Peripheral nerve injury (PNI) is common in clinical injuries, which can be caused by multiple traumas such as avulsion, stretch, transaction, and crush injuries, as well as local ischemia and tumor conditions. It is estimated that PNI occurs 2.8% to 5% of all trauma patients. PNI associated with trauma is a devastating complication that can cause partial or complete damage to the nervous system resulting in irreversible impairment, loss of function, and other neurological diseases, which leads to high disability rate and brings heavy economic burden to the family and society. The major challenges for PNI recovery are the distance required by axons to reach the lesion site and the delay in nerve regeneration, and prolonged deprivation of distal stumps impairs their regenerative capability. Given the importance of nerve regeneration in recovery of peripheral nervous system, further studies are needed to focus on improvement of nerve generation.
Adipose tissue‐derived mesenchymal stem cells (ADSCs) are pluripotent stem cells derived from adipose tissues. ADSCs are considered as an attractive cell source in tissue engineering and regenerative medicine, due to their capabilities of self‐renewal, multilineage differentiation, low immunogenicity, high proliferation, and lack of ethical issues, as well as their availability and abundance. In the early stage of this project, ADSCs were used as seed cells and transplanted with acellular nerve allografts into a rat model of sciatic nerve deficit. Our results suggested that ADSCs promoted repair of peripheral nerve injury via synthesis and secretion of neurotrophic factors including nerve growth factor, brain‐derived neurotrophic factor, neurotrophin‐3, glial cell‐derived neurotrophic factor, and ciliary neurotrophic factor. Intrathecal transplantation of autologous ADSCs showed mild improvements in neurological function in patients with spinal cord injury. Extensive efforts have been made to promote ADSCs' capacity of neural differentiation potential, which contribute to its clinical application in human diseases. Therefore, it is important to explore effective nerve regeneration‐promoting approaches to nerve tissue damages and gain better understanding of related mechanisms.
Ghrelin, a 28‐amino‐acid peptide hormone secreted from the stomach, stimulates the release of growth hormone from the pituitary. Ghrelin is involved in a variety of physiological and biological activities, such as enhancing food intake, maintaining energy homeostasis, as well as regulating cell proliferation, apoptosis, glucose metabolism, and immunity. Ghrelin has recently been shown to have neuroprotective effects, including increasing neuronal survival, reducing inflammation and oxidative stress, improving mitochondrial function, and promoting the proliferation, differentiation, and migration of neural cells. Studies indicated that ghrelin inhibited experimental diabetic neuropathy in mice, and prevented cisplatin or paclitaxel‐induced peripheral neuropathy. These findings suggest neuroprotective effects of ghrelin on periphery nervous system. The diverse functions of ghrelin raise the possibility of its clinical significance in treatment of neurodegenerative disease.
In the present study, we focused on the effects of ghrelin on neural differentiation of ADSCs. We induced neurogenesis of ADSCs and different concentrations of ghrelin were applied to intervene neural differentiation of ADSCs. The influence of ghrelin on AKT/m‐TOR and β‐catenin signaling pathways will be examined to explore the molecular mechanism by which ghrelin regulates neurogenesis.
All animal studies were in accordance with the Guide for the Care and Use of Laboratory Animals and approved by Mudanjiang Medical College.
Male Sprague‐Dawley rats were purchased from Liaoning Changsheng Biotechnology Co. Ltd. (Benxi, China). Rats were anesthetized and immobilized, and inguinal fat pads were collected. After washing twice with phosphate‐buffered saline (PBS), the samples were digested with 0.1% collagenase (Gibco Life Technologies, Carlsbad, California) at 37°C for 10 minutes to remove fibrous tissue precipitation, followed by a second digestion with 0.1% collagenase at 37°C for 50 minutes to dissociate viable cells. Digestion was terminated by adding Dulbecco's modified Eagle medium (DMEM/F12, Gibco Life Technologies) supplemented with 10% fetal bovine serum (FBS; Hyclone, Logan, Utah). After centrifugation and removal of supernatant, the cell pellet was resuspended in DMEM/F12 medium containing 10% FBS and cultured at 37°C under 5% CO2 in a humidified incubator. Cell cultures were passaged after reaching confluence. Third‐passage and the fifth‐passage cells were kept and used for subsequent experiments. Their morphological changes were observed under a light microscope.
