Essential role of the synaptic vesicle protein synapsin II in formalin-induced hyperalgesia and glutamate release in the spinal cord

Pain 115 (2005) 171–181 www.elsevier.com/locate/pain

Essential role of the synaptic vesicle protein synapsin II in formalin-induced hyperalgesia and glutamate release in the spinal cord Achim Schmidtkoa, Domenico Del Turcob, Ovidiu Costea, Corina Ehnerta, Ellen Niederbergera, Peter Ruthc, Thomas Dellerb, Gerd Geisslingera, Irmgard Tegedera,* a

Pharmazentrum Frankfurt, Institut fu¨r Klinische Pharmakologie/ZAFES, Klinikum der Johann Wolfgang Goethe-Universita¨t Frankfurt, Theodor-Stern-Kai 7, 60590 Frankfurt am Main, Germany b Institut fu¨r Klinische Neuroanatomie, Johann Wolfgang Goethe-Universita¨t Frankfurt, Theodor-Stern-Kai 7, 60590 Frankfurt am Main, Germany c Pharmazeutisches Institut, Pharmakologie und Toxikologie, Auf der Morgenstelle 8, 72076 Tu¨bingen, Germany Received 26 September 2004; received in revised form 7 February 2005; accepted 22 February 2005

Abstract The synaptic vesicle protein synapsin II plays an important role in the regulation of neurotransmitter release and synaptic plasticity. Here, we investigated its involvement in the synaptic transmission of nociceptive signals in the spinal cord and the development of pain hypersensitivity. We show that synapsin II is predominantly expressed in terminals and neuronal fibers in superficial laminae of the dorsal horn (laminae I–II). Formalin injection into a mouse hindpaw normally causes an immediate and strong release of glutamate in the dorsal horn. In synapsin II deficient mice this glutamate release is almost completely missing. This is associated with reduced nociceptive behavior in the formalin test and in the zymosan-induced paw inflammation model. In addition, the formalin evoked increase in the number of c-Fos IR neurons is significantly reduced in synapsin II knockout mice. Touch perception and motor coordination, however, are normal indicating that synapsin II deficiency does not generally disrupt sensory and/or motor functions. Antisense-mediated transient knockdown of synapsin II in the spinal cord of adult animals also reduced the nociceptive behavior. As the antisense effect is independent of a potential role of synapsin II during development we suggest that the hypoalgesia in synapsin II deficient mice does involve a direct ‘pain-facilitating’ effect of synapsin II and is not essentially dependent on potentially occurring developmental alterations. The distinctive role of synapsin II for pain signaling probably results from its specific localization and possibly from a specific control of glutamate release. q 2005 International Association for the Study of Pain. Published by Elsevier B.V. All rights reserved. Keywords: Synaptic vesicle protein; Spinal cord; Pain; Microdialysis

1. Introduction Stimulation of nociceptors causes a release of glutamate and other excitatory neurotransmitters in the dorsal horn of spinal cord. Ongoing stimulation and sustained glutamate release may result in the development of a state of hyperexcitability of nociceptive neurons, clinically manifesting as hyperalgesia and allodynia. Various, possibly sequential mechanisms are involved in the sensitization process starting with the activation of various * Corresponding author. Tel.:C49 69 6301 7621; fax: C49 69 6301 7636. E-mail address: [email protected] (I. Tegeder).

kinases including protein kinase A (PKA) (Aley and Levine, 1999; Malmberg et al., 1997a), protein kinase C (PKC) (Malmberg et al., 1997b) and MAP kinases (Ji et al., 1999), subsequent release of nitric oxide (Aley et al., 1998; Meller and Gebhart, 1993) and prostaglandins (Geisslinger et al., 2000; Malmberg and Yaksh, 1995) and ultimately long lasting changes of gene expression (Costigan et al., 2002; Woolf and Costigan, 1999). Synaptic vesicle proteins including synapsins have been recognized as targets of PKA and other kinases (Benfenati et al., 1992; Greengard et al., 1993; Hosaka et al., 1999) and phosphorylation of synapsins probably changes synaptic efficiency. The role of synapsins for nociceptive processing is unknown so far.

