Acylated Carrageenan Changes the Physicochemical Properties of

Article pubs.acs.org/JPCB

Acylated Carrageenan Changes the Physicochemical Properties of Mixed Enzyme−Lipid Ultrathin Films and Enhances the Catalytic Properties of Sucrose Phosphorylase Nanostructured as Smart Surfaces Jefferson M. Rocha,† Adriana Pavinatto,‡ Thatyane M. Nobre,‡ and Luciano Caseli*,† †

Institute of Environmental, Chemical and Pharmaceutical Sciences, Federal University of Sao Paulo, Diadema, SP 04021-001, Brazil São Carlos Physics Institute, University of São Paulo, São Carlos, SP 13566-590, Brazil

‡

S Supporting Information *

ABSTRACT: Control over the catalytic activity of enzymes is important to construct biosensors with a wide range of detectability and higher stability. For this, immobilization of enzymes on solid supports as nanostructured films is a current approach that permits easy control of the molecular architecture as well as tuning of the properties. In this article, we employed acylated carrageenan (AC) mixed with phospholipids at the air−water interface to facilitate the adsorption of the enzyme sucrose phosphorylase (SP). AC stabilized the adsorption of SP at the phospholipid monolayer, as detected by tensiometry, by which thermodynamic parameters could be inferred from the surface pressure−area isotherm. Also, infrared spectroscopy applied in situ over the monolayer showed that the AC−phospholipid system not only permitted the enzyme to be adsorbed but also helped conserve its secondary structure. The mixed monolayers were then transferred onto solid supports as Langmuir−Blodgett (LB) films and investigated with transfer ratio, quartz crystal microbalance, fluorescence spectroscopy, and atomic force microscopy. The enzyme activity of the LB film was then determined, revealing that although there was an expected reduction in activity in relation to the homogeneous environment the activity could be better preserved after 1 month, revealing enhanced stability.

1. INTRODUCTION Achievement of molecular architectures in nanostructured films that are able to conserve the biological activity of biomacromolecules, especially enzymes, is a current challenge. For this, matrices constituted of materials able to accommodate enzymes with low loss of catalytic activity and higher stability are desired. Considering the systems that can be considered as nanostructured, the Langmuir−Blodgett (LB) technique is an approach that permits obtaining ultrathin films with high control over molecular architecture and thickness. However, immobilization of enzymes onto solid supports is not straightforward because many polypeptides do not form coherent Langmuir monolayers at the air−water interface, and for this reason, they cannot be subsequently transferred onto solid supports using the LB methodology. Although enzymes have been spread alone at bare interfaces,1,2 some strategies have been reported not only to facilitate the adsorption of enzymes at the air−water interface but also to allow better molecular accommodations, which may influence the geometrical conformation of the macromolecules, preserving at least part of their catalytic properties. Some of these strategies include the use of high ionic strengths in the aqueous subphase.3 Another strategy involves spreading lipids that form stable Langmuir monolayers, which permits the adsorption of © 2016 American Chemical Society

enzymes from the aqueous subphase, forming mixed enzyme− lipid assemblies. The literature shows that lipids can serve as protector agents for enzymes, giving them stability in terms of conservation of catalytic properties.4−6 Usually, the enzyme activity of enzymes immobilized on solid supports is lower than that of enzymes in solution,7,8 most commonly attributed to a higher restriction in terms of molecular mobility as enzymes strive to achieve a lower surface energy or may not adequately expose their catalytic sites. This can be convenient for sensing because sometimes a very high reaction rate hampers the estimation of enzymatic activity and determination of the analytic curve. However, there are some cases that report an increase in catalytic activity for LB films, especially when the catalytic site is sensitive to hydrophobic environments, such as those provided by lipid matrices.9,10 With these implications in mind, new molecular architectures have been developed to enhance the molecular accommodation of enzymes for sensing properties. A suitable strategy has been the use of polysaccharides. As they are flexible and hydrophilic Received: April 6, 2016 Revised: May 6, 2016 Published: June 1, 2016 5359

