Evaluation of different commercial hydrophobic supports for the immobilization of lipases: tuning their stability, activity and specificity

Abstract

Five different commercial supports (Lifetech™ ECR1061M (styrene/methacrylic polymer), Lifetech™ ECR8804M (octadecyl methacrylate), Lifetech™ ECR8806M (octadecyl methacylate), Lifetech™ ECR1090M (styrene) and Lifetech™ ECR1030M (DVB/methacrylic polymer)) were compared to octyl agarose in their performance in the immobilization of four different lipases (from Rhizomucor miehie (RML), from Thermomyces lanuginosus (TLL) and the forms A and B from Candida antarctica, (CALA and CALB)) and of the phospholipase Lecitase Ultra™ (LU). The new enzymatic derivatives were evaluated and compared with the commercial biocatalyst (Novozym 435 (CALB), Lipozyme RM IM and Lipozyme TL IM). Textural properties, loading capacity, enzyme stability under different conditions, and activity versus different substrates were analyzed. Although all of the supports reversibly immobilized lipases via interfacial activation of lipases versus the hydrophobic surface of the support, some of them permitted a significant improvement in the final biocatalyst compared to the reference support or the commercial preparations. Enzyme specificity depended strongly on the used support (e.g., the new ones gave almost null activity versus p-nitrophenyl butyrate). However, there is not a universal optimal support; the “best” support depends on the enzyme, the parameter studied and the substrate utilized. Nevertheless, under the conditions utilized, the preparations showed a very good performance in a diversity of reactions and permitted their reuse (both the biocatalyst and the supports after eliminating the enzyme by washing the enzyme with triton X-100). These supports will permit enlarging the library of immobilized lipase biocatalyst, being supports useful for aqueous or organic medium.

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1. Introduction

Lipases are among the most utilized enzymes in biocatalysis because nature offers a wide variety of these enzymes with different properties^1–7 and they are able to recognize a wide range of substrates,^8–10 catalyzing many different reactions^1–10 some of them very far from the physiological function, as the so-called promiscuous reactions.^11–13 Moreover, they are very stable in different reaction media, from organic solvents^14–16 to neoteric media.^17–22

Lipases are esterases able to work at the interface of their natural substrates, the drops of oils.^23–29 This is possible thanks to the mechanism of interfacial activation: lipases have the capability to become adsorbed on the hydrophobic surface of these oil drops via their active center.^23–29 The lipase active center and its surroundings are hydrophobic. In aqueous homogenous media, this area is usually isolated from the medium by a polypeptide chain called lid or flap, (closed form). The lid is mobile and has a hydrophobic internal face and a hydrophilic external face, the pocket formed by the internal side of the lid plus the active center surroundings make available a very large hydrophobic area, unfavorable in aqueous medium, that becomes stabilized after adsorption on the hydrophobic substrate drop (open form).^23–29 The lipase active form shows the lid shifted (e.g., it is adsorbed to the hydrophobic surface of the substrate drop), exposing the active center to the medium and originating the active form of the lipase.^23–29 The lid may be very small, unable to fully isolate the lipase active center from the medium, like in the case of the lipase B from Candida antarctica,³⁰ or very large and complex, like the double lid presented in a lipase from Geobacillus themocatenulatus.³¹ The movement of the lid causes structural changes along the whole enzyme structure. This generates a great flexibility of the lipase active center and makes that the lipase properties may be easily altered without destroying its activity.^32–39

This tendency of lipases to become adsorbed on hydrophobic surfaces makes their handling complex, e.g., lipases tend to form aggregates involving two open lipase molecules,^40,41 or may become adsorbed on hydrophobic areas of other proteins,⁴² altering their properties. However, it has permitted to develop a strategy for the one step immobilization, purification, stabilization and hyperactivation of lipases involving the open form of the lipase.⁴³ The protocol is rather simple: lipases are the only water soluble proteins that become almost fully immobilized on hydrophobic supports at very low ionic strength.⁴⁴ There are many new supports that have enabled several new approaches for immobilization of lipases (nanorods and nanoparticles, core–shell, mesoporous silicates),⁴⁵ but the hydrophobic supports are mostly used to immobilize lipases via a reversible interfacial activation versus their hydrophobic surface.

Using this immobilization strategy, the high flexibility of the lipase active center^23–31,34 permits that the final properties of the lipase depend on the support features even although the same immobilization mechanism is occurring and the same enzyme orientation is achieved: the support hydrophobicity, its surface topography and the presence of different groups on its surface alter the final catalytic properties of the immobilized lipase.^46–49 In some instances, some lipases only are immobilized on a certain hydrophobic support, and this selective adsorption is not always a function of just higher or lower support hydrophobicity, in some instances this has permitted to separate different lipase isoforms.^50–53

Therefore, the availability of a large diversity of commercially hydrophobic supports having very different properties is very interesting. Many different hydrophobic supports have been used, in some cases supports coated with acyl groups (e.g., silica^54–59 or agarose^{44,45,50–53} coated with acyl groups), in other instances supports with a hydrophobic core (e.g., styrene divinylbenzene) have been employed.^48,49

In this paper, we have compared 6 different commercial supports to immobilize several lipases, and compared them to the commercial immobilized preparations when available. Octyl-agarose beads support has been utilized as a reference support, as this support has been described as very useful to produce very active and stabilized lipase immobilized preparations (even more stable than those immobilized via multipoint covalent attachment).^60,61 This support has been compared with 5 new ones, Lifetech™ ECR1061M (styrene/methacrylic polymer), Lifetech™ ECR8804M (octadecyl methacrylate), Lifetech™ ECR8806M (octadecyl methacylate), Lifetech™ ECR1090M (styrene) and Lifetech™ ECR1030M (DVB/methacrylic polymer), new supports scarcely used in literature and never in a comparison like the one presented in this paper.⁶¹ As enzymes, we have used some of the most utilized in literature: lipases A and B from Candida antarctica (CALB^62,63 and CALA⁶⁴), lipases from Thermomyces lanuginosus (TLL)⁶⁵ and from Rhizomucor miehei (RML).^66,67 Lecitase Ultra (LU), a commercial artificial chimeric phospholipase A1,^68,69 has been also utilized, as this enzyme is able to suffer artificial activation as a standard lipase.⁷⁰ This enzyme is the result of the fusion of the gen of the lipase from Thermomyces lanuginosus and that of the phospholipase from Fusarium oxysporum.^66,67 All supports immobilize the lipases via interfacial activation versus their hydrophobic surfaces.

2. Experimental section

2.1. Materials

Supports were kindly supplied by Purolite® ECR Enzyme Immobilization Resins (Wales, UK). Soluble lipases from Candida antarctica (isoform A and B, CALA and CALB, respectively), Thermomyces lanuginosus (TLL), Rhizomucor miehie (RML) and phospholipase Lecitase Ultra as well as Novozym® 435, Lipozyme® RM IM and Lipozyme® TL immobilized enzymes were a kind gift from Novozymes (Spain). Commercial and home-made immobilized enzymes were washed with distilled water before its use, to ensure that we employ fully wet enzyme derivatives. Methyl phenylacetate, methyl mandelate, phenylacetic acid, mandelic acid, diacetin, triacetin and p-nitrophenyl butyrate (p-NPB) were purchased from Sigma-Aldrich (St. Louis, USA). Octyl-agarose CL-4B beads were from GE Healthcare (Uppsala, Sweden). Acetonitrile (HPLC grade) was purchased from Fisher Scientific (Leicestershire, UK). Electrophoresis reagents were obtained from Bio-Rad (Hercules, USA). Other reagents were of analytical degree.

