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Simple adsorption of Candida rugosa lipase onto multi-walled carbon nanotubes for sustainable production of the flavor ester geranyl propionate
Accepted Manuscript Title: Simple Adsorption of Candida rugosa Lipase onto Multi-walled Carbon Nanotubes for Sustainable Production of the Flavor Ester Geranyl Propionate Author: NurRoyhaila Mohamad Nor Aziah Buang Naji A. Mahat Joazaizulfazli Jamalis Fahrul Huyop Hassan Y. Aboul-Enein Roswanira Abdul Wahab PII: DOI: Reference:
S1226-086X(15)00352-4 http://dx.doi.org/doi:10.1016/j.jiec.2015.08.001 JIEC 2605
To appear in: Received date: Revised date: Accepted date:
6-1-2015 28-6-2015 2-8-2015
Please cite this article as: N.R. Mohamad, N.A. Buang, N.A. Mahat, J. Jamalis, F. Huyop, H.Y. Aboul-Enein, R.A. Wahab, Simple Adsorption of Candida rugosa Lipase onto Multi-walled Carbon Nanotubes for Sustainable Production of the Flavor Ester Geranyl Propionate, Journal of Industrial and Engineering Chemistry (2015), http://dx.doi.org/10.1016/j.jiec.2015.08.001 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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Simple Adsorption of Candida rugosa Lipase onto Multi-walled Carbon Nanotubes for
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Sustainable Production of the Flavor Ester Geranyl Propionate
3 NurRoyhaila Mohamad1, Nor Aziah Buang1, Naji A. Mahat1, Joazaizulfazli Jamalis1, Fahrul
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Huyop2, Hassan Y. Aboul-Enein3 and Roswanira Abdul Wahab1,*
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Department of Chemistry, Faculty of Science, Universiti Teknologi Malaysia, 81310 UTM Johor
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Bahru, Johor, Malaysia 2
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Department of Biotechnology and Medical Engineering, Faculty of Biosciences and Medical
Engineering, Universiti Teknologi Malaysia, 81310 UTM Johor Bahru, Johor, Malaysia
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Pharmaceutical and Medicinal Chemistry Department, Pharmaceutical and Drug Industries Research Division, National Research Centre, Dokki, Cairo 12311, Eqypt
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*Corresponding author:
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Roswanira Abdul Wahab, Department of Chemistry, Faculty of Science, Universiti Teknologi
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Malaysia, 81310 UTM Johor Bahru, Johor, Malaysia, Telephone: +607 5534148; Fax +607
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5566162; Email: [email protected]
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Highlights: 1. The CRL−MWCNTs improved yield of geranyl propionate by 2−fold
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2. The CRL−MWCNTs showed a 2−fold improved thermal stability
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3. CRL−MWCNTs as cheap biocatalyst to produce geranyl propionate
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Abstract
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In this study, geranyl propionate was enzymatically synthesized from geraniol and propionic acid
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using Candida rugosa lipase immobilized on acid functionalized multi−walled carbon nanotubes.
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The efficiency of the CRL−MWCNTs biocatalysts to catalyze the esterification production of
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geranyl propionate (solvent log P, alcohol:acid molar ratio and thermal stability) was compared
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with the free CRL for parameters. The use of CRL−MWCNTs in n-heptane (log P 4.0) and
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alcohol:acid molar ratio of 5:1 resulted in a 2−fold increased conversion frequency as compared to
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the free CRL for the production of geranyl propionate, in addition to a noteworthy 2−fold enhanced
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thermal stability.
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Keywords: Candida rugosa lipase; biocatalysts; immobilization; multi−walled carbon nanotubes;
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esterification; geranyl propionate
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1.
Introduction Terpene esters of short-chain fatty acids are essential oils with significant applications in
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food, cosmetic and pharmaceutical industries as flavors and fragrances; geraniol and citronellol,
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being the two foremost substances [1]. Conventionally, these esters are produced via various
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methods viz. chemical synthesis, extraction from natural products as well as fermentation [2].
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Nonetheless, chemical synthesis is often the preferred choice to mass produce such esters even
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though the method entails the use of strong acid catalyst and hazardous chemicals, considerably
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lengthy reaction time while providing low conversion rate [1]. Additionally, the downstream
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processing of such method incurs tedious separation processes, involving extreme exposure of
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toxicants as well as superfluous harmful by−products [3]. Considering the numerous disadvantages
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that are typically linked to the current chemical approach for producing these terpene esters [4,5],
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alternative methods that are more environmentally friendly, economically attractive and improve
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reaction yield would be significant.
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In addition, the approval of REACh (Registration, Evaluation, Authorization and Restriction
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of Chemicals) regulation by the European Parliament and of the Council as well as regulation and
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the environmental restrictions imposed by US, Japan and E.U. have opened up new challenges for
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utilizing Green Chemistry philosophy in industrial processes [6]. The philosophy emphasizes on the
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industrial usage of chemicals acquired from biomass, the use of green solvents and more sustainable
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industrial processes [7]. The key aspect of ‘‘green’’ industrial processes is the effective
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incorporation of catalytic technologies (chemical or enzymatic) into a general organic synthesis
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scheme. In this sense, biocatalysis offers many attractive features in the perspective of green
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chemistry [7]. The biocatalytic route embraces gentle reaction conditions (physiological pH and
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temperature) and environmentally friendly catalysts (an enzyme or a cell) that display high
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activities and chemo-, regio− and stereoselectivities in multifunctional molecules. Additionally, the
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use of biocatalysts usually circumvents the need for functional group activation and therefore,
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affords processes that are shorter, produce less waste and economically smarter [5, 8].
82 Lipases (triacylglycerol ester hydrolysis EC 3.1.1.3) are medicinally, industrially and
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commercially pertinent biomolecules that have acquired popularity as biocatalysts [9] over
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inorganic biocatalysts for reasons such as high specificity, efficient reaction rate, non-toxicity and
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biodegradability, reproducibility under normal laboratory conditions [10]. Also, they are able to
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convert a large number of substrates with their high stereospecificity [11]. In this context, Candida
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rugosa lipase (CRL) remains one of the most versatile biocatalysts due to its high activity and broad
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specificity in reaction medium [10]. In view of (a) the poor stability of the free form CRL, (b) low
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activity in organic solvents, (c) high tendency of deactivation in prolonged exposure to high
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temperature and extreme pH [10], immobilization of CRL onto various nanosupports offer many
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benefits; one of them being large surface-to-volume-ratio [12-13]. The use of nanosupports such as
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multi−walled carbon nanotubes may be a practical solution [14−15] for improving stability and
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activity as well as extending the reaction life of the enzyme.
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Enzyme immobilization confers the advantages of structural stability, improved activity,
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specificity and selectivity as well as reduced inhibition [16]. Moreover, immobilization of enzyme
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facilitates facile separation of the enzyme from product, hence simplifying enzyme application for a
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reliable and efficient reaction technology. Since enzymes are expensive and required in large
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quantities for industrial purposes [17], immobilization of such enzymes onto solid support matrices
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may prove beneficial in permitting reusability of the biocatalysts, often an essential prerequisite for
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establishing an economically viable enzyme catalyzed process [18]. In this context, the physical
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adsorption method may have higher commercial values than other methods [19] because it remains
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as one of the simplest and cheapest immobilization methods available and importantly, in most
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cases the enzyme productivity remains unaffected [20]. 4 Page 4 of 34
Immobilized enzymes are currently the subject of considerable attention for their numerous
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advantages over soluble enzymes. Consequently, the current study was intended at investigating the
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potential use of physically adsorbed CRL− multi-walled carbon nanotubes biocatalysts
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(CRL−MWCNTs) as alternative economical and sustainable biocatalysts for the enzymatically
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based esterification production of geranyl propionate. Performance of the CRL−MWCNTs
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biocatalysts for catalyzing production of geranyl propionate was compared with that of the free
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CRL. Both forms of biocatalysts were evaluated for the effects of solvent log P, molar ratio
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alcohol:acid as well as thermal stability in rendering the highest conversion of geranyl propionate.
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2.
Materials and methods
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2.1
Chemicals
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MWCNTs were received as-synthesized from the Department of Chemistry, Faculty of
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Science. Lipase from Candida rugosa (EC 3.1.1.3), Type VII (1410 U mg-1) and Bradford reagent
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were procured from Sigma Aldrich Co. (St. Louis, USA). For the esterification reactions, substrates
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geraniol (98%) and propionic acid (99%) were also purchased from Sigma Chemical Co. (St. Louis,
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USA). Solvents, diethylether, benzaldehyde, toluene, n-heptane and decane were purchased from
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Merck (Darmstadt, Germany). Nitric acid, HNO3 65% (v/v); hydrochloric acid, HCl 37% (v/v) and
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sulphuric acid, H2SO4 95−97% (v/v) from Merck (Darmstadt, Germany) were used as acid solvent
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systems in the purification and oxidation processes of MWCNTs. Other chemicals, namely sodium
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hydroxide (NaOH) and phosphate buffer (Merck, Germany) were used without further purification.
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Distilled water was produced in the lab and was used in all experiments. All chemicals were of
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analytical grade unless specified otherwise.
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2.2
Purification and functionalization of MWCNTs For the purification process, the synthesized−MWCNTs (0.5 g) were transferred into a 100 5 Page 5 of 34
mL flask that contained 4 M HCl (20 mL) and refluxed with stirring at 80°C for 5 h. After cooling
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to room temperature, the liquid was decanted, the MWCNTs washed with distilled water until no
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remaining acid was detected and dried in an oven at 60°C for 24 h. The purified MWCNTs were
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refluxed at 120°C by stirring in a mixture of concentrated HNO3 and H2SO4 (1:3 v/v) for 24 h. After
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cooling to room temperature, the mixture containing the acid functionalized MWCNTs
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(F−WMCNTS) was decanted, washed with distilled water until no remaining acid was present and
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dried at 60°C for 24 h.
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2.3
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Adsorption immobilization of lipase and its characterization
The F−MWCNTs (10 mg/mL) were first sonicated to homogeneity in aqueous buffer (pH 7)
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for 30 min. Then, the MWCNTs were transferred into a 50 mL flask of phosphate buffer (50 mM,
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solution pH 7) that contained CRL (10 mg/mL) and incubated at 20°C with constant stirring at 150
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rpm. Following 3 h incubation period, the flask containing CRL MWCNTs was cooled to room
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temperature and stored at 4°C for 24 h. After the stipulated period, the unbound proteins were
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removed by washing with phosphate buffer (pH 7) until no evidence protein was detected in the
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supernatant. The supernatant was analyzed for its protein content by Bradford method [21].
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2.3.1 Fourier Transform Infrared Spectroscopy (FTIR) A ratio of 1:100 mass of sample was ground thoroughly with potassium bromide and the
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resulting powder was pressed into a transparent pellet by a hydraulic press. The FTIR spectra
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obtained using a BOMEM spectrophotometer (Quebec, Canada) in transmission mode between 400
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and 4000 cm-1 at a resolution of 4 cm were analyzed.
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2.3.2 Thermal Gravimetric Analysis (TGA)
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Thermal gravimetric analysis (TGA) for F−MWCNTs and CRL−MWCNTs was conducted
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in a compact ceramic alumina crucible and heated from 30°C to 800°C under a nitrogen atmosphere
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using a Thermogravimetric Analyzer 4000 (Perkin Elmer) at a heating rate of 10°C/min.
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2.3.3 Transmission Electron Microscopy (TEM) and Field Emission Scanning Electron Microscopy (FESEM)
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Morphological analysis for the MWCNTs, F−MWCNTs and CRL−MWCNTs were
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determined using transmission electron spectroscopy (TEM) and field emission scanning electron
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microscopy (FESEM). The measurement was conducted by heating the sample (5.0−5.2 mg) in a
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platinum pan (5mm in diameter), under oxygen flow in temperature range of 35−1000oC, ramped at
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the rate of 10oC min-1. TEM analysis was executed using electron source as W-emitter and LaB6
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that operated at an accelerating voltage of 200 Kv. The analysis employed an objective lens (S-
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Twin) having a point resolution of 2.0 nm or better with a 25x to 7,500,200x or higher
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magnification and a single tilt holder with LCD camera. The individual sample (0.5 mg) of
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MWCNTs, F−MWCNTs and CRL−MWCNTs was ultrasonicated for 2 h in deionized water and a
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drop of each sample was positioned on a copper grid, observed after drying in vacuum. For the
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analysis of FESEM, a sample which consisted of a drop of the dispersed MWCNTs was placed on a
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silicon wafer and allowed to dry in vacuum for 30 min prior to analysis using JEOL JEM−6700F.
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Determination protein loading and lipase activity
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The concentration of protein in the enzyme solution, prior and post immobilization, in the
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washing buffer was quantified by the Bradford method using BioRad protein dye reagent
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concentrate and bovine serum albumin (BSA) as the protein standard [21]. Different concentrations
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of BSA were prepared using a BSA stock solution (0.1 mg in 10 mL of deionized water) and the
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absorbance was measured at 595 nm with a spectrophotometer (HITACHI U-3210) using 7 Page 7 of 34
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preparations without BSA as the blank. Analyses were performed in triplicate. The activity of CRL−MWCNTs was determined according to the method suggested by
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Langone and Sant'Anna (1999) [22]. Lipase activity was quantified by the consumption of lauric
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acid in the esterification reaction with glycerol (lauric acid/glycerol, 1:3) at 50oC, with the enzyme
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concentration of 5 mg/mL. One esterification unit of the lipase (U) was defined as 1 μ mole of
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lauric acid consumed/min and the CRL−MWCNT biocatalysts had an esterification activity of 792
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U mg-1. All determinations were performed in triplicate.
