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Key Words: adenosine triphosphate • pharmacology; brown fat • physiology; electrophysiology; potassium channels • physiology; receptors, purinergic • metabolism
Abbreviations: HP, holding potential; IKv, voltage-dependent K+ current
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; Russ et al., 1993). Brown adipocyte IKv is similar to delayed rectifier potassium currents in other nonexcitable cells. IKv activates rapidly with depolarizations positive to approximately –35 mV, inactivates slowly, and deactivates rapidly when the cell is returned to the holding potential (Lucero and Pappone, 1989). The current is blocked by quinine, verapamil, 4-aminopyridine, and high tetraethyl ammonium concentrations, but not by apamin or charybdotoxin (Lucero and Pappone, 1989; Pappone and Ortiz-Miranda, 1993).
Sympathetic nerve stimulation potently regulates brown fat cell function. Acute adrenergic stimulation activates membrane conductances, elevates cytosolic calcium, and increases thermogenesis by activating lipolysis and mitochondrial uncoupling (Nedergaard and Lindberg, 1982; Girardier and Schneider-Picard, 1983; Horowitz et al., 1983; Lee et al., 1993). Chronic sympathetic stimulation promotes hyperplasia, hypertrophy, and differentiation of brown fat, resulting in increased thermogenic capacity (Geloen et al., 1988; Himms-Hagen, 1989; Bronnikov et al., 1992). Norepinephrine released by sympathetic nerve activity stimulates both 
- and β-adrenergic receptors to depolarize brown adipocytes, potentially activating IKv. -Adrenergic stimulation activates a depolarizing Ca2+-dependent Cl– conductance (ICl,Ca) (Dasso et al., 1990; Pappone and Lee, 1995). β-Adrenergic stimulation activates a nonselective cation conductance (Horwitz et al., 1989; Lucero and Pappone, 1990). In addition, cell swelling activates a second depolarizing Cl– conductance (Pappone and Lee, 1995). These sympathetically induced membrane depolarization events could activate IKv.
IKv is important for brown adipocyte function. IKv is necessary for proliferation of brown adipocytes in culture (Pappone and Ortiz-Miranda, 1993), but not for norepinephrine-activated heat production (Pappone and Lucero, 1992). The mechanism of IKv's effects on brown adipocyte growth are not known. Possibly sympathetic activation of IKv is important for cell volume regulation or Ca2+ responses required for cell proliferation. Simultaneous sympathetic activation of ICl,Ca and IKv in brown fat cells could result in loss of KCl from the cell and decrease cell volume, as in lymphocytes, epithelial cells, and hepatocytes (Cahalan and Lewis, 1988; Felipe et al., 1993; Strange, 1994; Graf and Haussinger, 1996). IKv activation could also facilitate Ca2+ entry into the brown adipocyte by increasing the Ca2+ electrochemical gradient, akin to the mechanism described in lymphocytes (Lin et al., 1993).
The effects of adrenergic stimulation in vitro do not fully explain the effects that cold exposure or overfeeding have on brown adipose tissue in vivo (Geloen et al., 1988; Yamashita et al., 1994), suggesting that additional agents may contribute to sympathetic actions. Norepinephrine and ATP are colocalized within sympathetic nerve terminals and may be released simultaneously with sympathetic neuronal activity (Lagercrantz, 1976; Westfall et al., 1990). Moreover, brown adipocytes respond to extracellular ATP. P2 receptor stimulation increases cytosolic Ca2+ and increases membrane turnover and membrane conductances, but does not alter norepinephrine-elicited thermogenesis (Pappone and Lee, 1996; Lee and Pappone, 1997a).
Channel activity in other cell types can be modulated by extracellular ATP. P2 receptor stimulation decreases the activity of voltage-gated K+ currents in hippocampal neurons (Nakazawa and Inoue, 1994) and enhances delayed rectifier K+ current activity in atrial myocytes (Matsuura et al., 1996b; Matsuura and Ehara, 1997). Structurally related voltage-gated Ca2+ currents are inhibited by purinergic stimulation in ventricular myocytes (Qu et al., 1993a,b; Von zur Muhlen et al., 1997) and potentiated in hippocampal neurons (Dave and Mogul, 1996). We therefore examined whether extracellular ATP affects IKv in rat brown adipocytes. We find that brown adipocyte IKv has two components with different gating properties. One component (IKv-A) is activated at physiological membrane potentials, while the other (IKv-B) is inactive under normal conditions. Extracellular ATP modulates the gating of IKv-A, and converts IKv-A into IKv-B. A preliminary account of this work has been presented (Wilson and Pappone, 1996).
