Nanopatterned Adhesive, Stretchable Hydrogel to Control Ligand

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A Nanopatterned Adhesive, Stretchable Hydrogel to Control Ligand Spacing and Regulate Cell Spreading and Migration Jie Deng, Changsheng Zhao, Joachim P. Spatz, and Qiang Wei ACS Nano, Just Accepted Manuscript • DOI: 10.1021/acsnano.7b03449 • Publication Date (Web): 11 Jul 2017 Downloaded from http://pubs.acs.org on July 12, 2017

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A Nanopatterned Adhesive, Stretchable Hydrogel to Control Ligand Spacing and Regulate Cell Spreading and Migration Jie Deng,a,b Changsheng Zhao,b Joachim P. Spatz,a* Qiang Weia* a

Department of Cellular Biophysics, Max Planck Institute for Medical Research, Heidelberg,

and Laboratory of Biophysical Chemistry, University of Heidelberg, Jahnstraße 29, 69120 Heidelberg, Germany b

College of Polymer Science and Engineering, State Key Laboratory of Polymer Materials and

Engineering, Sichuan University, Chengdu 610065, China. KEYWORDS: Stretchable hydrogel, nanopattern, interligand spacing, heterogeneous interface, cell adhesion, cell migration.

ABSTRACT

Spatial molecular patterning enables the regulation of adhesion receptor clustering, and can thus play a pivotal role in multiple biological activities such as cell adhesion, viability, proliferation, and differentiation. A wide range of nanopatterned, adhesive interfaces have been designed to

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decipher the essence of molecular-scale interactions between cells and the adhesive interface. Although an interligand spacing of less than 70 nm is a proven prerequisite for the formation of stable focal adhesions, there is a paucity of data concerning how cells behave on substrates featuring heterogenous adhesiveness. In this study, a stretchable hydrogel functionalized with a quasi-hexagonally arranged nanoarray was stretched along one direction, resulting in ligands periodically arranged in a pattern resembling a centered rectangular lattice with an interligand spacing smaller than 70 nm in one direction and greater than 70 nm in the orthogonal direction. This substrate was utilized to modulate interligand spacing and investigate cell adhesion and migration. An interligand spacing larger than 70 nm – even in just one direction – prevented the establishment of stable focal adhesions. The stretched interface promoted dynamic remodeling at cell contacts, resulting in higher cellular mobility. Our nanopatterned stretchable hydrogel permits reversible control over cell adhesion and migration on nanopatterned ligand interfaces.

Numerous surface chemical and physical properties including surface chemistry,1, 2 stiffness,3, 4 topography,5-8 and combinations of these properties9-11 have been employed to control cell behavior at cell-material interfaces. Concerning topographical cues, it is well known that twodimensional molecular patterning enables the regulation of adhesion receptor clustering. In this way nanoscale topographical structures on adhesive interfaces can effectively modulate multiple cell behaviors including cell adhesion,12-14 differentiation,15-18 dedifferentiation,19 proliferation,20 and migration.6, 7 Moreover, the extracellular matrix (ECM) surrounding cells possesses intricate nano-topographic structures.21 The activation of specific transmembrane receptors such as integrins, which give rise to the assembly of adhesion sites known as focal adhesions (FA), is a


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key step during the adhesion of tissue cells to the ECM.22 The spatial organization and activation of integrins is mainly directed by epitopes that feature an Arg-Gly-Asp (RGD) sequence and which are situated on various adhesive ECM proteins.23 Besides the specific chemical sequence of the adhesive epitope, recent studies have also indicated that cells respond to many different environmental parameters such as stiffness,3, 24, 25 geometry,26, 27 dimensionality,28 and precise epitope spacing.29 These properties play paramount roles in modulating receptor-mediated adhesion formation and signaling, and thus influence cell characteristics like morphology, migration, differentiation, proliferation, and apoptosis.30 It has been verified that αvβ3 integrinmediated cell adhesion is significantly affected by how far ligands are separated apart.31 Specifically, Arnold et al.29 showed that a universal interligand spacing of less than 70 nm was essential for successful integrin clustering and subsequent cell-ECM adhesion. It was verified that nanopatterned cyclic RGDfK peptides with interligand spacings ranging from 25 to 120 nm affected cell adhesion, spreading, FA assembly, and migration in different ways.31 More precisely, when the adhesive nanodots are spaced ≥ 73 nm apart, cell adhesion, spreading, and FA formation are highly restricted. Conversely, interligand distances of ≤ 58 nm support effective cell adhesion. This phenomenon is attributed to the prevention of integrin clustering, rather than an insufficient number of adhesive ligands.29 Building on this principle, cell polarization and migration at adhesive interfaces – using nanoparticle spacing gradients between 50 to 250 nm and different gradient strengths – were further investigated. At around 80 nm distance between nanoparticles, cells tend to polarize and migrate in the direction of the smaller interligand distances, in other words, towards a higher density of adhesive ligands.31 Cell responses on a variety of nanopatterned adhesive interfaces have been extensively investigated.32 Recently, some heterogeneous nanoribbons, silicon nanoparticle arrays,


