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Small laccase-catalyzed synthesis of a caffeic acid dimer with high antioxidant capacity
Accepted Manuscript Title: Small laccase-catalyzed synthesis of a caffeic acid dimer with high antioxidant capacity Authors: Blessing Nemadziva, Marilize Le Roes-Hill, Neil Koorbanally, Tukayi Kudanga PII: DOI: Reference:
S1359-5113(17)31852-4 https://doi.org/10.1016/j.procbio.2018.03.009 PRBI 11294
To appear in:
Process Biochemistry
Received date: Revised date: Accepted date:
4-12-2017 11-3-2018 13-3-2018
Please cite this article as: Nemadziva Blessing, Le Roes-Hill Marilize, Koorbanally Neil, Kudanga Tukayi.Small laccase-catalyzed synthesis of a caffeic acid dimer with high antioxidant capacity.Process Biochemistry https://doi.org/10.1016/j.procbio.2018.03.009 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.
Small laccase-catalyzed synthesis of a caffeic acid dimer with high antioxidant capacity
Blessing Nemadzivaa, Marilize Le Roes-Hillb, Neil Koorbanallyc, Tukayi Kudangaa*
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Department of Biotechnology and Food Technology, Durban University of Technology, P.O.
Biocatalysis and Technical Biology Research Group, Institute of Biomedical and Microbial
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b
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BOX 1334, Durban 4000, South Africa.
Biotechnology, Cape Peninsula University of Technology, Bellville Campus, Symphony Way,
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School of Chemistry and Physics, University of KwaZulu-Natal, Private Bag X54001,
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c
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P.O. Box 1906, Bellville 7535, South Africa.
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Durban, 4000, South Africa.
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Declarations of interest: none
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Corresponding author: Tukayi Kudanga
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Department of Biotechnology and Food Technology, Durban University of Technology, P.O. Box 1334, Durban, 4000, South Africa Tel: +27 31 373 5286 Fax: +27 31 373 3758 E-mail addresses: [email protected], [email protected]
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Graphical abstract
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Highlights
First report on the use of a small laccase in biocatalysis.
A β-β caffeic acid dimer, phellinsin A was synthesized in a biphasic system.
The dimer exhibits enhanced antioxidant capacity when compared to caffeic acid.
The dimer exhibits improved water solubility properties compared to caffeic acid.
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Abstract
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The increase in the demand for antioxidant compounds for application in industries such as food and pharmaceutical, has necessitated the need to investigate new production methods. The application of laccase for antioxidant synthesis is gaining notable attention as a viable approach
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to address this need. In this study, a β-β caffeic acid dimer, phellinsin A, was synthesized using the small laccase, SLAC, a two-domain bacterial laccase native to Streptomyces coelicolor A3(2). Phellinsin A showed a 1.5-fold increase in 2,2′-diphenyl-1-picrylhydrazyl (DPPH) radical scavenging capacity and a 1.8-fold improvement in trolox equivalence antioxidant capacity (TEAC) when compared to caffeic acid. Phellinsin A also showed improved solubility 2
properties in aqueous media and remarkable stability at acidic pH (pH 2.2 and pH 5.5). This study demonstrates the potential of the bacterial laccase, SLAC, as a catalyst for the biotransformation of natural phenolics into value-added compounds.
Keywords
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Small laccase; caffeic acid; coupling reactions; antioxidant activity; physicochemical
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properties
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1. Introduction
Mediterranean plant based diets have gained popularity among consumers since they contain
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high contents of bioactive compounds such as plant phenolics. Scientific research has linked plant phenolics with therapeutic properties such as improvement of plasma antioxidant capacity
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[1], anticarcinogenicity [2] and minimizing oxidative stress in general [3]. Plant phenolics are a group of phenol based bioactive compounds, which exist in plants as secondary metabolites.
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Caffeic acid is one of the major hydroxycinnamic acids found in plants. In particular, it is one
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of the major phenolic acids found in potatoes, wine and coffee [4].
The use of phenolic compounds for their antioxidant properties has been limited due to their low solubility in aqueous media [5] and susceptibility to heat, oxygen and light. Simple monomeric phenolics were reported to have a short half-life in the body [6] and may be metabolized to less active metabolites [7]. Caffeic acid, for example, is converted into
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conjugates such as glucuronide, sulphated or methylated derivatives during in vivo metabolic processes [7]. Piazzon et al. [8] showed that some of these structural alterations have a significant impact on antioxidant properties of caffeic acid: close to 20-fold decrease in antioxidant capacity of a caffeic acid metabolite (caffeic acid 4’-O-glucuronide) was reported in comparison to the parent compound. Caffeic acid is a simple phenolic compound consisting
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of only two hydroxyl groups on its aromatic ring and a C=C sidechain which are pivotal for its antioxidant properties. This may explain why metabolic conjugation significantly affects its
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bioactive properties. Modification processes that increase the number of hydroxyl groups could
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mitigate the reduction in activity resulting from these conjugation reactions.
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Consequently, interest in biocatalytic approaches of improving the properties of phenolic
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compounds is increasing. The enzymatic modification of phenolic compounds were also
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motivated by various other factors. These include the poor antioxidant capacity of some natural phenolics [9]; some monomeric forms acting as pro-oxidants [10]; the high cost of extraction
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of natural phenolics [11]; inefficiencies of chemical synthesis; and the need to enhance the bioavailability of natural phenolics, which is usually reduced due to poor solubility properties
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[12]. On the other hand, high molecular weight phenolics have been shown to have better bioactive properties compared to low molecular weight phenols [13]. Of the biocatalytic
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approaches, laccases have generated the most interest in improving antioxidant capacity through dimerization [9, 12, 14-16] or polymerization [6] to higher molecular weight
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compounds.
Laccase (E.C. 1.10.13.2) is a multi-copper oxidase that uses oxygen to abstract a single electron from a phenolic substrate to produce phenoxy radicals with water as a by-product [11]. Its ability to produce phenoxy radical intermediates has been applied in the formation of coupling
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products of therapeutic value. Although several researchers have reported the synthesis of compounds with enhanced bioactive properties using laccases (as reviewed in [11]), the work has employed only plant and fungal laccases. However, bacterial laccases would be attractive industrial catalysts due to the ease of expression in hosts such as Escherichia coli [17] and some favourable biochemical properties. For example, Machczynski et al. [18] characterized
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and overexpressed a two domain (small) laccase (SLAC) native to Streptomyces coelicolor A3(2) in E. coli. This enzyme has several attributes that make it potentially valuable for
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industrial application. It is thermally stable, recorded optimal activity at pH as high as 9.5 and
is resistant to denaturation by detergents [18]. Despite these favorable properties, there is no
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record of its application in biocatalysis. This is the first report of the application of SLAC in
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the synthesis of a dimeric antioxidant using caffeic acid as substrate. The bioactive and
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2. Materials and methods
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physicochemical properties of the synthesized compound were also investigated.
2.1 Chemicals and enzymes
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Chemicals were purchased from Sigma Aldrich (South Africa). The E. coli clone used for the expression of SLAC (wild type) was kindly provided by Prof. Gerard Canters, Leiden Institute
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of Chemistry, University of Leiden (Netherlands).
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2.2 Production of SLAC The production of SLAC was performed according to the protocol described by Prins et al. [19] with a few modifications. E. coli BL21(DE) cloned with a pET-20b(+) vector (Novagen) containing the gene coding for SLAC (pSLAC) was cultured in 2xYT media (16.0 g/L tryptone, 10.0 g/L yeast extract and 5.0 g/L NaCl) overnight. Overnight culture (1%, v/v) was inoculated
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in 2xYT media containing 100 µg/mL ampicillin and incubated at 37°C, 180 rpm until the OD600 reached 0.8. Induction for the overexpression of SLAC was achieved through the addition of isopropyl β-D-1-thiogalactopyranoside (IPTG) to a final concentration of 0.4 mM. The culture was further incubated at room temperature (22±3°C), 160 rpm for 20 h. Cells were harvested by centrifugation (5000 rpm, 10 min, 4°C) and resuspended in sodium phosphate
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buffer (pH 7.3). Cells were sonicated to release SLAC, which is produced intracellularly. The soluble fractions were collected by centrifuging (13 500 x g, 40 min, 4°C) and incubated in 1
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mM CuSO4 for 20 h at 4°C. The fractions were then dialyzed for four 1 h sessions at room
temperature (22 ± 3°C) with sodium phosphate buffer (10 mM, pH 7.3) and 1 mM EDTA added
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for the second session to chelate excess copper. The enzyme was further purified using size
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exclusion chromatography as reported in Prins et al. [19].
