Biosynthesis of butyl esters from crude oil of palm fruit and kernel using halophilic lipase secretion by Marinobacter litoralis SW-45

Abstract

Initially, a new moderate halophilic strain was locally isolated from seawater. The partial 16S rRNA sequence analysis positioned the organism in Marinobacter genus and was named ‘Marinobacter litoralis SW-45’. This study further demonstrates successful utilization of the halophilic M. litoralis SW-45 lipase (MLL) for butyl ester synthesis from crude palm fruit oil (CPO) and kernel oil (CPKO) in heptane and solvent-free system, respectively, using hydroesterification. Hydrolysis and esterification of enzymatic [Thermomyces lanuginosus lipase (TLL)] hydrolysis of CPO and CPKO to free fatty acids (FFA) followed by MLL-catalytic esterification of the concentrated FFAs with butanol (acyl acceptor) to synthesize butyl esters were performed. A one-factor-at-a-time technique (OFAT) was used to study the influence of physicochemical factors on the esterification reaction. Under optimal esterification conditions of 40 and 45 °C, 150 and 230 rpm, 50% (v/v) biocatalyst concentration, 1:1 and 5:1 butanol:FFA, 9% and 15% (w/v) NaCl, 60 and 15 min reaction time for CPO- and CPKO-derived FFA esterification system, maximum ester conversion of 62.2% and 69.1%, respectively, was attained. Gas chromatography (GC) analysis confirmed the products formed as butyl esters. These results showed halophilic lipase has promising potential to be used for biosynthesis of butyl esters in oleochemical industry.

Electronic supplementary material

The online version of this article (10.1007/s13205-019-1845-y) contains supplementary material, which is available to authorized users.

Introduction

The academia and indeed industrial sector have over quite a period of time now consistently devoted a considerable amount of time to developing novel and more efficient biocatalysts that are capable of tolerating harsh conditions of most industrial and biotechnological processes. There has been a change in course from conventional process to a greener approach for the synthesis of bioproducts. This has been continuously attributed to several attractive advantages of enzymatic processes over the traditional chemical reactions. These advantages include biodegradability, derivatization from renewable resources, the capability to function under relatively mild-temperature conditions, reduced secondary reactions, and low energy demand (Jemli et al. 2016). However, as mentioned earlier, most industrial processes often function in harsher conditions of extreme pH, elevated temperature and pressure, non-aqueous media, and oxidative conditions that normally inactivate mesophilic microbial enzymes (Jemli et al. 2016). Therefore, enzymes from extremophiles particularly halophiles have been proposed as suitable candidates in the aforementioned harsh reaction conditions (Veit 2004).

Recently, attention has been drawn towards halophilic enzymes as ideal alternative to catalyse reactions in harsh industrial conditions because of their polyextremophilic nature, that is, the capacity to show activity and stability in multiple extreme conditions such as increased salt concentrations, wide-range pH tolerance, thermal denaturation, and organic solvent tolerance (Delgado-García et al. 2012; Munawar and Engel 2013). Halophilic hydrolases such as halophilic lipases are reportedly advantageous especially with regard to their elevated temperature tolerance and organic solvent compatibility and stability, together with their halophilic traits. These enzymes can also be applied in industrial settings for flavour formation and synthesis of polyunsaturated fatty acid-enriched oils in the food industry, biodiesel production, synthesis of oleochemicals and other biotechnological processes requiring these unique characteristics (Salihu and Alam 2015).

Lipase-catalysed reactions in recent times have been conducted in organic solvents because of the need to realize the enzyme’s full potential, and because of the limitations associated with reactions performed in aqueous media such as high-enzyme-volume requirement, microbial contamination, and poor solubility of substrates (Kumar et al. 2012). Lipase biocatalytic reaction in organic solvents offer several merits such as high solubility of non-polar compounds, change of thermodynamic equilibrium towards synthetic reaction than hydrolytic one, limited aqueous-dependent undesired side reactions, no contamination by microbes, controlled change of enzyme specificity towards certain compounds, absence of racemization, reusability and ease of enzyme recovery, possible improvement of thermal stability as well as possibility of catalysing reactions regarded impossible in organic media (Karan et al. 2012). Unfortunately, many lipases are known to lose their activity under harsh organic solvent condition; hence, steps are being taken to obtain lipases that are compatible and stable in such condition. Halophilic lipases have been considered important biocatalyst in aqueous/organic or completely non-aqueous condition for varieties of industrial processes involving transesterification, hydrolysis, and ester synthesis (Fuciños et al. 2012). This is because the enzyme functions naturally under extremely high salt concentrations, that is, in low-water environments. A wide variety of fatty acid esters (FAEs) is common in cosmetic preparations that are synthesized by enzyme-catalysed esterification as well as transesterification in non-aqueous systems (Hills 2003). Such fatty acid esters known as speciality esters which are fatty acid derivatives play a major role as emulsifiers, emollients (butyl oleate, myristyl myristate, oleyl erucate, cetyl ricinoleate), detergents (butyl stearate), flavour and fragrance (butyl caprylate, butyl valerate) and thickeners in the cosmetic industry (Abdelmoez and Mustafa 2014). Nevertheless, the major limitation of lipase-catalysed ester production is their instability under harsh unfavourable reaction conditions of temperature, pH extremes and some organic solvents, and subsequently leading to lipase deactivation (Hills 2003).

Thus, in this investigation, a halophilic extracellular lipase from M. litoralis SW-45, which is highly solvent tolerant, was employed as enzyme for esterification of FFA derived from CPO and CPKO in heptane and solvent-free system, respectively. The application of crude halophilic lipase from M. litoralis SW-45 to butyl ester synthesis from renewable raw materials such as CPO and CPKO has not been investigated. Moreover, there is a dearth of studies on halophilic lipase application for the synthesis of value-added products. Thus, the notion of this investigation is to produce halophilic lipase from M. litoralis SW-45, then followed by characterisation of the enzyme based on salt, temperature and solvent tolerance, and finally to explore its performance for the synthesis of butyl esters.

