Neuroblastoma

ABSTRACT We imaged the intramembrane potential (a combination of transmembrane, surface, and dipole potential) on N1E-115 neuroblastoma cells with a voltage-sensitive dye. After activation of the B^sub 2^ bradykinin receptor, the electric field sensed by the dye increased by an amount equivalent to a depolarization of 83 mV. The increase in intramembrane potential was blocked by the phospholipase C (PLC) inhibitors U-73122 and neomycin, and was invariably accompanied by a transient rise of [Ca^sub 2+^]^sub i^. A depolarized inner surface potential, as the membrane loses negative charges via phosphatidylinositol 4,5bisphosphate (PIP^sub 2^) hydrolysis, and an increase in the dipole potential, as PIP^sub 2^ is hydrolyzed to 1,2-diacylglycerol (DAG), can each account for a small portion of the change in intramembrane potential. The primary contribution to the measured change in intramembrane potential may arise from an increased dipole potential, as DAG molecules are generated from hydrolysis of other phospholipids. We found bradykinin produced an inhibition of a M-type voltage-dependent K^sup +^ current (/^sub k(M)^)- This inhibition was also blocked by the PLC inhibitors and had similar kinetics as the bradykinin-induced modulation of intramembrane potential. Our results suggest that the change in the local intramembrane potential induced by bradykinin may play a role in mediating the /^sub K(M)^ inhibition.

INTRODUCTION

Although the behavior of the voltage-gated ion channels in cell membranes is physiologically modulated by the transmembrane potential, the regulatory elements in the channels (e.g., gating charges) are fundamentally responding to the electric field within the plasma membrane-the intramembrane electric field, which is generated by a complicated sum of transmembrane potential, surface potential, and dipole potential (for reviews, see Honig et al, 1986; Loew, 1993). Surface potential is the electric potential difference between the membrane-aqueous interface and the bulk aqueous solutions (for reviews, see McLaughlin, 1977, 1989). The difference between the negative surface potential on the outer and inner surfaces of a cell membrane contributes to the intramembrane electric field. Cell membranes also possess a substantial dipole potential arising from the oriented dipoles of lipid ester groups or immobilized water molecules at the membrane-water interface (Szabo, 1974, 1977; Clarke, 1997; Flewelling and Hubbell, 1986; Franklin and Cafiso, 1993; Gawrisch et al., 1992; Zheng and Vanderkooi, 1992). This potential is on the order of several hundred mVs, positive in the interior of the membrane. Like transmembrane potential, membrane surface potential (Chandler et al., 1965; Gilbert and Ehrenstein, 1969; McLaughlin, 1989) can modulate the activity of voltagedependent ion channels by changing the intramembrane electric field. The effect of dipole potential on ion channel activity is less well studied. However, there is evidence indicating that modulators of dipole potential such as phloretin and cholesterol do affect the potassium current in the squid giant axon (Strichartz et al, 1980) and voltagedependent calcium channels in several cell types (Locher et al., 1984; Zhou et al, 1991; Sen et al., 1992). In differentiated N1E-115 cells, it has been found in this lab that the voltage-gated sodium channels of the growth cone have a faster activation rate than those in the soma (Zhang et al., 1996), possibly because the growth cone region has a more positive dipole potential than the soma membrane (Bedlack et al., 1994).

Whether the activation of membrane receptors can modulate intramembrane potential, especially surface potential and dipole potential, has not been investigated, perhaps because appropriate methods for monitoring the intramembrane potential in cells have not been well established. In particular, we are interested in the possible change in intramembrane potential after receptor activation of phospholipase C (PLC). Activation of PLC leads to hydrolysis of phosphatidylinositol 4,5-bisphosphate (PIP^sub 2^) to the second messengers, diacylglycerol and inositol 1,4,5-trisphosphate (IP^sub 3^) (Berridge, 1993). Whereas IP^sub 3^ causes Ca^sup 2+^ release from intracellular Ca^sup 2+^ stores by activating IP^sub 3^ receptors, diacylglycerol activates protein kinase C (PKC) thereby modulating a wide range of cellular responses. Recently, it has been suggested that changes in PIP^sub 2^ concentration could play an important role in regulating cellular processes (Lee and Rhee, 1995). The anionic PIP^sub 2^ comprises 0.1%-5% of total lipids in the inner leaflet of the membrane of most cells (Kleinig, 1970; Tran et al, 1993). We hypothesize that the surface potential and dipole potential profile of the membrane may be changed after hydrolysis of PIP^sub 2^ based on the following two findings: first, a decrease in PIP^sub 2^ concentration could lead to a change in the surface potential (McLaughlin, 1989); secondly, generation of 1,2-diacylglycerol (DAG) from phosphatidylinositol could increase dipole potential, since the dipole potential of a DAG monolayer is 127 mV more positive compared to that of a phosphatidylinositol monolayer (Smaby and Brockman, 1990).

