[a] Diblock copolymer where x and y give the numbers of monomer repeat units. [b] Obtained from scanning force microscopy and high resolution electron microscopy. [c] Obtained from scanning electron microscopy. [d] Optical phase microcopy. [e] Vinculin or integrin clustering observed by confocal microscopy. [f] In a microsquare pattern.
Activation of Integrin Function by Nanopatterned Adhesive Interfaces
Article first published online: 10 MAR 2004
Copyright © 2004 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
Volume 5, Issue 3, pages 383–388, March 19, 2004
How to Cite
Arnold, M., Cavalcanti-Adam, E. A., Glass, R., Blümmel, J., Eck, W., Kantlehner, M., Kessler, H. and Spatz, J. P. (2004), Activation of Integrin Function by Nanopatterned Adhesive Interfaces. ChemPhysChem, 5: 383–388. doi: 10.1002/cphc.200301014
- Issue published online: 10 MAR 2004
- Article first published online: 10 MAR 2004
- Manuscript Received: 17 OCT 2003
- cell adhesion;
To study the function behind the molecular arrangement of single integrins in cell adhesion, we designed a hexagonally close-packed rigid template of cell-adhesive gold nanodots coated with cyclic RGDfK peptide by using block-copolymer micelle nanolithography. The diameter of the adhesive dots is <8 nm, which allows the binding of one integrin per dot. These dots are positioned with high precision at 28, 58, 73, and 85 nm spacing at interfaces. A separation of ≥73 nm between the adhesive dots results in limited cell attachment and spreading, and dramatically reduces the formation of focal adhesion and actin stress fibers. We attribute these cellular responses to restricted integrin clustering rather than insufficient number of ligand molecules in the cell-matrix interface since “micro-nanopatterned” substrates consisting of alternating fields with dense and no nanodots do support cell adhesion. We propose that the range between 58–73 nm is a universal length scale for integrin clustering and activation, since these properties are shared by a variety of cultured cells.
Cell–cell and cell–extracellular matrix (ECM) adhesion are complex, highly regulated processes that play a crucial role in most fundamental cellular functions including motility, proliferation, differentiation, and apoptosis.1, 2 Focal adhesions and related structures are major cellular sites responsible for cell-ECM attachment and adhesion-mediated signaling. These complex multimolecular assemblies consist of transmembrane integrin receptors that are linked to the actin cytoskeleton via cytoplasmic anchor proteins, such as vinculin, paxillin, and focal adhesion kinase.2, 3 Integrins are heterodimers formed by noncovalent association of α and β subunits that bind to a RGD (arginine-glycine-aspatate) motif, a sequence present in many ECM proteins. It had been shown that the assembly of the molecular complexes at focal adhesion sites require both occupancy and clustering of the integrin receptors, though the molecular properties of these clusters have not been defined.4, 5
To address the precise molecular topology of focal adhesions one needs to create model patterned surfaces where molecularly well-defined adhesive spots are separated by nonadhesive regions so that individual integrins are positioned in the cell membrane by the interaction of a cell with such surfaces. This approach is quite powerful since it enables the testing of the effect of ligand presentation on specific cellular responses entirely to the interaction with specific ligands, and how lateral positioning of single integrins effects cell functions. An essential requirement for such cell adhesion assays is the lateral control of specific adhesive and inert sites on surfaces. Polyethylene glycol (PEG) based substrates are widely used as biologically inert interfaces.6–10 The surface concentration and spatial distribution of cell-adhesive ligands may be controlled by mixing bioactive macrosystems with unsubstituted molecules,9, 11 dynamically by electrochemical control of ligand release12 or by the use of lithographic techniques.13 So far, an understanding of how adhesion and signaling of cells depend on size and distribution of focal adhesions was limited to sub-micrometer patches, due to technological limitations. Using microcontact printing, surfaces with adhesive and nonadhesive domains at length scales down to the micrometer level have been prepared. Such surfaces have been successfully used to geometrically control cell shape and viability.13 Even smaller patterns with cell adhesive patches of 200 nm diameter and 700 nm separation, prepared by dip-pen nanolithography, still showed attachment of cells.14 This resolution is rather low compared to the length scale at which protein clustering in focal adhesion occurs, which is in the range of 5–200 nm. More recently, it was demonstrated that nanotopography roughness of substrates affects cell function.15
Integrin-mediated cell adhesion to interfaces modified with appropriate ligands depends on many factors such as affinity and specifity of the ligands to the particular integrin, mechanical strength of the ligand support, spacer length, overall ligand concentration, surface topography and ligand density.16 Until now, methods for controlling all of these variables were limited, and usually suffered from nonhomogeneous ligand distribution on a molecular scale.17 Novel macromolecular designs of PEG molecules have the advantage that large and flexible chains may be responsible for different cell-binding activities mediated by anchoring compliance. However, such PEG macromolecules allow only for control of an average number of RGD peptides per macromolecule and only an average RGD-to-RGD distance can be calculated if RGD-coupled PEG macromolecules are mixed with unmodified ones.9, 10 Thus, this approach does not provide an accurate positioning of ligand molecules at surfaces, as is demonstrated here.
