The Journal of Neuroscience, March 30, 2005, 25(13):3400-3413; doi:10.1523/JNEUROSCI.3231-04.2005
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Cellular/Molecular
Relationship between Membrane Phosphatidylinositol-4,5-Bisphosphate and Receptor-Mediated Inhibition of Native Neuronal M Channels
Joanna S. Winks,1
Simon Hughes,2
Alexander K. Filippov,2
Lucine Tatulian,2
Fe C. Abogadie,2
David A. Brown,2 and
Stephen J. Marsh2
1Ion Channel Pharmacology Group, IPC 388, Pfizer Global Research and Development, Sandwich Laboratories, Sandwich, Kent CT13 9NJ, United Kingdom, and 2Department of Pharmacology, University College London, London WC1E 6BT, United Kingdom
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Abstract
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The relationship between receptor-induced membrane phosphatidylinositol-4'5'-bisphosphate (PIP2) hydrolysis and M-current inhibition was assessed in single-dissociated rat sympathetic neurons by simultaneous or parallel recording of membrane current and membrane-to-cytosol translocation of the fluorescent PIP2/inositol 1,4,5-trisphosphate (IP3)-binding peptide green fluorescent protein-tagged pleckstrin homology domain of phospholipase C (GFP-PLC
-PH). The muscarinic receptor agonist oxotremorine-M produced parallel time- and concentration-dependent M-current inhibition and GFP-PLC-PH translocation; bradykinin also produced parallel time-dependent inhibition and translocation. Phosphatidylinositol-4-phosphate-5-kinase (PI5-K) overexpression reduced both M-current inhibition and GFP-PLC-PH translocation by both oxotremorine-M and bradykinin. These effects were partly reversed by wortmannin, which inhibits phosphatidylinositol-4-kinase (PI4-K). PI5-K overexpression also reduced the inhibitory action of oxotremorine-M on PIP2-gated G-protein-gated inward rectifier (Kir3.1/3.2) channels; bradykinin did not inhibit these channels. Overexpression of neuronal calcium sensor-1 protein (NCS-1), which increases PI4-K activity, did not affect responses to oxotremorine-M but reduced both fluorescence translocation and M-current inhibition by bradykinin. Using an intracellular IP3 membrane fluorescence-displacement assay, initial mean concentrations of membrane [PIP2] were estimated at 261 µM (95% confidence limit; 192-381 µM), rising to 693 µM (417-1153 µM) in neurons overexpressing PI5-K. Changes in membrane [PIP2] during application of oxotremorine-M were calculated from fluorescence data. The results, taken in conjunction with previous data for KCNQ2/3 (Kv7.2/Kv7.3) channel gating by PIP2 (Zhang et al., 2003
), accorded with the hypothesis that the inhibitory action of oxotremorine-M on M current resulted from depletion of PIP2. The effects of bradykinin require additional components of action, which might involve IP3-induced Ca2+ release and consequent M-channel inhibition (as proposed previously) and stimulation of PIP2 synthesis by Ca2+-dependent activation of NCS-1.
Key words: PIP2; M current; neuronal excitability; G-protein-coupled receptors; PLC; sympathetic neurons
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Introduction
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M channels are low-threshold K+ channels that regulate neuronal excitability in both the CNS and the PNS (Brown and Adams, 1980; Brown, 1988; Marrion, 1997). They are composed of subunits of the KCNQ (Kv7) gene family (Wang et al., 1998; Jentsch, 2000). The channels are inhibited by stimulating Gq/11-coupled receptors, for example in sympathetic neurons by M1 muscarinic acetylcholine receptors (M1-mAChRs) (Marrion et al., 1989; Bernheim et al., 1992) and by bradykinin (BK) B2 receptors (Jones et al., 1995). The activated G-protein subunit [principally G
q in the rat sympathetic neuron (Haley et al., 1998)] does not appear to gate the M channels directly but instead induces their closure by some indirect mechanism (Selyanko et al., 1992; Marrion, 1993).
