Research ArticleDIABETIC NEUROPATHY

Hyperpolarization-activated cyclic nucleotide–gated 2 (HCN2) ion channels drive pain in mouse models of diabetic neuropathy

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Science Translational Medicine  27 Sep 2017:
Vol. 9, Issue 409, eaam6072
DOI: 10.1126/scitranslmed.aam6072

Taking diabetic pain to heart

People with diabetes can, over time, develop painful diabetic neuropathy, a chronic pain condition induced by nerve damage. The molecular bases of diabetes-induced pain are poorly understood, and currently, there are no effective treatments. Tsantoulas et al. show that HCN2 channels, known for their role in neuronal excitability and heart rate modulation, are overactivated in the nociceptive neurons that detect pain. Ivabradine, an HCN inhibitor used for treating heart conditions, reduced chronic pain in mice with diabetes, suggesting that HCN2-selective inhibitors might be a valuable therapeutic strategy for treating diabetic neuropathies.

Abstract

Diabetic patients frequently suffer from continuous pain that is poorly treated by currently available analgesics. We used mouse models of type 1 and type 2 diabetes to investigate a possible role for the hyperpolarization-activated cyclic nucleotide–gated 2 (HCN2) ion channels as drivers of diabetic pain. Blocking or genetically deleting HCN2 channels in small nociceptive neurons suppressed diabetes-associated mechanical allodynia and prevented neuronal activation of second-order neurons in the spinal cord in mice. In addition, we found that intracellular cyclic adenosine monophosphate (cAMP), a positive HCN2 modulator, is increased in somatosensory neurons in an animal model of painful diabetes. We propose that the increased intracellular cAMP drives diabetes-associated pain by facilitating HCN2 activation and consequently promoting repetitive firing in primary nociceptive nerve fibers. Our results suggest that HCN2 may be an analgesic target in the treatment of painful diabetic neuropathy.

INTRODUCTION

Diabetes causes a characteristic degeneration of peripheral sensory nerves, frequently associated with continuous pain. Painful diabetic neuropathy (PDN) affects about one in four diabetic patients and typically manifests itself as a range of unpleasant positive symptoms such as spontaneous pain, mechanical allodynia (a painful sensation caused by light touch), and paresthesias (tingling, shooting pain), as well as negative symptoms such as thermal hyposensitivity (1). Despite its high prevalence, the pathophysiology of PDN remains poorly understood. Both anatomical changes (demyelination, loss of epidermal nerve density) and functional changes (reduced nerve conduction velocity) are characteristics of PDN and of other “die-back” neuropathies (14). The pathologies observed in the peripheral nervous system suggest that peripheral nerve damage is the driver of PDN and that the associated ongoing pain is likely to be due to repetitive discharge of action potentials in nociceptive (pain-sensitive) nerve fibers (57). However, the molecular basis of peripheral nociceptor hyperexcitability remains elusive.

Hyperpolarization-activated cyclic nucleotide–gated (HCN) ion channels have recently emerged as crucial determinants of nociceptive excitability [reviewed in (8, 9)]. HCN channels are unusual in that they are activated by hyperpolarization in the range −60 to −90 mV, in contrast to all other voltage-activated channels that are activated by depolarization. There are four HCN isoforms (HCN1 to HCN4) expressed in sensory neurons. More than half of small nociceptive neurons express HCN2 channels (10), whereas in large sensory neurons the fast HCN current (Ih) is mediated mainly by HCN1 (11, 12). HCN3 is widely expressed across dorsal root ganglion (DRG) neurons of all sizes (11), whereas HCN4, which has a key pacemaking role in the heart (13, 14), shows limited expression in somatosensory neurons (15, 16). Elevations of intracellular cyclic adenosine monophosphate (cAMP) cause a strong shift in the voltage dependence of activation of HCN2 and HCN4 to more positive membrane voltages, causing an increase in the inward current carried by these channels at resting membrane voltage, whereas HCN1 and HCN3 are relatively insensitive to cAMP and so have less influence in modulating neuronal excitability (17). In nociceptive neurons, inflammatory mediators such as prostaglandin E2 (PGE2) and bradykinin activate adenylate cyclase via a Gs protein–coupled pathway, thus causing a rise in intracellular cAMP, HCN2 activation, and increased spontaneous firing of action potentials (10).

A critical role for HCN2 in inflammatory pain and in the neuropathic pain caused by direct mechanical damage to sensory nerves has been demonstrated by the potent analgesic actions of specific HCN channel blockers and by targeted deletion of the Hcn2 gene in nociceptive neurons in mice (10). Pharmacological inhibition of Ih prevents pain in chemotherapy-induced neuropathy (18), which also features a die-back denervation pattern, and circumstantial evidence has hinted at an involvement of unidentified HCN family members in autonomic diabetic neuropathy (19, 20).

Here, we expand our understanding of the critical role of HCN2 in chronic pain by showing that cAMP-mediated HCN2 activation in a mouse model of diabetic neuropathy can trigger repetitive activity in small nociceptive fibers, leading to central sensitization and ongoing pain. Pharmacological or genetic block of HCN2 activity exerts potent analgesic effects in animal models of both type 1 and type 2 diabetes.

RESULTS

Streptozotocin treatment results in symptoms indicative of PDN

Diabetes was induced in wild-type (WT) mice by a single injection of streptozotocin (STZ). Selective accumulation of STZ in pancreatic islet β cells causes DNA alkylation, cell death, and consequent loss of insulin production, resulting in an elevation of blood glucose. Mice treated with STZ exhibited significantly increased blood glucose concentrations by day 7 compared to the pre-STZ baseline (21.9 ± 2.1 mM versus 8.4 ± 0.2 mM, P < 0.001; Fig. 1A). STZ-treated mice remained hyperglycemic thereafter, whereas blood glucose concentration did not change in vehicle-treated mice (10.1 ± 0.7 mM). The small number of mice that did not exhibit hyperglycemia after STZ injection was excluded from analysis (see Table 1).

Fig. 1. The HCN channel blocker ivabradine (IVA) is analgesic in a mouse model of type 1 diabetes.

