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Research Paper

Pharmacological characterization of a rat Nav1.7 loss-of-function model with insensitivity to pain

Chen, Lubina,b,c; Effraim, Philip R.b,c,d; Carrara, Jennifera,b,c; Zhao, Penga,b,c; Dib-Hajj, Fadia B.a,b,c; Dib-Hajj, Sulayman D.a,b,c; Waxman, Stephen G.a,b,c,*

Author Information
doi: 10.1097/j.pain.0000000000001807

1. Introduction

Congenital insensitivity to pain (CIP) is an extremely rare genetic disorder characterized by the complete absence of nociception. In most cases, painlessness is inherited in an autosomal recessive manner, suggesting disruptive mutations of a single gene can lead to such phenotype.43 So far, genes associated with CIP include those controlling function of nociceptors, such as SCN9A,13 and those affecting nociceptor neurodevelopment, such as NTRK1,30NGF,20PRDM12,12 and CLTC1.42 Among all the identified causative genes for CIP, SCN9A, which encodes the voltage-gated sodium channel Nav1.7, is validated as an important participant in controlling nociception.6,19 Since early reports of truncating SCN9A mutations,2,13,24 more than 30 nonsense and mis-sense SCN9A mutations have been identified in individuals with CIP.14,21,55

Nav1.7 is preferentially expressed along the first-order neurons within the nociceptive pathway, extending from intraepidermal nerve fibers to presynaptic central terminals.7 Slow closed-state inactivation kinetics of Nav1.7, together with relatively hyperpolarized activation, render it a threshold channel, amplifying subthreshold depolarizations to trigger action potential firing in nociceptors.16,47 In addition, presynaptic Nav1.7 channels contribute to control of neurotransmitter release at terminals of both dorsal root ganglion (DRG) neurons4,38 and olfactory sensory neurons.56 These actions of Nav1.7 imply that SCN9A-related CIP may result from reduction of excitability and impairment of neurotransmitter release.

It has been proposed that loss of pain sensibility in Nav1.7-mediated CIP may be related to upregulation of endogenous opioid preproenkephalin (PENK) and can be substantially reversed by the opioid antagonist naloxone.39 Recently, study of a female CIP patient with loss of function of fatty acid amide hydrolase revealed that pain insensitivity can be caused by elevated circulating endocannabinoids.26 Although this case is not Nav1.7-related, it suggests that changes in modulatory systems can induce a painless phenotype as strong as seen in Nav1.7-mediated CIP.

Global and conditional Nav1.7 KO mouse models have provided valuable insights for understanding Nav1.7 function in nociception.23,28,38,44,50 Importantly, global Nav1.7 KO mice, with special husbandry strategies, can survive to adulthood and fully recapitulate the clinical presentation of Nav1.7-mediated CIP, including anosmia and complete loss of pain sensation.23 Adult-onset deletion of Nav1.7, which avoids any potential developmental deficits in genetic KO models, leads to profound insensitivity to pain, further confirming the pivotal role of Nav1.7 in nociception.50 Grubinska et al.25 have recently described a rat Nav1.7 knock-in model expressing humanized Nav1.7 protein. Owing to unexpected tissue-specific splicing, the full-length chimeric Nav1.7 protein is not detectable in DRG neurons, although it is expressed in olfactory neurons, essentially generating a rat Nav1.7 loss-of-function model.25

In this study, using the newly established Nav1.7 loss-of-function rat model, we further characterize the behavioural response of Nav1.7 loss-of-function rats to multiple modalities of noxious stimuli. Using pharmacological approaches and multielectrode array (MEA) recordings, we evaluate the contribution of endogenous opioids and cannabinoids as well as changes in DRG neuronal excitability to the observed painless phenotype.

2. Materials and methods

2.1. Animals

WT and Nav1.7 loss-of-function rats (termed “HOM-KI” rats by Grubinska et al. 2019) were group-housed at an Association for Assessment and Accreditation of Laboratory Animal Committee–accredited facility in nonsterile solid bottom polycarbonate cages with corn cob bedding; bedding in all cages was removed and replenished at least once per week. Animals had ad libitum access to pelleted feed (Teklad 2018; Envigo, Madison, WI) and water. Animals were maintained on a 12:12-hour light:dark cycle (6 am:6 pm) at temperature 21 ± 3°C, humidity 50 ± 20%, and had access to enriched environment. All animal procedures were conducted in accordance with the NIH Guide for the Care and Use of Laboratory Animals and were approved by the IACUC of the Veterans Administration Connecticut Healthcare System. Adult rats (age range strictly 7.5-8.5 weeks) of both sexes (female weight range 195 to 235 g, male weight range 265 to 370 g) were used in the behavioural tests. Juvenile rats (age range 5-7 weeks) of both sexes were used in the MEA recordings.

2.2. Drugs

Naloxone hydrochloride dihydrate (N7758, Sigma, St Louis, MO) was dissolved in saline to 2-mg/mL or 8-mg/mL solution and was injected intraperitoneally (i.p.) in volume of 1 mL/kg. Morphine sulphate (10-mg/mL in 1-mL single-dose vials, WEST-WARD Pharmaceuticals, Eatontown, NJ) was injected i.p. in volume of 1 mL/kg. SR141716A (0923, Torcris, Bristol, United Kingdom; or Rimonabant hydrochloride, SML 0800, Sigma) and SR144528 (5039, Torcris) were dissolved in ethanol:tween80:saline (1:1:18) solution to 3.3 mg/mL and were injected i.p. in volume of 3 mL/kg. All drug solutions were made fresh on the same day of experiment.

Naloxone (2 mg/kg or 8 mg/kg), SR141716A (10 mg/kg), or SR144528 (10 mg/kg) was administered 30 minutes before the hot plate test (20 minutes before the formalin test). Morphine (10 mg/kg) was administered 15 minutes before naloxone injection or 45 minutes before the hot plate test (35 minutes before the formalin test). Twenty-five–gauge needles (1-mL Vanish Point Syringe, Retractable Technology, Little Elm, TX) were used for all i.p. injections. Twenty-six–gauge needles (1-mL BD syringe) were used for intraplantar injections.

2.3. Behavioural assays

All behavioral assays were performed and scored by trained experimenters blinded to the genotype or drug treatment. To minimize visual cues for identifying Nav1.7 loss-of-function rats, animals with severe wounds were excluded from behavioral assays.

2.3.1. Hot/cold plate

Animals were placed on a metal plate (Hot Plate Analgesia Test Meter, IITC, Woodland Hills, CA) uniformly heated to a constant temperature of 52°C (or 0.5°C for cold plate). The response latency for first licking or flinching of the hind paws was recorded. The rats were immediately removed from the hot plate upon showing nocifensive behaviors or if no response occurred within the 30-second cutoff time. To avoid any potential nerve damage caused by repetitive noxious heat stimuli, animals were tested no more than twice (once for baseline and then after drug treatment) in the hot plate test.

