Article pubs.acs.org/Langmuir
Cellulase and Alcohol Dehydrogenase Immobilized in Langmuir and Langmuir−Blodgett Films and Their Molecular-Level Effects upon Contact with Cellulose and Ethanol Dilmer Rodrigues, Fernanda Ferraz Camilo, and Luciano Caseli* Rua Sao Nicolau, 210, Laboratorio de Materiais Hibridos, Diadema, SP 09913-030, Brazil S Supporting Information *
ABSTRACT: The key challenges for producing devices based on nanostructured films with control over the molecular architecture are to preserve the catalytic activity of the immobilized biomolecules and to provide a reliable method for determining the intermolecular interactions and the accommodation of molecules at very small scales. In this work, the enzymes cellulase and alcohol dehydrogenase (ADH) were coimmobilized with dipalmitoylphosphatidylcholine (DPPC) as Langmuir−Blodgett (LB) films, and their biological activities were assayed by accommodating the structure formed in contact with cellulose. For this purpose, the polysaccharide was dissolved in an ionic liquid, 1-buthyl-3-methylimidazolium chloride (BMImCl), and dropped on the top of the hybrid cellulase-ADH-DPPC LB film. The interactions between cellulose and ethanol, which are the catalytic substrates of the enzymes as well as important elements in the production of second-generation fuels, were then investigated using polarization-modulation infrared reflection−absorption spectroscopy (PM-IRRAS). Investigation of the secondary structures of the enzymes was performed using PM-IRRAS, through which the presence of ethanol and cellulose was observed to highly affect the structures of ADH and cellulase, respectively. The detection of products formed from the catalyzed reactions as well as the changes of secondary structure of the enzymes immobilization could be carried out, which opens the possibility to produce a means for producing second-generation ethanol using nanoscale arrangements.
1. INTRODUCTION The incorporation of biomolecules into nanostructured films can be utilized for several applications, such as biosensing, bioremediation, and heterogeneous catalysis. Of particular relevance for nanobiotechnology has been the investigation of new methods to immobilize enzymes, whose activity may then be preserved for long periods of time. In this sense, such methods may be effectively achieved if appropriate matrices are employed. When enzymes are immobilized in ultrathin films, such as those produced with the Langmuir−Blodgett (LB) technique,1,2 accurate control over the thickness, composition, surface elasticity, and molecular density is achieved. As a result, LB films composed of lipids and enzymes have been proposed in recent years not only for producing structures with architectural control at the molecular level, but also because the amphiphilic nature of lipids may help to preserve the conformation of active enzymes3,4 and to induce the polypeptide moiety to adopt an adequate orientation for access to the catalytic substrates.5,6 The LB methodology requires the formation of stable monolayers at air/liquid interfaces, known as Langmuir films, and their subsequent transfer to solid supports that vertically intercept the interface.2 The incorporation of the enzyme into the monolayer from the aqueous subphase depends on the specific affinity between the polypeptide and the functional groups of the lipid. Usually, enzymes present some surface activity due to their amphiphile nature;7 however, it has been © 2014 American Chemical Society
widely reported that the presence of lipids induces the adsorption of enzymes at the air−water interface.8−10 Thus, as contact with a bare air−water interface can cause polypeptide denaturing,11 lipid monolayers can act as a protector matrix for the enzyme at the interface.3,6 Because the preservation of activity depends on molecular-level interactions, a detailed investigation of the properties of Langmuir films is necessary, and determining how the enzyme conformation is affected by the lipid environment or by contact with other liquids of interest is of importance. These issues are particularly important for biotechnology, as many enzymes are currently employed for catalysis or for analyte identification in biosensing. In this study, we address significant issues associated with the immobilization of the enzymes cellulase and alcohol dehydrogenase (ADH) in solid lipid LB films. Considering the technological importance of these enzymes in biotechnology, especially in the production, identification, and/or control of second-generation ethanol, we intend to investigate how the conformations of these enzymes into lipid LB films are affected in the presence of cellulose and ethanol. To the best of our knowledge, this is the first report on this issue in the literature. For this study, tensiometry and polarization-modulation infrared reflection absorption spectroscopy (PM-IRRAS) were Received: November 15, 2013 Published: January 28, 2014 1855
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containing LB films, ethanol (Synth) was dropped on top of the solid substrate, and PM-IRRAS spectra were recorded before and after contact with the film. All spectra of the LB films were recorded with a minimum of 1200 scans. Cellulase enzyme activity assay was determined according to the method described by Gosh et al.,14 with 0.05 mol/L sodium citrate (Sigma-Aldrich) and a filter paper strip (0.5 × 0.5 cm2) inserted together with the LB film into a cuvette with temperature elevated to 50 °C, and the absorbance at 540 nm measured with time. For ADH, the enzyme activity was determined according to the method previously described in the literature,15 using NADP and following ethanol oxidation with the absorbance evolution at 340 nm. Enzyme activities were determined for 3 kinds of film: ADH−DPPC, cellulase−DPPC, and ADH−cellulase−DPPC.