2.2. Standard determination of enzyme activity

This assay was performed by measuring the increase in absorbance at 348 nm produced by the released p-nitrophenol in the hydrolysis of 0.4 mM p-NPB in 25 mM sodium phosphate at pH 7.0 and 25 °C (ε under these conditions is 5150 M⁻¹ cm⁻¹). Reaction was started by adding 50–100 μL of lipase solution or suspension to 2.55 mL of substrate solution. Spontaneous hydrolysis of p-NPB was monitored under identical conditions without enzyme. One unit of activity (U) was defined as the amount of enzyme that hydrolyzes 1 μmol of p-NPB per minute under the conditions described previously. Protein concentration was estimated by the Bradford dye binding method,⁷¹ recording the absorbance at 595 nm and using bovine serum albumin as the reference.

2.3. Immobilization of enzymes

2.3.1. Wetting of the Purolite supports. Purolite supports are so hydrophobic that water can hardly penetrate into their pores. Therefore, these supports were submitted to a treatment to remove air and fill the pores with water. A sample of 10 g of each support was suspended in 50 mL of methanol for 1 h under mild stirring; after that, 50 mL of distilled water were added to have a 50% water solution. After 15 additional minutes of mild stirring, the supports were filtered under vacuum and washed 5 times in a glass funnel with 5 volumes of water. Finally, supports were stored at 4 °C, in flask with enough distilled water to avoid the dehydration.

2.3.2. Immobilization of lipases on octyl agarose (OC) and Purolite supports. Conditions previously described were used.⁴⁵ 10 g of each support was suspended in an enzyme solution (0.1–0.5 mg protein per mL) in 5 mM sodium phosphate at pH 7 and 25 °C and left in continuous agitation at 200 rpm. The enzyme concentration and volume were adjusted to obtain the desired enzyme loading on the final biocatalyst. Activity of both supernatant and suspension was followed using the pNPB assay. After immobilization the suspension was filtered and the supported enzymes were washed several times with distilled water, and stored at 4 °C.

2.3.3. Determination of loading capacity of the different supports. The necessary volume of lipase solution (0.2 mg protein per mL in 5 mM sodium phosphate at pH 7 and 25 °C) were added to 1 g of the each supports, to reach an amount of protein of 1–80 mg protein per g support. Supernatant samples were taken periodically to measure its activity by the pNPB (see above) assay and immobilization was considered to be complete when no significant changes in the activity on the supernatant were detected after 8 h under continuous agitation.

2.4. Study of the stability of the different lipase biocatalysts

2.4.1. Thermal inactivation of different enzyme immobilized preparations. 0.25 g of immobilized enzyme was suspended in 5 mL of 50 mM sodium acetate at pH 5, sodium phosphate at pH 7 or sodium bicarbonate at pH 9 at different temperatures. Periodically, samples were withdrawn and the activity was measured using methyl mandelate as described below.

2.4.2. Inactivation of different preparations in the presence of organic co-solvents. Enzyme preparations were incubated in mixtures of acetonitrile or 1,4-dioxane/100 mM Tris–HCl (pH 7) at different temperatures. After some time, samples were withdrawn and the activity was measured using methyl mandelate as described below. The organic co-solvents presented in the samples did not have a significant effect on the enzyme activity (results not shown).

2.5. Determination of the hydrolytic activity of the biocatalyst versus different substrates

2.5.1. Hydrolysis of triacetin. Solutions of 100 mM triacetin in 500 mM sodium acetate containing 20% acetonitrile at pH 5 were prepared (substrate was fully soluble). Then, we added 100 mg of the different biocatalysts to 10 mL of this substrate solution and the reaction suspensions were gently stirred at 25 °C. Samples were periodically withdrawn from these reaction suspensions and the biocatalyst was discarded by centrifugation. In order to determine the concentration of reaction products, supernatants were analyzed by RP-HPLC (JASCO PU-2085) coupled with a UV-1575 Intelligent UV/VIS Detector. Detection was performed at 230 nm using 10% acetonitrile/water (v/v) as mobile phase and a Kromasil C18 column (15 cm × 0.46 cm). Retention times were 34.5 min for triacetin and 6.5 min for 1,2 diacetin. Concentrations of triacetin and diacetin were calculated from calibration curves using analytical standards. Activity was determined by triplicate with a conversion of 20–30%, and data are given as average values.

2.5.2. Hydrolysis of methyl phenylacetate. For these experiments, 20–200 mg of the immobilized preparations were added to 0.5–3 mL of 20 mM substrate in 50 mM sodium phosphate at pH 7 containing 50% acetonitrile and 25 °C under continuous stirring. The conversion degree was analyzed by RP-HPLC (JASCO PU-2085 coupled with a UV-1575 Intelligent UV/VIS Detector) using a Kromacil C18 (15 cm × 0.46 cm) column. Samples (20 μL) were injected and eluted at a flow rate of 1.0 mL min⁻¹ using acetonitrile/10 mM ammonium acetate (35 : 65, v/v) at pH 2.8 as mobile phase and UV detection was performed at 230 nm. The ester presented a retention time of 3.5 minutes while the acid had a retention time of 9.6 minutes. One unit of enzyme activity was defined as the amount of enzyme necessary to produce 1 μmol of phenylacetic acid per minute under the condition described above. Activity was determined by triplicate with a maximum conversion of 20–30%, and data are given as average values.

2.5.3. Hydrolysis of the R- and S-methyl mandelate. Enzyme activity was also determined using R or S-methyl mandelate as substrate. For these experiments 50–300 mg of the immobilized preparations were added to 1–2 mL of 50 mM substrate in 50 mM sodium phosphate at pH 7 and 25 °C under continuous stirring. The conversion degree was analyzed by RP-HPLC (JASCO PU-2085 coupled with a UV-1575 Intelligent UV/VIS Detector) using a Kromacil C18 (15 cm × 0.46 cm) column. Samples (20 μL) were injected and eluted at a flow rate of 1.0 mL min⁻¹ using acetonitrile/10 mM ammonium acetate (35 : 65, v/v) at pH 2.8 as mobile phase and UV detection was performed at 230 nm. The ester presented a retention time of 4.2 minutes while the acid had a retention time of 2.4 minutes. One unit of enzyme activity was defined as the amount of enzyme necessary to produce 1 μmol of mandelic acid per minute under the condition described above. Activity was determined by triplicate with a maximum conversion of 20–30%, and data are given as average values.

2.6. SDS-PAGE analysis

SDS-polyacrylamide gel electrophoresis was performed according to Laemmli⁷² using a Miniprotean tetra-cell (Bio-Rad), 14% running gel in a separation zone of 9 cm × 6 cm, and a concentration zone of 5% polyacrylamide. To analyze the amount of proteins adsorbed on supports, a sample of 100 mg of the support was re-suspended in 1 mL of rupture buffer (2% SDS and 10% mercaptoethanol), boiled for 8 min and a 10 μL aliquot of the supernatant was used in the experiments. The samples were run at 80 volts until the lowest marker reached the lower edge of the gel. Gels were stained with Coomassie brilliant blue. A low molecular weight calibration kit for SDS electrophoresis (GE Healthcare) was used as a molecular weight marker (14.4–97 kDa).

