Stabilization against Denaturation at OilWater Interfaces - American

Chapter 19

Exploiting Favorable Silicone-Protein Interactions: Stabilization against Denaturation at Oil-Water Interfaces Downloaded by CORNELL UNIV on September 16, 2016 | http://pubs.acs.org Publication Date: March 10, 2003 | doi: 10.1021/bk-2003-0838.ch019

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Paul M. Zelisko, Vasiliki Bartzoka, and Michael A. Brook

Department of Chemistry, McMaster University, 1280 Main Street West, Hamilton, Ontario L8S 4M1, Canada

Water-in-silicone oil emulsions were prepared using either silicone surfactants containing pendant polyethylene oxide (PEO) side chains (DC3225C) or terminal Si(OEt) groups. The aqueous solutions contained either the enzymes α -chymotrypsin or alkaline phosphatase (DC3225C emulsions) or the surface active protein human serum albumin (TES -PDMS emulsions). Labelled albumin was shown to reside almost exclusively at the oil/water interface in the emulsion. The activity of the enzymes was followed over time by first breaking the emulsion and then performing standard enzyme assays on the aqueous phase. The enzymes were observed to undergo denaturation, as measured by reduced enzymatic activity, as a result of the mechanical energy used to make and break the emulsion. However, the rate of enzyme denaturation in the emulsions was lower than that observed for the aqueous control which had not been exposed to silicone. These results are consistent with favorable interactions between PEO or Si(OEt) groups (or hydrolytic byproducts in the latter case) and the proteins that not only stabilize the interface of a water -in-oil emulsion, but also the protein, which normally would otherwise undergo efficient denaturation in the presence of silicone oil. 3

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© 2003 American Chemical Society

Clarson et al.; Synthesis and Properties of Silicones and Silicone-Modified Materials ACS Symposium Series; American Chemical Society: Washington, DC, 2003.

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Introduction Silicones are exceptionally hydrophobic materials, which is partly the source of their impressive surface activity. By contrast, proteins are generally very hydrophilic, at least at their external surface, with die more hydrophobic amino acid residues typically being sequestered in the core of the folded protein. Relatively little energy is normally needed to perturb protein tertiary structure. Favorable interactions between the more hydrophobic protein core with silicone oils have been attributed to the facile denaturation that can occur if proteins are exposed to dimethylsilicone fluids (e.g., during emulsification): in order for the protein and silicone to favorably interact, the protein must change its conformation such that the hydrophobic moieties of its core are available at the external surface. In such a case, the hydrophobic domains on both polymers can associate. However, this change in conformation typically results in the catastrophic denaturation of the protein. Silicones are not, however, always deleterious to protein structure. Microparticles consisting of a starch/protein mixture, to which a surface coating of functional silicone ([(EtO)3Si(CH2)nSiMe (OSiMe2)m]20, TES-PDMS) had been added, served as a vehicle to deliver the entrapped protein to mice. That is, the entrapped protein elicited an immune response when administered orally. No such response was observed when the particles were coated with Me Si-capped silicone oils of comparable molecular weight. It was extremely surprising to learn that such a subtle difference in structure (two Si(OEt) groups on a polymer of MW 28000!) could have such a profound effect on the bioavailability of the protein in these microparticles. However, it was not clear whether the silicone was simply acting as macromoleeular protecting group, or whether a more profound interaction was taking place between the biological and the synthetic polymer. Assessing these interactions was particularly difficult with the starch microparticles, because of the presence of many solid/liquid and liquid/liquid interfaces. Therefore, we have examined, and describe below, the rate of denaturation of various proteins, including enzymes, when contained in water-in-silicone oil emulsions that contain a single liquid/liquid interface. We also examine the affinity of the protein for the oil/water interface. 2

Downloaded by CORNELL UNIV on September 16, 2016 | http://pubs.acs.org Publication Date: March 10, 2003 | doi: 10.1021/bk-2003-0838.ch019

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Experimental Section Reagents. D (Me SiO) and DC3225C were obtained from Dow Corning and were used without further purification. Two structurally different functional polymers were utilized in this study; commercially available Dow Corning 3225C, and (EtO) Si(CH ) -modified silicones, TES-PDMS. The latter polymers, with functional groups ((EtO) Si) at the tenriini, were prepared using methodology previously reported with molecular weights ranging from about 500-30000. 4

