A Self-Powered Piezo-Bioelectric Device Regulates Tendon Repair-Associated Signaling Pathways through Modulation of Mechanosensitive Ion Channels

www.advmat.de 2008788 (1 of 18) © 2021 The Authors. Advanced Materials published by Wiley-VCH GmbH ReseaRch aRticle A Self-Powered Piezo-Bioelectric Device Regulates Tendon Repair-Associated Signaling Pathways through Modulation of Mechanosensitive Ion Channels Marc A. Fernandez-Yague,* Alexandre Trotier, Secil Demir, Sunny Akogwu Abbah, Aitor Larrañaga, Arun Thirumaran, Aimee Stapleton, Syed A. M. Tofail, Matteo Palma, Michelle Kilcoyne, Abhay Pandit, and Manus J. Biggs* Dr. M. A. Fernandez-Yague, A. Trotier, S. Demir, Dr. S. A. Abbah, Dr. A. Larrañaga, Dr. A. Thirumaran, Prof. M. Kilcoyne, Prof. A. Pandit, Prof. M. J. Biggs CÚRAM SFI Research Centre for Medical Devices National University of Ireland Galway H91W2TY, Ireland E-mail: [email protected]; [email protected] DOI: 10.1002/adma.202008788 place a considerable burden on healthcare systems (> $2 billion annually, and post-surgery complications result in nearly 1 million additional days of inpatient care each year).[1–3] Surgical intervention via direct end-to-end repair using sutures and biological or synthetic grafts represents the gold standard in treatment, and despite the relative success, these repairs frequently fail to restore full tendon functionality. Following injury, disorganized tissue deposition leads to scar tissue formation, proteoglycan accumulation, and calcification, resulting in poor biomechanical properties and impaired function that triggers chronic inflammatory signaling pathways and progresses into tendinopathy. Hence, to achieve long-term repair, innovative functional solutions that focus on the activation of endogenous tissuerepair signaling pathways represents a paradigm shift in the field of biomedical devices and regenerative medicine (RM).[4] Many studies confirm that resident tendon cell populations are highly mechanosensitive and are responsible for orchestrating the repair processes after injury through specialized sensory machinery, including mechanosensitive ion channels.[5–8] Critically, mechanotherapy (i.e., low-level exercise or extracorporeal shock waves) has been reported to promote tendon Tendon disease constitutes an unmet clinical need and remains a critical challenge in the field of orthopaedic surgery. Innovative solutions are required to overcome the limitations of current tendon grafting approaches, and bioelectronic therapies show promise in treating musculoskeletal diseases, accelerating functional recovery through the activation of tissue regenerationspecific signaling pathways. Self-powered bioelectronic devices, particularly piezoelectric materials, represent a paradigm shift in biomedicine, negating the need for battery or external powering and complementing existing mechanotherapy to accelerate the repair processes. Here, the dynamic response of tendon cells to a piezoelectric collagen-analogue scaffold comprised of aligned nanoscale fibers made of the ferroelectric material poly(vinylidene fluoride-co-trifluoroethylene) is shown. It is demonstrated that motionpowered electromechanical stimulation of tendon tissue through piezobioelectric device results in ion channel modulation in vitro and regulates specific tissue regeneration signaling pathways. Finally, the potential of the piezo-bioelectronic device in modulating the progression of tendinopathyassociated processes in vivo, using a rat Achilles acute injury model is shown. This study indicates that electromechanical stimulation regulates mechanosensitive ion channel sensitivity and promotes tendon-specific over non-tenogenic tissue repair processes. The ORCID identification number(s) for the author(s) of this article can be found under https://doi.org/10.1002/adma.202008788. © 2021 The Authors. Advanced Materials published by Wiley-VCH GmbH. This is an open access article under the terms of the Creative Commons Attribution License, which permits use, distribution and reproduction in any medium, provided the original work is properly cited. Dr. A. Larrañaga University of the Basque Country Department of Mining-Metallurgy Engineering and Materials Science and POLYMAT Barrio Sarriena Bilbao 48013, Spain Dr. A. Stapleton, Prof. S. A. M. Tofail University of Limerick Department of Physics Limerick V94 T9PX, Ireland Prof. M. Palma Queen Mary University of London Materials Research Institute and School of Biological and Chemical Sciences Mile End Road, London E1 4NS, UK 1. Introduction Severe tendon injuries resulting from athletic or repetitive activity affect more than 102.5 million adults every year and Adv. Mater. 2021, 33, 2008788 www.advmat.dewww.advancedsciencenews.com 2008788 (2 of 18) © 2021 The Authors. Advanced Materials published by Wiley-VCH GmbH regeneration and provide a reliable route for appropriate postoperative management.[9] Additionally, piezoelectricity-derived electric fields produced during physiological locomotion may provide additional bioelectric signaling cues to activate tendonspecific regenerative pathways (Figure1A). Several recent studies have revealed the significant potential of electrical fields in mediating cell migration, promoting collagen synthesis, and inducing successful wound healing by externally applied direct current electrical stimulation (ES).[10,11] In the last decade, DC stimulators (i.e., Zimmer Biomet SpF and OsteoGen) have improved the success rate of spinal fusion procedures and nonhealing tissue injuries.[12] Despite the proven clinical effect, a significant roadblock to wide-scale clinical adoption of ES is the infection risk to penetrating electrodes, potential nonspecific off-target effects, and the cumbersome design of ES units, making patient compliance a concern.[13] The need for more safe, efficient, and less invasive electrically stimulating systems drives the development of novel bioelectric strategies that control somatic cell functions and enhance specific tissue regenerative processes.[14] The discovery of piezoelectricity in bone and more recently in its constituent collagen type I, has spurred new research into the role of bioelectricity in tissue regeneration and the development of self-powered electrical stimulation technologies.[15,16] A promising strategy for developing biomimetic electromechanical stimulation (EMS) devices has been enabled by synthesizing compliant ferroelectric polymers (i.e., PVDFTrFE), conformed into a nanofibrous scaffold that mimics the intrinsic electrical, mechanical, and morphological properties of collagen type I (Figure 1B). Mechanical actuation of these structures or piezo-bioelectric devices through repetitive physiological loading and unloading (i.e., monotonic stretching) can produce biologically relevant electrical fields, enabling studies into the role of “mechanically-induced” electrical cues on musculoskeletal tissue function. Significant efforts from the biomedical community have demonstrated the potential of Adv. Mater. 2021, 33, 2008788 Figure 1. Overview of the tendon electromechanical environment and the piezoelectricity origin analogy of collagen and PVDF-TrFE fibers. Tendon is a dynamic tissue that connects muscle to bone and is under continuous mechanical loading and unloading. A) This mechanical stress is borne by a highly anisotropic extracellular matrix composed of collagen type I, a high-tensile, piezoelectric material which undergoes electrical polarization in response to mechanical loading. B) Schematic representation of the crystal structure of the collagen triple helix consisting of three polypeptides chains stabilized via hydrogen bonding. A-A: Cross-sectional schematic of an individual alpha-chain showing the triple Gly (glycine)-X (proline)-Y (hydroxyproline) residuesi (i). Polar bonds between carbonyl (CO groups) and amine (peptide NH group of glycine residues) groups along the backbone of collagen result in a dipole moment or electrical polarization (ii). Cross-section of multiple individual collagen molecules resulting in a non-centrosymmetric hexagonal fibril arrangement (iii). B-B: A cross-section of the collagen triple-helix core; here, the three helical chains and glycine residues are observable (iv). Representation of an individual electric dipole (v). C) Representation of the PVDF-TrFE all-trans (TTTT) zigzag planar configuration top view (vi). Crosssection of the crystal and along the orthogonal axis of the all-trans (TTTT) zigzag planar configuration (vii). Representation of an individual electric dipole composed of fluorine (green sphere), carbon (blue sphere), and hydrogen (orange sphere) atoms (viii). PVDF-TrFE has a non-centrosymmetric structure. Dipoles are generated by the highly electronegative difference between hydrogen and fluorine atoms. Representation of the molecular chain structure of the electroactive β-phase of PVDF-TrFE showing a resultant dipole moment (ix). 