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hierarchical assembled indium tin oxide nanowire arrays, vertical nanopillar arrays, and nanoneedles have been designed for cell capture, sorting, and isolation.33 Nanostructured fibrous hydrogel can be used to support the three-dimensional cell culture of chondrocytes.34 Moreover, nanoimprint lithography was implemented to fabricate a nanopattened interface with predesigned protein binding sites.14 Our previous work demonstrated that an interligand spacing of less than 70 nm has been identified as a prerequisite for the induction of integrin clustering and FA assembly.29 Nevertheless, it remains unknown how cells would behave on an interface with a pattern that is heterogeneous with regard to its adhesiveness due interligand spacings of less than 70 nm in one direction and greater than 70 nm in the orthogonal direction. Previous reports mainly focused on adhesive interfaces with nanodots periodically arranged in a quasi-hexagonal lattice pattern, resulting in similar interligand distances between all neighbouring adhesion sites. We are curious to know whether cells can form stable adhesions on an interface like the one described above. Does the formation of FAs require densely arranged ligands (≤ 70 nm) only in a single dimension? If not, can we reversibly regulate cell adhesion and migration through variably tunable interligand spacing? We established a highly stretchable hydrogel system with nanopatterned adhesive ligands to address the above questions. Poly(N-acryloyl glycinamide) (PNAGA) hydrogels have previously been reported to be a stable and highly stretchable type of hydrogel. They don’t swell in water and maintain mechanical strength in a variety of aqueous solutions.35 Moreover, PNAGA-based hydrogels can prevent unspecific cell adhesion (Figure S1). To obtain an interface with heterogenous – in this case, periodic stripe-patterned – adhesiveness we stretched a cyclic RGDfK nanopatterned PNAGA hydrogel (Figure 1A) and utilized this as a platform for the study


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of cell behavior. We examined the effects of heterogeneously adhesive interfaces on cell spreading, FA formation, cell polarization, and cell migration.

RESULTS AND DISCUSSION In a first step, we fabricated a cyclic RGDfK nanopattern with an interligand spacing of 35 nm on a PNAGA hydrogel using a transfer strategy illustrated in Figure S2. Given that it was difficult to directly generate stable RGD nanoarrays on a polymeric interface,36 nanopatterns were first fabricated on glass slides via block copolymer micelle nanolithography (BCMN) and then transferred to the PNAGA hydrogel. BCMN technology makes it possible to position gold nanoparticles with a size of 1-15 nm in a quasi-hexagonal pattern with a tunable inter-particle distance between individual ligands ranging from 15-250 nm by varying the molecular weight of diblock copolymers.37 The detailed fabrication procedure is described in materials and methods. Previous research has demonstrated that gold nanoparticles anchored to the surface through adhesive cyclic RGDfK can bind no more than one integrin per particle each.38 Thus, local changes in integrin clustering upon cell adhesion mirror the cellular response sensitivity to spatial distances between individual adhesion molecules. First, the presence of the gold nanopattern on the glass slides was confirmed by scan electron microscopy (SEM). Figure S3A shows the homogeneously distributed gold nanoparticles in a quasi-hexagonal pattern. The average distance between the nanoparticles was about 35 nm. A cryo SEM image of the nanopatterned PNAGA hydrogel is shown in Figure 1C. The slightly deformation of the soft hydrogel during drying made the gel surface to be rough.10 The out-offocus nanodots made it difficult to distinguish the quasi-hexagonal pattern. As a result, the


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nanodots on the hydrogel seem random. To further confirm the transfer of the nanodots, the glass slides were imaged after transfer (see Figure S3B). No gold nanodots were left on the surfaces, suggesting their successful transfer to the PNAGA hydrogel surface. To test the stability of the nanodots on the hydrogel surface under stretching, we stretched the hydrogel up to 3.2 times its original length for 24 h followed by 24 hours of relaxation. For the most part the hydrogels recover to their original length. As shown in Figure 1D, the distribution of the gold nanodots after `stretching and relaxing´ was similar to before stretching. This confirms their stability on the hydrogel surface during stretching.

