A novel nerve guide conduit applied with conductive polymer PEDOT: PSS for repair of long sciatic nerve gap

Abstract

Despite that artificial nerve guide conduits (NGC) have been widely applied in nerve tissue repair for treating peripheral nerve injury (PNI), those apparatuses face great challenges in repairing long-gap nerve defects due to their non-conductive nature. Herein, we report that such clinical issue can be addressed by a bio-mimic polycaprolactone (PCL)-based nerve conduit doped with conductive mixtures (denoted as PP) of polyethylene dioxythiophene (PEDOT) and polystyrene sulfonate (PSS). The PCL-PP nerve conduits demonstrated porous reticular fibrous networks with biocompatible and physicochemical properties. The structure novelty endows the material with a robust mechanical character with a Young’s modulus of 0.32 MPa and a tensile strength of 2.9 MPa. Benefited from the capability of conducting endogenous electrical stimulation due to the high conductivity of 5.8 × 10^-3 S/m, the PCL-PP nerve conduits can regulate the biological behavior of Schwann cells (SCs), and remarkably promote the myelin sheath growth and regeneration of nerve tissues in a 10 mm sciatic nerve defect SD rat model. Compared to the contrast nerve conduits without PP compositions, the PCL-PP nerve conduits accelerated the recovery rate of extremity motor function of SD rats by a factor of 1.25-fold. These findings prove that our reported novel PCL-PP composite nerve conduit with functional integration of bioelectrical stimulation is a promising therapeutic approach toward PNI therapy.

Keywords

peripheral nerve injury nerve guide conduit conductive polymer nerve regeneration

1. Introduction

Peripheral nerve injury (PNI) leads to impaired sensation and decreased motor activity functions of patients,¹ and in severe cases, it also results in permanent disability. Although autologous graft implantation is the current gold standard for repairing nerve defect,² it faces several challenges including donor scarcity, potential complications of donor site implantation, compatibility problems between donor and recipient, and incomplete recovery.³ As an alternative, nerve guide conduits (NGCs) have attracted widespread attention due to their potential to provide sufficient physical support and axon regeneration guidance,⁴ and to allow neurotrophic factor diffusion for rapidly bridging the damaged nerve stumps.^5–7

Owing to the advantages of stability, hydrophilicity, biocompatibility, and processability, polycaprolactone (PCL) has been widely applied as tissue scaffolds.^8,9 The PCL-based nerve scaffold prepared by electrospinning provides robust mechanical support and can effectively promote nerve regeneration process.¹⁰ For example, Ozcicek et al. prepared conductive nerve conduits with functional channels and gold nanoparticles (AuNPs), using polycaprolactone (PCL) and poly(D,L-lactic-co-glycolic acid) (PLGA) as the matrix materials, which were applied for the regeneration of spinal cord injury (SCI) in rats.¹¹ Notably, peripheral nerve regeneration is a highly complex pathophysiological process,¹² involving the establishment of specific microenvironment and channels suitable for axon regeneration,¹³ sprouting and extension of axons,^14,15 myelination and maturation of regenerating nerves.^16–18 According to reported research, the presence of conductive media and the successful construction of functional conductive pathways are key prerequisites for ensuring the efficient progress of nerve repair.^19,20 The lack of an effective conductive pathway formation may directly lead to the blockage of nerve electrical signal transmission, significantly inhibit directional growth of axons and process of myelination, ultimately resulting in incomplete nerve regeneration and impaired functional recovery.^21–23 This is particularly prominent in the repair of long-segment nerve defects, which seriously restricts the achievement of repair effects.^24,25

