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1Department of Oral Surgery and Diagnostic Sciences, Division of Neurosciences, College of Dentistry, 2Department of Physiological Sciences, College of Veterinary Medicine, and 3Department of Neuroscience, College of Medicine and University of Florida McKnight Brain Institute, Gainesville, Florida
Submitted 8 June 2004; accepted in final form 4 October 2004
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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nociceptors (types 4, 6, and 9) expressed slowly decaying nAChR. Three major forms of nicotinic currents were identified. Specific agonists and antagonists were used to demonstrate the presence of 
7 in two classes of capsaicin-sensitive, unmyelinated nociceptors (types 2 and 8). In type 2 cells, 7-mediated currents were found in isolation. Whereas 7 was co-expressed with other nAChR in type 8 cells. These were the only classes in which 7 was identified. Other nociceptive classes expressed slowly decaying currents with 
4 pharmacology. Based on concentration response curves formed by nicotinic agonists [ACh, nicotine, dimethyl phenyl piperazinium (DMPP), cytisine] evidence emerged of two distinct nAChR differentially expressed in type 4 (34) and types 5 and 8 (34 5). Although identification could not be made with absolute certainty, patterns of potency (type 4: DMPP > cytisine > nicotine = ACh; type 5 and type 8: DMPP = cytisine > nicotine = ACh) and efficacy provided strong support for the presence of two distinct channels based on an 34 platform. Studies conducted on one nonnociceptive class (type 3) failed to reveal any nAChR. After multiple injections of Di-I (1,1'-dilinoleyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate) into the hairy skin of the hindlimb, we identified cell types 2, 4, 6, 8, and 9 as skin nociceptors that expressed nicotinic receptors. We conclude that at least three nicotinic AChR are diversely distributed into discrete subclasses of nociceptors that innervate hairy skin. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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). The influence of these agents has been extensively investigated, yet our understanding of the interplay among the pain, vascular, immune, and endocrine systems over the course of trauma and healing remains fairly limited. Acetylcholine (ACh) is a prototypical pro-inflammatory agent that contributes to both pain and healing. ACh is liberated from fibroblasts, keratinocytes, epithelial, endothelial, and immune cells after trauma ( Buchli et al. 1999; Grando et al. 1993; Parnavelas et al. 1985; Wessler et al. 1998, 1999). Once liberated, ACh can activate A and C fiber nociceptors in superficial and deep tissues ( Bernardini et al. 2001; Bessou and Perl 1969; Fjallbrant and Iggo 1961; Haegerstam et al. 1975; Lang et al. 2003; Schmelz et al. 2003; Steen and Reeh 1993; Tanelian 1991) and activate CNS relays ( Carstens et al. 2000). Either ACh or nicotinic agonists can be painful to humans when applied at a variety of superficial and deep tissues sites including blister base, arteries, cornea, and tongue ( Armstrong et al. 1953; Dessirier et al. 1998, 1999; Keele and Armstrong 1964; Wilson and Stoner 1947).
Heteromeric neuronal nicotinic receptors (nAChR) can be formed from the pentameric assembly of 7 alpha (2, 3, 4, 5, 6, 9, 10) and 3 beta (2, 3, 4) subunits. Functional nAChR heteromers manifest distinct pharmacology, permeability, and kinetics ( Albuquerque et al. 1997; Dani 2001; McGehee and Role 1995; Papke 1993). Additional alpha subunits (7, 8, 9) combine to form homomeric channels with exceptional Ca2+ permeability ( Couturier et al. 1990; Elgoyhen et al. 1994; Gerzanich et al. 1994; Seguela et al. 1993). The composition and distribution of heteromeric nAChR, in vivo, is not known with any certainty, but the predominant forms in the mammalian CNS and PNS are believed to be the 42 and 34, respectively ( Flores et al. 1992; Genzen et al. 2001; Liu et al. 1998; Wada et al. 1989). All of these alpha and beta subunits are expressed in sensory ganglia (2, 3, 4, 5, 6, a7, 9, 10, 2, 3, 4) (Boyd et al. 1991; Flores et al. 1996; Genzen et al. 2001; Lips et al. 2002; Liu et al. 1998). These subunits could assemble into a variety of functional nAChR. The distribution of nAChR in nociceptive and nonnociceptive populations is not known, but it is reported that 
50% of DRG cells respond to nicotinic agonists ( Genzen et al. 2001; Liu et al. 1993; Sucher et al. 1990). A portion of these neurons are likely to be nociceptive.