For induction, third‐passage ADSCs were seeded according to experimental groups. After the cells adhered to the plate wall, they were induced with appropriate differentiation medium. (a) For induction of adipogenesis, the cells were cultured with adipogenic differentiation medium (Cyagen Biosciences, Santa Clara, California) for 9 days. Then the cells were fixed with 4% paraformaldehyde and stained with Oil Red O for 30 minutes to detect lipid droplets. The staining was then observed under a light microscope. (b) For induction of osteogenesis, the cells were cultured with osteogenic differentiation medium (Cyagen Biosciences) for 3 weeks. Then the induced cells were fixed and stained with Alizarin Red dye to detect calcium deposits found in osteocytes. The staining was then observed under a light microscope. (c) For induction of neurogenesis, the cells were induced by neural differentiation medium (Gibco Life Technologies) for 3 days. After induction, they were cultured in the presence of different concentrations of ghrelin with or without addition of 1 μM XAV‐939 (β‐catenin antagonist) or 10 nM MK‐2206 (AKT antagonist). At post‐induction days 3, 6, and 10 , the cell cultures were photographed and morphological changes were observed. The proportion of neuron‐like cells and the number of primary dendrites and branching dendrites per cell were calculated. The cells were collected for subsequent immunofluorescence and western blot at day 10 after induction.
For ghrelin treatment, ghrelin was added to the culture medium at final concentration of 0, 3, 30, or 300 nM (Dalian Meilun Biotechnology, Dalian, China) when third‐passage ADSCs grew to 70% confluence. After being cultured in the presence of ghrelin, cells in each group were collected for further experiments.
Flow cytometry was used to assess the cell surface characterization of ADSCs by detecting CD90, CD45, CD44, CD34, CD29, and CD11b/2f/c. Briefly, third‐passage ADSCs were harvested after centrifugation and PBS washes. The cells were resuspended in 100 μL of flow cytometry staining buffer. They were then labeled with fluorescein isothiocyanate (FITC)‐conjugated antibodies by addition of anti‐CD90 (0.06 μg/test), CD45 (0.5 μg/1 × 106), CD44 (1 μg/1 × 106 cells), CD29 (1 μg/1 × 106 cells), CD11b/2f/c (0.1 μg/1 × 106 cells), or CD34 (2.5 μg/1 × 106 cells) followed by anti‐Sheep antibody and Alexa Fluor 488 (5 μg/mL). ADSCs treated with FITC‐conjugated mouse IgG, hamster IgG, or sheep IgG served as a negative control. All antibodies were purchased from Thermo Fisher Scientific (Waltham, Massachusetts). The cells in each group were incubated in the dark at 4°C for 10 minutes. After centrifugation and removal of the supernatant, the precipitate was washed twice with PBS, resuspended in 500 μL of staining buffer and then subjected to flow cytometric analysis.
After 10 days of neural differentiation (7 days with ghrelin treatment), the induced cells were subjected to western blotting analysis. Total protein was extracted from cells with RIPA lysis buffer (Solarbio Science & Technology, Co., Ltd., Beijing, China), and quantified using a BCA protein assay kit (Solarbio Science & Technology, Co., Ltd.) according to manufacturer's instruction. Equal amounts of protein were loaded and separated by SDS‐PAGE, and then transferred onto PVDF membranes (Millipore, Billerica, Massachusetts). After blocking with 5% (M/V) skim milk at room temperature (RT) for 1 hour, the membranes were probed with primary antibodies, including glial fibrillary acidic protein (GFAP, 1:1000), β‐Tubulin III (1:1000), phosphorylated form of glycogen synthase kinase (GSK)‐3β (p‐GSK‐3β, 1:1000), GSK‐3β (1:500), P‐AKT (1:1000), AKT (1:500), p‐mTOR (1:400), mTOR (1:1000), and β‐catenin (1:500) were purchased from Cell Signaling Technology (Danvers, Massachusetts), microtubule‐associated protein 2 (MAP2, 1:400) from Bioss (Beijing, China), Nestin (1:500) and GAPDH (1:10 000) from Proteintech Group (Wuhan, China), and Histone H3 (1:5000) from Gene Tex at 4°C overnight. After washing with Tris‐buffered saline and tween20, the membranes were incubated with secondary antibodies goat anti‐rabbit IgG‐HRP (1:3000) or goat anti‐mouse IgG‐HRP (1:3000) obtained from Solarbio Science & Technology, Co., Ltd. at 37°C for 1 hour. The protein bands were detected by ECL kit and analyzed using Gel‐Pro‐Analyzer.
