Chronic softening spinal cord stimulation arrays

Journal of Neural Engineering

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Journal of Neural Engineering J. Neural Eng. 15 (2018) 045002 (17pp)

https://doi.org/10.1088/1741-2552/aab90d

Chronic softening spinal cord stimulation arrays Aldo Garcia-Sandoval1, Ajay Pal2 , Asht M Mishra2,6, Sydney Sherman4, Ankit R Parikh1, Alexandra Joshi-Imre3, David Arreaga-Salas3, Gerardo Gutierrez-Heredia3,8, Adriana C Duran-Martinez4, Jordan Nathan2, Seyed Mahmoud Hosseini5, Jason B Carmel2,6 and Walter Voit1,3,4,7 1

Department of Mechanical Engineering, The University of Texas at Dallas, Richardson, TX 75080, United States of America 2 Motor Recovery Laboratory, Burke Medical Research Institute, White Plains, NY 10605, United States of America 3 Department of Materials Science and Engineering, The University of Texas at Dallas, Richardson, TX 75080, United States of America 4 Department Bioengineering, The University of Texas at Dallas, Richardson, TX 75080, United States of America 5 Department of Chemistry, The University of Texas at Dallas, Richardson, TX 75080, United States of America 6 Brain and Mind Research Institute and Departments of Neurology and Pediatrics, Weill Cornell Medicine, New York, NY, United States of America 7 Center for Engineering Innovation, The University of Texas at Dallas, Richardson, TX 75080, United States of America 8 Centro de Investigaciones en Optica, Leon Guanajuato 37150, Mexico E-mail: [email protected] Received 14 October 2017, revised 15 March 2018 Accepted for publication 23 March 2018 Published 15 May 2018 Abstract

Objective. We sought to develop a cervical spinal cord stimulator for the rat that is durable, stable, and does not perturb the underlying spinal cord. Approach. We created a softening spinal cord stimulation (SCS) array made from shape memory polymer (SMP)-based flexible electronics. We developed a new photolithographic process to pattern high surface area titanium nitride (TiN) electrodes onto gold (Au) interconnects. The thiol-ene acrylate polymers are stiff at room temperature and soften following implantation into the body. Durability was measured by the duration the devices produced effective stimulation and by accelerated aging in vitro. Stability was measured by the threshold to provoke an electromyogram (EMG) muscle response and by measuring impedance using electrochemical impedance spectroscopy (EIS). In addition, spinal cord modulation of motor cortex potentials was measured. The spinal column and implanted arrays were imaged with MRI ex vivo, and histology for astrogliosis and immune reaction was performed. Main results. For durability, the design of the arrays was modified over three generations to create an array that demonstrated activity up to 29 weeks. SCS arrays showed no significant degradation over a simulated 29 week period of accelerated aging. For stability, the threshold for provoking an EMG rose in the first few weeks and then remained stable out to 16 weeks; the impedance showed a similar rise early with stability thereafter. Spinal cord stimulation strongly enhanced motor cortex potentials throughout. Upon explantation, device performance returned to pre-implant levels, indicating that biotic rather than abiotic processes were the cause of changing metrics. MRI and histology showed that softening SCS produced less tissue deformation than Parylene-C arrays. There was no significant astrogliosis or immune reaction to either type of array. Significance. Softening SCS arrays meet the needs for research-grade devices in rats and could be developed into human devices in the future. 1741-2552/18/045002+17$33.00

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© 2018 IOP Publishing Ltd Printed in the UK

A Garcia-Sandoval et al

J. Neural Eng. 15 (2018) 045002

Keywords: bioelectronics, spinal cord stimulation, shape memory polymers, neural interfaces, electrochemistry, titanium nitride, paired plasticity S Supplementary material for this article is available online (Some figures may appear in colour only in the online journal)

Abbreviations

damage, and painful stimulation, and current devices have been found to migrate from initial implant location [10, 11]. New devices have been recently developed which are responsive to the constraints of stimulation in the rodent spinal cord. Courtine et al [12] used a soft polydimethylsiloxane (PDMS) SCS lead with platinum composite electrodes. These devices were implanted in the lumbosacral segments for up to six weeks maintaining stable electrochemical properties. Parylene-C SCS leads have also been studied to treat similar spinal cord injury conditions [13, 14]; however, chronic capabilities have not been demonstrated. Parylene-C, although it is a polymeric material, presents a mechanical mismatch of 6 orders of magnitude relative to the neural tissue [15]. PDMS is a soft elastic material that helps reduce such mechanical mismatch by 3 orders of magnitude relative to Parylene-C [16, 17]. Nevertheless, the low modulus of PDMS devices leads to implantation challenges as they become very difficult to handle and insert in the epidural space due to their lack of rigidity. However, in general, polymer-based neural interfaces are prefered over rigid implants mainly due to their higher flexibility [18]. Implantable bioelectronics is an active field with many competing approaches for chronically interfacing with the body. Much effort has gone into the field of softening electronics, pioneered by Capadona et al, with an inspiring study on the cellulose/poly(vinyl alcohol) composites which mimic the sea cucumber: stiff while dry and soft when implanted [19]. Our research group has carried on these efforts in softening bioelectronics, moving to characterize implantable devices made on different SMPs based on acrylates [20], thiols [21] and various photolithographic approaches to pattern novel structures on such polymers [22–24]. The electrochemistry at the neural interface itself and ensuring that the properties of this information exchange remain stable over chronic use points is also important for implants. In 2008, a descriptive overview of this space was presented [25], which detailed safe water limits, design metrics and helpful advice about a host of electrode materials including TiN, platinum, platinum–iridium, sputtered iridium oxide and electrodeposited iridium oxide. We chose TiN for this study because of its fractal morphology, its ease of processing via magnetron sputtering, and sufficient electrochemical properties, even at layer thicknesses as thin as 200 nm. Much of the design space and inspiration comes from studying the effects of different stimulation parameters, different materials, and different device configurations [26, 27]. To meet the demands of cervical SCS in rats, we sought to develop and characterize a softening polymeric SCS array. These arrays should be stiff at room temperature facilitating easy implantation by sliding the device in the epidural space following a single laminectomy. The implant should soften

SCS spinal cord stimulation SCI spinal cord injury PDMS Polydimethylsiloxane EIS electrochemical impedance spectroscopy CV cyclic voltammetry EMG electromyogram MEP motor evoked potentials VT voltage transients CSC charge storage capacity CIC charge injection capacity TiN titanium nitride PBS phosphate buffered saline RIE reactive-ion etching 1. Introduction Spinal cord stimulation (SCS) has been used for more than a half century with a focus on the mitigation of chronic back pain [1]. Recent studies have shown that using SCS is possible to treat not only pain but also other conditions, such as partial or complete paraplegia, tetraplegia, reflex sympathetic dystrophy and others [2–6]. Most SCS research conducted to date has been performed on the thoracic and lumbar segments where the epidural space is large enough for thick devices to be implanted, leaving stimulation of the cervical spine a much less studied area. Few research efforts have focused on cervical stimulation due, in part, to the smaller epidural space and the extensive movement and rotation of the neck in comparison with the thoracic or lumbar segments [7] which would induce large spinal deformation and strain to the implants and lead to premature failure due to device breakage. The limited cervical epidural space also limits the implant size, and specifically the thickness. Therefore, these implants need to be thin to fit within the epidural space and soft to prevent deformation and damage to the spinal cord [8, 9], putting constraints on device design. In fact, many standard materials may not perform adequately under these criteria because they are either too stiff, chronically, or too soft, initially, to allow for a successful implantation when the overall device measures only several hundred microns in thickness. Current devices are not well-suited for the demands of cervical SCS. Commercially available SCS leads are typically made of silicone rubber materials in paddle shapes with platinum–iridium electrodes and cobalt alloys as conductive wires. Even these semi-soft devices, however, have complications when they are implanted in the thoracic or lumbar spinal cord. These complications include epidural hematoma, neural 2


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once it is exposed to physiological conditions, and conform to the spinal cord allowing for a secure and effective interface and electrical stimulation. In this study, a comparison of the in vivo and in vitro performances of softening SMP-based SCS arrays is undertaken with a focus on electrochemical and physiological tests. An in vitro accelerated aging test was completed to simulate the amount of time the devices were implanted and estimate the lifetime of the softening SCS arrays. Devices were implanted in the cervical spinal cord of rats and their spinal threshold was monitored throughout 16 weeks to determine their stability and efficacy. Parallel to this study, Parylene-C SCS arrays were also fabricated, tested, and used as controls to determine the improvements of the softening arrays over Parylene-C. In this work, we demonstrate the ability to design, fabricate, test, and validate a robust and useful SCS array for use in small animal models that can survive multiple months and show stable stimulation thresholds.

