Dominant albumin-surface interactions under independent control of

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Dominant albumin-surface interactions under independent control of surface charge and wettability Shanshan Guo, Dicky Pranantyo, En-Tang Kang, Xian Jun Loh, Xiaoying Zhu, Dominik Ja#czewski, and Koon-Gee Neoh Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.7b04117 • Publication Date (Web): 10 Jan 2018 Downloaded from http://pubs.acs.org on January 11, 2018

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Dominant albumin-surface interactions under independent control of surface charge and wettability Shanshan Guo†, Dicky Pranantyo‡, En-Tang Kang‡, Xian Jun Loh ǁ,⊥, #, Xiaoying Zhu*,§, Dominik Jańczewski*,∇, Koon Gee Neoh*,†,‡ †

NUS Graduate School for Integrative Science and Engineering, National University of Singapore, Kent Ridge, Singapore 117576 ‡

Department of Chemical and Biomolecular Engineering, National University of Singapore, 4 Engineering Drive 4, Singapore 119260. E-mail: [email protected]; Tel: +65 6516 2176; Fax: +65 6779 1936 §

Department of Environmental Science, Zhejiang University, Hangzhou, China, 310058. E-mail: [email protected]; Tel: +86 571 88982651.

ǁ

Institute of Materials Research and Engineering, A*STAR (Agency for Science, Technology and Research, 2 Fusionopolis Way, Singapore 138634.

⊥

Department of Materials Science and Engineering, National University of Singapore, 9 Engineering Drive 1, Singapore 117576 #

Singapore Eye Research Institute, 11 Third Hospital Avenue, Singapore 168751

∇Laboratory

of Technological Processes, Faculty of Chemistry, Warsaw University of Technology,

Noakowskiego 3, 00-664, Warsaw, Poland. E-mail: [email protected]; Tel: +48 22 234 5583; Fax: +48 22 234 5504 KEYWORDS: layer-by-layer assembly, protein adsorption, protein conformational changes, protein adhesion force, bovine serum albumin, protein-polymer interaction

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ACS Paragon Plus Environment

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Abstract Understanding protein adsorption behaviors on solid surfaces constitutes an important step towards development of efficacious and biocompatible medical devices. Both surface charge and wettability have been shown to influence protein adsorption attributes including the kinetics, quantities, deformation and reversibility. However, determining the dominant interaction in these surface-induced phenomena is challenging due to the complexity of inter-related mechanisms at the liquid/solid interface. Herein, we reveal the dominant interfacial forces in these essential protein adsorption attributes under the influence of a combination of surface charge and wettability using a quartz crystal microbalance with dissipation monitoring (QCM-D) and atomic force microscopy (AFM)-based force spectroscopy on a series of model surfaces. These surfaces were fabricated via layer-by-layer assembly, which allowed two-dimensional control of surface charge and wettability with minimal cross parameter dependency. We focused on a soft globular protein, bovine serum albumin (BSA), which is prone to conformational changes during adsorption. The information obtained from the two techniques shows that both surface charge and hydrophobicity can increase protein-surface interaction forces and adsorbed amount. However, surface hydrophobicity triggered a greater extent of deformation in the adsorbed BSA molecules, leading to more dehydration, spreading and resistance to elution by ionic strength changes regardless of surface charge. The role played by surface charge in adsorbed protein conformation and extent of desorption induced by changes in ionic strength is secondary to that of surface hydrophobicity. These findings advance the understanding of how surface chemistry and properties can be tailored for directing protein-substrate interactions.

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1. Introduction Protein adsorption at aqueous-solid interfaces plays an important role in various areas of contemporary technology such as biosensors,1 medical implants2-4 or protein purification.5 For example, preadsorbed proteins on the implant surfaces mediate subsequent cell adhesion and tissue integration and thus affect healing. Protein adsorption is a complex process involving hydrophobic, electrostatic and van der Waals interactions, as well as hydrogen bonding.6 In addition, initial attachment is commonly followed by conformational rearrangements (e.g. unfolding and spreading) of proteins,7,8 due to changes in the thermodynamic state of the system involving the surface, proteins and solvent.2,9 Thus, proteins can attach to a surface in diverse quantities and conformations, depending on the local conditions. Both the adsorbed quantities and conformations are reported as important factors in implant biocompatibility.10 Surface wettability (or hydrophobicity), among the various surface properties, has been the subject of many studies. A number of proteins, including lysozyme3,11 and bovine serum albumin (BSA),11 adsorbed in higher amounts and were more strongly bound on hydrophobic polystyrene11 or methyl-coated silica surfaces3 compared to hydrophilic silica surfaces as a reference. Typically, proteins, e.g. lysozyme and α-lactalbumin, also deformed faster on hydrophobic silane-coated surfaces than on hydrophilic silica surfaces.7 However, these studies used a bare silica surface, which was negatively charged at physiological pH. Thus, it is difficult to distinguish the hydrophobic interaction of proteins with the surface from that caused by electrostatic interaction. Other studies of protein adsorption as a function of surface wettability showed that protein adsorption (e.g. BSA) did not increase monotonically with surface hydrophobicity,8 likely because the model surfaces used were not well-defined in terms of other possible relevant surface parameters such as surface charge and roughness.12 Self-assembled monolayers (SAMs) have been used as well-defined model systems to study protein adsorption on surfaces. The wettability of such surfaces is controlled by the terminal functional group of the SAM e.g. polar functional group OH and nonpolar group CH3.8-10,13 A higher amount of protein (e.g. collagen13 and BSA9) adsorbed on hydrophobic CH3 than on hydrophilic OH surfaces. These studies also showed that the degree of 3


