(-)-Hardwickiic acid and Hautriwaic acid induce antinociception via

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(−)-Hardwickiic Acid and Hautriwaic Acid Induce Antinociception via Blockade of Tetrodotoxin-Sensitive Voltage-Dependent Sodium Channels Song Cai,† Shreya S. Bellampalli,† Jie Yu,†,⊥ Wennan Li,† Yingshi Ji,†,# E. M. Kithsiri Wijeratne,§ Angie Dorame,† Shizhen Luo,† Zhiming Shan,†,∇ May Khanna,†,∥ Aubin Moutal,† John M. Streicher,†,‡ A. A. Leslie Gunatilaka,§ and Rajesh Khanna*,†,‡,∥ Downloaded via UNIV OF SOUTH DAKOTA on December 22, 2018 at 06:22:25 (UTC). See https://pubs.acs.org/sharingguidelines for options on how to legitimately share published articles.

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Department of Pharmacology and ‡Neuroscience Graduate Interdisciplinary Program, College of Medicine, §Natural Products Center, School of Natural Resources & the Environment, College of Agriculture & Life Sciences, The University of Arizona, Tucson, Arizona 85724, United States ∥ The Center for Innovation in Brain Sciences, The University of Arizona Health Sciences, Tucson, Arizona 85724, United States ⊥ College of Basic Medical Science, Zhejiang Chinese Medical University, Hangzhou 310058, P.R. China # Department of Pharmacology, College of Basic Medical Sciences, Jilin University, Changchun, Jilin 130021, P.R. China ∇ Department of Anesthesiology, Shenzhen People’s Hospital & Second Clinical Medical College of Jinan University, Shenzhen 518020, P.R. China ABSTRACT: For an affliction that debilitates an estimated 50 million adults in the United States, the current chronic pain management approaches are inadequate. The Centers for Disease Control and Prevention have called for a minimization in opioid prescription and use for chronic pain conditions, and thus, it is imperative to discover alternative non-opioid based strategies. For the realization of this call, a library of natural products was screened in search of pharmacological inhibitors of both voltage-gated calcium channels and voltage-gated sodium channels, which are excellent targets due to their well-established roles in nociceptive pathways. We discovered (−)-hardwickiic acid ((−)-HDA) and hautriwaic acid (HTA) isolated from plants, Croton californicus and Eremocarpus setigerus, respectively, inhibited tetrodotoxin-sensitive sodium, but not calcium or potassium, channels in small diameter, presumptively nociceptive, dorsal root ganglion (DRG) neurons. Failure to inhibit spontaneous postsynaptic excitatory currents indicated a preferential targeting of voltage-gated sodium channels over voltage-gated calcium channels by these extracts. Neither compound was a ligand at opioid receptors. Finally, we identified the potential of both (−)-HDA and HTA to reverse chronic pain behavior in preclinical rat models of HIV-sensory neuropathy, and for (−)-HDA specifically, in chemotherapy-induced peripheral neuropathy. Our results illustrate the therapeutic potential for (−)-HDA and HTA for chronic pain management and could represent a scaffold, that, if optimized by structure−activity relationship studies, may yield novel specific sodium channel antagonists for pain relief. KEYWORDS: Natural products, (−)-hardwickiic acid, hautriwaic acid, voltage-gated sodium channels, non-opioid, HIV-associated sensory neuropathy, chemotherapy-induced peripheral neuropathy

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most long-term pain management4−6 given the inefficacy of opioids and the host of debilitating side effects, including tolerance, addiction, and overdose leading to death accompanying their long-term use.7 Thus, novel therapeutic options are needed to provide alternative drugs for pain that are readily usable by clinicians and patients. In the context of disease, plants have always been key players as bioactive compounds with potential as therapeutics, and this remains true even in the disease of pain. Opium, the natural

INTRODUCTION The International Association for the Study of Pain defines pain as an emotional and sensory experience that is largely unpleasant resulting from actual or potential tissue damage. Chronic pain is then said to be pain that continues past the point of normal healing. This form of persistent pain affects about 20% of the world’s population.1 Multimodal therapy is recommended for chronic pain management and includes medications from different classes (e.g., NSAIDs, selective COX-2 inhibitors, topical capsaicin, corticosteroid, gabapentin,2 or amitriptyline3). In favor of safer alternatives, The Centers for Disease Control and Prevention (CDC) have recommended the reduced use and prescription of opioids for © XXXX American Chemical Society

Received: November 7, 2018 Accepted: December 7, 2018 Published: December 7, 2018 A

DOI: 10.1021/acschemneuro.8b00617 ACS Chem. Neurosci. XXXX, XXX, XXX−XXX

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ACS Chemical Neuroscience

in DRGs by challenging them with veratridine (30 μM), a functional agonist of voltage-gated Na+ channels used in screens for Na+ channel blockers because it opens Na+ channels and prevents them from entering into the inactivated state.16 In contrast to the modest degree of inhibition of KCltriggered efflux observed with some extracts, we noted a significant block of veratridine-triggered sodium channel activity with >15 compounds (data not shown). This functional screening campaign led to the selection of (−)-Hardwickiic acid ((−)-HAD) (Figure 1A), isolated from

source from which many opiates have been extracted, can be dated back to B.C. periods, as early as 3400 B.C., to the poppy plants of lower Mesopotamia, but it was not until 1805, when Friedrich Wilhelm Adam Sertürner isolated the archetypal opioid morphine,8 that campaigns for discovery of painrelieving substances from natural sources were kick-started. An exemplar of the search for novel non-opioid therapeutics for severe pain is ziconotide (Prialt), a peptide isolated from the fish-hunting cone snails, Conus geographus,9 that blocks the Ntype voltage-gated calcium (CaV2.2) channel.10 Prialt was approved by the United States Food and Drug Administration (FDA) in 2004 and is used readily in treatment of various pain conditions, for example, neuropathic and nociceptive pain.11 We recently reported the antinociceptive efficacy of betulinic acid, a bioactive natural product isolated from the desert plant Hyptis emoryi, in preclinical models of neuropathic pain via action on N- and T-type voltage-gated calcium (CaV3.2) channels.12 Here, our screening of a library of natural products for antinociceptive compounds identified two: (i) (−)-hardwickiic acid ((−)-HDA) from the aerial part of a Salvia wagneriana plant Croton californicus (Euphorbiaceae),13 and (ii) hautriwaic acid (HTA) from Eremocarpus setigerus (Croton setigerus) (Euphorbiaceae).14 C. californicus is an herb native to the Mohave Desert.15 Native Americans have used application of its powdered leaves for treating rheumatism.16 E. setigerus (C. setigerus) is an ornamental plant native to western North America, including California.17 Similarly, Native Americans have applied the fresh leaves of this plant as a counter-irritant for pain, and a weak decoction of the plant was both taken internally for chills and fever and used for treatment of typhoid and other fevers.18 Both extract compounds inhibited voltagegated sodium (NaV1.x) channels, which are well established as key players in chronic pain.13,14 While structurally similar, (−)-HDA and HTA showed different specificities among the subtypes of voltage-gated sodium channels tested. Neither compound was a ligand at opioid receptors. Failure to inhibit spontaneous postsynaptic excitatory currents indicated a preferential targeting of voltage-gated Na+ channels over voltage-gated Ca2+ channels by these extracts. Finally, we identified the potential of both (−)-HDA and HTA to reverse chronic pain behavior in preclinical rat models of HIV-sensory neuropathy and of (−)-HDA specifically, in chemotherapyinduced peripheral neuropathy. Our results illustrate the therapeutic potential for (−)-HDA and HTA for chronic pain management and could represent a scaffold, that, if optimizable by structure−activity relationship studies, may yield novel specific sodium channel antagonists for pain relief.