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Figure 1. A 3D computer-aided design (CAD) model reconstruction of the cervical spinal column houses a softening spinal cord stimulation (SCS) array, implanted in the spinal canal with electrodes on the C5 and C6 segments from (a) side and (b) top down views. (c) An optical photograph shows an assembled, photolithographically defined, connected SCS array.

2. Methods We describe the underlying technical procedures to fabricate, deploy, characterize, and use the SCS array as shown from a side view and a top view in figures 1(a) and (b). A 3D computer-aided design (CAD) model reconstruction of the cervical spinal column houses a softening SCS array implanted in the spinal canal with electrodes on the C5 and C6 segments. Data to generate the 3D model were obtained from a CT scan of the rat spinal cord and column that was reconstructed to demonstrate the ideal insertion of our devices. The devices consist of polymers designed and synthesized described in section 2.1. Device electronics were built using photolithography following processes described in section 2.2. Materials and devices were characterized mechanically in section 2.3 while the electrodes themselves were optically and electrically characterized in vitro in sections 2.4 and 2.5, respectively. Devices were then implanted following procedures described in section 2.6 and characterized in vivo. Histological methods are described in section 2.7, while methods to establish the stability of the paired plasticity stimulation paradigm are described in section 2.8.

synthesis of the softening polymer has been previously published by Ware, et al [20]. Following synthesis, the solution was spin coated on a glass slide at an acceleration of 1000 RPM s−1 and a speed of 650 RPM for 60 s, yielding approximately a 50 µm thick film. The glass slides with the spin coated monomer solution were then placed in a UV oven and polymerized using 365 nm light for 1 h. The polymer was then post cured in a vacuum oven at 115 °C for 12 h. 2.2. Fabrication process

The fabrication of the softening SCS array (figure 1(c)) was performed using a glass slide as a carrier substrate and the softening polymer as the mechanical substrate with no special treatment or adhesion layer in a 10 000 class clean room using standard semiconductor fabrication tools and photolithographic processes. All fabrication process steps are shown in figures 2(a)–(l). Optical microscopy was performed throughout the fabrication process to verify success at each every step. A 300 nm gold layer, used for interconnects was deposited using a Temescal electron-beam evaporator. The electrode material, approximately 200 nm thick TiN, was deposited on top of the gold layer using a RF-sputtering system (AJA Magnetron 1500). Following TiN deposition, a 20 nm protective layer of aluminum was evaporated on top of the entire TiN layer. Lithography processes and wet etching were used to pattern the TiN electrodes, aluminum protective layer, gold interconnects, and connection pads. The lithography processes included deposition by spin coating of positive photoresist (MICROPOSIT S1813) on top of the metal layers. The spin coating parameters were 3000 RPM s−1 of acceleration and 2000 RPM of speed for 60 s. Then the photoresist was baked for 10 min at 85 °C. Afterwards, a mask aligner (Karl Suss MA6/BA6) and

2.1. Polymer synthesis and deposition

1,3,5-Triallyl-1,3,5-triazine-2,4,6(1H,3H,5H)-trione (TATATO), Tricyclo[5.2.1.02,6]decanedimethanol (TCMDA) and 2,2-Dimethoxy-2-phenylacetophenone (DMPA) were purchased from Sigma-Aldrich. Tris[2-mercaptopropionyloxy) ethyl]isocyanurate (TMICN) was purchased from Bruno Bock. All chemicals were used as purchased without further processing. TATATO (0.345 mol%) and TCMDA (0.31 mol%) were poured in a glass vial and mixed at 3000 RPM for 5 min. 0.1 wt% DMPA (photoinitiator) of total monomer solution was dissolved into the solution by mixing for 10 min. The vial was covered using aluminum foil to avoid exposure of the monomer solution to light. TMICN (0.345 mol%) was then added to the covered vial and mixed for 5 min. Detailed 3


J. Neural Eng. 15 (2018) 045002

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Figure 2. Renderings show the microfabrication process of the SCS arrays on a softening shape memory polymers (SMPs) integrating gold interconnects and titanium nitride (TiN) electrodes. (a) A carrier glass slide is (b) spin-coated with a 50 µm thick layer of co-monomers which are photo cured under 365 nm UV light into a smooth polymer film. (c) A 300 nm thick Au film is e-beam evaporated on the polymer, (d) subsequently coated with a 200 nm thick fractal TiN layer using an RF Magnetron Sputtering tool (e) and then protected with an e-beam evaporated Aluminum layer 20 nm thick. Positive photoresist, S1813, is spun onto the samples, patterned (using its corresponding mask) and developed. Then the whole sample is immersed in titanium nitride etchant resulting in (f) the 500 µm in diameter TiN electrodes. (g) The exposed gold layer is then treated with a similar photolithography process (different mask set) to pattern the Au interconnects. (h) Another 50 µm thick layer of co-monomers is spun and cured onto the samples as before and then (i) coated with a 150 nm thick silicon nitride (SiN) layer which is used as hard mask (protective layer). (j) Another photolithography process is done to pattern the SiN then the exposed SiN film is etched using a RIE (dry etch) process resulting in the hard mask that protects the device while (k) a following RIE oxygen plasma process removes excess polymer opening the electrode vias, wire connection pads and outline the SCS arrays. The removal of the excess polymer is followed by a SiN and Al wet etching process. (l) The resulting SCS array is removed from the glass carrier and ready for assembly, characterization, and use.

encapsulate the connection points to prevent leakage currents and short circuits. A similar fabrication procedure was used to fabricate the two previous generations of SCS arrays shown in figures S7(a1) and (a2). These previous generations of arrays had different dimensions (they were longer, exposing more of the thin-film electrode connector to the mechanical stresses inside the body) than our most successful (third generation) SCS arrays. The first generation arrays were fabricated utilizing a thinner softening polymer substrate (25 µm) and were encapsulated with a 500 nm thick Parylene-C layer. Also, a different connector was fixed to the gold pads during array assembly. The second generation arrays were fabricated on a 25 µm softening polymer substrate and then encapsulated with a second 25 µm softening polymer layer.

the corresponding photomask were used to pattern the photoresist by exposure to UV light (g-line = 435 nm). Once patterned, the samples were immersed in a developer solution (MF-319) to dissolve the exposed portion of the photoresist. Then, the samples were immersed in the corresponding wet metal etchant dissolving the uncovered metal or ceramic. A 50 µm second layer of softening polymer was spin coated and cured, encapsulating the metal patterns. A 200 nm SiN layer was deposited by plasma enhanced chemical vapor deposition (PECVD) and used as hard mask to allow for opening vias, electrode sites, and to outline the final devices. Once this film was patterned, using a similar lithography process as the one previously explained, a reactive-ion etching (RIE) process of oxygen plasma was used to etch the exposed softening polymer resulting in the finished softening SCS arrays. A dip in diluted hydrofluoric acid (20:1) removed SiN and Al coatings. Devices were then removed from the glass slide and assembled (figure S1(a) (stacks.iop.org/JNE/15/045002/mmedia)) using polytetrafluoroethylene (PTFE) insulated stainless steel wires and a 2-channel connector. The wires were purchased from A-M Systems (Sequim, WA, USA) and the connectors from Plastics One (Roanoke, VA, USA). A conductive silver epoxy (Electron Microscopy Sciences, Hatfield, PA, USA) was used to fix the wires to the SCS arrays and to the two channel connector. Once all parts were soldered together, silicone epoxy was used to

2.3. Mechanical characterization of materials and devices

The softening polymer was mechanically characterized using two different tools: a universal tensile machine (UTM) with a BioPuls temperature controlled liquid bath (Instron) and a dynamic mechanical analyzer (DMA) with a temperature controlled liquid bath (TA instruments RSA-G2). The softening samples were prepared by depositing and curing a 50 µm softening monomer solution on a glass slide. Then, a SiN film was 4


