Introduction
The peripheral nervous system (PNS) has a fundamental capacity for repair and intrinsic regeneration following injury [1]. Unlike the central nervous system (CNS), the PNS exhibits great regenerative potential, which attributes to the distinct responses of their respective glial cells to injury. The process of peripheral nerve regeneration following injury is affected by variable factors including genetic makeup, hormones, growth factors, neurotransmitters, aging, and environmental influences. Specially, the regenerative potential of the PNS results from interaction between growth factors, cellular elements and extracellular matrix.
The currently available drugs for treating neurodegenerative diseases provide only limited symptomatic relief [2]. Combination drug therapy or high-dose chemotherapy, aimed at eliminating degenerated neurons, poses a significant challenge for patients, but is toxic due to its targeting of healthy neurons in both the CNS and PNS [3]. Stem cells, with their self-renewal capabilities, can replicate and maintain their undifferentiated state [4], a property vital for creating a sustainable source of cells for transplantation and therapeutic applications [5]. Their potential leading differentiation into specialized cell types, with the ability to self-renew, positions stem cells as a formidable tool in regenerative medicine, which offers new hope for treating neurodegenerative diseases. Therefore, stem cell transplantation therapy could act as a potential alternative to drug therapy, albeit with limited toxicity to healthy peripheral nerves [6]. Moreover, stem cell transplantation has shown beneficial effects on certain neurodegenerative diseases, including Alzheimer’s disease, Parkinson’s disease, and amyotrophic lateral sclerosis [7]. Previous studies have demonstrated that mouse embryonic stem cells (mESCs)-derived dopamine (DA) neurons can alleviate symptoms of Parkinson’s disease in rat or monkey models. Also, the transplantation of human embryonic DA neurons into patients with Parkinson’s disease was reported in 2003 [8].
Human dental pulp stem cells (hDPSCs) are readily accessible postnatal stem cells that can be isolated from the dental pulp tissue of the third molar or exfoliated teeth [9]. These cells exhibit high proliferative potential and self-renewal capabilities and can differentiate into various cell types, including multiple progenitor populations within the pulp such as osteoblasts, adipocytes, skeletal muscle cells, endothelial cells, and neuronal cells [10]. hDPSCs characterized by the presence of various cell surface markers, including STRO-1 (BioLegend), CD105, or CD146 (Abcam), indicative of early mesenchymal progenitor cells, are readily accessible postnatal stem cells that previous reports show that STRO-1 expression indicates cells with strong abilities to grow and differentiate into different cell types, which is important for tissue regeneration [11]. Notably, STRO-1 is present in pericytes, odontoblasts and dental pulp cells and may be a useful tool to identify hard tissue forming cells. Enhanced STRO-1 expression is observed during the development of rat tooth crowns and roots, as well as in cases of inflamed pulp. CD105 regulates cells to become bone, fat, or cartilage cells [12] and CD146, a cell adhesion molecule and integral membrane glycoprotein at the intercellular junction, promotes angiogenesis and tissue repair [13].
The isolation of purified stem cell populations may augment the efficacy of hDPSCs for their diverse potential clinical applications. The administration of hDPSCs has shown therapeutic promise in various disease models, including cerebral stroke, and nervous system repair [14]. The beneficial effects of stem cell transplantation in the CNS have been extensively researched [14,15]. For example, stem cell transplantation has shown the improvement in multiple sclerosis by enhancing the oligodendrocyte lineage in glial cell-derived neurotrophic factor (GDNF)-modified neural stem cell-grafted mouse models [15]. Moreover, other studies have shown that dental pulp stem cells (DPSCs) can repair neuronal damage by stimulating nerve regeneration, including the facial nerves and peripheral nerve injuries [16,17]. Nevertheless, limited research has delved into the application of stem cell transplantation therapy for peripheral nerve injuries.
