Contribution of presynaptic HCN channels to excitatory inputs of spinal substantia gelatinosa neurons

Accepted Manuscript Contribution of Presynaptic HCN Channels to Excitatory Inputs of Spinal Substantia Gelatinosa Neurons S.-C. PENG, J. WU, D.-Y. ZHANG, C.-Y. JIANG, C.-N. XIE, T. LIU PII: DOI: Reference:

S0306-4522(17)30451-7 http://dx.doi.org/10.1016/j.neuroscience.2017.06.046 NSC 17858

To appear in:

Neuroscience

Received Date: Revised Date: Accepted Date:

13 April 2017 19 June 2017 23 June 2017

Please cite this article as: S.-C. PENG, J. WU, D.-Y. ZHANG, C.-Y. JIANG, C.-N. XIE, T. LIU, Contribution of Presynaptic HCN Channels to Excitatory Inputs of Spinal Substantia Gelatinosa Neurons, Neuroscience (2017), doi: http://dx.doi.org/10.1016/j.neuroscience.2017.06.046

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CONTRIBUTION

OF

PRESYNAPTIC

HCN

CHANNELS

TO

EXCITATORY INPUTS OF SPINAL SUBSTANTIA GELATINOSA NEURONS S.-C. PENG,

a†

J. WU,

a†

D.-Y. ZHANG,

b

C.-Y. JIANG,

c

C.-N. XIE

a

AND T. LIU

a,c,d* a

Department of Pediatrics, the First Affiliated Hospital of Nanchang University,

Nanchang, Jiangxi 330006, China b

Department of Pain Clinic, the First Affiliated Hospital of Nanchang University,

Nanchang, Jiangxi 330006, China c

Jisheng Han Academician Workstation for Pain Medicine, Nanshan Hosptial,

Shenzhen 518052, China d

Jiangxi Key Laboratory of Molecular Diagnostics and Precision Medicine,

Nanchang, Jiangxi 330006, China *

Corresponding author. Present address: Department of Pediatrics, the First

Affiliated Hospital of Nanchang University, Nanchang, Jiangxi 330006, China. Tel: +86-791-88692139; fax: +86-791-88623275. Email address: [email protected] (T. Liu). †

These authors contributed equally to this work.

Abbreviations: DRG, dorsal root ganglia; FSK, forskolin; GAD67, glutamic acid decarboxylase-67;

GFP,

green

fluorescent

protein;

HCN

channels,

hyperpolarization-activated cyclic nucleotide-gated channels; Ih, h-current; IC50, half-maximal

inhibitory

concentration;

IR,

immunoreactivity;

KS

test,

Kolmogorov–Smirnov test; mEPSCs, miniature excitatory postsynaptic currents; PB,

phosphate buffer; SD rats, Sprague–Dawley rats; sEPSCs, spontaneous

excitatory postsynaptic currents; SG, substantia gelatinosa; TTX, tetrodotoxin; 1

VGLUT2,

vesicle

glutamate

transporter

type

II;

ZD7288,

4-ethylphenylamino-1,2-dimethyl-6- methylaminopyrimidinium chloride. Abstract—Hyperpolarization-activated

cyclic

nucleotide-gated

(HCN)

channels are pathological pain-associated voltage-gated ion channels. They are widely expressed in central nervous system including spinal lamina II (also named the substantia gelatinosa, SG).

Here, we examined the

distribution of HCN channels in glutamatergic synaptic terminals as well as their role in the modulation of synaptic transmission in SG neurons from SD rats and GAD67-GFP mice. We found that the expression of the HCN channel isoforms was varied in SG. The HCN4 isoform showed the highest level of co-localization with VGLUT2 (23 ± 3%). In 53% (n = 21/40 neurons) of the SG neurons examined in SD rats, application of HCN channel blocker, ZD7288 (10 M), decreased the frequency of spontaneous (s) and miniature (m) excitatory postsynaptic currents (EPSCs) by 37 ± 4% and 33 ± 4%, respectively. Consistently, forskolin (an activator of adenylate cyclase) significantly increased the frequency of mEPSCs by 225 ± 34%, which could be partially inhibited by ZD7288. Interestingly, the effects of ZD7288 and forskolin on sEPSC frequency were replicated in non-GFP-expressing neurons, but not in GFP-expressing GABAergic SG neurons, in GAD67-GFP transgenic C57/BL6 mice. In summary, our results represent a previously unknown cellular mechanism by which presynaptic HCN channels, especially HCN4, regulate the glutamate release from presynaptic terminals that target excitatory, but not inhibitory SG interneurons. Key words: Hyperpolarization-activated cyclic nucleotide-gated channels; substantia

gelatinosa;

excitatory

postsynaptic

currents;