(A) Time course of blood glucose concentrations after a single injection of vehicle (VEH; blue) or STZ (red; 150 mg/kg, intraperitoneally) in mice. BL, pre-injection baseline blood glucose. ***P < 0.001 versus BL (VEH, n = 6 mice; STZ, n = 11 at BL and n = 9 at 52 days, due to STZ-induced weight loss over the experimental time course) (see Table 1). ***P < 0.001 versus BL, one-way repeated-measures (RM) analysis of variance (ANOVA) followed by Student-Newman-Keuls (SNK) post hoc [VEH, n = 6 mice; STZ, n = 11 (BL) and n = 9 (52 days), due to STZ-induced lethality over the experimental time course]. (B) Time course of sensitivity to mechanical stimulation, shown as the von Frey force threshold for withdrawal, in VEH- or STZ-treated mice. *P < 0.05 and *P < 0.001 versus BL [VEH, n = 6; STZ, n = 18 (BL) and n = 12 (8 weeks)]. (C) Time course of the thermal withdrawal latency in VEH- or STZ-treated mice. **P < 0.01 versus vehicle [VEH, n = 12; STZ, n = 13 (BL) and n = 10 (8 weeks)]. (D) Time course of sensitivity to mechanical stimulation, shown as the von Frey force threshold for withdrawal in control (WT) or diabetic (STZ-injected) mice treated with different doses of IVA or with vehicle. Dotted lines show time of injection of STZ and IVA. *P < 0.05 versus saline, #P < 0.05 versus post-STZ, +++P < 0.001 versus BL (10 mg/kg, n = 8; 2.5 mg/kg, n = 8; 5 mg/kg, n = 6; saline, n = 14 mice). (E) Effect of repeated injection of IVA (5 mg/kg) on the von Frey force threshold in STZ mice. Dotted lines show the time of STZ (red) and IVA (blue) injection. *P < 0.05 and ***P < 0.001 versus saline; ##P < 0.01 and ###P < 0.001 versus post-STZ; +++P < 0.001 versus BL (n = 6 per group). (F) Effect of two daily injections of IVA (5 mg/kg) for 4 days on the von Frey force threshold in STZ mice. Dotted lines show the time of injection of STZ (red) and IVA (blue). IVA was injected twice daily over 4 days, 3 hours before and 3 hours after von Frey threshold testing. *P < 0.05 and *P < 0.01 versus saline; ##P < 0.01 and ###P < 0.001 versus post-STZ; +++P < 0.001 versus BL (n = 6 per group). Statistical analysis was performed using two-way RM ANOVA followed by SNK post hoc tests.

Table 1. WT mouse usage and percentages that developed diabetes and PDN.
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We examined whether the diabetic state was accompanied by altered pain sensation, similar to that encountered in human PDN. Diabetic mice showed a progressive hypersensitivity in response to mechanical stimulation (Fig. 1B). The mechanical pain threshold was significantly lower than baseline at 2 weeks, as indicated by the mean force required to elicit paw withdrawal to mechanical stimulation (4.1 ± 0.1 g versus 3.5 ± 0.1 g, P = 0.012) and continued to decrease until 8 weeks (2.3 ± 0.1 g, P < 0.001), representing a 44% increase in mechanical pain sensitivity (Fig. 1B). Mechanical hypersensitivity persisted for at least 18 weeks after STZ (2.7 ± 0.1 g, P < 0.001). In contrast, mechanical pain thresholds of vehicle-treated mice remained at baseline levels for the duration of the study (Fig. 1B). Testing with a thermal heat stimulus revealed thermal hyposensitivity 6 to 8 weeks after induction of diabetes, as indicated by increased latency of paw withdrawal (6 weeks, 12.0 ± 0.6 s versus 9.2 ± 0.6 s, P = 0.002; 8 weeks, 12.7 ± 0.5 s versus 9.4 ± 0.5 s, P < 0.001 compared to vehicle-treated mice) (Fig. 1C). The dual manifestation of mechanical hypersensitivity and thermal hyposensitivity mirrors the clinical presentation of diabetic neuropathy in humans. Activation of transient receptor potential ankyrin 1 (TRPA1) by STZ may be responsible for some of the manifestations of pain for 10 days after administration of STZ (21), and for this reason, we carried out behavioral pain measurements at later time points (8 to 18 weeks after STZ). Notably, Fig. 1 (B and C) includes all mice that developed diabetes, but for unknown reasons, only 70 to 90% of these mice developed substantial pain hypersensitivity (see Materials and Methods and Table 1).

Blocking peripheral HCN channels reverses mechanical hypersensitivity

We next investigated whether HCN function is involved in the mechanical pain hypersensitivity in PDN. To address this, we selected mice with the most robust pain phenotypes at 8 weeks after STZ and injected them intraperitoneally with ivabradine (IVA), which inhibits all four HCN isoforms with approximately equal potency (22). Note that IVA is excluded from the central nervous system (CNS) (18), so its actions can be attributed to blockade of HCN ion channels in the periphery. A single injection of IVA (2.5 mg/kg) increased diabetic pain thresholds [from 2.2 ± 0.2 g after STZ to 3.1 ± 0.2 g at 30 min (P = 0.006) and 3.2 ± 0.2 g at 60 min (P = 0.002) after IVA injection] (Fig. 1D). This effect was also observed with higher IVA doses [5 mg/kg: from 2.3 ± 0.1 g after STZ to 3.2 ± 0.5 g at 30 min (P = 0.018) and 3.1 ± 0.3 g at 60 min (P = 0.024) after IVA injection; 10 mg/kg: from 2.1 ± 0.1 g after STZ to 3.1 ± 0.3 g at 30 min (P = 0.001) and 3.3 ± 0.3 g at 60 min (P < 0.001) after IVA injection]. At all doses, the IVA-mediated analgesic effect on pain threshold was significant compared to the vehicle group at 60 min after IVA injection (2.5 mg/kg, P = 0.014; 5 mg/kg, P = 0.016; 10 mg/kg, P = 0.021) and subsided after 120 min. IVA had no effect on acute mechanical and thermal pain thresholds of control mice (Fig. 1D and fig. S1), confirming previous observations that HCN channels are not engaged during normal acute pain processing (10). In addition, thermal hypoesthesia in mice treated with STZ was unaltered by IVA (fig. S1), suggesting that thermal hyposensitivity is unrelated to changes in HCN ion channel function.

Because IVA did not completely restore normal pain sensitivity, we asked whether the analgesic effect could be enhanced by repeated administration of the drug. Two consecutive injections (5 mg/kg; separated by 2 hours) increased pain thresholds beyond those of a single administration, and a third injection completely restored thresholds to prediabetic levels [from 3.9 ± 0.2 g before STZ to 4.0 ± 0.2 g at 30 min (P = 0.789), 3.8 ± 0.3 g at 60 min (P = 0.843), and 3.7 ± 0.3 g at 120 min (P = 0.960) after IVA injection] (Fig. 1E). Note that there is a small change in threshold after each saline injection, likely to be due to analgesia caused by the stress of handling and injection, but the analgesia caused by IVA injection is significantly greater, particularly after repeated injection.