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2.5
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Production of geranyl propionate by free CRL and CRL−MWCNTs A typical esterification reaction was executed in propionic acid (0.25 M) and geraniol (1.25
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M) dissolved in n-heptane (unless specified otherwise) as solvent in a 100 mL round bottom flask.
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The free CRL (10 mg/mL) or CRL−MWCNTs (5 mg/mL) were reacted in a 30 mL volume of this
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mixture, respectively. The reaction mixture was stirred with continuous stirring at 200 rpm in a
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paraffin oil bath. Aliquots (1 mL) of the reaction mixture were withdrawn periodically for 12 h and
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analyzed by titration with NaOH (0.05 M) with phenolphthalein as indicator. The geranyl
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propionate obtained was expressed in terms of percent conversion i.e. percent of propionic acid
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converted versus the total acid in the reaction mixture. Each measurement was performed in
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triplicate and the standard error was calculated. The percent conversion was calculated according to
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the prescribed equation [1].
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% Conversion = ((Vo − Vt) /Vo) x 100
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whereby, Vo is volume of NaOH at initial time (t = 0) and Vt is volume of NaOH at stipulated hour
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(t = t1, t2, t3 ...).
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2.5.1 Effect of Solvents (Log P)
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Stability of biocatalysts in organic solvents is an important characteristic to be considered
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for a biosynthetic reaction. The effect of solvents in the enzymatic production of geranyl propionate 8 Page 8 of 34
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was determined using solvents of increasing log P: diethylether (0.8), benzaldehyde (1.48), toluene
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(2.50), n-heptane (4.0) and decane (6.0), respectively. The solvents were added to the reaction
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mixtures containing the free CRL and CRL−MWCNTs at a molar ratio of geraniol:propionic acid
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of 5:1, unless specified otherwise.
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It has been described that relative proportions of the various substrates in a reaction mixture
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is a factor that defines the physical and chemical properties of a reaction system and subsequently
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influences the yield of product. The reaction mixtures were prepared at increasing molar ratio of
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geraniol: propionic acid (molar ratio = 1:1, 2:1, 3:1, 4:1 and 5:1).
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The thermal stability of CRL−MWCNTs was examined by incubating the biocatalysts in
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sealed vials for 1 h at various temperatures ranging from 50 to 80°C, at increasing intervals of 10oC
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each. After the stipulated period, the CRL−MWCNTs were left to cool to room temperature. Such
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evaluation was attempted by introducing the thermally treated CRL−MWCNTs into the reaction
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mixture that consisted of propionic acid and geraniol dissolved in n-heptane and incubated at 40°C
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with constant stirring. The relative activities were determined and expressed as percentage of the
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enzyme activity at varying temperatures.
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Analysis of Geranyl Propionate using Gas Chromatography (GC) and Nuclear
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Magnetic Resonance (NMR) Spectroscopy
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Identification of geranyl propionate in liquid samples was carried out by Gas
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Chromatography (GC) (Agilent Technologies 6890N) equipped with flame ionization detector
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using a capillary column of fused silica INOWAX (30 m × 250 μm × 0.25 μm). Helium was used as
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the carrier gas (1.0 mL/min) at constant pressure, the detector at 1.0 kV, split mode (1:20), and the 9 Page 9 of 34
injector whose temperature was maintained at 250°C. The temperature program used was as
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follows: 50°C for 1 min; increment of 5°C/min up to 300°C; and the temperature was kept constant
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for 1 min. Each sample was filtered prior to GC analysis. Nuclear magnetic resonance spectra
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(NMR) were run on a Bruker Avance III-HD 700 MHz. spectrometer. CDCl3 was employed as the
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solvent for all experiments. Samples were obtained by dissolving about 0.2 g of methyl oleate in
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about 0.55 mL of CDCl3 and transferred to a standard 5 mm tube. Standard acquisition parameters
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were used for all samples.
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Result and Discussion
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3.1
Rationale for the functionalization of MWCNTs
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Stirring the MWCNTs in acid mixture of H2SO4 and HNO3, introduces the polar groups
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(COO-) to the non-polar surface and tubular ends of the MWCNTs supports. This is to ensure that
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the surfaces of the MWCNTs are primed for interaction with other polar moieties (NH2, O−H)
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present on the CRL protein. The COO- moiety on the surface of MWCNTs anchor to the CRL
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protein by attractions between the oppositely charged carboxyl moiety on the MWCNTs and the
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hydrogen of the back-bone or side-chain of polar amino acids present on the outer surface of the
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CRL protein [23]. On the other hand, poor attachment of the CRL proteins to the non-polar non-
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functionalized MWCNTs was observed, an inference made from the high protein concentrations in
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the phosphate buffer washings after immobilization of the free CRL. Conversely, the physical
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adsorption of CRL to the F−MWCNTs was reasonably strong as the amount of detached proteins in
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the reaction mixtures was not observed or negligible following the 6 h of stirring. The
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CRL−MWCNTs recorded a lower enzyme activity than the free CRL corresponding to 792 U mg-1.
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However, the CRL−MWCNTs biocatalysts were anticipated to be more stable and undergo a slower
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decline in enzymatic activity when utilized under extended reaction period and high temperature.
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Also, the acid functionalized CRL−MWCNTs were observably well dispersed when stirred in the
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reaction vessel as compared to the non-functionalized ones. The facile dispersability of the
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CRL−MWCNTs is suggestive of proper mixing between the biocatalysts with the substrates, hence,
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increases the propensity of effective enzyme−substrate collision to improve the esterification yield
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[23].
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3.2
Characterization of Immobilized Lipase (CRL−MWCNTs)
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3.2.1 Fourier Transform Infra−Red Spectroscopy (FT-IR)
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FTIR is principally employed as a qualitative technique to assess the presence of functional
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groups in a particular compound. The FT−IR spectra acquired for the as-synthesized MWCNTs,
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F−MWCNTs and CRL−MWCNTs as well as the resultant wavenumbers are presented in Table 1.
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The emergence of a broad band at 3428.01 cm-1 corresponding to the O−H stretching of the surface
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group of the as-synthesized MWCNTs may be attributable to ambient atmospheric moisture that
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was firmly bound to the MWCNTs [24]. A weak band which appeared at 1630.82 cm-1 in the
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spectrum of as-synthesized MWCNTs is associated with the existence of conjugated C=C bonds
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[25]. Following oxidation, the presence of a C−O stretching due to the successful oxidation of sp2
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hybridized carbon in the as-synthesized MWCNT to sp3 [26] is established by the emergence of a
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new peak at 1112.73 cm-1 in the spectra of the F−MWCNTs. Correspondingly, the O−H stretch
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moved from 3428.01 cm-1 to 3445.22 cm-1, strongly implying the development of covalent bonds
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between the O−H and the sidewalls of the MWCNTs [27]. In addition, the C=O stretching vibration
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of the carboxylic group represented by a peak (1634.70 cm-1) was observed attributable to
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adaptation of additional single bond character through resonance by the C=O of the introduced
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acidic surface groups. Conjugation of O=C with the C=C of the MWCNTs had resulted in the
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formation of a predominantly C−O functional group, an observable fact that eventually led to the
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decline in the vibration frequency of the C=O group in the spectra of F−MWCNTs. Furthermore,
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the appearance of a new peak at 1202.46 cm-1 is attributed to the C−O of the COOH moiety [28].
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In addition, a band (1634.70 cm1) for C=O stretching vibration of the carboxylic group was
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detected in the F−MWCNTs. Such band was apparently substituted by a new band at lower
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wavelength (1620.71 cm-1) that corresponded to the C=O stretching of amide for CRL−MWCNTs,
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suggesting excellent adsorption bands of proteins [25]. The existence of a cyano functional group
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(C−N) was further substantiated by the appearance of a new band (1032.96 cm-1). Additionally, the
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CH2 and N−H functional groups were characterized by explicit bands appearing at 2929.41 cm-1 and
290
3420.59 cm-1, respectively. It was revealed that the differing spectra for MWCNTs and
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CRL−MWCNTs were seen within the spectral region for carboxylate functional groups, in the
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range of 900−1200 cm-1 [29]. The FT−IR results further illustrated that following immobilization, a
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rather strong peak detected at 1125.04 cm-1 in the spectrum for free CRL correlated with the
294
vibrations of C−O of the carboxylate moiety was found expanding, reducing and shifting towards
295
lower frequency (1116.03 cm-1). Such finding was in accordance with the previously described
296
work [25], proving that the CRL was successfully immobilized onto the outer face of the
297
F−MWCNTs. The fact that an adsorption band at 2929.41 cm-1 was present in the CRL−MWCNTs
298
alone (Figure 1c), while similar band was evidently missing in the free CRL as well as
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F−MWCNTs, it can be interpreted that the immobilization process of CRL onto the surface of the
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F−MWCNTs had been successful.
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3.2.2 Thermal Gravimetric Analysis (TGA)
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The degree of alterations in physical and chemical properties of the biocatalysts as a function of
304
temperature or time was determined by TGA. The method is often used to monitor any reaction that
305
involves oxidation or dehydration. By measuring the decomposition or weight loss of the sample,
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the analysis via TGA would reveal the change in thermal stability of the support used in the
307
immobilization process, indicating if it has been modified [30]. In this investigation, TGA in the
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range of 30−800°C was employed to examine the mass loss percentage of F−MWCNTs and
309
CRL−MWCNTs, respectively (Figure 1). In general, combustion of the carbon nanotubes and the 12 Page 12 of 34
carbonaceous contaminants occurs at temperatures above 300oC. Any mass loss observed at
311
temperature below 400oC is usually associated with the evaporation of absorbed water or the
312
presence of surface functional groups. The thermogram for F−MWCNTs (Figure 1a) exhibited a
313
steady, uniform and multistage mass loss over the range of 30−150°C, 150−350°C and 350−800°C.
314
These changes corresponded to a 30% reduction in mass which strongly indicated the degradation
315
of various functional groups present on the surface of F−MWCNTs. Correspondingly, the first mass
316
loss (30−150°C) of 5% reduction may be attributable to the evaporation of hydrated water group
317
[31]. Further increment in temperature (150−350°C) rendered further decline in the mass of
318
F−MWCNTs by as much as 25%, implying degradation of surface groups on the MWCNTs
319
following acid treatment [32], confirming successful attachment of the –COOH groups on the walls
320
of the MWCNTs [31,33]. It has been reported that ccombustion of carbon materials would only
321
transpire when temperature reaches at least 400°C [33]. Finally, no further mass loss was found
322
beyond 350°C, indicative of thermally stable carbon that was created during removal of graphitic
323
and catalytic metal particles during the oxidation of the purified MWCNTs [33].
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As for the CRL−MWCNTs (Figure 1b), a 38% mass loss was recorded as a result of the
326
decomposition of the adsorbed CRL protein on the surface of the F−MWCNTs. An initial 6.5%
327
mass loss (30−150°C) was due to the elimination of both surface hydroxyl groups and organic
328
structure of lipase [34]. In comparison to F−MWCNTs, the decomposition curve of
329
CRL−MWCNTs was distinctively dissimilar between 250−400°C when compared with that of
330
F−MWCNTs. This was attributable to the thermal decomposition of lipase [35] present on the
331
surface of CRL−MWCNTs biocatalysts. Based on the calculations, approximately 13 wt% of CRL
332
protein was successfully attached to the surface of the F−MWCNTs following the immobilization
333
process.
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3.3
Transmission electron microscopy (TEM) and field emission scanning electron microscopy (FESEM)
338
Analysis on the morphological surface of the as-synthesized MWCNTs, F−MWCNTs and
339
CRL−MWCNT biocatalysts observable through TEM and FESEM are depicted in Figure 2. The
340
image of the as-synthesized MWCNTs illustrated closely attached open-ended tubular structures
341
with smooth surfaces that revealed the presence of contaminants on the surface of nanotubes
342
(Figure 2a). Since the contaminants on the surface of the as-synthesized MWCNTs (graphitic, metal
343
catalytic and amorphous carbon) may interfere and hamper the potential characters of the carbon
344
nanotubes, it was pertinent that these contaminants were removed [36].
us
cr
ip t
337
an
345
Following functionalization with the H2SO4 and HNO3 acid mixtures, the diameter of
347
MWCNTs was significantly increased, revealing noticeable rough surfaces that were attributable to
348
acid related eroded nanotubes [37]. The increased diameter of the F−MWCNTs strongly signified
349
the successful attachment of acid functional groups (COO-) on the surface of MWCNTs (Figure
350
2b), hence, corroborating previous findings reported by other studies [38]. In addition, the evidently
351
shorter length of the F−MWCNTs after treatment with the acid mixture was consistent with the
352
earlier studies [36, 39]. The perceived change in the highly tangled long MWCNTs into shorter,
353
open ended pipes strongly indicated that acid treatment on the nanotubes can potentially create
354
copious number of COO- at the open ends for linkage with chemical and biological [40]. According
355
to Khani and Moradi (2013), increased oxidation time and elevated temperatures were most likely
356
to destroy the structure of MWCNTs [40], generating surface imperfections on the MWCNTs that
357
subsequently affected the properties of the nanotubes. The increased diameter of CRL−MWCNTs
358
as compared to the F−MWCNTs CRL was physical an evidence of CRL on the surface MWCNTs,
359
(Figure 2c) and therefore, confirming the successful immobilization of the CRL on the MWCNTs
360
support.
