Introduction
Traumatic spinal cord injury (SCI) results in disability with frequently devastating consequences that affect long-term health and employment status [
1]. SCI pathophysiology is initiated by mechanical tissue damage disrupting anatomical and functional integrity of the spinal cord, which contributes to secondary detrimental events. Neuroinflammation plays an important role in secondary neuronal damage following SCI [
2]. The inflammatory reaction begins at the lesion site and then expands to surrounding tissue during the secondary phase [
3]. A persistent inflammatory microenvironment and the presence of reactive oxygen species (ROS) are potential factors that impede injury repair following SCI [
4,
5].
Microglia are major contributors to the inflammatory response after central nervous system (CNS) injury [
6]. The voltage-gated proton channel (Hv1) maintains physiological intracellular pH and is abundantly expressed on the surface of microglia [
7]. Although Hv1 is specifically expressed in microglia in the nervous system, Hv1 can influence neurons by regulating microglial functions under pathological conductions. Previous studies found that Hv1 was implicated in pathological NADPH-oxidase-mediated ROS generation leading to neuronal apoptosis in ischemic stroke [
8,
9]. Our previous study has shown that ROS levels and pro-inflammatory cytokines are reduced in Hv1-deficient (Hv1
−/−) microglia compared to those in wild-type (WT) microglia after oxygen-glucose deprivation in vitro [
10]. However, the contribution of microglial Hv1 in neuronal damage and the underlying molecular mechanisms following SCI remain unknown.
Neuronal death via apoptosis, pyroptosis, or necroptosis is regulated through activities of various host proteins that induce different biological outcomes [
11,
12]. In contrast to apoptosis, neuronal pyroptosis is a form of cellular death induced by inflammatory caspases (e.g., caspase-1, caspase-4 and caspase-5, or caspase-11) implicated in ischemic stroke and SCI [
13,
14]. However, little is known regarding the spatial and temporal distributions of neuronal pyroptosis and apoptosis following SCI. Cleavage and activation of the pore-forming effector protein, gasdermin D (GSDMD), has been shown to determine pyroptosis initiated by inflammasomes [
15]. The nod-like receptor 3 (NLRP3) inflammasome, the well-studied inflammasome, is a multiprotein complex consisting of nod-like receptor 3 (NLRP3), apoptosis-associated speck-like protein (ASC), and an effector pro-caspase-1 [
16,
17]. It is unknown whether microglial Hv1 can promote neuronal pyroptosis through the NLRP3 inflammasome after SCI. Here, we used a mouse model to analyze spatial and temporal patterns of neuronal apoptosis and pyroptosis following SCI. We examined whether Hv1 deficiency exerts neuroprotective effects through modulating inflammasome activity and neuronal pyroptosis. Our findings may contribute to the identification and development of a potential therapeutic strategy for individuals with SCI.
Materials and methods
Animals
C57BL/6 female mice (wild-type, WT) were obtained from the Laboratory Animal Facilities of Hubei Center (Wuhan, China). Hv1
−/− mice were kindly provided by Prof. Long-Jun Wu (Department of Cell Biology and Neuroscience, Rutgers University) and were generated as previous study [
18]. The Hv1
−/− genotype was confirmed using PCR, Western blotting, and immunofluorescent analysis of tail DNA, spinal cord protein, and tissue sections, respectively. Mice were maintained under a 12-h light/12-h dark cycle at 22 °C and were provided food and water ad libitum. All female mice were 10–12 weeks of age and weighed 20–25 g in weight. WT and Hv1
−/− mice were randomly distributed into sham-operation and SCI groups. All experiments and procedures were approved by the Institutional Animal Care and Use Committee of Tongji Medical College, Huazhong University of Science and Technology.
Spinal cord injury model
Prior to surgery, mice were anesthetized with 2% isoflurane. A T9-11 laminectomy was performed, and the spinal cord was exposed at T10. A 5-g weight was dropped from a height of 11 mm onto the exposed dorsal surface of the spinal cord using a modified NYC impactor (J$K Seiko Electronic, China) [
19]. Sham-treated mice underwent a laminectomy at T9-11 without spinal cord injury. The bladder was voided twice daily until the recovery of urinary function post-spinal cord injury (SCI). Locomotor function was evaluated using the Basso Mouse Scale (BMS) [
20].
