Evelin A. Manoel‡
abc,
Martina Pinto‡d,
José C. S. dos Santos‡ce,
Veymar G. Tacias-Pascaciobf,
Denise M. G. Freireb,
José Carlos Pinto*d and
Roberto Fernandez-Lafuente*b
aDepartamento de Biotecnologia Farmacêutica, Faculdade de Farmácia, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil
bDepartamento de Bioquímica, Instituto de Química, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil. E-mail: rfl@icp.csic.es
cDepartment of Biocatalysis, ICP-CSIC, Campus UAM-CSIC, Cantoblanco, 28049, Madrid, Spain
dPrograma de Engenharia Química, COPPE, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil. E-mail: pinto@peq.coppe.ufrj.br
eInstituto de Engenharias e Desenvolvimento Sustentável, Universidade da Integração Internacional da Lusofonia Afro-Brasileira, CEP 62785-000, Acarape, CE, Brazil
fUnidad de Investigación y Desarrollo en Alimentos. Instituto Tecnológico de Veracruz, Calzada Miguel A. de Quevedo 2779, 91897 Veracruz, Mexico
First published on 27th June 2016
Different core–shell polymeric supports, exhibiting different morphologies and composition, were produced through simultaneous suspension and emulsion polymerization, using styrene (S) and divinylbenzene (DVB) as co-monomers. Supports composed of polystyrene in both the core and the shell (PS/PS) and the new poly(styrene-co-divinylbenzene) support (PS-co-DVB/PS-co-DVB) were used for the immobilization of three different lipases (from Rhizomucor miehie (RML), from Themomyces lanuginosus (TLL) and the form B from Candida antarctica, (CALB)) and of the phospholipase Lecitase Ultra (LU). The features of the new biocatalysts were evaluated and compared to the properties of commercial biocatalysts (Novozym 435 (CALB), Lipozyme RM IM and Lipozyme TL IM) and biocatalysts prepared by enzyme immobilization onto commercial octyl-agarose, a support reported as very suitable for lipase immobilization. It was shown that protein loading and stability of the biocatalysts prepared with the core–shell supports were higher than the ones obtained with commercial octyl-agarose or the commercial lipase preparations. Besides, it was shown that the biocatalysts prepared with the core–shell supports also presented higher activities than commercial biocatalysts when employing different substrates, encouraging the use of the produced core–shell supports for immobilization of lipases and the development of new applications.
Lipases are among the most used enzymes in biocatalysis, due to their characteristic wide specificity and the wide range of reactions that these enzymes can catalyze (including many promiscuous reactions).13–15 Besides, lipases show very high enantioselectivity16–19 and are very robust, being successfully employed in different reaction media (e.g., aqueous media, organic solvents, neoteric media).20,21
The active centers of most lipases are secluded from the reaction media by a polypeptide chain (the lid), which is isolated from the medium by the large hydrophobic pocket where the active center is located (closed form).23–27 The lid can move and exposes this hydrophobic pocket to the medium, generating the open and active form of the lipase. This open lipase form readily adsorbs onto hydrophobic surfaces, including oil drops,22,28 hydrophobic supports,29 other open lipase molecules,30,31 other hydrophobic proteins.32
In fact, a strategy that is becoming very popular for the immobilization of lipases is based on the interfacial activation of the enzyme on hydrophobic support surfaces.33 This strategy allows the immobilization, purification and stabilization of the open lipase form (usually leading to hyper-activation), also producing an increase in the lipase stability for this reason.29 It has been reported that the internal morphology or physical properties of the hydrophobic support may tune the lipase properties, including its activity, stability and specificity.34–36 Therefore, there is a great interest in the new development of new hydrophobic matrices that can further improve lipase properties. Among these new materials, core–shell polymeric particles, consisting of “large” particles (core) coated with small nano-particles (shell) may play a pivotal role.37–40
