Solvent-Induced Nanotopographies of Single Microfibers Regulate

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Biological and Medical Applications of Materials and Interfaces

Solvent Induced Nanotopographies of Single Microfibers Regulate Cell Mechanotransduction Abdolrahman Omidinia Anarkoli, Rahul Rimal, Yashoda Chandorkar, David Gehlen, Jonas Rose, Khosrow Rahimi, Tamás Haraszti, and Laura De Laporte ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.8b17955 • Publication Date (Web): 29 Jan 2019 Downloaded from http://pubs.acs.org on February 4, 2019

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ACS Applied Materials & Interfaces

Solvent Induced Nanotopographies of Single Microfibers Regulate Cell Mechanotransduction Abdolrahman Omidinia-Anarkoli1, Rahul Rimal1, Yashoda Chandorkar1, David B. Gehlen1, Jonas C. Rose1, Khosrow Rahimi1, Tamás Haraszti1, Laura De Laporte1,2* 1- DWI-Leibniz Institute for Interactive Materials, Aachen, 52074, Germany. 2-ITMC-Institute of Technical and Macromolecular Chemistry, RWTH Aachen University, Aachen, 52074, Germany. Keywords: Dry spinning, Fibers, Solvent properties, Topography, YAP, Mechanotransduction Abstract: The extracellular matrix (ECM) is a dynamic 3-dimensional (3D) fibrous network, surrounding all cells in vivo. Fiber manufacturing techniques are employed to mimic the ECM but still lack the knowledge and methodology to produce single fibers approximating cell size with different surface topographies to study cell-material interactions. Using solvent assisted spinning (SAS), the potential to continuously produce single microscale fibers with unlimited length, precise diameter, and specific surface topographies demonstrated. By applying solvents with different solubility and volatility, fibers with smooth, grooved, and porous surface morphologies are produced. Due to their hierarchical structures, the porous fibers are the most hydrophobic, followed by the grooved and smooth fibers, respectively, while the fiber diameter is increased by increasing the polymer concentration or decreasing the collector rotational speed. Moreover, 1 ACS Paragon Plus Environment

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SAS offers the advantage to control the inter-fiber distance and angle to fabricate multilayered 3D constructs. This report shows for the first time that the micro- and nano-scale topographies of single fibers mechanically regulate cell behavior. Fibroblasts, grown on fibers with grooved topographical features stretch and elongate more compared to smooth and porous fibers, while both porous and grooved fibers induce nuclear translocation of yes-associated protein (YAP). The presented technique, therefore, provides a unique platform to study the interaction between cells and single ECM-like fibers in a precise and reproducible manner, which is of great importance for new material developments in the field of tissue engineering. 1. INTRODUCTION: In their natural microenvironment, cells are surrounded by the extracellular matrix (ECM), which can be organized in complex random or aligned architectures, depending on the tissue type. Biochemical and mechanical signals direct cell-matrix interactions and adhesion, subsequently controlling cell signaling, migration, proliferation, and differentiation, and thus tissue morphogenesis.1-3 Physical cues from outside the cell are transmitted through cell-matrix contact and affect cellular and nuclear mechanics via several mechanotransduction pathways, which result in rapid remodeling of the cytoskeleton.4 As cells respond to their microenvironment at different micro- and nanoscopic length scales, it is of great importance to design and select an appropriate matrix that directs cell fate during cell culture and tissue regeneration. In spite of recent findings of the role of 3-dimensional (3D) matrix stiffness on mechanotransduction pathways, studies of the interplay between surface topography and cell mechanotransduction have been limited to biophysical cues present on 2-dimensional (2D) surfaces. Over the last decade, many top-down and bottom-up patterning techniques have been applied to fabricate structured substrates in order to mimic the native ECM and investigate the effect of various 2 ACS Paragon Plus Environment

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topographic features on cellular behavior.5 Common fabrication techniques include photolithography,6 electron-beam lithography,7 nanoimprint,8 polymer phase separation,9 and direct writing methods, such as laser writing10 and ink-jet printing.11 For instance, cells grown on pillars, fabricated by thermal nanoimprinting, adopt a less spread morphology in between the pillars and undergo stronger nuclear deformation when the feature size increases from nano (500 nm × 2 μm : diameter × height) to micro (2 μm × 10 μm) scale.12 On the other hand, multi-grooved surfaces with micro or nanoscale ridges have shown great potential in promoting the differentiation of pluripotent cells. Nano-grooved patterned arrays (350 nm ridges) stimulate human embryonic stem cells (hESCs) into neuronal lineage without any differentiation-inducing agents,13 while another study demonstrates that neuron differentiation and groove pitch are inversely proportional, with a ~ 1.7 fold increase in neuronal growth when seeded on a microgroove surface with 2 µm ridges compared to flat surface.14-15 Using mathematical algorithms, surfaces with several thousand random topographies, called TopoChips, have been developed to elucidate the interplay between cells and substrate topography.16 However, despite the recent advances and developments, conventional surface patterning techniques are mostly 2D and lack the 3D fibrillar structure of the native ECM.17 Micro- and nanoscale fibers have been produced with different fiber manufacturing techniques for various applications, such as textiles, electronics, or healthcare products.18-19 One of the most versatile and widely applied methods to fabricate nanofibrous mats is solution electrospinning (SES).20-21 To introduce nanoscale topography on the surface of electrospun fibers, different methods have been applied: i) employment of post-treatment, such as salt extraction and calcination,22 nano-imprint lithography,23 and selective removal of one component,24 or ii) variation of the solvent properties and relative humidity.25 While the first approach requires an 3 ACS Paragon Plus Environment