Recent evidence suggests that receptor-induced closure results from hydrolysis of the membrane phospholipid, phosphatidylinositol-4'5'-bisphosphate (PIP2) (Suh et al., 2004). As with a number of other channels (Hilgemann et al., 2001; Runnels et al., 2002; Loussouarn et al., 2003), PIP2 is required to maintain KCNQ channels in their open state (Zhang et al., 2003), and resynthesis of PIP2 is necessary for recovery from M-channel inhibition produced by mAChRs (Suh and Hille, 2002) and by nucleotide receptors (Ford et al., 2003). Thus, mAChR-induced closure might result directly from the local depletion of PIP2 consequent on its hydrolysis. In contrast, BK-induced M-channel inhibition more likely stems from the action of a downstream product of PIP2 hydrolysis, inositol 1,4,5-trisphosphate (IP3), which releases Ca2+ from intracellular stores and thereby closes channels via binding to calmodulin (Selyanko and Brown, 1996; Cruzblanca et al., 1998; Bofill-Cardona et al., 2000; Delmas et al., 2002; Gamper and Shapiro, 2003). Another product of PIP2 hydrolysis, diacylglycerol, might contribute to mAChR-induced inhibition of M channels through activation of protein kinase C and subsequent channel phosphorylation (Hoshi et al., 2003). These alternative scenarios are summarized by Delmas et al. (2004).
Because both muscarinic agonists and bradykinin are clearly capable of activating phospholipase C (PLC) in sympathetic neurons (Del Rio et al., 1999), we examined more closely the relationship between PIP2 hydrolysis and M-channel closure in these neurons. For this, we used a fluorescent IP3/PIP2-binding peptide [the green fluorescent protein-tagged pleckstrin homology domain of PLC (GFP-PLC-PH)] (Stauffer et al., 1998; Varnai and Balla, 1998; Nahorski et al., 2003). We then asked what happens when the synthesis of PIP2 is enhanced by overexpressing the enzyme phosphatidylinositol-4-phosphate-5-kinase (PI5-K) (Ishihara et al., 1996) or the neuronal calcium sensor-1 protein (NCS-1; frequenin), which increases the activity of phosphatdidylinositol-4-kinase (PI4-K) (Burgoyne et al., 2004). Finally, we compared these effects of PI5-K overexpression with its effects on receptor-induced inhibition of G-protein-gated inward rectifier (GIRK; Kir3.1+Kir3.2) channels, which has been attributed to PIP2 depletion (Kobrinsky et al., 2000; Lei et al., 2001; Meyer et al., 2001; Bender et al., 2002).
Preliminary reports of our observations have been published previously (Winks and Marsh, 2003; Winks et al., 2003)
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Materials and Methods
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Neuron cultures. Superior cervical ganglia (SCGs) were isolated from 17-d-old Sprague Dawley rats (either gender) after terminal anesthesia using CO2, followed by decapitation in accordance with United Kingdom Home Office regulatory procedures. Ganglia were digested in collagenase solution (800 international units of collagenase class 2; Sigma-Aldrich, Dorset, UK), 12 mg of bovine serum albumin (BSA; Sigma-Aldrich), and 2 ml of HBSS plus HEPES for 15 min at 37°C. Ganglia were then rinsed with HBSS plus 10 mM HEPES and placed in trypsin solution [2 mg of trypsin, bovine type XII S (Sigma-Aldrich), 12 mg of BSA, 2 ml of HBSS plus 10 mM HEPES) for 30 min at 37°C. Digested fragments were then transferred into 2 ml of growth medium [42 ml of L-15 plus GlutaMAX (Sigma-Aldrich), 8.5 ml of 1.26% NaHCO3 (24 mM; Invitrogen, San Diego, CA), 5 ml of fetal calf serum (Sigma-Aldrich), 1.15 ml of 30% glucose, 0.61 ml of 200 mM L-glutamine (2 mM; Invitrogen), 50 ng/ml NGF 7S (Sigma-Aldrich)], and the mixture was triturated and then centrifuged five times, after which the remaining cell suspension was spun for 5 min at 800 rpm. The supernatant was then discarded, and the cells were resuspended in 5 ml of growth media. For electrophysiological recording/imaging, 
500 µl of cell solution was added to each sterile culture/recording chamber consisting of a glass ring (diameter, 16 mm; height, 3 mm) attached to a laminin substrate-coated borosilicate glass coverslip (22 x 22 mm; thickness, 0.13-0.17 mm; BDH Chemicals, Poole, UK) placed inside a 35 mm Petri dish. Chambers were then incubated at 37°C in 95% O2/5% CO2 for at least 4 h to allow neurons to adhere to the coverslip. Neurons were usually transfected/recorded between 1 and 4 d after plating.