Third‐passage ADSCs were cultured in neural differentiation medium and then treated with different concentrations of ghrelin. After induction for 10 days, the cells in each group were collected for immunofluorescence staining. Cells cultured on coverslips were fixed in 4% paraformaldehyde for 15 minutes. After washing with PBS, cells were permeabilized using 0.1% triton X‐100 (Beyotime Institute of Biotechnology, Haimen, China) and blocked with goat serum (Solarbio Science & Technology, Co., Ltd.) at RT for 15 minutes. The cells were then incubated with primary antibodies against GFAP (1:50, Proteintech Group), Nestin (1:50, Proteintech Group), MAP2 (1:200, CST), β‐Tubulin III (1:100; Abcam, Cambridge, Massachusetts), and β‐catenin (1:100, CST) at 4°C overnight, and immunolabeling was visualized by incubation with secondary antibodies Cy3‐labeled goat anti‐rabbit IgG or Cy3‐labeled goat anti‐mouse IgG (1:200; Beyotime Institute of Biotechnology) at RT for 1 hour. After counterstaining with DAPI (Beyotime Institute of Biotechnology) for nuclei visualization, cells were mounted on slides with fluorescence quenching agent (Solarbio Science & Technology, Co., Ltd.) and observed under a fluorescence microscope at ×200 magnifications (Olympus, Tokyo, Japan).
All data were presented as mean ± SD. Statistical analysis was performed with the use of GraphPad Prism 7.0. The difference between two groups was analyzed by a Student t test and differences among more than two groups using one‐way analysis of variance (ANOVA) followed by a Bonferroni's multiple comparisons test. A P < .05 was defined as statistically significant.
MSCs derived from adipose were isolated and cultured in DMEM/F12 medium containing 10% FBS at 37°C under 5% CO2 in a humidified incubator. Cell cultures were passaged after reaching confluence. Characterization of ADSCs was determined using flow cytometry and multilineage differentiation. Third‐passage ADSCs were subjected to flow cytometry to assess cell surface antigens including CD90 (Figure 1A), CD45 (Figure 1B), CD44 (Figure 1C), CD34 (Figure 1D), CD29 (Figure 1E), and CD11b/2f/c (Figure 1F). The flow cytometric analysis revealed that ADSCs were positive for MSCs markers including CD90 (98.50%), CD44 (95.77%), and CD29 (98.13%), whereas negative for hematopoietic stem cell markers including CD45 (1.71%), CD34 (2.96%), and CD11 (1.27%).
Enlarge this image.Identification of isolated rat ADSCs. Flow cytometry analysis of CD markers including CD90 (A), CD45 (B), CD44 (C), CD34 (D), CD29 (E), and CD11b (F) in third‐passage ADSCs. FITC‐conjugated mouse IgG, hamster IgG or sheep IgG was used as a negative control. ADSCs were positive for MSCs markers including CD90 (98.50%), CD44 (95.77%), and CD29 (98.13%), whereas negative for hematopoietic stem cell marker including CD45 (1.71%), CD34 (2.96%), and CD11 (1.27%). ADSCs were isolated from rat adipose tissues and cultured in DMEM/F12 medium containing 10% FBS at 37°C under 5% CO2 in a humidified incubator. Representative images of cell culture in the third passage (G) and fifth passage (H). I, Osteogenesis of third‐passage ADSCs was induced using osteogenic differentiation medium, and confirmed by Oil Red O staining to detect lipid droplets after 9 days of induction. J, Adipogenesis of third‐passage ADSCs was confirmed by Alizarin Red staining of calcium deposits found in osteoblasts after induction with adipogenic differentiation medium for 3 weeks
Morphological changes of third‐passage and fifth‐passage ADSCs were observed under a light microscope. Third‐passage ADSCs (Figure 1G) presented long fusiform and fibroblast‐like cell morphology and cells in the fifth‐passage (Figure 1H) were uniform with parallel arrangement.