DOI: 10.1021/acs.jpcb.6b03468 J. Phys. Chem. B 2016, 120, 5359−5366

Article

The Journal of Physical Chemistry B

For the preparation of monolayers at the air−water interface, a Langmuir trough (model: Mini; KSV Instruments) was initially filled with phosphate buffer solution. DMPA solutions were then spread onto the buffer−air interface to obtain an area per molecule of ∼140−150 Å2. A time of 20 min was assigned for chloroform evaporation, and the monolayers were compressed with two movable barriers at a rate of 5 Å2 molecule−1 s−1, with the surface pressure values monitored using a Wilhelmy plate made of filter paper that intercepted the interface in the center of the trough. When the monolayer collapsed, compression was stopped. For the preparation of mixed AC−DMPA monolayers, predetermined aliquots of AC were co-spread with DMPA. After allowing the surface pressure to stabilize, the interface was compressed, with the surface pressure monitored as a function of the molecular area. For the preparation of mixed enzyme−AC− DMPA monolayers, after spreading AC and DMPA and subsequent evaporation of the solvent, predetermined aliquots of the enzyme solution were carefully injected below the interface into the buffer. After allowing the surface pressure to stabilize, the interface was compressed. The surface pressure was monitored as a function of the molecular area. PM-IRRAS was performed on a KSV PMI 550 instrument (KSV Instrument Ltd, Helsinki, Finland) for the Langmuir monolayers. An incidence angle of 80° for the incoming light, which was continuously modulated between p and s polarizations at a frequency of 1500 cm−1, was employed. The PM-IRRAS signal is obtained from the reflectivities of both the p and s fractions of the light at the same angle of incidence. The difference between the signals divided by the sum gives the socalled PM-IRRAS signal. In this way, the effects of water vapor and carbon dioxide are significantly reduced so that information on vibrational groups present at only the air−water interface is provided. For such measurements, the monolayers were compressed until the desired surface pressure was attained, and the spectra were recorded for a minimum of 6000 scans. During measurement of the PM-IRRAS spectra, the monolayer was kept with a constant surface pressure owing to the constant action of the barriers, which compressed the monolayer continuously in case of a decrease in surface pressure. BAM (model: micro BAM3; KSV-Nima Instruments) was employed to obtain images of the monolayer at desired values of surface pressure. For additional measurements, aliquots of the aqueous subphase of the SP−AC−DMPA monolayer were withdrawn and analyzed by dynamic light scattering on Malvern Zetasizer Nano S, equipped with a 633 nm laser and detector, at a scattering angle of 173°, operating at 25 °C. The floating monolayers were then transferred onto solid glass supports. These supports, previously cleaned with KOH and ethanol, were immersed in the aqueous solutions contained in the trough before monolayer preparation. The monolayers were compressed to 40 mN/m, and the solid supports were vertically withdrawn across the air/film interface at a rate of 5 mm min−1. For multilayer films, the solid supports were subsequently immersed in the aqueous subphase under the same conditions employed for the withdrawn rending Y-type LB films. This procedure was repeated depending on the number of layers desired, with transfer ratio values ranging between 0.9 and 1.0 for further analysis. For all transfers, a constant surface pressure of 30 mN/m was maintained during passage of the substrate. For analysis of the LB films and examination of co-transfer of the enzyme, fluorescence spectroscopy (Spectrophotometer model RF-5301PC; Shimadzu) was employed at an excitation wave-

macromolecules, polysaccharides usually entrap a high content of water in their structures, which can help the enzyme to preserve its activity. It is reported, for instance, that polysaccharides from the microalgae Cryptomonas tetrapirenoidosa influence the adsorption and enzyme activity of urease on cationic lipid monolayers.11 Carrageenans are another class of polysaccharides with the potential to serve as matrices for enzymes. They serve as cell-wall components in several species of seaweeds, constituting sulfated linear polymers of β-D-galactose alternatively linked by α-1,3 and β-1,4 linkages. They are employed in the pharmaceutical and food industries because of their ability to form gels, increasing the viscosity of the medium. It is reported that different classes of carrageenans interact with lipid monolayers.12 Also, it is reported that carrageenans interact with lipid−enzyme systems, such as alkaline phosphate with a negatively charged lipid.13 As it is well established that carrageenans are suitable as part of lipid/polysaccharide systems, with or without enzymes, it would be interesting to employ carrageenans with a higher capacity of adsorption at the air−water interface, forming stable Langmuir monolayers. As they are soluble in water, their adsorption from the aqueous subphase can generate instabilities because processes of diffusion toward the interface, lateral diffusion at the interface, and molecular reaccommodation at the lipid monolayer are expected.12 As a result, the use of acylated carrageenans (ACs) may help overcome these problems, permitting the formation of highly stable carrageenan films at the air−water interface. In this present work, we employed ACs to form stable mixed lipid−carrageenan−enzyme monolayers. The enzyme chosen was sucrose phosphorylase (SP), an intracellular enzyme responsible for catalysis in the conversion of sucrose to Dfructose and α-D-glucose-1-phosphate. Therefore, this enzyme can be employed for sucrose sensing. Also, the ability to form lipid−enzyme LB films was previously reported by our group,7 motivating us to employ such a system. Lipid dimyristoylphosphatidic acid (DMPA) was employed as the most stable element for the formation of floating monolayers at the air−water interface, therefore being the auxiliary structural matrix. The structural and thermodynamic properties of the monolayers were studied with surface pressure−area isotherms, Brewster angle microscopy (BAM), and polarization-modulation infrared reflection−absorption spectroscopy (PM-IRRAS). The floating films were transferred onto a solid support through the LB technique, and the catalytic properties of the enzyme under these conditions were investigated.