Materials and methods

Chemicals and raw materials

p-Nitrophenyl esters [p-nitrophenyl acetate, p-nitrophenyl butyrate, p-nitrophenyl valerate, p-nitrophenyl octanoate, p-nitrophenyl decanoate, p-nitrophenyl laurate, p-nitrophenyl myristate and p-nitrophenyl palmitate] as lipase substrate, p-nitrophenol (p-NP), butyl ester standards (butyl hexanoate, octanoate, decanoate, laurate, and stearate), methyl heptadecanoate internal standard, triacylglycerol substrates, glyceryl trioctanoate, glyceryl tristearate, glyceryl trioleate, glyceryl trilinoleate and Thermomyces lanuginosus lipase (≥ 100,000 U/g) were bought from Sigma (St. Louis, USA). The lipase is in liquid form and was used after pretreatment with 25 mM potassium phosphate buffer to make half of its initial concentration. The organic solvents used were of analytical grade, which was procured from Merck (Darmstadt, Germany). All natural oils including olive oil, coconut oil, jatropha oil, and soybean oil were acquired from local markets except castor oil that was bought from Sigma (St. Louis, USA). CPO and CPKO with a molecular weight of 256 g/mol and 200 g/mol, respectively, were acquired from Sime Darby Plantation Sdn Bhd (Selangor, Malaysia). The Malaysian CPO fatty acid composition (wt%) is as follows: 0.2% lauric acid, 1.1% myristic acid, 44% palmitic acid, 4.5% stearic acid, 39.2% oleic acid, 10.1 linoleic acid and 0.4 linolenic acids, while Malaysian CPKO has the following composition (wt%): 0.3% caproic acid, 4.4% caprylic acid, 3.7% capric acid, 48.3% lauric acid, 15.6% myristic acid, 7.8% palmitic acid, 2.0% stearic acid, 15.1% oleic acid, 2.7% linoleic acid and 0.2% linolenic acid.

Microbial strain isolation and identification

The bacterial strain utilized in this investigation was recently isolated from seawater of Tanjung Piai National park (Latitude 1.2681°N, Longitude 103.5087°E), Johor Bahru, Malaysia. Different serial dilutions (10³–10⁶) of the seawater sample were incorporated into halophilic isolation medium and incubation was done at 37 °C for 96 h. The halophilic medium [Modified Sehgal and Gibbons complex (SGC) medium] composition (g/L) is as follows: casein peptone 7.5; yeast extract 10.0; KCl 2.0; sodium citrate 3.0; MgSO₄·7H₂O 20.0; FeSO₄·7H₂O 0.01; agar 20.0 supplemented with 7% (w/v) NaCl. The medium was set to pH 7.0 using 1 M NaOH (Anisha et al. 2012). A pure growth of the strain was kept on the aforementioned medium agar slant and kept in the cold room at 4 °C until further needed. The strain’s lipolytic potential both on a tributyrin-agar plate and in halophilic fermentation broth supplemented with olive oil as inducer has been confirmed in our previous preliminary analysis (Musa et al. 2018b).

Microbial characterization at morphological and physiological levels

The bacterial strain was characterized morphologically, biochemically and physiologically, but molecular characterization technique was used to confirm the identity of the organism as M. litoralis SW-45. Morphological characterization (shape and colour of colony) and biochemical characteristics of the strain such as Gram’s reaction, catalase, gelatine liquefaction, casein hydrolysis, starch hydrolysis, motility, citrate utilization and sugar fermentation were studied following the method of Okanlawon et al. (2010). In furtherance, micro-morphological characterization of the strain was done by viewing the cell shape in Motic-BA200 light microscope (Hong Kong, China) as well as in a scanning electron microscope (SEM). Physiological characterization such as optimum growth pH, temperature and salt (NaCl) tolerance was equally determined for the strain.

Microbial identification at molecular level

The strain was outsourced to Macrogen, Seoul, Republic of Korea, for molecular identification by conducting PCR amplification using primer sequence set “27F (5′ AGA GTT TGA TCM TGG CTC AG 3′) and 1492R (5′TAC GGY TAC CTT GTT ACG ACT T 3′)”, and performing partial 16S rRNA sequencing. The partial gene (16S rRNA) sequence of the organism was conducted by sequencing the amplified DNA templates using highly specific universal primers “785F (5′ GGA TTA GAT ACC CTG GTA 3′) and 907R (5′ CCG TCA ATT CMT TTR AGT TT 3′)”. The gene sequence has been submitted to the Gene Bank with Gi: 265678537 accession number. The strain was kept in Microbiology Culture Collection Centre of Universiti Malaysia Perlis. The strain type can also be accessed at the Culture Collection Centre with JCM 11547 as the registration number. The obtained sequences were annotated and integrated into the database using the automatic alignment tool. The 16S rRNA partial gene sequence was analysed and similarity search was conducted using the BlastN program and the information contained in the GenBank of the National Centre for Biotechnology Information (NCBI) website (www.ncbi.nih.gov). The phylogenetic and molecular analyses of the 16S rRNA nucleotide sequences were done for sequence alignments by utilization of ClustalW and MEGA5 software programs. Construction of the phylogenetic tree was done using maximum likelihood algorithm technique and the tree topology was evaluated by means of Bootstrap analysis (Yoon et al. 2003).

Production of extracellular halophilic lipase under optimized conditions

Lipase secretion by the halophilic M. litoralis SW-45 was performed in optimized halophilic culture medium (modified SGC) obtained from our previous study (Musa et al. 2018a), and with composition as follows (g/L): casein peptone 3.5; yeast extract 10.0; KCl 2.0; maltose 3.0; MgSO₄·7H₂O 10.16; FeSO₄·7H₂O 0.014; enriched with 1% (v/v) olive oil and 6.83% (w/v) NaCl. The medium pH was set to 7.0 using 1 M NaOH (Anisha et al. 2012). The strain was inoculated into 50 mL modified SGC medium contained in 250-mL Erlenmeyer flask and incubation was carried out at 37 °C for 102 h with shaking (150 rpm). The crude halophilic lipase was collected after incubation by growth broth centrifugation at 8000 rpm for 25 min at 4 °C, and the collected supernatant free of bacteria cell was utilized for halophilic lipase activity assay.