employed to obtain information on enzyme-catalytic substrate interactions at the molecular level.
2. EXPERIMENTAL SECTION The water used in all experiments was purified using a Milli-Q system (resistivity of 18.2 Ω cm, pH ≈ 6.0). Dipalmitoylphosphatidylcholine (DPPC), cellulase from Aspergillus niger, and ADH recombinant from Escherichia coli were obtained from Sigma-Aldrich. A DPPC solution was dissolved in chloroform (Synth) to a concentration of 0.88 mg/ mL. ADH and cellulase solutions with concentrations of 7.2 mg/mL were obtained by dissolving each enzyme in an aqueous buffer solution of K2HPO4 (Sigma-Aldrich) and KH2PO4 (Sigma-Aldrich) with a salt concentration of 0.01 mol/L and pH ≈ 7.2. BMImCl (1-butyl-3methylimidazolium chloride) was prepared as previously described in the literature.12 For the preparation of monolayers at the air−water interface, a Langmuir trough (KSV Instruments, model: Mini) was initially filled with the phosphate buffer solution. The DPPC solution was then spread onto the buffer/air interface to obtain an area per molecule of ≈100−120 Å2. After the evaporation of chloroform for 20 min, the monolayers were compressed with two movable barriers at a rate of 5 Å2 molecule−1 s−1. The surface pressure values were monitored with a filter paper Wilhelmy plate that intercepted the interface in the center of the trough. The sensitivity of this measurement is 0.1 mN/m. Compression was stopped either when the monolayer collapsed or when the minimum area allowed by the equipment had been attained. For mixed enzyme (cellulase or ADH)−lipid monolayers, after spreading the DPPC and evaporating the solvent, predetermined aliquots of the enzyme solution were carefully injected below the interface. After allowing the surface pressure to stabilize, the interface was compressed. The surface pressure was then monitored as a function of the film surface area. Polarization-modulation infrared reflection absorption spectroscopy measurements were performed using a KSV PMI 550 instrument (KSV Instrument Ltd., Helsinki, Finland) for the Langmuir monolayers. With an incidence angle of 80°, the incoming light was continuously modulated between p- and s-polarizations, allowing for the simultaneous measurement of the spectra for the two polarizations. The PM-IRRAS signal is obtained from the reflectivities of both the p and s fractions of the light. The difference between the spectra provides surface-specific information, and the sum provides the reference spectrum. The ratio between the difference and the sum gives the PM signal, which reduces the effects of water vapor and carbon dioxide, providing information on vibrational groups present only at the air−water interface. For such measurements, the monolayers were compressed until the desired surface pressure was achieved, and PM-IRRAS spectra were recorded for a minimum of 600 scans. Images of the air−water interface were obtained using a Brewster Angle Microscope (KSV-Nima MicroBAM) adapted to the Langmuir trough. The Langmuir films were then transferred onto solid glass supports, which had previously been cleaned with KOH and ethanol, by vertically withdrawing the support across the air/film interface with an immersion speed of 5 mm min−1 and at a constant surface pressure of 30 mN m−1. The results indicated that transfer ratio values in the interval of 0.9−1.0 permitted effective deposition. Then, the resulting Langmuir−Blodgett (LB) films were analyzed. Four types of LB films were produced depending on the origin of the Langmuir monolayer: pure DPPC, ADH−DPPC, cellulase−DPPC, and ADH−cellulase− DPPC. All LB films had a thickness of one single layer. For further analysis of the cellulase-containing LB films, microgranular cellulose (Sigma-Aldrich) dissolved in BMImCl was dropped onto the cellulase−DPPC films. The dissolution was conducted according to the literature.13 Briefly, 200 mg of cellulose was added to a vial containing 2.0 mL of BMImCl (10 wt %), and the mixture was placed in a conventional microwave oven and heated with 2 s pulses at full power for 8 cycles, after which the complete dissolution of cellulose was observed. PM-IRRAS spectra of the LB film were recorded before and after the contact with cellulose. For the ADH-
3. RESULTS AND DISCUSSION 3.1. Tensiometry. Figure 1 shows the surface pressure− area isotherms for DPPC with different concentrations of ADH