Transfection. Neurons were transfected with cDNA 1-2 d after culture by intranuclear microinjection of cDNA plasmids. Sharp electrodes (>50 M
resistance) were filled with plasmid dissolved in injection solution (in mM: 154 NaCl, 5 HEPES, 2.5 KCl, and 0.5 MgCl2, pH adjusted to 7.4 using NaOH) at a concentration of 100 ng/µl of each cDNA plasmid to be injected. Injections were performed using an Eppendorf Scientific (Westbury, NY) automated microinjector (Transjector 5246) or (for Kir 3.1/3.2 cDNAs) a Narishige (Tokyo, Japan) micromanipulator. Neurons were incubated for 1 d before use unless otherwise stated. The following plasmids were used: pleckstrin-homology domain of phospholipase C-1 in enhanced GFP (eGFP)-C1 vector (GFP-PLC- PH) [obtained from T. Meyer (Stanford University, Stanford, CA) via K. Young (University of Leicester, Leicester, UK)]; rat neuronal calcium sensor protein-1 in pcDNA 3.1 vector (obtained from J. Weiss, University of Sheffield, Sheffield, UK); murine phosphatidylinositol 4-phosphate-5-kinase-I
in pcDNA 3.1 vector containing GFP sequence (GFPmPIP5K-I; here, PI5-K; obtained from J. N. Davis, Louisiana State University Health Sciences Center, Shreveport, LA); mutant phosphatidylinositol 4-phosphate-5-kinase-I, lacking kinase activity, in pcDNA 3.1 vector containing GFP sequence [GFPmPIP5K-I(K179M); here, mutant PI5-K; obtained from J. N. Davis); Kir3.1 and Kir3.2 (GIRK1 and GIRK2) in pCI and pRc/cytomegalovirus (CMV) vectors, respectively (from F. Lesage, Sophia Antipolis, Valbonne, France); enhanced green fluorescent protein in pcDNA 3.1 vector (eGFP) and red fluorescent cloning vector (pDSRed2-N1; Clontech, Cambridge, UK). All plasmids used contained the CMV promoter. Plasmids were propagated in DH5 (Invitrogen) Escherichia coli and purified using Qiagen (Hilden, Germany) maxiprep columns.
Electrophysiological recording. The perforated-patch configuration of the patch-clamp technique was used (except where otherwise stated) with amphotericin B as the pore-forming agent (Rae et al., 1991). Neurons were superfused via a gravity-driven system (10-15 ml/min; bath volume, 500 µl) with an external solution containing the following (in mM): 136 NaCl, 11.1 glucose, 5 HEPES, 3 KCl, 2.5 CaCl2, and 1.5 MgCl2, pH adjusted to 7.4 with NaOH. For inward rectifier (Kir 3.1/3.2, GIRK) current recording, the KCl concentration in the bath solution was increased to 6 mM (KCl substituted for NaCl) and tetrodotoxin (0.5 µM) added to block the Na+ current. Pipettes with 2-3 M resistance were filled with an internal solution containing the following (in mM): M current, 30 KCl, 100 K acetate, 20 HEPES, 10 EGTA, and 3 MgCl2, pH7.2 with KOH; Kir/GIRK current, 20 KCl, 90 K acetate, 3 MgCl2, 40 HEPES, and 0.1 BAPTA, pH 7.4 with KOH; and N-type Ca2+ current, 140 CsCl, 1 MgCl2, 0.1 BAPTA, 0.1 CaCl2, and 10 HEPES, CsOH to pH 7.4. All recordings were made at room temperature (19-22°C) unless otherwise stated. Currents were recorded using an Axopatch 200A or 200B amplifier (except where otherwise stated), filtered at 1 kHz, and digitized at 1-10 kHz. Pipette series resistances were compensated 60-90%.
M currents. Cells were held at -20 mV to preactivate the current. Currents were then deactivated at intervals of 10-15 s by 1 s hyperpolarizing steps to -50 mV. Current amplitudes were measured from extrapolated deactivation current tails using a biexponential curve-fitting procedure.
Ca2+ currents. N-type Ca2+ currents were recorded by holding the cell at -70 mV and depolarizing to +5 mV for 200 ms. Currents were leak subtracted by blockade with 5 mM CoCl2. Ca2+ charge entry was determined from the integral of the current (after leak subtraction) as described by Wanaverbecq et al. (2003).