Figure 1. (A) Schematic illustration of cell adhesion studies on nanopatterned interfaces with homogeneous or heterogeneous adhesiveness. (B) Photographs of PNAGA hydrogels after stretching to different lengths. Morphological observations of the nanopatterned hydrogel surfaces prior to stretching (C) and after `stretching and relaxing´ (D). Next, the cyclic RGDfK with a spacer was anchored to the gold nanoparticles, resulting in a cyclic RGDfK nanopatterned PNAGA hydrogel with an interligand spacing of 35 nm. Given that the diameter of integrin in the cell membrane is between 8-12 nm and each nanodot can anchor up to one integrin molecule, the uniformly patterned surface can provide an accurate length scale for inter-integrin spacing.38 To produce interfaces with differing interligand spacings in orthogonal directions, the cyclic RGDfK nanopatterned PNAGA hydrogels were stretched in one direction to predetermined lengths (Figure 1B; unstretched hydrogels are referred to as L0, stretched hydrogels as Lx, where x represents the stretched:original length ratio). As illustrated in Figure 1A, the increase in the ratio between stretched:original interligand distance in the


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direction of stretching should be equal to the stretched:original hydrogel length ratio. Accordingly, stretching leads to a decreased ligand spacing in the direction orthogonal to stretching. Namely, the width for L1.5, L1.9, and L3.2 hydrogels became 91.7% 83.4%, and 62.5%, respectively, of its original width. Since the hydrogel is not completely elastic, the poisson’s ratio is not constant. The poisson’s ratios for L1.5, L1.9, and L3.2 hydrogels were 0.167, 0.184, and 0.170, respectively. Additionally, we also theoretically illustrated the distributions of the nanodots after being stretched to different lengths via ImageJ (Figure S4) to give a general impression of the distribution of the nanodots on the stretched hydrogels. For cell adhesion studies, the cyclic RGDfK nanopatterned PNAGA hydrogels were stretched to different lengths, namely L1.5, L1.9, and L3.2. MC3T3 cells were then seeded to the surface and cultured under standard cell culture conditions. In the first 3 h most cells on these surfaces exhibited a round shape without any morphological changes or polarization (Figure S5). After 15 h culture cell spreading was studied. Figure 2A shows optical images of the cells adhering to the surfaces of stretched hydrogels of different lengths. Cell adhesion behavior was influenced by the degree of stretching. Cells spread very well on both the unstretched hydrogel with an interligand spacing of 35 nm as well as on the L1.5 hydrogel with an interligand spacing of approx. 52 nm in the direction of stretching. In contrast, the spreading area of cells adhering to the surface of the L1.9 hydrogel (with approx. 67 nm interligand spacing in the direction of stretching) decreases (see Figure 2B). Hydrogel stretching to 3.2 times the original length (approx. 112 nm interligand spacing in the direction of stretching) leads to a decrease in the spreading area of the adhered cells to about 700 µm2 per cell. In comparison, cells on the L0 hydrogel spread to about 3100 µm2 per cell.


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Cells on the L3.2 hydrogel also exhibited an elongated shape. This lead us to also quantify the cell aspect ratio,31 a morphometric parameter of adherent cells defined as the axial ratio of the most extended length of the cell body and the most extended width of the cell body. Figure 2C shows that the mean aspect ratio of the cells adhering to the L3.2 hydrogels was about 6, much higher than that of the cells cultured on L0, L1.5, or L1.9 hydrogels. This proves that the cells on the L3.2 hydrogel are much more polarized. Moreover, these cells also display more filopodial structures. The orientation angle of the cells (defined in the insert in Figure 2G) was quantified in Figure 2D-G. There was no significant difference in the orientation angle between the cells adhering to L0 or L1.5 hydrogels. Cells preferred the unstretched direction over the stretched direction on the L1.9 and L3.2 hydrogels. Control cells, which were cultured on the surfaces of unstretched and stretched hydrogels without cyclic RGDfK, exhibited no cell spreading (Figure S6). This proves that cell spreading on the cyclic RGDfK nanopatterned interfaces is attributable to the presence of adhesive ligands. Increased substrate stiffness has also been implicated in facilitating cell spreading.4 Therefore, we investigated the stiffness of unstretched and stretched hydrogels (Figure S7). The initial stiffness of the hydrogel was about 64 kPa. After stretching to 3.2 times the original length stiffness increased to 73 kPa. After 2 h continuous stretching the stiffness of the stretched hydrogel decreased to about 38 kPa, due to damage to the hydrogel’s dynamic hydrogen-bond crosslinking. However, since hydrogen bond damage is reversible, the stiffness of the hydrogel can return to its original value after 12 h of relaxation. It is therefore unlikely that changes in hydrogel stiffness should play a significant role in affecting cell adhesion on these hydrogels. The change of the material properties induced by stretching is very complicated and it is also very difficult to keep constant. To further confirm the crucial influence of interligand spacing on


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cell spreading area and polarization, we covalently immobilized a confluent layer of type I collagen onto the unstretched and L3.2 stretched hydrogels. As shown in Figure S8, cells are able to spread perfectly on both hydrogel surfaces without any preferred direction for polarization. This confirms that the decreased spreading area and the polarization of the cells on the stretched hydrogels can be ascribed to the increased interligand spacing in the stretched direction, rather than variations in the physical properties of the hydrogel substrates. In addition to unidirectional stretching, the ligand nanopatterned hydrogels can also be stretched in two dimensions, thereby maintaining homogenous interligand distances. For example, stretching hydrogels 2.5 times along two orthogonal directions produces a surface with all equal interligand distances of approx. 88 nm. In full agreement with our previous publication,29 cell spreading was inhibited on this surface (Figure S9).

