Accepted Manuscript Title: “In vitro antioxidant and antidiabetic activities of biomodified lignin from Acacia nilotica wood” Author: Anand Barapatre Keshaw Ram Aadil Bhupendra Nath Tiwary Harit Jha PII: DOI: Reference:
S0141-8130(15)00015-X http://dx.doi.org/doi:10.1016/j.ijbiomac.2015.01.012 BIOMAC 4819
To appear in:
International Journal of Biological Macromolecules
Received date: Revised date: Accepted date:
5-6-2014 26-11-2014 8-1-2015
Please cite this article as: A. Barapatre, K.R. Aadil, B.N. Tiwary, H. Jha, “In vitro antioxidant and antidiabetic activities of biomodified lignin from Acacia nilotica wood”, International Journal of Biological Macromolecules (2015), http://dx.doi.org/10.1016/j.ijbiomac.2015.01.012 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
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Title:
lignin from Acacia nilotica wood”
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Authors:
Department of Biotechnology, Guru Ghasidas Vishwavidyalaya
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Affiliation/Institute:
(A Central University), Bilaspur, Chhattisgarh, India. 495009
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Address for correspondence: Department of Biotechnology, Guru Ghasidas
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Anand Barapatre, Keshaw Ram Aadil, Bhupendra Nath Tiwary and Harit Jha *
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“In vitro antioxidant and antidiabetic activities of biomodified
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Vishwavidyalaya (A Central University), Bilaspur,
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Chhattisgarh, India. 495009
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Telephone No. :
+91-9826630805
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Email ID:
[email protected] Ac ce p
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Abstract The antioxidant and antidiabetic activity of biomodified alkali lignin extracted
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from a deciduous plant Acacia nilotica, was evaluated in vitro. The extracted alkali
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lignin was subjected to microbial biotransformation by ligninolytic fungus Aspergillus
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flavus and Emericella nidulans. These modifications were done under varying
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concentration of carbon to nitrogen sources. The structural feature of the lignin
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samples were compared by FTIR, functional group analysis and 13C solid state NMR.
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All lignin samples were tested for antioxidant efficiency, reducing power and H2O2
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scavenging power. Modifications in all lignin samples showed correlation with their
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antioxidant scavenging activity and reducing power. Antidiabetic properties were
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evaluated in terms of in vitro glucose movement inhibition and α-amylase inhibition
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assay. Modified samples exhibited increased glucose binding efficiency as
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demonstrated by the decreased glucose diffusion (55.5–76.3%) and 1.16-1.18 fold
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enhanced α-amylase inhibition in comparison to their control samples. The results
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obtained demonstrate that the structure and functional modifications in lignin
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significantly affects its bioefficacy in term of antioxidant and antidiabetic activities.
Key Word: Acacia lignin, biotransformation, antioxidant, antidiabetic, α-amylase, In vitro glucose movement.
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1. Introduction Plant phenolics and polyphenols have been increasingly enticing the attraction due to their beneficial therapeutic values including antioxidant,
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inflammatory, cardio-protective, anticancerous, chemo-preventive and neuro-protective
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properties. Polyphenols have been considered, a health food supplement and are claimed
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to possess health promoting or disease-preventing properties [1]. Lignin, the second
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most abundant natural macromolecule (polyphenolic in nature and 10–35% of dry wt. of
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lignocellulosic biomass), is a natural polymerized product of optically active p-
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hydroxycinnamyl alcohol monomers and related monolignols (p-coumaryl, coniferyl,
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and sinapyl) formed by oxidative reactions. This polymer is the result of various inter
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unit linkages in the monomer and monolignols (e.g. β-O-4, β-5, β−β, biphenyl (5-5), 4-
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O-5) [2]. The precise chemical structure of lignin is not known because of its complex
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polymeric nature and due to many random coupling. Alkali lignin is currently the largest
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produced among all lignin classes and a less valuable co-product of biofuel and paper
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pulp industries, which is separated from fibres by a chemical pulping (mainly soda and
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antimicrobial, anti-
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sulphite) process [3].