Different techniques have been employed for the production of core–shell particles. Among them, the combined suspension and emulsion polymerization process has special interest. This technique typically comprises two fundamental steps. In the first step (suspension polymerization), the particle cores are synthesized. When the monomer conversion reaches a certain value in the core formation, the second step is initialized. To this goal, the elements of a typical emulsion polymerization are fed into the reaction vessel. The new nanoparticles coagulate over the previously prepared particle cores to form the shell. During the second reaction step, the suspension and emulsion polymerization processes are conducted simultaneously. At the end of the process, micrometric, porous and (in some cases) functionalized polymer particles are obtained.40,41
In the present manuscript, distinct polymeric supports presenting core–shell morphology were produced through simultaneous suspension and emulsion polymerization, using styrene (S) and divinylbenzene (DVB) as co-monomers. S and DVB are hydrophobic monomers, while DVB can also promote chain crosslinking, leading to modification of the morphology and mechanical resistance of the obtained polymer particles. The first step of this study comprised the determination of how the co-monomers feed flow rate of S and DVB affects the specific area and porosity of the synthesized core–shell particles. Afterwards, among the different supports that were produced, one of them was selected for the enzyme immobilization procedure: the support with the highest specific area (PS-co-DVB/PS-co-DVB). The previously described core–shell polystyrene support (PS/PS), that has been successfully employed in the immobilization of the lipase B from Candida antarctica, was also employed for comparison.41
The prepared polymeric supports were used for the immobilization of three lipases: lipases from Rhizomucor miehei, RML,42 from Thermomyces lanuginosus, TLL,43 and the form B from Candida antarctica, CALB.44 While CALB has a very small lid, which does not completely isolate its active center from the reaction medium,45 TLL and RML exhibit very large lids.46,47 The polymeric supports were also used for immobilization of the chimeric artificial phospholipase Lecitase Ultra (LU).48,49 PS/PS core–shell supports had been previously and successfully used for CALB immobilization, but this is the first attempt to immobilize the other enzymes on this support. The high specific area PS-co-DVB/PS-co-DVB support is used to immobilize enzymes for the first time in this paper.
The new biocatalysts were compared with commercial biocatalysts (Novozym 435 (CALB), Lipozyme RM IM and Lipozyme TL IM) and biocatalysts prepared through enzyme immobilization onto commercial octyl-agarose, a very popular support successfully used in many instances for lipase immobilization.33,50 Lipases immobilized on octyl-agarose have been reported to be much more stable than the free enzyme, and even more than some lipases immobilized via multipoint covalent attachment.51,52 This has been explained by the higher stability of the adsorbed open form of the lipase compared to the lipase in conformational equilibrium.53,54 Lipases tend to form dimeric aggregates with altered properties and that may alter the results of the activity and stability studies and suggests that the use of free lipase to compare the properties with immobilized enzymes may not be very adequate.55–59
Styrene supplied by Sigma Aldrich (USA) (minimum purity of 99.5% (wt/wt)) was used as monomer for the production of PS/PS particles. For the production of PS-co-DVB/PS-co-DVB particles, styrene was provided by INOVA (Brazil) and distilled under vacuum before its use and DVB was supplied by Merck (USA).
Other reagents and solvents were of analytical grade and were used as received, without any purification step.
Reactions were carried out in an open 1 L jacketed glass reactor (FGG Equipamentos Científicos, São Paulo, Brazil) equipped with a thermostatic bath (Haake Phoenix II model, Thermo Scientific, Karlsruhe, Germany) that was employed to keep the reactor temperature at 85 °C. For the production of PS/PS particles, styrene was used as the only monomer in the suspension and emulsion processes.60 For the production of PS-co-DVB/PS-co-DVB, copolymerization of styrene (75% (wt/wt) and DVB (25% wt/wt)) was conducted during the suspension and emulsion polymerization steps. For production of PS-co-DVB individual core particles, classic suspension copolymerization was performed, without addition of the emulsion feed.