Figure 1. Surface pressure−area isotherms for DPPC on phosphate buffer (0.01 mol/L, pH 7.2) with different ADH concentrations (indicated in the inset).
in the buffer subphase. For DPPC spread on a subphase in the absence of an enzyme, the curve presents a typical profile, as previously reported in the literature.16 A typical transition between the liquid-expanded and the liquid-condensed states is identified by a plateau located in the surface pressure range 4−8 mN/m, which corresponds to a mean molecular area range 78− 52 Å2/molecule. With further compression, the monolayer attains the liquid-condensed state and collapses at 44 Å2, with a corresponding surface pressure of approximately 70 mN/m. With the presence of ADH in the subphase, the isotherm is progressively shifted to large areas with enzyme concentrations varying from 0.8 to 4.2 μg/mL. For instance, with an ADH concentration of 4.2 μg/mL, the isotherm shifted from 42 to 53 Å2/molecule at a surface pressure of 30 mN/m. This result indicates that the enzyme was incorporated in the air−water interface, which leads to expansion of the monolayer. Considering the total number of DPPC spread at the interface, this shift represents an increase of 2.42 × 1017 Å2 of total area. Taking into account that all ADH molecules incorporated into the lipid monolayer and considering a molecular weight of ca. 141 kDa, the molecular area of ADH is estimated in 7 Å2. Obviously this area is too small for a protein with this molecular weight, and this fact could be explained either because of a lower amount of enzyme adsorbed, or because of the fact that the adsorption of the enzyme occurred below the polar heads of DPPC molecules, and therefore it did not penetrate between 1856
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observed when higher concentrations of macromolecules, such as proteins22 or polysaccharides,23 are subsequently added to the monolayer. Figure 2 shows the effect of cellulase on the DPPC monolayers. For enzyme concentrations in the range 42−196
the lipid alkyl chains. With an isoelectric point of 5.4, ADH has a net positive charge, and dipole−ion interactions may occur with the polar head of DPPC. Also, when the air−ADH aqueous solution interfaces are compressed, i.e., with no DPPC present on the interface, no significant variation in the surface pressure is observed, indicating poor surface activity of the enzyme. The surface activity of ADH as Gibbs monolayers is reported in the literature only with higher concentrations, such as 5 mg/mL.17 Therefore, the shift of DPPC isotherms to higher areas reflects the affinity of this enzyme toward the lipid monolayer. Furthermore, we can observe a remarkable difference of the effect of ADH on the DPPC monolayer when the enzyme concentration varies from 1.2 to 2.8 μg/mL. The initial surface pressure (with a DPPC molecular area of 112 Å2) increases significantly from zero to approximately 8 mN/m. This result suggests a high adsorption of the enzyme on the lipid monolayer. With further compression, the plateau that characterizes the liquid-expanded to liquid-condensed phase transition is no longer well-defined, and between 8 and 28 mN/ m, the monolayer is, by visual contrast, more compressible, which indicates that the surface pressure varies less as the monolayer area decreases. For higher pressures, the monolayer expansion is less evident, which can indicate some expulsion of the incorporated ADH molecules toward the buffer aqueous subphase or toward the monolayer subsurface. The increase in the compressibility of the monolayers is associated with the compressional modulus or with the in-plane elasticity of the monolayer,18 which is defined as −A(∂π/∂A)T, where A is the molecular area, π is the value of surface pressure, and T is the absolute temperature. A higher value of this quantity indicates that the monolayer is highly sensitive to area compression. This sensitivity is reflected in the high rate of increase in surface pressure, and it is usually associated with the degree of packing and organization of the monolayer. Therefore, as ADH caused a decrease in the monolayer elasticity, there was a disruption of the well-packed structure of DPPC at the air−water interface. This result is expected because a new component is being introduced at the interface, which reflects an entropy gain and therefore results in a decrease