Immobilisation of Candida rugosa lipase on polyhydroxybutyrate via a combination of adsorption and cross-linking agents to enhance acylglycerol production

Journal Pre-proof Immobilisation of Candida rugosa Lipase on Polyhydroxybutyrate Via a Combination of Adsorption and Cross-linking Agentsto Enhance Acylglycerol Production Narisa Binhayeeding, Tewan Yunu, Nisa Pichid, Sappasith Klomklao, Kanokphorn Sangkharak

PII:

S1359-5113(19)31680-0

DOI:

https://doi.org/10.1016/j.procbio.2020.02.007

Reference:

PRBI 11925

To appear in:

Process Biochemistry

Received Date:

4 November 2019

Revised Date:

29 January 2020

Accepted Date:

6 February 2020

Please cite this article as: Binhayeeding N, Yunu T, Pichid N, Klomklao S, Sangkharak K, Immobilisation of Candida rugosa Lipase on Polyhydroxybutyrate Via a Combination of Adsorption and Cross-linking Agentsto Enhance Acylglycerol Production, Process Biochemistry (2020), doi: https://doi.org/10.1016/j.procbio.2020.02.007

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1 Immobilisation of Candida rugosa Lipase on Polyhydroxybutyrate Via a Combination of Adsorption and Cross-linking Agentsto Enhance Acylglycerol Production Narisa Binhayeedinga ,Tewan Yunub, Nisa Pichidb, Sappasith Klomklaoc and Kanokphorn Sangkharakb,* a

Biotechnology Program, Faculty of Science, Thaksin University, Phatthalung, Thailand,

93210 b

Department of Chemistry, Faculty of Science, Thaksin University, Phatthalung, Thailand,

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93210

Department of Food Science and Technology, Faculty of Agro- and Bio-Industry,

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Thaksin University, Phatthalung, Thailand, 93210

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*Corresponding author

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Phone: +66-76-609-634

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Graphical abstract

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E-mail address: [email protected]

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Highlights



●

Lipase was immobilized on polyhydroxybutyrate via a combination of adsorption and

crosslinking techniques. 

● Immobilization increased the stability and reusability of lipase enzyme.

 ● Application of immobilized lipase in monoacylglycerol process are determined.

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 High purity of monoacylglycerol was produced using immobilized lipase.

ABSTRACT

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In the present study, a combination of immobilisation processes was utilised to prepare robust biocatalysts. First, lipase from Candida rugosa was adsorbed on polyhydroxybutyrate (PHB)

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particles, followed by cross-linking with glutaraldehyde. Conditions for creating immobilised lipase involved the addition of 0.6 M glutaraldehyde and 45 U mL-1 lipase while mixing at 150

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rpm (4°C) for 30 min. These conditions produced the highest yield of immobilised lipase (92%)

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and the highest levels of activity (1.94 mg g- 1 support). At 40°C and pH 9 the immobilised enzyme was optimally active with a Km and Vmax at 1. 2 mM and 2. 5 × 10- 3 mmol min- 1, respectively. The use of immobilised lipase improved thermal stability, storage stability, and reusability.The immobilised lipase retained 80% of its activity after incubation at 30–60°C for 2 h and 4°C for 30 d in 0.2 M sodium phosphate buffer (pH 7.0). Moreover, the immobilised

3 enzyme retained 50% of its activity after more than 14 cycles under optimal conditions. The immobilised lipase was used to produce monoacylglycerol (MAG). The existence of a carbonyl group at 1,743 and 1,744 cm- 1 was identified using attenuated total reflectance (ATR)-Fourier transformed infrared spectroscopy. Results showed that 48% MAG was produced. Keywords:

Immobilised

enzyme;

Glycerolysis;

Lipase;

Monoacylglycerol;

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Polyhydroxyalkanoate

4 1. Introduction Polyhydroxybutyrate ( PHB) is an organic polymer with the potential to be used commercially as a biodegradable thermoplastic and biomaterial. PHB is well known as a carbon and energy reserve. The material is produced by a variety of microorganisms, and its synthesis is promoted by environmental stresses such as nitrogen, phosphate, or oxygen limitation [1]. PHB is synthesised and deposited intracellularly in the form of granules, accounting for up to

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90% of the dry cell weight. Recently, investigations into the production of PHB have received increased attention as a result of its potential use in the packaging, medical, agricultural, and fishery industries [2]. However, there are few reports assessing the use of PHB as a support for

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immobilised enzymes. PHB could serve as a good alternative for enzyme immobilisation

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support due to its biocompatibility, biodegradability, strength, easy reabsorption, non-toxicity and eco-friendliness [1].

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Lipases (triacylglycerol hydrolases, E.C. 3.1.1.3) stand out for their enantioselectivity,

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high stability and broad subatrate recognition. Lipases are among the enzymes most widely used in enzyme technology due to their broad specificity for substrates capable of catalysing

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many different reactions, such as hydrolysis or synthesis of esters, esterifications, aminations, trans-esterifications, and alcoholysis, among others. Due to the substrate versatility of lipase,

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the application of these enzymes extends to various sectors, such as synthesis reactions and production of pharmaceutical drugs and biofuels. Standardising lipase methodologies can be difficult, as some of their properties may vary depending on the genus and species of microorganisms or other organisms used. The active site of lipase is composed of a serine, an aspartate or glutamate, and a histidine. Lipases are very unique enzymes that process a peculiar

5 pathway called interfacial activation. In homogenous media, the active centre of a large percentage of lipase molecules is covered by a polypeptide chain called lid, which may isolate it from reaction medium (closed form). As a result of interfacial activation, lipases can manifest lf in two different conformations, open (active) and closed (inactive) forms. The open form is thought to be more stable than the closed form. Typically, the open form of lipases occurs with the movement of the lid in the presence of hydrophobic surfaces, increasing enzyme activity.

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However, this movement does not significantly change the final properties of lipases, such as specificity and selectivity. These properties are readily modulated by genetic manipulation or physio-chemical modifications (including immobilisation) [3–6]. Lipases are the most widely

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used enzymes in bio-catalysis, and the most utilised method for enzyme immobilisation is using

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hydrophobic supports at low ionic strength. This method allows the one step immobilisation, purification, stabilisation, and hyperactivation of lipases, which is the main cause of their

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popularity [7].

The lipase produced by Candida rugosa is rapidly becoming one of the most

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industrially used enzymes as a result of its high activity in terms of hydrolysis and synthesis

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and its resulting broad applicability [8]. C. rugosa secretes a mixture of enzymes, possessing at least five isoenzymes encoded in a ‘lipase minigene family’ as demonstrated by gene cloning.

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Each enzyme has a single polypeptide chain, consisting of 543 amino acids and an apparent molecular weight of 60 kDa, with a well-defined catalytic triad and an overarching flap at the active site [9]. C. rugosa lipases have a broad spectrum of specificities. The specificity of lipase is controlled by the molecular properties of the enzyme, structure of the substrate and factors affecting binding of the enzyme to the substrate. Benjamin and Pandey [ 10] isolated and

6 characterised three distinct forms of lipases from Candida rugosa (DM-2031), produced by solid-state fermentation. Three distinct forms of extra-cellular lipase (lipA, lipB and lipC) were isolated by ammonium sulphate precipitation, dialysis, ultra-filtration, and gel filtration using Sephadex-200. The purification was 43-fold with specific activity of 64.35 mg/ml. SDS-PAGE analysis of purified lipase revealed three distinct bands indicating the existence of three isoforms with apparent molecular weights of 64, 62, and 60 kDa. All three forms had optimal

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activity at 35–40oC and pH 7–8. Ag++ and Hg++ strongly inhibited the activity of all iso-forms, whereas Ca++ and Mg++ enhanced lipase activity. The activity of all three forms was completely inhibited

by

the

serine

protease

inhibitors

dichloroisocoumarin

and

pefabloc.