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). Cells were incubated at 37°C and 5% CO2 in Dulbecco's modified Eagle's medium supplemented with 5% fetal calf/horse serum, 0.2 U/ml insulin, 100 µg/ml penicillin, 0.1 mg/ml streptomycin, and 0.25 µg/ml amphotericin B. Cells were maintained in culture for 1–14 d before patch clamping. Mature brown adipocytes for voltage clamp experiments were identified by their many fat droplets.
Electrophysiology
Voltage clamp currents were measured using nystatin perforated-patch (Horn and Marty, 1988) or standard whole-cell patch clamp techniques (Hamill et al., 1981) as previously described (Lucero and Pappone, 1989, 1990). Thick-walled borosilicate capillaries (Sutter Instruments, Inc.) were used to manufacture pipettes with resistances of 3–4 Mohm. Typically, 50% of the 7–10-Mohm access resistance was compensated at the clamp amplifier. The voltage offset between the patch pipette and the bath solution was nulled immediately before patch formation. Membrane currents were recorded, filtered through a four-pole bessel filter at 5 kHz, and membrane capacity currents nulled with the patch clamp amplifier (3900; Dagan Corp., or Axopatch 200A; Axon Instruments) connected to a computer via a TL-1 (Axon Instruments) or ITC-16 (Instrutech) interface. Unstimulated cell capacitance ranged from 20 to 40 pF. Resting membrane potentials measured in current clamp were –20 to –40 mV.
Several different voltage protocols were used. The voltage dependence of activation was determined from currents during step depolarizations from the –60-mV holding potential (HP) or after a 3-s conditioning prepulse to –100 mV. The depolarizations ranged from –50 or –60 to +60 mV in 10-mV increments and were 400-ms long. The time course of ATP action was determined from currents during depolarizations to +40 mV. These 400-ms depolarizing steps were repeated every 20 s or 1 min from the –60 mV HP or –100 mV prepulse potential. Peak instantaneous tail current amplitude was determined from currents recorded during repolarization from a +40 mV, 400-ms depolarization. The cell was repolarized to potentials from +40 to –60 mV in 10-mV increments. The voltage dependence of steady state inactivation was determined from currents during depolarizations after 3- or 6-s conditioning prepulses from 0 to –120 mV. Linear leak and membrane capacitative currents were subtracted using a P/4 or P/5 procedure (Armstrong and Bezanilla, 1974; Lucero and Pappone, 1989) in the activation, time course, and tail current protocols. Pulse protocols were delivered and data collected and analyzed using either pClamp, version 6 (Axon Instruments) or Pulse, version 8.07 (Instrutech) software.