Such intramembrane potentials are impossible to measure using conventional electrodes placed in the bulk solutions, since the surface potentials arising from the negative charges on both surfaces of the plasmalemma decay with a Debye length of under 10 A in 0.1 M monovalent salt solutions (McLaughlin, 1989; Cevc, 1990; Langner et al, 1990), and the dipole potentials are localized to an ~5 A thick region within the plasmalemma just below both surfaces (Szabo, 1974, 1977; Clarke, 1997; Flewelling and Hubbell, 1986; Franklin and Cafiso, 1993; Gawrisch et al., 1992; Zheng and Vanderkooi, 1992). However, certain voltage-sensitive dyes can report the local electric field at their location within the membrane. We have shown that the dual wavelength ratio of styryl dyes provides a consistent measure of transmembrane potential (Montana et al., 1989; Zhang et al., 1998). In earlier studies, others and we characterized the ability of di-8-ANEPPS to measure dipole potential changes and variations along the membrane (Bedlack et al, 1994; Cladera and O'Shea, 1998; Clarke, 1997; Clarke and Kane, 1997; Gross et al., 1994). In a recent study (Xu and Loew, 2003), we showed that the styryl dye ratiometric measurements could be used to monitor changes in surface potential.

In this paper, dual wavelength ratiometric imaging of di-8-ANEPPS was used to determine the change in the intramembrane potential of a cell during receptor-mediated activation of the PLC signaling pathway. We used bradykinin (BK) to activate PLC in N1E-115 neuroblastoma cells, a mouse neuroblastoma cell line that expresses a number of G-protein-coupled receptors, including B^sub 2^ receptors (Coggan and Thompson, 1995; 1997). BK has also been shown to modulate the M current in these cells, but the mechanism of this modulation remains unclear (Higashida and Brown, 1987). The dye ratio reports a large slow change in intramembrane potential that has the same time course as the modulation of the M current by B^sub 2^ receptor activation in these cells. Because I^sub K(M)^ has been described as a voltage-dependent K^sup +^ current (Constanti and Brown, 1981; Brown and Constanti, 1980), this raises the possibility that changes in intramembrane potential may underlie the modulation of I^sub K(M)^.

METHODS

Cell preparation

N1E-115 mouse neuroblastoma cells were grown and maintained in Dulbecco's modified Eagle's medium containing 10% fetal bovine serum. Cells were grown until 50% confluent before seeding on uncoated glass coverslip. Differentiation was initiated by replacing the media with one containing 0.5% fetal bovine serum and 1% dimethylsulfoxide. Cells were cultured for a further 2-5 days to complete differentiation and achieve neuronal morphology.

Dye loading

For di-8-ANEPPS staining, each coverslip was washed in Eagle's balanced salt solution (EBSS) buffered to pH 7.35 with 20 mM HEPES, and then incubated for 20 min at 4[degrees]C in the buffered EBSS supplemented with l-(3-sulfonatopropyl)-4-[[beta][2-(di-n-octylamino)-6-naphthyl]vinyl]pyridinium betaine (di-8-ANEPPS; 0.5 [mu]M) and Pluronic F-127 (BASF, Wyandotte, MI) (PF-127;0.05%). For double staining with Calcium Green and di-8-ANEPPS, Calcium Green-1-AM (Molecular Probes, Eugene, OR) was diluted to 8 [mu]M in 2 ml EBSS. A single coverslip of cells was immediately added, incubated 30 min at 22-25[degrees]C in the dark, and then transferred to fresh EBSS to allow recovery at 37[degrees]C for 30 min. The cells were then transferred and incubated in the 4[degrees]C buffered EBSS containing di-8-ANEPPS and Pluronic F-127 for another 20 min.