Sciatic nerve injury (SNI) manifests as weakness in knee flexion and foot movements. The crush injury model is a commonly used SNI model for assessing post-traumatic motor function impairment, providing certain benefits over the nerve transaction model [18]. Previous research has shown that peripheral nerve injury acutely elevates pro-inflammatory cytokines in the proximal region, resulting in transient movement deficits on the sciatic functional index (SFI) in rodent models of SNI [19]. Additionally, it has been observed that movement impairment might be due to motor neuron depletion in the sciatic nerve, as SNI markedly impacts motor neurons [20].
In this study, we investigated the transplantation of hDPSCs, including sorted STRO-1+ or CD146+ populations, influence nerve regeneration and functional recovery after SNI. We also detected the expression of neurotrophic factors (NTFs) such as neuronal differentiation 1 (NeuroD1), GDNF, and insulin-like growth factor (IGF)-2 and assessed the impact on functional recovery. Through these analyses, we aimed to validate the regenerative potential of hDPSCs with specific stem cell markers and clarify their specific roles in promoting neuronal regenerative capacity following SNI.
Materials and Methods
1. Isolation and culture of hDPSCs and mESCs
hDPSCs were obtained from the third molar of dental patients at Chonnam National University Dental Hospital. Teeth extracted without caries or infection were immediately placed in phosphate-buffered saline (PBS; Thermo Fisher Scientific) with added antibiotics (100 μM/mL penicillin and 100 μg/mL streptomycin; Thermo Fisher Scientific) and 0.25 μg/mL fungizone (Thermo Fisher Scientific). These teeth were transported to the laboratory on ice within 15 minutes post-extraction. Horizontally sectioned 1-mm below the cementoenamel junction using a fissure bur attached to a high-speed handpiece with air-water spray, the teeth were then split open. The pulp tissues, gently harvested and minced into small fragments with a blade under aseptic conditions, were placed in a 6-well cell culture plate. The plate was then incubated in Dulbecco’ s Modified Eagle Medium (DMEM; Gibco-BRL) supplemented with 10% fetal bovine serum (FBS; Gibco-BRL) and antibiotics. The cultures were maintained at 37℃ in a humidified atmosphere with 5% CO2. Cell cultures from the third to fourth passage were utilized for this study. The study was approved by the Institutional Review Board of Chonnam National University Dental Hospital (approval no. CNUDH-2024-003). All procedures involving human-derived materials were conducted in accordance with the ethical standards outlined in the Declaration of Helsinki. Written informed consent was obtained from all participants.
mESCs (SCRC-1002 cell line) were procured from the American Type Culture Collection (ATCC). These cells were cultured in DMEM enriched with FBS, leukemia inhibitory factor (Sigma- Aldrich), penicillin-streptomycin (Thermo Fisher Scientific), β-mercaptoethanol (Sigma-Aldrich), and L-glutamine (Thermo Fisher Scientific). Cultures were kept at 37℃ in a humidified atmosphere containing 5% CO2. To preserve the undifferentiated state, cells were passaged every three days and underwent regular quality control checks.
2. Flow cytometry assay
The cultured hDPSCs were identified based on surface antigens through a flow cytometry method. After trypsinization, cells were incubated in PBS with 0.1% FBS for 45 minutes with fluorescein-conjugated monoclonal antibodies targeting STRO-1 and CD146. A flow cytometry assay was then conducted using a fluorescence activated cell sorting (FACS) Calibur flow cytometer (BD Biosciences), facilitating the quantification and characterization of cells based on their fluorescence properties.
3. FACS of hDPSCs
To isolate hDPSCs, a minimum of 1 × 105 cells were suspended in cold PBS containing 0.1% bovine serum albumin (BSA; Sigma-Aldrich), followed by incubation for four hours in the dark with mouse anti-human monoclonal antibodies against STRO-1 and CD146. The cells were then washed with 0.1% BSA in cold PBS to eliminate unbound antibodies and contaminants. FACS Calibur instrument to sort the cells based on the presence or absence of the specific surface markers targeted by the antibodies. The data from FACS were analyzed using Win MDI Version 2.9 (Biology Software Net), specialized biology software, to confirm the successful isolation of hDPSCs expressing the targeted markers.