2.3 Enzyme activity
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SLAC activity was determined using the 2,2'-Azinobis (3-ethylbenzthiazoline-6-sulfonic acid) (ABTS) activity assay [20]. SLAC oxidation of ABTS was monitored by spectrophotometry at
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420 nm (ABTS, ε420 = 36 mM-1cm-1). The reaction mixture constituted citrate buffer (833 µL, 50 mM, pH 4.5), ABTS (110 µL, 5 mM) and laccase (57 µL). The reaction was conducted at
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25°C. One unit of laccase activity was defined as the amount of enzyme required to oxidise 1
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µmol of ABTS in 1 min at 25°C.
2.4 Oxidation of caffeic acid by SLAC Oxidation reactions were carried out in a buffer-organic solvent mixture. The reaction mixture contained caffeic acid (10 mM final concentration) and SLAC (0.64 U) in 50 mM sodium phosphate buffer (pH 7.5) (conditions determined from preliminary experiments). Optimum
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product yield was determined by comparing four organic solvents: the water miscible solvents, methanol and ethanol, and the water immiscible solvents, ethyl acetate and hexane at 50, 70, 80 and 90% organic solvent concentration (v/v) using 50 mM sodium phosphate (pH 7.5) as a buffer. The reaction was conducted for 3 h (25°C, 250 rpm). Since it was previously observed that SLAC was affected by organic solvents, 0.64 U/ mL enzyme was added at 90-min intervals
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to maintain enzyme activity.
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2.5 Analysis of products 2.5.1 Thin layer chromatography (TLC)
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Product formation was monitored by thin layer chromatography using aluminum backed silica
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gel 60 F254 TLC plates (Sigma, South Africa). A solvent system containing ethyl acetate,
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methanol, water and formic acid (8:1:1:0.02 v/v) was used as the mobile phase for the
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chromatographic separation of products. Plates were viewed under UV light at 254 nm.
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2.5.2 High Performance Liquid Chromatography (HPLC) HPLC was used for the analysis of oxidation products. For the preparation of samples, an equal
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volume of ice cold methanol was added to stop enzyme activity. The samples were kept on ice for 20 min, followed by centrifugation (13 500 x g, 10 min, 4°C) to precipitate out the enzyme
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as well as removing polymeric precipitates. Samples were filtered through 0.22 µm microfilters, transferred to clean HPLC vials and analyzed using a Waters HPLC system
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(Massachusetts, USA). Separation was achieved on a Waters HSS T3 column, 2.1x100 mm with 1.7 um particles. A linear gradient was applied using 0.1% formic acid (solvent A) and acetonitrile (solvent B). The linear gradient started at 100% solvent A for 1 min and changed to 50% B over 22 min. It then went to 100% B after 23 min where it was held until 24.5 min,
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followed by re-equilibration to initial conditions for 4 min. The flow rate was 0.25 mL/min and the column was kept at 60 ºC. The injection volume was 3 µL.
2.6 Purification of products Purification of the oxidation product was performed by preparatory thin layer chromatography
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glass plates (500 µm, silica gel 60 F254, Sigma, South Africa) using ethyl acetate, methanol,
water and formic acid (8:1:1:0.02 v/v) as the mobile phase. Pure bands were observed under
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UV light at 254 nm and were marked with a pencil. The product bands were scraped off the plate and re-dissolved in methanol. After filtering off the silica using 0.22 µm syringe filters,
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the product was washed with acetone, water and acetone again before being dried in clean glass
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vials.
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2.7 Characterization of products
2.7.1 Liquid Chromatography-Mass Spectrometry (LCMS)
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A Waters Synapt G2 quadrupole time-of-flight mass spectrometer (Massachusetts, USA) was used for LC-MS analysis. It was fitted with a Waters ACQUITY Ultra Performance Liquid
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Chromatography (UPLC) system and photo diode array detection. Separation was achieved
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on a Waters HSS T3 column as described in Section 2.5.2.
Data was acquired in MSE mode which consisted of a low collision energy scan (6V) from m/z
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150 to 1500 and a high collision energy scan from m/z 40 to 1500. The high collision energy scan was done using a collision energy ramp of 30-60V. The photo diode array detector was set to scan from 220-600 nm.
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The mass spectrometer was optimized for best sensitivity, a cone voltage of 15 V, desolvation gas was nitrogen at 650 L/hr and desolvation temperature 275ºC. The instrument was operated with an electrospray ionization probe in the negative mode. Sodium formate was used for calibration and leucine encephalin was infused in the background as lock mass for accurate
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mass determinations.
2.7.2 Nuclear Magnetic Resonance (NMR)
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For NMR analysis, 5-10 mg samples were dissolved in 0.5 mL CD3OD. 1H, 13C and 2D NMR spectra were recorded on a Bruker Avance instrument operating at 400 MHz. Chemical shifts were reported as δ values (ppm) relative to the solvent line of CD3OD (3.34 ppm for 1H; 49.0
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ppm for 13C).
4-(3,4-dihydroxybenzylidene)-2-(3,4-dihydroxyphenyl)-5-oxotetrahydrofuran-3-carboxylic
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acid P1, 1H NMR (CD3OD, 400 MHz): H 7.50 (d, J = 1.9 Hz, H-7'), 7.22 (d, J = 2.0 Hz, H2'), 7.09 (dd, J = 8.2, 2.0 Hz, H-6'), 6.83 (d, J = 8.2 Hz, H-5'), 6.76 (d, J = 8.1 Hz, H-5), 6.75
Hz, H-8);
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(d, J = 2.1 Hz, H-2), 6.69 (dd, J = 8.1, 2.1 Hz, H-6), 5.60 (d, J = 2.5 Hz, H-7), 3.94 (d, J = 2.3 C NMR (CD3OD, 100 MHz): C 178.2 (C-9), 175.5 (C-9'), 149.4 (C-4'), 146.8,
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146.7, 146.6 (C-3', C-3 and C-4); 139.9 (C-7'), 133.9 (C-1), 127.6 (C-1'), 125.7 (C-6'), 123.3
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(C-8'), 118.3 (C-6), 117.8 (C-2'), 116.6 (C-5), 116.4 (C-5'), 113.5 (C-2), 84.8 (C-7), 58.7 (C8).
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2.8 Determination of antioxidant properties Antioxidant activities of caffeic acid and the oxidation product were determined using the following methods: Trolox equivalent antioxidant capacity (TEAC) assay and 2,2′-diphenyl-1picrylhydrazyl (DPPH) assay.
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2.8.1 Trolox equivalent antioxidant capacity (TEAC) assay The ABTS radical scavenging activity of caffeic acid and the reaction product was determined according to the protocol by Re et al. [21] with slight modifications. ABTS•+ solution was prepared 12-16 h before use by reacting 7 mM of ABTS salt with 5.95 U SLAC and then storing in the dark until the assay was performed. The ABTS•+ solution was diluted with methanol to
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give an absorbance (Libra S21 spectrophotometer, Biochrom, Cambridge, United Kingdom)
of 0.70 ± 0.002 at 734 nm. Each sample (100 μL) prepared at different concentrations was
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mixed with 1100 μL ABTS•+ solution and the absorbance was read after 30 min incubation at 25°C.
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2.8.2 2,2′-diphenyl-1-picrylhydrazyl (DPPH) scavenging effect
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DPPH was prepared by dissolving DPPH in methanol to a final concentration of 0.025 mg/mL.
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Test samples for both caffeic acid and reaction product were prepared in methanol at various concentrations. To conduct the assay, 0.1 mL of test sample was mixed with 3.9 mL DPPH.
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The mixture was shaken vigorously and incubated in the dark at 25°C for 60 min. After incubation, the absorbance was measured by spectrophotometry at 517 nm. The EC50 of the
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test samples were determined by plotting the percentage remaining DPPH against sample concentration. Percentage remaining DPPH was calculated by calculating the percentage of the
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concentration of DPPH at T60 against DPPH concentration at T0. EC50 value is defined as the
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concentration (mM) of substrate that results in 50% loss of DPPH• [14].
2.9 Physicochemical properties The physicochemical properties of caffeic acid and the oxidation product were compared by investigating pH stability, temperature stability and photo-stability. The physicochemical
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properties were studied in hydro-alcohol solutions due to the solubility characteristics of caffeic acid, which is not soluble in aqueous medium.
2.9.1 pH stability pH stability was investigated at pH 2.2, 5.5 and 7.5 following the protocol described by
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Amendola et al. [22]. The test samples were dissolved in buffer-alcohol solution (85:15 v/v) using 50 mM citrate-phosphate buffer at pH 2.2, 5.5 and 7.5 to a final concentration of 0.5 mM.