Assay of halophilic lipase activity

p-Nitrophenyl laurate (p-NPL) is a specific substrate for the halophilic lipase, MLL, because of its preference for short-to-medium-sized acyl chain length. In the studies involving determination of lipase fatty acid specificity, the correct reaction conditions and substrates need to be utilized to confirm that the changes encountered during the reaction are due to structural alteration of the fatty acid group and not as a result of other unknown effects (Macrae and Hammond 1985). Activity assay of the halophilic lipase was assessed using p-NPL according to Gutarra et al. (2009). The assay was done by the introduction of halophilic lipase (0.05 mL) to a 25 mM potassium phosphate buffer (2.2 mL) solution (pH 7.0) and 2.5 mM p-NPL (0.25 mL). The p-NPL degradation reaction was performed at 50 °C for 20 min, then 0.25 mL of 0.1 M Na₂CO₃ was included to terminate the assay, and the activity was monitored at 412 nm using a UV–Vis spectrophotometer (Shimadzu, Japan). One unit of lipase activity by the producing strain is defined as the quantity of enzyme that releases p-nitrophenol (1 mol) per min from p-NPL substrate in 20 min using the assay conditions.

Halophilic lipase characterization

Influence of salt and temperature on MLL activity

To evaluate the simultaneous influence of NaCl and temperature on MLL activity, experiments were conducted by measuring activity in assay solution containing 100 mM potassium phosphate buffer (pH 8.0) and different NaCl concentrations (0–30% w/v) at 3% interval (Samaei-Nouroozi et al. 2015). The assay mixture was then incubated at a temperature range of 30–70 °C (10 °C intervals) for 20 min using p-NPL standard assay method as described earlier.

Determination of MLL activity in organic solvents at different salt concentrations

The MLL activity in selected organic solvents (acetonitrile, sec-butanol, diethyl ether, heptane, hexane and toluene) at different salt (NaCl and KCl) concentrations was assessed by pre-incubating the halophilic lipase mixture in buffer (100 mM at pH 8.0) with varying salt concentrations (0–30% w/v) and selected organic solvents (to make 75% (v/v) concentration (Alsafadi and Paradisi 2013). The reaction was incubated at 37 °C for 1 h in a rotary shaker (Sartorius, Germany) at 150 rpm. Sampling was done by withdrawing from aqueous phase after the incubation period, and residual activity of the lipase was assessed following standard assay conditions. Lipase pre-incubated without salt but with selected organic solvents was considered as control.

Hydrolysis of CPO and CPKO using Thermomyces lanuginosus lipase (TLL)

Hydrolysis reaction of CPO and CPKO was performed in 125-mL stoppered conical flasks that were initially filled with 1.5 g of CPO and CPKO and 15 mL diethyl ether and heptane solvent, respectively. A 15 mL buffer solution (25 mM, pH 7.0) was introduced into individual flasks so that the oil to buffer ratio was 1:1, wherein a two-layered mixture was formed (Serri et al. 2008). Hydrolysis reaction was initiated by adding 5% (v/v) TLL to each of the flasks and continuously agitated at 50 °C and at 150 rpm for 1 h. The objective of using TLL was to attain sufficient hydrolysis level so as to obtain FFA in considerable amount for esterification to butyl esters in the follow-up step using MLL as a biocatalyst. The stoppered conical flasks with the reaction mixture but in the absence of biocatalyst were considered as the control flask. Samples (1 g) were removed from the organic solvent–oil phase of the mixtures using a micropipette, and then transferred into 125-mL conical flasks for analysis (that is, initial acid value determination). Experiments were conducted in triplicate. Ten millilitres of ethanol:diethyl ether (1:1, v/v) was introduced into the oil phase sample to inactivate the lipase, thereby completely terminating the reaction. The released FFAs in the mixture were titrated with 0.1 M NaOH using 25 µL indicator (phenolphthalein), which was prepared by dissolving 0.2% in ethanol. The rate of hydrolysis (%) was calculated using the following formula (Bressani et al. 2015):

$$\text{Hydrolysis}\left(\%\right)=\left(\frac{v\times M\times {\text{MM}}_{\text{FA}}}{W_{\text{t}}\times f}\right)\times 100,$$ 1

where v equals NaOH volume needed for titration (mL), M equals concentration (0.1 M) of NaOH solution, MM_FA is the average molar mass of FFA derived from CPO (256 g/mol) and CPKO (200 g/mol), w_t equals mass of the sample withdrawn (g) and f is the mass ratio of oil to buffer at the beginning of hydrolysis (% m/m).

Several hydrolysis batches of CPO and CPKO reaction mixtures sufficient enough for esterification step were prepared, and the FFAs formed in the reaction mixture were concentrated and purified.

Concentration of FFAs in the reaction mixture

Fifteen millilitres each of diethyl ether and heptane solvents was added to CPO and CPKO reaction mixture, respectively, upon complete hydrolysis of the oil substrates. The mixtures obtained thereof were subjected to centrifugation at 5000 rpm and at 25 °C initially for 15 min, and the superior phases containing organic solvents and FFAs were differentiated from the non-solvent phase (lipase, glycerol and buffer solution) below. The aqueous phases were centrifuged again for 5 min to differentiate the remaining superior phase from the non-solvent phase. Anhydrous Na₂SO₄ was then added to the FFA–solvent phase to dehydrate residual water remaining in the superior solvent phase. The organic solvent phase containing the FFAs was vacuum filtered using Buchner funnel, and the respective solvents (diethyl ether and heptane) were removed by a rotary evaporator (Chowdhury et al. 2013).

Butyl ester synthesis by esterification reaction using MLL as biocatalyst

Esterification of the purified FFAs obtained from several hydrolysis batch experiments of the two oil substrates was performed in 125-mL stoppered conical flasks containing equimolar butanol to the FFA molar ratio of 1:1 dissolved in heptane medium. Esterification was initiated by the addition of 3 mL (30% v/v) MLL to the reaction medium (10 mL) contained in individual conical flasks. The CPO and CPKO reaction flasks were both incubated at 45 °C for a period of 1 h with rigorous stirring on a shaking incubator at 200 rpm. Control experiment performed was similar to the above esterification mixture but without enzyme. Esterification was done using the methodology of Bressani et al. (2015) with little changes. Ester yield was evaluated by determining the concentration of the remaining fatty acids in the esterification medium using titration method (Silva et al. 2014). Purified FFA was taken (1 mL) and dissolved in 10 mL ethanol/acetone equimolar ratio mixture and titrated. All reactions were conducted in triplicates. The percentage ester conversion of FFAs based on the quantity of fatty acid utilized was estimated using the following formula (Silva et al. 2014):

$$\text{Ester conversion}\left(\%\right)=\left(\frac{\text{Avo}-\text{Avt}}{\text{Avo}}\right)\times 100,$$ 2

where Avo is the initial acid value of the solvent phase sample withdrawn from the reaction medium and Avt is the acid value of the solvent phase sample at time t.

