
==== Front
Regen Ther
Regen Ther
Regenerative Therapy
2352-3204
Japanese Society for Regenerative Medicine

S2352-3204(24)00140-8
10.1016/j.reth.2024.08.005
Original Article
Subpial transplantation of adipose-derived stem cells alleviates paraplegia in a rat model of aortic occlusion/reperfusion-induced spinal cord infarction
Takahara Eisaku a
Kamizato Kota b
Kakinohana Manabu b
Sunami Hiroshi c
Kise Yuya d
Furukawa Kojiro d
Ntege Edward Hosea a
Shimizu Yusuke yyssprs@gmail.com
a⁎
a Department of Plastic and Reconstructive Surgery, Graduate School of Medicine, University of the Ryukyus, Okinawa 903-0215, Japan
b Department of Anesthesiology, Graduate School of Medicine, University of the Ryukyus, Okinawa 903-0215, Japan
c Center for Advanced Medical Research, School of Medicine, University of the Ryukyus, Nishihara 903-0215, Japan
d Department of Thoracic and Cardiovascular Surgery, Graduate School of Medicine, University of the Ryukyus, Okinawa 903-0215, Japan
⁎ Corresponding author. Department of Plastic and Reconstructive Surgery Graduate School of Medicine, University of the Ryukyus, Okinawa, Japan. yyssprs@gmail.com
21 8 2024
6 2024
21 8 2024
26 611619
7 6 2024
21 7 2024
2 8 2024
© 2024 The Author(s)
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
Background

Thoracoabdominal periprocedural occlusion/reperfusion injury of the spinal cord (SCII/R) can lead to devastating paraplegia, underscoring the critical need for effective interventions. However, our knowledge of optimal medical strategies and their efficacy remains limited. Preclinical investigations have shown promise in harnessing adult stem cells, including pluripotent and multipotent stem cells such as mesenchymal stem cells (MSCs), to address SCII/R by enhancing neuro-inflammation, axonal growth, and myelination. Particularly, growth factors derived from adipose tissue-derived MSCs (ADSCs) have been proposed to facilitate recovery. Despite advancements, achieving complete recovery remains a formidable challenge. Therefore, gaining a more profound insight into the role of ADSCs in alleviating SCII/R-induced paraplegia, including optimizing the delivery systems for therapies, is imperative.

Materials and methods

In this study, we assessed the impact of subpial allogeneic rat adipose tissue-derived MSCs (rADSCs) transplantation on paraplegia using a rat SCII/R model induced by ephemeral aortic occlusion, known as the Taira-Marsala model. rADSCs were isolated from adipose tissue of male Sprague-Dawley rats, cultured, characterized, and cryopreserved. One week following the induction of paraplegia, rADSCs (n = 6) or physiological saline (n = 6) were transplanted. Hind limb motor function was evaluated before treatment and at 3-, 7-, and 14-days post-treatment using the Basso-Beattie-Bresnahan scoring system.

Results

The rADSC-treated group demonstrated a significant improvement in hind limb motor function compared to the saline-treated group (p < 0.05), with 5 out of 6 rats exhibiting enhanced motor function following treatment.

Conclusions

Our findings suggest that subpial rADSC engraftment may enhance SCII/R-induced paraplegia recovery. These initial results drive further research to validate this potential, understand the molecular mechanisms, and optimize therapies.

Highlights

• SCII/R poses challenging treatment and significant public health concerns.

• rADSCs therapy advances indicate potential in SCII/R paraplegia mitigation.

• Subpial delivery optimizes rADSCs therapeutic prowess in SCII/R.

• Subpial rADSCs delivery holds promise in SCII/R, validation is ongoing.