Since many traditional nerve conduits are short of conductive media, which helps to establish electrical signal transduction pathways in nerve defects and guiding nerve regeneration,²⁶ they face great challenges in complete regeneration and functional recovery of nerve tissues, especially in repairing long-gap nerve defects.^27,28 Fortunately, many studies began to introduce conductive materials into nerve conduits to better transmit electrical signals. The introduction of electrical stimulation can obviously simulate the body’s neural electrophysiological microenvironment, conduct electrical signals and activate Schwann cell activity, thus promoting Schwann cell proliferation and migration, accelerating axonal directional growth and remyelination; at the same time, the electroactive microenvironment can weaken the inflammatory response, reduce the breeding of scar tissue, and improve the efficiency of peripheral nerve regeneration and repair from multiple dimensions.^29,30 Among the conductive materials, polyethylene dioxythiophene (PEDOT) and polystyrene sulfonate (PSS) mixture (short for PEDOT: PSS) is widely applied in bioelectrical materials due to its steady conductivity, good water-solubility and ideal biocompatibility.³¹ Here, PEDOT is a polythiophene derivative with high electrochemical stability, conductivity, and thermal stability.^32–34 However, PEDOT itself is insoluble and infusible with poor processability, making it difficult to be directly used in the preparation of nerve conduits and other biomedical materials. While PSS, as a water-soluble polyanion, can form stable ion pairs with the positively charged backbone of PEDOT through its sulfonic acid groups (SO₃^-), enabling steady dispersion of PEDOT in aqueous solutions.^35–37 Therefore, loads of research on applying PEDOT: PSS to conductive nerve conduits have been reported.^38,39 For instance, Liu et al. invented a poly (lactic co glycolic acid) (PLGA) nerve conduit (NGC) modified with PEDOT: PSS/chitosan (CS) and combined with exogenous electrical stimulation to enhance PNI recovery in rats. The incorporation of CS into PEDOT: PSS coating could synergistically improve the surface bioactivity and conductivity, promote cell adhesion and create a more favorable microenvironment for nerve regeneration.⁴⁰ In cooperation with the contribution of conductive materials, the three-dimensional structures also play a crucial role in affecting the biomechanical properties of the nerve grafts.^41–43 Chen et al. proposed a piezoelectric biodegradable NGC, which was composed of aligned polycaprolactone (PCL)-β-glycine composite nanofibers, generating electrical stimulation in response to low-frequency mechanical vibration, thereby achieving wireless nerve regeneration. This NGC exhibited high-voltage electrical output under mechanical stress, including low-frequency vibration of a massage gun. The results of in vitro and in vivo experiments showed that piezoelectric stimulation significantly enhanced Schwann cell myelination and neurite outgrowth, endowed NGC the ability to perform robust structural repair of 10 mm sciatic nerve defects, with 99% recovery of motor function and 96% recovery of nerve conduction, which was comparable to the performance of autografts.⁴⁴ Qi et al. developed MXene/polydopamine (PDA)/PLLA conductive nerve conduits, which were fabricated by laser additive manufacturing. The results showed that MXene was uniformly wrapped around PLLA, forming a three-dimensional continuous conductive network in the conduit. The enhanced current promoted cellular neural differentiation, with a 2.4-fold upregulation of Nestin and a 4.3-fold increase in Ca²⁺ influx, indicating that this conductive NGC represented great advantages in nerve repair due to its high current that effectively promoted cell nerve differentiation.⁴⁵ Those studies clearly demonstrated that conductive biomaterials that closely mimic the structural characteristics of nerve tissues can regulate the cellular behavior^46,47 and microenvironment,^48–50 as well as deliver bioactive growth factors,^51,52 which is also beneficial for promoting the proliferation, adhesion, and differentiation of nerve cells.^53–55

In this study, to solve the clinical challenges in repairing long-gap nerve defects, we prepared a bio-mimic PCL-based nerve conduit doped with conductive PEOT: PSS mixtures (denoted as PP mixtures) based on electrospinning technique. Here, the PP mixtures with a hydrophilic nature have steady conductivity, water solubility, and biocompatibility. The brand-new PCL/PP composite nerve conduit was delicately designed with a porous network microstructure mimicking that of living nerve tissues, which could induce directional growth of nerve tissue along the functional tunnels. The addition of PP mixtures with abundant charged groups not only improved the surface bioactivity and conductivity of the scaffold, but also formed continuous conductive pathways in the scaffolds. As a result, the PCL/PP nerve conduit could regulate the biological behavior of Schwann cells (SCs), exhibited a higher repair efficiency in a 10 mm sciatic nerve defeat of SD rats, and contributed to a faster recovery of extremity motor function of SD rats than its nonconductive counterparts.

2. Materials and methods

2.1. Preparation of PCL and PCL coated with PEDOT: PSS conduits

As shown in Figure 1, PCL electrospinning solution was prepared by dissolving polycaprolactone (M_w = 80000) in hexafluoroisopropanol (HFIP) (the mass fraction of hexafluoroisopropanol solution of polycaprolactone was 5%, 7% and 9%, respectively). The releasing agent (K90) was loaded into a syringe equipped with a blunt-end needle. The syringe pump controls the release of K90. The releasing speed of K90 was 1.5 mL/h, and the applied DC voltage was +15 kv/-3 kv. The distance between the needle and the collector was 18 cm. The ejected fibers were collected on the receiver. Similarly, the prepared PCL spinning solution was electrospun under above mentioned conditions. The nerve conduits with an inner diameter of 1.6 mm were successfully prepared and named PCL-5, PCL-7, PCL-9 respectively.

Figure 1.

Schematic diagram of preparation of PCL electrospun nerve conduit coated with PEDOT: PSS and the promoting process of nerve regeneration.

The conductive nerve conduit PCL-PP with a length of 1.1 cm was fabricated in following method: PCL-7 was soaked in absolute ethanol. After 3 minutes, it was taken out, rinsed with purified water, then soaked in 0.5% mass fraction sodium hydroxide aqueous solution for 30 minutes and rinsed with purified water. The pretreated PCL-7 was put into 2 mg/ml polydopamine hydrochloric acid buffer, protected from light and ultrasonicated for 4 hours. The solution was removed and the nerve conduit was rinsed with purified water. Different mass fraction (0.5%, 1%, 1,5%) of PEDOT: PSS in aqueous solution and 2 mg/ml polydopamine hydrochloric acid buffer were uniformly mixed in a brown bottle. After that, the nerve conduit was loaded in the same brown bottle and ultrasonicated for 20 hours, then it was taken out and rinsed with purified water and dried. The conductive nerve conduit made from different mass fraction of PEDOT: PSS PCL-0.5PP, PCL-PP, PCL-1.5PP was successfully prepared. The best comprehensive performance of conductivity and mechanical properties of above conductive conduits was further used in animal experiments.