0304-3959/$20.00 q 2005 International Association for the Study of Pain. Published by Elsevier B.V. All rights reserved. doi:10.1016/j.pain.2005.02.027

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Synapsins are a family of neuron-specific synaptic vesicle phosphoproteins that have been implicated in the regulation of neuronal development and transmitter release. In vertebrates, synapsins are encoded by three distinct genes (synapsin I, II and III) that produce at least five proteins (synapsin Ia, Ib, IIa, IIb, and IIIa) (Valtorta et al., 1992). Synapsin I is preferentially localized at inhibitory synapses (Terada et al., 1999), whereas synapsin II was primarily found in excitatory synapses (Mandell et al., 1992). Synapsin I and II probably have distinct roles in synapse formation (Ferreira et al., 1995, 1998; Han et al., 1991), neurite outgrowth (Kao et al., 2002) and neurotransmitter release (Hilfiker et al., 1999b; Hosaka and Sudhof 1998a,b). It was shown by videomicroscopy that unphosphorylated synapsins anchor synaptic vesicles to the actin-based cytoskeleton thereby maintaining a reserve pool of vesicles (Ceccaldi et al., 1995). Upon phosphorylation, synapsins dissociate from synaptic vesicles and lose their actin binding property (Bahler and Greengard, 1987; Chi et al., 2001; Greengard et al., 1993). The resulting mobilization of reserve vesicles allows to maintain the release of neurotransmitters during strong neuronal activity (Humeau et al., 2001). High activity of nociceptive neurons evoked by strong or persistent nociceptive stimulation (Yaksh et al., 1999) may therefore require a synapsin-mediated supply with reserve vesicles. Synapsin II deficiency in knockout mice causes synaptic depression upon repetitive stimulation in hippocampal neurons and a decrease of post-tetanic potentiation, i.e. a defect of short-term synaptic plasticity (Rosahl et al., 1995). The number of synaptic vesicles distant from the plasma membrane and calcium-evoked glutamate release is also reduced in mice lacking synapsins (Hilfiker et al., 1999a; Li et al., 1995). Based on these characteristics and the similarities between short-term synaptic plasticity and sensitization of nociceptive neurons we hypothesized that synapsins, and particularly synapsin II, may be involved in the synaptic transmission of nociceptive stimuli. We therefore studied the localization and regulation of synapsin II expression in the dorsal horn of the spinal cord and assessed the effects of complete and transient synapsin II deficiency on the nociceptive behavior in various models.

2.2. Intrathecal delivery of synapsin II antisense oligonucleotides Because of the difficulty of a continuous spinal delivery in mice, antisense experiments were done with rats. Synapsin II protein expression in the lumbar spinal cord was suppressed with an antisense oligonucleotide (5 0 -CGA CCA AAG GTG GTC CGC GTC TC-3 0 ) previously shown to knockdown synapsin II protein in vitro (Ferreira et al., 1994). The oligonucleotide was continuously delivered onto the lumbar spinal cord through a spinal catheter (polyethylene tube, ID 0.28 mm, OD 0.61 mm, length 5–7 cm). The tip of the catheter was placed above L4/L5. The catheter was attached to an osmotic mini pump (Alzet model 2001, Durect Corp., Cupertino, USA) which was implanted into the subcutaneous space in the tail region. Pumps contained either oligonucleotide (sense or antisense 5 mg/ml, BioSpring, Frankfurt, Germany) or artificial cerebrospinal fluid (ACSF) which was used as vehicle. The infusion rate was 1.0 ml/h and was continued for 6 days. During the last hour of the infusion the formalin test was performed. At completion, the position of the catheter was confirmed, the lumbar spinal cord (L3–L5) dissected, snap frozen in liquid nitrogen and kept at K808C until Western Blot analysis. 2.3. Spinal cord microdialysis in mice Microdialysis was employed to assess glutamate release in the dorsal horns of the spinal cord in wild type and synapsin II knockout mice. A dialysis probe was constructed from a polyacrylonitrile hollow fiber (AN69, : 200 mm; molecular mass cutoff w40 kDa; Hospal, Nu¨rnberg, Germany), a polyethylene (PE) inlet tube (inner : 0.26 mm, outer : 0.6 mm) and a micropin for insertion. The catheter was inserted during isoflurane anesthesia through the intervertebral joints between vertebra Th13 an L1 as previously described (Ates et al., 2003). For dialysis, anesthesia was switched to urethane (1.2 g/kg i.p.). The catheter was connected to a CMA 100 microdialysis pump and perfused with artificial cerebrospinal fluid (ACSF, 141.7 mM NaC, 2.6 mM KC, 0.9 mM Mg2C, 1.3 mM Ca2C, 122.7 mM ClK, 21.0 mM 2K HCOK 3 , 2.5 mM HPO4 , and 3.5 mM dextrose, bubbled with 5% CO2 in 95% O2 to adjust pH to 7.2) at a flow rate of 1.5 ml/min. Dialysates were collected directly from the hollow fiber. After equilibration for 2 h, baseline dialysates were sampled for 12! 5 min intervals. Formalin was then injected into the hindpaws and dialysates were collected for further 24!5 min. Dialysates were kept on ice and analyzed for glutamate within 15 min of sampling. A CMA 600 Analyzer (CMA, Stockholm, Sweden) was used according to the manufacturer’s instructions. At completion, the correct placement of the microdialysis catheter (dorsal horn, L4) was confirmed by microscopic inspection.

