Neuropathic pain is an often crippling condition with a prevalence of approximately 7% to 8% within a western population.1 It is induced by damage to central or peripheral neural structures with diverse underlying pathophysiologic causes such as metabolic, infectious, autoimmune, or mechanical injury. The disease usually develops with a temporal latency after the eliciting event and its clinical presentation is characterized by spontaneous pain, hyperalgesia, and allodynia. Typical examples of the disease caused by mechanical injury to the peripheral nerve system are radiculopathies after disk herniation, and phantom pain and nerve entrapment syndromes. Complex regional pain syndrome (CRPS) displays similar symptoms but is considered a separate entity, because the classic presentation (CRPS I, M. Sudeck, reflex sympathetic dystrophy) is defined by the absence of verifiable neuronal damage.2 The pharmacologic therapy for neuropathic pain syndromes differs from that of acute pain insofar that mainly anticonvulsants, antidepressants, and long-acting opioids are used.3 However, although there are efficient medications, some patients do not benefit from them or are forced to discontinue the drugs because of side effects, thus creating a demand for better understanding of the underlying pathophysiology and development of new therapeutic strategies.
A widely used animal model for the study of mechanically induced neuropathic pain is the chronic constriction injury (CCI) model of the rat sciatic nerve, which was introduced and established by Bennett and Xie4 in 1988. Using this CCI model, our group confirmed alterations in pain sensation, namely, spontaneous pain as well as responses to innocuous and painful stimuli (mechanical hypersensitivity and thermal allodynia).5 However, we were able to show that muscle tissue of the affected hindlimb presented with caspase 3 activation and marked cell apoptosis.5,6 This novel finding prompted us to investigate whether muscle cell apoptosis contributes to neuropathic pain. Herein, we demonstrate that CCI animals treated with a pan-caspase inhibitor lack not only muscle cell apoptosis, but also pathologic pain, suggesting a possible contribution of muscle cell apoptosis in mediating neuropathic pain.
The experiments were performed on male Sprague-Dawley rats (body weight bw 225–250 g; Charles River Laboratories, Sulzfeld, Germany) provided with water and standard laboratory chow ad libitum. The experimental protocol was approved by the local animal rights protection authorities and followed the Public Health Service Policy on Humane Care and Use of Laboratory Animals and the Guide for the Care and Use of Laboratory Animals.
Induction of CCI of the Sciatic Nerve
Neuropathy was induced according to the procedure, previously described by Bennett and Xie in 1988. Briefly, rats were anesthetized with 6% pentobarbital-sodium (55 mg/kg bw, intraperitoneally, Narcoren; Merial GmbH, Hallbergmoos, Germany). Under aseptic conditions, the right common sciatic nerve was exposed at the level of the middle thigh by blunt dissection. Proximal to the trifurcation, the nerve was carefully freed from the surrounding connective tissue and 4 chromic cat gut ligatures (4-0; Ethicon, Norderstedt, Germany) were tied loosely around it with approximately 1-mm spacing. After hemostasis was confirmed, the incision was closed in layers (n = 11). Sham-operated rats served as controls and underwent the same surgical procedure with exposure, but not ligation of the sciatic nerve (n = 6). Animals were then allowed to recover from anesthesia and surgery and were kept 1 per cage with free access to water and standard laboratory chow.
CCI animals received either the pan-caspase inhibitor zVAD (OMe)-fmk (n = 5) (3.3 mg/kg bw, intraperitoneally; Alexis, Grünberg, Germany)7,8 or equivalent volumes of vehicle (n = 6) postoperatively and every 24 hours up to day 4 after sciatic nerve ligation via intraperitoneal injection.
All animals were allowed to habituate to the experimental conditions by allowing them to spend several hours in the laboratory on days −3 to 0 preceding any test. Behavioral observation tests were performed on the day before surgery as well as on days 1 and 4 after the operation.