Acacia nilotica, locally known as “Babul”, is a multipurpose deciduous tree of
Mimosaseae family predominantly found in central India. This plant contains a variety of bioactive components such as ellagic acid, isoquercitin, leucocyanadin, kaempferol-7diglucoside, derivatives of (+)-catechin, apigenin derivatives etc. [4]. Traditionally in the
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central region of India leaves, pod, bark and root of A. nilotica is used for the treatment
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of various diseases related to oral, bone and skin, like cold, bronchitis, diarrhoea,
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dysentery, biliousness, bleeding piles and Leucoderma [5].
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Oxidative damages creates by free radicals, play a substantial role in the evolution of
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human diseases. Toxicity of free radicals contributes to proteins and DNA damage,
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inflammation, tissue injury and subsequent cellular apoptosis, which finally leads to
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cancer, emphysema, cirrhosis, arteriosclerosis and arthritis [6]. Oxidative stress is
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created in the body due to a disruption in the equilibrium between the production of
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reactive oxygen/nitrogen species (ROS/RNS) and the removal via the antioxidant
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defence system [7]. Antioxidants can interfere with the oxidative processes by reacting
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with free radicals, chelating catalytic metals and also by acting as oxygen scavengers
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thus helping the human body to reduce oxidative damage. From various epidemiological
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studies, it is proved that polyphenolic compounds possess an excellent antioxidant
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properties [6, 8]. Previous studies also reported that the polyphenols found in Acacia sp.
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plants having good antioxidant power [4, 5, 9, 10]. As a complex phenolic polymer,
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lignin also possess a respectable medicinal properties [2, 3, 11].
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Over the last century changes in human behaviour and life style have resulted in a
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dramatic increase in the incidence of diabetes world over. Presently, it is estimated that
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more than 220 million suffer from diabetes in which 90% is from type 2 diabetes Type 2
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diabetes is the results of ineffectiveness of insulin and the primary cause of complications linked to cardiovascular disease, renal failure, blindness, neurological complications, and so on [12]. Currently available conventional therapies for the treatment of diabetes include insulin and oral antidiabetic agents such as sulfonylurea, biguanides, and alpha-glucosidase inhibitors. Traditionally, many active compounds of
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plant origin including several Acacia sp. plants, have been also employed in the
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treatment of diabetes, mostly the secondary metabolites including alkaloids, flavonoids,
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phenolics, steroids, carbohydrates, glycopeptides, terpenoids etc. [13]. The inhibition of
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enzyme like α-amylase as well as the delay in glucose absorption to be an important
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strategy in the management of blood glucose level in type 2 diabetic [14]. Based on the fact that structural heterogeneity will affect the chemical properties of
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lignin, we report the bioefficacy of biologically modified lignin in terms of the
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antioxidant, antiradical and hydrogen peroxidase scavenging property and also evaluate
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the effectiveness of modified lignin as an inhibitor of α-amylase activity and in vitro
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glucose movement.
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2. Material and methods
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2.1. Chemical and reagents
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Gallic acid, D-glucose, α-amylase (EC 3.2.1.1), 1, 1-diphenyl-2-picrylhydrazyl
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(DPPH), neocuproine (Nc), catalase from bovine liver (966 U mg-1) were purchased
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from Sigma–Aldrich Inc. (Mumbai, India). All other chemicals and reagents used were
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of high purity analytical grade and purchased from Merck Pvt. Ltd. (India). Ultrapure
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water (Elix, Merck Milipore, India) was used throughout the experiment. Wood dust (18
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mesh size) of A. nilotica hardwood was procured locally from the saw mill of Bilaspur,
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Chhattisgarh, India.
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2.2. Extraction and characterization of alkali lignin Alkali extraction of lignin was achieved by treatment of wood dust with an aqueous
solution of NaOH (1.2% w/v) in a 1 L glass flask for 1 h at 120 ºC, 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 by slow heating at 60 ºC
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in an oven and acidified up to pH 5.5 with 6 M HCl. The water-soluble hemicellulosic
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fraction was removed by precipitation, after adding two volumes of 95% ethanol (v/v).
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The precipitated hemicellulosic were removed by gravity filtration. The remaining
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filtrate was concentrated to 20–30 mL, and the pH was adjusted to 1.5–2.0 with 6 M
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HCl. The alkali lignin was precipitated and sediments by centrifugation at 10,000 rpm
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(Remi R-24, India) for 10 min. The pellet was dried and stored at room temperature for
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further study [15].