2. Material and methods 2.4. Formalin test 2.1. Animals Sex and age matched, 4–6-week-old synapsin II knockout and wild type mice (strain B6,129S-Syn2tm1Sud and B6129SF2/J, respectively; JAX Research Systems, Bar Harbor, USA), and male Sprague–Dawley rats (Charles River, Sulzfeld, Germany) were used. The ethics guidelines for investigations in conscious animals were obeyed and the experiments were approved by the local Ethics Committee for Animal Research.

Animals were habituated to the observation chamber. In mice, 15 ml of a 5% formaldehyde solution (formalin) was injected subcutaneously (s.c.) into the dorsal surface of one hindpaw. The time spent licking the formalin-injected paw was recorded in 5 min intervals up to 45 min after formalin injection. In rats, 50 ml 5% formaldehyde was used and flinches were counted in 1 min intervals up to 60 min starting directly after formalin injection. Flinches of 5 min intervals were summarized as mean flinches per

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minute. The observer was unaware of treatments or genetic background. 2.5. Zymosan-induced mechanical hyperalgesia The mechanical sensitivity of the plantar side of the hindpaw was assessed with a series of calibrated von Frey hairs (8, 20, 40, 70, 160, 400, 600, 1000, 1400, 2000, 4000 mg/mm2; Stoelting Corp., Wood Dale, USA). Stimuli were applied in increasing order (five repetitions each) until a strong and immediate withdrawal occurred. This threshold was confirmed after a short break. For statistical analysis, mechanical thresholds were log transformed. Following four baseline measurements, 15 ml zymosan suspension (5 mg/ml in 0.1 M PBS pH 7.4) was injected into the plantar subcutaneous space of one hindpaw and von Frey thresholds were determined at 1, 2, 3, 4, 5, 6, and 8 h after zymosan injection by a blinded observer. 2.6. Rotarod test Motor coordination was assessed with a Rotarod Treadmill for mice (Ugo Basile, Comerio, Italy) at a constant rotating speed of 32 rpm. All mice had three training sessions before the day of the experiment. The fall-off latency was averaged from three tests. The cut-off time was 120 s. 2.7. Western Blot analysis Tissue samples were homogenized in buffer containing 10 mM Tris/HCl, pH 7.4, 20 mM 3-((3-cholamidopropyl)-dimethylammonio)-1-propane-sulfonate (CHAPS), 0.5 mM ethylene-diaminetetraaceticacid (EDTA), 1 mM dithiothreitole (DTT), 0.5 mM phenylmethylsulfonylfluoride (PMSF) and 1 mM Pefabloce SC (Alexis, Gru¨nberg, Germany). Extracted proteins (30 mg) were separated by SDS-polyacrylamide gel electrophoresis, transferred onto nitrocellulose membranes (Amersham Pharmacia Biotech, Freiburg, Germany) by electro-blotting and incubated with primary antibodies directed against synapsin II (1:1000, StressGen, Victoria, Canada), cGMP dependent protein kinase I (PKG-I, 1:500, Huber et al., 2000) or extracellular signal-regulated kinase 2 (ERK-2, 1:10000, Santa Cruz, Heidelberg, Germany) according to standard procedures. ERK-2 was used as loading control (Ji et al., 1999; Tegeder et al., 2001). After incubation with the secondary antibody conjugated with Alexa Fluor 680 (dilution 1:10,000, Molecular Probes, Eugene, USA), blots were visualized on the Odyssey Infrared Imaging System (LI-COR Biosciences, Bad Homburg, Germany).

