant extracts exhibited a concentration dependent inhibitoryeffect on glucose movement. While in another study Büyükbalciand Nehir [21] f ound that there was no significant effect of herbal

teas and infusions (traditionally used in the treatment of diabetesin Turkey) on in vitro glucose diffusion.

In the present study, control and modified lignin samples weretested for inhibitory effects on glucose movementout of dialysis

tube. No significant inhibition in glucose movement was observedby both control lignin samples as compared to blank (Table 2).But HCLN F10 and HNLC F10 lignin samples appeared to be themost potent inhibitorin glucose movement out of the dialysis tube,

decreasing movement upto 76.3% and 68.1% ( p< 0.001) respec-tively, in comparison to the blank. The other two lignin samples,HCLN APF4 and HNLC APF4, inhibited glucose movement nearly55% ( p< 0.001, as compared to blank). Our results provide ample

evidence that functional groups modification might be the possi-ble cause of increased glucose movement inhibiting properties of samples.

To date, no systematic scientific studies on the inhibitory effect

of lignin on in vitro glucose movement is available. We hypothesizethat the inhibition of glucose movement might occur by formationof lignin-glucose complex. This complex formation may be causedby hydrogen bond formation between the hydrophilic groups of

lignin and hydroxyl groups of glucose molecules. An increase in theintensityof hydrophilic functional group (like –OH) in phenolic andaliphatic structure (asobserved in the FTIR spectrum) may increasethe chance to form hydrogen bonding between glucose molecule

and modified lignin samples.

3.7. In vitro ˛-amylase inhibition assay

Alpha-amylase is responsiblefor the breakdown of complex car-bohydrate like starch to more simple sugars like glucose. Thus, theinhibition of this enzyme can delay the carbohydrate digestion and

reduce the rate of glucose absorption [12]. Adisakwattana et al.[41] f ound that theligninmodelcompounds, including11 cinnamicacid derivatives, caffeic acid, ferulic acid, and isoferulic acid did notexhibit any inhibitory action on

-amylase.

The -amylase inhibitory activity (expressed in percent) of the unmodified and biomodified lignin was investigated at rangesconcentrations of 0.1, 0.5, 1 and 5 mg/mL (Table 3). All lignin sam-ples significantly inhibited -amylase activity, but the modifiedlignin samples showed much higher inhibition than unmodi-fied ones. At the lower concentration, i.e. 0.1 and 0.5mg/mL, alllignin samples showed a similar inhibition activity, but in higherconcentration, i.e. 5 mg/mL, all modified lignin samples showed

higher inhibition than their respective unmodified lignin samples.At 5 mg/mL concentration, HNLC F10 exhibited the highest inhi-bition (78.2±0.97%) from all other samples followed by HNLCAPF4, HCLN APF4 and HCLN F10.The inhibition (%) of -amylaseactivity was approximately four times higher in the presence of 5 mg/mL than 0.1 mg/mL sample concentration. The inhibition of

-amylase activity was high in HNLC modified possibly be due tothe presence of high amount of methylated lignin fragments as

compared HCLN samples. Xiao et al. [42] have reported that themethylation of the hydroxyl group enhances the affinity of thepolyphenols for-amylase. Based on the finding they have opinedthat the increase in the number of hydrogen atom donor/acceptor

in the polyphenol might have decreased the affinity for towards-amylase.

Our finding also suggests that the biomodification in ligninenhances its inhibitory effect on -amylase. Some of the otherworkers reported that inactivation of this enzyme could be due toprecipitation by formation of inactive enzyme–inhibitor complexor enzyme–inhibitor–substrate complex [43]. The main drivingforces in the complex formation are hydrophobic interactions

and hydrogen bonding. In hydrophobic interaction, aromatic ringof phenolic compound and a hydrophobic part of proteins areinvolved, whereas hydrogen bonding occurs between hydroxylgroups of polyphenol and H-acceptor groups (NH, OH and COOH)

of proteins [42,44]. Appearance of new aromatic ring conjugatedhydrophilic group in biomodified samples as observed in the FTIR analysis supports our contention that chemically modified ligninis capable to hydrophobic and hydrogen bonding with enzymes

thus inhibiting its activity. With the increase in concentration of test samples, the chance of interaction is also increased, whichleads to diminished enzyme activity (upto 78.2%). The dimin-ished in enzyme activity are most likely by the formation of

enzymes–lignin–substrate or enzyme–lignin type complex forma-tion.

Table 3

% inhibition activity of -amylase enzyme by control and modified lignin samples at different concentration.

Samples % Inhibition

0.1mg/mL 0.5mg/mL 1 mg/mL 5 mg/mL

HCLN Control 14.95±2.49 58.24±1.74 63.27±1.57 62.69±1.70

HCLN F10 23.14±1.92*** 54.58±1.63* 60.20±1.22* 71.33±1.52***

HCLN APF4 14.89±1.62* 57.37±0.92* 61.96±3.08* 72.59±2.12***

HNLC Control 15.87±1.44 56.77±1.38 61.50±1.67 66.41±3.72

HNLC F10 19.70±1.99*** 56.53± 0.98* 60.52±0.63* 78.20±0.97***

HNLC APF4 17.46±0.20** 57.17±2.61* 60.00±1.57* 75.30±1.75***

Results were presented in mean±SD (n =3). Mean±SD, significantly different from their respective control at *** p


9/9


4. Conclusion

Ourresult reveals that thealkali lignin, extracted from A. nilotica,has free radical scavenging activity which is a structure depend-ent property, and was significantly altered when subjected to

bimodification. The DPPH and H2O2 scavenging assay showedremarkable potential of biomodified lignin as antioxidant. In thisstudy the degree of biomodification was influenced by the nutri-tional variations which were provided during biomodification.

In addition, the modified alkali lignin also showed a significant

in vitro -amylase inhibitory activity indicating its potential anti-hyperglycemic properties. Functional and structural modificationsin alkali lignin altered its binding efficiency towards glucose

molecule, which affected its movement across the membrane andimproving glycemic control by limiting the postprandial glucoseabsorptions. In conclusion the antioxidant,-amylase and in vitroglucose movement inhibition properties of A. nilotica lignin may

offer a potential therapeutic source for the treatment of oxidativestress and diabetes.

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