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deposited by PECVD on top of the polymer, patterned with a lithographic process, and used as hard mask for the polymer RIE plasma etching process to cut out the samples. For the DMA tensile tests, the samples were actual softening SCS arrays (100 µm thick) and for the UTM tensile tests, the softening polymer and Parylene-C samples were plasma etched into a dog bone (Type V from D638-10) shape and tested following the ASTM standard D638-10 [28]. The UTM was used to perform quasi-static tensile tests utilizing a strain rate of 0.01 s−1 while the DMA was used to perform dynamic tensile tests at 1 Hz in both dry and wet conditions. Figures S3(a) and (b), show three different tensile tests performed with the UTM. Parylene-C and softening polymer samples were tested in dry conditions at room temperature (~24 °C), and other softening polymer samples were tested in PBS at 37 °C. The softening samples tested in PBS were placed in a vial full of PBS at 37 °C for 24 h prior to the test to allow the polymer to soak up PBS and plasticize. The dynamic mechanical analysis tests (figures S4(a) and (b)) were performed utilizing the RSA-G2 tool with its immersion bath in tension mode to measure the storage modulus (E′) and tan δ of the softening rectangular samples in wet and dry conditions. The clamping distance used was 15 mm with a preload force of 0.2 N, a frequency of 1 Hz, and a deformation ampl itude of 0.275% strain. Dry experiments were run from 20 °C to 100 °C using a heating rate of 2 °C min−1. Soaking experiments were run using the immersion system of the RSA-G2 filled with PBS. The samples were clamped and the immersion chamber was filled with PBS. In the first step, the PBS was heated to 37 °C using a hotplate before being poured in the DMA chamber followed by an isothermal oscillation for 60 min at 37 °C. In the second step, the PBS was cooled down to the starting temperature of 20 °C at a rate of 3 °C min−1, and then the temperature was increased to 75 °C at a rate of 2 °C min−1. After the fabrication of the softening SCS arrays, they were prepared for a tensile test, figure S4(c), using the DMA tool (RSA-G2) instead of the neat polymer samples which were tested in figures S3(a), (b) and S4(a), (b). Three different tests were performed on the fully fabricated devices: in dry conditions at 37 °C, in PBS at 37 °C with not soaked samples and, in PBS at 37 °C using samples that were soaked for 1 h prior to the test. The clamping distance used was 10 mm for the quasistatic tensile test (0.01 s−1 strain rate).

force microscopy (AFM) was performed to scan through the surface of the TiN electrode to analyze its morphology (figure 3(d)) and determine its roughness. AFM was performed using a Veeco Dimension 5000 SPM system in tapping mode at 1 Hz in a 1 × 1 µm area. Electron microscopy of explanted devices was performed using a Zeiss EVO environmental SEM. 2.5. In vitro electrode electrochemical characterization

An electrochemical workstation (CH Instruments CHI660D Potentiostat/Galvanostat) was used to perform basic electrochemical characterization, such as cyclic voltammetry (CV) and electrochemical impedance spectroscopy (EIS). A threeelectrode setup was used to perform the tests in which a platinum wire serves as the counter electrode, a Ag|AgCl as the reference electrode, and the TiN electrodes immersed in the air-equilibrated phosphate buffered saline (PBS) as a working electrode. The in vitro analysis was done in PBS at 37 °C with a pH of 7.14. CV was measured at a 50 mV s−1 sweep rate between potential limits of −0.8 V and 0.8 V, remaining in the safe water electrolysis voltage window for TiN. EIS was performed to deliver 5 mV RMS sinusoidal signal to the working TiN electrodes between frequencies of 1 Hz to 100 kHz, and the magnitude and phase impedance were measured to determine the electrode properties. A Sigenics monophasic stimulator and a Tektronix TPS2014B oscilloscope were used to conduct voltage transient (VT) tests. VT measurements were taken to find the charge injection limits of the TiN electrodes. Similar to the CV and EIS tests, a Pt wire and Ag|AgCl were used as counter and reference electrodes, respectively. During VT testing, cathodal current pulse widths between 100 µs to 500 µs were applied to the TiN electrodes at 50 Hz frequency. At the end of each pulse, there was a 400 µs interphase delay when the electrode polarization was monitored, after which the electrode was returned (pulled) to zero potential by a voltage-controlled anodic discharge phase. In order to calculate maximum charge injection capacity, the level of current pulse was slowly increased manually until a maximum electrode polarization was recorded at −0.9 V (Emc). To estimate the lifetime of the softening SCS arrays, five devices (ten electrodes) were subjected to an accelerated aging test by immersing them in a vial filled with PBS. The vial was then placed inside an oven at 75 °C for up to 350 h. Electrochemistry tests were done at 37 °C every 10 h of aging. According to equation (1), every 24 h of aging time simulates two weeks at in vivo conditions [29].

2.4. Electrode microscopy

Optical, scanning electron, atomic force, and magnetic resonance microscopy pictures were obtained to assess the physical properties of the spinal cord stimulators and electrodes. Figure 1(c), taken with a Canon T4i DSLR, shows the assembled device. A Leica DM4000M LED optical microscope was used to analyze the finished arrays up close. Scanning electron microscopy (SEM) of the electrode surface (figure 3(c)) was performed using a Zeiss Supra 40 SEM with a resolution capability up to 2 nm. The SEM micrographs were obtained in vacuum using an electron acceleration potential of 5 KV, a magnification of 200 KX, and a working distance of 5.4 mm. Atomic

∆T/10 (1) t37 = tT × Q10 .

Here, ΔT = T − 37 °C, where T is the experiment temperature and tT represents the duration that the samples were exposed to the experiment temperature. An aging factor (Q) of 2.0 was used as a conservative measure to test polymeric devices and polymers in general. Therefore, in 350 h, the samples were aged for a total of 29 simulated weeks. CV tests with three different scan rates, 50 V s−1, 5 V s−1, and 50 mV s−1, were performed alongside EIS tests to characterize the performance of the aging devices throughout the test. 5


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Figure 3. Optical pictures express detail at 20× magnification of the SMP-based SCS array showing (a) the serpentine 100 µm wide Au interconnects and (b) one of the device’s two high charge injection capacity TiN electrodes. (c) A scanning electron micrograph (SEM) shows the inherently rough, nanostructured TiN surface that enables large electrochemical surface area per geometric surface area. (d) An atomic force micrograph (AFM) image quantifies average surface roughness of the TiN coating as 2.83 nm root mean squared (RMS). SCS arrays are characterized electrochemically to benchmark performance before implant and aging studies: (e) cyclic voltammetry (CV) shows a characteristic TiN electrode curve performed at 50 mV scan rate in PBS 1× using a Ag|AgCl reference electrode and Pt wire as a counter electrode. Cathodic charge storage capacity (CSCc) is 1.81 mC cm−2; and (f) electrochemical impedance spectroscopy (EIS) measures impedance and phase angle as a function of frequency of the TiN electrodes. Impedance of 764 ohms at 1kHz and between −60° and −90° phase lag suggests capacitive behavior at low frequencies as expected. (g) Voltage transient signals in response to (h) current pulses of varying width are measured to investigate charge injection capability of the TiN electrodes. Monophasic cathodal current pulses resulting in a maximum electrode polarization (Emc) of −0.9 V are presented in the figure. While the shortest pulse (at 100 µs pulse width) allows the highest current, its charge density is the smallest compared to larger pulse widths up to 500 µs. (i) Charge injection capacity (Qinj) with Emc limit of −0.9 V and electrode bias at 0 V is plotted as a function of pulse width, as calculated from the results in (g) and (h).

2.6. Device implantation and in vivo characterization

CT, USA) to maintain body temperature of 37.5 °C as measured continuously by a rectal probe (PhysioSuite). In a single procedure, motor cortex screw electrodes and EMG wires were placed, and the head connectors were brought to the common head cap. Rats were anesthetized and head-fixed in a stereotaxic frame (David Kopf Instruments, Tujunga, CA, USA). The skull was exposed and burr holes were made using a Jobber Drill (number 60, Plastics One, Roanoke, VA, USA) without disturbing the dura mater. Stainless steel screw electrodes (1.19 mm diameter with a flat tip; Plastics One) were implanted over the forelimb area of the motor cortex in one hemisphere at two locations as described in the literature [30, 31]. The screw electrodes were attached in advance to a head connector (Plastics One) that was secured with skull screws and dental acrylic. To measure cortical motor evoked responses (MEPs), we inserted stainless steel braided wire (Cooner Wire Co., Chatsworth, CA, USA) electrodes into the biceps muscle to record an EMG signal. The other end is tunneled subcutaneously and attached in advance to a head connector, mounted on the skull, and