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additional, sometimes harsh, post-fabrication step, the second technique enables tuning surface topography during the electrospinning process in situ. For instance, increasing the ambient humidity results in porous electrospun fibers with higher surface roughness, leading to enhanced osteogenesis of human mesenchymal stromal cells (hMSCs), whereas a lower surface roughness mainly triggers chondrogenic differentiation.26 There are, however, limitations related to the high voltage present during the SES process, where fiber formation primarily occurs by axial tensile forces induced on the charged polymer jet in the direction of the flow. This results in an uncontrolled whipping motion and a tightly packed mesh with small pore sizes, which hinders studying cell-single fiber interactions as cells are in contact with multiple fibers at once in a 2D manner.27-28 To overcome this issue, alternative techniques produce 3D fibrous construct in a more controlled manner, such as solvent cast printing,29 melt extrusion additive manufacturing,30 microfluidic spinning,31 and melt electrospinning.32 However, these techniques produce fibers with diameters of tens to hundreds of micrometers and often lack the ability of changing the fiber surface topography. In one exception, the fiber morphology is varied by using patterned nozzles for extrusion.33 Nevertheless, the complex process of fabricating patterned nozzles and the resulting large fiber diameter (~ 0.6 – 1 mm) limit further application of this technique in studying single fiber-cell interactions. As an alternative, dry spinning based techniques have shown promising results in controlling the fiber diameter, alignment, spacing, and multilayer deposition.34-37 Compared to SES, no electrical field is applied and the produced fibers do not experience pulling and stretching forces due to the presence of surface charges. Fiber formation in dry spinning is caused only by the axial force applied by the rotating drum. This provides the ability of controlling the fiber diameter in the micrometer range, which is more similar to the size of a single cell.38 In addition,

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dry spinning offers accurate control over fiber deposition to create 3D constructs with sufficiently large pores in between the fibers for cell infiltration and a defined interconnected architecture, enabling studying cell behavior in 3D when in contact with single fibers. For example, different cell migration and elongation behavior is observed depending on the architecture of the fiber structures, i.e. inter-fiber distance and angle, for fibers produced using a spinneret-based tunable engineered parameters (STEP) technique.35, 39-40 While the dry spinning technique holds great potential towards the production of 3D fibrous constructs that mimic the native ECM, this technique still lacks knowledge and methodology to tailor the surface topography of the single fibers to investigate its effect on cellular behavior. Microscale topographical cues on the level of a single cell typically result in cell guidance and polarized cell morphology. Cells undergo linear actin polymerization parallel to an aligned pattern, such as oriented fibers,41 and nanoscale morphological features at the scale of single cell receptors enable control of cell responses by targeting receptor-driven pathways involving integrin clustering.3 Early studies have shown that the hierarchical combination of nano- and microscale topography of fibers has a superior effect compared to using each single scale separately.38 These fibers do not only provide a higher specific surface area but also specific integrin-size domains for cellular attachment, which will further affect the mechanobiology of these systems. In this study, we systematically investigate the effect of solvent properties on single fiber surface topography using a dry spinning-based technique, called solvent assisted spinning (SAS). We demonstrate the transformation between different topographies, ranging from smooth to porous to grooved, and levels of hydrophobicity by varying the composition of the binary solvent systems. Moreover, we highlight the ability and reliability of this technique to assemble multiple 5 ACS Paragon Plus Environment


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layers of these fibers with different dimensions, inter-fiber distance and angle, and different structures in a hierarchical manner. Ultimately, for the first time, the effect of single fiber surface topography on cellular behavior and mechanotransduction in an ECM-like fibrous system is reported. 2. EXPERIMENTAL SECTION: Experimental details can be found in the Supporting Information. 3. RESULTS AND DISCUSSION: 3.1. SES versus SAS Crucial differences between SES and SAS lead to different fiber topographies, depending on a wide range of solvent conditions and spinning parameters. Despite the numerous attempts on exploring the effect of solvent systems in SES, these processes have not yet been investigated for SAS. For both techniques, the solution hemisphere formed at the spinneret tip is subjected to constant solvent evaporation from its surface prior to jet ejection. In the case of SES, the charged hemisphere polymer solution forms a Taylor cone, from where the polymer jet travels with a high speed toward the collector. Three main forces are exerted on the polymeric jet during its trajectory toward the collector (Figure 1A): i) the Coulomb force (FC) due to the applied voltage, ii) the viscoelastic force (FV) of the polymer solution, and iii) the axial pulling force (FT) applied by the rotating drum. Depending on the applied voltage and the solvent properties, the average velocity of the jet can reach up to 15 m/s.42 Together with fast solvent evaporation, stretching of the polymeric jet due to the electrostatic field and the charged jet instabilities ultimately lead to the solidification of the final nanofibers.


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In contrast to SES, the initial polymer hemisphere in SAS is manually drawn toward the contact point of a low-speed rotating collector. As there is no Coulomb force, the polymeric jet mainly experiences two main forces: i) the viscoelastic force (F´V) of the polymer solution and ii) the axial pulling force (F´T) applied by the rotating drum. At the same polymer concentration, the viscoelastic forces in SAS and SES can be assumed to be similar (F´V = FV), while the axial force during SAS is significantly lower than for SES due to the absence of electric voltage and the low speed rotating drum, resulting in a reduced speed of the polymeric jet (F´T > F´T). Depending on the technique (SES and SAS), the same polymer solution (15 wt% PCL in CF-ACE (50-50)) results in fibers with either confined small pores (B) or elongated grooves (C), respectively. Scale bar: 5 µm


