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The Effect of Fluid Flow on Selective Protein Adsorption on Polystyrene-block-Poly(methyl methacrylate) Copolymers Qin Li,*,†,‡ K. H. Aaron Lau,‡ Eva-Kathrin Sinner,‡ Dong Ha Kim,§ and Wolfgang Knoll*,‡, †
Department of Chemical Engineering, Curtin University of Technology, Perth, WA 6845, Australia, Max Planck Institute for Polymer Research, Ackermannweg 10, 55128 Mainz, Germany, §Department of Chemistry and Nano Science, Ewha Womans University, Seoul 120-750, Republic of Korea, and Austrian Institute of Technology, Donau-City-Strasse 1, 1220 Vienna, Austria )
‡
Received May 10, 2009. Revised Manuscript Received July 29, 2009 The effect of fluid flow on protein patterning and the stability of the adsorbed protein array templated by polystyreneblock-poly(methyl methacrylate) (PS-b-PMMA) has been investigated. Protein nanoarrays can be formed on a PS-bPMMA copolymer surface by physisorption and rinsing with an open flow. The protein arrays inherit the original hexagonal pattern generated by the phase separation of PS-b-PMMA with the proteins only residing on the PS domains. Subsequent analysis of the protein array stability under confined flow field reveals that the array integrity is strongly dependent on the flow velocity and the duration time. The interplay between the intermolecular forces and the hydrodynamic shear force has been discussed in detail. A simplified model has been proposed to explain the site-selective adsorption and protein migration under the shear of fluid flow. This study provides valuable information on the formation mechanism and stability of physisorbed protein nanopatterns under conditions concerning biosensing applications. It also represents an explorative study of protein adsorption under hydrodynamic flow conditions, which may assist in better designs of fluidic devices relevant to protein studies.
Introduction The adsorption of proteins from solution onto solids surfaces is of vital importance to many areas in biotechnology and biomedical applications, such as biofouling, immunosensors, proteomics, microreactor, and tissue engineering. One of the particular interests in site-selective protein immobilization is greatly driven by the development of high-density immunosensor arrays, which is highly desirable for investigations in cellular functions, diseases, and therapeutic methods.1-4 Controlled immobilization of proteins in terms of site-selectivity and orientation is also the key to providing a functional platform for parallel arrays of microreactors and chemo/biosensors. Protein adsorption on solid surfaces is influenced by a broad range of factors, including surface chemistry, protein structure, protein concentration, temperature, and solution properties such as pH and ionic strength.5-7 Essentially, this complex phenomenon is controlled by a dynamic interplay of intermolecular forces between the protein and the surface mediated by the solvent. For example, it has been shown experimentally that protein folding changes over time after adsorption onto solid surfaces, and that the orientation of adsorbed proteins can vary according to the charge and polarity of the surface.8 This sensitivity of protein adsorption to the surface chemistry obviously can afford an additional route for patterning protein micro/nanoarrays. *Corresponding author. E-mail:
[email protected] (Q.L.); wolfgang.
[email protected] (W.K.). (1) Lee, K. B.; Kim, E. Y.; Mirkin, C. A.; Wolinsky, S. M. Nano Lett. 2004, 4 (10), 1869–1872. (2) Lynch, M.; Mosher, C.; Huff, J.; Nettikadan, S.; Johnson, J.; Henderson, E. Proteomics 2004, 4(6), 1695–1702. (3) Wilson, D. S.; Nock, S. Curr.Opin. Chem. Biol. 2002, 6(1), 81–85. (4) Wilson, D. S.; Nock, S. Angew. Chem., Int. Ed. 2003, 42(5), 494–500. (5) Norde, W. Cells Mater. 1995, 5, 97–112. (6) Clarke, M. L.; Wang, J.; Chen, Z. J. Phys. Chem. C 2005, 109, 22027–22035. (7) Kim, J.; Somorjai, G. A. J. Am. Chem. Soc. 2003, 125(10), 3150–3158. (8) Mermut, O.; Phillips, D. C.; York, R. L.; McCrea, K. R.; Ward, R. S.; Somorjai, G. A. J. Am. Chem. Soc. 2006, 128(11), 3598–3607.