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The test flasks were covered with aluminum foil to eliminate light interference and were kept
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at 4°C for 30 days. Samples were taken for HPLC analysis at 5-day intervals.
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2.9.2 Temperature stability
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Storage temperature stability was analyzed for 30 days at 4°C, 25°C and 37°C according to the
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method adopted and modified from Amendola et al. [22]. The test samples were dissolved in a hydro-alcohol solution of water and methanol (85:15 v/v) to a final concentration of 0.5 mM.
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The test flasks were covered with aluminum foil and kept at 4°C, 25°C and 37°C for 30 days.
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Samples were taken for HPLC analysis at 5-day intervals.
2.9.3 Photo-stability
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Photo-stability was investigated using fluorescent light. The test samples were prepared by dissolving the compounds in a hydro-alcohol solution of water and methanol (85:15 v/v) to a final concentration of 0.5 mM. The test flasks were exposed to fluorescent light (Osram L18W/640 with a luminous flux of 1200 lm, 18 W and luminance of 508 542 lx) for 30 days. Samples were taken for HPLC analysis at 5-day intervals.
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3. Results and discussion 3.1 Structural characterization of SLAC oxidation product The SLAC catalyzed oxidation of caffeic acid resulted in the formation of one major product designated as P1 (tR=5.29), (Fig. 1a). LC-MS analysis conducted in the negative mode
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indicated that the product was a dimer m/z 357, [M] 358.07 (Fig. 1b). The in situ fragmentation pattern showed dominant signals at m/z 313 (– 1 carboxyl group) and m/z 269 (– 2 carboxyl
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groups). Substrate conversion to the dimer was monitored by HPLC over a 12 h oxidation
period. It was observed that at 6 h, optimum dimer was produced (yield 32.8%), after which
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the yield started decreasing. This may be due to a combination of non-specific side reactions
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as well as polymerization reactions, usually associated with laccase-mediated radical reactions
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[11].
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The NMR results of P1 indicated that it is a dimer of caffeic acid through a β-β linkage (Fig.
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2). Since the molecule was not symmetrical, the resonances for each of the aromatic protons in the rings could be seen separately. Ortho coupling was seen for H-5 and H-6 at H 6.76 (d,
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J = 8.1 Hz) and H 6.69 (dd, J = 8.1, 2.1 Hz) and H-5' and H-6' at H 6.83 (d, J = 8.2 Hz) and
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H 7.09 (dd, J = 8.2, 2.0 Hz). H-6 and H-6' were meta coupled to H-2 and H-2' respectively at H 6.75 (d, J = 2.1 Hz) and 7.22 (d, J = 2.0 Hz). The olefinic H-7' resonance was deshielded at H 7.50 appearing as a doublet with J = 1.9 Hz due to long range coupling with H-8, which
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appeared as a triplet at H 3.94 (J = 2.3 Hz) due to the double doublet coalescing. H-7 was seen at H 5.60 as a doublet with J = 2.5 Hz. The small J value of H-7 and H-8 indicated that the dihedral angle was approaching 90. The carbon resonances for the two carbonyl groups could be seen at C 178.2 (C-9) and 175.5 (C-9'), assigned due to HMBC correlations with H7 and H-8 (for C-9) and H-7' (for C-9'). C-4' was distinctly separated from the other aromatic 12
oxygenated resonances at C 149.4, and assigned due to HMBC correlations with H-2', H-6' and H-5'. The other three aromatic C-O resonances, C-3', C-3 and C-4 appeared at C 146.8, 146.7 and 146.6 and can be interchanged. Both C-1 and C-1' at C 134.0 and 127.6 were identified by HMBC correlations with H-5 and H-5' respectively. The remaining singlet carbon
HMBC spectra, please refer to Fig. S1-Fig. S8 (supplementary data).
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resonance at C 123.3 was assigned to C-8'. For details of 1H and 13C NMR, COSY, HSQC and
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The dimer has been reported in culture broth of Phellinus sp. and was identified as phellinsin A [23] and has also been previously synthesised from Lycopus europaeus catechol oxidase [24]
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and the cis form from Momordica charantia peroxidase-catalysed processes [25]. However,
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the SLAC-catalysed process may potentially be the cheapest method compared to previously
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described protocols [24, 25] which employed plant-derived biocatalysts thus making the
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processes potentially cost-intensive. In addition, the dependence of peroxidases on H2O2 makes
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the synthetic process expensive as well as a health hazard [26].
Caffeic acid is one of the putative lignin monomers. Although β-O-4 linkages are the
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predominantly formed linkages in polymer formation in nature [27], it has been observed that
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dimerization usually results in β-5 and β-β linkages [28]. The suggested mechanism for the formation of the β-β dimer is shown in Fig. 3 [24, 25]. A radical is formed from the SLAC oxidation of the hydroxyl group occupying the para position of caffeic acid. The lone electron
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on the subsequent radical can occupy different positions through resonance stabilization. C–C linkage of two phenoxy radicals with unpaired electrons at β positions results in the formation of the β-β caffeic acid dimer, phellinsin A. A similar reaction pathway was proposed by Adelakun et al. [9] for the formation of a laccase catalyzed β-β ferulic acid dimer, suggesting that the β-β linkage is one of the most preferred radical coupling positions for phenolic acids. 13
3.2 Effect of solvent on the formation of phellinsin A Laccase catalysis of phenolic compounds is usually performed in buffer-organic solvent reaction systems. This is because of the sparse solubility of most phenolic compounds (including caffeic acid) in aqueous media, It has also been observed that increasing solvent
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concentrations help minimize polymerization [9, 14]. For this study, four solvents, methanol, ethanol, ethyl acetate and hexane were investigated at four different concentrations (50, 70, 80
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and 90%) to determine the best solvent conditions for the production of phellinsin A.
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Results indicated that there was an increase in product formation as the polarity of the solvent
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decreased; Hexane> ethyl acetate > methanol > ethanol (Fig. 4). There was a general increase
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in phellinsin A formation as the solvent concentration increased from 50% to 80%, after which
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a sharp decline in product formation at 90% co-solvent, was observed. Although 80% hexane resulted in the highest yield of phellinsin A, ethyl acetate, which yielded the second highest
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concentration of phellinsin A, offered a cleaner reaction with less side reactions, thus making it easier for purification. Therefore, scale up reactions were carried out using 80% ethyl acetate
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as co-solvent. There was a significant loss of enzyme activity as the oxidation reaction progressed; by 90 minutes, 94.7% of activity was lost (Fig. S9, supplementary data). Therefore,
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0.64 U enzyme was periodically added at 90-minute intervals for the reaction to proceed. Organic solvents were reported to affect laccase activity by direct interaction with the enzyme
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molecule or through modifying the thermodynamic water activity of the enzyme [29, 30].
3.3 Bioactive properties Antioxidant capacity of phellinsin A was compared to caffeic acid using DPPH and TEAC assays. The DPPH and TEAC assays are based on the radical scavenging ability of an
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antioxidant to reduce DPPH• and ABTS•+ to their more stable states (1,1-diphenyl-2-picryl hydrazine and 2,2'-azino-bis-3-ethylbenzthiazoline-6-sulphonic acid, respectively) [31]. Both assays showed an improved antioxidant capacity of the dimer; 1.5-fold and 1.8-fold increase in antioxidant capacity for DPPH and TEAC assays, respectively compared to caffeic acid (Table 1). Studies of structure-activity relationships (SAR) of antioxidants have attributed the
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antioxidant capacity of a compound to factors such as the availability of electron donating
groups attached to the aromatic ring [11, 32] as well as ability to form stable radicals. Caffeic
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acid contains two hydroxyl groups, as well as an unsaturated C=C bond in the side chain.
Phellinsin A, on the other hand has four hydroxyl groups as well as the unsaturated C=C bond,
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which explains its improved antioxidant capacity. It has been reported that effective
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antioxidants are able to form stable phenoxy radicals through either hydrogen bonding [33] or
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delocalization of lone pair electrons into the aromatic ring [11, 34]; since the structure of
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phellinsin A does not consist of adjacent aromatic rings which enable expanded electron delocalization [32], enhanced antioxidant activity may be attributed to increased number of
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hydroxyl groups which facilitates radical stability through hydrogen bonding.
