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C: Surfaces, Interfaces, Porous Materials, and Catalysis
Critical Domain Sizes of Heterogeneous Nanopattern Surfaces with Optimal Protein Resistance Yun Li, Wenhao Qian, Jin Huang, Xianjing Zhou, Biao Zuo, Xinping Wang, and Wei Zhang J. Phys. Chem. C, Just Accepted Manuscript • DOI: 10.1021/acs.jpcc.8b01022 • Publication Date (Web): 17 Apr 2018 Downloaded from http://pubs.acs.org on April 18, 2018
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The Journal of Physical Chemistry
Critical Domain Sizes of Heterogeneous Nanopattern Surfaces with Optimal Protein Resistance
Yun Li, Wenhao Qian, Jin Huang, Xianjing Zhou*, Biao Zuo, Xinping Wang*, Wei Zhang Department of Chemistry, Zhejiang Sci-Tech University, Hangzhou 310018, China
*
Corresponding author. Xinping Wang & Xianjing Zhou Email:
[email protected] ;
[email protected] Tel/fax: +86-571-8684-3600 1
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ABSTRACT: In order to investigate protein-resistant surfaces with heterogeneous nanopatterns, V-shaped polymer brushes composed of a hydrophilic methoxypoly(ethylene glycol) (mPEG) arm and a hydrophobic polystyrene (PS) or fluorinated poly(methyl methacrylate) (PMMA-b-PFMA) arm were prepared in which the surface structure and phase-separation behavior were controlled by altering the relative lengths of the two arms. The protein resistance of these amphiphilic brushes was better than that of pure poly(ethylene glycol) (PEG) brushes, and when the domain size of the phase-separated structures was about twice the size of the protein molecules, the surfaces exhibited optimal protein repellence. At the same time, the amount of protein adsorption was well related to both the adhesion and the relative friction coefficient of the protein on the brush surface. A heterogeneous surface with phase-separated domains twice the size of protein molecules may be beneficial for minimizing protein adsorption through the synergistic effect of hydrophobic and water-soluble domains. These results provide an important way for designing and preparing protein-resistant materials with heterogeneous surfaces.
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1. INTRODUCTION Protein adsorption on solid surfaces is an ubiquitous problem that induces a series of undesirable effects, such as contamination blockage of filtration membranes, negative biosensor signals, inflammation on implants, reduction in the operating efficiency or shortened run times of various devices and thrombosis of indwelling medical devices in vivo,1 in the fields of food packaging,2 medical devices,3 ships,4 biosensors5 and other areas.6,7 During marine fouling growth, the key step is that proteins are adsorbed physically to form a conditioning film, which is also a major problem for in vivo nanomedicine applications. Therefore, it is of great significance to understand the mechanism of the interaction between proteins and material surfaces,8 which can ultimately lead to better design and preparation of protein-resistant materials. Antifouling materials are generally prepared according to the principle of minimization of interaction forces between a protein and a solid surface. Among the many approaches investigated, a number of hydrophilic molecules for surface modification, such as poly(ethylene glycol) (PEG),9 heparin10 and polysaccharides,11 can reduce adsorption of proteins due to a hydration layer that prevents the proteins from adsorbing to the surface. However, when these hydrophilic surfaces are used in complex media, they are usually considered to lose their protein-resistance performance due to their susceptibility to oxidation.12-14 Hydrophobic coatings (e.g., fluoropolymers15 and polysiloxanes16) also exhibited excellent fouling-release performance due to their low surface free energy and low Young’s modulus, which 3
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minimizes the adhesion work of the proteins, promoting the release of accumulated foulants with the aid of hydrodynamic forces generated by movement through water. Hydrophobic polymers are the most commonly used fouling-release moieties, while the chemical and physical effects of the poorly biocompatible compounds on the environment and human health have become a major concern. Meanwhile, proteins possess different adsorption mechanisms due to their inherently amphiphilic nature,17 which results in various affinities to hydrophobic and hydrophilic surfaces.18 Therefore, homogeneous surfaces (solely hydrophilic or hydrophobic) with single type functionality are often inadequate for repelling proteins upon permanent exposure to complex environments.19 Thus far, much work has focused on amphiphilic polymer materials. These polymers, which possess an environmental-response characteristic such that they can restructure their surfaces in water, have been demonstrated to improve protein resistance.20 In contrast to homogeneous surfaces, amphiphilic surfaces with both hydrophilic and hydrophobic components are considered to permanently maintain their protein resistance21 and prevent proteins from adopting conformations allowing them to adsorb on the surface.22 Similarly, amphiphilic surfaces have greatly improved biocompatibility,23 which can provide the functional groups and morphological changes for surface modification and incorporation of other functional materials. At the same time, amphiphilic polymer surfaces are frequently complex, and microphase separation usually occurs due to the incompatibility of the hydrophilic and hydrophobic components. It is thus of critical importance to know the 4