Spontaneous behavior, i.e., different positions of the lesioned hindpaw were observed and rated in a custom-made plexiglas cage 16 × 15 × 22 cm without intervention of the observer, as described by Attal et al.9 After 10-minute habituation, different positions of the lesioned hindpaw were rated 3 times for 300 seconds during a 15-minute period according to a numerical scale: 0 = the operated paw is pressed normally on the floor; 1 = the paw rests lightly on the floor and the toes are in a ventroflexed position; 2 = only the internal edge of the paw is pressed on the floor; 3 = only the heel is pressed on the floor and the hindpaw is in an inverted position; 4 = the whole paw is elevated; and 5 = the animal licks the operated paw. The score was recorded over a 5-minute period and provides an index of spontaneous pain intensity for each rat. It was calculated by the following formula: t1 + 2t2 + 3t3 + 4t4 + 5t5/300 seconds, where t1, t2, t3, t4, and t5 are the durations in seconds spent in the categories 1, 2, 3, 4, or 5, respectively.
By changing the bottom layer of the cage to a plate set to either 40°C or 4°C, responses of the lesioned hindpaw to thermal stimuli were additionally analyzed, as described above.
For assessment of tactile allodynia, the hindlimb withdrawal threshold evoked by stimulation of the hindpaw by von Frey filaments was determined while the rat was placed on a metal mesh floor with 0.6 × 0.6 cm cells. After a 10-minute acclimation period, mechanical stimuli were applied to the affected foot pad with 6 calibrated filaments (Semmes-Weinstein monofilaments; North Coast Medical, Inc., Morgan Hill, CA) ranging from 0.6 to 15.0 g (5.83–147 mN) until the filament bent. A single trial of stimuli consisted of 6 applications of filaments at a frequency of 1/second. Five trials were performed at 3-minute intervals. The paw withdrawal response frequency in each of these 5 trials was expressed as a percent response frequency: foot withdrawals/5 × 100 = % response frequency.10 The same procedure was repeated for each filament in ascending order starting from the weakest.
Analysis of Skin Temperature and Edema
Under brief isoflurane anesthesia, skin temperature and volume of the hindpaw were assessed by placing a surface electrode (Mon-a-therm Skin Temperature Probe 90045; Mallinckrodt Medical, St. Louis, MO) on the plantar region of the hindpaw and submersion of the paw in a calibrated cylinder filled with water, so the water level just reached the proximal tip of the glabrous surface of the heel, respectively. Data are presented as differences in plantar skin temperature and hindlimb volume of the operated paws on days 1 and 4 after CCI compared with the respective values preoperatively (day 0). In addition, the extent of skeletal muscle edema was determined in biopsies obtained from the tibialis anterior muscle at the end of the 4-day observation period. After assessment of wet weight, the muscle was dried for 72 hours in a laboratory oven (60°C) and weighed again (dry weight) for calculation of the wet-to-dry ratios as an additional indicator of edema formation.
Intravital Fluorescence Microscopy
Pentobarbital-anesthetized animals were placed on a heating pad for maintenance of body temperature at 37°C, tracheotomized, and their lungs mechanically ventilated (tidal volume 1 mL/100 g bw; 50 breaths/min; PaO2 >75 mm Hg; PaCO2 30–35 mm Hg). Polyethylene catheters in the right carotid artery and left jugular vein served for injection of fluorescent dyes and monitoring of central hemodynamics (Sirecust; Siemens, Germany). The extensor digitorum longus (EDL) muscle was microsurgically prepared to allow direct access for in vivo high-resolution multifluorescence microscopy.11,12 NADH (nicotinamide adenine dinucleotide) autofluorescence of muscle tissue was monitored by ultraviolet epiillumination. For contrast enhancement of the microvascular network and for in vivo staining of leukocytes, a single bolus of fluorescein-isothiocyanate-labeled dextran (15 mg/kg bw; Sigma, Deisenhofen, Germany) and rhodamine 6G (0.15 mg/kg bw; Sigma) was injected IV.