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214 α-Chymotrypsin, alkaline phosphatase, human serum albumin (HSA), iVglutaryl-Z-phenylalanine-p-nitroanilide, and /7-nitrophenyl phosphate (Sigma, Inc.) were used without further purification. The HC1 and trishydroxymethylaminomethane (Tris) used for preparing buffers were obtained from BDH while K H P 0 and absolute ethanol were supplied by Anachemia. The buffers comprising the aqueous phase of the emulsion were formulated using deionized, organic-free, distilled water containing 1.25 χ 10" g mL" of sodium azide (Aldrich). Calcium chloride was obtained from Aldrich. 5-(4,6Dichlorotriazinyl)aminofluorescein (5-DTAF) was obtained from Molecular Probes and used without further purification. Allyltriethoxysilane, hydrideterminated polydimethylsiloxanes (28000 MW), and platinum divinyltetramethyldisiloxane complex in vinyl-terminated polydimethylsiloxane (Karstedt's catalyst) were obtained from Gelest, and used without further purification. Fluorescein-labelled HSA. The 5-DTAF (0.5 mg) was added to 20.0 mL of an HSA solution (0.5 mg mL" ) in 1.0 M Tris-HCl buffer (pH 8). The mixture was protected from light and allowed to stand for 1 hour at room temperature. Dialysis was then performed using the Tris-HCl buffer as the exchange medium until such time as the exchange medium no longer displayed an absorbance peak at 492 nm. Emulsion Formulation for Enzyme Stability Studies Using DC3225C. The silicone surfactant DC3225C (1.0 g) was dissolved in D (29.2 g) (continuous phase) at a concentration of 3.54 wt% and added to the mixing vessel. a-Chymotrypsin (0.0118 g, at a concentration of 1.09 χ 10" g mL" ) was dissolved in 0.08 M Tris-HCl buffer (pH 7.8)/1.0 M CaCl (108 mL). Alkaline phosphatase (8.7 χ 10" g, at a concentration of 8.06 χ 10' g mL" ) was dissolved in a 0.1 M Tris-HCl buffer solution (108 mL, pH 7.8). These solutions constituted the dispersed phase of their respective emulsions. All enzyme solutions were prepared on the day of emulsification. The aqueous dispersed phase (10.0 g), containing the desired enzyme, was added drop-wise to the silicone continuous phase under dual blade, turbulent mixing conditions at 2780 rpm using a Caframo mixer over a period of 10 min. Emulsions were made in a Pyrex 180 mL beaker model number 1140. The bottom of the upper mixing blade (pitched at a 45° angle) was positioned at a height of 2/3 the liquid depth (from the bottom of the mixing vessel), while the bottom of the lower blade (at a 90° angle) was positioned at a position equal to 1/3 the diameter of the mixing vessel (from the bottom of the mixing vessel). The total height of the fluid in the mixing vessel (diameter 4.5 cm) was 3.0 cm. The emulsion was allowed to stir for an additional 20 min following addition of the dispersed phase. The enzyme solution (3.0 g) was stored in a sealed container at room temperature as a control. ' Emulsion Formulation using HSA and TES-PDMS. HSA (0.9 g, at a concentration of 0.03 g mL' (dispersed phase)) was dissolved in 1.0 M Tris-HCl buffer (30.0 mL, pH 7.8). A portion of the aqueous dispersed phase was stored in a sealed container at ambient temperature as a control. (Triethoxysilyl)ethyl2