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effect of the surface micromorphology of these materials on the adsorption of proteins. The phase structure of materials that imitated the native glycocalyx was first investigated using amphiphilic comb poly(vinyl amine) with oligosaccharide side chains.24 It was observed that both the water contact angle and protein adhesion were significantly lower than those of the parent surface, since the microphase separation on the surface will occur in aqueous conditions due to the swelling of the side chains and the extension of the chain segments. Another interesting finding was that polymers with different chemical structures, from linear oligosaccharides to dendritic polysaccharides, showed similar biological antifouling properties because they self-assemble into well-ordered structures with 10 nm domains.25 Ober et al.26 reported that surface active polystyrene-b-poly(ethoxylated fluoroalkyl acrylate) with an approximately 35 nm domain exhibited a great enhancement in resistance to bovine serum albumin (BSA) adsorption. Shen et al.27,28 fabricated amphiphilic coatings by self-assembly of poly(styrene)-b-poly(2-hydroxyethyl methacrylate) (PS-b-PHEMA) and found that the PS200-b-PHEMA50 surface with PS cylinders (diameter ~ 20 nm) in a PHEMA matrix was strongly protein-repulsive.27,28 Until now, many studies have supported the hypothesis that amphiphilic surfaces with optimal nanoscale heterogeneity present energetically unfavorable substrates for protein adsorption, but the mechanism on phase-separated amphiphilic surfaces is relatively poorly understood, thus giving rise to certain questions. For example, what are the domain chemical components and the critical domain sizes of heterogeneous 5
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nanopatterns for different proteins on the surface that exhibit the best protein resistance? Herein, we developed a novel method to prepare phase-separated heterogeneous surfaces by preparing amphiphilic V-shaped polymer brushes with a water-soluble poly(ethylene glycol) arm and a hydrophobic fluorinated poly(methyl methacrylate) or polystyrene arm. The phase-separated structure and surface composition can be adjusted by changing the length of the V-shaped brush arm and the chemical component of the hydrophobic arm. The effects of the microphase structures on the corresponding protein adsorption were investigated. Our results revealed that microphase structures on the surface with a length scale twice that of protein molecules exhibited excellent protein-resistance capability. At the same time, the amount of protein adsorption was highly related to protein adhesion and the relative friction coefficient on the corresponding brush surfaces. Furthermore, the water-soluble PEG domain on these polymer brushes stretched out at the interface and the low-surface-free-energy domain with suitable size remained relatively stable in an aqueous environment. Thus, a heterogeneous surface with phase-separated domains twice the size of protein molecules may minimize the adsorption of proteins through a synergistic effect of the hydrophobic and water-soluble domains.
2. EXPERIMENTAL SECTION 2.1. Materials. Styrene (S, 99%), 2-perfluorooctylethyl methacrylate (FMA, 97%), methyl 6
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methacrylate (MMA, 99%), tert-butyl acrylate (tBA, 99%), 2-bromopropionyl bromide (BPB, 97%) and poly(ethylene glycol) monomethyl ethers (mPEG110, Mn = 5000 g/mol; mPEG42 Mn = 1900 g/mol) were purchased from Aldrich Co. (USA). Triethylamine (TEA, 99%) and 4-(dimethylamino) pyridine (DMAP, 99%) were purchased from Alfa Aesar Co. (UK). All of the above chemicals were purified according to methods described elsewhere.29,30 CuBr (99%) was purified according to a
reported
procedure.31
12-mercaptododecanoic
γ-Glycidoxypropyl-trimethoxysilane acid
(HS(CH2)11COOH,
96%),
(GPS,
98%),
N,N’,N’,N”,N”-
pentamethyldiethylenetriamine (PMDETA, 99%) and methylene dichloride (CH2Cl2, 99.8%) were purchased from Aldrich Co. (USA) and used as received. Bovine serum albumin (BSA, CP), human fibrinogen (HFg, CP), fluorescein isothiocyanate-labeled bovine serum albumin (FITC-BSA), and fluorescein isothiocyanate-labeled human fibrinogen (FITC-HFg) were purchased from Wanhuashi Biotech Co. (Beijing, China). The other chemicals were purchased from Shanghai Reagent Co. (China) and used as received.
2.2. Synthesis of Amphiphilic Block Copolymers by Stepwise ATRP. The amphiphilic block copolymers mPEG110-b-PAA7-b-PMMAn-b-PFMA5 were synthesized by stepwise atom transfer radical polymerization (ATRP) according to the previously reported method,29 and their structures were characterized by gel permeation chromatography (GPC), 1H NMR and fluorine elemental analysis. First, the mPEG110-Br macroinitiator was prepared by reacting the –OH at one end of 7
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mPEG110 chain with BPB.29,30 The mPEG (2.0 mmol) and DMAP (8.2 mmol) were mixed with TEA (14 mmol) in 100 mL of CH2Cl2. When the solution was cooled to 0 °C, 3 mL of BPB (24.3 mmol) in 37 mL of CH2Cl2 was added dropwise into the flask over 1 h under nitrogen. After stirring at room temperature for 24 h, the mPEG110-Br macroinitiator was obtained. The mPEG110-b-PtBA7-Br macroinitiator was synthesized according to the following procedure. A [tBA]/[PMDETA]/ [CuBr]/[mPEG110-Br] molar ratio of 10:2:1:1 was prepared in toluene under a N2 atmosphere. The syntheses of the mPEG110-b-PtBA7-b-PMMAn-Br macroinitiator and amphiphilic copolymer mPEG110-b-PtBA7-b-PMMAn-b-PFMA5 were similar to that of mPEG110-b-PtBA7-Br. Finally, the PtBA segment in mPEG110-b-PtBA7-b-PMMAn -b-PFMA5 was selectively hydrolyzed to form mPEG110-b-PAA7-b- PMMAnb-PFMA5 as per a previously described method.29,32 Both mPEG110-b-PAA9-b-PSn and mPEG42-b-PAA6-b-PSn block copolymers were prepared using a procedure similar to that used for the synthesis of mPEG110-b-PAA7-b-PMMAn-b-PFMA5.