Tissue preparation
Animals were anesthetized and then perfused transcardially with 0.01 M of phosphate-buffered saline (PBS) at 1, 3, 7, 14, and 28 days after treatment (n = 5 for each time point). For Western blotting and ROS detection, spinal cord tissues—including the injury core and approximately 2 cm of surrounding tissue—were rapidly excised and frozen in cooled isopentane. For immunofluorescence, mice were perfused with 4% paraformaldehyde (PFA) in PBS and spinal cord tissues—including the injury core and approximately 2 cm of surrounding tissue—were dissected, fixed in PFA overnight at 4 °C, and then transferred to 30% sucrose in 0.01 M of PBS for 3 days. All the tissue samples were stored at − 80 °C until further use. Ten-micron sections were then prepared for Luxol fast blue (LFB) staining, immunofluorescence, and terminal dexynucleotidyltransferase-mediated dUTP nick-end labeling (TUNEL) assays.
Luxol fast blue staining
Luxol fast blue (LFB) staining was used to assess lesion volumes after SCI. Briefly, spinal cord tissue sections at 14 and 28 days after treatment were incubated overnight in 0.1% LFB (Servicebio, Wuhan, China) at 60 °C. Sample observations and image acquisitions were performed using a light microscope (BX51, Olympus, Japan).
Anterograde tracing
Anterograde-tracing was performed to detect axonal regeneration as previously described [
21]. Biotinylated dextran amine (BDA; MW,10000; Invitrogen) 10% w/v in sterile PBS was injected into two sites (0.4 μl/ site) of the spinal cord by micropipette. One week later, mice were anesthetized and perfused transcardially with 4% paraformaldehyde. Thirty-micron sections were incubated with Alexafluor 488-conjugated streptavidin (1:500; Invitrogen) for 1 h at room temperature. The images were acquired with a confocal microscope (Olympus, BX51).
Cell culture
PC12 cells (Cell Storage Center of Wuhan University) were cultured in Dulbecco’s modified Eagle’s medium and Ham’s F-12 (DMEM/F12, Hyclone, USA) supplemented with 10% horse serum (Hyclone, USA), 5% fetal bovine serum (FBS) (Hyclone, USA), and 1% penicillin/streptomycin (Hyclone, USA). Cultures were maintained at 37 °C, 95% air, and 5% CO2 in a cell culture incubator (Thermo Fisher Scientific, USA).
Oxygen-glucose deprivation/reoxygenation and drug treatment
Oxygen-glucose deprivation/reoxygenation (OGD/R) was used to model ischemic injury in vitro, as described previously [
22]. Briefly, PC12 cell cultures were washed with ice-cold PBS and culture medium was replaced with glucose-free DMEM (Gibco, USA). Cells were incubated in a hypoxic incubator (Thermo Scientific; USA) at 94% N
2, 5% CO
2, and 1% O
2 for 2 h at 37 °C. Cells were then returned to standard culture medium and incubation conditions (37 °C, 95% air, and 5% CO
2). The ROS inhibitor,
N-acetyl cysteine (NAC, Sigma-Aldrich, USA), was dissolved in sterile H
2O prior to cellular treatments [
23].
Lactate dehydrogenase assay
PC12 cells were cultured in 96-well plates with or without NAC (at concentrations of 0, 10, 20, 30, 50, or 100 μM) for 3, 6, or 24 h. Culture supernatant was collected and analyzed using a commercial lactate dehydrogenase (LDH) kit (Beyotime, China) according to the manufacturer’s instructions.
Detection of reactive oxygen species
To determine reactive oxygen species (ROS) levels in the spinal cord, tissue samples were homogenized in 10 ml/g of ice-cold PBS as described previously [
24]. Homogenates were centrifuged at 10,000×
g for 10 min at 4 °C. Supernatants were collected and diluted 200-fold in PBS. Then, 100 μl of diluted supernatant was mixed with 100 μl of the molecular probe, 20,70-dichlorodihydrofluorescein diacetate (DCFH-DA, 10 μM; Sigma-Aldrich, USA), and was then transferred to a microplate culture well. To determine PC12 intracellular ROS levels, DCFH-DA (10 μM) was added to cells in serum-free medium. Cells and tissue sample supernatants were incubated with DCFH-DA at 37 °C for 1 h in the dark. ROS levels were measured at 492/520 nm using a multiplate reader (Synergy HT; Biotech, USA).