4. Animal model of sciatic nerve injury
We procured 7-week-old male Sprague-Dawley rats from Dae-Han Bio Link. The rats were kept under standard conditions with a temperature of 22℃, humidity at 55%, and a 12- hour light/dark cycle. They had ad libitum access to food and water. For the experimental procedures, we anesthetized the rats with an intraperitoneal injection of 10 mg/kg Rompun® (Bayer Co.) and 200 mg/kg Zoletil® (Vibac Laboratories). After shaving the right thigh with a hand shaver, a 1:100,000 solution of epinephrine with 2% lidocaine (Huons Co.) was injected, and the area was sterilized using povidone-iodine (Betadine ®; Mundipharma). Subsequently, we exposed the right sciatic nerve through a 30-mm skin incision and created a 10- mm incision in the epineurium at the mid-thigh level. We then closed the incision using a 16 gauge conduit tube and 10-0 Ethylon® microsurgical sutures (Ethicon). PBS, hDPSCs, STRO- 1+ hDPSCs, CD146+ hDPSCs, and mESCs were transplanted into the artificial conduit within 30 minutes of the sciatic nerve axotomy. After cells transplantation, we sutured the skin wound with 4-0 Nylon® (Ailee Co.) and allowed the animals access to food and water following full recovery from anesthesia. For immunohistochemistry and protein extraction post-SNI, we divided the rats into six groups: control, PBS transplantation, hDPSCs transplantation, STRO1+ hDPSCs transplantation, CD146+ hDPSCs transplantation, and mESC transplantation. Each group comprised 3–5 animals. The Chonnam National University Animal Care and Use Committee approved all experimental procedures.
5. Cell transplantation
Cell transplantation and medium controls were conducted immediately following suturing the epineurium. The cell density was approximately 105 viable cells per microliter (µL). The stem cells were precisely injected into the 10-mm gap between the sciatic nerve stumps, utilizing a silicone tube as a conduit for the procedure. Rats in the negative control group received PBS injections. Each subject received a total of 25 µL of the transplantation solution, administered at a rate of 1 µL per minute using a glass pipette connected to a Hamilton syringe with an approximately 80 µm tip (Hamilton Company). A waiting period of 2 minutes was observed to allow the stem cells to settle before the needle’s removal. The vehicle was administered similarly to the stem cells. To mitigate rejection of the hDPSCs, rats in all experimental groups were given cyclosporine A (15 mg/kg/day, subcutaneously; Sandoz Pharmaceutical Corp.), diluted in extra virgin oil (Sigma-Aldrich). Treatment started with a double dose one day before the surgery. Ten days posttransplantation, the dosage was decreased to 10 mg/kg/day and maintained until the day preceding sacrifice.
6. Tissue harvest and immunohistochemistry
Rats were anesthetized with a combination of 10 mg/kg Rompun® and 200 mg/kg Zoletil® after one day or three days following conduit implantation. The regenerated right sciatic nerves, including both proximal and distal sides, along with the 10-mm stumps, were meticulously harvested under an operating microscope (SZX7; Olympus). The harvested nerve samples underwent a series of procedures: they were soaked in 4% paraformaldehyde (Sigma-Aldrich) for overnight fixation of 12 hours, followed by a 12-hour rinse under running water. Subsequently, the samples were dehydrated in a gradient of alcohol, embedded in paraffin, and sliced into consecutive 4- μm-thick sections using a microtome (Leica CM3050 S; Leica Biosystems). For the immunostaining of specific markers, such as NeuroD1, β3 tubulin, IGF-2, and HuNu, the sections were initially pretreated at 37℃ and neutralized with PBS. This was followed by a one-hour blocking step with a solution of 5% horse serum (Sigma-Aldrich) in PBS containing 0.4% Triton X-100 (Sigma-Aldrich). The primary antibodies, including antirabbit NeuroD1 (1:200, Abcam), anti-mouse β3 tubulin (1:100, Cell Signaling Technology), anti-mouse IGF-2 (1:100, Chemicon), and anti-goat HuNu (1:100, Wako Chemicals USA, Inc.), were incubated overnight at 4℃. Subsequently, the sections were exposed to secondary antibodies conjugated to FITC or CY3 (Jackson Immuno-Research) for two hours, followed by a 30-minute incubation with 10 mg/mL 4′6′-diamidino- 2-phenylindole (Sigma-Aldrich). The resulting images were captured using a confocal microscope (LSM 880; Zeiss), which was equipped with an argon/krypton laser, two helium/neon lasers (Coherent), and a Chameleon two-photon laser.