Keywords

Adipose derived stem cells
Spinal cord ischemia
Reperfusion injury
Stem cell transplantation
Subpial transplantation
Abbreviations

ADSCs Adipose tissue-derived mesenchymal stem cells

BBB Basso, Beattie and Bresnahan

CNS central nervous system

CSF Cerebrospinal fluid

CD cluster of differentiation

DMEM Dulbecco's modified Eagle's medium

FBS Fetal bovine serum

IgG Immunoglobulin G

MSCs Mesenchymal stem cells

NSCs Neural stem cells

SCII/R Occlusion/reperfusion injury of the spinal cord

P passaging

PE Phycoerythrin

PBS Phosphate-buffered saline

NS Physiological/normal saline

rADSCs rat adipose tissue-derived MSCs

SCII spinal cord ischemic injury

SD Sprague–Dawley
==== Body
pmc1 Introduction

Spinal cord injury (SCI) constitutes a significant global health burden, resulting in either transient or permanent neurological impairments and substantial socioeconomic costs [1]. Estimates suggest a global prevalence of approximately 20.6 million cases and an annual incidence ranging from 250,000 to 500,000 individuals [2]. Trauma, encompassing falls, motor vehicle collisions, and falling object impacts, represents the primary etiology; however, non-traumatic contributions also exist [1,3]. Spinal cord ischemic injury (SCII), a specific subtype of SCI, is uncommon but carries significant morbidity. SCII accounts for roughly 1–2% of all neurovascular emergencies [4]. Despite the relatively low prevalence, SCII is associated with high rates of complications, and ongoing debate surrounds patient survival outcomes [5,6].

The pathophysiology of SCII involves a complex cascade of cellular events, including programmed cell death (apoptosis) primarily occurring within 48 h of ischemia [7]. Additionally, oxidative stress, characterized by excessive reactive oxygen species production, and necroptosis (another form of programmed cell death) contribute to the cellular damage observed in SCII [8,9]. These mechanisms ultimately culminate in severe neurological deficits, such as motor impairments, sensory loss, and autonomic dysfunction [10].

Paraplegia, a serious motor impairment complication of SCII, manifests in approximately 6.5% of cases, with a particularly notable rise in incidence following aortic repairs, such as thoracic endovascular aortic repair [11,12]. The motor dysfunction associated with paraplegia is closely linked to neuronal cell death occurring within the spinal cord's border zone, a region where blood flow is often compromised [[13], [14], [15]].

The adult central nervous system has limited capacity for self-repair, rendering SCII inherently challenging for recovery. Current therapeutic options for SCIs face numerous hurdles, including the need for injury stabilization, prevention of secondary injury cascades, and comprehensive rehabilitation support [16,17]. Various interventions, such as intravenous fibrinolysis, cerebrospinal fluid drainage, meticulous vascular risk factor management, antiplatelet/anticoagulation therapy, neurotrophic factors, neuroprotective agents, and spinal cord-like scaffolds, have demonstrated limited efficacy in promoting spinal cord regeneration [16,[18], [19], [20]]. The heterogeneity of these approaches and their inability to comprehensively address the complex pathophysiology of SCII have hampered their success. Consequently, there is a critical need to develop novel treatment strategies.

Stem cell therapy, particularly utilizing mesenchymal stem cells (MSCs), holds promise as a therapeutic approach for SCII [[21], [22], [23]]. MSCs can be derived from various tissues, such as adipose tissue, bone marrow, and perinatal tissues, offering a potentially safer and more practical approach compared to other stem cell types [[24], [25], [26]]. Among different MSC sources, adipose-derived MSCs (ADSCs) have emerged as a desirable candidate for CNS repair due to their abundant availability, relative ease of extraction, and demonstrated therapeutic potential [17,25,26].

A recent review by Qin et al. [27] highlights that ADSCs exert their therapeutic effects on SCI through multiple mechanisms. They secrete a cocktail of neurotrophic factors, including brain-derived neurotrophic factor, glial cell line-derived neurotrophic factor, and nerve growth factor, which promote neuronal survival and axonal regeneration. Additionally, ADSCs possess the potential to differentiate into neuron-like cells and glial cells, potentially replacing lost or damaged cells. These cells also possess immunomodulatory capabilities, acting to inhibit the activation and proliferation of various immune cells and secreting anti-inflammatory cytokines, such as interleukin-10 and transforming growth factor-beta, which can attenuate inflammation and reduce glial scar formation [27]. Furthermore, ADSCs promote angiogenesis by secreting angiogenic factors, such as vascular endothelial growth factor and hepatocyte growth factor, which stimulate the formation of new blood vessels, restore blood flow, and reduce ischemia [[28], [29], [30]].