Finally, we examined whether this cumulative effect could also be seen with a more clinically relevant, spaced-out dosing regimen. We treated the animals with IVA twice a day, 3 hours before and 3 hours after pain threshold measurements over four consecutive days. Mechanical thresholds on days 2 to 4 in mice treated with IVA were not different from those recorded before STZ injection and progressively increased compared to saline-treated animals (day 2, P = 0.012; day 3, P = 0.002; day 4, P = 0.014). The analgesia was fully reversible, and cessation of treatment for 4 days resulted in a drop of mechanical thresholds back to diabetic levels.

Because IVA inhibits all HCN isoforms, including HCN4, which is important for cardiac pacemaking, we investigated the effect of the drug on cardiac function using pulse oximetry. As expected, IVA (5 mg/kg) caused a reduction in basal heart rate in control mice (29.2% at 30 min versus baseline; P < 0.001) and a smaller reduction in diabetic mice (15.8% at 30 min; P = 0.009 versus baseline, P = 0.001 versus CTRL + IVA at 30 min; fig. S2). This lesser effect of IVA on diabetic cardiac function has also been documented in STZ rats (23). IVA reduced respiratory rate (fig. S2B) in both control and diabetic mice, but no change was detected in oxygen saturation (fig. S2C) or pulse distention at 30 min after injection (fig. S2D). There was no obvious effect on alertness or exploratory behavior as detected by visual observation. Together, the pulse oximetry data suggest that an analgesic dose of IVA is well tolerated in both control and diabetic mice, causing bradycardia but no other adverse effects on cardiac function.

Peripheral HCN block is analgesic in type 2 diabetes

Type 2 diabetes is the most common form of diabetes in humans; therefore, we tested the effect of HCN block in the db/db mouse model of type 2 diabetes, in which mice homozygous for a leptin receptor mutation (db/db) become obese in adulthood and show hyperglycemia together with an array of diabetic symptoms (24). At 18 weeks of age, all db/db mice were obese and hyperglycemic compared to heterozygous age-matched controls (db/+) (body weight: db/db, 48 ± 0.6 g; db/+, 29.3 ± 0.4 g, P < 0.001; blood glucose: db/db, 27 ± 1.3 mM; db/+, 9.4 ± 0.4 mM, P < 0.001) (Fig. 2A). The db/db mice exhibited significantly lower mechanical pain thresholds (3.6 ± 0.2 g versus 4.5 ± 0.2 g; P < 0.001) (Fig. 2, B and C). Note that both the elevation in blood glucose and the mechanical hyperalgesia were less than those of type 1 diabetic (STZ-injected) mice (compare Fig. 2 with Fig. 1, A and B). A single IVA administration (5 mg/kg, intraperitoneally) restored normal pain thresholds in db/db mice (from 3.6 ± 0.2 g to 4.6 ± 0.2 g; P < 0.001 versus saline at 60 min), whereas saline injection had no significant effect on pain threshold (P = 0.47 at 60 min) compared to baseline (Fig. 2C).

Fig. 2. IVA is analgesic in a mouse model of type 2 diabetes.

(A) Blood glucose concentration in mice heterozygous (db/+; controls) and homozygous (db/db; model of type 2 diabetes) for the db mutation. ***P < 0.001, Student’s t test (db/db, n = 12 mice; db/+, n = 6 mice). (B) Von Frey force threshold in control (db/+) or diabetic (db/db) mice. ***P < 0.001, Student’s t test (n = 6 mice per group). (C) Effect of IVA (5 mg/kg) or vehicle intraperitoneal injection on the von Frey force threshold in db/db mice. Dotted line shows the von Frey force threshold in control (db/+) mice. ***P < 0.001 versus saline, two-way ANOVA followed by Bonferroni test (n = 14 mice per group except for 120 min, where n = 6 mice per group).

Induction of spinal c-Fos in diabetic mice is reversed by HCN inhibition

We next sought a more direct way of demonstrating that enhanced excitability of peripheral nociceptive neurons is involved in diabetes-induced hyperalgesia. Activation of second-order neurons in the spinal cord by primary afferent input induces c-Fos, an immediate-early gene rapidly up-regulated by neuronal activity (25). Therefore, C-FOS protein expression in neurons of the outer layers of the dorsal horn, where nociceptive fibers terminate, may be indicative of sustained peripheral nociceptive input and ongoing pain in diabetes. Quantification of C-FOS immunoreactivity in lumbar spinal cord sections (laminae I and II) revealed an increased number of C-FOSpositive neurons in the dorsal horn at 8 weeks after STZ administration compared to controls (26.0 ± 2.6 versus 1.2 ± 0.2 neurons per section; P < 0.001) (Fig. 3). Although C-FOS activation was notably less compared to that induced by a strong noxious stimulus [fig. S3 and (26)], it agrees with previous studies in diabetes (27). IVA treatment (three intraperitoneal injections of 5 mg/kg every 2 hours, as in Fig. 1E) reduced diabetes-induced C-FOS activation 6 hours after the first injection (6.2 ± 0.4 neurons per section; P < 0.001 versus STZ). These data indicate that PDN increases activation of dorsal horn neurons, likely due to enhanced nociceptive input to the spinal cord, and that this activation can be reversed by pharmacological block of HCN in the periphery.

Fig. 3. Diabetes-induced C-FOS expression in spinal dorsal horn neurons is reduced by IVA.

(A) Representative images of the immunohistochemical analysis showing NeuN (left, red) and C-FOS (right, green) expression in spinal dorsal horn neurons in WT vehicle, STZ vehicle, and STZ + IVA–injected mice. Insets, higher magnification of dashed rectangles. Scale bars, 50 μm. Arrows indicate cells coexpressing NeuN and C-FOS. (B) Quantification of C-FOS–positive neurons in the three different experimental conditions depicted in (A). ***P < 0.001, one-way ANOVA followed by SNK post hoc tests (control, n = 4; STZ, n = 3; STZ + IVA, n = 3 mice).

