Transcranial Optogenetic Stimulation Promotes Corticospinal Tract Axon Regeneration to Repair Spinal Cord Injury by Activating the JAK2/STAT3 Pathway
Article information
Abstract
Graphical Abstract
Abstract
Objective
Regeneration of corticospinal tract (CST) axons after spinal cord injury (SCI) is a key element in rebuilding neuronal connections to restore voluntary motor function. However, it remains challenging owing to limited effective interventions. This study adopted a modified transcranial optogenetic technique to stimulate CST axon regeneration into the injury site of completely transected SCI and explore the underlying molecular mechanisms.
Methods
A novel optogenetic light emitting diode (LED) device was used to stimulate the brain motor cortex in channelrhodopsin-2–yellow fluorescent protein (ChR2-YFP) transgenic mice to observe the regeneration of CST axons in the injury site of a complete SCI. The LED device was also used in vitro to stimulate the motor cortex slices of the transgenic mouse brain for observing the outgrowth of their neurites.
Results
After transcranial optogenetic stimulation, the pyramidal neurons of bilateral cerebral motor cortices, in ChR2-YFP transgenic mice were activated, CST axons regenerated into the injury site of the spinal cord, and the motor function of the paralyzed hindlimbs improved. Proteomic analysis revealed that CST axon regeneration was associated with the activation of the Janus kinase 2/signal transducer and activator of transcription 3 (JAK2/STAT3) pathway in the cerebral motor cortices. In vitro LED blue light illumination enhanced the outgrowth of neurites from the brain slices of transgenic mice. Treatment with a JAK2/STAT3 inhibitor led to a significant attenuation of neurite outgrowth.
Conclusion
The modified transcranial optogenetic technique stimulated bilateral motor cortices, in the brains of ChR2-YFP transgenic mice. It increased the excitability of pyramidal neurons in the motor cortices, and promoted CST axon regeneration by activating the JAK2/STAT3 pathway, repairing complete SCI.
INTRODUCTION
The corticospinal tract (CST), the descending nerve fibers connecting the pyramidal neurons of the motor cortex and with the motor neurons in the anterior horn of the spinal cord, is essential for controlling voluntary movements of the body [1]. However, CST axons have a limited regenerative capacity after injuries such as spinal cord injury (SCI) [2]. Several putative mechanisms have been proposed to underlie the failure of CST axon regeneration after SCI, such as inadequate chemoattraction, [3], glial scar formation [4], the existence of inhibitory molecules [5] and the lack of intrinsic growth capacity [6]. A therapeutic strategy targeting the protein translation control latter has been developing rapidly since the discovery of conditional depletion of the phosphatase and tensin homolog (PTEN) to alleviate its inhibitory effect on the mammalian target of rapamycin (mTOR) pathway, enabling long-distance regeneration of CST axons in mice with hemisected SCI [7]. Additionally, retaining the embryonic transcriptional state of the motor cortex neurons by grafting spinal cord-derived neural progenitor cells at the injured site leads to robust regeneration of CST and restoration of forelimb function post-SCI [8]. Overall, these findings suggest that manipulating the genetic or functional state of cortical motor neurons might enhance their intrinsic regenerative power and aid in axon regeneration post-SCI.
Physical stimulation has been shown to effectively promote CST regeneration by altering the function or activities of the motor cortex. For example, electrical stimulation to the cerebral cortex of rats has been found to increase the collateral sprouting and regeneration of CST axons in the injured spinal cord [9]. Similarly, high-frequency repetitive transcranial magnetic stimulation has been shown to enable the collateral sprouting and regeneration of CST axons after dorsal hemisected SCI by activating the BRAF-MAP2K1/2 pathway in cortico-motor neurons [10]. Intermittent theta burst stimulation can produce lasting muscle evoked potential enhancement and be accompanied by CST axonal outgrowth and structural changes at the CST-spinal interneuron synapse [11]. However, the precise mechanisms underlying the impact of electrical or magnetic stimulation on CST axon regeneration remain unexplored since several cells could be responsive to such physical intervention, including, but not limited to, neurons and glial cells [12]. Therefore, the relationship between corticospinal neuron activation and CST regeneration after physical stimulation needs to be elucidated. Optogenetic technology offers potent resolution to assess the gain-of-function stage of specific neurons [8]. Optogenetic stimulation allows selective activation of neurons other than glial cells, highlighting the exclusive function of targeted neurons [9]. After transcranial optogenetic stimulation, the CST axons have been shown to develop robust collateral sprouting and synaptic plasticity in the midbrain red nucleus after contusive SCI [13] or hemisected SCI [14]. However, CST axon regeneration was not observed at the site of SCI, which might be attributed to the inadequacy of the irradiance power of the brain motor cortex treated with the transcranial optogenetic technique. However, despite the encouraging research progress, whether optogenetic stimulation leads to CST regeneration in a completely transected SCI remains unclear. Furthermore, key molecules underlying optogenetic stimulation-induced CST axon regeneration still need to be explored, although several putative candidates have been hypothesized [15].
Pharmacology and stem cell therapy have emerged as promising approaches in the treatment of SCI [16,17]. Optogenetics offers a precision and spatiotemporal control that is not always achievable with traditional pharmacological methods, which can suffer from issues of drug delivery, metabolism, and off-target effects. Similarly, while stem cell therapies hold promise for regenerative medicine, they are still subject to challenges such as cell survival, integration, and ethical considerations. Transcranial optogenetic stimulation, in comparison, provides a non-invasive strategy that can potentially activate central neural pathways, promoting functional recovery without the need for systemic drug administration or exogenous cell transplantation. However, each approach has its unique advantages and limitations, and a multimodal treatment strategy that combines the strengths of optogenetics, pharmacology, and stem cell therapy may ultimately provide the most effective path to recovery for individuals with SCI.
Therefore, in this study, we employed a modified transcranial optogenetic technique for stimulating the motor cortices of the mice brains to enhance the excitability of pyramidal neurons, promote CST axon regeneration into the injury site of completely transected SCI, and explore the possible molecular mechanisms underlying CST axon regeneration. The present study might provide a novel theoretical basis for enhancing the regeneration of CST axons at the injury site after complete SCI via a modified transcranial optogenetic technique.
MATERIALS AND METHODS
1. Ethics Statement
All experimental protocols and animal handling procedures were approved and supervised by the Animal Care and Use Committee of Sun Yat-sen University (IACUC# SYSU-2019-B551; SYSU-2019-B943) and aligned with the Laboratory Animal Regulations of Guangdong Province (2010 No. 41). Animal welfare complied with the Laboratory Animal Guidelines for Ethical Review of Animal Welfare, General Administration of Quality Supervision, Inspection and Quarantine of the People’s Republic of China/Standardization Administration of China (GB/T 35892-2018).