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361 14 Page 14 of 34
362
3.3
Enzymatic production of geranyl propionate by free CRL and CRL−MWCNTs
363
3.3.1 Effect of Solvents (Log P) The partition-coefficient (log P) represents the ratio of concentrations of a compound in a
365
mixture of two immiscible phases consisting of 1−octanol and water at equilibrium, a key factor
366
that can influence the relative solubility, activity and specificity of the substrates [41] in an
367
enzymatic reaction. Figure 3 represents the reaction profile based on the increasing log P values of
368
organic solvents; hydrophilic log P < 1.4, diethyl ether (0.85); medium hydrophobicity: 1.4 < log P
369
< 3.5 (benzaldehyde 1.48, toluene 2.50) and hydrophobic: log P > 3.5 (n-heptane 4.0, decane 6.0)
370
in the production of geranyl propionate catalyzed by the free CRL and CRL−MWCNTs.
us
cr
ip t
364
an
371
The results showed an almost 2−fold increased conversion of geranyl propionate with
373
increasing hydrophobicity of solvents for the free CRL and CRL−MWCNTs. The highest
374
conversion was achieved when n-heptane was used as solvent for both the free CRL (28.1%) and
375
CRL−MWCNTs (51.3%) catalyzed reactions, confirming good compatibility of log P > 4 solvents
376
in enzymatically catalyzed reactions [42]. It was observed that n−heptane (log 4.0) was a suitable
377
solvent for the enzymatic production of geranyl propionate catalyzed by both free CRL and
378
CRL−MWCNTs. The solvent n−heptane sufficiently dissolved and suspended the substrates in the
379
reaction vessel without adversely affecting the catalytically active form of the CRL protein.
380
Previous studies have described that the high log P solvents would render better disassociation of
381
weak organic acids and prevent the stripping of the essential water layer surrounding the enzyme
382
[42], during the course of reaction. In this context, it is pertinent to point out that using n−heptane
383
as solvent can sustain the protein structure and catalytically active conformation of the CRL
384
[42,43]. Hence, the observed higher product yield [41] for the free CRL and CRL−MWCNTs
385
catalyzed esterification production of geranyl propionate in n−heptane was obtained.
Ac ce p
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372
386
15 Page 15 of 34
It is generally accepted that hydrophilic, namely, polar solvents tend to strip off essential
388
water from enzyme molecules that lead to the inactivation of the enzyme activity [44]. Considering
389
the hydrophobicity of the solvents employed in this investigation, it was expected that the activity
390
of the enzyme to improve with the reduction of solvent hydrophilicity. Although, it has been
391
described in literature that polarity cannot be used as the only measure when the effects of solvents
392
on enzyme activity of immobilized lipases are tested, the results in this study proved otherwise. The
393
notably low enzyme activity in hydrophilic solvents (log P < 2) rendering lowest conversion of the
394
geranyl propionate strongly suggests the dehydrating effect of such solvents on the enzyme active
395
site [45]. Enzyme deactivation on the free CRL and CRL−MWCNTs was reduced, reflected in the
396
higher production of methyl oleate when hydrophobicity of solvents used in the reactions was
397
increased. Apart from increased penetration of solvent molecules into the lipase protein, hydrophilic
398
solvents also promoted deformation and increased flexibility of the lipase surface and loss of
399
enzyme stability. Correspondingly, poor predictability for various hydrophilic solvents (log P 2.0–
400
4.0) has been reported [46]. In this perspective, it is vital to highlight that the hydrophilic solvents
401
clearly had a negative effect on the free CRL and CRL−MWCNTs than the hydrophobic ones for
402
the esterification production of geranyl propionate.
404 405
cr
us
an
M
d
te
Ac ce p
403
ip t
387
3.3.2 Effect of Molar Ratio of Alcohol to Acid Previous studies have illustrated that equilibrium of an enzymatic reaction can be
406
manipulated to favor high conversion of the ester product, through proper selection of molar ratio of
407
the reacting alcohol or acids or by removal of the products from the reaction mixture [47]. In some
408
cases, the use of excess alcohol is required to reduce the intrinsic denaturing properties of acids on
409
lipases. Presence of excess acid in a reaction, to a certain extent, can deactive the lipase particularly
410
whenever the pH drops below pH 2. Considering that high concentrations of acid, alcohol or both
411
are significant factors to influence the equilibrium of enzymatic esterification reactions [48], such
412
factors were examined in this study. The effect of molar ratio of alcohol to propionic acid for the 16 Page 16 of 34
413
enzymatic production of geranyl propionate was monitored for molar ratio geraniol: propionic in
414
the range of 1:1 to 5:1.
415 Figure 4 depicts the optimum molar ratio of geraniol to propionic acid as 5:1 corresponding
417
to 28% and 47% of geranyl propionate for the free CRL and CRL−MWCNTs, respectively. The
418
conversion percentage of geranyl propionate for the CRL−MWCNTs significantly increased over
419
the free CRL by 2−fold, indicating higher activity of the CRL−MWCNTs following immobilization
420
onto the F−MWCNTs. The elevated activity of the CRL−MWCNTs over the free CRL was
421
anticipated as similar enhancements reported for the commercially immobilized lipase i.e.
422
Novozyme 435 [4]. Conversion of geranyl propionate was the lowest for both free CRL and
423
CRL−MWCNTs at the molar ratio of 1:1, corresponding to 13.6% and 32.4%, respectively. The
424
low conversion of geranyl propionate at equal proportions of geraniol to propionic acid can be
425
attributed to the inhibition of the CRL activity due to the presence of surplus acid in the reaction
426
medium that adversely decreased the initial rate of reaction [49]. However, the noteworthy higher
427
conversions of the ester in all CRL−MWCNTs catalyzed reactions confirm the enhanced stability of
428
the adsorbed CRL protein. The additional network of interaction i.e. van der Waals forces,
429
hydrophobic interactions and hydrogen bond formed between the surface of the CRL and the acid
430
groups on the surface of the MWCNTs reduce the flexibility of the CRL protein. Hence, the
431
structure of CRL is less predisposed to the inherently denaturing effects of the alcohol and acid
432
substrates that can disrupt the three-dimensional active form of the CRL protein, above all, during
433
prolonged reaction time.
Ac ce p
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d
M
an
us
cr
ip t
416
434 435
3.3.3 Thermal Stability
436
Thermal stability of an enzyme is a vital aspect of bioprocess reactions operating at high
437
temperatures in industrial setting. A high reaction temperature is often preferred as it promotes
438
substrates solubility and reduces viscosity to which mass transfer limitation can be avoided [50,51]. 17 Page 17 of 34
In this study, thermal stability of both free CRL and CRL−MWCNTs was investigated after
440
incubating both forms of CRL for 2 h at various temperatures (50−80oC) and the data are illustrated
441
in Figure 5. It was revealed that the thermal stability for CRL−MWCNTs was enhanced over the
442
free CRL by almost 2−fold with the former retaining at least 22.6−32.1% of activity even at the
443
highest temperature of 80°C when compared with that of free CRL (12.6−18.1%). Such observation
444
indicates the improved robustness of the biocatalysts to catalyzed reactions under elevated
445
temperature. The higher thermal stability of CRL−MWCNTs over the free CRL potentially
446
conferred by additional electrostatic interactions and hydrogen bonds between the enzyme and the
447
support, favorably influencing the thermal stability [52,53] of the immobilized lipase.
us
cr
ip t
439
On the other hand, the activity of the free CRL was diminished by 46%, retaining only
449
12.2% of enzyme activity. Again, the considerable improvement in thermal stability of
450
CRL−MWCNTs over the free CRL during reaction can be associated with the additional
451
interactions between the molecules of CRL and MWCNTs that rendered the protein structure of
452
CRL−MWCNTs to be more rigid. Therefore, the increased structural rigidity resulted in less
453
yielding to the temperature-induced change in the lipase structure. Conformational transitions of the
454
CRL are averted [54], resulting in a more stable structure even at high reaction temperatures.
455
Hence, immobilized CRL is able to retain its catalytically competent form for a longer period of
456
time particularly during extensive reaction times, even under elevated temperatures [50]. Similar
457
thermal stabilization has also been described for powdered layered double hydroxides (LDH)
458
known for its use as nanocarriers or catalyst support for functional molecules [12].
Ac ce p
te
d
M
an
448
459 460
3.4
Analysis of Geranyl Propionate Using Gas Chromatography (GC) and Nuclear
461
Magnetic Resonance spectroscopy
462
The production of geranyl propionate by the free CRL and CRL−MWCNTs (Schematic 1)
463
was monitored over a 12 h period and samples were withdrawn and identified by GC analysis.
464
Separations of the components in the sample are based on the interaction strength of compounds 18 Page 18 of 34
with the stationary phase as well as other factors i.e. boiling point and polarity of compounds [55].
466
Prior to analysis, the boiling point for each component was determined as: propionic < geraniol <
467
geranyl propionate. According to Figure 6, geranyl propionate has a longer retention time as
468
compared to geraniol and propionic acid. In Figure 6a, the initial composition of the reaction only
469
revealed a peak at 10.660 min. Following a 12 h incubation period, an existence of a new peak at
470
13.782 min (Figure 6b) was observed. The new peak represents the liberated geranyl propionate that
471
was enzymatically synthesized from geraniol and propionic acid in reactions catalyzed by both free
472
CRL and CRL−MWCNTs. Similar retention time for GC analysis of geranyl propionate has been
473
previously described [55].
us
cr
ip t
465
Analysis of 1H and 13C NMR revealed peaks that were characteristics of geranyl propionate
475
(Figure 7a and Figure 7b). The 1H NMR for geranyl propionate is as follows: (700 MHz, CDCl3): δ
476
(ppm): 1.15 (3H, t, -CH3), 1.60 (3H, s, -CH3), 1.69 (3H, s, -CH3), 1.71 (3H, s, -CH3), 2.15 (2H, t, -
477
CH2), 2.17 (2H, t, -CH2), 2.35 (2H, m, -COCH2), 4.60 (2H, d,-OCH2), 5.08 (1H, t, - =CH), 5.35 (1H,
478
t, - =CH). The
13
C NMR shows characteristic peaks i.e. (700 MHz, CDCl3): δ (ppm): (174.49 (C-
te
479
d
M
an
474
3=O), 142.03 (C6), 131.78 (C10), 123.74 (C9), 118.42 (C-5), 61.24 (C-4), 38.61 (C-7), 27.67 (C-2),
481
26.84(C-8 and C-11), 17.86 (C-13) and 8.12 (C-1). Hence, the 1H NMR and
482
geranyl propionate confirmed presence of a 13-carbon moiety that matched the structure of geranyl
483
propionate.
484 485
4.
Ac ce p
480
13
C NMR spectra of
Conclusions
486
The improved activity and stability of CRL−MWCNTs over the free CRL to catalyze
487
production of geranyl propionate was attributable to the enhanced structural integrity and
488
mechanical strength of the CRL protein conferred by the F−MWCNTs. In consequence, the CRL
489
protein became more rigid, resistant towards premature unraveling as well as sustaining its
490
catalytically competent structure for longer period of time. Such property is useful especially when 19 Page 19 of 34
dealing with extensive esterification process under harsh conditions such as extreme temperatures
492
and pH. The 2−fold enhanced activity and increased thermal robustness of the CRL−MWCNTs
493
biocatalysts subsequently resulted in a more efficient biosynthesis of geranyl propionate. This work
494
proves that physical adsorption of CRL onto F−MWCNTs has improved the stability and activity of
495
CRL as a biocatalyst. Considering the costs of 10 g of commercial enzymes i.e. Novozyme
496
(>USD900) and Lipozyme (USD400), the developed CRL−MWCNTs biocatalysts in this study
497
would potentially be an economical and sustainable alternative for the enzymatic production of
498
geranyl propionate as well as to produce other similarly important commercial esters.
us
cr
ip t
491
499
501
an
500 Acknowledgments
The authors wish to express their gratitude to the Faculty of Science, Universiti Teknologi
503
Malaysia for their facilities. This work was funded by the Ministry of Higher Education of Malaysia
504
under the Exploratory Research Grant Scheme (ERGS R.J130000.7826.4L132).
505 506 References
508
[1]
509 510
Ac ce p
507
te
d
M
502
S.M. Radzi, W.A.F Mustafa, S.S. Othman, H.M. Noor, World Acad. Sci. Eng. Technol. 59 (2011) 677–680.