whole-cell

patch-clamp. 2

INTRODUCTION Hyperpolarization-activated cyclic nucleotide-gated (HCN) channels belong to the pore-loop cation channel superfamily (Postea and Biel, 2011) and encompass four homologous isoforms (HCN1-HCN4) (Ludwig et al., 1998), in which HCN2 and HCN4 are cAMP-sensitive. They are broadly distributed throughout the nervous system including primary (located in the dorsal root ganglia, DRG) and secondary (located in the superficial spinal dorsal horn) nociceptive neurons (Wainger et al., 2001; Antal et al., 2004; Papp et al., 2006; Vasilyev et al., 2007; Papp et al., 2010; Liu et al., 2015a). Upon activation by either membrane hyperpolarization or cAMP, HCN channels generate an inward mixed Na+-K+ current (Ih) (Mayer and Westbrook, 1983; McCormick and Pape, 1990). Moreover, a fraction of HCN channels are tonically open near the resting membrane potential (RMP), indicating a critical role in modulation of neuronal excitability. Indeed, animal models of chronic pain are usually associated with an over-activation or an increase in expression of HCN channels (Takasu et al., 2010; Emery et al., 2011; Weng et al., 2012; Schnorr et al., 2014; Smith et al., 2015; Ding et al., 2016; Zhang et al., 2016). The use of HCN channel blocker 4-ethylphenylamino-1,2-dimethyl-6-methylaminopyrimidinium chloride (ZD7288) is efficient in the treatment of formalin-induced inflammatory pain and nerve injury-induced neuropathic pain, probably due to a reduction in excitatory inputs to the lamina II of spinal dorsal horn (also known as substantia gelatinosa, SG). For instance, monosynaptic excitatory postsynaptic currents (EPSCs) mediated by Aand/or C-fibers or miniature (m) EPSC frequency (a marker of presynaptic release of glutamate) was decreased by ZD7288 in a sub-population of SG neurons (Takasu et al., 2010). These findings raise the following question: Are HCN channels expressed in all the glutamatergic synaptic terminals in SG? If so, what type of neurons do these terminals innervate? Given the important role of SG in 3

nociception, a better understanding of the distribution and function of HCN channels in SG will be crucial to the development of pharmacologic therapies for chronic pain. Our previous results have indicated that Ih can be recorded from sub-populations of SG neurons (Liu et al., 2015b; Hu et al., 2016). In this study, we performed immunofluorescence staining with isoform-specific antibodies and whole-cell

patch-clamp

recordings

of

EPSCs

in

SG

neurons

from

Sprague–Dawley (SD) rats and GAD67-GFP mice. And importantly, we found that HCN channels were expressed in ~52% excitatory presynaptic terminals and their postsynaptic targets in SG were excitatory but not inhibitory neurons.

EXPERIMENTAL PROCEDURES Ethical approval All experimental procedures were approved by the Animal Ethics Committee of Nanchang University. Experiments were designed to minimize animal suffering and to reduce animal numbers. Animals In this study, juvenile (3–5 weeks old) male SD rats and GAD67-GFP transgenic C57/BL6 mice were used. In GAD67-GFP mice, Green Fluorescent Protein (GFP) is selectively expressed in GABAergic interneurons (Tamamaki et al., 2003; Pan et al., 2009; Song et al., 2013). All animals were fed ad libitum and housed in a 12:12–hour light–dark cycle. Cells from at least three different animals of the aforementioned genotype were used in each experiment. Spinal cord slice preparation

4

Spinal cord slices were prepared as previously described (Tong and MacDermott, 2014; Liu et al., 2015b; Hu et al., 2016; Peng et al., 2016) . Animals were deeply anesthetized with urethane (1.5 g/kg, i.p.) and were transcardially perfused with ice-cold carbogenated (95% O2 and 5% CO2) dissection solution containing (in mM) 240 sucrose, 2.5 KCl, 3.5 MgCl2, 0.5 CaCl2, 1.25 NaH2PO4, 0.4 ascorbic acid, 2 pyruvate, 25 NaHCO3, and 1 kynurenic acid. The lumbosacral sections were quickly extracted and immersed in the same solution. The animals were then sacrificed by exsanguination while still under anesthesia. After removing the pia-arachnoid membrane and the roots, the spinal cord was mounted on a vibratome (VT1000S; Leica, Nussloch, Germany) cutting stage. Transverse slices (400–500 m thick) were cut and placed in an incubator filled with carbogenated Krebs solution at 32°C for at least 30 min. The Krebs solution contained (in mM) 117 NaCl, 3.6 KCl, 2.5 CaCl2, 1.2 MgCl2, 1.2 NaH2PO4, 25 NaHCO3, 11 D–glucose, 0.4 ascorbic acid, and 2 pyruvate. Patch-clamp recordings Whole-cell voltage-clamp recordings from SG neurons were obtained as described previously (Peng et al., 2016). In brief, one spinal cord slice was moved to the recording chamber and continuously perfused with Krebs solution at room temperature (23–25°C) with a perfusion rate of 2–4 ml/min. Recording electrodes were pulled from borosilicate glass (1.5 mm OD, 1.12 mm ID; World Precision Instruments, Sarasota, FL, USA) with a micropipette puller (P-97; Sutter Instrument, Novato, CA, USA). The typical resistance of the pipette was 3–6 MΩ when filled with intracellular solution. The intracellular solution contained the following (in mM): 130 K-gluconate, 5 KCl, 10 phosphocreatinine, 0.5 EGTA, 10 HEPES, 0.3 Li-GTP, 4 Mg-ATP (pH = 7.3, adjusted with KOH, 295 mOsm). Signals were amplified with an EPC-10 amplifier and Patchmaster software 5