Despite the promising preclinical data on ADSC therapy for SCI, translating this approach into routine clinical practice necessitates overcoming challenges such as standardizing protocols for large-scale production of clinical-grade cells and optimizing transplantation methods to ensure safety and efficacy [21]. While intrathecal ADSC implantation has demonstrated an acceptable safety profile and modest efficacy in clinical trials, significant limitations have been observed due to adverse events [[30], [31], [32], [33]].

Preclinical studies utilizing rat models of SCII suggest that subpial delivery of therapeutic agents directly to the spinal cord parenchyma represents a promising strategy for treating paraplegia [[34], [35], [36], [37]]. This approach offers advantages such as targeted delivery, potentially improved cell survival rates, and a reduced risk of systemic side effects compared to other methods [36,37]. Based on this rationale, we hypothesized that subpial transplantation of allogeneic ADSCs could enhance functional recovery following SCII.

In the present study, we utilized rat models of spinal cord ischemic injury/reperfusion (SCII/R) induced by temporary aortic occlusion using the established Taira-Marsala model [34] to evaluate the impact of subpial rat adipose tissue-derived MSCs (rADSCs) transplantation. Our findings provide supportive evidence for the potential therapeutic efficacy of this approach in mitigating the functional consequences of SCII/R-induced paraplegia, as demonstrated by significant improvements in motor function.

2 Materials and Methods

2.1 Animals and surgical procedures

Adult male Sprague-Dawley (SD) rats (240–330 g) were used for this study following approval from the Institutional Animal Care and Use Committee at the University of the Ryukyus, Okinawa (A2023029). All procedures adhered to relevant guidelines for animal research. The experimental design is outlined in Fig. 1. Twelve animals were randomly assigned to two groups (n = 6/group): (1) rADSCs administration group and (2) control group receiving phosphate-buffered saline (PBS). General anesthesia was induced and maintained with 2–5% isoflurane (YAKUKENSHA CO, Japan) in oxygen during surgery. The surgical site was disinfected with chlorhexidine gluconate solution (SARAYA, Japan). Core body temperature was maintained at 37.5 °C throughout the procedure. All animals in the rADSCs group received immunosuppression with tacrolimus (1.0 mg/kg/day, subcutaneous) starting on the day of surgery and continuing throughout the observation period. Additionally, mycophenolate mofetil (30 mg/day) was administered intraperitoneally 1 day before, on the day of, and 1 day after surgery. At the study endpoint, animals were euthanized according to ethical guidelines.Fig. 1 Experimental Design. Following spinal cord ischemia/reperfusion injury, rats with severe motor dysfunction (BBB ≤3) were assigned to receive: rADSCs (n = 6) or NS (control, n = 6) via subpial injection one-week post-injury, and motor function recovery was assessed using the BBB score on days 0, 3, 7, and 14 after treatment.

Fig. 1

2.2 Post-injury motor function assessment

The Basso, Beattie, and Bresnahan (BBB) locomotor scale [38] was used to assess post-surgical motor function recovery. This validated scale (0–21 points) provides a quantitative measure of hindlimb function after SCII. The BBB scale evaluates aspects of movement including joint mobility, stepping, limb coordination, trunk stability, paw placement, and tail position. Based on their BBB scores, animals were categorized into three stages of recovery: Early Stage (0–7, limited hindlimb movement), Intermediate Stage (8–13, uncoordinated stepping), and Late Stage (14–21, coordinated movement).