[2]
511
F.X. Malcata, H.R. Reyes, H.S. Garcia, C.G. Hill, C.H. Amundson, J. Am. Oil Chem. Soc. 67 (1990) 890–910.
512
[3]
T.W. Charpe, V.K. Rathod, Waste Manage. 31 (2011) 85–90.
513
[4]
N. Paroul, L.P. Grzegozeski, V. Ciaradia, H. Treichel, R.L. Cansian, J. Vladimir Oliveira, D.
514 515 516
de Olivera, J Chem. Technol. Biotechnol. 12 (2010) 1636–1641. [5]
D.M. Solano, P. Hoyos, M.J. Hernaiz, A.R. Alcantara, J.M. Sanchez-Montero, Biores. Biotechnol. 115 (2012) 196−207. 20 Page 20 of 34
517
[6]
P. Anastas, N. Eghbali, Chem. Soc. Rev. 39 (2−010) 301–312.
518
[7]
B.M. Nestl, B.A. Nebel, B. Hauer, Curr. Opin. Chem. Biol. 15 (2011) 187–193.
519
[8]
J. Tao, R.J. Kazlaukas, 2011. Biocatalysis for Green Chemistry and Chemical Process
521 522
Development. John Wiley & Sons Ltd., Hoboken, New Jersey. [9]
R.B. Salah, H. Ghamghui, N. Miled, H. Mejdoub, Y. Gargouri, J. Biosci. Bioeng. 103 (2007)
ip t
520
368–372.
[10] G. Zhou, C. Wu, X. Jiang, J. Ma, H. Zhang, H. Song, Process Biochem. 47 (2012) 953−959.
524
[11] R.A. Wahab, M.B. Abdul Rahman, M. Basri, R.N.Z. Raja Abdul Rahman, A.B. Salleh, T.C.
Leow. Biotech. Biotech. Eq. 28 (2014) 1065−1072.
us
525
cr
523
[12] Y. Kuang, L. Zhao, S. Zhang, F. Zhang, M-D. Dong, S. Xu. Materials 3 (2010) 5220-5235.
527
[13] J. Song, M. Chen, V.R. Regina, C. Wang, R.K. Meyer, E. Xie, C. Wang, F. Besenbacher, M. Dong. Adv. Eng. Mat. 14 (2012) B240-B246.
M
528
an
526
[14] K.E. Jaeger, T. Eggert, Curr. Opin. Biotechnol. 13 (2001) 390–397.
530
[15] M. Shim, N.W.S. Kam, R.J. Chen, Y.M. Li, H.J. Dai, Nano. Letts. 22 (2002) 28−58.
531
[16] P. Zhang, DB. Henthorn, AIChE J. 56 (2010) 1610–1615.
532
[17] M. Cesar, M.P. Jose, F.L. Gloria, M.G. Jose, F.L. Roberto, Enzym. Microb. Technol. 40
te
Ac ce p
533
d
529
(2007)1451–1463.
534
[18] W. Tischer, V. Kascher, Trends Biotechnol. 17 (1999) 326−335.
535
[19] D. Norouzian, Iran. J. Biotechnol. 1 (2003) 197−206.
536
[20] P.C. de Oliveira, G.M. Alves, H.F. de Castro, Biochemical Eng. J. 5 (2000) 63−71.
537
[21] M.M. Bradford, Anal. Biochem. 72 (1976) 248–254.
538
[22] M.A.P. Langone, G.J. Sant'anna, G. J. Appl. Biochem. Biotechnol. 77-79 (1999) 759−770.
539
[23] N.R. Mohamad, N.A. Buang, N.A. Mahat, Y.Y. Lok, F. Huyop, H.Y. Aboul-Enein, R.A.
540
Wahab. Enzym. Microb. Technol. 72 (2015a) 49−55.
541
[24] T. Ramanathan, F.T. Fisher, R.S. Ruoff, L.C. Brinson, J. Mat. Chem. 17 (2005) 1290−1295.
542
[25] N.Z. Prlainoc, D.I Bezbradica, J. Ind. Eng. Chem.19 (2013) 279−285. 21 Page 21 of 34
543 544
[26] I.V. Pavlidis, T. Tsoufis, A. Enotiadis, G. Gournis, H. Stamatis, Advan. Eng. Mat. 12 (2010) 5.
545
[27] L. Zhang, V.U. Kiny, H. Peng, J.J. Zhu, Mat. Chem. 16 (2004) 2055−2061.
546
[28] M.F. Alkhatib, M.E.S. Mirghani, I.Y. Qudsieh, I.A.F. Husain, J. Appl. Sci. 10 (2010) 2705−2708.
ip t
547
[29] A. Natalello, D. Ami, S. Brocca, M. Lotti, S.M. Doglia. Biochem. J. 85 (2005) 511−517.
549
[30] N.R. Mohamad, N.H. Che Marzuki, N.A. Buang, F. Huyop, R.A. Wahab. Biotechnol. Biotec. Eq. 29(2015b) 205−220.
us
550
cr
548
[31] V. Datsyuk, M. Kalyva, K. Papagelis, J. Parthemios, D. Tasis, Carbon. 46 (2008) 833−840.
552
[32] J.L. Vicente, A. Albesa, J.L. Llanos, E.S. Flores, A.E. Fertutta, J Argent. Chem. Soc. 98
554 555
(2011) 29−38.
[33] T. Raghavendra, A. Basak, L.M. Manoca, A.R. Sha, D. Madamwar, Biores. Technol. 140
M
553
an
551
(2013) 103-110.
76
[34] H. F. Castro, M.L.C.P. Silva, G.L.J.P. Silca, Braz. J. Chem. Eng. 17 (2000) 4−7.
557
[35] Y. Shi, F, Jin, Y. Wu, F, Yan, X. Yu, Y. Quan, Dalian Institute of Light Industry. 13
559 560 561 562
te
(1997)111−113.
Ac ce p
558
d
556
[36] R. Yudianti, H. Onggo, R. Sudirman, Y. Saito, T. Iwata, J.I. Azuma, Open Mat. Sci. J. (2011) 5:242−247.
[37] M.F. Alkhatib, M.E.S. Mirghani, I.Y. Qudsieh, I.A.F. Husain, J. Appl. Sci. (2010) 10(21) 2705−2708.
563
[38] M.R. Joha, S.H.M. Suhaimy, Y. Yusof, Appl. Surf. Sci. (2014) 450−454.
564
[39] I. Mazova, V.L. Kuznetsova, I.A. Simonovaa, A.I. Stadnichenkoa,. Appl. Surf. Sci. (2012)
565
6272−6280.
566
[40] H. Khani, O. Moradi, J. Nanostruc. Chem. (2013) 3−73.
567
[41] S. Gogoi, N.N. Dutta, Ind. J. Chem. Technol. 16 (2009) 209−215.
568
[42] C. Laane, S. Boeren, K. Vos, C. Veeger, Biotech. Bioeng. 30 (1987) 81−87. 22 Page 22 of 34
569 570
[43] J. Song, D. Kahveci, M. Cheng, Z. Guo, E. Xie, X. Xu, F. Besenbacher, M. Dong. Langmuir 28 (2012), 6157−6162. [44] S.H. Krishna, N.G. Karanth, Catal Rev. 4 (2002) 499−591.
572
[45] A. Zaks, A.M. Klibanov, Science. 22 (1984) 1949−1951.
573
[46] L. Yang, J.S. Dordick, S. Garde, Biophys. J. 87 (2004) 812−821.
574
[47] M. Trani, F. Ergan, G. Andre, J. Am. Oil Soc. 68 (1991) 20−22.
575
[48] H. Stamatis, P. Christakopolous, D. Kekos, B.J. Macris, F.N. Kolisis, J. Mol. Catal. B:
cr
Enzym. 4 (1998) 229−236.
us
576
ip t
571
[49] P. Pires Cabral, M.M.R. Fonseca, S.F. Dias, Biochem. Eng. 43 (2009) 327−332.
578
[50] M.B. Abdul Rahman, N. Chaibakhsh, Basri M, R.N.Z. Abdul Rahman, A.B. Salleh, S. Md
580 581
Radzi, J. Chem. Technol. Biotechnol. 83 (2008) 1534−1540.
[51] R. Dave, D. Madamwar, in: C. Larroche, A. Pandey, C.G. Dussap, (Eds.), Current Topics on
M
579
an
577
Bioprocesses in Food Industry. Asiatech Publishers Inc., New Delhi, 2006, pp. 70–80. [52] L. Wang, L. Wei, Y. Chen, R. Jiang. J. Biotechnol. 150 (2010) 57–63.
583
[53] K. Zhu, A. Jutila, E.K.J. Tuominen, P.K.J. Kinnunen, Pro Sci. 10 (2001) 339–351.
584
[54] E. Yilmaz, K. Can, M. Sezgin, M. Yilmaz, Biores. Technol.102 (2001) 499−506.
585
[55] R.R. Thanighai, B. Nambikkairaj, P. Sivamani, Ind. J. Appl. Res. 3 (2013) 375−379.
te
Ac ce p
586
d
582
23 Page 23 of 34
586 List of Legends Table 1
The frequency table for FTIR analyses for a) Raw MWCNTs, b) FMWCNTs and c) CRL−MWCNTs Thermogram of the TGA performed on (a) F−MWCNTs and (b)
ip t
Figure 1
CRL−MWCNTs.
Transmission electron microscope (TEM) and field emission scanning
cr
Figure 2
F−MWCNTs and (c) CRL−MWCNTs.
The effect of solvent (log P): diethyl ether (0.8), benzaldehyde (1.48),
an
Figure 3
us
electron microscope (FESEM) images of (a) as-synthesized MWCNTs, (b)
toluene (2.5), n-heptane (4.0) and decane (6.0) in the enzymatic synthesis
M
geranyl propionate. [Temperature: 55°C, molar ratio propionic acid/geraniol (1:5), enzyme amount 5 mg/mL for CRL−MWCNTs and 10 mg/mL free
The effect of the molar ratio of geraniol to propionic acid (1:1 to 5:1) on the
Ac ce p
Figure 4
te
n-heptane].
d
CRL, molecular sieves: 100 mg, stirring speed 200 rpm, time: 12 h, solvent:
synthesis of geranyl propionate by CRL−MWCNTs at reaction conditions:
[Temperature: 55°C, enzyme amount: 5 mg/mL for CRL−MWCNTs and 10
mg/mL free CRL, molecular sieves: 100 mg, stirring speed: 200 rpm, time: 12 h, solvent: n-heptane].
Figure 5
The effect of thermal stability on the synthesis of geranyl propionate by CRL−MWCNTs. [Temperature: 40°C; molecular sieves: 100 mg, stirring speed: 200 rpm, time: 12 h, solvent: n-heptane].
Figure 6
Gas chromatogram of sample from the enzymatic esterification of geraniol and propionic acid catalyzed by the free CRL and CRL−MWCNTs at 0 h
24 Page 24 of 34
and 12 h. Figure 7
The a) H-NMR and b) C-NMR spectra of the purified esterification product geranyl propionate synthesized by the CRL-MWCNTs. The reaction scheme for the lipase-catalyzed esterification to produce geranyl propionate from geraniol and propionic acid.