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3.4 Physicochemical properties
Besides enhanced bioactivity, physicochemical profiles also qualify a compound for
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commercial development [35]. Conditions such as pH, temperature, air and light were reported to affect stability of phenolic compounds [36]. Despite significant progress being made in
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laccase synthesis of bioactive compounds, comprehensive characterization of products has been neglected. A literature survey of all laccase mediated coupling products synthesized since 2000 showed that only one article investigated physicochemical properties of the coupling products [37]. The extent of product characterization has narrowly focused mostly on bioactivity, however, an extensive analysis of coupling products is required for the constructive
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progression of laccase biotransformation technology. Therefore, the physicochemical properties of phellinsin A were investigated.
3.4.1 Phellinsin A solubility Caffeic acid is not soluble in aqueous media, therefore it was dissolved in at least 50% methanol
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for complete solvation at room temperature. However, phellinsin A was completely soluble in aqueous media at room temperature. This may be attributed to the increased number of
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hydrogen bond interactions between the compound and water [38]. Solubility is an important factor in drug development since it can give a theoretical indication of the bioavailability of a
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compound in vivo [39].
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3.4.2 Temperature stability
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Phellinsin A showed significant instability when incubated at body temperature (37°C) in a hydro-alcohol solution of water and methanol (85:15 v/v). In the first 5 days of incubation,
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there was 63.6% loss of phellinsin A and by day 20, it was completely degraded (Fig. 5). At storage temperatures (4°C and 25°C), phellinsin A exhibited relative stability, recovering
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85.5% and 39.3% of the compound, respectively after 30 days of incubation. These results have shown that the concentration of phellinsin A decreased with increasing temperature. Despite
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phellinsin A exhibiting significant instability at body temperature over prolonged periods, it can still be an effective antioxidant. Pharmacokinetic studies have shown that most phenolic
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compounds are metabolized within 24 h, after which the plasma concentration reaches basal levels [40, 41]. At body temperature, about 60-80% of phellinsin A was retained during the first 24 h (Fig. 5 and S10), when the compound is expected to be metabolized. Therefore, phellinsin A can be a useful antioxidant since it has shown relative stability at storage temperature (4°C) while exhibiting significant stability within the metabolism period. On the
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other hand, caffeic acid showed a contrasting trend; increasing stability as temperature increased. Caffeic acid exhibited the highest stability at 37°C and retained 76.5% of compound by day 30 (Fig. 5). Previous studies on temperature–antioxidant activity of phenolic acids determined in triacylglycerols of sunflower oil showed that cinnamic acid derivatives including caffeic acid were more effective antioxidants at 90°C than at 22°C, owing to their higher
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stability at high temperatures [34, 42].
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3.4.3 pH Stability
pH stability was investigated considering potential applications and the related environments.
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For instance, if phellinsin A is considered for the development of a nutraceutical, it would be
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interesting to observe the effect of gastrointestinal pH (pH 2.2 for the stomach and pH 7.5 for
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alkaline intestinal conditions) [43]; or if phellinsin A could be an ingredient in development of
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topical products, then its stability at skin pH (pH 5.5) [44] is important. pH stability tests were performed at 4°C to reduce any microbial contamination which is common especially at acidic
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buffer conditions [22]. Results showed that the stability of the SLAC synthesized phellinsin A is pH-dependent. It showed remarkable stability in acidic conditions but was extremely
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unstable under alkaline conditions. At pH 5.5, it was completely stable; no compound loss was observed after monitoring for 30 days. Therefore, the product may be considered for
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application in the development of skincare products. At pH 7.5, phellinsin A was completely degraded within 5 days of incubation (Fig. 6). Similarly, previous studies on stability profiles
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of phenolic compounds have demonstrated that phenolic compounds such as caffeic acid, gallic acid, flavonoids, chlorogenic acid and green tea catechins are pH-sensitive; exhibiting greater stability in acidic pH and are unstable under alkaline conditions [45, 46, 47]. It has been suggested that the effect of pH on stability of a phenolic compound is related to the resonance stabilization of its subsequent phenoxide ions which are mostly stable in acidic conditions than
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in alkaline medium [47]. It was therefore not surprising the dimer with a higher electron density was more stable than the monomeric unit.
3.4.4 Light stability Light significantly impacted the stability of phellinsin A with only 11.5% of the compound
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recovered after 30 days of incubation (Fig. 7). Photo-stability tests were carried out in conditions that, to a large extent, simulated normal storage environments at room temperature,
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oxygen and under normal fluorescent light. Photodegradation of phenolic compounds is induced when valence band electrons are excited in the presence of oxidants such as molecular
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oxygen causing a compound to be more susceptible to oxidative degradation [48]. The
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susceptibility of phellinsin A to degradation was not surprising since previous investigations
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have shown that polyphenols are generally light sensitive [49]. Therefore, the compound may
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be most suitable for application in topical product formulations since it has shown the highest stability at pH 5.5 and significant light stability (approximately 80% retained within 24 h, Fig
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S10) during the period in which it is expected to be metabolized. Due to the susceptibility of phellinsin A to light as well as high temperatures, the best storage conditions will be at 4°C in
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packaging material that excludes light (such as amber bottles).
4. Conclusion
This study has reported the application of SLAC as a biocatalyst in the synthesis of antioxidants
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for the first time. A β-β caffeic acid dimer, phellinsin A, was successfully synthesized and its bioactive and physicochemical properties elucidated. The dimer exhibits improved solubility properties and enhanced antioxidant properties compared to the parent compound, caffeic acid. This study indicates that SLAC could be a potential biocatalyst for the production of value added antioxidant products.
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5. Acknowledgements We express our sincere gratitude to Ms Devrani Naicker (Durban University of Technology) for her assistance with chromatography analysis, Mr Bibhuti Ranjan (Durban University of Technology) for his assistance with enzyme purification and Ms Thandokazi Andiswa for
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assistance with NMR. We are grateful to the National Research Foundation (NRF) for financial
assistance (Grant 105889, 112099 and 93362) and to the Council for Scientific and Industrial
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Research (CSIR) for student (Blessing Nemadziva) bursary support. Any opinion, findings, and conclusions or recommendations expressed in this material are those of the author(s), and
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therefore, the NRF does not accept any liability in regard thereto.
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Figure legends Fig 1a. HPLC chromatogram of caffeic acid oxidation product (tR = 5.29). Fig 1(b) mass spectra of SLAC oxidation product (P1). Fig 2. The β-β linked dimer (phellinsin A) formed during SLAC-catalyzed oxidation of
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caffeic acid. Oxidation reactions were carried out in buffer-organic solvent systems using 50 mM sodium phosphate buffer (pH 7.5) and varying concentrations of selected organic
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solvents. Dimer was purified using preparatory thin layer chromatography.
Fig 3. Proposed reaction pathway for the SLAC-catalysed synthesis of phellinsin A. A C–C bond is formed between two β-positioned caffeic acid radicals. The intermediate compound
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undergoes rearrangement reactions to form the β-β dimer, phellinsin A.
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Fig 4. Effect of organic solvents on the synthesis of phellinsin A. Caffeic acid was oxidized
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by SLAC in either biphasic or monophasic reaction systems at varying concentrations of the test organic solvent (ethanol, ethyl acetate, hexane or methanol). Phellinsin A formation was
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replicate determinations.
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monitored by HPLC analysis. All results are means ± standard deviation (SD) of three
Fig 5. Stability profile of caffeic acid and phellinsin A at 4°C, 25°C and 37°C. Test samples
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were prepared in hydro-alcohol solutions and incubated in the dark at the investigated temperature for 30 days. All results are means ± standard deviation (SD) of three replicate
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determinations.
Fig 6. Stability profile of caffeic acid and phellinsin A incubated in citrate-phosphate buffer at pH 2.2, 5.5 and 7.5. Test samples were prepared in buffer-alcohol solutions at varying pH and were incubated in the dark at 4°C for 30 days. All results are means ± standard deviation (SD) of three replicate determinations.
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Fig 7. Photo-stability profile of caffeic acid and phellinsin A. Test samples were prepared in hydro-alcohol solutions and were exposed to fluorescent light for 30 days. All results are
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means ± standard deviation (SD) of three replicate determinations.
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Table 1 Antioxidant activity of caffeic acid and phellinsin A. Standard assays were used to determine activity. All results are means ± standard deviation (SD) of three replicate determinations. Molecular weight (g.mol-1) EC50 DPPH (mM)
TEAC valuea (mM)
Caffeic acid
180.16
0.44 ± 0.008
2.17 ± 0.060
Phellinsin A
358
0.29 ± 0.004
3.88 ± 0.004
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a
Compound
TEAC value is defined as the concentration of Trolox (mM) with an antioxidant potential
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equivalent to 1 mM of compound under investigation [14]. EC50 value is defined as the
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concentration (mM) of substrate that results in 50% loss of DPPH• [14].
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Fig 1.