Results and Discussion
In earlier work,18–21 we described a substrate-patterning strategy based on self-assembly of diblock copolymer micelles. After the assembly of the Au-dot-containing micelles, the polymer is entirely removed by a gas plasma treatment, which results in extended and highly regular Au nanodots, deposited into a nearly perfect hexagonal pattern on substrates such as glass. Herein, Au-nanodot-patterned interfaces are prepared on glass cover slips or Si wafers by self-assembly of polystyrene-block-poly[2-vinylpyridine(HAuCl4)0.5] diblock copolymer micelles, that is, PS-b-P[2 VP(HAuCl4)0.5]. Regular hexagonally close-packed Au dots are shown as insets in Figure 1 a-d as bright spots in a scanning electron microscopy (SEM) image. Distances between Au dots are controlled by the molecular weight of diblock copolymers (Table 1). A Au nanodot of about 8 nm diameter provides a single anchor point to which only one integrin molecule can bind, since the diameter of integrin in the cell membrane is between 8–12 nm.22–24 Given that each nanodot can anchor only a single integrin molecule, a uniform patterning of extended substrate areas by self-organization of diblock copolymers25–27 can provide an accurate length-scale for inter-integrin spacing which is essential for cell adhesion. To serve as specific ECM-mimetic structures, Au dots are functionalized by “c(RGDfK)-thiols” (the cyclic peptide linked via the spacer aminohexanoic acid to mercaptopropione acid) that contain the cell-adhesive RGD sequence, which is recognized by αvβ3-integrin with high affinity.28, 29 To eliminate cell binding to the areas between the Au dots, these areas were passivated by PEG.
|PS(x)-b-P2 VP(y)[a]||Au dot diameter[b][nm]||Au dot separation[c][nm]||Au dot density [dots/μm2]||Cell spreading[d]||Focal adhesion formation[e]||Actin fiber formation[e]|
In Figure 1, MC3T3-osteoblasts were seeded on glass and examined after one day by optical bright-field microscopy. Only three-quarters of the glass substrate area was patterned with Au nanodots with different spacings between the dots. The Au dots were functionalized by c(RGDfK)-thiols and the free glass was passivated by PEG. A line of cells marks the borderline of the nano-pattern area (white arrows). The right side was entirely passivated against cell adhesion—thus, cell adhesion and attachment is only observed on the left side of the images. When plated on Au-nanodot patterns with various spacings, functionalized by c(RGDfK)-thiols, MC3T3-osteoblasts show different adhesion behaviors (Figure 1 a-d). It is evident that cells spread very well on the 28 (Figure 1 a) and 58 nm (Figure 1 b) patterns, appearing as they do on uniformly RGD- or fibronectin-coated surfaces. On the other hand, hardly any cell spreading is observed on substrates with 73 (Figure 1 c) and 85-nm spaced nanodots (Figure 1 d). Quiescent and migrating cells can be noted. Quiescent cells (yellow arrows) present a rounded shape which causes strong scattering of light, while migrating cells (green arrows) are usually characterized by extended filopodia. These observations have been repeated with additional cell types, that is, REF52-fibroblasts, 3T3-fibroblasts, and B16-melanocytes, indicating a universally characteristic cell-adhesion behavior. Figure 1 e shows MC3T3-osteoblasts on Au nanodots separated by 58 nm and not conjugated to c(RGDfK)-thiols. Cell spreading on these surfaces is rather poor. Only a few cells remain attached after gentle rinsing. Different adhesion properties of cells on these interfaces are monitored by displaying the cell number density of essentially attached cells in Figure 1 f.
Since c(RGDfK)-thiols have a high affinity for αvβ3-integrins,28, 29 these Au-nanodot/c(RGDfK)-thiol interfaces provide an ideal stiff matrix of adhesive patches to study this integrin clustering. The molecular flexibility of the c(RGDfK)-thiol molecules on an Au dot is weak as the spacer between the thiol and the c(RGDfK) is short. The largest diffusion amplitude of a c(RGDfK) molecule, as estimated from molecular dimensions of the peptide, is assumed to be smaller than 10 Å.
