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Synthesis of magnetic gold mesoporous silica nanoparticles core shell for cellulase enzyme immobilization: Improvement of enzymatic activity and thermal stability
Accepted Manuscript Title: Synthesis of Magnetic Gold Mesoporous Silica Nanoparticles Core Shell for Cellulase Enzyme Immobilization: Improvement of Enzymatic Activity and Thermal Stability Authors: Elaheh Poorakbar, Abbas Shafiee, Ali Akbar Saboury, Behzad Lame Rad, Kamyar Khoshnevisan, Leila Ma’mani, Hossein Derakhshankhah, Mohammad Reza Ganjali, Morteza Hosseini PII: DOI: Reference:
S1359-5113(17)31976-1 https://doi.org/10.1016/j.procbio.2018.05.012 PRBI 11348
To appear in:
Process Biochemistry
Received date: Revised date: Accepted date:
24-12-2017 14-5-2018 15-5-2018
Please cite this article as: Poorakbar E, Shafiee A, Saboury AA, Rad BL, Khoshnevisan K, Ma’mani L, Derakhshankhah H, Ganjali MR, Hosseini M, Synthesis of Magnetic Gold Mesoporous Silica Nanoparticles Core Shell for Cellulase Enzyme Immobilization: Improvement of Enzymatic Activity and Thermal Stability, Process Biochemistry (2018), https://doi.org/10.1016/j.procbio.2018.05.012 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Synthesis of Magnetic Gold Mesoporous Silica Nanoparticles Core Shell for Cellulase Enzyme Immobilization: Improvement of Enzymatic Activity and Thermal Stability
Elaheh Poorakbara,b,c, Abbas Shafieeb, Ali Akbar Sabouryc*, Behzad Lame Rada, Kamyar Khoshnevisand, Leila
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Ma’manie, Hossein Derakhshankhahf, Mohammad Reza Ganjalig, Morteza Hosseinih a
Department of Biology, Faculty of Sciences, Payame Noor University, P.O.Box: 19395-3697, Tehran, Iran
b
Department of Pharmaceutics, Tehran University of Medical Science, Tehran, Iran
c
Institute of Biophysics and Biochemistry, University of Tehran, Tehran, Iran
d
Biosensor Research Center, Endocrinology and Metabolism Molecular-Cellular Sciences Institute, Tehran University of
Medical Sciences, Tehran, Iran
Department of Nanotechnology, Agricultural Biotechnology Research Institute of Iran (ABRII), Karaj, Iran
f
Pharmacutical Sciences Research Center, Kermanshah University of Medical Sciences, Kermanshah, Iran.
g
Center of Excellence in Electrochemistry, Faculty of Chemistry, University of Tehran, Tehran, Iran
h
Department of Life Science Engineering, Faculty of New Sciences & Technologies, University of Tehran, Tehran, Iran.
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(*Corresponding Author: [email protected])
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Graphical abstract
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Highlights
Novel magnetic gold core shell nanoparticles as a support material for cellulase immobilization were fabricated.
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The binding efficiency of cellulase to the support matrix was 76%.
Thermal stability and activity of the immobilized cellulase was improved upon immobilization.
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The immobilized cellulase was reusable for five cycles.
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Abstract
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Magnetic gold mesoporous silica nanoparticle core shells (mAu@PSNs) were fabricated as a support and their size, morphology and structure was further characterized by X-Ray diffraction (XRD),
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vibrating sample magnetometer (VSM), scanning electron microscopy (SEM), energy dispersive X-ray
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analysis (EDAX), transmission electron microscopy (TEM), dynamic light scattering (DLS) and thermal gravity analysis (TGA). Cellulase (CEL) immobilization on mAu@PSNs was performed via
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covalent bonding. Fourier transform infrared (FTIR) spectroscopy confirmed the successful binding of
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enzyme to mAu@PSNs while Bradford assay determined the binding efficiency to be 76%. The enzyme activity was measured at different pHs and temperatures by FPase method using Whatman
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filter paper as the substrate. The immobilized enzyme maintained 58% of its initial catalytic activity after nine hours. In this research, a new nano-system was designed as a solid support for cellulase immobilization which enhanced its thermal stability and facilitated its long term storage. In addition, the immobilized enzyme can be applied in a broader temperature and pH ranges while enzyme separation can be simply carried out by an external magnet. 2
Keywords: cellulase activity; thermal stability; enzyme immobilization; magnetic nano-particles; covalent bonding.
1. Introduction
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Natural resources shortage as well as the environmental pollution caused by fossil fuels compels us to look for new renewable energy resources [1]. Cellulosic materials such as biomass have been employed as eco-friendly sustainable energy resources in biofuels production (for example bioethanol) [2]. To produce such materials, it is necessary to modify the biomass’s chemical and structural properties to prevent its disassembly [3, 4]. Cellulose, the insoluble polymer of glucoside units, is the main source of
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carbon and one of the most abundant resources of renewable biomass [5, 6]. It can be obtained from the
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forests as well as the agricultural wastes. Cellulose is a suitable low-cost substrate for obtaining
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fermentable sugars with various industrial applications [7].
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Several physical, chemical and biological methods have been employed for hydrolysis of cellulosic
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materials. The enzymatic hydrolysis of cellulose has gained much attention because of its mild reaction
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conditions [8-10]. Microorganisms such as fungi [11, 12], bacteria [13-15] and yeasts [16] produce cellulose hydrolyzing enzyme, cellulase. Fungal cellulases are the most powerful cellulose hydrolyzing
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enzymes [17-19]. Several studies have reported the industrial applications of cellulase since it plays a
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key role in bio-conversion of many bio-substrates to produce bio-fuels [20-22]. In the past decades, research on cellulases have increased due to the interest in producing bioethanol from lignocellulosic
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biomass [1, 2, 4, 19, 23]. Cellulase is a biomolecular complex including three different enzymes (endo-cellulase,
exo-cellulase or cellobiohydrolase, and β-glucosidase) which produces soluble sugar from cellulolytic substrates [24]. It is also the second most commonly used industrial enzyme in various fields including agriculture, food, animal feed, brewery and wine, textile, detergents, pulps and papers industries [25, 3
26]. As mentioned before, the biological hydrolysis of cellulose occurs in mild conditions of temperature and pressure, with low energy consumption, high specificity and less toxic products. Furthermore, selectivity of the enzymatic process makes it more preferable than chemical methods [27]. Despite their advantages, enzymatic methods endure some limitations such as separation
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difficulties, enzyme inactivation in organic solutions and their vulnerability to temperature and pressure which are the major hindrances in obtaining glucose from cellulose and subsequently the secondgeneration ethanol [28]. Further, the high cost of enzymes makes the process economically unattractive, as a result enzyme immobilization on supports becomes a valuable tool to overcome these limitations and confer numerous benefits, such as simple recovery and purification of immobilized
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enzyme [29, 30].
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Physical (ionic exchange, affinity adsorption and hydrophobic adsorption) and chemical (covalent
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binding) methods have been applied for enzyme immobilization on different supports [30-32].
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Comparatively, the covalent strategy eliminates or decreases enzyme leakage significantly due to
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increased enzyme bonding strength. Recently, immobilization of cellulase on different carriers such as
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nanoparticles (NPs), nanofibers, and nano-polymers has been studied [33-35]. Nanotechnology has opened new horizons in different fields especially in the enzyme world and in the context of enzymatic
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activity and stability improvements through enzyme immobilization [36, 37]. Improved
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characterization, easy separation of enzyme from the reaction mixture, microbial growth prevention and higher enzymatic activity are among the most important advantages of enzyme immobilization [23,
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38, 39].
The high surface area and pore volume, uniform pore size, simple surface functionalization,
hydrothermal stability accompanied biocompatibility of porous silica nanoparticles (PSNs) have made them a suitable support to immobilize a wide variety of enzymes [40, 41]. Well-defined pores having a narrow diameter distribution and simple surface functionalization are the vital characteristics which 4
allows the absorbance of molecules through covalent and hydrogen bonding as well as electrostatic interactions [42]. Pore diameter and organic surface modification of PSNs have a special role in efficacy enhancement of enzyme immobilization. Moreover, PSNs-based magnetic nanoparticles (MNPs) are the key players in molecular adsorption modulation and delivery systems [43-47].