15214095, 2021, 40, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/adma.202008788 by Johns Hopkins University, Wiley Online Library on [28/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License www.advmat.dewww.advancedsciencenews.com 2008788 (3 of 18) © 2021 The Authors. Advanced Materials published by Wiley-VCH GmbH piezoelectric materials to transduce mechanical into physiologically relevant electrical cues (direct piezoelectricity) and affect different cell types, including osteoblasts, mesenchymal stem cells, and neurons.[17–20] In this study, we hypothesized that a ferroelectric scaffold could serve as mechanical support for the regeneration of damaged tendon tissues and mimic the bioelectrical cues usually provided by collagen’s piezoelectricity to maintain tendon-cell phenotype and promote tendon regeneration (Figure1B,C). 2. Design and Fabrication of a Piezo-Bioelectric Device The physical properties of a scaffold, including its mechanical behavior, microstructure (fiber diameter and alignment), and crystallinity-dependent piezoelectric response, play a pivotal role in regulating the cellular events involved in tissue regeneration.[21] To develop a collagen piezoelectrical-analog device that can recapitulate the fibrous structure of extracellular tendon tissue an electrospinning process was implemented, and nanoscale fibers with varying diameter and alignment were produced through optimization of solution and processing parameters (Figure2). First, we analyzed the impact of fiber diameter on the biophysical properties of the scaffolds. By varying the concentration of the polymeric solution (1.1, 1.2, 1.3, 1.4, 1.5, 1.6, and 1.7 mg mL−1 in dimethylacetamide (DMAc) we could obtain fibers with distinct diameters. We used a 30 kV potential difference between collector and nozzle tip (Ø = 200µm), a flow rate of 1mL h−1, and a tip-to-collector distance of 6cm (shortdistance electrospinning). Importantly, a collector disk with an 8mm width was used to concentrate the electrical field while rotating at 29.3 m s−1 (linear speed) to draw and align the collected fibers. The resulting fibers ranged from 180 to 540nm in diameter (Figure 2A; Figure S1, Supporting Information), which is similar to the diameter of collagen fibers in tendon tissue 50–500nm.[22] It was observed that the fiber diameter governed the stiffness of the resulting scaffolds and constructs with lower fiber diameters possessed an increased Young’s modulus, as determined by tensile-test analysis (Figure2A). The observed enhancement of the mechanical properties of scaffolds with decreasing fiber diameter was a result of a high level of chain extension and orientation along the fiber axis as a consequence of fibers being drawn by centripetal forces during fiber collection. With semi-crystalline polymers (including poly(vinylidene fluoride-co-trifluoroethylene) P(VDF-TrFE)), it has been demonstrated that stress hardening does not depend on polymer crystallinity but rather, on the density of amorphous polymer chain entanglement, which in respect to electrospinning, is related to the distance between the needle and collector, the collector rotational speed and the polymer solution concentration.[23] As a result, scaffolds with a fiber diameter between 390 and 540 nm (Figure S1>D,E, Supporting Information) demonstrated increased fiber alignment. Finally, nanofibers with diameters <240 nm (Figure S1F, Supporting Adv. Mater. 2021, 33, 2008788 Figure 2. Cold drawing improves fiber alignment and enhances electroactive β-phase formation. A) Analysis of the relationship between fiber diameter and the relative Young’s modulus (Erel) indicated that the mechanical behavior of highly aligned scaffolds was inversely proportional to fiber diameter. B) The morphology of scaffolds obtained by electrospinning collected at low (4.2 m s−1 rpm) and high (29.3 m s−1) linear speeds. Fast fourier transform (FFT) spectra showed a broad distribution of intensities for low speed (4.2 m s−1) and a clear peak for high speed (29.3 m s−1) characteristic of highly anisotropic structures. Fiber alignment was significantly increased, and the morphology mimicked that of tendon collagen. C) Total crystallinity and β-phase content (calculated from FTIR and XRD spectra) increased as a function of collector distance and following cold drawing. D) Creep/stress relaxation was minimized after cold drawing and resulted in reduced fiber diameter and increased fiber alignment. SEM images of scaffolds with and without cold drawing. 15214095, 2021, 40, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/adma.202008788 by Johns Hopkins University, Wiley Online Library on [28/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License www.advmat.dewww.advancedsciencenews.com 2008788 (4 of 18) © 2021 The Authors. Advanced Materials published by Wiley-VCH GmbH Information) were prone to alignment disorganization due to the air currents generated during collection. To decouple the effect of piezoelectric stimulation from mechanical loading on the cellular response, a non-piezoelectric PTFE scaffold which maintained the topography (i.e., fiber diameter and organization) and was chemically analogous (i.e., a fluorinated polymer) to PVDF-TrFE was also fabricated. PTFE is non-piezoelectric due to the presence of a centrosymmetric structure with strong polar CF covalent bonds (C∂+—F∂−). Conversely, PVDF-TrFE presents non-centrosymmetric electrical dipoles, generated by the high electronegative difference between hydrogen and fluorine atoms (Figure S2, Supporting Information). The direct electrospinning of PTFE fibers, however, was not possible due to the unique chemical properties of PTFE. Therefore, a two-step short-distance electrospinning process was optimized to obtain scaffolds of PTFE fibers.[24] The tendon microstructure is highly hierarchical, with collagen fibers aligning to the direction of mechanical loading. To reproduce this critical feature in both PVDF-TrFE and control PTFE scaffolds the collector velocity was adjusted between 4.2–29.3 m s−1, and the concentration of the polymeric solution fixed at 1.7 mg mL−1. As evident in Figure 2B, the scaffolds obtained at a collector linear velocity of 29.3 m s−1 (≈4000rpm) displayed a highly organized fiber morphology, while a broad distribution of fiber orientation was observed for scaffolds formulated with a collector linear velocity of 4.2 m s−1 (500rpm). Additionally, as determined by X-ray diffractometry (Figure S3, Supporting Information and Table S1), the degree of crystallinity increased from 40% in randomly aligned PVDF-TrFE fibers to 49% in aligned fibers (Figure 2C), suggesting that mechanical drawing during fiber collection at high rotational speeds resulted in PVDF-TrFE fibers with a reduced diameter (590 ± 130nmfor 4.2 m s−1 and 540 ± 120nmfor 29.3 m s−1) and enhanced crystallinity (Table S1, Supporting Information). Our short-distance electrospinning configuration contrasted to previously reported electrospinning configurations for PVDFTrFE (i.e., long-distance electrospinning).[25] We have demonstrated that scaffolds produced using a short-distance configuration possessed increased fiber organization (Figure S1, Supporting Information) and crystallinity (Figure2C), supporting the hypothesis that fibers are significantly drawn when subjected to an electric-field increased intensity. The mechanical behavior of the scaffolds obtained by the short-distance configuration was improved compared to scaffolds obtained by a conventional (long-distance) configuration in terms of elastic modulus, creep resistance, and yield strength (Figure S2, Supporting Information), which is of vital importance for the device to retain the integrity of its macro and microstructure under loading following implantation.