Initially, 100 g of an organic solution (containing the monomer mixture and 3.8% (wt/wt) of the initiator benzoyl peroxide) were dispersed in 370 g of an aqueous solution (containing distilled water and 0.80% (wt/wt) of poly(vinyl alcohol), used as stabilizer). The dispersion was kept under continuous agitation (950 rpm in the PS/PS reaction and 800 rpm in PS-co-DVB and PS-co-DVB/PS-co-DVB polymerizations) at a constant temperature of 85 °C. After two hours of reaction, the emulsion constituents (the monomer mixture and the aqueous solution, containing distilled water, 0.13% (wt/wt) of the initiator potassium persulfate, 0.13% (wt/wt)% of sodium bicarbonate and 1% (wt/wt) of sodium lauryl sulfate) were added to the reaction medium. 30 g of the monomer mixture and 230 g of the aqueous solution were added as a single load. The remaining 70 g of the monomer mixture were added under continuous flow (0.026 L h−1) in the PS/PS reaction.60 For the production of PS-co-DVB/PS-co-DVB particles, different flow rates were employed, as shown in Table 1. After feeding, two additional hours of reaction were permitted to ensure the appropriate coverage of the core and formation of the shell. Then, the reactor was cooled down to room temperature and the obtained particles were filtrated and washed with cold water. Finally, the obtained polymer particles were dried in a vacuum oven at 30 °C until constant mass. The scheme of the polymerization process is shown in Fig. 1.
Operational condition | ||
---|---|---|
Reaction | Ratio of (S:DVB) | Comonomer feed flow rate (L h−1) |
1 | 3:1 | — |
2 | 3:1 | 0.032 |
3 | 3:1 | 0.076 |
4 | 3:1 | 0.122 |
5 | 3:1 | 0.019 |
6 | 3:1 | 0.037 |
7 | 3:1 | 0.069 |
Fig. 1 Scheme of the production of core–shell particles by simultaneous suspension and emulsion polymerization process. |
The morphological characterization of the supports (specific area, average pore diameter and volume of pores) was determined by nitrogen physisorption, using a surface analyzer (ASAP 2020 model, supplied by Micromeritics, Norcross, GA, USA) and the obtained values were adjusted using the BET model. Sample treatment was performed under vacuum at 60 °C. The average particle diameters were evaluated with a particle size distribution analyzer supplied by Malvern Instruments (Master sizer Hydro 2000S model). Measurements were performed in duplicates and the experimental errors were calculated with confidence level of 95%.
Polymer particles were also characterized by optical microscopy. The binocular microscope (Nikon, model SMZ800 with capacity of 50× magnification) was equipped with a digital camera (Nikon Coolpix 995), enabling the amplification and digitization of the images. A Scanning Electron Microscope (Fei Company, Model Quanta 200) was also used to characterize the obtained particles. Photomicrographs were processed in an image analyzer (Fei Company). Typical morphological features of PS/PS particles have been described in previous publications.60
Fig. 3 illustrates the influence of the co-monomer feed flow rate on the morphology of the polymeric particles. Considering the specific area (Fig. 3A) and the volume of pores (Fig. 3B) of the core–shell particles, there was a particular range on the monomer feed flow rate that resulted in particles with pronounced specific area and porosity. However, apparently the comonomer feed flow rate did not affect the average pore diameter of the particles (Fig. 3C). Probably, a low co-monomer feed flow rate (0.019 L h−1) provided a longer time for the coating of the cores (that could result in higher specific area and more porous particles). However, it may also cause greater stability of the emulsified particles, and that may result in lower core coatings and lower specific area of the core–shell particles. Moreover, a high co-monomer feed flow rate caused a destabilization of the emulsion, resulting in larger agglomeration of particles (both the core and the shell nanoparticles) and in a decrease of the specific area and the porosity. Therefore, regarding the enzyme immobilization procedure, only one support among the ones that were obtained was evaluated: the one having the highest specific area (produced on reaction 7), called PS-co-DVB/PS-co-DVB. The support PS/PS was used for comparison.