Dual wavelength imaging

Stained cells were mounted on an inverted Zeiss microscope (Axiovert 135 TV) and viewed with an oil objective (Plan-Apochromat 63 x 1.40). Dual wavelength image pairs of di-8-ANEPPS (excitation at 440 and 530 nm, emission >570 nm) were acquired using a cooled CCD camera (Model CH100, Photometries, Tucson, AZ) on the microscope, controlled by Cellscan Acquisition Control unit (Scanalytics, Marlborough, MA). An additional magnification of x 2.5 was used when fluorescent images were taken. Exposure time was 0.7 s for 440 nm and 1.4 s for 530 nm. A 2 x 2 binning of CCD pixels was used to improve the signal. Autofluorescence signals were negligible at both excitation wavelengths compared with di-8-ANEPPS fluorescence. All experiments were performed at room temperature (22-24[degrees]C). For cells that were double labeled for simultaneous calcium and membrane potential imaging, di-8-ANEPPS fluorescence was excited at 440 and 530 nm and detected at >570 nm, whereas Calcium Green fluorescence was excited at 485 nm and detected at 525 nm.

Patch clamp

Micropipettes with tip diameters of 1-2 [mu]m were pulled from glass capillaries (Drummond Scientific 100) using a Model P-80 Brown-Flaming micropipelte puller (Sutler Instrument, San Francisco, CA). The resistance of the pipettes was from 3 to 7 M[Omega]. The patch pipette solution contained (in mM): 80 potassium acetate, 30 KCl, 3 MgCl^sub 2^, 3 EGTA, 40 HEPES, 1 CaCl^sub 2^ (PH 7.3 with KOH). The whole-cell patch experiments were performed in N1E-115 neuroblastoma cells with an Axopatch ID amplifier (Axon Instruments, Foster City, CA). All electrophysiological recordings were performed at room temperature (22-24[degrees]C). Whole-cell current clamp was used to monitor membrane potential. We digitized the output signal by using the Clampex program (Axon Instruments).

Lipid vesicles

Solutions were buffered with 20 mM HEPES/Tris, pH 7.0 (21[degrees]C), and contained 140 mM NaCl in the absence of any divalent salts. Small unilamellar liposomes were made as described by Deamer and Uster (1983). Briefly, we prepared the lipid vesicles by drying chloroform solutions of either 10 mg egg phosphatidylcholine (PC) (Sigma Chemical, St Louis, MO; type XI-E), or bovine brain phosphatidylserine (PS) (Avanti Polar Lipids, Birmingham, AL), or a mixture of 90% PC and 10% l-palmitoyl-2-oleoyl-sn-glycerol (Avanti Polar Lipids) (PC:DAG), or a mixture of 90% PS and 10% 1-palmitoyl-2-oleoyl-sn-glycerol (PS:DAG), under a stream of argon for a minimum of 3 h, resuspending the lipids in 1 ml of the buffer and sonicating to clarity with a Laboratory Supplies (Hicksville, NY) G1128P1T sonifier under an argon atmosphere. For fluorescence measurements, 60 [mu]l of the vesicle suspension was diluted into 3 ml of buffer containing 0.2 [mu]M di-4-ANEPPS. The corrected excitation spectra of the vesicle suspension were measured with a SPEX CM dual-wavelength fluorescence spectrometer (SPEX Industries, Edison, NJ). All experiments were performed at 21[degrees]C.