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S1359-5113(17)31852-4 https://doi.org/10.1016/j.procbio.2018.03.009 PRBI 11294
To appear in:
Process Biochemistry
Received date: Revised date: Accepted date:
4-12-2017 11-3-2018 13-3-2018
Please cite this article as: Nemadziva Blessing, Le Roes-Hill Marilize, Koorbanally Neil, Kudanga Tukayi.Small laccase-catalyzed synthesis of a caffeic acid dimer with high antioxidant capacity.Process Biochemistry https://doi.org/10.1016/j.procbio.2018.03.009 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.
Small laccase-catalyzed synthesis of a caffeic acid dimer with high antioxidant capacity
Blessing Nemadzivaa, Marilize Le Roes-Hillb, Neil Koorbanallyc, Tukayi Kudangaa*
a
Department of Biotechnology and Food Technology, Durban University of Technology, P.O.
Biocatalysis and Technical Biology Research Group, Institute of Biomedical and Microbial
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b
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BOX 1334, Durban 4000, South Africa.
Biotechnology, Cape Peninsula University of Technology, Bellville Campus, Symphony Way,
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School of Chemistry and Physics, University of KwaZulu-Natal, Private Bag X54001,
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c
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P.O. Box 1906, Bellville 7535, South Africa.
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Durban, 4000, South Africa.
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Declarations of interest: none
*
Corresponding author: Tukayi Kudanga
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Department of Biotechnology and Food Technology, Durban University of Technology, P.O. Box 1334, Durban, 4000, South Africa Tel: +27 31 373 5286 Fax: +27 31 373 3758 E-mail addresses: [email protected], [email protected]
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Graphical abstract
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Highlights
First report on the use of a small laccase in biocatalysis.
A β-β caffeic acid dimer, phellinsin A was synthesized in a biphasic system.
The dimer exhibits enhanced antioxidant capacity when compared to caffeic acid.
The dimer exhibits improved water solubility properties compared to caffeic acid.
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Abstract
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The increase in the demand for antioxidant compounds for application in industries such as food and pharmaceutical, has necessitated the need to investigate new production methods. The application of laccase for antioxidant synthesis is gaining notable attention as a viable approach
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to address this need. In this study, a β-β caffeic acid dimer, phellinsin A, was synthesized using the small laccase, SLAC, a two-domain bacterial laccase native to Streptomyces coelicolor A3(2). Phellinsin A showed a 1.5-fold increase in 2,2′-diphenyl-1-picrylhydrazyl (DPPH) radical scavenging capacity and a 1.8-fold improvement in trolox equivalence antioxidant capacity (TEAC) when compared to caffeic acid. Phellinsin A also showed improved solubility 2
properties in aqueous media and remarkable stability at acidic pH (pH 2.2 and pH 5.5). This study demonstrates the potential of the bacterial laccase, SLAC, as a catalyst for the biotransformation of natural phenolics into value-added compounds.
Keywords
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Small laccase; caffeic acid; coupling reactions; antioxidant activity; physicochemical
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properties
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1. Introduction
Mediterranean plant based diets have gained popularity among consumers since they contain
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high contents of bioactive compounds such as plant phenolics. Scientific research has linked plant phenolics with therapeutic properties such as improvement of plasma antioxidant capacity
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[1], anticarcinogenicity [2] and minimizing oxidative stress in general [3]. Plant phenolics are a group of phenol based bioactive compounds, which exist in plants as secondary metabolites.
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Caffeic acid is one of the major hydroxycinnamic acids found in plants. In particular, it is one
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of the major phenolic acids found in potatoes, wine and coffee [4].
The use of phenolic compounds for their antioxidant properties has been limited due to their low solubility in aqueous media [5] and susceptibility to heat, oxygen and light. Simple monomeric phenolics were reported to have a short half-life in the body [6] and may be metabolized to less active metabolites [7]. Caffeic acid, for example, is converted into
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conjugates such as glucuronide, sulphated or methylated derivatives during in vivo metabolic processes [7]. Piazzon et al. [8] showed that some of these structural alterations have a significant impact on antioxidant properties of caffeic acid: close to 20-fold decrease in antioxidant capacity of a caffeic acid metabolite (caffeic acid 4’-O-glucuronide) was reported in comparison to the parent compound. Caffeic acid is a simple phenolic compound consisting
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of only two hydroxyl groups on its aromatic ring and a C=C sidechain which are pivotal for its antioxidant properties. This may explain why metabolic conjugation significantly affects its
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bioactive properties. Modification processes that increase the number of hydroxyl groups could
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mitigate the reduction in activity resulting from these conjugation reactions.
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Consequently, interest in biocatalytic approaches of improving the properties of phenolic
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compounds is increasing. The enzymatic modification of phenolic compounds were also
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motivated by various other factors. These include the poor antioxidant capacity of some natural phenolics [9]; some monomeric forms acting as pro-oxidants [10]; the high cost of extraction
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of natural phenolics [11]; inefficiencies of chemical synthesis; and the need to enhance the bioavailability of natural phenolics, which is usually reduced due to poor solubility properties
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[12]. On the other hand, high molecular weight phenolics have been shown to have better bioactive properties compared to low molecular weight phenols [13]. Of the biocatalytic
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approaches, laccases have generated the most interest in improving antioxidant capacity through dimerization [9, 12, 14-16] or polymerization [6] to higher molecular weight
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compounds.
Laccase (E.C. 1.10.13.2) is a multi-copper oxidase that uses oxygen to abstract a single electron from a phenolic substrate to produce phenoxy radicals with water as a by-product [11]. Its ability to produce phenoxy radical intermediates has been applied in the formation of coupling
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products of therapeutic value. Although several researchers have reported the synthesis of compounds with enhanced bioactive properties using laccases (as reviewed in [11]), the work has employed only plant and fungal laccases. However, bacterial laccases would be attractive industrial catalysts due to the ease of expression in hosts such as Escherichia coli [17] and some favourable biochemical properties. For example, Machczynski et al. [18] characterized
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and overexpressed a two domain (small) laccase (SLAC) native to Streptomyces coelicolor A3(2) in E. coli. This enzyme has several attributes that make it potentially valuable for
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industrial application. It is thermally stable, recorded optimal activity at pH as high as 9.5 and
is resistant to denaturation by detergents [18]. Despite these favorable properties, there is no
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record of its application in biocatalysis. This is the first report of the application of SLAC in
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the synthesis of a dimeric antioxidant using caffeic acid as substrate. The bioactive and
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2. Materials and methods
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physicochemical properties of the synthesized compound were also investigated.
2.1 Chemicals and enzymes
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Chemicals were purchased from Sigma Aldrich (South Africa). The E. coli clone used for the expression of SLAC (wild type) was kindly provided by Prof. Gerard Canters, Leiden Institute
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of Chemistry, University of Leiden (Netherlands).
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2.2 Production of SLAC The production of SLAC was performed according to the protocol described by Prins et al. [19] with a few modifications. E. coli BL21(DE) cloned with a pET-20b(+) vector (Novagen) containing the gene coding for SLAC (pSLAC) was cultured in 2xYT media (16.0 g/L tryptone, 10.0 g/L yeast extract and 5.0 g/L NaCl) overnight. Overnight culture (1%, v/v) was inoculated
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in 2xYT media containing 100 µg/mL ampicillin and incubated at 37°C, 180 rpm until the OD600 reached 0.8. Induction for the overexpression of SLAC was achieved through the addition of isopropyl β-D-1-thiogalactopyranoside (IPTG) to a final concentration of 0.4 mM. The culture was further incubated at room temperature (22±3°C), 160 rpm for 20 h. Cells were harvested by centrifugation (5000 rpm, 10 min, 4°C) and resuspended in sodium phosphate
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buffer (pH 7.3). Cells were sonicated to release SLAC, which is produced intracellularly. The soluble fractions were collected by centrifuging (13 500 x g, 40 min, 4°C) and incubated in 1
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mM CuSO4 for 20 h at 4°C. The fractions were then dialyzed for four 1 h sessions at room
temperature (22 ± 3°C) with sodium phosphate buffer (10 mM, pH 7.3) and 1 mM EDTA added
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for the second session to chelate excess copper. The enzyme was further purified using size
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exclusion chromatography as reported in Prins et al. [19].
2.3 Enzyme activity
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SLAC activity was determined using the 2,2'-Azinobis (3-ethylbenzthiazoline-6-sulfonic acid) (ABTS) activity assay [20]. SLAC oxidation of ABTS was monitored by spectrophotometry at
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420 nm (ABTS, ε420 = 36 mM-1cm-1). The reaction mixture constituted citrate buffer (833 µL, 50 mM, pH 4.5), ABTS (110 µL, 5 mM) and laccase (57 µL). The reaction was conducted at
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25°C. One unit of laccase activity was defined as the amount of enzyme required to oxidise 1
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µmol of ABTS in 1 min at 25°C.