[26] Regarding the piezoelectric response, a growing body of research suggests that the piezoelectric performance of PVDF (i.e., the base comonomer of the PVDF-TrFE copolymer) is affected by the degree of crystallinity and in particular the β-phase content (Table S2, Supporting Information).[27] As determined by Fourier-transform infrared spectroscopy (FTIR) (Table S2, Supporting Information), the short-distance configuration promoted the crystallization of the electroactive β-phase (Figure2C). Overall, these observations reinforce the argument for the potential use of short-distance electrospinning in the fabrication of PVDF-TrFE-based scaffolds. To further increase the mechanical properties and β-phase content of the scaffolds, while avoiding the well-reported crimping of the fibers under repetitive mechanical loading, two post-synthesis processes were investigated: i) strain-hardening (cold-drawing) and ii) thermal annealing (cold crystallization). Based on the initial properties of the scaffolds, we explored cold drawing at ≈12% to induce strain-hardening through plastic deformation of the structures. Overall, this process resulted in ≈8% plastic deformation, alignment of fibers (from 65% to 98%) and reduction in individual fiber diameter (from 540 to 516 nm). Importantly, fiber diameter reduction and better fiber alignment resulted in scaffolds with an improved elastic modulus (61.8± 5.1MPa, p< 0.05) and strength (31± 4.2 Mpa, p< 0.001) (Table1). We conducted extensive SEM inspections along the entire obtained scaffolds (0.8 × 30cm)to check the fiber direction. We observed that fibers were aligned with the collector rotating direction and parallel to the longitudinal axis of the scaffold. Only a few fibers far from the middle part of the scaffold presented a skewed orientation (<7± 6°) and were corrected before implantation of the devices during the subsequent cold drawing process.Concurrently, the stress relaxation/creep of the fibers was significantly reduced (Figure 2D). Thermalannealing around the Tc (90 °C for 1 h) was used to further increase the β-phase content. The amount of β-phase content in PDVF-TrFE samples subjected to post-synthesis cold-drawing and thermal annealing was higher than the non-post-treated counterparts as determined by FTIR analysis (Figure2C). The effect of alignment on the piezoelectric performance of individual fibers was demonstrated by switch-spectroscopy piezoresponse microscopy (SS-PFM). The piezoelectric response of fibers from randomly-aligned scaffolds was compared to fibers from mechanically drawn, aligned scaffolds. The piezoresponse or d33 value was found to be dependent on the fiber alignment. A strong correlation was observed between scaffolds with increased fiber alignment and a higher piezoelectric response (from −16.92 to −24.61 pm V−1) (Figure3). The elastic modulus of individual fibers was 350± 80MPa as measured using peak-force imaging, yet no significant differences were observed between random and aligned fibers. Interestingly, a cooperative piezoelectric effect previously described by Persano et al. was found in dense arrays of fibers (Figure 3A) due to electromechanical interactions between adjacent fibers and the Adv. Mater. 2021, 33, 2008788 Table 1. The physical properties of piezoelectric PVDF-TrFE and non-piezoelectric PTFE scaffolds. Scaffold type Elastic modulus [MPa] Strength [MPa] Elongation [%] Fiber diameter [nm] d33 [pC N−1] Non-piezoelectric with drawing 14.5± 1.7 16± 0.3 91.3± 10.4 690± 110 0 Piezoelectric w/o drawing 56.6± 7.6 15± 3.7 46.4± 6.1 540± 120 29.3± 2.7 Piezoelectric with drawing 61.8± 8.1 31± 4.2 39.4± 3.2 513± 80 36.5± 3.9 15214095, 2021, 40, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/adma.202008788 by Johns Hopkins University, Wiley Online Library on [28/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License www.advmat.dewww.advancedsciencenews.com 2008788 (5 of 18) © 2021 The Authors. Advanced Materials published by Wiley-VCH GmbH scaffold stiffness gradient, resulting in an overall enhancement of the scaffold piezoresponse (d33=−36.5 ± 3.8 pm V−1 and g33=−0.41V m N−1).[28] Differences between single fibers and multiple fibers (arrays) arise from local differences in stress distribution variations. Discrepancies between obtained values and reported values (d33=−29 pC N−1) may arise from differences in fiber diameter, orientation, geometry, and arrangement.[29] Mechanical and electromechanical stimulation of cells was performed in vitro using non-piezoelectric PTFE and piezoelectric PVDF-TrFE scaffolds under cyclic mechanical stretching. The physical and electrical properties of the scaffolds are shown in Table1. 3. The Effect of Electromechanical Stimulation on Tendon-Specific Gene Expression and Phenotypic Maintenance In Vitro We investigated the effect of topographical, mechanical, and electrical cues on tendon gene and protein expression. Due to the intrinsic non-adhesive properties of fluorinated polymers, the fibers were functionalized with an ECM molecule to enhance cell adhesion post-fabrication.[30] Fibronectin functionalization was chosen and characterized (Figure S4, Supporting Information) since it showed higher levels of cell adhesion and triggered a more significant cellular proliferation Adv. Mater. 2021, 33, 2008788 Figure 3. Fibrous aligned piezoelectric scaffolds demonstrated higher piezoelectrical performance than piezoelectric films due to geometrical boundary variables that control the piezoresponse of individual PVDF-TrFE fibers. PFM amplitude and phase images for single and multiple fibers. A) Random individual fibers (a) demonstrated lower piezoelectric performance relative to aligned individual fibers (b). Aligned superimposed fibers (c) displayed an enhanced piezoelectric response (in-plane), showing a cooperative effect due to the electromechanical interaction among adjacent fibers. Multiple dense layers (d) of adjacent fibers presented the highest piezoelectric coefficient. Commercial PVDF-TrFE films (e) display a lower piezoelectric coefficient. A non-piezoelectric platinum substrate showed no piezoresponse (f). B) Direct comparison of d33 values (piezoresponse) between samples (a–f). C) Individual fiber DMT elastic modulus (i) and electrical conductivity (iii). Electrical currents (ii) and DMT elastic modulus (iv) were measured on a platinum-coated glass surface as control, I= 300 pA. The relative stiffness difference between single fiber (350 ± 80MPa) and fiber arrays is responsible for the out-of-plane piezoresponse enhancement. No residual (I= 0 pA) electrical currents were measured in the individual fibers indicating no resistive mode of conduction characteristic of piezoelectric materials. (All measurements were obtained using N= 3 samples, r= 7 replicates per sample.) The values are presented as mean ± SD. Significant differences (one-way ANOVA) of piezoresponse of fibers (*p< 0.05, **p< 0.001) indicate results of the posthoc test (Bonferroni). 15214095, 2021, 40, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/adma.202008788 by Johns Hopkins University, Wiley Online Library on [28/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License www.advmat.dewww.advancedsciencenews.com 2008788 (6 of 18) © 2021 The Authors. Advanced Materials published by Wiley-VCH GmbH response compared with collagen type I or PLL coating.