Agrin requires specific proteins to selectively activate γ-aminobutyric acid neurons for pain suppression

Experimental Neurology 261 (2014) 646–653

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Agrin requires specific proteins to selectively activate γ-aminobutyric acid neurons for pain suppression Diana Erasso a, Gabriel Tender b, Roy C. Levitt a,c,d, Jian-Guo Cui a,⁎ a

Department of Anesthesiology, Perioperative Medicine and Pain Management, University of Miami Miller School of Medicine, Miami, FL, USA Department of Neurosurgery, School of Medicine, Louisiana State University, New Orleans, LA, USA John P. Hussman Institute for Human Genomics, University of Miami Miller School of Medicine, Miami, FL, USA d Bruce W. Carter Miami Veterans Healthcare System, Miami, FL, USA b c

a r t i c l e

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Article history: Received 1 April 2014 Revised 11 August 2014 Accepted 14 August 2014 Available online 21 August 2014 Keywords: Agrin Selective activation Neurofilament 200 Mitofisin 2 Neuropathic pain Spinal cord injury

a b s t r a c t Agrin, a heparan sulfate proteoglycan functioning as a neuro-muscular junction inducer, has been shown to inhibit neuropathic pain in sciatic nerve injury rat models, via phosphorylation of N-Methyl-D-aspartate receptor NR1 subunits in gamma-aminobutyric acid neurons. However, its effects on spinal cord injury-induced neuropathic pain, a debilitating syndrome frequently encountered after various spine traumas, are unknown. In the present investigation, we studied the 50 kDa agrin isoform effects in a quisqualic acid dorsal horn injection rat model mimicking spinal cord injury-induced neuropathic pain. Our results indicate that 50 kDa agrin decreased only in the dorsal horn of neuropathic animals and increased 50 kDa agrin expression in the dorsal horn, via intraspinal injection of adeno-associated virus serum type two, suppressed spinal cord injury-induced neuropathic pain. Also, the reason why 50 kDa agrin only activates the N-Methyl-D-aspartate receptor NR1 subunits in the GABA neurons, but not in sensory neurons, is unknown. Using immunoprecipitation and Western-blot analysis, two dimensional gel separation, and mass spectrometry, we identified several specific proteins in the reaction protein complex, such as neurofilament 200 and mitofusin 2, that are required for the activation of the NR1 subunits of gamma-aminobutyric acid inhibitory neurons by 50 kDa agrin. These findings indicate that 50 kDa agrin is a promising agent for neuropathic pain treatment. © 2014 Elsevier Inc. All rights reserved.

Introduction Spinal cord injury-induced neuropathic pain (SCI–NP) is a debilitating syndrome associated with pathologic changes in structure, biochemistry, and genes in the peripheral and central nervous system (Dworkin et al., 2003; Jarvis and Boyce-Rustay, 2009; Jensen et al., 2009; Julius and Basbaum, 2001; Scholz and Woolf, 2007; Wieseler et al., 2010; Yezierski, 2009). It is estimated that up to 85% of SCI patients suffer from neuropathic pain (NP) (Bell et al., 2009; Siddall, 2009; Ullrich et al., 2008). Current pharmacological and surgical therapies are often ineffective over time (Bryce et al., 2007; Hulsebosch, 2005). Therefore, SCI–NP decreases quality of life and represents a major barrier for these patients rehabilitation from SCI-induced physical, psychological, and social problems (Stormer et al., 1997; Ullrich et al., 2008). Recently, we found that agrin, a heparan sulfate proteoglycan, plays an important role in pain modulation (Cui and Bazan, 2010), although its function was initially described as a synaptic inducer (Bezakova ⁎ Corresponding author at: Department of Anesthesiology, Perioperative Medicine and Pain Management, University of Miami Miller School of Medicine, Miami, FL 33136, USA. Fax: +1 305 243 1373. E-mail address: [email protected] (J.-G. Cui).

http://dx.doi.org/10.1016/j.expneurol.2014.08.014 0014-4886/© 2014 Elsevier Inc. All rights reserved.