7. Walking track analysis for footprints
Footprints were analyzed based on three parameters (Supplementary Fig. 1): 1) the distance from the heel to the third toe, termed “paw length;” 2) the span between the first and fifth toes, toe spread (TS); and 3) the distance from the second to the fourth toes, named “intermediary TS.” These measurements were recorded from both the experimental (E) and normal (N) sides. Functional recovery was evaluated using the SFI formula: SFI = –38.3 (EPL–NPL) / NPL + 109.5 (ETS–NTS) / NTS + 13.3 (EIT–NIT) / NIT – 8.8, based on walking track analysis. Postoperative assessments were conducted every three days for two months. The investigators conducting the walking track analysis were blinded to the groupings of the animals. An SFI value of “0” indicates normal function, while “–” signifies complete dysfunction (EPL: experimental leg paw length, NPL: normal leg paw length, ETS: experimental leg toe spread between the first and fifth toes, NTS: normal leg toe spread, EIT: experimental leg intermediate toe spread between the second and the fifth toes, NIT: normal leg intermediate toe spread).
8. Western blot analysis
Sciatic nerve tissue was homogenized in RIPA buffer solution (comprising 150 mM NaCl, 1.0% IGEPAL CA-630, 0.5% sodium deoxycholate, 0.1% SDS, and 50 mM Tris; Sigma- Aldrich) for effective protein extraction. The proteins were then separated by electrophoresis using a 10% SDS-polyacrylamide gel. After electrophoresis, the proteins were transferred from the gel to a nitrocellulose membrane in a Tris-Glycine buffer containing 10% methanol (Sigma-Aldrich) to facilitate efficient protein transfer. The membrane was incubated with primary antibodies at specific dilutions: anti-NeuroD1 (1:100), anti- IGF-2 (1:100), anti-BDNF (1:100, Cell Signaling Technology), anti-GDNF (1:100, Cell Signaling Technology), and anti-Actin (1:100, Cell Signaling Technology). This incubation occurred overnight at 4℃ to ensure optimal antibody binding to the target proteins. Following the primary antibody incubation, the membrane was washed three times with Tris-buffered saline with Tween (Thermo Fisher Scientific). Subsequently, the membrane was exposed to horseradish peroxidase (HRP)- labeled secondary antibodies (Jackson Immuno-Research). For visualization and quantification of protein expression levels, Immobilon Western Chemiluminescent HRP Substrate (Millipore) was applied. The chemiluminescent signals emitted by the antibody-bound proteins were detected and analyzed using Fusion FX equipment (Vilber Lourmat).
9. Statistical analysis
Data are presented as means ± standard error of the mean. Statistical analyses involved one-way analysis of variance, followed by the Student–Newman–Keuls test for multiple comparisons or the Student’s t-test, using GraphPad Prism software, version 6 (GraphPad). A p-value of less than 0.05 was deemed statistically significant.