Figure 2. (A) Phase contrast images of MC3T3 cells after 15 h in culture on cyclic RGDfK nanopatterned hydrogels with differing ligand spacings; the arrow above the images represents the direction of stretching. The scale bar is 50 µm. (B) Cell spreading areas of the cells adhering to the nanopatterned interfaces. Mean values and standard deviations from fifty randomly selected cells are presented. (C) Cell aspect ratio of the cells cultured on the nanopatterned interfaces. Mean values and standard deviations from fifty randomly selected cells are presented. Cell orientation angles for the cells adhering to L0 (D), L1.5 (E), L1.9 (F), or L3.2 (G) hydrogels. Mean values and standard deviations from two hundred randomly selected cells are presented. *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001 by ANOVA in prism. Note: unstretched hydrogels are referred to as L0, stretched hydrogels as Lx, where x represents the stretched:original length ratio.


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Additionally, fluorescent immunostaining was carried out to verify the different types of cell adhesion on the different surfaces. Intracellular actin, paxillin, and zyxin39 were fluorescently stained and imaged using high-magnification microscopy. As shown in Figure 3A, cells spread well on the surfaces of L0 and L1.5 hydrogels. Filamentous actin bundles and FA protein points (made visible by fluorescent paxillin, a focal adhesion-associated adaptor protein) were observed as expected. On L1.9 hydrogels cell spreading was decreased, yet filamentous actin bundles and FA points were still clearly visible. The decrease of the spreading area of these cells is caused by the large interligand spacing in the direction of stretching – which is nearly 70 nm. Nevertheless, adhesion could be accomplished, even though cells seeded on the surface of the L3.2 hydrogels exhibited a much elongated, almost spike-like shape and filamentous structures like actin and FA points were not visible. Figure 3B shows the quantification of the length, width and area of paxillin on the stretched hydrogel surfaces. Strong FA points were formed on L1.0 and L1.5 hydrogels, while the length, width, and area of FA points decreased a little on L1.9 hydrogels. When the hydrogels were stretched to 3.2 times the original length – i.e., the interligand spacing in the stretched direction reached about 120 nm – FA formation was strongly inhibited. Another FA protein, zyxin, was also fluorescently labeled to confirm FA formation on these hydrogels (Figure 3C and D). The results were highly similar to the results obtained from stained paxillin. Although the interligand spacing was less than 50 nm in the direction orthogonal to stretching, FAs could not be observed. This was attributed to a lack of effective integrin clustering, which inhibits the formation of stable FAs and actin fiber networks. Although the integrin in the cell membrane can be activated by cyclic RGDfK via specific anchoring junctions,32 subsequent integrin crosslinking is highly restricted simply because crosslinking proteins like α-actinin and


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talin only have a size of approx. 70 nm.39 As a control, the cells adhering to the confluent type I collagen immobilized L3.2 hydrogel surfaces were stained as well (Figure S8). Both filamentous actin bundles and FA points were observed. For comparison, homogenously adhesive hydrogels with a quasi-hexagonal nanoparticle pattern separated by interligand distances of 120 nm were tested as well. As expected, almost no filamentous actin bundles or FA points could be detected on adhering cells (Figure S10). Therefore, the decrease in the number of FAs on the stretched hydrogels can be directly attributed to an increase of interligand spacing in the stretched direction. We conclude that the crosslinking of integrin in one dimension is not sufficient for successful integrin clustering, in other words, the formation of stable FAs requires interligand spacings ≤ 70 nm in two dimensions.

Figure 3. (A) Fluorescent immunostaining of cells adhering to the surfaces of cyclic RGDfK nanopatterned L0, L1.5, L1.9, and L3.2 hydrogels; the arrow above the images represents the direction of stretching. All scale bars equal 50 µm. (B) Paxillin fluorescent immunostaining staining was used to determine the length, width, and area of the focal adhesion points of MC3T3 cells adhering to the hydrogels. Mean values and standard deviations from one hundred values are presented. (C) Fluorescent immunostaining of zyxin in the cells adhering to the hydrogels. All scale bars are 50µm. Mean values and standard deviations from one hundred values are presented. (D) The area of the focal adhesion points (based on stained zyxin) of the cells adhering to the hydrogels. *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001 by ANOVA in prism.


