Int J Clin Exp Med 2017;10(10):14386-14393www.ijcem.com /ISSN:1940-5901/IJCEM0041440

Original ArticleMitochondrial biogenesis in ventral spinal cord and nerve following sciatic nerve axotomy

Wei Huang1*, Tianbing Wang1*, Baoguo Jiang1, Ming Zheng2

1Department of Trauma and Orthopaedics, Peking University People’s Hospital, Beijing, China; 2Department of Physiology and Pathophysiology, Health Science Center, Peking University, Beijing, China. *Equal contributors.

Received March 27, 2016; Accepted January 26, 2017; Epub October 15, 2017; Published October 30, 2017

Abstract: In the nerve system, mitochondria play pivotal roles in regulating various cellular processes and provide over 90% ATP supply to nerve activities. After nerve injury, cell bioenergetic status and energy metabolism related genes were significantly changed, but it is not clear whether mitochondrial biogenesis is involved in post-injury nerve degradation and regeneration processes. Here we show that mitochondrial biogenesis was down-regulated in spinal cord and up-regulated in proximal nerve after sciatic nerve injury, as indicated by the decreased or increased expressions of biogenesis biomarkers PGC-1α, NRF-1, and TFAM. Mitochondrial to nuclear DNA ratio decreased in spinal cord and increased in proximal nerve. Moreover, mitochondrial morphology and ATP production were changed accordingly after nerve injury, with less, irregular mitochondria and reduced ATP content in spinal cord, and more, condensed mitochondria and elevated ATP content in proximal nerve. Our study revealed important but distinct changes of mitochondrial biogenesis in ventral spinal cord and nerve after peripheral nerve injury, and may provide treatment strategies for peripheral nerve injury through adjusting mitochondrial biogenesis and metabolic status.

Keywords: Mitochondrial biogenesis, nerve, motoneuron, energy metabolism, nerve regeneration

Introduction

Peripheral nerves degenerate after injury by Wallerian degeneration, a process defined as the degeneration of the axon distal to the injury site [1, 2]. The neurons then re-initiate proximal axonal growth, known as regeneration, to repair the injured connection of the nerves with their targets. Enormous effort was made to promote nerve regeneration by improving the surgical technique, increasing neuronal survival and the rate of axon regeneration or retarding Schwann cell and basal lamina atrophy. However, emerg-ing evidence has shown that peripheral nerve injury also triggers a number of cellular and molecular events in neurons which might be essential for nerve regeneration [3, 4]. The study by Hu et al [4] revealed that some genes expression related to cell apoptosis increased and genes associated with neuronal survival or function decrease. Indeed, axonal regeneration is a coherent process of neurons, both proximal and distal axon, and even the Schwann cells, and the functional and metabolic status of these parts determine the degree of neurologi-cal recovery [5].

Mitochondria are important organelles, playing central roles in regulating various cellular pro-cesses including production of ATP, modulation of calcium signaling, generation of reactive oxy-gen species, and even determination of cell fates. In neuron, mitochondria provide more than 90% of total amount of ATP through oxida-tive phosphorylation [6], to supply the estab-lishment of membrane excitability, the synthe-sis of neurotransmitter and excitability condu- ction, and the regeneration of nerves [7]. Under different physiological and pathological condi-tions, energy demands are different, the mor-phology, function and amount of mitochondria in neurons are regulated accordingly. After root avulsion, most of the gene expression associ-ated with energy metabolism decreased in motor neurons [4]. However, it is not clear whether genes related to mitochondrial biogen-esis of the ventral neuron and proximal nerve stump area changed after peripheral nerve injury.

In this study, we analyzed the molecular, mor-phological, and bioenergetic changes of mito-chondria in proximal sciatic nerve and spinal

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cord area following sciatic nerve injury. We found that mitochondrial biogenesis biomark-ers PGC-1α, TFAM, NRF-1 and mitochondrial to nuclear DNA ratio were oppositely regulated in post-injury spinal cord and proximal nerve, with decreased expression in spinal cord and incre- ased expression in proximal nerve. Consistently, sciatic nerve injury reduced mitochondrial amount and ATP content in spinal cord while elevated mitochondrial amount and ATP con-tent in proximal sciatic nerve.