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terminated PDMS (4.0 g) was dissolved in D (16.0 g, continuous phase) in the mixing vessel. The aqueous dispersed phase (20.0 g) was added to the silicone oil phase in a continuous, drop-wise manner over a period of 2 h under dual blade, turbulent mixing conditions at 2780 rpm using a Caframo mixer. The emulsion was allowed to mix for an additional 2 h following the addition of the dispersed phase. Enzyme Extraction. The enzymes (α-chymotrypsin or alkaline phosphatase) were extracted from the emulsion prior to the assessment of the enzyme activity. To do so, an aliquot (5.0 mL) of the emulsion was centrifiiged at 2500 rpm for 60 min, and the silicone supernatant was discarded. The concentrated emulsion was transferred to the mixing vessel. The Tris-HCl buffer (2.0 mL) was added drop-wise to the concentrated emulsion at a rate of 0.1 mL min" while stirring at 3000 rpm. The mixture was then allowed to stir for an additional 30 min following the addition of the buffer. Once extracted, the aqueous phase was drawn into a 5.0 mL syringe, filtered through a 0.2 pm pore syringefilterinto a vial, and stoppered. iV-Glutaryl-X-Phenylalanine-p-Nitroanilide Solution. The substrate solution for α-chymotrypsin was prepared by combining JV-glutaryl-Iphenylalanine-p-nitroanilide (0.1 g) with Tris-HCl buffer (20.0 mL, pH 8.0). Fresh substrate solutions were prepared each day that an activity assay was performed. /j-Nitrophenyl Phosphate Solution. The substrate solution for alkaline phosphatase was preparedfromp-nitrophenylphosphate (0.0393 g, 3.93 mM) in


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Tris-HCl buffer solution (10 mL, pH 8.0). Fresh substrate solutions were prepared each day that an activity assay was performed. Activity Assay for α-Chymotrypsin. After the emulsions were broken, enzyme activity assays were performed on days 1, 3, 5, 7, and 9 using a Cary 400 spectrophotometer. Concentrations of enzyme extracted from the emulsion were calculated at 256 nm using Beer's Law and an extinction coefficient (ε) of 26520 M ' for α-chymotrypsin. The extinction coefficient for α-chymotrypsin at 256 nm was calculated using known concentrations of the enzyme. The concentrations of extracted enzymes were normalized with the control. Assays were performed by adding Tris-HCl (1.25 mL) to iV-glutaryl-Z-phenylalanine-/?nitroanilide solution (1.25 mL) in an absorbance cuvette and equilibrating at room temperature for 5-10 minutes. Distilled, deionized water (10.0 pL) was added to the blank, and an enzyme solution (extracted enzyme or control, 10.0 pL) was added to the sample cuvette. The increase in absorbance at 400 nm was measured for 10 min to determine enzyme activity. The slope of the initial linear portion of the curve was taken as being representative of enzymatic activity. Activity Assay for Alkaline Phosphatase. The hydrolytic activity of alkaline phosphatase was quantified by following spectrophotometrically the formation of /?-nitrophenoxide at 405 nm. The /?-nitrophenol substrate solution (1.25 mL) and Tris-HCl buffer (1.25 mL) were transferred to a 4.0 mL polystyrene absorbance cuvette. An enzyme solution (10 pL), either control or extracted from the emulsion, was added to the cuvette and assayed. The molar 1


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extinction coefficient of alkaline phosphatase at 278 nm, in Tris-HCl buffer at pH 8.0, was taken as 18000 M" cm' . 1

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Results and Discussion Water-in-silicone oil emulsions were formed by the simple expedient of slow addition of the aqueous protein-containing phase to the stirring silicone oil containing the silicone surfactant. The speed of a dual blade mixer was adjusted such that the tip of the blade had a velocity of approximately 4.6 m s' . With DC3225C (Chart 1), emulsions were readily formed with D in the absence of protein. Such emulsions were extremely stable and are, in fact, relatively difficult to break. By contrast, surfactant TES-PDMS (MW 28000) (Chart 1) only led to stable emulsions if a protein, such as HSA, was present. The ease of forming the emulsion and the subsequent emulsion stabilities were also dependent on the specific protein and the concentration of the two cosurfactants. Stable water-in-oil emulsions could be prepared with water in concentrations exceeding 80% using DC3225C (data not shown). Emulsion droplets ranged in diameter from 1-10 pm with the majority having a diameter between 3-5 pm (Figure 1 A).