Solutions
Cells for patch clamp experiments were placed in a 0.5-ml chamber and continuously perfused with either bicarbonate-buffered Krebs solution equilibrated with 95% O2 /5% CO2 at a rate of 
1 ml/min or, in most perforated-patch experiments, a low-chloride Ringer's solution with 500 nM apamin, to eliminate Ca2+-activated K+ and Cl– currents (ICl,Ca and IK,Ca). The Krebs solution contained (mM): 120 NaCl, 4.5 KCl, 0.5 MgCl2, 2 CaCl2, 0.7 Na2HPO4, 1.3 NaH2PO4, 25 NaHCO3, 10 glucose, pH 7.4. The low-chloride solution contained (mM): 135 Na-aspartate, 4 KCl, 2 CaCl2, 0.5 MgCl2, 10 HEPES, pH 7.4. In some experiments, a Ringer's solution was used (mM): 135 NaCl, 4 KCl, 2 CaCl2, 0.5 MgCl2, 10 HEPES, pH 7.4. The pipette solution for whole-cell experiments usually contained (mM): 100 K-aspartate, 25 KCl, 10 NaCl, 1 CaCl2, 10 EGTA, 10 MOPS, titrated to pH 7.2 with KOH. For perforated-patch experiments, the internal pipette solution usually contained (mM): 115 K-aspartate, 25 KCl, 10 NaCl, 10 MOPS, 0.25 mg/ml nystatin, and 0.1% pluronic F127, titrated to pH 7.2 with KOH. In some whole-cell experiments, 1 mM ATP and 0.5 mM MgCl2 were added to the pipette solution. The cytosolic Ca2+ concentration was calculated to be 20 nM for the whole-cell pipette solution with Patchers Power Tools (F. Mendez, Göttingen, Germany). Osmolarities of the solutions ranged from 273 to 310 mosM. All chemicals applied to cells were mixed with the bath solution from thawed frozen stock solutions immediately before application. ATP, 2-methylthio ATP (2-MeSATP), ,β methylene ATP (,β MeATP), ADP, and adenosine were made up as 1–100-mM stock solutions in H2O. Nystatin (50 mg/ml) and pluronic F127 (0.2 g/ml) stock solutions were made up in DMSO.
Chemicals
2-Methylthio ATP and ,β methylene ATP were from Research Biochemicals Int. or Calbiochem, pluronic F127 was from Molecular Probes, Inc. Nystatin dihydrate was from Fluka Biochemical. Apamin, charybdotoxin, β-dendrotoxin, quinine sulfate, and tetraethylammonium were from Alamone Labs. All other chemicals were from Sigma Chemical Co.
Analysis
Peak conductances (G) were calculated from peak current values for the voltage dependence of K+ current activation at each potential assuming that the reversal potential was equal to EK. The conductance values were fitted with the Boltzmann relation:
 | (1) |
where G max was the maximum conductance value, E1/2 the half-maximal activation voltage, Em the membrane voltage, and k the slope factor of the voltage dependence. The voltage dependence of steady state inactivation was described by fitting peak currents (I) during depolarizations from different prepulse potentials with the sum of two Boltzmann relations:
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 | (2) |
where I max was the maximum current, E1/2 the half-maximal inactivation voltage, Em the membrane voltage, and k the slope factor of voltage dependence. Values in tables are given as the mean ± SEM. Student's t test was performed using a P < 0.05 criterion to test for significance. Either Statview version 4.5 (SAS Institute) or Excel version 5 (Microsoft Corp.) software was used for statistical analysis.
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The steady state inactivation-voltage relation for these two voltage-gated K+ currents can be fit by the sum of two Boltzmann distributions (Eq. 2), as shown in Fig. 1 B. We have termed the component activated from a –60-mV HP IKv-A. The current component inactivating at more negative potentials is IKv-B. IKv-B inactivates at potentials that average 60–70 mV more negative than IKv-A, as summarized in Table I. Our previous studies of IKv (Lucero and Pappone, 1989) were confined to more physiological potential ranges, and so failed to activate this second component of IKv.
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50% of total IKv on average in whole-cell experiments as shown in Table III. However, the distribution between IKv-A and IKv-B varied widely from cell to cell, with 0–85% of total IKv carried by IKv-B. Table III also shows that the distribution of IKv between IKv-A and IKv-B changes with the recording condition. The apparent redistribution was due to a 2.5-fold increase in IKv-B magnitude in whole-cell compared with perforated-patch recordings. Neither the total K+ current nor the relative proportions of IKv-A and IKv-B varied with cell size or the culture age of the cell.
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P2 receptor stimulation caused smaller shifts in the activation voltage dependence of IKv-A in whole-cell experiments. Fig. 5 shows that the threshold for detectable IKv-A activation was shifted by approximately –10 mV and the midpoint of the conductance–voltage relation shifted by –13 mV in a cell exposed to 0.02 µM ATP for 10 min. IKv-A activation shifts with ATP were consistently approximately one third the magnitude of inactivation shifts in whole-cell recordings. IKv-B activation voltage dependence was not determined, since both IKv-A and IKv-B components were recovered at the –100-mV conditioning potential after ATP and they could no longer be separated.