The molecular formation of focal contacts and the assembly of actin stress fibers in MC3T3-osteoblasts and B16-melanocytes adhering on these nanopatterned substrates were investigated by immunohistochemical staining for integrin, vinculin, focal adhesion kinase (FAK), and actin in Figure 2. Co-localization of integrin and FAK were observed only when Au dots were separated by ≤58 nm and covered with c(RGDfK)-thiols (Figure 2 a). Separations of ≥73 nm yielded limited spreading (Figure 2 b). Focal adhesion is strongly expressed only at cell extension and around the nucleus. Apparently, cells detect chemical signals from the c(RGDfK)-thiol adhesive dots but stable focal adhesions cannot be formed when the interdot spacing exceeds 73 nm. In the case of pattern where Au nanodots were not covered by c(RGDfK)-thiols (respective insets in Figure 2 a and 2 b), GFP-integrin β3 is mainly localized at the cell periphery, however without clustering. In both patterns the expression of FAK is only detectable in the perinuclear region.
Integrin-mediated adhesion induces actin stress fiber formation and vinculin recruitment to focal adhesions as evidenced in cells plated on a homogenous surface coated with c(RGDfK) (Figure 2 c), or fibronectin (Figure 2 d) and on Au/RGD-dot pattern with ≈28 and ≈58 nm spacing (Figure 2 e,f). Well-constituted and rather long vinculin clusters (green) as well as organized actin stress fibers (red) can be noted in confocal micrographs, which indicate good cell adhesion. Nonorganized vinculin and actin localization are observed by rather blurred molecular distribution imaging when Au dots are not covered by c(RGDfK)-thiols (Figure 2 i,j,k,l) or the interdot distance is larger than 73 nm (Figure 2 g,h).
The increase in dot separation distances causes a decrease in the number of dots. Therefore, the observed limitation of cell adhesion at increased dot separation could be reasoned either on the number of c(RGDfK)-thiol covered Au dots (Table 1) or on the local dot-to-dot distance. In order to address this issue we created “micro-nanostructured” interfaces.18,21 This technique allowed the deposition of a defined number of Au nanodots in a confined area of the substrate. The surfaces were designed such that the average dot density was 90 dots μm−2 and thus significantly smaller than in all cases of extended Au dot pattern (Table 1). The local dot density organized in 2×2 μm2 patches of 58 nm spaced dots was 280 dots μm−2 (Figure 3 a). Figure 3 b shows a bright field optical micrograph 3 h after plating of MC3T3-osteoblasts on the substrate. Clearly, cells are confined to the structured area only, and the process of cell spreading advanced as indicated in the inset. After 24 h (Figure 3 c), well-spread cells are present in this area whereas cells located outside the frame (indicated by arrows) are poorly spread. Figure 3 d illustrates a confocal fluorescent micrograph after immunohistochemical staining for vinculin (green) and actin (red) demonstrating the confinement of focal adhesion to the square pattern and the origin of actin stress fibers from there. In this case the mean average focal adhesion length is 2.6±0.9 μm (Figure 3 e, green columns). This value is between the side length of one square pattern and its diagonal. The distribution in focal adhesion lengths (Figure 3 e, green column) is remarkably narrow as it has to be confined by a square. Cells do not adhere to all squares, but in some areas a separation distance between focal adhesions of 1.5 μm is recognized, as illustrated by the inset in Figure 3 d. If cultured on a pattern uniformly structured with dots separated by 58 nm (Figure 2 f), these cells form focal adhesion lengths with an average mean value of 5.6±2.7 μm (Figure 3 e, white hatched columns). Interestingly, the mean focal adhesion width on the “micro-nanopattern” is 1.4±0.4 μm and thus is broader than the mean focal adhesion width formed on the extended Au-nanodot pattern which is 1.0±0.3 μm. The mean focal adhesion area is 3.2±1.3 μm2 on the “micro-nanopatterned” substrates and 5.5±2.9 μm2 on the extended Au nanopattern. Since the focal adhesion length formation is restricted by the “micro-nanostructured” pattern, a focal adhesion cluster uses the full confined adhesive site of a square by increasing its width. This underlines the meaningful contribution of quantitative examination of focal adhesion by molecularly confined structures to the understanding of its formation and regulation.
These adhesion experiments indicate that local dot–dot separation, rather than total number of dots, was critical for inducing cell adhesion and focal adhesion assembly. Thus, for example, the dot density located under cells attached to the “micro-nanostructured” squares, consisting of 58-nm separated dots, is considerable lower than that of dots located underneath cells attached to a substrate, uniformly patterned by dots, separated by ≤73 nm. Nevertheless, the cells did form focal adhesions on the former surface and failed to do so on the latter. This is schematically summarized in Figure 2 m. The ligand pattern quality and rigidity are superior and thus enabled the measurement of this important length scale in focal adhesion.