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2.3. Biodegradation and characterization of alkali lignin
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2.3.1. Microorganism (fungus)
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The two potent ligninolytic fungus A. flavus and E. nidulans were used for the
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biotransformation of alkali lignin. The strains were isolated from soil samples collected
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from Guru Ghasidas Vishwavidyalaya campus, Bilaspur (C.G.) and near the effluent
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discharge site of the Orient paper mill situated in Amalai (M.P.), India. Two potentially
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ligninolytic strains were characterized and identified based on morphological
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characterization and partial gene sequencing of Internal Transcribed Spacer (ITS)
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regions as A. flavus (F10, NCBI accession no. KC91163) and E. nidulans (APF4, NCBI
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accession no. KC911632) respectively. The strains were maintained on malt agar slants
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for further use.
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2.3.2. Basal culture medium
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Biodegradation of lignin was performed under different carbon to nitrogen ratio i.e.
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low and high in Basal Salt Medium (BSM). The BSM contained gL-1 of KH2PO4, 0.2 g; MgSO4.7H2O, 0.05 g; CaC12, 0.01 g supplemented with a 1 mL mineral solution. The mineral solution contained (in gL−1) nitrilotriacetate, 1.5; MgSO4 .7H2O, 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
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low nitrogen (HCLN) medium contained 56 mM D-glucose as a carbon source and 2.4
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mM nitrogen (0.6 mM NH4NO3 and 0.6 mM L-asparagine) whereas the low carbon high
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nitrogen (HNLC) medium contained 8.8 mM D-glucose was and 24 mM nitrogen
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(NH4NO3 and L-asparagine, 6 mM each).The pH of BSM was maintained at 5.6-5.8 for
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biomodification of alkali lignin [16].
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2.3.3. Biomodification and characterization of alkali lignin Two hundred mililiter of HCLN and HNLC BSM medium maintained at pH 5.6-5.8
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were transferred to 500 mL erlenmeyer flasks. The autoclaved media were aseptically
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inoculated with 3 bores (6 mm diameter) of 7 days old culture of the strains F10 and
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APF4 and incubated at 28 °C for 21 days under static condition. The fungal mat was
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separated from the medium by filtration through sterilized Whatman filter paper No 4.
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The modified lignin was recovered from filtrate by the same method as described in
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section 2.2. Uninoculated media were used as negative control.
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Biotransformed alkali lignin was characterized by FTIR. A small fraction of the
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sample was ground properly with equal amount of KBr. The FTIR spectrum was
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obtained in the range of 400–4000 cm-1 using FTIR spectrophotometer Affinity A1,
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(Shimadzu, Japan).
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A high-resolution 1D solid-state
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C NMR spectra were obtained with the Cross-
Polarization Magic-Angle Spinning (CPMAS) technique on a Bruker spectrometer
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(Indian Institute of Science, Bangalore, India) operating at 100.525 MHz frequency for the 13C carbon nuclei. A total of 300 mg solid sample was used for the NMR spectra at 294 K. The 2000-6000 scan was performed to obtain a 1D spectra with a 29.10 ms acquisition time and 5 s relaxation delay. 2.4. Total polyphenol content (TPC)
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Total polyphenol content (TPC) was determined by reaction with Folin-Denis
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reagent [17]. One mL of each lignin sample (50 μg/mL) was added to 0.5 mL of Folin-
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Denis reagent. After 30 sec, 1 mL of 20% (w/v) sodium carbonate was added and the
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volume was made up to 5 mL with distilled water. The mixture was allowed to stand at
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room temperature for 10 min. The absorbance of resulting blue complex was measured
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at 765 nm against blank using UV-Visible double beam spectrophotometer (Shimatzu
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UV-1800, Japan). A calibration curve of gallic acid was prepared, and phenolic contents
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were determined from the linear regression equation of this curve. The results were
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expressed as μg gallic acid equivalents (GAE) per milligram of dry material.