Genetic deletion of HCN2 in nociceptive neurons prevents development of diabetic pain

IVA blocks all four HCN isoforms equally (22), so to decipher which HCN isoform causes hyperalgesia in diabetes, we used a conditional knockout mouse line in which HCN2 had been selectively deleted in nociceptive (NaV1.8-expressing) DRG neurons (HCN2 cKO) (10). STZ treatment of HCN2 cKO mice or their littermate controls (fHCN2) caused a similar hyperglycemia within 1 week (HCN2 cKO pre-STZ versus post-STZ, 7.8 ± 0.5 mM versus 21.5 ± 1.4 mM, P < 0.001; fHCN2 pre-STZ versus post-STZ, 8.1 ± 0.3 mM versus 25.2 ± 3.4 mM, P < 0.001) (Fig. 4A). High blood glucose concentrations were also present at 8 weeks (HCN2 cKO, 40.4 ± 0.6 mM; fHCN2, 34.2 ± 6.3 mM; P < 0.001 versus pre-STZ) (Fig. 4A). There was no significant difference between genotypes at any time point (1 week, P = 0.07; 8 weeks, P = 0.291), indicating that both genotypes become diabetic to an equal extent.

Fig. 4. HCN2 deletion from nociceptive neurons prevents development of pain and C-FOS induction in diabetic mice.

(A) Blood glucose concentration in mice in which HCN2 has been genetically deleted in NaV1.8-expressing sensory neurons (HCN2 cKO), as well as in their littermates (fHCN2+/+, effectively WT, used as control). ***P < 0.001 versus BL; P > 0.05 for fHCN2 versus HCN2 cKO, two-way RM ANOVA followed by SNK post hoc test (HCN2 cKO, n = 12; fHCN2, n = 20 mice). (B) Time course of the von Frey force threshold in fHCN2 and HCN2 cKO mice after STZ injection (indicated by dotted lines). *P < 0.05 and ***P < 0.001 versus fHCN2; ###P < 0.05 versus BL, two-way RM ANOVA followed by SNK post hoc test [HCN2 cKO, n = 17 (BL) and 10 (8 weeks) mice; fHCN2, n = 21 (BL) and 10 (8 weeks) mice]. (C) Representative immunohistochemical images showing NeuN (left, red) and C-FOS (right, green) expression in spinal dorsal horn neurons 8 weeks after STZ in fHCN2 and HCN2 cKO. Right: Quantification of C-FOS–positive neurons in fHCN2 and HCN2 cKO STZ-treated animals. ***P < 0.001, Student’s t test (n = 3 mice per group). Insets, higher magnification of dashed rectangles. Scale bars, 50 μm. Arrows show neurons coexpressing NeuN and C-FOS.

We next studied the development of PDN in these transgenic mice. Diabetic fHCN2 mice, which are effectively WT, developed a progressive mechanical hypersensitivity, evident as a significant drop in mechanical thresholds compared to prediabetic baseline (2 weeks, 3.7 ± 0.2 g; 4 weeks, 3.4 ± 0.2 g; 6 weeks, 3.1 ± 0.2 g; 8 weeks, 3.2 ± 0.3 g; P < 0.001 versus BL; 4.2 ± 0.1 g) (Fig. 4B). When we assessed the HCN2 cKO mice, however, we found no signs of mechanical hypersensitivity, despite the ongoing diabetes. Mechanical thresholds at 2, 4, 6, and 8 weeks after STZ (4.2 ± 0.1 g, 4.2 ± 0.2 g, 4.3 ± 0.1 g, and 4.4 ± 0.1 g, respectively) were not significantly different from prediabetic values (4.2 ± 0.1 g; P = 0.987, P = 0.964, P = 0.623, and P = 0.737, respectively) (Fig. 4B) and remained significantly higher than the corresponding values in the fHCN2 group (2 weeks, P = 0.037; 4 to 8 weeks, P < 0.001).

Using C-FOS immunoreactivity in the dorsal horn as a measure of nociceptive activity impinging on second-order neurons of the spinal cord, we observed an increase in the numbers of C-FOS–positive neurons in diabetic fHCN2 mice, whereas C-FOS expression was greatly reduced in diabetic HCN2 cKO mice (15.6 ± 0.8 versus 1.4 ± 0.2 neurons per section; P < 0.001) (Fig. 4C). Together, these data suggest that genetic deletion of HCN2 in NaV1.8+ nociceptors recapitulates the analgesia and suppression of peripheral nociceptive drive caused by pharmacological block of HCN channels and, therefore, that HCN2 expressed in nociceptive neurons is necessary for diabetes-induced pain in the mouse STZ model.

Diabetes reduces plantar skin innervation

Intraepidermal nerve fiber density is often used as a diagnostic criterion for peripheral neuropathy in human diabetic patients. We calculated intraepidermal nerve fiber density in plantar skin of the hindpaw by staining for PGP9.5, a pan-neuronal marker present in nerve terminals, and for collagen IV, which stains the basement membrane at the dermis-epidermis boundary. We also used 4′,6-diamidino-2-phenylindole (DAPI) stain to highlight the nuclei of basal layer cells of the basement membrane. With this combination of stains, nerve fibers crossing into the epidermis were visualized (Fig. 5A). Quantification of intraepidermal nerve fiber density revealed a significant reduction in WT diabetic mice at 8 weeks after STZ compared to nondiabetic WT controls (26.8 ± 2.0 versus 44.2 ± 3.5 fibers/mm; P = 0.016), and this reduction was also present 8 weeks after STZ in both fHCN2 mice (effectively WT) and HCN2 cKO mice (fHCN2: 25.0 ± 2.6 fibers/mm, P = 0.023 versus control; HCN2 cKO: 28.1 ± 6.7 fibers/mm, P = 0.014 versus control) (Fig. 5B). These results confirm the peripheral neuropathy in the STZ mouse model and demonstrate that the lack of a pain phenotype in the HCN2 cKO mice is not due to a change in the denervation pattern. To further characterize the identity of affected intraepidermal fibers, we stained for calcitonin gene-related peptide (CGRP), which identifies peptidergic fibers, including transient receptor potential cation channel subfamily V member 1 (TRPV1)–positive fibers that sense noxious heat. Diabetes reduced intraepidermal CGRP+ fibers from 17.4 ± 0.8 to 12.5 ± 0.6 fibers/mm (P = 0.006; Fig. 5C, bottom left graph). Because most TRPV1+ fibers in mouse coexpress CGRP (28), this reduction agrees with the observed thermal hyposensitivity in STZ mice (Fig. 1C). There was a significant inverse correlation between intraepidermal CGRP+ fiber density and heat pain thresholds in diabetic mice (R2 = 0.88, P = 0.006; Fig. 5C, bottom right graph).

Fig. 5. Diabetes decreases intraepidermal nerve fiber density in the skin of the hindpaw in mice.