2.7. Textural features of the different supports

The textural characterization of the supports was determined by Mercury Intrusion Porosimetry (MIP). In this technique approximately 0.1 g of sample, previously dried overnight at 60 °C, was accurately weighed into a sample holder and placed in a low-pressure porosimeter (Pascal 140, Thermo Scientific). The sample was then outgassed to a vacuum of 0.1 kPa and flooded with mercury. The pressure over the mercury was then slowly increased from vacuum to 400 kPa, and the intrusion data were collected as a function of the applied pressure. Subsequently, the pressure was reduced to ambient pressure, and the sample holder was removed and weighed before being placed in the high pressure porosimeter (Pascal 240, Thermo Scientific) in which the pressure was raised from atmospheric pressure up to 200 MPa. The combined pressure/volume data was converted into a cumulative pore volume versus pore diameter curve by use of the Washburn equation⁷³ with the recommended values of surface tension (484 dyne per cm) and contact angle (141) for mercury.⁷⁴ Thus, starting under vacuum conditions and increasing the pressure to 200 MPa, the textural characteristics of the materials over a range of approximately 120 μm down to 7.5 nm was determined. Analysis of the data gives rise to the cumulative pore volume, pore size distribution, bulk and skeletal densities. Assuming a cylindrical non-intersecting pore model, the intrusion data also provide an indication of the surface area of the materials by summation of the surface areas of the pore walls at each incremental pressure. However, as the low pressure data was due to the interparticulate porosity of the materials only the data below 1000 nm was considered for the calculation of the pore volume due the intraparticulate porosity within the support material. From the data below 1000 nm the first upward deviation in the cumulative curve is denominated as the “threshold diameter” and may be considered as the limiting diameter below which the porous structure of the support materials becomes accessible. From the derivative of the cumulative intrusion curve the peak maxima indicate the sizes of the most frequent pores in the material.

3. Results and discussion

3.1. Support characterization

Table 1 shows the main textural features of the 5 new supports used in this paper. Results point that all of them has porosity much larger than the lipases size (the largest one if RML with dimensions of: 71.6 Å × 75 Å × 55 Å).⁷⁵ Specific area is quite large for most supports (over 100 m² g⁻¹ for supports styrene and styrene methacrylate), the lower specific area is for support octadecyl methacrylate (25 m² g⁻¹). At first glance, all of them may be suitable for protein immobilization.

Table 1 Textural properties of commercial Purolite® supports. Experimental details are given in Section 2

Support	Pore volumen (cm^3 g^−1)	Bulk density (g cm^3)	Skeletal density (g cm^3)	Porosity (%)	Surface area (m^2 g^−1)	Threshold diameter (mm)	Maximum pore size (nm)
Styrene	1.36 ± 0.06	0.36 ± 0.03	0.91 ± 0.04	60 ± 2	129 ± 7	600 ± 22	108 ± 3
Styrene methacrylate	0.78 ± 0.02	0.55 ± 0.04	1.03 ± 0.03	47 ± 1	105 ± 6	225 ± 8	48 ± 3
Octadecyl methacylate	0.46 ± 0.02	0.66 ± 0.03	1.29 ± 0.05	49 ± 2	56 ± 3	150 ± 5	52 ± 2
DVB methacrylate	0.46 ± 0.03	0.69 ± 0.04	1.10 ± 0.04	38 ± 1	81 ± 5	80 ± 2	29 ± 1
Octadecyl methacrylate	0.19 ± 0.01	0.85 ± 0.05	1.14 ± 0.06	26 ± 1	25 ± 2	140 ± 5	40 ± 2

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3.2. Loading capacity of the different supports

Enzyme loading capacity is determined by the specific area of the support and the pore diameter (must be enough to enable the entry of the enzyme). At first glance, optimal support should be independent on the enzyme utilized provided that the pore is large enough to permit the entry of the enzyme and mainly related to that features (Table 1). However, Table 2 shows that this is not the case. Styrene methacrylate is the support enabling the lowest loading using CALA and CALB. However, immobilizing LU and TLL, octyl agarose offered the lowest loading and using RML the worst results were using 3 of the supports, octyl agarose, DVB methacrylate and octadecyl methacrylate. This suggests that some other factors may be affecting the loading capacity of the supports.

Table 2 Loading capacity of the different studied supports. Experiments were performed as described in Section 2., using the adequate volumes of 0.2 mg protein mL⁻¹ solution of the different commercial lipases in 5 mM sodium phosphate at pH 7 and 25 °C. Loading capacity is given in mg of protein per g of support

Supports	Lipases
	CALA	CALB	Lecitase Ultra	RML	TLL
Octyl agarose	9 ± 0.53	8 ± 0.16	4 ± 0.19	7 ± 0.07	5 ± 0.22
Macroporous styrene	5 ± 0.24	11 ± 0.61	25 ± 0.37	31 ± 0.56	12 ± 0.02
Styrene methacrylate	0.5 ± 0.01	4 ± 0.18	8 ± 0.45	24 ± 0.39	6 ± 0.22
Octadecyl methacylate	10 ± 0.41	13 ± 0.21	21 ± 0.62	47 ± 0.43	17 ± 0.83
DVB methacrylate	10 ± 0.14	10 ± 0.20	17 ± 0.37	16 ± 0.83	13 ± 0.30
Octadecyl methacrylate	5 ± 0.25	9 ± 0.19	14 ± 0.21	20 ± 0.39	10 ± 0.44

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The support with the highest loading also changed when changing the enzyme. Using CALA, differences are not clear using octyl agarose, octadecyl methacylate or DVB methacrylate. Using CALB, octadecyl methacylate allowed the highest loading, shortly followed by styrene and DVB methacrylate. For LU, styrene was the best matrix, followed by octadecyl methacylate and by DVB methacrylate. In the case of RML, octadecyl methacylate enabled the maximum loading, followed by styrene. TLL reached maximum loading when using octadecyl methacylate followed by DVB methacrylate.

Interestingly, except for CALA, always there are some supports that permit a significant higher loading than the reference support (octyl agarose): with CALB the best new support permitted a 50% higher loading, 6 folds using LU, almost 7 times using RML and more than 3 folds using TLL, although the best support regarding loading capacity is not the same for all enzymes as discussed above. Among this diversity, it is remarkable that octadecyl methacylate is in all cases first or second in loading capacity, styrene or DVB methacrylate are in one of these positions in 3 cases.