2. Experimental Animals
A total of 64 female thy-1 channelrhodopsin 2-yellow fluorescent protein (ChR2-YFP) transgenic adult mice and 5 wildtype (WT) adult C57 mice (6–8 weeks old, each weighing 18–22 g) were used in the present study. The ChR2 transgenic mice (B6. Cg-Tg [Thy1-COP4/EYFP] 18Gfng/J, Strain #: 007612) were purchased from the Jackson Laboratory. The WT mice were obtained from the Experimental Animal Center of Sun Yat-sen University.
3. Assembly and Implantation of the Optogenetic Stimulation Device
Our lab-made optogenetic stimulation device comprised 3 major components: a blue light (wavelength: 473 nm) LED bulb (Beike Ltd., China), a pair of electrodes (pure copper wire, 1.5 mm2; BVR, China) and the wires connecting them, and a waveform generator (JSD2900, Juntek Ltd.). The electrode was welded to the back of the LED bulb with a soldering iron soldering gun (P908, ELECALL, China). During the experiment, the electrodes on the LED bulb on the animal’s head were assembled by connecting the waveform generator with the alligator clip. At the end of the experiment, the crocodile clip can be removed, and the stimulation device can be restored and repositioned. The wire connection points of the LED bulb were sealed with ethylene vinyl acetate copolymer hot-melt adhesive (DL5042, Deli Ltd.).
After anesthesia with 1% isoflurane, the mice were fixed in a stereotaxic frame (68044, RDW, China), and an incision of 1.5 cm was made on the middle line of the scalp toward the top to expose the skull. After gently removing the connective tissue, the bregma was identified under a stereomicroscope. The skull was thinned by a dental drill till the underlying blood vessels on the brain surface could be clearly visualized (same criteria for 2-photon confocal microscopy [18]). The thinned area of the skull was 1 mm caudal to the bregma, with dimensions of 3×10 mm. The pre-fabricated LED bulb (3×3 mm2) was placed on top of the thinned skull and was fixed by a pair of stainless steel screws. The screws were fixed carefully to avoid penetrating the skull and damaging the brain parenchyma. The LED bulb and the screws were completely embedded in dental resin. The entire device was held with a tweezer while waiting for the resin to harden. The stimulation parameters were modified based on the reference [19] as follows: complementary metal oxide semiconductor (CMOS) pulse style; voltage 4.5 V; frequencies 5, 10, 20, 40, or 80 Hz; pulse width 3 msec. The duty cycle for each stimulation is 0.25, with a 1-minute stimulation period followed by a 3-minute rest period. The luminous flux was 4,500±200 lux, measured using a digital split-type illuminometer (AS823, Smart Sensor Ltd.). The maximum power of the LED bulb was 90 mW. For in vivo optogenetic stimulation, the mice were treated for 40 minutes daily at the same time for 14 days consecutively.
4. Optogenetic Stimulation of Organotypic Cultured Brain Slices
Organotypic cultured brain slices were prepared according to the previous studies [20,21], with slight modifications. Briefly, after anesthesia, the whole brain of a ChR2-YFP transgenic adult mouse was dissected and immersed in a cold Hank’s balanced salt solution. The motor cerebral cortex was separated under a stereomicroscope, cut into slices of 300- to 500-μm thickness using a pair of parallel razor blades, and then cut into smaller pieces of about 1×1 mm2. The brain slices from the ChR2-YFP mouse (ChR2 group and ChR2+OS group) and wild-type mouse (WT+OS group) were cultured in a 35-mm culture dish (tissue culture treated, Corning), containing neurobasal-A medium containing 2% fetal bovine serum, 20-ng/mL brain-derived neurotrophic factor, and 2% B27, with medium renewal every other day. In addition, some of the brain slices from the ChR2-YFP mouse in the OS+FLLL31 group were cultured in a medium containing FLLL31, a tetramethyl curcumin that specifically inhibits STAT3 phosphorylation [22]. For optogenetic stimulation, a 3.5×3.5-mm2 window was created on the lid of a culture dish, and an LED bulb was mounted into the window and fixed with hot-melt adhesive. The stimulation protocol was as follows: Wavelength of blue light: 473 nm; frequencies: 5, 10, 20, 40, or 80 Hz; duration of each stimulation: 3 msec; duty and interval of each cycle were 1 and 3 minutes, respectively; and total stimulation time: 40 minutes. To validate neuronal activity postoptogenetic stimulation, patch clamps were used to detect the action potential from pyramidal neurons in the cortex slices immediately after blue light stimulation in the WT+OS group and the ChR2+OS group or OS group. The blue light-evoked firing was recorded in the current-clamp mode using pipettes (2–5 MΩ) filled with potassium-gluconate-based internal resistance solution containing 130 mM potassium gluconate, 2 mM MgCl2, 5 mM KCl, 0.6 mM EGTA, 10 mM HEPES, 2 mM Mg-ATP, and 0.3 mM Na-GTP (pH 7.2–7.3; solution osmolarity: 270–285 mOsm).
5. SCI Model
Adult female ChR2-YFP transgenic adult mice and WT mice were used for the SCI model. On the day of surgery, the mice were anesthetized via inhalation of 1%–3% isoflurane. After fixation and skin disinfection, the skin and superficial fascia were cut under sterile conditions. The laminectomy was carried out at the T9−10 vertebral level, and the dura was cut using microscissors. The spinal cord was transected completely with angled microscissors, and a 1-mm segment of the spinal cord at the T10 level was removed. After adequate hemostasis, musculature and skin were sutured sequentially. After surgery, each animal was injected intramuscularly with 160,000 units of penicillin at 1 mL/day for 3 consecutive days. Meloxicam (2 mg/kg, subcutaneous injection) was used as an analgesic for 3 consecutive days. Single cage feeding was applied for each mouse after surgery to reduce stress. Artificial urination was induced with moderate pressure by hand in the bladder area, 1–2 times a day. After that, based on the recovery of bladder function, the frequency of urination gradually reduced untill automatic urination was restored. Two weeks after the surgery, a group of mice were subjected to optogenetic stimulation (the OS group) for 2 weeks. Another group of mice were not stimulated by optogenetics as the control group (the nOS group). All mice survived 6 weeks after the surgery.