and Ruegg, 2003; Bowe and Fallon, 1995). Agrin has multiple isoforms, due to different amino acid inserts at X, Y, and Z sites (Gesemann et al., 1995). These isoforms are expressed in many cell types and have broad functions in the central nervous system (CNS) (Bose et al., 2000; Hilgenberg et al., 2002; Kroger and Schroder, 2002; Mantych and Ferreira, 2001). However, the agrin isoforms with 4 amino acids inserted at the Y site and 8 (or more) amino acids inserted at the Z site at the C-termini are solely expressed in neurons, i.e., neuronal agrins (neuronal agrins will be simply called “agrin” henceforth) (Cohen et al., 1997; Ji et al., 1998). In the rat spinal cord, we identified two isoforms: the 50 kDa (Ag50) and the 25 kDa (Ag25) agrin (Cui and Bazan, 2010). We have demonstrated that, in sciatic nerve injury rat models, such as Bennett and Gazelius, agrin decreased only in the dorsal horn (DH) and dorsal root ganglia (DRG) of the rats that developed neuropathic pain (Cui and Bazan, 2010). When an agrin gene, coded for Ag50, was delivered into the DH sensory neurons via adeno-associated virus (AAV, serum type two—AAV2-Ag50) (McCarty, 2008), Ag50 was expressed and released to selectively activate gamma-aminobutyric acid (GABA) neurons via phosphorylation of GABA neurons' NMDA receptor NR1 subunits (Wang and Kriegstein, 2008; Xi et al., 2009) at serine residue 896/897, thus suppressing NP, while Ag25 did not phosphorylate NR1, nor did it suppress NP. However, several questions arise

D. Erasso et al. / Experimental Neurology 261 (2014) 646–653

from our previous study: 1. Does agrin decrease in SCI-induced NP? 2. Under what conditions does Ag50 only activate GABA neurons via NR1 subunits? NR1 subunit is a common, basic structure of NMDA receptors (Furukawa et al., 2005; Kalia et al., 2008; Plant et al., 1997). NMDA receptors are widely expressed on sensory neurons, glial cells, as well as GABA neurons (Schipke et al., 2001; Xi et al., 2009). Why does Ag50 solely activate the NR1 subunit of GABA neurons? It is reasonable to hypothesize that specific conditions, such as unique associated proteins to form certain activation structure, are required for GABA neuron activation by Ag50. In order to investigate the role of Ag50 in SCI–NP, we utilized the quisqualic acid (QA) induced SCI rat model. QA is a mixed AMPA and glutamate metabotropic receptor agonist (Johnson et al., 1985; Rainbow et al., 1984; Walker, 1976). When injected into DH superficial layers, QA overexcites/damages spinal sensory neurons and eventually induces neuropathic pain in rats a few weeks after injection, presenting as tactile allodynia and thermal hyperalgesia (Pisharodi and Nauta, 1985; Vierck et al., 2000; Yezierski, 2005). Our current results demonstrated that agrin was decreased in the DH of rats after QA-induced SCI–NP and intraspinal injection of AAV2Ag50 increased Ag50 expression and suppressed SCI–NP. More importantly, neurofilament 200 (NF200) and mitofusin 2 (Mfn2) were identified in the Ag50-induced reaction protein complex precipitated by agrin antibody from Ag50 treated dorsal spinal cords, suggesting that NF200 and Mfn2 act as the specific associated proteins to form activation structure. This activation structure, including NF200, Mfn2, and other specific associated proteins, is required by Ag50 to solely activate GABA neurons.

Materials and methods Animals All procedures and protocols were approved by the Animal Care and Use Committee of the University of Miami and the Louisiana State University. The animals were allowed to accommodate for 7 days before surgery, with free access to food and water and 12/12 h day/night cycle.

Surgery Surgical procedures were performed under anesthesia by 1.5% isoflurane with a mixture of 50% oxygen and 50% air delivered by means of an open-mask system at 1.5 L/min. Body temperature was maintained during the procedures at 38 ± 0.5 °C by an automatic heating device.

Quisqualic acid intraspinal injection Male Sprague–Dawley (SD) rats, with weights of 240–250 g, were used for QA injection. The vertebral column of the rats was stabilized by two clamps holding the T11 and T2 processes on a stereotaxic frame, leaving the chest to move freely. A small laminectomy was performed on the left T13 vertebra to expose the dura and spinal cord. A modified Hamilton G36 syringe with a finer, shorter tip (Length: 0.14 mm; OD: 0.14 mm) was inserted into the left L3–5 DH along the root entry zone at a depth of 220–250 μm with a micromanipulator (Wallin et al., 2002). QA (2 μl, 125 mM in saline) was injected slowly at three sites, 4 mm apart. After QA injection, the rats were examined for gait, muscular tone, stress, paw shaking, and paw licking as well as tactile allodynia and thermal hyperalgesia, every other day until a new procedure started. Sham surgery using saline instead of QA was performed on a group of rats as a control.