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Phenylmethanesulphonyl fluoride partially inhibited their activity. Potential industrial

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applications of C. rugosa lipase include the production of fatty acids and glycerol via hydrolysis of oils and fats, modification of composition and physical properties of triglyceride mixtures

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by inter-esterification and trans-esterification, and synthesis of inorganic solvents [11]. Diverse

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products ( e. g. juices, baked food, fermented vegetables, dairy enrichment, desirable interesterification of fats and oils to produce modified glycerides unobtainable by chemical inter-

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esterification) have been manufactured by free or immobilised C. rugosa lipases [12–14]. In the

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presence of a hydrophobic surface, the C. rugosa lipases will adsorb, establishing a new “open” structure where the active centre is fully exposed, thus allowing the lipases to hydrolyse drops of oils. This idea has been exploited to selectively immobilise C. rugosa lipases on a variety of hydrophobic supports via their open forms. Moreover, the open form of a lipase molecule can stabilise the open form of other lipases, creating dimers with altered catalytic features. This interaction is stable enough to use immobilised lipases to selectively adsorb other lipase

7 molecules. As a result, studying free lipases may become quite a complex task [3, 15– 19] . Moreover, the costs of producing lipases to catalyse versatile reactions which contribute to biotechnology processes are often prohibitive. Hence processes that do not require the physical presence of lipase in the final product and that use feedstocks that are fluids (or that can be treated as such) are more economic if the lipase is employed in an immobilised form. Polyhydroxybutyrates are widely used for enzyme immobilisation and have a plethora

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of applications in bio-catalysis, biosensor development, decontamination, and energy storage. Enzyme immobilisation is a simple and reproducible method for enzyme immobilisation that does not require sophisticated equipment. In this context, several supports can be adopted to

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perform enzyme immobilisation. In general, the carrier structure needs enough mechanical

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strength, resistance of chemical attack and microbial decomposition. Meanwhile, abundant reactive groups and hydrophilic chains are necessary on the surface of carriers. After

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immobilisation, other phenomena (desired or undesired) may occur, thus the researcher must

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detect these phenomena and develop tools to control them. For example, the first immobilisation may be through a one- point or a multipoint interaction, and after this

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immobilisation, the support may continue increasing the number (or even the quality) of the interactions involving new groups, as is the case for heterofunctional supports [ 7, 20, 21] .

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Mendes et al. [22] reported that lipase immobilised on PHB could produce higher bio-diesel yields than the free enzyme, due to increased catalytic activity. Biodiesel production yields have been compared using lipase immobilised on PHB versus lipase immobilised on other materials. Lipase immobilised on PHB yielded 98% bio-diesel, which was comparable to yields of enzymes immobilised on synthetic materials [22]. Candida rugosa lipase was successfully

8 immobilised on poly( 3- hydroxybutyrate- co- hydroxyvalerate) (PHBV) by physical adsorption with an efficiency of 30% [2]. However, the creation and application of immobilised Candida lipase on PHB has never been reported. PHB, from Breribacterium casei has been utilised to immobilise nattokinase. Deepak et al. [23] extracted and synthesized PHB nanoparticles using the nanoprecipitation technique. The authors observed a 20% increase in the enzymatic activity of nattokinase immobilised on PHB nanoparticles. Immobilisation of the enzyme also

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contributed to enhance enzyme stability. Examining the effects of temperature showed that the purified enzyme was stable up to 50°C, while the PHB-immobilised enzyme was stable at much higher temperatures (70°C) without loss of enzyme activity. However, no information regarding

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the production of monoacylglycerol (MAG) by lipase immobilised onto PHB has been reported.

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Therefore, we chose MAG production as our reaction model. Moreover, from an industrial

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point of view, monoacylglycerols are most widely used as raw materials for producing more lipophilic or more hydrophilic molecules for use in the cosmetic and food industries. The main

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disadvantage of the use of lipase in industrial applications is the high cost. To overcome this problem, lipase is employed in an immobilised form, allowing for continuous and repeated use

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of the enzyme. Normally, enzymes are immobilised using both chemical and physical methods [24]. Selecting an appropriate support material and technique for lipase immobilisation is very

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important, and therefore, the pursuit of suitable materials continues. Investigating the effectiveness of methods used to immobilise enzymes, which defines their potential use in industrial processes, is also ongoing. Cabrera- Padilla et al. [ 25] attempted to improve the efficiency of immobilised lipase on PHBV via physical adsorption using different ionic liquids as immobilisation additives. The results revealed that the activity of C. rugosa lipase on PHBV

9 increased from 30% to 78% in the absence of hydrophilic ionic liquids compared to when immobilisation additives were used [25]. Therefore, it is necessary to develop methods increase enzyme efficiency. The adsorption method of enzyme immobilisation is the most simple, easy and economical. A disadvantage of the adsorption immobilisation method is the weak interaction between the enzyme and the support that can results in low activity and stability [ 26] . Recently, lipase- catalysed glycerolysis for the biosynthesis of partial glycerides is

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receiving increased attention as a possible alternative to the classical method. The main reasons are the higher yields achieved and much milder reaction conditions, resulting in products of higher quality and less energy costs. However, the development of an easy, cheap and

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recyclable method is still a challenge. Physical adsorption may be a suitable method of

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immobilisation. In addition, utilisation of glutaraldehyde to modify the immobilised enzyme has been extensively studied. Glutaraldehyde, one of the most widely used chemical

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compounds, modifies primary amino groups of proteins but may eventually react with other groups (thiols, phenols, and imidazoles) [22]. It is a very effective cross-linker that is widely

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used for intermolecular cross-linking. Moreover, glutaraldehyde has been used to covalently

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immobilise enzymes to pre- existing supports. Glutaraldehyde is a reagent that can intermolecularly cross-link enzyme molecules immobilised on any type of supports, especially if

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the enzyme molecules are closely packed together [ 7, 28] . However, in most cases, glutaraldehyde is used to immobilise enzymes on supports bearing primary amino groups. Therefore, the combination of physical adsorption and glutaraldehyde-mediated cross-linking could be used to construct immobilised enzymes with better operational stability, avoiding the loss of adsorbed enzymes from the supports during the reaction. Moreover, glutaraldehyde acts

10 as a bi-functional reagent. This type of immobilisation forms a spacer arm between enzyme and carriers that is mediated by glutaraldehyde. The immobilisation of enzyme on carriers via a spacer arm seems to help avoid steric hindrance and increases enzyme activity [27]. With the introduction of a flexible spacer arm onto the supports, the enzyme can flexibly stretch and catch the substrate more easily. The process begins with enzyme adsorption on its support, followed by glutaraldehyde-mediated enzyme cross-linking. Glutaraldehyde is one of the most

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widely used reagents in the design of biocatalysts. It is a powerful cross-linker that has the advantage of being able to react with itself. Since we will u se pre-activated supports, we highlight the heterofunctional nature of the supports, as well as the drawbacks and advantages

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that heterofunctionality may have. Particular attention will be paid to the first event that causes

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the immobilization, which depends on experimental conditions that alter the enzyme orientation on the support surface. Thus, glutaraldehyde remains the most widely used

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compound for the design of biocatalysts and has the broadest application possibilities [28]. Hu et al [29] employed magnetic Fe3O4 nanoparticles modified with 3-aminopropyltriethoxysilane

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as carriers to immobilise lipase from Serratia marcescens ECU1010 (SmL) with glutaraldehyde

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as the coupling agent. The immobilised lipase showed a higher binding efficiency and activity recovery than that of lipase adsorbed directly onto the supports. Similar results have been

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reported for the introduction of an aminopropyl spacer arm as Bacillus licheniformis Larabinose isomerase (BLAI) was immobilised [30]. Many studies suggest that the spacer arm between enzyme and carriers could detach the enzyme from the carrier surface and prevent undesirable side attachment between enzyme molecules and the support. This immobilisation method favours the activity retention and improves the performance of the immobilised

11 enzyme [31]. The combination of physical adsorption and cross-linking for immobilising C. rugosa lipase on PHB particles has not been reported thus far. Therefore, this study focused on the use of PHB to immobilise lipase. The ability of a combination of immobilization processes to obtain higher immobilization yields was also evaluated immobilisation. Thereafter, immobilised enzyme was further characterised and used to produce acylglycerol. This is the first report to assess the combination of immobilisation techniques to immobilise Candida

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lipase on PHB via physical adsorption and glutaraldehyde. Moreover, the effectiveness of enzyme immobilisation using a combination of techniques was compared to other publications.