Adult female Sprague-Dawley rats (Charles River, body weight of 275 ± 25 g, n = 32) were used for the in vivo electrochemical and physiological tests. The animals were housed in individual cages with free access to food and water on a 12/12 h light/dark cycle. All experimental procedures were in full compliance with the approved Institutional Animal Care and Use Committee protocol of Weill Cornell Medical School. Rats were perfused after two weeks of implantation with 150 ml cold 0.1 mol l−1 PBS followed by 400 ml 4% paraformaldehyde in PBS for CT/MR imaging or at the end of the study for histological analysis. For all surgical procedures, rats were anesthetized via an intraperitoneal injection of a mixture of ketamine (90 mg kg−1) and xylazine (10 mg kg−1 body weight). −1 Buprenorphine (0.05 mg kg body weight) was administered before and after the survival surgery to alleviate pain. Anesthesia levels were monitored by respiration and heart rate and responses to foot pinch. Animals were placed on heating pads (PhysioSuite, Kent Scientific Corporation, Northwest 6


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Figure 4. Experimental timeline. Type of arrays are shown on left, with information about softening arrays above the timeline and Parylene C below the timeline. After two weeks of implantation one rat in each group was used for CT & MRI (figures 1(a), (b) and 6(a) and S4(a)). One rat in the softening group was used for ex vivo testing after four weeks (figure 5). In Parylene-C group 3 arrays failed at week 6, 8 and 12, and the spinal cord tissue used for histology (figures S4b1-3 and c1-3). Four devices in the softening group failed at week 10, 16,18 and 22, and the spinal cords were used for histology (figures 6, b2-5 and c2-5). One array failed at 22 weeks in the Parylene-C group. In the softening group, one of the array was active until 29 weeks; this was removed while it was active.

with 150 ml cold 0.1 mol l−1 PBS followed by 400 ml 4% paraformaldehyde in PBS. To visualize the SCS arrays implanted over the spinal cord, we performed ex vivo magnetic resonance imaging (MRI) of the spinal column with the array in situ in two rats, one with a softening array and other with Parylene-C array. After perfusion, the wires from the array connector to the head were removed. The cervical and upper thoracic spinal column with array electrodes in place and postfixed overnight in 4% paraformaldehyde in PBS and then left for 1 d in PBS at 4 °C. The spinal column was not dehydrated to limit the tissue shrinkage postfixation. The fixed spinal column was placed into 10 mmwide NMR tubes (Wilmad, Buena, NJ, USA) filled with nonprotonic solvent (Fluorinert Liquid FC-40, 3M Science, St. Paul, MN, USA). Air bubbles were removed from the tube. The sample was kept at 20 ± 1 °C during imaging. MRI was performed with a 7T Varian system. 2D images were acquired using an ultra-short echo time (UTE) imaging parameters with a field of view (FOV) of 12.5 × 12.5 mm and an acquisition matrix of 256 × 256. The spatial resolution was 49 × 49 µm in plane with a section thickness of 490 µm. A total of 15 sections were acquired in the axial plane, centered on the array. To investigate the 3D morphology of the spinal cord with implanted SCS arrays, the samples were scanned using the IVIS Spectrum CT scanner. The x-ray tube scanning parameters were set to 30 KV (peak) and 500 µA with a voxel resolution of 15 µm. Multiple projection images were obtained with a range of 360° of rotation of the sample. The exposure time was established as 7000 ms, and the total time to acquire the entire image was approximately 30 min. All of these images were reconstructed using slice reconstruction software to obtain the 3D images.

also secured with skull screws and dental acrylic (Lang Dental Manufacturing Co., Inc., Wheeling, IL, USA). In a second procedure, the arrays were implanted. The T1 spinous process was exposed and clamped to stabilize the spine. The C4 spinal cord segment was exposed by a laminectomy. The tip of the array (furthest from the connector) was introduced into the epidural space at the laminectomy site and advanced caudally until the connector was positioned over the C4 laminectomy site. The implantation procedure is shown in the supplementary movie S1. We ensured that the two electrodes were placed in the middle of the spinal cord over C5–C6 by observing the electrode position between the vertebrae. The array head connector was tunneled subcutaneously and secured to the skull with skull screws and dental acrylic. Muscles and skin were sutured in layers. A similar electrochemical setup was used to perform the in vivo electrochemical characterization, CV and EIS, as described in method section 2.4 related to the in vitro work. A three-electrode setup was used to perform these tests: a platinum wire serves as the counter electrode; a stainless steel needle electrode (BD needle #305106—0G × 1/2 in Becton, Dickinson and Company, Franklin Lakes, NJ, USA) serves as the reference placed subdermally on the trunk, and the TiN array electrodes placed on the C5–C6 spinal cord serve as the working electrode. CV was conducted at a 50 mV s−1 sweep rate between potential limits of −0.8 V and 0.8 V remaining in the safe water voltage window for TiN. EIS was performed to deliver 5 mV RMS sinusoidal signal to the working TiN array electrodes between 1 Hz to 100 KHz. First and second generation devices were tested daily for spinal thresholds and the time to failure was measured in days. The timeline of the in vivo experiments with the third generation devices is shown in figure 4. Above the timeline are the times that were examined for the softening arrays and below the line are the times for the Parylene-C arrays. Gray arrows indicate devices that failed and were explanted for inspection. At these times, the animals were perfused and the spinal cords removed for histology was performed. Rats were perfused

2.7. Histological analysis of spinal cord

Spinal cord tissue sections were analyzed for astrogliosis and inflammation. At the times indicated in figure 4, rats were perfused and the spinal cords were removed, post-fixed overnight 7


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in 4% paraformaldehyde in PBS and left for 1 d in PBS containing 30% sucrose. Axial 40 µm thick sections of the spinal cord spanning from C4 to T1 area were cut with a cryostat As a positive control, we included a rat with a chronic spinal cord injury. The spinal cord injury was produced by contusion of the C4 spinal cord using a 200 kdyn force applied through a 3 mm tip using the Infinite Horizons impactor. Eight weeks after spinal cord contusion, the rat was perfused and histology was performed. This animal did not receive a SCS array, only a spinal contusion. For immunohistochemistry, the following primary antibodies were used on free-floating sections: chicken-antiGFAP (for astrogliosis; # AB5541, Chemicon, Hofheim, Germany, at 1/500), and mouse anti-rat monocytes/macrophages (clone ED1; # MAB1435, Chemicon, Hofheim, Germany, at 1/300). The sections were blocked in PBS +3% donkey serum + 0.1% Triton-X, and incubated with the primary antibody in PBS + 3% donkey serum + 0.1% Triton-X overnight at 4 °C on a rotating platform. The following day, sections were incubated with the secondary antibodies (# A-11004; Goat anti-Mouse IgG (H + L) Cross-Adsorbed Secondary Antibody, Alexa Fluor 568 for ED1; 1:500, and # A11039; Goat anti-Chicken IgY (H + L) Secondary Antibody, Alexa Fluor 488 for GFAP; 1:1000 Thermo Fisher Scientific, Waltham, MA, USA). Sections were mounted onto gelatin-coated glass slides, air-dried, dehydrated, and coverslipped with Fluoroshield histology mounting medium (Sigma-Aldrich, St. Louis,MO, USA). A Nikon eclipse Ni microscope (Nikon Corp. Shinagawa, Tokyo, Japan) was used to perform the histologic analysis.

were quantified. First, the motor cortex was stimulated at varying intensities to determine the cortical stimulation threshold needed to evoke a short-latency MEP. Then, raw EMG signals were collected across 20 trials at 110% of cortical threshold intensity. MEPs were extracted, processed and quantified using a customized MATLAB (MathWorks, Natick, MA, USA) script. The area under the curve (AUC) was calculated for averaged MEPs. We measured the effects of subthreshold cervical SCS (90% of threshold) on cortical MEPs (110% of threshold), when SCS was delivered 10 ms after motor cortex stimulation for maximum augmentation of MEPs as described in Mishra et al [31]. The statistical analysis was done with SPSS (Version 22). Analysis of variance (ANOVA) was used to test changes in paired stimulation efficacy over time. 3. Results Using standard semiconductor fabrication processes, we developed and fabricated a softening SCS array with TiN microelectrodes capable of stimulating the cervical spine for long periods of time in a safe manner. We characterized the SCS array in vitro and in vivo for multiple weeks and observed changes in electrochemical properties and physiology of the underlying tissue. 3.1. The TiN electrodes: geometrical, morphological, and electrochemical properties