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Feasibility of MNPs fast separation from the mixture is their most advantageous characteristic [48-50]. Despite various cellulase immobilization techniques reported in the last decade, there is still demand for more efficient and reusable supports [51]. The aim of this study was to combine the benefits of AuNPs with PSNs in order to fabricate novel biocompatible separable MNPs as a supporting material for immobilization of cellulase enzyme and enhancing its stability. To this purpose,
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glutamic acid and polyethylene glycol were used as linkers to form the covalent bonds between
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cellulase and the surface of the support material.
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2. Materials and Methods
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2.1. Materials
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Cellulase (3–10 units/mg solid), penicillium funiculosum, dinitrosalicylic acid (DNS), bovine serum albumin (BSA), glucose, ascorbic acid (AA), n-BuOH and HAuCl4.4H2O were purchased from Potassium
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Sigma-Aldrich.
sodium
tartrate,
FeCl2.4H2O,
FeCl3.6H2O,
sodium
hydroxide,
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tetramethylammonium hydroxide, hexadecyltrimethylammonium bromide (CTAB), AgNO3, tetraethyl orthosilicate (TEOS), glutamic acid (Glu), octane, p-toluenesulfonyl chloride (p-TsCl), trimethylamine
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(Et3N) trisodiumcitrate dihydrate, PEG-600-Silicate and other chemicals used in this study were obtained from Merck. Enzyme samples were assayed by LAMBDA 35 UV/Vis spectrophotometer, Shimadzu, Japan. Synthesized NP’s morphology was studies using scanning electron microscopy (SEM) (Hitachi S-4800 II, Japan) equipped with energy dispersive X-ray spectroscopy (EDAX) and the size of synthesized 5
NPs was analyzed by transmission electron microscopy (TEM) (Hitachi H-7650, Japan) at 80 kV. X-Ray diffraction (XRD) was performed by a Philips X’pert 1710 diffractometer using Cu Kα (α=1.54056Å) in Bragg-Brentano geometry (θ-2θ) to study the structure of synthesized NPs while their magnetic properties were determined by vibrating sample magnetometery (VSM). The Fourier
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transform infrared spectroscopy (FTIR) spectra was studied by a Nicolet FT-IR Magna 550 spectrophotometer (USA) in the region of 4000–400 cm−1. The size of NPs was measured by dynamic light scattering (DLS) (Nano-ZS 90, Malvern Instrument, United Kingdom) at 25℃. NPs zeta potential was measured in folded capillary cells using Nano sizer (Zeta sizer Nano ZS90, Malvern Instruments
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Ltd., Malvern, UK).
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2.2. Methods
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2.2.1. Synthesis of magnetic gold nanoparticles (mAu@NPs)
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Fe3O4-NPs were synthesized using a simple co-precipitation method according to Roth et al. with slight
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modifications [52]. FeCl2.4H2O (3.7 mmol) and FeCl3.6H2O (7.4 mmol) were dissolved in deionized
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water (DW) (30 mL) under an inert atmosphere at room temperature and were added to a 25% NH4OH solution (10 mL) under vigorous stirring condition, where the drop rate of NH4OH was controlled
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precisely at ~ 1 mLmin-1. The prepared solution was stirred at 90ºC for 1 h. The supernatant was
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separated with an external magnet and the obtained black precipitate of Fe3O4-NPs was washed with DW to achieve a neutral Fe3O4 suspension and stored at 4ºC. Then mAu@NPs were prepared using
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Fe3O4-NPs as seed [53]. Prepared Fe3O4-NPs were sonicated for 20 minutes after which 10 mL of Au solution was added to 50 mg of the aforementioned nanoparticles in a dropwise manner. Prepared nanoparticle solution was then dispersed in a micellar solution containing 0.1 M HAuCl4.4H2O, 3 g CTAB, 2.5 g n-butanol, 15 g octane and incubated at 38ºC for three hours in a shaking incubator [Fig. 1]. NaBH4 was then added to the mixture as reducing agent and mixed at room temperature. NPs 6
were washed with DW until neutralization and then mAu@NPs were separated and dried under vacuum condition. 2.2.2. Synthesis of magnetic gold-mesoporous silica nanoparticles core shell (mAu@PSNs) 100 µL of 0.1 M AA and 2.5 mL of 0.01 M AgNO3 were added to a mixture of 50 mg of mAu@NPs
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dispersed in 10 mL DW, and pH was adjusted to 10. mAu@Ag-NPs were produced after 2 hours of vigorous stirring. Fabrication of mesoporous SiO2 shell on the surface of mAu@Ag-NPs was carried out according to the procedure described previously by Gorelikov and colleagues with slight modifications [54]. Briefly, 250 μL of 0.1 M NaOH was added to stirring mAu@Ag-NPs solution. Afterwards, 500 μL of TEOS dissolved in MeOH (10% v/v) was added drop-wise with gentle stirring.
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The mixture was stirred at room temperature for 24 hours and then 100 mL of H2O2 was added
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continuously into the stirring solution of 50 mL of mAu@Ag@mSiO2 overnight. Finally,
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mAu@mSiO2-NPs were separated by an external magnet, washed with EtOH and dried under vacuum
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condition [Fig.1].
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2.2.3. Synthesis of Glutamic acid functionalized mAu@PEGylated PSNs (Glu@PEGylated
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mAu@PSNs)
PEG600-silane (1.5 mmol) was added to a suspension containing 100 mg of mAu@mSiO2-NPs
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dispersed in 50 mL toluene and was refluxed for 24 h. Later, the solid residue was separated and
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washed using an external magnet and dried under the vacuum condition to provide PEGylated mAu@mSiO2NPs (mAu@mSiO2-PEG-OH). mAu@mSiO2-PEG-OH (150 mg) was added
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to a solution of 1.5 mmol p-TsCl and Et3N in 50 mL dry CH2Cl2 and was stirred for 12 hours at room temperature. Afterwards, the solid (mAu@mSiO2-PEG-OTs) was separated and washed by DW and EtOH and dried for 12 hours. In the next step, 1.5 g of Glu was added to 30 mL of 1 M sodium bicarbonate and the pH was adjusted to 8.5. In the next step, 2.0 g of mAu@mSiO2-PEG-OTs was added to the latter solution and allowed to stir at 60–65°C for 12 hours. The resultant product, 7
L-glutamic-functionalized PEGylated mAu@mSiO2 (Glu@PEGylated mAu@PSNs), was washed with water, 10% acetic acid and water again until it gained neutral pH and the final solution was dried overnight [Fig. 1]. 2.2.4. Enzyme Activity
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Several methods exist for assaying cellulase enzyme activity. Among those, carboxymethyl-Cellulase “CMCase” and filter paper activity (FPase) are the most commonly used methods which convert cellulose to glucose. In this research, we used the second technique. Filter paper units (FPU) is the procedure of measuring cellulase activity by International Union of Pure and Applied Chemistry (IUPAC) [55]. In this assay, 2.0 mg of reducing sugar (glucose) released from 50 mg Whatman filter
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papers (1×6 cm) in 60 minutes is the reaction product and its absorbance is measured by UV-visible
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spectroscopy. 250 mL of diluted enzyme solution, 1250 mL citrate buffer and 50 mg substrate were
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incubated for 1 hour under the optimum condition (50 mM sodium citrate buffer, pH 4.8 at 50ºC). After
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this time, 200 mL of the mixture was poured into another tube and 200 mL DNS reagent was added to
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the mixture. The tubes were kept in a boiling water bath for 5 minutes. Then, 1600 µL DW was added
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to the boiling tube and homogenized by vortexing. Finally, the absorbance of the solution was read at 540 nm.