(HEKA Electronik, Lambrecht, Germany). The holding potential for recording EPSCs was –70 mV (at which voltage GABAergic and/or glycinergic inhibitory postsynaptic currents are negligible). The SG neurons in transverse spinal slices were visualized with an infrared differential interference contrast (IR-DIC) camera (IR-1000; Dage, Michigan City, IN, USA). GFP positive neurons were identified by using a monochromator (Polychrome 5000, Till Photonics, Germany) with excitation light at 488 nm. Series resistances were typically measured between 10 and 30 MΩ and were monitored throughout the recording period. Data were excluded if the series resistance changed by >20%. The data were not corrected for liquid junction potential which was calculated to be 15.1–15.2 mV at room temperature. Only one neuron was examined per slice, and 4–6 slices were recorded per rat or mouse. Chemicals All drugs were obtained from Sigma-Aldrich (St. Louis, MO, USA), except for ZD7288, forskolin and tetrodotoxin (TTX), which were obtained from Tocris Bioscience (Bristol, UK). All chemicals were dissolved in distilled water except for forskolin, which was dissolved in dimethyl sulfoxide. All drugs were dissolved at 1000 times the concentration to be used and stored at −20°C unless otherwise mentioned. TTX was applied in a concentration of 1M for at least three minutes when recording mEPSCs. Other drugs were applied for around 10 min after control. Immunohistochemistry Male SD rats were deeply anesthetized and were transcardially perfused with 4% paraformaldehyde in 0.1 M phosphate buffer (PB) after 150 ml normal saline. L4−L5 spinal cord segments were removed and fixed in 4% paraformaldehyde at 6

4°C for 6 hours and then cryoprotected in 30% sucrose in 0.1 M PB at 4°C for three days. For free-floating sectioning, 30 m−thick transverse spinal cord slices were prepared in a freezing microtome (CM1950; Leica, Nussloch, Germany) and incubated in primary antibodies including the marker of excitatory presynaptic terminals, anti-glutamate vesicular transporter type 2 (VGLUT2) (Guinea pig, diluted 1:5000, Millipore), and anti-HCN 1-4 (Rabbit, diluted 1:200, Alomone Laboratories) at 4°C for three days. Secondary antibodies Cyanine 5.18 (Donkey Anti-Guinea pig, diluted 1:400, Jackson ImmunoResearch) and Alexa Fluro 488 (Donkey Anti-Rabbit, diluted 1:400, Invitrogen) were used (4°C overnight). The specificity of the primary antibodies of HCN1-4 was verified by pre-absorption of the antibodies with the same concentration of their antigen. All of the sections were mounted in Aqua-Poly/mount medium (Polysciences, Inc., Warrington, PA) and then observed with a confocal laser scanning microscope (LSM700; Carl Zeiss, Jena, Germany). Quantification of the co-localization between HCN1-4 and VGLUT2 was assessed by the co-localization function of the ZEN 2010 software equipped on the confocal microscope. The co-localization coefficient, which measures the fraction of Cyanine 5.18 pixels that are also positive for Alexa Fluro 488, was performed on the whole image containing lamina II under the microscope with 40× magnification and a 1.0 zoom. In addition, the predefined number of pixels of each frame was 1024×1024. Data were averaged from 2−3 sections/rat (N = 3 rats) and both left and right side of sections were included. A co-localization coefficient of 1 and 0 means complete co-localization and no co-localization, respectively. Statistical analysis The frequency and amplitude of EPSCs were analyzed off-line with MiniAnalysis (version 6.0.3; Synaptosoft Inc, Decatur, GA). Curve fitting was established by 7

GraphPad Prism 5.0 (GraphPad Software, USA). The dose-response curve was fitted with the Hill equation: y = Imax/[(1+(IC50/x)n], where Imax represents the maximal inhibitory response, IC50 is the half-maximal inhibitory concentration, x is the concentration of ZD7288 in M, and n is the Hill coefficient (Luo et al., 2014). Liquid junction potential was calculated with Clampex (version 9.2.0.09; Axon Instruments). Statistical analysis was performed by SPSS version 17.0 (SPSS Inc., Chicago, IL, USA). Shapiro–Wilk test was used to assess the normal distribution of data. Levene test was used to test the homogeneity of variance. One-way ANOVA followed by a post hoc Fisher’s least significant difference test was used for multiple comparisons. Paired or unpaired Student’s t-test was used as appropriate. The cumulative distributions were compared by using the Kolmogorov–Smirnov test (KS test). All data are given as the mean ± SEM. Significance was set at P < 0.05.

