Immobilized Lipases—A Versatile Industrial Tool for Catalyzing Transesterification of Phytosterols Solubilized in Plant Oils to Produce Their Fatty Acid Esters

Abstract

The conjugation of phytosterols (PSs) with fatty acids results in producing phytosterol esters (PSEs) characterized by enhanced lipophilicity and improved functional properties of major interest in food and nutraceutical applications. The use of immobilized lipases to catalyze direct transesterification reactions between PSs and plant oils to form PSEs as a green alternative to conventional chemical production methods has attracted interest during the last two decades. The low solubility of PSs in common plant oil triglycerides, typically below 3% at ambient temperatures, remains the main challenge for bringing lipase-catalyzed direct transesterification reactions of PSs and oil triglycerides to commercial scales. This study focuses on the enzymatic synthesis of PSEs starting from solubilized PSs at concentrations of up to 30% wt./wt. of oil mixtures comprising fatty acid ethyl esters (FAEEs), monoglycerides (MGs), diglycerides (DGs), and triglycerides (TGs) as a homogeneous medium for the direct transesterification reaction. The results of this study show for the first time that the addition of FAEEs into the reaction medium results in an alteration of the substrate preference of the enzyme, making MGs the favorite fatty acyl group donors for PSs amongst all other fatty acyl donors present in the reaction system. The proposed new enzymatic route allows starting with high concentrations of solubilized PSs, making the direct transesterification of oil glycerides attractive for the production of PSEs at industrial scales.

1. Introduction

PSs are naturally occurring in plants, typically found in seeds, vegetable oils, and nuts. Structurally, they belong to the same family of cholesterol derived from animal sources [1]. During the last two decades, the use of PSs in oils and fats has gained attention due to their potential health benefits. Once incorporated in oils and fats, PSs have been shown to reduce the absorption of dietary cholesterol in the intestine, which results in lowering blood cholesterol levels and maintaining a healthier lipid profile, therefore contributing to the reduction in the risk of heart diseases such as atherosclerosis, heart attacks, strokes, and other related body functional disorders [2,3,4,5,6]. Furthermore, many research studies have shown that PSs in their pure form or mixed in oils/fats offer a natural alternative to synthetic cholesterol-lowering drugs [5,7,8]. Despite the health benefits of the various formulations of PSs approved in many research studies, their use remains subject to regulatory guidelines and limitations, and these guidelines may vary in different countries [4,9,10].

PSs have limited solubility in fats and oils, typically 1–3% at ambient temperatures depending on the fatty acid profile of the oil medium, which means that high concentrations cannot be easily incorporated into functional food products [11]. Furthermore, PSs are susceptible to oxidation, especially under certain processing conditions, which is responsible for negatively altering the taste, odor, and nutritional quality of oils and fats [12,13]. To overcome the limited solubility of PSs as well as their low oxidative stability, many research studies have shown that esterified PSs offer an attractive solution for facilitating their applications in functional foods, cosmeceuticals, and pharmaceuticals [2,5,7,14,15]. One of the most common methods for synthesizing PSEs involves the esterification of PSs with free fatty acids. This process typically employs strong acids as catalysts, such as sulfuric and p-toluene sulfonic acids [2,16,17,18], or the use of native and immobilized lipases as biocatalysts [7,19,20]. The reaction can be carried out under various conditions, including solvent-based or solvent-free microaqueous systems. Solvent-free lipase-catalyzed esterification methods have gained popularity due to their eco-friendly nature and ease of product recovery [20]. Common fatty acids used for esterification of PSs include palmitic, stearic, and oleic acids, typically from vegetable oil sources [21,22,23]. Transesterification is another possible method for synthesizing PSEs. This reaction involves the direct donation of fatty acyl groups to PSs by using oil TGs or fatty acid short-chain alkyl esters, such as FAEEs, as fatty acyl group donors in the solvent as well as the solvent-free reaction medium. This reaction can be catalyzed by strong alkaline catalysts such as sodium methoxide/ethoxide, or by using lipases in their free or immobilized forms [15,20,24]. On the contrary to strong alkaline and acid catalysts, the major advantages of lipases are their capability to catalyze esterification and transesterification reactions simultaneously [25], operate under mild reaction conditions, are therefore low in energy consumption, and have fewer concurrent byproducts, as well as their tolerance to micro-amounts of water present in the reaction medium [15]. Free as well as lipases immobilized on an organic or inorganic polymer support matrix have been used for the esterification/transesterification of PSs and fatty acyl group donors [22,23]. In many cases, immobilization allows lipases to be more stable, reusable, and easily separated from the reaction mixture. It also improves their operational efficiency in industrial applications and allows the use of conventional industrial reactor configurations, including batch- and continuous-wise stirred-tank, fluidized- and fixed-bed reactors [17,19].

Due to the higher solubility of PSs in free fatty acid medium as compared to using oil TGs as fatty acyl group donors, most studies in the field have used native or immobilized lipases to catalyze esterification reactions between both substrates in solvent- and solvent-free systems. Most of these studies have found that lipases derived from Candida rugosa, Candida antarctica, Alcaligenes sp., and Rhizomucor miehei are the best-performing enzymes to catalyze the production of PSEs in stirred batch and continuous stirred-tank reactors [20,21,23,26]. After the desired esterification reaction conversion is achieved, the product PSEs, characterized by its high solubility in oil medium, can be separated from the reaction mixture. Common separation methods, including solvent extraction, distillation, crystallization, or chromatographic techniques, can be applied to obtain high-purity PSEs. Lipase-catalyzed direct transesterification reactions between PSs with either oil TGs or FAEEs have been less documented due to the low solubility of PSs in hydrophobic oil-based mediums [17,20]. To increase PSs concentrations above 3%, the transesterification reaction medium containing PSs at a maximum concentration of 10% wt./wt. of oil has been heated up to 90–130°C, which led to severe losses in enzyme activity, staining of the final product, and technical complexity in the removal of the free enzyme residues from the reaction medium [11,20]. Although direct enzymatic transesterification exhibits advantages such as one-step reaction under milder reaction conditions, its use to produce PSEs remains unresolved at industrial scales.

