Increased frequency of AMP‐activated protein kinase‐positive spinal motor neurons after sciatic nerve injury in a mouse model

Abstract

The role of AMP‐activated protein kinase (AMPK) in the regulation of energy metabolism and the control of skeletal muscle regeneration post injury has been described previously. It remains unknown whether this metabolic sensor plays a role in the mechanism of axonal regeneration post injury. In this study, we used a sciatic nerve crushed mouse model to detect the expression of AMPK in sciatic nerve and spinal motor neurons at 1 week, 2 weeks and 3 weeks after injury by immunofluorescence staining. Electrophysiological and histopathological studies were used to confirm the nerve injury and regeneration. Our results showed that frequency of AMPK‐positive spinal motor neurons was significantly higher on day 7 after sciatic nerve crush (SNC) and peaked on day 14. No expression of AMPK was detected in axons of the sciatic nerve before and after the injury. Taken together, our study suggested a possible role of AMPK in the mechanism of motor nerve regeneration after injury.

Introduction

Traumatic peripheral nerve injury is common and may result in limb weakness and loss of sensory function. Investigating the mechanisms involved in nerve repair and regeneration post injury is necessary to improve nerve regeneration and therefore function. Generally, after peripheral nerve injury, Wallerian degeneration firstly occurs, followed by migration of activated macrophages to clear the myelin and axon debris and axonal sprouting from spinal neurons [[1], [2]].

AMPK is a heterotrimeric serine/threonine kinase that exists in all eukaryotic cells and consists of a catalytic α subunit and two regulatory β and γ subunits [[3], [4]]. Accumulating evidence indicates that AMPK plays an essential role in a series of neurophysiological and pathological processes [[5], [6]]. Price et al. reported that AMPK activation could induce neuronal plasticity via mammalian target of rapamycin (mTOR) complex 1 [6]. AMPK signaling can inhibit tau phosphorylation and amyloid‐beta production thereby delaying the progress of Alzheimer's disease in transgenic mice [7]. Furthermore, Gong et al. have demonstrated that some microRNA's function as a protector of dopaminergic neurons during Parkinson's disease by regulating the AMPK pathway [8]. However, it remains unknown whether AMPK participates in the mechanism of post injury nerve regeneration. In this study, using a sciatic nerve crushed (SNC) mouse model, we explored the dynamic expression of AMPK in spinal motor neurons during the nerve regeneration process.

Materials and methods

Animals

Male C57BL/6 mice (8–10 weeks) were purchased from Pengyue Laboratory Animal Breeding Limited Company (Jinan, China) and housed in a temperature‐controlled facility with 12 h light/dark cycles and free access to food and water. The experiment was approved by the Institutional Animal Care and Use Committee of the Affiliated Hospital of Jining Medical University and performed in accordance with the Principles of Laboratory Animal Care.

Reagents

Phosphate‐buffered saline (PBS, 0.1 M, PH 7.4) and Tween‐20 were purchased from Boster (Wuhan, China). TritonX‐100 was purchased from Genview (TX, USA). Goat serum was purchased from Zhongshan Golden Bridge Company (Beijing, China). Anti‐neurofilament H nonphosphorylated antibody (SMI32) was purchased from Biolegend (CA, USA). Anti‐AMPK alpha 1 + AMPK alpha 2 antibody, Alexa‐568‐conjugated goat anti‐mouse IgG, Alexa‐488‐conjugated donkey anti‐rabbit IgG and 4′,6‐diamidino‐2‐phenylindole (DAPI) were purchased from Abcam (MA, USA).

Sciatic nerve crush

Twelve mice were randomly assigned to four groups (n = 3 per group): (1) normal control group, (2) SNC 7d group, (3) SNC 14d group and (4) SNC 21d group. The SNC group mice were anesthetized by intraperitoneal injection of 10% chloral hydrate (0.05 ml/100 g). The left hindquarter was then carefully shaved prior to skin cleansing with Betadine swabs. The skin was incised 1 mm posterior and parallel to the femur, and the biceps femoris was bluntly split to expose the sciatic nerve. The sciatic nerve was then crushed 5 mm proximal to its trifurcation with No. 5 Jeweler's forceps for 30 s as previously described [9]. The nerve crush was confirmed by the presence of a translucent band across the nerve and electrophysiological test. The incision was then closed in layers (muscle and skin) with absorbable sutures to allow recovery. The contralateral sciatic nerve was left intact as a surgical control.