Materials and methods

Animals and surgical procedures

All experimental programs were approved by Ethics Committee of Peking University People’s Hospital. Sprague Dawley adult male rats of 220-250 g were used in the present study. Firstly, rats were anesthetized using sodium pentobarbital, 30 mg/kg and then the right sci-atic nerves were exposed from the muscle gap and cut off 5 mm distal to the lower edge of piri-formis with a microscopic scissors. In order to inhibit the retraction of nerve, the proximal stump was sutured to adjacent muscles with 10-0 nylon sutures. Rats with the sciatic nerve exposed but without nerve axotomy were used as sham control. The spinal cord at lumbar enlargement and proximal sciatic nerve (5 mm proximal side to the lesions site of sciatic nerves) at the lesioned side were taken and quickly frozen with liquid nitrogen for further

measuring reagent. The light emission was determined with Luminometer (Bio-Rad). ATP concentration of the samples was calculated according to ATP standard curve. All data was normalized with the protein abundance by using the Bradford method.

Transmission electron microscopic examina-tion

The processing of ventral part of the spinal cord of lesioned side or sciatic nerves for electron microscopic analysis was performed as previ-ously reported [8]. Briefly, ventral cord speci-mens and the proximal stump of the sciatic nerves were obtained and fixed in 2.5% glutar-aldehyde with 0.1 M phosphate buffer (pH, 7.2) for four hours. After fixation in osmium tetrox-ide and dehydration by gradient acetone, tis-sues were embedded in Epon-812-resin, sec-tioned by ultramicrotome for semithin sec- tionsand stained with toluidine blue. The sec-tions were examined with a light microscope to locate motor neurons at ventral horn and nerve fibers. Then ultrathin sections were harvested and examined with a transmission electron microscope (JEM-1230, Tokyo, Japan) equipped with a digital photography.

RNA isolation and reverse transcription (RT), real-time PCR analysis

Total RNA extraction from ventral part of the spinal cord or proximal sciatic nerve was com-

Figure 1. Surgical procedures of generation of sciatic nerve injury model. A. The right sciatic nerve was exposed and sectioned 5 mm distal to the lower edge of piriformis. The ventral horn of the lesion side of cord and proximal sciatic nerve (5 mm proximal side to the lesions site of sciatic nerves) at the lesion side were taken. B. Ulcer of the limb at sciatic nerve injured side.

experiments, at time points of 3, 7, 14, 21, and 28 days after operation (Figure 1A).

ATP measurement

ATP contents in ventral horn of the right side of the cord or proximal sciatic nerve were determined by an ATP Assay System (Vigorous, China) acc- ording to the principle of lucif-erin-luciferase following the manufacturer’s instructions. Briefly, the spinal cord and sciatic nerves were harvest-ed, homogenized in tissue lysis buffer, and then centri-fuged at 10000 g for 15 min at 4°C. Supernatant was col-lected and added to the ATP

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pleted using TRIzol (Invitrogen, Carlsbad, CA, USA). Reverse transcription was performed with 3 ug total RNA at 25°C for 5 min, 42°C for 60 min, 70°C for 15 min using Oligo (dT) and GoScript II reverse transcriptase (Promega). The cDNA product was stored at -20°C. For Real-Time PCR, the reaction system of 10 ul was adopted containing 50 ng of cDNA. The sequences of primers were as following: PGC-1α: forward, ACGAAAGGCTCAAGAGGGACGAAT; Reverse, CACGGCGCTCTTCAATTGCTTTCT; NRF-1: forward, TGTCACCATGGCCCTTAACAGTGA; Reverse, TGAACTCCATCTGGGCCATTAGCA; TF- AM: forward, AAATGGCTGAAGTTGGGCGAAGTG; Reverse, AGCTTCTTGTGCCCAATCCCAATG; GA- PDH: forward, CGGAGTCAACGGATTTGGTCGTAT; Reverse, AGCCTTCTCCATGGTGGTGAAGAC.