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Basically, the FPase assay measures enzyme’s ability to hydrolyze both crystalline and amorphous
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regions of biomaterials. It measures the activities of both endo- and exo- types using a strip of filter paper (Whatman #1, 50 mg, 1X6 cm) for 1 hour reaction at 50°C. In this relation, one unit of enzyme
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activity is defined as the amount of enzyme producing 1 mmol of glucose equivalents per minute at 50°C and pH 4.8. Enzyme unit was calculated as follow [56]: Activity of cellulase (μmol/mLmin) = 1000W /MVt
(1)
Where W is the amount of released glucose equivalents, M is glucose’s molecular weight, V is the volume of measured sample and t is the reaction time, respectively. 8
2.2.5. Enzyme immobilization For enzyme immobilization on the fabricated NPs, following the optimization reactions 25 mg of prepared NPs was added to 4 mL of 0.25 mg/mL cellulose solution, stirring in a cold room at 4-6°C. After 24 hours, the supernatant was simply separated from the solution using an external magnet.
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2.2.6. Measurement of binding efficiency
The Bradford protein assay was used to determine the amount of bounded enzyme and immobilization efficacy [57]. The amount of protein in the supernatant was determined at 595 nm while BSA was used as standard. The amount of bounded enzyme was calculated as follow [56]: Immobilization efficiency (%) = Ci – (Cs/Ci) × 100
(2)
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Where Ci is the initial concentration of cellulase in the reaction, and Cs is the unbound cellulase
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concentration in each purification cycle.
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The standard curve was generated as a plot of absorbance versus protein concentration (μg) and the
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2.2.7. Determination of thermal stability
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immobilization efficiency was determined to be 76%.
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Most enzymes lose their activity due to structural changes caused by thermal denaturation. Here, in order to investigate thermal stability, equal concentrations of both free and immobilized cellulase was
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used for activity determination. The incubation temperatures were 35, 40, 45, 50, 55, 60, 65, 70 and
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75oC for 1 hour under the explained protocol for enzyme activity assay (2.2.6). After 1 hour the absorbance of free and immobilized cellulase was read at 595 nm.
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2.2.8. Determination of pH stability Substrate’s affinity to enzyme and enzyme activity are affected by pH. Effect of pH on cellulase activity was studied using phosphate buffer solution with different pHs (ranging from 2-8) at 50C [48]. Following one hour of incubation, enzyme activity was assayed as described earlier. 2.2.9. Recycling of immobilized cellulase 9
The same enzyme assay was used to determine the recovery and recycling stability of the immobilized cellulase [58]. The biocatalyst was incubated for 1 hour in the optimum condition (50 °C, pH 4.8 and 0.05 M citrate buffer). After each cycle CEL@Glu@PEGylated mAu@PSNs were magnetically separated and washed with DW to eliminate residual substrate or product and finally the enzyme assay
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was performed for each cycle. In this report, all experiments are carried out in triplicate.
3. Results and Discussion
Catalysts (including cellulase) are widely used in biotechnological processes. Despite its common applications, cellulase sensitivity to changes in the environmental temperature, pH, ionic concentration
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or solvents and its subsequent loss of enzymatic activity as well as its inability to be reused are always
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the main challenges that must be dealt with. Different approaches exist that can be used for enhancing
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enzymatic activity including immobilization and chemical modification of the enzyme. Among
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different materials that could be utilized for enzyme immobilization, nanoparticles are outstanding
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candidates due to their unique physico-chemical properties. Nanoparticle’s low toxicity has made them
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a promising support material candidate in preparing nano-biocatalysts [50]. The problem of enzyme separation from the reaction medium can be solved by utilizing magnetic nanoparticles. Magnetic
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nanoparticles have also been considered as support material for cellulase immobilization.
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Functionalization with gold and silica coating are two commonly used strategies that make nanoparticle’s surface biocompatible and chemically stable [51]. Here, we have designed a novel
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nanoparticle system containing L-glutamic-functionalized PEGylated mAu@PSNs as a supporting matrix for the improvement of cellulase stability. Following magnetic Au@NPs synthetization as mentioned in the materials and methods section, silver coating was applied using AA as the reducing agent to produce mAu@Ag-NPs. Mesoporous silica shell coating was then applied to NPs by hydrolysis and condensation of TEOS using CTAB to form mAu@Ag@mSiO2-NPs. Silica coating 10
improved NPs stability and biocompatibility. In the next step, in order to form a yolk like structure, the Ag shell was removed by H2O2 to obtain mAuNR@mSiO2-NPs. Next, the produced nano-system was targeted and specialized by covalent bonding of glutamic acid using PEG600-silane as a linker, decorating the surface of mesoporous SiO2 shell [Fig. 1]. The structure of NPs was fully characterized
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using different techniques such as TEM, SEM, DLS, Zeta, XRD, EDAX, VSM, FT-IR, and TGA. 3.1. Characterization of Glu@PEGylated mAu@PSNs
Magnetic core shell nanoparticles were synthesized according to co-precipitation method and the fabricated nano-system was characterized with different techniques. The particle size and morphology of the PSNs were characterized by TEM/SEM microscopy [Fig. 2 a and b]. A transmission electron
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microscope produces a black and white image as the results of the interactions between sample and
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energetic electrons in the vacuum chamber. TEM was used to study the physical characteristics of
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Glu@PEGylated mAu@PSNs and to confirm magnetic gold-silica core shell framework. The size of
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synthesized nano carrier (Glu@PEGylated mAu@PSNs) determined to be about 68 nm [Fig. 2a]. SEM
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image showed that (Glu@PEGylated mAu@PSNs) has a mono-dispersed spherical structure with the
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average diameter of 45-80 nm [Fig. 2b]. 3.2. Particle Size and zeta potential
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Since the enzyme activity is sensitive to environmental potential, it is critical to determine the surface potential of nanoparticles in the process of enzyme immobilization. For this purpose, DLS was used to
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determine the nanoparticle’s zeta potential. Data presented in table 1 shows the size and surface
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potential of bare and immobilized nanoparticles. The size changes from 68 nm to 78 nm, in bare and bounded nanoparticles respectively, showing that cellulase has successfully bonded to the surface of support matrix. As data shows, nanoparticle’s surface charge changed from +19.3 to + 15.7 mV as the result of enzyme binding to nanoparticles. Zeta potential determination by DLS as well as microscopic analysis confirms enzyme immobilization. [53, 59]. For each sample, the Zeta potential and particle 11
size were measured 4 times at triplicate [Table 1]. DLS data indicate that the average size of CEL@Glu@PEGylated mAu@PSNs was 78 nm. In addition, the particle size distribution was very narrow. Poly dispersity index (PDI) was about 0.388 and the zeta potential was determined +15.7 mV. Overall, there is consistency between DLS, TEM and SEM results.
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3.3. VSM
Magnetic properties of the nano-complex was studied by VSM. Magnetic hysteresis measurements for mAu@mSiO2-NPs and Glu@PEGylated mAu@PSNs was performed in an applied magnetic field with a sweeping rate from −8000 to +8000 kV at room temperature. As shown in Fig. 3, the M (H) hysteresis loops were completely reversible, showing that the NPs exhibit super-paramagnetic
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characteristics. Both particles showed high permeability in magnetization and that their magnetization
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is sufficient for magnetic separation induction using a conventional magnet. The reversibility in
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hysteresis loop confirms that no aggregation imposes to the nanoparticles in the magnetic fields [53].