Optimization of enzymatic esterification of FFA by OFAT approach

The OFAT approach was utilized to analyse the impact of reaction conditions on butyl ester yield. The esterification variables studied include temperature, agitation speed, enzyme concentration, butanol to fatty acid molar ratio, NaCl concentration, and reaction time. The reaction temperature was differed from 30 to 55 °C at 5 °C interval and the influence of agitation speed was investigated in the range of 150–250 rpm. Enzyme concentration was varied from 10 to 60% (v/v), while butanol to FFA molar ratio was studied from 1:1 to 5:1. The influence of various NaCl concentrations was also determined in the range of 0–18% (w/v). For the time course study (influence of reaction time), 1 mL of the CPO and CPKO esterification medium was taken at varying time intervals and ester conversion was measured by means of titration. At some point, the type and amount of butyl esters formed in the reaction medium were quantified by gas chromatography (GC) analysis.

Comparison of enzymatic esterification of CPKO-derived FFA catalysed by commercial lipase (TLL) and halophilic lipase (MLL)

This experiment was conducted to compare the effectiveness of halophilic lipase (MLL) with a commercial (TLL) lipase for application in esterification of CPKO-derived FFA. Esterification reaction catalysed by halophilic lipase was performed under optimal conditions of 45 °C temperature, 230 rpm agitation speed, 50% (v/v) biocatalyst concentration, 5:1 butanol:FFA molar ratio and 15% (w/v) NaCl for an optimum reaction time of 15 min. The reaction catalysed by the commercial lipase was conducted under similar reaction condition as the halophilic enzyme. As described earlier, esterification reactions were conducted in 10 mL reaction medium contained in 125-mL stoppered conical flasks, which were all triplicated. A control experiment similar to the aforementioned reaction conditions was conducted with the exception of a biocatalyst. Quantification of butyl esters formed in the reaction medium was done by titrimetric and confirmed by gas chromatographic methods.

GC analysis of the synthesized butyl esters

At some point, the percentage ester conversion was confirmed by gas chromatography. Butyl esters from CPO and CPKO were analysed by utilizing gas chromatography, GC-2010 plus model (Shimadzu, Tokyo, Japan), equipped with a flame ionization detector (FID) and a BPX-70 column (30 m × 0.32 mm × 0.25 µm). N₂ was utilized as the carrier gas with a flow rate of 25 mL/min. The temperature of the injector which was in a splitless mode and that of the detector was fixed at 250 °C. GC programming of temperature was done as follows: column temperature was maintained at 90 °C for 3 min, then increased up to 120 °C at 25 °C/min and maintained constantly for 10 min. The temperature was then set at 25 °C/min to 170 °C and maintained constantly for 15 min. 0.1% stock solutions of butyl ester standards (butyl hexanoate, butyl octanoate, butyl decanoate, butyl laurate and butyl stearate) were prepared in heptane, while 10 mg/mL of methyl heptadecanoate internal standard was prepared in heptane as stock solution. For qualitative analysis, GC samples were prepared in heptane at 1% (v/v), and filtered using a filter of 0.45 µm pore size before injection into GC machine. On the other hand, quantitative estimation involved taking 20 µL of the samples and diluting in 480 µL of the methyl heptadecanoate internal standard (IS) solution. Approximately 1 µL of the prepared samples and the standard mixture was then injected into a splitless mode. The ester contents were measured using the peak area of methyl heptadecanoate according to the following equation (Gasparini et al. 2011):

$$\text{Ester}\left(\%\right)=\frac{\sum A-\text{AC17IS}}{\text{AC17IS}}+\frac{\left(\text{CEI}\times \text{VEI}\right)}{m}\times 100,$$ 3

where ƩA is the sum of areas of the butyl ester peaks, AC17IS is the internal standard peak area, CEI equals concentration of methyl heptadecanoate solution (mg/mL), VEI equals methyl heptadecanoate volume mixed with the sample, and m is the mass of the sample (mg).

Results

Bacterial strain characterization and molecular identification

The bacterial strain is Gram negative, aerobic and rod shaped in nature. Colonies are creamy white, irregular in shape, opaque, and mucoid in consistency on modified SGC agar plate as seen in Fig. 1a. The scanning electron microscopy of the halophilic bacteria cells occurring singly, as chains and in clusters is also indicated in the figure. Catalase and citrate utilization tests by the strain were found positive, while motility, casein hydrolysis, starch hydrolysis, and gelatin liquefaction were all negative. Acid was produced from fructose but showed no fermentation in sucrose, lactose, maltose, myo-inositol, galactose, cellobiose and cellulose. Physiological characterization showed that the strain is capable of exhibiting growth with or without NaCl. It showed growth in media composed of 0–15% (w/v) NaCl but optimal growth was found at 7% (w/v) NaCl. This behaviour is quite typical of marine bacteria. Thus, the organism could be regarded as a moderately halophilic bacterium (Kushner 1993). Optimum strain growth was noticed at 37–40 °C whereas the highest strain growth was found to be at 45 °C. A neutral to alkaline pH range of 6.0–8.5 supported maximum growth of the strain. An organism with similar morphological and physiological traits was earlier isolated from seawater in South Korea by (Yoon et al. 2003), whereby the name M. litoralis SW-45 was proposed.

Fig. 1.

a Representation of halophilic Marinobacter. Culture plate and SEM photographs of M. litoralis SW-45 grown on modified SGC agar (7% w/v NaCl) at 37 °C for 48 h are shown. The arrows on the SEM micrograph indicate chains and cluster formation. b Phylogenetic relationship of a moderate halophilic M. litoralis SW-45 isolated from seashores of Tanjung Piai National Park, Johor Bahru, Malaysia, is also displayed. The phylogenetic topology of the 16S rRNA partial gene sequence was constructed using the neighbour-joining method. The lipase-producing strain is indicated

Phylogenetic evaluation of the 16S rRNA partial gene sequence comparison (Fig. 1b) showed that the strain is categorized in the genus Marinobacter. The analysis also revealed 99% gene sequence similarity with M. litoralis SW-45. Thus, the strain was tentatively named as M. litoralis SW-45 and its gene sequence has been kept in the Gene Bank, and assigned accession number Gi: 265678537.