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Intraspinal injection of adeno-associated viral–agrin-gene vector After SCI–NP was confirmed on three consecutive days, the rats were subjected to AAV2-Ag intraspinal injection. The same surgical technique as QA injection was used for AAV2-Ag injection. AAV2-Ag50, AAV2Ag25, or AAV2-null (5 × 108 GC in 3 μl) was injected into the superficial DH over 5 min. After completing the injection, the needle was kept in for 1 min. The tissue and skin were then closed in anatomical layers. Following AAV2-Ag injection, the rats were examined daily for tactile allodynia. Behavioral tests In a quiet room with daylight-like illumination, the animals were allowed to adapt for 30 min before data were collected. All of the data collections were performed between 9:30 am and 4:00 pm. If a rat would exhibit severe stress, autotomy, or paralysis, during these experiments, the rat would be euthanatized. Tactile allodynia test In an organic-glass chamber (20 × 20 cm) with mesh net floor, the animals were subjected to the tactile allodynia test, in which an innocuous touch stimulus may induce a brief paw withdrawal to escape from the stimulation. A set of the von Frey filaments from 0.01 to 26 g (Stoelting) was applied to the mid-paw of rats, starting from 1 g, and going in either direction. The filament was pressed on the plantar surface for 1 to 2 s, until it bended, starting with the right paw. If a filament induced a brief withdrawal of a paw for three times out of five stimuli, the value was recorded as the animal's withdrawal threshold (Cui et al., 2000, 1996, 1997). Allodynia was considered to be present if a withdrawal threshold decrease reached a difference of two standard errors compared to the threshold in pre-injury level (Cui et al., 1996; Tender et al., 2008). After QA intraspinal injection, the rats were tested for tactile allodynia daily for 4 weeks and every third day thereafter, until allodynia was resolved. Thermal hyperalgesia test The thermal hyperalgesia test device is composed of a chamber (20 × 20 cm) with glass bottom of adjustable temperature and a light beam focused on the paw plantar surface (LifeSci, CA). When a button is pushed, the light beam is changed to an intense one to induce paw withdrawal. The temperature of the glass bottom was set at 28 °C, and the intensity of the intense light beam at 35% of the light source. The latency time in seconds from the onset of the intense light beam to paw withdrawal was defined as the paw withdrawal latency. Two consecutive tests performed at an interval of 20 min were averaged to establish the latency. If a latency value of an injured paw was 30% lower than that of pre-injury baseline, the animal was considered to exhibit thermal hyperalgesia (Cui et al., 1996; Tender et al., 2008). After QA SCI, the rats were tested for thermal hyperalgesia daily for 4 weeks and every third day thereafter. Immunoprecipitation and Western blot Protein extraction At two-week post QA injection, or 2-day and 7-day post AAV2-Ag injection, the rats were deeply anesthetized with isoflurane and decapitated. Dorsal spinal cord L3–5 segments were quickly removed and put into RIPA lysis buffer (1 ml, Sigma). The lysis buffer contained protease inhibitors and phosphatase inhibitors (1:100, Sigma). The tissue was homogenized for 2 min and incubated on ice for 45 min. Protein concentration was determined by Agilent protein chip (Agilent 2100 Bioanalyzer, Germany) and the Bradford method (BioRad reagents).

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Immunoprecipitation Fifty microliters of IgG dynamic beads (Invitrogen) were incubated with agrin or p-NR1 antibody (2 μg) for 45 min. The supernatant was discarded and the beads were washed with binding buffer. Sample proteins (200–400 μg) were added and incubated with the beads for 4 h at 4 °C. The complex was washed 3 times, and the proteins in this complex were eluted out for two dimensional gel electrophoresis. Two dimensional difference gel electrophoresis and mass spectrometry Subsequently, Applied Biomics Company ran two dimensional (2D) gel and MS according to their standard protocols. The eluted protein samples were labeled with 1 μl Cy3 or Cy5 dye dilution (Amersham, Piscataway, NJ) on ice for 30 min, followed by the addition of 1 μl of 10 mM lysine to stop the labeling reaction. The Cy-Dye-labeled preparations were mixed and an equal volume of 2 × 2D sample buffer (8 M urea, 4% CHAPS, 20 mg/ml DTT, 2% pharmalytes) was added. This was followed by the addition of 100 μl destreak solution (GE Healthcare, Piscataway, NJ) and rehydration buffer (7 M urea, 2 M thiourea, 4% CHAPS, 20 mg/ml DTT, 1% pharmalytes and trace amount of bromophenol blue) to a total volume of 260 μl. Solutions were incubated at RT for 10 min on a shaker and centrifuged for 10 min at 16,000 g before loading 250 μl per immobilized pH gradient (IPG) strip (13 cm, pH 3–10 linear isoelectric focusing strip, Amersham). IEF was performed for a total of 25,000 V-h under standard conditions (Amersham). Each IPG strip was incubated with 10 ml of equilibration solution 1 (50 mM Tris–HCl, pH 8.8, containing 6 M urea, 30% glycerol, 2% SDS, trace amount of bromophenol blue, and 10 mg/ml DTT) for 15 min with gentle shaking followed by incubation in 10 ml of equilibration solution 2 (50 mM Tris–HCl, pH 8.8, containing 6 M urea, 30% glycerol, 2% SDS, trace amount of bromophenol blue, and 45 mg/ml iodoacetamide) for 10 min with gentle shaking. Strips were rinsed in SDS gel running buffer once and inserted into a 10.5% SDS gel prepared on low fluorescent glass plates (18 × 16 cm, 1 mm thickness) and sealed with 0.5% agarose sealing solution in SDS running buffer. Electrophoresis was performed at 16 °C. Each gel was scanned immediately following electrophoresis with a Typhoon Trio Scanner (Amersham). Images were analyzed with ImageQuant software. Western blot Protein (20 μg) from each sample were mixed with 2× electrophoresis buffer and boiled for 5 min. The samples were loaded on 4–15% acrylamide tris–glycine SDS gels and electrophoresed in a cell (Bio-Rad) for approximately 1 h. After the protein was transferred on to a PVDF, standard Western blot procedures were performed according to the protocol. The membrane was scanned with an Odyssey (LI-COR) scanner or ChemiDoc XRS imaging system (Bio-Rad) and analyzed by Image Lab software (Cui et al., 2010; Lukiw et al., 2005). For IP-Western blot, a half (10 μl) of the IP elute was loaded for electrophoresis, followed by the same western procedures with targeted protein antibodies. Immunohistochemistry (IHC) The animals were deeply anesthetized with isoflurane and perfused via left ventricle with 100 ml warmed saline, followed by 50 ml warmed 4% paraformaldehyde (pH 7.4) and 250 ml cold 4% paraformaldehyde. The L3–5 spinal cord segments were dissected out and cut into 20 μm sections with a cryostat (Leica, Germany). The sections were mounted on a slide and stored at −80 °C. When processed for staining, a standard IHC protocol was followed. In addition, negative and positive controls for an antibody were used, and antibody absorption by matched peptides was tested. Statistics All data are presented as mean + SEM. Comparisons between different treatment groups were subjected to one-way ANOVA or twoway ANOVA, followed by Tukey's multiple comparison test or post hoc Steel–Dwass's multiple comparison test. Changes of the paw withdrawal