2.3.2. Hargreaves test

Animals were acclimated on a glass plate (temperature maintained at 30°C) for at least 30 minutes, and plantar paw surface was exposed to radiant heat using a Hargreaves apparatus (IITC, Woodland Hills, CA), and the paw withdrawal latency was measured. The heat stimulation was repeated 3 times on both paws, with at least 5-minute intervals between consecutive tests. The cutoff value was set at 30 seconds to prevent tissue damage.

2.3.3. Dry ice test

Animals are acclimated on a 1/5″ thick Borofloat glass plate (S.I. Howard Glass Company, Worcester, MA) for at least 30 minutes. A dry ice pellet was made by compressing fine dry ice powder into a 3-mL syringe (with top cutoff) and was then firmly pressed against the glass surface underneath the rat hind paw until a withdrawal response resulted.10 The cold stimulation was repeated 3 times on both paws at least 15-minute interval between consecutive tests. The cutoff value was set at 60 seconds to prevent tissue damage.

2.3.4. Thermal preference test and gradient test

The thermal preference apparatus consisted of 2 heating/cooling plates (Bioseb, Chaville, France) placed side-by-side and enclosed in a plexiglass chamber. The reference plate was set to 25°C, and the test plate was set to 15°C or 33°C. Animal movements were recorded for 600 seconds by an automated infrared camera tracking system. The time spent on the reference and test plates was calculated. The thermal gradient apparatus maintained a stable temperature gradient from about 6°C to 58°C using the 2 heating/cooling devices positioned at each end of a metal floor. The floor area was divided into 10 zones, each with a stable temperature measured using a thermometer (TM-3 three-scale temperature monitor, Warner Instruments, Hamden, CT). Animals were placed onto the gradient plate on the cold end. Tracking was performed using software provided by the manufacturer and the time spent in each zone over the 60-minute test period determined.

2.3.5. von Frey test

Animals were placed on an elevated wire grid, and the plantar surface of each paw was presented with a series of calibrated von Frey filaments (Stoeling, Wood Dale, IL). Each filament was applied for 5 seconds; and a response was recorded when flinching, withdrawal, paw licking, or toe spreading was observed, with at least 5-minute interval between stimuli. The 50% withdrawal threshold was determined using the “up-down” method.

2.3.6. Pin-prick test

Animals were placed on an elevated wire grid, and the plantar surface of each paw was presented with an insect pin (tip diameter 0.03 mm, 26001-70, size 7, Fine Science Tools, Foster City, CA) attached to a 10-g von Frey filament. The pin was applied for up to 5 seconds, and a response was recorded when paw withdrawal was observed. The ability to detect mechanical pain was characterized as the number of paw withdrawals in a total of 10 stimulations with at least 5-minute interval between each stimulus.

2.3.7. Grimace scale

The facial expression of rats was scored for their prominence in still photographs taken from 10-minute videos (rats were recorded for 30 minutes after drug treatment, but the videos of the last 10 minutes were used) of WT or Nav1.7 loss-of-function rats after SR141716A or vehicle treatment. For each rat, 5 photographs were randomly selected. Four action units, including orbital tightening, nose/cheek flattening, ear changes, and whisker changes, were used in the assessment. Each action unit was scored on a 3-point (0 = not present, 1 = moderate, and 2 = severe), and the average score of all action units was analyzed.51

2.3.8. Formalin test

Rats were habituated to observation chambers for at least 30 minutes before experiments. Approximately 50 μL of 2.5% formalin was injected subcutaneously (s.c.) into the plantar surface of the left hind paw. After injection, rats were immediately returned to the observation chamber and nociceptive behaviours were recorded for 60 minutes. The duration of formalin-induced flinching and licking in 5-minute intervals was scored by an experimenter blinded to treatment or genotype.

2.4. Multielectrode array recordings

2.4.1. Primary sensory neuron isolation for multielectrode array

12-well MEA plates (Axion Biosystems, Atlanta, GA) were coated with poly-D-lysine (50 µg/mL) and laminin (10 µg/mL). Lumbar DRGs from 5- to 7-week-old WT or Nav1.7 loss-of-function rats were harvested, and neurons were dissociated as previously described.18 Briefly, lumbar DRGs were incubated for 20 minutes at 37°C in complete saline solution (in mM: 137 NaCl, 5.3 KCl, 1 MgCl2, 25 sorbitol, 3 CaCl2, and 10 N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid [Hepes], adjusted to pH 7.2 with NaOH) containing 1.5-mg/mL Collagenase A (Sigma) and 0.6-mM EDTA, followed by a 17-min incubation at 37°C in complete saline solution containing 1.5-mg/mL Collagenase D (Sigma), 0.6-mM EDTA, and 30-U/mL papain. Dorsal root ganglia were then centrifuged and triturated in 0.6 mL of DRG media (Dulbecco's modified Eagle's medium-F12 [1:1] with 100 U/mL penicillin, 0.1-mg/mL streptomycin [Invitrogen, Rockford, IL], and 10% fetal bovine serum [Hyclone] containing 1.5-mg/mL BSA [low endotoxin] and 1.5-mg/mL trypsin inhibitor [Sigma]). After trituration, 2 mL of DRG media was added to the cell suspension, which was filtered with 70-μm cell strainer (Falcon, 352350, Corning, NY). The mesh was washed twice with 2 mL of DRG media. The cell suspension was centrifuged (100 g for 3 minutes), and the cell pellet was resuspended in DRG media and seeded 60 µL per well. After neurons were settled in 95% air/5% CO2 (vol/vol) incubator at 37°C for 50 minutes, each well was topped up with 1.44 mL of DRG media containing nerve growth factor (50 ng/mL) and glial cell line–derived neurotrophic factor (50 ng/mL). Dissociated DRG neurons were maintained at 37°C in a 95% air/5% CO2 (vol/vol) incubator for 3 days before MEA recording.

2.4.2. Multielectrode array recording

Action potential firing was measured using a multiwell MEA system (Maestro, Axion Biosystems). Four 12-well MEA recording plates each with 768 electrodes (64 electrodes in each well) were used. In each experiment, neurons from one WT and one Nav1.7 loss-of-function rats (age- and sex-matched littermates) were seeded in a 12-well MEA recording plate (6 wells for each animal). The investigator was blinded to the identity of neurons. Precise temperature control of the MEA system was used to create temperature ramps and maintain temperatures at 30°, 33°, 37°, 40°, and 43°C each for 200 seconds. For experiments with capsaicin (1 µM), steady-state neuronal firing over 90-second periods before or after introduction of capsaicin was measured. A spike detection criterion of >6 SDs above background signals was used to separate action potential spikes from noise. Electrodes registering >5 recorded spikes over 200-second period were determined as active. Number of active electrodes and mean firing frequency were measured. Data were analysed using Axion Integrated Studio AxIS 2.1 (Axion Biosystems) and Neuro Explorer (NexTechnologies, Madison, AL).

2.5. Data and statistical analysis

Unless otherwise noted, statistical significance was determined using the Mann–Whitney U test. Data are expressed as mean ± SEM. A P value of < 0.05 was considered significant.