G-protein-gated inward rectifier potassium currents. GIRK currents were recorded from cells preinjected with Kir3.1/3.2 and eGFP cDNA plasmids and identified by eGFP fluorescence, as described by Fernandez-Fernandez et al. (1999) but with the differences that perforated-patch recording was used and the external K+ concentration was 6 mM. Currents were generated by applying 200 ms voltage ramps between -140 and -40 mV every 5 s from a holding potential of -40 mV and recorded using an Axoclamp 2B switch-clamp amplifier in discontinuous mode (sample rate, 6-8 kHz).
Fluorescence microscopy. Live neurons were imaged using a Nikon (Tokyo, Japan) Diaphot inverted microscope with a 40x oil-immersion lens. Excitation light was provided by a monochromator (Polychrome II; TILL Photonics, Martinsried, Germany) controlled via an Apple Macintosh G4 computer and Openlab imaging software (Improvision, Coventry, UK). Images were captured on a 12-bit grayscale digital camera (C488000; Hamamatsu, Shizouka, Japan) and pseudo-color palettes overlaid according to the dye imaged. Images were acquired with minimum delay, according to the exposure time of the camera, but usually more than one image per second.
Digital deconvolution. Images obtained with the above equipment are subject to a certain amount of blurring, caused by light "haze" emitted from above and below the plane of focus. To remove this blurring without the use of laser confocal microscopy, digital deconvolution was used. This technique corrects and removes the noise and haze originating from images in other focal planes using mathematical algorithms, the simplest of which is the "nearest neighbor algorithm" (Agard et al., 1989; Castleman, 1996). This algorithm calculates the contribution of light noise from the focal planes above and below the desired slice and subtracts them. An automation was designed in-house, using Openlab software, which automatically focused the microscope to the bottom of the cell, and then programmed it to acquire several optical slices (
20) in 0.5 µm steps through the cell. An image from a particular focal plane was corrected using the nearest five-neighbors algorithm (i.e., using five optical slices above and five below the chosen plane). The algorithm calculates the point-spread function of the microscope, which can be estimated once the objective, immersion solution, camera resolution, and binning are known. The images produced by this method were indistinguishable from those obtained from a "conventional" laser-scanning microscope (SP2; Leica, Nussloch, Germany), which essentially (at this resolution) gave the same results.
Simultaneous electrophysiological and fluorescence recording. After locating a fluorescent cell, it was then patched at a holding potential of -60 mV using only transmitted light. The light was then switched off while the patched membrane permeabilized, because amphotericin B is light sensitive. When the access resistance was suitably low (<15 M), the fluorescent light was switched on and the focus and camera exposure time were adjusted. The membrane potential was then set to the holding potential for M-current activation (-20 to -30 mV). Hyperpolarizing steps to -50 mV to deactivate M currents were initiated by a digital pulse, which was triggered by commencement of the imaging recording, to synchronize the two recordings.
Ca2+ imaging. Cells were incubated with 2 µM fura-2 AM for 30 min at 37°C and then washed for 20 min before commencing experimentation. Alternate images at 350 and 380 nm excitation wavelengths were recorded at 1 Hz and background subtracted, and the ratio images were calibrated using an algorithm formulated in a separated set of experiments. The equipment was calibrated by cell dialysis in the whole-cell configuration using intracellular solutions from a calcium calibration buffer kit (Molecular Probes, Eugene, OR) containing fura-2 acid (100 µM).
Cell volume measurements. Cell volume was determined using a Leica SP2 confocal-scanning microscope. A z-stack consisting of a series of 0.2 µm optical slices was taken through cells that had been preloaded with the intracellular fluorescent marker calcein (calcein-AM, 2 µM for 35 min at 37°C; excitation/emission wavelengths, 480/530 nm). These two-dimensional slices were then three-dimensionally reconstructed, and the cell surface was rendered using the Imaris software package (Bitplane, Zurich, Switzerland). Volumes of the isolated soma were determined by voxel counting of the entire calibrated and reconstructed volume.
