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Altered ATP release and metabolism in dorsal root ganglia of neuropathic rats



Adenosine 5'-triphosphate (ATP) has a ubiquitous role in metabolism and a major role in pain responses after tissue injury. We investigated the changes in basal and KCl-evoked ATP release from rat dorsal root ganglia (DRG) after peripheral neuropathy induction by unilateral sciatic nerve entrapment (SNE).


After SNE, rats develop long-lasting decreases in ipsilateral hindpaw withdrawal thresholds to mechanical and thermal stimulation. At 15–21 days after neuropathy induction, excised ipsilateral L4-L5 DRG display significantly elevated basal extracellular ATP levels compared to contralateral or control (naive) DRG. However, KCl-evoked ATP release is no longer observed in ipsilateral DRG. We hypothesized that the differential SNE effects on basal and evoked ATP release could result from the conversion of extracellular ATP to adenosine with subsequent activation of adenosine A1 receptors (A1Rs) on DRG neurons. Adding the selective A1R agonist, 2-chloro-N6-cyclopentyladenosine (100 nM) significantly decreased basal and evoked ATP release in DRG from naïve rats, indicating functional A1R activation. In DRG ipsilateral to SNE, adding a selective A1R antagonist, 8-cyclopentyl-1,3-dipropylxanthine (30 nM), further increased basal ATP levels and relieved the blockade of KCl-evoked ATP release suggesting that increased A1R activation attenuates evoked ATP release in neurons ipsilateral to SNE. To determine if altered ATP release was a consequence of altered DRG metabolism we compared O2 consumption between control and neuropathic DRG. DRG ipsilateral to SNE consumed O2 at a higher rate than control or contralateral DRG.


These data suggest that peripheral nerve entrapment increases DRG metabolism and ATP release, which in turn is modulated by increased A1R activation.


Certain neuropathic pain states that result from peripheral nerve injury are associated with hyperexcitability of neurons within sensory ganglia [1] and potentiated cross-excitation among neighboring sensory neurons in the absence of synaptic specializations [2]. Cross-excitation occurs when discharge in one sensory neuron leads to a depolarization in the cell bodies of adjacent passive neurons sharing the same ganglion. Cross-excitation was reported to be chemically mediated [3], but the identity of the chemical mediator(s) is unknown. One possible candidate is ATP which, in addition to its universal role in metabolism, is released from stimulated sensory nerves [4]. There is compelling evidence for exocytotic vesicular release of ATP from neurons [5] and we have demonstrated Ca2+-dependent ATP release within trigeminal ganglia after neuronal stimulation in vivo, as well as from isolated somata of trigeminal sensory neurons [6]. For ATP release from non-neuronal cells, a variety of transport mechanisms have been proposed, including but not limited to ATP-binding cassette transporters and connexin hemichannels, as well as vesicular release [5]. Extracellular ATP levels are controlled by both specific (ectonucleotidases) and nonspecific (ectoenzymes) enzymes with an extracellularly-oriented catalytic site [7]. Other ectoenzymes also contribute to interconversion of nucleotides, exerting control over the extracellular levels of various nucleotides under physiological and pathophysiological conditions [7]. Most studies examined ATP release after bulk equilibration in extracellular samples with the luciferin-luciferase assay (e.g., [6]). While highly sensitive, these measurements greatly underestimate the true [ATP] in the pericellular environment because of its rapid degradation by ectonucleotidases and delayed diffusion due to unstirred layer effects [8]. Recent studies have suggested that true levels of ATP at the plasma membrane are underestimated by >20-fold with bulk phase measurements [9]. This should be considered when relating the extracellularly measured concentrations of nucleotides to the effects on their receptors.

ATP and related purines interact with three families of purinergic receptors, the ionotropic P2X (7 receptors cloned) and metabotropic P2Y (8 receptors cloned) and P1 (4 receptors cloned) [8, 1012]. Given such tremendous diversity of receptors and natural ligands it is not surprising that ATP and related purines might be involved in a wide spectrum of physiological and pathophysiological activities, including nociception and chronic pain. For example, ATP is a known algogen whose application by iontophoresis to human skin elicits pain which is exaggerated by inflammation [13]. Application of ATP to dissociated neurons from rat sensory ganglia evokes inward currents via the P2X ion channels, mainly P2X2/3 purinoceptors (P2XRs) [14, 15]. Sensory neurons in culture form synapses that appear to utilize ATP as their neurotransmitter [16] and injury of skin cells near cultured neurons results in P2XR activation on identified nociceptors [17]. The P2X3Rs are especially concentrated in small diameter sensory neurons and their expression is differentially affected in injured and uninjured neurons after peripheral neuropathy induction [1820]. Increased P2XR-mediated signaling appears to contribute to sensory neuron hyperexcitability after spinal nerve ligation [21, 22], while functional down-regulation of P2X3Rs alleviates neuropathy symptoms [23]. The role of P2YRs in nociception and chronic pain is less clear. Several studies have suggested that ATP or uridine 5'-triphosphate (UTP) acting via P2Y2Rs increase the excitability and action potential discharge of sensory neurons, promote neuropeptide release and axonal transport, and facilitate increased expression of growth factors and neuropeptides by increasing the phosphorylation of cAMP responsive element binding protein [24]. However, there is also evidence that activation of P2Y1Rs on sensory neurons inhibits N-type voltage-gated Ca2+ channels (VGCCs) [25] and P2X3R channels [26], while intrathecal administration of preferential agonists for P2Y1 and P2Y2/4 receptor subtypes inhibits pain transmission [8, 26]. Aside from the reported increase in P2Y1R mRNA after sciatic nerve transection [27], little is known about the plasticity of P2YR signaling in neuropathic pain states.

Involvement of the P1 receptors in nociception and chronic pain is well-established. A1 and A3R activation usually results in antinociception and decreased neuropathic pain symptoms, whereas A2A and A2BR activation promotes nociception [10, 28]. These receptors respond selectively to extracellular adenosine produced from two sources: 1) bidirectional facilitated diffusion and 2) extracellular conversion of adenine nucleotides [10, 29]. The bidirectional transport of adenosine into and out of cells is dependent on the intra and extracellular levels of adenosine [30]. Blockade by transport inhibitors usually increases extracellular adenosine levels. Inhibition of cytosolic adenosine kinase also increases extracellular adenosine [10, 29]. Spinal nerve ligation enhances stimulus-evoked increases in extracellular adenosine [31]. Inhibitors of adenosine kinase appear to decrease pain symptoms of peripheral neuropathy in preclinical studies [32].

By contrast, neuropathy-induced changes in extracellular levels of ATP (an adenosine precursor), have not been addressed. We used extracellular sampling from an in vitro dorsal root ganglion (DRG) preparation [33] to assess possible changes in ATP release from DRG in a rat model of neuropathy induced by sciatic nerve entrapment [34] and examine possible mechanisms and consequences of such alterations. We found that basal extracellular [ATP] was increased while KCl-evoked ATP release was suppressed in the injured DRG. Selective blockade of A1Rs increased basal [ATP] and relieved the block of KCl-evoked ATP release. We also demonstrated that cellular metabolism was increased in the injured DRG, representing a potential source of neuropathy-induced increases in extracellular [ATP].

