Rheological Properties of Self-Healing and Electroconductive Hydrogels for Injured Spinal Cord Regeneration

Introduction: In this study, we studied the rheological properties of the self-healing hydrogels including the mechanical stiffness, gelation time, and injectability. Self-healing hydrogels were formulated using Schiff base chemistry between aldehyde-displaying oxidized polysaccharides (hyaluronic acid (HA) or Pectin) and amine-displaying chitosan. Poly(3,4-ethylenedioxythiophene) (PEDOT) derivatives were incorporated to promote electrical communication between the central nervous system neurons and help with cellular differentiation and axonal outgrowth [1]. Furthermore, a decellularized human spinal cord (DSC) based injectable solution was added to provide pro-regenerative cues for axonal development. The hydrogels had self-healing properties to mimic the dynamic nature of the spinal cord, as under higher strain (1000%) applied to the scaffold, the crosslinking structure breaks down but the crosslinking reforms under lower strain (1%). The mechanical stiffness (storage modulus) of the hydrogels was around ~300-600 Pa, sufficient to support neuronal cell outgrowth. PEDOT derivative incorporation resulted in around ~5.6 mS/cm electrical conductivity [2]. The Schiff base chemistry between the aldehyde-displaying polysaccharides (HA/Pectin) and amine-displaying chitosan resulted in swift gelation (within 5 minutes of incubation at 37°C); the hydrogels exhibited shear thinning properties demonstrating their potential for injectability into patients. Overall, the rheological studies show that hydrogels have self-healing properties with appropriate mechanical stiffness and electroconductivity appropriate for axonal development in the injured spinal cord. The success of this project may result in the demonstration of the self-healing and electroconductive hydrogels that would be suitable to be injected into patient-specific spinal cord injuries. Materials and Methods: The human spinal cord was procured from the National Disease Research Interchange (NDRI). The aldehyde functional groups on HA and Pectin were imparted by oxidation with sodium periodate [3]. The oxidized polysaccharides were solubilized at 1% wt in PBS and the amine-modified chitosan was solubilized at 2% wt in acidic water. Different PEDOT derivative concentrations, 0.5 and 1 mg/mL were added. The human spinal cord was decellularized by modifying the Hudson method. Shear strain, time sweep, frequency sweep, and shear thinning experiment were carried out using an Anton Paar Modular Compact Rheometer (MCR 302) using parallel plate geometry with a 1 mm gap and 8 mm sandblasted parallel plate. For time sweep, 125 μl of hydrogel solution was placed on a 37°C pre-warmed Peltier plate with a 1 mm gap at 1% shear strain and 1 Hz frequency. Storage (G’) and loss modulus (G’’) were recorded at 10-second intervals up to 1000 seconds. Following that frequency sweep study was carried out. The frequency varied from 0.01 to 100 Hz at a constant 1% strain at 25°C, G’ and G’’ were recorded. For shear strain experiments, the strain was alternatively varied from 1% and 1000% consecutively for 5 cycles. The angular frequency was kept constant at 1 rad/s. G’ and G’’ were recorded for 15 seconds at each cycle. For shear thinning experiment was conducted by placing 125 μL of hydrogel solution and the shear rate was varied from 0-150 Pa, the temperature was constant at 37°C, and viscosity was recorded. Results, Conclusions, and Discussions: As shown in Figure 1, the modified Hudson protocol used was able to efficiently remove the cellular and nuclear debris (evident by the absence of DAPI stain) and retain the extracellular matrix (presence of Collagen I, Collagen IV, and laminin). Previously, the presence of ECM-like Collagen I, Collagen IV, and laminin has shown pro-regenerative cues for axonal development. The time sweep study showed that the gelation times of the hydrogels showed rapid gelation, around 5 minutes at 37°C for both HA and Pectin-based hydrogels, ideal for tissue regeneration applications (Figure 2). The rapid gelation can be attributed to the Schiff base formation between oxidized polysaccharide (HA/Pectin) and amine-modified chitosan. The concentration of PEDOT derivatives affected the mechanical strength and the gelation time, the 1 mg/mL concentration of PEDOT derivatives took much longer to reach maximum gelation, yielding undesirable results compared to 0.5 mg/mL. Based on these results, for the remaining studies only 0.5 mg/mL of PEDOT derivatives was used. Furthermore, the hydrogels exhibited a decrease in viscosity with an increase in shear strain (Figure 3). Such shear thinning property is attributed to polymerization relaxation and mimics the action of a hydrogel expelled through a syringe. The hydrogels demonstrated self-healing properties due to the Schiff base chemistry with shear strains between 1000% and 1% (Figure 4). Also, the frequency sweep study showed that at 1% strain and 1 Hz frequency, DSC and PEDOT derivatives incorporated HA-based hydrogels had mechanical stiffness of around 300 Pa, while Pectin-based hydrogels had mechanical stiffness of around 600 Pa (similar to biological tissues). Overall, this study examines the gelation kinetics of polysaccharide-based hydrogels with the addition of electroconductive materials to be used in the treatment of spinal cord injuries and neuronal regeneration. The rheological analysis demonstrated that the hydrogels reached an optimal mechanical strength suitable for spinal cord applications with rapid gelation times, that they were injectable through a syringe, and they showed self-healing properties using dynamic covalent bonding. The developed hydrogels will be evaluated for their ability to promote axonal regeneration in a rodent spinal cord injury model. Acknowledgements (Optional): We acknowledge the use of tissues procured by the National Disease Research Interchange (NDRI) with support from US NIH grant U42OD11158. The authors would like to acknowledge the funding support received from the US NIH (P0225954) and Pruitt chair funding awarded to Christine E. Schmidt. We also acknowledge funding support from the UK Biotechnology and Biological Sciences Research Council (BBSRC, BB/L013762/1 and BB/L013819/1), European Regional Development Fund (03R19P03809) and Royal Society (RG160449) to John G. Hardy.