2. MATERIALS AND METHODS The water used in all experiments was purified using a Milli-Q system (resistivity 18.2 Ω cm, pH ∼6.0). The lipid DMPA and enzyme SP from Leuconostoc mesenteroides were obtained from Sigma-Aldrich. DMPA was dissolved in chloroform (Synth), with a concentration of 0.5 mg/mL. SP was dissolved in an aqueous buffer solution of K2HPO4 (Sigma-Aldrich) and KH2PO4 (Sigma-Aldrich), with a salt concentration of 0.01 mol/L and pH ∼7.0. The final concentration of the enzyme was 0.55 mg/mL. AC was obtained by acylation, with palmitoyl chloride and methasulfonic acid used as catalysts (see Supporting Information). NMR and Fourier transformed infrared spectroscopy (FTIR) were employed to characterize the compound, revealing that 14% of the hydroxyl groups of carrageenan was substituted by the palmitoyl chain. AC was then dissolved in chloroform to a final concentration of 0.47 mg/mL. 5360

DOI: 10.1021/acs.jpcb.6b03468 J. Phys. Chem. B 2016, 120, 5359−5366

Article

The Journal of Physical Chemistry B length of 285 nm, with the glass placed directly in the fluorimeter holder and a slit-width of 1 nm. The catalytic activity of the enzyme was measured according to a method previously described in the literature.14 For this purpose, a solution containing 50 mmol/L sucrose (Synth), 50 mmol/L 2-amino-2-hydroxymethyl-propane-1,3-diol (Synth), 1 ́ mmol/L ethylenediaminetetraacetic acid (Exôdo Cientifica), and 5 mmol/L magnesium sulfate (Synth) was prepared. Aliquots (1 μg) of β-NAD (Sigma-Aldrich), phosphoglucomutase (SigmaAldrich), and glucose-6-phosphate dehydrogenase (SigmaAldrich) were added to the solution, with a final volume of 1 mL, and the pH was adjusted to 7.0 using hydrochloric acid (Synth). After dispensing the enzyme solution or enzymecontaining LB film into the cuvette, the absorbance at 340 nm was measured with time. Determination of enzyme activity in homogeneous medium was carried out for an enzyme solution of concentration 1 μg/mL. To assure quality control of the measurement of catalytic activity, the protocol was first performed for the enzyme in a homogeneous environment and subsequently for the LB films. Each value of enzyme activity shown is an average of four independent measurements with four different films. Values of absorbance at 340 nm were obtained at 1 s intervals, and enzyme activity was obtained for the first 2 min of the reaction after a lag time, always assuring a linear increase in this interval. Atomic force microscopy (AFM) was also employed for further characterization, and images were obtained in the tapping mode, employing a resonance frequency of approximately 300 kHz, scan rate of 1.0 Hz, and scanned areas of 5.0 × 5.0 μm2 on films deposited on mica. Nanogravimmetry using a quartz crystal microbalance (model QCM200; Stanford Research Systems) was employed to estimate the surface density of the film deposited on a surface bound by gold electrodes on a thin disk of quartz as substrate. The surface density of the deposited film was determined according to the Sauerbrey equation.15 All experiments were performed at a temperature of 25.0 ± 0.2 °C.

Figure 1. Surface pressure−area isotherms for DMPA monolayers with or without AC (1:4 mass/mass) and SP with proportions in mol in relation to those of DMPA indicated in the inset. The inset is an independent graph showing the surface pressure isotherm for AC spread alone on the air−water interface in the same amount as that in mixed monolayers.

but also because it has been successfully utilized as an immobilization matrix for other enzymes,7,17,18 including SP.7 The isotherm for DMPA (Figure 1) shows a typical behavior for this lipid when spread on this kind of buffer. Because of compression of the monolayer, the surface pressure lifts off at 110 Å2, denoting the beginning of the liquid-expanded phase. A region that reassembles a plateau is present between 90 and 60 Å2 and represents the transition to the liquid condensed state. With continuous compression to lower molecular areas, surface pressures attain values as high as 58 mN/m, and the monolayer then collapses. With AC, the isotherms are shifted to higher molecular areas as a consequence of AC incorporation. It is important to say that the amount of AC employed for this case is the same as that employed to obtain isotherms for pure AC (inset of Figure 1), and the proportions in mass are 1:4 (AC/DMPA). Although the isotherms are shifted to higher areas, the possibility of forming aggregates of AC monomers in mixed AC−DMPA monolayers cannot be discarded. On gradually introducing SP in the mixed AC−DMPA monolayers, the isotherms are progressively shifted to lower areas, which indicates the condensation of the mixed AC− DMPA monolayer. Successive additions of SP lead the isotherms to progressively lower areas. For proportions higher than 10 mol % of SP, the monolayers are not shifted anymore, which indicates a saturation point of the effect of condensation. This is in contrast to the results obtained for SP introduced in the aqueous subphase of pure DMPA monolayers,7 which show a shift in the isotherms to higher areas. The presence of carrageenan groups covering the polar heads of the phospholipid may influence the adsorption of the enzyme, causing a possible molecular rearrangement among SP, DMPA, and AC, condensing the monolayer. Another explanation for this effect would be the loss of AC from the air−water interface to the aqueous subphase, owing to aggregation with SP. This fact would also lead to a shift of the isotherms to lower areas. For this, dynamic light scattering measurements were carried out for aliquots obtained from the aqueous subphase, and they showed the presence of particles of 300−400 nm, expected for aggregates formed with AC and SP. This may then explain the shift in the isotherms to lower areas.