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(1:500, Huber et al., 2000), substance P (1:500, Santa Cruz Biotechnology), calcitonin gene related peptide (CGRP, 1:1000, Chemicon, Hofheim, Germany), c-Fos (1:500, Santa Cruz), nuclear neuronal protein (NeuN, 1:1000, Chemicon), glial fibrillary acid protein (GFAP, 1:1000, Chemicon), neurofilament 200 (NF200, 1:1000, Chemicon) and neuronal nitric oxide synthase (nNOS, 1:500, Alexis Biochemicals, Gru¨nberg, Germany). For colocalization of synapsin II with substance P we used a mouse monoclonal antibody to SP (1:500, abcam, Cambridge, UK). Binding sites were visualized with Alexa Fluor 488 or Alexa Fluor 568 conjugated species specific secondary antibodies (Molecular Probes). After immunostaining, slides were incubated for 10 min at RT in 0.3% Sudan black B in 70% ethanol to reduce lipofuscin-like autofluorescence (Schnell et al., 1999). Griffonia simplicifolia isolectin B4 (IB4) conjugated to Alexa Fluor 568 (1:500, Molecular Probes) was used for staining of unmyelinated C-fiber afferents in lamina II (Ritter et al., 2000). Confocal images (optical sections: 1 mm) were obtained using a Zeiss LSM 510 confocal laser scanning microscope (Zeiss, Go¨ttingen, Germany). For quantification of c-Fos upregulation, immunoreactive nuclei in the ipsilateral dorsal horn were counted by a blinded observer. Five random sections spanning the L4 segment from each mouse (nZ5 in each group) were counted and the individual mean was used for statistical analysis. 2.9. Statistical analysis Statistical evaluation was done with SPSS 11.0.1 for Windows. Data are presented as the meanGstandard error (SEM). For the formalin test in mice, the licking time of phase one and two (1–10 min, 11–45 min, respectively) was analyzed with the Student’s t-test. In rats, the sum of flinches of phase one (1–10 min) and phase two (11–60 min) was submitted to univariate analysis of variance (ANOVA) and subsequent t-tests employing a Bonferroni alpha correction for multiple comparisons. For analysis of mechanical hyperalgesia, areas under the ‘von Frey threshold (log transformed)’ versus ‘time’ curve were calculated using the linear trapezoidal role and submitted to t-tests. t-Tests were also used to compare baseline mechanical von Frey thresholds and the number of c-Fos IR cells. Differences in glutamate release were assessed by submitting AUCs to t-tests. The rotarod performance was assessed using the Mann–Whitney U-test and expressed as median and interquartile range of the fall-off latency. For all tests, a probability value P!0.05 was considered as statistically significant.

2.8. Immunofluorescence

3. Results

Mice were intracardially perfused with 0.9% saline followed by 4% paraformaldehyde in 0.1 M phosphate buffered saline (PBS, pH 7.4) under deep pentobarbital anesthesia (300 mg/kg). The spinal cord (lumbar enlargement) was removed, post-fixed in the same fixative overnight (4 8C) and 30 mm thick transversal sections were cut on a vibratome. For c-Fos IR quantification, 18 mm sections were used. Sections were incubated in blocking buffer (5% normal goat serum, 0.3% Triton X-100 in 0.1 M PBS) for 30 min at room temperature and incubated overnight at 4 8C with primary antibodies directed against synapsin II (1:500, StressGen), PKG-I

3.1. Expression of synapsin II in mouse spinal cord Immunofluorescence studies of lumbar spinal cord sections show that synapsin II is primarily expressed in neuronal fibers in superficial laminae I and II of the dorsal horn (Fig. 1(A)) i.e. in the areas where A-delta (lamina I, V) and C-fibers (lamina II) terminate. Synapsin II immunoreactivity colocalizes with the binding of Griffonia simplicifolia isolectin B4 (IB4, Fig. 1(B) and (C)) which