[31] While fibronectin functionalization might increase integrinmediated forces (up to 30 nN[10]) producing deformations on the piezoelectric fibers and generating interfering electrical signals, using Multiphysics simulations we have found that these level of forces are negligible compared to 4% deformation signals (see Figure S5, Supporting Information) and do not induce any permanent plastic deformations. Therefore, based on these simulations, cell-adhesion does not change the electromechanical properties of the fibers and threshold for EMS (Figure S5, Supporting Information). First, we investigated the explicit effect of fiber alignment on cell morphology, cell organization, and expression of the tenocyte-specific marker Tenomodulin (TNMD) and its transcription factor Scleraxis (SCX) by seeding human TDCs (hTDCs) onto fibrous scaffolds (non-aligned and aligned) and planar PVDF-TrFE films for 1, 3, and 7 days (Figure4). Our observations revealed that early passage (P2) tenocyte cells cultured on planar or non-aligned fibers presented a polyhedral morphology and ovoid nuclei. Conversely, the fibrous aligned structure promoted increased cell alignment in the longitudinal direction of fibers validated by actin staining. Quantification of alignment over time showed that hTDCs cultured on aligned scaffolds presented cytoskeletal elongation by day 1 and expressed higher levels of SCX and TNMD when compared to cells seeded on both planar films and non-aligned scaffolds (Figure4A,B), thus confirming the beneficial effect of fiber-aligned structures for promoting hTDC phenotype maintenance. Next, we used a high force uniaxial actuator system to apply specific strain protocols to activate different piezoelectric responses of our devices, previously seeded with hTDCs. Live/ dead assay was performed after actuation to demonstrate that the mechanical stretching actuation did not negatively affect the viability of hTDCs in vitro (Figure 4C). To optimize the actuating conditions, we analyzed the piezoelectric performance of the devices, the hTDC viability, the nuclear deformation, and the TNMD and SCX expression (Figure4C,D; Figure S6, Supporting Information) under stretching conditions. As piezoelectric systems exhibit frequency-modulated performance (output charge) due to the frequency dependence of the elastic/piezoelectric constants, we assessed the effect of strain rate (0.25, 0.5, 1, 2, and 3 Hz) on the piezoelectric constant d31 using physiologically relevant strain magnitudes (1%, 2%, and 4%) (Figure 4E,G). In general, aligned PVDF-TrFE fibers produced a significantly increased piezoelectric performance when the application of force was parallel to the fiber orientation (Figure4F). As anticipated, voltage measurements showed a positive correlation between voltage output and strain magnitude, demonstrating the device’s piezoelectric nature (also confirmed using SS-PFM, Figure4F). At a fixed frequency (i.e., 0.25Hz) the voltage output increased linearly with increasing strain magnitude (from 0.28V at 1% to 1.21 V at 4%, respectively). At fixed strain magnitude (i.e., 4%), the voltage output ranged from 0.51V at 3Hz to 1.21V at 0.25Hz (Figure4G). We then tested the voltage output stability of the device up to 500 cycles of continuous dynamic strain, and no significant change to the device voltage output (Figure2D) was observed. Finally, we chose 0.5Hz frequency and 4% strain magnitude for subsequent genomic analyses (Table2) as cell viability was preserved (comparable to tissue culture plate, Figure4C, p> 0.05 for all days). As shown in our cell viability data (Figure S7, Supporting Information), we have observed a transient effect in cell viability at day 7 in dynamic (4% strain, 0,5Hz,8h per day)versus static conditions. Importantly, this effect was not material/treatment dependent as the cell viability was identical for piezoelectric and non-piezoelectric scaffold in both static and dynamic conditions. At the same time, the TNMD expression was significantly increased (fold-increase of ≈1.5 and ≈2, after 5 and 10 days of stimulation respectively, Figure 4D). Scaffolds demonstrate a high level of piezoelectric output (1 V),illustrating thepotential of our actuated piezoelectric devices (electromechanical stimulation) in sustaining hTDC proliferation and promoting a tendon-like phenotype. Tendons have a great ability to respond to mechanical forces by adapting their structure and biochemical composition and cyclic mechanical stretching of the tendon is a recognized method for treating tendon-related injuries.[32] Several studies have confirmed that tendon is subjected to 3–4% strain during normal activities[33] and, that strain rate (loading frequency) is important in modulating the cellular response in vitro.[34] Under physiological conditions, the strain rate of the tendon is around 0.1–0.5Hz; however, during intense activity, the frequency can be as high as 10Hz.[35,36] Recently, it has been demonstrated that tendon cells modulate their gene expression, protein synthesis and mitogenesis in vitro through activation of mechanotransductive signaling pathways under physiological mechanical stimulation.[37] Therefore, we compared the effect of mechanical and electromechanical stimulation on human hTDCs gene and protein expression (Table3, 4 and 5) (Figure5). Genomic analysis of human hTDCs cultured on piezoelectric PVDF-TrFE and non-piezoelectric PTFE scaffolds under both static (4% static strain) and physiologically relevant dynamic loading conditions (4% dynamic strain at 0.5Hz for 8 h per day) was performed at 1, 5, and 10 days. Overall, the analysis of gene expression correlated well with the results observed at the protein level (Figure 5D). Electromechanical stimulation (EMS) using actuated PVDF-TrFE scaffolds, induced a rapid and sustained up-regulation of tendon-related and bone-related genes relative to static controls at day 1. Interestingly, after 10 days of EMS bone-related genes returned to basal levels whereas tendonrelated genes remained upregulated, suggesting a strong effect of EMS toward tenogenic differentiation. Similarly, mechanical stimulation alone (MS) through PTFE scaffolds induced a significant up-regulation of bone and tendon-related genes at day 1 (relative to static counterpart), but unlike EMS conditions, bone-related genes remained up-regulated while tendon-related genes returned to control levels by days 5 and 10 (Figure5A,B; Figure S8, Supporting Information). Specifically, by day 10, bone-related genes were up-regulated in hTDCs subjected to mechanical stimulation (Figure 5A,B; Figure S8, Supporting Information), whereas SCX and TNMD were upregulated only in the EMS group (Figure 5E). Taken together, these observations indicate that trans-differentiation of hTDCs toward an osteogenic/chondrogenic lineage can be modulated by EMS and MS. Moreover, EMS offered sustained tenogenic differentiation capacity relative to MS alone. To unravel the signal transduction pathways associated with the regulation of tenogenic differentiation, ingenuity pathway Adv. Mater. 2021, 33, 2008788 15214095, 2021, 40, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/adma.202008788 by Johns Hopkins University, Wiley Online Library on [28/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License www.advmat.dewww.advancedsciencenews.com 2008788 (7 of 18) © 2021 The Authors. Advanced Materials published by Wiley-VCH GmbH Adv. Mater. 2021, 33, 2008788 Figure 4. Tenoycte phenotype is promoted by scaffold alignment under specific mechanical loading conditions. A) Human TDCs demonstrated an elongated morphology from day 1 when cultured on aligned PVDF-TrFE scaffolds. Inserts are higher magnification (red = actin, green = vinculin, and blue = nucleus, scale bar = 400 μm). Angle distribution radar analysis indicated that the percentage of aligned cells was significantly modulated by fiber orientation at day 1, 3, and 7 (0° and 90° correspond to alignment and no alignment, respectively). B) Cells demonstrated differential expression of tenospecific proteins and morphological changes when cultured on electrospun scaffolds or planar PVDF-TrFE films. hTDCs maintained their phenotype and expression levels of TNMD and SCX when cultured on aligned piezoelectric scaffolds; this effect was absent on 2D planar films (N= 3, r= 3, mean ± SD, ***p< 0.001). C) hTDCs cultured on piezoelectric scaffolds under dynamic stimulation subjected to different strain rates (0.5, 1, and 2Hz) exhibited proliferation rates inversely proportional to the strain frequency (N= 3, r= 3, mean ± SD). D) An increase in the expression of TNMD was observed in cells subjected to 0.5Hz electromechanical stimulation at days 5 and 10 (N= 3, r= 3, mean ± SD *p< 0.05). E). An overview of the mechanical loading