2.2. Characterization of PCL and PCL-PP

The morphologies of the obtained nerve conduits PCL-5, PCL-7, PCL-9 and PCL-PP-2 were characterized by scanning electron microscopy (SEM, apreo 2C Thermo Scientific), respectively. The molecular structure of PCL-PP-2 was tested by Fourier transform infrared spectroscopy (FTIR, Nicolet is50, Waltham, Ma, USA) to verify attachment of PEDOT: PSS to the surface of PCL substrate. The tensile strength of all conduits was evaluated using a universal testing machine. The conductivity of PCL-PP-2 was examined under st2643 ultrahigh resistance micro current tester. Water contact angles of all conduits were measured with a water contact angle meter (CA200, Kunshan Beidou Precision Instrument Co. Ltd.).

To investigate the degradation performance of the two groups of conduits, all samples were lyophilized for about 12 hours (n=8) and pre-weighed to record the initial weight (W1). Afterwards, the samples were immersed in 0.1 mol/L PBS solution (pH = 7.20-7.40) at 37 °C. The degradation medium was replaced with fresh PBS every week. On a weekly basis, the nerve conduits were taken out from the PBS solution, lyophilized and weighed to record the residual weight (W2). The above operation was repeated for four cycles.

Degradation rate (%) = \frac{(W 1 - W 2)}{W 1}

(1)

2.3. Evaluation of the cytocompatibility of PCL-7 and PCL-PP

Rat Schwann cells (RSC96; Xiamen Yimo Technology Co., Ltd.) were evaluated for cytotoxicity. The complete medium was prepared by adding 10% fetal bovine serum and 1% penicillin streptomycin (P/S; C3420-0100) in DMEM. PCL-7 and PCL-PP conduits were mixed with complete medium as the ratio of conduit surface area/volume of complete medium = 1.25 cm²/mL. In addition, a 96-well cell plate with RSC96 cell density of 1 × 10⁴ cells/well was obtained by cell counting. The conduits/complete medium mixed solution and cell plate were respectively placed in the incubator (carbon dioxide concentration = 5%, temperature = 37 °C) for 24 hours. Then the extract was diluted with DMEM to obtain the extracts with the concentration of 0%, 25%, 50%, 75% and 100%, respectively. About 100 μL of the above extracts with different concentrations were added to the corresponding cell wells of 96-well cell plates which were cultured for 24 hours, then the well-mixed cell plates were laid up in the incubator for 24, 48 and 72 hours respectively. The above cells were divided into two batches for testing cell viability and live dead staining, respectively. To detect the toxicity of live/dead cells, Calcein AM/PI (Product No. KGA9501-1000, Jiangsu Kaiji Biotechnology Co., Ltd.) was mixed with PBS (phosphate buffered saline) in 1/1/2000 ratio to incubate in the dark for 30 minutes, and the cell viability was observed by fluorescence microscope (Olympus DP70; Olympus, Tokyo, Japan): the survival cells were green, the dead cells were red. CCK-8 Kit (Wuhan Sanying Biotechnology Co., Ltd.) was used for cell viability test. 10 μL CCK-8 reagent was added to each well of 96-well cell plate, incubated for 1 hour, and the absorbance was measured at 450 nm wavelength with microplate reader (BioTek ELX800, VT, USA) to obtain the absorbance percentage of cells in PCL-7 and PCL-PP groups. Cell viability was calculated according to the following formula:

C e l l v i a b i l i t y (%) = \frac{[A] - [B]}{[C] - [B]} \times 100

(2)

where [A] is the absorbance of PCL-7 or PCL-PP samples, [C] is the absorbance of the control groups, and [B] is the absorbance of the blank groups.

2.4. In vivo study of PCL-PP

2.4.1. Peripheral nerve injury modeling and scaffold implantation

All in vivo experiments were approved by the Animal Ethics Committee of Shandong Anzhong Medical Device Inspection and Testing Co., Ltd. (Approval No. 202405, Dezhou, China). Adult male SD rats weighing 220.0-240.0 g were anesthetized intraperitoneally with 2.0% sodium pentobarbital solution (30 mg/kg). The right side of the rat was depilated with depilatory cream, sterilized with iodophor, the skin around the right thigh was cut, the sciatic nerve was exposed in a sterile environment, and the nerve with a length of about 10.0 mm was excised. The testing modules were divided into four groups, with 8 testing mice in each group. ①“Control” group: after nerve exposure, muscles and skin on both sides were sutured with 5/0 nylon suture; ②“Autograft” group: 10mm of nerve was cut and then directly sutured with 9/0 nylon to the bilateral nerve stumps; ③PCL-7 and ④PCL-PP groups: PCL-7 and PCL-PP with length of 11 mm were separately implanted into the bilateral nerve stumps, and then the proximal and distal adventitia of the nerve stumps were sutured to the conduit. After the incision was closed, the rats were intramuscularly injected with 5000 units of penicillin and placed under mild ventilation conditions.

2.4.2. Gait experiment of SD rats

Mouse gait experiments were performed at 4th and 8th week after surgery, respectively. The two hind feet of mice in different experimental groups were stained with ink, and the testing mice were put into pre-installed crawling boxes in turns (as shown in Figure S1). Food was placed at the outlet of the box to guide the mice to crawl, and white paper was placed at the bottom of the box to record the footprints left by the mice after crawling. After collecting the footprints, the EPL, NPL, ETS, NTS, EIT, NIT of tested mice were respective measured, and the sciatic nerve function index (SFI) was calculated according to the following Bain formula to evaluate the recovery of motor function.