As previously described, functional capillary density was defined as the total length of red blood cell–perfused capillaries per observation area and given in cm/cm2.5,13 For assessment of leukocyte-endothelial cell interaction in postcapillary venules, flow behavior of leukocytes was analyzed for free-floating, rolling, and adherent leukocytes. Rolling leukocytes were defined as those cells moving along the venular wall at a velocity <40% of that of leukocytes at the centerline and were expressed as a percentage of the total leukocyte flux. Venular leukocyte adherence was defined as the number of leukocytes not moving or detaching from the endothelial lining of the vessel wall during an observation period of 20 seconds and expressed as nonmoving cells per endothelial surface (n/mm2).5,6
Dorsal Root Ganglion Preparation and Immunohistochemistry
At the end of the experiment, the rats were deeply anesthetized with a pentobarbital overdose (200 mg/kg bw, intraperitoneally) and intracardially perfused with 100 mL 0.9% saline followed by 350 mL fixative solution (4% paraformaldehyde in phosphate-buffered saline). The ipsilateral dorsal root ganglia (DRG) of L4-5 were rapidly dissected under 25 × magnification and placed in the same medium for 48 hours. After embedding in paraffin, 4-μm sections were cut and exposed to an apoptosis-specific staining kit (indirect in situ DNA nick end labeling TUNEL assay, ApopTag; CHEMICON International, Inc., Temecula, CA) according to the manufacturer’s instructions. Analysis was conducted with a 400× magnification light microscope. Sham-operated animals served as controls.
Transmission Electron Microscopy
Sciatic nerves were rapidly dissected using the previously used surgical access, resected, and immersed overnight in 0.1 M Sorensen phosphate buffer containing 1% paraformaldehyde and 2% glutaraldehyde. Subsequently, small samples of the sciatic nerve were osmicated, dehydrated through a graded series of ethanol, embedded in Epon, and polymerized for 48 hours at 60°C. Ultrathin sections were mounted on mesh grids and contrasted with uranyl acetate and lead citrate. Sections were analyzed using a Zeiss 902 electron microscope (Zeiss, Oberkochen, Germany) at 80 kV. Photographs were taken with a CCD camera, and pictures were adjusted using Adobe Photoshop CS2 (Adobe Systems, San Jose, CA).
The EDL muscle was removed and fixed in 4% phosphate-buffered formalin for 2 to 3 days and then embedded in paraffin. From the paraffin-embedded tissue blocks, slices were cut and dyed using the technique mentioned above. Quantitative analysis was performed by counting the number of TUNEL-positive tissue-confined cells in 50 consecutive high-power fields (×400 magnification).
After approving or disproving the assumption of normality and equal variance across the groups, statistical evaluation was performed using analysis of variance followed by the Holm-Sidak test to compensate for multiple comparisons (SigmaStat; Jandel, San Rafael, CA). For the sake of clarity and comprehensiveness of this report, results are given as mean ± SEM. Statistical significance was set at P < 0.05.
Loose ligation of the sciatic nerve in rats led to cold- and heat-evoked pain (Fig. 1, A and B). By placing the rats on a chilled 4°C cold plate, a prolonged hindpaw withdrawal, i.e., an increase of nociception index was seen in the nerve-injured group, but not in sham-operated controls on days 1 and 4 (Fig. 1A). Heat-evoked pain-related behavior was tested on a 40°C warm plate and also revealed significantly higher pain scores of the affected hindlimb of nerve-ligated animals compared with controls (Fig. 1B). Hindpaw withdrawal responses upon cold and heat stimulation were drastically reduced in animals with sciatic nerve ligation and concomitant zVAD application.
The measure of mechanical nociceptive withdrawal response, involving the application of von Frey fibers over the plantar aspect of the hindpaw, showed a slight increase in response frequency with increased fiber strength before sciatic nerve ligation and on day 1 after surgery, however, without significant differences among animals in the 3 groups (Fig. 2, left and mid panels). In contrast, on day 4 after surgery, von Frey thresholds decreased in the nerve-ligated animals, presenting significantly higher response frequencies when stimulated with von Frey filaments ranging from 10 to 15g (Fig. 2, right panel). Again, animals that underwent nerve ligation and zVAD application seemed to benefit from pan-caspase inhibition with respect to mechanically induced pain. Withdrawal frequency in these animals reached only 20% upon 15-g von Frey filament challenge, revealing physiologic pain behavior as seen in animals before surgery or upon sham operation (Fig. 2).
Skin Temperature and Paw Edema
During the observation period, no statistically significant differences in paw temperature could be found. However, the nerve-ligated animals showed a tendency toward higher temperatures on the first postoperative day (Table 1).
As measured by water displacement, rats developed volume differences in the nerve-ligated hindpaw on days 1 and 4 after surgery (Table 1). Sham-operated controls and zVAD-treated nerve-ligated animals displayed no significant edema formation with an increase in paw volume below 0.1 mL, whereas nerve-ligated animals had hindlimb volume differences of approximately 0.24 mL (Table 1). Accordingly, wet-to-dry weight ratios of tibialis anterior muscle were highest in the nerve-ligated animals (Table 1).
Analysis of Muscle Tissue Microcirculation and Tissue Apoptosis
In line with previous experiments from our group, in vivo microscopy did not reveal nerve ligation–associated deteriorations of the EDL muscle microcirculation in all studied groups (Table 2). Leukocyte flow behavior revealed low interaction of inflammatory cells with the microvascular endothelial lining. Both the number of cells rolling along as well as those firmly adherent to the endothelial lining were within the range of physiologic values and did not differ among animals in the 3 groups (Table 2). Moreover, EDL muscle microcirculation in all animals showed homogeneous nutritive perfusion with absence of individual capillary perfusion failure, as reflected by physiologic values of functional capillary density (Table 2).
Analysis of Muscle Tissue Apoptosis
Assessment of apoptotic cells by immunohistochemical DNA nick end labeling demonstrated a 10-fold increase of apoptosis in muscle tissue of nerve-ligated versus sham-operated control animals (Fig. 3). zVAD completely suppressed tissue apoptosis similar to control values, underlining the efficacy of caspase inhibition to block the apoptotic pathway (Fig. 3).
Analysis of DRG Apoptosis
Animals in neither the CCI group nor sham-operated animals showed any apoptosis in ipsilateral DRG neurons. However, individual apoptotic satellite cells surrounding the neuron perikarya were found in both groups (Fig. 4).
Analysis of the nerve ultrastructure by transmission electron microscopy revealed a similar extent of nerve compression–like lesions (Fig. 5) in both experimental groups. In contrast to the sham-operated group (Fig. 5A) sciatic nerves of CCI-treated rats presented with axonal degeneration, myelin sheath intussusceptions (“tomacula”), formation of myelin figures within Schwann cells, and peri- and endoneural edema (Fig. 5, B and D) regardless of treatment with zVAD (Fig. 5, C and E). The distribution pattern of damaged nerves appeared comparable within both experimental groups, as can be seen in the semithin sections of the sciatic nerve (Fig. 5, B and C).
In this study, we were able to confirm our previous findings that painful neuropathy after loose ligation of the sciatic nerve is not associated with an inflammatory response, but is characterized by muscle cell apoptosis. We now show that blockade of myocyte apoptosis using the pan-caspase inhibitor zVAD is paralleled by a reduction in thermal and mechanical allodynia.
CCI of the sciatic nerve damages both myelinated and unmyelinated peripheral axons with the injury being more pronounced in the myelinated fibers. Moreover, central neurochemical alterations, such as a decrease of substance P and galanin in laminae 1 and 2 of the spinal cord dorsal horn, were found, possibly explaining the development of pain and impairment of muscle innervation.14
The effect of a peripheral nerve injury on the central nervous system is an issue for ongoing debate. Some studies suggest that the development of neuropathic pain depends on induction of apoptosis in neurons in the dorsal horn of the spinal cord,15–17 whereas others reject neuronal apoptosis as a consequence of peripheral nerve injury.18,19 By administering zVAD intrathecally, Scholz et al.17 observed prevention of spinal neuronal apoptosis leading to the absence of symptoms of neuropathic pain in a spared-nerve injury model. In contrast, we deliberately administered zVAD peripherally (intraperitoneally). Because zVAD does not cross the intact blood-brain barrier,20,21 and has a half-life of only 40 minutes,22 an action on central neurons seems unlikely in the present setting.
The DRG is another possible place of action for zVAD in our experiments. In contrast to other peripheral and central neural structures, the DRG has no substantial blood-brain/nerve barrier.23 However, in accordance with the most recent results by Schaeffer et al.,24 we could show that apoptosis of afferent neurons does not occur in the DRG during the early phase up to day 4 after CCI, which suggests that the antiapoptotic effect of zVAD is of minor pathophysiologic relevance for the development of neuropathic pain. Nonetheless, other yet unknown effects of zVAD could take place and have a role in the formation of neuropathic pain.
After nerve injury such as CCI, the permeability of the blood-nerve barrier is increased.23 Therefore, zVAD could affect peripheral nerves. To address this issue, we performed electron microscopic studies on the injured nerve. In contrast to normal nerve morphology in sham controls, nerves subjected to CCI treatment had typical compression-like lesions with no major differences in extent and pattern as in the case of zVAD treatment.25
In an earlier study, we reported that loose ligation of the sciatic nerve is associated with increased activation of caspase 3 protein and, as a consequence, with increased cell apoptosis within skeletal muscle tissue.5 This is substantiated by the fact that we can now demonstrate an absence of apoptotic myocytes in muscle tissue of CCI animals upon blockade of caspases by the pan-caspase inhibitor zVAD.
From a previous study, we know that spontaneous apoptosis of healthy muscle cells rarely occurs.26 Because the number of muscle cells undergoing apoptosis spontaneously is negligible, antiapoptotic effects brought about by zVAD treatment were not expected. Therefore, we did not include a control group that received a sham operation and zVAD treatment.
Defective muscle innervation, as found in patients with polyneuropathy, was shown to prompt muscle fibers to undergo apoptosis.27 Similarly, apoptotic DNA fragmentation, assessed by TUNEL technique, was reported for denervated muscle fibers in rats, and the absence of caspase 3, as the main executing enzyme of apoptosis, protected against denervation-induced apoptosis of skeletal muscle in vivo.28,29 It is important to state that loose nerve ligation with intraneural edema formation and not only complete nerve transection with muscle tissue denervation was found to cause muscle tissue apoptosis. Interestingly, muscle apoptosis lacked local signs of inflammation and invading cells. This is in line with the fact that phagocytosis of apoptotic cells is mostly described as a noninflammatory process, leading to active suppression of proinflammatory cytokine production by macrophages and immature dendritic cells, whereas the response toward cells that die by necrosis induces a proinflammatory cytokine response.30
The prevention of myocyte apoptosis by zVAD administration was paralleled by a remarkable attenuation of neuropathic pain symptoms. We are aware that the present study provides no pathogenetic explanation of how muscle cell apoptosis can lead to neuropathic pain. However, we could demonstrate in a previous study that a supernatant of traumatized muscle tissue is able to exaggerate pain syndromes in a CRPS model.6 Therefore, it is reasonable to suggest that proalgesic, paracrine-acting substrates are released by muscle cells undergoing apoptosis and elicit symptoms of neuropathic pain.
In summary, chronic nerve injury does not lead to inflammation in this model but is able to induce apoptosis in muscle cells and produce symptoms of neuropathic pain. Muscle cell apoptosis as well as concomitant symptoms of neuropathic pain are preventable by treatment with the pan-caspase inhibitor zVAD, suggesting a contributing role of skeletal muscle cell apoptosis in the development of neuropathic pain.
Name: Georg Gradl, MD, PhD.
Contribution: This author helped design the study, conduct the study, analyze the data, and write the manuscript.
Attestation: Georg Gradl has seen the original study data, reviewed the analysis of the data, approved the final manuscript, and is the author responsible for archiving the study files.
Name: Philipp Herlyn, MD.
Contribution: This author helped design the study, conduct the study, analyze the data, write the manuscript, and is co-first author.
Attestation: Philipp Herlyn has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
Name: Burkhard Finke, MD.
Contribution: This author helped conduct the study and analyze the data.
Attestation: Burkhard Finke has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
Name: Philip Gierer, MD, PhD.
Contribution: This author helped design the study, conduct the study, and analyze the data.
Attestation: Philip Gierer has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
Name: Andreas Wree, MD, PhD.
Contribution: This author helped conduct the study.
Attestation: Andreas Wree approved the final manuscript.
Name: Martin Witt, MD, PhD.
Contribution: This author helped conduct the study, analyze the data, and write the manuscript.
Attestation: Martin Witt has seen the original study data and approved the final manuscript.
Name: Thomas Mittlmeier, MD, PhD.
Contribution: This author helped design the study and write the manuscript.
Attestation: Thomas Mittlmeier has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
Name: Brigitte Vollmar, MD, PhD.
Contribution: This author helped design the study, analyze the data, and write the manuscript.
Attestation: Brigitte Vollmar has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
This manuscript was handled by: Quinn H. Hogan, MD.
1. Torrance N, Smith BH, Bennett MI, Lee AJ. The epidemiology of chronic pain of predominantly neuropathic origin: results from a general population survey. J Pain. 2006;7:281–9
2. StantonHicks M, Janig W, Hassenbusch S, Haddox JD, Boas R, Wilson P. Reflex sympathetic dystrophy: changing concepts and taxonomy. Pain. 1995;63:127–33
3. Attal N, Cruccu G, Haanpää M, Hansson P, Jensen TS, Nurmikko T, Sampaio C, Sindrup S, Wiffen PEFNS Task Force.. EFNS guidelines on pharmacological treatment of neuropathic pain. Eur J Neurol. 2006;13:1153–69
4. Bennett GJ, Xie YK. A peripheral mononeuropathy in rat that produces disorders of pain sensation like those seen in man. Pain. 1988;33:87–107
5. Gradl G, Gaida S, Gierer P, Mittlmeier T, Vollmar B. In vivo evidence for apoptosis, but not inflammation in the hindlimb muscle of neuropathic rats. Pain. 2004;112:121–30
6. Gradl G, Gaida S, Finke B, Gierer P, Mittlmeier T, Vollmar B. Exaggeration of tissue trauma induces signs and symptoms of acute CRPS I, however displays distinct differences to experimental CRPS II. Neurosci Lett. 2006;402:267–72
7. Yaoita H, Ogawa K, Maehara K, Maruyama Y. Attenuation of ischemia/reperfusion injury in rats by a caspase inhibitor. Circulation. 1998;97:276–81
8. Eipel C, Bordel R, Nickels RM, Menger MD, Vollmar B. Impact of leukocytes and platelets in mediating hepatocyte apoptosis in a rat model of systemic endotoxemia. Am J Physiol Gastrointest Liver Physiol. 2004;286:769–76
9. Attal N, Jazat F, Kayser V, Guilbaud G. Further evidence for ‘pain-related’ behaviours in a model of unilateral peripheral mononeuropathy. Pain. 1990;41:235–51
10. Chaplan SR, Bach FW, Pogrel JW, Chung JM, Yaksch TL. Quantitative assessment of tactile allodynia in the rat paw. J Neurosci Methods. 1994;53:55–63
11. Tyml K, Budreau CH. A new preparation of the rat extensor digitorum longus muscle for intravital investigation of the microcirculation. Int J Microcirc Clin Exp. 1991;10:335–43
12. Schaser KD, Vollmar B, Menger MD, Schewior L, Kroppenstedt SN, Raschke M, Lubbe AS, Haas NP, Mittlmeier T. In vivo analysis of microcirculation following closed soft-tissue injury. J Orthop Res. 1999;17:678–85
13. Gradl G, Gaida S, Finke F, Lindenblatt N, Gierer P, Menger MD, Mittlmeier T, Vollmar B. Supernatant of traumatized muscle induces inflammation and pain, but not microcirculatory perfusion failure and apoptotic cell death. Shock. 2005;24:219–25
14. Munglani R, Harrison SM, Smith GD, Bountra C, Birch PJ, Elliot PJ, Hunt SP. Neuropeptide changes persist in spinal cord despite resolving hyperalgesia in a rat model of mononeuropathy. Brain Res. 1996;743:102–8
15. Azkue JJ, Zimmermann M, Hsieh TF, Herdegen T. Peripheral nerve insult induces NMDA receptor-mediated, delayed degeneration in spinal neurons. Eur J Neurosci. 1998;10:2204–6
16. Maione S, Siniscalco D, Galderisi U, de Novellis V, Uliano R, Di Bernardo G, Berrino L, Cascino A, Rossi F. Apoptotic genes expression in the lumbar dorsal horn in a model neuropathic pain in rat. Neuroreport. 2002;13:101–6
17. Scholz J, Broom DC, Youn D, Mills CD, Kohno T, Suter MR, Moore KA, Decosterd I, Coggeshall RE, Woolf CJ. Blocking caspase activity prevents transsynaptic neuronal apoptosis and the loss of inhibition in lamina II of the dorsal horn after peripheral nerve injury. J Neurosci. 2005;25:7317–23
18. Polgar E, Gray S, Riddell JS, Todd AJ. Lack of evidence for significant neuronal loss in laminae IIII of the spinal dorsal horn of the rat in the chronic constriction injury model. Pain. 2004;111:144–50
19. Polgar E, Hughes DI, Arham AZ, Todd AJ. Loss of neurons from laminas IIII of the spinal dorsal horn is not required for development of tactile allodynia in the spared nerve injury model of neuropathic pain. J Neurosci. 2005;25:6658–66
20. FelderhoffMueser U, Sifringer M, Pesditschek S, Kuckuck H, Moysich A, Bittigau P, Ikonomidou C. Pathways leading to apoptotic neurodegeneration following trauma to the developing rat brain. Neurobiol Dis. 2002;11:231–45
21. Park S, Yamaguchi M, Zhou C, Calvert JW, Tang J, Zhang JH. Neurovascular protection reduces early brain injury after subarachnoid hemorrhage. Stroke. 2004;35:2412–7
22. GarciaCalvo M, Peterson EP, Leitling B, Ruel R, Nicholson DW, Thornberry NA. Inhibition of human caspases by peptide-based and macromolecular inhibitors. J Biol Chem. 1998;273:32608–13
23. Abram SE, Yi J, Fuchs A, Hogan QH. Permeability of injured and intact nerves and dorsal root ganglia. Anesthesiology. 2006;105:146–53
24. Schaeffer V, Meyer L, PatteMensah C, Eckert A, MensahNyagan AG. Sciatic nerve injury induces apoptosis of dorsal root ganglion satellite glial cells and selectively modifies neurosteroidogenesis in sensory neurons. Glia. 2010;58:169–80
25. Richardson PM, Thomas PK. Percussive injury to peripheral nerve in rats. J Neurosurg. 1979;51:178–87
26. Stratos I, Rotter R, Eipel C, Mittlmeier T, Vollmar B. Granulocyte-colony stimulating factor enhances muscle proliferation and strength following skeletal muscle injury in rats. J Appl Physiol. 2007;103:1857–63
27. Tews DS, Goebel HH, Meinck HM. DNAfragmentation and apoptosisrelated proteins of muscle cells in motor neuron disorders. Acta Neurol Scand. 1997;96:380–6
28. Yoshimura K, Harii K. A regenerative change during muscle adaptation to denervation in rats. J Surg Res. 1999;81:139–46
29. Plant PJ, Bain JR, Correa JE, Woo M, Batt J. Absence of caspase-3 protects against denervation-induced skeletal muscle atrophy. J Appl Physiol. 2009;107:224–34
30. Roos A, Xu W, Castellano G, Nauta AJ, Garred P, Daha MR, van Kooten C. Mini review: a pivotal role for innate immunity in the clearance of apoptotic cells. Eur J Immunol. 2004;34:921–9