2.4 Oxidation of caffeic acid by SLAC Oxidation reactions were carried out in a buffer-organic solvent mixture. The reaction mixture contained caffeic acid (10 mM final concentration) and SLAC (0.64 U) in 50 mM sodium phosphate buffer (pH 7.5) (conditions determined from preliminary experiments). Optimum
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product yield was determined by comparing four organic solvents: the water miscible solvents, methanol and ethanol, and the water immiscible solvents, ethyl acetate and hexane at 50, 70, 80 and 90% organic solvent concentration (v/v) using 50 mM sodium phosphate (pH 7.5) as a buffer. The reaction was conducted for 3 h (25°C, 250 rpm). Since it was previously observed that SLAC was affected by organic solvents, 0.64 U/ mL enzyme was added at 90-min intervals
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to maintain enzyme activity.
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2.5 Analysis of products 2.5.1 Thin layer chromatography (TLC)
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Product formation was monitored by thin layer chromatography using aluminum backed silica
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gel 60 F254 TLC plates (Sigma, South Africa). A solvent system containing ethyl acetate,
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methanol, water and formic acid (8:1:1:0.02 v/v) was used as the mobile phase for the
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chromatographic separation of products. Plates were viewed under UV light at 254 nm.
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2.5.2 High Performance Liquid Chromatography (HPLC) HPLC was used for the analysis of oxidation products. For the preparation of samples, an equal
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volume of ice cold methanol was added to stop enzyme activity. The samples were kept on ice for 20 min, followed by centrifugation (13 500 x g, 10 min, 4°C) to precipitate out the enzyme
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as well as removing polymeric precipitates. Samples were filtered through 0.22 µm microfilters, transferred to clean HPLC vials and analyzed using a Waters HPLC system
A
(Massachusetts, USA). Separation was achieved on a Waters HSS T3 column, 2.1x100 mm with 1.7 um particles. A linear gradient was applied using 0.1% formic acid (solvent A) and acetonitrile (solvent B). The linear gradient started at 100% solvent A for 1 min and changed to 50% B over 22 min. It then went to 100% B after 23 min where it was held until 24.5 min,
7
followed by re-equilibration to initial conditions for 4 min. The flow rate was 0.25 mL/min and the column was kept at 60 ºC. The injection volume was 3 µL.
2.6 Purification of products Purification of the oxidation product was performed by preparatory thin layer chromatography
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glass plates (500 µm, silica gel 60 F254, Sigma, South Africa) using ethyl acetate, methanol,
water and formic acid (8:1:1:0.02 v/v) as the mobile phase. Pure bands were observed under
SC R
UV light at 254 nm and were marked with a pencil. The product bands were scraped off the plate and re-dissolved in methanol. After filtering off the silica using 0.22 µm syringe filters,
U
the product was washed with acetone, water and acetone again before being dried in clean glass
A
N
vials.
M
2.7 Characterization of products
2.7.1 Liquid Chromatography-Mass Spectrometry (LCMS)
ED
A Waters Synapt G2 quadrupole time-of-flight mass spectrometer (Massachusetts, USA) was used for LC-MS analysis. It was fitted with a Waters ACQUITY Ultra Performance Liquid
PT
Chromatography (UPLC) system and photo diode array detection. Separation was achieved
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on a Waters HSS T3 column as described in Section 2.5.2.
Data was acquired in MSE mode which consisted of a low collision energy scan (6V) from m/z
A
150 to 1500 and a high collision energy scan from m/z 40 to 1500. The high collision energy scan was done using a collision energy ramp of 30-60V. The photo diode array detector was set to scan from 220-600 nm.
8
The mass spectrometer was optimized for best sensitivity, a cone voltage of 15 V, desolvation gas was nitrogen at 650 L/hr and desolvation temperature 275ºC. The instrument was operated with an electrospray ionization probe in the negative mode. Sodium formate was used for calibration and leucine encephalin was infused in the background as lock mass for accurate
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mass determinations.
2.7.2 Nuclear Magnetic Resonance (NMR)
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For NMR analysis, 5-10 mg samples were dissolved in 0.5 mL CD3OD. 1H, 13C and 2D NMR spectra were recorded on a Bruker Avance instrument operating at 400 MHz. Chemical shifts were reported as δ values (ppm) relative to the solvent line of CD3OD (3.34 ppm for 1H; 49.0
N
U
ppm for 13C).
4-(3,4-dihydroxybenzylidene)-2-(3,4-dihydroxyphenyl)-5-oxotetrahydrofuran-3-carboxylic
M
A
acid P1, 1H NMR (CD3OD, 400 MHz): H 7.50 (d, J = 1.9 Hz, H-7'), 7.22 (d, J = 2.0 Hz, H2'), 7.09 (dd, J = 8.2, 2.0 Hz, H-6'), 6.83 (d, J = 8.2 Hz, H-5'), 6.76 (d, J = 8.1 Hz, H-5), 6.75
Hz, H-8);
13
ED
(d, J = 2.1 Hz, H-2), 6.69 (dd, J = 8.1, 2.1 Hz, H-6), 5.60 (d, J = 2.5 Hz, H-7), 3.94 (d, J = 2.3 C NMR (CD3OD, 100 MHz): C 178.2 (C-9), 175.5 (C-9'), 149.4 (C-4'), 146.8,
PT
146.7, 146.6 (C-3', C-3 and C-4); 139.9 (C-7'), 133.9 (C-1), 127.6 (C-1'), 125.7 (C-6'), 123.3
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(C-8'), 118.3 (C-6), 117.8 (C-2'), 116.6 (C-5), 116.4 (C-5'), 113.5 (C-2), 84.8 (C-7), 58.7 (C8).
A
2.8 Determination of antioxidant properties Antioxidant activities of caffeic acid and the oxidation product were determined using the following methods: Trolox equivalent antioxidant capacity (TEAC) assay and 2,2′-diphenyl-1picrylhydrazyl (DPPH) assay.
9
2.8.1 Trolox equivalent antioxidant capacity (TEAC) assay The ABTS radical scavenging activity of caffeic acid and the reaction product was determined according to the protocol by Re et al. [21] with slight modifications. ABTS•+ solution was prepared 12-16 h before use by reacting 7 mM of ABTS salt with 5.95 U SLAC and then storing in the dark until the assay was performed. The ABTS•+ solution was diluted with methanol to
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give an absorbance (Libra S21 spectrophotometer, Biochrom, Cambridge, United Kingdom)
of 0.70 ± 0.002 at 734 nm. Each sample (100 μL) prepared at different concentrations was
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mixed with 1100 μL ABTS•+ solution and the absorbance was read after 30 min incubation at 25°C.
N
U
2.8.2 2,2′-diphenyl-1-picrylhydrazyl (DPPH) scavenging effect
A
DPPH was prepared by dissolving DPPH in methanol to a final concentration of 0.025 mg/mL.
M
Test samples for both caffeic acid and reaction product were prepared in methanol at various concentrations. To conduct the assay, 0.1 mL of test sample was mixed with 3.9 mL DPPH.
ED
The mixture was shaken vigorously and incubated in the dark at 25°C for 60 min. After incubation, the absorbance was measured by spectrophotometry at 517 nm. The EC50 of the
PT
test samples were determined by plotting the percentage remaining DPPH against sample concentration. Percentage remaining DPPH was calculated by calculating the percentage of the
CC E
concentration of DPPH at T60 against DPPH concentration at T0. EC50 value is defined as the
A
concentration (mM) of substrate that results in 50% loss of DPPH• [14].
2.9 Physicochemical properties The physicochemical properties of caffeic acid and the oxidation product were compared by investigating pH stability, temperature stability and photo-stability. The physicochemical
10
properties were studied in hydro-alcohol solutions due to the solubility characteristics of caffeic acid, which is not soluble in aqueous medium.
2.9.1 pH stability pH stability was investigated at pH 2.2, 5.5 and 7.5 following the protocol described by
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Amendola et al. [22]. The test samples were dissolved in buffer-alcohol solution (85:15 v/v) using 50 mM citrate-phosphate buffer at pH 2.2, 5.5 and 7.5 to a final concentration of 0.5 mM.
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The test flasks were covered with aluminum foil to eliminate light interference and were kept
U
at 4°C for 30 days. Samples were taken for HPLC analysis at 5-day intervals.