Caspase-1 activity
Caspase-1 activity was measured using the Caspase-1 Activity Assay Kit (RD, K111-100) according to the manufacturer’s instructions. Briefly, 2–5 × 106 cells were lysed in 50–100 μl of ice-cold lysis buffer for 10 min and were then centrifuged. Each specimen was incubated in a 96-well plate with Ac-YVAD-pNA (200 μM) at 37 °C for 2 h. The optical density of each specimen was read on a microplate reader (Synergy HT; Biotech, USA).
Scanning electron microscopy
PC12 cells were fixed in 2.5% glutaraldehyde for 3 h and were then rinsed with PBS. Cells were dehydrated through a graded ethanol series, dried using tertiary butanol, sputter coated with gold, and imaged via scanning electron microscopy.
Flow cytometry
The PC12 cell apoptotic rate was determined using flow cytometry with an annexin V-FITC/PI apoptosis detection kit (BD Biosciences, 556547) according to the manufacturer’s instructions. Briefly, single-cell suspensions were stained with annexin-V and propidium iodide (PI) at room temperature for 15 min in the dark and were subjected to flow cytometry (BD Biosciences, San Jose, CA). Data were analyzed using FlowJo (TreeStar, Ashland, OR).
Immunofluorescence
Immunofluorescent staining was performed as previously described [
25]. Briefly, tissue sections or cells were blocked using a 10% bovine serum albumin solution with 0.25% triton X-100 for 1 h at room temperature, followed by incubation overnight at 4 °C with primary antibodies. These primary antibodies included rabbit anti-NeuN (1:500, Millipore), rabbit anti-GFAP (1:500, Cell Signaling Technology), rabbit anti-IBA-1 (1:500, Wako Pure Chemical Industries), goat anti-IBA-1 (1:500, Wako Pure Chemical Industries), mouse anti-GSDMD (1:50, Santa-Cruz), mouse anti-ASC (1:200, Santa-Cruz), mouse anti-caspase-1 (1:200, Santa-Cruz), mouse anti-NLRP3 (1:200, AdipoGen), rabbit anti-Hv1 (1:50, Sigma), and mouse anti-8-OHdG (1:200, Abcam). Next, samples were washed in PBS and incubated at room temperature for 1 h with the appropriate secondary antibodies, as follows: FITC-conjugated goat anti-mouse immunoglobulin G (IgG), Cy3-conjugated goat anti-rabbit IgG, Cy3-conjugated rabbit anti goat IgG, Cy3-conjugated rabbit anti-rat IgG, and 488-conjugated donkey anti-goat IgG (Jackson ImmunoResearch, West Grove, PA, USA). Finally, sections were stained with 4,6-diamidino-2-phenylindole (DAPI). Samples were imaged using a confocal microscope (Olympus, BX51).
TUNEL assay
Apoptosis was detected using the In-Situ Cell Death Detection Kit (TUNEL fluorescence FITC kit, Roche) according to the manufacturer’s instructions. TUNEL staining was performed with fluorescein-dUTP for detection of apoptotic cell nuclei and DAPI for staining of cellular nuclei. The number of TUNEL-positive cells was analyzed via confocal microscopy (Olympus, BX51).
Western blotting
Tissue samples or PC12 cells were lysed in total protein lysis buffer and protein concentrations were determined using a BCA protein kit (Beyotime, China). Proteins (30 μg) were separated by sodium dodecyl sulfate-polyacrylamide gels (SDS-PAGE) and were transferred to nitrocellulose filters (NCs) or polyvinylidene-difluoride (PVDF) membranes. Membranes were blocked for 1 h at room temperature using 5% non-fat milk in Tris-buffered saline containing 0.1% Tween-20, then incubated overnight at 4 °C with primary antibodies. The following primary antibodies were used: rabbit anti-GSDMD (1:1000, Cell Signaling Technology), mouse anti-Caspase-1 (1:500, AdipoGen), mouse anti-NLRP3 (1:1000, AdipoGen), rabbit anti-ASC (1:1000, Cell Signaling Technology), rabbit anti-IL-18 (1:1000, Abclonal), rabbit anti-Hv1 (1:1000, Sigma), rabbit anti-NOS2 (1:1000; Abclonal), mouse anti-NF-L and anti-NF-H (1:1000; Cell Signaling Technology), rabbit anti-TUJ1 (1:1000, Abcam), mouse anti-MAG (1:1000, Santa Cruz), rat anti-MBP (1:1000, Millipore), rabbit anti-β-actin (1:1000, Servicebio), and rabbit anti-GAPDH (1:1000, Servicebio). Membranes were subsequently incubated with horseradish peroxidase-labeled anti-rabbit, anti-mouse, or anti-rat secondary antibodies (1:5000, Servicebio). Bands were visualized using a Bio-Rad Chemidoc XRS+ imaging system with enhanced chemiluminescent kits (Advansta). Protein levels were determined based on O.D. value using ImageJ software. Protein expression levels were normalized to β-actin or GAPDH internal controls.