2.3. Preparation of Amphiphilic V-shaped Polymer Brushes. Amphiphilic V-shaped polymer brushes with two highly incompatible arms were prepared by reacting -COOH groups in mPEG110-b-PAA7-b-PMMAn-b-PFMA5 and mPEG-b-PAA-b-PS with epoxy groups on a SiO2 surface from the melt according to a previously reported method,29,33 in which GPS was used to obtain epoxy group-functionalized SiO2 substrates. A thin polymer film was spin-coated on a 8
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modified SiO2 substrate using 2% polymer chloroform solution, and then heated for 24 h at 150 °C under vacuum. The thin film was washed with chloroform by Soxhlet extraction for 12 h to remove the nongrafted polymer. The resulting brush was dried in a vacuum oven for 12 h at 110 °C. A pure PEG brush was prepared for purposes of comparison using mPEG110-b-PAA7. Since the midblock PAA in the block copolymer was an anchor and was reacted with epoxy group on the substrate to form V-shaped polymer brushes with two arms. Therefore, when the block copolymer was used to prepare V-shaped brush, ‘PAA’ in the block copolymer was replaced with ‘V’ to indicate corresponding V-shaped polymer brushes.
2.4. Characterization. Characterization of Amphiphilic Block Copolymers. Gel permeation chromatography (GPC) was performed using a Waters 1515 apparatus equipped with a sequence of Waters Styragel HR1 and HR4 columns and a Waters 2414 Refractive Index Detector. THF was used as mobile phase at a flow rate of 1 mL/min at 35 °C and standards were monodispersed polystyrene. 1H NMR spectra were recorded using a Bruker Avance AMX-400 NMR spectrometer with CDC13 as the solvent. The FMA content in the copolymers was determined by fluorine elemental analysis.34 Characterization of the Polymer Brushes. The contact angles of 3 µL liquid drops of water or diiodomethane on the brushes were measured by the Sessile drop method with a Krüss DSA10-MK2 contact angle goniometer (Hamburg, Germany). The surface free energy of the V-shaped amphiphilic brushes was estimated using 9
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contact angles of water and diiodomethane according to the theory of Owens and Wendt.35 X-ray photoelectron spectroscopy (XPS) was performed to detect the surface composition of the various brushes using a PHI-5000C ESCA System with an Mg Kα X-ray source (1253.6 eV) at a detection angle of 30°. For each sample, four different sites or locations on the brushes were measured with a spot size of 100 µm diameter. Each measurement included a survey consisting of three sweeps and a high-resolution measurement for N1s with eight sweeps. The errors for quantitative determination of the elemental surface composition were less than ±10%. The brush thicknesses were evaluated in air by EP3SW imaging ellipsometry (Accurion GmbH Co., Germany) at a fixed wavelength (λ = 658 nm) with an incidence angle of 60°. In order to make sure that the results were reliable, an AOI (Angle of incidence) spectral measurement (from 70° to 45°, in 5° steps) was also performed and it was found that the result was the same as that obtained with a single angle within the error of 0.2 nm. Alignment is important for every ellipsometry measurement. The EP3SW imaging ellipsometer has an alignment laser and related mechanics to ensure that every measurement of the sample is perfectly aligned. All measurement conditions were carefully controlled to reduce measurement errors. The reported thickness for each sample is the average of eight measurements at different positions. The phase images and height images of the V-shaped polymer brushes were obtained in air and in PBS solution with a MultiMode-8 atomic force microscope (Bruker Co., USA) in the tapping modulated mode using rectangular silicon cantilevers with a resonant frequency of 300 kHz and a spring constant of about 40 N/m. The phase images and height images of the 10
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samples in PBS solution were obtained by applying the tapping mode in liquid form with silicon cantilever probes mounted in a fluid cell. The average domain size of the brushes was obtained from a power spectrum, which was performed by the 2D fast Fourier transform of the AFM images and the circular average of the resultant spinodal rings using Scion image analysis software (Scion Corporation, USA). Protein Adsorption on the Brushes. The protein adsorption experiments were carried out according to reported methods36 using HFg (length = about 46 nm), BSA (length = about 14 nm), FITC-HFg and FITC-BSA as model proteins. Briefly, the different brushes were immersed in PBS solution (pH = 7.4) containing protein (0.1 mg mL-1) at room temperature for 60 min. The brushes were gently washed with PBS solution in a shaking bath at room temperature with 100 rpm for 5 min, followed by washing three times with deionized water, and then dried with nitrogen. Protein adsorption measurements were performed qualitatively by fluorescence microscopy (Olympus, Japan) using FITC-HFg and FITC-BSA, and quantitatively by XPS using HFg and BSA. FITC was observed with a 480 nm excitation and 525 nm emission filter set. The protein adsorption was calculated by integration of the peak area of the N1s peak from the corresponding XPS spectrum. Measurement of Adhesion and the Relative Friction Coefficient between the Protein and the Brushes. The pyramid-shaped silicon AFM tips (Bruker Co., USA) with a cantilever nominal spring constant of 0.5 N/m and a tip radius of 10 nm were surface-modified by protein according to a reported method.36 The cantilevers were coated with approximately 30 nm of gold. The gold-coated cantilevers were 11
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surface-modified by 1 mM HS(CH2)11COOH solution at 37 °C in dark conditions for 12 h and then rinsed thoroughly with absolute ethanol. The resulting modified cantilevers were added to a 20 mL vial containing a solution of 9.58 mL DMF, 0.28 mL TEA and 0.14 mL trifluoroacetic anhydride and were incubated at 25 °C for 20 min. After the incubation period, the probe tips were treated with 0.1 mg/mL protein in PBS solution at 25 °C for 30 min. The cantilevers were then rinsed with water and dried under nitrogen. The AFM tip surface immobilized with protein molecules was characterized with an Ultra55 Scanning Electron Microscope and Energy Dispersive Spectrometer (Carl Zeiss SMT, Germany). The results indicated that the probe did not change noticeably before and after AFM measurements, as shown in Figure S1 (Supporting Information).