(A) Representative skin section from a control mouse stained for PGP9.5 (marker of nerve fibers), collagen IV (Col IV; marker of basement membrane separating dermis from epidermis), and DAPI (marker of cell nuclei). Merged signal is shown in the far right panel. Multiple free nerve endings crossing into the epidermis are visible in the merged image (arrows). (B) Top: Representative immunohistochemical images of skin innervation in WT control, WT diabetic (WT STZ), fHCN2+/+ (effectively WT) diabetic (fHCN STZ), and NaV1.8-Cre/HCN2 KO (HCN2 cKO) mice. Bottom: Quantification of intraepidermal nerve fiber (IENF) density. *P < 0.05, one-way ANOVA followed by SNK post hoc test (n = 3 mice per group except for HCN2 cKO STZ, where n = 6). (C) Top: Representative immunohistochemical skin images showing CGRP, collagen IV, and DAPI staining. Bottom left: Quantification of CGRP+ fiber density in control and STZ mice. **P < 0.01, Student’s t test (control, n = 6; STZ, n = 3 mice). Bottom right: Correlation of CGRP+ fiber density and threshold for noxious heat in diabetic mice. **P < 0.01, Pearson’s correlation test (n = 6 mice). Scale bars, 40 μm.

Diabetes does not affect HCN2 expression in sensory neurons

The preceding experiments show that enhanced peripheral HCN2 function plays a critical role in pain and mechanical hypersensitivity in diabetes. To determine whether this is due to an up-regulation of HCN2 expression in sensory neurons, we used immunohistochemistry to visualize HCN2 in lumbar DRG sections (Fig. 6A). In control animals, HCN2 immunoreactivity was detected in 73.2 ± 4.5% of all neurons, as revealed by colocalization with the pan-neuronal marker β3-tubulin (examples denoted by arrows). As previously reported, HCN2 was detected in a range of cell sizes, including most small-diameter neurons, the majority of which are nociceptive, as well as in larger neurons with a characteristic membrane ring staining (29). In diabetic animals at 8 weeks after STZ, the percentage of HCN2-positive neurons was 72.5 ± 7.0%, not significantly different from control (P = 0.934; Fig. 6B, left). In addition, the distribution of HCN2+ neurons among subclasses of DRG neurons (small-, medium-, and large-diameter classes) was also not different between control and diabetic (P = 0.116; Fig. 6B, middle). When examining staining intensity in HCN2-positive neurons, we found no effect of diabetes among all neurons [control, 3.5 ± 0.2 arbitrary units (AU), versus diabetic, 3.9 ± 0.1 AU; P = 0.102] or among DRG subpopulations (P = 0.802; Fig. 6B, right). The specificity of the HCN2 antibody was confirmed by detection of HCN2 in small NaV1.8+ nociceptive DRG neurons of WT mice but not of HCN2 cKO mice, as well as by the complete lack of HCN2 immunoreactivity in DRG from HCN2 global KO (gKO) mice (Fig. 6C).

Fig. 6. HCN2 expression in sensory neurons is not regulated by diabetes.

(A) Representative immunohistochemical staining for the pan-neuronal marker β3-tubulin (green), HCN2 (red), and the merged signal in transverse sections of lumbar DRG from control or diabetic (STZ) WT mice. Examples of HCN2-positive neurons are indicated by arrows. Scale bar, 40 μm. (B) Left: Percentage of HCN2+ neurons in control and STZ mice (Student’s t test). Middle: Distribution of HCN2 + neurons among DRG size classes (χ2 test). Right: Intensity of HCN2 immunoreactivity as a function of neuronal size (linear regression analysis comparing slopes). Control, n = 4 mice; STZ, n = 4 mice. ns, not significant. (C) Representative immunohistochemical staining in transverse sections of lumbar DRG for NaV1.8+ (marker of small nociceptive DRG neurons, green), HCN2 (red), and the merged signal in WT, HCN2 cKO, and HCN2 gKO. Examples of NaV1.8+ neurons are indicated by arrows. Scale bars, 40 μm.

We investigated whether enhanced HCN2 function could be due to changes not reflected in the cell soma (such as axonal trafficking), by examining HCN2 expression in the sciatic nerve. The intensity of axonal HCN2 staining was similar between control and STZ mice at 8 weeks (2.6 ± 0.1 AU versus 2.6 ± 0.6 AU; P = 0.871), suggesting that there is no change in axonal HCN2 expression in diabetes (fig. S4A). We did not detect any HCN2 in epidermal nerve fibers of control mice, probably because the channel is present in low quantities in the very fine peripheral nerve endings (fig. S4B).

We also examined expression and regulation by diabetes of the other HCN isoforms in DRG neurons. HCN1 was predominantly localized in medium-large neurons (fig. S5), with no difference between control and diabetic conditions in overall proportion of HCN1+ neurons, distribution of HCN1+ neurons among size classes, or intensity of stain as a function of cell size. The HCN3 isoform was detected in a mixture of neurons of all sizes (fig. S6), and there was no effect of diabetes on either proportion of HCN3+ neurons or stain intensity as a function of size. HCN4 was found mainly in medium-large neurons (fig. S7), and again, there was no effect of diabetes on any parameter. Overall, the immunohistological data show that diabetes does not induce up-regulation of any HCN isoform at the protein level. Note that a previous study reported that diabetes increases HCN expression and currents in parasympathetic neurons of the nodose ganglion, suggesting that HCN channels may be differentially regulated in autonomic neuropathy (19).

Diabetes does not change electrophysiological properties of isolated nociceptive neurons

We next examined the in vitro electrophysiological properties of small DRG neurons [<20 μm diameter, predominantly nociceptive in nature (30)] from control and STZ mice to determine whether diabetes alters HCN ion channel properties. In response to steady current injection, neurons from STZ and nondiabetic mice showed a similar increase in firing frequency with increasing injected current, reaching a similar plateau level (14.58 ± 1.87 versus 16.45 ± 2.99 spikes per second; Fig. 7, A and B). Exposure to 50 μM forskolin, which increases intracellular cAMP and shifts the activation curve of HCN2 to a more positive membrane voltage, increased firing frequency after current injection, from 13.51 ± 2.26 to 21.68 ± 2.72 spikes per second (40 pA; P < 0.001) and from 14.58 ± 1.87 to 24.0 ± 2.94 spikes per second (50 pA; P = 0.009). The cAMP-dependent sensitization of neuronal firing was also reflected in a change in the pattern of firing, from phasic (steady current elicits one or a few spikes) to tonic [firing continues throughout current injection; see Fig. 7C and (31)]. The proportion of phasic/tonic firing neurons was similar between STZ and nondiabetic mice at baseline (P > 0.05; Fig. 7D) and cAMP augmented tonic firing (P = 0.043 and P = 0.081, respectively) with no significant difference between groups (P = 0.652; Fig. 7D).