cr
587
ip t
Schematic 1
Ac ce p
te
d
M
an
us
588
25 Page 25 of 34
588 Table 1: Functional group
a)
1630.82 3428.01
conjugated C=C bonds O−H stretching
b)
1112.73 1202.46 1634.70 3445.22
C−O stretching C−O stretching C=O stretching O−H stretching
c)
1032.96 1116.03 2929.41 3420.59
C−N stretching C−O stretching CH2 bending N−H stretching
an
us
cr
ip t
Frequency (cm-1)
Ac ce p
te
d
M
589 590
26 Page 26 of 34
Schematic 1
O O + OH geraniol
OH
O
CRL-MWCNTs
+ H2O
or free CRL geranyl propionate
propionic acid
Ac
ce pt
ed
M
an
us
cr
ip t
Schematic 1:
Page 27 of 34
Figure 1
(
M
an
us
cr
ip t
a)
(
Ac
ce pt
ed
b)
Figure 1:
Page 28 of 34
Ac c
ep te
d
M
an
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Figure 2
Figure 2:
Page 29 of 34
Figure 3
60 FREE-CRL CRL-MWCNTs
40
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30
20
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Conversion (%)
50
0
1.48
2.5
4
6
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M
Solvent (Log P)
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Figure 3:
Page 30 of 34
Figure 4
60 FREE CRL CRL−MWCNTs
50
40
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30
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20
0 1:2
1:3
1:4
1:5
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1:1
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Molar ratio acid:alcohol
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Figure 4:
Page 31 of 34
Figure 5
40 FREE CRL CRL-MWCNTs
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10
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Residual activiry (%)
30
0
60°C
70°C
80°C
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50°C
Temperature
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Figure 5:
Page 32 of 34
Figure 6
Sample at 12 h
Geranyl propionate
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(b)
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(a) Sample at 0 h
Figure 6:
Page 33 of 34
Figure 7
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a)
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Figure 7:
Page 34 of 34
S1226-086X(15)00352-4 http://dx.doi.org/doi:10.1016/j.jiec.2015.08.001 JIEC 2605
To appear in: Received date: Revised date: Accepted date:
6-1-2015 28-6-2015 2-8-2015
Please cite this article as: N.R. Mohamad, N.A. Buang, N.A. Mahat, J. Jamalis, F. Huyop, H.Y. Aboul-Enein, R.A. Wahab, Simple Adsorption of Candida rugosa Lipase onto Multi-walled Carbon Nanotubes for Sustainable Production of the Flavor Ester Geranyl Propionate, Journal of Industrial and Engineering Chemistry (2015), http://dx.doi.org/10.1016/j.jiec.2015.08.001 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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Simple Adsorption of Candida rugosa Lipase onto Multi-walled Carbon Nanotubes for
2
Sustainable Production of the Flavor Ester Geranyl Propionate
3 NurRoyhaila Mohamad1, Nor Aziah Buang1, Naji A. Mahat1, Joazaizulfazli Jamalis1, Fahrul
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Huyop2, Hassan Y. Aboul-Enein3 and Roswanira Abdul Wahab1,*
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Department of Chemistry, Faculty of Science, Universiti Teknologi Malaysia, 81310 UTM Johor
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Bahru, Johor, Malaysia 2
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Department of Biotechnology and Medical Engineering, Faculty of Biosciences and Medical
Engineering, Universiti Teknologi Malaysia, 81310 UTM Johor Bahru, Johor, Malaysia
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Pharmaceutical and Medicinal Chemistry Department, Pharmaceutical and Drug Industries Research Division, National Research Centre, Dokki, Cairo 12311, Eqypt
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*Corresponding author:
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Roswanira Abdul Wahab, Department of Chemistry, Faculty of Science, Universiti Teknologi
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Malaysia, 81310 UTM Johor Bahru, Johor, Malaysia, Telephone: +607 5534148; Fax +607
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5566162; Email: [email protected]
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Highlights: 1. The CRL−MWCNTs improved yield of geranyl propionate by 2−fold
23
2. The CRL−MWCNTs showed a 2−fold improved thermal stability
24
3. CRL−MWCNTs as cheap biocatalyst to produce geranyl propionate
25 26 27 28 1 Page 1 of 34
Abstract
30
In this study, geranyl propionate was enzymatically synthesized from geraniol and propionic acid
31
using Candida rugosa lipase immobilized on acid functionalized multi−walled carbon nanotubes.
32
The efficiency of the CRL−MWCNTs biocatalysts to catalyze the esterification production of
33
geranyl propionate (solvent log P, alcohol:acid molar ratio and thermal stability) was compared
34
with the free CRL for parameters. The use of CRL−MWCNTs in n-heptane (log P 4.0) and
35
alcohol:acid molar ratio of 5:1 resulted in a 2−fold increased conversion frequency as compared to
36
the free CRL for the production of geranyl propionate, in addition to a noteworthy 2−fold enhanced
37
thermal stability.
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Keywords: Candida rugosa lipase; biocatalysts; immobilization; multi−walled carbon nanotubes;
41
esterification; geranyl propionate
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1.
Introduction Terpene esters of short-chain fatty acids are essential oils with significant applications in
57
food, cosmetic and pharmaceutical industries as flavors and fragrances; geraniol and citronellol,
58
being the two foremost substances [1]. Conventionally, these esters are produced via various
59
methods viz. chemical synthesis, extraction from natural products as well as fermentation [2].
60
Nonetheless, chemical synthesis is often the preferred choice to mass produce such esters even
61
though the method entails the use of strong acid catalyst and hazardous chemicals, considerably
62
lengthy reaction time while providing low conversion rate [1]. Additionally, the downstream
63
processing of such method incurs tedious separation processes, involving extreme exposure of
64
toxicants as well as superfluous harmful by−products [3]. Considering the numerous disadvantages
65
that are typically linked to the current chemical approach for producing these terpene esters [4,5],
66
alternative methods that are more environmentally friendly, economically attractive and improve
67
reaction yield would be significant.
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In addition, the approval of REACh (Registration, Evaluation, Authorization and Restriction
70
of Chemicals) regulation by the European Parliament and of the Council as well as regulation and
71
the environmental restrictions imposed by US, Japan and E.U. have opened up new challenges for
72
utilizing Green Chemistry philosophy in industrial processes [6]. The philosophy emphasizes on the
73
industrial usage of chemicals acquired from biomass, the use of green solvents and more sustainable
74
industrial processes [7]. The key aspect of ‘‘green’’ industrial processes is the effective
75
incorporation of catalytic technologies (chemical or enzymatic) into a general organic synthesis
76
scheme. In this sense, biocatalysis offers many attractive features in the perspective of green
77
chemistry [7]. The biocatalytic route embraces gentle reaction conditions (physiological pH and
78
temperature) and environmentally friendly catalysts (an enzyme or a cell) that display high
79
activities and chemo-, regio− and stereoselectivities in multifunctional molecules. Additionally, the
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use of biocatalysts usually circumvents the need for functional group activation and therefore,
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affords processes that are shorter, produce less waste and economically smarter [5, 8].
82 Lipases (triacylglycerol ester hydrolysis EC 3.1.1.3) are medicinally, industrially and
84
commercially pertinent biomolecules that have acquired popularity as biocatalysts [9] over
85
inorganic biocatalysts for reasons such as high specificity, efficient reaction rate, non-toxicity and
86
biodegradability, reproducibility under normal laboratory conditions [10]. Also, they are able to
87
convert a large number of substrates with their high stereospecificity [11]. In this context, Candida
88
rugosa lipase (CRL) remains one of the most versatile biocatalysts due to its high activity and broad
89
specificity in reaction medium [10]. In view of (a) the poor stability of the free form CRL, (b) low
90
activity in organic solvents, (c) high tendency of deactivation in prolonged exposure to high
91
temperature and extreme pH [10], immobilization of CRL onto various nanosupports offer many
92
benefits; one of them being large surface-to-volume-ratio [12-13]. The use of nanosupports such as
93
multi−walled carbon nanotubes may be a practical solution [14−15] for improving stability and
94
activity as well as extending the reaction life of the enzyme.
96
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Enzyme immobilization confers the advantages of structural stability, improved activity,
97
specificity and selectivity as well as reduced inhibition [16]. Moreover, immobilization of enzyme
98
facilitates facile separation of the enzyme from product, hence simplifying enzyme application for a
99
reliable and efficient reaction technology. Since enzymes are expensive and required in large
100
quantities for industrial purposes [17], immobilization of such enzymes onto solid support matrices
101
may prove beneficial in permitting reusability of the biocatalysts, often an essential prerequisite for
102
establishing an economically viable enzyme catalyzed process [18]. In this context, the physical
103
adsorption method may have higher commercial values than other methods [19] because it remains
104
as one of the simplest and cheapest immobilization methods available and importantly, in most
105
cases the enzyme productivity remains unaffected [20]. 4 Page 4 of 34
Immobilized enzymes are currently the subject of considerable attention for their numerous
107
advantages over soluble enzymes. Consequently, the current study was intended at investigating the
108
potential use of physically adsorbed CRL− multi-walled carbon nanotubes biocatalysts
109
(CRL−MWCNTs) as alternative economical and sustainable biocatalysts for the enzymatically
110
based esterification production of geranyl propionate. Performance of the CRL−MWCNTs
111
biocatalysts for catalyzing production of geranyl propionate was compared with that of the free
112
CRL. Both forms of biocatalysts were evaluated for the effects of solvent log P, molar ratio
113
alcohol:acid as well as thermal stability in rendering the highest conversion of geranyl propionate.
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2.
Materials and methods
117
2.1
Chemicals
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MWCNTs were received as-synthesized from the Department of Chemistry, Faculty of
119
Science. Lipase from Candida rugosa (EC 3.1.1.3), Type VII (1410 U mg-1) and Bradford reagent
120
were procured from Sigma Aldrich Co. (St. Louis, USA). For the esterification reactions, substrates
121
geraniol (98%) and propionic acid (99%) were also purchased from Sigma Chemical Co. (St. Louis,
122
USA). Solvents, diethylether, benzaldehyde, toluene, n-heptane and decane were purchased from
123
Merck (Darmstadt, Germany). Nitric acid, HNO3 65% (v/v); hydrochloric acid, HCl 37% (v/v) and
124
sulphuric acid, H2SO4 95−97% (v/v) from Merck (Darmstadt, Germany) were used as acid solvent
125
systems in the purification and oxidation processes of MWCNTs. Other chemicals, namely sodium
126
hydroxide (NaOH) and phosphate buffer (Merck, Germany) were used without further purification.
127
Distilled water was produced in the lab and was used in all experiments. All chemicals were of
128
analytical grade unless specified otherwise.
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2.2
Purification and functionalization of MWCNTs For the purification process, the synthesized−MWCNTs (0.5 g) were transferred into a 100 5 Page 5 of 34
mL flask that contained 4 M HCl (20 mL) and refluxed with stirring at 80°C for 5 h. After cooling
133
to room temperature, the liquid was decanted, the MWCNTs washed with distilled water until no
134
remaining acid was detected and dried in an oven at 60°C for 24 h. The purified MWCNTs were
135
refluxed at 120°C by stirring in a mixture of concentrated HNO3 and H2SO4 (1:3 v/v) for 24 h. After
136
cooling to room temperature, the mixture containing the acid functionalized MWCNTs
137
(F−WMCNTS) was decanted, washed with distilled water until no remaining acid was present and
138
dried at 60°C for 24 h.
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2.3
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Adsorption immobilization of lipase and its characterization
The F−MWCNTs (10 mg/mL) were first sonicated to homogeneity in aqueous buffer (pH 7)
142
for 30 min. Then, the MWCNTs were transferred into a 50 mL flask of phosphate buffer (50 mM,
143
solution pH 7) that contained CRL (10 mg/mL) and incubated at 20°C with constant stirring at 150
144
rpm. Following 3 h incubation period, the flask containing CRL MWCNTs was cooled to room
145
temperature and stored at 4°C for 24 h. After the stipulated period, the unbound proteins were
146
removed by washing with phosphate buffer (pH 7) until no evidence protein was detected in the
147
supernatant. The supernatant was analyzed for its protein content by Bradford method [21].
149 150
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2.3.1 Fourier Transform Infrared Spectroscopy (FTIR) A ratio of 1:100 mass of sample was ground thoroughly with potassium bromide and the
151
resulting powder was pressed into a transparent pellet by a hydraulic press. The FTIR spectra
152
obtained using a BOMEM spectrophotometer (Quebec, Canada) in transmission mode between 400
153
and 4000 cm-1 at a resolution of 4 cm were analyzed.
154
6 Page 6 of 34
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2.3.2 Thermal Gravimetric Analysis (TGA)
156
Thermal gravimetric analysis (TGA) for F−MWCNTs and CRL−MWCNTs was conducted
157
in a compact ceramic alumina crucible and heated from 30°C to 800°C under a nitrogen atmosphere
158
using a Thermogravimetric Analyzer 4000 (Perkin Elmer) at a heating rate of 10°C/min.
160
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2.3.3 Transmission Electron Microscopy (TEM) and Field Emission Scanning Electron Microscopy (FESEM)
162
Morphological analysis for the MWCNTs, F−MWCNTs and CRL−MWCNTs were
163
determined using transmission electron spectroscopy (TEM) and field emission scanning electron
164
microscopy (FESEM). The measurement was conducted by heating the sample (5.0−5.2 mg) in a
165
platinum pan (5mm in diameter), under oxygen flow in temperature range of 35−1000oC, ramped at
166
the rate of 10oC min-1. TEM analysis was executed using electron source as W-emitter and LaB6
167
that operated at an accelerating voltage of 200 Kv. The analysis employed an objective lens (S-
168
Twin) having a point resolution of 2.0 nm or better with a 25x to 7,500,200x or higher
169
magnification and a single tilt holder with LCD camera. The individual sample (0.5 mg) of
170
MWCNTs, F−MWCNTs and CRL−MWCNTs was ultrasonicated for 2 h in deionized water and a
171
drop of each sample was positioned on a copper grid, observed after drying in vacuum. For the
172
analysis of FESEM, a sample which consisted of a drop of the dispersed MWCNTs was placed on a
173
silicon wafer and allowed to dry in vacuum for 30 min prior to analysis using JEOL JEM−6700F.
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Determination protein loading and lipase activity
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The concentration of protein in the enzyme solution, prior and post immobilization, in the
177
washing buffer was quantified by the Bradford method using BioRad protein dye reagent
178
concentrate and bovine serum albumin (BSA) as the protein standard [21]. Different concentrations
179
of BSA were prepared using a BSA stock solution (0.1 mg in 10 mL of deionized water) and the
180
absorbance was measured at 595 nm with a spectrophotometer (HITACHI U-3210) using 7 Page 7 of 34
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preparations without BSA as the blank. Analyses were performed in triplicate. The activity of CRL−MWCNTs was determined according to the method suggested by
183
Langone and Sant'Anna (1999) [22]. Lipase activity was quantified by the consumption of lauric
184
acid in the esterification reaction with glycerol (lauric acid/glycerol, 1:3) at 50oC, with the enzyme
185
concentration of 5 mg/mL. One esterification unit of the lipase (U) was defined as 1 μ mole of
186
lauric acid consumed/min and the CRL−MWCNT biocatalysts had an esterification activity of 792
187
U mg-1. All determinations were performed in triplicate.