Bain formula:

S F I = - 38.3 (\frac{E P L - N P L}{N P L}) + 109.5 (\frac{E T S - N T S}{N T S}) + 14.3 (\frac{E I T - N I T}{N I T}) - 8.8

(3)

In Bain formula, N: nonsurgical foot; E: Experimental foot; Fingerprint length (PL): distance from hindfoot to the top of the third toe; Middle toe span (IT): the distance between the second toe and the fourth toe; Toe span (TS): the distance between the first toe and the fifth toe.

2.4.3. Wet weight and histologic assessment of gastrocnemius muscles

The gastrocnemius muscles of the experimental side (GM (E)) and the normal side (GM (N)) of the hind legs of mice at 4th and 8th week after surgery were taken and weighed, respectively. The percentage of muscle weight (GM weight (%)) was calculated according to the following formula:

G M w e i g h t (%) = \frac{G M (E)}{G M (N)} \times 100 %

(4)

The middle abdomen of mouse gastrocnemius muscle was fixed and stained with HE and Masson trichrome, respectively, and observed under a microscope. At least five random fields were selected for each section and analyzed with Image J (Media Cytometics, USA) software.

2.4.4. Morphometric evaluation of peripheral nerve fiber regeneration

For immunofluorescent staining, the cross-sectional cryosection sections of nerve stump of “Control”, “Autograft” and PCL-7, PCL-PP group were washed three times with PBS at room temperature, followed by adding dropwise 10% normal goat serum and 0.2% Triton-x blocking solution, respectively. After incubation at room temperature for 1 hour, the diluted primary antibody mixture IgG fraction of antiserum (NF200, 1:200, ab254348, Abcam, UK) and Mouse Anti-S100B (1:100-1:300, sc-53438, Santa Cruz, USA) was added dropwise, followed by incubation overnight at 4 °C in a wet box. The next day, the sections were rewarmed at room temperature for 30 minutes, rinsed with PBS for three times × 5 minutes, then the fluorescent secondary antibody mixture of Goat anti rabbit IgG H&L (CoraLite®488, 1:100, SA00013-2, Proteintech Group, Wuhan, China) and Goat anti mouse IgG H&L (CoraLite® 594, 1:100, SA00013-3, Pro-teintech Group, Wuhan, China) was added, followed by incubation in the dark at room temperature for 1 hour. After the incubation, the sections were washed with PBS in dark for 3 times × 5 minutes, 1 μg/mL DAPI (1:500, ab228549, Abcam, UK) staining solution was dropped to stain nuclei in dark at room temperature for 5 minutes, the solution was washed with PBS for another 2 times × 3 minutes. Finally, confocal microscope (TCS Sp8, Leica, Wetzlar, Hesse, Germany) was used to observe and take pictures, and the fluorescence intensity was measured with Image J software.

2.4.5. Biosafety analysis

All samples of heart, liver, spleen, kidney and lung of SD rats in “Autograft”, PCL-7, PCL-PP groups at 8th week after surgery were fixed in 4% paraformaldehyde for 24 hours, followed by dehydration through gradient ethanol solutions of 70%, 80%, 90%, and 95% (15-20 minutes per step), then dehydrated twice with 100% ethanol (10-15 minutes each time). After dehydration, the samples were cleared with xylene and embedded in paraffin. The embedded tissue blocks were sectioned and baked at 60 °C for 4 hours (section thickness: 5-6 μm). The prepared paraffin sections were dewaxed, rinsed with distilled water, and then subjected to hematoxylin-eosin (HE) staining. After staining, the tissue sections were dehydrated in 70%, 80%, 90%, and 95% gradient ethanol solutions for 30 seconds, followed by twice dehydration with 100% absolute ethanol (1 minute each time), then transferred to xylene for 10 minutes clearing, followed by sealed with neutral gum. Finally, the tissue morphology was observed under optical microscope (KF-SCAN-PL; Ningbo Jiangfeng Bioinformatics Technology Co., Ltd.).

2.5. Statistical analysis

All data were presented as mean±standard deviation (SD). Results of one-way analysis of variance was performed with graphpad prism software (version 9.5, San Diego, USA). Statistical significance was set as *p<0.05, **p<0.01, ***p<0.001, ***p<0.0001, ****p<0.0001, respectively; “ns” indicates no significance.

3. Results and discussion

3.1. Physical properties of PCL scaffolds and PCL-PP

As shown in Figure 2(a), Fourier transform infrared (FTIR) was used to analyze the chemical composition in the conduit. The stretching vibration of C=O bond, C-O-C ester bond in PCL, inferred to obvious absorption peaks at 1720 cm^-1 and 1000-1200 cm^-1, respectively. Compared with the PCL-7, the presence of stretching vibration of C-S in thiophene rings, SO^3-, C-O-C ether bonds in PEDOT: PSS significantly enhanced the absorption peaks of PCL-PP at 721 cm^-1, 1043 cm^-1, 1 300 cm^-1 and 1400 cm^-1, respectively. The above results proved that the conductive polymer PEDOT: PSS is indeed effectively attached to the PCL scaffold.

Figure 2.