2.3 Isolation and culturing of rADSCs

rADSCs were carefully isolated and cultured following established scientific protocols [24]. Adipose tissue was carefully harvested from the scapular region of the SD rats under general anesthesia, ensuring the preservation of tissue integrity and minimal disruption of blood flow. The freshly procured adipose tissue was then thoroughly rinsed with PBS and subsequently sectioned into small fragments. The isolation of rADSCs was conducted utilizing a slightly modified enzymatic digestion approach, as previously outlined [39]. This process involved the utilization of a precisely prepared solution containing 1.0 WU/mL liberase MNP-S, a good manufacturing practice-grade enzyme mixture comprising collagenase types I and II, alongside medium-content thermolysin (Roche Diagnostics). The digested tissue was subsequently filtered through a 150-μm cell strainer, followed by centrifugation at 1500 rpm for 5 min. Primary rADSCs were harvested and cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin (Gibco). Cells displaying plastic adherence characteristics were designated as Passage (P) Zero (P0) rADSCs. Following an initial 24-h incubation period (5% CO2; 37 °C), P0 rADSCs were gently washed with PBS and subsequently maintained in the DMEM containing 10% FBS and 1% penicillin/streptomycin, with media replenished every alternate day.

Upon achieving sub-confluent states, adherent colonies were dissociated utilizing TrypLE™ Express (Thermo Fisher Scientific) and subsequently seeded at a density of 1.1–1.4 × 103 cells/μl. This phase of the culture process was repeated twice, resulting in P3 cultures, which were harvested and cryopreserved until further use.

2.4 Characterization of rADSCs

The cell-surface antigen expression of the cultured rADSCs was analyzed using flow cytometry, following established phenotypic profiling methods [40]. The rADSCs previously cryopreserved at P3 were re-cultured, harvested, and resuspended in PBS. A 100 μL rADSC suspension (1 × 106 cells/mL) was prepared, with one set of cells stained with specific antibodies and another set left unstained as a negative control. The following phycoerythrin (PE)-conjugated antibodies were used for staining: anti-cluster of differentiation (CD) 19 (1D3, NBP2-24965 PE), anti-CD45 (30-F11, NB100-77417 PE), anti-CD34 (ICO115, NBP2-33076 PE), anti-CD105 (MEM-226, NB500-452 PE), anti-CD90 (551401), and anti-CD29 (562801). Corresponding isotype controls included PE-conjugated rat Immunoglobulin G (IgG) 2b (149/10H5, NBP1-43323 PE), mouse IgG1 (NBP1-43319 PE), mouse IgG1, κ (550617), and hamster IgG2, λ1 (553965). All antibodies and isotype controls were sourced from Novus Biologicals and BD Biosciences. The cells were incubated with the antibodies for 30 min at 4 °C in the dark. For the analysis, a total of 8.28 × 106 cells from ten 10 cm dishes were utilized, with the average cell size determined to be 15.85 μm. Dead cells were excluded from the analysis using the viability dye 7-amino actinomycin D (Thermo Fisher Scientific). Flow cytometry was conducted using a BD FACSCalibur™ flow cytometer (Becton Dickinson, San Jose, CA, USA), and the data were analyzed with FlowJo v10 software. Gates were set based on the unstained control and isotype controls to accurately identify positively stained cells. All samples were assessed within 24 h of preparation to ensure accuracy in cell-surface antigen profiling.

The flow cytometry analysis demonstrated that the cultured rADSCs were positive for the MSC phenotype markers CD29 and CD90, while negative for the hematopoietic cell markers CD19, CD34, and CD45. Interestingly, the cells were also found to be negative for CD105, another commonly used MSC marker (Fig. 2).Fig. 2 rADSC Characterization. Flow cytometry analysis revealed: (A–C) rADSCs are negative for hematopoietic markers (CD19, CD34, CD45), (D–E) rADSCs are positive for mesenchymal stem cell markers (CD29, CD90), (F) rADSCs lack CD105, another common MSC marker.