N
2.9.2 Temperature stability
A
Storage temperature stability was analyzed for 30 days at 4°C, 25°C and 37°C according to the
M
method adopted and modified from Amendola et al. [22]. The test samples were dissolved in a hydro-alcohol solution of water and methanol (85:15 v/v) to a final concentration of 0.5 mM.
ED
The test flasks were covered with aluminum foil and kept at 4°C, 25°C and 37°C for 30 days.
CC E
PT
Samples were taken for HPLC analysis at 5-day intervals.
2.9.3 Photo-stability
A
Photo-stability was investigated using fluorescent light. The test samples were prepared by dissolving the compounds in a hydro-alcohol solution of water and methanol (85:15 v/v) to a final concentration of 0.5 mM. The test flasks were exposed to fluorescent light (Osram L18W/640 with a luminous flux of 1200 lm, 18 W and luminance of 508 542 lx) for 30 days. Samples were taken for HPLC analysis at 5-day intervals.
11
3. Results and discussion 3.1 Structural characterization of SLAC oxidation product The SLAC catalyzed oxidation of caffeic acid resulted in the formation of one major product designated as P1 (tR=5.29), (Fig. 1a). LC-MS analysis conducted in the negative mode
IP T
indicated that the product was a dimer m/z 357, [M] 358.07 (Fig. 1b). The in situ fragmentation pattern showed dominant signals at m/z 313 (– 1 carboxyl group) and m/z 269 (– 2 carboxyl
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groups). Substrate conversion to the dimer was monitored by HPLC over a 12 h oxidation
period. It was observed that at 6 h, optimum dimer was produced (yield 32.8%), after which
U
the yield started decreasing. This may be due to a combination of non-specific side reactions
N
as well as polymerization reactions, usually associated with laccase-mediated radical reactions
A
[11].
M
The NMR results of P1 indicated that it is a dimer of caffeic acid through a β-β linkage (Fig.
ED
2). Since the molecule was not symmetrical, the resonances for each of the aromatic protons in the rings could be seen separately. Ortho coupling was seen for H-5 and H-6 at H 6.76 (d,
PT
J = 8.1 Hz) and H 6.69 (dd, J = 8.1, 2.1 Hz) and H-5' and H-6' at H 6.83 (d, J = 8.2 Hz) and
CC E
H 7.09 (dd, J = 8.2, 2.0 Hz). H-6 and H-6' were meta coupled to H-2 and H-2' respectively at H 6.75 (d, J = 2.1 Hz) and 7.22 (d, J = 2.0 Hz). The olefinic H-7' resonance was deshielded at H 7.50 appearing as a doublet with J = 1.9 Hz due to long range coupling with H-8, which
A
appeared as a triplet at H 3.94 (J = 2.3 Hz) due to the double doublet coalescing. H-7 was seen at H 5.60 as a doublet with J = 2.5 Hz. The small J value of H-7 and H-8 indicated that the dihedral angle was approaching 90. The carbon resonances for the two carbonyl groups could be seen at C 178.2 (C-9) and 175.5 (C-9'), assigned due to HMBC correlations with H7 and H-8 (for C-9) and H-7' (for C-9'). C-4' was distinctly separated from the other aromatic 12
oxygenated resonances at C 149.4, and assigned due to HMBC correlations with H-2', H-6' and H-5'. The other three aromatic C-O resonances, C-3', C-3 and C-4 appeared at C 146.8, 146.7 and 146.6 and can be interchanged. Both C-1 and C-1' at C 134.0 and 127.6 were identified by HMBC correlations with H-5 and H-5' respectively. The remaining singlet carbon
HMBC spectra, please refer to Fig. S1-Fig. S8 (supplementary data).
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resonance at C 123.3 was assigned to C-8'. For details of 1H and 13C NMR, COSY, HSQC and
SC R
The dimer has been reported in culture broth of Phellinus sp. and was identified as phellinsin A [23] and has also been previously synthesised from Lycopus europaeus catechol oxidase [24]
U
and the cis form from Momordica charantia peroxidase-catalysed processes [25]. However,
N
the SLAC-catalysed process may potentially be the cheapest method compared to previously
A
described protocols [24, 25] which employed plant-derived biocatalysts thus making the
M
processes potentially cost-intensive. In addition, the dependence of peroxidases on H2O2 makes
ED
the synthetic process expensive as well as a health hazard [26].
Caffeic acid is one of the putative lignin monomers. Although β-O-4 linkages are the
PT
predominantly formed linkages in polymer formation in nature [27], it has been observed that
CC E
dimerization usually results in β-5 and β-β linkages [28]. The suggested mechanism for the formation of the β-β dimer is shown in Fig. 3 [24, 25]. A radical is formed from the SLAC oxidation of the hydroxyl group occupying the para position of caffeic acid. The lone electron
A
on the subsequent radical can occupy different positions through resonance stabilization. C–C linkage of two phenoxy radicals with unpaired electrons at β positions results in the formation of the β-β caffeic acid dimer, phellinsin A. A similar reaction pathway was proposed by Adelakun et al. [9] for the formation of a laccase catalyzed β-β ferulic acid dimer, suggesting that the β-β linkage is one of the most preferred radical coupling positions for phenolic acids. 13
3.2 Effect of solvent on the formation of phellinsin A Laccase catalysis of phenolic compounds is usually performed in buffer-organic solvent reaction systems. This is because of the sparse solubility of most phenolic compounds (including caffeic acid) in aqueous media, It has also been observed that increasing solvent
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concentrations help minimize polymerization [9, 14]. For this study, four solvents, methanol, ethanol, ethyl acetate and hexane were investigated at four different concentrations (50, 70, 80
SC R
and 90%) to determine the best solvent conditions for the production of phellinsin A.
U
Results indicated that there was an increase in product formation as the polarity of the solvent
N
decreased; Hexane> ethyl acetate > methanol > ethanol (Fig. 4). There was a general increase
A
in phellinsin A formation as the solvent concentration increased from 50% to 80%, after which
M
a sharp decline in product formation at 90% co-solvent, was observed. Although 80% hexane resulted in the highest yield of phellinsin A, ethyl acetate, which yielded the second highest
ED
concentration of phellinsin A, offered a cleaner reaction with less side reactions, thus making it easier for purification. Therefore, scale up reactions were carried out using 80% ethyl acetate
PT
as co-solvent. There was a significant loss of enzyme activity as the oxidation reaction progressed; by 90 minutes, 94.7% of activity was lost (Fig. S9, supplementary data). Therefore,
CC E
0.64 U enzyme was periodically added at 90-minute intervals for the reaction to proceed. Organic solvents were reported to affect laccase activity by direct interaction with the enzyme
A
molecule or through modifying the thermodynamic water activity of the enzyme [29, 30].
3.3 Bioactive properties Antioxidant capacity of phellinsin A was compared to caffeic acid using DPPH and TEAC assays. The DPPH and TEAC assays are based on the radical scavenging ability of an
14
antioxidant to reduce DPPH• and ABTS•+ to their more stable states (1,1-diphenyl-2-picryl hydrazine and 2,2'-azino-bis-3-ethylbenzthiazoline-6-sulphonic acid, respectively) [31]. Both assays showed an improved antioxidant capacity of the dimer; 1.5-fold and 1.8-fold increase in antioxidant capacity for DPPH and TEAC assays, respectively compared to caffeic acid (Table 1). Studies of structure-activity relationships (SAR) of antioxidants have attributed the
IP T
antioxidant capacity of a compound to factors such as the availability of electron donating
groups attached to the aromatic ring [11, 32] as well as ability to form stable radicals. Caffeic
SC R
acid contains two hydroxyl groups, as well as an unsaturated C=C bond in the side chain.
Phellinsin A, on the other hand has four hydroxyl groups as well as the unsaturated C=C bond,
U
which explains its improved antioxidant capacity. It has been reported that effective
N
antioxidants are able to form stable phenoxy radicals through either hydrogen bonding [33] or
A
delocalization of lone pair electrons into the aromatic ring [11, 34]; since the structure of
M
phellinsin A does not consist of adjacent aromatic rings which enable expanded electron delocalization [32], enhanced antioxidant activity may be attributed to increased number of
ED
hydroxyl groups which facilitates radical stability through hydrogen bonding.
PT
3.4 Physicochemical properties
Besides enhanced bioactivity, physicochemical profiles also qualify a compound for
CC E
commercial development [35]. Conditions such as pH, temperature, air and light were reported to affect stability of phenolic compounds [36]. Despite significant progress being made in
A
laccase synthesis of bioactive compounds, comprehensive characterization of products has been neglected. A literature survey of all laccase mediated coupling products synthesized since 2000 showed that only one article investigated physicochemical properties of the coupling products [37]. The extent of product characterization has narrowly focused mostly on bioactivity, however, an extensive analysis of coupling products is required for the constructive
15
progression of laccase biotransformation technology. Therefore, the physicochemical properties of phellinsin A were investigated.