in the molecular order; i.e., the monolayer is more flexible under compression and may adopt other molecular arrangements to respond to the compression through the barriers. This behavior has been extensively reported for proteins adsorbing in Langmuir monolayers of lipids with saturated alkyl chains.19−21 This order is provided probably because the enzyme is incorporated adsorbing on the polar heads of the phospholipids rather than penetrating into the alkyl chains. The molecular order is further evidenced in this paper by analyzing the ratio between the intensity of the PM-IRRAS bands related to C−H stretches. The reported effect of ADH affecting the molecular organization of the DPPC monolayer can also be identified by observing the surface pressure of collapse, which is decreased compared to that for a monolayer of pure DPPC. This result indicates that a mixed monolayer is less stable under compression. With an ADH concentration of 5.6 μg/mL in the aqueous subphase, the isotherm, within the experimental error, is almost identical to the isotherm for the pure DPPC monolayer, except that the collapse occurs at a surface pressure of approximately 40 mN/m. This result can be attributed to the possible aggregation of ADH in the aqueous subphase, which disfavors enzyme adsorption at the monolayer. Similar behavior is
Figure 2. Surface pressure−area isotherms for DPPC on phosphate buffer (0.01 mol/L, pH 7.2) with different cellulase concentrations (indicated in the inset).
μg/mL, a progressive shift to higher areas is observed. For an enzyme concentration of 280 μg/mL, no significant monolayer expansion is observed compared to the previous enzyme concentration assayed (196 μg/mL). Moreover, for surface pressures greater than 20 mN/m, the monolayer becomes more compressible, which is a consequence of the decrease in the monolayer surface elasticity, causing the isotherm to shift to lower areas when compared to the isotherm for a cellulase concentration of 196 μg/mL. Remarkably, it required a higher amount of cellulase compared to ADH to affect the DPPC isotherm, which may be related to its lower hydrophobicity. With a further increase of the enzyme concentration in the aqueous subphase (500 μg/mL), no monolayer expansion is observed compared to the pure DPPC film. In a general way, this behavior for mixed cellulase−DPPC monolayers is similar to that observed for a mixed ADH−DPPC monolayer; i.e., the progressive shift to larger areas as long as high enzyme concentrations are employed is interrupted by a given concentration in which no significant expansion is observed. As in the previous case, the effects caused by cellulase on DPPC monolayers are the same: higher enzyme concentrations may favor their presence in the buffer subphase, not at the air−water interface, due to aggregation of the polypeptide chains in the aqueous phase. However, even at this concentration, some effect of the enzyme is observed on the lipid film because the monolayer collapse occurs at a lower surface pressure. The excess of enzyme located in the subsurface of the lipid layer may destabilize the monolayer, hampering the film from becoming structurally well-packed. Notably, the initial surface pressure of the cellulase−DPPC monolayer was zero for all cases, even for the highest enzyme concentration employed in this work. This result indicates a lower tendency for enzyme aggregation at large areas. Furthermore, the plateau that characterizes the transition from the liquid-expanded to the liquid-condensed phase is still more evident than for the ADH−DPPC isotherms. These results indicate that ADH has a greater effect compared to cellulase regarding the disruption of the surface-packing 1857
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structure for DPPC monolayers. However, taking into account its molecular weight of 42 kDa, the shifts to higher areas also represent a value too small considering that all cellulase molecules penetrate in the lipid monolayer. As its isoelectric point is 4.8, the net positive charge of the enzyme also may induce the adsorption below the polar heads of the phospholipid. Figure 3 shows the effect on the DPPC monolayer when both enzymes are present in the aqueous subphase. Aiming for
Figure 3. Surface pressure−area isotherms for DPPC on phosphate buffer (0.01 mol/L, pH 7.2) with ADH (4.2 μg/mL) and cellulose (196 μg/mL).