Figure 1. Chemical structures of (−)-Hardwickiic acid ((−)-HDA) and Hautriwaic acid (HTA). (A) Chemical structure of (−)-HDA. (B) Chemical structure of HTA.

C. californicus, and hautriwaic acid (HTA) (Figure 1B), extracted isolated from a Dodonaea viscosa plant E. setigerus, for further investigation into their potential as novel natural inhibitors of voltage-gated Na+ channels. It is interesting that both HDA and HTA have the same carbon skeleton and belong to the clerodane ditepenoids class on natural products. However, structurally they are different as the tertiary methyl group at C-5 in (−)-HDA has been oxidized to a hydroxy methylene group in HTA. (−)-HDA and HTA Inhibit Total Na+ Currents in DRG Neurons. Using whole-cell patch clamp electrophysiology, we next tested the inhibitory potential of the extracts on total Na+ currents in rat DRG sensory neurons. DRG neurons express both tetrodotoxin (TTX)-sensitive rapidly activating and inactivating currents upon membrane depolarization (NaV1.1, NaV1.3. NaV1.6 and NaV1.7) and TTX-resistant slowly activating currents (NaV1.8, NaV1.9, and NaV1.10), with NaV1.7 accounting for ∼75% of the total Na+ current in small-diameter (i.e., nociceptive) sensory neurons.17 we held DRG neurons at −60 mV and used a current (I)−voltage (V) protocol, which consisted of 150 ms step depolarizations ranging from −70 to +60 mV (in +5 mV increments) to test the ability of the extracts to inhibit peak inward sodium currents. Figure 2A displays representative peak sodium currents, recorded from DRG neurons treated with 0.1% dimethyl sulfoxide (DMSO; control) or 20 μM (−)-HDA or HTA. In our initial screen, there was no difference in sodium channel inhibition when we used 50 or 20 μM of the compounds. We could not achieve higher concentrations because of the solubility limit of the compounds. We used 20 μM for our experiment as it was the lowest concentration achieving the maximum inhibition of sodium currents that we could observe. Currents were obtained by a step to −5 mV. Both compounds significantly inhibited peak inward sodium currents (Figure 2B). To account for population diversity in terms of neuronal size, these peaks were normalized by cell capacitance and subsequently displayed as peak current density (pA/pF). Peak currents were inhibited by ∼50% with

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RESULTS AND DISCUSSION Selective Natural Product Extracts Preferentially Inhibit Voltage-Gated Na+ Channels. To discover novel compounds with therapeutic potential for pain relief, we screened a library of natural product extracts for small molecules capable of targeting voltage-gated Ca2+ and voltage-gated Na+ channels. This was accomplished using Fura2-acetoxymethyl, a ratiometric high-affinity intracellular calcium indicator, in rodent sensory dorsal root ganglion (DRG) neurons. First, we challenged the neurons with either 40 mM KCl (to trigger Cav3.x channels) or 90 mM KCl (to open high-voltage activated Ca2+ channels, i.e., Cav1.x and Cav2.x)15 and then treated the neurons with extracts from the natural product library. Second, we screened the same extracts B

DOI: 10.1021/acschemneuro.8b00617 ACS Chem. Neurosci. XXXX, XXX, XXX−XXX

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ACS Chemical Neuroscience

tion properties of sodium currents in DRG neurons. Comparing the midpoints (V1/2) and slope factors (k) in response to changes in command voltages (see Methods) of whole-cell ionic conductances allowed for measurement of changes in activation and inactivation of sodium currents for the DRG cells treated with the compounds. Representative Boltzmann fits for all treatment conditions are shown in Figure 2D and Table 1. There were no changes in the steady-state activation properties of sodium currents, per analysis of V1/2 and k values, between DRG neurons of any treatment condition (Table 1). There was a hyperpolarizing shift of ∼7 mV in the V1/2 of steady-state inactivation in the presence of HTA (Table 1), while the slope was unaffected. (−)-HDA and HTA Inhibit TTX-Sensitive, but Not TTXResistant, Sodium Currents. To further understand the interaction of (−)-HDA and HTA on voltage-gated Na+ channels, we dissected TTX-sensitive and TTX-resistant sodium current subtypes in DRG cells. Because of differential inactivation kinetics of TTX-resistant and TTX-sensitive channels, the fast-inactivation protocol (see Methods) allowed subtraction of electrically isolated TTX-R (current available after −40 mV prepulse) from total current (current available after −120 mV prepulse), as previously described.18 Neurons were subject to overnight treatment of 20 μM (−)-HDA or HTA, or control (0.1% DMSO) as indicated and TTXresistant and TTX-sensitive Na+ currents were subsequently recorded and isolated (Figure 3). Both compounds inhibited TTX-sensitive Na+ currents (Figure 3A, B, 0.1% DMSO: −383.9 ± 73.9 mV; (−)-HDA: −125.4 ± 22.4 mV, p = 0.02; HTA: −234.9 ± 41.6 mV, p = 0.05), but not TTX-resistant Na+ currents (Figure 4A, B, 0.1% DMSO: −206.1 ± 22.1 mV; (−)-HDA: −182.5 ± 50.3 mV; HTA: −172.3 ± 35.7 mV). These results show that the extracts (−)-HDA and HTA preferentially inhibit TTX-sensitive Na+ channels. (−)-HDA and HTA Have Varied Effects on NaV1.x Currents in HEK Cells. To test whether the extracts (−)-HDA and HTA had effects on different sodium channels subtypes, we examined the effects of these compounds at a 20 μM concentration on peak current, steady-state activation, and fast inactivation in human embryonic kidney 293 (HEK293) cells that stably expressed cardiac (hNav1.5) and central nervous system (hNav1.1, rNav1.3) channels using voltage protocols previously reported.19 We observed that (−)-HDA had inhibitory effects on all NaV1.x currents tested; representative currents are shown in Figure 5A, E, and I (Figure 5B, C, NaV1.1:0.1% DMSO: −57.2 ± 9.2 mV versus (−)-HDA: −30.1 ± 5.1 mV; Figure 5F, G, NaV1.3:0.1% DMSO: −99.8 ± 14.6 mV versus (−)-HDA: −45.4 ± 6.3 mV; Figure 5J, K, NaV1.5:0.1% DMSO: −571.1 ± 86.6 mV versus

Figure 2. (−)-HDA and HTA inhibit total Na+ currents in DRG sensory neurons. (A) Representative traces of Na+ currents from DRG sensory neurons treated with 0.1% DMSO (control) or 20 μM (−)-HDA or HTA. Currents were evoked by 150 ms pulses between −70 and +60 mV. Summary of the normalized (pA/pF) sodium current density versus voltage relationship (B) and peak Na+ current density at −5 mV (mean ± SEM) (C) from DRG sensory neurons treated as indicated. (D) Boltzmann fits for normalized conductance G/Gmax voltage relations for voltage dependent inactivation and activation of sensory neurons treated as indicated. V1/2 values for activation and inactivation are presented in Table 1. Asterisks indicate statistical significance compared with cells treated with 0.1% DMSO (*p < 0.05, unpaired two-tailed Student’s t test, n > 8 cells per condition).