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3.4. XRD and EDAX
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The powder X-Ray diffraction experiment was used to confirm the existence of iron oxide-gold core-
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shell NPs. Three bands were observed in the XRD pattern which correspond to gold atoms at 2θ = 44.67, 51.72 and 76.71 (JCPDS #89-3697) [Fig. 4 a and b]. However, the characteristic bands of
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magnetic core do not appear in the XRD of the final product. This absence is due to the fact that all iron
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oxide NPs are successfully coated and passivated with a gold shell [60]. To confirm the presence of Au and Fe in the support matrix, energy dispersive X-ray (EDAX) spectroscopy was performed. The spot-
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profile EDAX of nanoparticles showed strong signals for gold atoms along with weak signal from carbon (Fig. 4c). 3.5. TGA TGA results show that the 2% weight loss of mAu@mSiO2 NPs is due to the removal of water from its structure. The functionalized mAu@mSiO2 showed 30% weight loss that is related to the separation of 12
organic linkers from the Glu@PEGylated mAu@PSNs. Moreover, 58% weight loss in CEL@Glu@PEGylated mAu@PSNs indicates that 28% of nano-system weight is related to the cellulase existence [Fig. 5]. 3.6. Binding Confirmation
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The FT-IR spectra confirmed the presence of cellulase on NP’s surface [Fig. 6]. Figure 6 shows FT-IR data for bare NPs, free and immobilized cellulase. Infrared radiation of frequencies less than 800 cm-1 are attributed to the FeO band vibration of Magnetite. Typical peaks of SiO are presented at about 1080 cm-1. The intense peak observed about 1600cm-1 (amide peak) is obvious in both immobilized and free enzyme which confirms amide bond creation between the N-H groups of NPs and O-H bonds of the
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enzyme. The confirmation of binding inter-molecular hydrogen bonds emerging from N-H and O-H
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confirms cellulase binding to nano-system [61].
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groups appears at about 3000-3400cm-1 and indicates the presence of cellulase in the support matrix and
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3.7. Effect of temperature on enzyme activity and stability
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Temperature affects the stability and activity of free and immobilized enzymes [Fig.7]. As presented in
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figure 8, at the optimum temperature the free enzyme shows greater activity than the immobilized enzyme. However, when the temperature increases to 65, 70 and 75ºC, the immobilized enzyme shows
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slightly greater activity. In addition, immobilized cellulase activity was enhanced compared to unbounded cellulase when the temperature was dropped to 40ºC and 35ºC. Also, the activity of the
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immobilized enzyme was less than the free enzyme at 50◦C probably due to covalent binding. On the
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other hand, enzyme immobilization enhances enzyme activity under temperature changes due to increased stabilization which is one of the important advantages of binding enzyme to the support matrix [62]. These results show that the thermal stability of CEL@Glu@PEGylated mAu@PSNs is higher than the free enzyme. Enhanced thermal stability of immobilized enzyme might be related to the presence of 13
more covalent cross-links between amino functionalized magnetite nanoparticles and enzyme molecules which result in conformational stabilization of CEL in support [55, 63]. 3.8. Effect of pH on enzyme activity It is obvious that cellulase as an amphoteric molecule in nature is affected by the pH of its surrounding
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environment. Here, we studied the effect of pH on enzyme activity of free and immobilized cellulase in the pH range of 2-8. [Fig. 8]. Immobilized cellulase showed a significant increase of stability at higher pH compared to free cellulase. It can be suggested that the binding of cellulase along with the presence of amino bonds on nanoparticle’s surface might provide a greater resistance to higher pH levels. Being bounded to a support matrix allows the formation of new interactions between functional groups on the
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support’s surface and the enzyme residues upon structural and conformational changes in the enzyme
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molecule induced by small changes in acidic or basic properties of the environment near the active site,
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3.9. Effect of Time on enzyme activity
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leading to increased stability.
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The initial activity of free and immobilized enzyme was measured under the same conditions. Then,
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the activity of both forms was measured every two hours for 24 hours under optimum condition. Data presented in figure 9 shows that the activity of immobilized cellulase is 57% more than free cellulase
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[Fig. 9]. The result showed that the enzyme activity has decreased following immobilization compared
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to free enzyme. This loss of enzymatic activity could be related to the protein’s conformation that changes with time due to denaturation [62].
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3.10. Recycling study
In any industrial field, reusability of enzymes is a key factor in their application. Re-usability of CEL@Glu@PEGylated mAu@PSNs was studied for five cycles. The relative immobilized enzyme activity in reaction medium was 91%, 85%, 74%, 57% and 28% after five cycles. These results suggest that the cellulase activity loss was due to enzyme leakage in the reaction medium [Fig. 10]. 14
Nanoparticles have a high surface area that can be coated with a large number of amino groups. It seems that not all enzyme molecules bounded to the NP’s surface and that they might have been excluded from the reaction medium [63].
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4. Conclusion
Despite several reports on different carriers and immobilization strategies in the past decade, the demand for more efficient and eco-friendly nano-biocatalysts still exist. Recent advances in nanotechnology has led to development of new nanoparticle systems, as well as novel linkers and a
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variety of nanoparticles with different shapes and sizes. In the present study, we have successfully
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fabricated a novel, biocompatible and separable magnetic nanoparticle system exploiting the
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advantages of AuNPs and PSNs simultaneously. A new nano-support matrix was produced and
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thoroughly characterized by XRD, VSM, TGA, TEM, and SEM. The produced nano-system showed improved enzyme stability against heat and pH changes upon immobilization. The formation of
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covalent bonding between the N-H groups of the nano-system and O-H groups of the enzyme was
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confirmed by FT-IR spectroscopy. Enzyme stability was also enhanced 57% upon immobilization. Our nano-support provides a suitable matrix for cellulase immobilization and further industrial applications
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especially in the biofuels field. Furthermore, assessing the reusability showed that the immobilized
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cellulase remained active at least for five cycles. Many reports exist in the literature on fabrication of magnetic nanoparticles as support materials for enzyme immobilization including chitosan-coated
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paramagnetic nanoparticles [32], magneto-responsive graphene [34] and amine functionalized Fe2O3 [37]. In the present study, our fabricated support nanomaterial for cellulase immobilization showed greater enzyme activity and stability.
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Research on more compatible and efficient support biomaterials for cellulase immobilization is ongoing in our laboratory with the aim of reducing application costs by improving enzyme stability and reusability.
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Acknowledgments
Financial support from Payame Noor University and University of Tehran is gratefully acknowledged.
Conflict of interest
The authors declare that there are no conflicts of interest regarding the publication of this
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manuscript.
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Legends Table 1. Size and zeta potential measurements of immobilized enzyme and support nanoparticles by dynamic light scattering.
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Figure 1. The schematic illustration of the procedure for preparing CEL@Glu@PEGylated mAu@PSN.
Figure 2. TEM/ SEM images of Glu@PEGylated mAu@PSNs. a) Scale bar for TEM image is 100 nm. b) Scale bar for SEM image is 750 nm.
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Figure 3. VSM magnetization curve of Glu@PEGylated mAu@PSNs.
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Figure 4. (a and b) the XRD patterns of Fe3O4@NPs and Fe3O4@Au nanoparticles, respectively and (c)
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the EDAX profile of synthesized Fe3O4@Au nanoparticles.
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Figure 5. TGA curves of (a) the support matrix (mAu@mSiO2) (b) functionalized support matrix Glu@PEGylated mAu@PSNs and (c) immobilized cellulase (CEL@Glu@PEGylated mAu@PSNs)
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Figure 6. The FTIR spectroscopy of (a) free cellulase enzyme, (b) Glu@PEGylated mAu@PSNs (bare
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nanoparticles) and (c) CEL@Glu@ PEGylated mAu@PSNs (immobilized enzyme). Figure 7. The temperature dependence of enzyme stability. Effect of temperature on the thermal
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stability of immobilized versus free cellulase was measured at 35-75°C and pH= 4.8.
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Figure 8. Effect of pH on the activity of immobilized and free cellulase in the pH range of 3-8 at 50 °C was measured.
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Figure 9. Changes in the enzymatic activity of free versus immobilized cellulase with time at pH= 4.8 and 50 °C. Figure 10. Recycling diagram of immobilized cellulase. Re-usability of CEL@Glu@PEGylated mAu@PSNs was studied for five cycles.
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Table 1.