The differences may be explained considering several facts. It has been previously shown that hydrophobicity is not the only feature that produces lipase adsorption (e.g., pancreatic lipase immobilization in phenyl but not octyl supports).⁵² Moreover, there are a competition between lipase–lipase interaction and interaction of the lipase with the support.^40–42 Finally, the immobilization rate of the enzyme in the support may produce different packing of the lipases: if it is very rapid, distance between lipases may be very small, while if immobilization is slow enough, the distance between lipase molecules may be similar to the protein size.^76–80

3.3. Thermal stability of the different lipase preparations

Table 3 shows the stability studies of all enzyme preparations at different pH values, showing the residual activity after specific incubation times (those where the octyl-agarose gave 40–60% using pNPB as substrate). This strategy was used due to the extremely low activity found using pNPB of the enzymes immobilized on the new supports, low activity that as not related to adsorption of the substrate to the matrix. This result was also obtained using styrene/divinylbenzene.⁴⁹

Table 3 Residual activity (%) of the different studied lipase biocatalysts after thermal inactivation at different pH values and in the presence of organic cosolvents. Experiments have been performed as described in Section 2., determining the residual activity using S-methyl mandelate as substrate

Lipase	Enzymatic derivative	pH 5	pH 7	pH 9	Acetonitrile	Dioxane
CALA	Octyl agarose	24 ± 1.1	42 ± 1.4	78 ± 2.1	65 ± 2.0	55 ± 2.2
	Styrene	36 ± 1.6	44 ± 1.1	73 ± 2.6	52 ± 1.1	42 ± 1.3
	Styrene methacrylate	56 ± 1.8	57 ± 1.5	75 ± 2.0	57 ± 1.9	55 ± 1.7
	Octadecyl methacylate	14 ± 0.5	29 ± 0.6	54 ± 1.9	19 ± 0.3	28 ± 0.9
	DVB methacrylate	9 ± 0.4	12 ± 0.6	51 ± 2.1	15 ± 0.4	19 ± 0.7
	Octadecyl methacrylate	28 ± 0.9	42 ± 1.0	53 ± 1.5	31 ± 1.3	43 ± 1.6
CALB	Octyl agarose	18 ± 0.8	12 ± 0.3	11 ± 0.2	15 ± 0.6	15 ± 0.8
	Styrene	19 ± 1.0	31 ± 0.8	49 ± 1.8	6 ± 0.3	5 ± 0.6
	Styrene methacrylate	24 ± 0.6	25 ± 0.6	19 ± 0.5	12 ± 0.7	22 ± 1.8
	Octadecyl methacylate	25 ± 0.9	49 ± 1.3	60 ± 2.2	24 ± 0.9	15 ± 0.9
	DVB methacrylate	15 ± 0.4	16 ± 0.2	29 ± 0.7	9 ± 0.6	9 ± 0.5
	Octadecyl methacrylate	52 ± 1.9	79 ± 2.6	77 ± 2.4	13 ± 0.5	15 ± 0.9
	Novozym® 435	27 ± 1.1	51 ± 1.8	75 ± 2.9	16 ± 1.8	21 ± 1.0
LU	Octyl agarose	74 ± 2.8	79 ± 2.1	46 ± 1.7	73 ± 3.1	67 ± 1.9
	Styrene	41 ± 1.9	80 ± 3.8	43 ± 2.2	66 ± 1.5	64 ± 1.8
	Styrene methacrylate	76 ± 3.0	63 ± 1.9	42 ± 1.5	62 ± 1.9	59 ± 1.1
	Octadecyl methacylate	59 ± 2.2	79 ± 3.1	40 ± 1.8	75 ± 2.8	70 ± 1.9
	DVB methacrylate	41 ± 1.5	69 ± 2.4	44 ± 0.7	60 ± 2.0	51 ± 0.9
	Octadecyl methacrylate	71 ± 3.1	71 ± 2.7	58 ± 1.6	48 ± 1.1	43 ± 0.3
RML	Octyl agarose	35 ± 1.2	76 ± 3.5	54 ± 1.4	72 ± 2.2	67 ± 2.9
	Styrene	49 ± 1.0	66 ± 2.1	42 ± 0.8	49 ± 1.5	25 ± 0.6
	Styrene methacrylate	54 ± 1.7	76 ± 2.9	34 ± 1.0	47 ± 0.3	30 ± 0.9
	Octadecyl methacylate	24 ± 0.5	49 ± 2.0	43 ± 1.7	44 ± 1.3	27 ± 0.2
	DVB methacrylate	31 ± 0.7	65 ± 2.2	60 ± 2.5	41 ± 0.2	51 ± 1.7
	Octadecyl methacrylate	33 ± 0.2	58 ± 1.7	56 ± 1.6	55 ± 1.5	26 ± 0.3
	Lipozyme® RM IM	38 ± 0.9	63 ± 3.0	56 ± 2.2	52 ± 0.9	51 ± 2.1
TLL	Octyl agarose	41 ± 1.7	79 ± 3.3	48 ± 2.0	53 ± 1.8	22 ± 1.3
	Styrene	55 ± 1.2	77 ± 1.9	52 ± 1.5	41 ± 1.1	52 ± 3.4
	Styrene methacrylate	40 ± 2.0	75 ± 3.7	48 ± 1.9	28 ± 0.8	4 ± 0.3
	Octadecyl methacylate	24 ± 0.9	87 ± 3.1	69 ± 2.9	23 ± 0.9	50 ± 2.2
	DVB methacrylate	49 ± 1.2	59 ± 1.4	76 ± 3.3	49 ± 2.3	57 ± 1.6
	Octadecyl methacrylate	51 ± 1.7	88 ± 3.0	53 ± 2.1	41 ± 1.4	37 ± 1.1
	Lipozyme® TL	20 ± 0.8	82 ± 3.3	27 ± 1.0	9 ± 0.6	4 ± 0.2

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After 24 h of incubation of the enzyme preparations at 4 °C, their residual activities were determined using S methyl mandelate as described in methods section.

At pH 7, the most stable CALA preparation was styrene methacrylate (57% residual activity). Styrene CALA, octyl agarose CALA and octadecyl methacrylate CALA presented similar stabilities (just over 40% residual activities) and were the second most stable ones, while DVB methacrylate CALA was the least stable (12% residual activity). Moving to CALB, octadecyl methacrylate CALB (79% residual activity) was the only with higher residual activity compared to Novozym 435 (51%). The lowest stability using CALB biocatalysts were octyl agarose (12%) and DVB methacrylate. In the case of LU, octyl agarose LU and styrene LU were the most stable (80% residual activity) while styrene methacrylate was the less stable (63%). RML was most stable when immobilized on styrene methacrylate or octyl agarose (76% residual activity), and the lowest stability was observed using octadecyl methacrylate (49%), while the commercial RML IM retained 63% of the activity. Octadecyl methacrylate and octadecyl methacylate TLL were the most stable TLL preparations (87–88% remnant activity) while TLL IM presented 82% residual activity and octyl agarose TLL kept 79%. The least stable TLL preparation at this pH was DVB methacrylate.

The situation changes when the inactivation pH was altered, although not for all enzyme with the same intensity. Using CALA, the most stable preparation at pH 5 and 9 remained styrene methacrylate CALA, although at pH 9 octyl agarose CALA was very similar. The second most stable was styrene at both pH values, at pH 9 was near the value of styrene methacrylate CALA. The preparation with the lower stabilities also was similar, DVB methacrylate CALA, although at pH 9 octadecyl methacylate and octadecyl methacrylate CALA presented very similar residual activity.

Employing CALB, the results at pH 5 and 9 were not qualitatively very different to the results at pH 7. The most and the least stable preparations were the same, although slight differences could be appreciated. In the case of LU, differences were clear when the inactivation pH was altered. At pH 5, styrene LU moved from the most stable to be among the least stable preparations. At pH 9, octadecyl methacrylate LU was the most stable preparation.