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Protein patterning techniques can be categorized into two major streams: the first type utilizes a high-resolution protein dispensing system, represented by inkjet printing9-12 and dip-pen methods;1,13,14 therefore the pattern resolution is primarily determined by the dispensing instrumentations, whose best resolution is limited to ∼100 nm. The second type of technique utilizes an underlying chemically heterogeneous surface and the preferential immobilization of proteins. As examples of this latter category, widely used lithography methods, however, also suffer from limited resolution as well as laborious and expensive operations.9,15-18 Recent advances in block copolymer (BCP) self-assembly has provided a possibility to conveniently prepare chemically heterogeneous surfaces with genuine nanosize resolutions.19,20 Such BCPs are composed of chemically distinct polymer blocks with unfavorable interblock interactions that are covalently joined together. By controlling the molecular weight and block volume fractions, a variety of periodic nanoscale morphologies may be self-assembled via microphase (9) Xu, Q. C.; Lam, K. S. J. Biomed. Biotechnol. 2003, 5, 257–266. (10) MacBeath, G.; Schreiber, S. L. Science 2000, 289(5485), 1760–1763. (11) Zhu, H.; Bilgin, M.; Bangham, R.; Hall, D.; Casamayor, A.; Bertone, P.; Lan, N.; Jansen, R.; Bidlingmaier, S.; Houfek, T.; Mitchell, T.; Miller, P.; Dean, R. A.; Gerstein, M.; Snyder, M. Science 2001, 293(5537), 2101–2105. (12) Fang, Y.; Frutos, A. G.; Lahiri, J. J. Am. Chem. Soc. 2002, 124(11), 2394– 2395. (13) Lee, K. B.; Park, S. J.; Mirkin, C. A.; Smith, J. C.; Mrksich, M. Science 2002, 295(5560), 1702–1705. (14) Hyun, J.; Ahn, S. J.; Lee, W. K.; Chilkoti, A.; Zauscher, S. Nano Lett. 2002, 2(11), 1203–1207. (15) Fodor, S. P. A.; Read, J. L.; Pirrung, M. C.; Stryer, L.; Lu, A. T.; Solas, D. Science 1991, 251(4995), 767–773. (16) Senaratne, W.; Sengupta, P.; Jakubek, V.; Holowka, D.; Ober, C. K.; Baird, B. J. Am. Chem. Soc. 2006, 128(17), 5594–5595. (17) Pallandre, A.; De Meersman, B.; Blondeau, F.; Nysten, B.; Jonas, A. M. J. Am. Chem. Soc. 2005, 127(12), 4320–4325. (18) Bruckbauer, A.; Zhou, D. J.; Kang, D. J.; Korchev, Y. E.; Abell, C.; Klenerman, D. J. Am. Chem. Soc. 2004, 126(21), 6508–6509. (19) Fasolka, M. J.; Mayes, A. M. Annu. Rev. Mater. Res. 2001, 31, 323–355. (20) Hamley, I. W. Nanotechnology 2003, 14(10), R39–R54.
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separation.19,20 Their broad applications have been demonstrated in fabricating hierarchically ordered functional materials,21,22 lithography masks,23 and optical materials.24,25 For nanopatterning applications, BCP thin films with block domains oriented normal to the substrate surface, which present surface chemical patterns consisting of periodic lines or hexagonal arrays, are particularly useful. Recent reports have demonstrated submicroand nanoscale protein patterns templated by BCP substrates, which present a contrast in the affinity for protein adsorption between the different domains.26-30 In particular, the advantages of BCP self-assembly and the simplicity of protein physical adsorption may be combined to prepare protein nanoarrays with high resolution and retained bioactivity.27,29,30 We have prepared, for instance, immunoglobulin-G (IgG) and bovine serum albumin (BSA) nanoarrays consisting of protein clusters 30 nm in diameter templated by nanopatterned polystyrene-block-poly(methyl methacrylate) (PS-b-PMMA) BCPs.30 It is noted that most of the proteometric research and practice is carried out in aqueous environment under fluid flow, such as the microfluidic platforms. In some cases, the protein arrays are subjected to the buffer and bioassay flows for extended periods of time. Therefore, the protein arrays discussed above, where the proteins are immobilized only by noncovalent intermolecular forces, would need to be robust enough under a buffer flow field to be useful for sensing applications. Furthermore, in our previous demonstration of adsorbed protein nanoarrays, we have shown that both the surface chemistry and the application of a fluid flow field were important for directing proteins to exclusively adsorb on the particular BCP domains, i.e., the PS domains.30 Hence, it is important to understand the effects of fluid flow fields on the BCP-templated, physisorbed protein nanoarrays. Research on protein-surface interactions in the presence of fluid flow is also of fundamental scientific interest, and is pertinent to micro/nanofluidics design which involves the handling of proteins. In this paper we focus our investigation on the interactions between the adsorbed protein and the PS-b-PMMA copolymer surface and that between the protein and the fluid flow, in order to understand the patterning process and the stability of adsorbed protein arrays. In the past, the experimental investigation of protein-surface interactions under hydrodynamic conditions is complicated by the fact that the migration and rearrangement of adsorbed proteins on a surface with high protein coverage is difficult to monitor. However, such observations can be greatly aided by our ability to create absorbed protein nanopatterns, where flow-induced surface displacements of proteins are indicated by the geometric integrity of the adsorbed protein nanoarray. We first present experimental observations of the pattern evolution of the protein arrays being subjected to various flow fields, followed by a theoretical analysis to quantify the coopera(21) Ikkala, O.; ten Brinke, G. Science 2002, 295(5564), 2407–2409. (22) Krausch, G.; Magerle, R. Adv. Mater. 2002, 14(21), 1579–þ. (23) Park, M.; Harrison, C.; Chaikin, P. M.; Register, R. A.; Adamson, D. H. Science 1997, 276(5317), 1401–1404. (24) Kim, D. H.; Lau, K. H. A.; Joo, W.; Peng, J.; Jeong, U.; Hawker, C. J.; Kim, J. K.; Russell, T. P.; Knoll, W. J. Phys. Chem. B 2006, 110(31), 15381–15388. (25) Kim, D. H.; Lau, K. H. A.; Robertson, J. W. F.; Lee, O. J.; Jeong, U.; Lee, J. I.; Hawker, C. J.; Russell, T. P.; Kim, J. K.; Knoll, W. Adv. Mater. 2005, 17(20), 2442–þ. (26) Kumar, N.; Hahm, J. I. Langmuir 2005, 21(15), 6652–6655. (27) Kumar, N.; Parajuli, O.; Hahm, J. I. J. Phys. Chem. B 2007, 111(17), 4581– 4587. (28) Neto, C. Phys. Chem. Chem. Phys. 2007, 9, 149–155. (29) Kumar, N.; Parajuli, O.; Dorfman, A.; Kipp, D.; Hahm, J. I. Langmuir 2007, 23(14), 7416–7422. (30) Lau, K. H. A.; Bang, J.; Kim, D. H.; Knoll, W. Adv. Funct. Mater. 2008, 18 (20), 3148–3157.