Real-time and semi-quantitative PCR
Spinal cord tissue samples were collected at 1, 3, or 7 days (n = 5 for each time point) after surgery and total RNA was isolated using Trizol reagent (Invitrogen). Next, cDNA was prepared using a ReverTra Ace qPCR RT Kit (TOYOBO, Japan). Quantitative real-time PCR (qRT-PCR) was performed using a Real-Time PCR system (BioRad) and SYBR Green PCR Master Mix (TOYOBO, Japan). Then, mRNA expression levels were normalized to those of an endogenous reference gene, glyceraldehyde 3-phosphate dehydrogenase (GAPDH). Semi-quantitative PCR was performed using Quick Taq HS Dye Mix (Japan). PCR products were electrophoresed on a 2% agarose gel and were visualized with GelRed (Biotium; Hayward, CA) and a Gene Genius Bio-Imaging System (Syngene; Cambridge, UK). The following primers were used: interleukin-18 (IL-18) (forward: 5′GGCCGACTTCACTGTACAACCG3′, reverse: 5′GGTCACAGCCAGTCCTCTTACTTC3′), GAPDH (forward: 5′GGTTGTCTCCTGCGACTTCA3′, reverse: TGGTCCAGGGTTTCTTACTCC), Hv1 (forward: 5′-GAGATCCATCTGCCTCCGTTATGAGTG-3′, reverse: 5′-CTGTGTCTCCCTGTGGCTGAG-3′), and β-Geo-R (5′-GACAGTATCGGCCTCAGGAAGATCG-3′).
Statistical analysis
All data are presented as the mean ± standard error of the mean (SEM). Data were evaluated by one-way or two-way analyses of variance (ANOVAs) with Tukey’s post-hoc tests, when necessary. A value of p < 0.05 was considered statistically significant. Statistical analyses were performed using GraphPad Prism 6 (GraphPad Software, Inc., La Jolla, CA).
Discussion
The present study found distinct spatial and temporal patterns of neuronal pyroptosis and apoptosis after SCI and showed that Hv1 deletion significantly attenuated NLRP3-inflammasome-mediated neuronal pyroptosis and apoptosis as well as facilitated myelin-axon regeneration paralleled by improved motor function. These findings suggest that the effects on ROS-mediated modulation of NLRP3 inflammasome activation underlie protection against neuronal damage from Hv1 deficiency. To our knowledge, the present study is the first to elucidate spatial and temporal differences in neuronal pyroptosis and apoptosis during early-stage SCI. Moreover, this work is the first to reveal the underlying mechanism by which Hv1 deficiency attenuates neuronal pyroptosis after SCI.
Apoptosis has long been recognized as the predominant in a wide variety of neurological conditions [
31]. Recently, findings found that another special death mechanism different from apoptosis participated in the CNS pathology, which has been named as pyroptosis in 2001 [
32,
33]. Pyroptosis is characterized by cellular swelling [
30], formation of 10–14-nm GSDMD pores [
34], caspase-1-dependent cell death, and release of inflammatory cytokines such as IL-18 [
33]. More recently, neuronal pyroptosis—a gasdermin-mediated programmed necrotic cell death—has been reported to be linked to traumatic brain injury [
33], cerebral ischemia/reperfusion injury [
25], and SCI [
14]. In the present study, we showed that neuronal expression levels of the pyroptosis marker GSDMD and the inflammatory factor IL-18 were significantly upregulated in vivo SCI model in mice and in vitro OGD/R model in neurons mimicking SCI, which are consistent with the previous opinion that neuronal pyroptosis contributed to neuroinflammation after SCI [
33]. Accumulating studies showed that apoptosis and pyroptosis could both occur in neuron during the pathology of CNS diseases [
25,
33]. However, few researches further focused on the spatiotemporal patterns of neuronal pyroptosis and apoptosis. Here, for the first time, we reported that the peak of neuronal apoptosis prior to that of neuronal pyroptosis and the duration of pyroptosis was much longer than that of apoptosis in vitro and in vivo experiments. These results suggest that in contrast to apoptosis, timely intervention to inhibit neuronal pyroptosis following SCI is feasible. Thus, identifying a means of regulating neuronal pyroptosis after SCI is potentially valuable.