Styrene RML remained the most stable one at pH 5, while at pH 8 the most stable RML preparation was DVB methacrylate RML. Octadecyl methacylate RML remained the least stable at pH 5, but styrene was less stable at pH 9.

TLL–styrene was the most stable TLL-preparation at pH 5 and DVB methacrylate TLL was the most stable one at pH 9. The least stable preparations also depended on the pH, at pH 5 was octadecyl methacylate TLL and at pH 9 was the commercial IM-TLL.

Qualitative differences in stability when changing the pH value were clear, but usually not very large. Considering that the immobilization is via interfacial activation and the orientation should be similar, the changes may derived from some ionic groups that could be introduced on the resins during preparation (e.g., some negative charges due to oxidations) or that different movements on the enzyme structure are produced under different conditions⁸¹ and can favored/difficulty some positive/negative enzyme-support interactions.⁸²

3.4. Solvent stability of the different lipase biocatalyst

Next, we have analyzed the stability of the different enzyme preparations in two organic cosolvents, acetonitrile and dioxane (Table 3). In most cases, the qualitative values were coincident using both solvents. Using CALA, the most stable preparations were those prepared using octyl agarose and styrene, similar to the results found in thermal inactivations. The least stable CALA preparation was that prepared using DVB methacrylate. Using LU, the most stable preparations were octadecyl methacylate and octyl agarose LU, and the least stable one was octadecyl methacrylate LU. Using TLL, DVB methacrylate was the most stable catalyst and IM-TLL the least stable one. Using acetonitrile, TLL–octyl agarose presented a similar high stability (but no in dioxane, where it was among the least stable ones).

With RML and CALB the situation is not so similar when changing the organic solvent. In the case of CALB, the most stable preparation was octadecyl methacylate in acetonitrile and styrene in dioxane. The least stable preparations were styrene and DVB methacrylate CALB. Octyl agarose RML was the most stable preparation in both solvents, but DVB methacrylate and octadecyl methacylate RML were the least stable ones in acetonitrile; the same behavior was observed with styrene, octadecyl methacrylate and octadecyl methacylate RML in dioxane. These differences may be related to changes in the inactivation route when changing the solvent,⁸¹ but also with a different release of the enzyme to the reaction media caused by the different conditions,⁸³ and even a different change in the swelling properties of the different solvents that could really alter the support internal geometry.

3.5. Analysis of enzyme desorption during thermal and solvent inactivation

One of the problems related to the immobilization of lipases on hydrophobic supports is the desorption of the enzymes to the medium after enzyme during incubation under drastic conditions.⁸³ Fig. 1–5 show the analysis of the enzyme-support via SDS-PAGE after inactivation under different conditions. The lower the intensity of the protein band compared to the protein band before inactivation, the lower the amount of enzyme that remain attached to the support.

Fig. 1 SDS-PAGE analysis of different CALB-biocatalysts preparations after thermal inactivation at different pH values and in the presence of organic cosolvents. Experiments have been performed as described in Section 2. The gel shows the enzyme that remains bound to the support after inactivation. Panels (a) octyl agarose, (b) styrene, (c) styrene methacrylate, (d) octadecyl methacylate, (e) DVB methacrylate, (f) octadecyl methacrylate, and (g) Novozym 435. Lane 1: molecular weight marker, lane 2: commercial free CALB, lane 3: CALB-biocatalyst, lane 4: incubation at pH 7, lane 5: incubation at pH 5, lane 6: incubation at pH 9, lane 7: incubation in 50% acetonitrile, and lane 8: incubation in 80% dioxane.

Fig. 2 SDS-PAGE analysis of different CALA-biocatalysts preparations after thermal inactivation at different pH values and in the presence of organic cosolvents. Experiments have been performed as described in Section 2. Panels (a) octyl agarose, (b) styrene, (c) styrene methacrylate, (d) octadecyl methacylate, (e) DVB methacrylate, and (f) octadecyl methacrylate. Lane 1: molecular weight marker, lane 2: commercial free CALA, lane 3: CALA-biocatalyst, lane 4: incubation at pH 7, lane 5: incubation at pH 5, lane 6: incubation at pH 9, lane 7: incubation in 50% acetonitrile, and lane 8: incubation in 80% dioxane.

Fig. 3 SDS-PAGE analysis of different LU-biocatalysts after thermal inactivation at different pH values and in the presence of organic cosolvents. Experiments have been performed as described in Section 2. Panels (a) octyl agarose, (b) styrene, (c) styrene methacrylate, (d) octadecyl methacylate, (e) DVB methacrylate, and (f) octadecyl methacrylate. Lane 1: molecular weight marker, lane 2: commercial free Lecitase Ultra, lane 3: LU-biocatalyst, lane 4: incubation at pH 7, lane 5: incubation at pH 5, lane 6: incubation at pH 9, lane 7: incubation in 30% acetonitrile, and lane 8: incubation in 60% dioxane.

Fig. 4 SDS-PAGE analysis of different RML-biocatalysts preparations after thermal inactivation at different pH values and in the presence of organic cosolvents. Experiments have been performed as described in Section 2. Panels (a) octyl agarose, (b) styrene, (c) styrene methacrylate, (d) octadecyl methacylate, (e) DVB methacrylate, (f) octadecyl methacrylate, and (g) Lipozyme RM IM. Lane 1: molecular weight marker, lane 2: commercial free RML, lane 3: RML-biocatalyst, lane 4: incubation at pH 7, lane 5: incubation at pH 5, lane 6: incubation at pH 9, lane 7: incubation in 30% acetonitrile, and lane 8: incubation in 35% dioxane.

Fig. 5 SDS-PAGE analysis of different TLL-biocatalysts preparations after thermal inactivation at different pH values and in the presence of organic cosolvents. Experiments have been performed as described in Section 2. Panels (a) octyl agarose, (b) styrene, (c) styrene methacrylate, (d) octadecyl methacylate, (e) DVB methacrylate, (f) octadecyl methacrylate, and (g) Lipozyme TL. Lane 1: molecular weight marker, lane 2: commercial free TLL, lane 3: TLL-biocatalyst, lane 4: incubation at pH 7, lane 5: incubation at pH 5, lane 6: incubation at pH 9, lane 7: incubation in 50% acetonitrile, and lane 8: incubation in 60% dioxane.

Styrene methacrylate is the support that has the lowest amount of enzyme in all cases fitting the previous results (Table 2). Moreover, it was the support that retained lest enzyme in all cases, suggesting that is the one with lower affinity for the assayed lipases. CALA and RML cannot be detected on the support after the incubation in all the inactivation conditions assayed.

Results were different when changing the enzyme, although some enzyme release could be observed from all supports and enzymes at least in some conditions. CALB in thermal inactivations seem to release a low amount of enzyme except for styrene methacrylate. In organic solvent inactivations, all preparations lost most of the enzyme, without significant improvements comparing to the results obtained using Novozym 435 even although stabilities were significantly improved in certain cases. Using TLL, several of the new supports offered higher enzyme retention compared to the octyl support or the IM-TLL preparation (although the immobilization in this case follows a different method), mainly in organic solvent inactivations. Lecitase is the enzyme that more strongly immobilized on all supports, although some enzyme release under certain conditions may be appreciated using divinylbenzene or octadecyl methacrylate. Thus, in some instances the new supports permit a better retention of the enzyme that the commercial octyl agarose or the commercial preparations, while in other case not. And that it is not fully correlated to the stability results.