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After validation of our initial hypothesis, the spinning conditions (e.g. the applied solvents) are systematically varied to control the PCL fiber surface topography. The absence of a high voltage electrostatic field in SAS greatly expands the variety of solvents that can be applied to spin fibers, as in SES, the polymeric jet surface charge is affected by the solvent dielectric constant, which limits the applicable solvents.44 3.2. Controlling the fiber topography by varying the solvent system Solvents with different PCL solubility and volatility are tested (Table 1), with binary solvent systems being categorized in three main groups: i) good solvent/good solvent, ii) good solvent/partial solvent, and iii) good solvent/non-solvent. In the single and binary systems containing only good solvents (CF, DCM, and toluene (TOL)), PCL pellets dissolve well within an hour under the magnetic stirring at room temperature (RT). However, in the binary solvent systems comprising 75 v% dimethyl sulfoxide (DMSO) (non-solvent), PCL does not dissolve, even after 24 h of mixing at RT. Table 1: A) Different properties of the solvents used in this study. B) The solution viscosity of different binary solvent systems that are categorized based on their PCL solubility and volatility (boiling point).

Depending on the solvent system, two mechanisms are explored to change the surface topography of the fibers: thermally induced phase separation (TIPS) and non-solvent induced phase separation (NIPS). For both of these phase inversion processes, the same thermodynamic 9 ACS Paragon Plus Environment


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principles are involved: a thermodynamically stable solution precipitates due to an unsteady condition, such as rapid solvent evaporation and its associated cooling effect (TIPS) or nonsolvent phase separation (NIPS).45 As the solution passes the binodal curve, entering the metastable region, liquid-liquid demixing takes place and polymer- and solvent-rich phases appear. Initially, the effect of three single good solvents, DCM, CF, and TOL, with decreasing volatility, respectively, is investigated (Figure 2A). Both 100% CF or DCM result in fibers with a specific surface topography, while fibers produced from 100% TOL exhibit a smooth surface. Considering the high volatility of CF and DCM, compared to TOL (Table 1A), the pores on the fibers are likely formed due to fast solvent evaporation leading to TIPS. Yet, fibers produced with CF show very thin and elongated furrows compared to the wider and ellipsoid shaped pores on DCM fibers. Strikingly, previous studies using PCL in CF during SES have demonstrated that smooth fibers or fibers with circular pores are obtained depending on the spinning environmental condition, which may be explained by the fundamental differences between SAS and SES, as earlier explained, which results in different rates of phase separation.46-47 After SAS, the average pore length for fibers, made with 100% CF, is 5.10 ± 4.89 µm, which is ~ 1.5-fold higher than for 100% DCM fibers (3.33 ± 2.07 µm) (Figure 3A). The higher volatility of DCM may lead to the formation of larger initial pores, which are then stretched less as they solidify more quickly. This also explains the wider pores on the DCM fibers compared to CF fibers. No significant difference in pore density is observed between 100% CF or 100% DCM fibers due to a similar number of initial voids, formed on the jet.


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Figure 2: Transitions of surface topography after SAS using solvent systems with different PCL solubility and volatility: single solvents with good PCL solubility (A), binary solvent systems with different volume ratios of two good solvents with different volatilities (B) a good and partial solvent (C), or a good (CF or DCM) and non-solvent (DMSO) (D). Scale bar A: 5 µm, scale bars B-D: 10 µm.

3.2.1. Good Solvent/Good Solvent The good solvent CF is mixed with two other good solvents, DCM (volatile) or TOL (less volatile), at three different volume ratios: 75-25, 50-50, or 25-75. Figure 2B depicts the effect of 11 ACS Paragon Plus Environment


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solvent volume ratios on the topography transition of CF-DCM and CF-TOL fibers. In the case of CF-DCM fibers, rapid evaporation leads to fast cooling of the polymeric jet and consequently TIPS.25 During the course of fiber movement towards the collector, the polymer rich phase solidifies while voids from the solvent rich phase stretch to form the elongated pores. By increasing the DCM volume ratio, the volatility of the binary solvent increases, leading to smaller pore lengths and widths, ranging from 2.78 ± 2.58 µm length and 0.44 ± 0.22 µm width for 25% DCM to 1.06 ± 0.74 µm and 0.24 ± 0.15 µm, respectively, for 75% DCM (Figure 3A). The increase in volatility results in less time for pore elongation from their initial nucleation until final solidification. The significant increase in the number of pores on CF-DCM (25-75) could also be attributed to an increased TIPS effect. In contrast, the absence of sufficient TIPS in the case of CF-TOL leads to fibers with smooth surface topography at all mixing ratios. Moreover, the low viscosity of PCL solutions in a CF-TOL binary solvent (Table 1B) may also contribute to the smooth fiber topography, as it has been shown that at low polymer solution viscosity, the polymer-rich phase exhibits large resistance against the solution interfacial tension in the liquidliquid separation region. This results in less phase separation compared to highly viscous solutions, which may lead to a smooth surface after fiber solidification.45, 48 3.2.2. Good Solvent/Partial Solvent To study the effect on fiber topography when PCL is dissolved in less effective solvents, CF is mixed with acetone (ACE), acetic acid (AA), or DMF at three different volume ratios: 75-25, 5050, or 25-75. These binary solvents are selected based on their solubility (AA≥ACE>>DMF) and volatility (ACE>AA>DMF). For this library of solvent systems, only the combination of CF-AA leads to smooth fibers, while all other binary solvents result in fibers with a specific surface topography (Figure 2C). Similar to the CF-TOL system, the smooth surface topography of CF-