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The inhibition of FA formation, as seen on the L3.2 hydrogels, makes cell adhesion unstable and dynamic, which might endow cells with a higher mobility.40 This means that by using our highly stretchable hydrogels, we are able to regulate cell migration on the surface. MC3T3 and REF 52 (a rat fibroblast line) cells were selected as model cells to investigate cell migration on our adhesive ligand-nanopatterned hydrogel surfaces. First, MC3T3 cells were seeded to the surface of L0, L1.5, L1.9, and L3.2 hydrogels. During the first 3 h cells retained their round shape and neither morphological changes nor polarization were observed. After 6 h of culture cells had become polarized and cell mobility had increased. To study this development in greater detail, we used phase-contrast time-lapse imaging in 10 min intervals over a time period of 6 h after the cells had fully adhered to the surfaces. As shown in Figure 4A, cells on the L0 and L1.5 hydrogels had low polarization and spread well on the surfaces. They were also rather static and did not migrate at a large scale. In contrast, cells on the L3.2 hydrogels were highly polarized and migrated much more. Although the spreading area of the cells on the L1.9 hydrogels was smaller than that of the cells on the L0 and L1.5 hydrogels, cells adhering to the L1.9 hydrogels still formed stable adhesions and migrated only slightly. In the case of the highly polarized cells on the L3.2 hydrogels, unstable adhesion led to a much higher migration velocity and a much larger area of migration. Figure 4C shows the quantified migration velocity. There was no significant difference between the cells on the L0, L1.5, and L1.9 hydrogels, but cells on the L3.2 hydrogels migrated 2-3 times faster. Migration trajectories of the cells on the nanopattened hydrogel surfaces are shown in Figure 4B. Cells on the L3.2 hydrogel had a much larger migration zone than cells on the other hydrogels. In addition, cells preferred to migrate towards the direction in which they polarized, i.e., the orthogonal direction to the direction of stretching (Figure S11). REF 52 cells were also analyzed on the same hydrogels to show that the observed effects are not


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specific to MC3T3 cells. The results, which are quite similar to those obtained with MC3T3 cells, are shown in Figure S12 and Figure 4D and E. Because the mobility of the REF 52 cells was a little bit higher on the L3.2 hydrogels, we employed REF 52 in experiments on reversible cell migration through substrate stretching.

Figure 4. (A) Time-lapse phase-contrast images of MC3T3 cells cultured on the adhesive nanopatterned surfaces of L0, L1.5, L1.9, and L3.2 hydrogels; the arrow above the images represents the direction of stretching. (B) Migration trajectories of 10 representative MC3T3 cells cultured on the L0, L1.5, L1.9, and L3.2 hydrogels. (C) Quantified migration velocity of the MC3T3 cells on these hydrogels. Mean values and standard deviations from fifty values are presented. (D) Migration trajectories of 10 representative REF 52 cells cultured on the L0, L1.5, L1.9, and L3.2 hydrogels. (E) Quantified migration velocity of the REF 52 cells adhering to these hydrogels. Mean values and standard deviations from fifty values are presented. ****P < 0.0001 by ANOVA in prism.

Above we demonstrated that cellular adhesion is unstable and cell migration is stimulated on the surface of nanopatterned L3.2 hydrogels, whereas cell adhesion was stable and featured mature FAs on the surfaces of nanopatterned L1.9 hydrogels. In a next step we tested whether we can reversibly regulate cell migration by stretching and afterwards relaxing stretched hydrogels back to their previous size. For example, the L3.2 hydrogel can be relaxed to L1.9 within 30 min. As mentioned above, cells seeded on these hydrogels exhibited low mobility for several hours before beginning to polarize and migrate. Figure 5B shows the time-dependent migration


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velocity of REF 52 cells cultured on L1.9 hydrogels. The migration velocity was approximately 0.05 µm/min during the first 6 h and began to increase afterwards. After 8 h the migration velocity leveled off at about 0.08 µm/min. These findings led us to monitor reversibly controlled cell migration before the 6 h time point was reached (hitherto referred to as the pre-6h period) and after the 8 h time point was reached (hitherto referred to as the post-8h period).

Figure 5. (A) Time-lapse phase-contrast images of REF 52 cells cultured on the surface of a L1.9 hydrogel, which was continuously stretched to L3.2 over a time period of 2 h, and then was relaxed back to L1.9. The corresponding fluorescent immunostaining images are shown on the right. The arrow above the images represents the direction of stretching. (B) Time-dependent migration velocity of the cells cultured on the L1.9 hydrogels. (C) Quantified reversible migration velocity of the cells seeded on the hydrogels for 1 h prior to `stretching and relaxing´. Mean values and standard deviations from fifty values are presented. (D) Quantified reversible migration velocity of the cells seeded on the hydrogels for 10 h prior to `stretching and relaxing´. Mean values and standard deviations from fifty values are presented. ****P < 0.0001 by ANOVA in prism. To monitor the pre-6h period phase-contrast time-lapse imaging in 10 min intervals over 2 h was performed after the cells adhered to the L1.9 hydrogels for 1 h. Afterwards, the hydrogels were stretched further until a 3.2 stretched:original length ratio was achieved and the cells on the surfaces were monitored for another 2 h, followed by relaxing the hydrogels to L1.9 and phasecontrast time-lapse imaging. Cell morphology on all three hydrogel states – the L1.9 hydrogel, the hydrogel stretched to L3.2, and the hydrogel relaxed back to L1.9 length – exhibited a round shape