The present work reveals a versatile production process based on the use of free or immobilized lipases for catalyzing direct transesterification reactions between oil glycerides as fatty acyl group donors, and solubilized PSs with initial concentrations of up to 30% wt./wt. of oil at temperature ranges tolerated by most enzymes. The proposed process demonstrates for the first time the use of a homogeneous direct transesterification reaction medium where MGs serve as the most favored fatty acyl group donor for PSs to produce PSEs in the presence of a lipase as a catalyst. Such a system would offer a versatile tool for paving the way for employing immobilized lipases to produce PSEs at commercial scales starting from oil glycerides with no limitations for the low solubility of PSs in the reaction medium as well as with no restrictions for choice of reactor type and configuration.

2. Materials and Methods

2.1. Materials

Commercially available lipases derived from various microbial species have been kindly contributed by different enzyme manufacturers, as detailed in

Table 1. Commercial lipases, their sources, forms, and manufacturers, which have been tested in this study.

Commercial Name	Species of Microorganism	Form	Manufacturer
Lipase F-AP	Rhizopus oryzae	Free, powder	Amano Enzymes, Nagoya, Japan
Lipase G	Penicillium camemberti	Free, powder
Lipase A	Aspergillus niger	Free, powder
Lipase PS	Pseudomonas cepacia	Free, powder
Lipase PF	Pseudomonas fluorescens	Free, powder
Lipase R	Penicillium roqueforti	Free, powder
Lipase F	Rhizopus niveus	Free, powder
Lipase AY	Candida (C.) rugosa	Free, powder
Lipase CR	C. rugosa	Free, powder	Enzyme Development Corporation (EDC), New York, NY, USA
Lipase AN	Aspergillus niger	Free, powder
Lipase QLM	Burkholderia (B.) ubonensis	Free, powder	Meito Sangio, Tokyo, Japan
Lipase OF	C. rugosa	Free, powder
Lipase TL	Pseudomonas stutzeri	Free, powder
Lipase SL	Pseudomonas (B.) cepacia	Free, powder
Novocor AD L	C. antarctica A	Free, liquid	Novozymes, Copenhagen, Denmark
Eversa Transform 2.0	Thermomyces lanuginosus	Free, liquid
Lipozyme CALB L	C. antarctica B	Free, liquid

. Edible oils derived from different sources have been purchased from local groceries (Israel). Distilled MGs (Verol IG-90, 95% as MGs) and a mixture of mono- and di-glycerides (Verol IG, 38–42% wt. as MGs) were both generous gifts from Lasenor (Barcelona, Spain). Oleic acid and ethyl oleate of 96% purity were purchased from Sigma (Jerusalem, Israel). A mixture of PSs with a purity of 95%, composed of 40% β-sitosterol, 30% stigmasterol, and 30% campesterol, was kindly contributed by NuSci Institute & Corp. (Walnut, CA, USA). Ethanol, n-hexane, sodium hydroxide, sodium phosphate salts, and other reagents of analytical grade were purchased from Sigma (Israel).

2.2. Analysis Methods

The concentrations of all reaction components, including FFAs, FAEEs, MGs, DGs, TGs, PSs, and PSEs, were determined by gas chromatography. A Hewlett–Packard gas chromatograph (Agilent 6890; Santa Clara, CA, USA) equipped with a flame-ionization detector and a capillary column Agilent DB-1HT, 0.25 mm i.d. × 5 m length, and 0.1 µm film thickness (USA), was used under the following separation conditions: Injector and detector temperatures were maintained at 360 °C; the initial column temperature was 120 °C for 1 min; thereafter, the oven temperature was raised at a rate of 20 °C/min to reach 360 °C for 5 mins. Water concentration in the reaction medium was measured by the Karl Fischer titrator 831 KF Coulometer (Metrohm, Herisau, Switzerland), and FFAs content (%) in the reaction system was also measured by the 848 Titrino plus using a 0.1 M KOH ethanolic solution (Metrohm, Herisau, Switzerland).

2.3. Preparation of Immobilized Lipases

Free lipases were immobilized following standard procedures where a lipase preparation derived from a certain microorganism (typically, 1 g of a powderous or liquid lipase preparation described in Table 1) was solubilized in a phosphate buffer solution (50 mL) of 0.1 M at a certain pH value, typically 7.5 [23]. A macroporous hydrophobic organic polymer resin (10 mL), such as polypropylene beads (Accurel MP1000, beads size 300–700 µm, and mean pore diameter 25 nm, Product of EVONIK, Obernburg, Germany), was first washed twice with three bed volumes of distilled water, and then the wet beads were washed with three bed volumes of ethanol. Ethanol was removed by filtration, and then the beads were added to the lipase solution. The mixture was shaken at room temperature for 8 h. Cold acetone was added to the mixture to increase the enzyme precipitation on the polymer resin. The immobilized enzyme beads were obtained after removal of the aqueous solution by filtration, followed by drying of the wet polymer beads by lyophilization to a water content of less than 3% wt./wt.

2.4. Enzymatic Transesterification/Esterification between PSs and Various Fatty Acyl Donors

A mixture of PSs (0.3 g) and fatty acyl donors (3 g) comprised of any combination of the following two compositions: either a mixture of plant oil TGs, ethyl oleate (as FAEEs), oleic acid (as FFAs), MGs, and DGs at different weight ratios, or partly transesterified plant oil with ethanol comprised mainly of FAEEs, MGs, DGs, TGs, and a minor amount of FFAs (less than 3%), with the weight ratios as described below. The oil mixtures before reactions were degassed by bubbling nitrogen gas, then shaken for 1 h at 60 °C. Reaction between the components of the oil mixture was initiated by the addition of a free (powder/liquid form) or an immobilized lipase preparation (150 and 300 mg, respectively) into the mixtures. The reaction mixtures were incubated at 60 °C and shaken at 160 rpm for 24 h. The water concentration in the reaction medium was in the range of 200–300 ppm based on Karl Fischer titrator analysis. Samples of 60 microliters were withdrawn periodically from the reaction mixtures for GC analysis to determine the concentrations (presented as GC peak area ratios %) of the reaction medium components, including FFAs, MGs, PSs, DGs, PSEs, and TGs. The relative activity of enzymes was calculated as the peak area ratios for the products PSEs divided by the sum of peak area ratios for PSs and PSEs after one hour of reaction. Under the above-described reaction conditions, PSs were not completely soluble in reaction mediums comprised of predominantly oil TGs as fatty acyl group donors, while they were completely soluble in reaction mediums comprising any combination of fatty acid acyl group donors in the presence of FAEEs prepared as detailed below.