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AA fibers is likely the result of the solution’s low viscosity (Table 1B), accompanied with the high boiling point of AA (118 °C) and therefore insufficient phase separation. This is further confirmed by the uniform internal structure across the fiber cross-section (Figure 4). As TIPS is highly dependent on the boiling point of both solvents, and CF and ACE have similar low boiling points (61 ˚C and 56 ˚C, respectively), TIPS occurs at all ratios leading to a porous surface. While increasing the ACE volume ratio results in an increase in the pore density and decrease in pore length, the pore width does not seem to be affected (Figure 3B). The increase in pore density for higher percentages of ACE is likely due to a lesser solvent-polymer interaction as PCL is only partially soluble in ACE, enhancing the NIPS effect. The higher pore density decreases the sizes of the polymer rich domains, which may fasten solidification, resulting in the shorter pores. Mixing CF with the partial solvent DMF, which has a high boiling point (153 °C) and a lower PCL solubility compared to AA and AC, leads to a grooved structure for a volume ratio of 75-25 (Figure 2C). During the rapid evaporation of CF, voids are formed on the surface of the polymeric jet, while DMF keeps it wet to allow for void elongation before complete jet solidification. As Figure 4B demonstrates, the grooved topography of fibers is confirmed by a distinct sawtooth cross-section. In contrast to single solvents, the difference between boiling points in binary solvents is a driving force for sequential solvent evaporation that affects the fiber solidification kinetics and, therefore, the final surface topography. Due to the large difference in boiling points between the good (CF, bp= 61 °C) and the partial solvent (DMF, bp=153°C), TIPS is expected to be the dominant reason for grooved structures on CF-DMF fibers. Yet, NIPS likely also contributes to the formation of a grooved texture, as the poor solubility of PCL in DMF may be responsible for the small pores across the fiber cross-section in Figure 4B. Due to 13 ACS Paragon Plus Environment


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the high evaporation rate of the good solvent, the polymer concentration in the polymer-rich phase continuously increases, while the partial-good solvent ratio gradually increases, enhancing NIPS. For CF-DMF 50-50, buckling instability is observed, resulting in a wrinkled surface, which could be attributed to less rapid solvent evaporation, compared to 75-25 (Figure 2C). The slower removal of solvent from the core of the polymeric jet and the consequent outwards radial stress, arising from the continuous stretching of the jet, are the main reasons why the cylindrical polymer shell can buckle and form wrinkle type topographies.49 While increasing the DMF content does not significantly change the average groove width, the larger distribution in groove width in the case of CF-DMF 50-50 fibers is likely ascribed to the wrinkle formation (Figure 3B). As illustrated in Figure 2C, the polymer solution containing 25-75 CF-DMF no longer results in continuous and uniform fiber formation, which is likely due to the decreased solubility of the solvent, leading to frequent breaks in the polymeric jet during the spinning process.50


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Figure 3: Effect of different solvent systems on pore length, width, and density. Binary solvent systems with different volume ratios of two good solvents with different volatilities (A), a good and partial solvent (ACE or DMF) (B), or a good (CF or DCM) and non-solvent (DMSO) (C).

3.2.3. Good Solvent/Non-Solvent NIPS, which has been widely used to fabricate porous membranes,51 can be exploited as an approach to change fiber topography using binary solvent systems comprising good and nonsolvents. Depending on the solvent and non-solvent miscibility and the incompatibility between the non-solvent and the polymer, the solution can pass the binodal curve and enter the two-phase 15 ACS Paragon Plus Environment


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region with polymer-rich and solvent-rich phases. It is believed that a solution close to the binodal curve region favors the formation of spongy structures.51 Induction of NIPS by mixing a non-solvent, DMSO, with CF, results in fibers with a heterogeneous porous-grooved structure (Figure 2D). Increasing the amount of non-solvent to 50% enhances the NIPS effect, resulting in fibers with distinct porous topography. A further increase in DMSO content fails to dissolve PCL even after 24h mixing. While there is no significant difference between the pore width of CF-DMSO 75-25 and 50-50, the average pore length significantly decreases from 4.62 ± 6.49 µm to 2.22 ± 1.84 µm (Figure 3C). This may first seem unexpected as DMSO has a much higher boiling point than CF, which could allow for more pore stretching, but is likely due to the dominant effect of NIPS in the core of polymer jet leading to a slightly higher pore density, resulting in faster solidification of the polymer-rich domains. To study the possibility of forming interconnected pores, CF is replaced by another good solvent, DCM, which has an even lower boiling point (40 ˚C) compared to CF (61 ˚C). As DCM will rapidly evaporate, the content of DMSO (non-solvent with high boiling point) will persistently increase in the polymeric jet, leading to faster phase separation and freezing of elongated pores instead of creating a porous-grooved topography, observed with CF-DMSO 75-25 (Figure 2D). Interestingly, when comparing the pores for CF-DMSO 50-50 and DCM-DMSO 75-25, isolated pores with similar length, width, and density are observed, which highlights the collective role of NIPS and TIPS in surface topography formation (Figure 2D). While increasing the DMSO content from CF-DMSO 75-25 to 50-50 enhances NIPS, replacing CF by the more volatile DCM increases the effect of TIPS. A higher amount of DMSO in combination with DCM also leads to more pronounced NIPS and voids nucleation, resulting in a 3.5-fold increase in pore density, 16 ACS Paragon Plus Environment

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smaller pores, and an interconnective pore structure (Figure 3C, Figure 4C). An amount of 75 vol% DMSO fails to dissolve PCL.

Figure 4: Cross-section of fibers with three distinct surface morphologies: smooth (A), grooved (B), and porous (C), produced from CF-AA (50-50), CF-DMF (75-25), and DCM-DMSO (50-50), respectively. Scale bars:3 µm. The lower panel shows the magnified images. Scale bars:1µm.