3.4.1 Phellinsin A solubility Caffeic acid is not soluble in aqueous media, therefore it was dissolved in at least 50% methanol
IP T
for complete solvation at room temperature. However, phellinsin A was completely soluble in aqueous media at room temperature. This may be attributed to the increased number of
SC R
hydrogen bond interactions between the compound and water [38]. Solubility is an important factor in drug development since it can give a theoretical indication of the bioavailability of a
N
U
compound in vivo [39].
A
3.4.2 Temperature stability
M
Phellinsin A showed significant instability when incubated at body temperature (37°C) in a hydro-alcohol solution of water and methanol (85:15 v/v). In the first 5 days of incubation,
ED
there was 63.6% loss of phellinsin A and by day 20, it was completely degraded (Fig. 5). At storage temperatures (4°C and 25°C), phellinsin A exhibited relative stability, recovering
PT
85.5% and 39.3% of the compound, respectively after 30 days of incubation. These results have shown that the concentration of phellinsin A decreased with increasing temperature. Despite
CC E
phellinsin A exhibiting significant instability at body temperature over prolonged periods, it can still be an effective antioxidant. Pharmacokinetic studies have shown that most phenolic
A
compounds are metabolized within 24 h, after which the plasma concentration reaches basal levels [40, 41]. At body temperature, about 60-80% of phellinsin A was retained during the first 24 h (Fig. 5 and S10), when the compound is expected to be metabolized. Therefore, phellinsin A can be a useful antioxidant since it has shown relative stability at storage temperature (4°C) while exhibiting significant stability within the metabolism period. On the
16
other hand, caffeic acid showed a contrasting trend; increasing stability as temperature increased. Caffeic acid exhibited the highest stability at 37°C and retained 76.5% of compound by day 30 (Fig. 5). Previous studies on temperature–antioxidant activity of phenolic acids determined in triacylglycerols of sunflower oil showed that cinnamic acid derivatives including caffeic acid were more effective antioxidants at 90°C than at 22°C, owing to their higher
IP T
stability at high temperatures [34, 42].
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3.4.3 pH Stability
pH stability was investigated considering potential applications and the related environments.
U
For instance, if phellinsin A is considered for the development of a nutraceutical, it would be
N
interesting to observe the effect of gastrointestinal pH (pH 2.2 for the stomach and pH 7.5 for
A
alkaline intestinal conditions) [43]; or if phellinsin A could be an ingredient in development of
M
topical products, then its stability at skin pH (pH 5.5) [44] is important. pH stability tests were performed at 4°C to reduce any microbial contamination which is common especially at acidic
ED
buffer conditions [22]. Results showed that the stability of the SLAC synthesized phellinsin A is pH-dependent. It showed remarkable stability in acidic conditions but was extremely
PT
unstable under alkaline conditions. At pH 5.5, it was completely stable; no compound loss was observed after monitoring for 30 days. Therefore, the product may be considered for
CC E
application in the development of skincare products. At pH 7.5, phellinsin A was completely degraded within 5 days of incubation (Fig. 6). Similarly, previous studies on stability profiles
A
of phenolic compounds have demonstrated that phenolic compounds such as caffeic acid, gallic acid, flavonoids, chlorogenic acid and green tea catechins are pH-sensitive; exhibiting greater stability in acidic pH and are unstable under alkaline conditions [45, 46, 47]. It has been suggested that the effect of pH on stability of a phenolic compound is related to the resonance stabilization of its subsequent phenoxide ions which are mostly stable in acidic conditions than
17
in alkaline medium [47]. It was therefore not surprising the dimer with a higher electron density was more stable than the monomeric unit.
3.4.4 Light stability Light significantly impacted the stability of phellinsin A with only 11.5% of the compound
IP T
recovered after 30 days of incubation (Fig. 7). Photo-stability tests were carried out in conditions that, to a large extent, simulated normal storage environments at room temperature,
SC R
oxygen and under normal fluorescent light. Photodegradation of phenolic compounds is induced when valence band electrons are excited in the presence of oxidants such as molecular
U
oxygen causing a compound to be more susceptible to oxidative degradation [48]. The
N
susceptibility of phellinsin A to degradation was not surprising since previous investigations
A
have shown that polyphenols are generally light sensitive [49]. Therefore, the compound may
M
be most suitable for application in topical product formulations since it has shown the highest stability at pH 5.5 and significant light stability (approximately 80% retained within 24 h, Fig
ED
S10) during the period in which it is expected to be metabolized. Due to the susceptibility of phellinsin A to light as well as high temperatures, the best storage conditions will be at 4°C in
CC E
PT
packaging material that excludes light (such as amber bottles).
4. Conclusion
This study has reported the application of SLAC as a biocatalyst in the synthesis of antioxidants
A
for the first time. A β-β caffeic acid dimer, phellinsin A, was successfully synthesized and its bioactive and physicochemical properties elucidated. The dimer exhibits improved solubility properties and enhanced antioxidant properties compared to the parent compound, caffeic acid. This study indicates that SLAC could be a potential biocatalyst for the production of value added antioxidant products.
18
5. Acknowledgements We express our sincere gratitude to Ms Devrani Naicker (Durban University of Technology) for her assistance with chromatography analysis, Mr Bibhuti Ranjan (Durban University of Technology) for his assistance with enzyme purification and Ms Thandokazi Andiswa for
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assistance with NMR. We are grateful to the National Research Foundation (NRF) for financial
assistance (Grant 105889, 112099 and 93362) and to the Council for Scientific and Industrial
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Research (CSIR) for student (Blessing Nemadziva) bursary support. Any opinion, findings, and conclusions or recommendations expressed in this material are those of the author(s), and
N
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therefore, the NRF does not accept any liability in regard thereto.
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Figure legends Fig 1a. HPLC chromatogram of caffeic acid oxidation product (tR = 5.29). Fig 1(b) mass spectra of SLAC oxidation product (P1). Fig 2. The β-β linked dimer (phellinsin A) formed during SLAC-catalyzed oxidation of
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caffeic acid. Oxidation reactions were carried out in buffer-organic solvent systems using 50 mM sodium phosphate buffer (pH 7.5) and varying concentrations of selected organic
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solvents. Dimer was purified using preparatory thin layer chromatography.
Fig 3. Proposed reaction pathway for the SLAC-catalysed synthesis of phellinsin A. A C–C bond is formed between two β-positioned caffeic acid radicals. The intermediate compound
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undergoes rearrangement reactions to form the β-β dimer, phellinsin A.
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Fig 4. Effect of organic solvents on the synthesis of phellinsin A. Caffeic acid was oxidized
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by SLAC in either biphasic or monophasic reaction systems at varying concentrations of the test organic solvent (ethanol, ethyl acetate, hexane or methanol). Phellinsin A formation was
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replicate determinations.
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monitored by HPLC analysis. All results are means ± standard deviation (SD) of three
Fig 5. Stability profile of caffeic acid and phellinsin A at 4°C, 25°C and 37°C. Test samples
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were prepared in hydro-alcohol solutions and incubated in the dark at the investigated temperature for 30 days. All results are means ± standard deviation (SD) of three replicate
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determinations.
Fig 6. Stability profile of caffeic acid and phellinsin A incubated in citrate-phosphate buffer at pH 2.2, 5.5 and 7.5. Test samples were prepared in buffer-alcohol solutions at varying pH and were incubated in the dark at 4°C for 30 days. All results are means ± standard deviation (SD) of three replicate determinations.
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Fig 7. Photo-stability profile of caffeic acid and phellinsin A. Test samples were prepared in hydro-alcohol solutions and were exposed to fluorescent light for 30 days. All results are
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means ± standard deviation (SD) of three replicate determinations.
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Table 1 Antioxidant activity of caffeic acid and phellinsin A. Standard assays were used to determine activity. All results are means ± standard deviation (SD) of three replicate determinations. Molecular weight (g.mol-1) EC50 DPPH (mM)
TEAC valuea (mM)
Caffeic acid
180.16
0.44 ± 0.008
2.17 ± 0.060
Phellinsin A
358
0.29 ± 0.004
3.88 ± 0.004
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a
Compound
TEAC value is defined as the concentration of Trolox (mM) with an antioxidant potential
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equivalent to 1 mM of compound under investigation [14]. EC50 value is defined as the
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concentration (mM) of substrate that results in 50% loss of DPPH• [14].
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