Figure 3 shows TiN electrodes at varying resolution (optical, SEM, AFM) and quantifies their electrochemical properties through EIS, CV and VT measurements. Figures 3(a) and (b) show portions of the fully fabricated SCS array at 20× magnification. The Au serpentine traces in figure 3(a) are designed to minimize stress concentrations on the low strain capacity metals and geometrically accommodate these stresses and strains during utilization of the array in mechanically constrained environments. Figure 3(b) shows the full diameter of the 50 µm deep etched opening through the SMP that exposes the 200 nm thick TiN electrode and enables current flow from the SCS array into the surrounding environment. The SEM image in figure 3(c) provides a top-down morphological representation of a small portion of one of the high-surface area, TiN electrodes, including its sub-20 nm pyramidal surface features. An AFM image shows a 1 µm2 portion of the TiN surface morphology, figure 3(d), from which the root mean square (RMS) roughness was calculated to be 2.83 nm that is directly related to the electrochemical surface area (ESA) of the electrodes. Figures 3(e)–(i) show electrochemical properties of one of the TiN electrodes from a representative softening SCS array. CV of one representative TiN electrode is shown in figure 3(e). Current density measured between −200 and 200 µA per cm2 is plotted against the potential between the TiN electrode and the Ag|AgCl reference electrode from −0.8 V to 0.8 V, which was chosen with a 0.1 V safety factor from the safe water electrolysis window of TiN of −0.9 V to 0.9 V, as is documented in the literature for TiN [25, 32]. This curve measured at a voltage scan rate of 50 mV s−1 allows

2.8. Electrophysiological testing of electrode stability

Spinal cord and motor cortex physiology was performed on awake and unrestrained rats. To test stability of the implanted SCS arrays we measured activation threshold of the biceps muscle in response to stimulation (known as the spinal threshold). A single biphasic pulse was applied between the two electrodes positioned on the dorsal surface of the cervical spinal cord in midline. The biphasic square pulse was 0.2 ms for each polarity with no interphase delay (the second phase immediately following the first). Stimulation intensity was adjusted to determine the spinal threshold for provoking a short-latency motor-evoked potential (MEP). The motor cortex threshold was similarly determined as the minimum intensity for provoking a MEP after brain stimulation. The spinal and cortical thresholds were measured at least three times per week for 16 weeks (n = 4) in rats with softening SCS arrays and for eight weeks in rats implanted with Parylene-C SCS arrays (n = 4). The magnitude of the neuromodulatory effect of SCS on motor cortex MEPs was used to determine the the efficacy of SCS. This paradigm is identical that described in Mishra et al [31]. The paradigm was designed to cause motor cortex stimulation and dorsal SCS to arrive synchronously in the cervical spinal cord. A train of three biphasic square wave pulses was used for motor cortex stimulation and EMG was continuously acquired using an integrated electrophysiology system with a differential AC amplifier system. Cortical motor potentials 8


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Figure 5. Electrochemical characterization of SMP-based SCS array was performed during both ((a)–(c)) in vitro accelerated ageing studies where devices were soaked in PBS 1× at 75°C up to 350 h simulating 29 weeks and during ((d)–(f)) implantation at various time points in vivo up to six weeks. EIS quantifies impedance and phase angle across 29 weeks (accelerated ageing) and six weeks (true ageing) as a function of frequency each from one representative array from more than ten electrodes characterized in this manner. CV quantifies current as a function of voltage bias in accelerated and true ageing environments. Fig. 5d looks saturated due to a measuring equipment limitation. The maximum current that could be measured was set to ±1 µA.

calculation of the cathodal charge storage capacity (CSCc) of the TiN electrodes via the time integral of the cathodal current from the cyclic voltammogram. The CSCc is 1.81 mC cm−2. In figure 3(f), EIS quantifies the impedance magnitude and phase angle of the softening SCS array, and correspondingly of the TiN electrode, which due to its size, contributes the most heavily from the interaction between the SCS array and the electrochemical cell, to the measured values across the frequency range of 1 Hz to 1 MHz. The magnitude impedance at 1 KHz was 1000 ± 200 Ω for all the devices tested including the representative array in figure 3(f). The phase angle indicates resistive behavior, near 0° phase lag at 10 kHz and shows increasing capacitive behavior below 10 KHz reaching above −80° phase lag below 10 Hz. Figure 3(g) presents the potential excursion of the TiN electrode referenced to a Ag|AgCl electrode, in response to monophasic current pulses of pulse widths from 100 to 500 µs presented in Figure 3(h). VT measurements enable determination of maximum charge injection capacity (Qinj) of stimulating electrodes, which is shown in figure 3(i) for a representative TiN electrode from the SCS array. As the pulse width varied from 100 µs to 500 µs in 100 µs increments, Qinj varied: 0.125, 0.151, 0171, 0.179, 0.183 mC cm−2 respectively. In figure 5, we show the in vitro (a)–(c) electrochemical behavior of the accelerated aged (75 °C in PBS 1×) TiN electrodes on the SCS arrays to a simulated in vivo performance of 29 weeks. CVs of one representative aged TiN electrode are shown in figure 5(a). The current measured between −400 and 180 nA, −2.68 and 1.68 µA, and −2.18 and 1.05 µA in the initial (black), intermediate (gray) and final (red) representative CVs, respectively, are plotted against the potential between the

TiN electrode and the Ag|AgCl reference electrode from −0.8 to 0.8 V. The gray intermediate lines track representative weeks between week 1 (black) and week 29 (red). All measurements of the aged devices show an increase of the current measured and their CSC. For the EIS measurements across this study, impedance and phase angle represent the properties of the biotic-abiotic interface along the implantation tracking any change in capacitance of the TiN electrode or an increase of the resistive component from the tissue [25]. In figure 5(b), the impedance magnitude of a representative aged TiN electrode on a softening SCS array is plotted versus the frequency (both logarithmic). The impedance magnitude of the TiN at 1 kHz is 1.3 kΩ initially (black) and increased up to 2.3 kΩ for the 29 week test (red) with intermediate (grey) measurements ranging between 1.1 and 3.6 kΩ at 1 kHz which shows the impedance magnitude was maintained low and stable. In figure 5(c), the impedance phase angle measurements during the accelerated aging test of the representative TiN electrode are plotted versus frequency. At 1 kHz, the initial (black) measurement showed a phase angle of 45°, while intermediate measurements ranged between 44° and 60° and the final measurement had a phase angle of 52°. However, the phase angle at lower frequencies (1 Hz) became less capacitive, from −77° to −56°. 3.2. In vivo and ex vivo electrochemistry of TiN electrodes on an SCS array

In figure 5, we also show the in vivo (d)–(f) behavior of a representative TiN electrode on a softening SCS array that was implanted in the cervical spinal cord of a rat for six weeks. The in vivo CV and EIS tests are shown in black for the in 9


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(a)

(b)

(c)

(d)

(e)

Figure 6. Electrochemical characterizations via (a) CV and (b) EIS quantify implantable SCS array behavior in vitro, in vivo after 30 d implanted, ex vivo before cleaning and ex vivo after cleaning. Tissue attached to the electrodes as seen in SEM images ((c) and (d)) is hypothesized to be the culprit for the shifts in electrochemical properties in vivo, which could also explain (e) performance of the explanted device, as seen in an SEM image. Electrochemistry shows that device performance ex vivo is similar to initial in vitro performance indicating biotic mechanisms, not device materials’ failure, as the primary drivers of electrochemical changes across the first 30 d after implantation.

vitro (before implantation), gray for weeks 0 to 5th and lastly week 6 in red. In figure 5(d), the CV measurements of one representative TiN electrode are plotted. The current measured between −348 and 277 nA, initially in vitro, is plotted against the potential between the TiN electrode and the Ag|AgCl reference electrode from −0.8 to 0.8 V. Also, the implanted, intermediate and final, CV measurements (grey and red) are plotted against the potential between the TiN electrode and a stainless steel needle used as the reference electrode. The CV curves of weeks 0 to 6 are truncated due to electrochemical workstation settings limiting current delivered to the living animal to ±1 µA. The CVs in vivo show similar behavior to the in vitro tests in which the resistive component of the environment becomes more dominant. Such changes are more noticeable in the in vivo tests. In figure 5(e), the impedance magnitude of one representative TiN electrode is plotted versus the frequency. The impedance magnitude at 1 kHz (physiologically relevant frequency [33] is approximately 1 kΩ then increased to 14 kΩ in week 6 of implantation with intermediate measurements ranging between 2.8 and 7.5 kΩ. Again, the shifting impedance spectra (upwards) show that the in vivo environ ment is becoming more resistive. Figure 5(f) shows the in vivo TiN electrode phase angle versus frequency. In vitro, the measured phase angle measured 6° (near-resistive) at the frequency range between 10 to 100 kHz. At 1 kHz, the phase turns capacitive, about −40°, and maximized at 10 Hz to 1 Hz having almost an 80° phase shift. The phase angle remained between −40° and −60°at 1 kHz during all the measurements. However, the phase angle decreased from −80° to about −25° in the last measurement at 1 Hz showing a similar change to the aged arrays but even more drastic. Figure 6 describes both the electrochemical and SEM characterization of one of the SCS arrays that was implanted for 30 d in the cervical spinal cord. The TIN electrodes were 10