Fig. 2

2.5 Spinal cord ischemia/reperfusion injury model

A SCII/R injury model was established in rats using the Taira-Marsala technique [19], as illustrated in Fig. 3A, B, and 3C. Fig. 3B illustrates the placement of the 20G polytetrafluoroethylene catheter and 2F Fogarty catheter used in the experimental setup. As shown in Fig. 3C–a 2F Fogarty catheter (model CV 1035, American V. Mueller) was carefully introduced through the left femoral artery to achieve aortic occlusion. A 20G polytetrafluoroethylene catheter was inserted through the neck into the left common carotid artery for blood withdrawal and pressure monitoring. The blood collection circuit was maintained at 37 °C using a heating coil and positioned at a height of 54 cm to control proximal arterial pressure at 40 mmHg. Simultaneously, the Fogarty balloon occluded the aorta, and blood was withdrawn from the left common carotid artery to maintain blood pressure at 40 mmHg. After 10 min, the aortic occlusion was released, and reperfusion was initiated. The surgical incision was then closed, and local anesthesia (0.75% bupivacaine, Sandoz Pharma K.K. Japan) was administered. Rats were allowed to recover consciousness and were monitored closely for 2 h at a controlled temperature of 28 °C.Fig. 3 SCII/R Injury Model. (A) Schematic cross-section of spinal cord showing interneurons (blue dots) and motor nerves (red dots) in the watershed zone. (B) Experimental setup showing catheter placements. (C) Anatomical representation of the SCII/R induction system. 2F Fogarty catheter introduced through left femoral artery for aortic occlusion. 20G catheter in left common carotid artery for blood withdrawal. Blood pressure maintained at 40 mmHg (54 cm height) during 10-min occlusion. Heating coil maintains blood at 37 °C.

Fig. 3

2.6 Rat laminectomy and subpial rADSCs transplantation

A week after SCII/R surgery, only rats exhibiting BBB locomotor scores of ≤3 points were selected for randomization (n = 12, 6 per group), as outlined in the experimental design (Fig. 1). All surgical procedures were performed under aseptic conditions using a surgical microscope. The surgical area encompassed the T12 to L1 vertebral segments. A laminectomy was performed using a dental drill to expose the spinal cord at the thoracolumbar junction. Subsequently, a small durotomy was made to access the spinal cord. A 30G needle specifically designed for subpial injections was used to create a micro-incision in the pia mater (Fig. 4A–C). As illustrated in Fig. 4E, the subpial injection was performed using a specially designed blunt needle to minimize tissue damage. The needle was carefully inserted into the subpial space, and the cell suspension was slowly injected to ensure even distribution and minimize leakage.Fig. 4 Subpial Delivery of ADSCs. Schematic illustration of the surgical procedure for subpial administration of ADSCs to spinal cord-injured rats. (A) Fixation device, (B) Injection needle and catheter setup, (C) Rat fixed to the device, (D) Microscopic visualization of subpial injection using a blunt needle, (E) Detailed view of the subpial injection site.

Fig. 4

Following confirmation of proper subpial needle placement, animals in the rADSC group received a 10 μL injection of allogeneic P3 rADSCs in PBS at a concentration of approximately 8.0 × 104 cells/mL. The control group received a 10 μL injection of PBS alone (Fig. 4D). The surgical incision site was infiltrated with 2 mL of 0.75% bupivacaine before closure. Following surgery, rats were allowed to recover from anesthesia. Postoperative motor function in the hindlimbs was assessed using the BBB scoring system on days 0, 3, 7, and 14 post-transplantation.

2.7 Statistical analysis

All data were expressed as mean ± standard deviation (Mean ± SD) and analyzed using GraphPad Prism 8.0 software (GraphPad Software, Inc., La Jolla, CA, USA). For comparisons of BBB locomotor scores between the rADSC and control groups at each time point (days 7 and 14 post-transplantation), a two-tailed Student's t-test was employed. A significance level of p < 0.05 was used. Significant differences were denoted by asterisks (∗) with the corresponding p-value reported.

3 Results

3.1 Efficacy of subpial rADSC transplantation in improving motor function

rADSCs were characterized by flow cytometry to confirm their mesenchymal stem cell phenotype before transplantation (Fig. 2).