[^sup 32^P]PIP^sub 2^ analysis

To examine the relative PIP^sub 2^ content of membrane, NlE-115 cells were grown to 30-40% confluency in 35 mm culture dishes; ~4 x 10^sup 5^ cells per culture dish were labeled to equilibrium (48 h) with 25 [mu]Ci/ml ^sup 32^P-PO^sub 4^ (carrier free, ICN) at 37[degrees] in EBSS. After washing twice to remove the unincorporated ^sub 32^P, the cells were scraped and suspended with EBSS. Cell lipids were extracted with acidic chloroform-methanol as described previously (Agranoff et al., 1983). An aliquot (20 [mu]l) of the lipid extract was subjected to thin layer chromatography (TLC), using Whatman LK5D plates impregnated with 1% potassium oxalate and 2 mM EDTA, and the mobile phase chloroform-methanol-4 N NH^sub 4^OH (45:35:10) (Gonzalez-Sastre and Folch-Pi, 1968). Radioactive bands were identified by cochromatography of a PIP^sub 2^ standard. TLC spots were transferred to scintillation vials, suspended in Cytoscint ES (ICN Biomedicals, Aurora, OH), and radioactivity determined by liquid scintillation counting (Packard 2000 CA scintillation counter). A second 20 [mu]l aliquot of the lipid extract was applied to a piece of silica gel scraped from a blank TLC plate with area similar to the scraped PIP^sub 2^ spots, and suspended in Cytoscint ES to determine radioactivity of the total lipids spotted.

RESULTS

Bradykinin increases intramembrane potential

The potential gradient inside the membrane (intramembrane potential), which is the sum of contributions from transmembrane potential, dipole potential, and the difference in the surface potentials at both sides of the membrane, was measured with the 440 nm/530 nm fluorescence ratios of di-8-ANEPPS. To calibrate the dye, we first measured the 440 nm/530 nm fluorescence ratios along the plasma membrane of a single cell as a function of the applied transmembrane potential in voltage-clamped NlE-115 neuroblastoma cells as described by Zhang et al. (1998). We found di-8-ANEPPS showed linear dependence of fluorescence ratio on potential with a slope of +13% per 100 mV (data not shown). To monitor the effect of bradykinin on intramembrane potential, we took a pair of fluorescence images every 10 s at the excitation wavelengths of 440 and 530 nm after addition of bradykinin. The correspondent ratios were normalized to the baseline fluorescence ratios taken before the addition of bradykinin to avoid cell-to-cell variation. We expressed the ratio change as a change in intramembrane potential, calculated as: [Delta]V^sub IM^ = (R- R^sub 0^)/R^sub 0^ x 100%/13%, where R is the 440/530 nm fluorescence ratio at different times after bradykinin and R^sub 0^ is the baseline fluorescence ratio in the absence of bradykinin. The intramembrane potential, calculated in this way, corresponds to the transmembrane potential change that would be required to produce the observed change in ratio. Note that the dye fundamentally measures the electric field at its site in the membrane. It does not measure a true potential difference between its location and the extracellular ground.

When stimulated with a saturating concentration of bradykinin (1 [mu]M), NlE-115 neuroblastomas showed a highly reproducible change in the fluorescence ratio of di-8-ANEPPS, which was characterized by an increase that returned to baseline even in the continued presence of bradykinin; the average peak increase in the ratio was 10.9 + or - 0.5% with a time to peak of 158.2 + or - 4.4 s (n = 11) (Fig. 1). If this ratio change were solely due to a change in transmembrane potential, it would be equivalent to a depolarization of 83.5 + or - 4.0 mV. During our recording period, no apparent phototoxicity was observed, and there was no significant change in fluorescence ratio in the EBSS control group across time (P > 0.05, n = 5; Fig. 1). Fig. 2 shows a typical recording of changes in intramembrane potential induced by bradykinin on an individual cell. Consistent with earlier studies (Zhang et al., 1998), we can see that the intramembrane electric field is heterogeneous along the cell surface; in Fig. 2, for example, a persistently higher than average ratio can be found in the membrane patch centered at 3 o'clock. Comparison between the changes in intramembrane potential along the whole cell and along the high ratio area reveals no significant difference (Fig. 2, B and C).