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60% decrease in IKv-A amplitude. ATP transiently increased the leak current measured at –55 mV also. This could be due to stimulation of Ca2+-activated conductances, since neither Ca2+-activated Cl– or K+ conductances were controlled. Alternatively, a nonselective cation conductance may have been activated by ATP (Lee and Pappone, 1997a). However, this is unlikely because in other perforated-patch experiments where the Ca2+-activated K+ and Cl– conductances were blocked, ATP effects on IKv-A were not necessarily associated with increases in leak currents. Fig. 6 B shows superimposed IKv-A currents during depolarization to +40 mV, before and after ATP. The initial exposure to ATP increased the rates of both activation and inactivation. Moreover, as shown in Fig. 6, C and D, ATP shifted the potential for maximal steady state IKv-A (window current) to more hyperpolarized potentials. The potential of the peak window current was shifted negative by 18 mV, through a –25-mV shift in inactivation and a –23-mV shift in activation. Thus, the effects of ATP on IKv-A in perforated-patch clamped cells are qualitatively similar to those in whole-cell experiments in that the voltage dependence of IKv-A gating is shifted negative. However, there are differences in the time course, concentration dependence, and magnitude of ATP's effects with recording configuration.
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IKv-A and IKv-B May Represent Two States of the Same Channel
Purinergic stimulation may convert IKv-A into IKv-B in whole-cell experiments. The total amount of IKv did not change significantly with purinergic stimulation, but the relative contributions of IKv-A and IKv-B to the total IKv did change. After ATP total IKv decreased 4 ± 4% in 11 whole-cell experiments and increased 20 ± 10% in 12 perforated-patch experiments. The decrease in IKv in the whole-cell recordings is similar to current run down without ATP exposure. Fig. 8 A shows steady state inactivation voltage relations before and after ATP in a whole-cell recording. Maximum IKv-A was reduced by 0.5 nA after ATP, and maximum IKv-B increased by 0.5 nA, leaving total IKv unchanged. In whole-cell recordings in which the two current components were clearly identifiable after ATP, such as that in Fig. 8 A, there was a correlation between the decrease in IKv-A and increase in IKv-B induced by ATP as shown in Fig. 8 B. The correlation was strongest in cells exposed to ATP in the whole-cell configuration. In perforated-patch experiments, total IKv increased after ATP, and in many cells this was due to an increase in IKv-A amplitude without any change in IKv-B.
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ATP (0.02–1 µM, n = 78) > ADP (10–100 µM, n = 5) ,β MeATP (100 µM, n = 3). Bath application of adenosine (100–1,000 µM, n = 3), UTP (40 µM, n = 4), or the P2X7 agonist BzATP (2'-3'-O-[4-benzoylbenzoyl]-ATP) (Lee and Pappone, unpublished observation) did not decrease the peak amplitude of IKv-A. Thus, the agonist sensitivity is consistent with modulation by P2Y1 or any of several P2X receptors, but not the P2X7 receptor (Rassendren et al., 1997). The concentration of ATP required to affect IKv-A depended on the recording configuration. In perforated-patch recordings, 5–10-fold higher ATP concentrations (1–5 µM, n = 21) were required compared with whole-cell experiments, and ATP had to be applied two to three times to cause similar decreases in IKv-A amplitude.
The P2 receptor signaling of IKv modulation has unusual properties. P2Y1 receptors normally act through G proteins, but the P2 receptor-mediated decrease in IKv-A in whole-cell recordings was not altered by standard manipulations of G protein function. Internal exposure to 1 mM GTP
S (guanosine 5'-O-[3-thiotriphosphate]; n = 6), AlF4, GDPβS (guanosine 5'-O-[2-thiodiphosphate]), pretreatment with pertussis toxin, or ATP-free solutions did not affect IKv-A or its response to externally applied ATP (Lee and Pappone, manuscript submitted for publication). P2X receptors usually act as ion channels and can trigger intracellular signals by carrying Ca2+ influx. However, in whole-cell experiments where extracellular Ca2+ was removed and intracellular Ca2+ was buffered with EGTA to <20 nM (n = 6), ATP stimulated IKv-A decreases similar to those seen when external Ca2+ was present and internal Ca2+ was unbuffered (n = 10). Thus, the signaling pathway does not fit nicely into either the G protein– linked P2Y or the channel-forming P2X family of P2 receptor.