Fig. 7. Diabetes does not affect electrical properties of isolated nociceptive neurons.

(A and B) Action potential firing frequency as a function of injected current (0 to 50 pA) in small DRG neurons (<20 μm diameter) from STZ (A) and nondiabetic (B) mice (n = 5 mice per group), in control conditions [control (■), n = 36; STZ (▲), n = 29] or exposed to 50 μM forskolin [control (●), n = 16; STZ (▼), n = 16]. Only neurons showing a tonic-type firing were analyzed [see (C)]. ***P < 0.001 for (A) and **P < 0.01 for (B), Student’s t test after combining the numbers of spikes per second for each neuron at 40- and 50-pA current injection. (C) Examples of the two firing patterns observed in small DRG neurons after 10-pA (left) and 50-pA (right) current injection. Cells showing only one spike or transient firing of a few spikes after 50-pA current injection were classified as strongly adapting or phasic (upper traces), whereas those showing a maintained train of action potentials were classified as tonic (lower traces). (D) Proportions of small DRG neurons from STZ and nondiabetic mice, classified as in (C), in control conditions and after increase in cAMP by forskolin (50 μM). Forskolin increased the ratio of tonic/phasic firing neurons (control, P = 0.043; STZ, P = 0.081), with no significant difference between the two groups (P > 0.05, χ2 test; cell numbers noted above graphs). (E) Maximum current densities in small DRG neurons from STZ and control mice, recorded using a hyperpolarizing voltage step from −60 to −140 mV for 1.5 s, without and with application of 50 μM forskolin. P > 0.05, one-way ANOVA (STZ, n = 39; control, n = 37). (F) Ih activation curves in small DRG neurons. Left: Curves from STZ mice in the absence (■, solid line, n = 32) or presence (●, dotted line, n = 15) of 50 μM forskolin. Right: Curves from nondiabetic mice without (▲, solid line, n = 23) or with (▼, dashed line, n = 16) forskolin. A significant shift to a more depolarized voltage is observed in both cases (***P < 0.001, two-way ANOVA followed by Bonferroni test), but there is no difference between STZ and control mice (P > 0.05, two-way ANOVA).

We used voltage clamp to compare maximum current densities in nociceptive neurons by fully activating HCN channels with a hyperpolarizing voltage step from −60 to −140 mV. Similar maximal current amplitudes were observed in STZ and control mice and in the presence or absence of forskolin, showing that maximum current density is not affected by diabetes or by cAMP increase (P = 0.545; Fig. 7E). This result supports the immunohistological data that showed no up-regulation of HCN2 expression in diabetic neuropathy.

We examined the effect of cAMP on the voltage dependence of HCN channel activation in small DRG neurons from STZ and nondiabetic mice. Forskolin caused a depolarizing shift in HCN half-activation voltage (V1/2) in both STZ (+10.83 mV, P < 0.001 versus pre-forskolin, −97.1 ± 1.6 mV) and control neurons (+12.36 mV, P < 0.001 versus pre-forskolin, −99.0 ± 1.1 mV) (Fig. 7F). However, the magnitude of the shift was not significantly different between diabetic and control neurons (P = 0.779). Finally, to investigate the possibility that the altered concentration of glucose in diabetes might affect the function of HCN channels, we compared Ih currents in neurons in extracellular solutions containing either low (5 mM) or high (25 mM) glucose. Glucose concentration had no effect on activation curves or current densities in either control or STZ neurons (fig. S8).

PDN is associated with increased cAMP in sensory neurons

The immunohistochemical and electrophysiological experiments above show that diabetes does not cause any change in expression of HCN2 or in the electrophysiological properties of isolated nociceptive neurons. The marked effect of diabetic neuropathy on HCN2 function in vivo could instead be due to factors that are present in diabetes and potentiate HCN2 function but are rapidly lost after neuronal isolation. The most likely way in which these unidentified factors might act is by causing an increase of intracellular cAMP concentration in nociceptive neurons, resulting in a potentiation of HCN2 activation through a positive shift in its activation curve on the voltage axis. We therefore examined the effect of diabetes on the cAMP content in whole DRG from control and diabetic mice. Analysis of DRG tissue from STZ-treated mice in which prominent mechanical hypersensitivity had been verified through behavioral tests showed that the cAMP content was increased more than eightfold (nondiabetic, 3.1 ± 1.6 pmol/mg; STZ pain, 26.6 ± 10.3 pmol/mg of total protein; P = 0.014). When samples from fully diabetic mice, which did not exhibit substantial mechanical hypersensitivity, were assessed (STZ no pain), cAMP concentrations were not significantly different from nondiabetic controls (P = 0.811 versus nondiabetic; Fig. 8A).

Fig. 8. Pain in diabetes is associated with increased cAMP in DRG.

(A) Bar graph showing cAMP concentration in lumbar DRG in control (nondiabetic, n = 6), STZ mice with no mechanical hyperalgesia (“STZ no pain,” n = 5), and STZ mice with mechanical pain hypersensitivity (“STZ pain,” n = 4). *P < 0.05, one-way ANOVA followed by SNK post hoc test. (B) Effect of systemic treatment with IVA or different doses of the PKA inhibitor H-89 on von Frey threshold (P < 0.05, IVA versus H-89 at any dose). *P < 0.05 and **P < 0.01, H-89 (5 mg/kg) or IVA versus post-STZ; ##P < 0.01 and ###P < 0.001, IVA versus saline; +++P < 0.001 versus BL, two-way RM ANOVA followed by SNK post hoc test (n = 12 per group).

Apart from binding directly to HCN ion channels, cAMP can also activate the cAMP-dependent protein kinase A (PKA), potentially causing phosphorylation of other targets involved in pain transduction, such as NaV1.8 and TRPV1. We therefore investigated the effect of the PKA inhibitor H-89 on diabetes-induced hyperalgesia at doses found to be effective in reducing PGE2-induced inflammatory pain (fig. S9) (32). Systemic treatment with H-89 (5 mg/kg) had some effect in increasing mechanical pain thresholds 8 weeks after STZ (P < 0.05 versus post-STZ), but the magnitude of this effect was smaller compared to the effect of IVA (P < 0.05 IVA versus H-89 at any dose) (Fig. 8B).