Bradykinin inhibits M current and depolarizes the membrane

The change in intramembrane electric field may arise from changes in transmembrane potentials, surface potentials, and dipole potentials. Bradykinin has been reported to be able to inhibit I^sub K(M)^ (Higashida and Brown, 1987) and depolarize the membrane in N1E-115 neuroblastoma cells (Tertoolen et al, 1987). To test to what extent the depolarization of transmembrane potential may contribute to the observed change in intramembrane potential, we monitored the transmembrane potential on N1E-115 cells under whole cell current clamp. Tertoolen et al. (1987) reported that addition of bradykinin (2 [mu]M) to N1E-115 cells induced an immediate hyperpolarization (10-15 s) followed by a depolarization of the membrane (5-10 mV, lasts for a few minutes) in all the cells tested using conventional electrophysiological techniques. The transient hyperpolarization was caused by activation of I^sub K(Ca)^ (Tertoolen et al., 1987; Higashida and Brown, 1987). I^sub K(Ca)^ was inhibited by intracellular Mg^sup 2+^ and could only be recorded reliably by using citrate but not acetate as the anion in the pipette, due to its high buffering capacity for intracellular Mg^sup 2+^ (Robbins et al, 1992). In our patch-clamp experiments, 3 mM MgCl^sub 2^ were included and acetate was used as the anion in the pipette solution. As a result, we found that under current clamp, addition of bradykinin (1 [mu]M) caused only a slow depolarization of the membrane potential of 7.2 + or - 0.9 mV (n = 9) that lasted for a few minutes without apparent immediate hyperpolarization. The time to peak for bradykinin-induced depolarization ranges from 62 to 220 s (Fig. 3 A, panel a and c), with an average of 140.6 + or - 17.6 s (n = 9). This is not significantly different than the time to peak for intramembrane potential modulation (158.2 + or - 4.4 s; Fig. 1). The depolarization was associated with decreased conductance, as monitored by the voltage response to transient hyperpolarizing current injections (0.15 nA) (Fig. 3 A, panel a). Under voltage clamp, we recorded an M-like current in N1E-115 cells, which is characterized by slow inward relaxations during membrane hyperpolarization, (Fig. 3 B, panel a). After addition of bradykinin, these relaxations were reduced (Fig. 3 B, panel b), consistent with an inhibition of the outward current. Current-voltage curves (Fig. 3 B, panel c) showed that bradykinin inhibits a normal outward rectification current recorded at potentials positive to -60 mV, with little effect on the current recorded at potentials negative to -60 mV. These results are compatible with the view that inhibition of the voltage-dependent I^sub k(M)^ accounts for the reduction of conductance and depolarization induced by bradykinin in N1E-115 neuroblastoma cells (Higashida and Brown, 1987). Therefore, the similar time to peak between the bradykinin-induced depolarization and intramembrane potential modulation suggests that M current inhibition and intramembrane potential modulation may be related.

of IP^sub 3^ receptor calcium channels in the ER. The increase in intramembrane potential is likely to also require activation of PLC, because we determined that: i), it is not found in the minor group of cells that do not display Ca^sup 2+^ release; ii), its amplitude was significantly correlated with the amplitude of [Ca^sup 2+^J^sub i^ elevation; and (iii), it was blocked by the PLC inhibitors U-73122 and neomycin. This Discussion will explore the possible biophysical origins of the change in intramembrane potential and its role in the modulation of I^sub KM[middot]^

The intramembrane potential is comprised of transmembrane potential, the difference between internal and external surface potential, and the internal and external dipole potential. The potential profile within the cell membrane can be derived from the sum of the potential profiles due to each of these contributions. In interpreting the dye signal, it is important to keep in mind that the ratio is fundamentally sensitive to the potential gradient (i.e., electric field) at its location rather than the actual value of potential relative to some distant arbitrary ground. First, from current-clamp measurement of membrane potential, we know that the measured 83 m V increase in intramembrane potential induced by bradykinin has only a minor contribution, 7 mV, from transmembrane depolarization. The remaining increase in intramembrane potential (~76 mV) may have significant contributions from changes in surface potential and dipole potential caused by the loss of PIP^sub 2^ and its replacement DAG. The second source of intramembrane potential is surface potential. Considering that the membrane loses ~3 negative charges for every PIP^sub 2^ that is hydrolyzed upon bradykinin-induced PLC activation, the increase in intramembrane potential may be partially explained by a decreased surface potential at the inner side of the plasma membrane. Thin layer chromatography analysis of the ^sup 32^P incorporation into the cellular lipids showed that PIP^sub 2^ comprised 0.43% of the total cellular phospholipids, which consist of phospholipids from both plasma membrane and intracellular membranes including endosomes, endoplasmic reticulum, lysosomes, mitochondria, nucleus, and Golgi. It has been found that ~50% of total cellular phospholipids are present in the plasma membrane of cultured cells (Warnock et al., 1993; Lange et al., 1989). Additionally, it has been found that ~40% of total cellular PIP^sub 2^ are present in the plasma membrane of cultured cells (Watt et al., 2002). Taking these factors into consideration, the basal level of PIP^sub 2^ comprises ~0.3% of phospholipids in the plasma membrane in our neuroblastoma cells. Furthermore, the percentage content of PIP^sub 2^ relative to the phospholipids in the inner leaflet of the plasma membrane would be 0.5%, since 80% of PIP^sub 2^ are found in the inner leaflet of the plasma membrane. Using the Gouy-Chapman-Stern theory (Chandler et al., 1965; Gilbert and Ehrenstein, 1969; McLaughlin et al., 1981), we can estimate the changes in surface potential at inner surface of the membrane after hydrolysis of all these PIP^sub 2^ molecules. If we assume the free negative charge density for the inner surface of the membrane is 0.14 e/nm^sup 2^ (Chandler et al., 1965), hydrolysis of such an amount of PIP^sub 2^ in the inner leaflet should lead to an increase in internal surface potential of ~4 mV (i.e., from -24 mV to -20 mV). Similarly, as described in the Results, the change in Ca^sup 2+^ is too small to significantly screen the internal surface charge and is in any case too fast to account for the change in intramembrane potential. Therefore, the major portion of the increase in intramembrane potential cannot be attributed to obvious mechanisms for modulation of surface potential.