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denaturation increased with surface hydrophobicity for proteins such as BSA,8,9,14 fibrinogen9,14 and collagen.13 Studies covering more extensive terminal functional groups including both ionic and nonionic chemical groups15,16 or mixture of two terminal groups14 demonstrated a direct correlation between the amount of protein adsorbed and surface hydrophobicity.15,16 However, when SAMs with different terminal groups are used to achieve variation of a single surface parameter, e.g. hydrophobicity, there is limited control over other properties. For example substitution of alkyl for hydroxyl group affects not only surface hydrophobicity but also surface zeta potential and the ability to participate in hydrogen bonding. The layer-by-layer (LbL) technique, consisting of self-assembled, sequential adsorption of oppositely charged polymers, offers opportunities to form coatings with tunable properties such as surface wettability17 and charge2,18-22 to manipulate protein adsorption. The effect of LbL surface charge on the adsorption behavior of different types of proteins, including BSA, fibrinogen and lysozyme has been studied. The results show that surfaces of charge opposite to that of the protein were more effective at promoting protein adsorption for different LbL systems.2,18-22 The effect of surface charge on the secondary structure of the adsorbed protein was also investigated. For LbL films assembled from poly(allylamine hydrochloride) and poly(styrenesulfonate), the structural changes of BSA were found to be larger when the charges of the LbL terminal layer and the protein were opposite.20 Surface wettability effects have been studied by altering the composition of the LbL films. For example, surface hydrophobicity was increased by increasing poly(styrene sulfonate) content in LbL films prepared from poly(styrene sulfonate) and poly(acrylic acid), and the adsorption of immunoglobulin G increased with surface hydrophobicity.17 Hydrophilic surfaces fabricated from LbL films of a diblock copolymer comprising a hydrophilic poly(ethylene oxide) block were shown to minimize protein (e.g. BSA and fibrinogen) adsorption.18 Less is known about the effect of LbL surface hydrophobicity on protein conformational aspects upon adsorption. Moreover, many studies focusing on engineering the surface wettability largely overlooked the influences on other surface parameters. For example, changes to surface

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wettability of LbL films assembled from poly(styrene sulfonate)/poly(acrylic acid)17 or alkylated polyethylenimine/poly(acrylic acid)23 were accompanied by significant change in surface roughness. Past studies using different engineered surfaces mainly focused on altering one parameter and much less is known about the relative and synergistic influence of surface charge and wettability on protein adsorption. Due to inter-related mechanisms involved in a multi-parameter study, a two-dimensional (2D) independent control over both charge and wettability is needed for minimizing cross-parameter influence. In our recent study, we developed a surface fabrication strategy that allowed us to independently adjust both surface charge and wettability using a LbL protocol24 with similar smoothness and chemical composition. In the current work, we used these model surfaces to study protein adsorption via quartz crystal microbalance with dissipation monitoring (QCM-D) and atomic force microscopy (AFM)-based force spectroscopy. Application of those techniques allowed us to study the essential protein adsorption attributes (e.g. kinetics, quantities, deformation and reversibility). By investigating the two relevant parameters collectively, the relative contribution of forces and dominant interactions acting between proteins and surfaces can be altered to provide a more complete picture of how the parameters affect protein adsorption. This will enhance our ability to predict or control protein adsorption attributes in a rational way. Our work is focused on a “soft” and globular protein, BSA, which is one of the most abundant blood proteins that potentially affect cell adhesion on medical implants.25 The effect of ionic strength and pH on BSA-surface interaction was also investigated.

2. Materials and methods 2.1

Materials

PEI (Mw 25 kDa, branched), poly(isobutylene-alt-maleic anhydride) (PIAMAn, Mw 60 kDa), 6-aminocaproic acid (Mw 131.17 Da), (3-aminopropyl)-trimethoxysilane (APTMS, 97%) and BSA (MW 66 kDa, lyophilized powder, ≥96%) were purchased from Sigma-Aldrich (St. Louis, MO, USA). Amine-PEG-carboxylic acid (NH2-

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PEG-CM, Mw 2000) was purchased from Laysan Bio Inc. Ethanol and toluene were purchased from Tedia. Silicon wafers were purchased from Latech Scientific Supply Pte. Ltd. 2.2

LbL film preparation and characterization

The polyanions PIAMA-C5 and PIAMA-PEG (chemical structures shown in Figure 1a) were synthesized as described previously.24 These polyanions were paired with branched PEI in the LbL assembly. Prior to LbL assembly, the silicon substrates were ultrasonicated in water and ethanol (10 min each), respectively. After drying with nitrogen, they were subjected to oxygen plasma treatment for 3 min at 160 W. The plasma-treated samples were coated with a precursor layer of positively charged amine by immersing in toluene containing 10 mM APTMS for 3 h. The polyions were then deposited by immersing the substrates in polycation and polyanion solutions (1.0 mg mL−1) for 5 min in a cyclic manner, up to 6.5 or 7 bilayers. 6.5 bilayers deposition resulted in films with a polyanionic outermost layer whereas 7 bilayers resulted in those with a polycationic outermost layer. Ultrapure water rinsing was performed between each immersion for 1 min. The pH of the polyelectrolyte solutions was adjusted by adding HCl or NaOH aqueous solution (0.1 M), and selected from a larger library of conditions used for fabricating LbL surfaces previously.24 The detailed fabrication conditions are presented in Table S1. Surface zeta (ζ) potential was measured by an electrokinetic analyzer (SurPass, Anton Paar). The electrolyte solution used was 0.001 M KCl. The pH of the KCl solution was controlled by auto pH titration with HCl (0.01 M). The average ζ potential value at a given pH was calculated from four repeats. Contact angles were measured using a goniometer (250-F1 from Ramé-Hart Instrument Co.) via the static sessile drop method. The average contact angle (θ) was calculated from four measurements on different locations. 2.3