3. RESULTS AND DISCUSSION 3.1. Air−Water Interface. Figure 1 shows the surface pressure−area isotherms for the components involved in this article. SP, being highly soluble in water, does not form Langmuir monolayers, and when spread alone at the air−buffer interface, the surface pressure does not increase significantly. However, on compressing the interface, the surface pressure increases up to 3 mN/m.7 This surface excess of protein is probably caused by its hydrophobic residues. AC, being less soluble in water, presents a higher surface activity, and when its solution in chloroform is spread on the interface and subjected to compression, the surface pressure increases up to 16 mN/m (inset of Figure 1). Also, when higher quantities of AC are spread, higher values of surface pressure are obtained (as high as 30 mN/m). However, the isotherms obtained are not reproducible, probably because of self-aggregation of the molecule. Although AC contains acyl chains, which give it a considerable surface activity, its polar region consists of a chain of polysaccharides, which may aggregate or rearrange molecularly, not allowing for the formation of coherent Langmuir monolayers. With these facts in mind, a compound able to form stable floating monolayers should be used as a matrix to assure the presence of AC and SP at the air−water interface in a coherent molecular arrangement. For that, DMPA was employed, not only because of the ease of this lipid to be transferred as multilayers16 5361

DOI: 10.1021/acs.jpcb.6b03468 J. Phys. Chem. B 2016, 120, 5359−5366

Article

The Journal of Physical Chemistry B

The morphology of the monolayers was investigated using BAM. At high values of surface pressures, DMPA monolayers do not present differentiation of phases because of the homogeneity presented by the liquid condensed state.20 At this phase, DMPA forms coherent monolayers, but with AC, some domains appear, with or without SP (Figure 3), pointing to the formation of

To investigate the stability of the monolayers, the air−water interface was compressed to 40 mN/m, and the surface pressure was followed with time after the stop of the barriers. Figure 2

Figure 3. BAM images for AC−DMPA monolayers at 40 mN/m without (left) and with SP (right). Field of view: 3600 × 4000 μm2.

aggregates upon incorporation of AC. With the enzyme incorporated in the DMPA monolayer without AC, a homogenous pattern is also observed. As a result, both images in Figure 3 contain domains that can be attributed to the aggregation caused by AC. The PM-IRRAS spectra for DMPA monolayers (Figure 4) show how AC and SP influence the vibrational transitions of the species that are part of the monolayer. It is important to emphasize that the formation of aggregates at the air−water interface could influence the vibrational bands. For this reason, spectra were taken several times for different monolayer preparations. As any relevant differences are observed when these spectra are compared, the data shown in this article are representative of each system, independent of the formation of aggregates. Panel A shows basically the region for the CH stretching mode, attributed mainly to the alkyl region of the phospholipid. The band centered at 2847 cm−1 represents the symmetric stretch for CH in CH2 groups, and the other band centered at 2921 cm−1 is attributed to the asymmetric stretch for CH in CH2 groups. No relevant changes in these bands are observed in terms of relative intensities upon incorporation of AC and SP. The band centered at 2921 cm−1 for pure DMPA is shifted to 2917 cm−1 with AC, and this shift is maintained after the incorporation of SP. Possible effects caused by this shift may be disregarded considering the resolution of the equipment (8 cm−1). The smaller bands centered at 2890 or 2886 cm−1 can be attributed to CH stretches in CH3 groups. Panel B shows the regions attributed to the CO stretching mode. The band centered at 1740 cm−1 is attributed to the CO stretching mode of the phospholipid. The bands at 1624 and 1651 cm−1 are attributed to H−O−H bends for the water molecules present on the surface, being an effect resulting from the difference in reflectivity of the air−water interface covered and uncovered by the monolayer. The bands centered at 1681 and 1698 cm−1 are attributed to the CO stretches in AC. The band at 1667 cm−1 is more evident for the monolayer with SP, and it is attributed to the amide I vibration related to the polypeptide moieties of the enzyme. Panel C shows a band centered at 1465 cm−1 for the pure DMPA monolayer. This band is attributed to the CH2 bending mode, and it is split into two bands (centered at 1459 and 1473 cm−1) for the monolayer at which the enzyme is present. The band centered at 1339 cm−1 can be attributed to SO stretches present in AC, and the band centered at 1315 cm−1 is attributed to the α-helix for amide III vibrations (in-phase combination of

Figure 2. Stability curves for DMPA monolayers compressed to 40 mN/ m. Panel A represents the surface pressure vs time curve after compression of the monolayer to 30 mN/m. Panel B represents the treatment of curves according to the equation proposed in ref 19

shows that the monolayer composed of phospholipid and enzyme has a lower stability in relation to that of the monolayer composed of pure DMPA. This can be a consequence of the rearrangement of the supramolecular structure when the film is compressed out of equilibrium. On introducing AC, however, this instability is diminished, and the rate of decrease in surface pressure with time is similar to that reached for the pure lipid film. These results can be treated using the following equation proposed by Magett-Dana19 ln