2.5. Preparation of Partially Transesterified Plant Oils via Lipase-Catalyzed Transesterification of Oil TGs and Ethanol

A mixture of canola oil (100 g) containing 10% wt. ethanol and 0.1% wt. water was incubated at 60 °C. Thermomyces lanuginosus lipase immobilized on Accurel MP 1000 beads (5 g) was added to the mixture. The reaction mixture was then incubated at 40 °C and shaken at 160 rpm for 6 h. Samples of 60 microliters were withdrawn periodically from the reaction mixture for GC analysis to determine the concentrations (as GC peak area ratios %) of FAEEs, FFAs, MGs, DGs, and TGs. The reaction mixture after 6 h of reaction was filtered off for the removal of the biocatalyst to obtain partially transesterified canola oil, residual water, and nonreacted ethanol. Residual ethanol and water were flash evaporated, and the remaining reaction mixture with the composition of 67.8% FAEEs, 1% FFAs, 12.4% MGs, 9.7% DGs, and 9.1% TGs (as GC peak area ratios) was used as a medium for solubilization of PSs at concentrations of up to 30% wt./wt. of oil medium and as a fatty acyl group donor in transesterification/esterification reactions of PSs.

3. Results and Discussion

3.1. Screening of Free Lipases of Direct Transesterification Activity to Produce PSEs

Table 2. The concentrations of the reaction components after 24 h of direct transesterification between PSs and canola oil TGs using different types of commercial free lipase preparations. Reaction conditions: See Section 2.4. Values correspond to the mean (n = 3) ± standard deviation.

		GC Peak Area Ratio (%)
Name of Lipase	Source of Lipase	FFAs	MGs	PSs	DGs	PSEs	TGs
Control	-	1.4 ± 0.03	0.3 ± 0.01	9 ± 0.18	5.1 ± 0.1	0 ± 0	84.2 ± 3.37
Lipase F-AP	Rhizopus oryzae	4.8 ± 0.1	3 ± 0.06	6.8 ± 0.14	7 ± 0.14	2.2 ± 0.04	76.2 ± 3.05
Lipase G	Penicillium camemberti	3.3 ± 0.07	1.3 ± 0.03	7 ± 0.14	2.3 ± 0.05	2 ± 0.04	84.1 ± 3.36
Lipase A	Aspergillus niger	2.5 ± 0.05	1.2 ± 0.02	7.5 ± 0.15	3.2 ± 0.06	1.5 ± 0.03	84.1 ± 3.36
Lipase PS	Pseudomonas cepacia	7.9 ± 0.16	3.6 ± 0.07	4 ± 0.08	11.9 ± 0.24	5 ± 0.1	67.6 ± 2.7
Lipase PF	Pseudomonas fluorescens	6.8 ± 0.14	2.6 ± 0.05	3 ± 0.06	12.5 ± 0.25	6 ± 0.12	69.1 ± 2.76
Lipase R	Penicillium roqueforti	8.4 ± 0.17	3.9 ± 0.08	7 ± 0.14	11.4 ± 0.23	2 ± 0.04	67.3 ± 2.69
Lipase F	Rhizopus niveus	3.7 ± 0.07	0.9 ± 0.02	7.1 ± 0.14	2.4 ± 0.05	1.9 ± 0.04	84 ± 3.36
Lipase AY	C. rugosa	7.4 ± 0.15	3.7 ± 0.07	0.3 ± 0.01	13.4 ± 0.27	8.7 ± 0.17	66.5 ± 2.66
Lipase CR	C. rugosa	7.6 ± 0.15	3.9 ± 0.08	0.1 ± 0	15 ± 0.3	8.9 ± 0.18	64.5 ± 2.58
Lipase AN	Aspergillus niger	8.6 ± 0.17	3.9 ± 0.08	2.8 ± 0.01	9.5 ± 0.19	6.2 ± 0.12	69 ± 2.76
Lipase QLM	B. ubonensis	9.5 ± 0.19	3.7 ± 0.07	0.1 ± 0	14.2 ± 0.28	8.7 ± 0.17	63.8 ± 2.55
Lipase OF	C. rugosa	8.4 ± 0.17	3.5 ± 0.07	0.3 ± 0.02	13.9 ± 0.28	8.7 ± 0.17	65.2 ± 2.61
Lipase TL	Pseudomonas stutzeri	8.2 ± 0.16	4.3 ± 0.09	0.2 ± 0	12.7 ± 0.25	8.6 ± 0.17	66 ± 2.64
Lipase SL	B. cepacia	11.3 ± 0.23	3.2 ± 0.06	0.8 ± 0.02	13.5 ± 0.27	8.2 ± 0.16	63 ± 2.52
Novocor AD L	C. antarctica A	17.5 ± 0.35	6.5 ± 0.13	6 ± 0.12	18.4 ± 0.37	3 ± 0.06	48.6 ± 1.94
Eversa Transform 2.0	Thermomyces lanuginosus	21.6 ± 0.43	9.9 ± 0.2	7.5 ± 0.15	16.6 ± 0.33	1.5 ± 0.03	42.9 ± 1.72
Lipozyme CALB L	C. antarctica B	3.3 ± 0.07	2 ± 0.04	7.5 ± 0.15	1.7 ± 0.03	1.5 ± 0.03	84 ± 3.36