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2. Materials and methods 2.1. Materials

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Lipase powder from Candida rugosa was purchased from Sigma-Aldrich (USA). Nominal

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lipase activity was 2,381 U g- 1. C. rugosa lipase was immobilised on the external surface of intracellular poly(3-hydroxybutyrate) (PHB) beads. Commercial PHB had a particle size of 300

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μm, pore size of 0.2–0.5 μm and surface area of 0.9–2 μm2. PHB naturally originating from several bacterial species was purchased from Sigma-Aldrich(Germany). All other reagents and

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solvents used in this study were obtained from Merck(Germany) and were of analytical grade.

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2.2. Immobilisation using a combination of adsorption and cross-linking techniques First, C. rugosa lipase was immobilised on PHB particles by adsorption. PHB, in

powdered form (0.5 g), was soaked in 50 mL of anhydrous ethanol for 2 h at room temperature [32]. After the swelling step, excess ethanol was removed and PHB was washed with 20 mL of 0.2 M sodium phosphate buffer (pH 7.0) using vacuum filtration. Afterwards, PHB was added to

12 5 mL of lipase solutions (0.75 mg protein mL-1 solubilised in 5 mL of 0.2 M sodium phosphate buffer, pH 7.0). For all lipase preparations a protein loading of 2 mg g- 1 support was used in order to avoid diffusional problems. The immobilisation technique was performed based on the interfacial activation of lipases on hydrophobic supports at low ionic strength [33]. PHB beads with enzyme solution were kept under mild stirring (150 rpm) in a shaker for 30 min at 4 °C. Thereafter, 20 mL of 0.2 M sodium phosphate buffer at pH 7.0 was added, and the suspension

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was filtered through a Buchner funnel. The immobilised enzyme was washed on filter paper with another 20 mL of 0. 2 M sodium phosphate buffer at pH 7. 0 and dried in a vacuum desiccator for 8 h [34]. A schematic of the enzyme preparation process is shown in Fig. 1. For

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this immobilisation study, enzyme activity and the immobilised yield were calculated using the

𝐼𝐼𝐼𝐼𝐼 𝐼𝐼𝐼𝐼𝐼𝐼𝐼𝐼𝐼𝐼𝐼 𝐼𝐼𝐼𝐼𝐼𝐼𝐼𝐼 (𝐼)

𝐼𝐼𝐼𝐼𝐼 𝐼𝐼𝐼𝐼𝐼𝐼𝐼 𝐼𝐼𝐼𝐼𝐼𝐼𝐼 𝐼𝐼𝐼𝐼𝐼𝐼 𝐼𝐼𝐼𝐼𝐼𝐼𝐼𝐼 (𝐼)

×

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𝐼𝐼𝐼𝐼𝐼𝐼𝐼𝐼𝐼𝐼𝐼 𝐼𝐼𝐼𝐼𝐼 (%) =

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following formula:

100.

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Afterwards, glutaraldehyde was applied to enhance immobilisation yield. The effect of glutaraldehyde concentration (0–1.0 M) was also determined using a single-factor experimental

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design. After glutaraldehyde was added to the mixture, the sample was incubated at room

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temperature while mixing at 100 rpm for 2 h. Finally, the immobilised enzymes were filtered and treated using the same procedure previously described in the adsorption method. 2.3. Determination of hydrolytic activity and protein concentration Hydrolytic activities of free and immobilised lipase were assayed by the hydrolysis of 0.5% (w/v) p-nitrophenyl palmitate (p-NPP), according to the methodology described by Kuepethkaew et al. [35]. The substrate solution was prepared by dissolving 1.12 mM p-NPP in 10 mL of 2-

13 propanol. The reaction mixture consisting of 50 mM sodium phosphate buffer, pH 7.0 with 0.3% of Triton X-100 containing 10 mg immobilised enzyme or 25 μL free lipase was initiated by adding 0.25 mL of substrate and mixed for 5 min at 37 °C. The reaction was stopped by heat shock (1 min, 90 °C) followed by the addition of 0.5 mL of 0.25 M Na2CO3. Thereafter, the sample was centrifugation at 10,000 xg (4 oC) for 5 min. The quantity of liberated p-nitrophenol (p-NP) was determined by measuring absorption at 410 nm using a UV–Vis spectrophotometer

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(UV-2450, Shimadzu, Japan). Enzyme activity was determined using a p-NP standard curve (ԑ = 3.50 M-1 cm-1). One unit of enzyme activity was defined as the quantity of enzyme that liberated 1 µmol of p-nitrophenol per minute under the assay conditions. The level of enzyme protein

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was determined using the Bradford method with bovine serum albumin (BSA) as a standard

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[36]. 2.4.Characterisation of immobilised lipase on PHB

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The morphology of the immobilised lipase on PHB was evaluated by SEM (Phenom-World

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BV, Eindhoven, Netherlands) equipped with an energy dispersive spectrometer (EDS), which was operated at 10.0 kV.Samples were fixed to the holders with double-sided adhesive tape,

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demagnetised and then sputter-coated with a thin over-layer of gold to prevent sample-charging

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effects before they were examined using a microscope. The infrared spectra of PHB and immobilised lipase on PHB were recorded using Fourier transformed infrared spectroscopy (FTIR-8300, Shimadzu, Japan). 2.5. Optimum pH, optimum temperature and the kinetic study of free and immobilised lipase Optimum pH of free and immobilised enzymes were assayed in the 50 mM of glycine-HCl, citrate phosphate, phosphate, NaHCO3-Na2CO3, Na2HPO4-NaOH and KCl-NaOH buffer by

14 testing activities at pH values ranging from pH 2–12 at 37°C. The buffers used to assess optimal pH were glycine-HCl buffer (pH 2), sodium citrate phosphate buffer (pH 3–5), sodium phosphate buffer (pH 6–8), NaHCO3-Na2CO3 buffer (pH 9–10), Na2HPO4-NaOH buffer (pH 11) and KClNaOH buffer (pH 12). To assess optimum temperature, enzymatic activity was assessed at temperatures ranging from 30 to 60 °C in 50 mM sodium phosphate buffer, pH 7. The effects of different substrate concentrations (0.1–4 mM) on the activity of the immobilised enzyme were

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assessed under optimum conditions. 2.6 Determination of thermal stability

Thermal stability of the immobilised enzyme was determined by estimating residual

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activity under optimal conditions. The thermal stability was tested by incubating 10 mg of

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immobilised lipase at temperature ranging from 30 to 60 °C in 0.2 M sodium phosphate buffer

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(pH 7) for 1 h and then cooled on ice, after which the enzyme activity was determined. The activity of enzyme that had not been pre-incubated were used to define 100% enzyme activity.