6. Tracing CST Axons Using a Biotinylated-Dextran Amine Injection
After optogenetic stimulation for 2 weeks, the biotinylated-dextran amine (BDA) was used to anterogradely trace the CST in the nOS and OS groups. Briefly, the head of a mouse was fixed to the stereotaxic apparatus, and the LED bulb device was gently removed. Following the approach of previous studies [23], BDA injection was conducted to trace the CST axons. The skull was trephined using a dental drill to create 2 circular bone windows, each approximately 3 mm in diameter. Then, a 10% BDA solution (molecular weight: 10,000, Invitrogen) was mounted with a 10 μL Hamilton microsyringe (Model 700 RN/33G/15MM, No. 21-2063). Six injection points were distributed on each side of the brain with the following coordinates: 1.0 or 2.0 mm posterior to the bregma; 0.5, 1.0 and 2.0 mm lateral to the sagittal suture; and 0.5 mm deep from the surface of the cerebral cortex. The blood vessels were avoided at the time of injection. The volume at each injection point was 0.4 μL. Therefore, each mouse received a total of 4.8-μL BDA. The microsyringe needle was retained at the injection point for 5 minutes after each injection and then slowly withdrawn to reduce backflow. After injection, the scalp was sutured, and the mice were routinely fed for 2 weeks. Each animal was injected intramuscularly with 160,000 units of penicillin at 1 mL/day for 3 consecutive days after the surgery. They survived 2 weeks after receiving the BDA injection. The brain and spinal cord tissue were harvested after the animals were perfused. The extracted tissues were stained with streptavidin-conjugated Alexa Fluor 555 (Supplementary Table 1).
7. Immunofluorescence Staining
After euthanasia, the mice were intracardially perfused with 4% paraformaldehyde. Then, their brains and spinal cords were quickly dissected out, fixed in the fixing solution for 72 hours at 4°C, and transferred to a series of 10%−30% (w/v) sucrose solutions (at 4°C) for dehydration for a week. The fixed tissues were cut into 20-μm-thick sections using a cryostat. Immunofluorescence staining was performed as described previously [24]. Briefly, the frozen sections were rewarmed, rinsed with 0.01 M phosphate-buffered solution (PBS), blocked with 10% goat serum for 30 minutes at 37°C, and incubated with different primary antibodies, supplemented with 0.3% Triton X-100, overnight at 4°C. Next day, the sections were rinsed with PBS and incubated with secondary antibodies for 1 hour at 37°C. Hoechst33342 was used to counterstain cell nuclei where necessary. Finally, the slides were observed under a confocal microscope (Zeiss LSM800).
All the antibodies and fluorescent dye reagents used in this study, including primary and secondary antibodies, are listed in Supplementary Table 1.
8. Western Blot Analysis
After intracardiac perfusion, the skull of each mouse was opened, and the brain was dissected on ice. The motor cortices, subjected to blue light stimulation, was carefully removed from the brain, transferred into the centrifugal tube, cut into small pieces with iris scissors and dissolved in a cell lysate cocktail (containing 1% protease inhibitor) on ice for 10 min. Ultrasonic homogenization was applied until no obvious tissue mass was visible. The lysate samples were centrifuged at 12,000 rpm for 30 minutes in a centrifuge pre-cooled at 4°C. The supernatant was carefully collected using a pipette. The bicinchoninic acid (BCA) method was used to measure protein concentration according to the BCA Protein Quantitation Kit (Beyotime, P0012S) instructions. Briefly, the protein samples were mixed with a loading buffer and denatured in boiling water for 5−6 minutes. Then, the samples were cooled and stored at -80°C for further analysis. Later, the protein samples were removed from the freezer, separated using 8% sodium dodecyl sulfate-polyacrylamide gel electrophoresis gel, and then transferred onto polyvinylidene difluoride (PVDF) membranes. The PVDF membranes were then blocked with 5% BSA dissolved in tris-buffered saline with Tween 20 (TBST) for 1 hour at room temperature. The membranes were then cut according to the molecular weights of the protein targets. The membranes were then washed thrice with TBST solution for 10 minutes. Next, the membranes were incubated overnight at 4°C with primary antibodies (anti-p-JAK2 and anti-p-STAT3) incubated with TBST. Then, the membranes were washed thrice with TBST for 10 minutes and incubated with the corresponding secondary antibodies (horseradish peroxidase) for 1 hour at room temperature. Again, the membranes were washed thrice with TBST washing solution for 10 minutes. Finally, the membranes were photographed. After exposure and development, the images were quantified using ImageJ software (https://imagej.nih.gov/ij/download.html, 1.53).
9. Proteomic Analysis
Mouse brain proteins were extracted from the OS (n=3) and the nOS group (n=3) and subjected to western blotting as described in Section 2.8. Liquid chromatography-mass spectrometry (MS)/MS analysis was performed, and the resulting MS/MS data were processed using the Proteome Discoverer search engine (v2.4). MaxQuant software Mus_musculus_10090_SP_20201214. fasta database was used to identify the protein sequence and assess the significance of the difference. The relative quantitative value in the comparison group was assessed using the t-test, and p<0.05 indicated significant differences. Differential expressions of >1.3 and <1.3 indicated significantly upregulated and downregulated differential proteins, respectively. The differential protein gene ontology analysis and Kyoto Encyclopedia of Genes and Genomes (KEGG) enrichment pathway analysis were performed using the Fisher exact test method, and the difference was statistically significant at p<0.05.
10. Electrophysiology and Behavioral Scores
Four weeks after the surgery, the mice in the nOS and OS groups were anesthetized by isoflurane. They were fixed to the stereotaxic apparatus with a backup position. The skin surrounding the femur was cut, and the sciatic nerve was exposed. The tips of the electrodes were bent into a hook shape and hooked onto the sciatic nerve. Then, the wires from the electrodes were connected to an electrophysiological instrument. When the cerebral cortex of mice was stimulated by light, this instrument was used to observe the response of the brain-derived descending neural pathway to transmit neural information.
Hindlimb motor function of mice was assessed according to the Basso mouse scale (BMS) scoring method proposed by Basso et al. [25] Briefly, the bladder of a mouse was emptied before assessment and placed in an open field one day before the surgery to familiarize them with the environment. The BMS scores for each mouse were evaluated 1 hour before the surgery and the third day after the surgery. The mice were assessed at the same time every week, from the seventh day after the surgery until the end of the sixth week. During this period, the mice were placed in the same open field and allowed to explore. Two independent investigators who did not know the specifics of the treatment observed the hindlimb movement frequency, range of joint movement, coordination, ankle movement, plantar and instep landing, trunk stability, and tail position of the mice. The data were recorded for 4 min each time, and the scores ranged from 0 to 9 (with 0 being completely paralyzed and 9 being completely normal).
11. Morphological Quantification
The proportion of activated neurons was analyzed by calculating the proportion of C-fos+ cells among the Map2+ cells. Five ChR2-YFP transgenic animals were stimulated by the LED bulb device and perfused after 90 minutes of optogenetic stimulation (n=5). Their brains were obtained and cryosectioned (in 30-μm-thick slices), and 5 sections (1 in 5 intervals) were selected from each animal. Five random fields (at ×200 magnification) were selected from each section. The number of Map2 and C-fos double-positive cells and the total number of Map2+ cells were calculated, and their percentage was assessed as the proportion of the activated neurons.