shows the concentrations of the reaction components FFAs, MGs, PSs, DGs, PSEs, and TGs after 24 h of direct transesterification reaction between PSs and canola oil TGs. In agreement with other research studies, the results presented in Table 2 show that lipases derived from C. rugosa, B. ubonensis, C. antarctica A, and Pseudomonas sp., are the most active enzymes for direct transesterification of oil TGs and PSs to produce PSEs, MGs, and DGs [17,20,23]. Since the maximum solubility of PSs in oil TGs is limited to 3% at ambient temperatures, the reaction medium was heterogeneous at the beginning and, with the progress of the reaction, became homogeneous once adequate amounts of PSEs and MGs were produced, which are responsible for the solubilization of PSs in the reaction medium. Because of the low solubility of PSs in oil TGs at ambient temperatures, the direct transesterification between the reaction components is limited to initial concentrations of PSs lower than 3% to ensure a homogeneous reaction medium as a precondition for the use of packed-bed column reactors. Furthermore, the results presented in Table 2 show that the concentrations of FFAs, MGs, and DGs have increased in the reaction medium after 24 h of reaction. This result is attributed to the competing hydrolysis side reaction catalyzed by most lipase preparations tested in this study in the presence of micro-amounts of water in the reaction medium. A substantial increase in the hydrolytic products was obtained when a commercial liquid lipase preparation containing water was used, which resulted in increasing the level of water in the reaction medium and therefore enhancing the hydrolysis side reaction.

3.2. Enzymatic Transesterification of PSs and Mixtures of Different Weight Ratios of Canola Oil TGs and FAEEs

In an attempt to increase the solubility of PSs in the reaction medium as well as determine the favored fatty acyl group donors, different combinations of oil TGs and ethyl oleate (as FAEEs) were prepared and reacted with PSs (10% wt./wt.) in the presence of two different lipases immobilized on a hydrophobic macroporous support (Accurel MP 1000), separately. Figure 1a–d shows the concentrations of the direct transesterification reaction components between PSs and mixtures of oil TGs and ethyl oleate at different weight ratios after 3 and 24 h of reaction using immobilized lipases derived from C. rugosa (Enzyme Development Corporation (EDC), Ucon, ID, USA), and B. ubonensis, respectively. The lipases derived from C. rugosa (EDC, USA), and B. ubonensis have been selected due to their high relative activity for the production of PSEs as well as because both enzymes are approved for food applications. The results show that the concentrations of TGs in any combination of the reaction medium composition described in Figure 1 were significantly reduced with time until reaching equilibria after 24 h of reaction (equilibria is not shown), while the concentrations of ethyl oleate remained approximately constant with no significant changes in both cases. These results indicate that both different lipases in their immobilized form prefer TGs rather than FAEEs as fatty acyl group donors for PS in the direct transesterification reaction process.

3.3. Transesterification/Esterification of PSs with Mixtures Comprised of Different Weight Ratios of Canola Oil TGs, MGs, and FFAs as Fatty Acyl Donors

Table 3. The concentrations of the reaction components comprised of PSs and a mixture of different weight ratios comprising canola oil (TGs), DGs, MGs, and FFAs after 3 and 24 h of reaction using a lipase derived either from B. ubonensis or C. rugosa immobilized on Accurel MP 1000. Reaction conditions: as described in Section 2.4. Values correspond to the mean (n = 3) ± standard deviation.