Comparing the cross-sections of CF-DMF and DCM-DMSO fibers confirms the primary role of TIPS and NIPS in groove and pore formation. As demonstrated in Figure 4B, TIPS takes place at the surface of the polymeric jet due to CF evaporation, while some isolated pores are formed inside the core due to DMF-induced NIPS. In the case of a non-solvent, NIPS-induced void nucleation occurs more profoundly in the core. When this is combined with the volatile DCM solvent, which evaporates rapidly from the jet surface, a unique interconnected porous network is formed throughout the cross-section of DCM-DMSO 50-50 fibers. (Figure 4C).



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3.3. Fiber characterization The diameter of fibers has previously demonstrated to significantly influence cell adhesion andmigration.52 Therefore, different techniques are employed to alter the diameter of the smooth (CF-AA, 50-50), grooved (CF-DMF, 75-25), and porous (DCM-DMSO, 50-50) fibers. Initially, the collector speed is varied as a higher rotation speed is known to stretch the polymer chains in a larger extent, reducing the diameter of the collected fibers.35 As Figure 5A illustrates, increasing the collector rotation speed 3-fold results in a decrease in diameter, depending on the binary solvent system used. The highest reductions in fiber diameter are observed for the binary systems CF-AA and CF-DMF with 59% and 71%, respectively. Noticeably, at a constant rotation speed, the diameter of porous fibers is larger than smooth or groove fibers, which can be explained by the higher viscosity of the DCM-DMSO solution. The largest differences in fiber diameter between the different topographies are observed for the higher rotation speeds, with the porous fibers portraying a diameter of approximately 10 µm and the smooth or grooved fibers demonstrating diameters around 2 µm. While increasing the rotation speed does not have a significant effect on the diameter of the porous fibers, using a smaller spinneret size significantly reduces the fiber diameter from 11.57 ± 3.88 µm to 5.06 ± 0.83 µm without any visible morphological change (Figure S1 A, Supporting Information), consistent with previous reports.53 The smaller needle size creates a smaller initial polymer hemisphere, leading to a more narrow initial jet radius and eventually smaller diameter of the final fiber. As smooth and grooved fibers show smaller diameters than porous fibers, the possibility to enhance their diameter by increasing the PCL concentration from 15 wt% to 17 wt%, is investigated. This adaptation leads to an increase in diameter from 5.26 ± 0.98 µm to 9.37 ± 1.35 µm for smooth fibers, and 5.32 ± 0.72 to 9.65 ± 1.18 for grooved fibers, without any visible morphological change (Figure S1 B-C, Supporting Information). 18 ACS Paragon Plus Environment

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Figure 5: A) Effect of the collector’s rotation speed on fiber diameter. B) Water contact angle both parallel and perpendicular to the direction of the fibers for three fiber morphologies: smooth, grooved, and porous. In both directions, the fibers become more hydrophobic when changing from smooth to grooved to porous topographies. C) DSC curves for the first heating scans of the samples. D) WAXS curves of the PCL before and after SAS.



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The hydrophobicity of solid surfaces depends on their topographical features.54 In order to evaluate the effect of secondary morphologies on the wettability of fibers, water contact angle (WCA) measurement are conducted for smooth (CF-AA, 50-50), grooved (CF-DMF, 75-25), and porous (DCM-DMSO, 50-50) fibers, both perpendicular and parallel to the direction of the fibers. As Figure 5B demonstrates, for all fiber morphologies, the WCA, parallel to the fibers, is significantly lower than the WCA, perpendicular to the fibers. This could be attributed to the alignment of fibers, which eases the movement of water droplet in the direction of the fibers, when fibers are spaced closely together.55 Due to their hierarchical structures, the porous fibers are the most hydrophobic with the highest WCA in both directions, followed by the grooved and smooth fibers, respectively. Interestingly, the wettability of grooved and smooth fibers is similar parallel to the fiber orientation, while perpendicular to the fibers, grooved fibers are more hydrophobic compared to smooth fibers. This can be explained by the grooved structures in the direction of the alignment of fibers, which further facilitates wetting and water droplet sliding. To investigate the effect of the SAS process on fiber crystallinity, differential scanning calorimetry (DSC), wide-angle x-ray scattering (WAXS), and grazing incidence wide-angle scattering (GIWAXS) analysis are performed. As Figure 5C and Figure S1 D (Supporting Information) demonstrate, SAS does not change the melting temperature of the PCL significantly. For PCL pellets before SAS, the melting temperature (Tm) of the first heating scan is recorded as 60.38 ± 0.31 °C, while a slight decrease to 59.50 ± 0.28 °C, 59.58 ± 0.03 °C, and 59.33 ± 0.25 °C is observed in the case of smooth, grooved, and porous fibers, respectively. The melting enthalpy (ΔHm), calculated from the endothermic curve of the first heating scan, exhibits higher values for SAS fibers compared to unprocessed PCL pellets, which suggest an increase in crystallinity for the SAS fibers compared to unprocessed PCL pellets, without significant


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differences between the fiber topographies (Figure S1 D, Supporting Information).56 Interestingly, as the full width at half maximum (FWHM) of the first endothermic heating scan suggests, the SAS process results in a sharp crystalline-amorphous transition indicating a smaller crystallite size distribution compared to unprocessed PCL pellets (Figure S1 D, Supporting Information).57 WAXS is further employed to evaluate the degree of crystallinity before and after the SAS process and the results confirm the higher degrees of PCL crystallinity for SAS fibers compared to unprocessed PCL pellets. Both SAS fibers and unprocessed PCL pellets show the diffraction peaks of (110) and (200) at 21.3° and 23.7°, respectively, on a wide halo peak. Using the deconvoluted peaks, the degree of crystallinity for PCL pellets before SAS is measured as 57.13 %, while for smooth, grooved, and porous fibers, the crystallinity is calculated to be 60.21, 60.72, and 59.16 %, respectively. Strikingly, previous reports that evaluated the crystalline structure of electrospun fibers often demonstrated a decrease in crystallinity due to a rapid solvent evaporation during spinning.47,