thresholds and paw withdrawal latency, before and after nerve injury or treatments with vector injection in the same rats were analyzed by paired t tests. Sample size of 8–10 rats will provide 80% power at a two-sided significance level of 0.05 to detect a mean difference of 30%.

Results Behavioral changes induced by intraspinal QA injection After quisqualic acid (QA) injection into the left superficial L3–5 spinal cord segments, the rats initially exhibited muscular hypertonia and spasticity in both legs (left more than right) and lower trunk, at 1- to 5-day post injection. The hypertonia and spasticity did not last longer than 7 days. Then, the rats gradually developed tactile allodynia and thermal hyperalgesia starting around day 9 post injection, as indicated by the abnormally low paw withdrawal threshold and significantly shorter paw withdrawal latency, respectively. In the majority of the QA-induced SCI rats (about 92%), both paws displayed an abnormally low withdrawal threshold and shorter withdrawal latency, as shown in Fig. 1. The average paw withdrawal threshold was 5.16 ± 1.02 g in the left paws and 5.88 + 1.42 g in the right paws at the time of peak allodynia (at the third week after QA injection), compared to a cutoff threshold of 26 g in the paws of sham operated rats. The average paw withdrawal latency was decreased to 48% in the left paws and 41% in the right paws, compared to that of sham operated rats. The signs of tactile allodynia and thermal hyperalgesia lasted 5 to 8 weeks in QAinduced SCI rats. The NP signs tended to disappear in majority of the rats between 5 and 8 weeks (data are not shown).

Biochemical alterations in the DH after QA injection Some of the rats that developed neuropathic pain after QA intraspinal injection were subjected to transcardial perfusion and immunostaining at the time of maximal neuropathic pain (the third week after QA injection, for the majority of rats). Immunohistochemistry (IHC) with agrin antibody (Agr510, Stressgen. 1:8000) revealed that agrin was decreased in the DH of SCI–NP rats (Fig. 2A; arrowhead). This decrease occurred in both sides of the DH in the majority of neuropathic rats. In the DH of normal and sham operated rats, agrin expression was mainly located in the laminas I, II, and III and symmetrical, except for a lucent gap between laminas I and II. The difference in agrin expression between SCI and sham or normal DH was significant (Fig. 2B). This staining was blocked by absorption with the immunizing agrin peptide (1 mg/ml; 1:2000). In addition, progenitor cells on a slide, as a negative control, could not be labeled by the agrin antibody. Western blot analysis further confirmed the agrin decrease in the SCI–NP rats, but not in sham-operated and normal rats (Figs. 2C and D). These results indicate that agrin decrease is associated with neuropathic pain after QA injection into the rat DH.