Time (h)	Immobilized Lipase	GC Peak Area Ratio (%)
		FFAs	MGs	PSs	DGs	PSEs	TGs
0	Control	24.25 ± 0.73	38.27 ± 1.15	25.14 ± 0.75	11.92 ± 0.36	0 ± 0	0.42 ± 0.01
	Control	14.01 ± 0.42	22.74 ± 0.68	20.29 ± 0.61	10.94 ± 0.33	0 ± 0	32.02 ± 0.96
	Control	9.36 ± 0.28	14.83 ± 0.44	16.75 ± 0.5	8.73 ± 0.26	0 ± 0	50.33 ± 1.51
	Control	5.1 ± 0.15	8.01 ± 0.24	15.04 ± 0.45	5.8 ± 0.17	0 ± 0	66.05 ± 1.98
	Control	4.5 ± 0.14	4.11 ± 0.12	13.67 ± 0.41	4.5 ± 0.14	0 ± 0	73.22 ± 2.2
	Control	2 ± 0.06	2.67 ± 0.08	10.59 ± 0.32	3.02 ± 0.09	0 ± 0	81.72 ± 2.45
	Control	0.86 ± 0.03	0.02 ± 0	10.14 ± 0.3	2.77 ± 0.08	0 ± 0	86.21 ± 2.59
3	B. ubonensis	29.55 ± 0.89	34.88 ± 1.05	23.38 ± 0.7	10.22 ± 0.31	1.76 ± 0.05	0.21 ± 0.01
	B. ubonensis	20.22 ± 0.61	22.47 ± 0.67	17.59 ± 0.53	22.55 ± 0.68	2.7 ± 0.08	14.47 ± 0.43
	B. ubonensis	12.66 ± 0.38	20.77 ± 0.62	14.11 ± 0.42	26.57 ± 0.8	2.64 ± 0.08	23.25 ± 0.7
	B. ubonensis	8.55 ± 0.26	15.46 ± 0.46	11.41 ± 0.34	28.66 ± 0.86	3.63 ± 0.11	32.29 ± 0.97
	B. ubonensis	7.2 ± 0.22	11.47 ± 0.34	9.34 ± 0.28	27.82 ± 0.83	4.33 ± 0.13	39.84 ± 1.2
	B. ubonensis	5.8 ± 0.17	6.6 ± 0.2	5.39 ± 0.16	23.94 ± 0.72	5.2 ± 0.16	53.07 ± 1.59
	B. ubonensis	4.6 ± 0.14	1.45 ± 0.04	3.74 ± 0.11	12.12 ± 0.36	6.4 ± 0.19	71.69 ± 2.15
	C. rugosa	28.89 ± 0.87	34.66 ± 1.04	23.03 ± 0.69	11.12 ± 0.33	2.11 ± 0.06	0.19 ± 0.01
	C. rugosa	19.88 ± 0.6	17.47 ± 0.52	16.48 ± 0.49	18.64 ± 0.56	3.81 ± 0.11	23.72 ± 0.71
	C. rugosa	16.25 ± 0.49	12.36 ± 0.37	12.42 ± 0.37	18.51 ± 0.56	4.33 ± 0.13	36.13 ± 1.08
	C. rugosa	12.48 ± 0.37	7.55 ± 0.23	10.27 ± 0.31	19.5 ± 0.59	4.77 ± 0.14	45.43 ± 1.36
	C. rugosa	9.55 ± 0.29	6.32 ± 0.19	8.86 ± 0.27	19.96 ± 0.6	4.81 ± 0.14	50.5 ± 1.52
	C. rugosa	6.3 ± 0.19	4.11 ± 0.12	5.5 ± 0.17	17.02 ± 0.51	5.09 ± 0.15	61.98 ± 1.86
	C. rugosa	4.6 ± 0.14	1.32 ± 0.04	3.93 ± 0.12	9.94 ± 0.3	6.21 ± 0.19	74 ± 2.22
24	B. ubonensis	31.22 ± 0.94	33.54 ± 1.01	19.19 ± 0.58	9.99 ± 0.3	5.95 ± 0.18	0.11 ± 0
	B. ubonensis	21.44 ± 0.64	20.69 ± 0.62	10.03 ± 0.3	22.52 ± 0.68	10.26 ± 0.31	15.06 ± 0.45
	B. ubonensis	17.1 ± 0.51	19.39 ± 0.58	7.42 ± 0.22	26.04 ± 0.78	9.33 ± 0.28	20.72 ± 0.62
	B. ubonensis	13 ± 0.39	14.72 ± 0.44	2.44 ± 0.07	28.17 ± 0.85	12.6 ± 0.38	29.07 ± 0.87
	B. ubonensis	11.3 ± 0.34	11.98 ± 0.36	4.55 ± 0.14	26.5 ± 0.8	9.12 ± 0.27	36.55 ± 1.1
	B. ubonensis	9.8 ± 0.29	8.1 ± 0.24	2.04 ± 0.06	24.68 ± 0.74	8.55 ± 0.26	46.83 ± 1.4
	B. ubonensis	8.5 ± 0.26	2.74 ± 0.08	0.59 ± 0.02	13.3 ± 0.4	9.55 ± 0.29	65.32 ± 1.96
	C. rugosa	31.92 ± 0.96	31.89 ± 0.96	22.79 ± 0.68	10.83 ± 0.32	2.35 ± 0.07	0.22 ± 0.01
	C. rugosa	21.01 ± 0.63	18.47 ± 0.55	14.97 ± 0.45	18.88 ± 0.57	5.32 ± 0.16	21.35 ± 0.64
	C. rugosa	15.77 ± 0.47	12.65 ± 0.38	10.81 ± 0.32	18.27 ± 0.55	5.94 ± 0.18	36.56 ± 1.1
	C. rugosa	12.91 ± 0.39	8.83 ± 0.26	6.84 ± 021	21.14 ± 0.63	8.2 ± 0.25	42.08 ± 1.26
	C. rugosa	12 ± 0.36	6.76 ± 0.2	6.62 ± 0.2	21.39 ± 0.64	7.05 ± 0.21	46.18 ± 1.39
	C. rugosa	9.9 ± 0.3	5.35 ± 0.16	2.71 ± 0.08	18.24 ± 0.55	7.88 ± 0.24	55.92 ± 1.68
	C. rugosa	5.31 ± 0.16	1.72 ± 0.05	1.38 ± 0.04	11.46 ± 0.34	8.76 ± 0.26	71.37 ± 2.14

depicts the results for determining the favorite fatty acyl group donor for both lipases derived from B. ubonensis and C. rugosa, immobilized on Accurel MP 1000 separately, to produce PSEs through transesterification/esterification reactions starting from PSs and a mixture of fatty acyl group donors comprised of FFAs, MGs, DGs, and TGs at different weight ratios. The presence of FFAs, MGs, and DGs in the reaction medium led to completely solubilizing PSs, producing a homogeneous reaction medium. The results presented in Table 3 show explicitly that TGs at any substrate composition ratio are better fatty acyl group donors than MGs, DGs, and FFAs for PSs to produce PSEs under the described reaction conditions. The results also show that there was an increase in the concentrations of FFAs, typically in the range of 4–8%, as well as for MGs and DGs after 3 and 24 h of reaction as compared to their initial concentrations. This increase is attributed to the competing hydrolysis side reaction catalyzed by both lipases in the presence of micro-amounts of water in the reaction medium.

3.4. Screening of Immobilized Lipases of Direct Transesterification Activity to Produce PSEs Using Partially Transesterified Plant Oils with Ethanol as a Reaction Medium

The results presented in

Table 4. The concentrations of the reaction components comprised of PSs and partially transesterified canola oil with ethanol, with the composition as described in Section 2.5, after 24 h of reaction using different lipase preparations immobilized on Accurel MP 1000. Reaction conditions: As described in Section 2.4, values correspond to the mean (n = 3) ± standard deviation.