Fig. 3 Effect of the comonomer feed flow rate on the morphological properties of the particles: (A) specific area; (B) volume of pores; (C) average pore diameter. |
Fig. 5 Scanning electron micrographs of the polymeric particles: (A) PS-co-DVB core particles; (B) core–shell PS/PS particles; (C) core–shell PS-co-DVB/PS-co-DVB particles. |
Considering the wide range of likely applications for the produced supports, the average particles diameter is important since it may condition their handling. Very small particles would require the use of complex methods for separation of biocatalysts from the reaction media at the end. Very large particles can intensify diffusional problems. Therefore, the production of micrometric support particles is desirable and can facilitate the industrial use of the biocatalysts. The average particle diameters of the supports that will be used on the enzyme immobilization process are shown in Table 2. The produced supports presented average particle diameters of approximately 100 μm. PS/PS particles were the largest ones. It can also be noticed that the average diameters of the new PS-co-DVB/PS-co-DVB core–shell particles were smaller than those of the corresponding core particles. This was due to coagulation of nanoparticles during the emulsion step, which shifted the average particle sizes towards smaller values. Nevertheless, the particles were still on the micrometric scale, which is advantageous for separation processes.
Table 3 shows the specific area, the average pore diameter and the volume of pores of the produced core–shell particles. These results clearly indicate the formation of the shell over the particle cores, as the core presents negligible specific area when compared to the core–shell supports.
Comparing the core–shell supports, the specific area and porosity were higher for the new PS-co-DVB/PS-co-DVB particles, indicating that the small modifications of the implemented operation procedures utilized in the preparation of these new materials allowed the production of more porous matrices. The presence of DVB in the reaction media leads to an increase in the particle porosity because DVB promotes chain crosslinking, changing the microstructure of polymeric particles, as discussed in previous works.41 Besides, the increase of the feed flow rate during the emulsion step (0.069 L h−1 instead of 0.04 L h−1, as in the previous study41) allowed the production of higher amounts of nanoparticles in shorter reaction times, increasing the desired core coverage without increasing the rate of undesired agglomeration of the support particles.
It is possible to observe in Table 3 that the average pore sizes of the supports ranged from 200 Å (PS-co-DVB/PS-co-DVB) to 290 Å (PS/PS), which are wide enough to permit the diffusion of even moderately large protein molecules. As the core compact particles exhibited negligible specific areas, they were not used for enzyme immobilization studies; they were used only to evaluate the shell coverage on the core–shell particles synthesis.
According to Cunha et al. (2014)41 all of these different supports should exhibit similar hydrophobicities. This indicates that distinct interactions between the polymeric supports and the enzymes should be mainly caused by differences of the surface characteristics of the particles (area, internal morphology). Moreover, as the synthesized supports are hydrophobic, lipases should tend to become interfacially activated versus their surface, and this should be the main cause for the immobilization procedure.29