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2.5
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Production of geranyl propionate by free CRL and CRL−MWCNTs A typical esterification reaction was executed in propionic acid (0.25 M) and geraniol (1.25
191
M) dissolved in n-heptane (unless specified otherwise) as solvent in a 100 mL round bottom flask.
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The free CRL (10 mg/mL) or CRL−MWCNTs (5 mg/mL) were reacted in a 30 mL volume of this
193
mixture, respectively. The reaction mixture was stirred with continuous stirring at 200 rpm in a
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paraffin oil bath. Aliquots (1 mL) of the reaction mixture were withdrawn periodically for 12 h and
195
analyzed by titration with NaOH (0.05 M) with phenolphthalein as indicator. The geranyl
196
propionate obtained was expressed in terms of percent conversion i.e. percent of propionic acid
197
converted versus the total acid in the reaction mixture. Each measurement was performed in
198
triplicate and the standard error was calculated. The percent conversion was calculated according to
199
the prescribed equation [1].
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% Conversion = ((Vo − Vt) /Vo) x 100
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whereby, Vo is volume of NaOH at initial time (t = 0) and Vt is volume of NaOH at stipulated hour
202
(t = t1, t2, t3 ...).
203 204
2.5.1 Effect of Solvents (Log P)
205
Stability of biocatalysts in organic solvents is an important characteristic to be considered
206
for a biosynthetic reaction. The effect of solvents in the enzymatic production of geranyl propionate 8 Page 8 of 34
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was determined using solvents of increasing log P: diethylether (0.8), benzaldehyde (1.48), toluene
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(2.50), n-heptane (4.0) and decane (6.0), respectively. The solvents were added to the reaction
209
mixtures containing the free CRL and CRL−MWCNTs at a molar ratio of geraniol:propionic acid
210
of 5:1, unless specified otherwise.
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It has been described that relative proportions of the various substrates in a reaction mixture
214
is a factor that defines the physical and chemical properties of a reaction system and subsequently
215
influences the yield of product. The reaction mixtures were prepared at increasing molar ratio of
216
geraniol: propionic acid (molar ratio = 1:1, 2:1, 3:1, 4:1 and 5:1).
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217 2.5.3 Thermal Stability
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The thermal stability of CRL−MWCNTs was examined by incubating the biocatalysts in
220
sealed vials for 1 h at various temperatures ranging from 50 to 80°C, at increasing intervals of 10oC
221
each. After the stipulated period, the CRL−MWCNTs were left to cool to room temperature. Such
222
evaluation was attempted by introducing the thermally treated CRL−MWCNTs into the reaction
223
mixture that consisted of propionic acid and geraniol dissolved in n-heptane and incubated at 40°C
224
with constant stirring. The relative activities were determined and expressed as percentage of the
225
enzyme activity at varying temperatures.
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2.6
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Analysis of Geranyl Propionate using Gas Chromatography (GC) and Nuclear
228
Magnetic Resonance (NMR) Spectroscopy
229
Identification of geranyl propionate in liquid samples was carried out by Gas
230
Chromatography (GC) (Agilent Technologies 6890N) equipped with flame ionization detector
231
using a capillary column of fused silica INOWAX (30 m × 250 μm × 0.25 μm). Helium was used as
232
the carrier gas (1.0 mL/min) at constant pressure, the detector at 1.0 kV, split mode (1:20), and the 9 Page 9 of 34
injector whose temperature was maintained at 250°C. The temperature program used was as
234
follows: 50°C for 1 min; increment of 5°C/min up to 300°C; and the temperature was kept constant
235
for 1 min. Each sample was filtered prior to GC analysis. Nuclear magnetic resonance spectra
236
(NMR) were run on a Bruker Avance III-HD 700 MHz. spectrometer. CDCl3 was employed as the
237
solvent for all experiments. Samples were obtained by dissolving about 0.2 g of methyl oleate in
238
about 0.55 mL of CDCl3 and transferred to a standard 5 mm tube. Standard acquisition parameters
239
were used for all samples.
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Result and Discussion
243
3.1
Rationale for the functionalization of MWCNTs
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Stirring the MWCNTs in acid mixture of H2SO4 and HNO3, introduces the polar groups
245
(COO-) to the non-polar surface and tubular ends of the MWCNTs supports. This is to ensure that
246
the surfaces of the MWCNTs are primed for interaction with other polar moieties (NH2, O−H)
247
present on the CRL protein. The COO- moiety on the surface of MWCNTs anchor to the CRL
248
protein by attractions between the oppositely charged carboxyl moiety on the MWCNTs and the
249
hydrogen of the back-bone or side-chain of polar amino acids present on the outer surface of the
250
CRL protein [23]. On the other hand, poor attachment of the CRL proteins to the non-polar non-
251
functionalized MWCNTs was observed, an inference made from the high protein concentrations in
252
the phosphate buffer washings after immobilization of the free CRL. Conversely, the physical
253
adsorption of CRL to the F−MWCNTs was reasonably strong as the amount of detached proteins in
254
the reaction mixtures was not observed or negligible following the 6 h of stirring. The
255
CRL−MWCNTs recorded a lower enzyme activity than the free CRL corresponding to 792 U mg-1.
256
However, the CRL−MWCNTs biocatalysts were anticipated to be more stable and undergo a slower
257
decline in enzymatic activity when utilized under extended reaction period and high temperature.
258
Also, the acid functionalized CRL−MWCNTs were observably well dispersed when stirred in the
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reaction vessel as compared to the non-functionalized ones. The facile dispersability of the
260
CRL−MWCNTs is suggestive of proper mixing between the biocatalysts with the substrates, hence,
261
increases the propensity of effective enzyme−substrate collision to improve the esterification yield
262
[23].
264
3.2
Characterization of Immobilized Lipase (CRL−MWCNTs)
265
3.2.1 Fourier Transform Infra−Red Spectroscopy (FT-IR)
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FTIR is principally employed as a qualitative technique to assess the presence of functional
267
groups in a particular compound. The FT−IR spectra acquired for the as-synthesized MWCNTs,
268
F−MWCNTs and CRL−MWCNTs as well as the resultant wavenumbers are presented in Table 1.
269
The emergence of a broad band at 3428.01 cm-1 corresponding to the O−H stretching of the surface
270
group of the as-synthesized MWCNTs may be attributable to ambient atmospheric moisture that
271
was firmly bound to the MWCNTs [24]. A weak band which appeared at 1630.82 cm-1 in the
272
spectrum of as-synthesized MWCNTs is associated with the existence of conjugated C=C bonds
273
[25]. Following oxidation, the presence of a C−O stretching due to the successful oxidation of sp2
274
hybridized carbon in the as-synthesized MWCNT to sp3 [26] is established by the emergence of a
275
new peak at 1112.73 cm-1 in the spectra of the F−MWCNTs. Correspondingly, the O−H stretch
276
moved from 3428.01 cm-1 to 3445.22 cm-1, strongly implying the development of covalent bonds
277
between the O−H and the sidewalls of the MWCNTs [27]. In addition, the C=O stretching vibration
278
of the carboxylic group represented by a peak (1634.70 cm-1) was observed attributable to
279
adaptation of additional single bond character through resonance by the C=O of the introduced
280
acidic surface groups. Conjugation of O=C with the C=C of the MWCNTs had resulted in the
281
formation of a predominantly C−O functional group, an observable fact that eventually led to the
282
decline in the vibration frequency of the C=O group in the spectra of F−MWCNTs. Furthermore,
283
the appearance of a new peak at 1202.46 cm-1 is attributed to the C−O of the COOH moiety [28].
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In addition, a band (1634.70 cm1) for C=O stretching vibration of the carboxylic group was
285
detected in the F−MWCNTs. Such band was apparently substituted by a new band at lower
286
wavelength (1620.71 cm-1) that corresponded to the C=O stretching of amide for CRL−MWCNTs,
287
suggesting excellent adsorption bands of proteins [25]. The existence of a cyano functional group
288
(C−N) was further substantiated by the appearance of a new band (1032.96 cm-1). Additionally, the
289
CH2 and N−H functional groups were characterized by explicit bands appearing at 2929.41 cm-1 and
290
3420.59 cm-1, respectively. It was revealed that the differing spectra for MWCNTs and
291
CRL−MWCNTs were seen within the spectral region for carboxylate functional groups, in the
292
range of 900−1200 cm-1 [29]. The FT−IR results further illustrated that following immobilization, a
293
rather strong peak detected at 1125.04 cm-1 in the spectrum for free CRL correlated with the
294
vibrations of C−O of the carboxylate moiety was found expanding, reducing and shifting towards
295
lower frequency (1116.03 cm-1). Such finding was in accordance with the previously described
296
work [25], proving that the CRL was successfully immobilized onto the outer face of the
297
F−MWCNTs. The fact that an adsorption band at 2929.41 cm-1 was present in the CRL−MWCNTs
298
alone (Figure 1c), while similar band was evidently missing in the free CRL as well as
299
F−MWCNTs, it can be interpreted that the immobilization process of CRL onto the surface of the
300
F−MWCNTs had been successful.
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302
3.2.2 Thermal Gravimetric Analysis (TGA)
303
The degree of alterations in physical and chemical properties of the biocatalysts as a function of
304
temperature or time was determined by TGA. The method is often used to monitor any reaction that
305
involves oxidation or dehydration. By measuring the decomposition or weight loss of the sample,
306
the analysis via TGA would reveal the change in thermal stability of the support used in the
307
immobilization process, indicating if it has been modified [30]. In this investigation, TGA in the
308
range of 30−800°C was employed to examine the mass loss percentage of F−MWCNTs and
309
CRL−MWCNTs, respectively (Figure 1). In general, combustion of the carbon nanotubes and the 12 Page 12 of 34
carbonaceous contaminants occurs at temperatures above 300oC. Any mass loss observed at
311
temperature below 400oC is usually associated with the evaporation of absorbed water or the
312
presence of surface functional groups. The thermogram for F−MWCNTs (Figure 1a) exhibited a
313
steady, uniform and multistage mass loss over the range of 30−150°C, 150−350°C and 350−800°C.
314
These changes corresponded to a 30% reduction in mass which strongly indicated the degradation
315
of various functional groups present on the surface of F−MWCNTs. Correspondingly, the first mass
316
loss (30−150°C) of 5% reduction may be attributable to the evaporation of hydrated water group
317
[31]. Further increment in temperature (150−350°C) rendered further decline in the mass of
318
F−MWCNTs by as much as 25%, implying degradation of surface groups on the MWCNTs
319
following acid treatment [32], confirming successful attachment of the –COOH groups on the walls
320
of the MWCNTs [31,33]. It has been reported that ccombustion of carbon materials would only
321
transpire when temperature reaches at least 400°C [33]. Finally, no further mass loss was found
322
beyond 350°C, indicative of thermally stable carbon that was created during removal of graphitic
323
and catalytic metal particles during the oxidation of the purified MWCNTs [33].
d
M
an
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cr
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310
te
324
As for the CRL−MWCNTs (Figure 1b), a 38% mass loss was recorded as a result of the
326
decomposition of the adsorbed CRL protein on the surface of the F−MWCNTs. An initial 6.5%
327
mass loss (30−150°C) was due to the elimination of both surface hydroxyl groups and organic
328
structure of lipase [34]. In comparison to F−MWCNTs, the decomposition curve of
329
CRL−MWCNTs was distinctively dissimilar between 250−400°C when compared with that of
330
F−MWCNTs. This was attributable to the thermal decomposition of lipase [35] present on the
331
surface of CRL−MWCNTs biocatalysts. Based on the calculations, approximately 13 wt% of CRL
332
protein was successfully attached to the surface of the F−MWCNTs following the immobilization
333
process.
Ac ce p
325
334 335 13 Page 13 of 34
336
3.3
Transmission electron microscopy (TEM) and field emission scanning electron microscopy (FESEM)
338
Analysis on the morphological surface of the as-synthesized MWCNTs, F−MWCNTs and
339
CRL−MWCNT biocatalysts observable through TEM and FESEM are depicted in Figure 2. The
340
image of the as-synthesized MWCNTs illustrated closely attached open-ended tubular structures
341
with smooth surfaces that revealed the presence of contaminants on the surface of nanotubes
342
(Figure 2a). Since the contaminants on the surface of the as-synthesized MWCNTs (graphitic, metal
343
catalytic and amorphous carbon) may interfere and hamper the potential characters of the carbon
344
nanotubes, it was pertinent that these contaminants were removed [36].
us
cr
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337
an
345
Following functionalization with the H2SO4 and HNO3 acid mixtures, the diameter of
347
MWCNTs was significantly increased, revealing noticeable rough surfaces that were attributable to
348
acid related eroded nanotubes [37]. The increased diameter of the F−MWCNTs strongly signified
349
the successful attachment of acid functional groups (COO-) on the surface of MWCNTs (Figure
350
2b), hence, corroborating previous findings reported by other studies [38]. In addition, the evidently
351
shorter length of the F−MWCNTs after treatment with the acid mixture was consistent with the
352
earlier studies [36, 39]. The perceived change in the highly tangled long MWCNTs into shorter,
353
open ended pipes strongly indicated that acid treatment on the nanotubes can potentially create
354
copious number of COO- at the open ends for linkage with chemical and biological [40]. According
355
to Khani and Moradi (2013), increased oxidation time and elevated temperatures were most likely
356
to destroy the structure of MWCNTs [40], generating surface imperfections on the MWCNTs that
357
subsequently affected the properties of the nanotubes. The increased diameter of CRL−MWCNTs
358
as compared to the F−MWCNTs CRL was physical an evidence of CRL on the surface MWCNTs,
359
(Figure 2c) and therefore, confirming the successful immobilization of the CRL on the MWCNTs
360
support.