Halophilic lipase characterization

Influence of salt temperature on MLL activity

In our previous study, it was quite evident that salt concentration had immense impact on the activity of MLL. As a follow-up to the previous work, the combined effect of NaCl and temperature on MLL was performed and the result is shown in Fig. 2. As NaCl concentration increases from 0 to 12% (w/v), the activity of the halophilic lipase was improved at all the temperature ranges (30–70 °C) investigated. The activity of the enzyme gradually increases by increasing salt concentration, but the enzyme became thermally deactivated at high salt concentrations beyond which the enzyme can tolerate. This phenomenon was more glaring at an elevated temperature range of 60–70 °C, in which the enzyme appeared to be completely inactivated but became reactivated upon introducing 9 and 12% (w/v) NaCl. The points in the figure displayed by the arrows indicate the position of halophilic lipase enhancement owing to the introduction of optimal NaCl concentrations required by the enzyme at respective temperatures.

Fig. 2.

Temperature effect on the activity of MLL at different NaCl concentration. The arrow shown in the diagram indicates the point of ‘reversible renaturation’ phenomenon that occurred upon introduction of 12% (w/v) NaCl

Activity of MLL in organic solvent systems at different salt concentrations

The result of the investigation is depicted in Fig. 3. It can be deduced that the enzyme activity in the individual aqueous–organic solvent system rises by increasing NaCl and KCl concentration. However, the activity later declined gradually at higher salt concentrations in the organic solvents studied. The suboptimal NaCl requirements of MLL in 75% (v/v) acetonitrile, 1-butanol, heptane, diethyl ether, hexane, and toluene solvent systems are 18%, 18%, 15%, 12%, 18%, and 9% (w/v), respectively (Fig. 3a). In Fig. 3b, a similar trend of salt-induced enzyme activity in the organic solvent system was observed for KCl, but MLL exhibits a different behaviour in hexane solvent system by responding negatively to increased KCl concentration; a decline in enzyme activity was noticed as the concentration of KCl increases from 3 to 30% (w/v). The different enzymatic performance observed in hexane solvent system compared with other solvents in the presence of increased KCl concentration may be due to a combination of several factors such as the influence of solvent water content and polarity, the active site polarity, and the kosmotropicity of the activating salt (Ru et al. 2001).

Fig. 3.

Organic solvent (acetonitrile, 1-butanol, heptane, diethyl ether, hexane and toluene) effect on the activity of MLL at different [0, 3, 6, 9, 12, 15, 18, 21, 24, 27 and 30% (w/v)] NaCl (a) and KCl (b) concentrations

Enzymatic hydrolysis of CPO and CPKO using TLL

The synthesis of FFA from CPO and CPKO using commercial TLL was conducted. The choice of this biocatalyst was based on the knowledge that the enzyme has high catalytic activity, and its efficiency in catalysing hydrolysis reaction in the organic solvent system. The enzyme-catalysed hydrolysis of CPO and CPKO was obtained under the following reaction conditions: 1.5 g of oil samples in 15 mL diethyl ether and heptane solvent, oil to buffer ratio is 1:1, 5% (v/v) enzyme, 50 °C, 150 rpm agitation speed and reaction time of 1 h. A high FFA conversion of 596.5% and 412% was obtained from CPO and CPKO substrates, respectively. Cavalcanti-Oliveira et al. (2011) also reported high catalytic activity of the biocatalyst when it was used to catalyse the hydrolysis of soybean oil. Since high hydrolysis rate was obtained for both CPO and CPKO, and also for the fact that the specific aim of this study was to investigate esterification using our crude halophilic MLL, the hydrolysis step was not studied in detail. The yield of FFA sufficient enough for esterification step was achieved by conducting several hydrolysis batches.

Optimization of enzymatic esterification of FFA by OFAT using MLL

Effect of temperature on enzymatic esterification

To assess the influence of temperature on the biosynthesis of butyl esters from CPO- and CPKO-derived FFA, esterification was done at a different temperature range of 30–55 °C, at fixed operating conditions. Butyl ester yields at a different temperature are presented in Fig. 4a. It could be observed that increasing reaction temperature resulted in a concomitant increase in butyl ester synthesis. The optimal working temperature of MLL for butyl ester synthesis from CPO- and CPKO-derived FFA is 40 °C and 45 °C, respectively. This result verifies that good ester yields are attainable at relatively low temperatures.

Fig. 4.

The effect of a varying reaction temperature (30–55 °C), b varying agitation speed (150–250 rpm), c varying enzyme concentration (10–60% v/v), d butanol:FFA molar ratio (1:1–5:1), and e varying NaCl concentration (0–18% w/v) on esterification of CPO- and CPKO-derived FFA using OFAT approach

Influence of agitation speed on enzymatic esterification

The result of the effect of agitation speed on esterification is depicted in Fig. 4b. It is quite obvious here that increasing the agitation speed beyond 150 rpm slightly decreased ester conversion of CPO-derived FFA, and a further increase in agitation speed had no significant effect on ester conversion. Thus, 150 rpm was chosen as the optimum agitation speed for the subsequent experiment. This shows that ester conversion was independent of agitation speed beyond 150 rpm, thereby indicating that the effect of external mass transfer was negligible. On the other hand, ester conversion percentage of CPKO-derived FFA increased slightly as agitation speed was increased. Increase in agitation speed from 150 to 230 rpm improved ester yield from 58.6 to 63.1%. Here, agitation speed of 230 rpm was considered optimum and was used for further esterification study.

Effect of enzyme concentration on esterification

Figure 4c depicts the influence of various enzyme concentrations on ester yield. As expected, the ester conversion percentage in both CPO- and CPKO-derived FFA reaction system increased steadily with the increase in enzyme concentration from 10 to 50% (v/v). But at higher enzyme concentration, that is, above 50% (v/v), butyl ester yield tends to fall abruptly. Maximal ester yield for both CPO- and CPKO-derived FFA esterification systems was achieved with 50% (v/v) MLL and was desired for further parameters’ study.