system used to measure scaffold voltage output under stimulated physiological strain conditions and in vitro analysis. F) Representative hysteresis and butterfly loops of the fiber piezoresponse and polarization. SS-PFM facilitated the measurement of the PFM phase and amplitude response loops as a function of the applied voltage. The amplitude image shows the magnitude of displacement response generated under a controlled applied voltage in drawn fibers, and the highest amplitude peak was shown to be around 300 pm in response to a 12V bias. The phase image shows the positive and negative values of antiparallel ferroelectric nanodomains. The phase measurements indicated a ≈180° switch of the dipoles between the applied positive and negative voltages. The piezoresponse obtained from PFM measurements showed that the electrical dipoles lie normal to the surface, characteristic of the organized electrical dipoles/chains of β-phase crystals. PTFE did not demonstrate piezoelectric behavior (gray line). The schematic indicates the voltage distribution on the fiber surface as a function of geometrical arrangement (fiber orientation) and interaction with adjacent fibers resulting in differing transverse deformation restriction and modulation of the piezoresponse the longitudinal fiber axis. G) Voltage measurements as a function of frequency (0.25, 0.5, 1, 2, and 3Hz) at a constant amplitude of 1%, 2%, and 4% strain.). 15214095, 2021, 40, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/adma.202008788 by Johns Hopkins University, Wiley Online Library on [28/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License www.advmat.dewww.advancedsciencenews.com 2008788 (8 of 18) © 2021 The Authors. Advanced Materials published by Wiley-VCH GmbH analysis (IPA) was used with gene expression profiles of hTDC exposed to MS and EMS stimulation. IPA attributes an activation z-score used to predict the activation state of biological functions and signaling pathways. Pathway analysis of human tendon derived cells exposed to either EMS or MS yielded significant modulation of broad functional signaling pathways, and a mechanistic network of genes and specific biological functions were differentially regulated (Figure S9, Supporting Information). Canonical signaling pathways with the most Adv. Mater. 2021, 33, 2008788 Table 2. QIAGEN genes list for customized gene array. Symbol Entrez gene name ABCB1 ATP binding cassette subfamily B member 1 ACAN Aggrecan ACTA1 actin, alpha 1, skeletal muscle ACVR1 activin A receptor type 1 AHSG alpha 2-HS glycoprotein ALPL alkaline phosphatase, liver/bone/kidney BGLAP bone gamma-carboxyglutamate protein BGN Biglycan BMP1 bone morphogenetic protein 1 BMP2 bone morphogenetic protein 2 BMP4 bone morphogenetic protein 4 BMP6 bone morphogenetic protein 6 BMP7 bone morphogenetic protein 7 BMPR1A bone morphogenetic protein receptor type 1A BMPR2 bone morphogenetic protein receptor type 2 CASP3 caspase 3 COL11A1 collagen type XI alpha 1 chain COL14A1 collagen type XIV alpha 1 chain COL1A1 collagen type I alpha 1 chain COL1A2 collagen type I alpha 2 chain COL2A1 collagen type II alpha 1 chain COL3A1 collagen type III alpha 1 chain COL4A1 collagen type IV alpha 1 chain COL5A1 collagen type V alpha 1 chain COL6A1 collagen type VI alpha 1 chain COMP cartilage oligomeric matrix protein DCN Decorin DLX5 distal-less homeobox 5 EGR1 early growth response 1 FGF10 fibroblast growth factor 10 GDF15 growth differentiation factor 15 GDF5 growth differentiation factor 5 GDF6 growth differentiation factor 6 GDF7 growth differentiation factor 7 HAT1 histone acetyltransferase 1 HDAC1 histone deacetylase 1 HNF1A HNF1 homeobox A IBSP integrin binding sialoprotein IGF1 insulin like growth factor 1 ITGA1 integrin subunit alpha 1 ITGA2 integrin subunit alpha 2 ITGA3 integrin subunit alpha 3 ITGA4 integrin subunit alpha 4 ITGA5 integrin subunit alpha 5 ITGAX integrin subunit alpha X ITGB1 integrin subunit beta 1 ITGB3 integrin subunit beta 3 Symbol Entrez gene name ITGB5 integrin subunit beta 5 KAT2B lysine acetyltransferase 2B KCNK2 potassium two pore domain channel subfamily K member 2 KCNK4 potassium two pore domain channel subfamily K member 4 KDR kinase insert domain receptor MGP matrix Gla protein MKX mohawk homeobox PIEZO1 piezo type mechanosensitive ion channel component 1 PIEZO2 piezo type mechanosensitive ion channel component 2 PIK3CG phosphatidylinositol-4,5-bisphosphate 3-kinase catalytic subunit gamma PTEN phosphatase and tensin homolog PTK2 protein tyrosine kinase 2 PTK2 protein tyrosine kinase 2 PXN Paxillin RUNX2 runt related transcription factor 2 SCX scleraxis bHLH transcription factor SMAD3 SMAD family member 3 SMAD4 SMAD family member 4 SMAD9 SMAD family member 9 SMURF1 SMAD specific E3 ubiquitin protein ligase 1 SMURF2 SMAD specific E3 ubiquitin protein ligase 2 SOX9 SRY-box 9 SP7 Sp7 transcription factor SPARC secreted protein acidic and cysteine rich SPP1 secreted phosphoprotein 1 TBX5 T-box 5 TGFB1 transforming growth factor beta 1 THBS4 thrombospondin 4 TLN1 talin 1 TNC tenascin C TNMD Tenomodulin TRPA1 transient receptor potential cation channel subfamily A member 1 TRPV1 transient receptor potential cation channel subfamily V member 1 TWIST1 twist family bHLH transcription factor 1 VCL Vinculin VEGFA vascular endothelial growth factor A ZYX Zyxin Table 2. Continued. 15214095, 2021, 40, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/adma.202008788 by Johns Hopkins University, Wiley Online Library on [28/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License www.advmat.dewww.advancedsciencenews.com 2008788 (9 of 18) © 2021 The Authors. Advanced Materials published by Wiley-VCH GmbH significant number of modulated gene expression were associated with day 1 and 10 in vitro (Figure S9, Supporting Information). The ERK/MAPK signaling pathway was consistently activated with both EMS and MS stimulation; however, this pathway was significantly upregulated in hTDCs subjected to MS at days 1 and 10 (Figure5E). Similarly, increased activation of cell transdifferentiation and differentiation toward the bone cell lineage and upregulations in osteogenic signaling pathways Wnt/β-Catenin and BMP were attributed to MS at day 10 (relative to EMS). Significantly, following 10 days in culture, the tendon-related signaling pathway IGF-1 and biological differentiation process associated with tenocyte function were activated in tendon derived cells subjected to EMS but downregulated under MS (Figure 5E). We looked at the protein expression profile of tendon-specific maturation markers and collagen synthesis to validate gene expression results. In agreement with the gene expression data, we observed a higher expression of tendon maturation markers (SCX and TNC) of tendon derived cells exposed to EMS relative to MS in addition to the increased synthesis of collagen type V (Figure5F,G). During tendon specific differentiation or trans-differentiation, highly regulated signal transduction events take place, leading to the expression of genes associated with the tendonspecific phenotype or phenotypic drift respectively. We hypothesized that mechanically activated and electrically gated membrane ion channels would undergo modulated expression in response to EMS. To investigate the role of these membrane sensors on signal generation and propagation, correlational analysis was conducted between significant modulations to tendon-specific functional pathways at day 1, 5, and 10 in the mechanistic network to differential changes in the expression of mechano-sensitive (TRP family ion channels, focal adhesion-related proteins and integrins β1, β3, and β5), electro-sensitive (KCNK family and Ca2+ L-type ion channel), and piezosensitive (Piezo family ion channels) receptors, as well as the BMP receptor BMPR1A[38,39] (Figure 5C). Critically, increased activation of tendon differentiation-related genes (Figure 5A) through the modulated activity of tendon-related transcription factors (i.e., SCX and EGR1, see Figure S8, Supporting Information) was correlated with significant modulation of the expression level of TRP family ion channels under stimulation (Figure5G). As shown in Figure5F, the expression of SCX and TNC was significantly increased under EMS conditions at both day 5 and 10, together with a significantly lower expression of KCNK4, TRPA1, and TRPV1 ion channels when compared to the MS group. These results were further validated at the protein level (Figure5G). Conversely under MS, a decrease in the expression of tendonrelated genes and an increase in the expression of genes associated with cartilage/bone differentiation was associated with higher expression of TRP ion channels (Figure5E,G; Figure S8, Supporting Information). Specifically, TRPA1 and TRPV1 ion channels underwent a sustained increase in expression which correlated with a positive regulation of transcription factors SOX9, RUNX2, and COMP, proteins that have a recognized role in the processes of osteogenesis (Figure 5B; Figure S6, Supporting Information). Overall, the gene expression results and pathways analyses, further validated using a custom-made protein array, demonstrated that MS of hTDCs in vitro resulted Adv. Mater. 