DISCUSSION

PDN is a typical die-back neuropathy, in which unmyelinated C-fibers and thinly myelinated Aδ nociceptive fibers retreat first (33, 34), and in which spontaneous firing develops in small fibers (5, 6, 35). The work reported here provides further support for the idea of spontaneous firing in small nociceptive nerve fibers, because in diabetic mice C-FOS, which is an index of nociceptive afferent activity (25), is induced in outer layers of the dorsal horn, where nociceptive afferents terminate. A critical role for the HCN2 ion channel in maintaining the firing in small-diameter afferent nerve fibers, and thus in promoting the pain of diabetes, is supported by the following: (i) the complete suppression of pain in mouse models of type 1 and type 2 diabetes by the HCN channel blocker IVA, which does not distinguish between HCN isoforms but is peripherally restricted (18), therefore localizing the site of HCN block to the peripheral nervous system; (ii) the similar suppression of the pain associated with diabetes caused by deletion of HCN2 in the NaV1.8-expressing population of nerve fibers, which are predominantly small unmyelinated C-fibers (36, 37); and (iii) the reduction in C-FOS expression in the dorsal horn achieved both by blocking HCN channels with IVA and by genetically deleting HCN2 in NaV1.8-expressing nerve fibers.

As many as 80% of diabetic patients with reduced small-fiber skin innervation, caused by nerve degeneration, also exhibit hyposensitivity to heat (3, 38, 39), which is consistent with the preferential expression of the heat-sensitive ion channel TRPV1 in small unmyelinated nerve fibers (40). An inverse correlation between TRPV1+ fiber skin innervation and heat detection has been directly demonstrated in diabetic patients (41), and we also found an association between degeneration of peptidergic intraepidermal nerve fibers (which include TRPV1+ fibers) and increased heat thresholds in STZ-treated mice.

The painful sensation caused by light touch, known as mechanical allodynia, which is one of the most unpleasant features of PDN, is likely to be conveyed by large myelinated fibers (42). In agreement with this, preventing conduction in large myelinated fibers alleviates mechanical allodynia in PDN (43). How can spontaneous activity in small HCN2-expressing unmyelinated fibers enhance the sensation of mechanical allodynia, which is conveyed via large myelinated fibers? A probable explanation lies in the phenomenon of central sensitization, in which activation of small nociceptive nerve fibers causes a sensation of mechanical allodynia, conveyed via large fibers. Central sensitization is thought to be a result of neural plasticity in the dorsal horn, in which afferent mechanoreceptor input is switched from second-order neurons signaling innocuous light touch to second-order neurons signaling a sensation of pain (44).

One possible explanation for increased firing in nociceptive fibers in the mouse model of diabetes could be an up-regulation of HCN2 expression in sensory neurons, but both immunohistochemical and electrophysiological approaches indicate that HCN2 protein expression and maximal current density were not different in neurons isolated from diabetic and nondiabetic mice. In sensory neurons, however, HCN2 function can be potentiated by an increased intracellular cAMP, which we found to be elevated in DRG from the mouse model of diabetes. Interaction of cAMP with the cyclic nucleotide binding domain in the C-terminal region of HCN2 results in a depolarizing shift in the voltage dependence of activation, leading to an enhanced inward current carried by HCN2 and thus to augmented spontaneous and evoked firing (10). In agreement, raising cAMP concentrations with forskolin in vitro shifted the voltage dependence of Ih toward more positive membrane voltages and increased firing rates in small neurons both in the animal model of type 1 diabetes and in control animals, and previous work has shown that both the shift in Ih and the increased firing rate depend on HCN2 (10).

The mechanisms by which cAMP is elevated in nociceptive neurons in diabetes have not been determined in the present study but are unlikely to involve only inflammatory mediators released at a peripheral location where neurodegeneration takes place, because we found an increase in cAMP in DRG cell bodies, some distance from the site of neurodegeneration. In other pain states, cAMP can be increased by binding of PGE2 or other inflammatory mediators to GPCRs (heterotrimeric guanine nucleotide–binding protein–coupled receptors), such as PGE2 receptor 4 (EP4), resulting in activation of adenylate cyclase and increased cAMP production (10). Several lines of evidence support the idea that diabetes is also associated with a similar proinflammatory phenotype. Diabetic neuropathy increases the amounts of inflammatory mediators in the DRG and sciatic nerve of rodents as well as in human blood (4548). Administration of inhibitors of adenylate cyclase restores mechanical thresholds in diabetic rats, whereas cAMP analogs have the opposite effect (49). The expression of cyclooxygenase-2 (COX-2), which is upstream of PGE2, is increased in diabetic DRG (50) and in sciatic nerve (51), and bradykinin-induced PGE2 secretion is also potentiated by diabetes (52). In support of a sensitizing role of cAMP in sensory neurons, COX-2 inhibitors are analgesic in diabetic mouse models when applied peripherally but not intrathecally (53, 54), and COX-2 KO mice have attenuated signs of peripheral neuropathy after induction of diabetes (55). Other proinflammatory pathways may also be involved in activating adenylate cyclase and so increasing levels of cAMP.

Increased cAMP also activates PKA, which in turn phosphorylates other ion channels in the pain pathway, such as NaV1.8 (56) and TRPV1 (57). Although an involvement of PKA-dependent sensitization of these channels in PDN is possible (58), the complete abolition of hypersensitivity either by an HCN blocker with few off-target actions (18) or by genetic deletion of HCN2 strongly suggests that HCN2 has the major influence in diabetes-induced hyperalgesia. This point is further substantiated by the much larger effect of IVA on diabetic mechanical pain thresholds compared to that of the PKA inhibitor H-89.

We propose that HCN2 sensitization, mediated by increased intracellular cAMP, mediates pain in diabetes both by causing spontaneous firing in nociceptive nerve fibers, due to HCN2 activation at the resting membrane potential, and by promoting higher spiking frequencies in response to a painful stimulus, due to a more rapid “pacemaker” depolarization rate between action potentials. Central sensitization is then activated by the enhanced firing in small nociceptive afferents and causes mechanical allodynia, a painful sensation initiated by activation of large fibers by tactile stimuli. The occurrence of spontaneous firing in C-fibers in STZ-treated rodents has been directly demonstrated using microneurography (6). We found evidence supporting the idea of increased peripheral nociceptive drive in diabetic mice by using C-FOS staining. In addition, diabetes-induced enhancement of firing frequency in C polymodal nociceptors has been reported using a skin-nerve preparation (35).

Diabetic rats treated with the potent TRPV1 agonist resiniferatoxin, which is neurotoxic to the TRPV1-expressing subpopulation of C-fibers, maintain mechanical allodynia (59), an observation that is often cited as indicating that C-fibers are not involved in PDN. However, two major classes of C-fibers are unaffected by resiniferatoxin, namely, nociceptive nonpeptidergic C-fibers expressing the receptor MAS-related GPR family member D (MRGPRD) and low-threshold mechanoreceptors (60). Activity in these C-fibers may be able to drive central sensitization and so cause mechanical allodynia.