The SCII/R injury model was established in twelve SD rats using the Taira-Marsala technique (Fig. 3). Blood pressure was maintained at 40 mmHg during the 10-min aortic occlusion.

One week post-SCII/R injury, rats with BBB scores ≤3 were randomized into two groups (n = 6/group): one receiving subpial rADSC transplantation and another receiving control injection at the T12-L1 level (Fig. 4).

BBB locomotor scores, assessed according to the experimental timeline, were significantly higher in the rADSC group compared to the control group at the two-week post-transplantation time point (p < 0.05) (Fig. 5). This finding suggests an improvement in motor function in animals treated with rADSCs. Notably, five out of six rats in the rADSC group demonstrated substantial improvements in hindlimb motor function, as further supported by the data presented in Supplementary Videos 1 and 2.Fig. 5 Improved Locomotor Function after rADSC Treatment. BBB scores were significantly higher in the rADSC group compared to the control group on days 7 and 14 post-treatment (P < 0.05), indicating enhanced motor function recovery. Data presented as means ± SD (n = 6 per group).

Fig. 5

4 Discussion

This study demonstrates that subpial rADSC delivery significantly improved lower limb motor function in a SCII/R paraplegic rat model, as evidenced by increased BBB scores. This approach aims to address critical challenges of efficient cell delivery and distribution in ADSC therapy for spinal cord injury [33,41,42]. Our findings highlight the efficacy of subpial delivery for rADSCs, suggesting potential advantages in targeting and functional outcomes. The technique may help overcome limitations observed in prior studies, such as low engraftment rates and poor spatial distribution of transplanted cells [33]. Moreover, the study contributes to the understanding of ADSC applications in ischemic spinal cord injuries, specifically within the SCII/R model.

Our experimental setup, as illustrated in Fig. 3, provides several advantages for studying SCII/R. The Taira-Marsala model allows for precise control of proximal arterial pressure during aortic occlusion, enabling consistent induction of spinal cord ischemia. Specifically, our setup maintained blood pressure at 40 mmHg using a blood withdrawal system positioned at a height of 54 cm and kept at 37 °C. This level of control is crucial for reproducibility and for understanding the relationship between perfusion pressure and ischemic injury. However, it's important to note that this model, while effective, may not fully replicate the complex pathophysiology of SCII/R in humans, particularly in cases of aortic surgery or trauma.

This study confirmed the MSC phenotype of rADSCs through the expression of CD29 and CD90 surface markers, and the absence of hematopoietic lineage markers CD19, CD34, and CD45. Consistent with prior studies reporting variability in MSC-specific marker expression in ADSCs [43,44], our rADSCs lacked CD105 (Endoglin), a marker often linked to differentiation and immunomodulation functionalities. The CD105-negative phenotype of our rADSCs warrants further investigation. While some studies suggest enhanced differentiation potential in CD105-negative MSCs [45,46], others have linked CD105 expression to improved angiogenic capacity [47]. The implications of this phenotype for SCII/R treatment remain unclear. Future studies comparing the efficacy of CD105-positive and CD105-negative rADSCs in SCII/R models could provide valuable insights into the role of this marker in therapeutic outcomes.

Subpial delivery offers several advantages compared to other cell therapy delivery routes, such as the intravenous and the intrathecal administration. By directly delivering rADSCs to the spinal cord parenchyma, subpial injection ensures targeted delivery and potentially enhances cell survival and integration due to the proximity of the transplanted cells to the lesion site [36,37]. Furthermore, subpial delivery may reduce the risk of systemic side effects associated with other delivery methods [[30], [31], [32], [33]]. The subpial injection technique we employed (Fig. 4) offers significant advantages over other cell delivery methods. The use of a specialized needle and fixation device enables precise targeting and minimizes tissue damage during injection. However, subpial delivery also has potential disadvantages. The procedure is more invasive than systemic delivery methods, requiring direct access to the spinal cord. This invasiveness could increase the risk of procedural complications such as infection or additional trauma to the spinal cord. Moreover, the technique requires specialized surgical skills, which may limit its widespread adoption. Future studies should focus on optimizing this technique to further reduce invasiveness and improve its translational potential. These advantages and limitations suggest that subpial delivery could be a promising approach for the clinical translation of rADSC therapy for SCII/R, although careful consideration and further research remain indispensable.