(π f − π t ) = kt (π f − π i )

where πf is the final pressure, πt is the surface pressure for time “t”, πi is the initial surface pressure (40 mN/m), and k is the rate constant. The values of k are shown as an inset in panel B of Figure 2. The values assume negative values as a result of the decrease in surface pressure, and the final surface pressure was established for the value attained in 60 min. Therefore, the higher the magnitude of this constant, the higher should be the stability of the monolayer. For the monolayer with AC, we observe two regimes of linearity, with the second one attaining significantly high values of k, which suggests a higher stabilization of the enzyme-containing monolayer in the presence of the polysaccharide. 5362

DOI: 10.1021/acs.jpcb.6b03468 J. Phys. Chem. B 2016, 120, 5359−5366

Article

The Journal of Physical Chemistry B

Figure 4. PM-IRRAS spectra for DMPA monolayers (40 mN/m) with or without AC (1:4 mass/mass) and SP, with proportions in mol in relation to those of DMPA indicated in the inset. Panel A represents the 2800−2950 cm−1 range. Panel B represents the 1600−1775 cm−1 range. Panel C represents the 1300−1500 cm−1 range. Panel D represents the 1125−1250 cm−1 range.

N−H in-plane bending and C−N stretching vibrations). Panel D shows a band centered at 1165 cm−1, attributed to PO stretches (present in DMPA). This band is probably influenced by the SO stretching mode, whose band also appears in this region. The band centered at 1234 cm−1 appears only when SP is present and can be attributed to β-sheet structuring in amide III vibrations. Changes in the secondary structure of proteins usually influence mainly amide I and amide III vibrations.21,22 Although amide I bands present stronger peaks than those of amide III bands, the region for amide I bands is probably hidden by other bands present in the spectra. Therefore, for our case, analyzing the spectra in the amide III region shall give clearer pieces of information about the structuring of SP at the air−water interface. Therefore, it is evident in these spectra that AC and SP are incorporated in the phospholipid monolayers, with the structuring of the enzyme in both α-helices and β-sheets. 3.2. LB Films. Transfer of the floating monolayers to solid supports was first characterized to determine the transfer ratio for each deposition. This ratio should be close to unity for further analysis. The mass deposited was estimated by the QCM technique and is presented in Figure 5, showing that the surface density increases linearly with the number of layers. For all films, the surface density for one layer was discarded because of the high roughness of the first monolayer, which should affect the linearity of the graph. This roughness is attributed to the direct contact of the phospholipid with the solid support in contrast to that in other transfers in which the monolayer is transferred onto a surface already covered by phospholipid molecules, pure or mixed with AC and SP. In Figure 6, we can observe that films containing AC and AC + SP present a surface density higher than that for DMPA. These differences must be a consequence of the incorporation of the polysaccharide and enzyme. As the tilts for

Figure 5. Surface density of films of monolayers of DMPA with or without AC (1:4 mass/mass) and SP (6% in mol) transferred from the air−water interface at 40 mN/m onto quartz crystals as a function of the number of deposited layers. The surface density of the first layer was arbitrarily omitted.

each curve are similar, one could suppose that the co-transfer of AC or SP is restricted to the first layers, and the increase in surface density for the sequential depositions is attributed to the transfer of only DMPA molecules. However, one may have in mind that the mass exclusively attributed to DMPA when it is as a mixed monolayer should be lower. As the surface pressure of deposition was fixed at 40 mN/m for all films, the area per molecule depicted in the surface pressure−area isotherms is higher, which indicates a lower surface density of DMPA at the air−water interface when the monolayer was transferred to the solid supports. Anyway, this result shows a higher amount of 5363

DOI: 10.1021/acs.jpcb.6b03468 J. Phys. Chem. B 2016, 120, 5359−5366

Article

The Journal of Physical Chemistry B

Figure 6. AFM images of films of monolayers of DMPA with or without AC (1:4 mass/mass) and SP (6% in mol) transferred from the air−water interface at 40 mN/m onto mica supports. (A) One-layer DMPA + SP ; (B) one-layer DMPA + AC; (C) one-layer DMPA + AC + SP; (D) seven-layer DMPA + AC + SP.

Figure 7 shows the fluorescence spectrum for mixed LB films and confirms the presence of the enzyme in the structure because DMPA and AC should not present fluorescence. The tryptophan group, present in some amino-acid residues of SP, was excited at

material transferred from the mixed monolayer, which suggests co-transfer of AC and SP. Figure 6 shows the morphology of the films. For pure DMPA LB films, the literature shows a homogeneous pattern,23 with the absence of domains at this scale. With AC and SP, we observe the presence of heterogeneous surfaces, with domains determining images with defects, such as ridges, agglomerates, and holes. Panel A shows that for one-layer DMPA−SP films, large holes and valleys are recurrent, with small higher points noted along the surface. For the film with AC co-incorporated with DMPA (Panel B), the surface is relatively homogenous, with a pattern of heterogeneity observed at only the bottom of the image. It is important to emphasize that these images were taken from other areas of the same film, and the pattern is persistent. When the three components are present (Panel C), the surface becomes rougher, but a homogeneous pattern of grouped agglomerates with a sphere-like shape is observed. When this film becomes thicker (Panel D), the surface turns noticeably rougher, and no geometrical pattern can be described.