The adhesion and the relative friction coefficient measurements were carried out using the PeakForce QNM mode and the Lateral Force mode in PBS solution in a fluid cell at room temperature, respectively. The pyramidally shaped AFM tips functionalized with BSA or HFg proteins were used to measure the adhesion and relative friction coefficient. For the measurement of adhesion, the vertical sensitivity was calibrated on a sapphire sample and the normal spring constant (0.68 N/m) was obtained by the thermal tune method. A single ramp was triggered with a maximum load of 20 nN to obtain a force-distance curve between the functionalized tip and the brush surface. The adhesion forces can be easily calculated from the curve, as the pull-off force. At least 100 curves were measured for each sample and the average adhesion value was obtained from them. 12
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The relative friction coefficient was measured according to a reported method.37,38 In this measurement method, a relatively high load of 2 V was applied to obtain lateral force maps in the trace and retrace scan directions, as separate images (Figure S2, Supporting Information). The induced voltage reflecting the true friction was calculated as half of the difference value between the two images.39 The relative friction coefficient was then roughly calculated as the specific value of the friction induced voltage and load voltage according to the AFM instruction manual (Figure S2, Supporting Information). Such a relative friction coefficient could qualitatively contrast the friction properties of different brush surfaces by using the same AFM probe in such a consistent manner.
3. RESULTS AND DISCUSSION 3.1. Structure of Amphiphilic V-shaped Polymer Brushes. The amphiphilic block copolymers mPEG110-b-PAA9-b-PSn, mPEG42-b-PAA6b-PSn and mPEG110-b-PAA7-b-PMMAn-b-PFMA5 were synthesized successfully as shown in Table 1. The V-shaped amphiphilic brushes were prepared via a “grafting to” approach from the melt by the epoxy groups on the epoxy group-functionalized SiO2 substrate reacting with –COOH groups in the block copolymers above. The structural characteristics of the V-shaped mPEG110-V-PMMAn-b-PFMA5, mPEG110-V -PSn and mPEG42-V-PSn brushes are listed in Table 1. The roughness of all the brushes was extremely low; in other words, all the brush surfaces were smooth, which indicates that the grafting density of the brushes on the substrates was homogeneous. 13
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Table 1. Characteristics of Amphiphilic Block Copolymers and the Resulting V-shaped Brushes Sample
Mna)
PDIb)
Thickness(nm)c)
mPEG110-b-PAA7-b-PMMA30-b-PFMA4
10600
1.20
8.2±0.4
mPEG110-b-PAA7-b-PMMA43-b-PFMA5
12500
1.25
8.9±0.6
mPEG110-b-PAA7-b-PMMA64-b-PFMA5
14600
1.19
10.2±0.5
σ(chain/nm2)d) γs(mJ/m2) e) Domain size(nm)f)d) 0.76
40.5
625
0.71
35.8
312
0.66
31.3
244
mPEG110-b-PAA7-b-PMMA82-b-PFMA5
16400
1.24
11.1±0.8
0.61
28.1
27
mPEG110-b-PAA7-b-PMMA93-b-PFMA4
16900
1.23
11.7±0.6
0.59
24.2
169
mPEG110-b-PAA7-b-PMMA109-b-PFMA5
19100
1.22
12.3±0.9
0.56
21.6
81
mPEG110-b-PAA7-b-PMMA128-b-PFMA5
21000
1.17
13.5±0.8
0.52
17.6
345
mPEG42-b-PAA6-b-PS43
6900
1.18
6.9±0.5
0.70
51.4
/
mPEG42-b-PAA6-b-PS78
10600
1.17
8.2±0.6
0.54
43.6
249
mPEG42-b-PAA6-b-PS128
15800
1.12
9.4±0.5
0.42
35.0
31
g)
mPEG110-b-PAA6-b-PS61
11800
1.13
8.7±0.4
0.51
58.8
134
mPEG110-b-PAA9-b-PS119
18000
1.15
12.6±0.6
0.49
46.1
76
mPEG110-b-PAA9-b-PS169
23200
1.18
14.7±0.7
0.44
38.2
111
mPEG110-b-PAA9-b-PS227
29300
1.17
16.6±0.8
0.40
34.5
152
a) Obtained from 1H NMR and elemental analysis; b) obtained by GPC; c) corresponding V-shaped brush thickness measured by ellipsometry; d) the densities and e) surface free energies of corresponding V-shaped brushes; f) the average domain size of V-shaped brushes obtained by Fourier transform of AFM images; g) no phase separation.
