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Generation of Functional Coatings on Hydrophobic Surfaces throughDeposition of Denatured Proteins Followed by Grafting fromPolymerizationKiran K. Goli,† Orlando J. Rojas,‡,§ A. Evren Ozcam,∥ and Jan Genzer*,∥

†Department of Materials Science and Engineering, North Carolina State University, Raleigh, North Carolina 27695-7907, UnitedStates‡Department of Forest Biomaterials, North Carolina State University, Raleigh, North Carolina 27695-8204, United States§Department of Forest Products Technology, Aalto University, FI-00076 Aalto, Espoo, Finland∥Department of Chemical and Biomolecular Engineering, North Carolina State University, Raleigh, North Carolina 27695-7905,United States

*S Supporting Information

ABSTRACT: Hydrophilic coatings were produced on flat hydrophobic substrates featuring n-octadecyltrichlorosilane (ODTS)and synthetic polypropylene (PP) nonwoven surfaces through the adsorption of denatured proteins. Specifically, physisorptionfrom aqueous solutions of α-lactalbumin, lysozyme, fibrinogen, and two soy globulin proteins (glycinin and β-conglycinin) afterchemical (urea) and thermal denaturation endowed the hydrophobic surfaces with amino and hydroxyl functionalities, yieldingenhanced wettability. Proteins adsorbed strongly onto ODTS and PP through nonspecific interactions. The thickness ofadsorbed heat-denatured proteins was adjusted by varying the pH, protein concentration in solution, and adsorption time. Inaddition, the stability of the immobilized protein layer was improved significantly after interfacial cross-linking withglutaraldehyde in the presence of sodium borohydride. The amino and hydroxyl groups present on the protein-modified surfacesserved as reactive sites for the attachment of polymerization initiators from which polymer brushes were grown by surface-initiated atom-transfer radical polymerization of 2-hydroxyethyl methacrylate. Protein denaturation and adsorption as well as thegrafting of polymeric brushes were characterized by circular dichroism, ellipsometry, contact angle, and Fourier transforminfrared spectroscopy in the attenuated total reflection mode.

■ INTRODUCTIONModification of hydrophobic polymer surfaces is of greatinterest in current materials research. Activities in the fieldinvolve the development of coatings with applications thatrange from enhancing wettability or adhesion to protectingsurfaces from biofouling.1,2 Polyolefins, such as polyethylene(PE) and polypropylene (PP), possess desirable propertiesincluding good mechanical strength, chemical resistance,thermal stability, and low cost. These attributes make thempromising candidates for various applications involving func-tional textiles, filtration devices, medical implants, and manyothers.3,4 Unfortunately, the major drawback of PE and PP istheir low surface energy, which results in inherent hydro-

phobicity and poor biocompatibility.3,5,6 For example, materialsdesigned for surgical implants must be biocompatible and resistadsorption of biospecies to avoid blood agglutination.7 Thegreater susceptibility of hydrophobic surfaces to bioadhesioncompared to hydrophilic surfaces has been generally recog-nized, and numerous studies have been dedicated to thehydrophilization of surfaces using various physical and chemicalmethods.8,9 Grafting of macromolecules to surfaces isparticularly attractive because it allows for precise control of

Received: January 13, 2012Revised: March 12, 2012Published: March 13, 2012

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© 2012 American Chemical Society 1371 dx.doi.org/10.1021/bm300075u | Biomacromolecules 2012, 13, 1371−1382

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the chemical composition of the interface by adjusting thechemistry of the coating.10 However, the lack of functionalgroups in polyolefins and their high chemical resistance makegrafting a difficult task without employing harsh and aggressivepretreatments, which may adversely affect their bulk proper-ties.2,9,11

To date, several methodologies have been reported thatprovide routes for preactivation of hydrophobic surfaces.2,11

These approaches typically involve surface modification,including plasma treatment,2,12,13 irradiation with UVlight,11,14,15 electron beam,16 ozone,17,18 or γ-rays,19 oradsorption of surface active amphiphiles (i.e., copolymers orsurfactants).20 Alternatively, surface modification can beachieved during the manufacturing process by premixing(melt-blending) polymer precursors with surface-active agents(i.e., stearyl alcohol ethoxylates) that segregate to the surface ofthe initially hydrophobic films or fibers.21 Relevant workincludes that of Wang et al.5 who reported on grafting ofpoly(N-vinyl-2-pyrrolidone) onto ozone-pretreated PP non-wovens to enhance the hydrophilicity and antifouling propertiesof PP nonwovens. Lavanant et al.22 grafted poly(poly(ethyleneglycol) methacrylate) brushes onto polyolefin surfacespremodified by photobromination. The work by Desai et al.23

demonstrated that alkyl bromide groups generated onpolypropylene through photobromination could be used togrow N-isopropyl acrylamide brushes by atom transfer radicalpolymerization (ATRP). While these methods are relativelyefficient in endowing surfaces with functional groups, involvingsuch harsh treatments might compromise the mechanicalproperties of the substrates or supports.9 This issue isparticularly problematic in the case of relatively fine fiber-based materials.24 Our approach toward modifying hydro-phobic substrates is inspired by the known affinity of proteinsto solid surfaces (both hydrophobic and hydrophilic) as a resultof nonspecific interactions.25

Proteins are macromolecules composed of amino acids; theyadopt characteristic micelle-like conformations with character-istics that depend on the hydrophobicity/hydrophilicity of thesurrounding medium.7,26 For example, in aqueous solutions, thehydrophobic amino acids tend to be concentrated inside thecore of coil conformations, whereas the hydrophilic residues arepresent on the periphery. This arrangement results from thetendency of proteins to minimize the exposure of the morehydrophobic residues to water. However, a small portion ofhydrophilic and hydrogen-bond forming residues may also bepresent in the core, and a fraction of hydrophobic amino acidscan be located in the corona.27,28

Proteins adsorb readily on both hydrophilic as well ashydrophobic surfaces. The driving force for protein adsorptionon hydrophilic surfaces originates from the tendency of thehydrophilic groups to contact the hydrophilic substrate viahydrogen bonding or electrostatic interactions. In contrast,adsorption on hydrophobic surfaces is driven primarily byhydrophobic and van der Waals interactions among thehydrophobic protein domains and the surfaces while thehydrophilic amino acid moieties, which are not in a closeproximity to the adsorbing hydrophobic domains, are allowedto dangle to the exterior and interact with the aqueousenvironment.8,26,29,30 Because the hydrophobic residues arepresent predominantly inside the protein coil, proteinpartitioning on hydrophobic surfaces involves generallydramatic conformational changes that may result in coilunfolding and permanent destabilization, that is, denaturation

of the proteins.7,26,31,32 While in most instances adsorption ofdenatured proteins is considered to be a nuisance, primarilybecause proteins often lose their biological function, in thiswork we take advantage of this phenomenon to generatefunctional surfaces on hydrophobic supports.26,32,33