		GC Peak Area Ratio (%)
Name of Lipase	Immobilized Lipase	FFAs	FAEEs	MGs	PSs	DGs	PSEs	TGs
Control	-	1.2 ± 0.02	56.5 ± 2.26	13.4 ± 0.27	10.5 ± 0.2	10.4 ± 0.2	0 ± 0	8 ± 0.16
Lipase F-AP	Rhizopus oryzae	8.3 ± 0.17	56.2 ± 2.25	6.9 ± 0.14	8.5 ± 0.17	10.3 ± 0.2	2 ± 0.04	7.8 ± 0.16
Lipase G	Penicillium camemberti	4 ± 0.08	57.2 ± 2.29	8 ± 0.16	9.6 ± 0.19	12.3 ± 0.25	0.9 ± 0.02	8 ± 0.15
Lipase A	Aspergillus niger	6.2 ± 0.12	56.8 ± 2.27	7.1 ± 0.14	9.5 ± 0.19	11.8 ± 0.24	1 ± 0.02	7.6 ± 0.16
Lipase PS	Pseudomonas cepacia	8.6 ± 0.17	56.8 ± 2.25	6.2 ± 0.12	6.3 ± 0.13	10.1 ± 0.21	4.2 ± 0.08	7.8 ± 0.16
Lipase PF	Pseudomonas fluorescens	9.5 ± 0.19	55.5 ± 2.22	6.3 ± 0.13	8.4 ± 0.17	10.2 ± 0.2	2.1 ± 0.04	8 ± 0.15
Lipase R	Penicillium roqueforti	7.3 ± 0.15	56.3 ± 2.25	8.1 ± 0.16	9.6 ± 0.19	10.1 ± 0.2	0.9 ± 0.02	7.7 ± 0.16
Lipase F	Rhizopus niveus	3.6 ± 0.07	57.5 ± 2.3	7.8 ± 0.16	9.7 ± 0.19	12.8 ± 0.26	0.8 ± 0.02	7.8 ± 0.15
Lipase AY	C. rugosa	14 ± 0.28	55.2 ± 2.21	5.2 ± 0.1	0.4 ± 0.01	8.2 ± 0.16	10.1 ± 0.2	6.9 ± 0.14
Lipase CR	C. rugosa	10.5 ± 0.21	55.2 ± 2.2	5.6 ± 0.11	0.7 ± 0.01	10.5 ± 0.21	9.8 ± 0.2	7.7 ± 0.15
Lipase AN	Aspergillus niger	6.6 ± 0.13	56.2 ± 2.25	7.7 ± 0.15	0.4 ± 0.01	11.2 ± 0.22	10.1 ± 0.2	7.8 ± 0.15
Lipase QLM	B. ubonensis	12.8 ± 0.26	55.6 ± 2.22	4.8 ± 0.1	0.5 ± 0.01	8.4 ± 0.17	10 ± 0.19	7.9 ± 0.16
Lipase OF	C. rugosa	13.9 ± 0.28	53.3 ± 2.13	5.7 ± 0.11	0.8 ± 0.02	8.6 ± 0.17	9.7 ± 0.21	8 ± 0.16
Lipase TL	Pseudomonas stutzeri	14.9 ± 0.3	55.2 ± 2.21	6.2 ± 0.12	0.2 ± 0	6.5 ± 0.13	10.3 ± 0.2	6.7 ± 0.13
Lipase SL	Pseudomonas (B.) cepacia	12.7 ± 0.25	56 ± 2.24	5.6 ± 0.11	0.4 ± 0.1	7.4 ± 0.15	10.1 ± 0.02	7.8 ± 0.16
Novocor AD L	C. antarctica A	9.7 ± 0.19	55.8 ± 2.23	5.8 ± 0.12	9.4 ± 0.19	10.6 ± 0.21	1.1 ± 0.02	7.6 ± 0.15
Eversa Transform 2.0	Thermomyces lanuginosus	10.7 ± 0.21	55.3 ± 2.21	7.4 ± 0.15	9.6 ± 0.19	8.1 ± 0.16	0.9 ± 0.02	8 ± 0.16
Lipozyme CALB L	C. antarctica B	9.9 ± 0.2	56 ± 2.24	7.4 ± 0.15	9.6 ± 0.18	8.4 ± 0.17	0.9 ± 0.02	7.8 ± 0.16

are in harmony with the results depicted in Table 2, where lipases derived from C. rugosa, B. ubonensis, C. antarctica A, Pseudomonas sp., and C. cylindracea in their free as well as immobilized forms exhibited the highest activity for catalyzing the direct transesterification of PSs and a mixture comprised of FAEEs and oil glycerides (mono-, di- and tri-glycerides). The results clearly show that the concentrations of FAEEs and DGs remained approximately unchanged before and after the transesterification reaction. Furthermore, the results show that the concentrations of TGs increased after 24 h of reaction, which can be attributed to the synthetic capability of most of the immobilized enzymes used to esterify MGs and DGs to TGs. The results show explicitly that the concentrations of MGs have dropped significantly after reaction due to two main competing reactions: the first is its conversion to TGs by most lipase preparations, and the second is its transesterification with PSs, which results in producing PSEs. These results show, surprisingly, that MGs are predominantly the most favored fatty acyl group donors amongst all other reaction medium components under the described reaction conditions. Based on the results presented in Table 2, Table 3 and Table 4, it can be concluded that the preference of lipases for MGs as fatty acyl group donors compared to TGs, DGs, and FFAs in direct transesterification reactions is ascribed to the addition of FAEEs into the reaction medium. This result can be interpreted as less steric hindrance for MGs molecules, which contain one fatty acyl group, as compared to TGs molecules, which contain three fatty acyl groups on the glycerol backbone. One more probable reason for such substrate preference could be due to the facilitated hydrophobic interactions between the fatty acyl groups of MGs molecules and the active site of the enzyme in the presence of FAEEs in the reaction medium. To the best of our knowledge, this is the first time to show that the addition of FAEEs into the reaction medium for lipase-catalyzed direct transesterification of PSs and fatty acyl donors, including FFAs, MGs, DGs, and TGs, results in altering substrate preference for the enzyme. Due to the addition of FAEEs into such reaction medium composition our results show that MGs become predominantly the favored fatty acyl group donors for PSs amongst the other tested possible fatty acyl group donors.

3.5. Enzymatic Transesterification of PSs and Partially Transesterified Oils with Ethanol Containing Low Levels of FAEEs

Table 5. The concentrations of the reaction components comprised of PSs and partially transesterified canola oil containing low concentrations of FAEEs (prepared as described in Section 2.5) at different time intervals using lipases from different sources immobilized on Accurel MP 1000 beads. Reactions were performed without and with the addition of water at weight ratios of 0.5% of the reaction medium after bringing the oil mixture to a neutral pH with a sodium hydroxide solution of 3 M. Reaction conditions: As described in Section 2.4, values correspond to the mean (n = 3) ± standard deviation.