(−)-HDA and ∼54% with HTA compared to control-treated DRGS (Figure 2C). As this current decrement could be as a result of changes in channel gating, we next asked if the extracts could alter voltage-dependent activation and inactiva-

Table 1. Effects of (−)-HDA and HAT on Gating Properties of Voltage-Gated Sodium Channels in DRG Neuronsa activation V1/2 k inactivation V1/2 k

control

(−)-HDA

HTA

−25.6 ± 0.7 (23) 5.7 ± 0.6 (23)

−16.9 ± 0.6 (16) 6.1 ± 0.6 (16)

−16.8 ± 0.7 (22) 6.6 ± 0.6 (22)

−29.9 ± 1.5 (24) −14.2 ± 1.2 (24)

−30.7 ± 0.8 (14) −7.7 ± 0.7 (14)

−37.0 ± 1.0 (21)*b −10.9 ± 0.9 (21)

Values are means ± SEM calculated from fits of the data from the indicated number of individual cells to the Boltzmann equation; V1/2, midpoint potential (mV) for voltage-dependent activation or inactivation; k, slope factor. bSignificantly different from the value for control (*P < 0.05; Student’s t test). a

C

DOI: 10.1021/acschemneuro.8b00617 ACS Chem. Neurosci. XXXX, XXX, XXX−XXX

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Figure 3. (−)-HDA and HTA have inhibitory effects on TTX-sensitive currents. (A) Representative traces of Na+ currents from DRG sensory neurons treated with 0.1% DMSO (control) or 20 μM (−)-HDA or HTA. Currents were evoked by 1000 ms prepulses between −120 and −40 mV. Summary of the normalized (pA/pF) sodium current density versus voltage relationship (B) and peak TTX-sensitive Na+ current density (C) at −120 mV (mean ± SEM) from DRG neurons treated with 0.1% DMSO (n = 17) (vehicle), 20 μM (−)-HDA (n = 9), or 20 μM HTA (n = 17). Asterisks indicate statistical significance compared with cells treated with 0.1% DMSO (*p < 0.05, unpaired two-tailed Student’s t test).

Figure 4. (−)-HDA and HTA do not affect TTX-resistant sodium currents. (A) Representative family of TTX-resistant current traces from DRG sensory neurons treated with 0.1% DMSO (control) or 20 μM (−)-HDA or HTA. (B) Summary of the normalized (pA/pF) sodium current density from TTX-resistant sodium currents. (C) Bar graph showing peak TTX-sensitive Na+ current density at −10 mV (mean ± SEM) from DRG neurons treated with 0.1% DMSO (vehicle), 20 μM (−)-HDA or 20 μM HTA as indicated. No asterisks are present, indicating no statistical significance compared with cells treated with 0.1% DMSO (n > 8 cells per condition) (*p < 0.05, unpaired two-tailed Student’s t test).

(−)-HDA: −330.8 ± 60.9 mV). In contrast, HTA did not affect any of the NaV1.x currents (Figure 5B, C, NaV1.1:0.1% DMSO: 57.2 ± 9.2 mV versus HTA: −51.7 ± 7.7 mV; Figure 5F, G, NaV1.3:0.1% DMSO: −99.8 ± 14.6 mV versus HTA: −104.3 ± 26.5 mV; Figure 5J, K, NaV1.5:0.1% DMSO: −571.1 ± 86.6 mV versus HTA: −607.8 ± 106.2 mV). In addition, (−)-HDA and HTA did not affect the biophysical properties of sodium channels, NaV1.1, NaV1.3, or NaV1.5 (Figure 5D, H, L). These data show the inhibitory function of (−)-HDA on NaV1.1, NaV1.3, and NaV1.5 while HTA had no effect on these Na+ channel subtypes. (−)-HDA and HTA Have No Effect on Potassium Currents in DRG Sensory Neurons. Potassium currents have an important role in modulating neuronal excitability, and potassium channel reduction is thought to contribute to the increased excitability and generation/patterning of spontaneous activity in sensory neurons following peripheral nerve injury. Therefore, we next tested whether (−)-HDA or HTA could affect potassium currents (IK). DRG neurons were treated with 20 μM of either (−)-HDA or HTA, or 0.1% DMSO as a vehicle control, and total potassium current (IK, including fast inactivating IK,A and slowly inactivating IK,S) was recorded (Figure 6A). The current density−voltage and the peak current density for both IK,A (Figure 6B, C) and IK,S (Figure 6E, F) were not different between 0.1% DMSO and (−)-HDA or HTA treated neurons. Plotting voltage relation to the I/Imax ratio did not show any alteration of the inactivation properties for IK,A currents (Figure 6D). Thus, we conclude

that neither (−)-HDA or HTA has an effect on potassium currents. Heterogenous Receptor-Based Fingerprinting of (−)-HDA and HTA on DRG Neurons. The results presented thus far collectively demonstrate the potential for (−)-HDA and HTA to inhibit TTX-sensitive voltage-gated Na+ channels. These results do not uncover cell-specific neuronal classes that may be targeted by (−)-HDA and HTA. We used the previously described constellation pharmacology protocol20,21 to explore cell-specific functional consequences of (−)-HDA and HTA treatment. The constellation pharmacology assay uses six successive stimulations, each 6 min apart, to compare Ca2+ influx due to activity of Ca2+-associated membrane proteins: Ca2+ permeable ligand-gated ion channels, metabotropic receptors and voltage-gated Ca2+ channels. Following these 6 challenges, KCl-evoked response due to membrane depolarization is used to assess viability of neurons; neurons not responsive to KCl are excluded from analysis. The heterogeneity of neuronal responsivity is demonstrated via example traces of sensory neurons treated with control (0.1% DMSO) and with a 20 μM concentration of (−)-HDA or HTA (Figure 7A−C) as indicated. Notably, inhibition of KCl-evoked Ca2+ influx was seen as a result of treatment with (−)-HDA and HTA; this slight inhibition is consistent with our screening data, and also suggests a heterogeneity in effects of these compounds on voltage-gated ion channels. Sensory neurons were incubated overnight with the specified treatment, control (n = 1109), (−)-HDA (n = 1390) or HTA (n = 1812), and subsequently imaged with the constellation pharmacology D

DOI: 10.1021/acschemneuro.8b00617 ACS Chem. Neurosci. XXXX, XXX, XXX−XXX

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Figure 5. Analysis of (−)-HDA and HTA on electrophysiological properties of Nav1.1, Nav1.3, and Nav1.5 currents in HEK293 cells. (A) Representative traces of currents from HEK293 cells expressing Nav1.1, Nav1.3, and Nav1.5 in the presence of the indicated treatents are shown. Summary of the normalized (pA/pF) sodium current density versus voltage relationships (B, F, J) or peak Na+ current density at 0 mV (NaV.1 and NaV1.3) or −20 mV (for NaV1.5) (mean ±SEM) (C, G, K) from HEK293 cells stably expressing NaV1.1., NaV1.3, and NaV1.5 and treated with 0.1% DMSO (control, n = 9), 20 μM (−)-HDA (n = 7), or 20 μM HTA (n = 6). (D, H, L) Representative Boltzmann fits for steadystate fast inactivation and activation for HEK293 cells treated with 0.1% DMSO (control) or 20 μM of extracts. Values for V1/2, the voltage of halfmaximal inactivation and activation, and the slope factors (k) were derived from Boltzmann distribution fits to the individual recordings and were averaged to determine the mean (±SEM) voltage dependence of steady-state inactivation and activation, respectively. Asterisks indicate statistical significance compared with cells treated with 0.1% DMSO (*p < 0.05, unpaired two-tailed Student’s t test).