3. Results

3.1. Nav1.7 loss-of-function rat model

As reported previously,25 we observed that Nav1.7 loss-of-function rats did not respond to noxious heat (52°C) or cold (0.5°C) in hot/cold plate tests, reaching cutoff time for every rat (Figs. 1A and B). Interestingly, in contrast to the complete absence of responses in the hot plate test, some Nav1.7 loss-of-function rats, particularly about 50% of females, still displayed withdrawal behavior in response to radiant heat in the Hargreaves test, although the withdrawal latency was significantly elevated (Fig. 1C). A similar phenomenon was seen in the dry ice test, where Nav1.7 loss-of-function rats still responded to cooling despite significantly increased latency (Fig. 1D). These observations suggested that the Nav1.7 loss-of-function rats might retain some ability to sense a change in temperature. As might be expected from this reasoning, in the thermal preference test, Nav1.7 loss-of-function rats could differentiate temperature differences of 8° to 10°C (33° vs 25°C, or 15° vs 25°C) and displayed warmth seeking and cold avoiding behavior similar to WT (Fig. 1E). Nav1.7 loss-of-function rats displayed significantly elevated mechanical threshold measured by the von Frey test, although about half of the Nav1.7 loss-of-function rats tested had mechanical threshold within the normal range observed for WT (Fig. 1F). In the pin-prick test, WT rats responded to each stimulus (10/10) as soon as the 10-g von Frey fiber with pin was applied to the plantar skin, while Nav1.7 loss-of-function rats displayed minimal response (Fig. 1G).

Figure 1.
Figure 1.:
Nav1.7 loss-of-function rats show insensitivity to acute noxious stimuli and exhibit increased spontaneous scratching. (A) Hot plate test, 52°C. Nav1.7 loss-of-function (LOF) rats lose heat nociception (WT, 7.9 ± 0.6 seconds, n = 21; Nav1.7 LOF, 30 seconds, n = 30). (B) Cold plate test, 0.5°C. Nav1.7 LOF rats lose cold nociception (WT, 94.5 ± 9.7 seconds, n = 6; Nav1.7 LOF, 120 seconds, n = 6). (C) Hargreaves test. Nav1.7 LOF rats display elevated withdrawal latency (male: WT, 14.4 ± 0.7 seconds, n = 6; Nav1.7 LOF, 29.9 ± 0.1 second, n = 9; female: WT, 13.5 ± 0.8 seconds, n = 7; Nav1.7 LOF, 27.0 ± 1.1 second, n = 9). (D) Dry ice test. Latency to paw withdrawal is increased in Nav1.7 LOF rats (WT, 12.8 ± 1.1 second, n = 6; Nav1.7 LOF, 26.4 ± 3.5 seconds, n = 6). (E) Thermal preference test. No difference is seen between WT and Nav1.7 LOF rats (WT, n = 7; Nav1.7 LOF, n = 7). (F) von Frey test. Average paw withdrawal threshold is increased in Nav1.7 LOF rats (WT, 7.5 ± 0.9 g, n = 12; Nav1.7 LOF, 12.6 ± 1.5 g, n = 15). (G) Pin-prick test. Nav1.7 LOF rats show minimal response to pin-prick (WT, n = 6; Nav1.7 LOF, n = 6). (H) Representative images of lesions seen in Nav1.7 LOF rats. Left, a severe case in which a large painless skin lesion can be seen at the dorsal head region. Right, a mild case in which lesions can be seen at cheek, head, and shoulder regions. (I) Nav1.7 LOF rats display significantly increased spontaneous hind paw scratching behaviour (WT, 6.5 ± 2.5/30 minutes, n = 5; Nav1.7 LOF, 78.4 ± 38.6/30 minutes, n = 8). For all, **P <0.01, ***P < 0.001, ****P < 0.0001, WT vs Nav1.7 LOF, Mann–Whitney U test.

We noticed that self-inflicted lesions started to appear as early as 3 weeks in Nav1.7 loss-of-function rats and were often associated with excessive scratching. The presentation of the tissue lesions was variable among individuals, with some rats showing severe skin lesions and others barely noticeable. The lesions appeared to be confined to the facial, head, neck, and shoulder regions (Fig. 1H). Nav1.7 loss-of-function rats, with or without skin lesions, showed significantly increased scratching behaviours (Fig. 1I). It was unclear whether there was a causative link between severity of wounds and spontaneous scratching behaviour.

3.2. Naloxone does not restore acute pain sensitivity in Nav1.7 loss-of-function rats

It has been proposed that endogenous opioids contribute to insensitivity to pain in humans and mice lacking Nav1.739 and it has been suggested that µ- and δ-opioid receptors are involved.45 We therefore examined whether the opioid antagonist naloxone could restore acute pain sensation in this Nav1.7 loss-of-function model. We observed that naloxone (2 mg/kg, i.p.) administration in Nav1.7 loss-of-function rats did not restore heat pain in the 52°C hot plate test (Fig. 2A) or affect nocifensive behaviours in the formalin test (Figs. 2B and C). At a dose of 2 mg/kg, naloxone successfully reversed analgesia caused by morphine (10 mg/kg, i.p.) in wild-type controls (Fig. 2A; and supplementary Fig. 1, available at A higher dose (8 mg/kg) of naloxone was used to ensure blockade of opioid receptors but still could not restore acute thermal pain sensation in Nav1.7 loss-of-function rats (Fig. 2A).

Figure 2.
Figure 2.:
Effects of naloxone on acute pain sensitivity in Nav1.7 loss-of-function rats. (A) Hot plate test, 52°C. Paw withdrawal latency in wild-type rats is not affected by naloxone (2 mg/kg, i.p.) (WT baseline, 8.3 ± 0.8 seconds; WT naloxone, 8.8 ± 0.8 seconds; n = 9) but significantly increased after morphine administration (10 mg/kg, i.p.) (WT morphine, 21.1 ± 2.9 seconds, n = 9). Naloxone can reverse the analgesia caused by morphine (WT morphine + naloxone, 9.5 ± 0.8 seconds, n = 12). Neither 2-mg/kg nor 8-mg/kg naloxone induces paw withdrawal response in Nav1.7 LOF rats. (B and C) Formalin test. Naloxone treatment in Nav1.7 LOF rats does not affect licking and flinching durations in either phase I (saline, 79.7 ± 28.8 seconds, n = 3; naloxone, 96.0 ± 20.5 seconds, n = 4) or phase II (saline, 220.3 ± 157.6 seconds, n = 3; naloxone, 223.5 ± 90.4 seconds, n = 4) of the formalin test. **P <0.01, *** P < 0.001, Mann–Whitney U test.