Pain suppression by AAV2-Ag50 intraspinal injection Behavioral alterations after intraspinal AAV2-Ag50 injection When AAV2-Ag50 (GC 5 × 108 in 3 μl; viral protein wrapped vector) was injected into the superficial DH of SCI–NP rats on day 15, the paw withdrawal thresholds of both left and right sides started to rise at 24 h post-injection. SCI–NP was suppressed in the rats by 48 h post injection, as indicated by the normalization of the withdrawal threshold. In contrast, intraspinal AAV2-Ag25 and AAV2-null injections did not change the withdrawal threshold (Fig. 3). Sham intraspinal injection did not affect the paw withdrawal thresholds either. These results suggest that Ag50 is the functional molecule responsible for pain suppression.

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Fig. 1. Intraspinal quisqualic acid (QA) injection into the L3–5 spinal cord segments induced rear paw hypersensitivity in rats. After the QA injection into the superficial layers of the dorsal horn, the rats exhibited bilateral tactile allodynia (left) and thermal hyperalgesia (right). Sham injections had no effect. There is a significant difference in paw withdrawal thresholds/paw withdrawal latencies between before and after QA treatment (n = 12, *P b 0.05; **P b 0.01; paired t test). The comparison for paw withdrawal thresholds/paw withdrawal latencies is also significantly different between QA treated and sham operated rats (*P b 0.05; **P b 0.01; F = 19.52; two-way ANOVA, followed by the Tukey's multiple comparison test; Data are presented as mean ± SEM).

Ag50 expression and increased p-NR1 in the DH The protein samples from spinal cords of sham-treated, AAV2-Ag50 treated, AAV2-Ag25 treated, and AAV2-null treated rats were subjected to Western blot analysis. AAV2-Ag50 and AAV2-Ag25 increased Ag50 and Ag25 proteins, respectively, in the DH, whereas sham and AAV2null treated samples did not exhibit increased expression of these proteins. Also, AAV2-Ag50, AAV2-Ag25, and AAV2-null treated samples were myc positive, suggesting that the Ag50 and Ag25 proteins were derived from the viral protein wrapped vectors (Fig. 4). Western blot showed that there is no obvious difference among these protein samples. GADPH labeling suggests protein sample loading was even. However, western blot after agrin antibody precipitated p-NR1 protein complex, p-NR1 bands showed a clear difference between allodynic, AAV2-Ag25 treated and AAV2-Ag50 treated samples, suggesting that p-NR1 was increased in AAV2-Ag50 treated DH.

Using the same immunoprecipitation method, we eluted protein complex from AAV2-Ag50 treated, AAV2-null treated, and normal DH. Half of each protein complex was loaded on Western blot for neurofilament 200 (NF200) or mitofusin 2 (Mfn2) antibody detection. On the western membrane, AAV2-Ag50 treated samples displayed a strong positive NF200 band and a strong positive Mfn2 band, while AAV2null treated samples showed a faint NF200 band and a faint Mfn2 band, close to background. Normal, untreated samples displayed a positive Mfn2 band and a weak positive NF200 band. The molecular weights of detected NF200 and Mfn2 in the protein complex were around 110 kDa and 55 kDa, respectively, instead of 200 kDa and 75 kDa, as would be expected for the entire protein. Although IP test is not quantitative, the results suggest that there is a significant difference between AAV2Ag50 and AAV2-null treated animals. These data indicate that NF200 and Mfn2 participate in Ag50-induced GABA neuron activation. Discussion

Immunoprecipitation with agrin antibody and 2D gel electrophoresis The L3–5 dorsal spinal cord segments, treated with AAV2-Ag50 and AAV2-null, were dissected out at 48 h post treatment for protein extraction. After the protein samples were incubated with agrin antibody beads, the eluted protein complex was subjected to twodimensional electrophoresis on a gel. Individual proteins were well separated (Fig. 5A). The AAV2-Ag50 treated samples showed 6 unique proteins, different from those of AAV2-null treated (Fig. 5A, left, circled), while AAV2-null treated samples showed 12 different proteins (Fig. 5A, right, circled). LC-MS identified the 6 proteins as neurofilament heavy chain, mitofusin 2, etc., according to the unique peptide sequence molecular weight. In contrast, p-NR1 antibody immunoprecipitation did not show any difference for AAV2-Ag50 and AAV2-null treated samples.

Spinal cord injury induced neuropathic pain occurs in up to 85% of these patients, and its treatment remains a challenge in clinic. Therefore, finding an effective, satisfactory treatment for SCI–NP represents a major priority. Animal models can only partially mimic the SCI condition in humans. Although there is no perfect SCI model, there are several animal models available for research, such as spinal cord contusion, spinal cord semitransection, and quisqualic acid intraspinal injection (Hulsebosch et al., 2009; Yezierski, 2005). While the contusion model could be used to study SCI–NP, injury magnitude and neuropathic pain incidence are more difficult to control and predict than the QA model. QA causes overexcitation and excitotoxic injury of spinal sensory neurons that produces SCI–NP, presenting as tactile allodynia and thermal hyperalgesia a