the greatest amount of enzyme molecules incorporated in the monolayer, the ADH and cellulase concentrations were chosen on the basis of the highest expansion of the lipid monolayers that they provided, as shown in Figures 1 and 2. This strategy therefore permits monolayers to be obtained below the limit of formation of enzyme aggregates in the aqueous subphase. The isotherm presented in Figure 3 shows effects of the combination of both enzymes, with a shift of the isotherms to larger areas, an increase of the initial surface pressure, a decrease of the surface elasticity in the liquid-condensed phase, a decrease in surface pressure of collapse, and a transition plateau that is still roughly defined. This result therefore indicates that the enzymes can be coincorporated in the air− water interface. 3.2. Vibrational Spectroscopy for Langmuir Monolayers. To better identify the effect of incorporating the enzymes in the lipid monolayer on the secondary structure of the polypeptide chains, PM-IRRAS spectroscopy was performed. Figure 4 shows the PM-IRRAS spectrum for a DPPC monolayer at a surface pressure of 30 mN/m. This surface pressure value was selected because it corresponds to that of a natural cell membrane,24 and it was also the surface pressure value chosen for the transfer to solid supports as LB films. The bands at 2921 and 2855 cm−1 are attributed to the asymmetric and symmetric C−H stretches for CH2, respectively. With the incorporation of the enzymes, the intensity of the symmetric (Is) band compared to the intensity of the asymmetric band (Ia) is decreased. The same approximate trend occurs for the relative integrated areas below the bands. The relative area value for pure DPPC is 1.86, and it decreases to 1.45, 1.30, and 1.05 for the monolayers with ADH, cellulase, and ADH + cellulase, respectively. This result indicates a decrease in the order parameter,25 which is consistent with the results of the surface pressure−area isotherms, in which the adsorption of the
Figure 4. PM-IRRAS spectra for DPPC monolayers with enzymes adsorbed as indicated in the inset. Baseline is the line of the zero level of the PM-IRRAS signal.
enzymes decreases the order of the lipid monolayer. In panel B, the band at 1719 cm−1, which is present in all cases, represents the CO stretches for the lipid. The large band centered at 1680 cm−1 appears for all spectra, being negative to baseline (for DPPC and DPPC−ADH) or positive (for the other films). This band is a result of the difference in reflectivity between the interface covered and uncovered with the monolayer and results from the bending vibration of surface water.26 This band may appear in the spectra as a consequence of the restructuring of water molecules at the air−water interface after monolayer formation. As the background spectrum is taken after the formation of the monolayer, this band can appear in case of restructuring of water molecules at the surface. With the incorporation of enzymes, amide I (CO stretches) bands appear in the 1600−1690 cm−1 region. Amide II bands (CN and NH bends) should appear at 1500−1580 cm−1, but they are not evident. It has been reported in the literature that when the amide I bands are more intense than the amide II ones, the protein structure is not completely denatured at the interface.27 The band at 1652 cm−1 indicates an α-helix structure, whereas a shoulder at 1621 cm−1 indicates the presence of β-sheet structures. For cellulase, the band at 1621 cm−1 is more defined, indicating a higher extension of β-sheets. When both enzymes are present at the interface, all of these bands continue to appear, with a slight shift for the α-helix band to higher wavenumbers, indicating a cooperative behavior between the enzymes. For comparison we show the FTIR spectra obtained for the enzyme in KBr pellets (Supporting Information), in which the band amide I can be 1858
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Figure 5. BAM images for DPPC monolayers at 30 mN/m: (A) without enzymes incorporated, (B) cellulase−DPPC, (C) ADH−DPPC, (D) ADH−cellulase−DPPC.