There may be functional diversity among the nAChR expressing peripheral afferent pool. In addition to its clear nociceptive role, nicotinics may be able to confer a degree of analgesia ( Carstens et al. 2001; Kesingland et al. 2000; Reuter et al. 2003). These influences might arise from modulation of peptide release ( Bannon et al. 1998a, b; Donnelly-Roberts et al. 1998), cross desensitization ( Sudo et al. 2002), tachphylaxis ( Dessirier et al. 2000; Jinks and Carstens 1999), or direct interaction with voltage-dependent channels ( Liu et al. 2004). Contrasting nicotinic influences could also arise from separate pools of afferents that contribute distinct analgesic or algesic influences or could reflect a segregated distribution of algesic and analgesic conferring afferent pools to distinct tissue sites. Little is known about the distribution of nicotinic channels in nociceptors or the sites that they innervate.
Our laboratory has been concerned with primary afferent sub-specializations within the pain system. Scroggs and colleagues originally devised a method of classifying dorsal root ganglion cells, in vitro, that was based on the distribution of hyperpolarization-activated currents and voltage-activated Ca2+ channels ( Cardenas et al. 1995). Using methods derived from Scroggs and colleagues, we have recently shown that patterns of hyperpolarization and depolarization activated currents form signatures that could identify nine distinct afferent subpopulations with internally uniform properties and histochemistry ( Petruska et al. 2000a, b, 2002). Based on a variety of evidence, eight of these cell populations were likely to be nociceptive and represent members of the distinct subtypes of nociceptive afferents that have been characterized in vivo (C polymodal, C mechanoheat, C high-threshold mechanoreceptor, C cold, C-silent, A high-threshold mechanoreceptor, A mechanoheat, A polymodal, A cold, A-silent (mechanically insensitive afferent) ( Ringkamp et al. 2001; Treede et al. 1992).
Consistent with the substantial diversity of the nociceptive population in vivo, investigations in vitro have revealed that the sensory cells of the DRG were composed of discrete, internally homogenous, classes of capsaicin-sensitive (types 1, 2, 5, 7, 8 and 9) and -insensitive (types 3, 4, 6) populations with distinct capacities to respond to 5HT, PGE2, protons, and ATP ( Cardenas et al. 1995, 1997, 1999; Cooper et al. 2004; Petruska et al. 2000a, 2002). Moreover, they are composed of histochemically uniform groups of IB4-positive (types 1, 2, 5, 7, 8) and -negative (type 3, 4, 6, and 9) cells that expressed CGRP (type 6 and 9), co-expressed CGRP and SP (types 5, 7, and 8), CGRP and somatostatin (type 1) or failed to express any of these peptides (types 2–4). It is likely that some of these cell classes represent those groups that express nAChR and mediate the afferent discharge and pain that are a consequence of nicotinic agonists. In the studies presented in the following text, we examined the distribution and peripheral representation of nAChR in subclassified nociceptive cells.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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Adult male rats (Harlan Sprague-Dawley, 90–150 g) were anesthetized with halothane and rapidly decapitated (Braintree Scientific, RG-100). The spinal cord was quickly removed, and 10–15 dorsal root ganglia were dissected free. Dissected cervical, thoracic, and lumbar ganglia were placed in a heated bath (35°C for 70 min) containing dispase (neutral protease, 5 mg/ml; Boehringer Mannheim) and collagenase (2 mg/ml; Sigma type 1). After wash and trituration, recovered cells were plated on 12 polylysine-coated petri dishes. Recordings were made at room temperature, 2–10 h after plating. Plated cells were maintained in a rat Tyrode's solution containing (in mM) 140 NaCl, 4 KCl, 2 MgCl2, 2 CaCl2, 10 glucose, and 10 HEPES, adjusted to pH 7.4 with NaOH). The pipette solution contained (in mM) 120 KCl, 5 Na2-ATP, 0.4 Na2-GTP, 5 EGTA, 2.25 CaCl2, 5 MgCl2, and 20 HEPES, adjusted to pH 7.4 with KOH. These methods were consistent with the Panel on Euthanasia of the American Veterinary Medical Association. All animal purchases, housing, and veterinary care was provided by Animal Care Services. A local IACUC committee reviewed and approved all procedures involving animals prior to any experimentation.
Cell classification and testing
Recordings were made exclusively from cells with diameters between 17 and 45 µm. Cell diameter was estimated from the average of the longest and shortest axis as measured through an eyepiece micrometer scale. All recordings were made at room temperature. After the whole cell mode was achieved, series resistance was compensated 30–60%. A junction potential error of 4 mV was not corrected. Other cell and membrane properties are presented in Table 1.
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, 2002).