An effective coating of proteins on hydrophobic surfaces isachieved when a sufficient amount of protein is adsorbed fromsolution and van der Waals and other nonspecific interactionsbetween the protein and the substrate is maximized.7,34

Adsorption of proteins is a complex process that dependsstrongly on multiple variables, including those related to thedeposition solution (e.g., pH, ionic strength, temperature), theprotein (e.g., primary and secondary structure, which determinethe charge density and distribution and structural stability), andthe substrate (e.g., surface energy, charge, topography, mobilityof surface groups).28,30,32,35 Depending on the solution pH,protein molecules may contain either an effective positive or anegative charge. At the isoelectric point (pI), the net charge ofthe molecule is zero, although pockets of negative or positivecharges may be present locally. Under these conditions, theamount of protein adsorbed onto the surface is maximizedbecause electrostatic repulsion among protein moleculesadsorbing from solution is minimized.28,32 The density ofhydrophobic contacts between the protein and the surface canbe further maximized by not only relying on substrate-induceddenaturation, but also by denaturing the protein in solutionprior to protein adsorption. This denaturation can beaccomplished by adding well-known denaturants, such as ureaor guanidine hydrochloride, or by heating the protein solutionsto high temperatures.34,36,37

Denatured proteins attach to hydrophobic substrates viarelatively weak physical van der Waals and other nonspecificinteractions.26 Although the number of contacts between thesurface and the protein molecules is relatively high, thesecontacts may not be strong enough to hold the adsorbedprotein in place permanently. To the best of our knowledge,surface coating with denatured protein and the enhancement oftheir stability by cross-linking on hydrophobic substrates havenot yet been considered. More specifically, we propose toimpart protein layer stability by chemically coupling neighbor-ing adsorbed protein molecules with a cross-linker, such asglutaraldehyde (GA).38,39 Our approach has the advantage thateven a single layer of protein will alter the surface propertiesconsiderably and, thus will be an enabling platform to introducenew functionalities. For instance, the affinities of variousfunctional groups in proteins can be utilized (even afterdenaturation) to bind or scavenge metals from solution. Metalnanoparticles, i.e., silver-containing, can be attached to impartantibacterial properties, and polymers or amphiphilic macro-molecules can be installed to impart antifouling or otherproperties. Furthermore, surface modification with proteins issimple and, in contrast to traditional methods such as layer-by-layer (LbL) deposition of polyelectrolytes, it does not requiresequential adsorption steps. In addition, it is not subject tolimitations such as interlayer penetration and stability depend-ence on solution pH and ionic strength, which otherwise mayexist in LbL modification.40,41

This paper focuses on the formation of hydrophilicfunctional coating layers on model hydrophobic n-octadecyltri-chlorosilane (ODTS) surfaces by adsorbing proteins such as α-lactalbumin (LALBA), lysozyme (LYS), fibrinogen (FIB), andsoy globulins (glycinin and β-conglycinin). Prior to adsorptionon the ODTS surfaces, proteins were denatured using urea or

Biomacromolecules Article

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by heating to elevated temperatures. After adsorption, theprotein layer was immobilized by cross-linking with GA in thepresence of NaBH4. These protein coatings were used asprimers for further chemical modification. Specifically, wedecorated the outer surfaces of the protein coatings withpolymerization initiators that served as active sites for “graftingfrom” polymerization. Concurrently, we prepared hydrophiliccoatings on PP nonwoven surfaces by replicating thetechnological steps used on the flat ODTS surfaces. As proof-of-principle, we demonstrated this process by employing atomtransfer radical polymerization (ATRP) of 2-hydroxyethylmethacrylate (HEMA) to form dense PHEMA brushes;however, this technology is applicable to various otherpolymerization methods involving a wide range of monomers.

■ MATERIALS AND METHODSDeionized water (DIW; resistivity >16 MΩ cm) was produced using aMillipore water purification system. Silicon wafers (orientation [100])were supplied by Silicon Valley Microelectronics. ODTS was obtainedfrom Gelest (Morrisville, PA). PP nonwoven fiber sheets wereobtained from the Nonwovens Institute pilot facilities at NC StateUniversity and were cleaned with isopropanol prior to use. Lysozyme(from chicken egg white, Mn = 14.3 kDa, pI=11.3), fibrinogen (fromhuman plasma, Mn = 340 kDa, pI=5.5), CuCl (99.99%), CuCl2(99.99%), 2,2′-bipyridine (bpy, 99%), 2-bromopropionyl-bromide (2-BPB), 2-hydroxyethyl methacrylate (HEMA, 98%), glutaraldehyde(GA), sodium borohydride (NaBH4), and poly(allylamine-hydro-chloride) (PAH, Mw = 56 kDa) were obtained from Sigma-Aldrich andwere used as received. Poly(styrene sulfonate) (PSS, Mn = 75 kDa)was obtained from Polysciences, Inc. α-Lactalbumin (from bovinemilk, Mn = 14.2 kDa, pI = 4.3) was received from Davisco FoodsInternational Inc. Soy proteins were supplied by Archer DanielsMidland Company and glycinin (11S) and β-conglycinin (7S) wereobtained by standard fractionation methods.42 Phosphate-bufferedsaline (PBS) solution, HPLC-grade methanol, and toluene wereobtained from Fisher Scientific and used as received. Tetrahydrofuran(THF) and triethylamine were distilled from Na/benzophenone andcalcium hydride, respectively.Hydrophobic ODTS Layers Assembled on Silicon Wafers.