While ADSCs have shown promise in various therapeutic applications, potential side effects of their transplantation must be considered. A systematic review by Toyserkani et al. [48] of clinical trials using adipose-derived cell therapy found a generally favorable safety profile. However, they reported cases of pulmonary thromboembolism, myocardial infarction, and cerebral infarction in studies involving systemic or cardiac administration. In the context of our study, these risks may be mitigated by the localized subpial delivery method. Nevertheless, potential side effects could include immune reactions, despite the immunomodulatory properties of ADSCs, particularly when using allogeneic cells. There's also a theoretical risk of ectopic tissue formation or uncontrolled cell growth, although this has not been widely reported in ADSC studies [48]. Local inflammation at the injection site is another possible side effect. Additionally, the long-term effects of ADSC transplantation, including potential interactions with resident cells and impact on the local microenvironment, require further investigation.

Our hypothesis focused on the role of interneurons in SCII-induced paraplegia pathophysiology. Damage to these interneurons disrupts the complex neurochemical signaling within the spinal cord, leading to paralysis [49]. The significant motor function improvement observed after rADSC transplantation suggests that ADSCs may modulate the damaged interneuronal environment. However, further investigation is necessary to confirm this and elucidate the specific mechanisms involved.

The mechanisms underlying rADSC-mediated functional recovery in SCII/R are complex and multifaceted. Assinck et al. [50] have outlined five primary mechanisms through which transplanted cells, including mesenchymal stem cells, may promote repair and functional improvements in spinal cord injury: neuroprotection, immunomodulation, axon regeneration, neuronal relay formation, and myelin regeneration. In our study, the observed motor function improvement suggests that rADSCs may be acting through one or more of these mechanisms. In addition to the neuroprotection, immunomodulation, and neuroregeneration previously discussed [27,28], rADSCs may also contribute to angiogenesis, remyelination, and modulation of the glial scar [27,29]. The subpial delivery method may enhance these effects by allowing for more direct interaction with damaged neural tissue and potentially increasing cell survival rates [36]. These effects are likely mediated by the secretion of various growth factors, cytokines, and extracellular vesicles that modulate the local microenvironment and support the survival and regeneration of damaged neural tissue [29,30]. Future studies should aim to elucidate which of these mechanisms are most prominent in subpially delivered rADSCs and how they contribute to functional recovery in SCII/R.

While our results are promising, several limitations are acknowledged. The small sample size limits the generalizability of our findings. Additionally, the short-term nature of our study does not address potential long-term effects or the sustainability of improvements. The use of a single animal model may not fully represent the complexity of human SCII/R. Furthermore, the lack of mechanistic investigations in this study leaves open questions about the precise pathways through which rADSCs exert their effects. Moreover, while our experimental setup and subpial injection technique offer precise control and targeted delivery, they also introduce variables that may not be present in clinical scenarios. The translation of these techniques to human patients will require careful consideration and adaptation.

Future research should address several key areas. Long-term studies are needed to assess the sustainability of functional improvements and monitor potential late-onset side effects. Mechanistic studies, including in vivo imaging and molecular analyses, could elucidate the pathways through which rADSCs promote recovery. Dose-response studies and investigations into the optimal timing of administration would inform clinical translation efforts. Finally, comparative studies with other cell types and delivery methods would help position subpial rADSC delivery within the broader landscape of cell-based therapies for SCII/R.