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(Figure S13). As shown in Figure 5C, a significant increase of the cell migration velocity was observed during hydrogel stretching from L1.9 to L3.2. The velocity decreased again after the hydrogels were relaxed back to L1.9 length. Figure 5A shows the reversible regulation of cell migration during the post-8h period. Cellular polarization was observed after 10 h culture on the L1.9 hydrogel surfaces. When the hydrogel was stretched to 3.2 times its original length (L3.2), cells on the surface became highly polarized and migrated more. Interligand spacings on the L3.2 hydrogel were approx. 112 nm in the direction of stretching and approx. 25 nm in the orthogonal direction. This was not sufficient to induce the formation of effective integrin clustering and mature FAs. As a result, cells dynamically and unstably adhered on the L3.2 hydrogels and showed a tendency to move. Interestingly, when the L3.2 hydrogels were relaxed back to L1.9 length, cells began to spread on the surface again. We quantified the cell migration velocity during this time period (shown in Figure 5D). The cell migration velocity significantly increased when hydrogels were stretched from L1.9 to L3.2. After the hydrogels were relaxed back to L1.9 length, the migration velocity decreased back to the same value as before stretching the hydrogel to L3.2. Fluorescent immunostaining was performed to investigate the formation of cellular FAs and filamentous actin bundles on the reversibly stretchable hydrogels (Figure 5A). Mature FA points and filamentous actin bundles were observed before stretching the L1.9 hydrogels and after relaxing back to L1.9 length. This suggests successful integrin clustering. In comparison, much fewer FA points and actin bundles were observed on the L3.2 hydrogels. Moreover, it is worth emphasizing that cell viability was not affected by the `stretching and relaxing´ process. In sum, our findings prove that our stretchable hydrogel system is highly suitable for regulating cellular adhesion and migration. As a control we also investigated cell migration on the surface of confluent type I


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collagen-coated hydrogels (see Figure S14). On collagen-coated hydrogels the migration velocity was not affected by stretching (L1.9 to L3.2) or re-relaxing (L3.2 to L1.9). This clearly proves that the change in migration velocity on the nanopatterned hydrogels was attributed to the change of the interligand spacing, rather than the physical properties of the hydrogel substrates. Additionally, this PNAGA-based hydrogel can endure multiple `stretching and relaxing´ cycles over tens of times without degeneration.35

CONCLUSIONS Taken together, these results demonstrate that cells are unable to stably adhere to a heterogeneously adhesive interface where the distance between neighbouring adhesion ligands is > 70 nm in one direction even when the interligand spacing between neighbouring ligands in the orthogonal direction is ≤ 70 nm. Cells adhering to this type of heterogeneously adhesive interface were highly polarized and preferred to orient in the direction of higher ligand density, i.e., the orthogonal direction to the direction of stretching. The inability to adhere strongly endowed the cells with a high mobility. Compared to the previously reported nanopatterned interfaces for cell capture, sorting, and isolation,33 we mainly focused on the molecular-scale interactions between cells and the adhesive interface. By using this stretchable hydrogel system, we can easily adjust the interligand spacing and obtain the heterogeneous interligand spacing, indicating that cells are unable to stably adhere to a heterogeneously adhesive interface where the distance between neighbouring adhesion ligands is > 70 nm in one direction even when the interligand spacing between neighbouring ligands in the orthogonal direction is ≤ 70 nm. Moreover, our ligand nanopatterned stretchable hydrogel system is highly suitable both as an


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efficient platform to investigate cell behaviors on heterogeneously adhesive interfaces and for application as a simple tool to reversibly regulate cell migration at an interface.

MATERIALS AND METHODS Materials. Polymers (PS(288)-b-P2VP(119) and PS(1056)-b-P2VP(671)) were purchased from Polymer Source Inc. (Canada) and chemicals were obtained from Sigma Aldrich (Germany), unless stated otherwise. Preparation of Nanostructured Substrates. Commercial glass slides (Carl Roth & Co. GmbH, Germany), size: 20 mm × 20 mm × 0.15 mm, were cleaned with a freshly prepared piranha solution (3 H2SO4 : H2O2) for 1 h, thoroughly rinsed with Milli-Q water (Milli-Q Millipore System, US) and then stored in Milli-Q water for future use. The substrates were dried with a N2 gun before coating them with the micellar solutions. Preparation of AuNP-decorated Glass Slides. Gold nanoarrays were prepared according to our previous report.37 Briefly, PS(288)-b-P2VP(119) was stirred overnight in dry o-xylene under atmospheric conditions. Then, the HAuCl3 × 3H2O precursor was added (Loading (L) = units (HAuCl4) / units (P2VP)) and the solution was stirred overnight again under the same conditions (L = 0.2) to obtain the Au-loaded micellar solution. After drying with the N2 gun, the glass slides were spin-coated with the micellar solution and plasma etched with W10 (hydrogen) in a Tepla PS210 microwave plasma system (PVA Tepla, Germany). The lateral distance between AuNPs was adjusted to 35 nm by varying the velocity of the spin coating process. PS(1056)-b-P2VP(671) (L