ODTS layers were prepared on silicon wafers as described in ourprevious work.43 The ellipsometric thickness of the obtained ODTSfilms were ≈25 ± 2 Å, which is consistent with the theoretical lengthof a covalently bound ODTS molecule of 26.2 Å.44,45 In addition,ODTS film thicknesses reported in the literature were between 20 and29 Å depending upon the tilt angles of the layers. Thus, we believe thatour ODTS layers were approximately a monolayer.46 Wettabilitymeasurements using DIW revealed that the contact angle was 108 ±2°.Denaturation and Adsorption of Proteins. α-Lactalbumin

(LALBA, pI = 4.3), lysozyme (LYS, pI = 11.3), and fibrinogen (FIB, pI= 5.5) solutions were denaturated using either urea treatment orthermal denaturation and adsorbed on surfaces as detailed below.For chemical denaturation, urea was added to PBS buffer (0.137 M

NaCl, 0.0027 M KCl, and 0.0119 M phosphates) and the solutionswere then filtered through a 0.2 μm filter followed by protein additionto obtain final concentrations of 0.01, 0.1, or 1 mg/mL. The solutionswere allowed to stand at room temperature for 6 h for completesolubilization. Protein adsorption was carried out at the isoelectricpoint of each protein (i.e., pH 4.3 for LALBA, 11.3 for LYS, and 5.5 forFIB); the solution pH was adjusted by adding either 0.1 N HCl orNaOH dropwise. Sodium azide (NaN3, 0.2%) was also added to thebuffer solution to prevent the growth of bacteria during adsorption.ODTS-coated wafers were used as model hydrophobic surfaces; theywere assumed to mimic the physicochemical characteristics of PE andPP in a flat geometry. ODTS-modified wafers and PP nonwovensheets were immersed in the chemically denatured protein solutionsfor 12 h at room temperature.

For thermal denaturation, LYS solutions of various concentrations(i.e., 0.01, 0.1, and 1 mg/mL) were prepared in PBS buffer solution atdifferent pHs of 7.4, 9, and 10. The solutions were allowed to stand atroom temperature for 6 h to solubilize the proteins and then kept in anoven preheated to 85 °C for 3 min before immersing the ODTSsubstrates for 15 and 30 min. Control experiments were run byimmersing ODTS substrates in protein solutions without heating forthe same duration. In addition, solutions of soy protein 7S and 11Sfractions of various concentrations (i.e., 0.01, 0.1, and 1 mg/mL) wereprepared at pH 7.4. Substrates were immersed in these solutions whileheating at 85 °C for 15 min. The substrates were then rinsed in PBSand DI water and blow-dried with nitrogen gas after removal fromprotein solution. The resultant flat ODTS substrates were charac-terized by ellipsometry and water contact angle measurements todetermine the thickness and wettability, respectively, of the substrateswith adsorbed protein coatings.

Circular Dichroism of Protein Solutions. Circular dichroism(CD) spectra for LYS and FIB proteins were measured by usingJASCO J-815 spectropolarimeter, equipped with 150 W xenon lamp.LYS and FIB were immersed into a quartz cell (path length 0.1 cm) atroom temperature with constant nitrogen gas purge. The CD spectrawere collected at a wavelength range between 195 and 250 nm with abandwidth of 2 nm, using steps of 0.2 nm, and a scanning speed of 50nm/min. Each spectrum reported here represents an average of at leastthree different scans. In all cases, the background contribution fromurea and buffer was subtracted.

Proteins were treated with urea or heat for chemical and physicaldenaturation, respectively. For denaturation using urea, LYS, and FIBprotein solutions were prepared at concentrations of 0.05 mg/mL attheir respective isoelectric points in PBS buffer. Different amounts ofurea (2, 4, and 8 M) were added to the resultant protein solutions.The resultant protein solutions were analyzed using CD to determinethe extent of protein denaturation. For thermal denaturation 0.05 mg/mL LYS and FIB solutions at pH 7.4 (PBS buffers) were heated in situfrom 40 to 90 °C with a heating rate of 2 °C/min while monitoringCD ellipticity at 222 nm, which is representative of secondary α-helixstructures.47

Cross-Linking of Adsorbed Protein Coatings and TheirStability. Protein-coated ODTS substrates and PP nonwovens wereimmersed in a solution containing 1% (w/v) cross-linker GA and0.01% NaBH4 (w/v). The interfacial cross-linking was allowed toproceed at room temperature for 6 h at pH 9, followed by rinsing withPBS buffer and DI water and blow-drying with nitrogen gas. Othershave reported on using GA to cross-link PEI and PAH, however, underdifferent conditions.48,49 The stability of the LYS protein coatings (1mg/mL, thermally denatured) was investigated by exposing theprotein-coated ODTS substrates (featuring either uncross-linked orcross-linked layers) to the following environmental tests: (1) dryingthe specimens at 85 °C for 12 h, (2) sonicating in DI water for 5 min,(3) incubating in THF solution for 12 h at room temperature, or (4)sonicating in 0.1 mg/mL nonionic Triton X-100 surfactant for 2 min.

Formation of Polyelectrolyte Layers. FIB-modified substrateswere placed in polycation solutions (0.02 M PAH, 0.5 M NaCl, pH 7)for 30 min, followed by rinsing in DIW and blow-drying with nitrogengas to form the first layer of PAH. The resultant PAH-functionalizedsubstrate was subsequently placed in a solution of PSS polyanion (0.02M PSS, 0.5 M NaCl, pH 7) for 30 min; this treatment provided asurface rich in negatively charged sulfonate groups. A second PAHlayer was deposited by dipping the PSS-functionalized substrate inPAH solution. In case of LYS-modified surfaces, PSS was deposited asthe first layer followed by the subsequent deposition of the PAH layer.The built-up of polyelectrolyte layers was carried out according to theprocedure as described by Balachandra and co-workers.50

Immobilization of ATRP Initiator on Protein-CoatedSurfaces. Initiator molecules were attached to amine and hydroxylgroups of protein-coated ODTS supports via amide and ester linkages.The substrate was immersed initially in a solution of 15 mL of dryTHF containing 0.363 g TEA. In another vial, 0.648 g of 2-BPBinitiator was added to 15 mL of dry THF. Both solutions were cooledto 0 °C prior to the addition of initiator solution. Initiator solution was



added dropwise to the TEA solution under continuous stirring withthe substrate present. The reaction was terminated after 2 min and thesubstrate was moved to a vial containing fresh THF followed byrinsing with ethyl acetate, ethanol, and DIW and blow-drying withnitrogen gas. TEA neutralizes the acid that would be released duringthe reaction between 2-BPB and the amine and hydroxyl groups.50 Asimilar procedure was used to immobilize initiator molecules over PPnonwoven surfaces.Preparation of PHEMA Brushes on Modified Surfaces via