RESULTS Expression of HCN1-4 subtypes in excitatory glutamatergic synapses of spinal lamina II A previous study has shown that blockade of HCN channels induces a decrease in mEPSC frequency in a subset of lamina II neurons (Takasu et al., 2010), indicating the expression of HCN channels at the site of excitatory synapses. To determine which isoform of HCN channels may play a role in excitatory synaptic transmission, we first detected their expression in lamina II by performing double-immunolabelling of four isoforms (HCN1-4) and VGLUT2, an excitatory presynaptic marker (Todd et al., 2003; Maxwell et al., 2007; Boulland et al., 2009; Yasaka et al., 2010). Lamina II was recognized by its relative transparent view 8

using bright field microscopy. All four isoforms of HCN channels were detected in lamina II. As shown in Fig. 1A-D, HCN1- and HCN4-immunoreactivity (IR) displayed a punctate staining pattern. In contrast, HCN2- and HCN3-IR were detected predominantly on the neuronal membrane or somata. Quantitative analysis of the extent of co-localization demonstrated that VGLUT2 and HCN4 exhibit a highest degree of co-localization (23 ± 3%, n = 17 frames from N = 3 rats), followed with a weaker co-localization of HCN1 (15 ± 2%, n = 17 frames, P < 0.05, One-way ANOVA, post hoc Fisher’s test) and HCN2 (14 ± 2%, n = 18 frames, P < 0.05, One-way ANOVA, post hoc Fisher’s test). There was almost no HCN3 co-localization detected with VGLUT2 (2 ± 0.4%, n = 17 frames, P < 0.01; Fig. 1E). In addition, the co-localization coefficients of HCN1 and HCN2 with VGLUT2 were not significantly different (P = 0.85, One-way ANOVA, post hoc Fisher’s test). These observations indicate a distinct expression pattern of HCN1-4 isoforms in excitatory synaptic terminals, among which HCN4 is the predominant isoform. Thus, HCN4 probably plays a major role in modulating presynaptic glutamate release in the spinal dorsal horn. ZD7288 dose-dependently decreases the frequency, but not the amplitude, of sEPSCs Although ZD7288, an HCN channel blocker, has been shown to decrease the mEPSC frequency in SG neurons under the neuropathic conditions (Takasu et al., 2010), little is known about the role of HCN channels in glutamate release under the physiological condition. To address this question, we performed whole-cell voltage-clamp recordings on SG neurons from acute spinal cord slice preparations. As shown in Fig. 2A, application of ZD7288 (10 M) for 10 min resulted in a reduction in the frequency of sEPSCs in 12 of 23 SG neurons examined. The suppression of sEPSC frequency by ZD7288 occurred within 3 −5 9

min, reached a peak at approximately 5−10 min, and persisted for at least 30 min after washout of ZD7288, which suggests an irreversible effect. No effects of ZD7288 on sEPSCs were detected in the remaining neurons. Furthermore, ZD7288 significantly prolonged the inter-event interval (P < 0.05, KS test), but did not alter the amplitude (P = 0.77, KS test) of sEPSCs obtained from the same neuron in Fig.2A, as indicated by Fig. 2B. This effect was confirmed in 11 other neurons. Next, to investigate whether the ZD7288-induced reduction in sEPSC frequency is dose-dependent, five different concentrations of ZD7288 (0.1-100 M) were used. The average sEPSC frequency relative to control were 101 ± 6% (0.1 M, n = 8/8 neurons, P = 0.51, paired Student’s t-test), 71 ± 8% (1 M, n = 4/10 neurons, P < 0.05), 63 ± 4% (10 M, n = 12/23 neurons, P < 0.01; Fig. 2C), and 47 ± 5% (50 M, n = 7/12 neurons, P < 0.01), respectively (Fig. 2D and Table 1). Surprisingly, 100 M of ZD7288 increased the sEPSC frequency to 179 ± 20% of control instead (n = 4/6 neurons; P < 0.05; Table 1). These contrary results were probably due to the nonspecific effect of ZD7288 at very high concentrations (Wu et al., 2012). The IC50 was 1.89 M when fitted with a Hill equation (Hill coefficient = −0.59; Fig. 2D). Changes in sEPSC frequency are generally used to reflect an alteration in presynaptic neurotransmitter release probability. Therefore, these results indicate that ZD7288 decreases glutamate release through a presynaptic mechanism of HCN channel inhibition. Postsynaptic HCN channel-driven spontaneous firing is not involved in the effect of ZD7288 on EPSCs Our previous studies have validated that HCN channels are expressed in sub-populations of SG neurons (Liu et al., 2015b; Hu et al., 2016). In addition, the activation of HCN channels in SG neurons is voltage-dependent and the reversal potential is about -40 mV (Liu et al., 2015b), indicating that postsynaptic HCN 10

channels are tonically open at -70 mV. Since bath application of ZD7288 also blocks postsynaptic HCN channels, which may lead to a hyperpolarization of the neuron and subsequent reduction in spontaneous firing rate (Deng et al., 2007), we next investigated the effect of ZD7288 on mEPSC frequency under the condition of TTX (1 M), a specific and potent blocker of sodium channels. Similar to sEPSC, ZD7288 still caused a significant decrease in mEPSC frequency by 33 ± 4% (n = 9/17 neurons, P < 0.05; Fig. 3A, C). As shown in Fig. 3B, cumulative distribution of the inter-event interval of mEPSCs was significantly increased (P < 0.01, KS test) without change in the amplitude (P = 0.09, KS test). Additionally, there was no significant difference of the reduction between the sEPSC and mEPSC frequency (P = 0.42, unpaired Student’s t-test; Fig. 3D). Consequently, these results indicate that the reduction of glutamatergic transmission is not due to the initiation of postsynaptic HCN channel-driven neuronal activity. Forskolin increases the frequency, but not the amplitude, of mEPSCs Since previous studies have shown that HCN2 and HCN4 channels can be activated by intracellular cAMP (Kaupp and Seifert, 2001; Biel et al., 2002), and our immunohistochemistry results indicated the enrichment of HCN4 in the excitatory presynaptic terminal (Fig.1), we hypothesized that activation of HCN channels by forskolin, an activator of adenylate cyclase (Huang and Hsu, 2006; Genlain et al., 2007), might increase the mEPSC frequency. As illustrated in Fig. 4A and C, the exposure of SG neurons to forskolin (50 M) for 10 min induced a dramatic increase in mEPSC frequency (325 ± 34% of control, n = 7/8 neurons, P < 0.01, paired Student’s t test), whereas the amplitude was unchanged (116 ± 8% of control, P = 0.13). Cumulative distribution analysis demonstrated that forskolin shortened the inter-event interval of mEPSCs (P < 0.01, KS test) but did not affect the amplitude (P = 0.30, KS test) (Fig. 4B). Furthermore, in order to confirm 11