When recording from small cells, application of CP1 revealed cells that expressed small amplitude H currents (<100 pA). These could be divided into types 3 and 7 using CP3. Type 3 cells had fast decaying, low-threshold inward currents. In contrast, type 7 cells exhibited high-threshold, slow decaying inward currents. (Fig. 1G). Both types 3 and 7 manifested similar CP2 patterns. Other small cells did not express H current and had distinct outward current patterns that were devoid of sharp A-current peaks (Fig. 1A). These cells were classified as type 1. A fourth group of small-diameter cells exhibited no IH but a strong transient outward current appeared on repolarization (Fig. 1B). Such neurons were classified as type 2. From the medium-diameter pool, we encountered many neurons that exhibited slow activating, large-amplitude hyperpolarization-activated currents (>500 pA). These were recognized as type 4 cells (Fig. 1D). Less frequently, we encountered medium-sized cells with fast-activating, weak hyperpolarization-activated currents (<200 pA at –110 mV). CP2 distinguished between patterns of outward currents that differentiated types 5 and 8. Both of these classes exhibited classic A-current peaks but differed in the number (threshold) of peaks revealed by CP2. If three peaks were observed, the cell was type 5 (Fig. 1E). If four peaks were observed, the cell was type 8 (Fig. 1H). If five peaks were observed, the cell was type 6 (Fig. 1F). The number of A-current peaks can also be defined as an A-current threshold (AT = 0, –20 and –40 mV). This is equivalent to counting peaks when using CP2 as defined in the preceding text. Another group of medium-sized cells were devoid of hyperpolarization-activated currents and exhibited intermediate threshold A-current peaks (4 peaks or AT = –20 mV). These cells were classified as type 9 (Fig. 1I). CP3 was not required to classify types 5, 6, 8, and 9. These cells classes have proven to have highly uniform properties ( Petruska et al. 2000a, 2002).
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BgTX (alpha-bungarotoxin), MLA (methyllycaconitine), mecamylamine, atropine, muscarine, and biccuculine]. BgTX, 4-OH-GTS21, and TC 2403 were kindly provided by Dr. R. Papke. All other agents were purchased from Sigma-Aldrich. Inhibitors were applied for 2 min prior to test application and were contained within test solutions. Inhibitors were washed out over a 2- to 3-min interval. Concentration response curves were formed from an ascending series of agonist superfusions that were separated by 2-min intervals. A maximal dose of ACh (640–1280 µM) was applied 4 min after the agonist peak response was determined. EC50s and efficacies were determined from these concentration response curves. All substances were presented by gravity fed sewer pipe positioned 1 mm distant from the recorded cell. Afferent tracing
Under aseptic conditions, five young adult male rats (80–100 g) were anesthetized with a mixture of ketamine and xylazine (ip injection; 80 mg/kg ketamine; 10 mg/kg xylazine). The following signs were monitored during surgery: heart rate, respiratory rate, ventilatory status (end-expired pCO2), and body temperature. Anesthetic depth was assessed by corneal, palpebral, and pinna reflexes. The animals were placed on a heating pad to maintain ideal body temperature (36–37°C). Anesthetic supplements were administered by ip injection when necessary. A small transverse incision was made through the hairy skin, lateral and caudal to the gastrocnemius muscle. Intradermal injections of the fluorescent tracer FastDiI oil (1,1'-dilinoleyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate; Molecular Probes) were then made several centimeters rostral to the incision site in the underside of the skin with a 33-gauge needle coupled to a Hamilton microsyringe (20 ml volume per animal divided into 10 injections per limb of 1 ml each). After each injection, the needle was slowly removed, any leakage was controlled by cotton-tipped applicators, and the site was rapidly sealed with n-butyl cryanoacrylate monomer glue (either Nexaband Liquid or SteriTac-B). After injections were completed in a limb, the incision was closed with cryanoacrylate. Rats were monitored daily and allowed to recover for 7 days. They were then killed for in vitro electrophysiological studies. Cells were plated in the usual manner but protected from ambient light. Dishes were mounted on a Nikon Diaphot inverted microscope with an epifluorescence attachment. Tracer-labeled cells were viewed with the appropriate Vivid filter set (XF102, Omega Optical), and ultraviolet light exposure to all fields was <1 min in duration. Only intensely fluorescent cells were considered positive. Cells were classified as type 1–9 or unknown (Fig. 1). Only one cell was recorded per dish. After a recording was completed, digital images of the brightfield and fluorescent fields of view were captured using a Dage MTI RC300 camera coupled to a PC running Scion Image 4.0.2.
To assess the possible spread of DiI from injection sites, injected tissue and underlying muscle tissues were harvested prior to plating the DRG cells. The tissues were placed in vials containing 4% paraformaldehyde in phosphate-buffered saline (PBS) for a 24-h period. Subsequently, this fixative solution was replaced by 30% sucrose in PBS for cryoprotection. Once the tissue equilibrated it was embedded in TBS Tissue Freezing Medium (Triangle Biomedical Sciences) and 10-µm sections were cut on a cryostat (HM 550; Microm). Sections were thaw-mounted onto slides and placed in a –20°C freezer until viewed under fluorescent microscopy. Cases in which DiI had leaked into underlying muscle tissue were not included.