protocol. Data was collected from 3 independent experiments, and individual neuronal responses to each constellation trigger were analyzed. Responses under 10% of baseline fluorescence were excluded from our analyses. We first determined how treatment with (−)-HDA or HTA would change overall functionality of sensory neurons. To test this, we assessed whether treatment with (−)-HDA or HTA would affect the functional cell subclasses (in number) present in populations of neurons (Figure 7D). In comparison to control-treated cells, the number of functional subclasses in (−)-HDA or HTA-neuronal populations were not altered. We then assessed whether the response of neurons to the number of stimulatory challenges, independently of which specific agonists triggered response, was altered in response to (−)-HDA or HTA treatment (Figure 7E). Compared to control-treated cells, more cells treated with (−)-HDA or HTA responded to three stimulatory challenges and less cells responded to only one challenge. HTA-treatment led to a decrease in cells responding to two stimulatory challenges and an increase in cells responding to four stimulatory challenges. Next, we asked if (−)-HDA or HTA affected the sensitivity of DRGs to the different constellation triggers, by analyzing the percent of cells responding to a specific receptor agonist, independently of any other constellation triggers these neurons may have responded to (Figure 7F). Cells treated with (−)-HDA or HTA exhibited decreased response to capsaicin challenge, but increased response to ATP. HTA-treated cells

also showed an inhibited response to allyl isothiocyanate (AITC) stimulus. This analysis showed that treatment with (−)-HDA or HTA increases sensitivity to ATP stimulation, and treatment with HTA additionally increases AITC sensitivity. Overall, both (−)-HDA and HTA increase functional competence of cells. In a more specific inquiry, we also explored how (−)-HDA and HTA altered the extent of Ca2+ influx following specific stimulation by each constellation trigger. Thus, we analyzed peak Ca2+ responses (Figure 7G) and area under the curve (AUC) of these responses in sensory neurons as a result of the treatment. Notably, treatment with (−)-HDA and HTA decreased peak Ca2+ responses due to stimulation by capsaicin, and only HTA increased peak Ca2+ responses due to stimulation by ATP; the AUC of Ca2+ response due to stimulation by capsaicin was decreased, and particularly, AUC of Ca2+ response due to stimulation by AITC was decreased in HTA-treated neurons (Figure 7H). Stimulation by histamine, menthol, ATP, or AITC, did not alter the AUC of Ca2+ response in sensory neurons after treatment with (−)-HDA or HTA. We then asked if (−)-HDA or HTA mediated inhibition of depolarization-evoked Ca2+ response would be altered in a functional class-specific manner. To test this, we assessed average peak Ca2+ response due to KCl challenge in sensory neurons that specifically responded to a particular constellation trigger, independent of any other constellation triggers they E

DOI: 10.1021/acschemneuro.8b00617 ACS Chem. Neurosci. XXXX, XXX, XXX−XXX

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physiological recording of substantia gelatinosa neurons (Figure 9) to test if either extract, (−)-HDA or HTA, could inhibit spontaneous excitatory postsynaptic currents (sEPSC). There was no significant decrease in spontaneous EPSC amplitude (postsynaptic effect) or spontaneous EPSC frequency (presynaptic effect) of neurons treated with 0.1% DMSO, (−)-HDA, or HTA (20 μM). This indicates that (−)-HDA and HTA do not likely affect the nociceptive pathway via targeting of calcium-dependent neurotransmitter release. Thus, it is more probable that these compounds achieve inhibitory effects on nociceptive signaling via preferentially targeting voltage-gated sodium channels over calcium channels. HIV-Induced Sensory Neuropathy Is Alleviated by Treatment with (−)-HDA or HTA. The above results identify (−)-HDA and HTA as novel compounds inhibiting TTX-S voltage-gated sodium channels. These channels are known to participate in neuropathic pain signal transmission.22−25 Sensory neuropathy is a frequent complication of Human Immunodeficiency Virus (HIV) infection.26 Since voltage-gated sodium channels have been shown to play a role in neuropathic pain,22−25 we explored the potential of (−)-HDA and HTA tocause antinociception in HIV-induced sensory neuropathy. To test this, we submitted rats to three intrathecal injections of the HIV-1 envelope glycoprotein (gp120) which resulted in the development of mechanical allodynia (Figure 10A), consistent with previous reports.27,28 Intrathecal (i.th.) injection of (−)-HDA or HTA (2 μg/5 μL) reversed mechanical allodynia 1−2 h postinjection and lasted for 2−3 additional hours (Figure 10A). This reversal of mechanical allodynia by (−)-HDA-treatment or HTA-treatment is supported by a significant increase of the area under the curve between the (−)-HDA treated animals or HTAtreated animals compared to vehicle (Saline) treated animals (Figure 10B). We conclude that both (−)-HDA and HTA have antinociceptive potential for HIV-induced sensory neuropathy. (−)-HDA, but Not HTA, Reverses ChemotherapyInduced Neuropathy Behavior in Rats. We next tested if sodium channel inhibition by (−)-HDA or HTA could be beneficial to reverse nociceptive behaviors in a rodent model of peripheral neuropathy due to paclitaxel injection. Rats received 4 injections of paclitaxel (2 mg/kg) and per a decreased paw withdrawal threshold, developed mechanical allodynia (Figure 10C). We injected (−)-HDA or HTA (2 μg/5 μL, i.th.) and observed significant relief of mechanical allodynia even just 0.5 h after injection for the (−)-HDA condition (Figure 10C). This effect lasted for 2−3 additional hours (Figure 10C) before returning back to predrug baseline. Area under the curve shows a significant antinociceptive effect of (−)-HDA-treated animals compared to vehicle (saline) injected animals (Figure 10D). In contrast, this reversal was not observed in HTA-treated animals (Figure 10C), and further confirmed by the AUC (Figure 10D). These results show that (−)-HDA, but not HTA, has the potential for pain relief in paclitaxel induced peripheral neuropathy.