AFM

Adhesion force measurements were performed using a JPK NanoWizard 3 NanoOptics AFM system in a liquid cell. Silicon dioxide colloidal probes (diameter 2.5 µm, Novascan Technologies, Inc.) were functionalized with BSA molecules according to a procedure reported previously.26 Briefly, the probes were treated with oxygen 6


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plasma at 120 W for 120 s, followed by vapor deposition of APTMS at 50 °C and curing at 90 °C to impart amine groups to the surfaces. The amine-functionalized probes were then immersed in glutaraldehyde solution (2.5 %) for 2 h, rinsed with phosphate buffered saline (PBS, pH 7.4), and immersed in BSA solutions (1 mg mL-1) in PBS for 3 h. BSA was chemically immobilized on the probe surface by formation of imine linkages between primary amine groups of BSA and aldehyde groups of glutaraldehyde. After rinsing with PBS, the BSA-coated probes were used immediately for AFM force spectroscopy measurement. Force maps of 64 force-versus-distance (f-d) curves were recorded on areas of 64 µm2, with a constant vertical scan rate of 1 Hz and approach and retraction speed of 1 µm s-1. Adhesion force histograms were obtained using ~250 f-d curves from 3 or 4 samples. The samples were tested on 3 separate occasions using a freshly prepared AFM probe on each occasion. Before force measurement, all systems were allowed to equilibrate for at least 1 h. The spring constant of the colloidal probe was measured using the thermal noise method, and the measured values were in the range of 0.08-0.12 N m-1. To obtain the f-d curves, the JPK SPM Data Processing software (v. 4.3.25) was used to process the raw AFM data acquired in terms of a photodetector signal in volts versus the relative piezo position (details on data processing are given in SI). The effects of ionic strength and pH on BSA adhesion forces on the LbL surfaces were investigated at three salt concentrations (10, 150, and 510 mM) and two pH values (3.6 and 7.4). The composition of buffer solutions used is summarized in Table 1. Table 1: Composition of buffer solutions used in AFM measurements Salt concentration pH

λD (nm)

Buffer type

NaCl (mM)

Total (mM)

Phosphate buffer (10 mM)

500

510

7.4

~0.4


140

150

7.4

~0.7


0

10

7.4

~2

Ionic strength variation

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Acetic acid buffer (10 mM)

140

150

3.6

~0.7


140

150

7.4

~0.7

pH variation

The integrity of the BSA coating on the AFM probe was assessed by comparing field emission scanning electron microscopy (FESEM) images of the probes before and after the AFM experiments (Figure S1). From a comparison of the FESEM images of an AFM colloidal probe fully coated with BSA prior to f-d measurements and a similar probe after ~1000 f-d measurements (Figures S1a and S1b, respectively), it can be concluded that the BSA protein remained on the probe after ~1000 f-d measurements. On the other hand, partial desorption of BSA molecules after ~2000 f-d measurements was observed (Figure S1c). Thus, the AFM measurements were kept within ~1000 f-d curves. In addition, the solutions and surfaces were exchanged in random order to check for any irreversible effect on the adsorbed protein layer and the measured forces showed random variation of adhesion forces without a systematic time-dependent change. This indicates that proteins did not desorb or undergo irreversible conformational changes during our AFM measurements. 2.4

QCM-D

QCM-D was performed using a Qsense E4 multichannel instrument. Quartz crystal disks coated with silicon dioxide (fundamental frequency 4.95 MHz) were used as sensor chips. The sensor surface was coated with LbL films prior to use according to the procedure described in Section 2.2. Protein adsorption experiment was conducted at pH 7.4 in buffer solution consisting of 10 mM phosphate buffer and 140 mM NaCl. Prior to protein adsorption measurement, the pure buffer solution was flowed into the QCM cell until a stable frequency (∆f) and dissipation (∆D) baseline reading was reached. Afterwards, the protein solution (100 µg ml-1) was passed through the measuring chamber. The adsorption process was carried out until surface saturation was achieved on each surface, and the required time differed among the surfaces. After surface saturation, the pure buffer solution was flowed into the cell to remove loosely attached molecules. Changes in ∆f and ∆D of the sensor chip were recorded in real time. The QCM experiments were performed on 2 or 3 independent samples. 8