Materials and methods

All experimental procedures were carried out in accordance with the guidelines of the NIH on animal care and University of California at Los Angeles animal research committee.

Sciatic nerve entrapment (SNE)

Surgery was performed as described previously [34]. Adult male Sprague-Dawley rats weighing 200–250 g were used. Rats were anesthetized (pentobarbital 50 mg/kg, i.p.) or alternatively with a mixture of isoflurane (1–3%), N2O (50%), and O2 (inhalation) and the subsequent surgical procedures were performed under sterile conditions. The hair of the lower back and thigh of the rats was shaved and the skin was sterilized with povidone-iodine solution. A skin incision was made on one thigh and the sciatic nerve exposed through blunt dissection of the overlying muscle. Three Tygon® cuffs (length = 1 mm, outer diameter = 2.28 mm, inner diameter = 0.76 mm) were placed around the exposed sciatic nerve. This tubing size allowed them to be fitted snugly around the sciatic nerve without constricting it. The muscle layer was closed with absorbable sutures (5.0 Vicryl, Ethicon, Johnson & Johnson, Irvine, CA) and the skin was closed with suture (3.0 Nylon, Ethicon). Triple antibiotic cream (Bacitracin, Neomycin and Polymyxin B, Rite Aid, Harrisburg, PA) was applied over the wound site. Buprenorphine (0.01 mg/kg, s.c.) was injected once daily for 2 days for post-surgical analgesia. Sutures were removed 7–10 days post-surgery under isoflurane/N2O anesthesia.

Behavioral testing

Mechanical sensitivity was assessed using an electronic von Frey hair pressure transducer (IITC Instruments, Model 1601C, Woodland Hills, CA). Baseline behavioral testing was performed one day before and on the same day prior to surgery. Testing was repeated on post-operative days 1, 2, 3, 4, 5, 7, 9, 11, 13 and 15. The rat was gently placed in a plastic-walled cage (10 × 20 × 13 cm) with a metal mesh floor (0.6 × 0.6 cm holes). A point in the middle of the hindpaw was tested with the tip of the electronic von Frey hair pressed on the spot until the animal withdrew the foot. The force (g) applied at the time of withdrawal was recorded. Each hindpaw was tested three times at 1 min intervals and the results were averaged for each paw for that day.

Thermal sensitivity testing was performed using the Hargreaves paw withdrawal apparatus (Hargreaves Model 390, IITC Instruments) which measures the withdrawal latency from a radiant heat source directed at the proximal half of the plantar surface of each hindpaw. The baseline and post-operative testing schedule was the same as for mechanical sensitivity. Prior to testing, rats were allowed to acclimate to the testing environment for 15 min, which consisted of translucent plastic-walled individual chambers (10 × 20 × 20 cm) and a 3 mm thick glass bottom (preheated to 30°C). A radiant heat source consisting of an adjustable infrared lamp and a built-in stopwatch accurate to 0.1 s were used to measure paw withdrawal latency. Each paw was tested three times at 25% maximal heat intensity allowing 5 min between each test. The test was performed only when a rat was stationary and standing on all four paws. Special care was taken to keep the glass bottom clean and dry during the testing. If the glass needed to be cleaned during the experiment, the rats were allowed 5–10 min to reacclimatize to the environment. The results of three tests were averaged for each paw for that day.

DRG extraction and sample collection

At 15–21 days after SNE rats were anesthetized with sodium pentobarbital (50 mg/kg, i.p.) or alternatively with a mixture of isoflurane (1–3%), N2O (50%), and O2 (inhalation). The L4 and L5 DRG on both sides were excised with attached spinal nerves and dorsal roots, and placed in cold (0–4°C) low-Na+ artificial cerebrospinal fluid (ACSF) composed of (in mM): KCl 2.5, NaH2PO4 1.25, CaCl2 0.5, MgCl2 3.5, NaHCO3 26, glucose 10 and sucrose 248. The ACSF was continuously bubbled with a 95/5% mixture of O2/CO2 to ensure adequate oxygenation of ganglia and pH 7.4. The DRG were cleaned of surrounding connective tissue and incubated for 5 minutes at room temperature in low-Na+ artificial cerebrospinal fluid (ACSF) composed of (in mM): NaCl 63, KCl 2.5, NaH2PO4 1.25, CaCl2 0.5, MgCl2 3.5, NaHCO3 26, glucose 10 and sucrose 124 before being placed in the sample collection chamber where they were slowly (160 μl/min) perfused with normal ACSF (NaCl 125 and no sucrose) at 35.0 ± 0.5°C. In initial experiments, we used an open sample collection chamber (custom-made), where DRG could be monitored for physiological integrity using extracellular suction electrode recordings of evoked compound action potentials [33]. In subsequent experiments with adenosine receptor ligands, we switched to a commercial closed chamber (RC-30WA, Warner Instruments) for better flow and temperature control. High KCl-containing (100 mM) ACSF was applied after the baseline collection. For high K+ solutions, NaCl content was reduced to maintain ACSF osmolality. 8-Cyclopentyl-1,3-dipropylxanthine (DPCPX, 10 μM in 10% dimethylsulfoxide) and N6-cyclopentyladenosine (NCPA,10 μM in H2O) from frozen (-20°C) stock were diluted in ACSF just prior to perfusion. In these experiments, the comparison perfusate without DPCPX contained 0.03% dimethylsulfoxide.

Samples were collected at 1 min intervals into polypropylene tubes (0.2 ml; Corning Incorporated, Corning, NY), immediately frozen in liquid nitrogen and stored at -80°C until analysis. All experimental samples were analyzed within 3 days of collection.

Measurement of ATP release

Release of ATP was determined directly using the firefly luciferin-luciferase (L/L) assay. In initial experiments, L/L reagent (30 μL, CLS II, Boehringer Mannheim, Mannheim, Germany) was added to samples (40 μL) in a cylindrical cuvette, mixed and placed into a luminometer (TD-20/20, Turner Designs, Sunnyvale, CA). Chemiluminometer settings were delay: 1 sec, integration: 10 sec. In experiments with adenosine receptor ligands, samples were placed in closed-bottom 96-well white polystyrene plates (Corning Life Sciences, Lowell, MA) in a microplate reader (Synergy HT, Biotek, Winooski, VT) which automatically dispensed and mixed L/L solutions (ATP Bioluminescence kit HS II, Roche Diagnostics, Indianapolis, IN) to individual wells. Several concentrations of ATP standard (100 pM – 1 μM) with ACSF were measured before analysis of each experimental sample set. Separate ATP standards containing appropriate KCl and/or drug concentrations were also measured when appropriate. Microplate reader settings were delay: 0 sec, integration: 15 sec.

