Lidocaine Enhances GαⁱProtein Function

Human A1R cDNA was subcloned into the pDoubleTrouble vector, and CHO-K1 cells were transfected as described previously, using the calcium phosphate precipitation method. 40Forty-eight hours after the transfection, cells were reseeded in Dulbecco's Modified Eagle Medium with nutrient mixture F12 (DMEM/F12 1:1) supplemented with 10% (v/v) fetal calf serum and with 2 mg/ml G-418. Transfected clones were selected by virtue of their viability in the neomycin analogue G-418 and carried in 0.6 mg/ml G-418. Expression of the receptors was verified by ligand binding and adenylate cyclase assays (see Adenylate Cyclase Assays section).

CHO-K1 cells stably transfected with the hA1R were routinely cultured in DMEM/F12, supplemented with 10% (v/v) fetal calf serum, penicillin (100 U), streptomycin (100 μg/ml), and 0.6 mg/ml G-418. Cells were grown at 37°C in a humidified 5% CO²/95% air atmosphere and subcultured two or three times weekly using trypsin (0.05% w/v)/ethylenediaminetetraacetic acid (EDTA) (0.02% w/v). 31 

For the measurement of hA1R-mediated cAMP accumulation, cells were grown as a confluent monolayer in 200-ml flasks. The culture medium was discarded, and the cells were washed once with phosphate-buffered saline (PBS) and then incubated in 5 ml PBS with 5 mm EDTA for 5–10 min. Dissociated cells were transferred to 30 ml PBS and centrifuged for 3 min at 1,000 rpm. The cell pellet was resuspended at a density of 100,000 cells/200 μl in N-(2-hydroxyethyl)piperazine-N-′(2-ethanesulfonic acid) (HEPES) (20 mm, pH 7.2), buffered DMEM/F12 containing 1 U/ml adenosine deaminase (ADA). Cells were allowed to recover for 30 min at room temperature. Drugs were added in a volume of 0.05 ml medium containing 1 U/ml ADA, the adenylate cyclase activator forskolin (final concentration, 10 μm), and, in some experiments, the phosphodiesterase inhibitor rolipram (final concentration, 10 μm). Incubation continued for 10 min in a 37°C shaking water bath, and then reactions were stopped by adding 0.5 ml 0.15 N HCl. Tubes were centrifuged for 10 min at 5,000 rpm and 0.5 ml was removed for measurement of cAMP levels, by the University of Virginia Diabetes Core Lab, using a radioimmunoassay method as described by Harper and Brooker. 41Cells used for cAMP determination had a viability of more than 95% as assessed by the exclusion of trypan blue.

Cells from at least 10 confluent dishes (20 cm in diameter) were washed twice with ice cold PBS, scraped into ice cold buffer A (10 mm HEPES, 10 mm EDTA, 0.1 mm benzamidine, 0.1 mm phenylmethylsulfonyl fluoride, and 2 μg/ml each of leupeptin, pepstatin A, and aprotinin, pH 7.4), and homogenized. The homogenate was centrifuged at 34,000 g  for 20 min at 4°C and then washed three times in ice cold buffer HE (10 mm HEPES, 1 mm EDTA, 0.02%[w/v] NaN³, 0.1 mm benzamidine, 0.1 mm phenylmethylsulfonyl fluoride, and 2 μg/ml each of leupeptin, pepstatin A, and aprotinin, pH 7.4). The pellet was resuspended in HE buffer plus 10% (w/v) sucrose at a concentration of 5 mg/ml, dounce homogenized, and stored in aliquots at −80°C. 31Protein concentration was determined by the Lowry method using bovine serum albumin as the standard and by linear regression.

Membrane preparations used for guanosine diphosphate (GDP)/guanosine triphosphate (GTP) γ³⁵S exchange assays were in addition treated with urea (i.e. , urea stripped membranes) to remove all endogenous G proteins of the CHO cells that would have interfered with the reconstitution of A1R with recombinant G protein subunits. Cells were treated exactly as described in the previous section, except that after the first centrifugation step, cells were resuspended in HE buffer with 7 M urea and incubated 30 min on ice followed by centrifugation at 142,000 g  for 30 min at 4°C. The pellet was then washed three times and finally resuspended in HE buffer plus 10% (w/v) sucrose. Aliquots at a concentration of 5 mg/ml were stored at −80°C.