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Chart 1 10 11 12

Proteins typically deposit on solid surfaces. ' ' To address how proteins behave at silicone-water interfaces, that is, to determine if they are similarly attracted to more flexible liquid/liquid interfaces, human serum albumin was labelled with a commercially available fluorescein derivative. Stable emulsions of HSA were made in D with linear TES-PDMS (MW 28000, di-terminally functional). Previous studies had demonstrated that optimum emulsion stabilization in such systems arises at HSA concentrations of 0.03wt%. At this protein concentration, HSA primarily resides at the interface; confocal 4

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microscopy (observing a single 2D plane) with the fluorescently-labelled protein shows a protein corona, indicating preferential adsorption of HSA at the watersilicone oil interface over the bulk aqueous solution (Figure IB). It was of interest to determine, during the formation and aging of the emulsion, to what extent the proteins were undergoing thermal denaturation. ' This question can be addressed, to some degree, by following the ability of the protein to process substrate. In preliminary experiments, alkaline phosphatase (AP) was entrapped in a TES-PDMS/D emulsion. After two months, the protein was extracted from the emulsion by the addition of water and the substrate /7-nitrophenyl phosphate was added to the aqueous phase. The solution immediately changed color due to the presence of p-nitrophenoxide. This indicated that the enzyme was still capable after two months of catalyzing the transformation of the substrate. With this result in hand, the rates of denaturation of AP and α-chymotrypsin were tested in DC3225C/D emulsions. Chromogenic assays are readily available for both of these enzymes (Scheme l). ' Approximately 1.1 χ 10" wt% in water of the α-chymotrypsin and 8.1 χ 10' wt% in water of the alkaline phosphatase were emulsified, as noted above, in DC3225C/D at room temperature. Aqueous solutions of the protein in buffer were left at room temperature as controls. Each second day, for 9 sequential days, an aliquot of the emulsion was broken and the enzyme activity was measured using chromogenic assays (vida supra). The rates of substrate processing, which reflect residual active protein, are shown against time (Figure 2A,B). Protein that was never exposed to silicone is indicated by triangles (σ), and protein extracted from the emulsion is indicated by squares (v). It is immediately apparent that there is initially much less active protein in the silicone emulsion than in the control. However, the rate of denaturation for both proteins, as shown by the respective negative slopes of the two lines, is far higher for the control. 8 13


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a-chymotrypsin

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alkaline phosphatase

p-nitrophenyl phosphate (clear, colorless) ' v i

p-nitrophenol (yellow)

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p-ni roanil.ne (yellow)

A/-glutaryl-L-phenylalanine-p-nitroanilide (clear, colorless)

Scheme 1



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Figure 1: A water-in-oil emulsion ofHSA/Tris-buffer in 28000 MW (EtO)$Si(CHi) modified silicones/D . Average size of small (stable) droplets is about 3-5 microns. A: optical microscopy, B: confocal microscopy. r

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The mechanical energy used to form an emulsion is detrimental to protein structure. The classic example of this is denaturation of egg whites (ovalbumin) when whipped. Proteins (e.g., α-chymotrypsin) in buffer solutions that were subjected to the same mixing stresses involved in the emulsification process, but in the absence of silicone oil and silicone surfactants, were irreversibly denatured (data not shown). Similar degradation can be observed in Figure 2 although the magnitude is lower. The damage done to the proteins during emulsification and in breaking the emulsion manifests itself in an initially lower concentration of active protein. However, over time, little further change in enzyme activity is observed with the emulsified protein. By contrast, the control protein denatures quite rapidly. This suggests that once emulsified, the protein that is in contact with the silicone/water interface is relatively stabilized against denaturation. That is, the amount of damage by mechanical energy is a constant, and otherwise the proteins in the emulsion degrade at a very slow rate. Proteins can be stabilized by hydrophilic species. Thus, van Alstine and.co-workers have shown that grafting PEO onto proteins retards denaturation, a process that has been commercialized. The hydrophilic compounds . act as osmolytes, controlling structure by controlling swelling by water. It is likely that such stabilization is also occurring at the PEO-protein/water interface. That is, the surface-active components, both protein and DC3225C, migrate to the interface. There, the protein sits against the PEO, rather than the silicone, and benefits from the stabilizing environment. The situation is less clear for TES-PDMS. Such compounds should hydrolyze to initially give hydrophilic Si-OH (or Si-O") groups that can behave 16


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Β Figure 2: A: Reaction of alkaline phosphatase with p-nitrophenyl phosphate; B: reaction of α-chymotrypsin with N-glutaryl-L-phenylalanine p-nitroanilide.