Statistics
EC50s were determined by fit of the normalized data to a function of the form: I = Imax/[1 + (EC50/[Ag])n], where Imax is the peak current and [Ag] is the agonist concentration. Student's t-test was used to compare EC50's and efficacy (agonistmax/AChmax) derived from concentration response curves of distinct nociceptive cell types. The alpha level was set at 0.05. In some instances, a 
decay was also determined. The exponential decay constants () were derived from the expression A1exp[–(t – k)/1]...+ C (Clampfit 6.0). Fits were made at points between 10% of the peak current and 90% of the return to baseline using Clampfit software (Axon Instruments).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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). All other classes exhibited some form of ACh evoked current, and in some cases, multiple currents could be demonstrated (Fig. 2C). In reactive classes, application of ACh produced large-amplitude currents that were highly consistent, in form, within each cell class. Across all classes, the proportion of cells responding to ACh approached 100 or 0% in each cell class (Fig. 2, E–M). The main exception was the type 5 cell class in which a portion of cells (13/58) failed to respond to ACh. We believe this represents a genuine subphenotype of the type 5 class.
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7 or muscarinic receptors, detailed characterization of the large, slowly adapting currents were carried out in the presence of methyllycaconitine (MLA, 50 nM) and atropine (500 nM). These antagonists were preapplied (2 min) and were also contained in agonist solutions. In selected cases, we replicated capsaicin sensitivity after application of nicotinic agonists. As previously reported, all type 1, 2, 5, and 8 cells were capsaicin sensitive (n = 7/7, 25/25, 20/20, and 12/12, respectively). In contrast, type 4 cells with nAChR were never capsaicin sensitive (0/21).
Fast currents with 7-like properties
Two afferent neuron subclasses expressed currents consistent with those of 7 homomeric channels. In the type 8 cell, 7 like currents were present with other nAChR, but they were expressed in isolation in the type 2 cell. Presentation of ACh (640 µM; n = 33) to the capsaicin sensitive type 2 cells consistently evoked a small-amplitude current that rapidly decayed with monophasic kinetics (tau = 165 ± 0.009 ms; Fig. 2B). Fast currents in type 2 cells could be activated by choline (500 µM) or the 7-specific agonist 4-OH-GTS21 (Fig. 3A–C), blocked by MLA (n = 3; 50–200) or BGTX (3 µM, n = 3; not shown). When activated by ACh or choline, 7-like currents were classic, fast decaying currents. When activated with 4-OH-GTS21, the 7-like currents were slowly desensitizing. The 9 agonist oxotremorine (100 µM) failed to evoke any currents in type 2 cells (n = 2; not shown). Therefore choline-activated currents were unlikely to be mediated by 9 containing channels ( Verbitsky et al. 2000).
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7-like currents, but these could only be demonstrated with specific agonists as large, slowly decaying, nicotinic currents were also present (see following text). Application of 4-OH-GTS21 confirmed that 7 was present in the type 8 cell (Fig. 3, F and G). These were fast decaying currents. In contrast, there was little indication of 4-OH-GTS21-sensitive currents in type 4 or 5 cells (0/8 and 1/6 cases, respectively). We did not apply 4-OH-GTS21 to cell types 1, 6, or 9.
Slow currents with 4-like properties
Combinations of alpha and beta subunits can yield nicotinic AChR with varying potency, efficacy, kinetics, and permeability ( Albuquerque et al. 1997; Dani 2001; McGhee and Role 1995). Relative to channels formed from 7 proteins, those formed as or heteromers exhibit currents that decay relatively slowly. Many DRG cell classes, particularly medium-sized capsaicin-sensitive (type 5, 8, and 9) and -insensitive cells (type 4 and 6), expressed slowly decaying currents that could be evoked by a variety of nicotinic agonists (Fig. 4). These large-amplitude currents were completely blocked by mecamylamine (40 µM; not shown; n = 6, 5, and 3, respectively, types 4, 5, and 8).
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7 was present in type 8, and this channel (as well as muscarinic receptors) could distort the apparent ACh potency in this cell classes by binding ACh or through initiating secondary messenger or Ca2+-dependent regulatory pathways. Although 7 was absent in types 4 and 5 (see preceding text), the presence of atropine and MLA significantly shifted the EC50 for ACh in both of these cell classes (Table 2; Fig. 5B). When MLA was excluded (ACh atropine-MLA vs. ACh-atropine), there was little indication of any influence of MLA (Fig. 5C). It was possible that muscarinic receptors were present and responsible for the shifted potency. Accordingly, all concentration response curves formed on members of these cell classes were carried out in the presence of atropine and MLA (ATR-MLA).