Characterization of physical structure and properties of PCL/PCL-PP: (a) FTIR spectrum of PCL-PP; (b) stress strain curves of nerve conduits in each group; (c) Young’s modulus data of nerve conduits in each group; (d) surface water contact angle data of nerve conduits in each group. All statistical data are represented as mean ± SD. (n=5; *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001, respectively and “ns” indicates no significance).

As shown in Table 1, conductivities of PCL-0.5PP, PCL-PP and PCL-1.5PP was 3.5×10^-4 S/m, 5.8×10^-3 S/m and 5.5×10^-3 S/m. It could be concluded that as mass fraction of PEDOT: PSS increased from 0.5% to 1%, conductivity of nerve conduits increased swiftly, while from 1% to 1.5%, it increased much slower. As shown in Figure SX, the TEM pictures of PCL-0.5PP and PCL-1.5PP demonstrated similar aligned nanofibers compared with PCL-PP. Taking mechanical properties and hydrophobicity of three conductive nerve conduits into consideration, Young’s modulus and water contact angels demonstrated no obvious differences. Thus PCL-PP was chosen as candidate to carry on animal experiments.

Table 1.

Physical properties of PCL nerve conduit in different groups.

Sample	Mass fraction of PCL in HFIP	Conductivity/S/m	Young’s modulus/MPa	Water contact angle/°	Porosity/%	Dimension/mm
Sample	Mass fraction of PCL in HFIP	Conductivity/S/m	Young’s modulus/MPa	Water contact angle/°	Porosity/%	Internal diameter	Outer diameter	Wall thickness
PCL-5	5%	-	0.60±0.02	76.30±0.76	56.75±5.50	1.5	1.9	0.15
PCL-7	7%	-	0.69±0.03	81.53±0.77	56.91±1.70	1.6	2.0	0.21
PCL-9	9%	-	0.95±0.04	84.21±0.83	50.72±4.46	2.0	2.6	0.40
PCL-0.5PP	7%	3.5×10-4	0.31±0.02	61.34±0.46	49.26±2.46	1.6	2.0	0.21
PCL-PP	7%	5.8×10-3	0.32±0.02	60.26±0.55	50.53±3.86	1.6	2.0	0.21
PCL-2PP	7%	5.5×10^-3	0.32±0.03	59.10±0.45	52.52±4.81	1.6	2.0	0.21

As shown in Figure S4, the continuous conductivities of PCL-PP were obtained with ultrahigh resistance micro current tester from 0 to 8th week. The results showed that the conductivities of PCL-PP from 0 to 8th week were 5.8 × 10^-3, 5.2 × 10^-3, 4.8 × 10^-3, 5.0 × 10^-3, 4.8 × 10^-3, 4.7 × 10^-3, 4.7 × 10^-3, 4.5 × 10^-3and 4.4 × 10^-3 S/m respectively, in line with the conductivity of sciatic nerve in SD rats, which was about 3 × 10^-4- 6 × 10^-3 S/m, indicating that conductivity of PCL-PP met the conduction requirements of bioelectric signals,² supplying a endogenous electric stimulation to accelerate differentiation of nerve tissue and regeneration of myelin sheath. Moreover, with the passage of time, the conductivity of PCL-PP decreased by 24% at 8th week, indicating that the long-term conductivity of PCL-PP is relatively stable, which could meet the needs of long-term nerve defect repair in animals.

The mechanical properties of PCL-PP, PCL-5, PCL-7, and PCL-9 were studied by tensile test. On the strain stress curve of nerve conduit obtained from the test (as shown in Figure 2(b)), Young’s modulus of nerve conduit in each group was calculated by tangent method. The results showed that the Young’s modulus of PCL-PP, PCL-5, PCL-7, and PCL-9 were 0.32 ± 0.02 MPa, 0.60 ± 0.02 MPa, 0.69 ± 0.03 MPa and 0.95 ± 0.04 MPa, respectively. It could be concluded that with the increase of PCL solution concentration, the Young’s modulus of PCL nerve conduit showed a slight upregulation. The increased Young’s modulus of PCL-9 led to enhancement of hardness in this group (as shown in Figure 2(c)), which may lead to a result in difficult suturing after surgery. In order to facilitate the implementation of surgery in subsequent animal experiments, PCL-7 was selected to combined with PEDOT: PSS in this work. Fortunately, after adding PEDOT: PSS, the Young’s modulus of conductive nerve conduit PCL-PP still matched the elastic modulus of mouse sciatic nerve (0.1-1 MPa), demonstrating that the composite nerve conduit met the basic mechanical requirements of subsequent sciatic nerve repair experiments.

Figure 2(d) and Figure S2 revealed the water contact angle test results of PCL-5, PCL-7, PCL-9 and PCL-PP nerve conduits, which were 76.30 ± 0.76°, 81.53 ± 0.77°, 84.21 ± 0.83° and 60.26 ± 0.55°, respectively. The above data showed that with the increase of PCL mass fraction, the hydrophobicity of the material was slightly enhanced. However, the introduction of PEDOT: PSS could significantly improve the hydrophilicity of PCL conduit. As nerve regeneration depends on Schwann cells, the surface of PCL-PP with better hydrophilicity was more prone to adsorb extracellular matrix (ECM) components (such as fibronectin), which may improve the adhesion efficiency and spreading area of Schwann cells, thus promoting their proliferation, and reducing the aggregation of inflammatory cells and the formation of fibrotic envelope.