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Fig. 1. (A) Dorsal horn of the lumbar spinal cord in wild type mice immunostained for synapsin II (red) and the neuron-specific nuclear protein NeuN (green). (B, C) Colocalization of synapsin II (green) and Griffonia simplicifolia isolectin B4 (IB4, red). (D, E) Colocalization of synapsin II (green) and substance P (SP, red). C and E are confocal higher magnifications of B and D, respectively. Double labeled terminals (fibers) are coded in yellow (arrows). Scale bars: 100 mm (A, B, D); 10 mm (C, E).

specifically labels unmyelinated primary afferents in a dense sheet of the inner lamina II (IIi). C-fiber afferents in this region have been associated with pain signaling (Ritter et al., 2000). Synapsin II expression, however, is not restricted to this dense area but also extends to the outer lamina II and lamina I. Some synapsin II immunoreactive primary afferent terminals also show immunoreactivity for substance P (SP) (Fig. 1(D) and (E)) which labels peptidergic primary afferents of nociceptive neurons (Todd et al., 2002).

Western Blot analysis of synapsin II shows a significant increase of synapsin II protein levels in lumbar spinal cord in response to zymosan injection into a hindpaw (Fig. 2) suggesting that synapsin II contributes to the maladaptive response that occurs in the dorsal horn in response to ongoing painful inflammatory stimuli. Changes in gene expression are considered to contribute to the development of chronic pain (Woolf and Costigan, 1999). Only the b-isoform of synapsin II was detectable in protein extracts of the spinal

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cord employing the commercially available synapsin II antibody that is directed against an N-terminal peptide (AA52-65) of synapsin IIa (74 kDa) and IIb (54 kDa). 3.2. Immunofluorescence of marker proteins in synapsin II knockout mice Fig. 2. Western Blot analysis of synapsin IIb (54 kDa) in lumbar spinal cord in naive mice (left) and after zymosan injection into a hindpaw (right). Synapsin IIa (74 kDa) was not detectable. ERK-2 is used as loading control.

To further assess the role of synapsin II in nociceptive pathways, we analyzed the cyto- and fiber architecture of the spinal cord in synapsin II deficient mice using immunostaining for several marker proteins. The lack of synapsin II in knockout mice was confirmed (Fig. 3(A) and (B)). Immunostaining of neuronal nuclei, fibers and astrocytes

Fig. 3. Immunostaining for synapsin II and ‘pain-associated’ marker proteins in lumbar spinal cord that are not altered in synapsin II knockout mice. The left panel shows wild type mice (A, C, E, G); the right panel shows synapsin II knockout mice (B, D, F, H). (A, B) Synapsin II, (C, D) Substance P; (E, F) calcitonin gene related peptide (CGRP); (G, H) neuronal nitric oxide synthase (nNOS). Scale bars: 100 mm.

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employing antibodies directed against neuronal nuclear protein (NeuN), neurofilament 200 (NF200) and glial fibrillary acid protein (GFAP), respectively, did not reveal any differences between wild type and synapsin II knockout mice (data not shown). We observed also no differences in substance P, calcitonin gene related peptide (CGRP) and neuronal nitric oxide synthase (nNOS) immunoreactivity between synapsin II knockout and wild type mice (Fig. 3(C)–(H)). Isolectin B4 staining of unmyelinated C-fiber afferents in the inner lamina II (IIi) appears to be reduced in synapsin II knockout mice and shows a slightly different pattern (Fig. 4(A) and (B)). Immunoreactivity for cGMP dependent protein kinase-1 (PKG-I) which labels primary afferents in the dorsal horn (Qian et al., 1996; Schmidt et al., 2002) is also reduced in synapsin II knockout

mice (Fig. 4(C) and (D)). Western Blot analysis confirms a quantitative reduction of PKG-I protein expression in synapsin II knockout mice (Fig. 4(E)). In addition, we show that the reduction of PKG-I protein in synapsin II knockout mice only occurs in the spinal cord but not in the cerebellum and heart (Fig. 4(F)). 3.3. Sensory and motor function in synapsin II knockout mice We next assessed effects of synapsin II deficiency on somatosensory functions and motor coordination. The von Frey threshold of naı¨ve synapsin II knockout mice (3.14G0.12 g) is similar to that of naı¨ve wild type mice