Integrin-mediated cell adhesion to interfaces depends on many factors such as affinity and specifity of ligands, mechanical strength of ligand support and linkage, spacer length, overall ligand concentration, and ligand clustering.16 Up to now, methods for variation ligand concentration on surfaces have suffered from nonhomogeneous ligand distribution on a molecular scale as shown, for example, in refs. 7, 9, 17. Smart macromolecular designs of PEG molecules such as PEG stars allow control of an average number of RGD peptides per star.9, 10 But only an average RGD cluster-to-cluster distance can be calculated if RGD-occupied PEG macromolecules are mixed with plain ones.9 This macromolecular approach has the advantage that large and flexible chains may be responsible for different cell-binding activity, mediated by cell membrane receptors and anchoring compliance. However, it does not control ligand clustering, as the ligand template is not well-ordered and too flexible.
Studies on the clustering of individual integrin molecules demand nanoscale surface patterning with a rigid ligand template that limits the possibility of integrin arrangements. Herein, we present the first demonstration of such patterned surfaces, consisting of a hexagonally packed template for c(RGDfK)-thiol peptides by self-assembly of diblock copolymers. Dots coated with c(RGDfK)-thiol peptides are positioned with high precision at 28, 58, 73, and 85 nm distance from each other. The size of nanodots (<8 nm) is small enough that only one integrin can bind to one dot. Thus, the template offers an ideal rigid c(RGDfK)-thiol nanopattern to which arrays of single integrins should bind. We show here that when adhesive dots are separated by ≥73 nm, cell adhesion and spreading, as well as the formation of focal adhesions, are aberrant, whereas separation of ≤58 nm between the dots allows effective adhesion. This feature is not attributable to insufficient number of ligand molecules but to the restriction of integrin clustering. We found the maximum spacing between c(RGDfK) occupied integrins necessary for adhesion and focal adhesion formation to be between 58 and 73 nm, not just for MC3T3-osteoblasts, but also for other cells, including B16-melanocytes, REF52-fibroblasts, and 3T3-fibroblasts (data for the last two cell lines are not shown). Thus, this length scale appears to be a universal spacing for effective adhesion mediated by integrins (Figure 2 m).
These first studies offer a unique opportunity to define the length scales of multimolecular complexes within focal adhesions with an unprecedented resolution as small as a single protein. Variations in nanodot organization, including alterations in ligand template pliability, changes in the patterns of nanodot fields, and presentation of small dot clusters (e.g., pairs or triplets) may shed light on the minimum molecular number of an effective integrin cluster necessary to obtain cell attachment, spreading or migration and of possible “pattern specific features” that trigger cell-adhesion-based signaling.
Materials and Methods
For the Au-nanodot pattern from diblock copolymer micelles, the preparation method is described in ref. 20, 21. The molecular characteristics of the polystrene-block-poly(2-vinylpyridine), that is, PS-b-P2 VP, diblock copolymers varied according to Table 1.
Immobilizing linear polyethylene glycols to glass interfaces on glass coverslips or Si-wafers (Crystec, Berlin) was performed by first chemically activating the substrates (H2SO4/H2O2=1/1). The substrates were immersed in a 1 mmol solution of a linear polyethylene glycol (CH3-(O-CH2-CH2)17-NH-CO-NH-CH2-CH2-CH2-Si(OEt)3) in dry toluene (p.a.) under nitrogen atmosphere for 5 days. Finally, substrates are rinsed intensively with toluene, methanol and ethyl acetate (all p.a. from Aldrich). The thickness of the layer was approx. 20 Å in the dry state as determined by multi-wavelength ellipsometry.