(Glu@PEGylated mAu@PSN)
(CEL@Glu@PEGylated mAu@PSN)
68 nm
78 nm
+19.3 mV
+15.7 mV
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Charge
Immobilized Enzyme
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Size
Functionalized Support
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Fig. 1
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S1359-5113(17)31976-1 https://doi.org/10.1016/j.procbio.2018.05.012 PRBI 11348
To appear in:
Process Biochemistry
Received date: Revised date: Accepted date:
24-12-2017 14-5-2018 15-5-2018
Please cite this article as: Poorakbar E, Shafiee A, Saboury AA, Rad BL, Khoshnevisan K, Ma’mani L, Derakhshankhah H, Ganjali MR, Hosseini M, Synthesis of Magnetic Gold Mesoporous Silica Nanoparticles Core Shell for Cellulase Enzyme Immobilization: Improvement of Enzymatic Activity and Thermal Stability, Process Biochemistry (2018), https://doi.org/10.1016/j.procbio.2018.05.012 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Synthesis of Magnetic Gold Mesoporous Silica Nanoparticles Core Shell for Cellulase Enzyme Immobilization: Improvement of Enzymatic Activity and Thermal Stability
Elaheh Poorakbara,b,c, Abbas Shafieeb, Ali Akbar Sabouryc*, Behzad Lame Rada, Kamyar Khoshnevisand, Leila
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Ma’manie, Hossein Derakhshankhahf, Mohammad Reza Ganjalig, Morteza Hosseinih a
Department of Biology, Faculty of Sciences, Payame Noor University, P.O.Box: 19395-3697, Tehran, Iran
b
Department of Pharmaceutics, Tehran University of Medical Science, Tehran, Iran
c
Institute of Biophysics and Biochemistry, University of Tehran, Tehran, Iran
d
Biosensor Research Center, Endocrinology and Metabolism Molecular-Cellular Sciences Institute, Tehran University of
Medical Sciences, Tehran, Iran
Department of Nanotechnology, Agricultural Biotechnology Research Institute of Iran (ABRII), Karaj, Iran
f
Pharmacutical Sciences Research Center, Kermanshah University of Medical Sciences, Kermanshah, Iran.
g
Center of Excellence in Electrochemistry, Faculty of Chemistry, University of Tehran, Tehran, Iran
h
Department of Life Science Engineering, Faculty of New Sciences & Technologies, University of Tehran, Tehran, Iran.
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(*Corresponding Author: [email protected])
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Graphical abstract
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Highlights
Novel magnetic gold core shell nanoparticles as a support material for cellulase immobilization were fabricated.
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The binding efficiency of cellulase to the support matrix was 76%.
Thermal stability and activity of the immobilized cellulase was improved upon immobilization.
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The immobilized cellulase was reusable for five cycles.
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Abstract
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Magnetic gold mesoporous silica nanoparticle core shells (mAu@PSNs) were fabricated as a support and their size, morphology and structure was further characterized by X-Ray diffraction (XRD),
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vibrating sample magnetometer (VSM), scanning electron microscopy (SEM), energy dispersive X-ray
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analysis (EDAX), transmission electron microscopy (TEM), dynamic light scattering (DLS) and thermal gravity analysis (TGA). Cellulase (CEL) immobilization on mAu@PSNs was performed via
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covalent bonding. Fourier transform infrared (FTIR) spectroscopy confirmed the successful binding of
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enzyme to mAu@PSNs while Bradford assay determined the binding efficiency to be 76%. The enzyme activity was measured at different pHs and temperatures by FPase method using Whatman
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filter paper as the substrate. The immobilized enzyme maintained 58% of its initial catalytic activity after nine hours. In this research, a new nano-system was designed as a solid support for cellulase immobilization which enhanced its thermal stability and facilitated its long term storage. In addition, the immobilized enzyme can be applied in a broader temperature and pH ranges while enzyme separation can be simply carried out by an external magnet. 2
Keywords: cellulase activity; thermal stability; enzyme immobilization; magnetic nano-particles; covalent bonding.
1. Introduction
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Natural resources shortage as well as the environmental pollution caused by fossil fuels compels us to look for new renewable energy resources [1]. Cellulosic materials such as biomass have been employed as eco-friendly sustainable energy resources in biofuels production (for example bioethanol) [2]. To produce such materials, it is necessary to modify the biomass’s chemical and structural properties to prevent its disassembly [3, 4]. Cellulose, the insoluble polymer of glucoside units, is the main source of
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carbon and one of the most abundant resources of renewable biomass [5, 6]. It can be obtained from the
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forests as well as the agricultural wastes. Cellulose is a suitable low-cost substrate for obtaining
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fermentable sugars with various industrial applications [7].
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Several physical, chemical and biological methods have been employed for hydrolysis of cellulosic
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materials. The enzymatic hydrolysis of cellulose has gained much attention because of its mild reaction
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conditions [8-10]. Microorganisms such as fungi [11, 12], bacteria [13-15] and yeasts [16] produce cellulose hydrolyzing enzyme, cellulase. Fungal cellulases are the most powerful cellulose hydrolyzing
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enzymes [17-19]. Several studies have reported the industrial applications of cellulase since it plays a
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key role in bio-conversion of many bio-substrates to produce bio-fuels [20-22]. In the past decades, research on cellulases have increased due to the interest in producing bioethanol from lignocellulosic
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biomass [1, 2, 4, 19, 23]. Cellulase is a biomolecular complex including three different enzymes (endo-cellulase,
exo-cellulase or cellobiohydrolase, and β-glucosidase) which produces soluble sugar from cellulolytic substrates [24]. It is also the second most commonly used industrial enzyme in various fields including agriculture, food, animal feed, brewery and wine, textile, detergents, pulps and papers industries [25, 3
26]. As mentioned before, the biological hydrolysis of cellulose occurs in mild conditions of temperature and pressure, with low energy consumption, high specificity and less toxic products. Furthermore, selectivity of the enzymatic process makes it more preferable than chemical methods [27]. Despite their advantages, enzymatic methods endure some limitations such as separation
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difficulties, enzyme inactivation in organic solutions and their vulnerability to temperature and pressure which are the major hindrances in obtaining glucose from cellulose and subsequently the secondgeneration ethanol [28]. Further, the high cost of enzymes makes the process economically unattractive, as a result enzyme immobilization on supports becomes a valuable tool to overcome these limitations and confer numerous benefits, such as simple recovery and purification of immobilized
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enzyme [29, 30].
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Physical (ionic exchange, affinity adsorption and hydrophobic adsorption) and chemical (covalent
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binding) methods have been applied for enzyme immobilization on different supports [30-32].
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Comparatively, the covalent strategy eliminates or decreases enzyme leakage significantly due to
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increased enzyme bonding strength. Recently, immobilization of cellulase on different carriers such as
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nanoparticles (NPs), nanofibers, and nano-polymers has been studied [33-35]. Nanotechnology has opened new horizons in different fields especially in the enzyme world and in the context of enzymatic
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activity and stability improvements through enzyme immobilization [36, 37]. Improved
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characterization, easy separation of enzyme from the reaction mixture, microbial growth prevention and higher enzymatic activity are among the most important advantages of enzyme immobilization [23,
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38, 39].
The high surface area and pore volume, uniform pore size, simple surface functionalization,
hydrothermal stability accompanied biocompatibility of porous silica nanoparticles (PSNs) have made them a suitable support to immobilize a wide variety of enzymes [40, 41]. Well-defined pores having a narrow diameter distribution and simple surface functionalization are the vital characteristics which 4
allows the absorbance of molecules through covalent and hydrogen bonding as well as electrostatic interactions [42]. Pore diameter and organic surface modification of PSNs have a special role in efficacy enhancement of enzyme immobilization. Moreover, PSNs-based magnetic nanoparticles (MNPs) are the key players in molecular adsorption modulation and delivery systems [43-47].