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4 containing heteromers ( Chavez-Noriega et al. 1997; Luetje and Patrick 1991; Papke and Heineman 1994). All subclassified DRG nociceptors exhibited a high efficacy and potency to cytisine over an application range of (5–300 µM; Fig. 4; Table 2). Once again, cytisine evoked currents were significantly right-shifted in type 4 cells relative to those from types 5 and 8. Efficacy shifts were also apparent with the same pattern obtained (Fig. 5F; Table 2). It is likely that these shifts in potency and efficacy for cytisine reflected distinct composition of channels expressed in nociceptive cells and, in particular, the number of 4 subunits. Difference in the relative potency of these nicotinic agonists in capsaicin-insensitive (type 4) and sensitive types 5 and 8 cells further emphasized the distinct distribution of nAChR (type 4: DMPP > cytisine > nicotine = ACh; types 5 and 8: DMPP = cytisine > nicotine = ACh). A series of experiments were performed to further specify the nAChR of the type 4 nociceptor. Despite the relatively smooth curves that were obtained, it was possible that these cells contained multiple functional nAChR, the presence of which were responsible for shifts in potency and efficacy relative to other subclassified cells. We used specific agonists and select antagonists to examine this possibility. To study the influence of antagonists, we used a procedure of repeated application of ACh in the presence and absence of selected agents (ACh 640 µM; 3-min intervals). After the first application, there was a significant tachyphylaxis. However, reactivity stabilized from application two through four so that highly consistent responses could be obtained (Fig. 6B; n = 5). The selected antagonist was applied during the interval between the second and third ACh presentation. The third application of ACh (also containing the selected antagonist) served as a test, and the fourth application was used as a wash. Neither atropine nor MLA was present in these experiments.
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9 or 910 like protein. These proteins have been reported in trigeminal and dorsal root ganglion neurons ( Lips et al. 2002; Liu et al. 1998). Pretreatment (2 min) with bicuculline (20 µM) significantly and reversibly reduced ACh- evoked current in type 4 cells (Fig. 6C; n = 8). These reductions were consistent with 9 or 910, (but not inconsistent with 34 or 24) ( Demuro et al. 2001). Although a relatively small portion of the total current was reduced (<33%), it was possible that a weakly co-expressed channel imparted the distinct properties found in this cell class. Further and more specific examinations failed to confirm the presence of 9 containing nAChR. Muscarine is an antagonist at 9 containing AChR ( Elgoyhen et al. 2001; Verbitsky et al. 2000). We were unable to demonstrate that any portion of the stable ACh-evoked current could be inhibited by muscarine (Fig. 6E). Moreover, 9 and 910 agonists choline (500 µM, n = 3) or oxotremorine (100 µM; n = 8) also failed to evoke any current in type 4 cells (Fig. 6D) ( Elgoyen et al. 1994, 2001; Rothlin et al. 1999; Verbitsky et al. 2000). The specific 42 agonist TC 2403 was also ineffective in type 4 or 5 cells (120 µM, n = 2 and 3, respectively) ( Papke et al. 2000). Therefore we could not demonstrate a co-expressed nAChR in type 4 cells (or type 5) that could impart distinct characteristics observed in concentration response curves. Projection fields of nAChR-expressing nociceptors
It has been reported that pain and/or nociceptor activity occurs after application of nicotinic agonists to skin (see INTRODUCTION). One or more of those nociceptive populations we have characterized could mediate these influences. Accordingly, we made a series of small (1 µl) injections of the tracer, Di-I, into the hairy skin overlying the gastrocnemius. After a 2-wk period to allow for transport to the ganglion, the rat was killed, and ganglia were harvested in the usual manner. Recordings made exclusively from intensely fluorescent cells revealed that populations of skin afferents corresponded to our nociceptive cell class types 2, 4, 6, 8, and 9 (n = 13, 7, 3, 2, and 3, respectively; Fig. 7). Fluorescent cells that were exposed to ACh (640 µM) responded with characteristic fast or slow decaying currents (Fig. 7). Other fluorescent cells were observed, and a portion of these were also ACh responsive (2/8 small and 5/7 medium sized neurons; not shown). Most of these noncategorized cells were capsaicin sensitive (6/8 small and 4/7 medium-diameter cells; 1 µM) and likely to be nociceptive.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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; Liu and Simon 1996; Liu et al. 1993; Papadopolou et al. 2004; Sucher et al. 1990). We now know that DRG neurons can be classified by physiological current signatures into distinct groups that exhibit highly uniform reactivity to algesic substances. Subpopulations of myelinated and unmyelinated afferents that uniformly express receptors for capsaicin, ATP, and protons have been extensively characterized using these methods ( Cardenas et al. 1995; Petruska et al. 2000a, b, 2002). We now demonstrate that these same populations also differentially expressed nAChR (Table 3) and include subfamilies that innervate hairy skin. These nicotinic receptors were observed both in small- and medium-sized cell classes and in both capsaicin-sensitive and -insensitive cells. It is possible that portions of such afferent groups contribute to nociceptor discharge, nocifensive behavior and pain report after application of ACh or nicotinic agonists to skin ( Bernardini et al. 2001; Lang et al. 2003; Schmelz et al. 2003; Steen and Reeh 1993).