The results show that the thicknesses of the mPEG110-V-PMMAn-b-PFMA5 brushes increased from 8.2 nm to 13.5 nm with increasing degree of PMMA polymerization (DP, n) from 30 to 128. Similar to mPEG110-V-PMMAn-b-PFMA5 brushes, the thicknesses of the mPEG110-V-PSn and mPEG42-V-PSn brushes increased with increasing length of the PS arm, i.e., the degree of PS polymerization. However, almost no residual polymer was observed when mPEG-b-PtBA6-b-PSn and mPEG110-b-PtBA7-b-PMMAn-b-PFMA5 (no –COOH group in copolymer) were used. These results indicate that the grafting reaction was successful. The grafting density 14
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(σ) of the V-shaped brushes could be estimated from their thickness according to equation (1):40 ܣܯܯܲܯ
σ = dܰ ܣ/ ቀ
ߩ ܲܣܯܯ
+
ܩܧ ܲܯ ߩ ܲ ܩܧ
ቁ
(1)
where d is the thickness of the brush, and NA is Avogadro’s constant. MPMMA and MPEG are the molecular weights of the PMMA (or PS) and PEG respectively, and ρPMMA (or ρPS) and ρPEG are the densities of PMMA (or PS) and PEG, respectively. The grafting density of all brushes was approximately 0.4~0.7 chains/nm2, which was similar to the reported value of the V-shaped brushes PS-V-PEO.40 Table 1 indicates that the surface free energy of the mPEG110-V-PMMAn -b-PFMA5 brushes sharply decreases from 40.5 to 17.6 mJ/m2 with increasing degree of PMMA polymerization from 30 to 128 with PFMA5 at the chain end, which is consistent with the variation in fluorine content on the brush surfaces. Figure 1 shows that both F/C and CF3/CF2 ratios obviously increase from 0.11 to 0.45 and from 0.11
Figure 1. F/C and CF3/CF2 ratios of mPEG110-V-PMMAn-b-PFMA5 brushes.
15
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to 0.28, respectively, in which an increase in CF3/CF2 ratio indicates that a well-oriented structure of the fluorinated groups was formed, resulting in low surface free energy.41 The amphiphilic block copolymer surface was highly complicated due to the chemical incompatibility of each block, which usually resulted in obvious microphase separation.42 The AFM phase images in Figure 2 presented the chemical information of the brush surface, in which the bright domains corresponding to crystalline phases of the PFMA component,29,36 increased gradually with increasing length of the PMMAn-b-PFMA5 arm. The AFM height images indicated that all the sample surfaces were smooth with a roughness of about 0.3~1.3 nm (shown in Figure S3, Supporting Information). The V-shaped polymer brushes with n = 82 and
Figure 2. AFM phase images (2.0 µm × 2.0 µm) of mPEG110-V-PMMAn-b-PFMA5 brushes. (A - F) n = 30, 64, 82, 93, 109 and 128, respectively.
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109 exhibited an alternating phase-separation structure with a domain size of 25~30 nm and 75~80 nm, respectively, as shown in Figure 2C, E. Since it was difficult to measure the domain size when the phase structure on the V-shaped brush surface was not well-ordered, the average domain size could be obtained via Fourier transform of the AFM images43,44 as shown in Figure S4 (Supporting Information), in which 1/qm corresponds to the average domain size as shown in Table 1. The domain sizes of
mPEG110-V-PMMA82-b-PFMA5
and
mPEG110-V-PMMA109-b-PFMA5
are
obviously identical to those shown in Figure 2C, E, respectively. At the same time, it was observed that the domain size showed an approximate bowl-shaped relationship with the arm length of PMMAn-b-PFMA5. When the length of the PMMAn-b-PFMA5 arm was much shorter than that of the PEG arm, the surface of the brushes was dominated by PEG, and the PEG domain decreased with increasing length of the PMMAn-b-PFMA5 arm. When the length of the PMMAn-b-PFMA5 arm was longer
Figure 3. Structural representation of the mPEG110-V-PMMAn-b-PFMA5 V-shaped polymer brushes with increasing DP of PMMA (n). 17
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than that of the PEG arm, the brush surface was dominated by the PFMA component, resulting in an increase in the bright domain, as shown in Figure 3. These findings are identical to the results obtained from the XPS and the surface free energy measurements.
3.2. Protein Adsorption on V-shaped Brush Surfaces. The amount of protein adsorbed on the surface of the materials was determined with XPS, which is a simple and effective method for analyzing materials that do not contain elemental nitrogen.45 Note that none of the V-shaped polymer brushes used in this study contain nitrogen atoms. Therefore, the N1s peak from the XPS spectra, the source of which could only be the protein, was used to detect the protein adsorption. The relative atomic N1s% on various V-shaped brush surfaces after protein adsorption is shown in Figure 4A. It was found that all of the V-shaped polymer brushes
Figure 4. (A) Atomic % N1s of adsorbed protein versus degree of PMMA polymerization (n) in mPEG110-V-PMMAn-b-PFMA5 brushes. (B) Atomic % N1s of adsorbed proteins on mPEGm-V-PSn brushes. The dashed lines are visual guides.