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Feasibility of MNPs fast separation from the mixture is their most advantageous characteristic [48-50]. Despite various cellulase immobilization techniques reported in the last decade, there is still demand for more efficient and reusable supports [51]. The aim of this study was to combine the benefits of AuNPs with PSNs in order to fabricate novel biocompatible separable MNPs as a supporting material for immobilization of cellulase enzyme and enhancing its stability. To this purpose,
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glutamic acid and polyethylene glycol were used as linkers to form the covalent bonds between
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cellulase and the surface of the support material.
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2. Materials and Methods
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2.1. Materials
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Cellulase (3–10 units/mg solid), penicillium funiculosum, dinitrosalicylic acid (DNS), bovine serum albumin (BSA), glucose, ascorbic acid (AA), n-BuOH and HAuCl4.4H2O were purchased from Potassium
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Sigma-Aldrich.
sodium
tartrate,
FeCl2.4H2O,
FeCl3.6H2O,
sodium
hydroxide,
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tetramethylammonium hydroxide, hexadecyltrimethylammonium bromide (CTAB), AgNO3, tetraethyl orthosilicate (TEOS), glutamic acid (Glu), octane, p-toluenesulfonyl chloride (p-TsCl), trimethylamine
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(Et3N) trisodiumcitrate dihydrate, PEG-600-Silicate and other chemicals used in this study were obtained from Merck. Enzyme samples were assayed by LAMBDA 35 UV/Vis spectrophotometer, Shimadzu, Japan. Synthesized NP’s morphology was studies using scanning electron microscopy (SEM) (Hitachi S-4800 II, Japan) equipped with energy dispersive X-ray spectroscopy (EDAX) and the size of synthesized 5
NPs was analyzed by transmission electron microscopy (TEM) (Hitachi H-7650, Japan) at 80 kV. X-Ray diffraction (XRD) was performed by a Philips X’pert 1710 diffractometer using Cu Kα (α=1.54056Å) in Bragg-Brentano geometry (θ-2θ) to study the structure of synthesized NPs while their magnetic properties were determined by vibrating sample magnetometery (VSM). The Fourier
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transform infrared spectroscopy (FTIR) spectra was studied by a Nicolet FT-IR Magna 550 spectrophotometer (USA) in the region of 4000–400 cm−1. The size of NPs was measured by dynamic light scattering (DLS) (Nano-ZS 90, Malvern Instrument, United Kingdom) at 25℃. NPs zeta potential was measured in folded capillary cells using Nano sizer (Zeta sizer Nano ZS90, Malvern Instruments
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Ltd., Malvern, UK).
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2.2. Methods
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2.2.1. Synthesis of magnetic gold nanoparticles (mAu@NPs)
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Fe3O4-NPs were synthesized using a simple co-precipitation method according to Roth et al. with slight
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modifications [52]. FeCl2.4H2O (3.7 mmol) and FeCl3.6H2O (7.4 mmol) were dissolved in deionized
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water (DW) (30 mL) under an inert atmosphere at room temperature and were added to a 25% NH4OH solution (10 mL) under vigorous stirring condition, where the drop rate of NH4OH was controlled
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precisely at ~ 1 mLmin-1. The prepared solution was stirred at 90ºC for 1 h. The supernatant was
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separated with an external magnet and the obtained black precipitate of Fe3O4-NPs was washed with DW to achieve a neutral Fe3O4 suspension and stored at 4ºC. Then mAu@NPs were prepared using
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Fe3O4-NPs as seed [53]. Prepared Fe3O4-NPs were sonicated for 20 minutes after which 10 mL of Au solution was added to 50 mg of the aforementioned nanoparticles in a dropwise manner. Prepared nanoparticle solution was then dispersed in a micellar solution containing 0.1 M HAuCl4.4H2O, 3 g CTAB, 2.5 g n-butanol, 15 g octane and incubated at 38ºC for three hours in a shaking incubator [Fig. 1]. NaBH4 was then added to the mixture as reducing agent and mixed at room temperature. NPs 6
were washed with DW until neutralization and then mAu@NPs were separated and dried under vacuum condition. 2.2.2. Synthesis of magnetic gold-mesoporous silica nanoparticles core shell (mAu@PSNs) 100 µL of 0.1 M AA and 2.5 mL of 0.01 M AgNO3 were added to a mixture of 50 mg of mAu@NPs
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dispersed in 10 mL DW, and pH was adjusted to 10. mAu@Ag-NPs were produced after 2 hours of vigorous stirring. Fabrication of mesoporous SiO2 shell on the surface of mAu@Ag-NPs was carried out according to the procedure described previously by Gorelikov and colleagues with slight modifications [54]. Briefly, 250 μL of 0.1 M NaOH was added to stirring mAu@Ag-NPs solution. Afterwards, 500 μL of TEOS dissolved in MeOH (10% v/v) was added drop-wise with gentle stirring.
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The mixture was stirred at room temperature for 24 hours and then 100 mL of H2O2 was added
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continuously into the stirring solution of 50 mL of mAu@Ag@mSiO2 overnight. Finally,
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mAu@mSiO2-NPs were separated by an external magnet, washed with EtOH and dried under vacuum
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condition [Fig.1].
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2.2.3. Synthesis of Glutamic acid functionalized mAu@PEGylated PSNs (Glu@PEGylated
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mAu@PSNs)
PEG600-silane (1.5 mmol) was added to a suspension containing 100 mg of mAu@mSiO2-NPs
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dispersed in 50 mL toluene and was refluxed for 24 h. Later, the solid residue was separated and
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washed using an external magnet and dried under the vacuum condition to provide PEGylated mAu@mSiO2NPs (mAu@mSiO2-PEG-OH). mAu@mSiO2-PEG-OH (150 mg) was added
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to a solution of 1.5 mmol p-TsCl and Et3N in 50 mL dry CH2Cl2 and was stirred for 12 hours at room temperature. Afterwards, the solid (mAu@mSiO2-PEG-OTs) was separated and washed by DW and EtOH and dried for 12 hours. In the next step, 1.5 g of Glu was added to 30 mL of 1 M sodium bicarbonate and the pH was adjusted to 8.5. In the next step, 2.0 g of mAu@mSiO2-PEG-OTs was added to the latter solution and allowed to stir at 60–65°C for 12 hours. The resultant product, 7
L-glutamic-functionalized PEGylated mAu@mSiO2 (Glu@PEGylated mAu@PSNs), was washed with water, 10% acetic acid and water again until it gained neutral pH and the final solution was dried overnight [Fig. 1]. 2.2.4. Enzyme Activity
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Several methods exist for assaying cellulase enzyme activity. Among those, carboxymethyl-Cellulase “CMCase” and filter paper activity (FPase) are the most commonly used methods which convert cellulose to glucose. In this research, we used the second technique. Filter paper units (FPU) is the procedure of measuring cellulase activity by International Union of Pure and Applied Chemistry (IUPAC) [55]. In this assay, 2.0 mg of reducing sugar (glucose) released from 50 mg Whatman filter
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papers (1×6 cm) in 60 minutes is the reaction product and its absorbance is measured by UV-visible
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spectroscopy. 250 mL of diluted enzyme solution, 1250 mL citrate buffer and 50 mg substrate were
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incubated for 1 hour under the optimum condition (50 mM sodium citrate buffer, pH 4.8 at 50ºC). After
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this time, 200 mL of the mixture was poured into another tube and 200 mL DNS reagent was added to
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the mixture. The tubes were kept in a boiling water bath for 5 minutes. Then, 1600 µL DW was added
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to the boiling tube and homogenized by vortexing. Finally, the absorbance of the solution was read at 540 nm.
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Basically, the FPase assay measures enzyme’s ability to hydrolyze both crystalline and amorphous
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regions of biomaterials. It measures the activities of both endo- and exo- types using a strip of filter paper (Whatman #1, 50 mg, 1X6 cm) for 1 hour reaction at 50°C. In this relation, one unit of enzyme
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activity is defined as the amount of enzyme producing 1 mmol of glucose equivalents per minute at 50°C and pH 4.8. Enzyme unit was calculated as follow [56]: Activity of cellulase (μmol/mLmin) = 1000W /MVt
(1)
Where W is the amount of released glucose equivalents, M is glucose’s molecular weight, V is the volume of measured sample and t is the reaction time, respectively. 8
2.2.5. Enzyme immobilization For enzyme immobilization on the fabricated NPs, following the optimization reactions 25 mg of prepared NPs was added to 4 mL of 0.25 mg/mL cellulose solution, stirring in a cold room at 4-6°C. After 24 hours, the supernatant was simply separated from the solution using an external magnet.