56

In contrast, this report

demonstrates that the relatively lower rates of solvent evaporation during SAS, compared to SES, results in a higher degree of crystallinity, which may be explained by the longer time available for the polymer chains to rearrange into a crystalline structure. Nevertheless, varying the binary solvent system does not change the crystallinity of fibers significantly. To evaluate the orientation of the crystalline domains, GIWAXS is performed on highly aligned SAS fibers. The geometry-based measurement is performed at three different angles (0°, 45°, 90°) in respect to the fiber orientation and beam direction. Interestingly, the (200) diffraction plane (2ϴ = 23.7°) is always more pronounced (i.e. smaller FWHM) when the long axis of fibers is perpendicular to the beam direction, independent of the fiber surface topography (Figure S2 A-D, Supporting Information). This suggests that the (200) diffraction plane is preferably oriented normal to the



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fiber axis and that the c-axis of the crystal lattices formed by the molecular backbone is preferably oriented along the fiber axis (Figure S2 E, Supporting Information). This can be attributed to the axial tension induced by rotating the drum during solvent evaporation. A similar effect has been shown previously where electrospinning aligned fibers increases the stretching force on the fibers, inducing higher degrees of PCL molecular chain alignment along the fiber axis, acting as row nuclei for subsequent crystallization.56 These oriented electrospun nanofibers thus lead to higher degrees of crystallinity, compared to randomly oriented fibers, still lower but close to that of unprocessed PCL pellets. 3.4. Multilayered structured fibers Unlike SES, the SAS technique enables the formation of a single stable fiber jet, which can be collected in a precise manner due to the absence of a high electric voltage. Fibers with a uniform and adjustable diameter and desired surface topography can, therefore, be aligned or deposited at a defined angle, with variable spacing on a rotating collector. Figure 6 (A-D) depicts the effect of the collector’s rotation speed on the inter-fiber space, while keeping the rate of the transitional movement constant (1mm/min).

Figure 6: A-C) Aligned fibers with different inter-fiber distances can be produced using SAS. Scale bars: 200 µm. D) Effect of the collector’s rotation speed on the inter-fiber distance, while keeping the 22 ACS Paragon Plus Environment

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transitional speed of the rotating drum constant (1mm/min). E-H) Multilayer fibers at different angles by changing the transitional speed (E-F) or the collector angle (G-H). Scale bars E and G: 500 and 50 µm respectively. F and H) Higher magnifications of E and G, respectively. Scale bars: 10 and 20 µm, respectively.

Increasing the collector speed 5-fold decreases the inter-fiber distance more than 80%. Furthermore, the angle in between fibers can be varied by increasing the transitional speed of the collector. This allows for the fabrication of 3D porous structures, comprising fibers with unique topography. Figure 6 (E-F) demonstrates fibers with grooved topography (CF-DMF, 75-25), collected on a drum with 50 rpm rotational speed and 300 mm/min translational speed, result in an angle of ~ 30° between fibers. Alternatively, changing the collector angle after each layer of collection can result in a multi-layer fiber network array with aligned fibers rotated 90° in each layer (Figure 6 G-H). 3.5. Cell spreading and elongation on fibers Although the effect of different 2D topographies on cells has been investigated,38,

58

the

mechanisms by which cells sense topographical cues induced by fibrillar structures are poorly understood. Depending on the cell type and size, fiber diameter, topography, and chemistry can play an important role in cell behavior and morphology.59-60 Importantly, the majority of these types of studies are performed with electrospun fiber mats and do not enable analysis of single fiber-cell interaction. The SAS described here, provides a unique platform to control fiber topography and inter-fiber distance, and investigate single fiber-cell interactions in a more precise manner. L929 mouse fibroblasts are cultured on smooth (CF-AA, 50-50), grooved (CFDMF, 75-25) and porous (DCM-DMSO, 50-50) fibers with the same average diameter (~ 10 µm) to decouple the effect of fiber curvatures and surface topography.



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First, the cells are seeded on closely packed, aligned fibers to explore the effect of two types of hierarchical structures: the grooves in between two adjacent fibers and the topography of the fibers themselves (Figure 7A). As shown in Figure 7 (A-C), all fibers induce cell elongation parallel to the fibers and thus the material anisotropy, in spite of their surface topography type. This is likely due to the linear structure of the fibers, inducing cell contact guidance and preventing efficient cell protrusion perpendicular to the fiber alignment.41 Interestingly, the secondary structural features, created by the topography of the individual fibers, do have an effect on the nucleus and cytoskeleton aspect ratios, as demonstrated in Figure 7 (D-E). Fibers with a grooved topography lead to more elongated cytoskeleton and nuclei, compared to smooth or porous fibers, which can be explained by the secondary ridges present on grooved fibers.


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Figure 7: Effect of three different fiber morphologies on fibroblasts elongation: (A) smooth, (B) porous, and (C) grooved. Fiber orientation is indicated by arrows. Scale bars: 50 µm. D) Nucleus aspect ratio. E) 25 ACS Paragon Plus Environment


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Cytoskeleton aspect ratio. F) MTS cell proliferation assay of fibroblasts on smooth, grooved, and porous fibers. Values are normalized to smooth fibers.