whether the enhancement effect of forskolin on mEPSCs is due to HCN channel activation or not, we pretreated the slices with 10 M ZD7288 (Fig. 4D). To our surprise, in the presence of ZD7288, forskolin still caused an increase in mEPSC frequency (216 ± 31% of control, n = 7/7 neurons, P < 0.05; Fig. 4E and Table 1). However, the extent of increase in mEPSC frequency was significantly less in ZD7288-pretreated group than that in forskolin alone-treated mEPSC (P < 0.05, unpaired Student’s t-test; Fig. 4F). These data suggest that forskolin-induced increase in mEPSC frequency is partially subject to the activation of presynaptic HCN channels. In addition, we cannot exclude the possibility that other cAMP targets, such as cyclic nucleotide-gated (CNG) channels, might lead to the enhancement of forskolin on mEPSC frequency. Together, these results demonstrate that activation of HCN channels can increase presynaptic glutamate release. Both ZD7288 and forskolin have no effect on sEPSC frequency in GFP-positive SG neurons Spinal lamina II mainly contains both excitatory (glutamatergic) and inhibitory (GABAergic and glycinergic) interneurons. Because only a proportion of SG neurons respond to ZD7288, we then considered the possibility that this subset of neurons might belong to either type of SG neurons. Therefore, we performed the same experiment on spinal cord slices prepared from GAD67-GFP mice (Fig. 5A). We first examined the effects of ZD7288 (10 M) and forskolin (50 M) on sEPSCs recorded from GFP-labelled GABAergic interneurons (Fig. 5B and D). Surprisingly, in all of the neurons examined, neither ZD7288 (103 ± 5% of control, n = 9/9 neurons, P = 0.54) nor forskolin (113 ± 6% of control, n = 7/7 neurons, P = 0.07) had any observable effect on sEPSC frequency (Fig. 5C and E). Since ZD7288-induced reduction in sEPSC frequency from SD rats was significantly 12

greater in 50 M than that in 10 M (P = 0.03, unpaired Student’s t-test), we then perfused the GFP-labelled neurons with 50 M ZD7288. Similarly, 50 M of ZD7288 still exerted no detectable effect on sEPSC frequency (107 ± 4% of control, n = 9/9 neurons, P = 0.11; Table 1). These results indicate that glutamatergic synaptic terminals expressing HCN channels seldom target to GABAergic interneurons. Next we investigated the effects of ZD7288 and forskolin on sEPSC frequency recorded from non-GFP-expressing SG neurons, more than 80% of which have been identified as excitatory interneurons (Li and Baccei, 2014). Interestingly, distinct from the GFP-labelled neurons, the sEPSC frequency of non-GFP-expressing SG neurons was significantly decreased by ZD7288 from 5.1 ± 0.9 Hz to 3.3 ± 0.8 Hz (n = 6/8 neurons, P < 0.01; Fig. 6A−B, Table 1), however drastically increased by forskolin from 4.3 ± 1.1 Hz to 8.9 ± 1.4 Hz (n = 9/9 neurons, P < 0.01; Fig. 6C−D, Table 1). Taken together, these findings indicate that HCN channels are primarily expressed in excitatory synaptic terminals innervating glutamatergic but not GABAergic interneurons in the SG.

DISCUSSION Although HCN channels have been identified to be critical in modulating excitatory transmission in SG neurons, there are still about 50% of SG neurons who do not respond to HCN channel blockers. Here, we provided the first direct evidence for the distinct expression of HCN channel isoforms in glutamatergic presynaptic terminals, and we further unveiled their specific role in excitatory synaptic processing within spinal lamina II under physiological status. First, our current studies indicate that HCN4 is the most enriched isoform, followed by HCN1 and HCN2, whereas HCN3 is less detected in glutamatergic terminals. Second, the blockade of the HCN channels by ZD7288 leads to a 13