ATRP. A typical “grafting from” polymerization of 2-hydroxyethylme-thacrylate (HEMA) included dissolving 9.09 mL (75 mM) of HEMAin a mixture of 9.09 mL (225 mM) of methanol, 1.82 mL (101 mM) ofDI water, and 0.54 g (3.39 mM) bpy. This mixture was degassed bythree freeze−thaw cycles to remove oxygen, followed by the additionof 0.172 g CuCl (1.74 mM) and 0.013 g CuCl2 (0.095 mM) undernitrogen gas atmosphere. The molar ratios of the various reactantswere [HEMA]/[bpy]/[CuCl]/[CuCl2] = 790:36:18:1. The resultantmixture was subjected to an additional freeze−thaw cycle. Theinitiator-immobilized ODTS substrate and PP nonwoven surfaces wereimmersed in this polymerization solution. The polymerization reactionwas allowed to proceed for 9 h at room temperature. Afterpolymerization, the substrates were removed from the reactionmixture, promptly rinsed with methanol and deionized water, anddried gently with nitrogen gas.51

Ellipsometry. The thickness of bare ODTS films before and afterthe formation of protein coating was determined by variable anglespectroscopic ellipsometry (VASE, J.A. Woolam Co.). In addition,ellipsometry measurements were carried out to monitor the “dry”thickness (H) of protein coatings, polyelectrolyte films, and PHEMAbrushes. Ellipsometric data were collected at an incidence angle of 75°for SAMs and protein coatings or 70° for PHEMA brushes usingwavelengths ranging from 400 to 1100 nm in 10 nm increments. Foreach coating layer, thickness values were determined for at least threedifferent spots and then averaged. The thicknesses were calculatedusing a Cauchy layer model assuming the index of refraction as 1.5 forODTS52 and 1.54 for protein coatings.51 A refractive index of 1.465was assumed for all polyelectrolyte layers.53 The thickness of PHEMAbrushes on protein coatings was calculated using the method used byArifuzzaman et al.51

Contact Angle Measurements. Contact angles (CAs) weremeasured with Rame-Hart CA goniometer (model 100-00) usingdeionized water (DIW) as the probing liquid at room temperature todetermine the wettability of protein coatings, polyelectrolyte films, andPHEMA brushes. Static DIW contact angles were recorded byreleasing 8 μL droplets of DIW on the surface. DIW contact anglemeasurements were carried out on at least three different positionsover the surface and then averaged.Fourier Transform Infrared Spectroscopy. The Fourier trans-

form infrared (FTIR) spectra for ODTS and modified PP nonwovensurfaces were recorded using a Bio-Rad-Digilab FTS-3000 Fouriertransform infrared spectrometer equipped with crystalline ZnSe in anattenuated total reflection (ATR) mode with a continuous nitrogengas purge. The nonwoven fiber was pressed against the crystal under auniform pressure of ≈700 psi using the micrometer pressure clamp.

The spectra reported represent an average of 5 accumulations of 64scans with a resolution of 4 cm−1. The data were analyzed using theBio-Rad Win IR Pro software.

Atomic Force Microscopy. Surface topography after proteindeposition on ODTS substrates was examined using Asylum ResearchMFP3D atomic force microscope (AFM). The AFM was operated inthe tapping mode in AC mode using Al-backside-coated Si cantileverswith a force constant of ≈5 N/m and a resonance frequency in therange of 120−180 kHz. During imaging, care was taken to keep the tipin the repulsive mode. The root-mean-square (RMS) surfaceroughness was calculated from height images using the softwareprovided by Asylum Research.

■ RESULTS AND DISCUSSION

Urea and Thermal Denaturation of Proteins. Proteindenaturation (thermal or chemical) and adsorption is illustratedschematically in Figure 1; the schematic indicates the disruptionof the native protein conformation by breaking hydrogen bondsthat stabilize the protein’s secondary structure (thus, unfoldingthe α-helix and/or β-sheet sections of the protein mole-cule).27,54

Figure 2a,b shows the far-UV CD spectra for native and urea-denatured LYS and FIB solutions at three different ureaconcentrations. Data reported here refers to the wavelengthrange 200−250 nm. The addition of urea prevents CD spectrameasurements below 210 nm for 2 and 4 M urea and below

Figure 1. Schematic representation of protein denaturation in solution and its subsequent deposition on a hydrophobic substrate. The latter processresults in the formation of a coating layer whose surface is comprising predominantly hydrophilic moieties present originally in the corona section ofa native protein.

Figure 2. CD spectra of urea-denatured protein solutions collected infar UV region for LYS (a) and FIB (b) proteins. The colored spectrumlines represent the native protein (black), protein with 2 M urea (red),protein with 4 M urea (green), and protein with 8 M urea (blue). CDspectra of LYS and FIB (c) with increasing urea concentration at 222nm.



215 nm for 8 M urea due to the high absorbance of the urea-solvent mixture. The CD spectra of native LYS and FIB arecharacterized by two minima located at 208 and 222 nm, whichcorrespond to the proteins’ secondary structures.47,55 The CDspectra of native protein obtained are in a good agreement withprevious reports.56,57 In general, molar ellipticity measurementsat 222 nm are considered to be the index of proteindenaturation and at this wavelength, α-helix has maximaloptical activity.47,56−58 To put these data into context, ellipticitymeasurements at 222 nm for a range of urea concentrations arereplotted in Figure 2c. The monotonous increase of negativeellipticities toward zero with increasing urea concentrationsindicates that both proteins under investigation exhibited areduction in the α-helix content due to the progressiveunfolding of LYS and FIB; the degree of unfolding is mostpronounced at urea concentrations of 8 M.Analogous results were reported by Barnes et al.37 for BSA

protein denatured with 8 M urea. Fernandez-Sousa et al.59 alsoobserved similar alterations in ellipticity values for LYS (frominsect eggs) proteins, denatured with 4% sodium dodecylsul-fate. The differences noticed in CD spectra for FIB and LYS areattributed to structural stability characteristics as well asdifferent amounts of secondary structures present in theindividual proteins; the α-helix contents present in LYS andFIB reported in the literature are 42 and 30−35%,respectively.60,28,61,62

We also unfolded protein structures using thermaldenaturation, which involves heating protein solutions.Thermal denaturation leads often to permanent conformationalunfolding of proteins, in contrast to denaturation using urea(because, in the latter case, protein can fold back partially withtime).38,54,63 As can be observed from Figure 3, the negative

ellipticities of FIB and LYS protein solutions decrease withincreasing temperature thus implying unfolding of proteins bybreaking the α-helices. Analogous reductions in ellipticity valueswere observed for FIB protein solutions heated from 40 to 110°C by Chen et al.61 The results clearly suggest that proteindenaturation increases with increasing protein solutiontemperature.