For confirmation of multilineage differentiation, third‐passage ADSCs were induced with osteogenesis and adipogenesis. After induction with osteogenic differentiation medium, there was an increase in Alizarin Red staining of calcium deposits found in osteoblasts (Figure 1I). After induction with adipogenic differentiation medium, Oil Red O staining of induced cells showed presence of lipid droplets suggesting differentiation of ADSCs into adipocytes (Figure 1J). Taken together, these data confirmed that the isolated ADSCs displayed surface expression characterization of MSCs and showed the capability of multilineage differentiation.
Third‐passage ADSCs were cultured in neural differentiation medium and treated with different concentrations of ghrelin (0, 3, 30, and 300 nM). After induction for 3, 6, and 10 days, the cells in each group were photographed and morphological changes were observed (Figure 2A). As the induction time increased, the induced cells displayed a neuronal appearance with extended processes and dendrite‐like projections, which are referred to as neuron‐like cells. In addition, the proportion of neuron‐like cells (Figure 2B) and the number of primary dendrites and branching dendrites per cell were calculated (Figure 2C). After treatment with ghrelin, there were more obvious cell morphological changes into neuronal phenotype as evidenced by and increased number of neuron‐like cells and branching dendrites.
Enlarge this image.Ghrelin promoted the proliferation and neural differentiation of ADSCs. Third‐passage ADSCs were induced to neurogenesis by incubation with neural differentiation medium. After reaching 70% confluence, the cells were treated with 0, 3, 30, or 300 nM ghrelin, and cultured in a humidified incubator with 5% CO2 at 37°C. A, At day 3, 6, and 10 after induction, cells in each group were photographed and morphological changes were observed. B, Quantification of percentage of neuron‐like cells. C, Quantification of the number of primary and branching dendrites per cell. Data were present as mean ± SD (n = 3) and analyzed by a Student t test. *P < .05, **P < .01 vs 0 nM ghrelin
After 10 days of neural differentiation (7 days with ghrelin treatment), the cells were collected and subjected to subsequent detection of neural markers. The differentiation potential of ADSCs into neurons was validated by detecting the expression of neural markers including astrocyte marker, GFAP; neuroectodermal stem cell marker, Nestin; mature neurons marker, MAP2; and immature neurons marker, β‐Tubulin III using immunofluorescence staining and western blot. Immunofluorescence revealed that the positive staining for neural‐specific markers GFAP, Nestin, MAP2, and β‐Tubulin III was significantly enhanced after treatment with 300 nM ghrelin compared to the controls (Figure 3A). Western blotting analysis showed that the protein expression of GFAP, Nestin, MAP2, and β‐Tubulin III was upregulated with increasing concentrations of ghrelin (Figure 3B,C). Compared to those in controls, there was a significant increase in the expression of these neural markers after treatment with 30 and 300 nM ghrelin. The results suggested that ghrelin promoted the neural differentiation of rat ADSCs.
Enlarge this image.Ghrelin enhanced the expression of neural markers in neurogenic differentiated ADSCs. Third‐passage ADSCs were cultured with neurogenic differentiation medium. After induction for 10 days (7 days with treatment of 300 nM ghrelin), the cells were collected and subjected to immunofluorescence (A) and western blot (B and C) for evaluating the expression of neural markers including GFAP, Nestin, MAP2, and β‐Tubulin III. DAPI counter‐staining was performed for nuclear visualization. Data were present as mean ± SD (n = 3) and analyzed by a Student t test. **P < .01 vs 0 nM ghrelin
Nuclear β‐catenin is a key effector of the canonical WNT pathway, and Wnt/β‐catenin signaling has been reported to play an important role in regulation of multi‐lineage differentiation of ADSCs. Thus, we assessed whether it was involved in the effects of ghrelin on ADSCs neural differentiation. After induction of neurogenesis and treatment with ghrelin for 10 days, immunofluorescene and western blot were performed to detect the activation of β‐catenin signaling. Immunofluorescence for analysis of nuclear β‐catenin localization showed that treatment with 300 nM ghrelin significantly enhanced the expression of total β‐catenin, compared to the controls (Figure 4A). Further, western blotting analysis revealed a concentration‐dependent increase in the protein expression of nuclear β‐catenin and p‐GSK‐3β, an inhibited form of the negative regulator GSK‐3β in Wnt/β‐catenin signaling, in ghrelin‐treated groups (Figure 4B,C). There was no difference in GSK‐3β expression among all groups. These results indicated the activation of the Wnt/β‐catenin signaling pathway after treatment with ghrelin.