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After protein adsorption, the adsorbed proteins on the chips were subjected to two cycles of buffer solution rinse at pH 7.4 using buffer solutions consisting of 10 mM phosphate buffer with different concentrations of NaCl. The proteins were first stabilized in buffer solutions with 10 mM phosphate buffer and 140 mM NaCl (the condition under which the proteins were adsorbed on the chips). The first cycling was between a buffer solution with high ionic strength (10 mM phosphate buffer and 500 mM NaCl, pH 7.4) and the adsorption buffer solution. The second cycling was between a buffer solution with low ionic strength (10 mM phosphate buffer without NaCl, pH 7.4) and the adsorption buffer solution. Each injection was carried out for 30 min to 1 h to reach stable frequency readings. The flow rate was controlled at 200 µL/min, which was high enough to allow a quasi-constant bulk concentration in the vicinity of the surface.27 The temperature was kept at 20.0 ± 0.1 °C. To calculate the hydrated mass of adsorbed proteins, the Sauerbrey equation was used for rigid adsorbed layers (∆D/∆f 100 nm, Table 2) during retraction. In contrast, the f-d curves on the hydrophilic surfaces LbL A and C returned to zero force at shorter distances. Such step-like features and large pull-out distances on LbL B and D suggest that the protein molecules were stretched and probably deformed during retraction.41,42 This is consistent with previous studies on protein unfolding triggered by surface hydrophobicity,9,14 which increases surface area of protein molecules available for interacting with the surface.43 Thus, the BSA structure can be easily deformed on the hydrophobic surfaces LbL B and D, leading to a variety of binding sites. This also accounts for the wider variability in the adhesion forces for the hydrophobic surfaces in comparison to the similarly charged hydrophilic surfaces as shown in Figure 2.

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Figure 3: Three representative f-d curves acquired with BSA-coated colloidal probes on different LbL films in a buffer solution of 10 mM phosphate buffer and 140 mM NaCl, pH 7.4. Dotted and solid lines represented the approaching and retracting traces, respectively. Insets provide a pictorial depiction of the interactions between the BSA-coated probe and the different surfaces. Table 2: Summary of AFM force measurements and QCM-D adsorbed mass and dissipation changes.a AFM Sample

Average adhesion forces (nN)

QCM-D Pull-out distance (nm)b

Adsorbed wet mass (ng mm-2)

|D/f| (10-6 Hz-1)c

Final |∆D/∆f| (10-6 Hz-1)d

|D/f |I=0.025; LbL A

3.40±0.46

89.8±35.9

12.00±1.95

0.091 |D/f |II=0.179

LbL B

5.81±1.76

167.7±63.29

7.07±0.33

LbL C

0.16±0.15

70.5±25.23

5.93±0.18

|D/f |=0.019

0.035

|D/f |I=0.180; 0.425 |D/f |II=0.760 LbL D

0.40±0.22

171.4±65.8

1.05±0.22

|D/f |=0.050

0.178

a

Buffer solution: 10 mM phosphate buffer and 140 mM NaCl, pH 7.4. Pull-out distance is defined as the separation distance at which the force returns to the zero force line in the retracting traces of f-d curves. c D/f values were obtained by fitting a line with r2 values >0.96 for all cases except for LbL D with r2~0.6 (drawn in Figure 5, at 3rd overtone). A good fit was difficult to acquire for small signal changes such as those on LbL D.44 Subscripts I and II refer to the two regimes exhibited by the ∆D/∆f curves in Figure 5. d Final |∆D/∆f| values were calculated for the 3rd overtone.

b

Considering the surface charge effect, on the negatively charged surfaces LbL C and D, electrostatic repulsion led to a small area of contact and weak adhesion. For the positively charged surfaces LbL A and B, their net charge is opposite to that of BSA. The resulting electrostatic attraction led to an increase in the area of contact and a concomitant increase in the adhesion forces. For LbL A, the profile of the f-d curves tends to be sharp, with few sequential and multiple rupture events and strong electrostatic attraction. This indicates that dominant electrostatic attraction has a limited influence on protein unfolding in AFM measurements. On the other hand, surface hydrophobicity changes the interaction landscape substantially, leading to accelerated unfolding of BSA and larger adhesion forces. This can be seen (Figure 3) when a comparison is made in the two pairs, LbL A vs 14


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B and LbL C vs D, where increasing surface hydrophobicity induced a larger area of contact and deformation due to the entropy gain from the dehydration of hydrophobic surfaces and reduction in the unfolding free energy barrier.9 Elevated attraction forces observed on LbL B, which is more hydrophobic than LbL A and has net charge opposing that of BSA, is an indication of the combined influence of electrostatic attraction and hydrophobic interaction. The unfolding of BSA under this combined influence allows for multiple site contact, leading to additional hydrophobic and electrostatic interactions. All four surfaces exhibit nanoscale roughness with LbL A, B and C having RMS roughness of ~1 nm and LbL D of ~4 nm (Table S2). The surface forces generated from rough surfaces may deviate from smooth surfaces due to changes in the local distance between two surfaces and nonuniform surface charge density.45,46 This in turn may affect the profile of the approach/retracting curves in the AFM measurements on a hard substrate.45,47,48 However, for measurement of adhesion forces on a soft polymer film, due to deformability of the material, nanoscale asperities can be squeezed and flattened out during adhesion in the force measurements to such a degree that the probe-substrate contact area is not significantly different compared to smooth surfaces.49-51 This reduces the effect of roughness on the measured adhesion forces,49-51 and hence, we expect such effect to be negligible in comparison to the influence of surface charge and wettability. Another technique used to investigate BSA interaction with the model surfaces was QCM-D. QCM-D detects both the adsorbed layers’ mass and energy-dissipating characteristics simultaneously and in real time, making it particularly useful for probing both protein affinity and layer properties.10,32 A solution containing BSA was passed over the QCM sensor coated with LbL A-D. The mass change (∆f) and viscoelastic change (∆D) were monitored as depicted in Figure 4. The injection of protein solutions was preceded by passing a buffer solution (pH 7.4) consisting of 10 mM phosphate buffer and 140 mM NaCl over the sensor until equilibrium, allowing the LbL films to be fully hydrated. Thus, the change in f and D signals after protein injection can be interpreted as adsorption of protein layers. The BSA solution was passed over the QCM sensor until equilibrium adsorption was reached. 15