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tive effects of the intermolecular and hydrodynamic forces in the system. A comparison is made between the experimental and theoretical results in order to develop an understanding of the protein-surface interactions under a fluid flow.
Experimental Methods PS-b-PMMA Template Preparation. The copolymer template used for patterning the protein nanoarray was prepared by a previously reported method.30,31 A diblock PS-b-PMMA (Polymer Source, Dorval, Canada) with a number molecular weight of 94 kg 3 mol-1 and a PS/PMMA volume ratio of 3:7, was used to prepare PS-b-PMMA thin films with a surface nanopattern composed of hexagonally arranged, 30 nm wide PS surface domains surrounded by a PMMA matrix, which have an average periodicity of 53 nm. The films were 25 nm thick, and were prepared by first spin coating a 1 wt % solution of the PS-bPMMA in toluene (3000 rpm, 20 s) on a “surface-energy-neutral” surface,31 and then annealing the copolymer above its glass transition temperature (180 C, 48 h in vacuum). Both silicon and glass substrates were used to support the surface-energyneutral surface, which was a thin layer of cross-linked PS-random-PMMA copolymer with a PS/PMMA ratio specifically tuned for the self-assembly of PS-b-PMMA copolymer thin film nanopatterns.31 Protein Pattern Formation. To create protein nanopatterns, the PS-b-PMMA templates were first immersed in protein solution for several minutes to allow for protein adsorption, then rinsed for 10 s in an open stream of deionized water. Biotinylated goat antirabbit IgG solutions in phosphate-buffered saline (PBS) at pH 7.4 were used throughout this study. The IgG was purchased from Sigma-Aldrich, and was affinity-purified and conjugated to biotin-N-hydroxysuccinimide ester as received. The optimal patterning conditions were a biotin-IgG concentration of 8.5 μg/mL and an immersion time of 6 min. Unfunctionalized IgG could also be patterned using a concentration of 14 ug/mL and an immersion time of 10 min. As reported elsewhere,30 after rinsing, we obtained a patterned protein array that matched the positions where the hexagonally arranged PS domains were exposed on the PS-b-PMMA surface. For characterization of the resulting pattern by atomic force microscopy (AFM) in air, the protein nanoarrays were rinsed in water and dried under a nitrogen flow. Flow Cell Study. Buffer flow was applied over the copolymer surface for a certain length of time at various flow rates. A polydimethylsiloxane (PDMS) liquid flow cell designed for in situ surface plasmon resonance (SPR) measurements was used for the flow experiments.30,32,33 The configurations and dimensions are as depicted in Figure 1: The flow chamber was formed by placing a 0.5 mm thick PDMS “o-ring” seal in between the protein adsorbed copolymer sample at the bottom and a quartz plate at the top. The inlet and outlet were short pieces of steel needles (0.8 mm inner diameter) placed within holes drilled into the quartz plate. The PDMS “o-ring” flow chamber was elliptical in shape. The major axis was 8 mm, and the minor axis was 3 mm. Tygon tubing (0.76 mm inner diameter) was fitted to the steel needles at the inlet and outlet, and the buffer flow rate was controlled by a peristaltic pump (Reglo digital, Ismatec, Germany). Flow rates between 60 and 3600 μL/min, corresponding to flow velocities of 0.04-2.4 m/ min, were accessible with our setup. AFM Characterizations. Tapping-mode AFM was performed with a Nanoscope IIIa Multimode (Digital Instruments/ Veeco Metrology, USA) using a 15 15 um scanner and 1.8 N/m, 70 kHz tetrahedral silicon microcantilevers (Olympus, Japan).
(31) Ryu, D. Y.; Shin, K.; Drockenmuller, E.; Hawker, C. J.; Russell, T. P. Science 2005, 308(5719), 236–239. (32) Knoll, W. Annu. Rev. Phys. Chem. 1998, 49(1), 569–638. (33) Yu, F.; Persson, B.; Lofas, S.; Knoll, W. J. Am. Chem. Soc. 2004, 126(29), 8902–8903.
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Figure 1. A schematic drawing of the flow cell geometry and dimensions.