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= 0.3) was used for the nanoarray-decorated glass slides with a lateral distance of 120 nm between AuNPs. Synthesis of N-acryloyl Glycinamide (NAGA). NAGA monomer was synthesized according to a previous report.35 6.3 g of glycinamide hydrochloride, 6 mL of Milli-Q water, 33.6 mL of 2 mol L-1 cold potassium carbonate and 18 mL of diethyl ether were added into a 100 mL bottom rounded flask, which was then placed in an ice-water bath. Subsequently, a solution of 5.70 g of acryloyl chloride in 24 mL diethyl ether was added dropwise under stirring at 0 ºC over one hour. Then, the mixture was further stirred for another 4 hours at room temperature. Afterwards, the pH of the obtained mixture was adjusted to 2 using a 6 mol L-1 HCl solution. The mixture was then washed with 150 mL of diethyl ether for three times to remove the organic phase and the remaining diethyl ether was evaporated under vacuum. Next, a 2 mol L-1 NaOH solution was utilized to adjust the pH of the mixture to neutral, after which it was freeze dried. The raw product was washed with 150 mL of an ethanol/methanol mixture (4/1, V/V) for three times. Then, the ethanol and methanol were removed by rotary evaporation and the remaining mixture was recrystallized at 0 ºC, filtered, and dried under vacuum to obtain the purified NAGA monomer. The molecular structure of the NAGA is depicted in Figure S1. Nuclear Magnetic Resonance Spectrometry (NMR, 500MHz, Varian INOVA) and mass spectroscopy were used to confirm the successful synthesis of the NAGA monomer. 1H NMR (D2O): δ=3.7 (Hc, -NH-CH2-), 5.6 (Ha, CH2=CH-), and 6.1-6.3 (Hb, CH2=CH-) ppm (Figure S15); m/z = 128 (Figure S16). Preparation of AuNP-nanopatterned PNAGA Hydrogels. The fabrication of PNAGA hydrogels decorated with AuNPs was performed according to a protocol similar to our


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previously published method.36 Glass slides decorated with quasi-hexagonal gold nanoarrays were stacked in a 1 mM solution of N,N’-bis(acryloyl)cystamine (BAC) in ethanol at room temperature for 1 h. After rinsing and N2 drying, the substrates were incorporated in a mold with a depth of 0.8 mm. The polymerization precursor was prepared by dissolving 0.175 g of NAGA monomer, 5 µL of 0.2 M ammonium persulfate aqueous solution, and 1 µL of the accelerator (N,N,N',N'-tetramethylethylenediamine) in 500 µL of Milli-Q water. This solution was then used cover to the surface of the AuNP-nanopatterned glass slides. Polymerization was carried out under a UV light (365 nm, 8W) for 2 h. Finally, the hydrogels were incubated in water until they detached from the rigid substrates, and then stored in Milli-Q water for future use. Stiffness Measurement. The stiffnesses of the hydrogels was measured at room temperature using a rheometer (Kinexus, Malvern, US) with a preload of 0.18 N. Preparation of Cyclic RGDfK-nanopatterned PNAGA Hydrogels. AuNP-nanopatterned PNAGA hydrogels were immersed in an aqueous solution of 25 µM cyclic RGDfK with OEG spacer (PSL Peptide Specialty Laboratories GmbH, Figure S17). The reaction was carried out overnight at 4 ºC. Afterwards, the hydrogels were washed in Milli-Q water for 6 h with at least six water exchanges. Preparation of Type I Collagen-coated Hydrogels. The PNAGA hydrogels were first rinsed with 200 mM Hepes (Sigma, pH 8.5), and then blot dried. Afterward, 200 µL of 50 mM sulfosuccinimidyl 6 (4′-azido-2′-nitrophenyl-amino) hexanoate (SANPAH: Pierce) in 200 mM Hepes, pH 8.5, was pipetted onto the hydrogel surfaces. The hydrogels were then immediately exposed to the UV light of a sterile hood at a distance of 6 inch for 5 min. The sulfo-SANPAH solution was removed, and the hydrogels were rinsed with Hepes. The activation procedure was