To assess the specificity of axon growth induced by the optogenetic stimulation, 5 random fields (at ×200 magnification) were selected and sampled from the brain slice, and the percentage of the number of neurofilament-200 (NF) and ChR2-YFP (ChR2) double-positive neurites among the total number of neurites observed in bright field was calculated (n=5). The number (points) of neurites and the maximum length of neurite extension in the brain slice were analyzed by directly counting the neurite number and measuring the neurite length of the dorsal root ganglion as described previously [26]. Briefly, some circles were drawn around the brain slice culture, and the crossover points between the neurites and circles were counted to obtain the number of neurites. Meanwhile, a circle was drawn around the culture, and another circle was drawn near the farthest neurites. The distance between the 2 circles indicated the maximum length of neurites.
CST axon regeneration in the nOS and OS groups was quantitatively analyzed. Considering the individual differences of each animal and the heterogeneity of the injection procedures, the CST axon number in the lesion area and caudal area were normalized by the total number of BDA-labeled CST axons in the same animal and converted into an axon number index for analysis. The axon number index, defined as the number of BDA-labeled (BDA+) CST axons at the injury site and the area caudal to the injury site of the spinal cord divided by the total number of BDA+ CST axons in a transected slice of the cervical segment 1 (C1) [27]. Transverse slices (20 μm thick) of the spinal cord C1 segment were obtained at intervals of one in three. Five slices were obtained from each mouse, and the mean value was calculated. All horizontal sections of the T10 segment containing CST axons of a mouse were obtained, and the number of BDA+ CST axons with a length greater than 20 μm in the injury site and caudal area (within 2 mm) of the lesion area was counted. The axon number index was obtained by dividing the number of CST axons in the lesion area and caudal area by the number of CST axons in segment C1. The axon number indexes for both groups were compared and analyzed (n=6 for each group).
12. Statistical Analysis
All quantitative data were reported as mean±standard deviation and analyzed using IBM SPSS Statistics ver. 22.0 (IBM Co.). All data were examined for normality using the Shapiro-Wilk test. When the assumptions of a normal distribution were met, the data were analyzed using Student t-test for comparisons between the 2 groups and 1-way or 2-way analysis of variance for comparisons between multiple groups. When the variance was homogeneous, the Student-Newman-Keuls post hoc test was employed for multiple comparisons. Otherwise, Dunnett’s T3 post hoc test was used. The nonparametric test (Kruskal-Wallis) was used for nonnormally distributed variable values. The significance level was set at p<0.05.
RESULTS
1. Transcranial Optogenetic Stimulation Activated the Cerebral Motor Cortexes
In this study, we developed a novel LED-based optogenetic stimulation device to activate bilateral cerebral motor cortexes (Fig. 1A). The device can be noninvasively without inflicting structural damage to the brain parenchyma and is thus suitable for a chronic functional study. It is pivotal to grind the skull as thin as possible to achieve a sufficient flux of photons. Once the blood vessels in the meninges were clearly visible (Fig. 1B), approximately 95% of photons (4,300–4,500 lux) could pass through the skull. ChR2-YFP transgenic mice with channel rhodopsin-2 expression in the fifth layer of the motor cortexes, the point of origin of CST axons, were used for the present study. The mice tolerated well with the mounted LED light stimulation device fixed firmly on the skull surface without discernible signs of stress (Fig. 1B). The pyramidal neurons were marked by green fluorescence emission from transgenic ChR2-YFP fusion protein (ChR2) (Fig. 1C). Ninety minutes after optogenetic stimulation of the brain of the mice, coronal sections of the brain were made to verify the LED blue light activation on the motor cortexes (Fig. 1D). Immunofluorescence staining showed that a large portion of ChR2+/Map2+ pyramidal neurons in the fifth layer of the motor cortexes expressed immediate early gene product C-fos (a cell marker for robust activities) (Fig. 1E). From the results of quantitative analysis, 90.90%±2.50% of Map2+ neurons expressed C-fos (Supplementary Fig. 1). As a control experiment, in the absence of the blue light stimulation, the expression of C-fos was not observed in the ChR2+/Map2+ pyramidal neurons on the coronal sections of the brain of ChR2-YFP transgenic mice (Supplementary Fig. 2). The above results show that the ChR2+ pyramidal neurons can be activated by LED blue light specifically.
Development of LED-based optogenetic stimulation device and the motor cortex neurons activated by blue light. (A) LED bulbs and electrodes. Blue light (473 nm) was emitted when the LED bulb was connected to the waveform generator. (B) A mouse skull before and after bone grinding. Meningeal blood vessels can be clearly observed (white arrow, upper on the right). The LED light stimulation device was installed on the mouse head (lower on the left), and blue light is visible when the device is on (lower on the right). (C) Pyramidal neurons in the fifth layer of motor cortexes of the ChR2-YFP transgenic mice showing green fluorescence (the image on the right is an enlargement of the image on the left). (D) A schematic diagram showing bilateral motor cortexes illuminated by LED blue light and the corticospinal tracts (red) connecting bilateral motor cortexes and spinal cord. (E) Co-localization of C-fos (red) and Map2 (white) within ChR2+ pyramidal neurons after LED blue light stimulation. (E) Panels E1 and e showing C-fos expressed in a Map2+ neuron (white arrow in the box). Scale bars=1 mm in (C), 150 μm in (E), 80 μm in (E1), and 10 μm in (e). LED, light emitting diode.