Results and Discussions Surface Patterning and Antibody Adsorption in an Open Flow Field. The as-formed PS-b-PMMA copolymer film exhibited hexagonally arranged, circular, 30 nm PS domains with an average periodicity of 53 nm, which were surrounded by a PMMA matrix. Both silicon and gold-coated glass substrates were used. This structure is schematically shown in Figure 2A, and experimentally verified by tapping-mode AFM phase imaging (Figure 2B), which reveals the difference in mechanical properties of the PS and PMMA domains.34 Topography imaging did not reveal any regular height difference (see Supporting Information). Phase contrast measures the difference in energy dissipation of the tapping AFM tip on different polymers35 and revealed PS domains as dark regions surrounded by a lighter colored PMMA matrix surface.24,36 The simultaneously recorded AFM topography image shows an essentially flat surface with little height contrast to distinguish the nanopattern (data not shown).30 In accordance with the 30:70 PS/PMMA volume ratio applied, the measured PS surface fraction was about 29%. To create protein nanopatterns, the PS-b-PMMA templates were first immersed in an IgG solution for a set length of time to allow for protein adsorption (see Experimental Methods), and then rinsed for 10 s in an open stream of pure PBS or deionized water at a flow rate of 1-5 104 μL/min (without a confined flow chamber). Protein pattern formation was not sensitive to the range of flow rate used nor to the length of rinsing within a range of a few seconds to a few minutes. The effects of solution concentration and immersion time on pattern definition were reported elsewhere.30 A comparison between the AFM phase image of the initial PS-b-PMMA template (Figure 2B) and the AFM topography image after protein adsorption (Figure 2C) clearly shows the same domain/cluster diameter and repeat period on both surfaces. The adsorbed protein clusters are identified as aggregates, 3-5 nm raised in height above the flat PS-b-PMMA substrate surface, and ∼30 nm in diameter. Compared to the nominal size of an IgG (14 12 5 nm3),37 the measured protein cluster height suggests that the IgG’s were adsorbed with their long axes parallel to the surface. Moreover, the reproduction of the PS domain nanopattern by the adsorbed proteins demon(34) Lau, K. H. A.; Bang, J.; Craig, H.; Kim, D. H.; Knoll, W. Biomacromolecules 2009, 10, 1061. (35) Magonov, S. N.; Reneker, D. H. Annu. Rev. Mater. Sci. 1997, 27, 175–222. (36) Xu, T.; Kim, H. C.; DeRouchey, J.; Seney, C.; Levesque, C.; Martin, P.; Stafford, C. M.; Russell, T. P. Polymer 2001, 42(21), 9091–9095. (37) Harris, L. J.; Larson, S. B.; Hasel, K. W.; McPherson, A. Biochemistry 1997, 36(7), 1581–1597.
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Figure 2. (A) Schematic drawing of the PS-b-PMMA pattern on a neutral surface with proteins adsorbed on the PS domains; (B) AFM phase image showing the PS-PMMA ordered pattern. (C) AFM topography image showing that the IgG proteins are selectively adsorbed on the PS domains.
strates that the proteins had adsorbed exclusively on the PS domains. It should be stressed that our previous study has shown that, during immersion of the PS-b-PMMA surface in the IgG solution, the proteins were attached on both the PS and the PMMA domains;30 then the proteins were selectively retained on the PS domains after rinsing with a buffer flow, resulting in a protein nanoarray with a high-fidelity to the original copolymer pattern. In contrast, if a static wash was applied by gradual immersion of the sample in a bath of pure buffer, a considerable amount of adsorbed proteins were retained on the PMMA. Thus the selective adsorption is a result of the chemical contrast between PS and PMMA surfaces, as well as the shear force induced by the rinsing fluid flow applied after the adsorption step.30 In particular, it is important to appreciate that the PS domains are more hydrophobic than the PMMA surfaces.38 This difference in the chemical potential between the polymers is also the cause of the microphase separation that enables the BCP to self-assemble and form the chemically heterogeneous nanopatterned surface. Both experimental observations5,7,8,39 and computational deductions40 have demonstrated that a hydrophobic surface exhibits a higher affinity for protein adsorption than a hydrophilic surface, due to long-range (0-10 nm) attractive hydrophobic intermolecular interactions.41,42 On the contrary a hydrophilic surface provides a repulsive energy toward the hydrated protein counterbalancing the ubiquitous van der Waals attraction. On a hydrophobic surface, the fragments of proteins in contact with the surface experience a partial loss of secondary structure, in particular R-helices, owing to the strong hydrophobic attraction during the initial stage of protein adsorption.43,44 This leads to a (38) Mansky, P.; Liu, Y.; Huang, E.; Russell, T. P.; Hawker, C. J. Science 1997, 275(5305), 1458–1460. (39) Li, L.; Hitchcock, A. P.; Robar, N.; Cornelius, R.; Brash, J. L.; Scholl, A.; Doran, A. J. Phys. Chem. B 2006, 110(33), 16763–16773. (40) Raffaini, G.; Ganazzoli, F. Macromol. Biosci. 2007, 7(5), 552–566. (41) Israelachvili, J. N. Intermolecular and Surface Forces; Academic Press Limited: London, 1992. (42) Li, Q.; Jonas, U.; Zhao, X. S.; Kappl, M. Asia-Pac. J. Chem. Eng. 2008, 3 (3), 255–268. (43) Raffaini, G.; Ganazzoli, F. Langmuir 2003, 19(8), 3403–3412. (44) Raffaini, G.; Ganazzoli, F. Langmuir 2004, 20(8), 3371–3378.