Results
1. Characterization of isolated hDPSCs population for transplantation
To isolate stem cells from the pulp of the third molar, primary cells cultured from this pulp were subjected to FACS analysis using antibodies specific for CD146 or STRO-1, markers indicative of hDPSCs. The proportion of the stem cell population, encompassing hDPSCs sorted with CD146 and STRO-1, was assessed using flow cytometry. The FACS analysis showed that 34.41% of hDPSCs were positive for CD146, 20.61% for STRO-1, and 40.31% exhibited co-expression of CD146 and STRO-1 (Fig. 1A and 1B). Additionally, hDPSCs sorted with CD146 demonstrated 97.9% positivity, while those sorted with STRO-1 showed 88.8% positivity, in comparison to cells incubated without antibodies (Fig. 1C). hDPSCs sorted for either CD146 or STRO-1 were characterized using immunocytochemistry with antibodies against CD146 (Fig. 1D) or STRO-1 (Fig. 1E), respectively.
2. Histological identification of NTFs after hDPSCs transplantation in SNI rats
NTFs, a family of growth factors including GDNF, BDNF, and IGF, play a crucial role in nerve maintenance and regeneration post-injury in the PNS. Mesenchymal stem cells (MSCs)- derived NTFs immunoreactive signals are predominantly modulated by paracrine or autocrine molecules, secreted or encapsulated in exosomes or microvesicles by MSCs [21]. To investigate whether transplanted hDPSCs in the SNI model affect on neural regeneration, the expression of nerve growth factors as indicators of active regeneration was examined in PBS (control) or hDPSCs-injection group post-SNI. The expression of HuNu, a human cell-specific marker, and IGF-2, a nerve growth factor, was detected by immunofluorescence in hDPSCs, STRO-1+ hDPSCs, and CD146+ hDPSCs-injected the injured sciatic nerves of rats. Transplanted hDPSCs survived in the SNI region, and HuNu+ cells from the injection sites adjacent to the injury three days post-transplantation were colocalized with IGF-2+ cells. Transplantation of CD146+ hDPSCs, STRO-1+ hDPSCs or hDPSCs post SNI increased the number of IGF-2+ cells, compared to the SNI region without hDPSCs transplantation (Fig. 2A). Additionally, mESCs, used as a positive control and tagged with green fluorescent protein, exhibited green fluorescence at 1, 3, and 5 days, confirming their survival at the SNI site (Fig. 2B). These results demonstrate that transplanted hDPSCs contribute to nerve regeneration by enhancing NTFs such as IGF-2. Furthermore, mESCs also survive at the injury site, indicating their potential for future studies or applications in nerve regeneration.
To investigate the effect of hDPSCs transplantation on neural regeneration after SNI, we performed immunohistochemical analysis for NeuroD1 (a marker of neuronal differentiation) and β3 tubulin (a marker of early neuron) in the SNI region. The β3 tubulin+ cells were reduced in the crushed sciatic nerve compared to the control sciatic nerve. However, three days post-SNI, cells double-positive for NeuroD1 and β3 tubulin increased in the hDPSCs transplantation groups compared to PBS group post SNI without transplantation. Similarly, ESCs transplantation in the SNI region showed an augument in NeuroD1 and β3 tubulin double-positive cells compared to the PBS injection group (Fig. 3A and 3B). These results demonstrate that transplantation of both hDPSCs and ESCs positively affects neural regeneration after SNI, underscoring their potential in regenerating peripheral nerve injuries.