Measurement of O2 consumption

The L4 and L5 DRG were obtained from anesthetized rats as described above and the attached nerves were cut at 10 mm from each side of the DRG. After 15 min incubation in cold normal ACSF, both DRG were transferred into a closed chamber (Multi-Port Measurement Chamber; World Precision Instrument, Sarasota, FL) filled with 2.5 ml of N-2-hydroxyethylpiperazine-N'-2-ethanesulphonic acid (HEPES, 26 mM)-buffered ACSF. Oxygen consumption was measured with an oxygen probe (ISO2; World Precision Instruments, Sarasota, FL) which continuously measured dissolved oxygen in HEPES-buffed ACSF. Once the chamber temperature reached 34.5 ± 0.4°C, recording continued for 60 min and later analyzed off-line using the pCLAMP9 software (Molecular Devices, Union City, CA).

Statistical analysis

All data are presented as mean ± S.E.M. Statistical analyses (one-way repeated measures ANOVA with post hoc comparisons, one-way ANOVA and two-way ANOVA) were used to compare the data.


Hindpaw withdrawal thresholds after sciatic nerve entrapment (SNE)

SNE is a variation on chronic constriction injury in which the loose ligatures placed around the sciatic nerve are replaced by fixed-diameter polyethylene cuffs, resulting in decreased variability across animals [34]. The use of chemically inert polyethylene also prevents potentially confounding inflammatory responses that can occur in response to chromic catgut sutures. Stable baseline measurements were obtained 1 day pre-operatively and on the day just prior to the operation to both mechanical and thermal stimuli. Post-operatively (PO), measurements revealed that the SNE animals developed increased sensitivity of ipsilateral hindpaw to both types of stimuli (Fig. 1). For mechanical sensitivity, the baseline threshold for withdrawal was 40–45 g prior to surgery, and this was maintained PO for the contralateral paw and in naïve (untreated) rats (Fig. 1A). The threshold for the ipsilateral side dropped significantly to ~20 g by 2 days PO; a change that was maintained throughout the testing period. Significant decreases in latency to hindpaw withdrawal from a thermal stimulus developed by 3 days PO, and the change in latency was maintained throughout the testing period (Fig. 1B). There were also signs of spontaneous pain behaviors which included guarding behavior and changes in the posture of the affected hindpaw such as plantar flexion and toe-clenching, typical of this model [34].

Figure 1

Increased mechanical and thermal sensitivity after SNE induction. A: hindpaw withdrawal thresholds (mean ± S.E.M.) to mechanical stimuli on the ipsilateral side (n = 6 rats) are significantly decreased compared to contralateral side and naïve rats (n = 6) at 2 days after SNE surgery. Note that the decreases persist to the last day of measurements. B: hindpaw withdrawal latency (mean ± S.E.M.) from thermal stimuli on the ipsilateral side is also significantly reduced by the 3rd day after SNE. *, significant difference from other group means.

Basal ATP release from L4/L5 DRG after SNE

Sciatic nerve entrapment leads to axonal injury of neurons with cell bodies in both L4 and L5 DRG [34], prompting us to measure ATP release in both L4 and L5 DRG. In these initial studies, samples collected for the first few minutes of DRG perfusion had very high ATP content (not shown) and declined to stable levels by 12–15 min of perfusion. These initial high levels of ATP are probably due to mechanical deformation of the DRG preparation [35] when it was placed in the collection chamber and therefore are not included in the experimental analysis. In some of these experiments we also confirmed the physiological integrity of the DRG preparations by measuring evoked compound action potentials. The fitting of suction recording electrodes to spinal nerves and dorsal roots likely also contributed to the initial high ATP levels. After establishing a stable baseline, perfusion of naive rat L4 and L5 DRG (n = 20, 10 rats) with 100 mM KCl evoked a reversible increase in the amount of ATP (Fig. 2A). In a separate set of experiments, we could not detect KCl-evoked increases in ATP release from preparations containing either spinal nerves alone or dorsal roots alone (data not shown), indicating that DRG neuronal somata are required for KCl-evoked ATP release. Also, we demonstrated previously that KCl-evoked ATP release in vivo is dependent on calcium influx through voltage-gated calcium channels because it could be blocked by cadmium application [6]. Comparison of the in vitro experimental data indicated that baseline samples from ipsilateral DRG of SNE rats (n = 19, 11 rats) had significantly (p < 0.05) higher ATP levels than samples from control DRG. However, 100 mM KCl perfusion did not evoke ATP release in ipsilateral DRG as it did in naive rat DRG (Fig. 2B, D). By contrast, samples from contralateral DRG had basal levels of ATP similar to DRG from naive rats and exhibited robust increases in ATP content after 100 mM KCl perfusion (Fig. 2C, D).

Figure 2

Altered ATP release from DRG after SNE injury. A: ATP release from naive rat DRG is increased by KCl (100 mM) stimulation. * denotes p < 0.05 compared with basal ATP levels. Basal release was defined as the average of 4 samples obtained 1 min prior to onset of KCl stimulation. For clarity, samples 1–9 are not shown. B: Basal ATP release is higher than in naive DRG, whereas KCl stimulation no longer evokes ATP release from ipsilateral DRG. C: The pattern of ATP release from contralateral DRG is similar to naive DRG. * denotes p < 0.05 compared with basal ATP level. D: Summary graph of basal and evoked ATP release in DRG from naive and SNE neuropathic rats. Data were binned as illustrated in A-C. Note the increased basal and loss of KCl-evoked ATP release in ganglia ipsilateral to the SNE.

Effect of selective A1R activation on basal and KCl-evoked ATP release

The differential effects of SNE on basal and evoked ATP release suggested a possible involvement of ATP metabolites. Normally, extracellular ATP is regulated by the activity of various ectonucleotidases which convert ATP to di- and mono-phosphates and subsequently into adenosine. DRG neurons are endowed with A1Rs whose activation, in turn, leads to decreased voltage-gated Ca2+ channel (VGCC) activity [36]. Since KCl-evoked ATP release in sensory neurons is dependent on VGCC activation [6], we hypothesized that the block of evoked ATP release after SNE could result from the conversion of extracellular ATP to adenosine with subsequent activation of A1Rs on DRG neurons. To test this hypothesis we first had to demonstrate that A1R activation can lead to altered ATP release within DRG. To that end, we compared basal and KCl-evoked ATP release in DRG from naïve rats in the presence and absence of the selective A1R agonist, NCPA (100 nM). In the absence of NCPA (n = 6 DRG, 4 rats), KCl (100 mM) application reliably evoked a reversible increase in the sample ATP levels (Fig. 3A, B). However, in the continuous presence of NCPA (n = 7 DRG, 4 rats), there was a significant decrease in both basal and KCl-evoked ATP release (Fig. 3A, B), suggesting functional modulation of ATP release by A1R activation.

Figure 3

Selective A1 receptor activation decreases basal and KCl-evoked ATP release. A: Data are presented as mean ± SEM of sample ATP levels from naïve rat DRG in normal perfusate (white bars) and in the continuous presence of NCPA (100 nM, black bars). When DRG are perfused in the presence of NCPA (100 nM) both basal and evoked ATP release is visibly decreased. * denotes p < 0.05 compared with basal ATP levels. B: Summary graph of basal and evoked ATP release in DRG from naive rats with and without NCPA. Data were binned as illustrated in A.