Figure 6. (−)-HDA and HTA do not affect potassium currents in DRG neurons. (A) Representative total, IK,A, and IK,S currents recorded from DRGS neurons treated with 0.1% DMSO (control) or 20 μM (−)-HDA or HTA. The current density−voltage relationship (B) and the peak current density (C) for IK,A was not different between 0.1% DMSO and (−)-HDA or HTA treated neurons. There were no differences in the inactivation of IK,A (D) in neurons from any of the groups. The current density−voltage relationship (E) and the peak current density (F) for IK,S was not different between 0.1% DMSO and (−)-HDA or HTA treated neurons (7−10 per condition).

might have responded to. Treatment with (−)-HDA or HTA significantly decreased KCl-evoked Ca2+ response in sensory neurons that responded to an ATP or capsaicin stimulus; HTA additionally decreased KCl-evoked Ca2+ response in sensory neurons that responded to an ACh stimulus. However, DRGs that responded to AITC, histamine, menthol, and capsaicin did not have altered KCl-evoked response due to either drug treatment (Figure 7I). These results help in uncovering the mechanism of inhibition of nociceptive signaling by (−)-HDA and HTA. Neither (−)-HDA nor HTA Binds to the Orthosteric Site of the Opioid Receptors. To confirm that (−)-HDA and HTA do not produce potential antinociception through off-target binding to the opioid receptors, we performed competition radioligand binding at all three opioid receptors in vitro. We competed (−)-HDA or HTA and a positive control compound (naloxone for MOR and DOR, U50,488 for KOR) vs a static concentration of 3H-diprenorphine in Chinese hamster ovary (CHO) cells containing the human μ (MOR), δ (DOR), or κ (KOR) opioid receptor. We found that neither compound bound to any opioid receptor up to a 10 μM concentration (Figure 8). In contrast, the positive control compounds bound to all three targets with expected affinity (Figure 8). These results strongly suggest that both (−)-HDA and HTA are not engaging the opioid receptors, and that potential antinociception would thus not be a result of opioid receptor association. (−)-HDA and HTA Do Not Alter Spontaneous Excitatory Postsynaptic Currents. To further interrogate the preferential targeting of sodium currents over calcium currents by (−)-HDA and HTA, we performed electro-

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CONCLUSIONS Here, we report that (−)-HDA and HTA, two structurally related plant-derived natural products, inhibit voltage-gated sodium channels that are sensitive to tetrodotoxin. The extracts had modest to no effects on voltage-gated calcium and voltage-gated potassium channels, respectively. Notably, F

DOI: 10.1021/acschemneuro.8b00617 ACS Chem. Neurosci. XXXX, XXX, XXX−XXX

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Figure 7. Functional “fingerprinting” of sensory neuron subclasses following treatment with (−)-HDA and HTA. (A) Representative traces of sensory neurons treated with 0.1% DMSO (vehicle), (−)-HDA (20 μM) (B), and HTA (20 μM) (C) responding to constellation pharmacology triggers (menthol (400 nM), histamine (50 μM), ATP (10 μM), AITC (200 μM), acetylcholine (1 mM), capsaicin (100 nM), and KCl (90 mM)) during Ca2+ imaging. Each trace represents an individual neuron; a typical experimental trial records the responses of >200 neurons concurrently. The x-axis represents time in seconds, and the y-axis shows the relative intracellular calcium [Ca2+] in each DRG neurons (i.e., the F340/F380 ratio). (D) Number of overall functional DRG sensory neuronal classes as a result of treatment with vehicle or drug (20 μM). (E) Percentage of DRG sensory neurons that responded to indicated number of triggers. “0” indicates those neurons that only responded to none other than KCl stimulus. (F) Percentage of sensory neurons responding to major classes of constellation triggers. Average peak response (G) and area under the curve (H) is shown for calcium response in sensory neurons post indicated treatment, after stimulation by major classes of constellation triggers. Area under the curve was calculated with Graphpad Prism software using the trapezoid rule. (I) Average peak KCl-evoked response of sensory neurons post indicated treatment. Statistical significance compared with cells treated with 0.1% DMSO are indicated by asterisks (*p < 0.05; Student’s t test). Abbreviations for constellation triggers are as follows: ACh, acetylcholine; AITC, allyl isothiocyanate; ATP, adenosine triphosphate; Hist, histamine; Ment, menthol; Cap, capsaicin; KCl, potassium chloride. Data was acquired from a total of three independent experiments with an overall n of 1109−1812 neurons (from three coverslips) for vehicle (0.1% DMSO) and (from three coverslips) for (−)-HDA (20 μM) and (from three coverslips) for HTA (20 μM).

additional to currents from NaV1.7, this includes sodium currents from NaV1.1 and NaV1.6.17 All of these channels have been linked to painful phenotypes. For example, Salvatierra and colleagues reported that inhibition of NaV1.1 channels reduces visceral hypersensitivity associated pain and chronic abdominal pain linked to irritable bowel syndrome.40 Likewise, mice lacking NaV1.6 channels exhibit relief from neuropathic pain models.41 Since our data shows that HTA did not affect NaV1.1 currents, it is possible that its inhibition of total sodium currents in DRG neurons, and subsequent inhibition of nociceptive behavior, arises from actions largely on NaV1.6 and NaV1.7; (−)-HDA appears to be involved in inhibition of pan-TTX-sensitive sodium subtypes. These data suggest that derivatives of the natural product (−)-HDA could be useful for pain relief in neuropathic pain and visceral hypersensitivity. In small-diameter sensory neurons, the NaV1.7 subtype seems to contribute to the majority of TTX-sensitive currents.17 It has been reported that although loss of neuronal

(−)-HDA and HTA did not interact with opioid receptors. The block of tetrodotoxin-sensitive pan-NaV1.X channels supported the antinociceptive value of the tested compounds in rodent neuropathic pain models, setting the stage for their development as naturally derived therapeutics for chronic pain. Voltage-gated sodium channels are critical targets in the nociceptive pathway due to their electrical properties and roles in propagating electrical signaling through allowing influx of Na+ ions during the climbing segment of the action potential. NaV1.7, NaV1.8, and NaV1.9 channels are preferentially found in the peripheral nervous system.29 NaV1.7, also known as PN1, is localized to the sympathetic ganglia, dorsal root ganglia and trigeminal ganglia, but only at low levels within the brain.30 This high level of NaV1.7 expression has been reported in multiple species including rodents, nonhuman primates, as well as humans.31−36 While NaV1.7 produces rapidly inactivating and activating TTX-sensitive currents,37 NaV1.8 and NaV1.9 produce TTX-resistant currents.38,39 Our results show that (−)-HDA and HTA act on TTX-sensitive sodium channels; G

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DRG NaV1.7 does not alter peak amplitude of action potentials (APs), the AP rising phase is slowed, preventing about a third of DRG all neurons from NaV1.7 knockout animals from being able to generate APs.42 Thus, targeting this voltage-gated sodium channel may also slow down nociceptive signal transmission. In the context of pain, NaV1.7 has been implicated in potentiating pain signaling. A link between NaV1.7 and inflammatory pain signaling was established when conditional elimination of NaV1.7 channels in sensory neurons led to loss in pain due to inflammatory challenges.43 The participation of NaV1.7 channels in potentiating inflammatory pain signaling supports the action of (−)-HDA) in animal pain models. Moreover, (−)-HDA) has been previously linked to anti-inflammatory and antinociceptive properties.44 NaV1.7 gain-of-function mutations in humans are implicated in painful conditions such as inherited Erythromelalgia (IEM),45 a chronic painful condition that is typified by pain attacks. Stephen Waxman’s group showed that many IEM-associated mutations lead to increased excitability of NaV1.7 channels.46 These data provide the basis for development of new analgesic pharmacotherapies like (−)-HDA and HTA that could be free of typical opioid-like side effects. There is also confirmation that NaV1.7 channels contribute to chemotherapy-induced peripheral neuropathy phenotypes. Li and colleagues showed upregulated NaV1.7 channels caused by paclitaxel-induced neuropathy in rats and humans.47 These data again provide support for the efficacy of (−)-HDA and HTA on TTXsensitive currents. The two compounds tested here are subtly different structurally (the tertiary methyl group at C-5 in (−)-HDA has been oxidized to a hydroxy methylene group in HTA), but they had very different pharmacological profiles; the reason for this is currently unclear but will be pursued through extensive structure−activity relationship (SAR) studies to achieve NaV1.x selective compounds. While we show that (−)-HDA and HTA work via action through voltage-gated, TTXsensitive channels, (−)-HDA also appears to interact with NaV1.1 and NaV1.5. NaV1.1 has been implicated in potentiation of mechanical pain,40 and NaV1.3 inhibition has been shown to alleviate diabetes-related allodynia.48 However, inhibition of NaV1.5 may pose a cardiac liability as this channel is key to propagating cardiac APs.49 Thus, our findings support the use of both (−)-HDA and HTA as lead compounds lending their structural scaffold for modification to create more selective and thus highly effective antinociceptive drugs.