Patients with neuropathic pain exhibit a variety of positive symptoms, for example, hyperalgesia and spontaneous pain, together with negative symptoms such as sensory loss, and different treatments may be appropriate depending on the symptoms displayed (61). In that context, our results may be particularly relevant for diabetic patients in whom ongoing pain, likely to be driven by spontaneous activity in peripheral nerves, is a prominent feature. Microneurography studies in humans have shown that peripheral nerve activity is routinely present in neuropathic pain syndromes involving peripheral neuropathy (6264), including diabetic neuropathy (65), and that the incidence of spontaneous activity is higher when accompanied by reports of ongoing pain (7). We note, however, that there may not be an invariant link between spontaneous activity in peripheral nerves and ongoing pain because a number of studies have proposed that changes in CNS processing can also influence the pain of diabetic neuropathy. Potential CNS events affecting neuropathic pain include microglial activation in the spinal cord (66), reduced thalamic activity and communication with cortex (67), loss of gray matter in the thalamus and somatosensory cortex (68), and impaired descending modulation (69).

We found that repeated treatment with IVA provided superior analgesia compared to single administration. Although the mechanism underlying this cumulative effect is not clear, it has been documented before (70) and may indicate the existence of a long-lived IVA metabolite, similar to the morphine metabolite morphine-6-glucuronide, which is 10-fold more potent than morphine itself (71). IVA is already used clinically to treat angina pectoris via inhibition of HCN-dependent firing in cardiac pacemaker cells, which slows heart rate and reduces the oxygen demand of cardiac muscle. The established clinical experience and available pharmacological profiling means that IVA could be swiftly assessed in PDN trials. One consideration will be the bradycardic effect due to HCN4 inhibition in the sinoatrial node; nevertheless, a previous study of IVA for diabetes-associated coronary artery disease showed a good safety profile with no notable bradycardia, visual disturbances, or adverse effects on glucose metabolism (72). In addition, we demonstrate here that targeted genetic deletion of HCN2 in the periphery gives effective pain relief without detectable side effects. Therefore, the discovery of CNS-excluded and HCN2-selective inhibitors is likely to provide marked analgesia in PDN, together with a safe pharmacological profile.

HCN2 channels are closed and generate no inward current over the normal range of the action potential, because their voltage range of activation is negative under nonpathological conditions (9). An important consequence of that is that blockade of HCN2 channels has no effect on baseline neuronal excitability, nor does it modulate pain thresholds under normal conditions. This is in contrast to the effects of blocking NaV channels with local anesthetic, which produces total insensibility to pain, or of human genetic deletion of NaV1.7, which causes complete analgesia even under normal conditions, with consequent self-injury (73). We propose here that in pathological pain states, including PDN, HCN2 channels are recruited as initiators of spontaneous activity only when the voltage range of activation undergoes a positive shift after interaction of inflammatory mediators with Gs-coupled receptors and a consequent elevation of intracellular cAMP. Thus, HCN2 channels are an ideal analgesic target, because their blockade has an analgesic effect only under pathological conditions and should have no effect on normal nociception.

MATERIALS AND METHODS

Study design

The main objective of this study was to test the effect of pharmacological or genetic inhibition of HCN2 on pain in diabetic mice. For behavioral assessment, animals were randomly chosen from multiple cages, provided that they satisfied the inclusion criteria outlined in detail in Supplementary Materials and Methods (“Diabetes models” section). Sample sizes for animal behavior, histology, biochemistry, and electrophysiology were chosen on the basis of previous experience with these assays as the minimum number of independent observations expected to achieve statistical significance. All behavioral testing was conducted during the day in a quiet temperature-controlled room by an experimenter blinded to the identity of drug treatment and/or mouse genotype.

Statistical analysis

All data are presented as means ± SEM, and group sizes are noted in each figure legend. All replicates refer to biological replicates. Statistical significance was determined with Prism (GraphPad Software) or SigmaPlot (Systat Software Inc.) using unpaired, equal variance, two-tailed Student’s t test, linear regression analysis, χ2 test, one-way ANOVA, two-way ANOVA, or two-way ANOVA with RM SNK or Bonferroni tests (for post hoc analysis), as indicated in the corresponding figure legends. Significance is indicated by *P < 0.05, **P < 0.01, and ***P < 0.001. A P value of <0.05 was considered significant.

SUPPLEMENTARY MATERIALS

www.sciencetranslationalmedicine.org/cgi/content/full/9/409/eaam6072/DC1

Materials and Methods

Fig. S1. No effect of IVA on heat thresholds in naïve or diabetic mice.

Fig. S2. Effect of IVA on cardiac parameters in vivo.

Fig. S3. C-FOS induction in the spinal cord 2 hours after complete Freund’s adjuvant.

Fig. S4. No regulation of axonal HCN2 expression by diabetes.

Fig. S5. HCN1 expression in control and diabetic DRG neurons.

Fig. S6. HCN3 expression in control and diabetic DRG neurons.

Fig. S7. HCN4 expression in control and diabetic DRG neurons.

Fig. S8. No effect of high glucose on properties of Ih in nociceptive neurons.

Fig. S9. Reduction of PGE2-induced inflammatory pain by systemic H-89 treatment in naïve mice.

REFERENCES AND NOTES

  1. Acknowledgments: We thank C. Hobbs, L. G. Pinto, and T. Guegan for assistance with experimental procedures and T. Buijs for proofreading the manuscript. Funding: This study was supported by the Medical Research Council (MR/J013129/1, C.T.), the Biotechnology & Biological Sciences Research Council (BB/J009180/1, S.L.; BB/L002787/1, B.V.), and the Wellcome Trust (099259/Z/12/Z, C.T.). Author contributions: C.T. and P.A.M. planned the study and wrote the manuscript. C.T. carried out all in vivo experiments, immunohistochemistry, and biochemistry. S.L. performed DRG culture and HCN2 electrophysiology (Fig. 7). S.W. and I.M. carried out cAMP enzyme-linked immunosorbent assay and HCN2 immunohistochemistry, respectively. B.V. did patch-clamping (fig. S8). Competing interests: P.A.M. is involved in a drug discovery program, funded by the Wellcome Trust, to develop HCN2-selective molecules as analgesics. C.T., S.L.V., S.W., I.M., and B.V. declare no competing interests. Data and materials availability: All data supporting the findings of this study are available within the paper. Transgenic animals are available upon request from the corresponding author.
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