3.3. Cannabinoid receptor type 1 (CB1) blocker SR141716A (rimonabant) does not restore acute pain sensitivity but induces pain-like behaviors in Nav1.7 loss-of-function rats

A recent case report linked loss of pain sensibility in 1 CIP patient (not Nav1.7-related) to elevated circulation concentrations of endocannabinoids.26 In this study, we tested whether endogenous cannabinoid system contributes to the lack of pain sensitivity in the Nav1.7 loss-of-function model, using CB1 blocker SR141716A (rimonabant) and CB2 blocker SR144528. SR141716A (10 mg/kg, i.p.) produced scratching in both wild-type and Nav1.7 loss-of-function rats minutes after administration. Compared with wild-type controls, CB1 blocker induced significantly more profound scratching behaviors in Nav1.7 loss-of-function rats (Fig. 3A). In addition to increased number of scratching events and intensified scratching movements, Nav1.7 loss-of-function rats showed enhanced wet dog shakes and caudal biting behaviours (supplementary Video, available at Nav1.7 loss-of-function rats but not wild-type controls displayed abnormal facial expressions (which could be interpreted as pain-like) after treatment with SR141716A (Fig. 3B). Grimace scale (GS) score in Nav1.7 loss-of-function rats was significantly increased after SR141716A injection (Fig. 3C). However, when tested in acute pain assays, Nav1.7 loss-of-function rats retained their insensitivity to pain. In the hot plate test, Nav1.7 loss-of-function rats had no response to noxious heat (52°C) 30 minutes after SR141716A administration (Fig. 3D). Although Nav1.7 loss-of-function rats appeared to display increased formalin-induced behaviours after administration of SR141716A in phase II (10-60 minutes), the difference between control and SR141716A-treated groups was not statistically significant (Figs. 3E and F). CB2 blocker SR144528 did not restore acute thermal pain sensitivity in Nav1.7 loss-of-function rats (supplementary Fig. 2A, available at or cause any scratching or abnormal facial expression. SR141716A did not have a statistically significant effect on pain sensitivity in wild-type controls in hot plate (Fig. 3D) or formalin tests (supplementary Fig. 2B and C, available at

Figure 3.
Figure 3.:
Effects of SR141716A (rimonabant) in Nav1.7 loss-of-function rats. (A) Hind paw scratching events after SR141716A treatment (10 mg/kg, i.p.). SR141716A induces profound scratching behaviour in both WT (vehicle, 4.4 ± 1.4/10 minutes, n = 5; SR141716A, 103.0 ± 22.4/10 minutes, n = 7) and Nav1.7 LOF (vehicle, 34.3 ± 19.9/10 minutes, n = 5; SR141716A, 201.0 ± 32.4/10 minutes, n = 8) rats. (B) Representative images of facial expressions in WT (upper) and Nav1.7 LOF (lower) rats 30 minutes after SR141716A treatment. Nav1.7 LOF rats display pain-like facial expression. (C) Grimace test. Mean grimace scale score is significantly increased in Nav1.7 LOF (vehicle, 0.20 ± 0.05, n = 6; SR141716A, 0.91 ± 0.11, n = 7) but not WT (vehicle, 0.13 ± 0.07, n = 3; SR141716A, 0.15 ± 0.05, n = 4) rats. (D) Hot plate test, 52°C. Paw withdrawal latency in wild-type rats is not affected by SR141716A (10 mg/kg, i.p.) (WT baseline, 8.7 ± 0.8 seconds; WT SR141716A, 9.7 ± 1.1 second; n = 9). SR141716A does not induce paw withdrawal response in Nav1.7 LOF rats. (E and F) Formalin test. SR141716A treatment in Nav1.7 LOF rats does not affect licking and flinching durations in either phase I (vehicle, 88.2 ± 9.9 seconds, n = 9; SR141716A, 85.0 ± 15.0 seconds, n = 10) or phase II (vehicle, 112.3 ± 54.0 seconds, n = 9; SR141716A, 187.6 ± 66.3 seconds, n = 10) of the formalin test. *P <0.05, ** P < 0.01, Mann–Whitney U test.

3.4. Excitability of dorsal root ganglion neurons from Nav1.7 loss-of-function rats in response to heat or capsaicin is significantly reduced

Previous whole-cell patch clamp recordings demonstrated that small DRG neurons from these Nav1.7 loss-of-function rats show a ∼40% reduction in tetrodotoxin-sensitive (TTX-S) Na+ currents and a ∼50% increase in current threshold for action potential firing.25 Using MEA recordings, we further examined the impact of loss of function of Nav1.7 on excitability of DRG neurons from Nav1.7 loss-of-function rats (2 male and 2 female) in response to noxious heat and the TRPV1 ligand capsaicin. Dorsal root ganglion neurons from male and female rats did not display any statistically significant differences in excitability, and we grouped them together for analysis. When DRG neurons from WT and Nav1.7 loss-of-function rats were cultured on MEA plates, the number of cells counted morphologically was the same between groups. Representative heat map plots showed temperature-dependent increases in the number of active electrode (colored circles) and the number of action potentials (color scale) fired by DRG neurons from Nav1.7 loss-of-function as well as WT rats, when temperature was raised from 30°C to 43°C (Fig. 4A). The increase was more profound in neurons from WT rats (Fig. 4A). Compared with wild-type controls, Nav1.7 loss-of-function group had significantly fewer active electrodes per well and decreased firing frequency (Figs. 4B and C). In addition, we observed that neurons from WT rats show bursts (>5 spikes) and continuous repetitive firing in response to elevated temperature, while neurons from Nav1.7 loss-of-function rats rarely display these firing patterns (Fig. 4D). When temperature was raised from 30°C to 43°C, there was a nearly 7-fold increase in the number of bursts per well in the WT group. Compared with the WT group, Nav1.7 loss-of-function group showed significantly less bursts (Fig. 4E). Nav1.7 loss-of-function neurons also produced significantly fewer spikes per burst (Fig. 4F).

Figure 4.
Figure 4.:
Dorsal root ganglion (DRG) neurons from Nav1.7 loss-of-function rats show reduced response to heat. (A) Representative heat-map plots of MEA recordings of DRG neurons from WT and Nav1.7 LOF rats at 30°C, 33°C, 37°C, 40°C, and 43°C. Each colored circle represents an active electrode within an 8 × 8 electrode array. More active electrodes and higher firing frequency are recorded at higher temperatures. There are fewer active electrodes in wells containing Nav1.7 LOF cells, and neurons from Nav1.7 LOF rats fire at a significantly lower frequency. (B) Number of active electrodes in WT and Nav1.7 LOF groups at different temperatures (30°C, 33°C, 37°C, 40°C, and 45°C). (C) Weighted firing frequencies of neurons from WT and Nav1.7 LOF rats. (D) Representative traces of action potential firing from a single electrode recording DRG neurons from WT and Nav1.7 LOF rats at 43°C. Both short bursts and continuous repetitive firing are common in WT neurons at 43°C. Neurons from Nav1.7 LOF rats show significantly less firing. (E) Number of bursts in WT and Nav1.7 LOF neurons. (F) Number of spikes per burst in WT and Nav1.7 LOF neurons. For all, *P <0.05, **P <0.01, ***P < 0.001, ****P < 0.0001, WT vs Nav1.7 LOF, Mann–Whitney U test.

We next examined the changes in neuronal excitability in response to capsaicin (1 μM) at 37°C. After capsaicin treatment, both WT and Nav1.7 loss-of-function groups showed a strong response (Fig. 5A). Quantitative analysis revealed that the numbers of active and bursting electrodes in the Nav1.7 loss-of-function group were significantly reduced compared with the WT group, with a ∼30% reduction in the number of active electrodes and a ∼40% reduction in the number of bursting electrodes (Figs. 5B and C). The active electrodes in the Nav1.7 loss-of-function group also recorded a ∼50% reduction in firing frequency compared with the WT group (Fig. 5D).