Fig. 4. (A, B) Griffonia simplicifolia isolectin B4 (IB4) staining of unmyelinated neuronal fibers in the dorsal horn of the spinal cord in wild type (A) and synapsin II knockout mice (B). (C, D) Immunostaining for cGMP dependent protein kinase I (PKG-I) in wild type (C) and synapsin II knockout mice (D). PKGI immunoreactivity is slightly reduced in synapsin II knockout mice. Scale bars: 100 mm. (E) Western Blot analysis of PKG-I in lumbar spinal cord in naı¨ve wild type (left) and naı¨ve synapsin II knockout mice (right). ERK-2 is used as loading control. The densitometric analysis reveals a significant difference with P!0.05, indicated with the asterisk. (F) Western Blot analysis showing the expression of synapsin II and PKG-I in spinal cord (SC), cerebellum (C) and heart (H) in wild type and synapsin II knockout mice.

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(2.95G0.06 g; PZ0.102) indicating normal perception of touch stimuli. The performance in the rotarod test is also not altered in synapsin II knockout mice indicating an intact motor coordination. The median fall-off latency is 53.3 s in knockout mice (interquartile range 23.0–96.8 s, nZ5) and 42.3 s in wild type mice (interquartile range 24.0–76.3 s, nZ11; PZ0.73. 3.4. Glutamate release and nociceptive behavior in synapsin II knockout mice To evaluate the role of synapsin II for nociceptive transmission we investigated: (i) the formalin-evoked glutamate release in the dorsal horn, (ii) formalin-evoked nociceptive behavior and (iii) formalin-induced c-Fos upregulation in the dorsal horn. c-Fos expression in neurons is a general indicator of neuronal activity. Its upregulation in

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dorsal horn neurons following nociceptive stimulation is considered to correlate with pain intensity. To ascertain that effects are not restricted to the formalin stimulus, we additionally studied inflammatory hyperalgesia in a model of chronic paw inflammation. Baseline glutamate release is similar in synapsin II knockout and wild type mice (Fig. 5(A)). As expected, formalin injection into the hindpaw causes an immediate and strong release of glutamate in the dorsal horn of wild type mice. In contrast, formalin evokes almost no glutamate increase in knockout mice. The area under the ‘glutamate versus time’ curve comprising the first 45 min after formalin injection (AUC0–45 min) is 575.9G77.1 mmol/L min in wild type mice and 283.5G41.9 mmol/L min in knockout mice. The difference is statistically significant (PZ0.01). In line with the inhibition of formalin-evoked glutamate release, synapsin II knockout mice spend less time licking the formalin injected paw than wild type mice in both phases of

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80 60 40 20 0 0

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Fig. 5. (A) Time course of glutamate concentrations in spinal cord microdialysates in wild type (:) and synapsin II knockout mice (B) before and after formalin injection into the hindpaws. Formalin was injected at time ‘zero’. Data are expressed as meanGSEM. nZ5 in each group. (B) Time course of the licking behavior of synapsin II knockout and wild type mice after formalin injection into a hindpaw. Data are expressed as mean licking timeGSEM. nZ8 in each group. (C, D) Immunostaining of c-Fos in the ipsilateral dorsal horn of the lumbar spinal cord (L4) 2 h after formalin injection into a hindpaw in wild type (C) and synapsin II knockout mice (D). (E) Number of c-Fos IR nuclei in the ipsilateral dorsal horn (L4) after formalin injection (nZ5 in each group). The asterisk indicates a statistically significant difference with P!0.05.