			GC Peak Area Ratio (%)
Time (h)	Immobilized Lipase	Water (wt./wt.%) of Oil	FAEEs	FFAs	MGs	PSs	DGs	PSEs	TGs
0	Control	0	20.8 ± 0.62	1.2 ± 0.02	27.3 ± 0.82	14.0 ± 0.28	20.8 ± 0.62	0.0 ± 0	15.9 ± 0.48
3	B. ubonensis	0	20.2 ± 0.61	2.3 ± 0.05	17.2 ± 0.52	10.6 ± 0.21	19.5 ± 0.59	5.3 ± 0.1	24.8 ± 0.74
	B. ubonensis	0.5	21.1 ± 0.63	6.5 ± 0.13	16.2 ± 0.49	11.2 ± 0.22	20.8 ± 0.62	4.9 ± 0.1	19.3 ± 0.58
	C. rugosa	0	20.2 ± 0.61	4.0 ± 0.08	20.4 ± 0.61	8.6 ± 0.17	23.1 ± 0.69	8.7 ± 0.17	15.0 ± 0.45
	C. rugosa	0.5	22.0 ± 0.66	7.3 ± 0.15	12.5 ± 0.38	3.5 ± 0.07	19.2 ± 0.58	16.9 ± 0.34	18.6 ± 0.56
6	B. ubonensis	0	20.5 ± 0.62	1.5 ± 0.03	14.5 ± 0.44	8.1 ± 0.16	25.0 ± 0.75	9.0 ± 0.18	21.3 ± 0.64
	B. ubonensis	0.5	20.8 ± 0.62	6.8 ± 0.14	14.5 ± 0.4	9.3 ± 0.19	19.6 ± 0.59	7.9 ± 0.16	21.1 ± 0.63
	C. rugosa	0	20.7 ± 0.62	3.2 ± 0.06	18.9 ± 0.57	9.1 ± 0.18	23.7 ± 0.71	8.9 ± 0.18	15.5 ± 0.47
	C. rugosa	0.5	21.0 ± 0.63	7.7 ± 0.15	10.8 ± 0.32	3.1 ± 0.06	20.3 ± 0.61	17.4 ± 0.35	19.7 ± 0.59
24	B. ubonensis	0	20.1 ± 0.6	1.6 ± 0.03	11.9 ± 0.36	4.2 ± 0.08	25.0 ± 0.75	15.7 ± 0.31	21.5 ± 0.65
	B. ubonensis	0.5	20.3 ± 0.61	6.8 ± 0.14	12.2 ± 0.37	4.9 ± 0.1	21.4 ± 0.64	14.4 ± 0.29	19.9 ± 0.6
	C. rugosa	0	21.0 ± 0.63	3.5 ± 0.07	18.8 ± 0.56	9.1 ± 0.18	22.9 ± 0.69	8.8 ± 0.18	15.9 ± 0.48
	C. rugosa	0.5	21.3 ± 0.64	8.3 ± 0.17	11.0 ± 0.33	3.0 ± 0.06	18.5 ± 0.56	17.1 ± 0.34	20.7 ± 0.62

shows the concentrations of the transesterification reaction components at different time intervals using PSs dissolved in partially transesterified canola oil with ethanol but containing 20.8% instead of 57.7% of FAEEs, using lipases from different sources immobilized on Accurel MP 1000 beads. Contrary to the results reported in former research studies [17,20], the results presented in Table 5 show explicitly that MGs, in the presence of FAEEs in the reaction medium, serve as the favored fatty acyl group donor for PSs as compared to TGs, DGs, and FFAs. The results also show that the addition of water (0.5% by wt.) to the reaction system leads to increased direct transesterification activity of both enzymes as well as higher concentrations of PSEs after 24 h of reaction. The increase in the concentrations of FFAs in all experiments where water was added is attributed to the occurrence of the competing hydrolysis side reaction to form FFAs as a hydrolytic byproduct for the partial hydrolysis of MGs, mainly. Furthermore, there was an increase in the concentrations of TGs, which can be attributed to the esterification activity of both lipases to partly convert MGs and FFAs to TGs.

3.6. Enzymatic Transesterification of PSs with Partially Enzymatically Transesterified Oil Comprised of FAEEs, MGs, and TGs/or with Oil Mixtures Prepared by Mixing Appropriate Weight Ratios of FAEEs, MGs, DGs, and TGs

The following are two oil mixtures (A and B) with approximately similar composition:

Mixture A: Partially enzymatically transesterified canola oil with ethanol with the following composition, which was prepared as described in Section 2.5:

FAEEs % ± SD	MGs % ± SD	DGs % ± SD	TGs % ± SD
67.8 ± 2.71	12.4 ± 0.25	9.7 ± 0.19	9.1 ± 0.18

Mixture B: An oil mixture prepared by mixing appropriate weight ratios of ethyl oleate (as FAEEs), MGs, DGs, and canola oil (as TGs):

FAEEs % ± SD	MGs % ± SD	DGs % ± SD	TGs % ± SD
68.4 ± 2.74	12.7 ± 0.25	9.6 ± 0.2	9.3 ± 0.2

Both oil mixtures have been enzymatically transesterified with 10% wt./wt. PSs separately under the same reaction conditions.

Table 6. The concentrations of the reaction components at different time intervals for the transesterification of PSs and oil mixtures A and B, separately, using different lipases immobilized on Accurel MP 1000 beads. Reaction conditions: As described in Section 2.4, values correspond to the mean (n = 3) ± standard deviation.