As shown in Figure 3, the effects of different mass fractions of PCL spinning solution on the morphology and properties of electrospun fibers were analyzed by SEM. It could be concluded from Figure 3(a) and Figure 3 that the fibers of PCL-5, PCL-7 and PCL-9 all presented porous network structures and highly ordered arrangement, which were benefitted to simulate the topology of the neural basement membrane (ECM) and provide a similar natural microenvironment for Schwann cell adhesion and axon growth. On the other hand, there are obvious differences in the size of nerve conduits prepared with different concentrations of PCL solution. As shown in Table 1 and it can be seen from SEM photos that with the increase of PCL solution concentration, its inner diameter increases from 1.5 mm to 2.0 mm, its outer diameter increases from 1.9 mm to 2.6 mm, and its wall thickness increases from 0.15 mm to 0.40 mm. In addition, because the amount of conductive polymer is less, the size of nerve conduits attached with different amounts of conductive polymer changes little. It could be concluded that outer and inner diameter of PCL-PP conduit was about 2.0 mm and 1.6 mm, respectively, with wall thickness was about 0.21mm, which closely fitted the size of sciatic nerve of SD rat. Further tests as shown in Figure 3(b) revealed that the fiber diameters corresponding to PCL-5, PCL-7 and PCL-9 were 1.71 ± 0.0076 μm, 2.09 ± 0.019 μm and 3.51 ± 0.0045 μm, respectively, and the porosities were 56.75 ± 5.50%, 56.91 ± 1.70% and 50.72 ± 4.46%, respectively. These results showed that with the increase of PCL solution concentration, the diameter of electrospun fibers gradually upgraded, while the porosity showed a slow downward trend. After attaching PEDOT: PSS, the porosity of PCL-PP was 50.53 ± 3.86%, which had no obvious change compared with PCL-7, indicating that the attachment of PEDOT: PSS had little influence on the porous structure of PCL substrate, furthermore presaged that comparing with nonconductive never conduit PCL-7, PCL-PP maintained the similar bio-mimic three-dimensional networks, which was benefit for adhesion and proliferation of nerve cells.

Figure 3.

Characterization of the surface morphology of nerve conduits in each group: (a) scanning electron microscopy photographs of nerve conduits in each group; (b) fiber size data of PCL-5, PCL-7, PCL-9; (c) porosity data of nerve conduits in each group.

The results of degradation rate of PCL-7 and PCL-PP in 8 weeks can be seen in Figure S5. It could be concluded that the average degradation rate at 8th week of PCL-7 and PCL-PP were 2.65% and 3.20%, respectively, indicating that the slow degradation rate endowed above mentioned nerve conduits a relatively stable mechanical support performance during whole testing periods.

3.2. Effects of PCL-7 and PCL-PP nerve conduits on cell proliferation

After incubation with extracts of different concentrations for 24, 48, and 72 hours, respectively, the cell viabilities of RSC96 and L929 cells were tested and the results were shown in Figure 4 and Figures S6-S7. Fluorescence images of live and dead RSC96 cells on 72 hours were shown in Figure 4(a). Compared with Control group, PCL-7 and PCL-PP group showed equivalent RSC96 proliferation. From the fluorescence image, the samples had obvious cell adhesion and spreading phenomenon, indicating that the cell growth conditions were reasonable. According to GB/T16886.5 toxicity classification standard, the material could be considered non-toxic as long as the cell activity reached above 75%. The results showed that with the extension of culture time, the optical density (OD) of cells in each group showed a steady upward trend. The cell activity of all samples cultured in the extract was more than 90% (Figure 4(b)). This indicated that the cell biocompatibility of all samples met the standards of animal experiments. Calmodulin AM/PI staining was used to detect the effects of different PCL-7 and PCL-PP extracts on the proliferation of RSC96 and L929 cells. The results showed that RSC96 and L929 cells cultured in 100% extraction solution still maintained a good proliferation state, indicating that PCL-7 and PCL-PP had no obvious toxic effects.

Figure 4.

The activity of RSC96 cells cultured with different concentrations of extract; (a) the images of live/dead RSC96 cells cultured with nerve conduit extract of different concentrations of PCL-7/PCL-PP for 72 h; (b) RSC96 cell activity data of the extracts from PCL-7/PCL-PP group after 1-3 days of incubation. (n=5; “ns” indicates no significance).

3.3. Experimental analysis of gait in SD rats

The sciatic nerve function index (SFI) was calculated by walking trajectory analysis combined with footprint parameters to quantify the degree of neurological dysfunction (Figure 5 and Figure S8). As shown in Figure 5(a), under normal conditions, SFI values range from -100 to 0, where 0 represents normal neurological function and -100 represents complete loss of neurological function. As shown in Table 2, the SFI values of PCL-PP group and autograft group at 4th week after operation were -75 and -53, respectively, which were higher than -78 of PCL-7, and the difference between the groups was statistically significant (p < 0.01). At 8th week after operation, the SFI values of the two groups further increased to -59 and -45, which were still significantly higher than those of the control group. The above results showed that PCL-PP could more effectively promote the recovery of peripheral nerve function compared with PCL-7, while its nerve-repairing effect was closer to that of the autologous transplantation group.

Figure 5.