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exhibited a better protein-resistance performance than that of a pure mPEG110 brush surface; the amount of protein adsorbed on the pure mPEG110 brush surface was similar to the reported value46. The adsorbed protein amount noticeably decreased with increasing DP (n) of PMMA, and the lowest protein adsorption on the V-shaped brushes was observed for BSA at n = 82 and for human fibrinogen (HFg) at n = 109. To intuitively visualize the amount of protein adsorbed on the brush surfaces, we used fluorescein isothiocyanate-labeled proteins (FITC-protein) as test proteins and examined the degree of protein adsorption using fluorescence microscopy;47 the
A
B
C
D
E
F
G
H
J
Figure 5. Fluorescence microscopy images of mPEG110-V-PMMAn-b-PFMA5 brushes after FITC-BSA adsorption. (A-G) n = 30, 43, 64, 82, 93, 109 and 128, respectively. (H) Pure mPEG110 brush. (J) Glass plate. Scale bar: 100 µm. Highlighted represented the optimal BSA repellence. 19
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results are presented in Figure 5 and Figure 6. Since both the PEG and fluorinated PMMA brushes showed good protein-resistance performance, no significant differences were observed between fluorescent micrographs in Figures 5 and 6. However, the difference in fluorescent micrographs is clearly evident. The variation in fluorescence intensity on all sample surfaces clearly presented the same trends of N1s% on the brush surfaces obtained with XPS. The above results indicate that the mPEG110-V-PMMAn-b-PFMA5 V-shaped brushes presented optimal BSA and HFg repellence (highlighted in Figures 5 and 6) when the surfaces had an obvious
Figure 6. Fluorescence microscopy images of mPEG110-V-PMMAn-b-PFMA5 brushes after FITC-HFg adsorption. (A - G) n = 30, 43, 64, 82, 93, 109 and 128, respectively. (H) Pure mPEG110 brush. Scale bar: 100 µm. Highlighted represented the optimal HFg repellence. 20
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alternating phase-separation structure with domain sizes of approximately 25~30 nm and 75~80 nm, which are about twice the length of BSA (14 nm × 4 nm × 4 nm)48 and HFg (45 nm × 5 nm × 5 nm)49, respectively. Figure 7 indicate atomic % N1s of adsorbed HFg protein versus domain sizes in mPEG110-V-PMMAn-b-PFMA5 brushes. It was obvious that HFg adsorption was the lowest when the domain sizes were approximately 75~80 nm, while for BSA the domain sizes were approximately 25~30 nm (Figure S5, Supporting Information). Baier and Schrader50,51 suggested that when the surface free energies of materials are in the range of 20~30 mJ/m2 the minimum protein adhesion would be presented instead of the lowest surface free energy. Since the surface free energies of the mPEG110-V-PMMA82-b-PFMA5 and mPEG110-V-
0.18
1.2
4.0 3.5
0.16 1.0 0.14 0.12
0.8
0.10 0.6
0.08
3.0 2.5 2.0
At.N%
Relative Friction Coefficient
PMMA109-b-PFMA5 V-shaped brushes were 28.1 and 21.6 mJ/m2, respectively, both
Adhesion (nN)
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60
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1.5 1.0
HFg
0.06 10
0.5 0.4 1000
100
Domain size (nm) Figure 7. Atomic % N1s of adsorbed protein, average measured adhesion force and the relative friction coefficient of HFg versus domain sizes in mPEG110-V-PMMAn-b-PFMA5 brushes. The dashed lines are visual guides. 21
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lying within Schrader’s suggested range, interfacial wetting may be another reason that these heterogeneous surfaces possess excellent protein-resistance capability. To further confirm the role of domain size on the effect of protein adsorption on the heterogeneous surfaces, V-shaped brushes with a PS arm and a mPEG arm were prepared by employing two types of block copolymers, namely, mPEG42-b-PAA6 -b-PSn and mPEG110-b-PAA9-b-PSn. Figure 4B indicates that the mPEG42-V-PS128 brushes exhibited the lowest BSA adsorption and mPEG110-V-PS119 exhibited the lowest HFg adsorption, which was identical to the fluorescence intensity in their fluorescence microscopy images after FITC-HFg and FITC-BSA adsorption, as shown in Figure S6 and Figure S7 (Supporting Information). The domain size and
Figure 8. AFM phase images (2.0 µm × 2.0 µm) of V-shaped brushes with two arms of mPEG and PSn. (A) mPEG42-V-PS43, (B) mPEG42-V-PS78, (C) mPEG42-V-PS128, (D) mPEG110-V-PS61, (E) mPEG110-V-PS119, and (F) mPEG110-V-PS169. 22
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phase images of both mPEG42-V-PSn and mPEG110-V-PSn brushes are presented in Table 1 and Figure 8. The domain size of the phase-separated structure was found to be
approximately
25~30
nm
for
mPEG42-V-PS128
and
75~80
nm
for
mPEG110-V-PS119, which presented the lowest adsorption of BSA and HFg, respectively. It could also be observed in the relationship between atomic % N1s of adsorbed protein and domain sizes in mPEG-V-PS brushes shown in Figure S8 (Supporting Information). Since the surface free energies of the mPEG-V-PSn brushes were in the range of 34.5~58.8 mJ/m2 as shown in Table 1, which is out of the range of 20~30 mJ/m2, the above results confirm that the domain size of the phase-separated amphiphilic brushes was the main reason why they exhibited the best protein-resistance performance, rather than their surface free energies.