			GC Peak Area Ratio (%)
Time (h)	Reaction Mixture	Immobilized Lipase	FAEEs	FFAs	MGs	PSs	DGs	PSEs	TGs
0	Mixture A	T = 0	62 ± 2.48	1.4 ± 0.03	11 ± 0.22	10.6 ± 0.21	7.9 ± 0.16	0 ± 0	7.1 ± 0.14
	Mixture B	T = 0	61.1 ± 2.44	1.7 ± 0.03	11.2 ± 0.22	10.6 ± 0.2	7.8 ± 0.15	0 ± 0	7.6 ± 0.15
3	Mixture A	B. ubonensis	60.9 ± 2.4	5.1 ± 0.1	8.9 ± 0.18	4.7 ± 0.09	7.7 ± 0.16	5.9 ± 0.12	6.8 ± 0.14
	Mixture B	B. ubonensis	59.8 ± 2.38	5.5 ± 0.11	9.5 ± 0.19	5.6 ± 0.11	7.5 ± 0.15	5 ± 0.1	7.1 ± 0.14
	Mixture A	C. rugosa	58.9 ± 2.36	7.1 ± 0.14	9.9 ± 0.2	1.8 ± 0.04	7.5 ± 0.17	8.8 ± 0.18	6 ± 0.12
	Mixture B	C. rugosa	59.2 ± 2.37	7.9 ± 0.16	8.7 ± 0.17	3.5 ± 0.07	6.8 ± 0.13	7.1 ± 0.14	6.8 ± 0.13
6	Mixture A	B. ubonensis	58.8 ± 2.35	5.3 ± 0.11	10.5 ± 0.21	2.8 ± 0.06	7.8 ± 0.16	7.8 ± 0.16	7 ± 0.14
	Mixture B	B. ubonensis	58.4 ± 2.34	6.8 ± 0.14	10.1 ± 0.2	4.4 ± 0.09	7.5 ± 0.15	6.2 ± 0.12	6.6 ± 0.12
	Mixture A	C. rugosa	59 ± 2.36	7.1 ± 0.14	9.7 ± 0.19	0.6 ± 0.01	7.2 ± 0.14	10 ± 0.2	6.4 ± 0.13
	Mixture B	C. rugosa	58.3 ± 2.33	7.7 ± 0.15	8.7 ± 0.17	1.8 ± 0.04	7.5 ± 0.15	8.8 ± 0.18	7.2 ± 0.16
24	Mixture A	B. ubonensis	59 ± 2.36	6.6 ± 0.13	9.8 ± 0.2	1.1 ± 0.02	7.6 ± 0.15	9 ± 0.18	6.9 ± 0.14
	Mixture B	B. ubonensis	59.3 ± 2.37	6.2 ± 0.12	10.5 ± 0.21	2.2 ± 0.04	7 ± 0.14	8.4 ± 0.17	6.4 ± 0.13
	Mixture A	C. rugosa	58.2 ± 2.33	6.8 ± 0.14	9.5 ± 0.19	0.1 ± 0	7.8 ± 0.16	10.5 ± 0.21	7.1 ± 0.14
	Mixture B	C. rugosa	59.2 ± 2.37	7.9 ± 0.16	9.2 ± 0.18	1.6 ± 0.03	7.6 ± 0.15	9.5 ± 0.19	5 ± 0.1

shows the concentration profiles of the reaction components after 3, 6, and 24 h of reaction using lipases from two different sources immobilized on Accurel MP 1000, separately. The results presented in Table 6 show clearly that MGs were primarily the preferred donors of fatty acyl groups to PSs rather than TGs or FAEEs under the described direct transesterification reaction conditions. Furthermore, the results presented in Table 6 support our hypothesis that the presence of FAEEs in the direct transesterification reaction system, besides enhancing the solubility of PSs, leads to altering the substrate preference for both lipases, making MGs the favored fatty acyl group donors for PSs.

3.7. The Effect of Phytosterol Concentration

The results presented in Figure 2 illustrate the effect of PSs concentration on the relative direct transesterification activity of B. ubonensis and C. rugosa lipases, separately immobilized on Accurel MP 1000. Phytosterols at concentrations of up to 30% wt./wt. of canola oil glycerides (mono-, di-, and tri-glycerides) mixture containing 20% FAEEs could be completely dissolved at the reaction temperature of 60°C, making a homogeneous reaction medium. Unlike the results reported in other research studies [17,20], the results illustrated in Figure 2 show that the enzyme transesterification activity did not decrease either due to increasing the PSs concentrations or due to an increase in the rate of formation of the byproduct DGs when the initial concentrations of PSs were raised in the range of 5–30%. These results show for the first time the applicability of lipase-catalyzed direct transesterification of phytosterols in homogeneous oil media with no restrictions for their initial concentrations up to 30% wt., as well as no enzyme activity inhibition caused by the byproducts MGs and DGs. Furthermore, the GC analysis results in these tests show explicitly that MGs present in the oil medium are the favorite fatty acyl group donors amongst all other possible used fatty acyl group donors for PSs to produce their fatty acid esters.

4. Conclusions

The present work demonstrates an industrially feasible solvent-free direct transesterification process for oils/fats and PSs using lipases immobilized onto a support matrix as an alternative method to conventional chemical processes to produce PSEs for different industrial applications. The findings of this study show for the first time that the presence of FAEEs in lipase-catalyzed direct transesterification reaction medium comprised of PSs and mixtures of different ratios of FFAs, MGs, DGs, and TGs leads to altering of substrate preference for the enzyme. Our study reveals that under such reaction conditions MGs would predominately become the favored fatty acyl group donors for PSs amongst all other tested fatty acyl group donors. This result can be interpreted due to the steric hindrance effect imposed by TGs molecules containing three fatty acyl groups, which are of higher average molecular weight than MGs containing only one fatty acyl group on the glycerol backbone. It is also anticipated that such substrate preference could be due to the amphiphilic nature of MGs molecules, which results in facilitating the hydrophobic interactions between the fatty acyl groups of MGs molecules in the presence of FAEEs in the reaction medium and the active site of the enzyme. Furthermore, this method offers several advantages over traditional methods using chemical catalysts or free enzymes, such as avoiding the limited solubility of PSs in oil TGs, ease of separation and reuse of the immobilized enzymes, and improved efficiency in PSE production processes. Another major advantage of these findings is the allowance for the use of homogeneous direct transesterification reaction medium comprised of various combinations of FAEEs, MGs, DGs, TGs, and up to 30% wt. of solubilized PSs, a prerequisite for utilizing recyclable immobilized enzymes in batch and continuous packed-bed reactors to avoid mass transfer limitations in the vicinity of the immobilized enzyme as well as blockage of packed-bed column reactors.

5. Patents

A US patent has been filed based on the results reported in this manuscript.