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2.5. Functional group analysis
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2.5.1. Phenolic hydroxyl groups by ultraviolet-spectroscopy
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The content of hydroxyl phenolic units in lignin fractions was determined by UV
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spectroscopy as described Aadil et al. [9]. This method is based on the difference in
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absorption of lignin samples at pH 6 (495 mL of 0.2 N potassium dihydrogen phosphate
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solution mixed with 113 mL of 0.1 N NaOH and diluted to 2 L with distilled water) and
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at alkaline buffer solution pH 12 (0.1 N of boric acid in 0.1 N NaOH solution). The
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difference in spectra was obtained by taking the absorbance of the alkaline solution
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relative to that of the neutralized solution in the range of 200–400 nm. The phenolic
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hydroxyl group content of lignin samples was calculated using ∆amax.
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% phenolic hydroxyl = ∆amax X 17/41
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2.5.2. Carboxyl groups determination by aqueous titration method An amount of 25 mg of recovered lignin samples was suspended in 25 mL of an
alkaline 0.1 N NaOH solution and stirring for 3 h. The pH was adjusted to 12 with 0.1 N NaOH. Followed by potentiometric titration with 0.1 N HCl, as described Aadil et al. [9].
2.6. Total reducing power assay
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Reducing the power of all samples was determined according to the method
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of Oyaizu [18]. Briefly, different concentrations of lignin samples were prepared in
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sodium phosphate buffer (200 mM, pH 6.6) and 2.5 mL of each sample was separately
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mixed with 2.5 mL of 1% (w/v) potassium ferricyanide. The mixture was incubated at
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50 °C for 20 min. A mixture containing all the reaction reagents except the test material
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serves as the control. The reaction was stopped by adding 2.5 mL of 10% (w/v) TCA and
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the mixture was centrifuged at 1,750 rpm for 10 min. The upper layer (2 mL) was mixed
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with 1 mL of 0.1% (w/v) of ferric chloride and made up to 8 mL with deionised water.
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The perl’s prussian blue colour formed due to reduction in Fe3+ was measured at A700.
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The EC50 of extracts were calculated from the graph of A700 versus extracts
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concentration.
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2.7. DPPH free radical scavenging activity
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The antiradical activity of lignin samples was measured based on their reaction with
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stable free radical DPPH* and subsequent reduction in λmax of DPPH* [19]. In this
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reaction 1.5 mL sample, (100 µg/mL, prepared in 50 mM phosphate buffer, pH 7.5) was
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allowed to react with 1.5 mL of 100 µM methanolic solution of DPPH* for 30 min in
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darkness at room temperature. The decrease in absorbance was measured at A515.
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DPPH* radical scavenging capacity was calculated using the following equation
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where A0 and A1 are absorbance of DPPH* radical at 515 nm in the absence and presence of the samples.
2.8. Hydrogen peroxide scavenging (HPS) assay HPS capacity of lignin samples was estimated by cupric reducing antioxidant
capacity method according to Ozyurek et al. [20]. In this method, hydrogen peroxide
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incubation solution and scavenger solutions were prepared. The hydrogen peroxide
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incubation solution (used as a reference) contained 0.7 mL of phosphate buffer (0.2 M,
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pH 7.4), 0.4 mL of 1 mM H2O2, 0.4 mL of 0.1 mM CuCl2.2H2O, whereas scavenger
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solutions (I and II) were prepared in two test tubes containing 0.2 mL of test samples,
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0.5 mL of phosphate buffer (0.2 M, pH 7.4), 0.4 mL of 1 mM H2O2 and 0.4 mL of 0.1
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mM CuCl2.2H2O (identical up to this step). The mixtures were incubated for 30 min at
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37 ºC in water bath. After incubation 0.4 mL of H2O was added to the reference and
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scavenger solution-I and 0.4 mL of catalase solution (268 U mL-1) in scavenger solution-
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II. The mixture was vortexes for 30 sec. From the above incubated mixtures, 1 mL
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solution was mixed with 1 mL of Nc (0.0075 M, freshly prepared in ethanol), 1 mL of
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0.1 mM CuCl2.2H2O and 2 mL of ammonium acetate buffer (1 M, pH 7). After 30 min,
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the absorbance of the final solution was taken at 450 nm against the reagent blank. The
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HPS activity (%) of samples was calculated using the following formula:
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HPS (%) = [A0 – {A1 – A2}] / A0 X 100
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where A0 is the absorbance of reference hydrogen peroxide incubation solution, A1 and
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A2 are the absorbance of scavenger solution-I and –II, respectively.