The levels of expression of A1Rs in stably transfected CHO cells were determined in saturation binding experiments using the specific A1R antagonist [³H]-8-cyclopentylxanthine (CPX). Increasing concentrations of [³H]-CPX (0.1–10 nm) were incubated with 20 μg of membrane protein in a total volume of 100 μl HE buffer (10 mm HEPES, 1 mm EDTA, 0.02% NaN³, and 1 U/ml ADA, pH 7.4) for 90 min at room temperature. By using a Brandel cell harvester (Brandel, Gaithersburg, MD), membranes were collected onto Whatman GF/C glass fiber filters by washing them three times with ice cold binding buffer (10 mm Tris and 5 mm MgCl², pH 7.4). Radioactivity trapped on filters was measured by using a scintillation counter (Beckman LS6500; Beckman Coulter, Inc., Fullerton, CA). Nonspecific binding was measured in the presence of 100 μm 8-cyclopentyltheophylline (CPT). All reactions were performed in triplicate. Specific binding was fit to a single-site binding model using nonlinear curve fitting to calculate receptor density expressed as the maximum binding capacity (B^max) and the dissociation constant K^d(Prism 3.0; GraphPad Software, San Diego, CA). 31 

To investigate interactions of lidocaine with agonist or antagonist binding, competition binding experiments were performed. To determine interactions with specific binding of the antagonist [³H]-CPX, membrane protein aliquots (20 μg) were incubated with various concentrations of lidocaine (10⁻⁹–10⁻²M) and with [³H]-CPX (2 nm) in a total volume of 100 μl HE buffer (10 mm HEPES, 1 mm EDTA, 0.02% NaN³, and 1 U/ml ADA, pH 7.4) for 90 min at room temperature. Radioligand binding was determined as described in the previous section, by using a cell harvester, glass fiber filters and by measuring radioactivity in a scintillation counter. To determine interactions with specific binding of the agonist [¹²⁵I]-N⁶-aminobenzyladenosine ([¹²⁵I]-ABA), 20 μg of membrane protein was incubated with various concentrations of lidocaine (10⁻¹⁰–10⁻²M) and with [¹²⁵I]-ABA [1 nm] in HE buffer (10 mm HEPES, 1 mm EDTA, 10 mm MgCl², 0.02% NaN³, and 1 U/ml ADA, pH 7.4) for 180 min at room temperature. Radioligand binding was again determined as described in the preceding section for [³H]-CPX, except that [¹²⁵I]-ABA was counted dry in a gamma counter.

Gαⁱand βγ subunits were expressed using the baculovirus/Sf9 insect cell system and purified to homogeneity as described previously. 42,43 

Urea stripped membrane protein (approximately 200 fmol measured as [³H]-CPX binding sites) was centrifuged for 10 min at 12,000 rpm in a microcentrifuge at 4°C. The pellet was resuspended in 450 μl reconstitution buffer (25 mm HEPES, 100 mm NaCl, 1 mm EDTA, 5 mm MgCl², 0.1% bovine serum albumin, 1 μm adenosine-5′-[(β,γ)-imido]triphosphate, 1 mm dithiothreitol, and 1 U/ml ADA) and incubated 30 min on ice. After 30 min, Gαⁱ¹and β^1γ2dimers and GDP were added, so that the final concentrations in a volume of 500 μl were as follows: 20 nm Gαⁱ¹, 40 nm β^1γ2, 500 nm GDP. The receptor–G protein mixture was incubated another 30 min on ice, followed by a 10-min equilibration at 25°C. The assay was started by the addition of 50 μl reconstitution buffer containing GTP γ³⁵S (final concentration, 7 nm). The ligand R-N⁶-phenylisopropyladenosine (final concentration, 100 nm) was added 8 min later (t = 0). The reaction was stopped by removing membrane aliquots for filtration through nitrocellulose filters (HAWP-025; Millipore, Billerica, MA) at t = 2, 4, 6, and 7.5 min. The wash buffer consisted of 10 mm Tris and 5 mm MgCl²(pH 7.4). Radioactivity trapped on filters was measured using a scintillation counter (Beckman LS6500). 44 

Results are reported as mean ± SEM. Student t  test was used to determine statistically significant differences between paired groups, with P < 0.05 considered significant. Concentration–response curves were fitted to the following function, derived from the Hill equation: y = y^min+ (y^max− y^min)[1 − xⁿ/(x⁵⁰ⁿ+ xⁿ)], in which y^maxand y^minare the maximal and minimal response obtained; n is the Hill coefficient; and x⁵⁰is the half-maximal effect concentration (i.e. , EC⁵⁰for an agonist and IC⁵⁰for an antagonist). The slopes of the agonist-induced GTPγS binding were calculated by using linear regression (Prism 3.0).