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METHODS

Animals. Adult Sprague−Dawley rats (male and female, 225− 250g; Harlan Laboratories) were kept in light-controlled (12 h light/ 12 h dark cycle; lights on 07:00−19:00) and temperature-controlled (23 ± 2 °C) rooms with access to rodent chow and water as needed. The University of Arizona’s College of Medicine Institutional Animal Care and Use Committee (IACUC) sanctioned all experiments. All experiments were performed per guidelines recommended published by National Institutes of Health Guide for Care and Use of Laboratory Animals and adhered to International Association for the Study of Pain ethical guidelines. For the behavioral experiments, rats were randomly assigned to control or treatment conditions. Animals were initially housed three/cage and singly after the intrathecal cannulation on a 12 h dark−light cycle with ad libitum food/water. Experimenters performing the behaviors were kept blinded to the experimental treatment conditions.

Figure 8. (−)-HDA and HTA do not bind to opioid receptors. Competition radioligand binding was performed in CHO cells expressing the human MOR, DOR, or KOR (see Methods for details). (−)-HDA, HTA, or a positive control compound was competed against 3H-diprenorphine in all three cell lines. Curves reported as the mean ± SEM of the mean value from each individual experiment in n = 3 independent experiments. KI also reported as the mean ± SEM of the individual value from each of n = 3 independent experiments. (−)-HDA and HTA did not produce competition binding up to 10 μM in any cell line. (A) DOR: naloxone KI = 140 ± 35 nM. (B) KOR: naloxone KI = 27.5 ± 3.8 nM. (C) MOR: KI = 57.7 ± 8.1 nM. H

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Figure 9. Perfusion of 20 μM (−)-HDA or HTA had no effect on spontaneous excitatory synaptic transmission in substantia gelatinosa neurons. (A) Photomicrograph of the spinal cord slice preparation showing that the substantia gelatinosa (SG) can be identified as a translucent pale band in the superficial dorsal horn (lamina I/II) enabling positioning of the recording electrode to this region. Substantia gelatinosa neurons recorded. Infrared differential interference contrast images of the same cell (indicated by a red box in middle panel) with part of the recording electrode after whole-cell configuration. (B) Representative recordings of spontaneous excitatory postsynaptic currents (sEPSCs) from control, (−)-HDA and HTA-treated groups. Amplitude of sEPSCs (C) and frequency of sEPSCs (D) of cells recorded from the indicated groups (n = 12−14). (E) The input resistance of the cells was not changed between the conditions. Data are expressed as means ± SEM (p > 0.05 (versus baseline); one-way ANOVA followed by Tukey’s post hoc test). Materials. Unless noted otherwise, chemicals were purchased from Sigma (St. Louis, MO). The natural products used in this study, (−)-HDA and HTA were isolated respectively from C. calcifornicus13 and E. setigerus14 as previously described. Preparation of Acutely Dissociated Dorsal Root Ganglion Neurons. Dorsal root ganglia (DRG) from all levels were acutely dissociated using methods as described previously.50,54 Approximately 150,000 dissociated DRG neurons were plated onto poly-D-lysineand laminin-coated glass 12 or 15 mm coverslips and cultured for up to 48 h in media consisting of DMEM (1% penicillin/streptomycin sulfate from 10,000 μg/mL stock, 10% fetal bovine serum (Hyclone)), and 30 ng/mL nerve growth factor). Calcium Imaging in Acutely Dissociated Dorsal Root Ganglion (DRG) Neurons. Dorsal root ganglion neurons were bathed for 30 min at 37 °C with a concentration of 3 μM Fura-2AM (Cat# F1221, Thermo Fisher, stock solution prepared at 1 mM in DMSO, 0.02% pluronic acid, Cat#P-3000MP, Life Technologies) to survey changes in intracellular calcium([Ca2+]c) as described before.52 The changes in [Ca2+]c changes were examined with a ratio of F340/ F380, calculated after subtracting background from both channels. Constellation Pharmacology. These experiments were done as reported earlier,21,54 but with the following modifications. Dorsal root ganglia neurons were loaded at 37 °C with a concentration of 3 μM Fura-2AM for 30 min in Tyrode’s solution. After a 1 min baseline measurement, Ca2+ influx was stimulated by the addition of the following receptor agonists: 400 nM menthol, 50 μM histamine, 10 μM adenosine triphosphate (ATP), 200 μM allyl isothiocyanate (AITC), 1 mM acetylcholine (ACh), and 100 nM capsaicin diluted in

Tyrode’s solution. At the end of the constellation pharmacology protocol, cell viability was assessed by depolarization-induced Ca2+ influx using an excitatory KCl solution as described earlier.21,54 After the 1 min baseline measurement, each trigger was applied for 15 s in the order indicated above in 6 min intervals. After each trigger, bath solution was continuously perfused over the cells to wash off excess of the trigger. This process was automated using the ValveBank II perfusion system that controlled the perfusion of the standard bath solution and triggers (Automate Scientific). For the (−)-HDA and HTA conditions, DRGs were incubated overnight with a 20 μM concentration of either compound. Fluorescence imaging was performed under the same conditions noted above for calcium imaging. A cell was defined as a “responder” if its fluorescence ratio of 340/380 nm was greater than 10% of the baseline value calculated using the average fluorescence in the 30 s preceding application of the trigger. Whole-Cell Electrophysiological Recordings of Sodium Currents in Acutely Dissociated DRG Neurons. Recordings were obtained from acutely dissociated DRG neurons as described by us before.51,55,56 The internal solution consisted of (in mM): 140 CsF, 10 NaCl, 1.1Cs-EGTA, and 15 HEPES (pH 7.3, mOsm/L = 290−310) and external solution contained (in mM): 140 NaCl, 30 tetraethylammonium chloride, 10 D-glucose, 3 KCl, 1 CaCl2, 0.5 CdCl2, 1 MgCl2, and 10 HEPES (pH 7.3, mosM/L = 310−315). DRG neurons were interrogated with current−density (I−V) and activation/inactivation voltage protocols as previously before.18,50 The voltage protocols were as follows: (a) I−V protocol: from a −60 mV holding potential, cells were depolarized by 150 ms voltage steps from I