Figure 5.
Figure 5.:
Dorsal root ganglion (DRG) neurons from Nav1.7 loss-of-function rats show reduced response to capsaicin. (A) Whole well raster plots of DRG neurons from WT and Nav1.7 LOF rats after 1-μM capsaicin, added at the time point indicated by the black arrow heads. Each row in the raster plots represents a single electrode with 64 electrodes per well, and all recordings were performed at 37°C. (B–D) Number of active electrodes (B), number of bursting electrodes (C), and average firing frequency (D) in WT and Nav1.7 LOF groups before and after capsaicin treatment. After capsaicin treatment, the Nav1.7 LOF group has significantly fewer active electrodes, fewer bursting electrodes, and reduced average firing frequency compared with the WT group. For all, *P <0.05, *** P < 0.001, WT vs Nav1.7 LOF, Mann–Whitney U test.

4. Discussion

The recently established rat loss-of-function model provides a unique tool for investigation of the molecular mechanisms underlying Nav1.7-mediated CIP.25 Using this tool, we examined the contribution of endogenous opioids and cannabinoids to the painlessness seen in the Nav1.7 loss-of-function rats. We found that blocking the endogenous opioid system or the endogenous cannabinoid system does not restore acute pain sensitivity in the Nav1.7 loss-of-function rats. However, we observed that CB1 blockade results in enhanced scratching behaviours and altered facial expression, although it is unclear whether these changes are pain-related. We demonstrated that DRG neurons from Nav1.7 loss-of-function rats display overall reduced excitability but are still able to fire action potentials in response to noxious heat and capsaicin. Taken together, these observations suggest that neither activity of the endogenous opioid and cannabinoid systems nor total loss of excitability in DRG neurons is required for loss of pain sensation in this Nav1.7 loss-of-function rodent model.

We observed that the Nav1.7 loss-of-function rats lose pain sensation in response to a wide range of noxious stimuli, including noxious heat, cold, and pinprick. The abilities to differentiate non-noxious temperature changes and to detect tactile input are retained. In this regard, this Nav1.7 loss-of-function rat model recapitulates the somatosensory symptoms in human CIP individuals, who have complete loss of pain, hyposensitivity to non-noxious thermal input but normal detection of touch and vibration.13,24,37

It has been proposed that the endogenous opioid system contributes to the Nav1.7-related CIP phenotype.31,39,45 Loss of functional Nav1.7 is shown to increase PENK mRNA expression39 and to potentiate opioid receptors,31 which are likely μ- and δ-opioid receptors.45 The hypothesis of Nav1.7-mediated PENK mRNA increase, however, has been challenged by a recent study showing no upregulation of PENK mRNA in Nav1.7 KO human iPSCs and very little basal expression in healthy control neurons,37 which is consistent with single-cell RNA sequencing profiling of mouse DRG neurons showing barely detectable baseline PENK expression.53 In the same study, in vitro treatment with naloxone failed to normalize hypoexcitability of Nav1.7 KO iPSCs.37 In this study, we used the Nav1.7 loss-of-function rat to examine the involvement of endogenous opioids in vivo and found that systemic naloxone administration does not restore the ability to perceive pain, suggesting that endogenous opioid system is not required for the complete analgesia seen in these rats. Our observations contrast with previous findings in the human Nav1.7-related CIP individual and Nav1.7 KO mice models.31,39,45 The discrepancy could be due to species differences. Alternatively, although structural deficit is unlikely in the peripheral terminals in this Nav1.7 loss-of-function rat model,25 we cannot rule out the possibility that there is a structural or functional abnormality in central terminals of the DRG or changed connectivity in the spinal cord and higher brain centers. Such changes might overshadow the effect of pharmacological blockade of endogenous opioids.

Our observations on pharmacological blockade of endogenous cannabinoids depict a complex association between loss of Nav1.7 and function of the endogenous cannabinoid systems. Loss-of-function of Nav1.7 in the rat model induces spontaneous itch-like scratching and potentiates SR141716A (rimonabant, a CB1 antagonist)-induced scratching. The SR141716A-induced scratching has been reported in several previous studies,17,32,48,49 indicating a potential role for endocannabinoid system in tonic modulation of the itch pathway. Pruritus has not been noted as a symptom in previously described human subjects with Nav1.7-related CIP.13,24,37 On the contrary, perception of itch has been reported to be lost in these individuals.37 Global Nav1.7 KO mice and sensory-specific adult-onset Nav1.7 KO mice both show loss of scratching behaviours in response to histamine.23,50 Blocking Nav1.7 with a monoclonal antibody effectively suppresses acute and chronic itch.35 However, self-inflicted scratching is a common phenotype seen in Nav1.7 KO mouse models and this Nav1.7 loss-of-function rat model.23,25,50 Consistent with these observations, rats treated with potent Nav1.7 blockers JNJ63955918 and ProTX-II show skin lesions associated with excessive scratches.22 Taken together, the evidence suggests that the lack of behavioural responses to pruritogens in both mouse and human may be due to the impaired excitability of itch-sensing neurons, the majority of which express Nav1.7,53 while the spontaneous scratching or enhanced SR141716A-induced scratching may possibly result from changes within the central nervous system. Emerging data indicate the existence of specific pruriceptors and distinctive chemical and mechanical itch-processing circuitries,9,27,40,52 and there is evidence that pain suppresses itch through an inhibitory circuitry in the spinal cord.3,36 Our observation suggests that the removal of nociceptive input may augment itch input. It remains unclear, however, why spontaneous itch-like symptom is seen in rodents but not in humans.

A recent report suggested that loss of function of fatty acid amide hydrolase can result in elevated anandamide concentrations and pain insensitivity in humans.26 In this study, we examined the hypothesis that the endogenous cannabinoid system may be involved in the painless phenotype in the Nav1.7 loss-of-function rat model. Interestingly, we observed that blockade of CB1 but not CB2 leads to abnormal facial expression, excessive scratching, and caudal biting behaviours in Nav1.7 loss-of-function rats. The involvement of CB1 in the pain pathway has been debated. Pharmacological studies have reported mixed results on the effects of CB1 blocker SR141716A on responses to acute thermal pain, with some studies claiming no effect5,15 and others suggesting a hyperalgesic effect.11,46 Mixed results are also seen in global CB1 knockout mice models. One study suggested that CB1 KO mice have normal pain sensitivity in various nociceptive tests,34 another study showed normal thermal pain threshold but mechanical hyperalgesia,29 while a third study reported hypoalgesia.57 Nociceptor-specific knockout of CB1 receptor has been reported to lead to significant hyperalgesia.1 Our results suggest that the function of endogenous cannabinoid system may be altered in Nav1.7 loss-of-function rats through a CB1-mediated pathway because the phenotype after CB1 blockade, which includes changed facial expression, caudal biting, and enhanced scratching, is not seen in wild-type control littermates. However, details of the mechanism underlying the observed phenotype are not clear. Notably, SR141716A fails to restore acute pain sensitivity measured by withdrawal latencies or formalin-induced licking and flinching in Nav1.7 loss-of-function rats. We suggest that the observed phenotype could be due to exaggerated itch perception upon removal of pain input. Alternatively, there could be changes in CB1-mediated endogenous cannabinoid function in Nav1.7 loss-of-function rats. Further investigation in Nav1.7-related CIP individuals will be needed to test this possibility in humans.