Western blot analysis

Tissue samples were homogenized in lysis buf-fer (containing 0.15 mol/l NaCl, 0.1 mol/lTris-Cl, 1 mmol/l EDTA, 1 mmol/l PMSF, 10 μg/ml Aprotinin, 10 μg/ml Pepstatin A, 1% TritonX- 100); lysates were then centrifuged at 12,000 g for 10 min at 4°C. The supernatant was col-lected and total protein was measured using Bradford Assay Kit. An equal volume of 2× sam-ple buffer (100 mM Tris-HCl pH 6.8, 2.5% SDS, 20% glycerol, 0.006% bromophenol blue and

10% β-mercaptoethanol) was added. Samples were boiled and then separated by 8-12% SDS-PAGE and transferred to polyvinylidenedifluo-ride membranes. Membranes were blocked with 5% non-fat milk for 2 hour at room tem-perature, then probed with primary antibodies (PGC-1α, NRF-1, TFAM or GAPDH) at 4°C over-night, and then incubated with horseradish-peroxidase-conjugated secondary antibodies. GAPDH was used as a loading control.

DNA isolation and real-time PCR analysis

Genome DNA from ventral part of the spinal cord or proximal sciatic nerve was isolated by using the Universal Genome DNA Kit (Beijing Zoman Biotechnology, China). The mtDNA copy number was determined by PCR through the relative number of Cyt B to genome β-actin [9, 10].

Statistical analysis

SPSS 17.0 software (SPSS, Chicago, IL, USA) was used for data analysis. Data were present-ed as mean ± SEM. Differences in the means between control group and experimental gr- oups were analyzed using Student’s t test. P<0.05 was considered statistically signifi- cant.

Figure 2. mRNA and protein changes of mitochondrial biogenesis biomarkers in spinal cord. mRNA levels of PGC-1α (A), TFAM (B), and NRF-1 (C) were suppressed at indicated days after sciatic nerve injury. D. Representative western blot showing the decreased protein level of PGC-1α, TFAM, and NRF-1 at 7 days after injury, and (E) the average data of (D). n=6 sides for each group. *P<0.05, **P<0.01 vs. control.

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Results

Generation of sciatic nerve injury model

Comparing with sham limbs, the operated limbs after sciatic nerve transection displayed dener-vation, characterized by dropping ankles and claudication. Among rats under sciatic nerve injury, some rats bitted their toes and the limbs appeared ulcer, even all of the toes disap-peared 1 week after operation (Figure 1B). The amount of rats with limb ulcers increased with time and most ulcers gradually aggravated. The operated limbs couldn’t walk 4 weeks after sur-gical operation, indicating the effective model-ing. In control group, the rats could walk normally.

Suppressed mitochondrial biogenesis in spinal cord after sciatic nerve injury

To determine if sciatic nerve axotomy disturbs mitochondrial biogenesis in spinal cord, we examined the gene expression levels of several mitochondrial biogenesis biomarkers, PGC-1α, TFAM, and NRF-1, in tissue samples taken from ventral cord (L4-L6) of axon injury side or sham control side, by real time PCR (RT-PCR). Interestingly, we found that after sciatic nerve surgery, the mRNA levels of PGC-1α, TFAM, and

NRF-1 in spinal cord from injury side showed similar downward trend as comparing with that before operation (Figure 2A-C). The mRNA lev-els of three mitochondrial biogenesis biomark-ers slightly decreased at as early as 3 days after nerve injury, and significantly decreased 7 days after injury, to a level of 70.5±5.0% for PGC-1α (Figure 2A), 66.3±7.1% for TFAM (Fi- gure 2B), and 53.6±11.6% for NRF-1 (Figure 2C), respectively, as compared to tissues from sham control rats. And at 14 days after sciatic nerve injury, the mRNA levels of the three mito-chondrial biogenesis biomarkers decreased more, to 42.7±5.9% for PGC-1α (Figure 2A), 44.5±8.5% for TFAM (Figure 2B), and 47.2± 6.0% for NRF-1 (Figure 2C), comparing with control sides. Moreover, the low mRNA levels kept till at least one month after surgery.