2021, 33, 2008788 Table 4. Signaling array: proteins associate to different signaling pathways (MAPK, FAK, TGF-B, BMP, and WNT). Proteins Abbreviation Purchased from Cat. no Smad 1 SMAD1 Cell Signaling 6944S 6944S Smad 5 SMAD3 Cell Signaling 12534S Phosphorylated Smad 1 SMAD1/5/8 Cell Signaling 5753S Phosphorylated Smad 1/5/8 pSMAD158 Cell Signaling 9516s 9516s Focal adhesion kinase FAK MBL D061-3 12G4 Phosphorylated FAK pFAK Cell Signaling 3284S 3284S MAPK ERK Cell Signaling 4696S L34F12 p44/42 MAPK pERK Cell Signaling 4377S Wnt/β-catenin β-Catenin Millipore 2 858 901 Active β-catenin, clone 8E7 Active β-Catenin Millipore ABC β-actin β-actin WAKO 019-19741 Table 3. Tenogenesis array: proteins associated with tendon regeneration or hTDC function. Proteins Abbreviation Purchased from Cat. no Scleraxis SCX Abcam ab58655 Tenomodulin TNMD Abcam ab203676 Byglican BGN Abcam ab49701 Decorin DCN Abcam ab175404 Thrombospondin 4 THBS-4 Abcam ab176116 Tenascin C TNC Abcam ab88280 Collagen I COLI Abcam ab138492 Collagen II COLI Abcam ab185430 Collagen III COLIII Abcam ab7778 Collagen V COLV Abcam ab7046 Table 5. Receptors array: proteins associate to cell membrane receptors. Proteins Abbreviation Purchased from Cat. no TRPV1 TRPV1 SantaCruz sc-20813 Piezo1 Piezo1 SantaCruz sc-164319 Piezo2 Piezo2 SantaCruz sc-84763 TRPA1 TRPA1 SantaCruz sc-32353 KCNK2 KCNK2 SantaCruz sc-11557 KCNK4 KCNK4 Abcam ab81367 L-type Ca2+L-type Ca2+SantaCruz sc-25686 BMPR1A BMPR1A ThermoFisher PA5-11856 Integrin1 ITG1 Abcam ab134179 Integrin3 ITG3 Abcam ab34409 Integrin5 ITG5 Cell Signaling 3629S 15214095, 2021, 40, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/adma.202008788 by Johns Hopkins University, Wiley Online Library on [28/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License www.advmat.dewww.advancedsciencenews.com 2008788 (16 of 18) © 2021 The Authors. Advanced Materials published by Wiley-VCH GmbH Adv. Mater. 2021, 33, 2008788 bigger is the size the higher is the expression level and the rest of dots are scaled proportionately. High-confidence values were represented by a red outline corresponding to a p-value <0.05orby a black outline corresponding to a p-value <0.01,whereasfor p-values >0.05theoutline was faint. Protein Expression: Protein antibody microarray was custom made. Nexterion slide H microarray slides were purchased from Schott AG (Mainz, Germany). Alexa Fluor 555 carboxylic acid succinimidyl ester was obtained from Life Technologies (Carlsbad, CA, USA). Protein samples were labeled with Alexa Fluor 555 carboxylic acid succinimidyl ester according to manufacturer’s instructions. The excess label was removed, and the buffer was exchanged with PBS, pH 7.4, by centrifugation through 3kDa molecular weight cutoff filters. Absorbance at 555 and 280nm was measured for labeled samples and calculations were performed according to manufacturer’s instructions using an arbitrary extinction coefficient of 100 000 and molecular mass of 100 000 to enable quantification of relative protein concentration and label substitution efficiency. All commercial antibodies (Table1) were buffer exchanged into PBS and quantified by a bicinchoninic acid (BCA) assay. Antibodies were diluted to print concentration in PBS and printed in six replicates on Nexterion H amine-reactive, hydrogel-coated glass slides using a SciFLEXARRAYER S3 piezoelectric printer (Scienion, Berlin, Germany) under constant humidity (62% +/− 2%) at 20°C. Each feature was printed using ≈1 nL of diluted antibody using an uncoated 90µm glass nozzle with eight replicated subarrays per microarray slide. After printing, slides were incubated in a humidity chamber overnight at room temperature to facilitate complete conjugation. The slides were then blocked in 100 × 10−3 m ethanolamine in 50 × 10−3 m sodium borate, pH 8.0, for 1 h at room temperature. Slides were washed in PBS with 0.05% Tween 20 (PBS-T) three times for 2 min each wash followed by one wash in PBS, dried by centrifugation (470 × g, 5min), and then stored with a desiccant at 4°C until use. Incubations were carried out in the dark. Microarray slides were incubated as previously described. Initially, one labeled sample was titrated (2.5–15µg mL−1) for optimal signal to noise ratio and all samples were subsequently incubated for 1 h at 23°C at 9µg mL−1 in Tris-buffered saline (TBS; 20 × 10−3 m Tris-HCl, 100 × 10−3 m NaCl, 1 × 10−3 m CaCl2, 1 × 10−3 m MgCl2, pH 7.2) with 0.05% Tween 20 (TBS-T). All microarray experiments were carried out using three replicate slides. Alexa Fluor 555 labeled cells lysate (10µg mL−1) were incubated in two separate subarrays on every slide to confirm retained antibody performance and printing, respectively (Figure 1). After incubation, slides were washed three times in TBS-T for 2min per wash, once in TBS and then centrifuged dry as above. Dried slides were scanned immediately on an Agilent G2505 microarray scanner using the Cy3 channel (532nm excitation, 90% photomultiplier tubes (PMT), 5µm resolution) and intensity data were saved as a .tif file. Antibody microarrays were verified to remain active for at least 2 weeks after printing and all incubations were carried out within that time frame. Data extraction from .tif files was performed mainly as previously described. Data were normalized to the mean of three replicate microarray slides (subarray-by-subarray using subarray total intensity, n= 4, 24 data points). Unsupervised hierarchical clustering of normalized data was performed using Hierarchical Clustering Explorer v3.0 (http://www. cs.umd.edu/hcil/hce/hce3.html) using the parameters no prefiltering, complete linkage, and Euclidean distance. All data presented here were confirmed using at least four replicates for each of the test groups and control group. The results are expressed as the mean of the values ± standard error of the mean. Different types of arrays were fabricated to investigated tendon regeneration or phenotype maintenance (tenogenesis), intracellular molecular pathways (signaling) or membrane proteins (receptors). In Vivo Animal Model: The animal care research ethics committee at the National University of Ireland, Galway and Health Products Regulatory Authority (AE19125/P055) approved all the animal procedures used in this study. Also, animal care and management followed the Standard Operating Procedures of the Animal Facility of the National University of Ireland, Galway. Animals were allowed to acclimatize for at least 7 days before any surgical procedures. Subsequently, animals were acclimated to the treadmill running for 1 week, and their behavior was analyzed. A total of 105 Female Lewis rats aged 6–8 (220g)weekswereused in this study. The animals were anaesthetized by isofluorane inhalation (5% induction reducing to 1–2% for maintenance during procedures). The right leg was shaved and swabbed with iodine to minimize the risk of bacterial contamination. An incision was created through the skin (≈1cm)fromthe myotendinous junction distally to the osteotendinous junction. The incision provided ample exposure to the Achilles tendon. The fascia surrounding the Achilles was transected longitudinally and carefully retracted to expose the Achilles tendon. Before implantation and