3.3. The Origin of Domain Size Dependent Protein Resistance on Surface with Heterogeneous Nanopatterns. Recently, some research groups have demonstrated that the microphase separation structure of amphiphilic polymer surfaces affects the adsorption behavior of proteins to a large extent.26-29 Ober et al.26 found that amphiphilic block copolymer film with about 35 nm domain size presented the lowest BSA adhesion, which was identical to the results in this work. Bhushan reported52 that due to the difference in morphologies and domain spacing, diblock PMMA-b-PAA and triblock PAA-bPMMA-b-PAA copolymers with similar compositions exhibited different protein adhesion. The adhesion of proteins on the surface in the PBS buffer solution was 23
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employed to evaluate the possibility of protein adsorption by AFM using a probe tip immobilized with protein.53 Recently, many studies have focused on the adhesion and friction properties of polymer brushes54-57 and have considered that they are primarily an effect of the surface coverage, chain end mobility and overlap between chains. In order to provide mechanistic insight into the results above, the adhesion and friction
Relative Friction Coefficient
1.2
Adhesion(nN)
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60
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1.0
0.8 0.6
0.4 0
30
60
90
120
150
0.18 0.14 0.10 0.06 0
DPPMMA(n)
30
60
90
120
DPPMMA(n)
Figure 9. Average measured adhesion force and the relative friction coefficient for a protein-coated AFM tip on the mPEG110-V-PMMAn-b-PFMA5 V-shaped brush surfaces. BSA, red; HFg, blue. The dashed lines are visual guides.
properties of polymer brushes with the proteins were investigated. Figure 9 summarizes the corresponding adhesion and relative friction coefficient values between proteins and the V-shaped mPEG110-V-PMMAn-b-PFMA5 brushes. The observed variation in the trend of the adhesion and relative friction coefficient on mPEG110-V-PMMAn-b-PFMA5 brush surfaces with the length of the PMMAnb-PFMA5 arm is consistent with the trend of the amount of protein adsorbed on the 24
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corresponding brushes. The lowest adhesion and relative friction coefficient for BSA were observed on the mPEG110-V-PMMA82-b-PFMA5 V-shaped brush surface and for HFg on the mPEG110-V-PMMA109-b-PFMA5 brush surface, in which corresponding domain sizes were approximately 25~30 nm and 75~80 nm, respectively (shown in Figure 7, Figure S5). The same result was obtained for the mPEG-V-PSn brushes shown in Figure 10 and Figure S8 (Supporting Information), in which the lowest
Figure 10. Averaged measured adhesion force and the relative friction coefficient for a protein-coated AFM tip on the mPEG-V-PSn V-shaped brush surfaces. BSA, red; HFg, blue. Highlight represented the lowest adhesion and relative friction coefficient.
protein adsorption amount corresponded to the lowest protein-brush interaction in an aqueous environment. The results above confirm that weak protein-brush interaction resulted in the brushes exhibiting an enhanced protein-resistant performance. This finding is identical to the reported result26 that the amount of BSA adsorbed on an amphiphilic polymer (PS-b-PAA-AMP) surface was substantially lower than that of a thermoplastic elastomer (SEBS) and a hydrophilic silicon wafer because protein 25
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adhesion was reduced by 50-fold and 8-fold, respectively. Yang et.al57 reported that the super low friction surface of zwitterionic poly(3-(1-(4-vinylbenzyl)-H-imidazol3-ium-3-yl)propane-1-sulfonate) exhibited a super low fouling surface. Several excellent reviews explain that the adsorption of proteins on the surface proceeds through two processes.58-60 First, the initial contact between proteins and the surface forms a moving precursor state, where the proteins are weakly adsorbed on the surface. This precursor molecule then diffuses along the material surface while the native 3D protein conformation reorganizes into the final adsorbed state and tightly binds to the surface. It is clear that the second step plays an important role in the adsorption of proteins, in which the protein molecules will maximize their interaction with the surface. However, the protein movement along the surface is closely related to the material surface chemistry.59 The protein molecules “roll” along the hydrophilic surface, while the movement of proteins on a hydrophobic surface is a “sliding” mode to reach a favorable interaction through reorientation. Thus, when the length scale of the heterogeneous nanopatterns on the amphiphilic surface is similar to that of the protein, the protein interaction with the surface would be disrupted because the material surface could not provide sufficient area for the precursor protein to form a stable adsorbed state, resulting in repelling of proteins. According to the above mechanism, it is presumed that protein adhesion on the surface corresponds to the first stage and the protein friction coefficient corresponds to the second stage. The mPEG110-V-PMMAn-b-PFMA5 and mPEG-V-PSn V-shaped brushes had different hydrophobic domains in which the surface free energy of PFMA on the 26
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mPEG110-V-PMMAn-b-PFMA5 surface was much lower than that of PS on the mPEG-V-PS
surface.