In consistent with mRNA levels, protein levels of PGC-1α, TFAM, and NRF-1 in spinal cord from injury side also decreased at 7 days after sci-atic nerve transection comparing with control side, as indicated by western blotting (Figure 2D). And highly comparable to the decreased mRNA levels, protein levels of PGC-1α, TFAM, and NRF-1 in the injury side of spinal cord decreased to 51.8±8.6%, 44.1±4.4%, and 58.7±9.1% to that of the control side, respec-tively (Figure 2E).

Figure 3. mRNA and protein changes of mitochondrial biogenesis biomarkers in proximal nerve stump. Real time PCR (RT-PCR) analysis for PGC-1α (A), TFAM (B), and NRF-1 (C) in proximal nerve stump at indicated days after sci-atic nerve injury. (D) Representative western blot showing the increased protein levels of PGC-1α, TFAM, and NRF-1 at 3 days after injury. (E) The average data of (D). n=6 sides for each group. *P<0.05, **P<0.01 vs. control.

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Elevated mitochondrial biogenesis in proximal sciatic nerve after sciatic nerve injury

Next, we sought to know if mitochondrial bio-genesis was disturbed in nerve after sciatic nerve injury. We also examined the mRNA lev-els of PGC-1α, TFAM, and NRF-1 by RT-PCR (Figure 3A-C). Surprisingly, in contrast to that in spinal cord, we found the mRNA levels of PGC-1α, TFAM, and NRF-1 in proximal nerve signifi-cantly increased from as early as 3 days after sciatic nerve injury, to 1.49±0.17, 1.41±0.13, and 1.94±0.22 folds of control nerve. The increased mRNA levels of PGC-1α, TFAM, and NRF-1 persisted at 7, 14, 21 28 days after sci-atic nerve injury, to 2.21±0.14, 1.72±0.12, 2.10±0.13, and 1.91±0.13 folds for PGC-1α, 2.17±0.21, 2.41±0.28, 2.22±0.14, and 2.28± 0.17 folds for TFAM, 2.14±0.23, 2.57±0.25, 2.18±0.27, and 2.34±0.20 folds for NRF-1, respectively, as compared to that of control nerve.

Similarly, protein levels of PGC-1α, TFAM, and NRF-1 increased comparably to mRNA levels (Figure 3D), to 1.44±0.12, 1.54±0.15, and 1.41±0.09 folds of that in control nerve fibers, at 7 days after sciatic nerve transection (Figure 3E).

Mitochondrial to nuclear DNA ratios changes after sciatic nerve injury

In order to evaluate mitochondria biogenesis from another aspect, we tested mitochondria DNA. At spinal cord, the mitochondrial to nucle-ar DNA ratio was 665.27±41.43 in control

group, and significantly decreased 7 days after injury, to a level of 390.56±42.29 (Figure 4A). The decreased mitochondrial to nuclear DNA ratio persisted at 14, 21 28 days after sciatic nerve injury, to 300.46±54.21, 374.35±39.98 and 346.62±60.27 (Figure 4A). At proxiaml nerve, the mitochondrial to nuclear DNA ratio was 24.79±5.64 in control group, and signifi-cantly increased 3 days after injury, to a level of 68.10±7.13 (Figure 4B). The increased mito-chondrial to nuclear DNA ratio persisted at 7, 14, 21, 28 days after sciatic nerve injury, to 62.86±10.3, 72.10±8.30, 61.27±8.11 and 62.10±8.39 (Figure 4B).

Ultrastructural alterations of mitochondria after sciatic nerve injury

We then performed transmission electron mi- croscopic (TEM) examination on ventral horn neurons and proximal sciatic nerve to visualize the ultra-structural changes of mitochondria. While numerous mitochondria with roughly sim-ilar length around 1 μm was observed in ventral horn neurons from control side (Figure 5A), less but enlarged mitochondria with length ranging from 1 μm to 3 μm was found in ventral horn neurons from injury side at 7 days after sciatic nerve transection (Figure 5B), support-ing the finding of declined mitochondrial bio-genesis, and further suggesting the dysfunc-tion of mitochondrial in neuronal bodies, even though the injury occurred in peripheral nerve fiber. In contrast to the decreased number of mitochondria in neuronal bodies, the number of mitochondria in Schwann cells around proxi-

Figure 4. Mitochondrial to nuclear DNA ratios changes on ventral horn and proximal sciatic nerve area after sciatic nerve injury. A. Mitochondrial to nuclear DNA ratio in ventral horn at indicated days after injury. B. Mitochondrial to nuclear DNA ratio in the injured proximal sciatic nerve area at indicated days after injury. n=6 samples from differ-ent rat for each group. *P<0.05 vs. control.