Influence of butanol:FFA molar ratio on enzymatic esterification

To assess the effect of butanol:FFA ratio on butyl ester (from CPO-derived FFA) yield, the reaction was conducted by varying substrate (butanol:FFA) molar ratio from 1:1 to 5:1. The reaction temperature, agitation speed, and enzyme concentration set at an optimum level were employed for 1 h reaction time. The results of the influence of butanol:FFA molar ratio on enzymatic esterification are presented in Fig. 4d. From the CPO-derived FFA esterification system, lower ester conversion was observed by an increment in substrate molar ratio, while higher ester conversion of 60.5% in 1 h was recorded for smaller substrate molar ratio. For the CPKO-derived FFA (Fig. 4d), the influence of butanol:FFA molar ratio on butyl ester conversion was determined by performing esterification reaction at five different ratios (1:1 to 5:1) while temperature was maintained at 45 °C, 230 rpm agitation speed and enzyme concentration at 50% (v/v) was kept fixed. Here, a gradual increment in ester conversion percentage with an increase in butanol:FFA molar ratio was observed. Maximum ester conversion of 68.4% was attained at butanol:FFA molar ratio of 5:1 in 1 h. Further increase in the substrate ratio from 5:1 to 6:1 (68.6%) did not produce any significant increase in ester yield, hence, 5:1 was desired as the optimal substrate ratio. Subsequent OFAT studies concerning butyl esters were performed at 1:1 butanol:FFA molar ratio, 40 °C, 150 rpm and 50% (v/v) enzyme concentration for CPO-derived FFA esterification, and 5:1 butanol:FFA molar ratio, 45 °C, 230 rpm and 50% (v/v) enzyme concentration for CPKO-derived FFA esterification reaction system. It can be said from this point onwards that the CPKO-derived FFA esterification has been performed in a solvent-free system since heptane was excluded at this stage due to fivefold increase of the acyl acceptor (butanol) added to the reaction medium.

Influence of various NaCl concentrations on enzymatic esterification

In this investigation, the influence of various NaCl concentrations [0–18% (w/v)] on butyl ester synthesis from both CPO- and CPKO-derived FFA was analysed, while keeping other optimized parameters fixed as shown in Fig. 4e. It was found in the CPO-derived FFA esterification system that increasing NaCl concentration from 0 to 9% (w/v) resulted in a simultaneous increase in ester conversion percentage until the maximal value of 62% was attained. This result is not surprising as it was earlier observed in this study that salt significantly improved the activity of MLL in various solvents investigated, which was attributed to right folding of proteins in organic solvents and subsequently, maintenance of enzyme activity and stability. Similarly, in the CPKO-derived esterification reaction system, butyl ester conversion percentage increased gradually with the increase of NaCl concentration from 0 to 15%, reaching a maximum of 69%.

Effect of reaction time on enzymatic esterification/time course study

The time needed to attain highest butyl ester yield for both esterification systems under optimum experimental conditions was determined by the periodic withdrawal of butyl ester samples and quantification using both titrimetric and gas chromatographic methods. Under optimal conditions of reaction temperatures 40 and 45 °C, agitation speed 150 and 230 rpm, enzyme concentration 50% (v/v), 1:1 and 1:5 butanol:FFA molar ratio, NaCl concentration 9% and 15% (w/v) for CPO- and CPKO-derived FFA esterification system, respectively, esterification of CPO- and CPKO-derived FFA was performed for a maximum reaction time of 150 min. The time course profiles of butyl esters production are depicted in Fig. 5a. For the CPO-derived FFA esterification system, butyl ester conversion showed a sharp increase to reach maximum conversion percentage of around 62.2% after 60 min of reaction, before showing a drastic decline in ester yield. While for the CPKO-derived FFA esterification system, ester conversion also showed a drastic increase to 69.1% until 15 min, and remained almost constant from 15 min onwards. The optimum esterification reaction time for this study was chosen as 60 and 15 min for CPO- and CPKO-derived FFA esterification system, respectively. These short reaction times required to attain a considerably high ester conversion percentage could translate to an economic advantage in the industrial production process.

Fig. 5.

a Time course profiles of butyl ester synthesis catalysed by crude halophilic MLL. The reactions were conducted at 40 °C and 45 °C, 150 rpm and 230 rpm agitation speed, 50% (v/v) enzyme concentration, 1:1 and 5:1 butanol:FFA, 9% and 15% (w/v) NaCl optimum conditions for purified CPO- and CPKO-derived FFA in heptane and solvent-free reaction systems, respectively. b Enzymatic esterification of purified CPKO-derived FFA in a solvent-free system using non-halophilic commercial lipase (TLL) and crude halophilic lipase (MLL), respectively, as a biocatalyst. Esterification reactions catalysed by TLL and MLL were performed under optimal conditions of 45 °C reaction temperature, 50% (v/v) biocatalyst concentration, butanol:FFA of 5:1 and 15% (w/v) NaCl in a solvent-free system. 1—MLL (halophilic lipase); 2—TLL (commercial lipase) with 15% NaCl; 3—control (without enzyme). c Gas chromatogram of (i) butyl ester standards (1—butyl hexanoate; 2—butyl octanoate; 3—butyl decanoate; 4—butyl laurate; 5—butyl stearate), (ii) butyl esters derived from crude palm oil (1—butyl laurate; 2—butyl stearate) and (iii) butyl esters derived from crude palm kernel oil (1—butyl hexanoate; 2—butyl laurate)

Comparison of enzymatic esterification of CPKO-derived FFA catalysed by normal (commercial) lipase and halophilic lipase

The specific objective of this experiment is to compare and assess the effectiveness of halophilic lipase (MLL) and normal, commercial lipase (TLL) in the esterification of CPKO-derived FFA conducted in a solvent-free condition and heptane medium, respectively. As seen in Fig. 5b, maximum butyl ester conversion percentage of around 69.1% for MLL-catalysed esterification was reached after a reasonably short period of 15 min. On the other hand, for TLL-catalysed esterification with 15% (w/v) NaCl, ester conversion by commercial lipase was only able to reach a maximum conversion of 69% after 150 min. It can be seen that ester conversion percentage by the commercial enzyme in the occurrence of 15% NaCl was slightly lower compared to that catalysed by the halophilic enzyme. This shows that the commercial lipase is non-halophilic, and as such inactivated in the presence of salt by stripping of hydration water surrounding the enzyme (Sinha and Khare 2014), thus culminating in the reduction of percentage ester conversion. It can, therefore, be inferred that our indigenous halophilic MLL exhibited superior esterification capability under saline esterification reaction condition when compared with the commercial lipase (TLL).