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Figure 4: Typical measurements of ∆f (left axis) and ∆D (right axis) shifts of BSA adsorption onto LbL coated silicon dioxide sensors. The ∆f plotted was the normalized frequency. Plots include the 3rd, 5th and 7th overtones. For LbL A and C, which show well-separated ∆f and ∆D curves under the three overtones, |∆f| and ∆D values increase with decrease in overtone number. Protein adsorption was carried out by flowing protein solution with a concentration of 100 µg mL-1 at time t = 0. Protein exposure was terminated by exchanging the protein solution for a pure buffer solution in the rinsing step. Buffer solution: 10 mM phosphate buffer and 140 mM NaCl, pH 7.4. For all four surfaces, BSA injection led to ∆f decrease and ∆D increase, indicating BSA adsorption on all the LbL coated QCM sensors.10 The saturated mass adsorbed on each surface can be calculated from the Sauerbrey equation or Voigt model using QTools software (Q-Sense).28 A greater adsorbed mass was observed on the positively charged surfaces (12.00±1.95 ng mm-2 for LbL A and 7.07±0.33 ng mm-2 for LbL B) than on the negatively charged surfaces (5.93±0.18 ng mm-2 for LbL C and 1.05±0.22 ng mm-2 for LbL D). 16


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The QCM-D adsorption results complemented the AFM protein adhesion results and our earlier work in which adsorption of fluorescein isothiocyanate (FITC)-labelled proteins was studied (Table S3).24 Both the direct adhesion forces and the adsorbed mass increased with positive surface charge, confirming electrostatic interactions as important components of BSA-surface interactions. However, the QCM-D and FITClabelled protein adsorption results are different when surface wettability is considered as a parameter. The FITC-labelled BSA assay indicates that a larger amount of BSA was adsorbed on the hydrophobic surfaces while QCM result shows slightly higher mass on the hydrophilic surfaces. This difference in the results can be explained by the hydration effect of BSA molecules.35,52 The FITC-labelled BSA assay measures the ‘dry’ protein mass adsorbed whereas QCM-D measures the total hydrated mass, which includes both protein and the associated water. The lower mass on the hydrophobic surfaces compared to the hydrophilic surfaces as measured by QCM-D suggests that surface hydrophobicity triggered BSA denaturation and reduced hydration of BSA adsorbed on the surface. This is also reflected by a minimal increase in ∆D (softness) upon BSA adsorption and overlapping ∆f and ∆D curves under different overtones53,54 for the hydrophobic surfaces LbL B and D (Figure 4). This is consistent with earlier reports that BSA undergoes conformational changes on hydrophobic surfaces and its hydration is reduced upon surface contact.10,29 The mass adsorption kinetics on the hydrophilic and hydrophobic surfaces show different patterns (Figure 4). On the hydrophobic surfaces LbL B and D, adsorption attained saturation rapidly within several minutes, whereas on the hydrophilic surfaces LbL A and C, the initial rapid adsorption was followed by a slower increase in mass before finally levelling off after typically several hours. This behavior is independent of protein adsorption concentrations (Figure S4), indicating that it is insensitive to molecular packing over a range of concentrations. To further analyze the differences in the BSA adsorption kinetics, ∆D vs ∆f curves (D-f plots) were plotted to present the dissipation change per unit of adsorbed mass on each surface (Figure 5).15,55 The slopes of the D-f plots (D/f) reflect the dynamic changes in the protein layers’ conformation.55 The two hydrophobic surfaces LbL B and D exhibit a low D/f value (Table 2), denoting the formation of a rigid protein layer, with protein 17


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adopting a more spread out and deformed conformation.55,56 This is also consistent with previous studies on methyl-terminated SAMs.9,29 In contrast, for the two hydrophilic surfaces LbL A and C, the D-f plots can be divided into two regimes: an initial fast regime with relatively low D/f and a second slow regime with linear and high D/f. The high D/f value in the second regime indicates a significant increase in the softness of the adsorbed protein, denoting a conformation transition in accordance with the adsorption of a second protein layer which is more readily penetrated by water, more loosely bound, and more native-like.55 The differences observed in the D-f plots may be due to the different adsorption kinetics on the hydrophobic and hydrophilic surfaces. As BSA adsorption progressed rapidly toward a deformed state on the hydrophobic surfaces, further structural rearrangements to accommodate more protein adsorption were inhibited.10 In contrast, on the hydrophilic surfaces, the slower and weaker adsorption allowed a small extent of structural rearrangements to accommodate additional protein, giving rise to a second adsorption regime. Moreover, the transition takes place when the adsorbed mass reaches a critical f (surface coverage) indicating that the interactions between the adsorbed protein molecules play a role in this transition. The differences in the conformation rearrangements of the protein layers led to different final ∆D/∆f ratios (Table 2), which indicate the ultimate viscoelastic nature of the adsorbed protein layers. The final ∆D/∆f ratios were also lower for the more hydrophobic surfaces than the hydrophilic ones with similar charges, indicating the formation of a less hydrated and stiffer protein layer on the hydrophobic surfaces.57 Thus, the QCM-D results show that surface hydrophobicity increased adsorbed protein rigidity due to surface-induced deformation regardless of surface charge. On the hydrophobic surfaces, entropy gains from exclusion of water allow a greater proportion of the protein to interact with the surface,9 which may help to explain the greater conformational change of BSA on the hydrophobic surfaces LbL B and D. Upon binding to these hydrophobic surfaces, as a result of protein conformational changes, the exposure of inner regions of the protein allows for additional hydrophobic and electrostatic interactions. On the other hand, surface charge opposing that of the protein increased adsorbed protein mass, but its role in adsorbed protein conformation is secondary to surface hydrophobicity.