Therefore,
protein
adhesion
on
the
mPEG110-V-
PMMAn-b-PFMA5 brushes was much lower than that on the mPEG-V-PS brushes. The relative friction coefficient of the proteins was measured by moving the protein molecules on the brush surface. This process likely involves molecular reorganization of the native 3D protein conformation. Thus, the relative friction coefficient of proteins may be more favorable for studying the effect of heterogeneous nanopatterns on the adsorption behavior of proteins. It is well known that the reconstruction of the heterogeneous surface will take place during immersion of the V-shaped brushes in an aqueous solution. Under an aqueous environment, the hydrophobic PS or PFMA domains on V-shaped brushes with low surface free energy remain unchanged due to their high Tg and water-insoluble properties. However, the chains of the water-soluble PEG domain will stretch, and the height, which is related to the hydrophobic domain in water, changes with the relative length of the two arms, which also results in a change in the domain size on the phase-separated surfaces. This was confirmed by AFM phase images and height images of the brushes in PBS solutions (shown in Figure S9 and Figure S10, Supporting Information), in which the phase images were almost similar and height images became smoother compared with that in the air. At the same time, the extended length of the hydrophilic PEG chains in an aqueous environment, which determine the height of the PEG chain over the hydrophobic domain, may be crucial in determining the protein adsorption. When the domain size of the phase-separated 27
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structures on the heterogeneous surface is two times larger than the size of the protein molecules, the PEG chain will stretch over the PS or PFMA domain in a liquid-like state in water with rapid motion and prevent adsorption or adhesion of protein molecules. Moreover, the hydrophobic domain size was not in favor of the proteins being able to adopt a conformation to adsorb to the surface. This synergistic effect resulted in excellent protein resistance of the V-shaped polymer brushes and the lowest adhesion and relative friction coefficient between the protein and the brush surface.
4. CONCLUSIONS Since proteins possess different adsorption mechanisms due to their inherently amphiphilic nature,17 homogeneous surfaces (solely hydrophilic or hydrophobic) with single type functionality are often inadequate for repelling proteins upon permanent exposure to complex environments.19 Until now, many studies have supported the hypothesis that amphiphilic surfaces with optimal nanoscale heterogeneity present energetically unfavorable substrates for protein adsorption.19-21 Ober et al.26 reported that amphiphilic block copolymer films with about 35 nm domain size presented the lowest BSA adhesion. However, it is unclear what critical domain sizes of heterogeneous nanopatterns surfaces for different proteins exhibit the best protein resistance. In this paper, amphiphilic V-shaped polymer brushes with a PMMAnb-PFMA5 or PSn arm and a mPEG arm were prepared, in which their surface structure and morphology were controlled by altering the relative length of the two arms. The 28
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results showed that the V-shaped brushes exhibited the lowest protein adsorption only when the size of their phase-separated domains was about twice the length of the protein molecules. This important relationship was first observed herein. At the same time, the amount of protein adsorption was well related to the adhesion and relative friction coefficient of the corresponding proteins on the brush surfaces, in which the lowest amount of protein adsorption was observed when the heterogeneous surface has the lowest adhesion and relative friction coefficient. The reason for these observations may be attributed to the adhesion and friction coefficient of the proteins corresponding to different stages in the protein adsorption process, namely, protein-surface contact and subsequent diffusion, respectively. Thus, the friction coefficient of the proteins may be more favorable for studying the influence of phase structure on protein adsorption behavior. Meanwhile, the extended length of the hydrophilic PEG chains in an aqueous environment, which determined the height of the PEG chains over the hydrophobic domain, may be crucial in determining the protein adsorption. When the PEG chains extended and stretched out at their interface and the hydrophobic domain remained unchanged to prevent disruption of its phase structure under an aqueous environment, the desirable domain size favored preventing the protein molecules that make initial contact from forming a moving precursor state and reorganizing into a thermodynamically stable adsorbed state. This phase -separated structure resulted in maximum enhancement of the synergistic effect between the hydrophilic and hydrophobic domains of the brush surfaces and disrupted the protein-surface interaction when the size of the phase-separated domains was 29
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double that of the protein. These observations provide a way to design and prepare phase-separated amphiphilic surfaces with excellent protein-resistant performance in the future.
ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jpcc.xxxxxxx. SEM and EDS of AFM functionalized tips; the representative lateral force trace and retrace maps of mPEG110-V-PMMA82-b-PFMA5 V-shaped brush; calculation method
of
relative
friction
coefficient;
AFM
height
images
of
mPEG110-V-PMMAn-b-PFMA5 brushes in air and in PBS solution; power spectrum of mPEG110-V-PMMAn-PFMA5 brushes; fluorescence microscopy images of mPEG-V-PSn brushes after FITC-BSA and FITC-HFg adsorption; Atomic % N1s of adsorbed protein, average measured adhesion force and the relative friction coefficient
versus
domain
sizes
in
mPEG110-V-PMMAn-b-PFMA5
mPEG-V-PSn brushes. Figures S1-S10 (PDF)
AUTHOR INFORMATION Corresponding Authors *E-mail:
[email protected] or
[email protected]
30
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and
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*E-mail:
[email protected]
ORCID Xinping Wang: 0000-0002-9269-3275 Xianjing Zhou: 0000-0001-7703-7555
Notes The authors declare no competing financial interest.
ACKNOWLEDGMENTS This work was funded by the National Natural Science Foundation of China (NSFC, Nos. 21504081 and 21704092) and the Natural Science Foundation of Zhejiang Province (No. LQ16B040001).
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