Effect of selective A1R blockade on basal and KCl-evoked ATP release in DRG ipsilateral to SNE

We next tested our hypothesis by measuring basal and KCl-evoked ATP release in DRG from neuropathic rats in the presence and absence of the selective A1R antagonist, DPCPX. DRG ipsilateral to SNE exhibited higher basal ATP concentrations than contralateral DRG (compare Fig. 4A with 4C). Also, there was no significant KCl-evoked ATP release in ipsilateral DRG compared with the contralateral DRG, confirming our initial findings (see Fig. 2). As we suspected, in the presence of the selective A1R inhibitor, DPCPX (30 nM), both basal and KCl-evoked ATP release was significantly increased in DRG contralateral to SNE (compare Fig. 4A with 4B). Blockade of A1Rs in DRG ipsilateral to SNE further increased basal ATP levels compared to untreated ipsilateral SNE DRG (compare Fig. 4C with 4D). In support of our hypothesis, we observed a robust KCl-evoked release of ATP in DRG ipsilateral to SNE during A1R blockade (Fig. 4D and 4E).

Figure 4

Selective A1 receptor blockade increases basal ATP levels and relieves the blockade of KCl-evoked ATP release in DRG ipsilateral to SNE. A: Data are presented as mean ± SEM of sample ATP concentration from L4/L5 DRG contralateral to SNE before and after KCl (100 mM) application. B: Data from DRG contralateral to SNE during continuous presence of DPCPX (30 nM). C: Data from DRG ipsilateral to SNE before and after KCl (100 mM) application. Note the absence of KCl-evoked ATP release. D: Data from DRG ipsilateral to SNE during continuous presence of DPCPX (30 nM). E: Summary graph of basal and evoked ATP release in DRG from neuropathic rats in the presence and absence of DPCPX. Data were binned as illustrated in A-D. Note the DPCPX-induced increases in basal release and recovery of KCl-evoked release of ATP in DRG ipsilateral to SNE.

Altered metabolism after SNE

Increases in basal ATP release observed in ipsilateral L4/L5 DRG after SNE suggested possible changes in metabolism. We reasoned that increased ATP levels could arise from the demonstrated increases in excitability of both injured and spared sensory neurons [1], which in turn would increase the metabolic demand for ATP needed for the maintenance of ionic gradients. Increased neuronal excitability has been previously linked to increased metabolic activity in spinal cord neurons of normal rats [37], as well as after sciatic nerve ligation [38, 39]. However, metabolic changes within sensory ganglia were not examined.

To test whether oxidative metabolism is increased in DRG ipsilateral to injury, we adapted the in vitro DRG preparation for measurements of O2 consumption. Since O2 consumption is greatly influenced by factors such as temperature, etc., particular care was taken to ensure similarity of conditions for testing ganglia from different rats. After the ganglia were placed in the testing chamber, the one-hour recording began when temperature increased to a stable level of ≥ 34.5°C. The average temperatures during O2 consumption measurements of 34.66 ± 0.15°C (naive DRG) and 34.53 ± 0.20°C (DRG ipsilateral SNE) were not significantly different. DRG ipsilateral to SNE consumed O2 faster than control rat DRG (Fig. 5A). The rate of O2 consumption (expressed as the rate of negative redox current increase produced by the O2 probe) in DRG from naive rats (-24.42 ± 1.14 nA/hr) was significantly different (p = 0.005) from that of DRG ipsilateral to SNE (-33.18 ± 2.58 nA/hr) (Fig. 5A). In a separate set of experiments, no significant differences were found in the O2 consumption of DRG contralateral to SNE and naive rat DRG (data not shown).

Figure 5

Increased oxygen consumption of neuropathic DRG. A: the O2 reduction is described as the mean (thick lines) ± SEM (thin lines) redox current (nA) from naive (n = 8) and SNE (n = 6) ganglia. All data were adjusted to 0 nA at time point zero. The gradient of O2 reduction (consumption) was significantly faster in DRG ipsilateral to SNE compared to naive DRG (p < 0.05).


We measured nanomolar ATP levels with the luciferin-luciferase assay after bulk equilibration of perfusate samples with the DRG preparation, permitting relative comparisons of changes in basal and evoked ATP release after neuropathy induction. However, these measurements greatly underestimate the true ATP concentration in the pericellular environment of DRG either during stimulated or basal release, because of its rapid degradation by ectonucleotidases and delayed diffusion due to unstirred layer effects. Recent studies have suggested that true levels of ATP at the plasma membrane are underestimated by >20-fold with bulk phase measurements [9]. Thus, based on measured basal ATP levels in control DRG (Fig. 2A) we would expect true ATP concentration at the neuronal plasma membrane to be >10 nM. At this extracellular [ATP], some degree of P2X3 receptor desensitization would be expected [40, 41]. This might explain the paucity of ATP (1–10 mM) responsiveness via P2X3Rs in neurons within intact DRG preparations compared to the robust responsiveness in acutely dissociated neurons [42].

Several possible explanations could account for the increased basal ATP release from DRG ipsilateral to SNE. Previous studies have shown that peripheral denervation (axotomy) leads to progressive degeneration of ~35% of DRG neurons [43, 44]. Thus, it is conceivable that breakdown of the plasma membrane of degenerating neurons could lead to spillover of intracellular ATP into the extracellular space where it would contribute to the measured increases in ATP levels. However, after sciatic nerve constriction injury both myelinated and unmyelinated axons are intact proximal to the constriction site despite damage to distal axons, indicating the survival of the DRG neurons whose axons are interrupted [45]. Another possibility which remains to be examined is a neuropathy-induced decrease in the ectonucleotidase-mediated degradation of extracellular ATP [7]. A more plausible explanation might be that higher extracellular ATP levels result from increased excitability of both injured and uninjured neurons [1]. Increased neuronal excitability is clearly linked with increased metabolism [46], which leads to increases in intracellular ATP levels. In yeast [47], as well as mammalian pancreatic β-cells [4850] and adrenal chromaffin cells [51], increased metabolism and intracellular ATP have been linked with increased extracellular ATP release. Surprisingly, this relationship has not been addressed in central or peripheral neurons. However, we suspect that the hyperexcitability-induced increases in ATP production might result in increased vesicular [52] and non-vesicular release [5] of ATP from somata of DRG neurons.

Evidence in support of this explanation was obtained by demonstrating that O2 consumption was increased in L4/L5 ganglia ipsilateral to SNE compared to ganglia from control rats. Almost all of ATP is produced by mitochondria through aerobic respiration. Since O2 is not stored to any significant degree by tissues, O2 consumption is proportional to energy generation/metabolism. To our knowledge, this is the first time this method was used to examine metabolic changes in sensory ganglia. Previously, the [14C]-2-deoxyglucose (2-DG) method was used to study effects of peripheral stimulation in sensory ganglia, but only in normal rats [37]. In these studies, electrical stimulation produced a frequency-dependent increase of glucose utilization in the dorsal horn of the spinal cord, but not within the ipsilateral DRG, leading authors to suggest that axon terminals and not the cell bodies of sensory neurons are the sites of enhanced metabolic activity during increased functional activity of this pathway [37]. Later, spinal cord metabolism was investigated using the 2-DG method in the rat monoarthritis [53] and sciatic nerve ligation [38, 39] models. Interestingly, these studies demonstrated increased metabolic activity bilaterally in the spinal cord of arthritic and neuropathic rats, but apparently sensory ganglia were not examined [38, 39, 53].