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A Michaelis-Menten model was used to determine the values of Km and Vmax as shown in the equation:

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V = Vmax x [S] / km + [S]

where v is the reaction rate, Vmax is the maximum reaction rate achieved by the system, at

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saturating substrate concentration (U mL- 1), [S] is the substrate concentration (mmol L- 1), and Km is the Michaelis-Menten constant (mM). Then, the residual activities were determined for the half- life of the immobilised lipase as the percentage yield of activity compared to the activity of the untreated control. The half-life (t1/ 2) was calculated by the ratio ln 2/Kd. The thermal inactivation constant (Kd) was determined by the equation:

15 lnA = lnA0 – Kd x t where Ao and A are the initial activity and the activity after time t (min) respectively. 2.7. Storage stability The stabilities of the free and immobilised lipases were measured by calculating the residual activity. Free lipase was stored in a 0.2 M sodium phosphate buffer (pH 7) and the immobilised lipase was stored in wet form in a similar buffer at 4°C. Enzyme activity was

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assayed every 10 d for 60 d. 2.8. Enzyme reusability

The ability of the immobilised enzyme to hydrolyse p-NPP was examined under optimum

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conditions. Each batch consisted of a hydrolysis reaction for 10 min at 37 oC and pH 7.0. At the

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end of the reaction, the immobilised enzyme was separated by filtration with Whatman paper (pore size 0.2 μm), and washed three times with 10 mL 0.2 M sodium phosphate buffer (pH 7).

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The enzyme was then desiccated for 12 h at room temperature. The dried immobilised lipase

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was used in a new reaction using new substrate. Each reaction was initiated by the addition of an immobilised lipase wash in new reaction medium composed of 0. 25 mL of p- NPP as

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described above. The repeated reactions were run several times to measure the residual activity of immobilised lipase. The residual activity determined after the complete reaction was express

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as relative conversion. The conversion achieved in the first batch was set to 100. This procedure was repeated for 15 cycles. 2.9. Use of PHB-immobilised lipase to produce acylglycerol Glycerolysis experiments were carried out in batch systems. Palm olein and glycerol containing 4.0% (w/w) water were used as substrates for MAG. Ten grams of reaction mixtures

16 containing approximately 9 g of palm olein and 1 g of glycerol were prepared [37]. Mixtures were first incubated 10 min before the enzyme was added. Next, the substrate was mixed with either soluble or immobilised lipase. Reactions were incubated in an orbital shaker at 40 °C and 250 rpm for 48 h. Aliquots of 50 μL were taken from the reaction mixture at different time points, dissolved in 5 mL of methyl tert-butyl ether (MTBE) and filtered through a 0.45-μm syringe filter to remove the immobilised enzyme. Aliquots from these solutions were

dissolved in MTBE at a final concentration of 6 mg mL-1.

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evaporated using nitrogen until a residue of constant weight was obtained. The residue was re-

Preliminary products were obtained by analysing acylglycerol released using thin layer

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chromatography ( TLC) . The reaction was carried out on silica gel G- 60( Merck Millipore,

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Germany) . The migration solvent consisted of a mixture of petroleum ether: diethyl

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ether:methanol:acetic acid (90:7:2.5:0.5). Chromatograms were detected under a UV lamp. In some cases, the bands did not clearly appear under the UV lamp. In these cases, the TLC plate

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was exposed to iodine vapour, which allowed for the direct visualisation of bands. Moreover, ATR- FTIR was also used to investigate the chemical structures of MAG, DAG and

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triacylglycerol (TAG) using frequencies within the range of 4,000-400 cm-1 [38].

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For quantitative analysis, approximately 0.2 μL of the final transparent solution was analysed using gas chromatography. Separations were performed using a Hewlett-Packard 5890 series II gas chromatograph with on-column injection using a 7-m HP-5MS capillary column, 0.25 mm I.D. (Agilent Technologies, USA). Injector and detector temperatures of 40 and 340 °C were utilised, respectively. The temperature programme was as follows: initial temperatures were 40 °C and were heated 42 °C min-1 to 250 °C and held 15 min. Then temperatures increased

17 15 °C min- 1 to 250 to 325 °C, after which they were held 20 min. Calibration curves for MAG, DAG, TAG and FFAs were carried out using monoolein (99%), diolein (99%), triolein (99%) and oleic acid (C18:1) (99%), from Sigma-Aldrich (USA). The peaks were computed using GC Chemstation software [37]. 2.10. Statistical analysis All experiments were carried out in triplicate and the data are presented as mean ± standard

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deviation (SD). A completely randomised design was used throughout this study. Data was subjected to an analysis of variance (ANOVA) and a means comparison was carried out using Duncan’s multiple range tests. Statistical analysis was performed using the Statistical Package

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for Social Sciences (SPSS for Windows, SPSS Inc., USA).

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3. Results and discussion

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3.1 Effect of enzyme concentration on the immobilisation of lipase on PHB Lipase was immobilised on PHB using a combination of immobilisation processes. First,

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lipase was adsorbed onto PHB particles and the effective enzyme concentration (20-45 U mL-1 enzyme) was determined. Through physical adsorption for 30 min, the immobilised yield was

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27-43%, and thus, loading was determined to be 0.43-0.79 mg g- 1 support. The immobilisation yield increased as enzyme concentration increased. The highest immobilisation yield (43%) and

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loading onto the supports ( 0. 79 mg g- 1 support) was achieved when the initial lipase concentration was 45 U mL- 1 enzyme. For all lipase preparation, a protein loading of 2 mg g- 1 support were offered in order to avoid diffusional problems. Moreover, loading more than 2 mg g-1 support enzyme may generate steric hindrance that prevents the accessibility of the large substrate to the enzyme if the orientation towards the reaction medium is imperfect. The enzyme

18 loading somehow affects the enzyme stability. Table 1 showes yield and expressed activity when growing the amount of offered to lipase onto PHB. The high yield and activity demonstrates that some molecules may be packed together close enough to interact with each other, altering the final stability in a positive or a negative way, depending on the enzyme and inactivation conditions. However, this may not be true when the immobilisation rate is much higher than the diffusion rate [ 39] . In those cases, the enzyme molecules may be packed

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together, and some inter- molecular interactions may be produced. This is the case of C. antarctica lipase B (CALB) immobilised on octyl agarose, where the immobilisation is so rapid that the enzyme molecules are very close, enabling a small reagent such as glutaraldehyde to

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produce inter-molecular cross-linking [58]. This “crowding” effect is reported to stabilise using

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free enzymes [39], it was found that when using CALB immobilised on octyl agarose and the support was fully loaded (1 mg g- 1 support), the enzyme stability decreased [58]. This negative

lP

effect was explained by interactions between nearby partially unfolded enzyme molecules (perhaps exhibiting hydrophobic pockets) that prevented the recovery of enzyme activity when

na

measure under mild conditions during inactivation experiments. Stabilising effects of crowded immobilised enzyme may be derived from immobility of the enzyme structure. If one

ur

enzyme is closely immobilised to another, they will collide when parts of the enzyme try to

Jo

move and thus conformational changes may be avoided. This suggests that PHB could be utilised as a promising hydrophobic support. Advantages of physical adsorption include ease of use, reasonable cost and the capacity for recycling of support materials when enzymes are no longer viable. However, changes to enzyme desorption are the main disadvantage of this immobilisation procedure [2, 40]. Therefore, after physical adsorption, cross-linking of enzymes

19 to the support surface was evaluated via treatment with various glutaraldehyde concentrations. An initial lipase concentration of 45 U mL-1 enzymes was used in subsequent experiments.