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mal sciatic nerve after injury increased signifi-cantly 3 days after sciatic nerve transection (Figure 5D), as compared with those in Schwann cells from control side (Figure 5C), indicating an essential role of mitochondria in proximal Schwann cells during axon regeneration. Toge- ther, our TEM data here are in general agree-ment with above mentioned down-regulated mitochondrial biogenesis in ventral spinal cord and up-regulated mitochondrial biogenesis in proximal nerve after sciatic nerve injury.

Bioenergetic changes after sciatic nerve injury

Mitochondria are the main source of intracellu-lar ATP production, so we measured ATP con-tents in both spinal cord and in proximal sciatic nerve. In parallel with the decreased mRNA and protein levels of mitochondrial biogenesis bio-markers, ATP contents in ventral horn of injured side reduced from two weeks after injury, to 57.2±11.0% of that in control side, and the declined ATP content persisted till at least one month (Figure 5E). Moreover, ATP content in the injured proximal sciatic nerve increased to

1.33±0.11 fold of control nerve at 3 days after injury, up to 1.77±0.17 fold at 7 days after inju-ry, and keep at the high level till at least one month (Figure 5F).

Discussion

In this study, we found that mitochondrial func-tion was oppositely regulated in post-injury ven-tral spinal cord and nerve proximal to the site of injury at least one month after nerve transec-tion, with a lower bioenergetic level in spinal cord and higher level in proximal nerve area. In post-injury spinal cord, mitochondrial biogene-sis biomarkers PGC-1α, NRF-1, TFAM and mito-chondrial to nuclear DNA ratio were down-regu-lated at both mRNA and protein levels, and consistently, less mitochondria and decreased ATP content were observed. In contrast with the changes in spinal cord, mitochondrial biogene-sis markers and ATP production were up-regu-lated in post-injury sciatic nerve area, and more mitochondria were found. In addition, the changes of mitochondria at proximal nerve area were earlier than at neuronal cell bodies,

Figure 5. Transmission electron microscope examination and bioenergetic changes on ventral horn and proximal sciatic nerve area after sciatic nerve injury. Ultra structures of mitochondria in control neuron (A), and in neuron 7 days after sciatic nerve transection (B). (C) Mitochondria in control Schwann cells, and (D) in Schwann cells around proximal sciatic nerve 3 days after sciatic nerve transection (E) ATP content in ventral horn at indicated days after injury. (F) ATP content in the injured proximal sciatic nerve area at indicated days after injury. n=6 samples from different rat for each group. *P<0.05, **P<0.01 vs. control.

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3 days compared with 7 days after axotomy. Collectively, our data indicated the important but distinct roles of mitochondria in regulating degradation and regeneration in neuronal cell body and nerve after peripheral nerve injury.