Enlarge this image.Activation of β‐catenin signaling after treatment with ghrelin. Third‐passage ADSCs were cultured with neural differentiation medium, and different concentrations of ghrelin (0, 3, 30, and 300 nM) were added after 3 days. After 10 days of induction (7 days with ghrelin treatment) (A) immunofluorescence analysis of β‐catenin was performed in both non‐treated and 300 nM ghrelin‐treated groups. DAPI counter‐staining was performed for nuclear visualization. B and C, The protein expression of p‐GSK‐3β, GSK‐3β, and nuclear β‐catenin after treatment with different concentrations of ghrelin was detected by western blot. Data were present as mean ± SD (n = 3) and analyzed by a Student t test. *P < .05, **P < .01 vs 0 nM ghrelin
To further prove that the neural differentiation of ADSCs was achieved through β‐catenin signaling pathway, ADSCs after neural induction by neural differentiation medium for 3 days were cultured in the presence of 300 nM ghrelin with or without 1 μM XAV‐939 (β‐catenin antagonist). Western blot showed that ghrelin‐induced β‐catenin expression was lowered by XAV‐939 (Figure 5A). As shown in Figure 5B‐D, morphological analysis revealed that inhibition of Wnt/beta‐catenin pathway by XAV‐939 suppressed ghrelin‐induced neural differentiation of ADSCs as evidenced by decreased neuron‐like cells and branching dendrites. Immunofluorescence for β‐Tubulin III (Figure 5E), a typical neural marker, showed that the number of β‐Tubulin III‐positive cells was elevated by the ghrelin treatment, whereas the number was lowered by XAV‐939 in the presence of ghrelin. These results suggested that ghrelin induced neural differentiation of ADSCs via activation of β‐catenin signaling pathway.
Enlarge this image.Ghrelin promoted neurogenesis via activation of β‐catenin signaling. After neural induction, ADSCs were cultured in the presence of 300 nM ghrelin with or without 1 μM XAV‐939 (β‐catenin antagonist). A, Western blotting analysis of β‐catenin expression. B, Morphological analysis of cell differentiation at day 10 after induction. C, Quantification of percentage of neuron‐like cells. D, Quantification of the number of primary and branching dendrites in a cell. E, Immunofluorescence for β‐Tubulin III with counterstaining by DAPI and quantification of β‐Tubulin III‐positive cells. Data were present as mean ± SD (n = 3) and analyzed by one‐way ANOVA test. **P < .01 vs 0 nM ghrelin; ##P < .01 vs 300 nM ghrelin
To further exploring the possible underlying mechanism of ghrelin‐induced neurogenesis of ADSCs, we examined the expression of p‐AKT, AKT, p‐mTOR, and mTOR in ADSCs following neural induction in the presence of different concentrations of ghrelin. Western blotting analysis showed that the ghrelin treatment resulted in a concentration‐dependent increase in both levels of p‐AKT (Figure 6A,B) and p‐mTOR (Figure 6C,D). There was no difference in AKT and mTOR expression among groups. Our data suggested the activation of AKT/mTOR signaling pathway following the ghrelin treatment.
Enlarge this image.Ghrelin activated AKT/mTOR signaling pathway. After induction of neurogenesis together with different concentrations of ghrelin treatment (0, 3, 30, and 300 nM), the differentiated cells were harvested for western blotting analysis. A‐B, The expression of p‐AKT and AKT was detected by western blot. C and D, The expression of p‐mTOR and mTOR was detected by western blot. Data were present as mean ± SD (n = 3) and analyzed by a Student t test. *P < .05, **P < .01 vs 0 nM ghrelin
We further assess whether ghrelin‐related activation of AKT/mTOR signaling contributes to neurogenesis. After neural induction by neural differentiation medium for 3 days, ADSCs were incubated in the presence of 300 nM ghrelin with or without 10 nM MK‐2206 (AKT antagonist). The inhibitory efficiency of MK‐2206 was detected by Western blot, and the result showed that the expression levels of p‐AKT and p‐mTOR were upregulated in the presence of ghrelin, while downregulated by addition of MK‐2206 (Figure 7A,B). Analysis of changes in cell morphology showed that ghrelin resulted in higher number of neuron‐like cells with increased primary and branching dendrites, which were lowered after inhibition of AKT/mTOR pathway by MK‐2206 (Figure 7C‐E). Additionally, immunofluorescence for β‐Tubulin III revealed an increase in neuronal cells that were stained positive after ghrelin treatment, while administration of MK‐2206 together with ghrelin decreased β‐Tubulin III‐positive cells (Figure 7F). These findings indicated that ghrelin promoted differentiation of ADSCs into neuronal cells through activation of AKT/mTOR signaling pathway.