Lipases | Supports | Inactivation conditionsa | |
---|---|---|---|
pH 7, 70 °C | 90% DMF, pH 7, 25 °C | ||
a The number of replicates of these analyses were 6 (n = 6) and the experimental errors were calculated with confidence level of 95%. | |||
CALB | PS/PS | 10.0 ± 0.5 | 45 ± 1.0 |
PS-co-DVB/PS-co-DVB | 30 ± 0.5 | 25 ± 1.0 | |
Octyl-agarose | 28 ± 1.5 | 8 ± 1.0 | |
RML | PS/PS | 5.0 ± 0.2 | 5.0 ± 1.0 |
PS-co-DVB/PS-co-DVB | 20 ± 0.2 | 18 ± 0.8 | |
Octyl-agarose | 10 ± 0.5 | 13 ± 2.0 | |
TLL | PS/PS | 120 ± 1.0 | 20 ± 1.0 |
PS-co-DVB/PS-co-DVB | 40 ± 0.1 | 40 ± 0.5 | |
Octyl-agarose | 165 ± 5.0 | 22 ± 3.0 | |
LU | PS/PS | 40 ± 0.5 | 15 ± 1.0 |
PS-co-DVB/PS-co-DVB | 5 ± 0.2 | 40 ± 1.0 | |
Octyl-agarose | 19 ± 4.0 | 8 ± 1.5 |
Therefore, the relative stability of the preparations depends on the enzyme and also on the inactivating agent, although, in general, in the presence of organic co-solvents the better performance of the new core–shell biocatalysts seems clear. Perhaps the high hydrophobicity of the produced particles permits a stronger enzyme adsorption on the support, when compared to octyl-agarose biocatalysts. This may reduce the release of the enzyme to the medium in the presence of organic solvents, being the main reason for lipase inactivation in organic solvent during incubation in high organic cosolvent concentration solutions.6
Hydrolysis of triacetin | ||
---|---|---|
Biocatalyst | Activity (μmol (min g)−1) | |
a Analyses were conducted in triplicate (n = 3) and the experimental errors were calculated with confidence level of 95%. | ||
CALB | Novozym 435 | 207 ± 1.2 |
PS/PS | 214 ± 1.0 | |
PS-co-DVB/PS-co-DVBa | 105 ± 0.8 | |
Octyl-agarose | 89 ± 0.5 | |
RML | RM-IM | 28.1 ± 0.5 |
PS/PS | 47.0 ± 0.3 | |
PS-co-DVB/PS-co-DVB | 50.0 ± 0.6 | |
Octyl-agarose | 42.8 ± 0.5 | |
TLL | TL-IM | 15.1 ± 0.2 |
PS/PS | 32.1 ± 0.6 | |
PS-co-DVB/PS-co-DVB | 32.2 ± 0.5 | |
Octyl-agarose | 27.3 ± 1.1 | |
LU | PS/PS | 20.8 ± 1.1 |
PS-co-DVB/PS-co-DVB | 31.3 ± 0.8 | |
Octyl-agarose | 28.3 ± 0.5 |
Hydrolysis of R-methyl mandelate | |||
---|---|---|---|
Biocatalyst | Activity (μmol (min g)−1) | VR/VS | |
a Analyses were conducted in triplicate (n = 3) and the experimental errors were calculated with confidence level of 95%. | |||
CALB | Novozym 435 | 140 ± 1.2 | 1.35 |
PS/PS | 210 ± 2.2 | 1.65 | |
PS-co-DVB/PS-co-DVB | 260 ± 1.6 | 2.65 | |
Octyl-agarose | 210 ± 2.8 | 2.30 | |
RML | RM-IM | 0.51 ± 0.3 | 0.32 |
PS/PS | 0.57 ± 0.6 | 0.31 | |
PS-co-DVB/PS-co-DVB | 0.82 ± 0.1 | 0.28 | |
Octyl-agarose | 0.74 ± 0.1 | 0.24 | |
TLL | TL-IM | 1.0 ± 0.2 | 2.3 |
PS/PS | 1.1 ± 0.1 | 1.64 | |
PS-co-DVB/PS-co-DVB | 2.3 ± 0.1 | 1.67 | |
Octyl-agarose | 1.1 ± 0.2 | 1.57 | |
LU | PS/PS | 0.9 ± 0.2 | 0.81 |
PS-co-DVB/PS-co-DVB | 2.6 ± 0.2 | 1.45 | |
Octyl-agarose | 0.95 ± 0.1 | 0.95 |
Using CALB biocatalysts in the hydrolysis of triacetin, the least active biocatalyst was octyl-agarose-CALB followed by PS-co-DVB/PS-co-DVB-CALB. PS/PS-CALB was slightly more active than the commercial and widely used Novozym 435 with this substrate (Table 5). However, in the hydrolysis of R methyl mandelate, Novozym 435 had the lowest activity while the new PS-co-DVB/PS-co-DVB-CALB was the most active, with octyl-agarose-CALB and PS/PS-CALB exhibiting similar activities (Table 6). All preparations preferred the R isomer with a moderate enantiopreference, although PS-co-DVB/PS-co-DVB-CALB doubled the ratio VR/VS compared to Novozym 435.