covered with tissue after explant. Upon explantation, electrodes were cleaned with tergazyme to remove tissue growth on the TiN electrodes. Figures 6(a) and (b) show the CV and EIS measurements of a newly assembled SCS array (black), in vivo after 30 d implanted (red), after explantation (green), and after cleaning of the electrodes (blue). In figure 6(a) the CV measured currents are plotted against the potential between the TiN electrode and the Ag|AgCl reference electrode and a stainless steel needle, for the in vitro and in vivo tests respectively. The measured current in vitro is between −127 and 74 nA, between −241 and 330 nA in vivo, −400 and 189 nA for ex vivo, and finally −400 and 117 for the cleaned TiN electrode. Figure 6(a) shows how the implantation and tissue growth on the electrode affects the array electrochemical performance. Figure 6(b) shows both the impedance magnitude (top) and phase angle (bottom) of the representative TiN electrode versus frequency. The impedance magnitude for all the in vitro tests is approximately the same along all frequencies and at about 1 kΩ at 1 kHz. The impedance magnitude of the in vivo test increased to about 10 kΩ at high frequencies from 100 Hz and above. Figures 6(c) and (d) demonstrate the organic tissue filling the opening of the encapsulation for the TiN electrodes. Figure 6(e) is a SEM picture of the explanted SCS array attached to a silicon wafer piece used as conductive base. It is noticeable that the SCS array is covered with tissue after initial explant and that impedance returns to pre-implant levels after explant, but is almost one order of magnitude higher after 30 d implanted. 3.3. Comparison of softening and Parylene-C SCS arrays

The SMP and Parylene-C based SCS arrays were designed to allow long-term stimulation of the cervical spinal cord in awake behaving rats. A group of animals with implanted


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Figure 7. Safety, stability and efficacy are demonstrated by the SMP-based SCS arrays. (a) Ex vivo T2-weighted magnetic resonance imaging (MRI) of the cervical spinal cord implanted with softening array at two weeks after implantation shows that the softening array takes the shape of spinal dorsal surface. Immunohistochemistry of cervical spinal cord stained for (b) astrocyte marker GFAP and (c) macrophage marker ED1. The softening SCS arrays sometimes deform the shape of the cord very little (animal 3, (b4)) and sometime more dramatically (animal 4, (b5)), but create less disturbance than the stiffer Parylene-C arrays (see Supplementary Information). Scale bar = 1mm.

softening SCS arrays was allowed to survive for 16 weeks. During this period, we tested: (i) how the implanted arrays were tolerated by animals, (ii) durability of the arrays, (iii) stability of spinal cord evoked responses, and (iv) how effective the arrays were in chronic neuromodulation over time after repeated stimulation. Our results show that most of experimental animals tolerated the invasive implantation well, leading to stable thresholds throughout the whole survival period. In none of the animals did the implantation cause noticeable neurological changes; in general, the overall locomotion remained unaffected. Softening SCS arrays conform to the spinal epidural surface post implantation: an axial scan via MRI through the ex vivo spinal cord to visualize the spinal column shows that the softening SCS array conformed to the spinal dorsal surface instead of flattening the column. A clear distinction between the typical butterfly appearance of the spinal cord gray matter and the surrounding hypointense white matter can be seen. Figure 7(a) shows the spinal cord with an implanted softening SCS array which has molded the cord post-implantation and has taken the shape of the spinal dorsal surface. The cord is false highlighted in yellow and the device in red. The spinal cord with an implanted stiff Parylene-C array (colored in red) is deformed into a flatter shape on top of spinal dorsal surface as shown in supplementary figure S5(a). To determine if the SCS arrays caused inflammation in the underlying spinal cord, we performed histological analysis on spinal cord sections of five different animals, one without an implant (control) and four others, each with an SCS array implant. The performance of our SCS arrays was analyzed using immunostaining of spinal cord tissue sections using two markers: (1) glial fibrillary acidic protein (GFAP) to determine if there was astrogliosis, and (2) ED1, which is a pan-macrophage marker. We also examined the shape of the spinal cords that were processed for histology to examine for underlying compression of the spinal cord. Figures 7(b) and (c) demonstrates safety via a lack of compression of the dorsal surface of spinal cord and limited observable tissue reactions to two of the implanted softening array electrodes (figures 7(b2), (b3) and (c2), (c3)), and slight compression to

the dorsal surface of the spinal cord and no injury/inflammation (figures 7(b4), (b5), (c4) and (c5)) post implantation with the other two SCS arrays. The staining appears similar to what we observed with spinal tissue with no implant (figures 7(b1) and (c1)). Similar results were observed in spinal cord tissue implanted with Parylene-C arrays with slightly more compression (supplementary figures S7(b1)–(b3) and (c1)–(c3)). For comparison, a spinal cord that was purposefully injured shows intense tissue reaction for both GFAP and ED1 antibodies; supplementary figures S7(b4) and (c4) respectively. 3.4. In vivo stability and efficacy of TiN electrodes on SCS arrays

In all, we tested three different generations of our chronic arrays. SCS arrays from the 3rd generations were more durable, stable and effective in neuromodulation. One of the 3rd generation softening arrays remained active for up to 29 weeks with a stable stimulation thresholds below 1 mA, as shown in figure S9. First generation (n = 8) and second generation (n = 9) SCS arrays were patterned such that in excess of 30 mm from the electrodes were patterned on the flexible substrate as seen in (figures S7(a) and (b). As clearly shown in supplementary figure S7(a1), we started with these arrays with longer stretch of SMP or Parylene-C materials throughout the connector; these arrays were 50 µm thick and the two electrodes were 5 mm apart that covered more than two segments of spinal cord. Post implantation these devices failed very early within approximately 10 d as shown in figure S7(b). Post ex-implantation it was observed that they sheared from two major regions: one at the point where they enter inside the spinal canal; and the other at the head connector point (figure S7(a3)). To overcome these failures we increased the SCS array thickness (100 µm) in our future generations of the arrays (figure S7(a2)) and also decreased the inter electrode distance to 3 mm. These modifications did not achieve chronic time points as the second generation arrays failed after 3 weeks of implantation (figure S7(b)). When we analyzed ex-implanted arrays, we observed that they sheared similarly as our first generation arrays (figure S7(a3)). For the 11


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Figure 8. (a) The electrode configuration and experimental setup is shown. Screw electrodes are placed over motor cortex, and the SCS arrays are placed in the epidural space between C5 and C6. Electromyogram (EMG) electrodes measure responses in the biceps brachii muscle on the rat’s forelimbs. (b) Spinal stimulation intensity is adjusted to determine the threshold for provoking a short-latency motor evoked potential (MEP) in >50% of trials. In four rats, spinal thresholds remain stable and below 600 µm throughout 16 weeks. (c) Thresholds to evoke motor response are determined via the cortical electrodes alone and then used at 90% of the threshold current such that coupled threshold currents to evoke forelimb motor response can be measured from the SCS arrays. The effect of pairing suprathreshold motor cortex stimulation and subthreshold spinal cord stimulation compared to motor cortex only stimulation yields large enhancement at lower, stable thresholds for 16 weeks.

third generation of arrays, we reduced the total size of photolithographic region to less than 25 mm and introduced a wire attachment which leads to the head connector (figure S7(a4)). After in vivo CV and EIS measurements, arrays were tested for SCS thresholds for at least 16 weeks post implantation in awake behaving rats (n = 4; figure 8(b)). The threshold for spinal evoked responses increased significantly in the first two weeks after implantation (F = 10.108, p ⩽ 0.003). Results after that threshold were very consistent and stable over 16 weeks (p ⩽ 0.113) in all four rats. We also tested the impedance of these arrays in vivo at 1 KHz. As seen in figure S8, the impedance increased in the first two weeks after implantation and remained stable over the following 14 weeks in all four rats. Further, these arrays were utilized in a paired plasticity experiment to determine post implant electrical stability following a representative use case experiment. The goal was to measure the chronic stability of cervical SCS threshold in uninjured rats. The experimental setup is shown in figure 8(a). Electrodes over motor cortex were used to evoke motor responses in the biceps muscle. The motor threshold to stimulation through the cortical screw electrodes was tested across 16 weeks. These thresholds were very consistent throughout, as shown in figure S10. In other experiments, the impedance of cortical screw electrodes was measured at the time of implantation and also after up to 14 weeks, and there was no difference (data not shown). The motor evoked potential curves of motor cortex stimulation alone (no spinal stimulation) were compared against the responses to paired motor cortex and SCS. In awake behaving rats with implanted arrays, we tested the immediate effect of paired motor cortex and cervical SCS over 16 weeks post implantation to assess the stability of these motor evoked responses over time. In the paired stimulation experiments, the spinal cord was stimulated at the C5–C6 spinal level with an intensity that was 90% of the spinal threshold for provoking a biceps motor evoked potential and cortical stimulation at 110% of cortical threshold intensity. There was no significant difference in cortical motor evoked potentials