Butyl ester identification

The gas chromatographic separations of butyl ester standard mixtures (butyl hexanoate, butyl octanoate, butyl decanoate, butyl laurate and butyl stearate) and those obtained from free fatty acids of crude CPO and CPKO are depicted in Fig. 5c. Comparison of the elution times between standard butyl esters [Fig. 5c(i)] and butyl esters derived from CPO [Fig. 5c(ii)] and CPKO [Fig. 5c(iii)] indicates that butyl laurate and butyl stearate were identified from crude palm oil, while butyl hexanoate and butyl laurate were identified from crude palm kernel oil, respectively. The unidentified peaks in the chromatogram of the CPO- and CPKO-derived butyl esters could be other palm-based butyl esters that were not detected due to unavailability of the respective standards as at the time of conducting this experiment.

Discussion

Halophilic lipase characterization for salt, temperature and organic solvent tolerance

Thermal stability of halophilic enzymes and proteins was earlier found to be controlled by the availability of salt (Sinha and Khare 2014). In this study, the reversible renaturation of the enzyme whose activity suddenly reappears (upon introduction of saline condition) after it was initially lost to thermal denaturation occurred. Similar ‘reversible renaturation’ phenomenon induced by salt (0.2 M) was documented when β-lactamase from moderately halophilic Chromobacter sp. maintained 75% of its activity after exposure to 100 °C for 5 min (Tokunaga et al. 2004). In another study by Gunny et al. (2014), higher thermal stability was imparted on the salt-tolerant cellulase from Aspergillus terreus UniMAP AA-6 when 3 M NaCl was introduced to the reaction. The ‘Irreversible aggregation’ of the halophilic enzyme in the current study may have probably been averted due to a combination of factors such as the availability of acidic amino acid residues on the enzyme-protein structure, helix stabilization, improved core packing, and increased ionic interactions (Sinha and Khare 2013). Haki and Rakshit (2003) also reported that certain specialized proteins referred to as ‘chaperonins’ which are synthesized by some extremophilic organisms especially thermopiles help in refolding enzymes into their native form and restore their catalytic functions after denaturation.

Since salt concentration was found to influence the thermal stability of MLL, the organic solvent tolerance of the enzyme at different salt (NaCl and KCl) concentrations was equally investigated (Figs. S1–S3). The gradual decline of the enzyme activity after initial improvement could be attributed to enzyme precipitation or denaturation as a result of a further increase in surface tension of hydration water caused by the presence of higher salt concentration (Danson and Hough 1998). Studies of organic solvent tolerance when high salt concentrations are present offer the opportunity to identify novel enzymes that can thrive in harsh unfavourable operating conditions (Stepankova et al. 2013). High salt concentration significantly reduces water activity (Sinha and Khare 2014). Halophilic enzymes are known to uniquely perform in low-water/non-aqueous media (Kumar and Khare 2012). The special acidic amino acid (mainly aspartate and glutamate) compositional properties of halophilic enzymes and proteins are the major reasons for their increased solubility in aqueous suspensions having elevated salt concentration, as well as their tolerance to organic solvents. Moreover, inorganic salts such as NaCl and KCl are salting-out agents that increase the surface tension of hydration water layer formed around the enzyme structure based on their position in Hofmeister series (Amoozegar et al. 2017). This salt-induced increase in surface tension of hydration water layer around the enzyme promotes the formation of compact enzyme structure, thereby maintaining its activity and stability (Danson and Hough 1998). Overall, the fact that the enzyme (MLL) retained 100% of its activity in most of the organic solvent tested at 75% (v/v) concentration, which notably improved in the presence of salt, shows that the enzyme is highly solvent tolerant and stable. This particular feature is of immense biotechnological significance and makes the enzyme highly attractive to industrial application.

Optimization of enzymatic esterification of FFA by OFAT using MLL

Temperature significantly impacts enzyme-catalysed esterification. In the current work, butyl ester synthesis by esterification of the concentrated FFA obtained via enzymatic hydrolysis of CPO and CPKO and butanol was performed at fixed conditions: equimolar ratio of butanol:FFA, enzyme concentration of 30% (v/v) in a shaker operating at 200 rpm for the highest time of 1 h. The temperature was differed at the range of 30–55 °C. Since esterification reaction is said to be endothermic, the rise in temperature usually results in an increase in final ester conversion (Marchetti and Errazu 2008). This may be attributed to the fact that high temperature increases the effective collisions (kinetic energy) of the reacting species in the system, thereby leading to improved interactions between the enzyme molecules and substrates, hence boosting the reaction rate (Chowdhury et al. 2014). Temperature rise usually lessens system viscosity, improves substrate solubility and enhances substrate diffusion process, thereby causing a reduction in mass transfer limitations and favouring yield (Ceni et al. 2010). A steady decline in butyl ester production was noticed when the optimum temperature of the enzyme-catalysed reaction was exceeded, which can be attributed to biocatalyst denaturation (Miranda et al. 2014). The utilization of moderate temperature (40 °C and 45 °C) translates to a reduction in energy utilization and prolongation of working stability of the enzyme, which are the relevant traits for industrial application of esterification process (Kleinaitė et al. 2014).

The impact of agitation on mass transfer was evaluated by agitation speed variation from 150 to 250 rpm while maintaining other reaction conditions [30% (v/v) enzyme concentration, butanol:FFA 1:1, 40 °C and 45 °C for CPO- and CPKO-derived FFA esterification] constant. In this study, the effect of increased agitation speed leads to reduction of droplet size, caused by the considerable increase in specific interfacial area between the substrates and the biocatalyst in the organic solvent phase of the reaction system (Basri et al. 2013). Usually, higher agitation speed enhances effective number of collisions between reacting species, thereby resulting in improved interaction of enzyme and substrate (Basri et al. 2013). Ester yield and productivity decreased steadily at 250 rpm in the CPKO-derived FFA esterification system. This might be attributed to unfavourable shear rate caused by more agitation speed. Normally, when the agitation speed goes beyond the optimal level, reaction yield usually declines because of the shearing effect on the enzyme (Yadav and Lathi 2005).