Using MEA assays, we showed that although the excitability of DRG neurons is significantly impaired due to loss of Nav1.7, these neurons from Nav1.7 loss-of-function rats are still excitable in response to noxious heat or capsaicin. This is consistent with our previous current-clamp recordings showing that small-diameter DRG neurons from Nav1.7 loss-of-function rats, which include nociceptors, are capable of firing action potentials, although with elevated current threshold.25 The reduction of excitability in nociceptors lacking functional Nav1.7 channels is also supported by dynamic-clamp experiments demonstrating that action potential threshold increases, but only by about 35%, after electronic removal of all Nav1.7 current.54 Finally, DRG neuron hypoexcitability has been observed in other Nav1.7 loss-of-function models and in nociceptors derived from iPSCs from human subjects with Nav1.7-related CIP.23,25,37,50 There is an apparent discrepancy between the partial reduction of excitability and the complete loss of thermal pain or capsaicin response. One possibility is that temporal summation of peripheral nociceptive input at the spinal cord level is required for pain sensation, and partial reduction in sensory neuron excitability impairs such summation. Alternatively, there is evidence that the presence of Nav1.7 in the central terminals of DRG neurons is required for action potential invasion of their central terminals and for transmitter release within the spinal cord,4,38 and it is possible that the probability of secure transmission is reduced at sites of low safety factor such as branch-points of preterminal axons in the dorsal horn in Nav1.7 loss-of-function rats. Finally, there is evidence that Nav1.7 is selectively expressed in neurons within the rodent hypothalamus and some brainstem nuclei.8,25,33,41 These brain regions in the painless Nav1.7 loss-of-function rats are devoid of Nav1.7 expression.25 We cannot rule out the possibility that Nav1.7 function in the CNS may be involved in pain perception, with loss of Nav1.7 in these brain regions contributing to the painless phenotype.

In conclusion, we have evaluated the contribution of endogenous opioid and cannabinoid systems as well as reduction in DRG neuron excitability to the painless phenotype in a Nav1.7 loss-of-function rat. The complete loss of pain sensation in Nav1.7 loss-of-function rats is not a result of total loss of responsiveness of DRG neurons and is unlikely dependent on the endogenous opioid system. Our observations on pharmacological blockade of endogenous cannabinoids suggest a complex association between loss of Nav1.7 and function of the endogenous cannabinoid system. Alternative molecular mechanisms underlying the Nav1.7-related painless phenotype remain to be explicated.

Conflict of interest statement

The authors have no conflicts of interest to declare.

Appendix A. Supplemental digital content

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The authors thank Pamela Zwinger for technical assistance.

This work was supported by Center Grant B9253-C from the US Department of Veterans Affairs Rehabilitation Research and Development Service. The Center for Neuroscience and Regeneration Research is a Collaboration of the Paralyzed Veterans of America with Yale University.