observed for cellulase in 1656 cm−1, and for ADH in 1662 cm−1. Obviously, the enzyme secondary structure may be slightly different in KBr when compared with its structure in the buffer solution. 3.3. Brewster Angle Microscopy for Langmuir Monolayers. Figure 5 shows the BAM images for the monolayers studied. At 30 mN/m, the surface seems relatively homogeneous due to the fact that the film attained the liquid condensed phase. However, we still observe some black interstices as a result of noncomplete packing of the monolayer. With the introduction of the enzymes, the interface becomes more heterogeneous as a consequence of the distribution of the enzymes at the interface and further formation of aggregates between enzymes and lipids. These results confirm that the enzymes were incorporated in the DPPC monolayers. 3.4. Characterization of LB Films. Figure 6 shows the spectra when the monolayers are transferred from the air−
monolayers supported at the water surface, which results in a clearer spectrum. Furthermore, the interaction with the glass surface may influence the conformation of the amide bands. The carbonyl stretch of DPPC is observed at different positions, depending on the enzyme employed, indicating that the enzymes may be interacting strongly with this polar group when water is absent. For ADH−DPPC and cellulase− DPPC, the band for the α-helix structure is absent, and others at 1678 and 1663 cm−1, related to unordered structures, were evident for LB films. The band for β-sheet structures continues to be evident but is located at different wavenumbers (1656, 1610, and 1625 cm−1 for ADH−, cellulase−, and mixed ADH/ cellulase−DPPC films, respectively). Amide II bands can now be better identified, and for the ADH−DPPC LB film, an amide II band centered at approximately 1555 cm−1 is clearly more intense than that for any amide I bands. All of these results indicate that the conformations of these enzymes are highly dependent on their surrounding environments. However, this result is not necessarily an indication of denaturation or the total loss of their catalytic activity. It has been extensively reported in the literature that enzymes immobilized in lipid LB films maintain part of their biological activity and that they may be widely employed as potential biosensors.28,29 Even when marked changes in their secondary structures are observed through PM-IRRAS, their catalytic activity is still partially maintained. Another important result worth mentioning is that the band in ∼1700−1800 cm−1, corresponding to the CO stretching mode for the phospholipid, is highly sensitive to the enzyme incorporated in the LB film. With ADH, this band is centered at 1751 cm−1, whereas for cellulase this band is centered at 1744 cm−1. Interestingly, for both enzymes, this band splits in two, at 1772 and 1717 cm−1, representing therefore the synergistic mutual contribution of the enzymes. This result confirms the information given by the surface pressure−area isotherms, in which we assumed that the main interaction between enzymes and DPPC occurs through the polar heads of the phospholipid. 3.5. Effect of the Enzymes Immobilized upon Contact with Cellulose and Ethanol. Because cellulase acts directly in the hydrolysis of cellulose, contact of the cellulase−DPPC LB film with cellulose could directly affect the secondary structure of the enzyme. Therefore, cellulose was dissolved in BMImCl, as described in the Experimental Section. Due to the hydrogen-bonded supramolecular structure present in cellulose, this polymer is known to be insoluble in water and most common organic liquids. However, its solubility in specific ionic liquids using microwave opens the opportunity to work with dissolved cellulose, which was not previously possible. The dissolution of cellulose in an ionic liquid in this work was necessary to study
Figure 6. PM-IRRAS spectra for DPPC−enzyme LB films. The enzymes coadsorbed with the lipid are indicated in the legends. Baseline is the line of the zero level of the PM-IRRAS signal.