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120 SynII -/Wild type

% of threshold

100 80 60 40 20 0

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Time [h] Fig. 6. Time course of the mechanical nociceptive threshold of synapsin II knockout and wild type mice following zymosan injection into a hindpaw. Mechanical thresholds were assessed with von Frey hairs. Data are expressed as percent of the baseline threshold set at 100% (meanGSEM). nZ6 in each group.

the formalin test (Fig. 5(B)). The mean licking time in the early phase is 109.7G18.5 s (wild type) and 59.2G11.8 s (knockout), P!0.05. In the second phase the licking time is 328.0G57.2 s (wild type) and 128.9G86.2 s (knockout), P!0.001, nZ8 in each group. Interestingly, the time of the formalin-evoked nociceptive behavior exactly coincides with the increase of glutamate release. As expected, the injection of formalin into a hindpaw causes a strong increase of the number of c-Fos immunoreactive nuclei in the ipsilateral dorsal horn in wild type mice. The number of c-Fos positive neurons is significantly reduced in synapsin II knockout mice (Fig. 5(C)–(E)). In the zymosan induced paw inflammation model synapsin II knockout mice develop only minor mechanical hyperalgesia, i.e. the threshold to elicit a withdrawal response drops only by about 20% as compared to baseline. In wild type mice the nociceptive threshold decreases by about 70% (Fig. 6). AUCs are significantly different (wild type mice 437.7G5.58% h versus knockout mice 680.0G 22.8% h; P!0.001, nZ6 in each group). 3.5. Nociceptive behavior in rats treated with synapsin II antisense oligonucleotides To evaluate the relative contribution of developmentdependent and -independent effects we additionally assessed the formalin-evoked nociceptive behavior in adult rats where synapsin II protein expression in lumbar spinal cord was knocked down by means of antisense treatment. As a continuous spinal infusion over several days is not feasible in mice, we performed these experiments in adult rats. The localization of synapsin II in the dorsal horn and its upregulation following nociceptive stimulation is similar in rats and mice (data not shown). Synapsin II antisense treatment results in a clear reduction of synapsin II protein levels in rat lumbar spinal cord compared with ACSF treated control animals (Fig. 7(A)). Sense treatment

Fig. 7. Effects of a transient knockdown of synapsin II protein in rat lumbar spinal cord with continuous spinal infusion of a synapsin II antisense oligonucleotide for 6 days. Control animals were treated with vehicle (ACSF, artificial cerebrospinal fluid) or the corresponding sense oligonucleotide. A: Western Blot analysis of synapsin IIb (54 kDa) in the spinal cord following antisense infusion. Synapsin IIa (74 kDa) is not detected. ERK-2 is used as loading control. Representative results of three experiments. The densitometric analysis reveals a significant difference with P!0.05, indicated with the asterisk. (B) Time course of the formalin-induced flinching behavior. The formalin test was performed during the last hour of the 6-day i.t. infusion. Groups consisted of six (antisense, ACSF) or five (sense, ACSF) animals. Data are mean flinches per minuteGSEM. (C) Western Blot analysis of cGMP dependent protein kinase-I (PKG-I) in the spinal cord following oligonucleotide infusion. ERK-2 is used as loading control.

has no effect. Antisense treated rats show significantly reduced flinching behavior in the second phase of the formalin test (sum of flinches 237.3G43.2), compared with controls (535.6G37.1) and sense-treated animals (456.8G

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26.3). Differences between antisense and control or sense are statistically significant with P!0.05 (Fig. 7(B)). The sum of flinches in the early phase is similar between groups (ACSF, 57.6G8.7; sense, 76.6G9.8 and antisense 40.0G15.1). In contrast to synapsin II knockout mice, PKGI protein expression in the dorsal horn remains unaltered in synapsin II antisense treated rats (Fig. 7(C)).