The third source of intramembrane potential that must be considered is the dipole potential. The dual wavelength fluorescence ratio of di-8-ANEPPS has been shown to be particularly sensitive to dipole potential because the sensor chromophore is situated just within the membrane where the electric field set up by this potential is most intense (Gross et al., 1994). Most relevant here is an earlier monolayer study showing a striking dipole potential difference between monolayers containing phosphatidylinositol and monolayers containing its PLC mediated hydrolysis product, DAG (Smaby and Brockman 1990). In this study we measured the effect of 10% DAG on intramembrane potential in PC and PS vesicles. DAG increased intramembrane potential in both PC (27 mV) and PS vesicles (49 mV). Therefore, one might expect that generation of DAG from hydrolysis of PIP^sub 2^ at the inner leaflet of the plasma membrane could produce a large increase in dipole potential. But will a dye probe that is situated on the outer leaflet of the membrane be sensitive to a dipole potential change associated with the inner leaflet? The region of largest electric field (i.e., the steepest potential gradient) associated with dipole potential occurs within a few Aof the membrane surface, so the dye ratio will be much more sensitive to a change in lipid composition in its own leaflet of the bilayer (Gross et al., 1994). This might be expected from an equilibration of DAG across the membrane because it is known to have an extraordinarily high rate of spontaneous flipping-in lipid vesicles, the time constant for DAG equilibration across the bilayer is 70 ms compared to hours for phospholipids (Bai and Pagano, 1997). Our analysis of the IP^sub 3^ production necessary to evoke the observed calcium signals after saturating bradykinin (Fink et al., 2000) indicates that newly formed DAG reaches a level of 12,560 molcules/[mu]m^sup 2^, or ~0.9% of the internal membrane (this is similar to the initial concentration of PIP^sub 2^, but could also reflect some resynthesis of PIP^sub 2^ as a consequence of BK stimulation (Xu et al., 2003). Given that 10% DAG produces a dipole potential change of 30-10 mV in lipid vesicles, 0.9% DAG is still, therefore, insufficient to explain the observed BK-induced change in the neuroblastoma cell di-8-ANEPPS fluorescence ratio.

However, it has been reported that in both a dorsal root ganglion x neurotumor hybrid cell line F11 (Francel and Dawson, 1988) and fibroblasts (van Blitterswijk et al., 1991; Fu et al., 1992), bradykinin induces a biphasic increase in DAG, with an initial rapid, transient peak followed by a more slowly developing accumulation that increases cellular DAG level up to three- to fourfold. The first peak is caused by PIP^sub 2^ hydrolysis and is accompanied by an increase in IP^sub 3^ and [Ca^sup 2+^]^sub i[middot]^. The second increase reaches its peak after minutes and comes from hydrolysis of PC, by the phospholipase C that acts on PC (PC-PLC), or by the concerted action of phospholipase D (PLD) and phosphatidate phosphohydrolase (van Blitterswijk et al., 1991). The activation of PC-PLC and PLD in many cases is thought to follow the PKC activation by DAG originating from the initial hydrolysis of PIP^sub 2^ (Exton, 1994). In N1E-115 cells, therefore, activation of PLC by bradykinin could induce the subsequent activation of PC-PLC and/or PLD to generate substantial DAG in a prolonged phase, thereby producing a large change in intramembrane potential. Additional support for this scenario can be found in many published studies, e.g.:

1. It has been found that in NG108-15 cell line, bradykinin induces both generation of DAG from hydrolysis of PIP^sub 2^ (Yano et al., 1985) and a sustained increase in non-PIP^sub 2^ derived-DAG (Brami et al., 1991).

2. Bradykinin activates PLD in PC12 cells (Purkiss et al., 1991).

3. Dimethylsulfoxide induces a biphasic increase in DAG levels in N1E-115 cells (Clejan et al., 1996), with an initial rapid, transient peak followed by a slowly developing accumulation that increases cellular DAG level up to twofold; the first phase of DAG is generated through PIP^sub 2^ hydrolysis, whereas the second phase of DAG is generated through PC-PLC-mediated hydrolysis of PC. This result demonstrates that the coupling mechanism between hydrolysis of PIP^sub 2^ and activation of PC-PLC exists in N1E-115 cells.

4. There is a high basal level of DAG in N1E-115 neuroblastoma cells (1300 pmol/mg protein) (Clejan et al., 1996). Generally, neuroblastoma cells tend to have a high basal level of DAG. For example, the basal level of DAG is 2000 pmol/mg protein in NG 108-15 cells (Brami et al., 1991), 1500 pmol/mg protein in human SK-N-SH cells (Lee et al., 1991), and comprises 2.2% of cellular phospholipids in mouse NB41A3 cells (Yorek et al., 1994).

A typical differentiated N1E-115 cell used in our experiments has a diameter of 40 [mu]m, and 400 pg protein. Assuming the molecular area of a phospholipid is 70 A^sup 2^, the basal DAG would comprise ~4% of cellular lipids. If bradykinin-induced hydrolysis of PIP^sub 2^ did activate PC-PLC (or PLD) and induced subsequent increase in DAG up to threefold, that could produce an increase in DAG from ~4% to 12% of cell lipids. That could be sufficient to account for the observed increase in intramembrane potential, especially if we also consider the possibility of lipid domains that are enriched in both dye and DAG. Thus, only 15-20 mV of the observed change in intramembrane potential could arise from the measured change in transmembrane potential and the inferred changes in surface and dipole potential due to hydrolysis of PIP^sub 2^ to DAG; the remaining ~60 mV could arise from the dipole potential changes induced by the subsequent hydrolysis of PC to form DAG.

Bradykinin also produces two membrane current changes when applied to N1E-115 mouse neuroblastoma cells (Higashida and Brown, 1987), rat pheochromocytoma PC 12 cells (Villarroel et al., 1989), and NG 108-15 cells (Brown and Higashida, 1988; Robbins et al., 1993). It activates a transient Ca^sup 2+^-dependent K^sup +^ conductance that is associated with an increase in intracellular calcium; and it inhibits an M-type voltage-dependent K^sup +^ current (I^sub K(M)^) that develops more slowly and lasts for minutes. The M current (I^sub K(M)^) is a time- and voltage-dependent K^sup +^ current activated over the range -70 mV-0 mV. It plays an important role in regulating cell membrane excitability by stabilizing the resting membrane potential. M current inhibition by bradykinin is mediated by a pertussis toxin-insensitive G-protein (Villarroel, 1996), specifically by G^sub [alpha]q^ and/or G^sub [alpha]11^ (Jones et al., 1995) that activates PLC.