[1]. Agarwal N, Pacher P, Tegeder I, Amaya F, Constantin CE, Brenner GJ, Rubino T, Michalski CW, Marsicano G, Monory K, Mackie K, Marian C, Batkai S, Parolaro D, Fischer MJ, Reeh P, Kunos G, Kress M, Lutz B, Woolf CJ, Kuner R. Cannabinoids mediate analgesia largely via peripheral type 1 cannabinoid receptors in nociceptors. Nat Neurosci 2007;10:870–9.
[2]. Ahmad S, Dahllund L, Eriksson AB, Hellgren D, Karlsson U, Lund PE, Meijer IA, Meury L, Mills T, Moody A, Morinville A, Morten J, O'Donnell D, Raynoschek C, Salter H, Rouleau GA, Krupp JJ. A stop codon mutation in SCN9A causes lack of pain sensation. Hum Mol Genet 2007;16:2114–21.
[3]. Akiyama T, Carstens E. Neural processing of itch. Neuroscience 2013;250:697–714.
[4]. Alexandrou AJ, Brown AR, Chapman ML, Estacion M, Turner J, Mis MA, Wilbrey A, Payne EC, Gutteridge A, Cox PJ, Doyle R, Printzenhoff D, Lin Z, Marron BE, West C, Swain NA, Storer RI, Stupple PA, Castle NA, Hounshell JA, Rivara M, Randall A, Dib-Hajj SD, Krafte D, Waxman SG, Patel MK, Butt RP, Stevens EB. Subtype-Selective small molecule inhibitors reveal a fundamental role for Nav1.7 in nociceptor electrogenesis, axonal conduction and presynaptic release. PLoS One 2016;11:e0152405.
[5]. Beaulieu P, Bisogno T, Punwar S, Farquhar-Smith WP, Ambrosino G, Di Marzo V, Rice AS. Role of the endogenous cannabinoid system in the formalin test of persistent pain in the rat. Eur J Pharmacol 2000;396:85–92.
[6]. Bennett DL, Woods CG. Painful and painless channelopathies. Lancet Neurol 2014;13:587–99.
[7]. Black JA, Frezel N, Dib-Hajj SD, Waxman SG. Expression of Nav1.7 in DRG neurons extends from peripheral terminals in the skin to central preterminal branches and terminals in the dorsal horn. Mol pain 2012;8:82.
[8]. Black JA, Hoeijmakers JG, Faber CG, Merkies IS, Waxman SG. NaV1.7: stress-induced changes in immunoreactivity within magnocellular neurosecretory neurons of the supraoptic nucleus. Mol pain 2013;9:39.
[9]. Bourane S, Duan B, Koch SC, Dalet A, Britz O, Garcia-Campmany L, Kim E, Cheng L, Ghosh A, Ma Q, Goulding M. Gate control of mechanical itch by a subpopulation of spinal cord interneurons. Science 2015;350:550–4.
[10]. Brenner DS, Golden JP, Gereau RWt. A novel behavioral assay for measuring cold sensation in mice. PLoS One 2012;7:e39765.
[11]. Calignano A, La Rana G, Giuffrida A, Piomelli D. Control of pain initiation by endogenous cannabinoids. Nature 1998;394:277–81.
[12]. Chen YC, Auer-Grumbach M, Matsukawa S, Zitzelsberger M, Themistocleous AC, Strom TM, Samara C, Moore AW, Cho LT, Young GT, Weiss C, Schabhuttl M, Stucka R, Schmid AB, Parman Y, Graul-Neumann L, Heinritz W, Passarge E, Watson RM, Hertz JM, Moog U, Baumgartner M, Valente EM, Pereira D, Restrepo CM, Katona I, Dusl M, Stendel C, Wieland T, Stafford F, Reimann F, von Au K, Finke C, Willems PJ, Nahorski MS, Shaikh SS, Carvalho OP, Nicholas AK, Karbani G, McAleer MA, Cilio MR, McHugh JC, Murphy SM, Irvine AD, Jensen UB, Windhager R, Weis J, Bergmann C, Rautenstrauss B, Baets J, De Jonghe P, Reilly MM, Kropatsch R, Kurth I, Chrast R, Michiue T, Bennett DL, Woods CG, Senderek J. Transcriptional regulator PRDM12 is essential for human pain perception. Nat Genet 2015;47:803–8.
[13]. Cox JJ, Reimann F, Nicholas AK, Thornton G, Roberts E, Springell K, Karbani G, Jafri H, Mannan J, Raashid Y, Al-Gazali L, Hamamy H, Valente EM, Gorman S, Williams R, McHale DP, Wood JN, Gribble FM, Woods CG. An SCN9A channelopathy causes congenital inability to experience pain. Nature 2006;444:894–8.
[14]. Cox JJ, Sheynin J, Shorer Z, Reimann F, Nicholas AK, Zubovic L, Baralle M, Wraige E, Manor E, Levy J, Woods CG, Parvari R. Congenital insensitivity to pain: novel SCN9A missense and in-frame deletion mutations. Hum Mutat 2010;31:E1670–1686.
[15]. Cravatt BF, Demarest K, Patricelli MP, Bracey MH, Giang DK, Martin BR, Lichtman AH. Supersensitivity to anandamide and enhanced endogenous cannabinoid signaling in mice lacking fatty acid amide hydrolase. Proc Natl Acad Sci U S A 2001;98:9371–6.
[16]. Cummins TR, Howe JR, Waxman SG. Slow closed-state inactivation: a novel mechanism underlying ramp currents in cells expressing the hNE/PN1 sodium channel. J Neurosci 1998;18:9607–19.
[17]. Darmani NA, Pandya DK. Involvement of other neurotransmitters in behaviors induced by the cannabinoid CB1 receptor antagonist SR 141716A in naive mice. J Neural Transm (Vienna) 2000;107:931–45.
[18]. Dib-Hajj SD, Choi JS, Macala LJ, Tyrrell L, Black JA, Cummins TR, Waxman SG. Transfection of rat or mouse neurons by biolistics or electroporation. Nat Protoc 2009;4:1118–26.
[19]. Dib-Hajj SD, Yang Y, Black JA, Waxman SG. The Na(V)1.7 sodium channel: from molecule to man. Nat Rev Neurosci 2013;14:49–62.
[20]. Einarsdottir E, Carlsson A, Minde J, Toolanen G, Svensson O, Solders G, Holmgren G, Holmberg D, Holmberg M. A mutation in the nerve growth factor beta gene (NGFB) causes loss of pain perception. Hum Mol Genet 2004;13:799–805.
[21]. Emery EC, Habib AM, Cox JJ, Nicholas AK, Gribble FM, Woods CG, Reimann F. Novel SCN9A mutations underlying extreme pain phenotypes: unexpected electrophysiological and clinical phenotype correlations. J Neurosci 2015;35:7674–81.
[22]. Flinspach M, Xu Q, Piekarz AD, Fellows R, Hagan R, Gibbs A, Liu Y, Neff RA, Freedman J, Eckert WA, Zhou M, Bonesteel R, Pennington MW, Eddinger KA, Yaksh TL, Hunter M, Swanson RV, Wickenden AD. Insensitivity to pain induced by a potent selective closed-state Nav1.7 inhibitor. Sci Rep 2017;7:39662.
[23]. Gingras J, Smith S, Matson DJ, Johnson D, Nye K, Couture L, Feric E, Yin R, Moyer BD, Peterson ML, Rottman JB, Beiler RJ, Malmberg AB, McDonough SI. Global Nav1.7 knockout mice recapitulate the phenotype of human congenital indifference to pain. PLoS One 2014;9:e105895.
[24]. Goldberg YP, MacFarlane J, MacDonald ML, Thompson J, Dube MP, Mattice M, Fraser R, Young C, Hossain S, Pape T, Payne B, Radomski C, Donaldson G, Ives E, Cox J, Younghusband HB, Green R, Duff A, Boltshauser E, Grinspan GA, Dimon JH, Sibley BG, Andria G, Toscano E, Kerdraon J, Bowsher D, Pimstone SN, Samuels ME, Sherrington R, Hayden MR. Loss-of-function mutations in the Nav1.7 gene underlie congenital indifference to pain in multiple human populations. Clin Genet 2007;71:311–19.
[25]. Grubinska B, Chen L, Alsaloum M, Rampal N, Matson D, Yang C, Taborn K, Zhang M, Youngblood B, Liu D, Galbreath E, Allred S, Lepherd M, Ferrando R, Kornecook T, Lehto SG, Waxman S, Moyer BD, Dib-Hajj S, Gingras J. [EXPRESS] Rat NaV1.7 loss-of-function genetic model: deficient nociceptive and neuropathic pain behavior with retained olfactory function and intra-epidermal nerve fibers. Mol pain 2019;15:1744806919881846.
[26]. Habib AM, Okorokov AL, Hill MN, Bras JT, Lee MC, Li S, Gossage SJ, van Drimmelen M, Morena M, Houlden H, Ramirez JD, Bennett DLH, Srivastava D, Cox JJ. Microdeletion in a FAAH pseudogene identified in a patient with high anandamide concentrations and pain insensitivity. Br J Anaesth 2019;123:e249–e253.