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2.9. Antidiabetic assay by in vitro glucose movement
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To evaluate the effects of biotransformed lignin on glucose movement an in vitro
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model system was used according to Büyükbalci and Nehir [21] with slight
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modification. The dialysis tube (6 cm X 14.3 mm) (HiMedia, Mumbai, India; pore size
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2.4nm) was filled with a total volume of 6 mL test sample mixtures containing 1 mg/mL biotransformed lignin and 1.65 mM D-glucose (prepared in 0.15 M NaCl) in the ratio of 2:1 (v/v). The dialysis tube was sealed at both ends and placed in a flask containing 45 mL 0.15 M NaCl. Dialysis experiment was performed on an orbital shaker water bath (100 rpm) at 37 °C for 3 h to induce the movement of glucose into the external solution.
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Concentration of glucose outside the dialysis tubing was measured by DNS
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(dinitrosaliacylic acid) reagent. The control experiment was conducted in the absence of
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the test sample.
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2.10. In vitro α-amylase inhibition assay
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The α-amylase inhibitory activity was determined by the method of Quesille-
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Villalobos et al. [22]. A total of 500 µL of different concentrations of each sample (0.1,
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0.5, 1 and 5 mg/mL) and 500 µL of 0.02 M sodium phosphate buffer (pH 6.9 with 0.006
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M NaCl) containing α-amylase (0.5 mg/mL) was incubated for 10 min at 25°C. After
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preincubation, 500 µL of 1% (w/v) starch solution in 0.02 M sodium phosphate buffer
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(pH 6.9 with 0.006 M NaCl) was added to each of the pre-incubated tubes. The reaction
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mixtures were then incubated at 25 °C for 10 min and stopped with 1 mL of DNS
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reagent. The test tubes were further incubated in a boiling water bath for 10 min and
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cooled to room temperature. The reaction mixture was diluted with 10 mL distilled water
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and the absorbance was measured at 540 nm. The absorbance of blank samples (buffer
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instead of enzyme solution) and a control (buffer in place of the sample extract) was also
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recorded for comparison. The final activity of α-amylase was calculated by subtracting
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the final A540 of sample with its corresponding A540 blank.
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2.11. Statistical analysis
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Unless otherwise stated experiments were performed in triplicate and statistical
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analysis was done in term of mean ± standard deviation (SD). Significance levels were
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calculated using Graph Pad Prism 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,
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Versatile Peroxidases (VPs) and dioxygenases enzymes, working in association with
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small molecules and radicals [23]. The production of these enzymes is associated with
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nutrient stress conditions. White rot basidiomycetes and brown rot ligninolytic fungi
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display a broad diversity in response to carbon and nitrogen source and their C/N ratio.
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In most of the ligninolytic fungi LiP, MnP and laccase production is primarily regulated
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by the nitrogen concentration [24, 25]. Biodegradation of lignin starts with
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depolymerizing activity of LiP and MnP followed by demethylation activity of laccase.
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It is reported that the side chain and aromatic rings of lignin model compounds and
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synthetic lignin (DHPs) were cleaved via aryl cation radical and phenoxy radical
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intermediates which is mediated by LiP/H2O2 and laccase/O2/mediator system. Whereas
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MnP catalyse demethylation, Cα-Cβ cleavage, alkyl-aryl cleavage and Cα oxidation of
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phenolic syringyl type β-1 and β-O-4 lignin structures [26].
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In present study bio-modified lignin was obtained by biomodification of lignin using
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two ligninolytic fungi F10 and APF4. During microbial transformation of lignin by F10
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and APF4, the onset of MnP and LiP activity in HCLN medium starts from 3rd day and
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continuously increase up to the 21st day, whereas in HNLC medium it starts from 3rd in
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F10 and 12th day and APF4. The activity of both enzymes is minimal in HNLC as
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compared to HCLN medium. On the other hand, the laccase activity was high in HCLN
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medium, whereas in HNLC medium it was very low (unpublished data). After
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biomodification recovered biomodified lignin samples were evaluated for their TPC,
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functional group analysis, reducing power, hydrogen peroxide scavenging capacity, DPPH free-radical scavenging capacity, α-amylase and in vitro glucose movement inhibition activity.