decrease, and the activation of HCN channels by forskolin results in an increase in EPSC frequency. Of note, both of the effects were observed in a sub-population of SG neurons. Third, these subsets of SG neurons are most likely excitatory interneurons. Hence, our current studies highly suggest that the HCN channel plays a crucial role in excitatory input of excitatory, but not in inhibitory interneurons in SG. Expression of HCN channels in the spinal dorsal horn Although HCN1-IR has been shown not to reside in lamina II of spinal cord (Tu et al., 2004; Milligan et al., 2006), our data demonstrated that all of the HCN channel isoforms are detected in the spinal dorsal horn (Fig. 1), which is consistent with a recent study showing a weak expression of HCN1-IR in the superficial of spinal dorsal horn (Weng et al., 2012). This discrepancy is likely attributed to the different ages of the rats recruited and/or the thickness of the sections used. However, the role of HCN1 in pathologic pain may be less important, as suggested by a study indicating no significant difference in HCN1-IR expression in the spinal cord between animals with CFA-induced inflammatory pain and the control group (Acosta et al., 2012). HCN2 is the most extensively studied subtype. Studies conducted by Papp et al. indicate that HCN2-IR is restricted to calcitonin gene-related peptide (CGRP, a marker of axon terminals of peptidergic nociceptive primary afferents)-positive axon terminals of primary afferents, and mostly exists in extrasynaptic sites (Antal et al., 2004). Furthermore, HCN2-positive terminals have been shown to be co-localized with substance P and contact principally excitatory, but not inhibitory interneurons (Papp et al., 2006). This is also verified by our current study (Fig. 5 −6). Interestingly, HCN2-IR has also been manifested to be co-localized with isolectin B4 (IB4, a maker of axon terminals of nonpeptidergic nociceptive primary afferents) (Weng et al., 14

2012). Regardless of the localization of HCN2, it is believed to play an essential role in inflammatory and neuropathic pain (Emery et al., 2011). HCN3-IR in the spinal cord seems to be undetectable according to a previous study (Weng et al., 2012), but surprisingly, our current studies demonstrate a distinct expression pattern of HCN3 that is mostly restricted to neuronal somata (Fig. 1C). HCN4-IR is broadly distributed in the spinal cord and displays a higher expression level in lamina I and II inner than in the deeper lamina (Hughes et al., 2013). Accordingly, we further validated the expression of HCN1-4 mRNA in the superficial spinal dorsal horn by RT-PCR approach (data not shown). In this study, we calculated the co-localization coefficient instead of the percentage of synaptic terminals expressing HCN1-4, by measuring the fraction of Cyanine 5.18 pixels that are also positive for Alexa Fluro 488. There are some limitations

of

determination.

the

immunofluorescence

Background

and

noise,

staining-based which

are

co-localization

inevitable

in

all

immunohistochemical images, are two major limitations of this approach and may generate some delusive signals (Swedlow et al., 2002; Murray et al., 2007; Wolf et al., 2007). In addition, because the co-localization analysis between HCN channels and VGLUT2 only relies on the spatial overlay indicated by the immunofluorescence signals, it is unclear if these two proteins can function by binding to each other directly or indirectly. Therefore, we further performed electrophysiological

experiments

to

prove

our

hypothesis.

Indeed,

our

electrophysiological results demonstrate that blockage of HCN channels by ZD7288 decreases, while activation of HCN channels by forskolin significantly increases the excitatory inputs of SG neurons (Fig. 2 and 4). Taken together, our results as presented here reveal for the first time that HCN4 is the most abundant subtype co-localized with VGLUT2, and suggest a predominant role of HCN4 in modulating excitatory synaptic transmission in the superficial spinal dorsal horn. 15

Modulation of glutamatergic synaptic transmission by HCN channels The role of HCN channels in regulating the function of pacemaking cells has been intensively studied. In contrast, the possible non-pacemaking functions of HCN channels, including control of basal synaptic transmission, have not been completely understood. Consistent with the observation that HCN channels are present at certain terminals of the central nervous system (He et al., 2014), marked reduction of sEPSC or mEPSC frequency by bath application of ZD7288 (10−50 M; 10−20 min) has been reported in hippocampal neurons (Neitz et al., 2011), periaqueductal gray neurons (Du et al., 2013), giant neurons in the cochlear nucleus (Rusznak et al., 2013), and SG neurons in the spinal dorsal horn (Takasu et al., 2010). In our study, the IC50 value of this ZD7288-mediated effect was 1.89 M (Fig. 2D). In DRG neurons, ZD7288 blocked Na+ currents (IC50 = 1.17 M) with a higher potency than that on Ih (IC50 = 15 M), which implicates that it may be necessary to reconsider the selectivity of ZD7288 for HCN channels. However in SG neurons, TTX, a sodium channel blocker, did not influence the effect of ZD7288 on mEPSC frequency (Fig. 3D). Thus, our data presented here suggest that the suppression of ZD7288 on EPSCs is not via postsynaptic HCN channel-driven tonic firing. Given that Ih can be recorded in certain types of SG neurons (presumably inhibitory interneurons) (Yasaka et al., 2010; Hu et al., 2016) and presynaptic terminals of SG neurons are technically inaccessible for patch-clamp, we cannot exclude the possibility of a subthreshold action of postsynaptic HCN channels. Nonetheless, direct evidence of presynaptic HCN channels modulating glutamatergic synaptic transmission has been obtained in a study by recording mEPSCs from a giant glutamatergic nerve terminal in the medial nucleus of the trapezoid body, calyx of Held, where cesium chloride reduces mEPSC frequency, and forskolin increased mEPSC frequency (Huang and Trussell, 2014). In line with this study, we also found that forskolin drastically 16