Analyzing RML biocatalysts, the most active one using both substrates was the new PS-co-DVB/PS-co-DVB-RML (Tables 5 and 6). The second most active biocatalyst depended on the employed substrate: it was PS/PS-RML using triacetin, while using methyl mandelate as substrate, the second most active was octyl-agarose-RML. The commercial preparation was the catalyst with the lowest activity in both cases. In this case, the preferred isomer was the S, with activity ratios between both isomers ranging from 3 to 4 (Table 6).
Considering TLL immobilized preparations, the most active biocatalysts in triacetin hydrolysis were both core–shell supports, shortly followed by octyl-agarose-TLL and both of them doubling the activity of the commercial preparation (Table 5). Using R methyl mandelate, PS-co-DVB/PS-co-DVB-TLL was the most active; octyl-agarose-TLL and PS/PS-TLL showed around half of that activity, shortly followed by the commercial preparation (Table 6). The preference for the R isomer was scarce (ranging from 1.57 using octyl-agarose-TLL to 2.3 employing the commercial preparation).
LU biocatalysts exhibited short differences using triacetin as substrate, being PS-co-DVB/PS-co-DVB-LU the most and PS/PS the least active one (Table 5). However, PS-co-DVB/PS-co-DVB-LU was 2.5 fold more active than the other two biocatalysts using R methyl mandate as substrate, (Table 6). The enantiopreference was very low, but while PS-co-DVB/PS-co-DVB-LU preferred the R isomer, the other two preparations preferred the S isomer (Table 6).
It was possible to observe generally better hydrolytic activities of the new biocatalysts, mainly the enzymes immobilized in the new PS-co-DVB/PS-co-DVB core–shell support, compared to the commercial ones. More interestingly, it was noticed that even though the immobilization of all home-made biocatalysts involved interfacial activation as immobilization mechanism, the final properties of each one were strongly modulated by the exact nature of the support. Thus, the most active biocatalyst produced using a specific support may exhibit a low hydrolytic activity when other substrate were investigated, as reported in many other cases.6
Another important feature of an immobilized enzyme is its operational stability. To this goal, each enzyme biocatalyst was employed on 5 consecutive cycles of hydrolysis of triacetin. After each cycle, the biocatalysts were washed 3 times with 3 volumes of 20 mM sodium phosphate. All the biocatalysts, including the commercial biocatalysts, those prepared using octyl-agarose and core–shell biocatalysts, showed a decrease of activity under 20% along the 5 reaction cycles, as shown in Fig. 7.
The new biocatalysts were much more stable than the commercial biocatalysts or the ones obtained using octyl-agarose in many instances but not always. This higher stability was mainly observed in organic solvents inactivations, where enzyme desorption play an important role in the biocatalyst stability and the more hydrophobic nature of the new PS-co-DVB/PS-co-DVB should reduce the enzyme release.67 However, considering each enzyme and each inactivation condition, it seems that there is not an absolute “optimal” immobilization support from those here studied regarding enzyme stability.
Finally, the activities of the new biocatalysts employing distinct substrates were much higher than the ones obtained with commercial products, except when triacetin was hydrolyzed by CALB. Moreover, in many cases, the enzymatic activities were also much higher than the ones observed when octyl-agarose was used as support. As it has been previously reported,35 the chemical and textural properties of the support surfaces alter the final lipase performance even if the immobilization mechanism is in all cases interfacial activation. Thus, the use of differently prepared hydrophobic core–shell supports may be a way to enrich a library of lipase biocatalysts.8
Based on the obtained results, it can be concluded that changing some operational conditions during the polymerization reaction, such as the co-monomer feed flow rate, it is possible to synthesize core–shell particles with distinct morphological aspects. Moreover, among the different core–shell supports that were produced, the new PS-co-DVB/PS-co-DVB constitutes a very promising support for lipase immobilization.
Footnotes |
† Electronic supplementary information (ESI) available. See DOI: 10.1039/c6ra13350a |
‡ These coauthors have contributed equally to this paper. |
This journal is © The Royal Society of Chemistry 2016 |