3.1. Characterization of modified lignin 3.1.1. FTIR of lignin samples
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The FTIR spectrum of the control and biomodified lining samples is presented in Fig
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1a and 1b. All modified lignin samples (B, C, E and F) showed a broad absorption band
300
at 3410–3460 cm−1, attributed to the O-H groups stretching in phenolic and aliphatic
301
structures and oscillation of the hydroxyl group. The relative intensity of this band
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stretching was more in the treated sample (both HCLN F10 and HCLN APF4) as
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compared to control sample, which indicated the phenolic ring modification by addition
304
of a hydroxyl group. Whereas in modified HNLC samples (HNLC F10 and HNLC
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APF4), the intensity of O-H groups stretching was almost same in comparison to
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modified HCLN samples. A strong bands at 2847 cm−1 arising from C-H stretching in
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the aliphatic methylene group was less intense in treated sample (B, C, E and F) as
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compared to their control samples (A and D). Bands centred on 2938 and 2842 cm−1,
309
predominantly arising from C-H stretching in aromatic methoxyl groups and in methyl
310
and methylene groups of side chains. The intensity of both peaks is reduced in treated
311
samples as compared to control samples. The increase in the intensity of -OH group and
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decrease in C-H stretching also an outcome of laccase activity as suggested in the
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literature [27,28].
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All samples displayed weak bands in the carbonyl/carboxyl region 1705–1720 cm−1,
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assigned for unconjugated ketone or unconjugated carbonyl stretching. The intensity of
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aromatic skeleton vibrations at 1600, 1515 and 1426 cm−1 characteristic of the aromatic
317
ring in alkali lignin showed significant decrease in treated samples as compared to
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controls, indicating the cleavage of the aromatic ring structure. The C–H deformation combined with aromatic ring vibration at 1462 cm−1 was observed in the untreated control samples, however, these peaks were absent in treated samples. These results suggested that the structural and functional groups of alkali lignin were altered by both the fungal strains [29].
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Fig. 1a.
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Fig. 1b.
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3.1.2. 13C NMR 14 Page 13 of 36
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A comparison of the
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C NMR spectra of an untreated control lignin samples with
biodegraded lignin sample under two different nutritional conditions is presented in
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Figure 2. In all three NMR spectra of lignin, there was an absence of signals between 90
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and 102 ppm which indicates that the samples were free of carbohydrate contamination.
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The signals for unconjugated carboxylic acids –COOH (178.0-167.5 ppm) were high in
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HNLC sample in comparison to control while low in HCLN sample. The relative
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increase in this signal could be attributed to the formation of aldehydes, acids and
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aroxiacetic structures probably because of the oxidation of the side-chains. These results
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suggest oxidative attack on the lignin by the microbial enzymatic system of fungus [30].
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The signal for the aliphatic (171–168.5 ppm) and phenolic (168.5–167 ppm) hydroxyl
337
groups were observed in the lignin samples treated with fungus (HCLN and HNLC)
338
whereas they are absent in control lignin. The three aromatic region signals for
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protonated aromatic specially unsubstituted aromatic carbons ortho or para to the
340
substituted carbon (125–103 ppm), the condensed aromatic mainly C-substituted
341
aromatic carbon (141–125 ppm), and the oxygenated aromatic mainly O-substituted
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aromatic carbon of guaiacol (160–141 ppm) were higher in control lignin as compared to
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treated lignin [31].
The region between 162 to 103 ppm belongs to aromatic carbon, a syringal units
produce strong signal at 153-151 ppm (C-3 esterified) whereas the guaicyl unit signals at 119 ppm (C-6), 115 ppm (C-5) and 111 ppm (C-2) were also observed in control and treated samples. The changes in the aromatic-C region was observed, notably the
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decrease in the syringyl and guaiacyl amount (signals at 153 ppm and 148 ppm
349
respectively) after the fungal degradation of the alkali-lignin. The relative decrease in
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syringyl units reflects an easier accessibility of the microorganisms to the less condensed
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syringyl units of the polymer [30, 31].