Figure 7. Fluorescence spectrum for 11 monolayers of DMPA with AC (1:4 mass/mass) and SP (6% in mol), transferred from the air−water interface at 40 mN/m onto mica supports as LB films. Emission wavelength: 285 nm. 5364

DOI: 10.1021/acs.jpcb.6b03468 J. Phys. Chem. B 2016, 120, 5359−5366

Article

The Journal of Physical Chemistry B 285 nm, and fluorescence emission was observed with a maximum at 333 nm. Other studies with mixed enzyme−lipid LB films also confirmed the presence of enzymes using fluorescence spectroscopy.7−11 On altering the number of layers, the amount of enzyme deposited did not influence the spectra but the presence of AC did. Films with DMPA−SP present an emission maximum at 366 nm, with broader bands.7 In general, this fact indicates that AC altered the molecular accommodation of SF in the LB film, causing emission spectra with narrower bands, with peaks at lower wavelengths. The enzyme activity of the systems employed in this work was then measured and shown in Table 1. These results primarily

conservation being even better than that for homogeneous environments. Denaturing of enzymes immobilized on solid supports is common, as enzymes strive to achieve a lower surface energy or may not adequately expose their catalytic sites.26 However, higher activities can be found if the enzyme is immobilized under mild conditions.27 Lipids can serve as protectors for enzymes,4,8 but lower enzyme activity areas are also reported for lipid− enzyme LB films,7,10 although preservation of catalytic activity can be improved for LB films.18 Milder conditions can be achieved with some strategies. For example, exopolysaccharides obtained from microalgae are reported as intermediates for urease coming from the aqueous subphase adsorbing onto cationic lipid monolayers at the air−water interface.11 Although these polysaccharides decrease the urease activity in solution, when co-immobilized with the cationic lipid and urease as LB films, they served to accommodate molecularly the enzyme, resulting in higher enzyme activities. Therefore, our result corroborates this idea, showing that AC acts as a matrix in the incorporation of SF in DMPA LB films. Obviously, additional experiments should be performed to employ these devices as biosensors. Different concentrations of sucrose may be exploited to estimate parameters such as sensitivity and enzymatic constants. Also, films with different number of layers could be explored. It is reported that enzyme activity either increases with the number of layers7 or is independent of the number of layers,8 with the activity restricted to the enzyme molecules incorporated in the outermost layer for the last case. The use of different amounts of enzyme immobilized per layer is also another kind of experiment that may be performed. As long as enhanced enzyme activity can be achieved for lower quantities of enzyme, a sensor with economy of materials can be obtained. At this point, these results help understand why some molecular architectures exhibit superior performance and which are the main molecular factors that influence the catalytic properties of the immobilized enzyme.

Table 1. Enzyme Activity for SP in a Homogeneous Medium or Immobilized in 11-layer AC−DMPA LB films (with Proportion in mol of SP Indicated in the Table)a relative proportion of SP in the monolayer (%)

lag time (min)

LB (4% SP) LB (6% SP) LB (8% SP) homogenous

32 ± 2 28 ± 2 28 ± 2 15 ± 1

activity (−ΔAbs340/min)

% to the homogeneous environment (1 μg/mL of SP)

% of relative activity after 30 days

27 ± 2 × 10−3 39 ± 2 × 10−3 40 ± 1 × 10−3 45 ± 1 × 10−3

60 87 89 100

74 87 87 67

a

Each value is an average of four independent measurements with four different films.

indicate that LB films are sensitive to the presence of sucrose, which is the first indication that they can be used as a sucrose sensor. The enzyme activity for a homogeneous medium is shown for comparison. Lower activities for enzymes immobilized on solid supports are commonly reported in the literature,7,24,25 and they are usually attributed to restrictions for the polypeptide structure in terms of molecular mobility, causing difficulty in the access of the substrate to the catalytic site of the enzyme. Also, a higher amount of enzyme in the medium could yield a higher enzyme activity. QCM data suggest that the amount of enzyme deposited is between 200 and 300 ng per layer, whereas for the homogeneous medium, 1 μg of SP as the total mass was employed. Also, comparing with the activities obtained for DMPA−SP films,7 the relative enzyme activity in relation to the homogeneous environment increases from the range of 4.2− 13.5% (for films not containing AC7) to 60−90% (for films containing AC). This fact indicates that AC enhances the properties of the enzyme for SP−DMPA LB films, possibly accommodating SP in a more favorable environment. It is important to note that a substantial increase in the enzyme activity was shown when the amount of enzyme incorporated in the floating monolayer was between 4 and 6%. However, no significant increase was observed when the amount of enzyme increased from 6 to 8%, indicating a saturation point. Interestingly, after 30 days, the enzyme activity was measured for a second time, and the conservation of the values was better for the LB film than that for the homogeneous environment. This fact indicates that the system was composed of AC, and DMPA promoted a mild environment as a matrix for the immobilization of SP. As a result, the enzyme activity of SP in the AC−DMPA matrix, detected in this proof-of-concept experiment, presents boosted properties such as higher values of catalytic activity when compared to those for DMPA−SP LB films. This fact may be associated with a better conservation of catalytic properties, with