Synthesis of c[RGDfK(Ahx-Mpa)]=“c(RGDfK)-thiol”: The protected thiol-anchor S-trityl-3-mercaptopropionyl-aminohexanoic acid was prepared by solid-phase peptide synthesis (SPPS) loading TCP resin in a silylated flask with Fmoc-Ahx-OH (protected ω-aminohexanoic acid; Fmoc=9-fluorenylmethyloxycarbonyl) in dry CH2Cl2 under addition of 2.5 eq. diisopropyl amine (DIPEA). After deprotection with 20% piperidine the peptide-resin was coupled with S-trityl-3-mercaptopropionic acid with HOBt⋅H2O (HOBt=N-hydroxybenzotriazole) and O-(benzotriazol-1-yl)-N,N,N′,N′-tetramethyluronium tetrafluoroborate (TBTU) in 15 mL NMP g−1 resin. The pH was maintained at 8–9 by addition of DIEA (diisopropyl ethyl amine). After washing with CH2Cl2 four times, 3 min cleavage from the resin was performed with AcOH/trifluoroethanol (TFE)/CH2Cl2 3:1:6 (60 min). Residues of acetic acid were removed by azeotropic distillation with toluene and the product was lyophilized. The product was characterized by NMR spectroscopy and ESI-MS. The protected cyclic peptide c[R(Pbf)GD(OtBu)fK] (Pbf=2,2,4,6,7-pentamethyl dihydrobenzofuran-5-sulfonyl) was prepared via solid-phase peptide synthesis of the linear precursor on TCP resin and, after cleavage from the resin, cyclized using (R)–hexadecanoic acid, 1-[(phosphonoxy)methyl]-1,2-ethanediyl ester, monosodium salt (DPPA) as described in ref. 29. S-trityl-3-mercaptopropionyl-aminohexanoic acid (0.2 mmol), 0.97 equiv N-[(dimethylamino)-1H-1,2,3-triazole[4,5-b]-pyridin-1-yl-methylene]-N-methylmethanaminium hexafluorophosphate (HATU), 1.1 equiv 7-aza-1-hydroxybenzotriazole (HOAt) and 10 equiv 2,4,6-collidine were dissolved in 2 mL N,N-dimethylformamide (DMF). After 1.5 h 1 equiv of the protected cyclic peptide with deprotected Lys was added and stirred for 24 h at room temperature. After removing the solvent in vacuo the residue was treated with water and washed twice with acetonitrile, once with saturated NaHCO3 and twice with water. The product was lyophylized from tBuOH (67 % yield). Deprotection was performed with a mixture of 90 % trifluoroacetic acid (TFA), 3 % water and 7 % tetra iso-propylsilane (TIPS). After 6 h the solvent was removed at 40 °C. After complete removal of TFA the peptide was dissolved in methanol, precipitated with ether, washed several times with ether, and purified by preparative HPLC; yield 33 %. ESI-MS m/z: 805.3 (100) [m+H+].
For immobilization of c(RGDfK)-thiols on Au dots, the PEG-functionalized substrates were immersed for 24 h in a 25 μmol c(RGDfK)-thiol–water solution to link the molecule via the thiol group to the Au nanodots. The substrates were then rinsed extensively with MilliQ water (R>18 MΩ) and shaken for 24 h with several water exchanges to remove noncovalently bound c(RGDfK)-thiols.
MC3T3-osteoblasts, REF52-fibroblasts and B16-melanocytes expressing GFP-β3-integrin receptors were cultured in DMEM media (Invitrogen, Germany) supplemented with 5 % fetal bovine serum and 1 % L-glutamine (Invitrogen) at 37 °C and 10 % CO2. Only cells at passage 5–12 were used. Before plating on the patterned glass substrates, cells were trypsinized with 0.25 % trypsin and 1 mM ethylenediaminotetraacetic acid (EDTA) in HBSS. The samples were sterilized in 70 % ethanol and washed with phosphate-buffered saline (PBS) at room temperature. Cells were plated at a density of 50.000–100.000/sample in DMEM containing 1% FBS for all the experiments.
After 12–24 h on the substrates, cells were washed with PBS and fixed with 1.5 % paraformaldehyde in PBS for 10 min. The cells were made permeable with 0.1 % Triton X-100, blocked with 1 % bovine serum albumin (BSA) in PBS for 20 min and incubated with a 1:50 dilution of rabbit anti-focal adhesion kinase and with a 1:50 dilution of mouse anti-human vinculin (Sigma Chemicals, Saint Louis, MO), respectively, for 1 h at 37 °C. Then the cells were labeled with a 1:100 dilutions of TRITC-IgG and FITC-IgG (Sigma Chemicals, Saint Louis, MO) in PBS for 1 h. The filamentous actin was labeled with a 1 μg mL−1 dilution of TRITC-conjugated phalloidin overnight at 4 °C. The cells were visualized with an inverted confocal microscopy (Zeiss, Axiovert) using the LSM software imaging program.
The authors are grateful to the interest and extended support from Prof. B. Geiger (Weizmann Institute of Science), Prof. M. Grunze (Angewandte Physikalische Chemie, U. Heidelberg) and Prof. M. Möller (RWTH Aachen). As important were intensive discussions with Dr. J. Curtis (Biophysical Chemistry, U. Heidelberg). The work was financed by the Deutsche Forschungsgemeinschaft (DFG, SP-520/5-1).
- 28J. Biol. Chem. 1994, 269, 20 233., , , , , , , ,