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The signals at 90-57.5 ppm display aliphatic C-O bonds and α, β, γ carbons on the lignin
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side chain. In which Cα, Cβ, and Cγ in β-O-4 can be identified in the regions 79.0-67.0,
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90.0-78.0, and 61.5-57.5 ppm, respectively [31]. A significant decrease was observed in
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Cγ in β-O-4 (61.8 ppm) both treated samples as compare to control, while the other two
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signals for Cα and Cβ in β-O-4 increased in HNLC sample as compared to control. Some
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new signal intensity (30.5, 28.98 and 26.08) were also observed in the 46-10 ppm
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spectral region, mainly in the case of HNLC lignin sample, suggesting accumulation of
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saturated alkyl structures (Aliphatic CH2, 35.9 ppm) in the biomodified lignin. One of
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the signal at 35.9 corresponding to Cα in arylpropanol unit was decreased in HCLN
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sample in comparison to control while it was unaffected in HNLC lignin sample [30].
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3.2. Total phenolic content and functional group analysis All treated samples exhibited significantly lower TPC as compared to their
365
respective controls (Table 1). It was detected that in F10 and APF4 modified lignin
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samples, the quantity of TPC was low under HCLN than HNLC condition. In lignin
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degradation when lignin is exposed to peroxidases, it undergoes decomposition into
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lower molecular weight fragments containing methoxyl groups. Laccase demethylates these fragments and peroxidases further degrade them into smaller fragments. These smaller fragments reduce into their respective phenols by MnP, LiP and VPs which undergoes ring cleavage and form keto acids. The keto acids enter through kreb’s cycle and are metabolised by the fungus [32]. Variations in TPC were observed possibly due
373
to difference in the activity of ligninolytic enzymes under varying nutrient conditions.
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The results obtained revealed that the TPC of samples was influenced by lignin
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degradation under carbon and nitrogen surplus/stress conditions. Under HCLN condition
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enzymatic activity was high due to which lignin depolymerized into smaller fragments 16 Page 15 of 36
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and subsequently metabolized by the fungus. Previous results also supported our
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contention that the ligninolytic enzymes activities of different fungi were low/diminished
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in nitrogen rich medium as compare to nitrogen deficient medium [16, 23]. Functional group analysis were also carried out by UV spectrophotometric and
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potentiometric titration methods and results were presented in Table 1. All treated
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sample have equal and high phenolic hydroxyl group in comparison to their control
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sample which was also confirmed by FTIR analysis. In our results a correlation was
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found in the phenolic and carboxyl group amount in control and biomodified samples.
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Higuchi [26] reported that, laccase oxidizes the phenolic hydroxyl groups on lignin to
386
carboxylic groups, this might be the possible reason of this correlation.
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3.3. Reducing power assay
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A reducing agent contributes to antioxidant activity by donating its electron to free
389
radicals, which result in neutralization of the reactivity of the radical, and the reduced
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species subsequently acquire a proton from the solution. It was previously reported that
391
the Acacia species contain a number of active phenol and polyphenolic contents which
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were implied in the reducing reactions [9, 10, 33, 34]. Reducing power of all samples
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was expressed in the form of EC50 value (Table 1), which ranged from 405.41–1056.34 µg. From the results, it was observed that no significant change occurred in reducing power of lignin samples biotransformed under HCLN condition as compared to control, it was almost equal to the control samples. On the other hand, reducing power of other two lignin samples modified in HNLC condition (HNLC F10 and HNLC APF4) showed
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a decrease in the reducing power. The reduction in reducing power activity might be due
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to the partial depolymerisation of lignin polymer into low electropositive products.
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Nevertheless, the alkali lignin (modified and unmodified) tested in this experiment
401
showed significant reducing power in the range of 0.4-1 mg/mL, whereas Sultana et al.
17 Page 16 of 36
[33] reported them ethanolic extract of A. nilotica bark contains reducing power at 10
403
mg/mL range. Kalaivani and Mathew [5] also reported that the different solvent extracts
404
of A. nilotica leaves showed good reducing ability which increased with the sample
405
concentration.
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3.4. DPPH free radical scavenging activity
ip t
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The radical-scavenging activity of the control and bio-modified lignin was tested
408
using a methanolic solution of the ‘‘stable DPPH*”. On accepting hydrogen from a
409
corresponding donor, it loses the characteristic purple colour. The radical scavenging
410
activity of phenolic compounds depends on the rate of abstracting the hydrogen atom
411
from a phenol molecule by a free radical and, also on the stability of the radical formed.