A cell proliferation assay shows that all fibers support cell growth (Figure 7F), while porous fibers promote cell proliferation the most. This is in agreement with previous reports, showing enhanced proliferation of hMSC on porous SES fibers, compared to smooth fibers,59 and a ~ 2-fold increase in human vascular smooth muscle cells (vSMCs) density when increasing the pore size on SES PLLA fibers from ~ 300 nm to 700 nm.61 The enhancement of cell proliferation on porous fibers may be due to the increase in specific fiber surface area, roughness, or the higher hydrophobicity that can increase protein adsorption, favoring cell attachment.26, 62 To take a closer look at the behavior of single cells, depending on the secondary topography, single fiber-cell experiments are performed, after which the cells are immunohistochemically stained for F-actin to visualize protrusions, vinculin to resolve focal adhesions, and two different mechanosensitive reporter proteins, the Yes-associated protein (YAP) and the myocardin related transcription factor A (MRTFA).63 After staining, the cells are imaged with confocal microscopy for fibers with sufficiently large interspace so only one fiber interacts with the cells, and the effect of the surface topography of the smooth, grooved, and porous fibers can be analyzed independently from the primary grooves governed by two adjacent fibers. As Figure 8B demonstrates, cells on porous fibers display a high number of spike-like structures, called filopodia, compared to smooth or grooved fibers. Filopodia have a diameter in the range of 250–400 nm and play an exploratory role in probing the surrounding during cell migration and extension.64 As the micron-scale porous fibers present a high number of pores with a width approximating the filopodia diameter, the pores may provide numerous adhesion sites and anchorage points for filopodia formation and growth. On a flat surface, cell protrusions 26 ACS Paragon Plus Environment

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extend in all directions after attachment but the presence of topographical features can mechanically restrict the formation of microfilament bundles, resulting in rapid retraction or extension of protrusions in specific directions.64 In the case of grooved fibers, significant cell elongation is observed parallel with the ridges, while protrusions perpendicular to the ridges are inhibited. As focal adhesions could be as large as 1 µm,65 and the average ridge width of the grooved fibers is ~ 1.5 µm, it is plausible that the cell protrusions and their underlying focal adhesions are highly affected by the ridges, and promote cell extension along their orientation (Figure 8C). Further analysis of the cell’s focal adhesions shows vinculin expression mainly in the cytosol in the case of smooth fibers as cells exhibit a more rounded morphology and lack defined focal adhesions (Figure S3, Supporting Information). For grooved and porous fibers, vinculin expression is more pronounced in the periphery of the cells, where vinculin is accumulated in focal adhesions at the end of actin stress filaments localized in the pores and ridges. Some of the key factors in shaping cell behavior are external physical and mechanical cues. Using mechanotransduction pathways, the mechanical inputs form the surrounding microenvironment are interpreted by cells via the formation of nascent focal adhesions and the exertion of contractile forces in the cytoskeleton.66 Such interactions induce several signaling pathways, which regulate various cellular responses, like cell adhesion, migration, and differentiation. For example, by regulating the focal adhesion and F-actin reorganizations via anisotropic geometries (300 to 1500 nm groove width), human neural stem cells (hNSC) differentiation toward neurons and astrocytes is enhanced.67 To gain more insight into the role of fiber surface topography on cell mechanotransduction, YAP and MRTFA translocation are examined. Different parameters of the ECM, such as matrix stiffness and fiber dimensions, as



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well as cytoskeletal tension, are known to regulate YAP and alter its intracellular distribution.4, 68 YAP shuttles from the cytoplasm (soft elastic substrates) to the nucleus (stiff elastic substrates) and regulates the activation of several target genes, which control and direct cellular responses to mechanical stress.4 YAP localization and transcriptional activity depend on the degree of cell spreading and extension, as increased cell spreading induces nuclear flattening and enlargement of nuclear pores, promoting nuclear translocation of YAP.69 Generally, such nuclear flattening is observed on stiff or soft stress relaxing substrates,70 where YAP accumulation in the nucleus increases with the cell spreading area.69 The effect of fiber topography in this report reveals that YAP is predominant in the nucleus when cells are grown on porous or grooved fibers, in contrast to smooth fibers, where the average ratio of fluorescence intensity (Fnucleus/Fcytoplasm) is ~ 1.5 times lower (Figure 8E). When comparing the YAP nuclear translocation with observation seen depending on the 2D substrate stiffness, most previous studies defined the ~ 100 kPa regime as the substrate elastic modulus (E-modulus),above which cells exhibit high ratios of YAP shuttling from the cytoplasm to the nucleus.71 For example, cells grown on stiff elastic substrates of 120 and 150 kPa demonstrate YAP signal ratios of ~ 2.5 and ~ 3.5, respectively.69,

72

Based on

atomic force microscopy (AFM) measurements performed on the fibers presented here, their stiffness appears to be in the MPa range without any significant differences between surface topographies (Figure S1 E, Supporting Information). However, AFM measurements on single fibers are very challenging due to the curvature, mobility, and heterogeneity of the fibers73, which together with the micro- and nanoscale surface topography of fibers lead to a large distribution of the data. Based on the YAP translocation on 2D substrates and the stiffness of the fibers here, one would expect YAP to already be mostly located in the nucleus but interestingly, while smooth fibers lead to a significantly lower FNucleus/FCytoplasm ratio of ~ 1.91, the signals in