Adsorption of Urea-Denatured Proteins. The adsorp-tion of LALBA, LYS, and FIB proteins on ODTS substrates wasfollowed by determining the dry thicknesses of protein layers.Proteins have been reported to adsorb onto surfaces asmonolayers, submonolayers, or multilayers depending on theprotein type and experimental conditions.7 The dry thicknesswas calculated and expressed as a “fractional surface coverage”and is summarized in Figure 4 (top; details on the fractional

surface coverage calculation are provided in SupportingInformation). At low protein concentrations (0.01 mg/mL),the surface coverages of LALBA and LYS are comparable andsignificantly larger than that of FIB, indicating that theadsorption of proteins on hydrophobic surfaces depends onprotein molecular weight and conformation. Haynes et al.60

reported that LALBA and LYS possess similar sizes, shapes, andspecific densities. Unlike the smaller, globular LALBA and LYS,the complex and elongated FIB protein structure forms largeaggregates and prevents close-packing. These properties mightresult in relatively low FIB surface coverage. Considering themolecular dimensions of native LALBA (25 × 32 × 37 Å3),27

LYS (30 × 30 × 45 Å3),7 and FIB (60 × 60 × 450 Å3)7

proteins, one must note that the observed thicknesses aresmaller than those predicted for end-on or side-on proteinadsorption (that is, smaller than the respective protein size),indicating protein denaturation. On the basis of the aboveresults, these thicknesses correspond to the length of a fewamino acids that form loose structures that do not pack ideally,leading to fractional surface coverage smaller than unity.The surface coverage increases with increasing concentration

of protein in solution. The adsorbed amount (i.e., surfacecoverage) of LALBA seems to nearly saturate the surface at 0.1mg/mL concentration. This observation is in contrast to thecase of LYS and FIB; here, surface coverage increases

Figure 3. Ellipticity at 222 nm determined from CD spectra of thermaldenatured LYS and FIB protein solutions as a function of increasedtemperature.

Figure 4. Ellipsometry (top) and DIW contact angle (bottom) profilesof urea-denatured protein coatings on ODTS surfaces at various bulkconcentrations of denatured proteins. The horizontal dotted line in thebottom spectrum denotes the wettability of ODTS.



progressively with protein concentrations increasing from 0.01to 1 mg/mL. The reasons for these pronounced differenceswarrants further investigation, for example, by using the notionof “soft” (LALBA)7 and “hard” (LYS and FIB)28,64 proteins.LALBA, being a “soft” protein, adsorbs to a larger extent evenat lower concentrations, undergoing considerable structuralchanges upon adsorption onto the surface as compared to“hard” proteins, which undergo limited structural rearrange-ments due to stronger internal cohesion. At 1 mg/mLconcentrations it appears that all proteins have adsorbed in acompact manner.Further insight into the structure of the adsorbed protein

layers can be gained from static water contact angles. Assummarized in Figure 4 (bottom), DIW contact angle valuesdecreased from 108 ± 2° (for ODTS) to 61 ± 3° afteradsorption of LALBA, LYS, and FIB proteins, regardless ofprotein type and concentration. Our results are in agreementwith the findings of Wosten et al.65 who reported thatadsorption of hydrophobin protein on a Teflon surface resultedin the reduction of DIW contact angles from 108 to 63°. TheDIW contact angles of ≈61° observed in this study suggest thatprotein adsorption on ODTS considerably reduced the initiallyhydrophobic ODTS surface by exposing many hydrophilic (aswell as some hydrophobic) amino acid residues to the exteriorenvironment. Because the molecular structure of the outer layerof any denatured protein used here is not known, we cannotcomment on which sections of each protein are likely to beexposed to the outer surface and which parts are likely to be incontact with the underlying hydrophobic substrate. While thecontact angles of adsorbed LALBA, LYS, and FIB are all ≈60°,Wosten and co-workers mention that high wettabilities withcontact angles as low as 22° are known for hydrophobinprotein coatings;65 unfortunately, a detailed discussion aboutthe wettability behavior upon the structure of amino acids wasnot provided in the aforementioned publication. As we willdiscuss later in the paper, the soy protein fractions exhibitwettabilities that are slightly better than those seen for LALBA,LYS, and FIB. Clearly, the chemical composition of the protein,

its conformation and coverage on the surface (including anyeffects associated with topographical variation due to proteinadsorption) play a role in determining the final wettability.Although, it is important to understand and address themicroscopic structure of the outer protein layer, this topic isoutside the scope of this paper. As we demonstrate later in thepaper, each of the protein coatings provides a sufficient numberof anchoring points, which can be employed for attaching theinitiator molecules.

Adsorption of Thermally Denatured Proteins. Thedegree of protein unfolding depends on the amount of thermalenergy delivered. With prolonged heating at elevated tempera-tures, more unfolding occurs and can potentially be irreversible.The disruption of the secondary or higher order structuresexposes internal hydrophobic groups to the aqueous environ-ment.27,66,67 At this stage, hydrophobic parts of the proteinmolecules adsorb onto the hydrophobic surface throughdehydration. However, far away from the hydrophobicinterface, denatured proteins may tend to aggregate due tothe formation of intermolecular interactions, primarily consist-ing of van der Waals and hydrophobic forces.66−68

In our work, we carried out protein adsorption with andwithout heating for different adsorption times, pHs, andconcentrations. Away from the isoelectric point of LYS,specifically at pH 7.4, protein coverage did not depend onprotein concentration, the incubation time, or the degree ofdenaturation (cf., Figure 5, left ordinate). This behavior is dueto charges present over the protein molecules that minimizeadsorption as well as aggregation. Lu et al.29 has also observedsimilar behavior during the adsorption of LYS at roomtemperature away from the isoelectric point. As the solutionpH approaches the isoelectric point, the global net charge ofproteins becomes neutral, which results in an increase inprotein adsorption. In addition, we observed that denaturedproteins, which contain a higher surface density of hydrophobicgroups, adsorbed more extensively as compared to nativeproteins.

Figure 5. Ellipsometry (left) and DIW contact angle (right) profiles of parent/native (black squares) and thermally denatured (red circles and greentriangles) LYS protein coatings on ODTS surfaces at various bulk concentrations.