Gait analysis experiments of mice in different experimental groups: (a) footprints of normal side and experimental side of SD rats at 8th week after operation; (b) SFI values of SD rats 4 weeks after surgery; (c) SFI values of SD rats at 8th week after surgery. All statistical data are represented as mean ± SD. (n=5; *p<0.05, **p<0.01, ****p<0.0001, respectively and “ns” indicates no significance).

Table 2.

SFI and wet weight ratio of gastrocnemius muscle of SD rats in different groups.

Sample	SFI		Wet weight ratio of gastrocnemius muscle
Sample	4 weeks	8 weeks	4 weeks	8 weeks
Control	-3	-3	0.96	0.95
Autograft	-53	-45	0.45	0.76
PCL-7	-78	-67	0.32	0.48
PCL-PP	-75	-59	0.37	0.60

3.4. Gastrocnemius atrophy and peripheral nerve fiber regeneration in SD rats

The histological and functional status of gastrocnemius muscle (GM) is the core functional evaluation index reflecting the effect of sciatic nerve regeneration. To evaluate the degree of gastrocnemius muscle atrophy after sciatic nerve injury, Masson trichrome staining and HE staining were used to observe the muscle fiber morphologies of the cross-sections of gastrocnemius muscles of SD rat in each group (Figures 6(a) and 6(b)).

Figure 6.

Morphometric analysis of gastrocnemius muscle in SD rats: (a) photographs of gastrocnemius muscle at 4th and 8th weeks after surgery; (b) HE staining and Masson staining of gastrocnemius muscle of SD rats at 4th and 8th week after surgery; (c) wet weight ratio of gastrocnemius muscle of SD rats at 4th week after surgery; (d) wet weight ratio of gastrocnemius muscle of SD rats at 8th week after surgery. All statistical data are represented as mean ± SD. (n=5; *p<0.05, ****p<0.0001, respectively and “ns” indicates no significance).

As shown in Figure 6(b), the microscopic images of Masson trichrome staining and HE staining of gastrocnemius muscle of SD rats at 4th and 8th week after surgery after operation showed that the gastrocnemius muscle fibers in PCL-PP group and autograft group were more regular and orderly than those in PCL-7 group, and the proportion of collagen fibers was lower than that in PCL-7 group. Through the detailed observation of the muscle fiber image at high magnification (under 100μm scale, Figure S9), it can be clearly observed that the muscle fiber of PCL-PP group is thicker than that of PCL-7 group, and the muscle areas of PCL-PP group and PCL-7 group are 51 ± 1.5% and 33 ± 2.5%, respectively. As shown in Figure 6(c), Table 2, the quantitative analysis results of gastrocnemius on wet weight rate (injured side/healthy side) showed that: at 4th week after surgery, the wet weight ratio of gastrocnemius muscle in PCL-PP group and autograft group were 37.00% ± 1.25% and 45.00% ± 3.20%, respectively, which were higher than 32.00% ± 2.36% in PCL-7 group; however, there was no significant difference between PCL-PP group and PCL-7 group at this time point. At 8th weeks after operation (Figure 6(d)), the wet weight ratio of gastrocnemius muscle in PCL-PP group and autograft group increased to 60.00% ± 2.61% and 76.00% ± 3.75%, respectively, which were 1.25-fold and 1.58-fold higher than 48.00% ± 4.36% in PCL-7 group. The underlying mechanism of the above differences might attribute to the PEDOT: PSS evenly coated PEDOT: PSS conductive layer, which was believed to furnish endogenous electrical stimulation, promoting the fractured nerve to upregulate neurotrophic factors, activating regeneration related signaling pathways, inhibiting glial scar and inflammation, and promoting oligodendrocyte precursor fine differentiation, so as to effectively stimulate axon regeneration and myelin maturation of the fractured nerve, thus enhanced the repairing process of PNI, and effectively slow the atrophy of gastrocnemius muscle.

Immunohistochemical staining was performed on the longitudinal and transverse sections of the middle part (as shown in Figures 7(a) and 7(b)), taking NF200 as marker of nerve fibers, and S-100B as marker of Schwann cells. At 8th week after surgery, compared with PCL-7, the regenerated nerve fibers (NF200 positive) and Schwann cells (S100B positive) in “Control”, “Autograft”, PCL-PP group were more evenly distributed, implying that the distribution of nerve axons in both autograft and PCL-PP groups was uniform. The positive area of S100B in the PCL-7 group (28.51 ± 1.07%) was lower than that in the autograft group (45.42 ± 0.80%) and PCL-PP group (42.10 ± 1.21%) (Figure 7(c)), and the positive area of NF200 in the PCL-7 group (34.43 ± 1.18%) was lower than that in the autograft group (61.44 ± 0.72%) and PCL-PP group (54.31 ± 1.34%) (Figure 7(d)). The results showed that compared with PCL-7, the nerve conduit with conductive material significantly increased the expression of S100B and NF200, hence demonstrated that PCL-PP nerve conduit showed good application potential as a carrier platform to promote peripheral nerve regeneration.

Figure 7.

Tissue changes at 8weeks after sciatic nerve repair: (a) immunofluorescence images of longitudinal sections of nerve fibers; (b) immunofluorescence image of a transverse sections of nerve fibers (c) fluorescence intensities of S100B of longitudinal sections of nerve fibers; (d) fluorescence intensities of NF200 of longitudinal sections of nerve fibers. All statistical data are represented as mean ± SD. (n=3; ***p<0.001, ****p<0.0001, respectively and “ns” indicates no significance).