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361 14 Page 14 of 34
362
3.3
Enzymatic production of geranyl propionate by free CRL and CRL−MWCNTs
363
3.3.1 Effect of Solvents (Log P) The partition-coefficient (log P) represents the ratio of concentrations of a compound in a
365
mixture of two immiscible phases consisting of 1−octanol and water at equilibrium, a key factor
366
that can influence the relative solubility, activity and specificity of the substrates [41] in an
367
enzymatic reaction. Figure 3 represents the reaction profile based on the increasing log P values of
368
organic solvents; hydrophilic log P < 1.4, diethyl ether (0.85); medium hydrophobicity: 1.4 < log P
369
< 3.5 (benzaldehyde 1.48, toluene 2.50) and hydrophobic: log P > 3.5 (n-heptane 4.0, decane 6.0)
370
in the production of geranyl propionate catalyzed by the free CRL and CRL−MWCNTs.
us
cr
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364
an
371
The results showed an almost 2−fold increased conversion of geranyl propionate with
373
increasing hydrophobicity of solvents for the free CRL and CRL−MWCNTs. The highest
374
conversion was achieved when n-heptane was used as solvent for both the free CRL (28.1%) and
375
CRL−MWCNTs (51.3%) catalyzed reactions, confirming good compatibility of log P > 4 solvents
376
in enzymatically catalyzed reactions [42]. It was observed that n−heptane (log 4.0) was a suitable
377
solvent for the enzymatic production of geranyl propionate catalyzed by both free CRL and
378
CRL−MWCNTs. The solvent n−heptane sufficiently dissolved and suspended the substrates in the
379
reaction vessel without adversely affecting the catalytically active form of the CRL protein.
380
Previous studies have described that the high log P solvents would render better disassociation of
381
weak organic acids and prevent the stripping of the essential water layer surrounding the enzyme
382
[42], during the course of reaction. In this context, it is pertinent to point out that using n−heptane
383
as solvent can sustain the protein structure and catalytically active conformation of the CRL
384
[42,43]. Hence, the observed higher product yield [41] for the free CRL and CRL−MWCNTs
385
catalyzed esterification production of geranyl propionate in n−heptane was obtained.
Ac ce p
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386
15 Page 15 of 34
It is generally accepted that hydrophilic, namely, polar solvents tend to strip off essential
388
water from enzyme molecules that lead to the inactivation of the enzyme activity [44]. Considering
389
the hydrophobicity of the solvents employed in this investigation, it was expected that the activity
390
of the enzyme to improve with the reduction of solvent hydrophilicity. Although, it has been
391
described in literature that polarity cannot be used as the only measure when the effects of solvents
392
on enzyme activity of immobilized lipases are tested, the results in this study proved otherwise. The
393
notably low enzyme activity in hydrophilic solvents (log P < 2) rendering lowest conversion of the
394
geranyl propionate strongly suggests the dehydrating effect of such solvents on the enzyme active
395
site [45]. Enzyme deactivation on the free CRL and CRL−MWCNTs was reduced, reflected in the
396
higher production of methyl oleate when hydrophobicity of solvents used in the reactions was
397
increased. Apart from increased penetration of solvent molecules into the lipase protein, hydrophilic
398
solvents also promoted deformation and increased flexibility of the lipase surface and loss of
399
enzyme stability. Correspondingly, poor predictability for various hydrophilic solvents (log P 2.0–
400
4.0) has been reported [46]. In this perspective, it is vital to highlight that the hydrophilic solvents
401
clearly had a negative effect on the free CRL and CRL−MWCNTs than the hydrophobic ones for
402
the esterification production of geranyl propionate.
404 405
cr
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Ac ce p
403
ip t
387
3.3.2 Effect of Molar Ratio of Alcohol to Acid Previous studies have illustrated that equilibrium of an enzymatic reaction can be
406
manipulated to favor high conversion of the ester product, through proper selection of molar ratio of
407
the reacting alcohol or acids or by removal of the products from the reaction mixture [47]. In some
408
cases, the use of excess alcohol is required to reduce the intrinsic denaturing properties of acids on
409
lipases. Presence of excess acid in a reaction, to a certain extent, can deactive the lipase particularly
410
whenever the pH drops below pH 2. Considering that high concentrations of acid, alcohol or both
411
are significant factors to influence the equilibrium of enzymatic esterification reactions [48], such
412
factors were examined in this study. The effect of molar ratio of alcohol to propionic acid for the 16 Page 16 of 34
413
enzymatic production of geranyl propionate was monitored for molar ratio geraniol: propionic in
414
the range of 1:1 to 5:1.
415 Figure 4 depicts the optimum molar ratio of geraniol to propionic acid as 5:1 corresponding
417
to 28% and 47% of geranyl propionate for the free CRL and CRL−MWCNTs, respectively. The
418
conversion percentage of geranyl propionate for the CRL−MWCNTs significantly increased over
419
the free CRL by 2−fold, indicating higher activity of the CRL−MWCNTs following immobilization
420
onto the F−MWCNTs. The elevated activity of the CRL−MWCNTs over the free CRL was
421
anticipated as similar enhancements reported for the commercially immobilized lipase i.e.
422
Novozyme 435 [4]. Conversion of geranyl propionate was the lowest for both free CRL and
423
CRL−MWCNTs at the molar ratio of 1:1, corresponding to 13.6% and 32.4%, respectively. The
424
low conversion of geranyl propionate at equal proportions of geraniol to propionic acid can be
425
attributed to the inhibition of the CRL activity due to the presence of surplus acid in the reaction
426
medium that adversely decreased the initial rate of reaction [49]. However, the noteworthy higher
427
conversions of the ester in all CRL−MWCNTs catalyzed reactions confirm the enhanced stability of
428
the adsorbed CRL protein. The additional network of interaction i.e. van der Waals forces,
429
hydrophobic interactions and hydrogen bond formed between the surface of the CRL and the acid
430
groups on the surface of the MWCNTs reduce the flexibility of the CRL protein. Hence, the
431
structure of CRL is less predisposed to the inherently denaturing effects of the alcohol and acid
432
substrates that can disrupt the three-dimensional active form of the CRL protein, above all, during
433
prolonged reaction time.
Ac ce p
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an
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416
434 435
3.3.3 Thermal Stability
436
Thermal stability of an enzyme is a vital aspect of bioprocess reactions operating at high
437
temperatures in industrial setting. A high reaction temperature is often preferred as it promotes
438
substrates solubility and reduces viscosity to which mass transfer limitation can be avoided [50,51]. 17 Page 17 of 34
In this study, thermal stability of both free CRL and CRL−MWCNTs was investigated after
440
incubating both forms of CRL for 2 h at various temperatures (50−80oC) and the data are illustrated
441
in Figure 5. It was revealed that the thermal stability for CRL−MWCNTs was enhanced over the
442
free CRL by almost 2−fold with the former retaining at least 22.6−32.1% of activity even at the
443
highest temperature of 80°C when compared with that of free CRL (12.6−18.1%). Such observation
444
indicates the improved robustness of the biocatalysts to catalyzed reactions under elevated
445
temperature. The higher thermal stability of CRL−MWCNTs over the free CRL potentially
446
conferred by additional electrostatic interactions and hydrogen bonds between the enzyme and the
447
support, favorably influencing the thermal stability [52,53] of the immobilized lipase.
us
cr
ip t
439
On the other hand, the activity of the free CRL was diminished by 46%, retaining only
449
12.2% of enzyme activity. Again, the considerable improvement in thermal stability of
450
CRL−MWCNTs over the free CRL during reaction can be associated with the additional
451
interactions between the molecules of CRL and MWCNTs that rendered the protein structure of
452
CRL−MWCNTs to be more rigid. Therefore, the increased structural rigidity resulted in less
453
yielding to the temperature-induced change in the lipase structure. Conformational transitions of the
454
CRL are averted [54], resulting in a more stable structure even at high reaction temperatures.
455
Hence, immobilized CRL is able to retain its catalytically competent form for a longer period of
456
time particularly during extensive reaction times, even under elevated temperatures [50]. Similar
457
thermal stabilization has also been described for powdered layered double hydroxides (LDH)
458
known for its use as nanocarriers or catalyst support for functional molecules [12].
Ac ce p
te
d
M
an
448
459 460
3.4
Analysis of Geranyl Propionate Using Gas Chromatography (GC) and Nuclear
461
Magnetic Resonance spectroscopy
462
The production of geranyl propionate by the free CRL and CRL−MWCNTs (Schematic 1)
463
was monitored over a 12 h period and samples were withdrawn and identified by GC analysis.
464
Separations of the components in the sample are based on the interaction strength of compounds 18 Page 18 of 34
with the stationary phase as well as other factors i.e. boiling point and polarity of compounds [55].
466
Prior to analysis, the boiling point for each component was determined as: propionic < geraniol <
467
geranyl propionate. According to Figure 6, geranyl propionate has a longer retention time as
468
compared to geraniol and propionic acid. In Figure 6a, the initial composition of the reaction only
469
revealed a peak at 10.660 min. Following a 12 h incubation period, an existence of a new peak at
470
13.782 min (Figure 6b) was observed. The new peak represents the liberated geranyl propionate that
471
was enzymatically synthesized from geraniol and propionic acid in reactions catalyzed by both free
472
CRL and CRL−MWCNTs. Similar retention time for GC analysis of geranyl propionate has been
473
previously described [55].
us
cr
ip t
465
Analysis of 1H and 13C NMR revealed peaks that were characteristics of geranyl propionate
475
(Figure 7a and Figure 7b). The 1H NMR for geranyl propionate is as follows: (700 MHz, CDCl3): δ
476
(ppm): 1.15 (3H, t, -CH3), 1.60 (3H, s, -CH3), 1.69 (3H, s, -CH3), 1.71 (3H, s, -CH3), 2.15 (2H, t, -
477
CH2), 2.17 (2H, t, -CH2), 2.35 (2H, m, -COCH2), 4.60 (2H, d,-OCH2), 5.08 (1H, t, - =CH), 5.35 (1H,
478
t, - =CH). The
13
C NMR shows characteristic peaks i.e. (700 MHz, CDCl3): δ (ppm): (174.49 (C-
te
479
d
M
an
474
3=O), 142.03 (C6), 131.78 (C10), 123.74 (C9), 118.42 (C-5), 61.24 (C-4), 38.61 (C-7), 27.67 (C-2),
481
26.84(C-8 and C-11), 17.86 (C-13) and 8.12 (C-1). Hence, the 1H NMR and
482
geranyl propionate confirmed presence of a 13-carbon moiety that matched the structure of geranyl
483
propionate.
484 485
4.
Ac ce p
480
13
C NMR spectra of
Conclusions
486
The improved activity and stability of CRL−MWCNTs over the free CRL to catalyze
487
production of geranyl propionate was attributable to the enhanced structural integrity and
488
mechanical strength of the CRL protein conferred by the F−MWCNTs. In consequence, the CRL
489
protein became more rigid, resistant towards premature unraveling as well as sustaining its
490
catalytically competent structure for longer period of time. Such property is useful especially when 19 Page 19 of 34
dealing with extensive esterification process under harsh conditions such as extreme temperatures
492
and pH. The 2−fold enhanced activity and increased thermal robustness of the CRL−MWCNTs
493
biocatalysts subsequently resulted in a more efficient biosynthesis of geranyl propionate. This work
494
proves that physical adsorption of CRL onto F−MWCNTs has improved the stability and activity of
495
CRL as a biocatalyst. Considering the costs of 10 g of commercial enzymes i.e. Novozyme
496
(>USD900) and Lipozyme (USD400), the developed CRL−MWCNTs biocatalysts in this study
497
would potentially be an economical and sustainable alternative for the enzymatic production of
498
geranyl propionate as well as to produce other similarly important commercial esters.
us
cr
ip t
491
499
501
an
500 Acknowledgments
The authors wish to express their gratitude to the Faculty of Science, Universiti Teknologi
503
Malaysia for their facilities. This work was funded by the Ministry of Higher Education of Malaysia
504
under the Exploratory Research Grant Scheme (ERGS R.J130000.7826.4L132).
505 506 References
508
[1]
509 510
Ac ce p
507
te
d
M
502
S.M. Radzi, W.A.F Mustafa, S.S. Othman, H.M. Noor, World Acad. Sci. Eng. Technol. 59 (2011) 677–680.
[2]
511
F.X. Malcata, H.R. Reyes, H.S. Garcia, C.G. Hill, C.H. Amundson, J. Am. Oil Chem. Soc. 67 (1990) 890–910.