The translation of subpial ADSC delivery from preclinical studies to clinical application faces several challenges, as highlighted by Assinck et al. [50]. These include optimizing cell survival and integration, addressing potential tumorigenicity, and overcoming the inhibitory environment of the injured spinal cord. Standardization of ADSC isolation, cultivation, and characterization protocols is crucial to ensure consistency and reproducibility of results. The optimal cell dose, timing of administration, and frequency of treatments need to be determined through further studies.

Ensuring safety in future clinical applications of ADSC therapy requires a comprehensive approach encompassing several critical measures. Rigorous cell characterization, including comprehensive phenotypic and functional assays, is essential to ensure the purity and potency of ADSC preparations. This should be complemented by regular genetic stability testing, such as karyotyping and genetic profiling, to detect any potentially harmful mutations in cultured ADSCs. Understanding the fate of transplanted cells through biodistribution studies is crucial for assessing potential off-target effects, while thorough immunogenicity testing is necessary to evaluate the potential for immune reactions, especially in allogeneic settings.

Comprehensive preclinical testing, including expanded animal studies in larger animals with anatomy more similar to humans, will be vital for better predicting clinical outcomes. As research progresses towards human trials, robust clinical trial design will be paramount, incorporating thorough safety monitoring protocols such as regular neurological examinations, imaging studies, and biomarker analyses. To address potential long-term effects, establishing a registry for extended follow-up of patients receiving ADSC therapy will be crucial for detecting any delayed adverse effects.

Throughout this process, adherence to current good manufacturing practices (cGMP) and regulatory guidelines for cell therapy products will be essential to ensure consistent safety and quality standards. By implementing these measures, researchers and clinicians can work towards maximizing the safety profile of ADSC therapy as it moves closer to clinical application.

Additionally, the development of advanced imaging techniques to track transplanted cells in vivo would be valuable for assessing treatment efficacy and safety. Galisova et al. [51] demonstrated the use of multimodal imaging, combining bioluminescence and magnetic resonance imaging, to track mesenchymal stem cells and assess their effects on local blood supply over an extended period. Similar approaches could be adapted for tracking subpially delivered rADSCs in spinal cord injury models, providing crucial information on cell survival, migration, and integration. Such imaging techniques could also help identify the optimal time window for cell transplantation and guide the timing of potential repeated treatments.

Regulatory challenges, including compliance with the cGMP standards for cell production, also need to be addressed for successful clinical translation. Furthermore, as Assinck et al. [50] point out, effectively modeling spinal cord injury in preclinical studies to accurately predict clinical outcomes remains a significant challenge that must be addressed to make cell transplantation a viable clinical option.

5 Conclusion

In conclusion, this study demonstrates the potential of subpial rADSC delivery for treating SCII/R-induced paraplegia, overcoming limitations of previous cell-based therapies. As highlighted by Assinck et al. [50], further research is crucial to understand the underlying mechanisms and overcome implementation obstacles. By advancing our understanding of SCII/R pathophysiology and rADSC potential, we can develop strategies to improve functional recovery and quality of life for individuals affected by SCII/R.

Ethics statement

All animal experiments and procedures were approved by the Institutional Animal Care and Use Committee of the University of Ryukyus.

Author contributions

ET, YS, KK, HS, and EHN performed the experiments, analyzed the data, prepared figures, and authored or reviewed drafts of the paper. ET, YS, and KK performed material preparation, performed the experiments. ET, YS, and KK conceived and designed the experiments. All authors reviewed the manuscript and agreed to be accountable for the content of the work.

Funding

This study was supported by the Japan Society for the Promotion of Science (JSPS) KAKENHI grant (No. 22K08940 ) to Prof. Yusuke Shimizu.

Data availability statement

The original contributions presented in this study are included in the article/Supplementary material, further inquiries can be directed to the corresponding author.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Appendix A Supplementary data

The following are the Supplementary data to this article:Multimedia component 1

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Acknowledgments

We would like to thank Editage (www.editage.com) for English language editing.

Peer review under responsibility of the Japanese Society for Regenerative Medicine.

Appendix A Supplementary data to this article can be found online at https://doi.org/10.1016/j.reth.2024.08.005.
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