Our studies do not differentiate between contributions of the various cell types to extracellular ATP release and metabolism of the entire DRG preparation. Thus, sympathetic fiber sprouting demonstrated by various groups to occur in the DRG in response to sciatic nerve injury, especially after axotomy and deafferentation, [54], could contribute to increased ATP levels via release from sympathetic nerve terminals. Furthermore, while neurons are expected to make a large contribution to extracellular ATP release and O2 metabolism, the involvement of other cell types is also likely. Satellite 'glial' cells which form a sheath around the somata of neurons in sensory ganglia undergo proliferation after axotomy or even mild skin abrasions [55]. It is therefore possible that proliferation of satellite cells after SNE contributes to the measured increases in extracellular ATP levels and the increases in O2 consumption.

One method used to study neuronal metabolism is the histochemical detection of cytochrome oxidase activity, which is tightly coupled to ATP production [56]. Using this method, chronic decreases in cytochrome oxidase activity were observed after axotomy of neurons within sensory ganglia [57, 58]. Loss of peripheral trophic factors and decreased depolarizing activity (action potential discharge) were proposed to contribute to decreased cytochrome oxidase activity in these cells [57]. These studies contrast with our data which indicate increased metabolism and basal ATP release in L4/L5 DRGs after SNE. They also contrast with the demonstrated hyperexcitability of neurons within axotomized DRG in electrophysiological studies [1, 5961]. One likely explanation for the differences is that axotomy results in the loss of a vast majority of the sensory neuron total cytoplasmic mass [62]. Those neurons which survive axotomy have a greatly decreased metabolic demand in maintaining a much shorter peripheral axon.

In addition to increased basal ATP levels, we no longer observed a KCl-evoked release of ATP after SNE. Previously, we used in vivo microdialysis in trigeminal ganglia to show that large increases in extracellular ATP levels result in the apparent loss of KCl-evoked ATP release [6]. Others have demonstrated in different tissues that increases in extracellular ATP lead to increased levels of its various metabolites, including adenosine [6365]. Also, the release of adenosine from primary sensory neurons is increased in the ipsilateral hindpaw after spinal nerve ligation [31]. Based on these data we reasoned that SNE-induced increases in extracellular ATP levels should also result in increased extracellular adenosine levels. KCl-evoked ATP release in vivo is dependent on the activation of voltage-gated Ca2+ channels (VGCCs) [6] and VGCCs are coupled to A1Rs in sensory neurons [36]. A1R activation leads to decreased Ca2+ influx through VGCCs and subsequent decreases in evoked transmitter release in these and other peripheral or CNS neurons [36, 66]. Therefore, we hypothesized that the differential SNE effects on basal and evoked ATP release could result from the conversion of extracellular ATP to adenosine with subsequent activation of A1Rs on DRG neurons. To test this hypothesis, we first demonstrated that selective neuronal A1R activation decreases both basal and evoked ATP release in DRG from naïve rats (Fig. 3). Then we demonstrated that selective A1R blockade was able to increase basal ATP levels in DRG both ipsilateral and contralateral to SNE, but more importantly relieved the blockade of KCl-evoked ATP release in DRG ipsilateral to SNE (Fig. 4). These data support our hypothesis that blockade of evoked ATP release is caused by increased A1R activation. An alternative that remains to be explored is that increased P2Y1 receptor activation also contributes to the neuropathy-induced blockade of KCl-evoked ATP release.

The findings of increased metabolism and ATP release in sensory ganglia after peripheral nerve injury have several implications for neuropathic pain mechanisms. Increased extracellular ATP may directly activate neighboring sensory afferents through P2XRs [14, 15] thereby increasing nociceptive drive to the CNS. ATP release and P2X-mediated signaling also participate in the enhancement of glutamatergic neurotransmission at the central terminals of primary sensory neurons [67, 68]. There is also evidence that peripheral increases in extracellular adenosine may contribute to hyperexcitability of human sensory axons via adenosine A2AR activation [69]. This may be tempered by increased activation of 'inhibitory' A1Rs at peripheral, spinal and supraspinal locations [28, 70, 71]. Increased activation of P2Y1Rs may also contribute to inhibition of neuronal hyperexcitability [24]. ATP may also potentiate TRPV1 activity, increase axonal transport and increase excitability via activation of P2Y2Rs which likely contributes to nociceptive signaling and hyperalgesia after peripheral nerve injury [7275]. Thus, SNE neuropathy-induced increases in ATP metabolism and release may contribute to enhanced generation and transmission of nociceptive signals at peripheral terminals, along the axons, within the DRG and at central terminals of primary sensory neurons.

In conclusion, we demonstrated that DRG neurons exhibit increased metabolic rates, increased extracellular ATP levels and increased A1R activation on the side of neuropathy induced by sciatic nerve entrapment. These data add to the literature supporting the use of selective purinoceptor ligands and purine metabolism inhibitors for the therapy of neuropathic pain symptoms [28, 76, 77].


  1. 1.

    Ma C, Shu Y, Zheng Z, Chen Y, Yao H, Greenquist KW, White FA, LaMotte RH: Similar electrophysiological changes in axotomized and neighboring intact dorsal root ganglion neurons. J Neurophysiol 2003, 89: 1588–1602.

    Article  PubMed  Google Scholar 

  2. 2.

    Devor M, Wall PD: Cross-excitation in dorsal root ganglia of nerve-injured and intact rats. J Neurophysiol 1990, 64: 1733–1746.

    CAS  PubMed  Google Scholar 

  3. 3.

    Amir R, Devor M: Chemically mediated cross-excitation in rat dorsal root ganglia. J Neurosci 1996, 16: 4733–4741.

    CAS  PubMed  Google Scholar 

  4. 4.

    Holton P: The liberation of adenosine triphosphate on antidromic stimulation of sensory nerves. J Physiol (Lond) 1959, 145: 494–504.

    CAS  Article  Google Scholar 

  5. 5.

    Burnstock G: Physiology and pathophysiology of purinergic neurotransmission. Physiol Rev 2007, 87: 659–797.

    CAS  Article  PubMed  Google Scholar 

  6. 6.

    Matsuka Y, Neubert JK, Maidment NT, Spigelman I: Concurrent release of ATP and SP within guinea pig trigeminal ganglia in vivo. Brain Res 2001, 915: 248–255.

    CAS  Article  PubMed  Google Scholar 

  7. 7.

    Zimmermann H: Ectonucleotidases in the nervous system. Novartis Found Symp 2006, 276: 113–28. discussion 128–30, 233–7, 275–81.:113–128.

    CAS  Article  PubMed  Google Scholar 

  8. 8.

    Abbracchio MP, Burnstock G, Boeynaems JM, Barnard EA, Boyer JL, Kennedy C, Knight GE, Fumagalli M, Gachet C, Jacobson KA, Weisman GA: International Union of Pharmacology LVIII: update on the P2Y G protein-coupled nucleotide receptors: from molecular mechanisms and pathophysiology to therapy. Pharmacol Rev 2006, 58: 281–341.