Fig. 2. Agrin expression decreased in the dorsal horn (DH) after quisqualic acid (QA) injection. (A) Agrin DH expression decreased in the rats undergoing QA injection, but not in normal and sham-operated rats. (B) Bar graph depicting the quantitative analysis of agrin fluorescent particles in normal, QA, and sham-operated rats. The difference between QA injected and normal or sham-operated rats is significant (n = 6, **P b 0.01; one-way ANOVA, followed by the Tukey's multiple comparison test. Data are presented as mean ± SEM). (C) Western blot analysis confirmed agrin decrease in the DH of QA injected, but not normal and sham-operated rats. (D) Bar graph depicting the quantitative analysis of agrin expression in normal, QA injected, and sham-operated rats. The difference between QA injected and normal or sham-operated rats is significant (n = 4, *P b 0.05; **P b 0.01; one-way ANOVA, followed by the Tukey's test).

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Fig. 3. 50 kDa agrin inhibited neuropathic pain in SCI–NP rat model. Ag50 gene delivery into the DH by adeno-associated virus serum type 2 (AAV2-Ag50) on day 15 (arrow) induced normalization of the paw withdrawal threshold, i.e., resolution of tactile allodynia (n = 12 rats). The Ag50 suppressive effects lasted for more than 12 days. Intraspinal AAV2-Ag25 (n = 8 rats) and AAV2-null (n = 6 rats) injections had no effect. Sham operation (saline injection) did not affect the paw withdrawal thresholds (n = 6 rats). Intraspinal AAV2-Ag50 injection did not affect the paw withdrawal thresholds in shamoperated rats either (n = 5 rats). There is a significant difference between AAV2-Ag50 treated and AAV2-Ag25 treated, or AAV2-null treated rats (*P b 0.05; **P b 0.01; F = 13.25; two-way ANOVA, followed by Tukey's multiple comparison test and post hoc Steel–Dwass's multiple comparison test).

few weeks after injection (Pisharodi and Nauta, 1985; Vierck et al., 2000; Yezierski, 2005). Precision localization and reproducible precise injection volumes allow for better control of spinal cord injury after QA treatment, and a high incidence of neuropathic pain (higher than 85%). Therefore, we selected the QA rat model for our study. Our experiments showed that initially, the rats exhibited muscular hypertonia and spasticity at 1- to 5-day post QA injection (these symptoms subsided in a few days), and then gradually developed tactile allodynia and thermal hyperalgesia (Fig. 1). These symptoms and signs suggest that there is a state of hyper-excitability in the spinal center, consistent with the mechanism of disease development that closely mimics that of SCI–NP patients (Yezierski et al., 1998). We further examined agrin expression in the DH of the SCI–NP rats. Although agrin has a synaptic protein function, it is located not only in synapses, but also in dentrites, axons, and soma. Various agrin isoforms derived from different amino acid inserts play multiple roles in

neuronal survival, growth, and synaptogenesis (Bezakova and Ruegg, 2003; Bose et al., 2000; Campanelli et al., 1991; Hilgenberg et al., 2002; Kroger and Schroder, 2002). Agrins with 4 and 8 amino acids inserted at Y and Z sites (or more amino acids inserted at the Z site), respectively, are solely expressed in neurons (Cohen et al., 1997; Cui and Bazan, 2010; Hoch et al., 1993; Ji et al., 2002). Our previous studies showed that neuronal agrin plays an important role in modulating pain signals in NP induced by peripheral nerve injury. In this study, we also observed that agrin immunoreactivity was decreased in the superficial DH of QA-induced SCI–NP rats. Agrin expression decreased bilaterally in the DH (Fig. 2A), consistent with the behavioral testing results (bilateral decrease in paw withdrawal thresholds). Western blot results further showed that Ag50 was significantly decreased in SCI–NP rats, compared to that in normal or sham-operated rats (Fig. 2C). These data indicate that Ag50 is involved in QA-induced SCI–NP. If Ag50 decrease results in SCI–NP after QA injection, can agrin upregulation suppress SCI–NP? When viral protein wrapped AAV2-Ag50 vector was injected into the superficial DH of SCI–NP rats, the paw withdrawal thresholds were elevated at 24 h and normalized at 48 h post injection, while AAV2-Ag25 and AAV2-null injections did not alter the paw withdrawal thresholds (Fig. 3). Although we did not examine whether Ag50 was increased in the DH at 24 h post AAV2-Ag50 injection in this study, our previous study suggested that Ag50 can be expressed at 24 h after AAV2-Ag50 injection and that Ag50 can be synthesized as quickly as 1.5 h after a drug treatment (Cui and Bazan, 2010). If Ag50 could not be produced as fast as physiologically required to modulate pain signals, it would have to be stored in large amounts, facts not supported by our DH staining results. Therefore, it is likely that Ag50 is quickly synthesized when dictated by the physiological needs, consistent with modulation requirements. In addition, sham injection and AAV2-Ag50 injection into normal rats did not affect their paw withdrawal thresholds (Fig. 3). Western blot analysis confirmed that 50 kDa and 25 kDa agrin (Ag25) were increased in the samples treated with AAV2-Ag50 and AAV2-Ag25 vectors, respectively (Fig. 4). However, only increases in DH Ag50 were associated with the normalization of paw withdrawal thresholds and increased p-NR1 in the precipitated protein complex (Fig. 4). These results demonstrate that Ag50 is the critical inhibitory molecule responsible in pain modulation. Ag25 may be a metabolite of Ag50, a possibility discussed in our previous publication (Cui and Bazan, 2010).