tendon transection, two looped sutured were inserted at the top (muscle) and bottom (bone). After the total tendon length was measured a 3 mm defect in proximal/distal extension (at 3mm from the calcaneus) was created using a positioning device and an 11 surgical blade, resulting in a 6mm gap after tissue retraction. The construct was then sutured (4-0 Ethicon) to both ends of the tendon to bridge the gap using a modified Kessler technique, and the skin was sutured. After a period of 2, 4, and 8 weeks the animals were euthanized and tendon tissue, as well as contralateral tendons, were harvested. Animals recovered for 2 weeks and were gradually exposed to treadmill running (see Table 6). Briefly, the treadmill running once a week for 5min to 30–45 min for 5 days a week. Histology: Repaired tendon tissues and contralateral tendons of all groups (n= 7) were dissected from the proximal myotendinous junction to the distal osteotendinous junction and processed for histological analysis. The samples were fixed in 10% neutral buffered formalin (24 h), dehydrated in gradient alcohols, cleared, and embedded in paraffin blocks, as reported previously. Histological sections (6 µm thick) were prepared using microtome sectioning (Leica Rotary Microtome). In order to distinguish between scar tissue or new tendon formation, polarization microscopy and picrosirius red staining were used. Also, for descriptive histology 6 µm, thick sections were stained using Hematoxylin & Eosin stain, Masson-Goldner’s stain, Alcian Blue, O-safranin stain, Red picrosirius stain, or Herovici’s polychrome stain according to the manufacturer’s guidelines. Areas of chondrification within the defect region at 4 or 8 weeks after surgery were measured using Safranin O staining and ImageJ (v1.52i). For each tissue, 3 different frontal-longitudinal sections were analyzed (1 section of the middle tendon, 1 section ventral to this middle part, and 1 section dorsal to this middle part). For each section5 consecutive images were captured Table 6. Exercise protocol used for the rat running group (treadmill). Time Duration [min] Speed [m min−1] Week 2 Day 14 5 8–9 Day 15 10 9–10 Day 16 15 9–10 Day 17 20 9–10 Day 18 30 9–10 Week 3 Day 19 45 9–11 Day 20 30 9–11 Day 21 45 9–12 Day 22 30 10–14 Day 23 45 10–14 Week 4 30 10–14 Week 5 45 10–14 Week 6 30 10–14 Week 7 45 10–14 Week 8 30 10–14 Week 9 45 10–14 Week 10–16 30 10–14 15214095, 2021, 40, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/adma.202008788 by Johns Hopkins University, Wiley Online Library on [28/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License www.advmat.dewww.advancedsciencenews.com 2008788 (17 of 18) © 2021 The Authors. Advanced Materials published by Wiley-VCH GmbH Adv. Mater. 2021, 33, 2008788 spanning the entire length of the tissue, omitting the transition zones of original tendon stumps to regenerated tissue. The volume fraction (VV) of tendon cells were used to estimate cell proliferation. A 192-point grid was overlaid on 40× images of H&E stained tissue sections. The number of tendon cells intersecting points of the grid was counted (PP), along with the total number of points on the tissue (PT). The volume fraction of tendon cells (VV) was calculated using the formula below: VP /P VPT = (4) Histology Scoring System: In order to highlight the effect of treadmill running and differences between groups on the progression of tendon repair and calcification, a score system was adopted previously described.[58] Briefly, the macroscopic score system included measurements on stained samples (Hematoxylin & Eosin, MassonGoldner’s stain, Alcian Blue, O-safranin, Red picrosirius, and Herovici’s polychrome) three animals per group (five sections of 6 μm per animal). A point-based scoring system was used, and the following parameters were scored (by two scorers): ECM organization of the whole repaired tendon, cellularity, proteoglycan content, cell alignment, organization of the tendon callus, integration of constructs to the normal tissue, vascularization, degenerative changes (osteochondral), and features of inflammation. A total of three scorers were used for the scoring and the slides were blinded by one of the authors (MFY). Functional Recovery Analysis: An animal treadmill (Exer 3/6, Columbus Instruments) track integrated with a video-based system was used to obtain spatiotemporal parameters of gait. The animal gait was analyzed through a clear plastic Lexan at the sides of the system consisting of a cage (50.8cm × 50.8cm × 33cm) with gates placed at each end of the walkway. A digital camera (8 M pixels and 120 frames per seconds) was positioned 30cm in front of the walkway to capture the sagittal view of the rat from the walkway. The data were analyzed and processed with Kinovea software (v0.8.27). For all groups, the system was calibrated using the same scale bar located in the image and modeled the leg motion using pairs of circle markers. The ankle, knee, hip, and MTP joint angles were measured using the automatic black ink marker feature recognition on the hip, knee, ankle, and 3rd metatarsal head at the four gait stages: initial contact, mid-stance, pre-swing, and mid-swing. The spatial and temporal gait parameters analyzed were step length, angle, and cycle time. The walking speed was calculated by dividing the step length by the cycle time. Each angle joint curve was normalized by cycle time before further analysis. To determine the change on gait angle, the maximum difference in amplitude (Ampl.) between initial contact and end of swing was measured during an entire step and was calculated according to following formula: ma x. min. Ampl. () () () °− °= ° (5) transectio nt ransection Fu nctionalRecovery Ampl.( )/Ampl.( ) post prior =° ° (6) COMSOL Simulations: To understand the influence of the potential distribution around the scaffold during electromechanical stimulation and how this might impact the cells near the scaffold, we calculated the potential distribution of the scaffold through a finite element method using electrostatic, piezoelectric, and solid mechanics interaction modulus in COMSOL. The finite element method (FEM) analysis uses an integrated device structure consisting of a tubular-shaped electrospun scaffold composed of PVDF-TrFE fibers under 4% deformation (0.2mm). The geometries and the electromechanical properties are set as the real measured values. Due to the complexity of the nanofibrous structure, we used a simplified PVDF-TrFE model consisting of a low-density film (density of 450kg m−3). The young modulus is 60MPa, Poisson’s ratio is 0.42, and the strain-charge coupling matrix is d31= 23 pC N−1, d32= 8pC N−1, and d33=−36.5pC N−1. In addition, we have investigated the effects of high adhesion forces (up to 30 nN[10]) applied through focal adhesion complexes (1–5 µm long) on a single PVDF-TrFE fiber. Statistical Analysis: Statistical analyses were performed using GraphPad Prism software version 8.4.3 (GraphPad Software, CA, USA). To test whether the data was normally distributed normality tests were applied (D’Agostino and Pearson). When data followed a normal distribution a one-way ANOVA analysis for the comparison of means between different groups was performed. Homogeneity of variances was tested using Bartlett’s tests, and in a case of unequal standard deviations (SD), Welch ANOVA test was applied. If data were not normally distributed, the comparison of medians between different groups was assessed by non-parametric Kruskal-Wallis test followed by Dunn post-hoc test. Since most of the data was not normally distributed, most of the results were expressed as median as central tendency characteristic and interquartile range (IQR) or range as dispersion characteristic and, p values of <0.05 were considered statistically significant. Supporting Information Supporting Information is available from the Wiley Online Library or from the author. Acknowledgements The work was supported by grants to MJPB from Science Foundation Ireland (16/BBSRC/3317), to MAFY from H2020 Marie Skłodowska-Curie Actions (898737) and grant from Science Foundation Ireland (SFI), co-funded under the European Regional Development Fund through Grant numbers 13/RC/2073 and 13/RC/2073_P2. The authors thank Dr. Oliver Carroll for technical assistance. SGIker technical services (UPV/EHU) are gratefully acknowledged for XRD and XPS support. The authors acknowledge the facilities and scientific and technical assistance of the Centre for Microscopy & Imaging at the National University of Ireland Galway, a facility that is funded by NUIG and the Irish Government’s Programme for Research in Third Level Institutions, Cycles 4 and 5, National Development Plan 2007–2013. Open access funding provided by IReL. Note: The acknowledgements section was updated on October 5, 2021, after initial publication online. Conflict of Interest The authors declare no conflict of interest. Author Contributions M.A.F.