3.6. Activity and specificity of the different enzyme immobilized on the different supports

The activity of the entire immobilized enzyme versus pNPB was almost fully suppressed except for octyl-agarose (results not shown). This result coincides with that obtained using styrene/divinylbenzene beads.⁴⁹ However, when using other substrates, the immobilized enzymes exhibited very high activity, in many instances overpassing those of the commercial preparations or the octyl-immobilized lipases.

3.6.1. Hydrolysis of triacetin. Table 4 shows the activity of all biocatalysts versus triacetin at pH 5 conditions adequate to produce 1,2 diacetin without suffering acyl migration⁸⁴ using the enzyme preparations at maximum loading.

Table 4 Activity of different lipase biocatalyst versus triacetin (100 mM) at pH 5 and 25 °C. Experiments were performed as described in Section 2. The activity (V) is given in μmoles of substrate hydrolyzed per min and per mg of immobilized enzyme and also and also per g of biocatalyst

Lipase	Enzymatic derivative	V (U mg^−1 protein)	V (U g^−1 enzymatic derivative)
CALA	Octyl agarose	0.0147 ± 0.0005	0.132 ± 0.0051
	Styrene	0.0146 ± 0.0005	0.071 ± 0.0032
	Styrene methacrylate	0.7104 ± 0.0322	0.341 ± 0.0156
	Octadecyl methacylate	0.0106 ± 0.0012	0.109 ± 0.0052
	DVB methacrylate	0.0422 ± 0.0019	0.438 ± 0.0214
	Octadecyl methacrylate	0.0003 ± 0.00001	0.002 ± 0.0001
CALB	Octyl agarose	0.69 ± 0.005	5.34 ± 0.04
	Styrene	0.27 ± 0.012	3.00 ± 0.12
	Styrene methacrylate	1.13 ± 0.055	4.96 ± 0.22
	Octadecyl methacylate	0.44 ± 0.019	5.69 ± 0.19
	DVB methacrylate	0.37 ± 0.016	3.71 ± 0.16
	Octadecyl methacrylate	0.44 ± 0.012	4.04 ± 0.12
	Novozym® 435	0.06 ± 0.002	5.21 ± 0.19
LU	Octyl agarose	0.020 ± 0.0009	0.079 ± 0.004
	Styrene	0.010 ± 0.0003	0.254 ± 0.012
	Styrene methacrylate	0.008 ± 0.0002	0.058 ± 0.002
	Octadecyl methacylate	0.079 ± 0.0031	1.665 ± 0.072
	DVB methacrylate	0.008 ± 0.0002	0.134 ± 0.006
	Octadecyl methacrylate	0.122 ± 0.0058	1.651 ± 0.062
RML	Octyl agarose	0.65 ± 0.008	4.76 ± 0.06
	Styrene	0.13 ± 0.005	3.87 ± 0.15
	Styrene methacrylate	0.14 ± 0.007	3.49 ± 0.16
	Octadecyl methacylate	0.29 ± 0.014	13.6 ± 0.65
	DVB methacrylate	0.10 ± 0.004	1.52 ± 0.07
	Octadecyl methacrylate	0.63 ± 0.032	12.5 ± 0.53
	Lipozyme® RM IM	0.03 ± 0.002	1.40 ± 0.17
TLL	Octyl agarose	0.50 ± 0.0247	2.25 ± 0.10
	Styrene	0.33 ± 0.0165	4.16 ± 0.11
	Styrene methacrylate	1.02 ± 0.0408	5.55 ± 0.27
	Octadecyl methacylate	1.16 ± 0.0483	19.15 ± 0.86
	DVB methacrylate	0.30 ± 0.0146	3.71 ± 0.15
	Octadecyl methacrylate	1.01 ± 0.0503	9.99 ± 0.46
	Lipozyme® TL	0.02 ± 0.003	1.43 ± 0.16

---

Using CALA, the highest activity per gram of biocatalyst was obtained using DVB methacrylate, 3.3 folds higher than using octyl-CALA. Styrene methacrylate permitted also a significant higher activity than octyl agarose using this enzyme and substrate. On the other side, octadecyl methacrylate was the least active (hundreds of folds). Considering the activity per mg of protein, that show how the immobilization affects the properties of the enzyme, styrene methacrylate permitted to achieve the highest specific activity (it was the one with lowest loading capacity for this enzyme) followed by DVB methacrylate (but this was 15 fold lower). The enzyme immobilized on octyl agarose had a specific activity around 3 fold lower than using DVB methacrylate.

Using CALB differences are not so large. Octadecyl methacylate CALB gave an activity per g of biocatalyst slightly higher than octyl-agarose CALB and Novozym 435, the enzyme immobilized in macroporous styrene showed the lowest activity (slightly over 50% of the activity of octadecyl methacylate CALB). Again styrene methacrylate gave the highest CALB specific activity, being the styrene CALB the one exhibiting the lowest specific activity.

In the case of LU, the immobilization on octadecyl methacylate and methacrylate gave the higher activity per g of catalyst, styrene methacrylate LU was the least activity shortly followed by octyl agarose LU. Considering the mg of enzyme, LU immobilized on octadecyl methacrylate presented clearly more specific activity than when immobilized on octadecyl methacylate and both were well over the other ones (6–15 folds).

Octadecyl methacrylate RML was almost 9 folds more active than the commercial preparation, and almost 3 folds more active than octyl agarose RML. The second least active was the DVD methacrylate. Considering the activity per mg of enzyme, RML immobilized on octyl agarose and on octadecyl methacrylate exhibited similar activities.

TL IM was the least active one, and was more than 13 folds less active than the most active biocatalyst, octadecyl methacylate TLL. The second most active had 50% of the activity and was octadecyl methacrylate, while octyl agarose TLL was the second least active. Regarding the specific activity of TLL, styrene methacrylate, octadecyl methacylate and octadecyl methacrylate presented a similar activity, doubling that obtained using octyl agarose.

Thus, considering the hydrolysis of triacetin, there are no a single “optimal” support to immobilize the enzymes of this study; results are different for each enzyme and sometimes it is not related to the maximum loading, as specific activity of the immobilized enzyme changes when changing the support. Thereby, the support producing the maximum activity using the lowest amount of enzyme should be selected, e.g., octadecyl methacrylate LU should be more adequate than octadecyl methacylate because the activity per g of support is similar, but the first has lower enzyme loading than the second biocatalyst.

3.6.2. Hydrolysis of methyl phenylacetate. Results of the hydrolysis of methyl phenylacetate are shown in Table 5. DVD methacrylate CALA and styrene methacrylate CALA were around twice more active than octyl agarose CALA. Styrene and octadecyl methacrylate were the least active ones, about four folds, this was the main difference comparing the results obtained triacetin, where octadecyl methacrylate was clearly the least active one. In specific activity, CALA was clearly the most active when immobilized on styrene methacrylate and the least active when immobilized on styrene and octadecyl methacylate.