Enlarge this image.Ghrelin promoted neurogenesis via activation of AKT/mTOR signaling. Third‐generation of ADSCs were induced to neural differentiation, followed by incubation in the presence 300 nM ghrelin with or without 10 nM MK‐2206 (AKT antagonist). After 10‐day induction, cells were collected for subsequent detection. A and B, Inhibitory efficiency of MK‐2206 on AKT/mTOR activation was determined by western blot. C, Representative images of cell differentiation at day 10 after neural induction. D, Quantification of percentage of neuron‐like cells. E, Quantification of the number of primary and branching dendrites in a cell. F, Immunofluorescence for β‐Tubulin III with counterstaining by DAPI and quantification of β‐Tubulin III‐positive cells. Data were present as mean ± SD (n = 3) and analyzed by one‐way ANOVA test. **P < .01 vs 0 nM ghrelin; #P < .05, ##P < .01 vs 300 nM ghrelin
ADSCs have been considered as ideal source for the treatment of nerve injuries or neurodegenerative disease. Administration of ADSCs improves peripheral nerve regeneration in a rat model of severe nerve injury. Indeed, knowledge of the importance of nerve regeneration has encouraged researchers to develop novel strategies to promote neural differentiation of ADSCs. In our previous study, we demonstrated that ADSCs promoted nerve regeneration and functional recovery in a rat model of sciatic nerve injury. In the present study, we aimed to study the effects of ghrelin on neural differentiation of ADSCs. Rat ADSCs were isolated and identified by positive for MSCs markers CD90, CD44, and CD29, and negative for hematopoietic stem cell markers CD45, CD34, and CD11b/2f/c, as well as multilineage differentiation potential into adipocytes and osteocytes. Different concentrations of ghrelin were applied to intervene in neurogenesis. The ghrelin treatment was found to promote neural differentiation of ADSCs along with increased neural marker expression in a concentration dependent manner. β‐Catenin and AKT/mTOR signaling pathways were activated after the ghrelin treatment. Thus, we demonstrate that ghrelin might promote neurogenesis of ADSCs through AKT/mTOR and β‐catenin signaling pathways.
Numerous studies have shown the differentiation potential of ADSCs into neuron‐like cells under neuronal induction conditions. Morphologically, our isolated rat ADSCs after induction of neurogenesis developed spherical or elliptical shapes with extended processes and dendrite‐like projections arising from cell bodies that is referred as neuron‐like cells, which was in accordance with previously reports. Based on morphological data, immunofluorescence and western blot were performed to examine the expression of multiple neural lineage markers including GFAP, Nestin, MAP2, and β‐Tubulin III. In the present study, ADSCs cultured in neural differentiation medium were clearly immunopositive for GFAP, Nestin, MAP2, and β‐Tubulin III after treatment with ghrelin, compared to the controls. Further, ghrelin‐induced neurogenesis of ADSCs was further examined by western blotting analysis as evidenced by upregulated protein expression of GFAP, Nestin, MAP2, and β‐Tubulin III after exposure to ghrelin. These findings indicated that the ghrelin treatment could significantly promote neural differentiation of ADSCs. Our data are in agreement with previous reports indicating that human ADSCs under neural induction conditions are able to develop a neuronal appearance and result in being positive for GFAP, Nestin, and type β‐Tubulin III. Therefore, the ghrelin treatment enhances neural differentiation of ADSCs.