Fig. 4. 50 kDa agrin expression increased in the dorsal horn (DH) after AAV2-Ag50 injection into the SCI–NP rats. A. Western blot analysis (Left) of the DH proteins showed that Ag50 and Ag25 expressions increased after AAV2-Ag50 and AAV2-Ag25 injections, respectively. B. Both Ag50 and Ag25 (Right) were myc positive, confirming their origin from the viral protein wrapped vectors. C. Bar graph (bottom) depicting the quantitative analysis of Ag50 and Ag25 expressions in the DH of sham-operated, AAV2-Ag50, AAV2-Ag25, and AAV2-null injected rats. The difference between AAV2-Ag50 and AAV2-Ag25 or AAV2-null injected rats is significant (n = 4; **P b 0.01; one-way ANOVA, followed by the Tukey's multiple comparison test). D. Western blot. The bands from left to right are: marker, normal, allodynic, AAV2-Ag25 treated, AAV2-Ag50 treated samples. There is no obvious difference among these protein samples. GADPH labeling suggests that protein sample loading was even. E. Western blot after agrin antibody precipitation. The samples were loaded in the same way as in A (50 μl beads, 2 μg agrin antibody, and 400 μg proteins were used for each sample precipitation). After agrin antibody precipitated p-NR1 protein complex, p-NR1 bands showed a clear difference between allodynic, AAV2-Ag25 treated and AAV2-Ag50 treated samples, suggesting that p-NR1 was increased in AAV2-Ag50 treated DH. Although IP-Western blot is not a quantitative test, thicker band of p-NR1 from AAV2-Ag50 treated samples suggests a clear increase.

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Fig. 5. 50 kDa agrin requires specific proteins for GABA neuron activation in the dorsal horn. (A) After incubating agrin antibody with the protein samples from AAV-Ag50 and AAV-null vector treated dorsal spinal cords, the precipitated protein complexes were subjected to two-dimensional gel electrophoresis. The spinal cords treated with AAV2-Ag50 (left) showed distinct proteins (6 circled: neurofilament 200, mitofusin 2, E3 ubiquitin, E3 ubiquitin-protein ligase, Ig gamma 2A, and Geranylgeranyl transferase) different from that treated with AAV2-null (right; 12 circled: beta tubulin 2A, beta tubulin 2B, keratin, dynein, complement C1q, actin alpha, tRNA pseudouridine synthase 1, glycerophosphocholine phosphodiesterase, alpha tubulin 1B, phosphatidylinositol transfer protein alpha, Forkhead box protein M1, and transport protein). (B) Precipitated protein complexes of the dorsal spinal cords treated with AAV2-Ag50, AAV2-null, and normal, were displayed on Western blot membrane, showing increased partial segments of neurofilament 200 and mitofusin 2 solely in the AAV2-Ag50 treated group. (C). Phosphorylated NR1 antibody antibody immunoprecipitation and 2D gel did not show any different protein spots, suggesting that the phosphorylation of NR1 subunits can come from other sources.

We have previously shown that 50 kDa agrin (Ag50) activates GABA neurons via phosphorylation of their NMDA receptor NR1 subunits at serine residue 896/897, to increase GABA expression in the DH (refer to Fig. 5G in J. Neurosci. 30, 15286–15297); and increased GABA suppresses sensory neuron excitation pre- and post-synaptically (Cui et al., 1996, 1997; Stiller et al., 1996; Watanabe et al., 2002), thus pain. To date it is unclear why Ag50 selectively activates GABA neurons and not sensory neurons despite the fact that the NR1 subunit of NMDA receptors is a basic, common structure that can be found in sensory neurons, glial cells, and GABA neurons (Schipke et al., 2001; Xi et al., 2009).