Activation of membrane receptors often initiates phosphorylation, phospholipid degradation, or alters cytoskeletal function, but manipulations to these pathways failed to mimic or affect the ATP-stimulated decrease in IKv-A in whole-cell recordings. 10-fold lower ATP concentrations were effective in decreasing IKv-A in whole-cell as compared with perforated-patch experiments, whether or not the internal solution contained ATP. Activation of protein kinase A with 1 mM 8-bromo-cAMP (n = 3) or 5 µM forskolin (n = 4) in perforated-patch recordings, or protein kinase C with 1.6 nM PMA (n = 13) in whole-cell experiments did not decrease IKv-A amplitude. Induction of nitric oxide production with Na-nitroprusside or a pipette solution containing cGMP (Lee and Pappone, unpublished observations) were also ineffective. In whole-cell recordings, the phospholipase products phosphatidic acid (Lee and Pappone, unpublished observation) or 1–100 µM arachidonic acid (n = 6) failed to mimic the ATP-induced decrease in IKv amplitude. Altering cytoskeletal function by pretreatment with cytochalasin or phalloidin (Lee and Pappone, unpublished observations) also did not affect the ATP-mediated decrease in IKv in whole-cell recordings. While negative results are not conclusive, the inadequacy of these many agents in affecting IKv modulation by ATP suggests that the signaling may occur through an unusual pathway.
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The continuous hyperpolarizing shift in the voltage dependence of IKv-A inactivation without any change in slope cannot be explained by a mechanism where ATP modulates only a few sites. A simple model, with four modulatory sites, each causing a –20-mV shift in inactivation voltage dependence, predicts that Kv current components with intermediate voltage dependence would be seen, and that the slope of voltage dependence would flatten. Neither occurred, indicating that the process controlling the IKv-A inactivation shift by ATP involves more than a few modulatory sites and/ or is quite complex. It is also unlikely that the hyperpolarizing shift in voltage dependence can be explained by ATP-dependent phosphorylation or dephosphorylation of the channel protein, since the hyperpolarizing shifts in IKv gating are similar with or without ATP in the pipette. Possibly the ATP-induced hyperpolarizing shift in IKv-A voltage dependence arises from changes in channel interactions with the cytoskeleton (Johnson et al., 1997), changes in the membrane lipid composition (Ji et al., 1993), or altered association of the channel with regulatory subunits (Jan and Jan, 1997). Cytoskeletal elements can modulate Ca2+ currents through phosphorylation-dependent pathways (Johnson et al., 1997) and fatty acids can bind to and affect the function of the cytoskeleton (Packham et al., 1991; Exton, 1997). ATP dramatically increases brown adipocyte membrane trafficking, resulting in up to a doubling of membrane capacitance (Pappone and Lee, 1996). The incorporation of new membrane may cause changes in the plasma membrane lipid composition, involve remodeling of the cytoskeleton, or insert subunits that modify IKv gating behavior. Increases in membrane trafficking could also change the number and type of IKv channels present in the membrane. ATP does not alter total IKv in whole-cell preparations but does in perforated-patch recordings. The increase in total IKv could be explained by an increase in the number of Kv channels, since exocytosis is more robust in perforated-patch than whole-cell recordings (Lee and Pappone, manuscript submitted for publication). P2 receptor stimulation can also activate phospholipases in other cell types (Boyer et al., 1989; el-Moatassim and Dubyak, 1992; Wang et al., 1992; Humphreys and Dubyak, 1996) and the generated fatty acids can cause hyperpolarizing shifts in the voltage dependence of voltage-sensitive K+ current gating (Honore et al., 1994).
In contrast to the shift in voltage dependence, the apparent conversion of IKv-A into IKv-B by ATP could be explained by a mechanism involving only a single site. Since the conversion is more prominent in whole-cell than in perforated-patch experiments, the conversion from IKv-A to IKv-B may depend on immobile factors while the conversion of IKv-B to IKv-A may depend on soluble components. One possibility is that purinergic stimulation may lead to a change in the gating mode of IKv-A by modifying interactions with regulatory β subunits, as seen in Na+ channels (Ji et al., 1994) and Maxi K+ channels (reviewed in Jan and Jan, 1997).