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2.2.6. Measurement of binding efficiency
The Bradford protein assay was used to determine the amount of bounded enzyme and immobilization efficacy [57]. The amount of protein in the supernatant was determined at 595 nm while BSA was used as standard. The amount of bounded enzyme was calculated as follow [56]: Immobilization efficiency (%) = Ci – (Cs/Ci) × 100
(2)
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Where Ci is the initial concentration of cellulase in the reaction, and Cs is the unbound cellulase
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concentration in each purification cycle.
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The standard curve was generated as a plot of absorbance versus protein concentration (μg) and the
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2.2.7. Determination of thermal stability
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immobilization efficiency was determined to be 76%.
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Most enzymes lose their activity due to structural changes caused by thermal denaturation. Here, in order to investigate thermal stability, equal concentrations of both free and immobilized cellulase was
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used for activity determination. The incubation temperatures were 35, 40, 45, 50, 55, 60, 65, 70 and
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75oC for 1 hour under the explained protocol for enzyme activity assay (2.2.6). After 1 hour the absorbance of free and immobilized cellulase was read at 595 nm.
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2.2.8. Determination of pH stability Substrate’s affinity to enzyme and enzyme activity are affected by pH. Effect of pH on cellulase activity was studied using phosphate buffer solution with different pHs (ranging from 2-8) at 50C [48]. Following one hour of incubation, enzyme activity was assayed as described earlier. 2.2.9. Recycling of immobilized cellulase 9
The same enzyme assay was used to determine the recovery and recycling stability of the immobilized cellulase [58]. The biocatalyst was incubated for 1 hour in the optimum condition (50 °C, pH 4.8 and 0.05 M citrate buffer). After each cycle CEL@Glu@PEGylated mAu@PSNs were magnetically separated and washed with DW to eliminate residual substrate or product and finally the enzyme assay
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was performed for each cycle. In this report, all experiments are carried out in triplicate.
3. Results and Discussion
Catalysts (including cellulase) are widely used in biotechnological processes. Despite its common applications, cellulase sensitivity to changes in the environmental temperature, pH, ionic concentration
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or solvents and its subsequent loss of enzymatic activity as well as its inability to be reused are always
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the main challenges that must be dealt with. Different approaches exist that can be used for enhancing
A
enzymatic activity including immobilization and chemical modification of the enzyme. Among
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different materials that could be utilized for enzyme immobilization, nanoparticles are outstanding
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candidates due to their unique physico-chemical properties. Nanoparticle’s low toxicity has made them
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a promising support material candidate in preparing nano-biocatalysts [50]. The problem of enzyme separation from the reaction medium can be solved by utilizing magnetic nanoparticles. Magnetic
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nanoparticles have also been considered as support material for cellulase immobilization.
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Functionalization with gold and silica coating are two commonly used strategies that make nanoparticle’s surface biocompatible and chemically stable [51]. Here, we have designed a novel
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nanoparticle system containing L-glutamic-functionalized PEGylated mAu@PSNs as a supporting matrix for the improvement of cellulase stability. Following magnetic Au@NPs synthetization as mentioned in the materials and methods section, silver coating was applied using AA as the reducing agent to produce mAu@Ag-NPs. Mesoporous silica shell coating was then applied to NPs by hydrolysis and condensation of TEOS using CTAB to form mAu@Ag@mSiO2-NPs. Silica coating 10
improved NPs stability and biocompatibility. In the next step, in order to form a yolk like structure, the Ag shell was removed by H2O2 to obtain mAuNR@mSiO2-NPs. Next, the produced nano-system was targeted and specialized by covalent bonding of glutamic acid using PEG600-silane as a linker, decorating the surface of mesoporous SiO2 shell [Fig. 1]. The structure of NPs was fully characterized
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using different techniques such as TEM, SEM, DLS, Zeta, XRD, EDAX, VSM, FT-IR, and TGA. 3.1. Characterization of Glu@PEGylated mAu@PSNs
Magnetic core shell nanoparticles were synthesized according to co-precipitation method and the fabricated nano-system was characterized with different techniques. The particle size and morphology of the PSNs were characterized by TEM/SEM microscopy [Fig. 2 a and b]. A transmission electron
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microscope produces a black and white image as the results of the interactions between sample and
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energetic electrons in the vacuum chamber. TEM was used to study the physical characteristics of
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Glu@PEGylated mAu@PSNs and to confirm magnetic gold-silica core shell framework. The size of
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synthesized nano carrier (Glu@PEGylated mAu@PSNs) determined to be about 68 nm [Fig. 2a]. SEM
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image showed that (Glu@PEGylated mAu@PSNs) has a mono-dispersed spherical structure with the
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average diameter of 45-80 nm [Fig. 2b]. 3.2. Particle Size and zeta potential
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Since the enzyme activity is sensitive to environmental potential, it is critical to determine the surface potential of nanoparticles in the process of enzyme immobilization. For this purpose, DLS was used to
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determine the nanoparticle’s zeta potential. Data presented in table 1 shows the size and surface
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potential of bare and immobilized nanoparticles. The size changes from 68 nm to 78 nm, in bare and bounded nanoparticles respectively, showing that cellulase has successfully bonded to the surface of support matrix. As data shows, nanoparticle’s surface charge changed from +19.3 to + 15.7 mV as the result of enzyme binding to nanoparticles. Zeta potential determination by DLS as well as microscopic analysis confirms enzyme immobilization. [53, 59]. For each sample, the Zeta potential and particle 11
size were measured 4 times at triplicate [Table 1]. DLS data indicate that the average size of CEL@Glu@PEGylated mAu@PSNs was 78 nm. In addition, the particle size distribution was very narrow. Poly dispersity index (PDI) was about 0.388 and the zeta potential was determined +15.7 mV. Overall, there is consistency between DLS, TEM and SEM results.
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3.3. VSM
Magnetic properties of the nano-complex was studied by VSM. Magnetic hysteresis measurements for mAu@mSiO2-NPs and Glu@PEGylated mAu@PSNs was performed in an applied magnetic field with a sweeping rate from −8000 to +8000 kV at room temperature. As shown in Fig. 3, the M (H) hysteresis loops were completely reversible, showing that the NPs exhibit super-paramagnetic
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characteristics. Both particles showed high permeability in magnetization and that their magnetization
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is sufficient for magnetic separation induction using a conventional magnet. The reversibility in
A
hysteresis loop confirms that no aggregation imposes to the nanoparticles in the magnetic fields [53].
M
3.4. XRD and EDAX
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The powder X-Ray diffraction experiment was used to confirm the existence of iron oxide-gold core-
TE
shell NPs. Three bands were observed in the XRD pattern which correspond to gold atoms at 2θ = 44.67, 51.72 and 76.71 (JCPDS #89-3697) [Fig. 4 a and b]. However, the characteristic bands of
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magnetic core do not appear in the XRD of the final product. This absence is due to the fact that all iron
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oxide NPs are successfully coated and passivated with a gold shell [60]. To confirm the presence of Au and Fe in the support matrix, energy dispersive X-ray (EDAX) spectroscopy was performed. The spot-
A
profile EDAX of nanoparticles showed strong signals for gold atoms along with weak signal from carbon (Fig. 4c). 3.5. TGA TGA results show that the 2% weight loss of mAu@mSiO2 NPs is due to the removal of water from its structure. The functionalized mAu@mSiO2 showed 30% weight loss that is related to the separation of 12
organic linkers from the Glu@PEGylated mAu@PSNs. Moreover, 58% weight loss in CEL@Glu@PEGylated mAu@PSNs indicates that 28% of nano-system weight is related to the cellulase existence [Fig. 5]. 3.6. Binding Confirmation
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The FT-IR spectra confirmed the presence of cellulase on NP’s surface [Fig. 6]. Figure 6 shows FT-IR data for bare NPs, free and immobilized cellulase. Infrared radiation of frequencies less than 800 cm-1 are attributed to the FeO band vibration of Magnetite. Typical peaks of SiO are presented at about 1080 cm-1. The intense peak observed about 1600cm-1 (amide peak) is obvious in both immobilized and free enzyme which confirms amide bond creation between the N-H groups of NPs and O-H bonds of the
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enzyme. The confirmation of binding inter-molecular hydrogen bonds emerging from N-H and O-H
A
confirms cellulase binding to nano-system [61].