Molecular biology reagents were purchased from Promega (Madison, WI), and other chemical reagents were obtained from Sigma (St. Louis, MO). Cell culture reagents were from Gibco Life Technologies, Inc. (Grand Island, NY), and radioligands were from NEN Life Sciences Products (Boston, MA). Stock solutions of lidocaine hydrochloride were freshly prepared in the required assay buffer and then diluted to the final concentrations.

Nontransfected CHO cells lack endogenous adenosine receptors as determined by two criteria. First, A1R binding in control cells was indistinguishable from nonspecific binding (data not shown). Second, no A1R-mediated inhibition of adenylate cyclase was observed in control cells (data not shown).

To use CHO cells stably expressing hA1R as a suitable in vitro  test system for LA effects on adenosine signaling, two requirements were necessary: A1Rs had to be expressed in sufficient numbers, and they had to be functional. We first determined the density of hA1R expressed in transfected CHO cells by radioligand binding studies using the specific antagonist [³H]-CPX. In a range from 0.1 to 10 nm, free drug specific binding was saturable. The saturation curve and Scatchard analysis (fig. 2A) conform closely to a single-site model with a dissociation constant K^dof 2.42 ± 0.21 nm and a B^maxof 2,755 ± 91 fmol/mg protein (n = 5), which confirmed sufficient expression. We then addressed the question whether transfected hA1 receptors were functionally active. If so, binding of an agonist should result in a decrease in cAMP (fig. 1). As shown in figure 2B, N⁶-cyclopentyladenosine (CPA) (10⁻¹²–10⁻⁵M), an hA1R agonist, decreased cAMP formation in a concentration-dependent manner. Fitting to the Hill equation yielded an EC⁵⁰of 2.8 × 10⁻¹⁰± 0.68 × 10⁻¹⁰M. Maximal cAMP reduction by 88.0 ± 1.36% of baseline was obtained with a concentration of 1 × 10⁻⁶M. Next we showed that addition of CPX (1 μm), a competitive antagonist, shifts the concentration response curve significantly to the right. EC⁵⁰for CPA in the presence of CPX was almost 1,000-fold higher (1.06 × 10⁻⁷± 0.83 × 10⁻⁷M) (fig. 2B). Thus, CHO cells possess the appropriate Gαⁱprotein to couple to the hA1R and use adenylate cyclase as an effector. These results were in accordance with previous findings. 29–32,35 

To determine whether lidocaine had any effect on the entire A1R-coupled signaling pathway, we used cAMP assays. Even in the absence of an agonist, lidocaine (10⁻⁵–10⁻¹M) reduced cAMP levels in a concentration-dependent manner with a calculated EC⁵⁰of 6.60 × 10⁻⁴± 3.24 × 10⁻⁴M (fig. 3A). Possible explanations of this effect may be that lidocaine acts as an agonist at the ligand binding site of the hA1 receptor or that it activates the pathway further downstream at the receptor itself or somewhere within the signaling cascade.

To characterize the underlying mechanism by which lidocaine enhanced hA1R-coupled signaling, we studied the inhibition of cAMP production by increasing concentrations of the hA1R agonist CPA (10⁻¹¹–10⁻⁷M) in the absence and presence of lidocaine at approximately EC⁵⁰(1 mm) (fig. 3B). Lidocaine significantly reduced cAMP production to approximately 50% of baseline (P < 0.001) in the absence of CPA and in the presence of CPA (10⁻¹¹–10⁻¹⁰M). Under control conditions (in the absence of lidocaine), cAMP concentrations range from around 10–20 pmol/ml at CPA concentrations from 10⁻¹¹–10⁻¹⁰M, and they were reduced significantly to 5–10 pmol/ml in the presence of lidocaine (at the same CPA concentrations). This shows the additional effect of lidocaine on cAMP reduction and therefore its ability to potentiate A1R signaling. At CPA concentrations greater than 10⁻⁹M, no additional effect of lidocaine was obtained, most likely because of the already low level of cAMP formation. The calculated EC^50’s for CPA (1.12 × 10⁻¹⁰± 0.22 × 10⁻¹⁰M in the absence of lidocaine and 1.58 × 10⁻¹⁰± 0.34 × 10⁻¹⁰M in the presence of lidocaine) did not differ significantly. In addition, lidocaine maintained its inhibitory effect on cAMP formation even in the presence of the A1R antagonist CPX, and at similar EC⁵⁰(11.15 × 10⁻⁴± 6.17 × 10⁻⁴M) (fig. 3A). Together, these findings suggest a site of action for lidocaine separate from the agonist binding site.