For protein bulk concentrations of 0.01 mg/mL and, in somecases, of 1 mg/mL, protein coverage is less than unity. Incontrast, protein coverage values exceed that of a singlemonolayer for protein adsorbed from solutions under theconditions of (a) pH closer to pI, (b) 1 mg/mL concentrations,and (c) adsorption times of 30 min at temperatures 85 °C (cf.,Figure 5). These conditions allow the proteins to aggregate insolution due to minimization of electrostatic repulsions andhigher probability of protein−protein contact. Similarly,Treuheit et al.67 reported that protein aggregation wasenhanced with increased protein concentration in bulk solution.In addition, they observed that unfolded proteins were morevulnerable to aggregation than native proteins. Under ourexperimental conditions, protein molecules adsorb as multi-layers either due to adsorption of aggregates formed in solutionor due to adsorption of individual protein molecules to surfacestempered with previously adsorbed proteins.Corresponding DIW contact angle measurements (cf., Figure

5, right ordinate) suggest that denatured protein layers possesshigher wettabilities as compared to undenatured protein layers.This effect might be attributed to higher concentrations of thehydrophilic protein residues present on the denatured proteincoating surfaces. As observed with the urea-denatured proteincoatings, the DIW contact angle of the absorbed LYS decreasedto 63 ± 3° (relative to 108 ± 2° measured for ODTS). Theonly exceptions are the native LYS protein layers and denaturedprotein layers formed with protein concentrations of 0.01 mg/mL at pH 7.4. We also observe that the contact angle increasesonce the fractional coverage is greater than one, wheremultilayers are present. The multilayers may consist of amixture of hydrophobic and hydrophilic patches present on thesurface, unlike monolayers, which are primarily composed ofhydrophilic groups oriented preferentially to the surroundingmedium. Overall, the DIW contact angle measurementsindicate that the wettablities of protein coatings on surfacesdo not depend on the mechanism of denaturation (at least forproteins investigated in this study).The application of protein coatings depends on the

availability and cost effectiveness of the protein molecules. 7S(β-conglycinin, 140−170 kDa) and 11S (glycinin, 340−375kDa) represent two major fractions of globulins, constitutingabout 75% of the total soy protein. Furthermore, 7S and 11Sglobulins have been the subject of work for modification ofhydrophilic surfaces (silica and cellulose).42 In the context ofthe present investigation, they are expected to be goodcandidates for modification of hydrophobic substrates. Toserve this purpose, thermally denatured soy proteins wereadsorbed onto ODTS surfaces to provide viable alternativescompared to relatively expensive functional proteins such asLALBA, LYS, and FIB. The denaturation temperatures of 7Sand 11S are ≈77.1 and ≈93.3 °C, respectively, as reported inthe literature.69 7S and 11S solutions were heated totemperatures of ≈85 °C for 15 min at pH 7.4. No visibleprecipitation of protein molecules was observed since the pH(=7.4) of protein solutions was away from the isoelectric pointsof the soy proteins (pI ≈ 4.5). Proteins were adsorbed onODTS substrates while maintaining protein solution temper-atures at 85 °C at pH 7.4. The resultant properties of soyprotein coatings are summarized in Figure 6. The coverage andDIW contact angle values of ODTS surfaces after adsorption of7S and 11S proteins are generally similar to those of LYScoatings. At low protein bulk concentrations of 0.01 mg/mL,the fractional coverage of 7S and 11S are smaller than those of

LYS, while at higher protein bulk concentrations, the proteincoverage of 7S and 11S are comparable to that of LYS.Wettability measurements reveal that, while at low proteincoverage, the DIW contact angle values are closer to 70 ± 3°,the DIW contact angle values reduce to ≈52 ± 5° withincreasing protein coverage. The latter values are lower thanthose observed in LALBA, LYS, and FIB. These results reiterateour earlier statements that the actual wettability depends on thelocal molecular features of the protein layer that are difficult toassess. Further investigation is needed to shed more light intothis phenomenon.The adsorption of soy proteins was demonstrated to confirm

that functionalizing the hydrophobic surfaces is not limited to aspecific protein. We have selected FIB and LYS proteins forfurther studies investigating protein layer stability andfunctionalization using polymer brushes, as discussed in detailin the next sections.

Cross-Linking and Stability of Protein Coatings.Adsorbed protein molecules were cross-linked to improve thestability of the adsorbed layers (see details about protein cross-linking in the Supporting Information). After cross-linking withGA, the thickness increased by 3 ± 1 Å due to theincorporation of GA as it reacted with the hydroxyl andamine groups of the protein. Cross-linking the protein layeralso resulted in a small increase in DIW contact angles. Thestability of the resultant protein layers (1 mg/mL LYS, pH = 9,thermally denatured) on ODTS substrates was investigated byexposing the substrates to conditions similar to those that maybe encountered in commercial applications. Specifically, LYSprotein coatings were selected for the stability tests since LYSdoes not form an inherent network structure, unlike FIB, whichforms networks even without cross-linking. The results,summarized in Figure 7, reveal that cross-linking the LYSlayer greatly improved the coating’s stability. The experimentsindicate that the fractional coverage and DIW contact angle didnot change when the cross-linked protein was exposed to these

Figure 6. Ellipsometry (top) and DIW contact angle (bottom) profilesof thermally denatured LYS and soy (7S and 11S) protein coatings onODTS surfaces at various bulk concentrations.



conditions. The uncross-linked protein layers showed areduction in coverage and increased DIW contact angle,especially in the presence of organic solvent and surfactant. Thelimited stability of un-cross-linked protein coatings may beattributed to the lack of in-plane cross-links among adsorbedprotein molecules. In contrast, penetration of surfactant andsolvent molecules and their ability to displace protein coatingsat the hydrophobic support will be limited in cross-linkedsystems.Deposition of Polyelectrolyte Layers and Surface-

Initiated ATRP of Polymer Brushes. Figure 8 showsschematically three methods employed based on FIB proteinprimers as substrates for surface-initiated polymerization ofHEMA. A stable cross-linked protein coating on top of ahydrophobic substrate was employed as a primer for attachingpolymerization initiator, 2-BPB, by reaction with hydroxyl andamine groups of protein molecules as represented in method(i). In addition, single (i.e., PAH1) or multilayers (i.e., PAH1/PSS/PAH2) of polyelectrolytes were formed through electro-static interactions on top of which the 2-BPB initiator wasattached, as shown in methods (ii) and (iii), respectively. Thereason for depositing polyelectrolyte layers was to create amore uniform and homogeneous distribution of functionalgroups throughout the substrate, to anchor the 2-BPB initiator.We considered that the usage of PAH as the outermost layer(instead of protein molecules) might make the composition offunctional −NH2 groups more uniform, as compared to theprotein-only primer, which possesses a heterogeneous dis-tribution of −OH and −NH2 groups. Furthermore, theaddition of polyelectrolytes may create stronger surfaces withfewer defects as well as improved chemical and mechanicalstability.70 The adsorption of each polyelectrolyte layer wasmonitored by ellipsometry and contact angle measurements.