Nerve transection causes a variety of modifica-tions in gene expression and cellular metabo-lism in neuron body, which likely are parts of the process to protect the cell, regenerate the injured axon and restore cellular functions [11-14]. However, whether mitochondria are invo- lved in this process is still unclear. The present study provides evidence showing that mito-chondria in both ventral spinal cord and proxi-mal nerve are profoundly involved in cellular modification after nerve transection. In spinal cord, less and enlarged mitochondria was fo- und at one week after sciatic nerve transec-tion, in consistent with previous reports of the swelling and irregularly arranged mitochondria in dorsal root ganglion and spinal cord moto-neuron after sciatic nerve axotomy [15, 16]. Moreover, decreased expressions of genes related to mitochondrial metabolism were also reported [3]. The changed mitochondrial mor-phology and metabolism related genes in neu-ron body after nerve transection suggest the dysfunction of mitochondria. Indeed, in the present study, we found that ATP content in spi-nal cord decreased two weeks after nerve inju-ry. The mechanisms underlying the dysfunc-tional mitochondria and the subsequent de- creased energy supply in spinal cord after nerve transection is currently unclear and needs further investigation. However, the sup-pressed mitochondrial function may prevent the neural regeneration after injury, and caus-ing poor functional recovery following nerve axotomy, particularly when surgical repair is delayed post injury. In contrast to the dysfunc-tional mitochondria in spinal cord, we found mitochondrial ATP production in nerveproximal to injury site was largely increased, indicating an active mitochondrial metabolism status responding to nerve injury, which may contrib-ute to the proximal axon regeneration. The find-ing that more condensed mitochondria app- eared in Schwann cells around nerve fiber area further supports this view.

Mitochondrial regeneration is a very complex process, which is regulated by mitochondrial and nuclear genes. While nuclear transcription factors, mainly nuclear respiratory factor 1 and

2 (NRF-1 and -2), and coactivator such as PGC-1α regulates the expression of nuclear-gene encoded mitochondrial proteins [17], the ex- pression of mitochondria-gene encoded pro-teins are mainly controlled by the mitochondrial transcription factor A (TFAM) [17-19]. In our present study, we found that, in spinal cord 7 days after nerve transection, both mRNA and protein levels of PGC-1α, NRF-1, and TFAM largely decreased, suggesting decreased mito-chondrial biogenesis after peripheral nerve injury. Less and enlarged or vacuolated mito-chondria with loose cristae in spinal cord sup-port the down-regulation of mitochondrial bio-genesis. Moreover, energy supply in neuron body concurrently down-regulated, indicating that the regulation and balance of these bio-genesis related genes determine mitochondrial oxidative capacity. In agreement with our find-ings, it has been reported that deprivation of each one of these genes leads to decreased mitochondria amount and lower neuronal activ-ity [20]. Interestingly, opposite changes occu- rred in nerve. Expression levels of PGC-1α, NRF-1, and TFAM were significantly increased companied with increased ATP content. There are two possible explanations for the increased ATP supply. One is that mitochondrial transpor-tation from neuron body to proximal nerve increased largely after axotomy [21]. Another is the up-regulated mitochondrial biogenesis in Schwann cells, which promote the re-growth of the axon. Anyway, the higher bioenergetic level at proximal nerve fiber area during early stage of sciatic nerve injury may serve as a protective response of sciatic nerve to injury.

In summary, our data have shown that mito-chondrial biogenesis were oppositely regulated in post-injury spinal cord and proximal nerve following sciatic nerve injury, with decreased level in spinal cord and increased level in proxi-mal nerve. And consistently, ATP content redu- ced in spinal cord while elevated in proximal sciatic nerve. These findings not only extend our understanding of changes of the mitochon-drial biogenesis and bioenergetic metabolism after peripheral nerve injury, but also may pro-vide treatment strategies for peripheral nerve injury through adjusting mitochondrial biogen-esis and metabolic status.

Acknowledgements

This work was supported by a grant from National Science Foundation of China (3137-

Mitochondrial biogenesis during sciatic nerve injury

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1210) and the National Key Basic Research Program of China (2014CB542200). The sup-porter had no role in the study design, datacol-lection and analysis, decision to publish, or preparation of themanuscript. We thank the electron microscope department, ultrastruc-tural center in Peking University First Hospital for the analysis of electron microscopy test.

Disclosure of conflict of interest

None.

Address correspondence to: Dr. Ming Zheng, Depart- ment of Physiology and Pathophysiology, Health Science Center, Peking University, Beijing 100191, China. Tel: 86-10-8280-2043; E-mail: zhengm@bjmu.edu.cn; Dr. Baoguo Jiang, Department of Trauma and Orthopaedics, People’s Hospital, Peking University, Beijing 100044, China. Tel: 86-10-8832-6550; E-mail: jiangbaoguo@vip.sina.com

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