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; Lang et al. 2003; Schmelz et al. 2003; Steen and Reeh 1993), and release of peptides by nicotinic agonists ( Benarroch and Low 1991; Parkhous and Lequesne 1988; Walmsley and Wiles 1990). These findings, for both rat and human, are consistent with our observations of nicotinic AChR in C-fiber peptidergic nociceptors in skin. As previously reported, capsaicin-sensitive cell types 5 and 8, bind the C fiber marker isolectin B4, and co-express SP and CGRP ( Petruska et al. 2002). While not directly demonstrated here, the Ca2+-permeable nicotinic currents would likely support release of peptides as would any action potentials that would occur after evocation of these powerful currents.
A restricted portion of cells expressed fast decaying currents that were physiologically and pharmacologically consistent with homomeric 7 channels. Time constants derived on the falling phase of the currents activated by ACh or choline were consistent with 7 ( Cuevas et al. 2000; McGehee and Role 1995). More significantly, MLA and Btx-sensitive currents could be reliably activated by the highly specific 7 agonist 4-OH-GTS21 ( Meyer et al. 1998; Papke et al. 2004). The fast decaying currents were consistent with classic 7 nAChR and differed from some 7 variants recently reported in superior cervical ganglion ( Cuevas et al. 2000). In DRG, we observed 7 in isolation from other nicotinics in the capsaicin-sensitive small-diameter, nonpeptidergic type 2 class as well as co-expressed with other nAChR in the capsaicin sensitive, peptidergic medium-diameter type 8 cell. These classes are likely to be C fiber nociceptors based on binding of isolectin B4 and capsaicin sensitivity ( Petruska et al. 2000b, 2002). Tracing experiments demonstrated that both of these cell classes were found in hairy skin. We did not observe 7 in myelinated, capsaicin insensitive nociceptors (type 4) or the unmyelinated capsaicin sensitive peptidergic type 5 cells. Nor did we observe 7 in other capsaicin-sensitive nociceptors of hairy skin. Therefore 7 was not uniformly expressed in C fiber nociceptors but was present in select, functionally distinct subgroups that differ not only with respect to peptide content, but also proton sensitivity ( Petruska et al. 2000b, 2002). The 7 channel is frequently found at presynaptic sites where it is believed to play a role in transmitter release in CNS neurons ( MacDermott et al. 1999; Wonnacott 1997). It may play the same role in central processes of some C fiber nociceptors. Type 2 cells are known to terminate in the lamina I and IIo of the dorsal horn ( Del Mar and Scroggs 1996). The central terminals of type 8 cells are not known, although peptidergic afferents are also known to terminate in lamina I, IIo as well as V ( Snider and McMahon 1998).
Most DRG nociceptive cells expressed a slowly decaying nicotinic current (type 1, 4, 5, 6, 8, 9). Although slowly decaying nicotinic currents have been previously reported in DRG, the molecular identity of this current(s), and its distribution among functional groups (i.e., nociceptors) is not known ( Genzen et al. 2001; Liu and Simon 1996; Liu et al. 1993; Sucher et al. 1990). A substantial variety of nicotinic subunits can combine to mediate slowly decaying currents. Many of these proteins have been identified in rat DRG or in central processes of primary afferents (3, 4, 5, 7, 9, 10, 2, 3, 4) ( Flores et al. 1996; Genzen et al. 2001; Lips et al. 2002; Vincler and Eisenach 2004). These subunits assemble into a diverse family of functional channels ( Albuquerque et al. 1997; Dani 2001; McGehee and Role 1995) and have been shown to co-elute in chick ganglia in heteromers consisting of up to four different protein subunits ( Conroy and Berg 1995). Because of the potential complexity, and the lack of specific agonists/antagonists, precise identification of a channel is not always possible. Nevertheless, many candidates can be ruled out. We were unable to evoked currents with the 42 specific agonist TC 2403 in types 4 or 5 ( Papke et al. 2000). Moreover, the patterns of potency and efficacy were inconsistent with the presence of 42 in subclassified DRG nociceptors (Table 4). In limited testing, we could not find any evidence of functional 9 or 910, in DRG nociceptors. The relative sensitivity of these channels to cytisine indicated that 4 expressing nAChR were present on all myelinated nociceptors we examined (types 4, 6, and 9). These include both capsaicin-sensitive (type 9) and -insensitive populations (type 4 and 6).