3.5. Histological study of regenerated sciatic nerve of SD rats

The functional integrity of gastrocnemius muscle is highly dependent on the innervation and regulation of motor nerves. In this study, the sciatic nerve tissue samples of the injured side of SD rats in each group were further collected for histological detection and analysis. As shown in Figure 8, the results of HE staining at 8th week after operation showed that the regenerated nerve fibers in PCL-PP group and “Autograft” group were regularly arranged and densely distributed. However, the nerve fibers in PCL-7 group were sparsely distributed and disorderly arranged, and the regeneration effect was poor. In addition, no obvious local inflammation or impaired microcirculation reconstruction was observed in all groups. The morphology of regenerated nerve tissue in PCL-PP group and “Autograft” group was similar to that of normal tissue, indicating that the introduced conductive material can promote nerve axon regeneration and myelination, further indicating that the introduction of conductive polymers in nerve conduit indeed promoted the differentiation and synaptic growth of primary neurons.

Figure 8.

Histological staining photos of sciatic nerve of SD rats in each experimental group at 8 weeks after operation; (a) transverse section of HE stained sciatic nerve after 8 weeks after operation; (b) longitudinal section of HE stained sciatic nerve after 8 weeks after operation.

3.6. Evaluation of organ toxicity in SD rats

In order to evaluate the biological safety of nerve conduit in vivo, the hearts, livers, spleens, lungs and kidneys of rats in autograft group, PCL-7 group, and PCL-PP group were collected at 8th weeks after operation, and histopathological examination was carried out (Figure S10). The micromorphological observation results showed that during the whole implantation, the cell morphology and tissue structure of the main organs of rats in each group remained normal, and there were no abnormal manifestations such as inflammatory cell infiltration, tissue necrosis or pathological damage. The above results fully confirmed that all nerve conduits used in this study not only played the role of peripheral nerve repair, but also its treatment and degradation process in 8 weeks would not trigger the body inflammatory reaction, and had no obvious toxicity to the main organs of rats, so that the biological safety in vivo of PCL-PP was acceptable.

4. Conclusion

In this study, a series of PCL nerve conduits were prepared by electrospinning method based on different concentrations of PCL/HFIP solution, and the optimal group PCL-7 was selected to combine with PEDOT: PSS due its reasonable mechanical properties. The water-soluble conductive polymer PEDOT: PSS was successfully attached to the surface of PCL-7 by ultrasonic adsorption, so that the conductive nerve conduit PCL-PP was prepared. Compared with PCL-7, the Young’s modulus of PCL-PP decreased slightly, but the water contact angle increased. With the optimization of the proportion of PEDOT: PSS, the conductivity of conductive nerve conduit met the bioelectrical conduction requirements. It showed that PCL-PP had suitable mechanical strength, good conductivity and ideal biocompatibility, integrated with its porous network structure, which was believed to be benefit for the myelin sheath growth and regeneration of nerve tissues. In vitro experiments showed that compared with PCL series nerve conduits, PCL-PP indeed promoted the growth and myelination of Schwann cells. In vivo experiments showed that PCL-PP promoted the regeneration and functional recovery of sciatic nerve in SD rats. However, PCL-PP still has some shortcomings, such as high hardness, slow nerve repair due to lack of nerve growth factor, and few animal cases of nerve repair. We will continue to study and improve these problems in the following-up work. In summary, the endogenous conductive nerve conduit reported in this paper demonstrated a sufficient conductivity to conduct bioelectricity, so it can provide continuous and effective electrical stimulation without external electrical stimulation, which can effectively promote nerve regeneration in rats. However, compared with autologous transplantation, the rate of nerve repair by conductive nerve conduits reported in this paper is still insufficient. We hope to continue our efforts in the future work to further improve the rate of nerve defect repair by optimizing the sheath structure, adding nerve growth factor, more carefully optimizing the ratio of conductive polymer and substrate, and the dispersion of conductive polymer in substrates.

Supplemental material

Supplemental material - A novel nerve guide conduit applied with conductive polymer PEDOT: PSS for repair of long sciatic nerve gap

Supplemental material for A novel nerve guide conduit applied with conductive polymer PEDOT: PSS for repair of long sciatic nerve gap by Duanqiang Xiao, Xueyu Pan, Enxiang Jiao, Kunshan Yuan, Kun Li, Yuechuan Li, Chengchen Deng, Hui Tang, Ziru Sun, Guiying Ren, Meihong Xu, Xiangfeng Cheng, Kai Guo, Yuanbiao Liu, Haijun Zhang in Polymers and Polymer Composites

Footnotes

Acknowledgements

The authors would like to acknowledge the support from the project “Construction and Product Development of Functional Artificial Blood Vessels” funded by National Key Research and Development Program of China (Grant No. 2023YFC2412400) and Shandong Provincial Natural Science Foundation (Grant No. ZR2023QE032).

ORCID iDs

Xiangfeng Cheng

Kai Guo

Funding

The authors disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This work was supported by the National Key Research and Development Program of China (Grant No. 2023YFC2412400) and Shandong Provincial Natural Science Foundation (Grant No. ZR2023QE032).

Declaration of conflicting interests

The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Supplemental material

Supplemental material for this article is available online.

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