512
[3]
T.W. Charpe, V.K. Rathod, Waste Manage. 31 (2011) 85–90.
513
[4]
N. Paroul, L.P. Grzegozeski, V. Ciaradia, H. Treichel, R.L. Cansian, J. Vladimir Oliveira, D.
514 515 516
de Olivera, J Chem. Technol. Biotechnol. 12 (2010) 1636–1641. [5]
D.M. Solano, P. Hoyos, M.J. Hernaiz, A.R. Alcantara, J.M. Sanchez-Montero, Biores. Biotechnol. 115 (2012) 196−207. 20 Page 20 of 34
517
[6]
P. Anastas, N. Eghbali, Chem. Soc. Rev. 39 (2−010) 301–312.
518
[7]
B.M. Nestl, B.A. Nebel, B. Hauer, Curr. Opin. Chem. Biol. 15 (2011) 187–193.
519
[8]
J. Tao, R.J. Kazlaukas, 2011. Biocatalysis for Green Chemistry and Chemical Process
521 522
Development. John Wiley & Sons Ltd., Hoboken, New Jersey. [9]
R.B. Salah, H. Ghamghui, N. Miled, H. Mejdoub, Y. Gargouri, J. Biosci. Bioeng. 103 (2007)
ip t
520
368–372.
[10] G. Zhou, C. Wu, X. Jiang, J. Ma, H. Zhang, H. Song, Process Biochem. 47 (2012) 953−959.
524
[11] R.A. Wahab, M.B. Abdul Rahman, M. Basri, R.N.Z. Raja Abdul Rahman, A.B. Salleh, T.C.
Leow. Biotech. Biotech. Eq. 28 (2014) 1065−1072.
us
525
cr
523
[12] Y. Kuang, L. Zhao, S. Zhang, F. Zhang, M-D. Dong, S. Xu. Materials 3 (2010) 5220-5235.
527
[13] J. Song, M. Chen, V.R. Regina, C. Wang, R.K. Meyer, E. Xie, C. Wang, F. Besenbacher, M. Dong. Adv. Eng. Mat. 14 (2012) B240-B246.
M
528
an
526
[14] K.E. Jaeger, T. Eggert, Curr. Opin. Biotechnol. 13 (2001) 390–397.
530
[15] M. Shim, N.W.S. Kam, R.J. Chen, Y.M. Li, H.J. Dai, Nano. Letts. 22 (2002) 28−58.
531
[16] P. Zhang, DB. Henthorn, AIChE J. 56 (2010) 1610–1615.
532
[17] M. Cesar, M.P. Jose, F.L. Gloria, M.G. Jose, F.L. Roberto, Enzym. Microb. Technol. 40
te
Ac ce p
533
d
529
(2007)1451–1463.
534
[18] W. Tischer, V. Kascher, Trends Biotechnol. 17 (1999) 326−335.
535
[19] D. Norouzian, Iran. J. Biotechnol. 1 (2003) 197−206.
536
[20] P.C. de Oliveira, G.M. Alves, H.F. de Castro, Biochemical Eng. J. 5 (2000) 63−71.
537
[21] M.M. Bradford, Anal. Biochem. 72 (1976) 248–254.
538
[22] M.A.P. Langone, G.J. Sant'anna, G. J. Appl. Biochem. Biotechnol. 77-79 (1999) 759−770.
539
[23] N.R. Mohamad, N.A. Buang, N.A. Mahat, Y.Y. Lok, F. Huyop, H.Y. Aboul-Enein, R.A.
540
Wahab. Enzym. Microb. Technol. 72 (2015a) 49−55.
541
[24] T. Ramanathan, F.T. Fisher, R.S. Ruoff, L.C. Brinson, J. Mat. Chem. 17 (2005) 1290−1295.
542
[25] N.Z. Prlainoc, D.I Bezbradica, J. Ind. Eng. Chem.19 (2013) 279−285. 21 Page 21 of 34
543 544
[26] I.V. Pavlidis, T. Tsoufis, A. Enotiadis, G. Gournis, H. Stamatis, Advan. Eng. Mat. 12 (2010) 5.
545
[27] L. Zhang, V.U. Kiny, H. Peng, J.J. Zhu, Mat. Chem. 16 (2004) 2055−2061.
546
[28] M.F. Alkhatib, M.E.S. Mirghani, I.Y. Qudsieh, I.A.F. Husain, J. Appl. Sci. 10 (2010) 2705−2708.
ip t
547
[29] A. Natalello, D. Ami, S. Brocca, M. Lotti, S.M. Doglia. Biochem. J. 85 (2005) 511−517.
549
[30] N.R. Mohamad, N.H. Che Marzuki, N.A. Buang, F. Huyop, R.A. Wahab. Biotechnol. Biotec. Eq. 29(2015b) 205−220.
us
550
cr
548
[31] V. Datsyuk, M. Kalyva, K. Papagelis, J. Parthemios, D. Tasis, Carbon. 46 (2008) 833−840.
552
[32] J.L. Vicente, A. Albesa, J.L. Llanos, E.S. Flores, A.E. Fertutta, J Argent. Chem. Soc. 98
554 555
(2011) 29−38.
[33] T. Raghavendra, A. Basak, L.M. Manoca, A.R. Sha, D. Madamwar, Biores. Technol. 140
M
553
an
551
(2013) 103-110.
76
[34] H. F. Castro, M.L.C.P. Silva, G.L.J.P. Silca, Braz. J. Chem. Eng. 17 (2000) 4−7.
557
[35] Y. Shi, F, Jin, Y. Wu, F, Yan, X. Yu, Y. Quan, Dalian Institute of Light Industry. 13
559 560 561 562
te
(1997)111−113.
Ac ce p
558
d
556
[36] R. Yudianti, H. Onggo, R. Sudirman, Y. Saito, T. Iwata, J.I. Azuma, Open Mat. Sci. J. (2011) 5:242−247.
[37] M.F. Alkhatib, M.E.S. Mirghani, I.Y. Qudsieh, I.A.F. Husain, J. Appl. Sci. (2010) 10(21) 2705−2708.
563
[38] M.R. Joha, S.H.M. Suhaimy, Y. Yusof, Appl. Surf. Sci. (2014) 450−454.
564
[39] I. Mazova, V.L. Kuznetsova, I.A. Simonovaa, A.I. Stadnichenkoa,. Appl. Surf. Sci. (2012)
565
6272−6280.
566
[40] H. Khani, O. Moradi, J. Nanostruc. Chem. (2013) 3−73.
567
[41] S. Gogoi, N.N. Dutta, Ind. J. Chem. Technol. 16 (2009) 209−215.
568
[42] C. Laane, S. Boeren, K. Vos, C. Veeger, Biotech. Bioeng. 30 (1987) 81−87. 22 Page 22 of 34
569 570
[43] J. Song, D. Kahveci, M. Cheng, Z. Guo, E. Xie, X. Xu, F. Besenbacher, M. Dong. Langmuir 28 (2012), 6157−6162. [44] S.H. Krishna, N.G. Karanth, Catal Rev. 4 (2002) 499−591.
572
[45] A. Zaks, A.M. Klibanov, Science. 22 (1984) 1949−1951.
573
[46] L. Yang, J.S. Dordick, S. Garde, Biophys. J. 87 (2004) 812−821.
574
[47] M. Trani, F. Ergan, G. Andre, J. Am. Oil Soc. 68 (1991) 20−22.
575
[48] H. Stamatis, P. Christakopolous, D. Kekos, B.J. Macris, F.N. Kolisis, J. Mol. Catal. B:
cr
Enzym. 4 (1998) 229−236.
us
576
ip t
571
[49] P. Pires Cabral, M.M.R. Fonseca, S.F. Dias, Biochem. Eng. 43 (2009) 327−332.
578
[50] M.B. Abdul Rahman, N. Chaibakhsh, Basri M, R.N.Z. Abdul Rahman, A.B. Salleh, S. Md
580 581
Radzi, J. Chem. Technol. Biotechnol. 83 (2008) 1534−1540.
[51] R. Dave, D. Madamwar, in: C. Larroche, A. Pandey, C.G. Dussap, (Eds.), Current Topics on
M
579
an
577
Bioprocesses in Food Industry. Asiatech Publishers Inc., New Delhi, 2006, pp. 70–80. [52] L. Wang, L. Wei, Y. Chen, R. Jiang. J. Biotechnol. 150 (2010) 57–63.
583
[53] K. Zhu, A. Jutila, E.K.J. Tuominen, P.K.J. Kinnunen, Pro Sci. 10 (2001) 339–351.
584
[54] E. Yilmaz, K. Can, M. Sezgin, M. Yilmaz, Biores. Technol.102 (2001) 499−506.
585
[55] R.R. Thanighai, B. Nambikkairaj, P. Sivamani, Ind. J. Appl. Res. 3 (2013) 375−379.
te
Ac ce p
586
d
582
23 Page 23 of 34
586 List of Legends Table 1
The frequency table for FTIR analyses for a) Raw MWCNTs, b) FMWCNTs and c) CRL−MWCNTs Thermogram of the TGA performed on (a) F−MWCNTs and (b)
ip t
Figure 1
CRL−MWCNTs.
Transmission electron microscope (TEM) and field emission scanning
cr
Figure 2
F−MWCNTs and (c) CRL−MWCNTs.
The effect of solvent (log P): diethyl ether (0.8), benzaldehyde (1.48),
an
Figure 3
us
electron microscope (FESEM) images of (a) as-synthesized MWCNTs, (b)
toluene (2.5), n-heptane (4.0) and decane (6.0) in the enzymatic synthesis
M
geranyl propionate. [Temperature: 55°C, molar ratio propionic acid/geraniol (1:5), enzyme amount 5 mg/mL for CRL−MWCNTs and 10 mg/mL free
The effect of the molar ratio of geraniol to propionic acid (1:1 to 5:1) on the
Ac ce p
Figure 4
te
n-heptane].
d
CRL, molecular sieves: 100 mg, stirring speed 200 rpm, time: 12 h, solvent:
synthesis of geranyl propionate by CRL−MWCNTs at reaction conditions:
[Temperature: 55°C, enzyme amount: 5 mg/mL for CRL−MWCNTs and 10
mg/mL free CRL, molecular sieves: 100 mg, stirring speed: 200 rpm, time: 12 h, solvent: n-heptane].
Figure 5
The effect of thermal stability on the synthesis of geranyl propionate by CRL−MWCNTs. [Temperature: 40°C; molecular sieves: 100 mg, stirring speed: 200 rpm, time: 12 h, solvent: n-heptane].
Figure 6
Gas chromatogram of sample from the enzymatic esterification of geraniol and propionic acid catalyzed by the free CRL and CRL−MWCNTs at 0 h
24 Page 24 of 34
and 12 h. Figure 7
The a) H-NMR and b) C-NMR spectra of the purified esterification product geranyl propionate synthesized by the CRL-MWCNTs. The reaction scheme for the lipase-catalyzed esterification to produce geranyl propionate from geraniol and propionic acid.
cr
587
ip t
Schematic 1
Ac ce p
te
d
M
an
us
588
25 Page 25 of 34
588 Table 1: Functional group
a)
1630.82 3428.01
conjugated C=C bonds O−H stretching
b)
1112.73 1202.46 1634.70 3445.22
C−O stretching C−O stretching C=O stretching O−H stretching
c)
1032.96 1116.03 2929.41 3420.59
C−N stretching C−O stretching CH2 bending N−H stretching
an
us
cr
ip t
Frequency (cm-1)
Ac ce p
te
d
M
589 590
26 Page 26 of 34
Schematic 1
O O + OH geraniol
OH
O
CRL-MWCNTs
+ H2O
or free CRL geranyl propionate
propionic acid
Ac
ce pt
ed
M
an
us
cr
ip t
Schematic 1:
Page 27 of 34
Figure 1
(
M
an
us
cr
ip t
a)
(
Ac
ce pt
ed
b)
Figure 1:
Page 28 of 34
Ac c
ep te
d
M
an
us
cr
ip t
Figure 2
Figure 2:
Page 29 of 34
Figure 3
60 FREE-CRL CRL-MWCNTs
40
ip t
30
20
cr
Conversion (%)
50
0
1.48
2.5
4
6
an
0.8
us
10
M
Solvent (Log P)
Ac
ce pt
ed
Figure 3:
Page 30 of 34
Figure 4
60 FREE CRL CRL−MWCNTs
50
40
ip t
30
cr
20
0 1:2
1:3
1:4
1:5
an
1:1
us
10
Molar ratio acid:alcohol
Ac
ce pt
ed
M
Figure 4:
Page 31 of 34
Figure 5
40 FREE CRL CRL-MWCNTs
ip t
20
cr
10
us
Residual activiry (%)
30
0
60°C
70°C
80°C
an
50°C
Temperature
Ac
ce pt
ed
M
Figure 5:
Page 32 of 34
Figure 6
Sample at 12 h
Geranyl propionate
Ac
ce pt
ed
(b)
M
an
us
cr
ip t
(a) Sample at 0 h
Figure 6:
Page 33 of 34
Figure 7
M
an
us
cr
ip t
a)
Ac
ce pt
ed
b)
Figure 7:
Page 34 of 34