Table 5 Activity of different lipase biocatalyst versus methyl phenylacetate (20 mM) at pH 7 and 25 °C. Experiments were performed as described in Section 2. The activity (V) is given in μmoles of substrate hydrolyzed per mg of immobilized enzyme and also per g of biocatalyst

Lipase	Enzymatic derivative	V (U mg^−1 enzyme)	V (U g^−1 enzymatic derivative)
CALA	Octyl agarose	0.008 ± 0.0002	0.08 ± 0.004
	Styrene	0.007 ± 0.0003	0.04 ± 0.001
	Styrene methacrylate	0.276 ± 0.0134	0.13 ± 0.006
	Octadecyl methacylate	0.008 ± 0.0003	0.08 ± 0.004
	DVB methacrylate	0.016 ± 0.0002	0.17 ± 0.002
	Octadecyl methacrylate	0.011 ± 0.0003	0.05 ± 0.002
CALB	Octyl agarose	13.3 ± 0.6	103 ± 5
	Styrene	7.78 ± 0.4	87 ± 4
	Styrene methacrylate	7.26 ± 0.3	32 ± 2
	Octadecyl methacylate	12.8 ± 0.6	166 ± 8
	DVB methacrylate	14.9 ± 0.7	149 ± 7
	Octadecyl methacrylate	15.2 ± 0.8	141 ± 7
	Novozym® 435	2.48 ± 0.1	204 ± 9
LU	Octyl agarose	0.009 ± 0.0004	0.04 ± 0.002
	Styrene	0.106 ± 0.0053	2.60 ± 2.131
	Styrene methacrylate	0.299 ± 0.0148	2.26 ± 0.112
	Octadecyl methacylate	0.004 ± 0.0001	0.09 ± 0.004
	DVB methacrylate	0.044 ± 0.0022	0.76 ± 0.033
	Octadecyl methacrylate	0.001 ± 0.0001	0.02 ± 0.0008
RML	Octyl agarose	0.24 ± 0.009	1.77 ± 0.06
	Styrene	0.32 ± 0.014	9.80 ± 0.33
	Styrene methacrylate	0.63 ± 0.004	15.4 ± 0.10
	Octadecyl methacylate	0.23 ± 0.011	10.8 ± 0.52
	DVB methacrylate	0.43 ± 0.021	6.61 ± 0.30
	Octadecyl methacrylate	0.30 ± 0.013	5.99 ± 0.27
	Lipozyme® RM IM	0.63 ± 0.031	29.7 ± 1.43
TLL	Octyl agarose	0.011 ± 0.0003	0.05 ± 0.001
	Styrene	<0.011	<0.05
	Styrene methacrylate	<0.011	<0.05
	Octadecyl methacylate	<0.011	<0.05
	DVB methacrylate	<0.011	<0.05
	Octadecyl methacrylate	<0.011	<0.05
	Lipozyme® TL	<0.011	<0.05

---

Novozym 435 was the most active CALB biocatalyst with this substrate, a 20% more than octadecyl methacylate or a 25% than DVD methacrylate CALB or octadecyl methacrylate. The least active CALB biocatalyst was styrene CALB. Agarose octyl CALB activity was 50% of that of Novozyme 435, and it was more active than styrene CALB and the least active one, styrene methacrylate CALB. In specific activity of CALB, octadecyl methacrylate, DVD methacrylate and octyl agarose gave similar values, shortly followed by octadecyl methacylate and doubling the values of styrene and styrene methacrylate.

Using LU, the most active biocatalyst was styrene LU, that was 65 folds more active that octyl agarose. Styrene methacrylate LU was the second most active (15% less active) while the least active was octadecyl methacrylate LU (half the activity of octyl agarose LU). The picture was very different to that obtained with triacetin (Table 2). Regarding the specific activity of LU, styrene methacrylate permitted the highest one, more than 30 folds that obtained using octyl agarose. The lowest one was observed using octadecyl methacrylate.

RML biocatalysts gave also very different results using this substrate compared to triacetin. IM RML was the most active with this substrate, doubling the activity of the second one, styrene methacrylate RML. The least active biocatalyst was octyl agarose LU, almost 17 folds less active than the commercial one. Regarding RML specific activity, styrene methacrylate gave the highest values, 2.5 folds that obtained using octyl agarose. Only octadecyl methacylate gave values similar to octyl agarose.

TLL only gave significant activity versus this substrate using octyl agarose TLL.

3.6.3. Hydrolysis of methyl mandelate. Table 6 shows the results with the last substrate, methyl mandelate. This is the complex one, a chiral molecule, and we have also included the activities ratios with both isomers. We supply the results used the S isomer, and the activity with S/activity with R ratio.

Table 6 Activity of different lipase biocatalyst versus R- or S-methyl mandelate (50 mM) at pH 7 and 25 °C. Experiments were performed as described in Section 2. The activity (V) is given in μmoles of substrate hydrolyzed per mg of immobilized enzyme and also per g of biocatalyst

Lipase	Enzymatic derivative	V (U mg^−1 enzyme)	V (U g^−1 enzymatic derivative)	VS/VR
CALA	Octyl agarose	0.13 ± 0.004	1.15 ± 0.05	1.51
	Styrene	1.09 ± 0.053	5.28 ± 0.16	2.08
	Styrene methacrylate	5.33 ± 0.232	2.56 ± 0.12	1.20
	Octadecyl methacylate	0.35 ± 0.015	3.61 ± 0.16	1.30
	DVB methacrylate	0.67 ± 0.032	6.98 ± 0.34	1.00
	Octadecyl methacrylate	0.39 ± 0.016	1.98 ± 0.06	1.61
CALB	Octyl agarose	10.3 ± 0.26	80.33 ± 2.00	0.17
	Styrene	5.93 ± 0.28	66.68 ± 2.26	0.43
	Styrene methacrylate	7.24 ± 0.34	31.77 ± 1.38	0.33
	Octadecyl methacylate	7.10 ± 0.32	91.76 ± 3.27	0.35
	DVB methacrylate	9.90 ± 0.39	99.38 ± 4.95	0.50
	Octadecyl methacrylate	6.96 ± 0.33	64.56 ± 2.23	0.30
	Novozym® 435	1.44 ± 0.05	118.3 ± 5.81	0.63
LU	Octyl agarose	0.28 ± 0.013	1.14 ± 0.05	5.18
	Styrene	0.25 ± 0.012	6.16 ± 0.28	5.76
	Styrene methacrylate	0.40 ± 0.016	2.99 ± 0.11	2.99
	Octadecyl methacylate	0.14 ± 0.005	2.88 ± 0.13	1.87
	DVB methacrylate	0.20 ± 0.008	3.46 ± 0.16	2.07
	Octadecyl methacrylate	0.20 ± 0.006	2.67 ± 0.12	5.24
RML	Octyl agarose	0.16 ± 0.004	1.20 ± 0.04	3.87
	Styrene	0.26 ± 0.011	7.90 ± 0.32	10.82
	Styrene methacrylate	0.19 ± 0.006	4.61 ± 0.22	13.97
	Octadecyl methacylate	0.12 ± 0.003	5.51 ± 0.23	1.94
	DVB methacrylate	0.39 ± 0.015	6.07 ± 0.23	3.25
	Octadecyl methacrylate	0.19 ± 0.003	3.71 ± 0.17	2.23
	Lipozyme® RM IM	0.04 ± 0.002	2.07 ± 0.09	3.00
TLL	Octyl agarose	0.28 ± 0.012	1.26 ± 0.06	5.25
	Styrene	0.15 ± 0.003	1.90 ± 0.08	1.09
	Styrene methacrylate	0.83 ± 0.022	4.51 ± 0.13	1.80
	Octadecyl methacylate	0.16 ± 0.008	2.55 ± 0.11	4.11
	DVB methacrylate	0.13 ± 0.002	1.61 ± 0.07	2.06
	Octadecyl methacrylate	0.24 ± 0.009	2.35 ± 0.08	1.48
	Lipozyme® TL	0.01 ± 0.001	1.24 ± 0.05	4.59