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the case of grooved and porous fibers increase to ~ 3.37 and ~ 2.91, respectively. This suggests that unlike stiff 2D substrates (e.g. TCPS), where only stiffness matters, YAP shuttling can be influenced by the topography of substrates in case of 2.5D single microfibers. As cells undergo a forced apical/basal polarity on 2D substrates but wrap around the fibers in a more 3D manner, the internal stresses of the F-actin filaments will change, which may lead to alterations in the mechanotransduction pathways. The topography-dependent difference in YAP nuclear localization might be explained by the fact that structural features on fibers cause stronger cell-fiber interactions, resulting in a greater cell spreading, which is visualized by the actin staining and the larger aspect ratio of the nucleus in the case of grooved fibers (Figure 7D-E, Figure 8G). Intriguingly, porous fibers, leading to enhanced YAP shuttling in the nucleus, do not result in larger cell areas but the spike-like filopodia formation and the more pronounced peripheral vinculin most likely cause additional internal tension inside the cells. This is in agreement with a previous study where an increase in unidirectional tensional stress is introduced as the reason for nucleus elongation, cell alignment, and a more contractile phenotype when vSMCs are cultured on an oriented electrospun fibrous mat with porous fiber surface topography.61 While another recent study indicates an influence of fiber diameter on cell mechanosensing, demonstrating higher YAP to nucleus shuttling for increasing diameter74, this is, to the best of our knowledge, the first report studying how single fiber topography affects cellular mechanotransduction. The YAP shuttling observed in this report is also in agreement with previous research using a high-resolution soft lithography technique to create patterns. Grooved surfaces with 15 nm ridges resulted in cytoplasmic YAP in the case of human neural stem cells, while 1 µm ridges triggered YAP nuclear translocation due to an increase in focal adhesion formation.75 Similarly, when cells grow on two adjacent smooth fibers



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with diameters of approximately 10 µm, resembling grooved topographical features, YAP nuclear shuttling is observed (FNucleus/FCytoplasm ~ 2.65), similar to cells grown on single porous fibers (FNucleus/FCytoplasm ~ 2.91), but larger than single smooth fibers (FNucleus/FCytoplasm ~ 1.91) (Figure 8D-F). This may be explained by the higher cell spreading area and elongation in the case of adjacent fibers. Another important mechanosensitive protein is MRTFA, which binds G-actin in the cytoplasm, and in response to force exertion on integrins, releases G-actin to prepare F-actin and shuttles to the nucleus as a direct sensor of the G-actin concentration.63 Quantification of the fluorescence intensity shows that MRTFA, which is 3 times larger in molecular weight than YAP, is localized in the cytoplasm, irrespective of the fibers topography (Figure 8F). This suggests two possibilities: (i) MRTFA, after releasing G-actin is unable to pass the nuclear pores into the nucleus or (ii) MRTFA is still bound to G-actin. Since we do not observe accumulation of MRTFA in the vicinity of the nucleus (Figure 8), it is plausible that MRTFA is bound to G-actin asno active forces are exerted on the cells. Notably, MRTFA translocation is also not affected by the adjacent smooth fibers and remains distributed in the cytoplasm. Since nuclear translocation of YAP is affected by the fiber topography, while the intracellular distribution of MRTFA is unaffected, this is most likely attributed to differential cell spreading on different fiber topographies and not due to other mechanical forces on cells. As the composition, stiffness, and crystallinity of fibers are similar between the surface topographies, cells seem to sense nanotopographies and spread accordingly, independent of the substrate stiffness. The observations, presented here, of cellular YAP shuttling on single, micron-scale fibers encourage future investigations to better understand the role of ECM like structures and fiber surface topography and dimensions (curvature) in directing cell mechanotransduction.


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Figure 8: Effect of different fiber topographies on fibroblast elongation and mechanotransduction: A) smooth, B) porous, C) grooved, and D) two adjacent smooth fibers. E-F) YAP and MRTFA translocation depending on fiber topography. G) Schematic illustration of nuclear flattening and enlargement of nuclear pores depending on fiber topography. Scale bars: 10 µm.

4. CONCLUSION: This study demonstrates a simple and one step technique to continuously control a single fiber surface topography in situ. By altering solvent properties, microfibers with smooth, grooved, and 31 ACS Paragon Plus Environment


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porous surface topographies can be achieved. Controlling the collector rotational speed, needle size, or polymer concentration allow for the formation of fibers with variable diameter. Furthermore, SAS enables the fabrication of multilayered fibrous structures with a distinct surface topography, precise inter-fiber distances, and variable angles. This provides a unique platform to study the synergistic effect of micro- and nanotopography on cellular behavior or single fiber-cell interactions. For the first time, the effect of single fiber topography on YAP nuclear translocation is observed, corroborating with our hypothesis that fiber topography influences mechanotransduction. We postulate that this mechanism is regulated by controlling the cell spreading area, with porous and grooved fibers inducing YAP shuttling from the cytoplasm to the nucleus. These results suggest that, unlike other fiber fabrication methods, SAS can be employed to study the effect of single fiber morphology on cell behavior at different hierarchical scales in a reliable and reproducible manner. This is of great importance when designing materials for different tissue engineering applications. Supporting Information The Supporting Information is available free of charge on the ACS Publications website. The experimental section. Figure S1. Figure S2. Figure S3. Corresponding Author *E-mail: [email protected]


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Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Acknowledgment We thank Dr. Vinokur Rostislav and Claudia Pörschke for experimental and data analysis assistance. This work was supported by the funding from the Max-Buchner Forschungsstiftung of Dechema and the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation program (ANISOGEL, grant agreement No 637853). This work was performed in part at the Center for Chemical Polymer Technology CPT, which was supported by the EU and the federal state of North Rhine-Westphalia (grant EFRE 30 00 883 02). We are grateful to Prof. Martin Möller for providing pivotal equipment to perform high quality experiments, such as the confocal microscope.



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