3.2 Effect of glutaraldehyde concentration on lipase immobilisation on PHB After physical adsorption, the crosslinking technique was applied to enhance the efficiency of enzyme immobilisation using glutaraldehyde as a cross- linking agent. The effect of glutaraldehyde at different concentrations (0-1.0 M) were evaluated. Glutaraldehyde treatment

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is normally performed to prevent enzyme leaching and maintain enzyme activity [41]. The effect of glutaraldehyde concentration on yield of immobilised lipase is shown in Table 1. When the

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concentration of glutaraldehyde increased from 0 to 1.0 M, the yield of immobilised lipase increased. The highest yield of immobilised lipase (92%) associated with an activity of 1.94 U

re

mg- 1 support was observed when 0. 6 M of glutaraldehyde was added. Glutaraldehyde

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concentration significant influenced enzymatic activity. As the concentration of glutaraldehyde surpassed 0.6 M, the immobilised activity gradually decreased. Based on the data in Table 2,

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0.6 M glutaraldehyde was selected. The optimum conditions for creating immobilised lipase involved the addition of 0.6 M glutaraldehyde and 45 U mL-1 enzyme while mixing at 150 rpm

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and 4°C for 30 min. In order to determine the approprite techniques for the immobilisation of

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C. rugosa lipase, enzyme activities and immobilisation yields were compared ( Table 3) . Combination techniques produced the highest levels of enzyme activity and the highest yields. The enzyme activity for combined techniques was 41.4 U mL-1 enzyme. The original free lipase had an activity of 45 U mL- 1, enzyme and the immobilisation yield was 92%. The standalone methods of physical adsorption and cross- linking onto PHB produced only 43% and 34% ,

20 immobilisation yields, respectively. Low immobilisation yields obtained for physical adsorption might be due to the weak bond between the enzyme and its support. The enzyme may leak during the recovery process (washing step). The addition glutaraldehyde of the crosslinker stabilise immobilised enzyme particles and helps prevent the enzyme leakage [42]. When glutaraldehyde is used as a cross- linker, the enzyme is primarily adsorbed by physical adsorption onto the PHB support, and then the enzyme- support compound is treated with

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glutaraldehyde under mild conditions [7, 17, 28]. In such a strategy, all the primary amino groups of the enzyme and the support are modified with just one molecule of the glutaraldehyde and an intense multipoint covalent enzyme- support attachment may be achieved [ 28] . de

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Oliveira et al. [43] demonstrated that the best immobilisation conditions for Rhizomucor miehei

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on chitosan beads consisted of first adsorbing lipase on chitosan beads at 4 °C for 1 h, 220 rpm followed by cross-linking the chitosan beads-enzyme complex with glutaraldehyde 0.6% v.v𝐼1 at

lP

pH 7.

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In this study, immobilised enzyme was recovered efficiently (> 92%) when compared to other studies. C. rugosa lipase immobilised on PHBV in aqueous solution with and without

ur

ionic liquid resulted in enzyme efficiencies of only 78% and 30% , respectively [ 22, 35] . Therefore, a combination of immobilisation methods shows the greatest potential for industrial

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applications. The physical adsorption of C. rugosa lipase on PHB cross- linked with glutaraldehyde enhanced enzyme efficiency. Moreover, de Oliveira et al. [43] suggested that glutaraldehyde cross- linking of lipases previously adsorbed on chitosan supports in the presence of Triton X-100 is an appropriate alternative to obtain derivatives with relatively good catalytically properties when using this surfactant to immobilise R. miehei lipase onto

21 chitosan support. Lipase activity can be improved in the presence of surfactants, which is mainly attributed to the breakdown of lipase aggregates (dimmers) and/or the shift towards the more active open form. It is believed that lipase- surfactant interactions are primarily hydrophobic and this may aid in immobilisation. However, the nature of the hydrophobic group on the surfactant also contributes to the contact, and the intensity of such interactions may be responsible for the breakdown of these self-assembled structures [43]. Moreover, it is important to

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consider that detergents may have complex effects on enzyme properties, in some cases affecting enzyme inactivation [44].

3.3 Characterization of immobilised lipase on PHB

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The examination of the surface of PHB and immobilised enzyme were performed using a scanning electron microscope which revealed important differences in the surface before and

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after immobilisation. The surface of PHB is highly regular and the addition of lipase covering

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its entire surface as can be seen in Figure 2. The surface texture after immobilisation was very rough. It is noteworthy to mention that the surface texture of immobilised lipase from Candida

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rugosa on PHB favours whole- surface functionalization, leading to coupling of yield and enzyme loading without losses in enzyme activity or specificity. Large surface areas and

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porosity may lead to higher enzyme loading per unit of mass. Moreover, the small PHB particles

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used in this study have larger surface areas than large PHB beads. The particle size of PHB is an important factor when used as support material. Mendes et al, [22] immobilised five microbial lipases by hydrophobic adsorption on smaller (75-90 μm) or large (750-1,180 μm) PHB beads. Their results showed that lipase was completely immobilised on small PHB beads. During immobilisation by physical adsorption on highly hydrophobic matrices, the lipase undergoes conformational changes in the presence of the support surface, and is adsorbed in the open

22 conformation ( monomolecular structure) [ 40] . However, the large PHB bead affected immobilised yield due to diffusion of the bimolecular lipases aggregates that may reduce the immobilisation rate with increasing particle size of the support. Generally, the surface area of small particles is higher than large particles [22]. Hence, more surface area on small PHB beads is available for the immobilisation of the lipases. The hydrolytic, specific and recovered activities of the biocatalysts were also strongly influenced by the particle size of the support.

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Small PHB derivatives displayed higher hydrolytic, specific and recovered activities than derivatives from large PHB. These results can be explained by steric effects of large PHB beads which significantly influence lipase conformation, leading to the changes in enzyme activity

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or diffusional limitations that restrict contact between the lipase active site and its substrate. It

of the support beads increases [22].

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is well known that diffusional limitations become more of a restraining factor as the diameter

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PHB and the immobilised enzyme were subjected to FT-IR analysis, and their spectra are presented in Figure 3. PHB beads and the enzyme were subjected to ATR-FTIR analysis and

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their spectra are presented in Figure 3. The spectrum of the support revealed a sharp adsorption band at 1,727 cm- 1, which corresponds to the ester carbonyl group. An additional band was

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apparent at 1,279 cm- 1, which corresponds to the CH group. A series of bands between 1,000

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and 1,300 cm- 1 indicate stretching of the C-O bonds in the ester group. The FTIR spectrum of PHB with immobilised enzyme shows the ester bands. Apart from the above-mentioned bands, other bands were present in the spectrum of lipase, such as the two signals at 1,700 and 1,490 cm- 1 , which were classified as the scissor vibrations of CONH. Further, a signal at 2,920 cm- 1 was generated by the C– H symmetric stretching vibrations of – CH2 and – CH3 groups. In

23 addition, the most important bands for lipase are the amide I and amide II bands seen at 1,645 and1,542 cm- 1, respectively [45, 46]. These results confirm successful adsorption on the PHB bead [47]. The presence of amide I and amide II bands also suggest that enzyme immobilisation did not interfere with its activity [48]. 3.4 Optimum pH, optimum temperature and kinetic parameters of free and immobilised lipase The effect of pH on the activity of free and immobilised lipase was investigated under pH

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ranging from 2-12. As illustrated in Figure 4a, the maximum activity was observed at pH 7 for free enzyme and pH 9 for immobilised enzyme. The optimum pH for immobilised enzyme (pH 9) shifted to a more alkali value compared to that of the free form (pH 7). This alkaline shift

-p

could be due to secondary interactions (e.g. ionic and polar interactions or hydrogen bonding)

re

between the enzyme and the hydrophobic support [49]. When the enzyme was immobilised, the surface of the support was negatively charged. After enzymes were immobilised, the negatively

lP

charged support increased the density of protons around the active site of the immobilised enzyme. Therefore, the pH of the bulk solution was higher than that of the immobilised enzyme.

na

To compensate for this effect, the optimum pH of the immobilised enzyme shifted to an increased pH value [50, 51]. A possible cause of this phenomenon is the interaction between the

ur

support and enzyme, which stabilised the conformation of the enzymes active site. Therefore,

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the immobilised enzyme can maintain its activity within a wider pH range than free enzyme. PHB-support may be too reactive with enzymes to use this strategy to direct enzyme

immobilisation, as it can covalently immobilise proteins in the range of pH 2-12. However, the reactivity between the protein groups and the support depends on pH, therefore the orientation of the enzyme molecules on the support may be altered by changing the immobilisation pH.