increased the mEPSC frequency (Fig. 4A-C), at least partially through an HCN channel-dependent mechanism as suggested by our observation (Fig. 4D-F). Moreover, ZD7288 has also been shown to increase paired-pulse facilitation, suggesting a presynaptic modulation of ZD7288 (Chevaleyre and Castillo, 2002). In contrast, previous studies indicate that ZD7288 can increase mEPSC frequency in entorhinal cortical layer III pyramidal neurons due to membrane hyperpolarization-induced activation of a T-type calcium channel through inhibition of the HCN1 channel (Huang et al., 2011). The discrepancy among these studies may be due to different types of neurons examined. Further work is required to unravel the exact mechanism by which presynaptic HCN channels control the glutamate synaptic release in SG neurons. Taken together, our current study provides a new evidence that presynaptic HCN4 plays a predominant role in modulating basal excitatory transmitter release. This will help us better understand how neural circuits function in pain signal processing. Postsynaptic targets of HCN channel-containing glutamatergic terminals in the SG Although the SG includes both excitatory and inhibitory interneurons, our results suggest

that

HCN

channel-expressing

glutamatergic

synaptic

terminals

predominantly synapse onto the excitatory interneurons. Indeed, neither ZD7288 nor forskolin affected the sEPSC frequency in the GFP-labelled GABAergic interneurons (Fig. 5). This is partially in line with a previous study in which most of the HCN2-positive terminals contain the opposite markers of excitatory interneurons (Papp et al., 2010). Another previous study also demonstrates that ZD7288 reduces mEPSC frequency in approximately 56% of the SG neurons examined (Takasu et al., 2010). In dorsal cochlear nucleus neurons, ZD7288 displays a more potent effect on suppressing glutamate release in neurons with a 17

higher initial sEPSC frequency, suggesting that local presynaptic HCN channels may exert a different effect. Given that peripheral noxious stimuli are conducted through excitatory interneurons in the superficial spinal dorsal horn, our results suggest that blockade of HCN4 in lamina II may control excitatory synaptic strength by reducing the presynaptic glutamate release probability most likely through hyperpolarization of the membrane potential.

CONCLUSION In summary, our study reveals a novel mechanism by which HCN channels, especially HCN4 isoform, are involved in regulation of the excitatory inputs of SG interneurons. Importantly, our findings indicate that HCN channels are primarily expressed in excitatory synaptic terminals innervating glutamatergic but not GABAergic interneurons in the SG, which could explain why ZD7288 decreases sEPSC frequency in a subset of SG neurons. In addition, our current study suggests that targeting the HCN4 channel may serve as a potential therapeutic approach for neuropathic pain.

CONFLICT OF INTERESTS The authors declare that they have no conflicts of interest.

AUTHOR CONTRIBUTIONS T.L. and S.C.P designed the experiments. S.C.P and J.W. performed the experiments. S.C.P, J.W., D.Y.Z., C.Y.J. and X.C.N. analyzed and interpreted the data. S.C.P. and T.L. wrote the manuscript. All authors approved the final version of the manuscript and agreed to be accountable for all aspects of the work. All persons designated as authors qualify for authorship, and all those who qualify for 18

authorship are listed. Acknowledgement—This work were supported by the National Natural Science Foundation of China (No. 31660289 to T.L. and 81560198 to D.Y.Z.) and the Innovation Fund of Nanchang University for Graduate Students (No. cx20160317). We thank professor Bing-xing Pan for GAD67-GFP transgenic mice.

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FIGURE CAPTIONS Fig. 1. Characterization of HCN1-4 isoforms and VGLUT2 in rat superficial spinal dorsal horn. (A-D): Representative images of co-immunostaining with anti-HCN1-4 (green) and anti-VGLUT2 (red) antibodies in superficial spinal dorsal horn. Arrows indicate the co-localization. (E): Quantitative analysis of the extent of co-localization between HCN1-4 and VGLUT2 shows that HCN4 exhibits a mild degree of co-localization, which is significantly larger than that of HCN1-3. In this and the following figures, error bar indicates SEM, * indicates P < 0.05, ** indicates P < 0.01, n.s. indicates not significant different.

Fig. 2. ZD7288 dose-dependently decreased the frequency but not the amplitude of sEPSCs in SG neurons. (A): Representative chart recordings of sEPSCs obtained before (left) and after (right) 10 min application of ZD7288 at a holding potential of −70 mV. Bottom panels: proportional enlargement of traces showing in an expanded time scale that correspond to the bar shown below the chart recording (the same in the following figures). (B): Cumulative probability of the sEPSC inter-event interval and amplitude plots from the same neuron in (A) showing a significant rightward shift in the distribution of the inter-event interval (P < 0.05) without any change in amplitude (P = 0.77). (C): Summary bar graphs of sEPSC frequency as well as amplitude in control and 10 M of ZD7288. In this and the following figures, dots mean the real values of the EPSC frequency or amplitude before and after the drug application. Line segments connect a pair of data. (D): The dose-response curve of the ZD7288-induced decrease in sEPSC frequency (IC50=1.89 M; Hill coefficient = −0.59). In this and the following figures, values in parentheses indicate number of cells examined.