3. Upregulation of NTFs after hDPSCs transplantation in SNI rats
Neural regeneration is profoundly influenced by the presence of essential NTFs such as GDNF, BDNF, and IGF-2. Western blot analysis was utilized to detect the expression of NeuroD1, GDNF, and IGF-2 in the transected sciatic nerve tissues following hDPSCs transplantation. Fig. 4A and 4B present the Western blot results, highlighting temporal changes in protein expression at one day and three days post-SNI. Fig. 4C quantifies the relative expression levels of these proteins across the experimental groups. The results indicate that the levels of GDNF and IGF-2 were significantly elevated in the PBS transplantation group at both one day and three days post-SNI compared to the normal control group. However, the expression of NeuroD1 showed no significant difference between the normal control group and the PBS transplantation group at either time point. In contrast, NeuroD1 expression was significantly increased in the CD146+ hDPSCs transplantation group one day post-SNI and in the hDPSCs transplantation group three days post-SNI compared to the PBS transplantation group. Enhanced NeuroD1 expression significantly decreased at three days post-SNI compared to one day post-SNI in the CD146+ hDPSCs transplantation group. Similarly, GDNF expression was significantly elevated in the PBS transplantation group compared to the normal control group at both one day and three days post-SNI. However, GDNF expression was markedly higher in the CD146+ hDPSCs and STRO-1+ hDPSCs transplantation groups compared to the PBS transplantation group one day post-SNI, with a significant reduction by three days post-SNI. The expression of IGF-2 was significantly increased in the PBS transplantation group compared to the normal control group three days post-SNI. Furthermore, IGF-2 expression was significantly enhanced in the CD146+ hDPSCs transplantation group at both one day and three days post-SNI compared to the PBS transplantation group. These findings indicate that the transplantation of hDPSCs, particularly CD146+ and STRO-1+ subpopulations, enhances the expression of NeuroD1, GDNF, and IGF-2, which are associated with peripheral neuron regeneration. Therefore, hDPSCs transplantation can promote neural repair and regeneration following SNI by upregulating proteins involved in neurogenesis and neuro-trophic support.
4. Improvement of motor dysfunction after hDPSCs transplantation in SNI rats
We also evaluated whether hDPSCs transplantation canrestore motor dysfunction in SNI rats. Initially, we analyzed SFI scores at various time points across different groups to explore the relationship between nerve regeneration and its functionality. All animals survived without trophic ulcerations on the operated leg during the regeneration period. Following transection of the right sciatic nerve, loss of motor function in the right hind limb was evident, characterized by tightly clenched toes and a dragging gait. Acupuncture on the right foot elicited no pain response. There were no instances of wound infection or ankle tumescence in any group. Three weeks postsurgery, ankle joint activity began to recover slightly, though tumescence persisted, and the toes started to separate. Acupuncture on the right foot caused a shrinking and escape response, but the injured side still displayed a noticeable limp. Fig. 5 illustrates that the mean SFI scores at 18 days after SNI were significantly improved in the hDPSCs and STRO-1+ hDPSCs transplantation group compared to the PBS transplantation group. Additionally, 30 days post-SNI, the hDPSCs transplantation group exhibited a significantly higher mean SFI value relative to the other groups. These findings indicate that hDPSCs transplantation ameliorates impairment of motor function following SNI.
Discussion
Stem cells offer potential in treating neurological disorders that affect both the CNS and the PNS [6,7]. Notably, peripheral nerves have significantly higher regenerative capacity than central nerves, with successful regeneration largely dependent on the suitability of microenvironment for nerve growth [17]. Introducing a graft bridge within a nerve defect serves as a conduit for regenerating axons, offering guidance and a conducive biological environment for nerve regeneration [22]. Prior research has reported that neural stem cells implanted into a silicone tube after rat peripheral nerve transection enhances the number of nerve fibers and improves the function of the recovering SNI [23].
DPSCs are a distinct subtype of MSCs, known for their selfrenewal capacity and multi-lineage differentiation capability [17]. DPSCs originated from the neural crest have shown potential in differentiating into neurons, expressing various NTFs for axonal regeneration, and performing immunomodulatory functions, proving their efficiency as a cell source for peripheral nerve regeneration [24]. Previous study has demonstrated that hDPSCs implanted in artificial tubes increase nerve fiber numbers and enhance spinal cord recovery [22]. In this study, rat peripheral nerve transection model was used to demonstrate that hDPSCs transplanted into a silicone tube as a conduit has the neuroregenerative potential to enhance the number of nerve fibers as well as the expression of NTFs near the injury site during recovery and improve the function of the recovering sciatic nerve.