    PubMed Central  CAS  Article  PubMed  Google Scholar 

  9. 9.

    Joseph SM, Buchakjian MR, Dubyak GR: Colocalization of ATP release sites and ecto-ATPase activity at the extracellular surface of human astrocytes. J Biol Chem 2003, 278: 23331–23342.

    CAS  Article  PubMed  Google Scholar 

  10. 10.

    Ribeiro JA, Sebastiao AM, de Mendonca A: Adenosine receptors in the nervous system: pathophysiological implications. Prog Neurobiol 2003, 68(6):377–392.

    Article  Google Scholar 

  11. 11.

    Jarvis MF, Khakh BS: ATP-gated P2X cation-channels. Neuropharmacology 2008, 56(1):208–215.

    Article  PubMed  Google Scholar 

  12. 12.

    Khakh BS, North RA: P2X receptors as cell-surface ATP sensors in health and disease. Nature 2006, 442: 527–532.

    CAS  Article  PubMed  Google Scholar 

  13. 13.

    Hamilton SG, Warburton J, Bhattacharjee A, Ward J, McMahon SB: ATP in human skin elicits a dose-related pain response which is potentiated under conditions of hyperalgesia. Brain 2000, 123(Pt 6):1238–1246.

    Article  PubMed  Google Scholar 

  14. 14.

    Bland-Ward PA, Humphrey PP: P2X receptors mediate ATP-induced primary nociceptive neurone activation. J Auton Nerv Syst 2000, 81: 146–151.

    CAS  Article  PubMed  Google Scholar 

  15. 15.

    Hamilton SG, McMahon SB: ATP as a peripheral mediator of pain. J Auton Nerv Syst 2000, 81: 187–194.

    CAS  Article  PubMed  Google Scholar 

  16. 16.

    Zarei MM, Toro B, McCleskey EW: Purinergic synapses formed between rat sensory neurons in primary culture. Neuroscience 2004, 126: 195–201.

    CAS  Article  PubMed  Google Scholar 

  17. 17.

    Cook SP, McCleskey EW: Cell damage excites nociceptors through release of cytosolic ATP. Pain 2002, 95: 41–47.

    CAS  Article  PubMed  Google Scholar 

  18. 18.

    Eriksson J, Bongenhielm U, Kidd E, Matthews B, Fried K: Distribution of P2X3receptors in the rat trigeminal ganglion after inferior alveolar nerve injury. Neurosci Lett 1998, 254: 37–40.

    CAS  Article  PubMed  Google Scholar 

  19. 19.

    Novakovic SD, Kassotakis LC, Oglesby IB, Smith JA, Eglen RM, Ford AP, Hunter JC: Immunocytochemical localization of P2X3purinoceptors in sensory neurons in naive rats and following neuropathic injury. Pain 1999, 80: 273–282.

    CAS  Article  PubMed  Google Scholar 

  20. 20.

    Tsuzuki K, Kondo E, Fukuoka T, Yi D, Tsujino H, Sakagami M, Noguchi K: Differential regulation of P2X3mRNA expression by peripheral nerve injury in intact and injured neurons in the rat sensory ganglia. Pain 2001, 91: 351–360.

    CAS  Article  PubMed  Google Scholar 

  21. 21.

    Zhou J, Chung K, Chung JM: Development of purinergic sensitivity in sensory neurons after peripheral nerve injury in the rat. Brain Res 2001, 915: 161–169.

    CAS  Article  PubMed  Google Scholar 

  22. 22.

    Kim C, Chung JM, Chung K: Changes in the gene expression of six subtypes of P2X receptors in rat dorsal root ganglion after spinal nerve ligation. Neurosci Lett 2003, 337: 81–84.

    CAS  Article  PubMed  Google Scholar 

  23. 23.

    Barclay J, Patel S, Dorn G, Wotherspoon G, Moffatt S, Eunson L, Abdel'al S, Natt F, Hall J, Winter J, Bevan S, Wishart W, Fox A, Ganju P: Functional downregulation of P2X3receptor subunit in rat sensory neurons reveals a significant role in chronic neuropathic and inflammatory pain. J Neurosci 2002, 22: 8139–8147.

    CAS  PubMed  Google Scholar 

  24. 24.

    Gerevich Z, Illes P: P2Y receptors and pain transmission. Purinergic Signal 2004, 1: 3–10.

    PubMed Central  CAS  Article  PubMed  Google Scholar 

  25. 25.

    Gerevich Z, Borvendeg SJ, Schroder W, Franke H, Wirkner K, Norenberg W, Furst S, Gillen C, Illes P: Inhibition of N-type voltage-activated calcium channels in rat dorsal root ganglion neurons by P2Y receptors is a possible mechanism of ADP-induced analgesia. J Neurosci 2004, 24: 797–807.

    CAS  Article  PubMed  Google Scholar 

  26. 26.

    Gerevich Z, Zadori Z, Muller C, Wirkner K, Schroder W, Rubini P, Illes P: Metabotropic P2Y receptors inhibit P2X3 receptor-channels via G protein-dependent facilitation of their desensitization. Br J Pharmacol 2007, 151: 226–236.

    PubMed Central  CAS  Article  PubMed  Google Scholar 

  27. 27.

    Xiao HS, Huang QH, Zhang FX, Bao L, Lu YJ, Guo C, Yang L, Huang WJ, Fu G, Xu SH, Cheng XP, Yan Q, Zhu ZD, Zhang X, Chen Z, Han ZG, Zhang X: Identification of gene expression profile of dorsal root ganglion in the rat peripheral axotomy model of neuropathic pain. Proc Natl Acad Sci USA 2002, 99: 8360–8365.

    PubMed Central  CAS  Article  PubMed  Google Scholar 

  28. 28.

    Sawynok J: Adenosine and ATP receptors. Handb Exp Pharmacol 2007, 309–328.

    Google Scholar 

  29. 29.

    Sawynok J, Liu XJ: Adenosine in the spinal cord and periphery: release and regulation of pain. Prog Neurobiol 2003, 69: 313–340.

    CAS  Article  PubMed  Google Scholar 

  30. 30.

    Gu JG, Foga IO, Parkinson FE, Geiger JD: Involvement of bidirectional adenosine transporters in the release of L-[3H]adenosine from rat brain synaptosomal preparations. J Neurochem 1995, 64: 2105–2110.

    CAS  Article  PubMed  Google Scholar 

  31. 31.

    Liu XJ, White TD, Sawynok J: Enhanced release of adenosine in rat hind paw following spinal nerve ligation: involvement of capsaicin-sensitive sensory afferents. Neuroscience 2002, 114: 379–387.

    CAS  Article  PubMed  Google Scholar 

  32. 32.

    McGaraughty S, Cowart M, Jarvis MF, Berman RF: Anticonvulsant and antinociceptive actions of novel adenosine kinase inhibitors. Curr Top Med Chem 2005, 5: 43–58.

    CAS  Article  PubMed  Google Scholar 

  33. 33.

    Spigelman I, Gold MS, Light AR: Electrophysiological recording techniques in pain research. In Methods in Pain Research. Edited by: Kruger L. CRC Press, New York; 2001:147–168.