We confirmed this finding using binding studies. Because agonist and antagonist binding kinetics differ, we measured binding of the antagonist [³H]-CPX or the agonist [¹²⁵I]-ABA at their dissociation constant K^d, in the presence of increasing concentrations of lidocaine (10⁻¹⁰–10⁻²M). Consistent with the functional data, lidocaine affected neither antagonist (fig. 4A) nor agonist (fig. 4B) binding to the hA1R significantly. At the functionally determined EC⁵⁰(6.60 × 10⁻⁴± 3.24 × 10⁻⁴M), only insignificant inhibition of binding occurred, which cannot explain the highly significant reduction of the cAMP levels. If anything, activation of hA1R signaling at the receptor level should have become evident in an increased binding of an agonist to the receptor rather than a decreased binding. These results make an interaction of lidocaine with the agonist–antagonist binding site highly unlikely. The site of action for lidocaine within the Gαⁱ-coupled signaling pathway therefore must be somewhere else at the receptor level or further downstream in the signaling cascade.

We next assessed lidocaine interference with A1R coupling to the Gαⁱprotein, GDP/GTP exchange at the Gαⁱsubunit, and interaction between Gαⁱand the β^1γ2dimers. For these experiments, the hA1R was reconstituted with pure Gαⁱ¹and β^1γ2subunits, and agonist-stimulated GTP γ³⁵S binding was measured as described in the Materials and Methods section. The data illustrated in figure 5support the ability of Gαⁱ¹and β^1γ2subunits to couple to the hA1 receptor, as indicated by the marked change in the rate of GTP γ³⁵S bound after addition of 100 nm of the specific A1R agonist R-N⁶-phenylisopropyladenosine. Because lidocaine (1 mm) did not significantly alter the slopes of the curves of unstimulated or R-N⁶-phenylisopropyladenosine–stimulated GTP γS binding (fig. 5), we concluded that LAs do not affect receptor–Gαⁱprotein coupling, GDP/GTP exchange, or the interaction between Gαⁱand β^1γ2subunits.

By adding the phosphodiesterase inhibitor rolipram (10 μm) to cAMP assays, we examined whether the significant reduction of cAMP levels by lidocaine was because of a stimulating effect on phosphodiesterase, which would lead to an accelerated breakdown of cAMP. As expected, cAMP levels were increased in the presence of the phosphodiesterase inhibitor. However, rolipram had no effect on the lidocaine-induced cAMP reduction, leaving its concentration–response curve (fig. 3A) and the EC⁵⁰(data not shown) unaffected. If lidocaine's cAMP reducing effects would have been caused completely or partly by stimulation of phosphodiesterase, this effect should have been abolished or reduced after the addition of rolipram. Thus, we can exclude phosphodiesterases, or at least those inhibited by the phosphodiesterase inhibitor rolipram, as a primary site of action for lidocaine.

In contrast, inhibition of cAMP formation by lidocaine was completely abolished after 24 h incubation of transfected CHO cells with PTX (1 μm). PTX inactivates the Gαⁱprotein and excludes a site of action for lidocaine downstream of the G protein in the signaling pathway. Again, the addition of rolipram only increased cAMP levels, but did not have any additional effect on the action of lidocaine (fig. 6). These data therefore indicate that stimulation of Gαⁱfunction is the most likely cause of lidocaine-induced decreases in cAMP levels.

As part of our program investigating LA interactions with G protein-coupled receptor signaling, we investigated lidocaine effects on Gαⁱprotein function by using the hA1R signaling cascade as a model. In contrast to previous studies showing inhibitory actions of LAs on Gα^q, lidocaine potentiates the hA1R signaling cascade, most likely by enhancing Gαⁱprotein function. Lidocaine did not significantly interact with the ligand binding site nor did the LAs affect the signaling pathway downstream of the Gαⁱprotein.

Agonist and antagonist binding were inhibited only marginally by high concentrations of lidocaine, making the ligand binding pocket as target site for the LAs unlikely. Nevertheless, the receptor as an additional target site for LA action cannot be excluded completely because lidocaine may bind to extracellular receptor domains other than agonist and antagonist binding sites and hence allosterically activate A1R signaling. However, one would expect this to result in altered receptor–G protein coupling. Because lidocaine did not have any effect on receptor–G protein interaction, as shown by the GDP/GTP exchange assays, this possibility appears less likely.