CD146, CD105, and STRO-1, known as distinctive markers of bone marrow-MSCs, have been used as stem-cell markers for periodontal ligament and dental pulp tissue [13]. The successful isolation and characterization of hDPSCs, employing CD146 or STRO-1 markers, represent pivotal steps in understanding the potential of these cells in transplantation studies. It was reported that the ability of hDPSCs positively immune-selected for STRO-1, c-kit and CD34 integrates and contributes to healing in rat SNI [11]. In this study, the notably high percentages of CD146+ cells or STRO-1+ cells, specifically 97.9% or 88.8%, in the sorted dental pulp cells, are in line with the findings of prior studies [13,25]. Immunocytochemistry further validated the presence of STRO-1 or CD146 in the sorted hDPSCs, reinforcing the credibility of our findings and emphasizing their characteristic identity as DPSCs, in accordance with related studies.
Previous studies have explored various conduit materials, focusing on biodegradable polymers such as polyglycolic acid, polylactic acid (PLA), polyphosphoester, and silicone [26]. These studies indicate that the conduit material itself does not significantly impact nerve repair. As a result, current nerve repair strategies emphasize the molecular biological manipulation of the conduit’s internal features. Silicone creates a conductive environment for cellular growth and migration. It also offers a protective shield for transplanted cells and maintains a microenvironment that promotes optimal cell survival and integration. The notable properties of silicone tubes include strength, biological inertness, elasticity, transparency, and malleability, allowing for easy observation of nerve regeneration [27]. In this study, we used a silicone tube as the conduit because of its ease of insertion at the injury site, which facilitated the guidance of transplanted hPDSCs. Additionally, a separate study demonstrated that neural stem cells, when transplanted within a PLA conduit, secreted neural inductive factors, including NGF and BDNF. This secretion led to nerve regeneration and functional recovery within six weeks in a rat model with a 10-mm nerve injury [28].
Neural maintenance and regeneration are intricately influenced by the presence of crucial NTFs such as GDNF, BDNF, and IGF. MSCs can effectively modulate the expression of these NTFs, which are released as part of the secretome or packaged into exosomes and microvesicles [21]. In the current study, immunofluorescence analysis using the human cell-specific marker HuNu revealed that transplanted hDPSCs, including STRO-1+ and CD146+ subsets, survived in the SNI region. Furthermore, the co-localization of HuNu+ cells with IGF-2+ cells underscores the involvement of hDPSCs in the modulation of key NTFs. This suggests their potential as a source of NTFs at the injury site. This observation aligns with previous study highlighting the role of IGF-2 in nerve regeneration [6,29]. Moreover, immunohistochemical analysis provided further insight into the regenerative potential of hDPSCs transplantation. A decrease in β3 tubulin+ cells, a marker of early neurons, was noted in the crushed sciatic nerve, but the hDPSCs transplantation resulted in an increase in double-positive cells for NeuroD1 and β3 tubulin [17]. Similar observations were made in the mESCs group, where an increase in doublepositive cells for NeuroD1 and β3 tubulin in the SNI region supports the idea that mESCs facilitate neuronal differentiation and regeneration. This study demonstrates that hDPSCs transplantation hold potential for promoting neural regeneration following SNI, similar to the effect of mESCs transplantation. This effect is likely mediated through the modulation of NTFs and the enhancement of neuronal differentiation. These findings contribute to our understanding of stem cell-based strategies for nerve regeneration and have significant implications for regenerative medicine and neural injury treatment.