412
This abstracting ability was increased if some additional conjugation with substituents
413
took place [11]. From results (Table 1) it was revealed that all alkali lignin samples
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(unmodified and modified) exhibited good scavenging activity, ranging from 37.94% to
415
71.44%. Both HCLN samples which were modified by F10 and APF4, displayed
416
increased antioxidant scavenging power i.e. 56.27% and 55.33% and both lignin samples
417
increase 21.2% and 12.6% of their free radical scavenging power with respect to HCLN
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control sample. Barclay et al. [35] reported that dimeric and tetrameric phenolic lignin model compounds have significantly higher antioxidant properties. Kiliç and Yesiloglu [34] and Rice-Evans et al. [37] also reported that hydroxycinnamtes (the basic units of lignin related compound) like p-coumaric acid, ferulic acids, chlorogenic acid, caffic acid also contain good antioxidant activity. Microbial depolymerisation of lignin in both
423
HCLN modified samples formed a high quantity of small monomeric, dimeric and
424
tetrameric compounds which might have increased the radical scavenging power, which
425
was also confirmed in NMR spectra.
18 Page 17 of 36
Lignin samples which were modified in HNLC condition displayed a decrease in
427
radical scavenging property. It was observed that HNLC F10 and HNLC APF4 lignin
428
brought a decrease of 32.6% and 23.9%, respectively, in their capability to scavenge the
429
free radical as compared to HNLC control. In HNLC condition, the produced
430
depolymerized lignin fractions by the activity of LiP and MnP, was probably less
431
demethylated due to suppression in laccase activity. Heim et al. [38] reported that O-
432
methylation decrease the antioxidant properties in flavonoid related compounds.
433
Dizhbite et al. [11] also reported that the due to extension of conjugation by carbonyl
434
groups in the propanoid side chain, radical scavenging activities diminishes significantly.
435
These above statements are the probable cause of reduction of antioxidant properties of
436
lignin sample which was modified in HNLC condition.
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3.5. Hydrogen peroxide scavenging assay
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Free radicals (especially OH*) are usually unstable, highly reactive, and energized
439
molecules generated from superoxide anion and hydrogen peroxide in the catabolic
440
system [7]. The results presented in Table 1, suggested that both modified HCLN
441
samples had increased hydrogen peroxide scavenging activity against control samples
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while in lignin samples modified under high nitrogen condition showed decrease in scavenging activity as compared to their control lignin. Sroka and Cisowski [39] studied hydrogen peroxide scavenging activities of some
water soluble phenolic acids and concluded that the hydrogen peroxide scavenging activity is positively correlated with the number of hydroxyl group bounded to the
447
aromatic ring, position of hydroxyl substitution in respect to each other and other
448
substituents (carboxyl and acetyl group) with their position in relation to the hydroxyl
449
group. They also concluded that there is a substantial correlation between antioxidant
450
and hydrogen peroxide scavenging activity of phenolic acids. Our studies also revealed a
19 Page 18 of 36
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strong correlation (r2 = 0.83, Fig 2) between antioxidant and hydrogen peroxide
452
scavenging activity of modified lignin samples.
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Fig. 3 3.6. Antidiabetic assay by in vitro glucose movement
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A surge in high post-prandial plasma glucose concentrations is associated with an
456
increased risk of developing type 2 diabetes and any reduction in the postprandial
457
glucose rush is potentially advantageous in avoiding diabetes. Gallagher et al. [40]
458
studied glucose diffusion inhibition activity of 10 traditional antidiabetic plants by in
459
vitro diffusion method. They reported agrimony and avocado as the most potent ( >60%
460
inhibition) inhibitor, whereas others inhibit the glucose diffusion in the range of 6–48%.
461
They also reported that the plant extracts exhibited a concentration dependent inhibitory
462
effect on glucose movement. While in another study Büyükbalci and Nehir [21] found
463
that there was no significant effect of herbal teas and infusions (traditionally used in the
464
treatment of diabetes in Turkey) on in vitro glucose diffusion.
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In the present study, control and modified lignin samples were tested for inhibitory
466
effects on glucose movementout of dialysis tube. No significant inhibition in glucose
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movement was observed by both control lignin samples as compared to blank (Table 2). But HCLN F10 and HNLC F10 lignin samples appeared to be the most potent inhibitor in glucose movement out of the dialysis tube, decreasing movement upto 76.3% and 68.1% (p