The adsorption of polyelectrolytes was conducted at pH 7,where FIB is negatively charged (pI = 5.5), PAH is positivelycharged (pKa = 8.5), and PSS is negatively charged (pKa = 1).The data in Figure 9 (top row) show thickness increasesassociated with the formation of LbL polyelectrolyte assembliesas well as polymer brushes on FIB-coated ODTS surfaces. Theamount of adsorbed polyelectrolytes depends strongly on thetype of the underlying surface. In particular, the adsorbedamount of PAH1 (9 Å) on top of FIB layer was less than thePAH2 (16.4 Å) adsorbed on a 16.4 Å thick PSS layer. Thesefindings are attributed to the greater anionic charge densityover a PSS film compared to that for FIB at pH 7. AlthoughFIB acquires anionic charge globally at the experimental pHconditions employed, it contains some positive charges locally.Lvov et al.71 reported that the direct assembly of oppositelycharged proteins is difficult. This is because the electrostaticattraction between globular proteins is not optimal due to thepatchy nature of protein. In contrast, the electrostaticinteractions between polyions are optimal due to uniformdistribution of charges and flexible nature of syntheticpolyelectrolytes. Hence, we presume that the electrostaticinteractions between protein and a polyelectrolyte are less thanoptimal. One can take advantage of the sensitivity of contactangle measurements to monitor surface changes effectively. Theaddition of a PAH1 layer on top of FIB protein layer resulted inan increase of DIW contact angles from 59 to 70°. In general,DIW contact angles measured from the respective individualPAH and PSS layers alternate depending on the topmost layer.Our results indicate that the PAH-terminated layers exhibitDIW of ≈70°, while PSS-terminated layers are more hydro-philic (θ ≈ 53°). While it is possible to grow or add morepolyelectrolyte layers onto ODTS surfaces modified withpreadsorbed proteins, we limited ourselves to only a fewpolyelectrolyte assemblies, as explained above.We also employed the LbL method to prepare functional

coatings on top of LYS supports. At pH 7, LYS is positivelycharged (pI = 11.3). Thus, PSS was deposited above the LYSlayer followed by deposition of oppositely charged PAH. Thisled to two methods to grow the PHEMA brushes on the LYS:method (i), where the PHEMA brushes were directlypolymerized on the LYS, and method (ii), where thePHEMA brushes were grown on polyelectrolyte multilayers(i.e., PSS/PAH). The resultant thickness and contact anglemeasurements are summarized in Figure 9 (bottom row).Specifically, the adsorbed amount of PSS was 7 Å on top of theLYS layer and the subsequent PAH was 10 Å. This resultfurther supports the presumption that interactions betweenpolyelectrolytes and proteins are not as strong as theinteractions between polyelectrolytes. No large variation inDIW contact angle values (60° for LYS and 57° for the PSSlayer) was noted after the adsorption of the PSS layer above theLYS layer, which is in accordance with the thicknessobservations.Coupling of 2-BPB initiator to protein and PAH-terminated

surfaces resulted in a 4 ± 1 Å increase in thickness as measuredby ellipsometry. XPS confirmed coupling of the 2-BPB initiatorto the protein substrate (see Supporting Information). Thegrowth of PHEMA brushes from 2-BPB from bare FIB, LYS,and their polyelectrolyte-modified analogs was confirmed withellipsometry measurements. The thickness of the resultantPHEMA brushes produced from bare FIB and LYS films andtheir respective polyelectrolyte layers was 440 ± 15 Å. Inaddition, control experiments carried out on bare ODTS

Figure 7. (a) Ellipsometry (top) and (b) DIW contact angle (bottom)profiles of thermally denatured LYS layers on ODTS surfaces afterstability studies. The DIW contact angle of ODTS is 108°.



surfaces without adsorbed (FIB and LYS) proteins revealed noincrease in thickness, indicating that the 2-BPB initiator did notattach. After the synthesis of PHEMA polymer brushes fromthe surface, DIW contact angle decreased to ≈49 ± 2°. Controlexperiments carried out on bare ODTS resulted in no increasein dry ellipsometric thickness suggesting no growth of PHEMAbrushes. In addition, DIW contact angle measurements ofcontrol substrates remained at ≈99° indicating the absence ofPHEMA brushes above the surface. Thus, adsorption ofproteins onto ODTS substrates provides the foundation forthe growth of PHEMA brushes as described. Surprisingly, theresults in Figure 9 reveal that the thickness and wettability ofthe final PHEMA layers prepared directly on top of the proteinprimers are comparable to those formed on top of LbL films.Morphologies of LYS and PHEMA-Coated ODTS

Layers. Surface topography of bare, protein- and protein/polymer brush-modified substrates were studied by AFM. InFigure 10 we present AFM images of surfaces coated withODTS only, LYS layers deposited onto ODTS precoatedsubstrates, and PHEMA grafts grown from ODTS/LYS-modified surfaces. Our AFM observations reveal that hydro-phobic ODTS surfaces possess an RMS roughness of 0.24 nm.Adsorption of LYS proteins results in an increase of surfaceRMS roughness to 1.89 nm with the appearance of a fewstructural features. Overall, the protein deposition is uniformacross the sample. PHEMA modified samples appear to berelatively smooth on a global scale as judged from the RMS

roughness value of 1.11 nm. Though the RMS roughness valuesof the PHEMA-modified surface is less than that of the LYS-modified surface, these differences are relatively small and canbe attributed to experimental variation. On the local scale, theAFM image of the PHEMA samples reveals the existence oflocal features of in-plane heterogeneity. These structuralfeatures are not very large, that is, the variation between theextremes in height is ≈3 nm, which is less than ≈7% of the totalbrush height. This thickness heterogeneity may originate frommultiple sources, including, inhomogeneous deposition of theinitiator or inaccessibility of all initiating sites for polymer-ization. To understand the detected roughness effects, one hasto monitor the overall modification process in situ, which cangive some insight into the possible rearrangements of thefunctional primers after (or even before) initiator attachment oreven during polymerization. Unfortunately, direct observationof LYS deposition and the preceding phenomena is a verychallenging task and it is outside the scope of this paper.Additional work would have to be done to shed more light onthese phenomena.