2. In Vitro Optogenetic Stimulation Enhanced Neurite Outgrowth From Brain Slices
Whole-cell patch clamps were used to assess electrophysiological changes in the brain slices of ChR2-YFP transgenic mice subjected to blue light stimulation (Fig. 2A). Indeed, the action potentials induced by recorded neurons in the brain slices were identical in frequency (5 Hz) to those subjected to blue light stimulation, suggesting that ChR2-expressing neurons in the brain slices could be efficiently activated by the device (Fig. 2B). Furthermore, a lab-made organotypic brain slice culture system was developed to detect the effect of optogenetic activation on axon growth in vitro (Fig. 2C). This system was used to evaluate the specificity effect of optogenetic stimulation on the neurite growth of the neurons. In the brain slices, most growing neurites induced by blue light stimulation appeared double-positive for NF (a marker of nerve fibers) and ChR2 (NF+/ChR2+, > 95%), indicating the specificity of optogenetic stimulation on the ChR2+ neurons (Fig. 2D–E). In the ChR2+OS group, the number and length of the neurites around the brain slices increased after exposure to blue light stimulation (Fig. 2F). Compared to the ChR2+OS group, fewer neurites were observed around the brain slices in the ChR2 (not exposed to blue light stimulation) and the WT+OS (exposed to 10-Hz blue light stimulation) groups (Fig. 2F). The growth of single neurite derived from the brain slices in the ChR2, WT+OS, and ChR2+OS groups was respectively observed under phase contrast microscopy for 8 hours in real time and the time interval for each frame is 40 minutes. The growth cone of the neurite from brain slices exhibited a distinct growth trajectory in the ChR2+OS group (Supplementary Fig. 3). However, the growth cone of the neurite showed stagnation in the ChR2 and WT+OS groups (Supplementary Fig. 3). To explore the proper frequency for neurite growth, the light stimulation protocol involved exposure of the ChR2+OS group to a series of escalating frequencies (5, 10, 20, 40, and 80 Hz). The groups exposed to blue light stimulation exhibited a significant increase in the number and length of neurites compared to the ChR2 and WT+OS groups (n=5, p<0.05) (Fig. 2G). Moreover, the ChR2+OS groups exposed to 10- to 80-Hz blue light stimulation exhibited significantly longer neurites than the group exposed to 5-Hz blue light stimulation (n=5, p<0.05) (Fig. 2G). However, the groups exposed to 10- to 80-Hz blue light stimulation did not exhibit significant differences in neurite length (n=5, p>0.05) (Fig. 2G). Taken together, these results indicated that blue light stimulation directly activated ChR2+ neurons in the brain slices, promoting neurite outgrowth. Since 10 Hz of blue light stimulation conferred good performance in enhancing neurite growth, this parameter was used for optogenetic stimulation during in vivo experiments.
Optogenetic stimulation promoted the outgrowth of neurites in vitro. (A) Giga seal of a ChR2+ cell (arrow) inside the brain slice with a glass micropipette. (a) showing electrophysiological instrument recording data from the brain slice exposed to blue light stimulation. (B) Evoked action potential recorded at a ChR2+ neuron derived from the brain slice. Note that the rhythmic waves were induced by the 5-Hz blue light stimulation. (C) A schematic diagram showing the lab-made organotypic brain slice culture system under non-light stimulation in the ChR2 group and the exposure of the WT+OS and ChR2+OS groups to blue light stimulation. (D) Bright field (BF) and immunostaining images revealing NF+/ChR2+ neurites (arrows) around the brain slice in the ChR2+OS group. (E) Pie chart showing the proportion of NF+/ChR2+ neurites among the total neurites. (F) Few neurites (arrows) around the brain slice in the ChR2 and WT+OS groups and a higher quantity of neurites (arrows) in the ChR2+OS group. (G) Bar charts displaying the number and length of neurites in the brain slices of the ChR2 group, WT+OS group (exposed to 10-Hz blue light stimulation), and various ChR2+OS groups (exposed to 5-, 10-, 20-, 40-, and 80-Hz blue light stimulation). For multiple comparisons, 1-way analysis of variance was used, followed by the Student-Newman-Keuls post hoc test. *p<0.05 when compared with the ChR2 group, #p<0.05 when compared with the WT+OS group, &p<0.05 when compared with the ChR2+OS group (5 Hz), and n=5 (brain slices) in each group. Scale bars=200 μm in (D) and 100 μm in (F). ChR2, channelrhodopsin-2; WT, wild-type; OS, optogenetic stimulation; NF, neurofilament-200.
3. Optogenetic Stimulation Enhanced CST Axon Regeneration in Mice With Completely Transected SCI
Two weeks after receiving optogenetic stimulation in a subacute phase of the complete SCI (2 weeks post-SCI) in the ChR2-YFP transgenic mice, BDA (a nontransynaptic neural trace) was injected into their bilateral motor cortexes for anterograde tracing of CST axons (Fig. 3A). Several BDA-labeled (BDA+) red fluorescent CST axons were detected in the cerebral cortex and spinal cord, signifying the effectiveness and specificity of the BDA labeling technique (Fig. 3B–C). Thus, this technique was used to assess CST axon regeneration post-complete SCI in the nOS and OS groups. The results revealed the presence of CST axons with BDA+ main trunks (a large bundle of red fluorescent nerve fibers) in the area rostral to the SCI site in the nOS and OS groups (Fig. 3D–E, D1, E1, d1, and e1). However, unlike the nonspecific red fluorescent spots in the nOS group (Fig. 3D, D2, and d2), numerous BDA+ CST axons with typical nerve fiber morphology were observed at the injury site of the spinal cord in the OS group (Fig. 3E, E2, and e2). More importantly, in the OS group, a few CST axons grew through the injury epicenter to the area caudal to the injury site (Fig. 3E, E3, and e3). Quantitative analysis revealed a significantly higher CST axon number index at the injury site in the OS group (9.75%±1.59%) than in the nOS group (0.49%±0.54%, p<0.05, n=6) (Fig. 3F). Moreover, the CST axon number index in the area caudal to the injury site in the OS group (5.56%±0.97%) was significantly higher than in the nOS group (0.34±0.52%, p<0.05, n=6) (Fig. 3D–F). Histological analyses revealed a higher number of BDA+ CST axons in the OS group than in the nOS group. The expression of growth-associated protein 43 (GAP43, an axon regenerationrelated protein) was observed in CST axon terminals in the area rostral to the SCI site (Fig. 3G and 3g). We further examined whether the spinal cord was completely transected to verify the reliability of the CST axon regeneration. Glial fibrillary acidic protein immunofluorescence staining displayed that the glial borders was formed at the areas rostral and caudal to the injury site in both the horizontal and sagittal spinal cord sections, suggesting that the transection injury was complete (Supplementary Fig. 4). The results suggested that the optogenetic stimulation of bilateral motor cortexes can promote CST axon regeneration in the ChR2-YFP transgenic mice with complete SCI.
Optogenetic stimulation enhanced corticospinal tract (CST) axon regeneration. (A) A schematic diagram illustrating that biotinylated-dextran amine (BDA) was injected into the motor cortexes to trace CST axon regenerating post-spinal cord injury (SCI). (B and C) Red fluorescence areas of bilateral BDA-labeled motor cortexes in the coronary section of the brain and of BDA+ CST axons in the horizontal section of the spinal cord of a ChR2-YFP transgenic mouse. (D and E) Showing 2 representative images (low magnification) of horizontal sections of the injured spinal cords of the mice in the nOS and OS groups, respectively. Moreover, 2 schematic diagrams of a horizontal section of the spinal cord are respectively showed in the lower right corners of panels D and E, highlighting the location and orientation of the sections, as well as displaying the direction of CST axon regeneration. (D1–D3) Higher magnification images for the boxed areas in (D), showing BDA+ CST axons (arrows) in the areas rostral and caudal to the injury site and the injury site. (d1–d3) Higher magnification images for the boxed areas in panels D1–D3, respectively. The dot-like profiles in panels d2 and d3 depict nonspecific staining of BDA. (E1–E3) Higher magnification images for the boxed areas in panel E showing BDA+ CST axons (arrows) in the areas rostral and caudal to the injury site and the injury site. (e1–e3) Higher magnification images for the boxed areas in panels E1–E3, respectively, depicting the specific staining of BDA+ CST axons (arrows). (F) The box plot showing the axon number index of CST in the nOS and OS groups (unpaired 2-tailed Student t-test was used for the analysis, *p<0.05, n=6). (G) Some BDA+/GAP43+ axons (white arrows) in a higher magnification image (g) from the box in (G). Scale bars=500 μm in panels B, D, and E; 50 μm in panels C, D1–D3, E1–E3, and G; and 20 μm in panels d1–d3, e1–e3, and g. LED, light emitting diode; YFP, yellow fluorescent protein; ChR2, channelrhodopsin-2; OS, optogenetic stimulation; nOS, nonoptogenetic stimulation.