This may suggest that there is a special protein–protein complex structure to ensure that Ag50 only interacts with NR1 subunits in the GABA neurons. Therefore, we explored those molecules that may contribute to the GABA neuron specific activation by Ag50. Using 2D gel and LC-MS, we discovered 6 specific Ag50 associated proteins (two of these have been identified/confirmed as neurofilament heavy chain, mitofusin 2) within the agrin antibody precipitated protein complex of the dorsal spinal cords treated with AAV2-Ag50 (Fig. 5A), while AAV2-null treated samples showed 12 different associated proteins (Fig. 5A). On the IP-western blot membrane, the neurofilament 200

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antibody and mitofusin 2 antibody yielded strong positive bands in the AAV2-Ag50 treated samples, while the two bands were barely detected in the AAV2-null treated samples (Fig. 5B). These data indicate that NF200 and Mfn2 are required by Ag50 to activate solely the GABA neurons. In addition, a light band of NF200 and a light band of Mfn2 were observed in normal rat DH, suggesting that there is naturally occurring phosphorylation of GABA neurons' NR1 subunits during sensory signals modulation under various conditions. Mitofusin 2 is a synaptic protein in control of mitochondrial fission and fusion for energy production (Hoppins et al., 2007), Ca2+ regulation, maintenance of plasma membrane potential, axonal and dendritic transport, and the release and re-uptake of neurotransmitters at synapses (Chan, 2006; Zhang and Chan, 2007). Mfn2 mutation results in axonal neuropathy, such as Charcot–Marie–Tooth (CMT) subtype 2A (CMT2A) (Calvo et al., 2009; Drogemuller et al., 2011) (axonal degradation in both sensory and motor neurons). Mfn2 reduction also induces neurodegenerative conditions, such as Alzheimer's disease (Manczak et al., 2011; Wang et al., 2009). During NR1 subunit activation by Ag50, Mfn2 may act to provide energy needed for phosphorylation and may also be the ‘adhesive’ in stabilizing the Ag50 complex conformation. NF 200 is a cellular structure found mainly in the neuronal axons and perikarya. Neurofilament heavy chains seem to be toxic to neurons, when irregularly accumulated in the axons (Gotow, 2000). NFs and phosphorylated NFs are often used as an indicator for diseases, such as amyotrophic lateral sclerosis (Boylan et al., 2013) and Alzheimer's disease (Rudrabhatla et al., 2010). From our study results, we consider that NF200 serves as a structure conformation adhesive for GABA neuron activation, because this NF200 has a modified structure of about 110 kDa, less than the entire NF of 200 kDa. This suggests that NF200 is reduced to fit the Ag50-induced activation structure. These unique associated proteins are required for Ag50 to solely activate GABA neurons. These roles of NF200 and Mfn2 in GABA neuron activation have not been previously reported. Fig. 6 summarizes the possible interaction between 50 kDa agrin and the associated proteins, such as NF200 and Mfn2, to solely activate GABA neurons. Since NR1 subunit is a basic, common structure of NMDA receptors and these NMDA receptors exist in the sensory neurons, glial cells,

Fig. 6. Proposed mechanism of Ag50 induced GABA neuron activation requires neurofilament 200 (NF200) and mitofusin 2 (Mfn2). When Ag50 interacts with the NR1 subunits of GABA neurons, NF200 and Mfn2 are required for this action. These specific associated proteins, such as NF200 and Mfn2, make a suitable structure to ensure that Ag50 selectively activates the NR1 subunit of the GABA NMDA receptor (this drawing is modified from our previously published figure in the Journal of Neuroscience 2010;30:15286).

and GABA neurons, NMDA receptor antagonists, when used for treating neuropathic pain, will block both sensory neuron and GABA neuron activations. GABA neuron suppression by NMDA receptor antagonists will reduce the inhibition of pain and/or excitation. This view is consistent with the fact that NMDA receptor antagonists used for pain treatment have shown a very limited effect in clinic (Gonda, 2012; Niesters and Dahan, 2012). Therefore, specific NMDA receptor antagonists that only target the NMDA receptor of sensory neurons need to be developed in the future for neuropathic pain treatment. Intra-spinal injections are technically changing and may be associated with complications. Therefore, this approach is typically used as a last resort to treat intractable neuropathic pain. Our further research will test whether peripheral nerve injections of AAV2-Ag50 vectors can express Ag50 in the dorsal horn to suppress pain. If this proves successful, neuropathic pain treatment can be achieved via a nerve branch injection with AAV2-Ag50, as a nerve branch block, a procedure routinely performed by neurosurgeons and anesthesiologists. In summary, 50 kDa agrin is a promising inhibitory molecule for the treatment of SCI and other forms of NP. Ag50 interacts with several specific associated proteins, such as NF200 and Mfn2, to selectively activate GABA neurons and inhibit neuropathic pain after peripheral nerve or spinal cord injury. This mechanism may also apply to other over-excitation states, such as tremor and epilepsy. This study was supported by Department of Anesthesiology, Miller School of Medicine, University of Miami, Wings for Life Spinal Cord Research foundation (WFL-US-002/11), and NIH grant (RO1 DEO2290301). We thank Dr. Zhengzheng Zhang for assisting statistical analysis.

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