The enzymatic esterification is hugely reliant on the quantity of biocatalyst. The influence of enzyme concentration on butyl ester synthesis was evaluated with butanol:FFA of 1:1 at 40 °C and 45 °C, 150 and 230 rpm for CPO- and CPKO-derived FFA, respectively, while varying the enzyme concentration from 10 to 60% (v/v) at 10% intervals. The increase in ester conversion with a high amount of MLL is probably as a result of more enzyme active sites available (Talukder et al. 2010). While the decline in ester conversion at higher enzyme concentration might be due to the diffusional limitation or bad dispersion of enzyme in the esterification medium which could not significantly improve any further conversion (Miranda et al. 2014). At higher enzyme concentration, the interfacial area is totally saturated with enzyme molecules, thereby forming aggregates and preventing movement of the substrates to the reactive site of the biocatalyst (Richetti et al. 2010). Therefore, any further rise in enzyme level in the esterification system would not improve the reaction rate.

It is a well-established fact that one of the most significant factors influencing enzymatic esterification process optimization is substrate (butanol:FFA) molar ratio. Sometimes, an enhancement in the quantity of a reactant particularly alcohol can move the equilibrium reaction to the right thereby leading to higher conversion (Richetti et al. 2010). However, the decline in ester conversion percentage experienced in this investigation could be due to increased viscosity of the esterification medium which can hinder the flow of reacting substrates from bulk reaction to the lipase catalytic sites (Friedrich et al. 2013). Another plausible reason for the decrease in ester conversion for the CPO-derived FFA esterification system is inhibition by excess butanol (Ilmi et al. 2018). Since butanol is the major reaction component at the high molar ratio, it may be gathered in the enzyme’s aqueous microenvironment due to its polar nature, thereby approaching a level sufficient to cause inactivation and modification of the protein, and subsequently inhibition (Trubiano et al. 2007). Excessive levels of alcohol usually cause enzyme inactivation by increasing medium polarity, which in turn reduces stabilization and enzyme’s hydration water layer (Ribeiro et al. 2011). Similar enzyme inhibition caused by alcohol accumulation has also been found by some authors (Kleinaitė et al. 2014; Silva et al. 2014) recently.

Extracellular halophilic lipases with elevated solvent tolerance are immensely beneficial to biotechnological processes where the simultaneous occurrence of elevated salt levels and non-polar organic solvents is common (Stepankova et al. 2013). Evidently, the availability of salt is a key prerequisite for optimum catalysis of the halophilic enzyme (Sinha and Khare 2014). NaCl and KCl always increase halophilic enzymes stability although some of them lower their catalytic ability at increased salt concentration (Ortega et al. 2011). It was noticed in this study that above the optimal NaCl concentrations of the enzyme in both reaction systems, an inhibition by NaCl occurred. Kosmotropic or salting-out salts such as NaCl and KCl improves protein stability, but at elevated levels, they rather favour precipitation (Amoozegar et al. 2017), which eventually leads to enzyme inactivation. Although quite a number of studies (Müller-Santos et al. 2009; Karan and Khare 2011; Ortega et al. 2011; Xue et al. 2014) have investigated the function of salt in modifying structures and regulating functions of moderate and extreme halophilic enzymes, to our knowledge, investigation on the influence of NaCl concentration on butyl ester synthesis from CPO and CPKO in relation to the biocatalytic function of the moderately halophilic lipase from M. litoralis SW-45 has never been performed.

Enhanced performance of the enzyme at high salt (15% w/v NaCl) concentration could be correlated to favourable interaction of the salt with the solvent (Kim and Dordick 1997). Halophilic proteins generally need salting-out medium to stabilize their poorly hydrophobic core. The salting-out nature of organic solvents prevents the poorly hydrophobic core of the halophilic enzyme from becoming favourably solvated by the medium, hence gives rise to a compact core and enhanced hydrophobicity which stabilizes the enzyme structure (Kim and Dordick 1997).

Butyl ester identification

The butyl esters synthesized were identified using gas chromatography by comparing the esters with known butyl ester standards. Gas chromatography is widely used for quantitative and qualitative analyses of compound mixture, and identification of reaction products and unknown volatile organic compounds (Al-Rubaye et al. 2017). Quantitative estimation of the butyl esters synthesized was carried out using methyl heptadecanoate as an IS with a retention time of 22.554 min. The percentage ester conversion calculated by both titrimetric method which is based on acid consumption and GC analysis was found to be in reasonable agreement. For the qualitative analysis, the butyl ester standards used were butyl hexanoate, butyl octanoate, butyl decanoate, butyl laurate and butyl stearate with the retention times of 5.24, 7.76, 12.69, 17.07 and 24.43 min, respectively. The butyl esters obtained in this study possess broad range of applications in cosmetic formulations as emollient esters (that is, pure oil for skin care), thickening and re-fatting agents, solubilisers, emulsifiers and multifunctional additives (Veit 2004). A number of butyl esters are utilized also as flavour and fragrance additives in the cosmetic industry for personal care purposes. Butyl hexanoate, for instance, is used in berry and fruity flavours as well as in fragrance additive, while butyl laurate holds fragrance and additive application in apple, orange, cape gooseberry, papaya and malt whiskey formulations (Green Biologics 2019). Butyl laurate is also extensively used as PVC plasticizer, diesel additive, textile wetting agent, and oiling agent in the leather industry.

In conclusion, a high percentage of butyl ester conversion of 62.2% and 69.1% from CPO- and CPKO-derived FFAs under optimum conditions of 40 °C and 45 °C, 150 rpm and 230 rpm, 50% (v/v) biocatalyst concentration, 1:1 and 5:1 butanol:FFA, 9% and 15% (w/v) NaCl concentration, 60 and 15 min reaction time, respectively, was achieved. The investigated physicochemical parameters substantially influenced butyl ester synthesis. One significant accomplishment of this investigation is that maximum ester conversion under minimum reaction time using the halophilic lipase was attained by the addition of salt, which improved the overall performance of the enzyme during esterification when compared with the commercial enzyme-catalysed reaction. This study has proven the applicability of the halophilic lipase as biocatalyst for the synthesis of environmentally benign value-added products, namely butyl hexanoate and butyl laurate that are of immense importance to the cosmetic industry.

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Conflict of interest

The authors declare that they have no conflict of interest.

Ethical approval

The current research work did not involve either of human or animal studies.