NTFs, including NeuroD1, GDNF, and IGF-2, are crucial for nerve maintenance and regeneration in peripheral nerve injuries [30]. Our current findings show that transplantation of hDPSCs significantly influences the expression of these NTFs, compared to the PBS-injected control group. As illustrated in Fig. 4, NeuroD1 expression was notably upregulated in the CD146+ hDPSCs transplantation group one day post-SNI and the hDPSCs transplantation group three days post-SNI, compared with the PBS transplantation group. Upregulation of NeuroD1 demonstrates enhanced neuronal differentiation, indicating the potential of hDPSCs in promoting this process. This finding is consistent with previous study on the roles of NeuroD1 in neuronal differentiation and regeneration post nerve injuries [29,31]. GDNF, known for its neuroprotective effects and its role in facilitating nerve regeneration. Similarly, GDNF was significantly upregulated in the CD146+ hDPSCs transplantation group one day post SNI relative to the PBS group [32]. Additionally, IGF-2 expression was significantly increased in the CD146+ hDPSCs group one and three days post-SNI compared to the PBS group, highlighting its role in nerve regeneration and repair. Prior studies have underscored the importance of IGF-2 in neural recovery and its potential therapeutic implications [33,34]. Together, these results suggest that SNI leads to a reduction in NTFs, but hDPSCs transplantation, particularly CD146+ hDPSCs, elevates the expression of NeuroD1, GDNF, and IGF-2, demonstrating that hDPSCs transplantation may mitigate the impairments caused by peripheral neuron damage in nerve injury models.
The evaluation of motor dysfunction after transplantation of hDPSCs in a rat model of SNI is crucial for understanding the functional impact of this treatment. We assessed the SFI scores at various time points to correlate changes in nerve regeneration with motor function, and our findings provide significant insights [35]. In this study, we explored the effectiveness of hDPSCs transplantation in the recovery of motor function with sciatic nerve injuries. The SFI was used as a measure to monitor improvements in leg function over time. We observed a marked improvement in leg function following hDPSCs transplantation, aligning with previous studies on the regenerative capacity of stem cells [36]. Notably, hDPSCs transplantation into the sciatic nerve injuries showed no adverse effects, and the rats’ leg movement gradually restored, indicating the safety and potential efficacy of this treatment for peripheral nerve injuries. The SFI in all groups was significantly reduced at 6 days, generally recovered by 18 days, and then declined 24 days. Specifically, the STRO1+ hDPSCs transplantation group exhibited significantly the increment of SFI scores at 6 days compared to the hDPSCs transplantation group, suggesting a high potential for rapid repair and regeneration in recovery from SNI. However, further long-term research is necessary to fully understand the mechanisms and optimize this approach for regenerative therapy [5,37].
These encouraging results suggest that transplantation of hDPSCs offers a promising therapeutic strategy for addressing motor deficits caused by peripheral nerve injuries, with significant potential for clinical application. Moreover, hDPSCs have demonstrated potential in a variety of regenerative applications, owing to their capacity to modulate the expression of critical NTFs, a key finding of our study. Additionally, analyzing the regenerative properties of hDPSCs in comparison with other stem/progenitor cell markers may prove beneficial in evaluating the regeneration potential in various stem/progenitor cells. Furthermore, in stem cell research, the potential risks such as cross-infection and graft-versus-host disease during the hDPSCs harvesting and transplantation process must be carefully considered. This emphasizes the need for rigorous protocols regarding manipulation techniques, donor screening data, as well as sterilization and safety measures [38].
While our study offers valuable insights, it also has limitations. The evaluation’s short-term nature and reliance on a solitary administration of hDPSCs without supplementary support materials limit our understanding of the long-term effects and alternative delivery methods. Although we noted changes in NTFs and motor function, a comprehensive investigation into the underlying molecular mechanisms is absent. Significantly, despite employing sorting markers, the intrinsic variability within the hDPSCs population might impact the accuracy of our results. Addressing these limitations is essential for enhancing and progressing stem cell-based methods for nerve regeneration.
In summary, our results demonstrate that the transplantation of hDPSCs leads to an increase in NeuroD1, GDNF, and IGF- 2, which are crucial for nerve maintenance and regeneration, in the SNI region. This is accompanied by improvements in motor function in the SNI model. The study shows that hDPSCs can stimulate the regeneration of damaged sciatic nerves. These findings introduce an innovative approach for hDPSCs transplantation in modulating the release of NTFs, offering potential as a therapeutic target for conditions related to neurodevelopment, neurodegenerative disorders, or neural injuries.