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34; however, there is only indirect evidence for this contention, and it is mainly based on single dose, relative cytisine potency ( Genzen et al. 2001). When nicotinic channels are reconstituted in expression systems, distinct patterns of potency and efficacy can be observed when diverse nicotinic agonists are presented. This approach can be applied to neurons when homogenous populations of cells can be identified. The classification procedure we employed made it possible to examine the nicotinic pharmacology of cells in a systematic fashion and in substantial detail in DRG neurons. We found that these slow currents exhibited properties that were highly consistent within cells classes but clearly distinct between some classes. Despite the substantial potential for diversity that was possible (based on a large number of subunits combinations predicted), the cell classes we examined expressed a limited subset of the slow decaying nAChR family. It was likely that mainly two isoforms were present. Based on the patterns of relative potency for nicotinic agonists, it was likely that the channel expressed by the type 4 cells (DMPP > Cyt > Nic = ACh) was significantly different from that of types 5 and 8 (DMPP = Cyt > Nic = ACh). Neither of these relative potency patterns are identical to that reported for reconstituted rat 34 (DMPP = Cyt = Nic > ACh; see Table 4 references). Nevertheless, an 34 platform (34 and possibly additional subunits) was likely given the pattern of potency and efficacy observed (Tables 2 and 4). Cell types 4, 5, 6, 8, and 9, all displayed a high potency and efficacy to cytisine and were likely to contain one or more 4 subunits (20–70 µM) ( Chavez-Noriega et al. 1997; Luetje and Patrick 1991; Papke and Heineman 1994). Moreover, both the absolute and relative potency of four nicotinic agonists were very consistent with 34 or a channel containing 34 and other protein. If two distinct nAChR were expressed by cell types 4, 5, and 8, then the simplest prediction is that one of these expressed 34 (e.g., type 4) and the other expressed 345 (e.g., types 5 and 8). This scenario is consistent with the observed a shift in cytisine potency in types 5 and 8 cells relative to type 4 and consistent with heteromerization of 5 with 34 as reported by Gerzanich and colleagues for human nAChR (70–20 µM) ( Gerzanich et al. 1998; see also Wang et al. 1996; Yu and Role 1998). We cannot rule out the possibility that addition subunits could be present or unrecognized posttranslational modifications might complicate interpretations. It is clear from our studies that nAChR are widely distributed in capsaicin-sensitive and -insensitive nociceptive cells, are present in both myelinated and unmyelinated cell classes in peptidergic and nonpeptidergic groups, and are present in several populations of skin nociceptors (types 2, 4, 6, 8, and 9). It is likely that the powerful currents of these nicotinic channels play a major role in algesic reactivity. As previously observed with respect to ATP and protons, the nociceptor population has evolved multiple strategies to detect each particular algesic. Quite distinct nicotinic, purinergic and proton-sensitive channels can be found in each nociceptor population (Table 3). The utility of this complexity is not certain but likely reflects particular nervous system adaptations required to detect the spectrum of algesics that are likely to be encountered in specialized tissues (skin, muscle, joint, etc.).
ACh is contained within a substantial variety of cells that are distributed into cutaneous and deep tissues. The release of ACh from keratinocytes, fibroblasts, endothelial cells, immune cells, and linings of viscera would result from local trauma or other release mechanisms ( Buchli et al. 1999; Grando et al. 1993; Parnavelas et al. 1985; Wessler et al. 1998, 1999). As noted, ACh-dependent nociceptor activation and pain have been demonstrated from cutaneous and deep tissues ( Bernardini et al. 2001; Bessou and Perl 1969; Fjallbrant and Iggo 1961; Haegerstam et al. 1975; Keele and Armstrong 1964; Lang et al. 2003; Schmelz et al. 2003; Steen and Reeh 1993; Tanelian 1991; Wilson and Stoner 1947). It is likely that select populations of nociceptors, expressing large powerful nAChR, would be closely associated with tissues populated by ACh-expressing cells. It is further possible that the mechanical allodynia that is common with wounds has a peripheral nicotinic component. Fragile tissue, in and around wound margins, are subject to mechanical disruption for several days following a trauma. Consequent to excessive movement or manipulation of healing tissues, the release of ACh from keratinocytes or endothelial cells would serve as a mechanical-chemo algesic signal. A mechano-chemo mechanism of this sort would include not only nociceptor activation and acute pain but also Ca2+-dependent upregulatory events through Ca2+-permeable nAChr ( Albuquerque et al. 1997; Dani 2001; McGehee and Role 1995). In this manner, nicotinic receptors could play a key role in the regulation of mechanical allodynia by contributing important sensory feedback relevant to the mechanical state of wound margins. This sensory component would complement known nicotinic contributions to wound healing ( Zia et al. 2000).
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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ACKNOWLEDGMENTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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FOOTNOTES |
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Address for reprint requests and other correspondence: B. Cooper, Dept. of Oral Surgery and Diagnostic Sciences, Div. of Neuroscience, Box 100416, JHMC, University of Florida College of Dentistry, Gainesville, FL 32610 (E-mail: Bcooper{at}dental.ufl.edu)
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REFERENCES |
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Armstrong DDRML, Keele CA, and Markham JW. Observations on chemical excitants of cutaneous pain in man. J Physiol 120: 316–351, 1953.