4. Optogenetic Stimulation Improved the Movement Function of Paralyzed Hindlimb
The motor function of paralyzed hindlimbs of the mice post-SCI was assessed using electrophysiology assessment and BMS scoring. The assessment of cortex motor evoked potential induced by optogenetic stimulation showed a significantly lower latency (representing nerve impulse conduction speed of CST axons activated by optogenetic stimulation) and a significantly higher amplitude (representing the number of CST axons activated by optogenetic stimulation) in the OS group (13.25±2.20 msec and 0.20±0.03 mV, respectively) than in the nOS group (21.25±2.17 msec and 0.04±0.02 mV, respectively; p<0.05; n=6) (Fig. 4A and B). Furthermore, the grid climbing test was performed to reflect the degree of sensory and movement ability recovery of the paralytic hindlimbs [28]. The results demonstrated that mice in the nOS group displayed few spontaneous placing reflexes, and most of mice dragged the hindlimbs behind them when their forelimbs climbed on the grid. On the contrary, the mice in the OS group exhibited a few spontaneous placing reflexes, and their hindlimbs occasionally stepped on metal grid during the climbing (Fig. 4C) (Supplementary video clip 1). In the later stages of optogenetic stimulation (after 10–14 days of blue light stimulation), occasional hindlimb movements were observed during the stimulation in the OS group (Fig. 4D). Open-field locomotor test25 also showed that most of mice in the OS group exhibited extensive hindlimb joint movement. On the contrary, the mice in the nOS group only dragged their hindlimbs, with occasional slight ankle movement of the hindlimbs (Fig. 4E) (Supplementary video clip 2). BMS analysis showed that the hindlimbs of the mice were completely paralyzed following T10 spinal cord transection in both groups 1-day post-SCI (Fig. 4F). Though both the nOS and OS groups exhibited minor improvement in hindlimb function from 3 weeks after SCI, the mean score of the OS group was signifi-cantly higher than that of the nOS group (2.8±0.3 vs. 1.3±0.3, respectively; p<0.05; n=6) (Fig. 4F), indicating that the optogenetic stimulation enabled mice to move the ankle joints of the hindlimbs extensively, carrying heavy feet to walk. However, the mice in the nOS group exhibited only minor ankle movement of the hindlimbs. Correlation analysis showed that the BMS scores were highly correlated with the CST axon number index in the injury site and the area caudal to the injury site (Fig. 4G). These results suggested that optogenetic stimulation promoted the recovery of movement function of the paralyzed hindlimbs in mice with complete SCI.
Analysis of electrophysiology and behavior. (A) A representative image of the cortex motor evoked potential (CMEP) activated by blue light stimulation in the nOS and OS groups 6 weeks after complete spinal cord injury (SCI). (B) Bar charts showing the latency and amplitude of CMEP in the nOS and OS groups (unpaired 2-tailed Student t-test was used for the analysis, *p<0.05 compared with the nOS group, n=6). (C and D) Showing grid climbing test and Basso mouse scale (BMS) assessment performed on the nOS and OS groups. (E) Hindlimb movement (arrows) of complete SCI mice during optogenetic stimulation. (F) Comparison of the BMS scores of the hindlimb movement functions of the mice in the nOS and OS groups (2-way repeated-measures analysis of variance followed by Student-Newman-Keuls post hoc test was used for multiple comparisons, *p<0.05 compared with the nOS group, n=6). (G) Correlation analysis showed that the BMS scores were highly correlated with the CST axon number index in the injury site (ρ=0.93) and the area caudal to the injury site (ρ=0.88). OS, optogenetic stimulation; nOS, nonoptogenetic stimulation.
5. Optogenetic Stimulation Activated the JAK2/STAT3 Pathway in Motor Cortexes
To further explore key molecules involved in CST axon regeneration after optogenetic stimulation, the motor cortexes of ChR2-YFP transgenic mice were subjected to proteomic analy-sis 6 weeks after complete SCI (Fig. 5A). The volcanic map in Fig. 5B showed the differential protein expression in the nOS group versus the OS group. The heat map in Fig. 5C showed a higher number of upregulated proteins in the OS group than in the nOS group. Furthermore, KEGG pathway enrichment analysis of differential proteins indicated the activation of a group of signaling pathways in the OS group. In particular, the Janus kinase 2/signal transducer and activator of transcription 3 (JAK2/STAT3) pathway, being closely associated with axonal regeneration, was combed out (Fig. 5D). Several proteins involved in the JAK2/STAT3 signaling pathway were detected and quantified using a protein-protein interaction network diagram (Fig. 5E). Western blot analysis showed a significant upregulation of p-JAK2 and p-STAT3 in the OS group than in the nOS group (p<0.05, n=3) (Fig. 5F and G). Enrichment analysis of the proteomics data showed that the enriched proteins in the OS group were associated with the 6 functional sets, including central nervous system (CNS) neuron development, axonogenesis, synaptic plasticity, neuron projection development, neuron projection organization, and regulation of neurogenesis (Fig. 5H).