24 Blocking of the support with nucleophiles ( e. g. , ethylenediamine) eliminates the chemical reactivity of the support, helping to avoid undesired enzyme- support covalent bonds [ 44] . Moreover, stabilisation and destabilisation can be affected by pH. The free enzyme may experience protein aggregation (mainly near to the isoelectric point). This may be caused by undesired enzyme-interactions that can stabilise incorrect enzyme structures. The tolerance of immobilised enzymes to alkali pH could enhance the feasibility of applying immobilised

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lipases to a variety of industries. Herein, covalent conjugation methods were implied for immobilisation. Figure 5 shows the activity of lipase on PHB. Enzyme activity was slow at all studied pH values. A significant decrease in enzyme activity was associated with free enzyme.

-p

Lipases from Candida rugose, molecular masse of 120,000, isoelectric point (pI) of

re

pH 4.5 and optimal activity between pH 6.5 and 7.5 [52]. The isoelectric point of the enzyme is

lP

important. During enzyme immobilisation, the buffer should have a pH value that favours electrostatic interactions with the carrier surface. Lipase enzyme preparation seems to

na

be very unstable in both alkaline ( pH>8) and highly acidic ( pH<4) solutions. This may be explained by the structural changes to the enzyme caused by pH variation. It is suggested that

ur

the variation in activity with varying pH (within a range of two to three pH units each side of

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the isoelectric points) is reversible for most enzymes [53]. Also, at acidic pH, the enzyme may form high molecular weight aggregates or nonspecific associations with other accompanying proteins due to its hydrophobic nature and proximity to its isoelectric point [54]. A protein with zero net charge (at pI) will have maximum hydrophobicity. However, at a pH where protein and adsorbent have similar charges, repulsion may occur which can reduce the interaction. Moreover, close to their isoelectric point, enzyme tend to spontaneously form less active

25 aggregates. As seen in Figure 4a, lipase immobilised at pH 2-4 showed only 65-70% activity. This result, therefore, can be explained by aggregate formation and enzyme-support repulsion near the isoelectric point of lipase from Candida rugosa (4.5). However, pH extremes would cause irreversible denaturation. At pH >8, activity is around 45–70%. This is mainly because the hydrophobic interactions are not greatly influenced by changes in the solution pH. However, if electrostatic forces are important for tadsorption, changes in pH over the isoelectric point of

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the protein will have a large impact on the protein binding constant [55]. The optimum temperature for lipase activity was also assessed, and the results are shown in Figure 4b. Immobilised lipase was maximally active at 40°C, while activity the soluble

-p

enzyme was optimal at 30°C. Moreover, the immobilised enzyme maintained relatively high

re

activity across a wider temperature range than free lipase. This suggests that enzyme immobilisation resulted in more sturdiess than its free form. The increase in the optimum

lP

temperature of the immobilised enzyme was caused by changes in the physical and chemical properties of the enzyme. Kaewthong et al. [34], working with Pseudomonas lipase, found that

na

immobilisation on Accurel promoted a shift in its temperature profile. This arose from a lower

ur

degree of restriction of substrate diffusion of and products at higher temperatures. The kinetic constants of the immobilised enzyme were calculated (Figure 6). The Km and Vmax of free lipase

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from C. rugosa were 3. 5 mM and 1. 4 × 10- 3 mmol min- 1, respectively. However, lipase immobilised on PHB had a Km value of 1. 2 mM and Vmax value of 2. 5 × 10- 3 mmol min1

. Kareem et al. [56] reported 1.7 mM and 6.0 × 10-3 mmole min-1 as the Km and Vmax for a lipase

isolated from Aspergillus niger immobilised on hydrogel beads, respectively. Km and Vmax values mainly depend on the enzyme source and reflect the specificity of the enzyme

26 toward its substrate. For instance, a decrease in the Km and an increase in the Vmax of an enzyme indicate that it has increased sensitivity toward the substrate [49]. 3.5 Thermal stability of free and immobilised lipase The thermal stability of soluble and immobilised lipase on PHB was determined at 30–60 °C. More than 80% of the immobilised enzyme activity remained after 2 h incubation at all temperature tested. Furthermore, the immobilized lipase was slightly more stable than the

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soluble enzyme (Figure 7). The resistance of immobilised lipase to temperature changes is an important advantage which can be exploited for commercial use. High temperatures can change enzyme conformation, resulting in decreasing enzyme activity [57]. On the other hand, nearby

-p

hydrophobic enzymes on the support may interact, preventing the reversibility of these

re

conformational changes with effects on activity [ 58] . The increased thermostability of the immobilised lipase suggests that hydrophobic immobilisation helps protect enzyme

lP

structure from conformational changes induced by temperature. For example, immobilisation

na

may fix the enzyme in such a way that reduces susceptibility to heat denaturation. Our results are consistent with others studies using lipase immobilisation on hydrophobic supports. Lipases

ur

from C. rugosa and Mucor miehei hydrophobically immobilised on octadecyl- Sepabeads respectively retain 100% and 60% of their activity after 2 h [33].

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The half-life of free and immobilised lipase was calculated. The free enzyme maintained

~74% activity up to 2 h at 40 °C and the half-life was 18.37 h. The half-life of the free enzyme at 30, 50 and 60 °C were 16.37, 13.22 and 13.10 h, respectively. Immobilised lipase showed higher thermal stability than the free enzyme, maintaining 80% activity after 2 h at 40 oC with a halflife of 25.97 h. At 30, 50, and 60 °C, the immobilised enzyme retained more than 62% activity

27 after 2 h, with half- lives of 17. 42, 20. 50 and 12. 35 h, respectively. Generally, enhanced hydrophobic interactions increase thermal stability [59]. The hydrophobicity of the support was increased upon enzyme immobilisation. However, aggregation of the enzyme on the support could weaken its hydrophobicity. Consequently, thermal stability of the enzyme was improved upon immobilisation [60-62]. 3.6 Storage capacity of immobilised enzyme

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The capacity of immobilised lipase to be stored for future use is one of the most significant measures of enzyme usability, and could suggest that immobilised lipase is advantageous to free lipase [49]. Candida lipase immobilised on PHB particles had an improved capacity to be

-p

stored relative to free enzyme. Immobilised lipase retained 80% activity after storage for 30 d

re

in 0.2 M sodium phosphate buffer (pH 7) at 4 °C. Afterward, immobilised enzymes began to gradually lose their activity, and 50% enzyme activity was observed after they were stored 60 d

lP

(Figure 8). In comparison, the free enzyme retained only 50% activity after 10 d and almost all of its activity was lost after being stored 20 d. Kaewthong et al. [34] found that Pseudomonas

na

lipase the immobilised on Accurel powder 𝐼90% activity after being stored 24 h at 45 °C. This

ur

confirms that physical adsorption and cross- linking lead to improved enzyme stability by preventing structural changes. In addition, multiple attachments between the enzyme and the