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Figure 5: Viscoelastic changes ∆D as a function of ∆f (3rd overtone). For the two hydrophilic surfaces (LbL A and C), the ∆D/∆f curves were divided into two regimes labeled as I and II with the time period indicated. Time increases from left to right as shown by the arrows. The fitted lines have r2 values >0.96 for all cases except for LbL D with r2~0.6. Buffer solution: 10 mM phosphate buffer and 140 mM NaCl, pH 7.4. Unlike the QCM-D adsorption experiment, AFM force spectroscopy measures the interaction strength between the protein and surface, which would not be affected by long-term conformational changes,37 lateral interactions between adsorbed protein,7 or multilayer formation processes58 that may be present in the former. Thus, it simulates the very early stage of protein adsorption.39 In general, the AFM force measurements agree with the macroscopic capacity of the different LbL surfaces to adsorb the BSA molecules as measured by QCM-D and FITC-labelled BSA assay. Both positive surface charge and hydrophobicity increased BSA-surface interaction 19


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forces and adsorbed amount. However, as suggested by the AFM and QCM-D experiments, BSA molecules were more deformed on the hydrophobic surfaces irrespective of surface charge, compared with those on the hydrophilic surfaces. The proposed mechanism also agrees with the protein layer thickness estimated using the Voigt model. On the hydrophobic LbL B and D, the estimated thickness values are lower, reaching ~5 nm and ~2 nm, respectively. On the hydrophilic LbL A and C, the estimated thickness is ~11 nm and ~9 nm, respectively. All the observed values fall within the range of published data.29,59 The low thickness values on the hydrophobic surfaces also suggest that BSA molecules adopted a more spread out and deformed conformation (Figure 6). BSA is known as a ‘soft’ protein prone to conformational changes during adsorption depending on surface properties.29 The results above are consistent with previous studies showing that protein deformation/spreading on hydrophobic surfaces occurs to a greater extent20,55 and at a higher rate than that on hydrophilic surfaces.7,14

Figure 6: Schematic of BSA molecular rearrangement on surfaces as a function of surface pI and θ. 3.3

BSA desorption

Removing preadsorbed proteins from a surface involves contributions from irreversible protein conformational changes and protein-protein interactions. To understand the role of the interfacial forces, protein unfolding and

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protein-protein interactions in protein desorption, the effect of ion concentration on desorption of the preadsorbed BSA molecules on the surfaces was investigated using QCM-D (Figure 7). As indicated in Figure 4, rinsing the BSA-loaded surfaces with a BSA-free physiological buffer solution generated negligible desorption, which is typical of adsorbed BSA on a variety of surfaces.60 To study BSA desorption at varying ionic strength, the BSA-loaded LbL surfaces were first equilibrated in buffer solutions with 10 mM phosphate buffer and 140 mM NaCl (the buffer in which BSA was adsorbed, λD ~ 0.7 nm, pH 7.4) to obtain the baseline. The BSA-loaded surfaces were then treated with buffer solutions characterized by high ion concentration (10 mM phosphate buffer and 500 mM NaCl, λD ~ 0.4 nm, pH 7.4) and low ion concentration (10 mM phosphate buffer without NaCl, λD ~ 2 nm, pH 7.4). As shown in Figure 7, for the two hydrophobic surfaces LbL B and D, the adsorbed mass (∆f) shows reversible changes upon exposure to high and low ion concentration solutions, indicating no appreciable protein desorption throughout the ionic strength cycle.61,62 This implies that the BSA molecules adsorbed on the hydrophobic surfaces were resistant to elution by ionic strength changes regardless of surface charge. This is consistent with a lower D/f value, i.e., a more spread out and deformed conformation of the adsorbed BSA molecules which maximized their footprint on the hydrophobic surfaces, as observed in previous sections. The reversible steep drops or rises observed in the adsorbed mass (∆f) during cycling (Figure 7) is due to the effects of salt mass and film swelling when added mobile ions disrupt some of the polycation-polyanion linkages in the film.63,64 Similar reversible mass changes (∆f) were observed on the native LbL films when exposed to the same ionic strength changes (Figure S5), indicating that the native LbL films maintained their structural integrity during the ionic strength cycling. In contrast, for the two hydrophilic surfaces LbL A and C, the changes in the adsorbed mass were irreversible, indicating mass losses from the adsorbed proteins triggered by changes in ion concentration. This is consistent with the more hydrated and native-like conformation of BSA molecules adsorbed in the second layer. The extent of mass losses was ~12 % and ~25 % for LbL A and C, respectively. BSA molecules desorbed from LbL C with increase in ion concentration. This desorption could be attributed to competitive adsorption of Clthat displaced BSA from the surface or screened localized electrostatic attraction.65 In contrast, on LbL A, the 21


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BSA molecules partially desorbed at lower ion concentration. Lowering ion concentration would increase the local electrostatic attraction between the surface and BSA molecules, which is expected to increase their binding and resistance to desorption. However, because of the large amount of BSA molecules adsorbed on LbL A, electrostatic repulsion among adsorbed BSA molecules could increase with decrease in ion concentration, which might lead to partial desorption of the outer protein molecules.66 The viscoelasticity changes of the adsorbed layers (∆D) were reversible upon ionic strength changes for all the surfaces (Figure 7). As changing ion concentration could cause rearrangements of protein layers66 and LbL films,63,64 and solvent/counterion expulsion/entrapment,62 which would be reflected in the dissipation loss/gain, the reversible changes in the viscoelasticity (∆D) of the adsorbed layers indicate that these processes were reversible upon ionic strength changes. For LbL A and C, this also indicates that the partial protein desorption caused minimal changes to the total viscoelasticity of the adsorbed layers.