Optogenetic stimulation activated the JAK2/STAT3 pathway in the motor cortexes. (A) Schematic diagram of proteins extracted from mouse cerebral motor cortexes for proteomic analysis and western blotting. (B) Volcano plot showing the upregulated proteins (orange), downregulated proteins (green), and unaffected proteins (gray) in the nOS and OS groups. (C) Heat map showing the differential protein expression ratios between the 2 groups. (D) Kyoto Encyclopedia of Genes and Genomes pathway enrichment analysis of differential proteins, showing that optogenetic stimulation impacted the JAK2/STAT3 signaling pathway (green boxed bar). (E) JAK2/STAT3 signaling pathway associated protein-protein interaction network diagram. Each node represents a protein. The colored nodes represent the detected proteins, while the gray ones represent the undetected proteins according to the ratio value. The size of a node represents the degree of correlation among the proteins. (F) Western blotting results showing p-JAK2 and p-STAT3 expression levels in the nOS and OS groups. (G) Bar chart exhibiting the standardized gray values of p-JAK2 and p-STAT3 in the nOS and OS groups (The data were analyzed by unpaired 2-tailed Student t-test and presented as mean±standard deviation, *p<0.05, n=3). (H) Network diagram showing the association of enriched proteins with 6 different functional sets in the OS group compared to the nOS group. The node size of the functional set represents the total number of candidate proteins identified by gene ontology analysis. LED, light emitting diode; OS, optogenetic stimulation; nOS, nonoptogenetic stimulation; p-JAK2, phosphorylated Janus kinase 2; p-STAT3, phosphorylated signal transducer and activator of transcription 3.
6. In Vitro Blocking of the JAK2/STAT3 Pathway Inhibited the Outgrowth of Neurites Activated by Optogenetic Stimulation
To assess the role of the JAK2/STAT3 signaling pathway in CST axon regeneration, the pathway was subjected to in vitro pharmacological blocking and neurite outgrowth around the brain slices from ChR2-YFP transgenic mice was observed after optogenetic stimulation in the OS group and OS+FLLL31 (FLLL31, a specific inhibition of STAT3 phosphorylation [22]) group (Fig. 6A). After exposure to 10-Hz blue light stimulation, a higher number and longer length of neurites were observed around the brain slices of the OS group (left, Fig. 6B). However, a marked decrease in the number and length of the neurites located around the brain slices were observed after treated with the FLLL31 in the OS+FLLL31 group compared to the OS group (right, Fig. 6B). After statistical analysis, the number of neurites in the OS+FLLL31 group (133.40±35.30 points) was significantly less than that in the OS group (281.40±46.40 points, *p<0.05, n=5) (Fig. 6C). Moreover, the length of neurites in the OS+FLLL31 group (260.50±80.30 μm) was also shorter than that in the OS group (680.70±94.70 μm, *p<0.05, n=5) (Fig. 6C). These results suggested that the suppression of the JAK2/STAT3 signaling pathway inhibited neurite outgrowth from the brain slices after optogenetic stimulation. In short, a novel LED optogenetic stimulation device could enhance the functional activities of pyramidal neurons in the cerebral motor cortexes of ChR2-YFP transgenic mice. By activating the JAK2/STAT3 signaling pathway of the neurons to increase their intrinsic regenerative power, it promotes the extension of regenerated CST axons into the damaged area of spinal cord (Fig. 6D).
Blocking the JAK2/STAT3 signaling pathway inhibited the outgrowth of neuronal neurites. (A) Schematic diagram showing the lab-made organotypic brain slice culture system under the blue light stimulation in the OS and OS+FLLL31 groups. (B) Showing high levels of neurites (arrows in the upper and lower images on the left) around the brain slice in the OS group and fewer neurites (arrows in the upper and lower images on the right) around the brain slice in the OS+FLLL31 group. (C) Bar chart showing the number and length of neurites from the brain slices in the OS and OS+FLLL31 groups (Data was calculated using unpaired 2-tailed Student t-test, *p<0.05, n=5). (D) Schematic diagram of transcranial optogenetic stimulation promoting CST axon regeneration to repair complete spinal cord injury by activating the JAK2/STAT3 pathway. Scale bars=200 μm in the left and right images on the top of (B) and 50 μm in the left and right images on the bottom of (B). LED, light emitting diode; OS, optogenetic stimulation; FLLL31, tetramethylcurcumin.
DISCUSSION
Optogenetics is an emerging promising technique for neuroscience [29]. The optogenetic techniques used to analyze the CNS are primarily divided into 2 categories: Implantable and nonimplantable. An implantable optogenetic technique is mainly used for studying deep cerebral nuclei to accurately detect the range of biological electrical signals of the nuclei [30]. However, craniotomy can cause damage to the brain. Nonimplantable optogenetic techniques, such as transcranial optogenetic technique, do not require craniotomy implantation and are primarily used to study superficial brain tissue. In the present study, an enhanced transcranial optogenetic device was developed to stimulate cerebral motor cortexes (Zeng X, et al. An optogenetic stimulation device for promoting CST regeneration to repair SCI CN Patent, CN201910103715.2.2023). This technique comprises a custom optogenetic LED device specifically designed to stimulate bilateral motor cortexes in ChR2-YFP transgenic mice. The blue light emitted by the LED device can pass through the artificially thinned skull, directly illuminating the motor cortexes. The key parameters of the LED device are the blue light wavelength of 473 nm, CMOS wave style, voltage of 4.5 V, and luminous flux of 4,500 lux. The results of the present study revealed that the transcranial optogenetic stimulation device enhanced CST axon regeneration by activating the JAK2-STAT3 pathway. The LED device allows contact-free blue light stimulation to the cerebral motor cortexes, without physical interference to the brain parenchyma, for obtaining reliable experiment results. Despite the non-implantable LED device requiring a skull thinning procedure, it does not cause damage to brain tissue. Additionally, the device was able to activate the bilateral motor cortexes compared to the optical fibers [13]. With this device and the corresponding light stimulation procedure, exploration of the function of specific cortex regions is warranted in the future.
Initially, in the present study, the modified optogenetic stimulation technique was found to enhance the activity of motor cortex neurons in the transgenic mice. Optogenetic stimulation can directly activate neurons, leading to altered neural function. Previous studies have also shown that optogenetic stimulation can directly modify synaptic plasticity [31]. Optogenetic stimulation enhanced the electrical activity of neurons by promoting oligodendrogenesis and subsequent myelination, leading to neural function improvement in mice [32]. Notably, optogenetic stimulation to the motor cortex of the Thy1 transgenic mice with incomplete SCI led to enhanced corticorubral axon sprouting and synapse formation with neurons in the red nucleus, suggesting that optogenetic stimulation might directly affect neural plasticity [13]. Though the activation of neurons by electrical stimulation has been shown to enhance neurite elongation [33], it is still unclear whether optogenetic stimulation promotes axonal regeneration in traumatic CNS. Indeed, the sprout of lateral branches is a compensatory mechanism of the spared interneurons in incomplete SCI [34,35]. Only the undamaged neurons can sprout [36]. Unlike collateral sprouting, axonal regeneration is a functionally related post-injury adaptation observed in neurons with severed axons. Spontaneous axon regeneration in adult CNS is limited [37], unless active interventions are applied, such as PTEN/SOCS3 deletion [38] stem cell transplantation [39,40], or bioactive material implantation [41]. In the present study, optogenetic stimulation was found to enhance CST axon regeneration in a complete SCI mice model. The widespread activation of cortical motor neurons was validated via the detection of C-fos in fifth-layer pyramidal neurons. With chronic, daily optogenetic stimulation, the regeneration of the pyramidal neuron-derived CST axons was observed through anterograde tracing. Therefore, our findings provided proof-of-principle evidence for optogenetic stimulation-based axonal regeneration in mammals with CNS trauma.