Modification of Hydrophobic Fibers. To further verifythe adsorption of FIB protein molecules and growth ofPHEMA brushes over hydrophobic surfaces, similar experi-ments of protein adsorption and polymer brush growth wererepeated on PP nonwoven fiber surfaces. The deposition ofprotein coating and PHEMA layers onto the fiber supports wascharacterized by using FTIR-ATR. In Figure 11 we plot IR

Figure 8. Schematic depicting a build-up of protein and polyelectrolyte layers followed by subsequent ATRP polymerization of PHEMA brushes. Inmethod (i), the 2-BPB initiator is attached directly to the cross-linked FIB protein layer. In methods (ii) and (iii), 2-BPB gets anchored to substratescomprising a single (i.e., PAH1) or a multilayer (i.e., PAH1/PSS/PAH2), respectively, of polyelectrolytes deposited by the LbL method on top ofprotein primer.



spectra of (a) PP nonwoven surfaces, (b) FIB-modified PPfibers, and (c) PP nonwoven surfaces with PHEMA brushes.The appearance of new peaks located between 1700 and 1550cm−1 corresponds to the presence of amide I (CO

stretching) and amide II bands (C−N stretching and N−Hbending). In addition, the emergence of amide A (N−Hstretching) and O−H stretching between 3420 and 3250 cm−1

confirms the attachment of protein molecules to PP supports.

Figure 9. (top row) Dry thickness (left) and DIW contact angle (right) of PHEMA brushes grown from functional layers deposited on top of flatODTS/FIB-coated silica substrates. The layer comprising denatured FIB was cross-linked with GA/NaBH4. (bottom row) Dry thickness (left) andDIW contact angle (right) of PHEMA brushes grown from functional layers deposited on top of flat ODTS/LYS-coated silica substrates. The layercomprising denatured LYS was cross-linked with GA/NaBH4. Data corresponding to fabrication methods (i−iii) denoted pictorially in Figure 8 areshown in the top row. The data corresponding to LYS protein primer are shown in the bottom row. For clarity, the order in the DIW data has beenreversed relative to that of the thickness.

Figure 10. AFM images, 3 × 3 μm2, of control ODTS (left), LYS-coated ODTS (middle), and PHEMA-coated ODTS (right) surfaces.



Because protein molecules contain several amino acids, thebands appeared to be featureless and broadened in nature.72,73

PHEMA growth is evident from the appearance of a strongcarbonyl peak at 1720 cm−1. The peaks at 1250 and 1080 cm−1

are assigned to C−O stretching and O−H deformation of theC−O−H groups, respectively. The characteristic hydroxylstretching vibration extended to lower frequencies, with asignificant increase in its intensity as well as broadness between3500 and 3100 cm−1 also confirms the growth of PHEMAbrushes.3,51,74 In addition, the wetting experiments presented asvideos in the Supporting Information indicate clearly that thesurface properties of PP are altered after each modification step.Specifically, water droplets placed on top of unmodified PPnonwoven sheets “roll away”. In contrast, the protein-modifiedPP nonwoven exhibits an improved wettability, as shown by thebehavior of the water droplet, which does not “roll away”.Finally, a considerable increase in wicking was noticed after thegrowth of PHEMA brushes due to the presence of polarhydroxyl groups on the surface.

■ CONCLUSIONSDenatured α-lactalbumin (LALBA), lysozyme (LYS), andfibrinogen (FIB) were used to alter the physicochemicalproperties including surface chemistry (functional groups) andsurface wettability of hydrophobic ODTS and PP nonwovensurfaces through physical deposition after denaturation. Theresults demonstrated that the wettability of the protein coatingsformed on hydrophobic surfaces improved significantly,regardless of protein type and the denaturation methodemployed. Surface modification of hydrophobic surfaces byprotein molecules is very appealing because it impartsnumerous inherent amino acid functionalities of proteinswithout involving any harsh treatments. In addition, theproperties of soy proteins (glycinin and β-conglycinin) weresimilar to those of the expensive functional model proteins,suggesting the application of the former ones as potentialalternative surface modifiers. Surfaces functionalized withdenatured protein layers were subsequently employed assupports for surface-initiated polymerization of HEMA. Theproperties of the resulting PHEMA grafts on PP are similar to

those grown from model flat surfaces, indicating a high densityand stability of polymeric grafts produced on top of protein-modified hydrophobic surfaces. Our surface functionalizationapproach is very versatile and can be employed to alter any kindof hydrophobic material. Selected applications of thesefunctional fibers that might be explored include capture ofmetals or other contaminants from waters, prevention ofprotein adsorption, attachment of metallic nanoparticles, andmany others.

■ ASSOCIATED CONTENT*S Supporting InformationInformation on XPS analysis of initiator immobilized surfaces.In addition, the results from the SDS-PAGE analysis arepresented that confirm cross-linking of proteins in solutionsafter the addition of GA. The growth of PNIPAAm brushes onFIB- and LYS-coated substrates is also reported. The videoshows wettability of water on parent PP fiber mats before andafter modification with protein and after attachment ofPHEMA brushes. This material is available free of charge viathe Internet at http://pubs.acs.org.

■ AUTHOR INFORMATIONCorresponding Author*E-mail: [email protected]. Tel.: +1-919-515-2069.

NotesThe authors declare no competing financial interest.

■ ACKNOWLEDGMENTSThe authors thank the NC State University’s NonwovensInstitute for supporting this work through Project Number 07-106. The United Soybean Board (USB) is also acknowledgedfor their support on soy protein research under Project Number0426. We thank Carlos Salas (NC State University) forproviding the 7S and 11S soy protein fractions. We alsoacknowledge the Laser Imaging and Vibrational SpectroscopyFacility located at NC State University in the Department ofChemistry for help with FTIR spectroscopy. We thank Dr. Johnvan Zanten from the Biomanufacturing Training and EducationCenter (BTEC) for help with CD experiments and Dr. AmitNaik from Department of Chemical and BiomolecularEngineering for help with SDS-PAGE experiments.

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