4. Discussion In previous studies, synapsin II deficiency caused defects in synaptic plasticity (Rosahl et al., 1995) that are associated with learning deficits (Silva et al., 1996). As the sensitization of the nociceptive system is generally considered to be a ‘memory-like’ process (Dubner and Ruda, 1992; Sandkuhler and Liu, 1998) we hypothesized that synapsin II deficiency may be associated with reduced pain. We show here that synapsin II is predominantly expressed in superficial laminae of the dorsal horn of the spinal cord where nociceptive A-delta and C-fibers terminate and its expression increases after nociceptive stimulation. The latter suggests that upregulation of synapsin II contributes to the maladaptive responses of dorsal horn neurons to ongoing peripheral painful stimuli caused by tissue injury. Synapsin II deficient mice show almost no formalin-induced glutamate release in the dorsal horn and much less c-Fos upregulation indicating that significantly fewer nociceptive neurons are activated by the formalin stimulus. Accordingly, synapsin II deficient mice show reduced nociceptive behavior. In contrast, touch perception and motor coordination are not altered in these mice. Hence, mechanoreceptive sensory and motor neuron synapses are apparently intact. We therefore suggest that the observed reduction of nociception is not simply part of a general defect of synaptic transmission. This is also supported by the mild phenotype of synapsin II knockout mice which are fertile with no gross anatomical abnormalities (Rosahl et al., 1995), normal body weight, growth and social behavior. The animals only show some learning deficits (Silva et al., 1996) and experience seizures, particularly at old age (O3 months) (Rosahl et al., 1995). Hence, the here observed alteration of nociceptive behavior appears to be caused by relative selective changes at nociceptive synapses. The subtle alteration of Griffonia simplicifolia isolectin B4 (IB4) binding and the slight reduction of cGMP dependent protein kinase I (PKG-I) but normal expression of other ‘pain-mediators’ including calcitonin gene-related peptide (CGRP), substance P, and neuronal nitric oxide synthase suggest that C-fiber synapses are primarily involved. Isolectin B4 is a marker for primary nociceptive neurons with unmyelinated axons that are glial cell-derived neurotrophic factor (GDNF)responsive (Ahluwalia et al., 2002; Bennett et al., 1998; Molliver et al., 1997). PKG-I also labels primary afferent C-fibers. The slight reduction of its expression might

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contribute to the reduced nociception in synapsin II knockout mice because PKG-I is important for axon growth and guidance of C-fibers in the dorsal horn (Schmidt et al., 2002) and the development of hyperalgesia (Schmidtko et al., 2003; Tegeder et al., 2004). However, our antisense experiments show that a knockdown of synapsin II without change of PKG-I expression is sufficient to reduce the nociceptive behavior in adult animals. Hence, the deficiency of synapsin II is essential for the observed hypoalgesia in synapsin II knockout mice, but not the PKG-I reduction. As synapsin II and PKG-I are both involved in the regulation of glutamate release deficiency of one may reduce requirements of the other. Interestingly, PKG-I expression does not decrease in the cerebellum where PKG-I mediates long-term depression. Hence, PKG-I is only reduced at the site where synapsin II and PKG-I cooperate. Previous studies show that synapsins regulate the supply with reserve synaptic vesicles (Chi et al., 2001, 2003) that are needed to maintain transmitter release in case of a sustained stimulation. In our experiments, synapsin II regulates the immediate and the protracted glutamate release. This may be mediated through a regulation of the number of available synaptic vesicles, their glutamate content or the release mechanism (Ferreira et al., 1994; Sugiyama et al., 2000). The latter might result from a cooperation of synapsin II with PKG-I. The upregulation of synapsin II following formalin or zymosan treatment may maintain or enhance the acute effects and may be involved in long-term hyperexcitability of nociceptive neurons. Possibly, more synapsin II allows for more glutamate release. Although we also find some colocalization of synapsin II with substance P, staining for SP was unaltered in synapsin II knockout mice suggesting that peptidergic synapses are less affected. However, we did not analyze SP release in the dorsal horn because microdialysis recovery is low and the available enzyme assays are not sensitive enough to reliably measure concentrations in dialysates. Hence, it is conceivable that SP and/or CGRP release is also reduced in synapsin II knockout mice. It is unclear at present how synapsin II facilitates glutamate release. Most mechanisms that were attributed to synapsins including actin bundling (Bahler and Greengard, 1987; Petrucci and Morrow 1987), ATP binding (Brautigam et al., 2004; Hosaka and Sudhof, 1998b), small G-protein interactions (Giovedi et al., 2004a,b) and RGS protein (regulator of G-protein signaling) inhibition (Tu et al., 2003) were demonstrated for synapsin I but not or not yet for synapsin II and it is unclear which functions are shared by all synapsins and which are specific. We show here that synapsin II plays a unique role for nociceptionevoked glutamate release, hyperalgesia and c-Fos expression in the dorsal horn. This is probably at least in part due to a special localization and possibly special function of this synapsin.

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Acknowledgements This study was supported by the Deutsche Forschungsgemeinschaft (SFB 553 (C6), DFG TE322_2-1 and SFB 269 (B7)).

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