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considerable degree of protein unfolding as well as a maximized number of contact amino residues between the protein and the hydrophobic surface of various metastable/stable orientations.43 Although it has been shown by computational studies that proteins can also adsorb and spread on hydrophilic surfaces as a result of cooperative effects, the attraction at each amino acid residue, or contact point, with a hydrophilic surface is not as strong as the hydrophobic interaction.43 For example, the attractive energy for each contacting amino acid residue in the adsorption of an albumin subdomain has been calculated to be 3 times higher on a hydrophobic graphite surface (57 ( 2 kJ/mol) than on a hydrophilic poly(vinyl alcohol) (PVA) surface (14.5 ( 2 kJ/mol, calculated with a cutoff distance of 5 nm).40 As mentioned above, the rinsing step has a defining role in inducing adsorption exclusively on PS domain surfaces. The hydrodynamic shear force exerted by the rinsing step was able to remove the proteins adsorbed on the PMMA domains but not those adsorbed on the PS. In other words, the buffer flow fulfills the task of discriminating the adhesion force for proteins adsorbed on the different domains. However, because the rinsing liquid flows over the solid substrate as a liquid film with ill-defined boundaries in this procedure, quantification of the shear rate of the flow is problematic. Protein Patterns under Confined Buffer Flow. A series of experiments have been conducted to test the stability of the protein nanopattern under the flow of PBS buffer solution (pH=7.4, room temperature), which often is the testing environment relevant to biosensing. Substrates with IgG nanopatterns adsorbed were placed at the bottom of the flow cell as shown in Figure 1. Constant flow rates of 600, 200, and 60 μL/min were applied. For each flow rate, the protein patterns were examined by AFM topography imaging after being subjected to the PBS flows for 30, 90, and 210 min. Figure 3 presents the typical IgG patterns after the different durations of rinsing under the three flow rates. The AFM images shown in Figure 3 were taken at the same relative position within the samples at all observation times (see Supporting Information). A close examination of the AFM topography images show that, after 30 min rinsing, the protein pattern under the 60 μL/min PBS flow remained essentially undisturbed (as compared to Figure 2C). However, in the cases of PBS flows for 30 min at 200 and 600 μL/min, although the regular hexagonal pattern was still discernible, rearrangements in the protein positions could be unmistakably detected. Moreover, the rearrangement was more pronounced in the case of 600 μL/min, as evidenced by the increased number of proteins spread over the surface in a random fashion. The patterns captured after 90 min rinsing show that, under the 60 μL/min PBS flow, the IgG pattern had been slightly changed. However, the hexagonal pattern was still obvious and largely intact. In the case of 200 μL/min, the additional 60 min PBS flow made a drastic rearrangement of the adsorbed proteins: instead of arranging in circular clusters in the original pattern, many of the proteins had been driven into ring arrangements that appeared to follow the surface interface between the PS and the PMMA domains, consistent with previous reports on the surface interface on a PS-b-PMMA nanopattern having a higher affinity for protein adsorption than on either PS or PMMA.34,45 In the case of 600 μL/min, the AFM image after 90 min shows a sweeping change in the protein arrangement with most of the ordering destroyed. It is also noted that, although there appeared a few (45) Parajuli, O.; Gupta, A.; Kumar, N.; Hahm, J.-i. J. Phys. Chem. B 2007, 111 (50), 14022–14027.
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Figure 3. The AFM topography images of the typical protein patterns under a constant PBS flow: upper row, 60 μL/min; middle row, 200 μL/min; bottom row, 600 μL/min; left column, after 30 min; middle column, after 90 min; right column, after 210 min. The flows applied are in the directions of up or down relative to the imaged samples as indicated by the arrow.
areas where the proteins formed into ringed arrangements, the majority of the protein clusters had coalesced into large formations somewhat elongated in shape. After 210 min rinsing at 60 μL/min, the AFM examination shows that the protein pattern had been mostly retained, although the ordering was noticeably compromised. The proteins also seemed to have expanded in area but flattened in height. The 60 μL/min pattern represents a stark contrast to the AFM images obtained for the other two flow rates. Prolonged 200 μL/min flow had further impaired the original hexagonal pattern. and the proteins had evidently been further rearranged by the PBS flow. The ring-type formations were still dominant, but they seemed to have been stretched wider, and more proteins had adsorbed on the PMMA areas. The AFM image of the 600 μL/min sample shows that the protein clusters have somewhat adopted a streamlined orientation induced by the PBS flow. It is seen that a number of IgG clusters coalesced into large clusters, with the rest stretched into more elongated forms (which could reduce the hydrodynamic drag). Taken together, the results shown in Figure 3 indicate that the fluid flow was able to rearrange the positions of the adsorbed proteins at sufficiently high flow rates or after a sufficiently long duration fluid shear. Higher flow rates (200 and 600 μL/min) caused more severe protein relocations, which destroyed the original hexagonal patterns, while the nanopatterns were not significantly influenced by a low enough flow rate (60 μL/min). Nonetheless, some protein migration from the inlet area of the flow cell to the outlet area was also observed (see Supporting Information). In summary, this set of experiments has demonstrated that the fluid flow rate and duration have a strong influence on the retention of a physisorbed protein pattern, and in general, on protein adsorption on heterogeneous surfaces. In view of the potential of adsorbed protein nanopatterns as a highthroughput biosensing platform, such a migration of proteins from their initially adsorbed sites deserves special attention for the optimization of protein nanoarray operation and substrate surface regeneration. An Analytical Analysis on the Forces. A theoretical analysis has been conducted to unravel the principal effects of the flow DOI: 10.1021/la901658w