β‐Catenin is a key target of the canonical WNT pathway that controls numerous developmental processes. Increasing studies reported that ghrelin regulates cell apoptosis, proliferation, and function via activation of Wnt/β‐catenin pathway in a variety of cells and tissues. Activation of β‐catenin signaling is important for neuronal differentiation, including dendritic development, axon growth and guidance, neuronal plasticity, and synaptic function. Mrak et al. revealed that ghrelin promoted osteoclastogenesis via enhancement of p‐GSK‐3β and β‐catenin levels. Ghrelin suppressed cell apoptosis by regulating Wnt/β‐catenin pathway. In consistent with previous studies, we have identified that the activation of β‐catenin pathway contributes to the beneficial effects of ghrelin on neurogenesis of ADSCs. Here we showed that the expression of nuclear β‐catenin and p‐GSK‐3β was increased with increasing concentrations of ghrelin. After treatment with ghrelin, an increase in p‐GSK‐3β, an inhibited form of the negative regulator GSK‐3β in Wnt/β‐catenin signaling, suggested that ghrelin might upregulate β‐catenin level by downregulation of its inhibitory pathway, and thus increased the accumulation of cytoplasmic β‐catenin leading to elevation in translocation of β‐catenin into nucleus. Further, inhibition of Wnt/β‐catenin signaling resulted in a suppression of ghrelin‐induced neurogenesis as evidenced by less presence of neuron‐like cells and reduced β‐Tubulin III expression. These results suggest an activation of β‐catenin signaling after the ghrelin treatment, which thus contributes to neurogenesis of ADSCs.
The AKT/mTOR signaling pathway is important for cell growth, survival, proliferation, angiogenesis, translation, transcription, and metabolism. mTOR is a downstream target of the P13K/AKT signaling pathway, which is activated through phosphorylation by activated AKT. Studies have reported that AKT/mTOR plays an important role in cell differentiation. For example, mTOR1 signaling promoted osteoblast maturation and differentiation. On the contrary, osteogenic differentiation was enhanced along with decreased phosphorylation level of mTOR. Inhibiting of the P14K/AKT/mTOR signaling pathway resulted in enhancement of muscle differentiation. The contradictory effect of the AKT/mTOR signaling pathway on cell differentiation remains unclear, and it is possible due to different characteristics of mTORC1 and mTORC2 resulting in different effect on different cell types or different differentiation stages. However, the role of AKT/mTOR in neural differentiation of ADSCs has not been reported yet. Our study showed that ghrelin could upregulated the phosphorylation levels of AKT and mTOR. Moreover, we observed a concentration‐dependent promotion effect of ghrelin on neural differentiation of ADSCs. A study showed that activation of P13K/AKT/mTOR pathway with an increase in the phosphorylation levels of AKT and mTOR contributes to osteoclast differentiation. In the present study, ghrelin‐induced neural differentiation of ADSCs was inhibited by deactivation of AKT/mTOR signaling. These findings indicate that ghrelin promote neurogenesis via activating AKT/mTOR pathway.
In conclusion, ghrelin promotes neural differentiation of ADSCs via activation of AKT/mTOR and β‐catenin signaling pathways, suggesting that ghrelin may be a promising candidate for improving the efficacy of MSCs‐based therapy in the treatment of PNI.
All authors declare no conflicts of interest.
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1 Department of Anatomy, School of Basic Medical Sciences, Mudanjiang Medical College, Mudanjiang, People's Republic of China; Institute of Neural Tissue Engineering, Mudanjiang Medical College, Mudanjiang, People's Republic of China
2 Department of Anatomy, School of Basic Medical Sciences, Mudanjiang Medical College, Mudanjiang, People's Republic of China; Key Laboratory of Cancer Prevention and Treatment of Heilongjiang Province, Mudanjiang Medical College, Mudanjiang, People's Republic of China
3 Department of Medical Imaging, Hongqi Hospital of Mudanjiang Medical College, Mudanjiang, People's Republic of China
4 Department of Infectious Diseases, Hongqi Hospital of Mudanjiang Medical College, Mudanjiang, People's Republic of China
5 Department of Anatomy, School of Basic Medical Sciences, Mudanjiang Medical College, Mudanjiang, People's Republic of China
6 Key Laboratory of Cancer Prevention and Treatment of Heilongjiang Province, Mudanjiang Medical College, Mudanjiang, People's Republic of China; Pathology Diagnosis Center, The First Clinical Medical School of Mudanjiang Medical College, Mudanjiang, People's Republic of China
7 Key Laboratory of Cancer Prevention and Treatment of Heilongjiang Province, Mudanjiang Medical College, Mudanjiang, People's Republic of China; Pathology Diagnosis Center, The First Clinical Medical School of Mudanjiang Medical College, Mudanjiang, People's Republic of China; Institute of Stem Cells, Mudanjiang Medical College, Mudanjiang, People's Republic of China