It is noteworthy that enhancement of Gαⁱfunction did not require the presence of an agonist (fig. 3). Endogenous adenosine cannot have acted as an agonist, because all endogenous CHO adenosine should have been inactivated by ADA. Lidocaine even maintained its cAMP-reducing effect in the presence of an A1R antagonist (fig. 3B). Taken together, these findings suggest that LA action does not require the A1R. This hypothesis is supported by a study by Hirota et al. , 34who found that lidocaine produced a concentration-dependent and naloxone-insensitive inhibition of cAMP formation in nontransfected CHO cells. The EC⁵⁰was 2.78 mm (1.63–4.73 mm), similar to the EC⁵⁰found by us (660 μm–1.6 mm) (fig. 3A). However, Hirota et al.  did not investigate the mechanism underlying this finding. Our results support the hypothesis that lidocaine interacts directly with the Gαⁱprotein, thereby leading to reduced cAMP formation. As to the exact mechanism by which LAs enhance Gαⁱfunction, several possible explanations can be ruled out by the GDP/GTP exchange assay results: First, LAs did not interfere with receptor–G protein coupling, neither by increasing the affinity of the G protein for the receptor, nor by enhancing uncoupling of the G protein from the receptor. Second, LAs did not affect GDP/GTP exchange at the Gαⁱsubunit. A direct activation of GDP-bound Gαⁱby lidocaine, for which the GDP/GTP exchange would be a prerequisite, can therefore essentially be excluded or is highly unlikely. Third, LAs did not affect interaction between Gαⁱand the β^1γ2dimers. Instead, the results suggest that lidocaine interacts with the pool of already activated Gαⁱpresent in the cytoplasm and thereby facilitates its ability to inhibit adenylate cyclase. At least two potential explanations for this facilitation of Gαⁱactivity may exist. First, LAs might inhibit Gαⁱ-related GTPase activity. Inhibition of GTPase activity could be a direct effect of LAs or could be the result of inhibitory action of LA on GTPase-activating proteins. Second, LAs might stabilize the Gαⁱ–adenylate cyclase complex. Both mechanisms would prolong the half-life of the activated Gαⁱ–GTP complex, resulting in reduced cAMP formation. Future experiments in isolated systems such as phospholipid vesicles, allowing reconstitution of the separate components being involved, including the various adenylate cyclase isoforms, will be necessary to define more precisely the interaction sites and the exact molecular mechanism. A direct inhibitory effect of lidocaine on adenylate cyclase can be excluded because PTX completely abolished the effect of lidocaine. The activated Gαⁱpresent in the cytoplasm, even in the absence of an agonist, could derive from at least two sources. First, G protein–coupled receptors have classically been viewed to have an inactive conformation (R) that requires an agonist-induced conformational change for receptor–G protein coupling and, thus, G protein activation to occur. New evidence suggests, however, a more complex two (or more)-state model, in which receptors are in equilibrium between the inactive conformation (R) and a spontaneously active state (R*). Classic agonists serve as catalysts and increase the concentration of R*. An equilibrium between R and R*, however, implies that at any time a certain proportion of receptors are in the R* state that can couple to the G protein, even in the absence of a ligand, thus resulting in release of Gα^I–GTP into the cytosol. Second, GDP/GTP exchange at the Gαⁱsubunit occurs to some extent, even in the absence of receptor stimulation. This spontaneous GDP release is reflected in our GDP/GTP γS exchange assays as the basal rate (fig. 5). 45,46 

Potentiation of A1 adenosine signaling by lidocaine independent of an agonist and receptor inevitably raises two questions: Does lidocaine also enhance hA1R signaling with an effector other than adenylate cyclase? Are other Gαⁱ-coupled receptor systems affected in the same way? A1Rs were originally characterized on the basis of their ability to inhibit adenylate cyclase via  the Gαⁱprotein (fig. 1). Meanwhile, a number of other PTX-sensitive effector mechanisms of A1Rs have been discovered, including increases in K⁺-channel conductance, decreases in Ca²⁺-channel conductance, and increases or decreases in phospholipase C activity. 33,39Testing the effects of LA on these effector systems would help to further clarify the mechanism by which LAs enhance Gαⁱfunction and whether it depends on cAMP. If further studies show that these effectors are also affected by lidocaine in a PTX-sensitive, agonist-independent manner, this would tremendously increase the number of potentially clinically relevant signaling pathways that are affected by LAs. A great number of receptors signal via  Gαⁱproteins (e.g. , α²-adrenergic, dopaminergic (D2, D3, D4), m2 and m4 muscarinic, GABA^B, and metabotropic glutamate receptors). 47,48If lidocaine affects other Gαⁱ-coupled signaling cascades in the same way as A1 signaling, a number of relevant pathways may be affected.