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Figure 7: Adsorbed BSA molecules on the LbL surfaces exposed to buffer solutions of different ionic strength. To obtain the baseline, BSA-loaded surfaces were stabilized in buffer solutions with 10 mM phosphate buffer and 140 mM NaCl (the condition under which the proteins were adsorbed). This is defined as the ‘rinse’ step in the figure. High ion concentration (10 mM phosphate buffer and 500 mM NaCl) is denoted as ‘500’ in the first step. Low ion concentration (10 mM phosphate buffer without NaCl) is denoted as ‘0’ in the second step. Buffer solutions of pH 7.4 were used in these experiments. The QCM desorption results show that whether the adsorbed BSA molecules on the surfaces could be removed by changes in ion concentration was primarily determined by the degree of protein deformation, dictated by surface hydrophobicity, regardless of surface charge or multilayer formation. We can conclude that fairly stable protein layers composed of denatured BSA molecules were formed on the hydrophobic surfaces, regardless of 23


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their surface ζ potential. This is also in agreement with earlier research suggesting that the effects of surface hydrophobicity could even dominate over protein-protein interactions on protein unfolding,67 thus extending this surface-induced effects to proteins adsorbed in multilayers. 3.4

Effect of ionic strength and pH on BSA-surface interaction

The effects of ionic strength and pH on protein-surface interaction were examined by determination of ‘pull-off’ forces between the protein and surfaces after immediate contact by AFM force spectroscopy. Information on the strength and nature of the interaction, and the changes in the dynamics of the interacting protein can be obtained. The effects of ionic strength on protein-surface interactions include contributions from ionic screening, ion binding to the charged protein and surface, and protein conformational changes.68 Thus, AFM force measurements performed over an extended ionic strength range helps in understanding the competitive contributions from various interfacial forces and the importance of protein conformation for protein binding.69 We examined changes in the adhesion forces at different ion concentrations (Figure 8) at pH 7.4. The ion concentration was adjusted by adding NaCl to 10 mM phosphate buffer up to 0, 140, 500 mM NaCl, which corresponds to λD of ~2 nm, 0.7 nm and 0.4 nm, respectively. Phosphate buffer at pH 7.4 was used to maintain structural stability of BSA during the measurements.70 As shown in Figure 8, for the positively charged and hydrophilic LbL A, the adhesion forces decreased with increasing ion concentration. At pH 7.4, BSA is negatively charged. Therefore, there is attractive electrostatic interaction between BSA and LbL A. Increasing ion concentration screened the attractive interactions between them, leading to lower adhesion forces. Moreover, high ion concentration screened the electrostatic repulsion between protein residues, leading to a more compact protein conformation, which impeded adhesive bond formation with the surface.71 The specific effect of NaCl as a weak salting-out (protein-stabilizing) agent also contributes to formation of a more compact conformation at high ion concentration.72 Thus, the decrease in the interaction forces with increasing NaCl concentration is a combination of ions screening the interactions

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between BSA and LbL A, and a more compact protein conformation. This result indicates that the adhesion forces were mainly derived from electrostatic interactions between BSA and LbL A. A rather different picture is observed for LbL B, where the BSA adhesion forces on the positively charged and hydrophobic LbL B were increased by elevated ion concentration. A likely explanation for this effect is that at low concentration, ions bind strongly to the charged protein residues due to limited shielding.68 As a result, there is a higher energy penalty associated with the release of ions that accompanies protein adsorption, thus lowering adhesion forces.73 However, at high ion concentration, the electrostatic attraction between ions and charged protein residues is reduced due to charge shielding. Water-ion pairs are formed which reduce the number of water molecules around the protein and disrupt the hydration layer of proteins.74 This facilitates the interaction between protein hydrophobic residues and the surface, thus enhancing protein affinity for hydrophobic surfaces.74 Moreover, across the ion concentration range tested, step-like behaviors during retraction and large pull-out distances were observed in the f-d curves (Figures 3, S6 and S7), consistent with greater deformation driven by surface hydrophobicity. On the other hand, on the negatively charged surfaces LbL C and D, the average adhesion forces remained weak over the ion concentration range investigated. Due to like charges of the protein and the surfaces, electrostatic repulsion dominated the interactions between the protein and the surfaces. The repulsive forces were reduced with increasing ion concentration, accounting for the slight increase in adhesion forces with increasing ion concentration. Despite the strong electrostatic repulsion, hydrophobicity effects on BSA deformation for the hydrophobic surface LbL D were evident, as shown by the longer pull-out distances in the retracting curves for LbL D than those for the hydrophilic surfaces LbL A and C (Figures 3, S6 and S7).