augmentation post implantation till 16 weeks (F = 1.424, p ⩾ 0.294) and the motor evoked potential remains consistent and stable post pairing (figure 8(c)) even after slight increase in spinal threshold of array electrodes. 4. Discussion Our aim is to create safe and effective, long-lasting softening cervical SCS arrays with stable electrical stimulation thresholds and limited lead migration. Our driving hypothesis is that softening bioelectronics will enable this paradigm with devices that are easy to insert, will conform around the spinal cord and be safe and effective during chronic stimulation. In this work, we have described the steps necessary to test and ultimately validate this hypothesis in small animal models and hope to use this information to drive further studies in larger animal models to someday create a more effective human clinical therapeutic and human clinical tool for neurological rehabilitation via paired plasticity. In this inquiry, we designed and fabricated three increasingly sophisticated generations of SCS arrays. The failure points in each generation helped us identify how to improve the design moving forward toward our chronic softening SCS array. 4.1. Analyses and hypotheses from most significant results

In our opinion, the most significant device results emerge when combining analysis from figures 5 and 6 together. Figure 5 describes electrochemistry in an in vitro accelerated aging paradigm and in a true ageing paradigm in a small animal model. Figure 6 shows the electrochemical properties of a device before implantation, during implantation and importantly, after the device is removed from the animal, before and after cleaning. Figure 5(a) shows the change in CV curves after soaking in cerebrospinal fluid (CSF) for 30 d. The change in shape of the CV curve is very similar to the change in shape of both the accelerated ageing curve and the true 12


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ageing curve (in vivo CV in figure 5(d)). This indicates that tissue response and biotic mechanisms play a role in the extent of the currents measured across the same voltage sweep, but that the shape of the curve remains consistent. More important are the EIS measurements from the device before, during and after surgery. We observe the explanted device returning to pre-implant levels in terms of the shape of the impedance and the change in phase angle both as a function of frequency. What this means is that the quality of TiN is not perturbed on the device during the implantation and explantation, but rather that tissue interactions are likely what alter the electrochemical properties at the biotic/abiotic interface. Often in these studies it is difficult to decouple biotic and abiotic failure mechanisms, and difficult to attribute changes to the physiological environment as a opposed to device breakdown and failure. The ex vivo results in figure 6(b) clearly point to a device with very similar properties to the pre-implanted tests indicating that the devices remain intact during implantation and after explantation. In fact, SEM images in figures 6(c)–(e) confirm that remnant tissue sticks to the high surface area TiN electrodes. The bits of tissue that remain on the device after explantation do not substantially affect the EIS curves of the TIN electrodes as evidenced by the fact that cleaning the SCS array electrode does very little to change the electrochemical properties. Once we confirm that the device itself is changing very little during at least the first 30 d of implant, the data in figure 5 take on exciting new meaning. We hypothesize that the differences between the in vivo studies and both the accelerated aging studies, figures 5(a)–(c) and the true ageing studies, figures 5(d)–(f), relate to both the effects of tissue growth on and around the electrodes, and to the effect of leakage pathways that are established through the device, but not abiotic (device) failure mechanisms. Figures 5(b) and (e) then confirm the results from figure 6. Accelerated aging shows very little change in EIS over the simulated 29 week time period, with small changes reflective of the slight fluid uptake of the softening substrate which likely lead to some additional parasitic capacitances between the surrounding fluid and the conductive traces. However this result is minimized with impedance at 1 kHz varying much less than an order of magnitude for the accelerated aging studies in figure 5(b). However in figure 5(e), the EIS confirms that order of magnitude increase in impedance at 1 kHz. Although this is out at six weeks and not just 30 d as shown in the figure 6, we hypothesize that this change is all biotic in nature and that impedance of the electrode would return to pre implant levels, indicating robustness of the TiN interface. Thus tissue is altering the interface with the neurons in the spinal cord, but importantly not in a way to preclude device functioning. What we observed in figure 8(b) is an increase in the stimulation threshold needed to evoke a motor response in the rat model. The increase in stimulation threshold maps nicely to the initial increase in impedance seen in figure 5(e). The electrode impedance at 1 kHz can also be seen in figure S8, showing a similar tendency as the spinal threshold in which both become stable after four weeks. Since we were able to

conduct the experiment in figures 8(b) and S8 out to 16 weeks and observed very stable thresholds near 450 µA and stable impedance at 1 kHz from four weeks onward, we believe that these properties would remain stable matching the in vitro aged arrays electrochemistry, had we conducted that test to a longer time point. However, the goal of the experiment as a whole was to develop a robust and useful way to stimulate the spinal cord over many months. We conclusively have designed, fabricated and tested a device lasting four months that is sufficient to conduct a paired plasticity study and deliver new neuroscience understanding. The results from figure 7 add additional power to the study by demonstrating how the devices do very little damage to the cord as evidenced by both MRI and immunohistochemistry. 4.2. Optimizing the electrochemistry of the TiN electrodes for stable stimulation

The microscopy characterization of the TiN electrode film shows its characteristic highly structured fractal surface which contributes to the measured CICc, good impedance and current density, especially for such a thin, ~200 nm, film. The TiN electrodes have sufficient electrochemical properties overcoming the small thickness of the TiN film and roughness, relative to much thicker films reported by other researchers. The CSC and CIC are moderately good relative to the roughness of the electrode. The impedance is in the 1 kΩ range which relates to the electrode size pre and post implant. The impedance of the full system after implant and tissue stabilization rises to approximately 10 kΩ, which is still viable for conducting long term studies with relatively low stimulation threshold, once the tissue stabilized around the device. The electrochemical properties of the in vitro aged TiN electrodes are overall stable with the most noticeable change in the CV tests in which the driving current range of the electrodes increased by one order of magnitude. The impedance remained close to 1 kΩ at 1 kHz and the phase angle showed the characteristic TiN behavior along all frequencies tested. The electrochemical properties of the implanted TiN electrodes appear very similar to those from the in vitro aged electrodes. The most noticeable changes are in the impedance magnitude which increased along all frequencies after implantation due, we hypothesize to the tissue interaction in and around the electrode area. The phase angle turned less capacitive (more resistive) at 100 Hz and lower frequencies. The CV behaves very similar to the previously shown (in vitro) after been implanted in which the current range increases one order of magnitude. The Impedance magnitude and phase angle appear to be only affected when the electrodes are implanted as the before and after implantation properties are unchanged, as shown in figures 6(a) and (b). SEM analysis (figure 3(c)) was done to the exposed TiN electrode to capture 2D pictures of its rough morphology. As the SEM only helps to visually inspect 2D features, AFM characterization (figure 3(d)) was used to measure the actual TiN electrode roughness, which is dependent mainly on the deposition thickness of the film.

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process that enables the electronics to remain near the neutral plane of the device to minimize forces on the metal-polymer interface during bending and shearing. This was surprisingly difficult with a 100 µm thick device because we had to find a precise processing technique that would be able to etch through 50 µm of softening polymer to expose the TiN electrodes and 100 µm to outline the SCS arrays. The directional dry etch used for the electrode opening is shown in figure 3(b), this was possible by using an RIE process with oxygen plasma. It was also used to open the connection pads for the soldering of the stainless steel wires, which bridge the array to the 2-channel connector, and to outline the final shape of the softening SCS arrays. While a model of the entire fabrication process is shown in figure 2, the critical steps that diverge from previous work are steps h, i, j and k, in which we were able to pattern through such a thick layer of polymer. Step e, was also instrumental in the full process, as the protective aluminum layer helped stabilize the 200 nm thick TiN layer during the oxygen plasma that was selectively etching the polymer to expose the electrode end and the gold connection pads. Steps a–d, f–g and l, are standard processing steps described in previous bioelectronics works by our group [20, 22] and with similarities to others in the field [36–39]. Enabling a TiN film that is embedded into the device with robust adhesion to the Au interconnects was critical. The acute results of this processing can be observed in figure 3, which detail the electrochemistry and morphology of the electrodes. The serpentine Au interconnects, figure 3(a), allow for a better stress distribution [40], which at the same time permits larger strains to be induced by the extensive rat’s neck movement and rotation without failing prematurely.