Whereas Cruzblanca et al. (1998) showed that bradykinin used intracellular Ca^sup 2+^ as second messenger to inhibit I^sub K(M)^ in rat sympathetic neurons, the mechanism underlying bradykinin suppression of I^sub K(M)^ in N1E-115 neuroblastoma cells, PC12 cells, and NG 108-15 cells remains unclear. It seems that Ca and IP^sub 3^ are not likely to be the "messenger", since in these cells i), inhibition of I^sub K(M)^ persists when the release of Ca and /or activation of I^sub K(Ca)^ is suppressed by buffering internal Ca with BAPTA (Villarroel et al., 1989); ii), inhibition of I^sub K(M)^ is not replicated by Ca^sup 2+^ injection (Higashida and Brown, 1986); and iii), introduction of IP^sub 3^ into the cells does not inhibit I^sub K(M)^ nor does it block the subsequent inhibition of I^sub K(M)^ by bradykinin, even though it does inhibit bradykinin-induced activation of I^sub K(Ca)^ (Villarroel et al., 1989; Higashida and Brown, 1986, 1987; 1988). Our finding that the IP^sub 3^-Ca^sup 2+^ release event has a much shorter duration and a much faster onset than M current inhibition also suggests that Ca^sup 2+^ elevation is not likely to be directly responsible for M current modulation in N1E-115 neuroblastoma cells, although M current inhibition requires activation of PLC, since it can be blocked by PLC inhibitors U-73122 and neomycin.

The dramatic change in the electric potential profile of the lipid bilayer that occurs after hydrolysis of PIP^sub 2^ could affect the voltage-dependence of ion channels including M-type potassium channels in the membrane. In the present work, we found that bradykinin reduced the M channel conductance in a voltage-sensitive manner. In addition, bradykinin increased intramembrane potential and inhibited I^sub K(M)^ with similar kinetics. These results are consistent with the hypothesis that an increase in intramembrane potential, possibly induced by PC-derived DAG, leads to an inhibition of I^sub K(M)^ channels in N1E-115 cells. Indeed, it has been found that 1-oleoyl-2-acetylglycerol, a membrane-permeable DAG analog (Kaibuchi et al., 1985), mimicked bradykinin-induced inhibition of I^sub K(M)^ (Brown and Higashida, 1988). DAG analogs also directly inhibit cloned voltage-gated Shaker IR, Kv 1.4, and Kv 1.6 potassium channels (Bowlby and Levitan, 1995), cardiac L-type calcium channels (Conforti et al., 1995; Schreur and Liu, 1996), and rod cyclic nucleotide-gated channels (Gordon et al., 1995) without a PKC-dependent phosphorylation reaction. Therefore, it is possible that the increase in intramembrane potential is the underlying mechanism for M current inhibition by bradykinin in these cells.

Finally, it should be noted that an increased intramembrane potential would not necessarily affect all the ion channels that reside in the membrane to the same extent. This is because the primary contribution to the measured change in intramembrane potential likely arises from an increase in dipole potentials, which are localized to an ~5 A thick region within the plasmalemma just below the lipid-aqueous interface. The gating sensors of different ion channels could be experiencing very different electric fields depending on whether they are located in this region.

In summary, we have shown that BK-mediated activation of PLC induces an increase in intramembrane potential in neurons. The increase in intramembrane potential has minor contributions from: a depolarized transmembrane potential; an increased surface potential at the inner surface, as the negatively charged PIP^sub 2^ is depleted; and an increased dipole potential, as PIP^sub 2^ is hydrolyzed to diacylglycerol. Its major contribution, we hypothesized, is an increased dipole potential produced by PC-derived DAG, as PC is hydrolyzed by PC-PLC or PLD after activation of PLC. This increase in intramembrane potential could be an underlying mechanism for M current inhibition by bradykinin, since the two events have similar kinetics and both can be blocked by PLC inhibitors.

We are pleased to acknowledge Jim Schaff, Dr. James Watras, and Dr. Mei-de Wei for providing advice and help in various aspects of this research.

We are grateful for the support of the National Institute of General Medical Science through grant No. GM35063.

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Chang Xu and Leslie M. Loew

Department of Physiology and Center for Biomedical Imaging Technology, University of Connecticut Health Center, Farmington, Connecticut 06030

Submitted May 16, 2002, and accepted for publication February 6, 2003.

Address reprint requests to Leslie M. Loew, Dept. of Physiology and Center for Biomedical Imaging Technology, University of Connecticut Health Center, Farmington, CT 06030. E-mail: les@volt.uchc.edu.

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