4. CONCLUSIONS We demonstrated that SP can be incorporated in AC−DMPA Langmuir monolayers, as detected with surface pressure−area isotherms, PM-IRRAS, and BAM measurements. The enzyme could be transferred along the mixed film onto solid supports as LB films, confirmed by the transfer ratio, fluorescence spectroscopy, and AFM. The enzyme activity could be detected for mixed enzyme−lipid LB films, thus demonstrating that this architecture is suitable for sensing sucrose. The performance for LB films could be compared to that for the enzyme in homogeneous medium, resulting in systems with a higher stability after 1 month. The enzyme activity for 11-layer SF−AC−DMPA LB films was 60−89% from the homogeneous environment, depending on the proportion of the enzyme, and they have a persisting activity of around 80% after 1 month, in contrast to the homogeneous environment, which preserved less than 70% of the original activity. Consequently, a matrix constituted of DMPA and AC may have provided a favorable environment for preserving the catalytic activity of the enzyme, which can be associated with the interaction of the polypeptide structure with the phospholipid and polysaccharide. Such interactions facilitate the access of the analyte to the catalytic site of the enzyme and allow for catalyzing the conversion of sucrose to other products, which may have an impact on the environment. The novelty of this article lies in the fact that enzyme activity could be better 5365

DOI: 10.1021/acs.jpcb.6b03468 J. Phys. Chem. B 2016, 120, 5359−5366

Article

The Journal of Physical Chemistry B preserved with immobilization as an LB film in a proof-ofconcept experiment.

■

(12) Lopez, R. F.; Nobre, T. M.; Accardo, C. M.; Pernambuco, P. C.; Nader, N. B.; Lopes, C. C.; Caseli, L. Effect of carrageenans of different chemical structures in biointerfaces: a Langmuir film study. Colloid Surf., B 2013, 111, 530−535. (13) Nobre, T. M.; de Sousa e Silva, H.; Furriel, R. P. M.; Leone, F. A.; Miranda, P. B.; Zaniquelli, M. E. D. Molecular view of the interaction between ι-carrageenan and a phospholipid film and its role in enzyme immobilization. J. Phys. Chem. B 2009, 113, 7491−7497. (14) De Winter, K.; Cerdobbel, A.; Soetaert, W.; Desmet, T. Operational stability of immobilized sucrose phosphorylase: continuous production of alpha-glucose-1-phosphate at elevated temperatures. Process Biochem. 2011, 46, 2074−2078. (15) Sauerbrey, G. Z. Verwendung von schwingquarzen zur wagung dunner schichten und zur mikrowagung. Z. Phys. 1959, 155, 206−222. (16) Haas, H.; Steitz, R.; Fasano, A.; Liuzzi, G. M.; Polverini, E.; Cavatorta, P.; Riccio, P. Laminar order within Langmuir−Blodgett multilayers from phospholipid and myelin basic protein: a neutron reflectivity study. Langmuir 2007, 23, 8491−8496. (17) Pizzolato, F.; Morelis, R. M.; Coulet, P. R. Enzymatic nanolayers from proteolipidic Langmuir−Blodgett films for biosensors. Quim. Anal. 2000, 19, 32−37. (18) Caseli, L.; Siqueira, J. R., Jr. High enzymatic activity preservation with carbon nanotubes incorporated in urease-lipid hybrid Langmuir− Blodgett Films. Langmuir 2012, 28, 5398−5403. (19) Maget-Dana, R. The monolayer technique: a potent tool for studying the interfacial properties of antimicrobial and membrane-lytic peptides and their interactions with lipid membranes. Biochim. Biophys. Acta 1999, 1462, 109−140. (20) Pérez-Morales, M.; Pedrosa, J. M.; Martín-Romero, M. T.; Möbius, D.; Camacho, L. Ellipsometric study of a phospholipid monolayer at the air−water interface in presence of large organic counter ions. Thin Solid Films 2005, 488, 247−253. (21) Volpati, D.; Aoki, P. H. B.; Alessio, P.; Pavinatto, F. J.; Miranda, P. B.; Constantino, C. J. L.; Oliveira, O. N., Jr. Vibrational spectroscopy for probing molecular-level interactions in organic films mimicking biointerfaces. Adv. Colloid Interface Sci. 2014, 207, 199−215. (22) Cai, S.; Singh, B. R. A distinct utility of the Amide III infrared band for secondary structure estimation of aqueous protein solutions using partial least squares methods. Biochemistry 2004, 43, 2541−2549. (23) Pavinatto, F. J.; Caseli, L.; Pavinatto, A.; dos Santos, D. S.; Nobre, T. M.; Zaniquelli, M. E. D.; Silva, H. S.; Miranda, P. B.; de Oliveira, O. N., Jr. Probing chitosan and phospholipid interactions using Langmuir and Langmuir−Blodgett films as cell membrane models. Langmuir 2007, 23, 7666−7671. (24) Pastorino, L.; Berzina, T. S.; Troitsky, V. I.; Fontana, M. P.; Bernasconi, E.; Nicolini, C. Biocatalytic Langmuir−Blodgett assemblies based on penicillin G acylase. Colloids Surf., B 2002, 23, 357−363. (25) Nakagawa, T.; Kakimoto, M.; Miwa, T.; Aizawa, M. New method for fabricating Langmuir−Blodgett films of water-soluble proteins with retained enzyme activity. Thin Solid Films 1991, 202, 151−156. (26) George, S. P.; Gole, A. M.; Sastry, M.; Rao, M. B. Interaction of xylanase I with a fatty lipid matrix: fabrication, characterization, and enzymatic activity of the enzyme-fatty lipid composite films. Langmuir 2002, 18, 9494−9501. (27) de Araújo, F. T.; Caseli, L. Rhodanese incorporated in Langmuir and Langmuir−Blodgett films of dimyristoylphosphatidic acid: physical chemical properties and improvement of the enzyme activity. Colloid Surf., B 2016, 141, 59−64.