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field on the migration of proteins on an adsorbing surface and the exclusive adsorption of IgG on PS domains on a PS-b-PMMA nanopattern. Because of the complex hierarchical structures of proteins, significant assumptions were made by neglecting the atomistic details. The dimensional data of the IgG’s used in this study are presented in Table 1. IgG’s are proteins produced by the vertebrate adaptive immune system for recognition of pathogens and other foreign objects.46 The molecular mass of an IgG is ∼160 kDa, and the proteins are generally idealized as Y-shaped molecular structures with nominal dimensions 14 12 5 nm3 (Table 1). With such a conformation, in general, there are both hydrophilic and hydrophobic functional groups on the native protein’s surface. As discussed earlier, AFM characterization shows that the height of the adsorbed IgG clusters was in the range of 3-5 nm (Figures 2 and 3), which is lower than the known dimensions of the IgG. This may be attributed to the protein unfolding when in contact with an adsorbing surface as well as the subsequent drying process. Investigations on the surface structure of adsorbed proteins from both experimental and molecular simulation approaches suggested that, when a protein is adsorbed on a flat surface at a low protein concentration, it tends to lie on the surface with its long axis parallel to the surface and undergo conformational changes at the interfacial plane, losing some of the secondary structures, in particular the R-helices.7,44 The adsorption is much more pronounced on a hydrophobic surface: as a result of the long-range hydrophobic attraction, it is likely that adhesion first occurs at hydrophobic residues distributed on the surface of the protein molecule as the protein reaches the vicinity of a hydrophobic surface. As the gap between the protein and the surface decreases, the protein is further adsorbed via van der Waals attraction between all residues of the protein and the surface. On the other hand, when a protein is in the vicinity of a hydrophilic surface, the hydrophilic residues on the protein are more prone to form the initial contacts with the surface. However, because of the hydration effect, the hydration repulsion can sometimes exceed the van der Waals attraction, which in fact is a contributing mechanism conferring some surfaces, e.g., poly(ethylene glycol) (PEG)-grafted surfaces, with nonfouling properties.48 When the system is subjected to a liquid flow parallel to the surface, the adsorbed protein particle is also under the work of shear stresses induced by the fluid flow. Depending on the fluid flow field and
the magnitude of the shear stress, the proteins may be “rolled” by the torque induced by the shear force in the direction parallel to the surface. Such phenomena have been observed and analyzed in detail for cell adhesion under shear.49,50 Although the two systems are of different length scales, they most likely share the same fundamental principles: shear force versus intermolecular adhesion forces. Current protein models calculated by either the energy minimization algorithm or molecular dynamics can be accurate down to the atomistic scale. However, it is cumbersome to combine these protein models with a fluid dynamics simulation to describe the motion of an adsorbed protein as a response to a hydrodynamic input, because of the wide differences in the relevant length and time scales between the microscopic (protein) and macroscopic (hydrodynamic) systems.40 Therefore, herein we propose a simplified analytical model to elucidate the experimental observations concerning protein adsorption on a chemically heterogeneous surface under a shear flow. We approximate the IgG protein as a spherical particle with a volumetric diameter of 7.3 nm (Table 1) adsorbed on a flat surface with small contacting asperities, i.e., the amino acid residues interacting with the surface. These surface-contacting residues are approximated by hemispherical protrusions 0.5 nm in radius. Since only the ones distributed at the bottom of the protein particle in contact with the substrate are of interest, a simplified sketch is given in Figure 4 to present the conceptual model. As a result of the short range of the effective distance, van der Waals forces only need to be considered at contacting amino acid residues.48 By assuming a hemispherical geometry for these contacting points, the van der Waals force can be estimated by F = Ar/6D2, where F is the adhesion force, A is the Hamaker constant, r is the radius of the small hemisphere, and D is the distance between the small hemisphere and the planar surface. Since the protein adsorption on PS domains was predominantly driven by hydrophobic interactions mediated by the hydrophobic amino residues of the proteins,8 at these hydrophobic-hydrophobic contact points, a D of 0.4 nm can be assumed.41 The Hamaker constant between a hydrophobic amino acid residue and PS in PBS buffer may be approximated as 1 10-21 J.41 Hence, the adhesion force is estimated to be on the order of 1.0 10-12 N. Finally, the total adhesion force experienced by a protein particle is nF, where n is the total number of the contacting amino acid residues. When a protein is adsorbed on a hydrophilic surface, e.g., the PMMA domains on the PS-b-PMMA surface, an adsorption model analogous to that presented above may be applied. As observed by Melmut and co-workers,8 weakly adsorbed proteins on a hydrophilic surface also undergo protein spreading but at a much slower pace. The Hamaker constant may remain the same as above, but the interfacial distance, D, is significantly enlarged as a result of the presence of a hydration layer between the protein and the surface. A D of 1 nm is estimated on the basis of previously reported data.51 The adhesion force of these hydrophilic amino acid residues in contact with PMMA is calculated to be 1.67 10-13 N, an order of magnitude lower than that on the hydrophobic PS surface. Since the free energy gain for the contact interaction on a hydrophobic surface is higher than that on a hydrophilic surface, a protein adsorbed on a hydrophobic surface may tolerate a higher level of protein unfolding in order to increase n, the total number of contacting amino acid residues.