Electrophysiology study

The electrophysiological study was performed to confirm the crush model and monitor the process of nerve repair. An electrophysiological surveillance system was setup and nerve function was recorded prior to the operation and at 1 week, 2 weeks and 3 weeks postoperatively (before the tissue processing). The left sciatic nerve in each mouse was exposed at the hip (proximal), and then one pair of needle electrodes was inserted into the sciatic notch (hip/proximal). The nerve stimulation parameters used to elicit compound muscle action potential (CMAP) were: 1 Hz pulses, with each pulse being 5 mA in amplitude and 0.3 ms in duration. The recording electrodes were positioned in the “belly” part of the gastrocnemius muscle to record evoked potentials from stimulating the sciatic nerve. The distance between stimulating electrodes and recording electrodes was 1.5 cm. The nerve conduction velocity of the surviving axons was determined by dividing the distance between the stimulation and recording points and the latency between stimulation and the appearance of the action potentials (V = m/s). A heating pad was used to maintain the body temperature of the mouse at 37 °C.

Tissue processing

Mice at different time points were deeply anesthetized using 10% chloral hydrate (0.05 ml/100 g) and were perfused through the left heart ventricle with 40 ml of ice‐cold PBS, followed by 40 ml of 4% paraformaldehyde in PBS. Then, sciatic nerve‐originated L3 L4 spinal cords were identified as previously described [10] and the left sciatic nerves were removed carefully and post‐fixed in 2% paraformaldehyde overnight at 4 °C. Half of the sciatic nerve from each mouse was processed for the electron microscopy analysis. Tissues were dehydrated with 30% sucrose solution in PBS. Cryostat sections of 10 μm thick were cut transversely for the immunofluorescence staining.

Electron microscopy

Sciatic nerves from different time points were fixed in a mixture of 2% paraformaldehyde and 3% glutaraldehyde in PBS overnight at 4 °C and the tissues were post‐fixed using 1% osmium tetroxide in PBS for 1 h, dehydrated in a gradient alcohol series and immersed in propylene oxide and a propylene oxide and embedding material mixture respectively. Transverse sections 70 nm thick (ultrathin) were obtained using an ultramicrotome (LKB‐V, Leica, Germany). Then, 70 nm thick sections were stained with 1% uranyl acetate and 3% lead citrate for 5 min and observed through a transmission electron microscope (JEOL‐1200EX, JEOL, Tokyo, Japan) attached with a side‐mounted transmission electron microscope charge coupled device camera (Morada G2, Olympus, Tokyo, Japan).

Immunofluorescence staining

Spinal cord and sciatic nerve sections were incubated for 2 h in PBS containing 0.3% Triton X‐100 and 10% goat serum at room temperature to block the unspecific binding. Subsequently, the sections were incubated overnight with anti‐SMI32 antibodies (1:2500 diluted) and anti‐AMPK alpha 1 + AMPK alpha 2 antibody (1:400 diluted) at 4 °C. The sections were then washed 3 times, two times with PBS‐T (PBS with 0.05% Tween‐20) and one time with PBS. Thereafter, donkey anti‐rabbit secondary antibodies conjugated with Alexa Fluor‐488 (1:1000 diluted) and goat anti‐mouse secondary antibodies conjugated with Alexa Fluor‐568 (1:1000 diluted) were added for an incubation of 2 h at room temperature. After another washing‐episode (two times with PBS‐T and one time with PBS), the sections were stained with DAPI (1:25 diluted in PBS) for 10 min to visualize the nuclear morphology. The sections were rinsed with PBS three times and mounted using anti‐fading medium. The stained spinal cord and sciatic nerve sections were observed using Observer Z1 confocal laser‐scanning microscope (Zeiss, Jena, Germany). Representative staining of SMI32 (red), AMPK (green), and DAPI (blue) were shown.

Image analysis

The frequency of AMPK‐positive motor neurons in the anterior spinal cord from the injury side was analyzed by counting ten sections (each separated from the next by an interval of about 100 μm) from each of L3‐L4 spinal cord segments. Lamina Ⅸ of the spinal cord containing motor neurons as selected as previously described [11]. Two rectangles were used to frame the lamina Ⅸ of spinal cord section. The frequency of AMPK‐positive motor neurons was calculated by the numbers of AMPK ‐SMI32‐DAPI positive cells divided by the numbers of SMI32‐DAPI positive cells. The results are shown as mean ± standard error. The analysis was performed using one‐way analysis of variance (ANOVA) using the SPSS 20.0 software by IBM (Armonk, NY). A p‐value<0.05 was considered significant.

Results

As shown in Fig. 1 and Fig. 2, the electrophysiological study and electron microscopic analysis confirmed the dynamic changes of sciatic nerve after injury. There was typical nerve regeneration at 3 weeks after injury.

Figure 1.

Electrophysiology studies of sciatic nerve after injury. The CMAP get to the nadir after SNC 7d injury, and improved after SNC 14d injury, and finally get back to normal after SNC 21d injury. The Latency was obviously prolonged after SNC 7d injury than normal, then improved after SNC 14d injury, and finally got back to normal after SNC 21d injury.

Figure 2.

Electron microscopic analysis of sciatic nerve after injury. Normal group (X 6000): The axon and myelin sheath are seen with normal structure. SNC 7d group (X 6000): Wallerian degeneration was occurred in the damaged nerve. SNC 14d group (X 6000): Wallerian degeneration and axon regeneration were co‐existed. SNC 21d group (X 6000): Myelinated fibers were well preserved, showing a nearly normal aspect.