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groups appears at about 3000-3400cm-1 and indicates the presence of cellulase in the support matrix and
M
3.7. Effect of temperature on enzyme activity and stability
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Temperature affects the stability and activity of free and immobilized enzymes [Fig.7]. As presented in
TE
figure 8, at the optimum temperature the free enzyme shows greater activity than the immobilized enzyme. However, when the temperature increases to 65, 70 and 75ºC, the immobilized enzyme shows
EP
slightly greater activity. In addition, immobilized cellulase activity was enhanced compared to unbounded cellulase when the temperature was dropped to 40ºC and 35ºC. Also, the activity of the
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immobilized enzyme was less than the free enzyme at 50◦C probably due to covalent binding. On the
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other hand, enzyme immobilization enhances enzyme activity under temperature changes due to increased stabilization which is one of the important advantages of binding enzyme to the support matrix [62]. These results show that the thermal stability of CEL@Glu@PEGylated mAu@PSNs is higher than the free enzyme. Enhanced thermal stability of immobilized enzyme might be related to the presence of 13
more covalent cross-links between amino functionalized magnetite nanoparticles and enzyme molecules which result in conformational stabilization of CEL in support [55, 63]. 3.8. Effect of pH on enzyme activity It is obvious that cellulase as an amphoteric molecule in nature is affected by the pH of its surrounding
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environment. Here, we studied the effect of pH on enzyme activity of free and immobilized cellulase in the pH range of 2-8. [Fig. 8]. Immobilized cellulase showed a significant increase of stability at higher pH compared to free cellulase. It can be suggested that the binding of cellulase along with the presence of amino bonds on nanoparticle’s surface might provide a greater resistance to higher pH levels. Being bounded to a support matrix allows the formation of new interactions between functional groups on the
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support’s surface and the enzyme residues upon structural and conformational changes in the enzyme
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molecule induced by small changes in acidic or basic properties of the environment near the active site,
M
3.9. Effect of Time on enzyme activity
A
leading to increased stability.
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The initial activity of free and immobilized enzyme was measured under the same conditions. Then,
TE
the activity of both forms was measured every two hours for 24 hours under optimum condition. Data presented in figure 9 shows that the activity of immobilized cellulase is 57% more than free cellulase
EP
[Fig. 9]. The result showed that the enzyme activity has decreased following immobilization compared
CC
to free enzyme. This loss of enzymatic activity could be related to the protein’s conformation that changes with time due to denaturation [62].
A
3.10. Recycling study
In any industrial field, reusability of enzymes is a key factor in their application. Re-usability of CEL@Glu@PEGylated mAu@PSNs was studied for five cycles. The relative immobilized enzyme activity in reaction medium was 91%, 85%, 74%, 57% and 28% after five cycles. These results suggest that the cellulase activity loss was due to enzyme leakage in the reaction medium [Fig. 10]. 14
Nanoparticles have a high surface area that can be coated with a large number of amino groups. It seems that not all enzyme molecules bounded to the NP’s surface and that they might have been excluded from the reaction medium [63].
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4. Conclusion
Despite several reports on different carriers and immobilization strategies in the past decade, the demand for more efficient and eco-friendly nano-biocatalysts still exist. Recent advances in nanotechnology has led to development of new nanoparticle systems, as well as novel linkers and a
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variety of nanoparticles with different shapes and sizes. In the present study, we have successfully
N
fabricated a novel, biocompatible and separable magnetic nanoparticle system exploiting the
A
advantages of AuNPs and PSNs simultaneously. A new nano-support matrix was produced and
M
thoroughly characterized by XRD, VSM, TGA, TEM, and SEM. The produced nano-system showed improved enzyme stability against heat and pH changes upon immobilization. The formation of
D
covalent bonding between the N-H groups of the nano-system and O-H groups of the enzyme was
TE
confirmed by FT-IR spectroscopy. Enzyme stability was also enhanced 57% upon immobilization. Our nano-support provides a suitable matrix for cellulase immobilization and further industrial applications
EP
especially in the biofuels field. Furthermore, assessing the reusability showed that the immobilized
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cellulase remained active at least for five cycles. Many reports exist in the literature on fabrication of magnetic nanoparticles as support materials for enzyme immobilization including chitosan-coated
A
paramagnetic nanoparticles [32], magneto-responsive graphene [34] and amine functionalized Fe2O3 [37]. In the present study, our fabricated support nanomaterial for cellulase immobilization showed greater enzyme activity and stability.
15
Research on more compatible and efficient support biomaterials for cellulase immobilization is ongoing in our laboratory with the aim of reducing application costs by improving enzyme stability and reusability.
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Acknowledgments
Financial support from Payame Noor University and University of Tehran is gratefully acknowledged.
Conflict of interest
The authors declare that there are no conflicts of interest regarding the publication of this
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manuscript.
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Legends Table 1. Size and zeta potential measurements of immobilized enzyme and support nanoparticles by dynamic light scattering.
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Figure 1. The schematic illustration of the procedure for preparing CEL@Glu@PEGylated mAu@PSN.
Figure 2. TEM/ SEM images of Glu@PEGylated mAu@PSNs. a) Scale bar for TEM image is 100 nm. b) Scale bar for SEM image is 750 nm.
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Figure 3. VSM magnetization curve of Glu@PEGylated mAu@PSNs.
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Figure 4. (a and b) the XRD patterns of Fe3O4@NPs and Fe3O4@Au nanoparticles, respectively and (c)
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the EDAX profile of synthesized Fe3O4@Au nanoparticles.
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Figure 5. TGA curves of (a) the support matrix (mAu@mSiO2) (b) functionalized support matrix Glu@PEGylated mAu@PSNs and (c) immobilized cellulase (CEL@Glu@PEGylated mAu@PSNs)
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Figure 6. The FTIR spectroscopy of (a) free cellulase enzyme, (b) Glu@PEGylated mAu@PSNs (bare
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nanoparticles) and (c) CEL@Glu@ PEGylated mAu@PSNs (immobilized enzyme). Figure 7. The temperature dependence of enzyme stability. Effect of temperature on the thermal
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stability of immobilized versus free cellulase was measured at 35-75°C and pH= 4.8.
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Figure 8. Effect of pH on the activity of immobilized and free cellulase in the pH range of 3-8 at 50 °C was measured.
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Figure 9. Changes in the enzymatic activity of free versus immobilized cellulase with time at pH= 4.8 and 50 °C. Figure 10. Recycling diagram of immobilized cellulase. Re-usability of CEL@Glu@PEGylated mAu@PSNs was studied for five cycles.
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Table 1.
(Glu@PEGylated mAu@PSN)
(CEL@Glu@PEGylated mAu@PSN)
68 nm
78 nm
+19.3 mV
+15.7 mV
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A
N
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Charge
Immobilized Enzyme
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Size
Functionalized Support
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Fig. 1
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A
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A
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(a)
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(b)
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D
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A
N
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Fig.2
A
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Fig. 3
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(a)
A
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D
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A
N
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(b)
Fig. 4 a, b and c
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(c)
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A
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A
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Fig.5
Fig. 6
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A
N
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Fig.7
A
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Fig. 8
Fig. 9
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D
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A
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N
A
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Fig. 10
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