Bannon AW, Decker MW, Holladay MW, Curzon P, Donnelly-Roberts D, Puttfarcken PS, Bitner RS, Diaz A, Dickenson AH, Porsolt RD, Williams M, and Arneric SP. Broad-spectrum, non-opiod analgesic activity by selective modulation of neuronal nicotinic acetylcholine receptors. Science 279: 77–81, 1998a.
Bannon AW, Decker MW, Kim DJ, Campbell JE, and Arneric SP. ABT-594, a novel cholinergic channel modulator, is efficacious in nerve ligation and diabetic neuropathy models of neuropathic pain. Brain Res 801: 158–163, 1998b.[CrossRef][ISI][Medline]
Benarroch EE and Low PA. The acetylcholine-induced flare response in evaluation of small fiber dysfunction. Ann Neurol 29: 590–595, 1991.[CrossRef][ISI][Medline]
Bernardini N, Sauer SK, Haberberger R, Fischer MJ, and Reeh PW. Excitatory nicotinic and desensitizing muscarinic (M2) effects on C-nociceptors in isolated rat skin. J Neurosci 21: 3295–3302, 2001.
Bessou P and Perl ER. Response of cutaneous sensory units with unmyelinated fibers to noxious stimuli. J Neurophysiol 32: 1025–1043, 1969.
Boyd RT, Jacob MH, McEachern AE, Caron S, and Berg DK. Nicotinic acetylcholine receptor mRNA in dorsal root ganglion neurons. J Neurobiol 22: 1–14, 1991.[CrossRef][ISI][Medline]
Buchli R, Ndoye A, Rodriguez JG, Zia S, Webber RJ, and Grando SA. Human skin fibroblasts express m2, m4, and m5 subtypes of muscarinic acetylcholine receptors. J Cell Biochem 74: 264–277, 1999.[CrossRef][ISI][Medline]
Cardenas CG, Del Mar LP, and Scroggs RS. Variation in serotonergic inhibition of calcium channel currents in four types of rat sensory neurons differentiated by membrane properties. J Neurophysiol 74: 1870–1879, 1995.
Cardenas CG, Del Mar LP, and Scroggs RS. Two parallel signaling pathways couple 5HT1A receptors to N- and L-type calcium channels in C-like rat dorsal root ganglion cells. J Neurophysiol 77: 3284–3296, 1997.
Cardenas CG, Mar LP, Vysokanov AV, Arnold PB, Cardenas LM, Surmeier DJ, and Scroggs RS. Serotonergic modulation of hyperpolarization-activated current in acutely isolated rat dorsal root ganglion neurons. J Physiol 518: 507–523, 1999.
Carstens E, Simons CT, Dessirier JM, Carstens MI, and Jinks SL. Role of neuronal nicotinic-acetylcholine receptors in the activation of neurons in trigeminal subnucleus caudalis by nicotine delivered to the oral mucosa. Exp Brain Res 132: 375–383, 2000.[CrossRef][ISI][Medline]
Carstens E, Andersen KA, Simons CT, Carstens MI, and Jinks SL. Analgesia induced by chronic nicotine infusion in rats: differences by gender and pain. Psychopharmacology 157: 40–45, 2001.[CrossRef][Medline]
Chavez-Noriega LE, Crona JH, Washburn MS, Urrutia A, Elliott KJ, and Johnson EC. Pharmacological characterization of recombinant human neuronal nicotinic acetylcholine receptors h alpha 2 beta 2, h alpha 2 beta 4, h alpha 3 beta 2, h alpha 3 beta 4, h alpha 4 beta 2, h alpha 4 beta 4 and h alpha 7 expressed in Xenopus oocytes. J Pharmacol Exp Ther 280: 346–356, 1997.
Conroy WG and Berg DK. Neurons can maintain multiple classes of nicotinic acetylcholine receptors distinguished by different subunit compositions. J Biol Chem 270: 4424–4431, 1995.
Cooper BY, Johnson RD, and Rau KK. Characterization and function of TASK channels in a rat nociceptive cell. Neuroscience 129: 209–224, 2004.[CrossRef][ISI][Medline]
Cooper BY, Rau K, and Johnson RD. Heat reactivity and TRP expression in capsaicin sensitive and insensitive subclassified sensory cells of the rat DRG. Soc Neurosci Abstr 381.11, 2003.
Cooper BY and Sessle BJ. Anatomy and physiology of the trigeminal system, In: The Headaches, edited by Oleson J, Tfelt-Hansen P, and Welch KMA. New York:Raven, 1993, p. 87–93.
Couturier S, Erkman L, Valera S, Rungger D, Bertrand S, Boulter J, Ballivet M, and Bertrand D. Alpha 5, alpha 3, and non-alpha 3. Three clustered avian genes encoding neuronal nicotinic acetylcholine receptor-related subu