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Figure 8: Average adhesion forces measured between BSA-coated AFM colloidal probes and LbL films under different ion concentrations. Buffer solution: 10 mM phosphate buffer with additional 0, 140 or 500 mM NaCl, pH 7.4. We further studied the BSA adhesion forces on LbL A and B by recording force curves at pH 3.6 and 7.4 while keeping the solution ion concentration constant at 150 mM. The charge of both the protein and the surface depends on the solution pH. BSA molecules carry a net positive charge at pH 3.6 and negative charge at pH 7.4. We have chosen LbL A and B (with pI~9) as model surfaces because they can maintain their positive surface charge at both pH values with limited variation (Figure S2). Thus, essentially only the charge of the protein was altered, allowing us to isolate the effect of protein properties from that of surface properties. The solution pH not only affects the charge of the protein but also its conformational state.75,76 26


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The average values and histograms of the adhesion forces between the BSA coated probe and the LbL films in media of different pH are shown in Figure 9. For both LbL films, the adhesion forces of BSA at pH 7.4 were greater than those at pH 3.6. The electrostatic forces between BSA and the LbL surfaces changed from attractive to repulsive as pH decreased from 7.4 to 3.6. Thus, electrostatic repulsive force weakened the adhesion forces of BSA with the LbL surfaces at pH 3.6. The hydrophobic LbL B exhibited an elevated adhesion force with BSA when compared with LbL A for both pH values. The increased surface hydrophobicity promoted BSA adhesion forces, despite pH changes. It is also possible that the BSA conformation exerted an effect on its adhesion forces, particularly at low pH. At pH 3.6, BSA is at its expanded state,75,76 and may expose more hydrophobic residues, thereby enhancing the interactions between BSA and hydrophobic surfaces.10 Moreover, for both pH conditions, multiple rupture events and step-like behaviors during retraction and large pull-out distances were observed in the f-d curves on the hydrophobic surfaces (Figures 3 and S8), consistent with a greater deformation driven by surface hydrophobicity.

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Figure 9: Average adhesion forces and force histograms measured between BSA-coated AFM colloidal probes and different LbL films (LbL A and B) at pH 7.4 and 3.6 with the same ion concentration. Buffer solution for pH 7.4: 10 mM phosphate buffer and 140 mM NaCl; Buffer solution for pH 3.6: 10 mM acetic acid buffer and 140 mM NaCl. Overall, solution pH and ionic strength affect the electrostatic interactions between protein and surfaces as well as the conformation of protein. The interplay of these effects is particularly obvious on LbL B, which involves both electrostatic and hydrophobic effects. On this surface, the protein conformation significantly affects the combined influence of these two effects: under attractive electrostatic effect (pH 7.4), a more easily deformed conformation (at higher ion concentration) led to much larger total adhesion forces due to additional hydrophobic and electrostatic interactions generated by unfolding. However, when electrostatic attraction rather than hydrophobic effects is the dominant force (LbL A), the profile of the force curves is sharp, with few 28


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sequential and multiple rupture events at all the solution pH and ionic strength investigated. This indicates that the effect of surface charge mainly alters the strength of protein-surface interaction, with less effect on protein deformation. On the other hand, it was consistently observed that protein deformation increased with surface hydrophobicity, which suggests that the role of surface charge is secondary to that of surface hydrophobicity in protein deformation on surfaces. 4. Conclusion In the present study, we utilized QCM-D and AFM force measurements to investigate BSA adsorption/desorption characteristics driven by a combination of surface charge and wettability using model surface platforms prepared via LbL technique that could simultaneously modulate the two surface variables. The AFM force spectroscopy results reveal the direct contribution of surface effects on protein-surface interaction, which complement the information provided by QCM-D on the adsorbed layer rigidity and packing. Through this combination of techniques, the influence of surface charge and wettability on the process of protein adsorption on the liquid/solid interface was delineated. Electrostatic attraction as the dominant force was found to have limited influence on protein deformation and extent of desorption by changes in ionic strength. On the other hand, surface hydrophobicity triggered rapid protein denaturation, leading to the formation of a rigid adsorbed layer and resistance to elution regardless of surface charge. The rigidity of the adsorbed protein layers as well as their desorption behavior on surfaces have strong impacts on their functionality and interaction with cells.25,77 The results obtained in this work may be important for understanding the varying functionality of the protein when adsorbed on different surfaces. For example, research has shown that the rigidity of adsorbed proteins on surfaces78 and the ease in displacing cell-repelling proteins such as BSA25 are important factors determining cell adhesion and tissue integration. We expect the results to contribute to the development of tailored-made surfaces to trigger more desired protein-substrate interactions for biomedical applications. Associated Content Supporting Information 29


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AFM data processing and colloidal probe (Figure S1); LbL film fabrication (Table S1); surface free energy and roughness (Table S2); representative surface ζ potential as a function of pH (Figure S2); film swelling measured by QCM-D (Figure S3); adsorbed mass measured by QCM-D and protein adsorption assay (Table S3); BSA adsorption at different concentrations (Figure S4); stability of polyelectrolyte layers in salt solutions (Figure S5); AFM force measurements in buffer solutions with 10 mM phosphate buffer and 500 mM NaCl (Figure S6), in buffer solutions with 10 mM phosphate buffer without NaCl (Figure S7) and in buffer solutions with 10 mM acetic acid buffer with 140 mM NaCl (Figure S8).

Author Information Corresponding Authors *(X.Z.) E-mail: [email protected]. Phone: +86 571 88982651. *(D.J.) E-mail: [email protected]. Phone: +48 22 234 5583. Fax: +48 22 234 5504. *(K.G.N.) E-mail: [email protected]. Phone: +65 6516 2176. Fax: +65 6779 1936. Notes The authors declare no competing financial interest. Acknowledgments We are grateful to the Agency for Science, Technology, and Research (A*STAR) for providing financial support and the A*STAR Graduate Academy for the PhD scholarship of S.G.

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Dominant albumin-surface interactions under independent control of

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