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solution (in millimolar): 140 KCL, 4 ATP, 5 EGTA, 10 HEPES, 5 MgCl2, 0.3 GTP, 2.5 CaCl2, and adjusted pH at 7.3 with KOH. Then the neurons were subjected to activation of IK, fast inactivating IK,A and inactivation IK,A protocols as previously described.57 Membrane holding was at −60 mV. IK activation was determined by incremental voltage steps of 300 ms each, applied at 5 s intervals from −80 to +60 mV (in +10 mV increments). In order to get the fast inactivating IK,A, a 4 s prepulse to −100 or −40 mV was applied then 0.5 s voltage steps ranging from −80 to +40 mV (increment = 20 mV/step). The fast inactivating IK,A was inferred by subtracting, digitally, the currents from the prepulse to −100 mV to the prepulse to −40 mV. Inactivation of IK,A was determined by using a series of 4 s prepulses that ranged from −100 to −40 mV with an increments of +10 mV per step that were immediately followed by a 200 ms step to +60 mV. Preparation of Spinal Cord Slices for Slice-Electrophysiological Recording. As described previously, postnatal rats (postnatal 12−21 days) were put under with isoflurane anesthesia.58 For spinal nerve blocking, 0.3 mL of 2% lidocaine was injected bilaterally into the vertebrae (L4 to 5 lumbar). A midthoracic to low lumbar level laminectomy was done, with the spinal cord being rapidly immersed into cold modified ACSF with 95% O2/5% CO2. The ACSF contained (in mM): 80 NaCl, 75 sucrose, 25 NaHCO3, 3.5 MgCl2, 3.0 sodium pyruvate, 2.5 KCl, 1.3 ascorbate, 1.25 NaH2PO4, and 0.5 CaCl2, with pH = 7.4 and osmolarity = 310 mOsm. Approximately 350 μm thick transverse slices were cut by using a vibratome (VT1200S; Leica, Nussloch, Germany). Slices were bathed for >1 h at room temperature in an oxygenated recording buffer containing (in mM): 125 NaCl, 26 NaHCO3, 25 D-glucose, 3.0 sodium pyruvate, 2.5 KCl, 2 CaCl2, 1 MgCl2, 1.25 NaH2PO4, 1.3 ascorbate, with pH = 7.4 and osmolarity = 320 mOsm. The slices were then positioned in a chamber for recording and uninterruptedly perfused, at a rate of 3 to 4 mL/min, with oxygenated solution before starting recordings. Whole-Cell Patch Recordings of Spontaneous Electric Postsynaptic Currents in Substantia Gelatinosa Neurons of Spinal Cord Slices. Substantia gelatinosa neurons were visualized and identified in the slices by means of infrared DIC video microscopy on a FN1 Nikon upright microscope (Nikon, Tokyo, Japan) outfitted with a 3.40/0.80 objective (water-immersion) and a charge-coupled device camera. Patch pipets with resistance ranging from 6 to 10 MΩ were fabricated from borosilicate glass (Sutter Instruments, Novato, CA) on a four-step micropipet P-90puller (Sutter Instruments, Novato, CA). The internal solution contained the following (in mM): 120 potassium gluconate, 20 HEPES, 20 KCl, 2 MgCl2, 2 Na 2 -ATP, 0.5 Na-GTP, 0.5 EGTA, with pH at 7.28 and osmolarity = 310 mOsm. The membrane was held at −60 mV using Patchmaster. In voltage-clamp mode, whole-cell configurations were obtained. To record spontaneous excitatory postsynaptic currents (sEPSCs), bicuculline methiodide (10 μM) and strychnine (1 μM) were added to the recording solution to block γ-aminobutyric acid-activated (GABA) and glycine-activated (GlyR) currents, respectively. Hyperpolarizing step pulses (5 mV in intensity, 50 ms) were periodically delivered to test the access resistance (15−25 MΩ), and recordings were stopped if the access resistance changed by >20%. For each recording, sEPSCs were acquired for a duration of 2 min. Currents were filtered and digitized at 3 and 5 kHz, respectively. Data were evaluated by the Mini-Analysis (Synatosoft Inc., NJ) and Clampfit 10.7 Program. The amplitude and frequency of sEPSCs were compared between neurons from animals in control and drug-treated groups. All data were collected in 1 h within drug perfusion. Cell Culture and Transient Transfection. Human derived HEK293 cells stably expressing hNaV1.1, rNaV1.3, or hNaV1.5 were cultured in standard cell culture conditions, 37 °C in 5% CO2, as previously reported.59 Implantation of Intrathecal Catheter. For drug administration via intrathecal (i.t.) route, rats were implanted with catheters as described by previously.60 Testing of Allodynia. The assessment of tactile allodynia was as before.61

Figure 10. (−)-HDA and HTA reduce gp120-induced and paclitaxelinduced mechanical allodynia. (A) Paw withdrawal threshold of adult male rats (n = 6) was measured 15 days after three intrathecal injections of gp120. Rats were treated with (intrathecally (i.th.) via catheter) saline (vehicle) or (−)-HDA (2 μg/5 μL) or HTA (2 μg/5 μL) as indicated. Asterisks indicate statistical significance compared with saline treatment (*p < 0.05; two-way ANOVA with a Student− Neuman−Kuels post hoc test). (B) Area under the curve was derived as indicated above using GraphPad Prism. Statistical significance is indicated by asterisks (*p < 0.05, Mann−Whitney test) in comparison to vehicle-treated rats. (C) Paw withdrawal threshold of adult male rats (n = 6) was measured 15 days after 4 intraperitoneal injections of 2 mg/kg paclitaxel. Rats were treated intrathecally with saline (vehicle) or (−)-HDA (2 μg/5 μL) or HTA (2 μg/5 μL) as indicated. Asterisks indicate statistical significance compared with saline treatment (*p < 0.05; two-way ANOVA with a Student− Neuman−Kuels post hoc test). (D) Area under the curve was derived again as indicated before using Graphpad Prism. Statistical significance is indicated by asterisks (*p < 0.05, Mann−Whitney test) in comparison to vehicle-treated rats. Experimenter was blinded to the treatment condition. −70 to +60 mV (5 mV increments) which permitted acquisition of current density values such we could analyze activation of sodium channels as a function of current vs voltage and infer peak current density (normalized to cell capacitance (in picofarads, pF)), which occurred between ∼0 to 10 mV; (b) inactivation protocol: from a −60 mV holding potential, cells were subjected hyperpolarizing/ repolarizing pulses for 1 s between −120 and 0 mV (+10 mV steps). This increment conditioned various proportions of channels into a state of fast-inactivation; in this case, 0 mV test pulse for 200 ms was able to reveal fast inactivation when normalized to maximum sodium current. Because of differential inactivation kinetics of TTX-resistant and TTX-sensitive channels, the fast inactivation protocol allowed subtraction of electrically isolated TTX-R (current available after −40 mV prepulse) from total current (current available after −120 mV prepulse), as previously described.18 To test for TTX-resistant currents, I−V protocol was conducted after 5 min of bathing in a concentration of 1 μM TTX. Pipettes with 1−3 MΩ resistance were used for all recordings. To isolate potassium currents (IK), DRG neurons were bathed in buffer composed of (in millimolar): 140 N-methyl-glucamine chloride, 10 glucose, 10 HEPES, 1 MgCl2, 2 CaCl2, and 5 KCl, pH adjusted to 7.4 with KOH. Recording pipets were filled with internal J