Potential Signaling Pathways
The potency order of different purinergic receptor agonists in decreasing IKv-A is consistent with stimulation of any of several P2 receptor types (Barnard et al., 1997; Burnstock, 1997). Brown adipocytes have several different P2 receptors (Lee and Pappone, manuscript submitted for publication) that cause increases in cytosolic Ca2+, membrane turnover, and conductance (Pappone and Lee, 1996; Lee and Pappone, 1997a). The increases in brown adipocyte cytosolic Ca2+ by ATP are consistent with G protein activation of phospholipase C, while the increases in membrane trafficking are resistant to G protein manipulations (Pappone and Lee, 1996; Lee and Pappone, 1997a; Lee and Pappone, manuscript submitted for publication). The purinergic receptor(s) active in modulating IKv show similar insensitivity to activation or inhibition of G protein function and to the absence of cytosolic ATP. The agonist sensitivity of IKv modulation is consistent with mediation by a P2Y1 receptor or any of several of the P2X receptors (Barnard et al., 1997; Burnstock, 1997). Although P2Y receptor actions usually involve G proteins, a G protein–independent pathway has been reported for heterologously expressed P2Y1 receptors (O'Grady et al., 1996). Signaling via a P2X receptor seems less likely. P2X receptors usually form ligand gated cation channels (Bean, 1992; Surprenant et al., 1995), but the changes in IKv function by ATP were not accompanied by increases in membrane conductance in whole-cell experiments or in perforated-patch recordings with Ca2+-sensitive conductances blocked. Nonionotropic mechanisms have been described for the P2X7 receptor (el-Moatassim and Dubyak, 1992; Humphreys and Dubyak, 1996; Zou et al., 1997). However, the potent P2X7 receptor agonist BzATP (Rassendren et al., 1997) is ineffective in modulating IKv in brown adipocytes (Lee and Pappone, unpublished observation). Thus, we cannot yet draw any conclusions as to which P2 receptor regulates IKv.
Our whole-cell results show that neither the ATP-induced shift in IKv-A voltage dependence nor the conversion of IKv-A into IKv-B require soluble factors, suggesting the signaling components for these effects are immobile. However, in perforated-patch recordings, the shifts in IKv voltage dependence are attenuated, suggesting that mobile components are involved in signaling the recovery reactions. A similar lack of Ca2+ or cytosolic ATP dependence of P2 receptor modulation of ionic currents has been observed previously (Qu et al., 1993a; Thomas and Hume, 1993; Matsuura and Ehara, 1996; Matsuura et al., 1996a). P2 receptor activation of muscarinic K+ currents in heart (Matsuura et al., 1996a) and K+ currents in skeletal muscle (Thomas and Hume, 1993) involve membrane delimited pathways. It will be interesting to determine whether new signaling pathways underlie the modulation of brown fat IKv by P2 receptor activation.
Possible Functions of IKv Modulation
Voltage-gated K currents are important for regulating cell proliferation, volume regulation, and Ca2+ signaling in a variety of nonexcitable cells (Nilius and Wohlrab, 1992; Deutsch and Chen, 1993; Lin et al., 1993). In brown adipocytes, IKv is necessary for proliferation of cultured cells (Pappone and Ortiz-Miranda, 1993), but not for norepinephrine-induced thermogenesis (Pappone and Lucero, 1992), suggesting that IKv may have a specific functional role in brown adipocyte growth and development. Proliferation of brown adipocytes is promoted by P2 receptor stimulation (Barsoum et al., 1998; Wilson, Barsoum, Wilson, and Pappone, manuscript submitted for publication), suggesting that modulation of IKv by ATP may be linked to the proliferative responses. As occurs in other cell types, IKv in brown adipocytes could be linked to mitogenesis or differentiation through volume regulatory pathways (Deutsch, 1990), or Ca2+ signaling (Lin et al., 1993; Verheugen et al., 1997). Thus, ATP modulation of IKv gating may regulate either cell volume responses or agonist-activated extracellular Ca2+ entry in brown adipocytes, and these changes in cell volume or cytosolic Ca2+ could be involved in the ATP effects on cell proliferation.