water interface to solid supports. Only the carbonyl vibration region is shown because, for C−H stretches (2800−3000 cm−1), little change is observed compared to the spectrum for the air−water interface (Figure 4A). Otherwise, the 1500−1800 cm−1 region in the spectrum changes completely compared to that of the air−water interface, mainly when enzymes are present. For pure DPPC (spectrum not shown for better clarity), a single band is observed at 1719 cm−1 as a result of CO vibrations. These changes are mainly attributed to the absence of water in the LB films, which may alter the secondary conformation of the polypeptide structure and also cause the absence of the band at 1680 cm−1, as observed for the 1859
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difficult to identify because its main vibrational groups (such as OH and CH) are also present in other components that were previously present in the LB film. Furthermore, after 30 min up to 150 min, no significant changes are observed with time. For the alkyl stretching region (2800−3000 cm−1), as shown in Figure 7, some bands appear that are related to only C−H stretches for CH2, with the primary bands centered at 2920 and 2850 cm−1 and others at 2960 and 2870−2910 cm−1. In contrast to pure DPPC LB films, in which the CH2 bands are thinner, after contact with the enzyme, broad bands are observed. This result can be attributed to the disorder caused by the interaction of the enzyme with the phospholipid, as previously discussed when describing the surface pressure−area isotherms and the PM-IRRAS spectra for the air−water interface.31,32 Cellulose was then dropped onto the LB films, and a progressive increase in the definition of the bands was observed with time. At equilibrium (longer than 120 min), we observe bands at 2960, 2925, and 2880 cm−1. Considering that DPPC is the main component responsible for the appearance of these bands in the PM-IRRAS spectra, all of these results together indicate that the interaction of cellulose with the enzyme also affects the lipid matrix. As expected, when the cellulose solution is dropped onto the ADH-DPPC LB films, a negligible effect on the secondary structure of the polypeptide chain upon contact with the polysaccharide is observed, indicating that ADH has little affinity for cellulose. When both enzymes are present (Figure 8), some changes are observed as a result of the cellulase−
the interaction between the enzyme and its catalytic substrate in a simple manner. It is important to mention that, before the measurements presented in Figure 6, an ionic liquid without dissolved cellulase was dropped onto the LB film, and a spectrum was recorded. No significant changes were observed in the spectrum, and the chemical groups of the ionic liquid were not detected in the regions of interest. This result most likely occurs because the ionic liquid molecules in the interface with the LB film do not vary in terms of their molecular orientation compared to the molecules in the inner part of the liquid above the solid interface. Figure 7 shows the PM-IRRAS spectrum for cellulose dropped onto a clean, solid glass surface, without any DPPC−
Figure 7. PM-IRRAS spectra for DPPC−cellulase LB films before and after contact with cellulose (contact time indicated in the inset). Baseline is the line of the zero level of the PM-IRRAS signal.
Figure 8. PM-IRRAS spectra for DPPC−ADH−cellulase LB films before and after contact with cellulose (contact time indicated in the inset). Baseline is the line of the zero level of the PM-IRRAS signal.