(46) Lesk, A. Antibody structure. In Introduction to Protein Science: Architecture, Function, and Genomics; Oxford University Press: New York, 2004; p 270. (47) Lindahl, E.; Hess, B.; Spoel, D. v. d. J. Mol. Model. 2001, 7, 306–317. (48) Ostuni, E.; Chapman, R. G.; Holmlin, R. E.; Takayama, S.; Whitesides, G. M. Langmuir 2001, 17(18), 5605–5620.
(49) Chang, K. C.; Hammer, D. A. Biophys. J. 2000, 79(4), 1891–1902. (50) Greenberg, A. W.; Brunk, D. K.; Hammer, D. A. Biophys. J. 2000, 79(5), 2391–2402. (51) Butt, H.-J.; Graf, K.; Kappl, M. Physics and Chemistry of Interfaces, 2nd ed.; Wiley-VCH: Weinheim, Germany, 2006.
Table 1. Characteristic Physical Data of IgG’s 46
molecular mass: 160 kDa approximate protein dimensionsa 14 12 5 nm3 b molecular volume 200.3 nm3 equivalent diameterc 7.3 nm 34 nm2 minimum footprintd,2 maximum footprintd,2 123 nm2 a IgG’s are “Y”-shaped proteins. The longest length is that between the tips of the “Y”, the shortest is the height of an IgG adsorbed “flat” on a surface, and the intermediate length is the height of the “Y”. However, the protein has been crystallized with a triclinic unit cell: a = 6.6 nm, b = 7.7 nm, c = 10.1 nm, R = 88.05, β = 92.35, γ = 97.23).37 b Calculated from the displaced volume of water molecules, using the molecular dynamics simulation package GROMACS,47 as the protein is immersed in a 20 20 20 nm3 volume. c The equivalent diameter is defined as that of a sphere occupying a volume equivalent to the molecular volume of the protein. d Minimum and maximum footprints: the areas projected by the protein adsorbed “standing up” and adsorbed flat on a surface, respectively.2
12148 DOI: 10.1021/la901658w
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Article
Figure 4. (a) A simplified model of a globular protein in contact with a solid surface. The asperities represent the amino acid residues. (b) Under a shear flow, when the shear force is large enough, it can break up the bonds one by one, therefore the protein can roll on the surface.
In other words, the value of n for an IgG adsorbed on a PS surface is greater than that on a PMMA surface, therefore, the difference in the total adhesion, nF, between the two scenarios would be greater than the order of magnitude difference calculated on the basis of a single contact point. This adhesion difference provides an operative window where an external force may be applied to remove the less strongly adhered proteins, which may be provided by a hydrodynamic shear force. Concerning the shear force, as a first approximation, we consider the protein particle as a sphere attached on a planar surface (Figure 4). The shear force of such a system in a flow channel can be calculated by52,53 Fs ¼ 32R2 τ where τ ¼ μ
4v¥ h
and v¥ ¼
3φ 2hw
ð1Þ
where Fs is the shear force on a single particle; R is the protein particle radius (assumed to be 3.7 nm, see Table 1); τ is the shear rate; μ is the fluid viscosity; ν¥ is the fluid velocity in the center of the fluid field; φ is the volumetric flow velocity; h is the height of the flow cell, i.e. 0.5 mm; and w is the width of the flow channel, averaged as 3 mm (Figure 1). As shown in eq 1, the shear stress is in linear proportion to the flow rate within the flow cell. When the flow rate φ is 600 μL/min, i.e., ν¥ = 1.3 10-4 m/s, the shear stress experienced by the protein particle is 4.43 10-13 N. At 200 μL/min, i.e., ν¥ = 4.4 10-5 m/s, the shear stress on the particle is 1.48 10-13 N. When the flow rate is reduced to 60 μL/min, the shear stress exerted on the protein becomes 4.43 10-14 N. From eq 1, we can also obtain a degree of understanding of the protein pattern formation induced by an open flow over a glass slide, although not in the most accurate manner: here w may be taken as the width of the glass slide, h theoretically can be considered as infinite because no physical upper boundary is given, thus v approaches 0, hence the Fs in this case is much smaller compared to the shear induced by a flow field in a confined channel. It should be noted that an exact calculation of the shear force under such an unconfined configuration is difficult. Experimentally, we note that the volumetric flow rate used in the open flow rinsing step to generate protein nanopatterns, at 1-5 104 μL/min, was 2 to 3 orders of magnitude higher than the rate used in the flow cell rinsing experiments described; Flushing the sample surface without using a flow cell appears to provide just the right magnitude of force required to rearrange the IgG adsorbed on the PMMA domains without disturbing the proteins on the PS domains. Attempts to replicate the shear force (52) Goldman, A. J.; Cox, R. G.; Brenner, H. Chem. Eng. Sci. 1967, 22(4), 653– &. (53) Forero, M.; Thomas, W. E.; Bland, C.; Nilsson, L. M.; Sokurenko, E. V.; Vogel, V. Nano Lett. 2004, 4(9), 1593–1597.