Potential Sources of ATP
There are several potential sources of purine nucleotides in brown fat, all of which may be activated with sympathetic stimulation. During periods of high sympathetic nerve activity, as occurs with cold exposure or overfeeding, ATP may be released with norepinephrine (Lagercrantz, 1976; Ellis and Burnstock, 1989; Westfall et al., 1990). Mast cells are abundant in brown fat tissue and possibly also release ATP (Rothwell et al., 1984; Osipchuk and Cahalan, 1992; Desautels et al., 1994). Lastly, sympathetic activation of metabolism or subsequent hypoxia may cause brown fat cells themselves to release ATP, as has been described for hepatocytes and epithelial cells (Abraham et al., 1993; Reisin et al., 1994; Prat et al., 1996; Wang et al., 1996). Extracellular ATP levels in brown fat are not known, but our results suggest that nanomolar concentrations of ATP could evoke significant responses.
Relation to White Fat
Recent findings indicate that brown adipocytes and white adipocytes have very similar functions, gene expression, and cell signaling systems. Both cell types can store, release, or waste metabolic energy. The main difference between the tissues is whether energy storage or wasting predominates (reviewed in Himms-Hagen, 1992; Lonnroth and Smith, 1992). The conductance properties and P2 receptor responses of white adipocytes have not been as well characterized as those of brown fat cells. However, to the extent that they are known, white fat resembles brown fat. Similar IKv is present in white fat, and ATP causes similar changes in IKv and activates membrane exocytosis as in brown adipocytes (Lee and Pappone, 1997b). Thus, purinergic stimulation may also regulate the function of white adipocytes.
Summary
We find that P2 receptor activation modulates the amount and voltage dependence of activatable voltage-gated K+ currents in brown adipocytes. Extracellular ATP is likely to be increased with sympathetic stimulation of brown fat and could either augment or reduce the available IKv, depending on the stimulus strength and duration. Since IKv is known to be essential for brown adipocyte proliferation, modulation of IKv by extracellular ATP could be an important component of sympathetic regulation of adipocyte number. P2 receptors and K+ channels could prove to be effective sites for pharmacological manipulation of adipocyte growth and development.
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ACKNOWLEDGMENTS |
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This work was supported by National Institutes of Health grant GM-44840.
Submitted: 7 July 1998
Accepted: 14 October 1998
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1-Adrenergic stimulation of Cl–efflux in isolated brown adipocytes, FEBS Lett, 1990, 262, 25–28.[Medline]-Adrenergic stimulation activates a calcium-sensitive chloride current in brown fat cells, J Gen Physiol, 1995, 106, 231–258.
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). For IKv-A (solid line), E1/2 = 7 mV, k = 10 mV; for IKv-B (dashed line), E1/2 = –7 mV, k = 18 mV (Eq. 
) amplitudes measured every 20 s. 0.025 or 0.05 µM 2-MeSATP was present during the times shown by the bars. Linear currents were subtracted from the +40-mV records using a P/4 protocol. The pipette contained a solution with K-aspartate with 
) in this cell. Smooth curves are the sum of two Boltzmann relations (Eq. 
) amplitudes measured during P/4 depolarizations to –55 mV from the –80-mV leak holding potential. 5 µM ATP was present during the times shown by the bars. Numbered time points 3 (control), 4, and 7 (after ATP) correspond to current traces in B, 1 and 2 to control normalized conductance–voltage measurements shown in C, and 5 and 6 to the conductance–voltage relations after ATP shown in D. (B) IKv-A measured at the times shown in A. 3 indicates control, 4 and 7 are after one and three exposures to ATP. (C) Normalized activation (g) and steady state inactivation (h
) conductance–voltage relations before ATP. Boltzmann relations (Eq. 
) experiments before and after ATP. Error bars represent ±SEM. For some conditions, the error is smaller than the symbol. *Significant difference between control and ATP-exposed conditions based on a paired t test (P < 0.05).