-Y., M.J.B., and A.P. conceived the experiments. M.A.F.-Y. performed the experiments. A.T., S.D., A.S., and A.L. performed research. M.K., A.P., T.S., and M.P. provided materials and expertise. M.A.F.-Y. analyzed the data and prepared the figures. A.P. and M.J.B. provided critique and context for the data. M.A.F.-Y. wrote the manuscript. All authors read and commented on the manuscript. Data Availability Statement The data that support the findings of this study are openly available in bioRxiv at https://doi.org/10.1101/2020.08.03.22786. Keywords bioelectronics, collagen, piezoelectrics, poly(vinylidene fluoride-cotrifluoroethylene), tendon regeneration Received: December 29, 2020 Revised: May 17, 2021 Published online: August 23, 2021 15214095, 2021, 40, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/adma.202008788 by Johns Hopkins University, Wiley Online Library on [28/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License www.advmat.dewww.advancedsciencenews.com 2008788 (18 of 18) © 2021 The Authors. Advanced Materials published by Wiley-VCH GmbH [1] A. J.Lomas, C. N. M.Ryan, A.Sorushanova, N.Shologu, A. I.Sideri, V. Tsioli, G. C. Fthenakis, A. Tzora, I. Skoufos, L. R. Quinlan, G. O’Laighin, A. M. Mullen, J. L. Kelly, S. Kearns, M. Biggs, A.Pandit, D. I.Zeugolis, Adv. Drug Delivery Rev. 2015, 84, 257. [2] J. G.Snedeker, J.Foolen, Acta Biomater. 2017, 63, 18. [3] A. C.Egger, M. J.Berkowitz, Curr. Rev. Musculoskeletal Med. 2017, 10, 72. [4] G.Nourissat, F.Berenbaum, D.Duprez, Nat. Rev. Rheumatol. 2015, 11, 223. [5] J.Zhang, J. H.-C.Wang, PLoS One 2013, 8, e71740. [6] C. Popov, M. Burggraf, L. Kreja, A. Ignatius, M. Schieker, D.Docheva, BMC Mol. Biol. 2015, 16, 6. [7] J.Zhang, T.Pan, Y.Liu, J. H. C.Wang, J. Orthop. Res. 2010, 28, 1172. [8] E. Gracey, A. Burssens, I. Cambré, G. Schett, R. Lories, I. B.McInnes, H.Asahara, D.Elewaut, Nat. Rev. Rheumatol. 2020, 16, 193. [9] M. L.Killian, L.Cavinatto, L. M.Galatz, S.Thomopoulos, J. Shoulder Elbow Surg. 2012, 21, 228. [10] M. S.Kim, M. H.Lee, B.-J.Kwon, H. J.Seo, M.-A.Koo, K. E.You, D.Kim, J.-C.Park, J. Tissue Eng. Regener. Med. 2017, 11, 862. [11] J.Song, G.Zhao, B.Li, J.Wang, Heliyon 2017, 3, e00377. [12] J. Bolander, Y. C. Chai, L. Geris, J. Schrooten, D. Lambrechts, S. J.Roberts, F. P.Luyten, Biomaterials 2016, 86, 106. [13] C. K.Kuo, J. E.Marturano, R. S.Tuan, BMC Sports Sci. Med. Rehabil. 2010, 2, 20. [14] B.Zhao, X.Cui, W.Ren, F.Xu, M.Liu, Z.-G.Ye, Sci. Rep. 2017, 7, 11319. [15] D. Denning, J. I. Kilpatrick, E. Fukada, N. Zhang, S. Habelitz, A.Fertala, M. D.Gilchrist, Y.Zhang, S. A. M.Tofail, B. J.Rodriguez, ACS Biomater. Sci. Eng. 2017, 3, 929. [16] D.Denning, M. T.Abu-Rub, D. I.Zeugolis, S.Habelitz, A.Pandit, A.Fertala, B. J.Rodriguez, Acta Biomater. 2012, 8, 3073. [17] G. G. Genchi, E. Sinibaldi, L. Ceseracciu, M.Labardi, A.Marino, S. Marras, G. De Simoni, V. Mattoli, G. Ciofani, Nanomedicine: Nanotechnology, Biology and Medicine 2018, 14, 2421. [18] S. M.Damaraju, Y.Shen, E.Elele, B.Khusid, A.Eshghinejad, J.Li, M.Jaffe, T. L.Arinzeh, Biomaterials 2017, 149, 51. [19] Y.-S.Lee, G.Collins, T. L.Arinzeh, Acta Biomater. 2011, 7, 3877. [20] G. G. Genchi, L. Ceseracciu, A. Marino, M. Labardi, S. Marras, F.Pignatelli, L.Bruschini, V.Mattoli, G.Ciofani, Adv. Healthc. Mater. 2016, 5, 1808. [21] Y.Li, Y.Xiao, C.Liu, Chem. Rev. 2017, 117, 4376. [22] J.Kastelic, A.Galeski, E.Baer, Connect. Tissue Res. 1978, 6, 11. [23] Z.Bartczak, Macromolecules 2005, 38, 7702. [24] R. D. Cardwell, L. A. Dahlgren, A. S. Goldstein, J. Tissue Eng. Regener. Med. 2014, 8, 937. [25] M. Baniasadi, Z.Xu, S.Moreno, S. Daryadel, J. Cai, M. Naraghi, M.Minary-Jolandan, Polymer (Guildf). 2017, 118, 223. [26] Frost & Sullivan, Artificial Intelligence in Health and Life Sciences Conference, Kisaco Research, 2016. [27] Y.Li, C.Liao, S. C.Tjong, Nanomaterials 2019, 9, 952. [28] L. Persano, C. Dagdeviren, C. Maruccio, L. De Lorenzis, D.Pisignano, Adv. Mater. 2014, 26, 7574. [29] J. Li, L. Kang, Y. Yu, Y. Long, J. J. Jeffery, W. Cai, X. Wang, Nano Energy 2018, 51, 728. [30] A.Poudel, M. A.Fernandez, S. A. M.Tofail, M. J. P.Biggs, Front. Chem. 2019, 7, 364. [31] D. Klee, Z. Ademovic, A. Bosserhoff, H. Hoecker, G. Maziolis, H. J.Erli, Biomaterials 2003, 24, 3663. [32] M. P.McHugh, C. H.Cosgrave, Scand. J. Med. Sci. Sports 2010, 20, 169. [33] J. H. C.Wang, Q.Guo, B.Li, J. Hand Ther. 2012, 25, 133. [34] N.Theis, A. A.Mohagheghi, T.Korff, J. Electromyogr. Kinesiol. 2012, 22, 947. [35] S.Chaudhry, D.Morrissey, R. C.Woledge, D. L.Bader, H. R. C.Screen, J. Appl. Biomech. 2015, 31, 69. [36] C. P.Udeze, E. R.Jones, G. P.Riley, D.Morrissey, H. R. C.Screen, Scand. J. Med. Sci. Sport. 2019, 29, 1511. [37] G.Yang, R. C.Crawford, J. H. C.Wang, J. Biomech. 2004, 37, 1543. [38] D. E.Ingber, Proc. Natl. Acad. Sci. U. S. A. 2003, 100, 1472. [39] S. S.Ranade, S. H.Woo, A. E.Dubin, R. A.Moshourab, C.Wetzel, M.Petrus, J.Mathur, V.Bégay, B.Coste, J.Mainquist, A. J.Wilson, A. G. Francisco, K. Reddy, Z. Qiu, J. N. Wood, G. R. Lewin, A.Patapoutian, Nature 2014, 516, 121. [40] K. M.Khan, A.Scott, Br. J. Sports Med. 2009, 43, 247. [41] H. Suzuki, Y. Ito, M. Shinohara, S. Yamashita, S. Ichinose, A. Kishida, T. Oyaizu, T. Kayama, R. Nakamichi, N. Koda, K.Yagishita, M. K.Lotz, A.Okawa, H.Asahara, Proc. Natl. Acad. Sci. U. S. A. 2016, 113, 7840. [42] J. H.-C.Wang, F.Jia, G.Yang, S.Yang, B. H.Campbell, D.Stone, S. L.-Y.Woo, Connect. Tissue Res. 2003, 44, 128. [43] K. A.Mace, J. C.Pearson, W.McGinnis, Science 2005, 308, 381. [44] L. N.Handly, A.Pilko, R.Wollman, Elife 2015, 8, e09652. [45] L. J. Leiper, P. Walczysko, R. Kucerova, J. Ou, L. J. Shanley, D.Lawson, J. V.Forrester, C. D.McCaig, M.Zhao, J. M.Collinson, BMC Biol. 2006, 4, 27. [46] T. W.Gilbert, S.Wognum, E. M.Joyce, D. O.Freytes, M. S.Sacks, S. F.Badylak, Biomaterials 2008, 29, 4775. [47] A.Islam, T.Mbimba, M.Younesi, O.Akkus, Acta Biomater. 2017, 58, 244. [48] A.English, A.Azeem, K.Spanoudes, E.Jones, B.Tripathi, N.Basu, K.McNamara, S. A. M.Tofail, N.Rooney, G.Riley, A.O’Riordan, G.Cross, D.Hutmacher, M.Biggs, A.Pandit, D. I.Zeugolis, Acta Biomater. 2015, 27, 3. [49] S. Ribeiro, A. C. Gomes, I. Etxebarria, S. Lanceros-Méndez, C.Ribeiro, Mater. Sci. Eng., C 2018, 92, 868. [50] Z. Goli-Malekabadi, M. Tafazzoli-Shadpour, M. Rabbani, M.Janmaleki, Biomed. Tech. 2011, 56, 259. [51] M.Younesi, A. Islam, V. Kishore, J. M. Anderson, O.Akkus, Adv. Funct. Mater. 2014, 24, 5762. [52] L. Li, W. Gu, J. Du, B. Reid, X.Deng, Z.Liu, Z. Zong, H. Wang, B. Yao, C. Yang, J. Yan, L. Zeng, L. Chalmers, M. Zhao, J. Jiang, Wound Repair Regener. 2012, 20, 840. [53] B.Reid, B.Song, C. D.McCaig, M.Zhao, FASEB J. 2005, 19, 379. [54] C. Halperin, S. Mutchnik, A. Agronin, M. Molotskii, P. Urenski, M.Salai, G.Rosenman, 2004. [55] Y.-S. Lee, S. Wu, T. Livingston Arinzeh, M. B. Bunge, Biotechnol. Bioeng. 2017, 114, 444. [56] M. U. Wagenhäuser, M. F. Pietschmann, B. Sievers, D. Docheva, M. Schieker, V. Jansson, P. E. Müller, BMC Musculoskelet. Disord. 2012, 13, 140. [57] J. D. R. Knight, H. Choi, G. D. Gupta, L. Pelletier, B. Raught, A. I.Nesvizhskii, A. C.Gingras, Nat. Methods 2017, 14, 645. [58] C.Stoll, T.John, C.Conrad, A.Lohan, S.Hondke, W.Ertel, C.Kaps, M. Endres, M. Sittinger, J. Ringe, G. Schulze-Tanzil, Biomaterials 2011, 32, 4806. Adv. Mater. 2021, 33, 2008788 15214095, 2021, 40, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/adma.202008788 by Johns Hopkins University, Wiley Online Library on [28/05/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons License