Mol Neurobiol DOI 10.1007/s12035-014-8706-9

Peripheral Nerve Injury Modulates Neurotrophin Signaling in the Peripheral and Central Nervous System Mette Richner & Maj Ulrichsen & Siri Lander Elmegaard & Ruthe Dieu & Lone Tjener Pallesen & Christian Bjerggaard Vaegter

Received: 23 December 2013 / Accepted: 1 April 2014 # Springer Science+Business Media New York 2014

Abstract Peripheral nerve injury disrupts the normal functions of sensory and motor neurons by damaging the integrity of axons and Schwann cells. In contrast to the central nervous system, the peripheral nervous system possesses a considerable capacity for regrowth, but regeneration is far from complete and functional recovery rarely returns to pre-injury levels. During development, the peripheral nervous system strongly depends upon trophic stimulation for neuronal differentiation, growth and maturation. The perhaps most important group of trophic substances in this context is the neurotrophins (NGF, BDNF, NT-3 and NT-4/5), which signal in a complex spatial and timely manner via the two structurally unrelated p75NTR and tropomyosin receptor kinase (TrkA, Trk-B and Trk-C) receptors. Damage to the adult peripheral nerves induces cellular mechanisms resembling those active during development, resulting in a rapid and robust increase in the synthesis of neurotrophins in neurons and Schwann cells, guiding and supporting regeneration. Furthermore, the injury induces neurotrophin-mediated changes in the dorsal root ganglia and in the spinal cord, which affect the modulation of afferent sensory signaling and eventually may contribute to the development of neuropathic pain. The focus of this review is on the expression patterns of neurotrophins and their receptors in neurons and glial cells of the peripheral nervous system and the spinal cord. Furthermore, injury-induced changes of expression patterns and the functional consequences in relation to axonal growth and remyelination as well as to neuropathic pain development will be reviewed. M. Richner : M. Ulrichsen : S. L. Elmegaard : R. Dieu : L. T. Pallesen : C. B. Vaegter (*) Danish Research Institute of Translational Neuroscience DANDRITE, Nordic EMBL Partnership, and Lundbeck Foundation Research Center MIND, Department of Biomedicine, Aarhus University, Ole Worms Allé 3, 8000 Aarhus C, Denmark e-mail: [email protected]

Keywords Neurotrophins . Peripheral nerve injury . Regeneration . Neuropathic pain

Introduction — Nerve Injury, Neurotrophins and Their Receptors Peripheral neuropathy is a condition where damage resulting from mechanical or pathological mechanisms is inflicted on nerves within the peripheral nervous system (PNS). Physical injury is the most common cause and may result in nerves being partially or completely severed, crushed, compressed or stretched. In contrast to the central nervous system (CNS), the PNS possesses a unique ability to regenerate, recognized since ancient times and initially described by surgeons and later methodologically investigated by modern neurobiologists including Camillo Golgi and Santiago Ramon y Cajal [1]. However, despite the considerable capacity for regrowth upon injury, PNS regeneration is far from complete and functional recovery rarely returns to pre-injury levels. Several biological factors influence the communication between peripheral nerves and tissues following injury including a variety of cytokines and growth factors. Similar conditions can be observed during the development of the nervous system where the PNS strongly depends on trophic stimulation for survival, differentiation, sprouting and maturation. As damage to the adult PNS induces cellular mechanisms that control neuronal differentiation as well as growth and connectivity during development, studies of neurotrophic factors are considered to represent one of the most promising areas of research aimed at finding new, more effective treatments for peripheral neuropathies [2]. Various families of neurotrophic factors are being studied; however, signaling by the neurotrophins is probably receiving most attention. The family of neurotrophins consists of nerve growth factor (NGF), brain derived neurotrophic factor

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(BDNF), neurotrophin-3 (NT-3) and neurotrophin-4/5 (NT-4/ 5), which can bind to two structurally unrelated receptors: the neurotrophin receptor p75NTR and the tropomyosin receptor kinases (TrkA, -B, and -C). Binding of neurotrophins to the Trk receptors is selective with NGF binding to TrkA, BDNF and NT-4/5 to TrkB and NT-3 to TrkC [3]. However, many neurons co-express Trk receptors and p75NTR, which together strengthen the affinity and specificity of the neurotrophins for the Trk receptors probably through the formation of a receptor complex [4–8]. Neurotrophins are synthesized as precursor proteins (proneurotrophins), which can be cleaved by furin or other proconvertases to produce the mature proteins of about 13– 15 kDa in size. Both pro-neurotrophins and neurotrophins are released as stable, non-covalent homodimers and are believed to elicit different signaling properties. The neurotrophins bind to their respective Trk receptors to mediate survival and differentiation via extracellular signal-regulated kinase (ERK), phosphatidylinositol 3-kinase (PI3K) and phospholipase C-γ (PLC-γ) pathways. Neurotrophins can also provide survival signaling via p75NTR and the NF-κB pathway or affect cytoskeletal organization and neurite outgrowth via downstream RhoA kinase activity (reviewed in [3, 9]). Yet, p75NTR is better recognized for its ability to induce cell death signaling via binding of various intracellular adaptor proteins, ultimately activating the c-Jun N-terminal kinase (JNK) pathway and subsequently p53 leading to apoptosis. For many years, p75NTR-mediated apoptosis was considered to be initiated by mature neurotrophins, whereas pro-

Spinal cord dorsal horn

neurotrophins were thought to be merely inactive precursors. Increasing evidence has, however, identified important biological functions of pro-neurotrophins, one of which is the ability to induce cell death by binding to the receptor sortilin (via the pro-domain) and p75NTR (via the mature domain), thereby creating a tertiary complex that results in activation of neuronal apoptosis [10–14]. To add to the complexity, it has recently been reported that TrkA and TrkC, but not TrkB, behave as dependence receptors during development, i.e., they may also trigger neuronal death on their own in absence of activation by their corresponding neurotrophins [15]. Hence, it is apparent that the consequences of neurotrophin signaling depend on the interplay between neurotrophins/proneurotrophins and their receptors in a very complex spatial and timely manner. Injury to peripheral nerves induces dramatic changes in the expression levels of neurotrophins and their receptors in all cellular components of the PNS (neurons, satellite glial cells [SGCs] and Schwann cells) as well as in some CNS cells of the spinal cord (motor neurons and microglia). In particular, altered neurotrophin signaling is believed to be essential for the numerous complex processes underlying peripheral nerve regeneration as well as in neuropathic pain development — two important biological responses to peripheral nerve injury. In the following sections, the expression of neurotrophins and their receptors as well as injury-induced changes in expression and signaling are described in three major structural areas of the nervous system: the dorsal root ganglia (DRG), the peripheral nerves and the spinal cord dorsal horn (Fig. 1).

Dorsal rootlets

Spinal cord ventral horn

Dorsal root

Peripheral nerve injury

Ventral rootlets Dorsal root ganglion

Ventral root

Pia mater

Arachnoid mater

Peripheral nerve

Dura mater

Fig. 1 The spinal cord and the peripheral nervous system (PNS). Peripheral nerve injury induces various changes in the PNS (peripheral nerve fibers and DRGs) but also in the central nervous system (CNS) (spinal

cord). Changes include altered expression levels of neurotrophins and their receptors with concomitant altered signaling processes in both the PNS and CNS

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each DRG fuses with the corresponding spinal ventral root at the same level of each vertebra to form a spinal nerve, which intermingles distally with other spinal nerves to form the peripheral nerves.

Dorsal Root Ganglia Anatomy at a Glance DRG neurons are primary afferent neurons responsible for transmitting sensory information from the periphery to the CNS. The DRGs are part of the PNS and are located in the neural foramen of the vertebrae, in close proximity to the CNS. The connective tissues surrounding the DRGs are continuous with the two outermost spinal meninges, the dura mater and the arachnoid mater [16]. For each cervical (C), thoracic (T), lumbar (L) and sacral (S) vertebra the DRGs are designated C, T, L or S and the corresponding number of the given vertebra in a rostro-caudal order. Thus, e.g., the DRGs of the fourth lumbar vertebra are named L4, while the second thoracic DRGs are named T2. The sciatic nerve, which is often the object in rodent nerve injury models [17], has its main part of neuronal somas in L3–L5 DRGs in mice and correspondingly in L4–L6 DRGs in rats [18]. Primary afferents have a unique pseudo-unipolar structure meaning that the soma of each neuron situated in a DRG projects one axon that bifurcates into a peripheral and a central branch. No other neuronal processes are formed by the soma (Fig. 2a). The peripheral branch extends distally towards the skin or viscera to receive sensory stimulation, whereas the central branch transmits these signals via synaptic terminals to the secondary sensory neurons of the spinal cord dorsal horns for central processing. This pseudounipolarity allows direct signal transmission from the site of stimulation in the periphery to the CNS, bypassing the neuronal soma. Even though afferent signals initiated in the periphery can bypass the soma, signal invasion to the soma provides necessary information for protein synthesis and for maintaining optimal levels of receptors and ion channels at the nerve terminals [19]. The distal root of

a

Glial Cells Glial cells are numerous in the DRGs and count Schwann cells, which wrap around the neuronal processes, and SGCs, which enclose each neuronal soma. Several SGCs, interconnected by gap junctions, together form a sheath around one neuronal soma (Fig. 2a), leaving a gap of just 20 nm between the SCGs and the neuron — the width of a synapse. This functional unit of soma and SGCs is enclosed by a layer of connective tissue [20–22]. The non-neuronal face of the slimly flattened SGCs has a basement membrane, which together with the endothelial basement membrane constitutes a blood– neuron barrier [23, 24]. Through expression of a number of receptors and channels, SGCs are believed to stabilize the neuronal microenvironment, supporting normal neuron function and providing electrical insulation (SGC protein expression reviewed in [24]). This intimate and isolated structural arrangement between SGCs and soma strongly implies that soma-SGC communication is a major determinant of neuronal activity, and that understanding injury-induced changes in this communication is important for understanding the neuronal changes and activity following peripheral nerve injury. SGCs may be compared to astrocytes of the CNS, as they have similar functions and express several similar biochemical markers, such as glutamine synthetase (GS) and, upon injury, glial fibrillary acidic protein (GFAP). However, similarity as such is confined to function, since SGCs originate from neural crest cells of the embryo, while astrocytes are derived from the neural tube [25].

b

Neuron cell body

Myelinated axons

Schwann cells

Node of Ranvier

Neuron cell body

Satellite glial cells

Satellite glial cells

Fig. 2 Dorsal root ganglia (DRG) neurons and encapsulating satellite glial cells (SGCs). a Each neuron is closely surrounded by multiple SCGs forming a neuron–SGC unit. Each pseudounipolar neuron projects one

axon that bifurcates into a branch that extends to the peripheral target tissue and a branch that extends to the spinal cord. b DRG cross-section illustrating primary sensory neurons surrounded by a single thin layer of SGCs

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Characterization of Primary Afferent Neuron Types and Modalities A widely used system to characterize the primary afferents is according to axon size, degree of myelination and, hence, conduction velocity. The large, heavily myelinated Aα- and Aβ-fibers transmit signals of proprioception and innocuous mechanoreception, respectively. Small myelinated fibers, designated Aδ-fibers, mostly convey fast sharp pain and temperature sensation, although some Aδ-fibers convey lowthreshold mechanosensation. Small unmyelinated fibers, designated C-fibers, convey sensation of temperature, slow dull pain and itch [26–29]. DRG neurons of different modality project axons to certain laminae of the spinal cord; nociceptive afferents terminate in dorsal horn laminae I and II, whereas mechanosensitive neurons mainly terminate in laminae III and IV. Proprioceptive neurons terminate in the ventral spinal cord where they synapse with motor neurons [30]. Neuronal somas in the DRG are divided into two groups according to morphology: large, light neurons with coarsely granular cytoplasm and small, dark neurons with a denser, more uniform cytoplasm [31–34]. The large, light neurons are referred to as type A neurons and the small, dark neurons as type B neurons [35, 36]. In rats, approximately 40 % of DRG neurons are type A neurons and approximately 60 % are type B neurons, while a small percentage of medium-sized neurons fall out of these categories [37–39]. The type A and B neurons can further be subdivided according to biochemical characteristics such as enzyme and neurotransmitter production [40, 41], which will not be discussed in this review. By combining intracellular recording and dye-injection, a direct correlation of soma size with peripheral nerve conduction velocity is possible [42]. Peripheral unmyelinated C-fibers originate from small, dark type B DRG neurons whereas peripheral Aα- and Aβ- fibers originate from large, light type A DRG neurons. Thus, type A neurons are largely proprio- and mechanosensory giving rise to A-fibers, while type B neurons are largely nociceptive and give rise to C-fibers. The average soma size of peripheral Aδ-fibers is between the average size of large, light and small, dark neuron populations, providing evidence of them being the out-of-category medium-sized neurons [42]. Trk Receptors in DRG Neurons and SGCs Distinct subpopulations of adult rat DRG neurons express TrkA, TrkB or TrkC mRNA [4, 5], although some report extendedly overlapping populations [43]. TrkA-positive neurons constitute about 40 % of DRG neurons [4, 38, 44–47] and are mostly small diameter type B neurons [38, 46, 48], as shown in Fig. 3, which depicts the DRG neuron populations

Aδ TrkC TrkA type A neurons TrkB

type B neurons

Ret Fig. 3 Expression profile of tropomyosin receptor kinase (Trk) receptors in DRG neurons. Large, light type A neurons constitute approximately 40 % of the DRG neurons (light gray area). Two distinct populations of type A neurons express TrkB and TrkC, respectively, and some TrkCpositive neurons also express TrkA. The small dark type B neurons constitute approximately 60 % of the DRG neurons (dark gray area). TrkA and glial cell line derived neurotrophic factor (GDNF) receptor Ret are expressed by distinct type B neuron populations, however, a distinct expression overlap is observed. Ret is also expressed by a small part of type A neurons. A small percentage of DRG neurons fit neither type A nor type B profiles and these neurons are likely the somas of Aδ-fibers that, at least partly, express TrkA

with respect to Trk receptor expression. A nociceptive function of the TrkA-positive population is supported by studies showing spinal cord TrkA immunoreactivity being restricted to laminae I and II [49] as well as by the abnormal pain phenotypes of NGF and TrkA knockout animals; Both homozygous NGF and TrkA knockout mice, which only survive for a few weeks postnatally, fail to respond to noxious mechanical and thermal stimuli. These mice suffer extensive loss of small and medium peptidergic neurons, i.e., type B neurons in DRGs [50, 51]. Heterozygous NGF knockout mice display a mild, albeit statistically significant phenotype, displaying a prolonged latency in the heat-sensitive tail flick test compared to wild-type mice [51]. The Aδ-fiber subpopulation of DRG neurons is suggested to be at least partially TrkA-positive as indicated by the significantly increased number of these mechanosensory Aδ-fibers in NGF-overexpressing mice [52]. Approximately 10–15 % of TrkA-positive neurons overlap with the TrkC-positive population [4]. The TrkB-positive neurons constitute 5-26 % of DRG neurons and display varied soma diameters [4, 46, 53] and are mainly belonging to the large, light type A neuron population [54]. Overall, 10–21 % of DRG neurons, almost exclusively of large size, are TrkCpositive [4, 46, 53] and are categorized as muscle afferents, i.e., proprioceptive [48, 49, 55]. P75NTR is co-expressed in most Trk receptor-positive neurons and does not seem to be expressed independently of the Trk receptors [4, 5]. A significant part of the DRG neurons does not express Trk receptors at all [5, 48]; The glial cell line derived neurotrophic factor (GDNF) receptor Ret is expressed in one third of DRG

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neurons, constituting about half of the small, dark type B neurons [47, 56], with only a limited overlap of Ret-positive and TrkA-positive populations (reported to 13 % of the TrkApositive neurons [38]). The Ret-positive neurons project axons to the inner lamina II (IIi) of the spinal cord [57], which is specifically linked to innocuous mechanoreception [58]. However, changes of innocuous input processing in lamina IIi dorsal horn neurons have been suggested to be critical for the establishment of post-injury neuropathic pain [57, 59], and some Ret-positive neurons are designated a nociceptive modality [29]. SGCs have also been reported to express neurotrophins and their receptors. Thus, TrkA and p75NTR are found in the SGC cytoplasm and in the SGC–neuron border [60, 61], and expression of TrkB [4], more specifically truncated TrkB without the cytosolic kinase domain, has also been described [48, 53]. Furthermore, TrkC mRNA and immunoreactivity in some SGCs surrounding large neurons has also been reported [61]. Finally, NT-3, NGF and BDNF have been observed in SGCs both at mRNA and protein levels [62, 63]. Injury-Induced Changes in DRGs Profound changes can be observed in the soma of DRG neurons and in spinal cord ventral horn motor neurons upon peripheral nerve injury, ranging from altered protein expression, structural changes and even cell death depending on the degree of injury and the neuronal subtype. In general, damage to the adult PNS induces cellular mechanisms that control neuronal differentiation, growth and connectivity during development. A series of morphologic changes known as chromatolysis can be observed as early as a few hours after injury, entailing cell body and nucleolar swelling as well as nuclear eccentricity. The purpose of these changes is to initiate an appropriate response to traumatic stress, switching the neurons from the steady-state production of neurotransmitters and housekeeping proteins to a “regrowth mode”. This involves production of components for cytoskeletal and axonal reconstruction and growth-related proteins including neuropeptides, neurotrophic factors and their receptors. Also, depending on the type of injury and the distance to the soma, a considerable fraction of primary afferent neurons contributing to the injured nerve will die. Whether this is a consequence of apoptotic signaling molecules, e.g., pro-neurotrophins, lack of trophic support from target organs or a more general cellular stress resulting from the injury is yet unclear (reviewed in [43, 64–66]). Functionally, neuronal somas in DRGs often develop lowered firing thresholds or persistent spontaneous firing following injury, which is assumed to intensify peripheral signals reaching the spinal cord and, thus, may contribute to the phenomenon of sensitization and neuropathic pain [67].

The mechanisms underlying spontaneous firing are not clarified but are proposed to involve an increase in neuronal ATP sensitivity, sensitization of neighboring uninjured neurons and altered expression of transmitters. Supporting these proposals, altered communication between DRG neurons and SGCs is likely to be involved. Of the seven identified purinergic ionotropic P2X receptors (P2X1–P2X7), P2X3 is exclusively expressed by neurons and P2X7 exclusively by SGCs in sensory ganglia, which has greatly facilitated the analysis of soma-SGC communication. Thus, it is now generally believed that the neuronal soma can communicate with surrounding SGCs through bidirectional release of ATP (reviewed in [20]). In accordance with this, increased neuronal expression levels and membrane trafficking of P2X3, as well as upregulation of P2X7 expression in SGCs, have been observed following nerve injury and following NGF treatment and both are believed to be important mechanisms underlying injury-induced sensitization of DRG neurons [20, 68]. SGCs can further communicate with each other via gap junctions, which under normal physiological conditions are only observed between adjacent SGCs within a structural unit, i.e., SGCs enveloping a single neuronal soma. Following nerve injury, however, electron microscopy and injection of the fluorescent dye lucifer yellow into individual SGCs have demonstrated the formation of new gap junctions between adjacent SGCs within individual envelopes as well as gap junction bridges between SGCs in neighboring envelopes. As gap junction blockers have been found to selectively reduce spontaneous activity of DRG neurons following nerve injury, this strongly supports the importance of neuron–SGC communication on neuronal function following nerve injury [21, 69, 70]. Neurotrophin Receptors in DRGs In agreement with the concept of an injury-induced change of DRG neurons to a “regrowth mode”, numerous reports over the past couple of decades have documented dramatic changes in the expression of neurotrophins and neurotrophin receptors in sensory ganglia neurons and SGCs following injury. However, as will be described in the following sections, their responses are quite complex and reflect the type of injury paradigm and neuronal subtype but also whether the neurons suffer direct injury to their distal axon or if the distal axons are intact but affected by neighboring injured axons. Trk receptors have been found to be selectively expressed in subpopulations of sensory ganglia neurons, and axotomy has generally been reported to result in downregulation of all Trk receptors on both mRNA and protein levels in adult rodents [71, 72]. The downregulation has been found to be more pronounced in younger rats compared to older rats, which might reflect the generally higher expression levels of Trk receptors observed in younger adult rodents [71]. While expression of all Trk receptors is generally reported to be

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downregulated in DRG neurons following injury, the situation is found to be more complex with the neurotrophins. BDNF in DRGs Several reports have demonstrated that nerve injuries greatly increase BDNF synthesis in primary afferents [56, 73–75], but it is also evident that different types of peripheral nerve injuries induce differential BDNF responses. BDNF does not appear to have an auto- or paracrine role within the DRG but is believed to be a trophic messenger that is anterogradely transported to the central terminals of primary afferents in the spinal cord dorsal horn where it may be released upon peripheral nociceptive activity inducing signal modulation and consequently pain symptoms [54, 73, 76–79]. Despite the neuronal upregulation of BDNF, only a subgroup of DRG neurons expresses BDNF prior to or as a result of nerve injury. In rats, BDNF mRNA and protein is found mainly, but not exclusively, in the subpopulation of DRG neurons that expresses the NGF receptor TrkA. TrkAnegative neurons that express BDNF include some large TrkC-positive neurons, a very limited fraction of mediumsized TrkB-positive neurons as well as a fraction of small nociceptive neurons [54]. Sciatic nerve axotomy and nerve crush have both been shown to induce a significant increase in the number of medium and large BDNF-positive neurons. However, whereas nerve crush has been found to result in both an increased number of small BDNF-positive neurons and increased BDNF intensity, nerve axotomy was found to reduce BDNF intensity in small neurons without affecting the number of BDNF-positive neurons [74]. Similarly, in a comparative study, the chronic constriction injury model on rat sciatic nerve has been observed to induce a significant increase in the percentage of small, medium and large BDNFpositive neurons in the ipsilateral L4 and L5 DRGs. On the other hand, L5 spinal nerve ligation (L5-SNL), which selectively allows injury of the L5 DRG, was shown to result in an increased number and intensity of BDNF-positive medium and large neurons in the ipsilateral L4 and L5 DRGs, but the number and intensity of small BDNF-positive neurons were found to decrease in the L5 DRG (injured), while it was found to be significantly increased in the L4 (uninjured) DRG [75, 80]. What might be the explanation for these differences? Axotomy or application of a firm ligation counteracts the communication of the proximal nerve stump with both the distal stump and the peripheral target. In contrast, both nerve crush and chronic constriction injury models allow continued contact and communication between the local neuroma and cell bodies of the injured neurons as well as with the peripheral part of the nerve and peripheral target tissue. Finally, L5-SNL induces Wallerian degeneration in the distal axons of the affected DRG neurons, which influences intact neighboring

axons and, thus, the neuronal soma in the uninjured L4 DRG. It is, therefore, likely that these aspects are causing the observed differential regulation of BDNF gene expression after cut/ligation versus crush/constriction injuries. NGF in DRGs The observation that BDNF is mainly expressed and upregulated in TrkA-positive neurons following nerve injury strongly implies the involvement of NGF as a signaling molecule in these events. This is supported by experiments in which stimulation with exogenous NGF has been found to increase BDNF mRNA and protein levels within TrkA-positive neurons, resulting in BDNF expressing in 90 % of TrkA-positive DRG neurons. As exogenous NGF has been shown not to increase BDNF expression in non-TrkA cells, it indicates that alternative signaling molecules are mediating BDNF regulation in these neurons following injury [54]. But what is the source of NGF in the DRGs? In contrast to BDNF, NGF appears to be downregulated in DRG neurons following injury [81]. However, a variety of peripheral inflammatory conditions result in locally increased NGF at both mRNA and protein levels and NGF is retrogradely transported from the injured nerve/periphery. In support of this, NGF-quenching antibodies or TrkA-Fc fusion proteins can prevent the translation of peripheral tissue inflammation into increased DRG responsiveness towards NGF [82]. The differential DRG response between cut/ligation and crush/constriction models as well as the disparity between injured and uninjured DRG responses in the L5-SNL model indicates that at least some NGF is derived from the periphery distal to the site of injury and retrogradely transported back to the DRGs via intact axons. The simplest model would be that NGF binds TrkA in the periphery to induce signaling in the TrkA-positive neurons both in the distal nerve as well as in the soma upon retrograde transport of the activated signaling complex. A competing model for NGF availability in the entire DRG might be that retrogradely transported NGF is re-released from the soma to act on neighboring neurons in the DRGs. However, such re-release is still to be demonstrated, and the tight envelope of SGCs surrounding the individual neuronal soma further argues against such a mechanism. Other evidence suggests that the periphery is not the only source of the observed NGF increase in the DRG. Following L5-SNL, a linear increase in NGF protein has been observed in the ipsilateral L4 DRG, reaching a plateau after 2 weeks. However, delayed increase in NGF mRNA in the injured L5 DRG, but not in the uninjured L4, has also been observed [75]. Combined with the observation that injured DRG neurons downregulate NGF mRNA, this suggests a non-neuronal source of NGF within the DRG upon direct injury to the distal axons. Indeed, several observations demonstrate that SGCs, but not neurons, increase NGF and NT-3 mRNA following

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injury [63, 81, 83] and, intriguingly, administration of gap junction inhibitors during injury was found to significantly decrease the number of NGF-positive SGCs [84, 85]. In general, the SGCs respond rapidly when injury is inflicted on the neuron, which they encapsulate. Dynamic neuron–SGC communication by P2X receptors, NMDA receptors and glutamate transporters in SGCs increases SGC– SGC communication via gap junctions and initiates a general increase in proliferation and cytokine communication [9, 20, 64, 86]. This SGC response suggests that SGC involvement in post-injury events, such as regeneration and the initiation and/ or maintenance of neuropathic pain, is a highly rational assumption. In accordance with this, knockdown of the gap junction protein Connexin43 as well as the use of pharmacological gap junction blockers have been demonstrated to decrease pain behavior in rodents [3, 87–89]. The observation of SGCs also being dynamically involved in neurotrophin signaling is being increasingly recognized. Presence of TrkA, TrkB, p75NTR, NGF and BDNF in SGCs under normal physiological conditions has been described, and in situ hybridization and immunohistochemical analysis have demonstrated upregulation of NGF and NT-3 starting as early as 48 h after nerve injury and lasting at least 2 months [53, 60, 63, 69, 90]. Sympathetic Sprouting into Injured DRGs Although the direct effects of SGC-derived NGF on DRG neurons are still to be demonstrated, other observations indicate involvement of SGCs in neurotrophin signaling. Sprouting of sympathetic axons into sensory ganglia following nerve injury is believed to underlie the phenomenon of sympathetically maintained pain, where increased pain is triggered when sympathetic activity is induced. Following nerve injury, sprouts of sympathetic terminals originating from the vasculature form synaptic terminals or “baskets” around selected large DRG sensory neurons. Sympathetic nerves express and respond to neurotrophins [11, 15, 91, 92], and the observed sympathetic sprouting clearly depends on NGF as antibodies against NGF or NT-3, delivered by an osmotic mini-pump to the DRG via the transected nerve, was found to reduce sprouting significantly [81, 93]. It has been reported that uninjured and even contralateral DRGs to some extent can be affected by sympathetic sprouting [16, 81], although this might reflect that any surgical procedure per se, to some extent, may initiates local tissue inflammation and, thus, may induce systemic responsiveness to released NGF. As axotomy decreases the availability of NGF from the peripheral nerve/target, the source of NGF within the DRG is believed to be the SGCs; a model which is consistent with the observed injury-induced increase of NGF in these cells. In general, the sympathetic sprouting around the axotomized neurons is associated with p75NTR-positive SGCs [20, 21,

63]. Based on the L5-SNL model, nerve injury results in reduced p75NTR expression in injured (L5) neurons, whereas the expression increases in uninjured but affected L4 neurons. However, whereas injured neurons decrease p75NTR expression, the surrounding SGCs experience a dramatic upregulation of p75NTR levels in response to injury. Increased expression of p75NTR is primarily found in SGCs surrounding large neurons, and co-localization studies further demonstrate that sympathetic sprouting mainly associates with the p75NTRpositive SGCs surrounding these large neurons. Furthermore, other findings show that sympathetic fibers wrap up around the SGCs rather than around the neurons, and that p75NTR knockout mice display impaired sympathetic sprouting into the DRGs [23, 24, 71, 72, 81, 93, 94]. Thus, increased expression of NGF and p75NTR by SGCs potentially provides an explanation for the abnormal growth of sympathetic fibers in DRGs after peripheral nerve injury. However, whether increased NGF expression by SGCs is restricted to the p75NTR-positive SGCs surrounding large injured neurons and how SGC p75NTR is involved in the supposed SGCderived NGF attraction of sympathetic sprouts is still to be explained. Only half of the p75NTR-positive neuron–SGC units attract sympathetic sprouts, and p75NTR-positive SGCs surrounding small diameter neurons do not attract sympathetic sprouts [24, 63], suggesting that sympathetic sprouting mechanisms include, but extend beyond, SGC-related increases in NGF and p75NTR expression. Neuronal Apoptosis in DRGs One of the effects of nerve injury is a progressive loss of neurons in the affected DRGs. The consequences of nerve injury on DRG neuron numbers have been studied in both mice and rats with up to 50 % total neuron loss observed, albeit with a rather high variation, likely depending on the injury model. The neuron loss predominantly affects the type B neuron population, whereas fewer type A neurons are lost [95–102]. As many type B neurons are TrkA positive, it has been proposed that these neurons die as a consequence of reduced NGF accessibility from peripheral targets. Attempts to prevent type B neuron loss by application of local or systemic NGF has, however, been conflicting, with observed neuronal rescue in some studies [26–29, 95], but without effect in others [30, 101]. Furthermore, crush injury, which is believed to allow retrograde transport and signaling by NGF on TrkA positive neurons (see section “NGF in DRGs”), still results in a significant loss of type B neurons [101]. It has also been suggested that p75NTR is involved in the injury-induced loss of neurons [96, 101, 103, 104]. This is based on the observations that no type B neurons are lost following axotomy in absence of p75NTR utilizing either p75NTR-deficient mice or by inhibiting p75NTR mRNA translation with antisense oligonucleotides. However, the fact that p75NTR-

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deficient mice develop serious degeneration of sensory ganglia with approximately 50 % neuron loss in the adult animal [101] might make this an inappropriate animal model to study further injury-induced neuron loss ascribed to p75NTR deficiency. A curious aspect is the finding that p75NTR is expressed in both type A and B neurons [40, 41, 105], but only type B neurons are seriously affected by injury-induced apoptosis. Furthermore, SGCs dramatically increase p75NTR expression following injury to their associated neurons, but whereas p75NTR is linked to neuronal death, SGCs proliferate following injury. It has been postulated that pro-neurotrophin-induced cell death could be the underlying cause of neuronal apoptosis, as sortilin and p75NTR are both widely expressed and are found with a high degree of co-expression in several DRG neurons [43, 105, 106]. ProNGF is also present in the DRGs [105], and the increase in neuronal BDNF synthesis following injury may provide a rich source of proBDNF in the DRGs. Only small-sized neurons co-expressing sortilin and p75NTR appear to be vulnerable to injury-induced neuronal death [38, 46, 48, 105]. Thus, the answer to the question why type B neurons are more vulnerable is currently not clear but might involve (pro)BDNF upregulation by small (TrkA positive) sortilin/ p75NTR-positive type B DRG neurons following nerve injury. Neurotrophin Signaling in DRGs Following Injury Accompanying the dynamic expression of neurotrophins and their receptors following nerve injury is a multiplicity of intracellular and cell–cell signaling in the DRGs. A general and widely used marker of neuronal injury is the induction of activating transcription factor-3 (ATF3), a member of the ATF/CREB transcription factor superfamily. After sciatic nerve axotomy in the rat, 82 % of L4 DRG neurons have been found to be ATF3-positive. It has been observed that intrathecal delivery of NGF for 2 weeks following nerve injury reduced ATF3 expression to 35 % of DRG neurons, preventing expression in a population of mainly small- to medium-sized neurons [55, 107]. This suggests that ATF3 expression may normally be induced by loss of target-derived neurotrophins. A general phenomenon following nerve injury is also the activation of the mitogen-activated protein kinase (MAPK) family of signaling molecules, which transduce various extracellular stimuli to intracellular mitogenic and differentiation signals. The MAPK family includes ERK1/2, p38, JNK and ERK5. Whereas ERK1/2 is involved in cellular growth and differentiation, p38 and JNK are involved in cellular stress such as apoptosis [9, 49]. In the L5-SNL model, axotomy was found to induce activation of ERK1/2, p38 and JNK in various populations of injured L5 DRG neurons. In contrast, only p38 activation was observed in TrkA-positive small neurons in the uninjured L4 DRG [108, 109]. Furthermore, increased phosphorylation of

TrkA was found in the uninjured L4 DRG neurons, but functional inhibition of p75NTR was found to suppress the injuryinduced phosphorylation of TrkA and p38 [4, 72]. These results demonstrate that nerve injury induces differential activation of MAPK in injured and uninjured DRG neurons, and that p75NTR might facilitate TrkA signaling in uninjured DRG neurons. Furthermore, as inhibition of p75NTR or injection of anti-NGF have been found to result in MAPK inhibition and as p38 inhibitors have been shown to suppress injury-induced neuropathic pain, MAPK activation in the DRG may participate in generating pain hypersensitivity after nerve injury [4, 46, 53, 72, 109, 110]. The increased NGF signaling in primary afferents following injury translates into potentiation of transient receptor potential cation channel subfamily V member 1 (TrpV1), an ion channel activated by heat as well as by various chemical substances such as capsaicin (the active ingredient in chili pepper) and hydrogen ions (lowered pH). In contrast to the findings that p75NTR facilitates TrkA signaling and injury-induced neuropathic pain [72, 109, 110], activation of TrpV1 by NGF appears to be mediated only by TrkA as NGF still induces TrpV1-dependent thermal hyperalgesia in p75NTR-deficient mice [54, 111–113]. However, other reports find that TrpV1 expression in small nociceptive DRG neurons is independent of TrkA expression [4, 46, 53, 114]. Similarly, the intracellular signaling cascades connecting NGF to TrpV1 activity are not clarified. Other studies have found that recruitment of PLC to TrkA is essential for NGF-mediated potentiation of TrpV1 activity whereas inhibition of protein kinase C (PKC) has been found not to have any effect [48, 49, 55, 112]. In contrast, other groups have found that inhibition of PLC does not block NGFinduced sensitization (albeit reducing the amplitude), whereas inhibition of PI3K or PKC completely abolishes the effect of NGF [4, 5, 113]. In addition, TrpV1 expression is described to be activated by the Ras/ERK pathway by some groups [5, 48, 115, 116], whereas others do not find this [47, 56, 113]. NGF is known to regulate several other pain-associated ion channels such as acid sensitive ion channels (ASICS2 and -3), voltage-gated sodium channels (Nav1.8), bradykinin receptors, endothelin receptors and various voltage-gated calcium channels (CaV 3.2, 3.3) [38, 117–119]. Thereby the present understanding of NGF involvement in hypersensitivity following nerve injury or inflammation is further complicated, prompting further investigation of the specific involvement of p75NTR and TrkA as well as intracellular signaling cascades in the DRGs.

The Peripheral Nerves Anatomy at a Glance The PNS is composed of parenchyma (nerve fibers, i.e., axons and surrounding Schwann cells) and stroma (the scaffold

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cytoplasmic processes surrounding and segregating groups of several small-diameter axons (C-fibers with diameters of 0.15–2 μm) thereby forming the Remak bundles. Myelin covers the A-fiber axon at intervals (internodes) of 150– 1,500 μm, leaving bare gaps on the axon — the nodes of Ranvier. Schwann cells produce a basal lamina that surrounds the Schwann cell itself and the associated axon. Extracellular matrix molecules in the basal lamina, such as laminin and collagen, regulate key aspects of Schwann cell development including the formation, architecture and function of myelin [65, 123].

made of several connective elements). Nerve fibers are wrapped in the endoneurium, a loose, soft layer of delicate connective tissue made up of endoneurial cells and a collagenous matrix that embeds and protects the nerve fibers [120] (Fig. 4a). The fibers are gathered into larger groups, the fascicles, each enveloped by the perineurium, a dense and mechanically strong sheath [121]. In mammalian nerve trunks, the perineurium consists of up to 15 layers of flattened polygonal cells, collagen fibrils and capillaries and functions as a metabolically active diffusion barrier [122]. Each fascicle may contain up to several thousand axons and several fascicles are usually bundled together with blood vessels within yet another sheath, the epineurium. The epineurium is the outermost, thick layer of dense irregular connective tissue surrounding a peripheral nerve and carrying the main supply channels of the intraneural vascular system. This connective tissue contains fibroblasts, collagen and variable amounts of fatty tissue and functions to protect and support the nerve fibers (reviewed in [65]).

Wallerian Degeneration When the normal functions of the peripheral nerves are compromised by various insults such as axotomy or nerve crush, the distal segments undergo distinct morphological and molecular changes known as Wallerian degeneration [123]. The term Wallerian degeneration refers to a series of slow processes in the distal nerve stump, setting the stage for regeneration of the nerve: distal axons and myelin sheaths degenerate, Schwann cells dedifferentiate and macrophages are recruited (Fig. 4b). Within hours after nerve fiber axotomy, the axon membrane fuses and seals followed by disintegration of the distal stump within the first days. A proposed mechanism of axonal self-destruction involves nerve insults leading to increased Ca2+ influx, calcium-dependent cytoskeletal disassembly, axonal protease activation and granular disintegration [122, 124–126]. Demyelination begins with the fragmentation of compact myelin into small structures called myelin ovoids [127]. Schwann cells in adult peripheral nerves provide a

Schwann Cells Schwann cells are the main nerve-ensheathing glial cells of the PNS and have a crucial role in maintaining normal nerve function and in mediating nerve repair following injury. Schwann cells exist as two types of cells in the adult PNS, the myelinating and the non-myelinating Schwann cells, both of which enclose the neuronal axons. The myelinating Schwann cells form a multi-layered membranous myelin sheath around a segment of a single large-caliber axon (Afiber) by spirally wrapping its plasma membrane around the axon (Fig. 4a). The non-myelinating Schwann cells form

a

Nerve fascicle Perineurium Endoneurium Schwann cells

b

Injured

Proximal axon

Distal axon stump

Wallerian degeneration

Macrophages Regenerating Axonal regrowth Axon Regenerated

Epineurium

Fig. 4 Peripheral nerve structure and regeneration following injury. a Cross-section of a peripheral nerve. The epineurium encapsulates the peripheral nerve, which consists of several nerve fascicles where each fascicle is bounded by the perineurium. Several axons covered by Schwann cells and by the endoneurium constitute a nerve fascicle. b Time course of Wallerian degeneration and axonal regeneration following nerve injury. Top panel: distal to the site of neuronal injury fragmentation of the axon and myelin sheaths occurs. Second panel: during Wallerian

degeneration, recruited macrophages engulf cellular debris and Schwann cells dedifferentiate. Third panel: the injured axon begins to regrow guided by the Schwann cells, which redifferentiate into myelinating cells and encapsulate the regenerating axon. Bottom panel: under optimal circumstances, redifferentiated Schwann cells encapsulate the regenerated axon and target reinnervation is restored, allowing normal signal transmission

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supportive milieu for axonal regeneration as they possess the ability to respond to nerve injury by dedifferentiating into their immature state and the ability to produce neurotrophic factors, two actions that facilitate successful regeneration of axons [25, 123]. Dedifferentiated Schwann cells phagocytize the generated myelin and axonal debris both independently and by recruiting circulating macrophages over a period of 3–6 weeks. In addition, Schwann cells provide a supportive cellular context for regenerative axon growth by upregulating their synthesis and secretion of neurotrophins to stimulate the regrowing axons [128–132]. During dedifferentiation, specific intracellular signaling molecules become activated and drive the dedifferentiation program: Schwann cells cease to express myelin genes, including myelin protein zero (P0) and Krox20/Egr-2, an essential transcription factor in myelination [133], and they reexpress genes found in immature states such as p75NTR [134]. Further, the transcription factor c-Jun plays a role in dedifferentiating Schwann cells not only in the demyelinating process, but also in the induction of neurotrophic factors such as GDNF and Artemin [135]. An essential process is the activation and recruitment of macrophages, which are involved in the clearance of axonal and myelin debris by phagocytosis. Activation of Toll-like receptors on Schwann cells is needed upon nerve injury to stimulate Schwann cells to produce proinflammatory mediators including TNFα, IL-1β and IL-6, which are important for recruitment of immune cells such as macrophages [136, 137]. Furthermore, secretion of metalloproteases and vasoactive mediators by resident macrophages and injured axons causes breakdown of the blood–nerve barrier, and circulating macrophages are recruited from the vasculature into the site of injury [137, 138]. Peripheral Nerve Regeneration Peripheral nerve regeneration requires activation of the intrinsic growth capacity of the injured neurons in order for axonal regrowth and target re-innervation to occur. Furthermore, remyelination of the regrowing axons is important for successful functional recovery after nerve injury. Following Wallerian degeneration, dedifferentiated Schwann cells proliferate and migrate, aligning in Bands of Büngner (Schwann cell tubes) within the basal lamina of the denervated distal endoneurial tubes. The Bands of Büngner are crucial for axonal regeneration as they provide trophic support, including secreted neurotrophins, for the regenerating axons and secrete extracellular matrix molecules like laminin and collagen as well as cell adhesion molecules (reviewed in [139]). Regrowing axons elongate from the intact proximal part of the nerve trunk across the injury site and into the Bands of Büngner. Consequently, axonal regeneration is severely compromised following axotomy, which involves damage to

the endoneurium [140, 141]. In contrast to axotomy, a crush injury usually leaves the endoneurium intact and axons can, therefore, regenerate in their native endoneurium, leading to improved regeneration and target reinnervation. The dedifferentiated Schwann cells redifferentiate upon contact with the regenerating axons, ensheath and subsequently remyelinate the axons or restore Remak bundles thereby facilitating functional recovery [139]. Neurotrophins are crucial in relation to axonal regrowth and remyelination, and their specific roles in these processes are beginning to be unveiled. Neurotrophins and Axonal Regrowth The spatial and temporal expression of neurotrophins and their receptors has been examined in the injured nerves and the main focus has been on elucidating their importance in relation to regeneration of the sciatic nerve and the median nerve, which both contain sensory and motor axons. Sensory and motor neurons have different requirements for neurotrophic support in intact as well as injured states, and techniques such as retrograde labeling allow separate analysis of neurotrophin requirements for regeneration of both axon types. As described in the section “Trk Receptors in DRG Neurons and SGCs”, sensory neurons can be divided into subpopulations depending on their need for neurotrophins. Accordingly, nociceptive and thermoceptive DRG neurons require NGF for survival [51], whereas proprioceptive neurons require NT-3 [142, 143]. BDNF and NT-4/5 are potent survival and trophic factors for intact as well as injured motor neurons in vivo [144–147]. BDNF in Axonal Regrowth Following nerve injury, a continuous slow monophasic increase of BDNF mRNA is observed in the distal segment of the rat sciatic nerve starting 3 days after axotomy and reaching maximal levels after 2–4 weeks [130, 148]. Several studies suggest that Schwann cells of the distal nerve stump synthesize BDNF and that they are responsible for the BDNF increase following injury [130, 148–150]. However, DRG neurons reportedly transport BDNF anterogradely in both peripheral and central processes [151], thus, potentially contributing to the BDNF increase as anterograde transport of BDNF is significantly enhanced after peripheral nerve injury [152]. BDNF-deficient mice die shortly after birth [153]. Thus, to examine the role of BDNF in axonal regrowth following peripheral nerve injury, research has been constrained to employ heterozygous BDNF mice and pharmacological manipulation of BDNF. The role of endogenous BDNF in axon regrowth from the sciatic nerve has been examined by depriving endogenous BDNF with BDNF-neutralizing antibodies. This treatment has been shown to result in reduced elongation of regenerating axons following sciatic nerve crush injury,

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which indicates that endogenous BDNF is important for stimulating axonal elongation [149]. Accordingly, axonal regeneration can be enhanced by repairing transected nerves of the common fibular nerve with nerve grafts to which exogenous BDNF has been applied [154]. The growth of axons through a common fibular nerve graft from a heterozygous BDNF knockout mouse is not significantly affected; however, it indicates that even reduced BDNF levels are sufficient to promote normal axonal growth [154]. These studies do not discriminate between sensory and motor neuron axonal regrowth when evaluating the role of BDNF. One approach to specifically study motor neuron axonal regeneration in mixed nerves is by applying retrograde tracers to transected nerve fibers and subsequently quantifying labeled motor neuron somas in the corresponding spinal cord segment. This approach has been employed to examine the role of various doses of exogenously applied BDNF on motor neuron axonal regrowth after transection of the tibial nerve. Low doses of exogenous BDNF has been shown to promote motor neuron axonal regeneration following repair of longterm transected nerves [155] whose capacity to regenerate is generally poor [140, 156]. In contrast, high doses of BDNF have been shown to significantly inhibit motor neuron axonal regrowth following nerve repair both immediately and after long-term transection. Thus, BDNF is suggested to exert dosedependent facilitation and inhibition on motor neuron axonal regrowth. The BDNF receptor TrkB appears to be critical for motor neuron axonal regeneration as axonal regrowth is significantly attenuated in heterozygous TrkB mutant mice. In contrast, function-blocking antibodies against p75NTR have been reported to increase motor neuron axonal regeneration and to reverse the inhibitory effect of BDNF, indicating an inhibitory role of p75NTR on motor neuron axonal regrowth [155]. Mature motor neurons express only low levels of p75NTR; however, expression is upregulated in regenerating motor neurons after peripheral nerve injury [157, 158]. Expression of p75NTR is triggered by retrograde transport of positive signals derived from axons regrowing through damaged or denervated peripheral nerve tissue [159], and it is downregulated when motor function is restored [158]. Similarly, p75NTR mRNA increases markedly in Schwann cells of the distal segment of the transected sciatic nerve, and the expression of p75NTR is downregulated to pre-injury levels when regenerating axons make contact with the Schwann cells of the distal nerve trunk [160, 161]. P75NTR expression in Schwann cells appears to be important for motor neuron axonal regrowth as a lower number of retrogradely labeled motor neurons is observed after sciatic nerve repair with a nerve-graft containing Schwann cells deficient of p75NTR [162]. The role of Schwann cell-derived BDNF in regeneration of motor neuron axons has recently been examined by analyzing regeneration of wild-type motor neuron axons through grafts

of BDNF-deficient Schwann cells. The length of regenerating axons into grafts of BDNF-deficient Schwann cells have been found to be significantly shorter than those growing into wildtype grafts. Addition of recombinant human BDNF at the site of repair were shown to rescue axon regeneration in absence of Schwann cell-derived BDNF as well as to enhance the length of regenerating axons into wild-type grafts [163], indicating that BDNF derived from Schwann cells is important for motor neuron axonal regeneration after nerve transection. It has been suggested that the enhancement of axonal regeneration is mediated via retrograde signaling of Schwann cell-derived BDNF acting on TrkB receptors on the growing axons [163]. In line with this hypothesis, Schwann cells express truncated non-catalytic TrkB receptors [147] and are suggested to bind and present BDNF to motor neuron axons. As described in the section “BDNF in DRGs”, BDNF is expressed by uninjured sensory neurons in the SNL model, and endogenous BDNF has, thus, been hypothesized to be responsible for activating the intrinsic growth capacity of DRG neurons following peripheral nerve injury. By neutralizing endogenous BDNF with BDNF siRNA or BDNF antibodies immediately prior to and following spinal nerve injury, respectively, a reduced injury/regeneration-associated gene expression and a reduced intrinsic capacity of these neurons to extend neurites in vitro have been detected. Endogenous BDNF is, however, not critical for maintaining the soma response to injury, as delayed infusion of BDNF antibody does not alter injury/regeneration-associated gene expression once it has been initiated. These results suggest BDNF as being critical for the initial induction of the regenerative response in injured sensory neurons [164]. Furthermore, BDNF enhances neurite outgrowth of DRG neurons in vitro, whereas exogenous and endogenous proBDNF collapses neurite outgrowth by activating the small GTPase RhoA and its downstream effector Rho kinase (ROCK) via p75NTR [165]. NGF, NT-3 and NT-4/5 in Axonal Regrowth The role of NGF, NT-3 and NT-4/5 with regard to axonal regrowth following peripheral nerve injury has, to the best of our knowledge, been less extensively investigated as compared to BDNF. NGF mRNA and protein are not detected in the intact adult rat sciatic nerve but expression is markedly induced in non-neuronal cells of the distal and the immediate proximal segment of transected nerves [131, 166]. The levels of NGF mRNA exhibit a biphasic response to axotomy with an immediate early rise in NGF mRNA peaking 6 h after injury. The second increase in NGF mRNA expression is slower and is believed to be caused by IL-1, released by macrophages invading the site of injury during Wallerian degeneration and lasting several weeks [131, 167]. Studies

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indicate Schwann cells as being the source of NGF [130, 148, 166, 167] in the transected nerve. The induction of p75NTR expression in Schwann cells distal to the site of axotomy [160, 161] has been proposed as a mechanism for Schwann cells to act as substitute targets for NGF-dependent neurons by concentrating and presenting NGF to regenerating NGFresponsive TrkA-positive sensory neurons [161, 168]. However, this hypothesis has been challenged by a study showing that anti-NGF treatment of crushed nociceptive axons does not affect axon regrowth or functional recovery [169]. The NT-3 level in nerve tissue has been shown to decrease shortly after nerve transection, but to return to the normal level 2 weeks post injury [130, 148]. As is the case for NGF, studies indicate Schwann cells as being the source of NT-3 in the transected nerve [170, 171]. Exogenously applied NT-3 has been found to prevent reduction of the conduction velocity of axotomized myelinated sensory neurons. Thus, NT-3 appears to promote regeneration of sensory neurons [84]. Moreover, NT-3 is important for survival of denervated Schwann cells in the distal segment of the injured nerve. Mature Schwann cells in adult nerves are able to survive in absence of axons by establishing an autocrine survival loop [170]. NT-3 is one of the important factors of the autocrine regulatory mechanism that support long-term Schwann cell survival [170, 171], nerve regeneration and remyelination after peripheral nerve injury [171]. However, prolonged denervation results in Schwann cell atrophy and, thus, a decreased axonal regeneration capacity due to loss of receptors and reduced expression of growth factors by Schwann cells [141, 172, 173]. Peripheral nerve injury has been reporter to result in downregulation of the NT-4/5 mRNA level shortly after injury. However, 2 weeks post injury the NT-4/5 mRNA level has been found to be upregulated in the distal nerve stump of the injured rodent sciatic nerve [130, 174]. Studies performed using grafts from homozygous or heterozygous NT-4/5 knockout mice incorporated into wild-type sciatic nerves have shown that even a small reduction in endogenous NT-4/5 results in a significant reduction of regenerating motor neuron axonal growth [154]. In line with these results, application of NT-4/5 to heterozygous NT-4/5 knockout mice at the time of surgical repair of a cut nerve has been found to restore motor axon growth [154]. Moreover, application of NT-4/5 to repaired rat sciatic nerves has been reported to result in enhanced axon regeneration observed by an increase in the number of regenerated axons and to improve axonal diameter and myelin thickness [175]. Schwann Cell Phenotype and Modality-Specific Regeneration Traditionally, Schwann cells are described as being either myelinating or non-myelinating [176]. However, studies indicate that Schwann cells express distinct sensory and motor

phenotypes according to growth factor expression and their ability to stimulate sensory and motor neuron axonal regrowth. Schwann cells of cutaneous nerves (containing axons of sensory neurons) and Schwann cells of ventral roots (containing axons of motor neurons) differ in growth factor profiles in both intact and injured nerves. For example, NGF and BDNF are upregulated in cutaneous nerves but minimally in ventral roots after nerve injury [177]. Accordingly, Schwann cells of sensory and motor phenotypes preferentially support cutaneous and ventral root axonal regeneration, respectively [177]. Recent evidence suggests that the Schwann cell phenotype varies even more as the profiles of growth factor expression by denervated Schwann cells have been found to vary in magnitude and pattern dependent on central or peripheral location and on the modality of their associated axons [178]. Neurotrophins and Myelination of Peripheral Nerves In addition to regrowth of axons, successful regeneration of injured peripheral nerves requires remyelination. The function of myelin is to maximize the efficiency and velocity of action potentials along the distance of axons, the importance of which becomes apparent from demyelination diseases such as multiple sclerosis and Charcot–Marie–Tooth disease [179, 180]. Peripheral myelin is formed by Schwann cells that wrap around regrowing axons in a complex, dynamic process that involves neuron–Schwann cell crosstalk. The process of peripheral myelination can be separated into three major phases of Schwann cell growth and differentiation. The first phase, the proliferative stage, is characterized by proliferation and migration of Schwann cells on axons. In the second phase, the premyelination stage, Schwann cells elongate and ensheath the axons. The third and final phase, the myelination stage, is characterized by formation and growth of the myelin sheath [181]. Neurotrophins are important for both migration of Schwann cells and for Schwann cell myelination during development and for regeneration of nerves following peripheral nerve injury. Neurotrophins and Schwann Cell Migration — the Proliferative Stage Schwann cells express, in addition to p75NTR [160, 161], TrkC and a truncated non-catalytic form of the TrkB receptor. The levels of Trk receptors are higher in the distal part of the transected sciatic nerve compared to the proximal part of the injured nerve [130]. BDNF and NT-3 are implicated in Schwann cell migration in co-cultures of Schwann cells and DRG neurons as the migration of Schwann cells is significantly enhanced by NT3 through activation of TrkC [182] and inhibited by BDNF signaling through p75NTR [183]. The differential effects of

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BDNF and NT-3 on Schwann cell migration are regulated through signaling pathways that depend on Rho GTPases. Specifically, NT-3 activation of TrkC enhances the migration of primary Schwann cells through the signaling pathway regulated by the Rho GTPases Rac1 and Cdc42. Endogenous BDNF acting through p75NTR inhibits Schwann cell migration dramatically by Src kinase-dependent activation of the guanine-nucleotide exchange factor Vav2 and RhoA [183]. In addition to NT-3, NGF also appears to facilitate Schwann cell migration, although in a p75NTR-dependent manner [184]. In line with these results, p75NTR is important for regulating Schwann cell migration during development of peripheral nerves [185]. Neurotrophins and Myelination — Developmental and In Vitro Studies Neurotrophins have been identified as important regulators of Schwann cell myelination both in vitro and in vivo during peripheral nerve development. NGF has been identified as a potent regulator of axonal signals that control myelination of TrkA-expressing DRG neuron subpopulations in vitro, thereby promoting myelination by Schwann cells [186]. The roles of BDNF and NT-3 on myelination have been examined in Schwann cell/DRG neuronal co-cultures. NT-3 levels are initially high, but are downregulated throughout the proliferation and the premyelination stages, whereas BDNF levels correlate with myelin formation. Moreover, exogenously applied BDNF enhances myelination, whereas application of NT-3 to the co-cultures inhibits myelination. In line with these results, removal of endogenous BDNF has been shown to inhibit myelination, whereas removal of endogenous NT-3 has been shown to increase myelination. Similar results have been observed in vivo in the developing mouse sciatic nerve [187]. Accordingly, adenoviral-mediated overexpression of BDNF in DRG neurons have been reported to enhance myelination in Schwann cell/DRG neuron co-cultures [188]. These results suggest that BDNF is a positive modulator of myelination and that NT-3 is a negative modulator of myelination [187]. The myelination-promoting effect of BDNF is mediated by p75NTR, whereas NT-3 mediates its myelination-inhibitory effect via TrkC. A truncated isoform of TrkB is upregulated during the myelination stage and acts as an inhibitory regulator of myelination by competing with p75NTR for the availability of endogenous BDNF. In agreement with these results, myelination is reduced upon treatment with an inhibitor of p75NTR and in homozygous p75NTR knockout mice [189]. Neurotrophins and Myelination — the Myelination Stage Considering the expression patterns of neurotrophins and neurotrophin receptors following peripheral nerve injury as

well as their roles in in vitro myelination and myelination during development, it is highly plausible that these factors play similar roles during remyelination of regenerating peripheral nerves. Especially BDNF and p75NTR have been examined and appear to be important players in remyelination after peripheral nerve injury. Endogenous BDNF can be neutralized by treatment with BDNF antibodies, resulting in an approximately 80 % reduction of the number and density of myelinated axons in the distal part of the injured sciatic nerve [149]. Likewise, p75NTR is important for remyelination after injury. Sciatic nerve crush experiments on p75NTR knockout mice have been found to result in a reduced number of myelinated axons, in reduced myelin sheath thickness as well as in reduced levels of myelin gene expression in regenerated nerves [190]. In particular, p75NTR expression in Schwann cells appears to be important for remyelination as poor myelination has been observed when p75NTR-deficient Schwann cells were grafted onto wild-type sciatic nerves [162]. Therapeutic Approaches Enhancing Peripheral Nerve Regeneration The PNS has significant regenerative potential allowing recovery after injury, although with varying degrees of success depending on type and severity of the injury. Peripheral nerve injury leads to a rapid and robust increase in neurotrophin synthesis in neurons and Schwann cells, which guide and support regenerating axons. Efforts are being made to take advantage of this to improve nerve regeneration. Various approaches of neurotrophin application have signified the important role of the neurotrophins during the early stages of regeneration and support the significance of developing combined gene and cell therapy for peripheral nerve repair. NGF — Therapeutic Approaches The expression of NGF in transected peripheral nerves increases in the distal and immediate proximal part within the first few weeks after injury [131], implying that initial basal levels of NGF could be a limiting factor, and that supplementation of NGF could have a beneficial effect on regeneration. Several experimental models have been used to investigate the effects of NGF on peripheral nerve regeneration in rodents, including direct NGF application [191], NGFcontaining fibrin sealants [192], injection of NGF-encoding lentiviral [191–194] or adenoviral [195] particles, microspheres [196] and modified scaffolds [197, 198]. These studies have shown positive effects on regeneration, but often with conflicting effects on both sensory and motor neurons. An alternative may be Schwann cell-based therapy as engineering Schwann cells for increased neurotrophin production facilitates their regenerative effect and renders them a potent tool to guide and support nerve regeneration. To

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optimize neurotrophin supply in the early stages of regeneration, a cell-based NGF gene therapy model has recently been tested in which Schwann cells robustly and continuously overexpressing NGF were transplanted onto the rat sciatic nerve upon transection. The transplanted Schwann cell were found to lead to significantly increased NGF levels and to a substantial improvement in axonal regeneration of a sub-class of sensory neuron populations resulting in reduced denervated muscle atrophy [199]. NT-3 — Therapeutic Approaches The ability of NT-3 to reverse axotomy-induced changes in adult motor and sensory neurons has been examined using the physiological measure of conduction velocity. Five weeks after axotomy, sensory and motor neuron conduction velocities have been found to be greatly reduced, however, application of NT-3 directly onto the transected nerve stump has been shown to largely prevent this and further to rescue motor neurons. This amelioration of physiological deficits in adult mammalian neurons suggests possible therapeutic applications of NT-3 [84]. NT-3 is an important component of the autocrine survival loop supporting Schwann cell survival and differentiation in absence of axons. Nerve regeneration deficiencies have been identified in NT-3 heterozygous knockout mice characterized by fewer Schwann cells in the regenerating nerve fibers of crushed sciatic nerves [171]. NT-3 heterozygous knockout mice display retardation of the myelination process, and this defect has been found to be associated with decreased Schwann cell survival and increased neurofilament packing density of regenerating axons. These observations indicate that the NT-3 status of the Schwann cells, but not of the axons, is responsible for impaired nerve regeneration and that NT-3 is essential for Schwann cell survival in early stages of regeneration-associated myelination in the adult rodent peripheral nerve [200]. Consistent with this hypothesis, the impact on axonal regeneration of the rat peroneal nerve after injury has been investigated using genetically modified peripheral nerve grafts repopulated ex vivo with Schwann cells modified to express NT-3, which has been found to result in a significant increase in the number of sensory fibers [201]. It has been demonstrated that xenograft transplants of Schwann cells from patients with Charcot–Marie–Tooth disease type 1A (CMT1A) into the sciatic nerve of nude mice delays myelination and impairs regeneration of axons passing through the grafted segments. The efficacy of NT-3 treatment using this model was evaluated by the number of myelinated fibers and Schwann cells, and NT-3 treatment was found to augment axonal regeneration. Furthermore, the effect of NT-3 on regeneration has been tested in patients with CMT1A by subcutaneous administration of NT-3. In this pilot clinical trial, it was shown that patients may benefit from NT-3

treatment, as regeneration in sural nerve biopsies before and after treatment was reported to increase the mean number of small myelinated fibers [171]. NT-4/5 and BDNF — Therapeutic Approaches The growth of regenerating axons is very poor if the surrounding Schwann cells are devoid of one or more of the neurotrophins or their receptors [155, 202] or if the Schwann cells are destroyed [203–205]. In line with this, topical application of either recombinant human BDNF or NT-4/5 to allografts from knockout mice at the time of surgical repair has been shown to overcome these deficits [154]. Similarly, electrical stimulation resulting in increased expression of BDNF and TrkB in motor neurons [206] and DRG neurons [203, 207] has been reported to result in enhancement of axonal regeneration following peripheral nerve transection [206, 208]. Furthermore, the regenerating axons have been found to grow more than twice as far during the first 2 postrepair weeks if the proximal stump of a cut nerve has been stimulated for merely 1 h at the time of surgical repair [203]. Finally, modest treadmill training has been reported to have a positive effect on axon regeneration [209] with a magnitude comparable to that obtained by electrical stimulation [210]. NT-4/5 in the proximal stump of the cut nerve is implicated in the treadmill training-induced enhancement of axon regeneration, while regeneration is independent of NT-4/5 presence in the tissue through which the axons grow [211]. Following peripheral axotomy, Schwann cells in the nerve segment distal to the injury synthesize and secrete BDNF into the Schwann cell basal lamina [130]. Initially, this Schwann cellderived BDNF is used by the regenerating axons entering endoneurial tubes in the distal segment to initiate axonal elongation. Spinal cord motor neurons contain BDNF and TrkB mRNAs [206, 212] with initially increased expression of BDNF following axotomy but decreasing again within 1 week [65, 130]. Schwann cells in the distal stump, however, can synthesize and secrete BDNF for 2 months or longer, and since dedifferentiated Schwann cells express the truncated non-catalytic TrkB receptor, stimulation of axon elongation is proposed to occur via a classic retrograde signaling [163, 213, 214]. In support of this, two smallmolecule TrkB agonists, 7,8-dihydroxyflavone [215] and deoxygedunin [216], have recently been found to enhance axon regeneration and to improved muscle reinnervation following peripheral nerve transection [217].

The spinal cord Anatomy at a Glance The murine spinal cord extends from the hindbrain at the foramen magnum to the lower part of the vertebral column

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ending at the level of the sixth lumbar vertebra [218]. Thus, the spinal cord does not reach the end of the vertebral canal, and the spinal nerves form the cauda equina in the caudalmost region of the vertebral canal. The general organization of the vertebral column is conserved among mammals, but interstrain and inter-species variations exist. In the light of the use of various mouse strains and rats for research, this may be important depending on experimental setup [18, 219–223]. As the spinal cord is shorter than the vertebral canal, only the rostral-most spinal segments correspond to the vertebrae numbers, whereas more caudally spinal cord segments are located more rostrally than the vertebrae numbers indicate. This discrepancy increases in rostro-caudal progression and increases from birth to maturity [224]. The spinal cord forms two clearly visible enlargements: the cervical enlargement (generally C5– T1) and the lumbar enlargement (generally L2–L6), which consist of the segments in which the nerves of the limbs connect to the spinal cord. The spinal cord is covered by three spinal meninges: the dura mater (outermost layer), the arachnoid mater (intermediate layer) and the pia mater (innermost layer). The subarachnoid space containing the cerebrospinal fluid is located between the arachnoid mater and the pia mater, while the epidural space is located between the dura mater and the periosteum of the vertebral canal. On each lateral side of the rodent spinal cord, the dorsal and ventral rootlets are attached and fuse to form the dorsal and ventral roots, respectively [225]. Each dorsal root enters a DRG, although dorsal rootlets in the cervical region may directly enter the DRG without forming a dorsal root [225, 226]. Each ventral root fuses with its corresponding dorsal root slightly distal to the DRG (Fig. 1). The dorsal roots consist of primary afferent sensory fibers and the ventral roots of efferent somatic motor axons [226]. The mature spinal dorsal horn can be divided into laminae I–V [227–229] and it is widely recognized that small, unmyelinated and thinly myelinated neurons, i.e., Aδ- and C-fibers that transmit pain and temperature, mainly terminate in the superficial laminae (I–II), while Aα- and Aβ-fibers that transmit proprioceptive information as well as innocuous touch and pressure mainly terminate in deeper laminae [230]. The connectivity in the spinal cord is rather complex with numerous inhibitory or excitatory interneurons modifying the afferent signals through GABAergic/glycinergic inhibition or glutamatergic excitation, respectively (reviewed in [231]). Hence, the final signal to the brain is a delicate balance between stimuli from the periphery modulated by excitatory and inhibitory signals in the spinal cord. Spinal Cord Neurotrophins and Neurotrophin Receptors P75NTR is expressed in spinal cord motor neurons, and following peripheral nerve injury, p75NTR expression increases

in their cell body as well as in their axons [232]. In addition, p75NTR is expressed in oligodendrocytes where it can induce cell death following stimulation by NGF [233, 234]. TrkA is in particular observed in the superficial dorsal horns (laminae I and II), but small amounts are also found in distinct areas of deeper laminae extending to laminae IVand V depending on the location along the spinal cord [49, 235]. TrkA expression is confined to a small subset of myelinated neurons, hence, TrkA and non-TrkA expressing neurons may represent functionally distinct nociceptor populations [49]. In addition, TrkA expressing neurons have been found to coexpress p75NTR [235]. NGF is expressed only at relatively low levels in the normal spinal cord but NGF transcript levels are upregulated as a response to peripheral nerve injury [158]. In addition to signaling via TrkA, NGF has been suggested to be important for spinal purinergic signaling in that intrathecal NGF administration has been shown to increase expression of purinergic receptor P2X3 on laminae I and II nociceptors as well as immediately ventrally to the central canal [236]. Hence, NGF may contribute to chronic pain development following peripheral nerve injury. In the normal spinal cord, BDNF transcripts have been found in lamina IX α-motorneurons and to a lower extent throughout the gray matter [237]. Its cognate Trk receptor, TrkB, is expressed on lamina I neurons but also in deeper laminae of the spinal cord [238], and TrkB transcripts are also observed in motor neurons [239]. Upon peripheral nerve injury, TrkB levels have been shown to be upregulated in both neurons and activated microglia in the spinal cord dorsal horn [78, 240]. BDNF is well known to be involved in modulation of pain processing in the CNS [241, 242] (reviewed in [82, 243]) and TrkB signaling is important for neuropathic pain development induced by peripheral nerve injury and inflammation [244, 245]. In line with this, abrogation of TrkB functioning has been shown to attenuate thermal and mechanical hypersensitivity [244, 245]. Following noxious stimuli, not only TrkB levels but also BDNF levels increase in the spinal cord dorsal horn [80, 246, 247] leading to phosphorylation of the downstream protein ERK1/2 in spinal cord dorsal horn neurons and the spinalothamic tract [78, 248]. Neurons have been shown to secrete BDNF [249] and activated microglia secrete both proBDNF and BDNF [250, 251]. Several alternatively spliced TrkB isoforms exist [252], where TrkB.T1 is the predominant isoform found in the adult mammalian nervous system. TrkB.T1 differs from the fulllength isoform by lacking the intracellular kinase domain, but extracellular high-affinity BDNF binding can still occur [252, 253]. Hetero-dimerization of TrkB.T1 with the full-length TrkB isoform inhibits TrkB autophosphorylation necessary for downstream signaling. Hence, it has been suggested that TrkB.T1 may function to reduce BDNF signaling [254–256] while others have suggested that TrkB.T1 may signal

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independently [257–260]. A study on inflammation has shown TrkB.T1 upregulation in the spinal cord dorsal horn [261] and in vitro studies on brain glial cells have revealed truncated TrkB receptor transcripts in astrocytes and oligodendrocytes but not in microglia [213]. A study on brain astrocytes has found expression of TrkB.T1 upon BDNF stimulation to be involved in calcium signaling [262]. Hence, truncated TrkB may participate in and possibly modify nociception. Transcripts of the other TrkB ligand, NT-4/5, are robustly observed in spinal cord dorsal horn laminae III– VII, to a lower extent in superficial laminae I and II and densely throughout the ventral horn of rat lumbosacral spinal cord. Furthermore, NT-4/5 transcripts are found in both neurons and in subpopulations of glial cells [237]. TrkC and its high-affinity ligand NT-3 are also found in the spinal cord [237, 239, 263]. TrkC expression has been found in spinal motor neurons [239], and following peripheral nerve injury TrkC transcript levels have been found to increase [130]. In normal adult rats, NT-3 mRNA expression has been observed to be restricted to the spinal cord gray matter in a similar pattern to BDNF expression [237] in that transcripts have been found most densely within lamina IX α-motorneurons and at lower levels elsewhere in the gray matter. Furthermore, NT-3 mRNA is also expressed by spinal cord glial cells but to a lesser extent compared to NT-4/5 mRNA [237].

Central Sensitization Following Peripheral Nerve Injury Primary afferents transmit signals from their peripheral sensory target tissue to the spinal cord dorsal horn [264]. Various changes in the spinal cord may contribute to spinal cord pathology following peripheral nerve injury, known as central sensitization. Such changes are believed to result from disinhibition, possibly caused by death of inhibitory interneurons in the dorsal horn following injury [265, 266] or by dysfunctional inhibitory signaling (reviewed in [267]) ultimately leading to the perception of non-noxious stimuli as being painful. Inflammatory mediators such as cytokines play a critical role in cell communication following nerve injury, and neuron– glia interactions are believed to be critical for establishing and maintaining injury-induced hypersensitivity. Microgliosis (outlined in the following section) induced by peripheral nerve injury is orchestrated through signals released by neurons as well as by astrocytes and immune cells. It appears that signals from injured neurons as well as from uninjured neighboring sensory neurons that share the same nerve trunk are critical for microgliosis development [268, 269], and signaling molecules including growth factors, a variety of cytokines, ATP, NO, complement components and reactive oxygen species facilitate direct neuron–microglia crosstalk (reviewed in [270]).

Microgliosis and Astrogliosis Spinal cord resident glial cells, i.e., microglia and astrocytes, play substantial roles in spinal cord pathogenesis as a response to peripheral nerve injury, ultimately affecting GABAergic and glycinergic inhibition and contributing to neuronal disinhibition. It has been demonstrated by injection of activated microglia into the spinal cord which was found to be sufficient to induce neuronal hypersensitivity [271]. Microglia constitute approximately 10 % of the total number of cells in the CNS and are considered CNS-resident immune cells originating from macrophages of the embryonal yolk sac [272]. Microglia can be divided into various groups according to their activation states: from the surveillance state to the “activated” response state with amoeboid phagocytic function. In the surveillance state, their morphological structure displays fine and long processes and a small soma. In case of altered homeostasis of the CNS, e.g., elicited by a peripheral nerve injury, they rapidly respond by changing their morphology to be amoeboid-like with retracted processes, changing their protein expression profile and increasing in number by proliferation and migration — collectively known as microgliosis [270, 273–275]. Numerous cells and signaling molecules are involved in microgliosis, including a variety of cytokines and the cytokine subfamily of chemokines (reviewed by [276, 277]). Other studies have shown that stimulation of microglial sortilin by neurotensin promoted microglial migration and their ability to secrete cytokines [278, 279]. Peripheral nerve injury triggers microgliosis that can be observed in the spinal cord dorsal horn as well as in the spinal cord ventral horn around the cell bodies of injured motor neurons ipsilateral to the injury [280, 281] as early as 24 h following nerve injury and being well established by day 3 post-injury [282] (Fig. 5). Microgliosis is confined to the spinal cord segments where injured primary afferent neurons fuse with the spinal cord [283]. Upon peripheral nerve injury, expression of a microglial activation marker, the purinergic P2X4 receptor, increases [271]. P2X4 receptor activation has been shown to be involved in mechanical allodynia development [284] by ATP-induced microglial BDNF release [251], albeit the origin of P2X4 receptor activating ATP is not clear [285, 286]. Not only BDNF but also proBDNF has been shown to be released by activated microglia [250, 251]. Astrocytes also become activated following injury [287] although at a slightly later time point than microglia [288, 289]. Astrocytes are the most numerous cells among CNS glial cells [290] and are essential for the maintenance of a functional environment in the CNS [291, 292]. Like microglia, astrocytes are able to change morphology and become more dense in response to neuronal injury, metabolic insults or inflammation [292]. Activated microglia release IL-18 that acts on astrocytes and plays an important role in generation

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a

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Fig. 5 a In the normal spinal cord, GABA(A) receptors mediate neuronal chloride influx while the potassium–chloride co-transporters KCC2 mediate chloride efflux. This maintains intracellular chloride concentrations below the equilibrium potential for chloride, i.e., neuronal hyperpolarization is achieved. b Upon peripheral nerve injury, microgliosis is

initiated (left). Activated microglia secrete BDNF, which acts on TrkB to initiate incompletely understood pathways leading to significantly decreased KCC2 levels (right). Consequently, intracellular chloride concentrations rise and GABA-mediated neuronal inhibition is attenuated, ultimately contributing to neuropathic pain development

of tactile allodynia [293]. Hence, microglia activation contributes to astrocyte activation, and continuous crosstalk between microglia and astrocytes involved in maintenance of spinal cord pathology following nerve injury is generally observed [290].

sortilin in the spinal cord [301] indicates a potential role of proBDNF in spinal cord signaling upon peripheral nerve injury. Under uninjured conditions, the neuron-restricted potassium–chloride co-transporter KCC2 establishes a chloride gradient by maintaining the intracellular chloride concentration below the equilibrium potential for chloride. Upon activation by GABA, the GABA(A) receptors mediate influx of chloride ions down their electrochemical gradient. This renders the neuronal membrane potential more negative relative to the resting membrane potential, i.e., hyperpolarization is achieved, and the chance of generating a successful action potential is diminished. Hence, KCC2 is critical for establishment of the chloride gradient that is needed for GABAergic and glycinergic inhibition in adult neurons [302, 303]. In the spinal cord, glycine receptors and GABA(A) receptors colocalize at postsynaptic densities [304], display similar chloride and HCO3− permeability [305] and both mediate inhibitory neurotransmission [306, 307]. Therefore, glycine receptors are believed to contribute to intracellular chloride concentrations in a similar manner to GABA(A) receptors both in normal conditions but also following nerve injury [302]. Upon nerve injury, microglia-derived BDNF binds to TrkB at the neuronal cell surface resulting in TrkB dimerization and

Effects of Spinal Cord BDNF Following Peripheral Nerve Injury Microglial BDNF released following peripheral nerve injury has been found to depend on microglial p38 MAPK activation [294–296]. In addition, in vitro studies suggest a role for the adenosine A2A receptor in controlling BDNF release from activated microglia [297]. To the best of our knowledge, no reports of proBDNF involved in spinal cord signaling following peripheral nerve injury have yet been made in relation to neuropathic pain, and it is BDNF that has been found at terminals of primary afferents in the superficial dorsal horn and to a lesser extent in deeper laminae [298]. However, in the hippocampus proBDNF as well as mature BDNF have been reported to be involved in hippocampal synaptic plasticity [299, 300]. Furthermore, secretion of proBDNF by activated microglia [250, 251] and expression of the proBDNF receptor

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autophosphorylation of tyrosine residues within the intracellular part of the receptor creating docking sites for various adaptor proteins. Phosphorylation of tyrosine residue Y816 enables docking of PLC-γ and phosphorylation of tyrosine residue Y515 enables docking of Src homology 2 domain containing transforming protein (Shc) and FGF receptor substrate 2 (FRS-2), all of which activate various signal cascades ultimately leading to decreased KCC2 levels [308] through yet unknown cellular sorting mechanisms (Fig. 5). Consequently, chloride ions accumulate in the neurons and neuronal hyperpolarization via activated GABA(A) receptors can no longer be sustained [302, 303, 309, 310]. Therapeutic Approaches in the Spinal Cord to Inhibit Neuropathic Pain Neuronal hypersensitivity induced by nerve injury is a complex process initiated and maintained by interplay of spinal cord substances released by neurons and activated glial cells. Various therapeutic approaches are being explored in order to reverse injury-induced hypersensitivity. Glial Inhibition Activation of microglial P2X4 receptors appears to play a critical role in tactile allodynia development as inhibition of P2X4 receptors has been reported to impair microglial BDNF release and to reverse tactile allodynia [251, 271]. Furthermore, it has been suggested that BDNF affects astrocyte activation [311]. This suggests a treatment strategy based on inhibition of BDNF release from microglia upon nerve injury, e.g., via a P2X4 receptor antagonist. As gliosis is only triggered by abnormal events such as nerve injury, modulation of glial responses to restore normal neuronal patterns might be a possible treatment method without interfering with normal neuronal function. TrkB Antagonists Because activated microglia respond by releasing BDNF, thereby activating neuronal TrkB with consequent KCC2 downregulation, TrkB can be considered a potential drug target. In accordance, systemic administration of the TrkB antagonist cyclotraxin-B has been proven to prevent and reverse cold allodynia in animal models [312]. However, the concurrent inhibition of systemic normal TrkB function may prove to result in numerous side effects.

Conclusion Neurotrophin signaling is essential for key phases of PNS development, including cellular survival and differentiation,

neuronal sprouting and for the myelination process. Following injury to peripheral nerves, neurotrophin signaling pathways are reactivated in neurons and glia to facilitate nerve regeneration and functional recovery. Unfortunately, nerve recovery is often far from complete, and unwanted side effects of neurotrophin signaling in the spinal cord further initiate processes that lead to chronic neuropathic pain symptoms. Thus, there is an immense need to increase our understanding of fundamental biological aspects of neurotrophins and their receptors in neuronal and glial biology as well as their involvement in cellular communication. Hopefully, future research will facilitate the development of better treatments to increase functional nerve recovery after injury and to prevent or alleviate neuropathic pain. Acknowledgments The authors thank the Lundbeck Foundation, Simon Fougner Hartmanns Familiefond, Familien Hede-Nielsens Fond, Aase og Ejnar Danielsens Fond and PAINCAGE (European Framework Programme 7).

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