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support preventing intermolecular process such as proteolysis and aggregation and therefore, creates a more rigid enzyme molecule [ 63] . Moreover, attachment also prevents conformational changes to the enzyme induced by distortion. The immobilized enzyme storage stability observed in this study is favourable with respect to its potential industrial use. 3.7 Enzyme reusability

28 In order to reduce the costs associated with enzymatic digestion, it is possible to reuse immobilised lipase for several successive rounds of hydrolysis, which would be extremely advantageous in an industrial setting [64]. Results here showed that immobilised lipase retained 50% activity for more than 14 cycles (Figure 9). The activity loss may be related to enzyme release from CTA, possibly caused by the detergent (0.25% v/v Triton X-100) used to prepare the p-NPP substrate [65]. Cabrera-Padilla et al. [2] reported that C. rugosa lipase immobilised on

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PHBV by physical adsorption retained 50% activity after 12 cycles of reuse. Ranieri et al. [66] reported that the lipase immobilised on asymmetric ceramic hollow fibre membranes could be used only six times. The capacity of an enzyme to be reused depends on two main parameters:

-p

storage stability and repeated use. These are commonly referred to as operational stability

re

parameters [49]. Biocatalyst reusability can be affected by several parameters including the properties of immobilisation matrix, immobilisation protocol, reaction type and reaction

lP

temperature. Diffusion caused by accumulation of the product on the surface of the support is also an important factor in enzyme reusability [ 67, 68] . Generally, physical adsorption of

na

immobilised lipase was considered to provide weak affinity between the carrier and lipase molecule, which does not support reusability. In this study, the high recycling stability of lipase

ur

immobilised on PHB was likely due to “affinity-like” hydrophobic interaction between the PHB

Jo

support and lipase. Moreover, the observed catalytic activity and reusability can be perhaps attributed to stabilisation of the open-form of the lipases by hydrophobic interactions [69]. In general, the lipase immobilisation of lipase by physical adsorption on hydrophobic supports not only stabilised lipase molecules in open conformation and promotes their hyperactivation but also enables the reuse of support by desorption of inactive enzymes using different detergents, thereby making this strategy especially suitable for lipase immobilisation Manoel

29 et al. [69] immobilised lipases from Thermomyces lanuginosus and Pseudomonas cepacea on octyl and cyanogen bromide agarose. The found that octyl agarose derivatives present a stabilised open form, while covalent preparations maintain closing/opening equilibrium. Since the product was highly soluble, we have not observed any product accumulation on the matrix surfaces [ 68] . Our reusability results indicate a high level of operational stability for immobilised lipase on PHB by a combinayion of physical adsorption and cross- linking

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methods increasing enzyme stability but also permits enzyme reuse, which suggests that the enzyme is suitable for large-scale industrial use.

3.8 Application of immobilised lipase on PHB to the production of acylglycerol

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Immobilised lipase adsorbed on PHB support and cross-linked with glutaraldehyde were

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subjected to a glycerolysis reaction. A control sample was prepared using the same assay conditions without enzyme. The finished product, which contained about 48% MAG, 32% DAG,

lP

10% TAG, 2% FFA and 8% glycerol was satisfactory. Similar to what is observed in most

na

manufacturing plants, the MAG concentration was only 35–50 %(w/w) [70]. A sample was taken for ATR-FTIR and TLC analysis to confirm the composition of obtained esterified products.

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Figure 10 shows a TLC plate on which estrified products produced by Candida rugosa free and immobilised lipases were examined. ATR-FTIR analysis showed the existence of a carbonyl

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group at wave numbers 1,743 and 1,744 cm- 1, indicating ester formation (Figure 11). These results indicate the existence of the three main products corresponding to MAG, DAG and TAG. Commercially available immobilised lipases are, in general, immobilised by physical adsorption on organic matrices such as Pseudozyma antarctica type B immobilised on Lewatit VPOC 1600 consisting of poly( methyl methacrylate- co- divinylbenzene) ( Novozym® 435)

30 and Mucor miehei immobilised on macroporous anion exchange resin (Lipozyme® RM IM) from Novozymes [22]. These biocatalysts have high costs which are mainly associated with the price of the support [71]. Therefore, the use of cheaper supports for lipase immobilisation is deal for large scale applications. According to our results, lipase immobilised on PHB displays promise in acylglycerol synthesis, which has vast applications in the cosmetic, pharmaceutical and food industries as an emulsifier in bakery products, chewing gum, shortening, whipped

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toppings, margarine, confections, beverages and ice creams. 4. Conclusions

Out finding suggest that PHB has the potential to be utilise as a hydrophobic support.

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Immobilisation of Candida lipase on PHB by adding 0.6 M glutaraldehyde and 45 U mL- 1

re

enzyme and mixing at 150 rpm at 4°C for 30 min produced the highest (92%) and activity (1.94 mg g- 1 support). The maximum activity of immobilised enzyme was observed at pH 9 and 40 o

lP

C, indicating that immobilised lipase was more tolerant to pH and temperature than free lipase.

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Enzymes immobilised on PHB particles and cross-linked with glutaraldehyde had improved thermal stability, storage stability and reusability. Additionally, lipase immobilised on PHB

ur

was suitable for acylglycerol production, while a 48% yield of monoacylglycerol was obtained by glycerolysis of palm olein with glycerol. Futher studies investigating the optimum conditions

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required for the continuous production of acylglycerol in a reactor will be required before these protocols are applied in an industrial setting.

Author statement Highlights Narisa Binhayeeding: Methodology, Data analysis, Writing- Original draft preparation

31 Tewan Yunu: Lab-assistant Nisa Pichid: Lab-assistant Sappasith Klomklao: Mentor Kanokphorn Sangkharak: Conceptualization, Methodology, Data analysis, Writing- Reviewing and Editing, Mentor Conflict of interest

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The authors declare no conflict of interest.

Acknowledgement

The authors would like to thank the Thailand Research Fund (TRF) for the Research and

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Researcher for Industries Grant (RRi) (project number PHD 61I0033), and the Research and Development Institute at Thaksin University for financial support.

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43 Table legends Table 1. Yield and enzyme activity of immobilised lipase. Influence of loading of enzyme. The activities were assayed by hydrolysing p-NPP in 50 mM sodium phosphate buffer (pH 7.0) at 37 °C. Table 2. The optimisation of condition for lipase immobilisation. Influence of glutaraldehyde concentration. The activities were assayed by hydrolysing p-NPP in 50 mM sodium phosphate

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buffer (pH 7.0) at 37 °C.

Jo

ur

na

lP

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-p

Table 3. Comparison of activity of immobilised lipase with different methods and materials.

44 Table 1. Yield and enzyme activity of immobilised lipase. Influence of loading of enzyme. The activities were assayed by hydrolysing p-NPP in 50 mM sodium phosphate buffer (pH 7.0) at 37 °C. Immobilised yield (%)

Immobilised activity

(U mL-1 enzyme)

(mg g-1 support)

20

0.43

20a

25

0.50

22b

30

0.57

35

0.65

40

0.72

45

0.79

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Enzyme activity

29c

34d

-p

40e

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na

lP

re

45f

45 Table 2. The optimisation of conditions for lipase immobilisation. Influence of glutaraldehyde concentration. The activities were assayed by hydrolysing p-NPP in 50 mM sodium phosphate buffer (pH 7.0) at 37 °C. Immobilised yield (%)

Immobilised activity

concentration (M)

(mg g-1 support)

0

0.79a

48a

0.2

1.82c

80c

0.4

1.82c

0.6

1.94d

0.8

1.80b

1.0

1.80b

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Glutaraldehyde

80c

92d

-p

79b

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79b

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na

lP

Different letters in the same column denote significant differences (p < 0.05).