Figure 3(d) also shows that most of the TiN features are smaller than 10 nm except for one feature which we hypothesize it is a defect or dirt on the surface at the time of the measurement. TiN high surface roughness is highly desired as this allows the ESA to be larger than the actual geometrical surface area (GSA) and higher charge injection capacity is obtained by increasing the ratio ESA:GSA. The CSC of the TiN electrode is another property that is improved by the electrode roughness. CSCc of these 200 nm (thickness) TiN electrodes is 1.81 mC cm−2 which is a good value relative to other TiN electrodes with CSCc of 2.47 mC cm−2 but an order of magnitude thicker (⩾1 µm) [32] which can interfere with the flexibility of the arrays and lead to cracking of the TiN, possibly delamination, and device failure. 4.3. Overcoming mechanical forces in the animal model to enable chronic studies

Unlike many other bioelectronic medicines, the SCS arrays are positioned in an area of the rat exposed to extreme mechanical bending and shearing forces, which present major design challenges for thin film electrodes. Commercially available spinal cord leads are typically large soft cylindrical structures with band electrodes or thick paddle leads, which are robust against these forces [34]. However, in commercial devices in humans, as noted above, lead migration and lead breakage contribute to more than 25% of devices failures [35]. We took on the task of developing a thin, minimally invasive epidural SCS array that can be threaded under the vertebrae onto the spinal cord. The device must be stiff enough to be pushed through the epidural space, and supple enough, after softening, to conform to the cord to prevent lead migration, and tough enough to withstand the large mechanical forces. The DMA tests in figure S4(b) show how long the polymer takes to soften once it is exposed to physiological conditions and also how much it softens. We can say the softening polymer softens one order of magnitude within the first 10 min of physiological conditions exposure. This prevents the arrays from damaging the spinal cord when implanted and allows the surgents to insert the array with ease. In comparison to the stiff arrays, Parylene-C does not soften and it actually has a higher Young’s modulus than the softening polymer, even in dry conditions as shown in figures S3(a) and (b). It is also important to note that when the softening arrays were stretched during the tensile tests, the gold interconnects did not crack. In fact, the only part of the softening arrays that showed cracks was near the TiN electrode site, as noted by optical microscope analysis (figure S4 (d)–(f)). The softening arrays are not expected to lose electrical conductivity when they are subjected to large mechanical strains and they remain conductive until complete device failure happens.

4.5. Understanding biological effects with predictable device properties

A device was explanted after 30 d of in vivo testing and SEM analysis was performed to detect indicators for signs of device failure. It was found that the device did not have any type of failure, such as cracks or delamination, and instead tissue was found covering most of the two TiN electrodes, which we hypothesize lead to increased impedance and current range while implanted. CV and EIS were performed afterwards to compare to the in vivo and in vitro characterization and it was found that properties were unchanged from the initial in vitro data leading to the conclusion that such tissue covering the TiN electrodes was driving the properties change. After this, the electrodes were cleaned with tergazyme and the array was characterized once again finding that the EIS was similar to the previous in vitro tests and the CV curve slightly less resistive than the ex vivo. An in vitro accelerated aging test of five softening SCS arrays was done to predict the long term performance of the arrays for in vivo stimulation. The electrochemical properties of the arrays were tracked during the length, looking for trends of change in impedance, phase angle, and CSC. The accelerated aging tests helped estimate the chronic performance of the arrays with limitations as the in vivo conditions are difficult

4.4. Overcoming fabrication challenges during the development process

The schematic and optical micrograph in figure 1, shows the proposed SCS array concept and the as fabricated and connected device. Figure 2 represents a new device stack that accomplishes several major objectives. First, we developed a 14


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to simulate in an in vitro setup due to the variability of the in vivo environment caused by the biotic/abiotic interaction of the arrays with the living tissues while implanted. Figure 5 shows a very similar behavior between the in vitro accelerated aging test and the implanted electrochemical tests. The in vivo and in vitro CV and phase angle graphs follow similar trends. In CV graphs, the measured current increased from hundreds of nA to the single digit µA levels. Even though the in vivo CV graph is cut off at the µA scale, due to test parameters selection (we wanted to limit currents seen in the body with large safety factors for animal safety), the trend of current increase is clear. In the EIS graphs, the impedance magnitude behavior does not have easily noticeable similarities as the impedance in vitro remains stable and does not increase as much as for the in vivo impedance. Again, the increase in impedance magnitude in the in vivo tests could be attributed to the living tissue in contact with the TiN electrode while in vitro the interaction is only with PBS which could present a slight change in salt concentration due to the high temperature to which it was exposed and evaporation of water. The phase angle also shows similar trends although the changes in the in vivo tests are larger with values changing from −80 to −25°. All of this suggests that in vivo environment contributes to the change in electrochemical properties in a great manner. We also hypothesize that the changes in electrochemical properties can be also due to an addition of a resistive pathway in the softening SCS arrays attributed to a possible leakage current through the softening polymer.

post implantation. There was a significant elevation of the spinal stimulation threshold current within four weeks in both types of SCS arrays (softening polymer and Parylene-C) that may result due to encapsulation of arrays in a fine membrane of connective tissue resulting in an increase in impedance. Spinal threshold stays almost constant after four weeks. Array electrodes were effective in modulating descending motor responses after paired SCS and the effect of this paired motor cortex and SCS was consistent and stable throughout 16 weeks in softening array and for eight weeks in Parylene-C group. The efficiency in chronic neuromodulation can be explored for further in recovery from corticospinal tract injury or higher cervical spinal lesions. 5. Conclusions We have introduced a novel process to manufacture chronically viable softening SCS arrays integrating TiN electrodes onto a softening polymer substrate which is stiff at room temperature for ease of insertion and softens once exposed to physiological conditions conforming around the spinal cord. Softening SCS arrays were successfully developed using standard semiconductor fabrication processes integrating Au serpentine interconnects with TiN stimulating electrodes. The accelerated aging test suggests that softening SCS arrays are viable for chronic treatments as their electrochemical properties remain within desired limits for up to 29 simulated weeks. In vivo tests prove they can stimulate and evoke EMG responses by applying cur rent pulses lower than 1 mA in the cervical spinal cord. Safety, stability, and efficacy of the SCS arrays in awake behaving rats was demonstrated. Softening SCS arrays are stable post implantation as observed in terms of their neuromodulatory effect by paired brain and SCS in awake behaving rats, though their spinal threshold increased in initial weeks post implant but that did not change the efficacy of modulatory effect. Softening SCS arrays can take the shape of spinal epidural surface post implantation and secure the arrays from migrating. Histological analysis shows the slight deformation in some rats but that is not leading to injury/damage to the spinal cord.

4.6. Stability of the SCS array

Post implantation, the deformation of the underlying spinal cord caused by the softening arrays was lower compared to the Parylene-C arrays. After the autopsy, the arrays were examined and they appeared to be encapsulated in a fine membrane of connective tissue. This membrane probably isolated the arrays from the spinal cord. There was no significant tissue injury or inflammation post-implantation for both the softening and Parylene-C arrays. Furthermore, no behavioral deficit was observed in the implanted rats within four weeks post-implantation [41]. It is difficult to provide an accurate comparison between the cervical arrays and the Parylene-C or electronic dura in previous studies, since there is a larger degree of neck movement in the case of cervical arrays as opposed to smaller movements for the lumbosacral spinal cord based stimulation [12, 42]. However, our softening arrays were active for a longer duration post-implantation (16 weeks) compared to the other two arrays (five weeks for the Parylene-C arrays; six weeks for the electronic dura). Softening electrode arrays were stable and effective in chronic neuromodulation in awake behaving rats. As far as we know, this is the first experimental study in the rat trying to register motor evoked responses after chronically stimulating cervical spinal cord repeatedly for 16 weeks or more. Spinal thresholds were measured two to five times per week to assess their reliability in stimulating spinal cord and to determine the potential for the array to be active chronically

Acknowledgments AGS and ACDM thank the Consejo Nacional de Ciencia y Tecnologia (CONACYT) of Mexico for the support under the graduate scholarship program. We acknowledge image acquisition by Eric Aronowitz, Dohyun Kim, and Henning U Voss at the Citigroup Biomedical Imaging Center, Weill Cornell Medicine. We also thank Shivakeshavan RatnaduraiGiridharan for his help with statistical methods. AG-S, JBC, and WV are co-inventors on a patent for softening spinal cord stimulators. Funding This work was funded by the NIH R21EB020318 grant.

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