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required to generate the exclusive patterning of proteins on the PS domains within a confined flow cell were not successful. Using ν¥ =1.3 10-5 m/s (60 μL/min) for the rinsing step, the lowest flow accessible with our equipment, no discernible difference in adsorbed protein concentration between the PS and PMMA domains could be observed, indicating that the shear force applied was already too high to produce an ordered nanopattern (data not shown). These estimated forces are consistent with observations of the rinsing experiment presented in Figure 3. Since the adhesion force at each contacting amino acid residue between the protein and PMMA (1.67 10-13 N) is an order of magnitude weaker than that between the protein and PS (1.04 10-12 N), a range of liquid flow rates are capable of removing the proteins from the PMMA domains while leaving the proteins on the PS domains unaffected. In fact, we have previously shown that the proteins removed from the PMMA domains actually readsorbed on the PS domains which were unoccupied by proteins in the initial adsorption step.30 Calculations based on eq 1 show that the estimated shear force (4.43 10-13 N) caused by the 600 μL/min flow on a single IgG protein is higher than the single amino residue contact point adhesion between IgG and the PMMA (1.67 10-13 N) and lower than the amino residue contact adhesion between IgG and PS (1.0 10-12 N). Therefore, in principle, the shear force is not strong enough to disrupt the IgG adsorption on the PS surface. This may explain the observation that the adsorbed protein pattern was still recognizable after being subjected to the 600 μL/min flow for 30 min. However, after being subjected to a longer period under the 600 μL/min flow (e.g., 90 min), it was observed that the fluid shear has broken off the bonds and moved the proteins away from the PS areas. This gradual disintegration of the adsorbed protein pattern may be attributed to the storable elastic energy: since the proteins are deformable macromolecular structures, under a constant shearing force the protein is able to store the applied mechanical energy as elastic deformations, hence change its relative position against the surface, as illustrated in Figure 4b. Such accumulated strains, together with the applied shear flow, may be able to break the adhesion points and continuously deform the protein. Also, due to the existence of multiple contact points constituting the adsorption of a protein, it takes time to disrupt the contact points one after another. As a result, the “migration” of proteins on the polymer surface was observed to be a slow, time-dependent process. With regards to the flow experiment at 60 μL/min, since the shear force was an order of magnitude weaker than at 600 μL/min, i.e., the exerted mechanical energy was at least an order of magnitude smaller, the flow had much less effect on the stability of the adsorbed IgG array over a long period of time (>90 min), in stark contrast to the other two flow rates tested (Figure 3). DOI: 10.1021/la901658w
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Conclusions In this paper we have investigated the influence of fluid flow on the patterning mechanism and the stability of an adsorbed protein nanoarray templated by a chemically heterogeneous polymer surface. We discussed in detail how the protein nanopattern formed as a result of the interplay between weak intermolecular forces and an applied hydrodynamic shear force. Because the IgG-PS adhesion at each amino acid residue in contact with the PS surface is roughly an order of magnitude stronger than the IgG-PMMA attraction, the hydrodynamic shear applied during an appropriate rinsing step was able to remove the IgG’s adsorbed on the PMMA domains while leaving the ones on the PS domains unaffected. However, when a stronger hydrodynamic shear was introduced, for example, by implementing a confined flow duct over the adsorption surface, the proteins attached on the PS domains could also be detached and rearranged by the flow shear over time. We demonstrated this by applying a range of PBS buffer flow rates over varying lengths of time in a geometrically well-defined flow cell over the protein patterned surface. A flow velocity on the order of 1 10-5 m/s was identified as a suitable operating condition for the adsorbed protein nanopattern, with little loss in the integrity of the nanopattern over a length of time relevant to biosensing applications (>120 min). However higher flow rates and longer periods of fluid flow were able to quickly disrupt the protein pattern. A simplified theoretical model was also developed to understand the hydrodynamic effects on proteins adsorbed on solid surfaces. The competition between protein-surface adhesion and
12150 DOI: 10.1021/la901658w
hydrodynamic shear force exerted on the protein particle was examined, and the model also alluded to the contribution of protein deformation to the gradual migration, possibly rolling movement, of proteins adsorbed on the surface. Moreover, despite the highly simplified approximations used, the model provides a quantitative evaluation on the force magnitudes of the competing van der Waals attraction and the hydrodynamic shear. In summary, this study provides valuable information on the formation mechanism and stability of physisorbed protein nanopatterns under conditions concerning biosensing applications. It also represents an explorative study of protein adsorption under hydrodynamic flow conditions, which may assist in better designs of flow cells and other fluidic devices relevant to protein studies. Acknowledgment. We gratefully acknowledge Dr. Sung Hyun Park in the Department of Biomedical Engineering at Northwestern University for GROMACS calculations. Q.L. acknowledges the support from the Australian Research Council (Project: DP0588727) and the hosting of the Max Planck Institute for Polymer Research; D.H.K. acknowledges the Korea Science and Engineering Foundation (KOSEF) grant funded by the Korea government (MEST) (No. R11-2005-008-00000-0). Supporting Information Available: Observations through AFM on protein migration from the inlet area of the flow cell to the outlet area. This material is available free of charge via the Internet at http://pubs.acs.org.
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