cellulase film deposited on it. Panel A shows that cellulose affects the amide groups of cellulase, altering not only its αhelix structure but also the β-sheets. The α-helix structure is identified with a more evident band at 1653 cm−1 upon contact with cellulose. Additionally, the β-sheet band is shifted from 1610 to 1621 cm−1. The amide II band is also defined with a maximum at 1561 cm−1, and the C−H bending bands at 1447 cm−1 are split into two bands after contact with cellulose. Overall, this result indicates an interaction between cellulose and the cellulase−DPPC LB film. As there is some specificity between cellulase and the enzyme, this is a particular case of molecular recognition intrinsic to this nanostructured film. Changes in the secondary structure of the polypeptide moiety of enzymes upon contact with their substrate have been reported in the literature.30 Note that the specific bands for cellulose incorporated in the LB film−liquid ionic interface are
cellulose interaction. The α-helix band at 1652 cm−1 is highlighted, which means that the secondary structure is affected for cellulase, even for the mixed ADH−cellulase system. Other bands highlighted are 1622 cm−1 for β-sheet structures, 1560 cm−1 for amide II vibrations, 1459 cm−1 for CH bending vibrations, and 1738 cm−1 for DPPC CO stretching. All of these bands broadly show a notable difference with the spectrum obtained prior to contact with cellulose, thus indicating the specific interaction of cellulose with the polypeptide moieties of the cellulase. As the effect is different from that observed for the cellulase−DPPC LB film (without ADH), interactions between the polypeptide moieties of both enzymes may be occurring. The spectra for the CH stretching bands (2800−3000 cm−1) also show some minor alterations, indicating changes in the lipid matrix structure. This 1860
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activity because they do not contain cellulase. For cellulasecontaining films, with or without ADH coimmobilized, the values of activity were similar, which indicates that ADH does not affect considerably the activity of cellulase. Compared to the homogeneous environment, the enzyme activity is lower when immobilized in solid matrices.22,28,29 For that, we show the values in percentage in relation to the homogeneous environment. Interestingly, the value of activity was not so low when compared to other enzymes immobilized as LB films.22,28,29 Furthermore, after 20 days, the cellulase activity is more preserved in the LB films than in homogeneous solution, which means that the devices provided a more stable environment for the enzyme. PM-IRRAS for the LB was also measured after 20 days, and no significant change in the spectra was observed, which indicates that the secondary structure of the enzyme was preserved. Similar behavior is observed for enzyme activity to ethanol (Table 2). Activity is negligible to the film without ADH, and it
result is omitted for clarity and is available in the Supporting Information file. To investigate the effect of ethanol on the ADH−cellulase LB films, ethanol was dropped on the films, and PM-IRRAS spectra were collected. Figure 9 shows the effect on the ADH−
Table 2. ADH Activity for the LB Films and Comparison to the Enzyme Activity in Solutiona Figure 9. PM-IRRAS spectra for DPPC−ADH−cellulase LB films before and after the contact with ethanol (contact time indicated in the inset). PM-IRRAS spectra for cellulose cast onto a bare substrate are also shown. Baseline is the line of the zero level of the PM-IRRAS signal.
LB film homogenoeus DPPC−ADH DPPC− cellulase DPPC− cellulase− ADH
cellulase−DPPC film upon ethanol contact. Ethanol cast onto a bare substrate presents a low peak/noise rate and is not shown. This result indicates that ethanol does not present an active band in PM-IRRAS spectra. In the amide band region, the 1692 cm−1 band was inverted to the baseline and became positive when in contact with ethanol. The bands for the α-helix structure (at ≈1650 cm−1) and for the β-sheet (at ≈1620 cm−1) are not well-defined, whereas the band at 1641 cm−1 (disordered structures) is more defined. This result indicates that ethanol alters the conformation of the enzymes present at the interface. We also can see that the band for CO stretches in DPPC, centered at 1731 cm−1, is enhanced after contact with ethanol. Other bands related to amide vibrations appear more clearly in the spectra, as those centered at 1612 cm−1 (β-sheets), 1448 cm−1 (C−H bending), 1553, 1501, and 1534 cm−1 (amide II modes). All these results show that the contact of the film with ethanol alters the conformation of the immobilized enzymes, which may alter in some manner the chemical groups of DPPC in contract with the enzyme. 3.6. Enzyme Activity. In order to evaluate the enzyme activity of cellulase in the LB films, we estimated, for all produced films, their catalytic activity to cellulose (Table 1). As expected, DPPC-ADH LB films present negligible enzyme
a
lag time
activity (ΔAbs340/ min/mg ADH)
% (homogeneous)
% activity preserved (after 20 days)