[27]. Han L, Ma C, Liu Q, Weng HJ, Cui Y, Tang Z, Kim Y, Nie H, Qu L, Patel KN, Li Z, McNeil B, He S, Guan Y, Xiao B, Lamotte RH, Dong X. A subpopulation of nociceptors specifically linked to itch. Nat Neurosci 2013;16:174–82.
[28]. Hoffmann T, Sharon O, Wittmann J, Carr RW, Vyshnevska A, Col RD, Nassar MA, Reeh PW, Weidner C. NaV1.7 and pain: contribution of peripheral nerves. PAIN 2018;159:496–506.
[29]. Ibrahim MM, Deng H, Zvonok A, Cockayne DA, Kwan J, Mata HP, Vanderah TW, Lai J, Porreca F, Makriyannis A, Malan TP Jr. Activation of CB2 cannabinoid receptors by AM1241 inhibits experimental neuropathic pain: pain inhibition by receptors not present in the CNS. Proc Natl Acad Sci U S A 2003;100:10529–33.
[30]. Indo Y, Tsuruta M, Hayashida Y, Karim MA, Ohta K, Kawano T, Mitsubuchi H, Tonoki H, Awaya Y, Matsuda I. Mutations in the TRKA/NGF receptor gene in patients with congenital insensitivity to pain with anhidrosis. Nat Genet 1996;13:485–8.
[31]. Isensee J, Krahe L, Moeller K, Pereira V, Sexton JE, Sun X, Emery E, Wood JN, Hucho T. Synergistic regulation of serotonin and opioid signaling contributes to pain insensitivity in Nav1.7 knockout mice. Sci signaling 2017;10:eaah4874.
[32]. Janoyan JJ, Crim JL, Darmani NA. Reversal of SR 141716A-induced head-twitch and ear-scratch responses in mice by delta 9-THC and other cannabinoids. Pharmacol Biochem Behav 2002;71:155–62.
[33]. Kanellopoulos AH, Koenig J, Huang H, Pyrski M. Mapping protein interactions of sodium channel NaV1.7 using epitope-tagged gene-targeted mice. EMBO J 2018;37:427–45.
[34]. Ledent C, Valverde O, Cossu G, Petitet F, Aubert JF, Beslot F, Bohme GA, Imperato A, Pedrazzini T, Roques BP, Vassart G, Fratta W, Parmentier M. Unresponsiveness to cannabinoids and reduced addictive effects of opiates in CB1 receptor knockout mice. Science 1999;283:401–4.
[35]. Lee JH, Park CK, Chen G, Han Q, Xie RG, Liu T, Ji RR, Lee SY. A monoclonal antibody that targets a NaV1.7 channel voltage sensor for pain and itch relief. Cell 2014;157:1393–404.
[36]. Liu Y, Abdel Samad O, Zhang L, Duan B, Tong Q, Lopes C, Ji RR, Lowell BB, Ma Q. VGLUT2-dependent glutamate release from nociceptors is required to sense pain and suppress itch. Neuron 2010;68:543–56.
[37]. McDermott LA, Weir GA, Themistocleous AC, Segerdahl AR, Blesneac I, Baskozos G, Clark AJ, Millar V, Peck LJ, Ebner D, Tracey I, Serra J, Bennett DL. Defining the functional role of NaV1.7 in human nociception. Neuron 2019;101:905–19.e908.
[38]. Minett MS, Nassar MA, Clark AK, Passmore G, Dickenson AH, Wang F, Malcangio M, Wood JN. Distinct Nav1.7-dependent pain sensations require different sets of sensory and sympathetic neurons. Nat Commun 2012;3:791.
[39]. Minett MS, Pereira V, Sikandar S, Matsuyama A, Lolignier S, Kanellopoulos AH, Mancini F, Iannetti GD, Bogdanov YD, Santana-Varela S, Millet Q, Baskozos G, MacAllister R, Cox JJ, Zhao J, Wood JN. Endogenous opioids contribute to insensitivity to pain in humans and mice lacking sodium channel Nav1.7. Nat Commun 2015;6:8967.
[40]. Mishra SK, Hoon MA. The cells and circuitry for itch responses in mice. Science 2013;340:968–71.
[41]. Morinville A, Fundin B, Meury L, Jureus A, Sandberg K, Krupp J, Ahmad S, O'Donnell D. Distribution of the voltage-gated sodium channel Na(v)1.7 in the rat: expression in the autonomic and endocrine systems. J Comp Neurol 2007;504:680–9.
[42]. Nahorski MS, Al-Gazali L, Hertecant J, Owen DJ, Borner GH, Chen YC, Benn CL, Carvalho OP, Shaikh SS, Phelan A, Robinson MS, Royle SJ, Woods CG. A novel disorder reveals clathrin heavy chain-22 is essential for human pain and touch development. Brain 2015;138:2147–60.
[43]. Nahorski MS, Chen YC, Woods CG. New mendelian disorders of painlessness. Trends Neurosciences 2015;38:712–24.
[44]. Nassar MA, Stirling LC, Forlani G, Baker MD, Matthews EA, Dickenson AH, Wood JN. Nociceptor-specific gene deletion reveals a major role for Nav1.7 (PN1) in acute and inflammatory pain. Proc Natl Acad Sci U S A 2004;101:12706–11.
[45]. Pereira V, Millet Q, Aramburu J, Lopez-Rodriguez C, Gaveriaux-Ruff C, Wood JN. Analgesia linked to Nav1.7 loss of function requires µ- and δ-opioid receptors. Wellcome open Res 2018;3:101.
[46]. Richardson JD, Aanonsen L, Hargreaves KM. SR 141716A, a cannabinoid receptor antagonist, produces hyperalgesia in untreated mice. Eur J Pharmacol 1997;319:R3–4.
[47]. Rush AM, Cummins TR, Waxman SG. Multiple sodium channels and their roles in electrogenesis within dorsal root ganglion neurons. J Physiol 2007;579:1–14.
[48]. Schlosburg JE, Boger DL, Cravatt BF, Lichtman AH. Endocannabinoid modulation of scratching response in an acute allergenic model: a new prospective neural therapeutic target for pruritus. J Pharmacol Exp Ther 2009;329:314–23.
[49]. Schlosburg JE, O'Neal ST, Conrad DH, Lichtman AH. CB1 receptors mediate rimonabant-induced pruritic responses in mice: investigation of locus of action. Psychopharmacology 2011;216:323–31.
[50]. Shields SD, Deng L, Reese RM, Dourado M, Tao J, Foreman O, Chang JH, Hackos DH. Insensitivity to pain upon adult-onset deletion of Nav1.7 or its blockade with selective inhibitors. J Neurosci 2018;38:10180–201.
[51]. Sotocinal SG, Sorge RE, Zaloum A, Tuttle AH, Martin LJ, Wieskopf JS, Mapplebeck JC, Wei P, Zhan S, Zhang S, McDougall JJ, King OD, Mogil JS. The Rat Grimace Scale: a partially automated method for quantifying pain in the laboratory rat via facial expressions. Mol pain 2011;7:55.
[52]. Sun YG, Chen ZF. A gastrin-releasing peptide receptor mediates the itch sensation in the spinal cord. Nature 2007;448:700–3.
[53]. Usoskin D, Furlan A, Islam S, Abdo H, Lonnerberg P, Lou D, Hjerling-Leffler J, Haeggstrom J, Kharchenko O, Kharchenko PV, Linnarsson S, Ernfors P. Unbiased classification of sensory neuron types by large-scale single-cell RNA sequencing. Nat Neurosci 2015;18:145–53.
[54]. Vasylyev DV, Han C, Zhao P, Dib-Hajj S, Waxman SG. Dynamic-clamp analysis of wild-type human Nav1.7 and erythromelalgia mutant channel L858H. J Neurophysiol 2014;111:1429–43.
[55]. Vetter I, Deuis JR, Mueller A, Israel MR, Starobova H, Zhang A, Rash LD, Mobli M. NaV1.7 as a pain target - from gene to pharmacology. Pharmacol Ther 2017;172:73–100.
[56]. Weiss J, Pyrski M, Jacobi E, Bufe B, Willnecker V, Schick B, Zizzari P, Gossage SJ, Greer CA, Leinders-Zufall T, Woods CG, Wood JN, Zufall F. Loss-of-function mutations in sodium channel Nav1.7 cause anosmia. Nature 2011;472:186–90.
[57]. Zimmer A, Zimmer AM, Hohmann AG, Herkenham M, Bonner TI. Increased mortality, hypoactivity, and hypoalgesia in cannabinoid CB1 receptor knockout mice. Proc Natl Acad Sci U S A 1999;96:5780–5.

Nav1.7; CIP; Nav1.7-related CIP; Endogenous opioid; Endogenous cannabinoid; SR141716A; CB1; Itch

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