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ACS Chemical Neuroscience Radioligand Binding in CHO Cells. The MOR-CHO cells was obtained from PerkinElmer (#ES-542-C). The DOR cell line was made and characterized as reported.62 The KOR cell line was also created in our lab; an N-terminal 3X-hemagglutinin tagged human KOR expression clone from Genecopoeia was electroporated into parental CHO cells and selected with 500 μg/mL G418. The resulting selected population was enriched for receptor expression by live cell labeling with anti-HA-Alexa488 antibody and separating the top 2% of the population by flow cytometry. This high expressing population was characterized by immunocytochemistry and Western blot to establish receptor expression and expected signal transduction activation. All 3 cell lines were characterized by saturation radioligand binding with 3H-diprenorphine, and the measured KD used in competition binding experiments to calculate the KI (MOR = 5.23 nM; DOR = 0.93 nM; KOR = 1.81 nM; all the mean of N ≥ 3 independent experiments). All cells were cultured as described before.62 Competition Radioligand Binding. Competition radioligand binding experiments were performed as before.62 HIV-Induced Sensory Neuropathy (HIV-SN). Mechanical allodynia is produced by i.th. injection of human immunodeficiency virus-1 (HIV-1) envelope glycoprotein, gp120.28,53 The compounds were assessed for their ability to affect mechanical allodynia at 10−14 days after the first injection of gp120. Paclitaxel-Induced Peripheral Neuropathy. Rats given paclitaxel (cat# P-925-1, Goldbio, Olivette, MO), based on the protocol described by Polomano et al., developed mechanical allodynia over the course of 7−10 days.63 The compounds were assessed for their ability to affect mechanical allodynia at 10−14 days after the first injection of paclitaxel. Statistical Analyses. All values represent the mean ± SEM. Data sets were was all tested with a D’agostino−Pearson test for normality (Graphpad Prism 7 Software). Following the result of the normality test, statistical significance was tested using the appropriate parametric or nonparametric Student’s t test or analysis of variance (ANOVA), after which we performed post hoc comparisons (Tukey). Von Frey behavioral data sets were analyzed by two-way ANOVA (Tukey posthoc test). Statistical significance was inferred for all p ≤ 0.05. GraphPad Prism 7 was used for all graphs. No data points were excluded in our studies.

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Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We are thankful to Dr. T. R. Cummins (IUPUI School of Science) for providing the sodium channel expressing cell lines.

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ABBREVIATIONS Ach, acetylcholine; AITC, allyl isothiocyanate; AUC, area under the curve; CaV2.2, N-type voltage-gated calcium channel; CDC, Centers for Disease Control and Prevention; CHO, Chinese Hamster Ovary; DOR, delta opioid receptor; DRG, dorsal root ganglion; gp-120, HIV-1 envelope glycoprotein 120 kDas; (−)-HDA, Hardwickiic acid; HEK 293, human embryonic kidney 293; HIV, Human Immunodeficiency Virus; HTA, Hautriwaic acid; KCl, potassium chloride; KOR, kappa opioid receptor; MOR, mu opioid receptor; NaV1.x, voltage-gated Na+ channel isoform 1.x; sEPSCs, spontaneous excitatory postsynaptic currents; TTX, tetrodotoxin; TTX-R, tetrodotoxin-resistant; TTX-S, tetrodotoxinsensitive

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REFERENCES

(1) Institute of Medicine Report from the Committee on Advancing Pain Research, Care, and Education (2011) Relieving Pain in America A Blueprint for Transforming Prevention, Care, Education and Research, The National Academies Press. (2) Crawford, P., Ghadiali, E., Lane, R., Blumhardt, L., and Chadwick, D. (1987) Gabapentin as an antiepileptic drug in man. J. Neurol., Neurosurg. Psychiatry 50 (6), 682−6. (3) Barsa, J. A., and Saunders, J. C. (1961) Amitriptyline (Elavil), a new antidepressant. Am. J. Psychiatry 117, 739−40. (4) Hassett, A. L., Aquino, J. K., and Ilgen, M. A. (2014) The risk of suicide mortality in chronic pain patients. Curr. Pain Headache Rep. 18 (8), 436. (5) Dowell, D., Haegerich, T. M., and Chou, R. (2016) CDC Guideline for Prescribing Opioids for Chronic Pain–United States, 2016. JAMA: the journal of the American Medical Association 315 (15), 1624−45. (6) Woolf, C. J., and Hashmi, M. (2004) Use and abuse of opioid analgesics: potential methods to prevent and deter non-medical consumption of prescription opioids. Curr. Opin. Investig. Drugs 5 (1), 61−66. (7) Benyamin, R., Trescot, A. M., Datta, S., Buenaventura, R., Adlaka, R., Sehgal, N., Glaser, S. E., and Vallejo, R. (2008) Opioid complications and side effects. Pain Physician 11 (2 Suppl), S105−20. (8) Klockgether-Radke, A. P. (2002) [F. W. Serturner and the discovery of morphine. 200 years of pain therapy with opioids]. Anasthesiol Intensivmed Notfallmed Schmerzther 37 (5), 244−9. (9) Safavi-Hemami, H., Brogan, S. E., and Olivera, B. M. (2019) Pain therapeutics from cone snail venoms: From Ziconotide to novel non-opioid pathways. J. Proteomics 190, 12. (10) McIntosh, J. M., Corpuz, G. O., Layer, R. T., Garrett, J. E., Wagstaff, J. D., Bulaj, G., Vyazovkina, A., Yoshikami, D., Cruz, L. J., and Olivera, B. M. (2000) Isolation and characterization of a novel conus peptide with apparent antinociceptive activity. J. Biol. Chem. 275 (42), 32391−7. (11) Webster, L. R., Fakata, K. L., Charapata, S., Fisher, R., and MineHart, M. (2008) Open-label, multicenter study of combined intrathecal morphine and ziconotide: addition of morphine in patients receiving ziconotide for severe chronic pain. Pain Med. 9 (3), 282−90. (12) Bellampalli, S. S., Ji, Y., Moutal, A., Cai, S., Wijeratne, E. M. K., Gandini, M. A., Yu, J., Chefdeville, A., Dorame, A., Chew, L. A., Madura, C. L., Luo, S., Molnar, G., Khanna, M., Streicher, J. M., Zamponi, G. W., Gunatilaka, A. A. L., and Khanna, R. (2018) Betulinic acid, derived from the desert lavender Hyptis emoryi,

AUTHOR INFORMATION

Corresponding Author

*Mailing address; Department of Pharmacology, College of Medicine, University of Arizona, 1501 North Campbell Drive, P.O. Box 245050, Tucson, AZ 85724, USA. Office phone: 520626-4281. Fax: 520-626-2204. E-mail: [email protected]. edu. ORCID

May Khanna: 0000-0002-9989-6374 John M. Streicher: 0000-0002-4173-7362 Rajesh Khanna: 0000-0002-9066-2969 Author Contributions

S.C. and S.S.B. are co-first authors. S.C., J.Y., W.L., and Z.S. conducted whole-cell electrophysiology. E.M.K.W. synthesized the compounds. S.S.B., Y.J., A.D., A.M. performed calcium imaging experiments. S.L. performed the behavioral experiments. J.M.S. supervised the opioid receptor binding experiments. A.A.L.G. and R.K. conceived the study. A.A.L.G., M.K., and R.K. designed/supervised the overall project and wrote the manuscript. Funding

This work is supported by a National Institutes of Health Award (1R01NS098772, 1R01DA042852, and 1R01NS098772) to R.K. A.M. was supported by a Young Investigator’s Award from the Children’s Tumor Foundation. K

DOI: 10.1021/acschemneuro.8b00617 ACS Chem. Neurosci. XXXX, XXX, XXX−XXX

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