    Google Scholar 

  34. 34.

    Mosconi T, Kruger L: Fixed-diameter polyethylene cuffs applied to the rat sciatic nerve induce a painful neuropathy: Ultrastructural morphometric analysis of axonal alterations. Pain 1996, 64: 37–57.

    CAS  Article  PubMed  Google Scholar 

  35. 35.

    Bodin P, Burnstock G: Purinergic signalling: ATP release. Neurochem Res 2001, 26: 959–969.

    CAS  Article  PubMed  Google Scholar 

  36. 36.

    Dolphin AC, Forda SR, Scott RH: Calcium-dependent currents in cultured rat dorsal root ganglion neurones are inhibited by an adenosine analogue. J Physiol (Lond) 1986, 373: 47–61.

    CAS  Article  Google Scholar 

  37. 37.

    Kadekaro M, Crane AM, Sokoloff L: Differential effects of electrical stimulation of sciatic nerve on metabolic activity in spinal cord and dorsal root ganglion in the rat. Proc Natl Acad Sci USA 1985, 82: 6010–6013.

    PubMed Central  CAS  Article  PubMed  Google Scholar 

  38. 38.

    Price DD, Mao JR, Coghill RC, d'Avella D, Cicciarello R, Fiori MG, Mayer DJ, Hayes RL: Regional changes in spinal cord glucose metabolism in a rat model of painful neuropathy. Brain Res 1991, 564: 314–318.

    CAS  Article  PubMed  Google Scholar 

  39. 39.

    Mao J, Price DD, Coghill RC, Mayer DJ, Hayes RL: Spatial patterns of spinal cord [14C]-2-deoxyglucose metabolic activity in a rat model of painful peripheral mononeuropathy. Pain 1992, 50: 89–100.

    CAS  Article  PubMed  Google Scholar 

  40. 40.

    Sokolova E, Skorinkin A, Moiseev I, Agrachev A, Nistri A, Giniatullin R: Experimental and modeling studies of desensitization of P2X3 receptors. Mol Pharmacol 2006, 70: 373–382.

    CAS  PubMed  Google Scholar 

  41. 41.

    Pratt EB, Brink TS, Bergson P, Voigt MM, Cook SP: Use-dependent inhibition of P2X3 receptors by nanomolar agonist. J Neurosci 2005, 25: 7359–7365.

    CAS  Article  PubMed  Google Scholar 

  42. 42.

    Stebbing MJ, McLachlan EM, Sah P: Are there functional P2X receptors on cell bodies in intact dorsal root ganglia of rats? Neuroscience 1998, 86: 1235–1244.

    CAS  Article  PubMed  Google Scholar 

  43. 43.

    Janig W, McLachlan E: On the fate of sympathetic and sensory neurons projecting into a neuroma of the superficial peroneal nerve in the cat. J Comp Neurol 1984, 225: 302–311.

    CAS  Article  PubMed  Google Scholar 

  44. 44.

    McKay Hart A, Brannstrom T, Wiberg M, Terenghi G: Primary sensory neurons and satellite cells after peripheral axotomy in the adult rat: timecourse of cell death and elimination. Exp Brain Res 2002, 142: 308–318.

    Article  PubMed  Google Scholar 

  45. 45.

    Munger BL, Bennett GJ, Kajander KC: An experimental painful peripheral neuropathy due to nerve constriction. I. Axonal pathology in the sciatic nerve. Exp Neurol 1992, 118: 204–214.

    CAS  Article  PubMed  Google Scholar 

  46. 46.

    Sokoloff L, Reivich M, Kennedy C, Des Rosiers MH, Patlak CS, Pettigrew KD, Sakurada O, Shinohara M: The [14C]deoxyglucose method for the measurement of local cerebral glucose utilization: theory, procedure, and normal values in the conscious and anesthetized albino rat. J Neurochem 1977, 28: 897–916.

    CAS  Article  PubMed  Google Scholar 

  47. 47.

    Boyum R, Guidotti G: Glucose-dependent, cAMP-mediated ATP efflux from Saccharomyces cerevisiae. Microbiology 1997, 143(Pt 6):1901–1908.

    CAS  Article  PubMed  Google Scholar 

  48. 48.

    Kennedy HJ, Pouli AE, Ainscow EK, Jouaville LS, Rizzuto R, Rutter GA: Glucose generates sub-plasma membrane ATP microdomains in single islet beta-cells. Potential role for strategically located mitochondria. J Biol Chem 1999, 274: 13281–13291.

    CAS  Article  PubMed  Google Scholar 

  49. 49.

    Rapaport E: Involvement of elevated intracellular and extracellular ATP in the regulation of insulin secretion: therapeutic targets in non-insulin-dependent diabetes mellitus. Am J Ther 1995, 2: 283–289.

    Article  PubMed  Google Scholar 

  50. 50.

    Eliasson L, Renstrom E, Ding WG, Proks P, Rorsman P: Rapid ATP-dependent priming of secretory granules precedes Ca2+-induced exocytosis in mouse pancreatic B-cells. J Physiol 1997, 503(Pt 2):399–412.

    PubMed Central  CAS  Article  PubMed  Google Scholar 

  51. 51.

    Morita K, Ishii S, Uda H, Oka M: Requirement of ATP for exocytotic release of catecholamines from digitonin-permeabilized adrenal chromaffin cells. J Neurochem 1988, 50: 644–648.

    CAS  Article  PubMed  Google Scholar 

  52. 52.

    Huang L-YM, Neher E: Ca2+-dependent exocytosis in the somata of dorsal root ganglion neurons. Neuron 1996, 17: 135–145.

    CAS  Article  PubMed  Google Scholar 

  53. 53.

    Schadrack J, Neto FL, Ableitner A, Castro-Lopes JM, Willoch F, Bartenstein P, Zieglgansberger W, Tolle TR: Metabolic activity changes in the rat spinal cord during adjuvant monoarthritis. Neuroscience 1999, 94: 595–605.

    CAS  Article  PubMed  Google Scholar 

  54. 54.

    Lee BH, Yoon YW, Chung K, Chung JM: Comparison of sympathetic sprouting in sensory ganglia in three animal models of neuropathic pain. Exp Brain Res 1998, 120: 432–438.

    CAS  Article  PubMed  Google Scholar 

  55. 55.

    Hanani M: Satellite glial cells in sensory ganglia: from form to function. Brain Res Brain Res Rev 2005, 48: 457–476.

    CAS  Article  PubMed  Google Scholar 

  56. 56.

    Wong-Riley MT: Cytochrome oxidase: an endogenous metabolic marker for neuronal activity. Trends Neurosci 1989, 12: 94–101.

    CAS  Article  PubMed  Google Scholar 

  57. 57.

    Ichikawa H, Helke CJ: Cytochrome oxidase activity in vagal and glossopharyngeal visceral sensory neurons of the rat: effect of peripheral axotomy. Brain Res 1992, 578: 311–316.

    CAS  Article  PubMed  Google Scholar 

  58. 58.

    Naito Y, Naito E, Honjo I, Newman A, Honrubia V: Effect of vestibular nerve section on cytochrome oxidase activity in the vestibular ganglion cells of the squirrel monkey. Hear Res 1995, 90: 72–78.