---

DVB methacrylate (7 U g⁻¹) and styrene (5.3 U g⁻¹) CALA biocatalysts were the most active ones, octyl agarose CALA was the least active biocatalysts, 6 folds less active. Octadecyl methacrylate CALA has 1.8 fold more activity than this one (in opposition with the results obtained using triacetin). CALA specific activity was the highest using styrene methacrylate, 5 folds more than using styrene; octyl agarose have the lowest specific activity. Most enzyme preparations preferred the hydrolysis of the S isomer, but the activity ratio with the R isomer was very small (a maximum of 2 using styrene) and for DVB methacrylate was identical.

Novozyme 435 was the most active CALB biocatalyst (120 U g⁻¹), followed by DVD methacrylate CALB (100 U g⁻¹) and for octadecyl methacrylate CALB (92 U g⁻¹). The least active preparation was styrene methacrylate (32 U g⁻¹). CALB highest specific activity was obtained using octyl agarose and DVD methacrylate, the lowest one was obtained using styrene (around 70% of the maximum). This enzyme preferred the R isomer, and octyl agarose CALB gave the best ratio (around 6) while Novozyme 435 gave the lowest one (less than 1.6).

Styrene LU was the most active preparation of LU with this substrate, almost doubling the second (DVB methacrylate), being octyl agarose LU the least active one (more than 5 times less active than the most active one). Styrene methacrylate produced the highest specific activity for this enzyme and substrate pair, while octadecyl methacrylate produced the lowest (almost 3 folds lower). All preparations prefer the isomer S, with activities ratios ranging from more than 5 (using octyl agarose LU, styrene LU or octadecyl methacylate LU) to around 2 (octadecyl methacrylate LU and DVB methacrylate LU).

IM RML was 40% more active than octyl agarose RML (the least active one), but almost 4 times less active than styrene RML, the most active one. DVB methacrylate RML was the second most active (75% of the activity of styrene RML), shortly followed by octadecyl methacrylate. RML specific activity was the highest using DVD methacrylate and the lowest using octadecyl methacrylate. Regarding the activity of both isomers ratio, the enzyme preferred the S isomer and values ranged from 14 using styrene RML to less than 2 using octadecyl methacrylate.

Using TLL biocatalyst, the highest activity was observed using styrene methacrylate TLL, almost doubling the activity of the second (octadecyl methacrylate TLL), while the least active ones were the commercial IM TLL and octyl agarose TLL (around 3.5 folds). The highest specific activity was observed using styrene methacrylate and almost 3 folds that of the second one (octyl agarose). All preparations preferred the S isomers, with activity ratios using both isomers ranging from around 5 (octyl agarose TLL) to 1.1 (using styrene TLL).

3.6.4. Discussion. The results above are a new and clear example on how changing the support the enzyme stability, activity and specificity may be strongly modulated even using similar immobilization protocol. This is directly visualized using R and S mandelic acids, but the comparison between all substrates show the strong alterations on enzyme activity and specificity. Moreover it is not possible to point an “optimal” supports for lipase immobilization that depends on the enzyme and the substrate. In some cases, a support and lipase pair is among the most active ones with some of the substrates, while is poorly active with other. It should be remarked the lack of significant activity using the new preparations and pNPB (we have discarded substrate adsorption by adding the support to the reaction mixture and after some time free enzyme, where we cannot visualize any alteration of the enzyme activity at short times). Thus, using only the conventional pNPB assay, these supports could be discarded, when with other substrate produced catalyst with activities far better than the commercial ones or octyl agarose.

3.7. Reuse of the biocatalyst

All the CALB biocatalysts could be reused in hydrolysis of 100 mM triacetin for 3 cycles without a significant decrease of activity during use or washing, even although 20% acetonitrile was present in the medium. This, the new supports showed excellent properties for their industrial implementation.

3.8. Reuse of the support

Another important question is the real possibility of reusing the supports. The reversibility of the immobilization protocol permits this. Thus, we washed the enzyme preparations 3 folds with 5 volumes 2% triton X-100 at 25 °C. Although most enzyme molecules were released, we could appreciate in some cases (even using the octyl support) that a small percentage of the protein remained immobilized (less than 5%) by SDS-PAGE. By increasing the washing T at 37 °C, we have been able to fully release all the enzyme molecules from the support, even using preparations submitted to previous thermal or solvent inactivation (this may affect the adsorption strength of inactivated enzymes to the support).⁸⁵ The fact that Triton X-100 is able to release most of the enzyme molecules confirm that the main force of the immobilization is a hydrophobic adsorption.

The washed supports presented identical maximum loading capacity and the immobilized enzyme identical stability that the initial preparations.

4. Conclusion

The new supports studied in these paper present very good properties to prepare immobilized lipases. However, there is not a universal optimal support; the “best” support depends on the enzyme, the parameter studied and the substrate used. That means even although all the supports immobilize the lipases via interfacial activation versus the hydrophobic surface of the support, the final properties of the biocatalyst will depend on the support features. For this reason, a large library of methods and supports to immobilized lipases become the only way to ensure the full utilization of the potential of immobilization to improve enzyme properties. The reversible mechanism of immobilization permits to re-use the supports after enzyme inactivation. However, it should be considered, as already described in other publications, that this may reduce the range of applicability: high cosolvent concentrations,^86,87 and even some substrates or products⁸⁸ may produce the release of the enzyme. Simple strategies to solve this problem has been already proposed, enlarging the range of application of this kind of immobilized enzymes.⁸⁹

Nevertheless, under the conditions utilized in this paper, the preparations showed a very good performance in a diversity of reactions and permitted their reuse for several cycles. These supports will enlarge the library of immobilized lipase biocatalyst and that way, they will enhance the prospects of finding suitable biocatalyst for a specific process.³⁴

Acknowledgements

We gratefully recognize the support from the MINECO from Spanish Government, (project numbers CTQ2013-41507-R and CTQ2016-78587-R). The predoctoral fellowships for Miss Peirce (Universita' degli Studi di Napoli Federico II) and Miss Tacias-Pascacio (CONACyT, Mexico) are also gratefully recognized. Dr Virgen-Ortíz thanks CONACyT Mexico for his Postdoctoral fellowship (No. 263815). The authors wish to thank Mr Ramiro Martínez (Novozymes, Spain) for kindly supplying some of the enzymes used in this research and Purolite for the kind donation of the supports. The advice and support of Drs A. Basso and S. Serban (Purolite) are also gratefully recognized.

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Footnote

† Permanent address: Catedrático CONACYT – Centro de Investigación en Alimentación y Desarrollo, A.C., Centro de Innovación y Desarrollo Agroalimentario de Michoacán, A.C., Antigua Carretera a Pátzcuaro s/n, 58341 Morelia, Michoacán, Mexico.

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