46 Table 3. Comparison of activity of immobilised lipase with different methods and materials. Immobilisation techniques

Free C. rugosa

Physical adsorption

lipase

Cross-linking

Physical adsorption and cross-linking

Enzyme activity

45

19.4

15.3

41.4

Not determined

0.79

0.63

1.94

Not determined

43

34

(U mL-1 enzyme) Immobilised activity

Immobilisation

yield

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lP

re

-p

(%)

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( mg g-1 support) 92

47 Figure legends Figure 1. A schematic of the enzyme preparation process. Candida rugosa lipase was immobilised on PHB beads with a protein loading of 1 mg g- 1 support at low ionic strength under 150 rpm, 30 min at 4 oC. Figure 2. SEM image of sample. SEM image of polyhydroxybutyrate surface (a) before and (b) after lipase immobilisation.

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Figure 3. ATR-FTIR spectra of samples. Spectra of polyhydroxybutyrate (a) before and (b) after lipase immobilisation.

Figure 4. Influence of pH and temperature on the activity of free and immobilised lipase. (a)

-p

Activity of (𝐼) free and (𝐼) immobilised lipase was determined at 37 oC under different pH

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conditions. ( b) Activity of ( 𝐼 ) free and ( 𝐼 ) immobilised lipase was evaluated at different

lP

temperature under pH 7.0.

Figure 5. Activity of lipase onto PHB. Activity of ( a) free and (b) immobilised lipase was

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determined at 37 oC under different pH conditions for 60 min. Figure 6. Kinetic parameters of free and immobilised lipase. ( a) Michaelis-Menten plot. ( b)

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Lineweaver-Burk plot.

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Figure 7. Thermal stability of free and immobilised lipase. Thermal stability of (a) free and (b) immobilised lipase was evaluated following incubation at (𝐼) 30 °C, (𝐼) 40 °C, (𝐼) 50 °C and (x) 60 °C. All lipase hydrolysis reactions were performed at pH 7.0. Figure 8. Operational stability of free and immobilised lipase. Operational stability of (𝐼) free and (𝐼) immobilised lipase operated under batch experiment at 4 oC and pH 7.0

48 Figure 9. Reusability of enzyme. Reusability of immobilised lipase was tested using p-NPP as a substrate at 37 °C under pH 7.0. Figure 10. TLC analyses of samples. TLC analyses of glycerolysis production liberated after glycerolysis of 9 g palm olein and 1 g glycerol containing 4.0% (w/w) water by free (lane 1) and immobilised lipase (lane 2) at 40 oC. Figure 11. ATR-FTIR spectra of samples. Spectra of acylglycerol liberated after glycerolysis

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na

lP

re

-p

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of palm olein and glycerol by (a) free and (b) immobilised lipase.

Jo

ur

na

lP

re

-p

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49

Figure 1. A schematic of the enzyme preparation process. Candida rugosa lipase was immobilised on PHB beads with a protein loading of 1 mg g- 1 support at low ionic strength under 150 rpm, 30 min at 4 oC.

ro of

-p

re

lP

na

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50

.

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51

Jo

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na

lP

re

-p

(a)

(b)

Figure 2. SEM image of sample. SEM image of polyhydroxybutyrate surface (a) before and (b) after lipase immobilisation.

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52

-p

Figure 3. ATR-FTIR spectra of samples. Spectra of polyhydroxybutyrate (a) before and (b) after

Jo

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na

lP

re

lipase immobilisation.

53

100 90

Relative activity (%)

80 70 60 50 40 30 20

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10 0 2

4

6

7

8

9

10

pH (a)

90

re

70

lP

60 50 40

20

ur

10

na

Relative activity (%)

80

30

12

-p

100

11

0

30

50

60

Temperature (oC)

Jo

(b)

40

Figure 4. Influence of pH and temperature on the activity of free and immobilised lipase. (a) Activity of (𝐼) free and (𝐼) immobilised lipase was determined at 37 oC under different pH conditions. ( b) Activity of ( 𝐼 ) free and ( 𝐼 ) immobilised lipase was evaluated at different temperature under pH 7.0.

ro of

-p

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lP

na

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54

55

100 90

pH2

70

pH4

60

pH6

Relative activity (%)

80

pH7 50

pH8

40

pH9

30

pH10

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pH11

20

pH12

10 0

15

30

45

(a)

re

100 90

lP

80 70

pH4 pH6

na

60

pH2

50 40

pH7 pH8 pH9

ur

Relative activity (%)

60

-p

0

30

Jo

20

pH10 pH11 pH12

10 0

0

(b)

15

30

45

60

Time (min)

Figure 5. Activity of lipase onto PHB. Activity of ( a) free and (b) immobilised lipase was

56 determined at 37 oC under different pH conditions for 60 min.

2

1.5

1

0.5

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Immobilized enzyme activity (U mL-1 )

2.5

0 0.1

0.3

0.6

0.8

1.2

1.6

2.5

4

Concentration (mM)

-p

(a)

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6

R² = 0.9991

lP 0

-10

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-15

2

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1/V (x10-3 mmole min-1)

4

-5

0

5

10

15

-2

-4

-6

1/S (mM)

(b)

Figure 6. Kinetic parameters of free and immobilised lipase. (a) Michaelis-Menten plot. (b) Lineweaver-Burk plot.

57

100 90

Relative activity (%)

80 70 60 50 40 30

10 0 0

15

30

45

60

75

Incubation time (h)

100

105

120

re

90 80

lP

70 60 50 40

na

Relative activity (%)

90

-p

(a)

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20

30 20

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10 0

Jo

0

15

30

45

60

75

90

105

120

Incubation time (h)

(b)

Figure 7. Thermal stability of free and immobilised lipase. Thermal stability of (a) free and (b) immobilised lipase was evaluated following incubation at (𝐼) 30 °C, (𝐼) 40 °C, (𝐼) 50 °C and (x) 60 °C. All lipase hydrolysis reactions were performed at pH 7.0.

58

100 90

70 60 50 40 30

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Relative activity (%)

80

20 10 0 10

20

30

40

50

-p

0

60

Incubation time (days)

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Figure 8. Operational stability of free and immobilised lipase. Operational stability of (𝐼) free

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na

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and (𝐼) immobilised lipase operated under batch experiment at 4 oC and pH 7.0

59

100 90

Relative activity (%)

80 70 60 50 40

20 10 0 2

3

4

5

6

7

8

Cycles

9

10

11

-p

1

ro of

30

12

13

14

15

Jo

ur

na

lP

a substrate at 37 °C under pH 7.0.

re

Figure 9. Reusability of enzyme. Reusability of immobilised lipase was tested using p-NPP as

ro of

60

-p

Figure 10. TLC analyses of samples. TLC analyses of glycerolysis production liberated after

Jo

ur

na

lP

immobilised lipase (lane 2) at 40 oC.

re

glycerolysis of 9 g palm olein and 1 g glycerol containing 4.0% (w/w) water by free (lane 1) and

61 (a) 110

% Transmittance

100 90 80 70 60

40 4000

3500

3000

ro of

50

2500 2000 1500 -1 Wavenumber (cm )

110

re

100

lP

90 80 70 60 50

Jo

ur

40 4000

na

% Transmittance

500

-p

(b)

1000

3500

3000

2500 2000 1500 -1 Wavenumber (cm )

1000

500

Figure 11. ATR-FTIR spectra of samples. Spectra of acylglycerol liberated after glycerolysis of palm olein and glycerol by (a) free and (b) immobilised lipase.