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repeated a second time. After photoactivation, the hydrogels were thoroughly rinsed with 200 mM Hepes. A 0.1 mg/mL solution of type I collagen was then layered onto the surface of the hydrogels. The reaction was carried out overnight at 4 ºC. Afterwards, the hydrogels were rinsed with Milli-Q water to obtain the collagen-coated hydrogels.41 Cell Experiments on Nanostructured Substrates. MC3T3 osteoblasts and REF52 fibroblasts were cultured in Dulbecco’s modified Eagle’s medium (DMEM) with 10% fetal bovine serum (FBS), 1% penicillin/streptomycin, and 1% L-glutamine (all from Gibco, Thermo Fisher Scientific, US) and maintained at 37 ºC in a humidified atmosphere with 5% CO2. If not stated otherwise, washings and solutions were performed in phosphate buffered saline (PBS; Sigma Aldrich, Germany). Prior to seeding cells, the nanostructured hydrogels were first sterilized with UV light for 5 min and then washed twice with sterile PBS and once with medium. After being treated with 0.05 % trypsin-EDTA (Gibco, Thermo Fisher Scientific, US), centrifuged, and resuspended in cell culture medium, cells were seeded to the surfaces of the hydrogels and maintained under standard culture conditions. Immunofluorescent Staining. Samples were washed once with the cell culture medium and twice with PBS (pH 7.4), followed by fixing with 4% paraformaldehyde (Sigma, Germany) at room temperature for 15 minutes. Then, the samples were washed thrice with cold PBS. For permeabilization purposes cells were treated with 0.25% (v/v) Triton-X 100 (Sigma, Germany) in PBS for 10 minutes at room temperature, followed by washing with PBS to remove the detergent. The samples were then incubated with 1% (w/v) Bovine Serum Albumin (BSA) in PBST (0.1% v/v Triton-X 100 in PBS) at room temperature for 45 minutes to block non-specific


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antibody binding. After briefly washing with PBST, the samples were incubated with the primary antibody – rabbit monoclonal [clone Y113] to Paxillin (ab32084, Abcam, Germany) diluted at a 1:200 (v/v) ratio or anti-zyxin antibody [EPR4302] (ab109316) diluted at a 1:100 (v/v) ratio in PBST with 1% BSA –for 1 h at room temperature. Afterwards, the samples were washed twice with PBST and thrice with PBS and were then incubated in the dark for another 1 h in a solution of Alexa Fluor® 488 phalloidin (1:500 dilution, Life Technologies, Thermo Fisher Scientific, US). Next, they were incubated in the anti-rabbit secondary antibody tagged with the fluorescent dye Alexa Fluor® 568 (1:200 dilution, A11011, Invitrogen, Germany) in PBST with 1% BSA. Finally, the samples were washed twice with PBS and were imaged under an inverted fluorescence microscope. Scanning Electron Microscopy (SEM). A Zeiss Ultra 55 SEM (Carl Zeiss AG, Germany), equipped with an in-lens detector operated at 5 kV and 3 kV was used for glass slides and hydrogels, respectively. Nanopatterned glass slides were measured without any further treatment, except for the required coating of graphite (approx. 8 nm). For the nanopatterned hydrogels, we performed SEM under low-temperature conditions (cryo SEM; toperation = -120 to -140 ºC). Low acceleration voltages of 3 kV were applied because of the low conductivity of the investigated samples. A BAL-TECH VLC100 shuttle and loading system and a BAL-TECHMED020 preparation device were utilized to cool down and transfer the 15 nm graphite-layered hydrogel samples into the SEM chamber. Fluorescence Microscopy. The samples were measured with an Axiovert 200 M or an Imager.Z1 (Carl Zeiss AG, Germany). All acquired images were handled as 16 bit files and processed with the software ImageJ to adjust image settings and create the overlay fluorescence images.


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ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website Figure S1-S17 AUTHOR INFORMATION Corresponding Author *Email: [email protected] (Q. Wei) and [email protected] (J.P. Spatz) Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes The authors declare no competing financial interest. ACKNOWLEDGMENT We thank Dr. Stephan Rauschenbach at the Max Planck Institute for Solid State Research for help analyzing the mass spectrum of NAGA monomer. C. Z. acknowledges the support from the National Natural Science Foundation of China (No. 51673125), and the State Key Research Development Programme of China (2016YFC1103000). The work was financially supported by the Max Planck Society. J.P.S. is the Weston Visiting Professor at the Weizmann Institute of Science and is a member of the Heidelberg cluster of excellence CellNetworks. REFERENCES


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Figure 1 85x51mm (300 x 300 DPI)


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Figure 2 177x130mm (300 x 300 DPI)


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Figure 3 177x150mm (300 x 300 DPI)


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Figure 5 177x128mm (300 x 300 DPI)


Nanopatterned Adhesive, Stretchable Hydrogel to Control Ligand

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