    CAS  Article  PubMed  Google Scholar 

  59. 59.

    Abdulla FA, Smith PA: Axotomy- and autotomy-induced changes in the excitability of rat dorsal root ganglion neurons. J Neurophysiol 2001, 85: 630–643.

    CAS  PubMed  Google Scholar 

  60. 60.

    Liu CN, Michaelis M, Amir R, Devor M: Spinal nerve injury enhances subthreshold membrane potential oscillations in DRG neurons: relation to neuropathic pain. J Neurophysiol 2000, 84: 205–215.

    CAS  PubMed  Google Scholar 

  61. 61.

    Li Y, Dorsi MJ, Meyer RA, Belzberg AJ: Mechanical hyperalgesia after an L5 spinal nerve lesion in the rat is not dependent on input from injured nerve fibers. Pain 2000, 85: 493–502.

    CAS  Article  PubMed  Google Scholar 

  62. 62.

    Devor M: Unexplained peculiarities of the dorsal root ganglion. Pain 1999, (Suppl 6):S27-S35.

  63. 63.

    Chen Y, Shukla A, Namiki S, Insel PA, Junger WG: A putative osmoreceptor system that controls neutrophil function through the release of ATP, its conversion to adenosine, and activation of A2 adenosine and P2 receptors. J Leukoc Biol 2004, 76: 245–253.

    CAS  Article  PubMed  Google Scholar 

  64. 64.

    Westfall DP, Todorov LD, Mihaylova-Todorova ST: ATP as a cotransmitter in sympathetic nerves and its inactivation by releasable enzymes. J Pharmacol Exp Ther 2002, 303: 439–444.

    CAS  Article  PubMed  Google Scholar 

  65. 65.

    Mohri K, Takeuchi K, Shinozuka K, Bjur RA, Westfall DP: Simultaneous determination of nerve-induced adenine nucleotides and nucleosides released from rabbit pulmonary artery. Anal Biochem 1993, 210: 262–267.

    CAS  Article  PubMed  Google Scholar 

  66. 66.

    Hasuo H, Shoji S, Gallagher JP, Akasu T: Adenosine inhibits the synaptic potentials in rat septal nucleus neurons mediated through pre- and postsynaptic A1-adenosine receptors. Neurosci Res 1992, 13: 281–299.

    CAS  Article  PubMed  Google Scholar 

  67. 67.

    Gu JG, MacDermott AB: Activation of ATP P2X receptors elicits glutamate release from sensory neuron synapses. Nature 1997, 389: 749–753.

    CAS  Article  PubMed  Google Scholar 

  68. 68.

    Nakatsuka T, Tsuzuki K, Ling JX, Sonobe H, Gu JG: Distinct roles of P2X receptors in modulating glutamate release at different primary sensory synapses in rat spinal cord. J Neurophysiol 2003, 89: 3243–3252.

    CAS  Article  PubMed  Google Scholar 

  69. 69.

    Lang PM, Tracey DJ, Irnich D, Sippel W, Grafe P: Activation of adenosine and P2Y receptors by ATP in human peripheral nerve. Naunyn Schmiedebergs Arch Pharmacol 2002, 366: 449–457.

    CAS  Article  PubMed  Google Scholar 

  70. 70.

    Lao LJ, Kawasaki Y, Yang K, Fujita T, Kumamoto E: Modulation by adenosine of Aδ and C primary-afferent glutamatergic transmission in adult rat substantia gelatinosa neurons. Neuroscience 2004, 125: 221–231.

    CAS  Article  PubMed  Google Scholar 

  71. 71.

    Maione S, de NV, Cappellacci L, Palazzo E, Vita D, Luongo L, Stella L, Franchetti P, Marabese I, Rossi F, Grifantini M: The antinociceptive effect of 2-chloro-2'-C-methyl-N6-cyclopentyladenosine (2'-Me-CCPA), a highly selective adenosine A1 receptor agonist, in the rat. Pain 2007, 131: 281–292.

    CAS  Article  PubMed  Google Scholar 

  72. 72.

    Molliver DC, Cook SP, Carlsten JA, Wright DE, McCleskey EW: ATP and UTP excite sensory neurons and induce CREB phosphorylation through the metabotropic receptor, P2Y2. Eur J Neurosci 2002, 16: 1850–1860.

    Article  PubMed  Google Scholar 

  73. 73.

    Moriyama T, Iida T, Kobayashi K, Higashi T, Fukuoka T, Tsumura H, Leon C, Suzuki N, Inoue K, Gachet C, Noguchi K, Tominaga M: Possible involvement of P2Y2 metabotropic receptors in ATP-induced transient receptor potential vanilloid receptor 1-mediated thermal hypersensitivity. J Neurosci 2003, 23: 6058–6062.

    CAS  PubMed  Google Scholar 

  74. 74.

    Sakama R, Hiruma H, Kawakami T: Effects of extracellular ATP on axonal transport in cultured mouse dorsal root ganglion neurons. Neuroscience 2003, 121: 531–535.

    CAS  Article  PubMed  Google Scholar 

  75. 75.

    Stucky CL, Medler KA, Molliver DC: The P2Y agonist UTP activates cutaneous afferent fibers. Pain 2004, 109: 36–44.

    CAS  Article  PubMed  Google Scholar 

  76. 76.

    Donnelly-Roberts D, McGaraughty S, Shieh CC, Honore P, Jarvis MF: Painful purinergic receptors. J Pharmacol Exp Ther 2008, 324: 409–415.

    CAS  Article  PubMed  Google Scholar 

  77. 77.

    Burnstock G: Pathophysiology and therapeutic potential of purinergic signaling. Pharmacol Rev 2006, 58: 58–86.

    CAS  Article  PubMed  Google Scholar 

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We thank Aaron Fang, Yuki Takehara, Tatsuya Aiyama, Kana Morimoto, Hiroyuki Takada, Chihiro Yoshida, Bryan Tran and Dr. Supanigar Ruangsri for help with the experiments We also thank Drs. Lawrence Kruger and Baljit Khakh for the many helpful discussions and critique of this manuscript. This research was supported by the NIH grant DE14573 and the Whitehall Foundation.

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Correspondence to Igor Spigelman.

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The authors declare that they have no competing interests.

Authors' contributions

YM and IS conceived of, designed, and coordinated the study. YM, SM, BS and IS prepared SNE animals. TO performed behavioral testing and O2 consumption experiments. YM, TO, HI, SM, TC, KSO YYNL and BS performed the extracellular sampling and ATP content analysis. YM, TO and BS performed statistical analyses. YM, TO, HI, SM, BS and IS interpreted the results. IS wrote the manuscript. All authors read and approved the final manuscript.

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Matsuka, Y., Ono, T., Iwase, H. et al. Altered ATP release and metabolism in dorsal root ganglia of neuropathic rats. Mol Pain 4, 66 (2008).

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  • Dorsal Root Ganglion
  • Dorsal Root Ganglion Neuron
  • Spinal Nerve Ligation
  • Evoke Compound Action Potential
  • Ipsilateral Dorsal Root Ganglion