Fig. 3. Sodium channel blocker did not affect ZD7288’s effect on EPSCs. (A): Exemplar continuous recording of effect of ZD7288 on mEPSCs in the presence of TTX. (B): ZD7288 induced a significant rightward shift in the distribution of the inter-event interval (P < 0.01) but did not affect the amplitude (P = 0.09) of mEPSCs. (C): Summary of the effects 23

of ZD7288 on mEPSC frequency (P < 0.05) and amplitude (P = 0.08). (D): TTX-pretreatment did not influence the effect of ZD7288 on EPSCs (P = 0.42).

Fig. 4. Forskolin increased the frequency but not the amplitude of mEPSCs and the enhancement could be partly blocked by ZD7288. (A): Representative mEPSC recordings before and after bath application of forskolin with TTX-pretreatment. (B): Forskolin induced a significant leftward shift in the distribution of the inter-event interval (P < 0.01) but did not alter the amplitude (P = 0.30) of mEPSCs. (C): Summarized data showing that forskolin dramatically increased the frequency of mEPSCs (P < 0.01) but did not affect the amplitude (P = 0.13). (D): Representative traces showing that pre-treatment of ZD7288 decreased the forskolin-induced enhancement of mEPSC frequency. (E-F): Summarized data showing that ZD7288 partially blocked forskolin’s effect on mEPSC frequency (P < 0.05).

Fig. 5. In GFP-labelled neurons, neither ZD7288 nor forskolin affected the sEPSCs. (A): Representative images of the SG region in a transverse spinal cord slice from a GAD67-GFP+ mouse at low (left) and high magnification (middle). Right is the same patched neuron in the middle under the observation of a monochromator. (B, D): Typical traces of sEPSCs in GFP+ neurons before (left) and after (right) application of ZD7288 (B) or forskolin (D). (C, E): Summary bar graphs showing that neither the frequency (ZD7288: P = 0.54; FSK: P = 0.07) nor the amplitude (ZD7288: P = 0.13; FSK: P = 0.64) of sEPSCs are affected by ZD7288 (C) or forskolin (E) in all GFP+ neurons tested.

Fig. 6. Similar results of ZD7288 and forskolin on sEPSC frequency can be obtained in non-GFP neurons. (A, C): Exemplar traces of sEPSCs in non-GFP neurons in the presence of ZD7288 (A) or forskolin (C). (B, D): Summary bar graphs showing that the frequency of sEPSCs is significantly decreased by ZD7288 (P < 0.01) and increased by forskolin (P < 0.01) in non-GFP neurons.

24

Table 1. Frequency and amplitude of EPSCs, relative to control Percentage of control Animal SD rat

Condition + ZD7288 (0.1 M) + ZD7288 (1 M) + ZD7288 (10 M) + ZD7288 (50 M) + ZD7288 (100 M) + TTX + ZD7288

Frequency

Amplitude

Frequency (Hz)

101 ± 6

Amplitude (pA)

Number of Cells

95 ± 8

2.7 ± 0.4

18.8 ± 2.3

8/8

71 ± 8

*

96 ± 4

2.5 ± 0.4

20.9 ± 4.1

4/10

63 ± 4

*

95 ± 3

3.3 ± 0.5

17.0 ± 1.5

12/23

47 ± 5

*

91 ± 10

4.0 ± 0.6

17.1 ± 2.2

7/12

97 ± 8

4.5 ± 0.4

19.6 ± 2.6

4/6

179 ± 20 *

93 ± 5

4.5 ± 1.2

19.9 ± 2.5

9/17

325 ± 34

116 ± 8

4.0 ± 0.5

22.7 ± 2.6

7/8

+ TTX + ZD7288 + forskolin

216 ± 31

*

100 ± 4

3.9 ± 0.9

16.9 ± 1.0

7/7

+ZD7288 (in GFP neurons, 10 M)

103 ± 5

94 ± 3

6.0 ± 1.2

18.6 ± 1.9

9/9

+ZD7288 (in GFP neurons, 50 M)

107 ± 4

98 ± 4

4.2 ± 0.9

17.0 ± 1.1

9/9

+ forskolin (in GFP neurons)

113 ± 6

100 ± 3

2.7 ± 0.7

16.3 ± 1.8

7/7

94 ± 3

5.1 ± 0.9

23.6 ± 2.2

6/8

99 ± 1

4.3 ± 1.1

17.7 ± 0.7

9/9

+ ZD7288 (in non-GFP neurons) + forskolin (in non-GFP neurons)

67 ± 4

*

*

+ TTX + forskolin GFP mouse

Control

61 ± 7

*

254 ± 33

*

Frequency and amplitude were measured around 10 min after beginning of drugs superfusion under various conditions. Values are means ± SEM. *P < 0.05.

25

26

27

28

29

30

31

32

HIGHLIGHTS ·HCN1-4 isoforms are heterogeneously expressed in glutamatergic terminals of the subtantia gelatinosa (SG). ·Suppression of HCN channels leads to reduction in EPSC frequency in a subset of SG neurons. ·Activation of HCN channels increases EPSC frequency. ·Glutamatergic terminals containing HCN channel most likely synapse onto the excitatory but not inhibitory SG neurons.

33