in two important ways. First, they can anchor the TES-PDMS at the water/silicone interface (i.e., amplify the surface activity) and also stabilize the proteins through hydrophilic and, possibly, hydrophobic interactions (with adjacent Si-alkyl groups). However, it might be expected that Si(OH) groups would undergo room temperature vulcanization (RTV) cure to give elastomers. We have seen no evidence for this. Future work will focus on preparing welldefined silicone/protein copolymers to see if such linked proteins are similarly stabilized against denaturation and to further probe the specific nature of the protein silicone interactions in TES-PDMS. 3


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Conclusions


Enzymes and proteins denature over time. Denaturation is even more prevalent when the enzyme comes into contact with non-functional hydrophobic silicones and/or when subjected to shear stress during mixing. However, proteins residing at silicone oil interfaces bearing PEO- or triethoxysilylfimctional silicone groups are stabilized against denaturation. It appears that entrapping enzymes in water-in-silicone oil emulsions is an effective means of protecting these biomolecules from denaturation by the external environment.

Acknowledgements Financial support of this work from the Natural Sciences and Engineering Research Council of Canada (NSERC) and the Canadian Institute for Health Research is gratefully acknowledged. We would like to thank Professor Paul Berti, McMaster University, for helpful discussions.

References 1

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Telefax: +1(905) 522 2509; email: [email protected] (a) Owen, M . J. Siloxane Surface Activity, In Silicon-based Polymer Science: A Comprehensive Resource, Zeigler, J. M.; Fearon, F. W. G., Eds., American Chemical Society (ACS Adv. Chem. Ser. 224): Washington, DC: 1990, Chap. 40, p. 705. (b) Owen, M . J. Surface Chemistry and Applications, In Siloxane Polymers, Clarson, S. J.; Semlyen, J. Α.; Eds., Prentice Hall: Englewood Cliffs, NJ, 1993, Chap. 7, p. 309. Sun, L.; Alexander, H.; Lattarulo, N.; Blumenthal, N. C.; Ricci, J. L.; Chen, G. Biomaterials 1997, 18, 1593. Brook, Μ. Α.; Loombs, L.; Heritage, P.; Jiang, J.; McDermott, M . ; Underdown, B. Microparticle Delivery System, US Patent 5571531 (to McMaster University), Nov. 5, 1996. Heritage, P. L.; Loomes, L.; Jiang, J.; Brook, Μ. Α.; Underdown, B. J.; McDermott, M. R. Immunology 1996, 88, 162. Bartzoka, V.; McDermott, M . R.; Brook, M . A. Protein-Silicone Interactions at Liquid/Liquid Interfaces, In Emulsions, Foams and Thin Films, Mittal, K. L.; Kumar, P., Eds., Dekker: New York, 2000, Chap. 21, pp. 371-380.


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Bartzoka, V.; Chan, G.; Brook, M. A. Langmuir 2000, 16, 4589. Brook, M . Α.; Zelisko, P. M . Polym. Prep. (Am. Chem. Soc., Div. Polym. Chem.) 2001, 42(1), 97. Anderson, R; Vallee, B. Proc. Nat. Acad. Sci. USA 1975, 72, 394. (a) Du, Y. J.; Cornelius, R. M.; Brash, J. L. Colloids Surf. B, 2000, 17, 59. (b) Sheardown, H.; Cornelius, R. M.; Brash, J. L. Colloids Surf. B, 1997, 10, 29. Chapman, R. G.; Ostuni, E.; Takayama, S.; Holmlin, R. E.; Yan, L.; Whitesides, G. M. J. Am. Chem. Soc. 2000, 122, 8303. Norde, W.; Favier, J. P. ColloidSurf.1992, 64, 87. With HSA, at least, confocal microscope studies of labelled protein indicated that the majority of protein resides at the silicone interface. Hummel, B. Can. J. Biochem. Physiol. 1959, 37, 1393. (a) Bloch, W; Schlesinger, M . J. Biol. Chem. 1974, 249, 1760. (b) Chlebowski, J; Coleman, J. J. Biol. Chem. 1974, 249, 7192. (a) Burns, N. L.; van Alstine, J. M.; Harris, J. M. Langmuir 1995, 11, 2768. (b) www.shearwatercorp.com.


Stabilization against Denaturation at OilWater Interfaces - American

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