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jpcb.6b03468. Synthesis and characterization of AC (FTIR and 1H NMR) (PDF)

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Tel: +55 11 3319-3568. Federal University of São Paulo, Rua São Nicolau, 210, Diadema, SP 04021-001, Brazil. Notes

The authors declare no competing financial interest.

■ ■

ACKNOWLEDGMENTS We are grateful to FAPESP (2015/23446-0), CNPq, and CAPES for sponsoring this study. J.M. Rocha was a CAPES fellow. REFERENCES

(1) Miyauchi, S.; Arisawa, S.; Arise, T.; Yamamoto, R. Study on the concentration of an enzyme immobilized by Langmuir−Blodgett films. Thin Solid Films 1989, 180, 293−298. (2) Dziri, L.; Boussaad, S.; Wang, S. P.; Leblanc, R. M. Surface topography of acetylcholinesterase in Langmuir and Langmuir− Blodgett films. J. Phys. Chem. B 1997, 101, 6741−6748. (3) Yano, Y. F.; Uruga, T.; Tanida, H.; Terada, Y.; Yamada, H. Rapid Xray reflectivity measurement using a new liquid interface reflectometer at SPring-8. J. Phys. Chem. Lett. 2011, 2, 995−999. (4) Girard-Egrot, A. P.; Godoy, S.; Chauvet, J. P.; Boullanger, P.; Coulet, P. R. Preferential orientation of an immunoglobulin in a glycolipid monolayer controlled by the disintegration kinetics of proteolipidic vesicles spread at an air−buffer interface. Biochim. Biophys. Acta 2003, 1617, 39−51. (5) Pavinatto, F. J.; Fernandes, E. G. R.; Alessio, P.; Constantino, C. J. L.; de Saja, J. A.; Zucolotto, V.; Apetrei, C.; Oliveira, O. N.; RodriguezMendez, M. L. Optimized architecture for Tyrosinase-containing Langmuir−Blodgett films to detect pyrogallol. J. Mater. Chem. 2011, 21, 4995−5003. (6) Jiao, J.; Leca-Bouvier, B. D.; Boullanger, P.; Blum, L. J.; GirardEgrot, A. P. A chemiluminescent Langmuir−Blodgett membrane as the sensing layer for the reagentless monitoring of an immobilized enzyme activity. Colloid Surf., A 2010, 354, 284−290. (7) Rocha, J. M.; Caseli, L. Adsorption and enzyme activity of sucrose phosphorylase on lipid Langmuir and Langmuir−Blodgett films. Colloid Surf., B 2014, 116, 497−501. (8) Zanon, N. C. M.; Oliveira, O. N., Jr.; Caseli, L. Immbolization of uricase enzyme in Langmuir and Langmuir−Blodgett films of fatty acids: possible use as a uric acid sensor. J. Colloid Interface Sci. 2012, 373, 69− 74. (9) Gazaryan, I. G.; Klaychko, N. L.; Dulkis, Y. K.; Ouporov, I. V.; Levashov, A. V. Formation and properties of dimeric recombinant horseradish peroxidase in a system of reversed micelles. Biochem. J. 1997, 328, 643−647. (10) Schmidt, T. F.; Caseli, L.; Viitala, T.; Oliveira, O. N., Jr. Enhanced activity of horseradish peroxidase in Langmuir−Blodgett films of phospholipids. Biochim. Biophys. Acta 2008, 1778, 2291−2297. (11) de Brito, A. K.; Nordi, C. S. F.; Caseli, L. Algal polysaccharides as matrices for the immobilization of urease in lipid ultrathin films studied with tensiometry and vibrational spectroscopy: physical-chemical properties and implications in the enzyme activity. Colloid Surf., B 2015, 135, 639−645. 5366

DOI: 10.1021/acs.jpcb.6b03468 J. Phys. Chem. B 2016, 120, 5359−5366