We used lidocaine as a prototypical compound; it is conceivable that other compounds might yield different results. We also realize that our findings and those of others 34might be biased by the in vitro  test system used (CHO cells). However, lidocaine and other LAs (e.g. , bupivacaine, mepivacaine, and ropivacaine) have all been found (albeit with varying potencies) to reduce cAMP levels in more physiologic preparations, such as cardiomyocytes, 13,49erythrocytes, 50and lymphocytes. 12,51However, the underlying molecular mechanisms were not investigated in detail or remained unclear. Some of the studies suggested the receptor as the target, others the adenylate cyclase system. For example, Roux et al.  13reported that the inhibitory effect of lidocaine (up to 10 mm) on isoproterenol- and forskolin-stimulated cAMP was reduced by 25% in the presence of cholera toxin, but it was completely abolished in the presence of PTX. The authors hypothesized that “lidocaine's interaction with the adenylate cyclase system” could contribute to the antiarrhythmic and the negative inotropic effects in vivo . However the mechanism underlying this interaction with the adenylate cyclase system was not further investigated. This issue may partly be associated with the heterogeneous background of these native cell preparations; for example, cAMP reduction might be the result of inhibition of Gα^sor enhancement of Gαⁱ-coupled signaling. Nevertheless, these observations in other models make it unlikely that our results simply represent an artifact of the transfection system or a unique property of lidocaine. In any case, additional experiments in various other physiologic preparations will be necessary to further define our findings and to test whether the data obtained with lidocaine can be extrapolated to other LAs and to other tissues.

The LA concentrations required to affect G protein function may appear to be relatively high. However, it is important to keep in mind that the concentrations of LA used in clinical practice vary widely depending on the method of application. 52After intravenous administration and similarly after epidural anesthesia, plasma concentrations in the low micromolar range are obtained. 53,54In contrast, LA concentrations routinely observed after topical application or tissue infiltration are approximately 1,000-fold greater. Similarly, millimolar LA concentrations are also present around the spinal nerves after spinal or epidural administration of an LA. 1,2,55,56Therefore, concentrations tested in our study are well within the range of those obtained in vivo . Interestingly, most in vitro  studies require much greater LA concentrations than in vivo  studies, in which the same or similar phenomenon often occurs at much lower concentrations. 2Although not fully understood, one reason for this discrepancy may well be that the multitude of molecular targets present in a complex in vivo  system allows interactions of diverse LA effects, which cannot be attained in a simplified in vitro  system. Thus, the previously mentioned cAMP-reducing effects of various LAs in cardiomyocytes 13,49or lymphocytes 12,51may represent the result of blockade of β-adrenergic receptors, and enhancement of Gαⁱfunction and even inhibition of Gα^s.

Adenosine administered systemically or injected spinally has been shown to exert long-lasting and significant antinociception under conditions of neuropathic pain, with no effect on acute pain. 57,58By using spinal nerve ligation as a model for neuropathic pain, it was reported that adenosine release in the spinal cord is stimulated and results in increased G protein activity. 59Our findings that lidocaine facilitates Gαⁱfunction thus may in part provide the molecular basis for the clinical application of lidocaine therapy in chronic pain states associated with hyperalgesia and allodynia. 60The combination of increased adenosine levels in addition to lidocaine's action might result in cAMP concentrations sufficiently low to provide antinociception.

In summary, our study shows that lidocaine interacts with hA1R-coupled signaling, most likely by facilitating Gαⁱprotein function. Expanding on previous findings, our data suggest that LA action depends on the specific G protein subunit affected and further support the hypothesis that specific G proteins represent an alternative target site of LA, separate from their well known actions on the Na⁺-channel.

The authors thank Heidi Figler, B.S. (Department of Internal Medicine, University of Virginia, Charlottesville, Virginia), for her excellent technical assistance and Dr. Norbert Roewer (Department of Anesthesiology, University of Wuerz-burg, Wuerzburg, Germany) for his support.