Growing evidence suggests that increasing neuronal electrical activity might promote axon outgrowth. Previous studies have shown that short-term electrical stimulation increases the speed of axon regeneration in peripheral nervous system (PNS) injury by activating the cell body [42]. A recent study achieved enhanced motor axon regeneration of PNS via optogenetic stimulation of the sciatic nerve [43] similar to electrical activation of PNS neurons. Both patterned optogenetic and chemogenetic activation promotes CST axon outgrowth and branching [44]. In practice, electrical stimulation activates axon growth-associated genes, supporting the recovery of motor and sensory circuits in both PNS and CNS injuries [45]. For example, the increase in neuronal cyclic adenosine monophosphate levels after electrical stimulation enhanced collateral sprouting of CST axons above the lesion site and led to better recovery of forelimb function compared to the recovery in unstimulated rats with SCI [46]. The present study showed widespread cortical motor neuron activation after optogenetic stimulation, which might be responsible for CST regeneration post-SCI. Next, we found that the JAK2/STAT3 pathway was involved in optogenetic stimulation-based CST regeneration in complete SCI. The JAK2/STAT3 signaling pathway plays a multimodal role in CNS homeostasis and injury repair. Previous studies reported that JAK2/STAT3 activation by binding to gp130 receptors after ciliary neurotrophic factor stimulation was essential for the maturation and myelination of oligodendrocytes [47]. Another study suggested that JAK2/STAT3 activation by cytokine hormone erythropoietin promoted neurite growth and elongation of in vitro adult retinal ganglion cells following optic nerve lesion [48]. Indeed, the effect of JAK/STAT on axonal regeneration might be independent of mTOR, which is essential for axonal outgrowth after CNS injury [7,49]. A previous study showed that JAK/STAT blockade attenuated the increase in C-fos-expressing neurons and the presynaptic terminals of CST axons after chronic electrical stimulation of the motor cortex [50]. It suggested the involvement of JAK/STAT signaling in activity-dependent CST regeneration.
However, although optogenetic stimulation enhanced CST regeneration, widespread nerve fiber regrowth was not achieved at the injury site of SCI, in contrast to the significant pro-regenerative effect of optogenetic stimulation in vitro. This finding suggested that an alternative mechanism might be involved in post-injury milieus. Indeed, the secondary injury of SCI profoundly alters the microenvironment of primary injury site, including putting up physical and biological barriers to the regenerating axons [51,52]. despite which they might have adopted an intrinsic growth capacity. Nonetheless, JAK2/STAT3 pathway activation could be one of the mechanisms underlying nerve fiber regrowth after optogenetic stimulation. Based on this finding, an in-depth exploration of other potential mechanisms and the development of a potent microenvironment modulation regimen are warranted in the future.
CONCLUSION
An optogenetic stimulation device was used to treat the brain parenchyma non-invasively and was found to activate the motor cortexes in the present study. This device was found to increase the electrical activity of cortical neurons and the excitability of pyramidal neurons in the bilateral motor cortexes of ChR2-YFP transgenic mice. In addition, it enhanced the intrinsic growth ability of injured CST axons and promoted CST axon regeneration by activating the JAK2/STAT3 pathway, improving the movement function of the mouse hindlimbs after complete SCI.
Supplementary Materials
Supplementary Figs. 1–4, Supplementary Table 1, and Supplementary video clips 1–2 can be found via https://doi.org/10.14245/ns.2449312.656.
Primary antibody, secondary antibody and fluorescent dye
Showing the percentage of C-fos* expression in the Map2* cells in the motor cortexes after the blue light stimulation
C-fos and Map2 immunofluorescence staining of ChR2 brain slices without blue light stimulation. (A) C-fos and Map2 immunofluorescence staining of ChR2 brain slices with non-stimulation. (B1) Showing higher magnification for the boxed area in panel A. (B2) Showing Map2+ neurons in panel B1. (B3) Showing C-fos negative staining in panel B1. Scale bars=150 μm in panel A; and 80 μm in panel B1–B3.
The growth of single neurite deriverd from the brain slices under phase contrast microscopy in real time. (A) One neurite from the brain slice in the ChR2 group. (B) One neurite from the brain slice in the WT+OS group. (C) One neurite from the brain slice in the ChR2+OS group. The green arrows showed the growth cones of the neurites. Scale bar=25 μm in panels A–C. ChR2, channelrhodopsin 2; WT, wild-type; OS, optogenetic stimulation.
Glial fibrillary acidic protein (GFAP) immunofluorescence staining of transgenic mice spinal cord tissue reveals the glial borders in horizontal and sagittal sections. (A) Panoramic view of the horizontal section of the spinal cord. (a1 and a2) Higher magnification images for the boxed areas in panel A showed the GFAP+ borders in the areas rostral and the caudal to the injury site, respectively. (B) Panoramic view of the sagittal section of the spinal cord. (b1 and b2) Higher magnification images for the boxed areas in panel B showed the GFAP+ borders in the areas rostral and the caudal to the injury site, respectively. Scale bars=500 μm in panels A and B; and 40 μm in panels a1–b2.
Notes
Conflict of Interest
The authors declare no conflict of interest.
Funding/Support
This study was financially supported by the National Natural Science Foundation of China (grant number: 818981003; 81891002; 81974357; and 31900975), the Natural Science Foundation of Guangdong Province (2023A1515030254; 2024A1515220057), Guangzhou Basic and Applied Basic Research Foundation (2023A03J0626), Guangzhou Municipal Science and Technology Program (202206010197; 2025A04J4286), the Startup Fund Projects of Guangzhou First People’s Hospital (KYQD20210017), and the Startup Grant of Guangdong Hospital of Chinese Medicine (2022KT1032).
Author Contribution
Conceptualization: YL, XZ, YSZ; Data curation: YHM, HYC, YL; Formal analysis: YHM, QSW, LZP, KJZ, QWD, LJW, ZL, BQL, YD, GL, BJ, YL, XZ, YSZ; Funding acquisition: YHM, BQL, YD, GL, YL, XZ, YSZ; Methodology: YHM, HYC, QSW, LZP, KJZ, QWD, LJW, ZL, XZ; Project administration: YHM, LJW, BQL, YD, GL, YL, XZ, YSZ; Visualization: YHM, HYC, QSW, LZP, KJZ, QWD, ZL; Writing – original draft: YHM, HYC; Writing – review & editing: BQL, YL, XZ, YSZ.