The frequency of AMPK‐positive spinal motor neurons in the SNC 7d group and SNC 14d group was higher than in the normal group (p < 0.001, 35.74 ± 2.90% vs 22.39 ± 3.03%; p < 0.001, 51.52 ± 2.50% vs22.39 ± 3.03%). The frequency of AMPK‐positive spinal motor neurons in the SNC 7d group and SNC 14d group was also significantly higher than in the SNC 21d group (p < 0.001, 35.74 ± 2.90%vs 24.71 ± 2.00%; 51.52 ± 2.50% ± 24.71 ± 2.00%). No significant difference of the frequency was found between the SNC 21d group and normal group (p > 0.05). Details are shown in Fig. 3. No expression of AMPK was detected in axons of normal and injured sciatic nerve (data not shown).

Figure 3.

Dynamic expression of AMPK in mouse spinal motor neurons after SNC. a. Representative confocal images of SMI32 (red), AMPK (green), and DAPI (blue) expression and merged images of three markers (down) at Normal, SNC 7d, SNC 14d and SNC 21d. Scale bar = 50 μm. b. As it was shown, the frequency of AMPK in SNC 7d group and SNC 14d group had a significant difference compared to the Normal group (***p < 0.001 versus the normal group). The frequency of AMPK in SNC 7d group and SNC 14d group had a significant difference compared to the SNC 21d group (^###p < 0.001 versus the SNC 21d group). V: Ventral; D: Dorsal.

Discussion

In this study a well‐established SNC model was used to investigate the dynamic expression of AMPK in spinal motor neurons. Our study firstly demonstrated that the frequency of AMPK positive motor neurons in spinal cords of mice with SNC significantly increased.

After peripheral nerve injury, soluble factors released by the injured nerve will recruit macrophages to clear myelin and axon debris, yielding nerve repair. After a lag period, the damaged axons form a growth cone and begin to regenerate along bands of Büngner formed by Schwann cells. If the axon can pass through the injury site, and its environment supports its growth along the entire distal stump, then the axon can connect with peripheral targets [1]. Previous studies have demonstrated that AMPK plays an important role in neuronal development by modulating neurite outgrowth. Dasgupta et al. recently demonstrated that activated AMPK in neurons promotes neurite outgrowth and mitochondrial biogenesis in an AMPK‐dependent manner [12]. Arsenic can inhibit neurite outgrowth targeting the Liver kinase B1‐AMPK signaling pathway [13]. We hypothesized that AMPK may also participate in the mechanism of nerve regeneration post injury. In our study, electrophysiological studies and electron microscopy analysis confirmed a dynamic process from nerve injury to repair. Consistently, the frequency of AMPK‐positive spinal motor neurons began to increase on day 7 after nerve injury and peaked on day 14, followed by a reduction on day 21 when significant nerve regeneration had occurred. Our data support that AMPK is involved in the mechanism of nerve regeneration after injury. AMPK is a key metabolic sensor of energy balance at the cell and organism level [14]. It has been demonstrated that AMPK in primary neurons originated from the brain could modulate the neuronal energy homeostasis and contribute to the neuroprotective effects of resveratrol in vitro [12]. Under metabolic stress, activation of AMPK in rat hippocampal neurons exerts a protective role by reducing neuronal excitability and thus conserving energy [15]. In hypothalamic neurons, AMPK responds to hormones (including leptin and adiponectin), nutrient cues, and cellular energy levels, participating in the hypothalamic control of feeding behavior and influencing whole‐body energy metabolism [16]. Given that nerve regeneration is a high‐energy consuming process, our study demonstrated that AMPK may participate in the mechanism of nerve regeneration after injury by modulating the energy metabolism of spinal neurons. Notably, the sciatic nerve contains a mixture of sensory and motor nerves. We hereby only investigated the AMPK expression in spinal motor neurons during nerve regeneration aiming to reveal whether there is a relation between AMPK and nerve regeneration. To fully understand the roles of AMPK in mechanism of nerve regeneration, further research is necessary to detect the expression of AMPK in sensory neurons of dorsal root ganglions as well as the Schwann cells in injured peripheral nerves.

In summary, our study firstly demonstrated that AMPK might be involved in the process of peripheral nerve regeneration after injury. Further research to investigate the mechanism of AMPK‐involvement in this process is necessary to explore its potentials in therapeutic strategies of peripheral nerve injury.

Acknowledgments

We thank Alex Doets from the Erasmus Medical Center, Rotterdam, the Netherlands for the edits. This work was supported by the National Natural Science Foundation of China [grant number 81572205].

Supporting information

Supplementary data

Supplementary data

Supplementary data related to this article can be found at https://doi.org/10.1016/j.kjms.2017.12.007.