Hv1 channel has been reported to regulate neuronal death in the pathogenesis of ischemic stroke [
8]. Our present study also showed that neuronal apoptosis following SCI was reduced in Hv1
−/− mice relative to that in WT mice. However, information on microglial Hv1-mediated neuronal pyroptosis remains unclear. Interestingly, our study is the first to discover that the expression of pyroptosis in neurons was lower in Hv1
−/− mice compared with that in WT mice after SCI. Microglia and macrophages, the first cells to be recruited in response to injury, are the most important modulators of secondary damage after SCI [
35] and activated microglia contribute to ROS generation [
36]. Hv1 channel, functionally expressed in microglia, has been reported to induce NADPH oxidase (NOX)-mediated ROS generation [
8]. In the current study, we observed upregulation of microglial ROS generation after SCI, which was significantly attenuated in Hv1
−/− mice in the acute period after SCI. This finding is consistent with our previous report that microglial Hv1 deficiency could attenuate ROS level [
10]. Emerging evidence showed that ROS signaling was the key in the initiation of pyroptotic death involving diverse diseases [
37,
38]. It was proposed that pyroptosis in hemorrhagic shock and resuscitation was downregulated through reducing ROS production by mitochondria [
37]. In another study, ischemia-reprerfusion-induced pyroptosis remarkably relied on ROS which was significantly inhibited by ROS scavenger NAC [
38]. In line with these previous researches, our in vitro data further showed that ROS-induced pyroptosis by OGD/R model in neurons was obviously rescued by NAC. Therefore, we reasonably believe that microglial Hv1 channel exacerbated neuronal pyroptosis through enhancing ROS production after SCI.
Unlike apoptosis, pyroptosis is an inflammatory cell death requiring the activation of caspase-1 [
39]. Caspase-1 activation is modulated by protein complexes termed inflammasomes. Inflammasomes can be divided into different types, such as NLRP1, NLRP2, NLRP3, NLRC4, NLRP6, NLRP7, and AIM2 [
14]. Among these inflammasomes, NLRP3 inflammasome expression is markedly upregulated following SCI, thus playing an important role in spinal cord tissue after SCI [
40]. Furthermore, Zendedel et al. [
41] found that NLRP3 was expressed in neurons, microglia, and astrocytes, and that neurons represented the major source of NLRP3. Here, we showed that the expressions of NLRP3, ASC, and caspase-1 p20 in neurons were significantly increased after SCI in mice, the findings of which are consistent with those of a previous study [
14]. In addition, NLRP3, ASC, and caspase-1 levels were lower in Hv1
−/− mice, suggesting that microglial Hv1 is a potential target for modulating the NLRP3 inflammasome. NLRP3 is triggered in response to diverse stimuli including microparticles, pore-forming toxins, ATP, and ROS [
42]. ROS acts as a second messenger that plays a significant role in NLRP3 inflammasome activation [
43]. The activation of NLRP3 recruits ASC and promotes the activation of caspase-1, thus processing cytokines to their active forms to ultimately induce pyroptotic death [
27]. We detected OGD/R activation of the NLRP3 inflammasome pathway in neurons in vitro, which again was inhibited by NAC, thus revealing a key role of ROS in this process. These results suggest that microglial Hv1-induced ROS generation regulates NLRP3-inflammasome-mediated pyroptosis after SCI.
Neuronal death in the injured area following SCI may contribute to disruption of neural circuitry, ultimately leading to functional impairment [
44]. Hence, inhibition of neuronal delayed death is critical to promoting axonal regeneration after SCI [
45]. Additionally, reducing neuroinflammation may create a favorable micro-environment for neuronal and myelin repair [
46]. Collectively, our findings suggest that inhibition of Hv1 can promote myelin/axonal regeneration and concomitantly improve motor function after SCI. Based on our current results, we propose that microglial Hv1 deficiency may serve as a promising strategy to improve SCI outcomes.
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