Free intracellular space. To determine the free intracellular space available to a small hydrophilic molecule such as inositol-1,4,5-trisphosphate (see Results), we compared the fluorescence intensity of known concentrations of a low-molecular-weight fluorophore [4,4-difluoro-4-bora-3a,4a-diaza-s-indacene fluorophoro (BODIPY-FL); molecular weight, 292; Molecular Probes] contained within aqueous vesicles with that of the same concentrations fully dialyzed into patch-clamped sympathetic neurons. Aqueous vesicles in a silicon elastomer (Sylgard 182; Dow Corning, Midland, MI) were fashioned after the methods of Canepari et al. (2001). BODIPY-FL was initially dissolved in intracellular solution and then 50 µl of this solution was rapidly stirred into 1 ml of uncured elastomer. A small aliquot was placed on a microscope slide and covered with a glass coverslip. The microscope slide was then heated on a hotplate at 60°C for 5 min to set, firmly embedding the vesicles within. The microscope was optimized for recording the fluorescence intensity of the highest concentration of BODIPY-FL (100 µM) contained within spherical aqueous vesicles of comparable size to sympathetic neurons (19-23 µm diameter) by adjusting the excitation intensity (480 nm) to minimize photobleaching (<5% in 10 min) and optimizing the 12-bit image capture by using multiple frame-averaging (n = 5) background subtraction and digital deconvolution. Once set, these parameters were used throughout the entire experiment. The dialysis of BODIPY-FL into SCG cells was monitored by taking brief fluorescent images (100 ms) every minute for 15 min, after which a digital deconvolved image was recorded. To minimize errors, we used the same BODIPY-FL-containing intracellular solutions for whole-cell dialysis and aqueous vesicle measurement. Fluorescence intensity was measured using a binary mask generated by image thresholding. This allows an average intensity within boundaries of both the dialyzed cells and aqueous vesicles to be measured. The free intracellular space was determined by subjecting the raw data to linear regression and calculating the ratio of the slopes of the lines generated (see Fig. 8 A).
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Figure 8. A, Determination of the free cytosolic volume. Graphs show the mean fluorescence intensity (ordinates) after the dispersion of different concentrations of a fluorescent low-molecular-weight compound (BODIPY-FL) in aqueous vesicles (squares) and mean cytosolic intensity of sympathetic neurons whole-cell patched with electrodes containing the same fluorescent compound (triangles) (see Materials and Methods). Mean slopes (fluorescence intensity per micromolar concentration [BODIPY]) ± SEM were as follows: aqueous vesicles, 26 ± 0.9; intracellular, 6.6 ± 0.66. The ratio of the slopes provides a measure of the free cytosolic space available to BODIPY, from which cytosolic concentration of IP3 generated by PIP2 hydrolysis could be estimated (see Results). B, Changes in distribution of GFP-PLC-PH between membrane and cytosol with progressive hydrolysis of PIP2, calculated from Equations 3-7, for a sympathetic neuron with a diameter of 20 µM, an initial membrane PIP2 concentration of 261 µM, an effective CMVR of 83, and binding constants of 0.1 and 2 µM for IP3 and PIP2, respectively (Table 1). The dashed line shows how the fractional M-channel availability would be expected to change during PIP2 hydrolysis, as calculated from the data in the study by Zhang et al. (2003).
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Immunocytochemistry. Cells for immunocytochemistry were cultured on glass coverslips in four-well plates (Nunclon; Nunc, Roskilde, Denmark). Cells were fixed with 0.2% gluteraldehyde/2% paraformaldehyde (in PBS; Sigma-Aldrich) for 20 min at room temperature. Fixed cells were then permeabilized with 0.1% Triton X-100 (Sigma-Aldrich) in PBS for 15 min. After washing the cells several times in PBS, they were incubated for 1 h in a blocking buffer consisting of 10 mg·ml-1 BSA (Sigma-Aldrich) in PBS and then for 1 h in the primary antibody, rabbit anti-NCS-1 (1:200). A secondary antibody (swine anti-rabbit IgG conjugated with tetramethylrhodamine isothiocyanate, 1:200) was then used for visualization. After several washes with PBS, the coverslips were removed from the four-well plates and mounted onto slides using a mounting medium, where they were sealed using clear nail varnish. Slides were refrigerated until examination. The level of auto-fluorescence was estimated by performing the above immunocytochemistry procedures in the absence of primary and secondary antibodies. Nonspecific binding was determined in the absence of the primary antibody.
Data. Results are expressed as means ± SEM (95% confidence limits). Significance levels were determined by one- or two-tailed t tests as appropriate, and differences were considered real at p < 0.05.
Drugs and chemicals. Reagents were obtained from BDH Chemicals unless otherwise stated. Oxotremorine-M (Oxo-M), bradykinin (triacetate salt), IP3 (D-myo-inositol 1,4,5-trisphosphate hexasodium salt), 1-(6-((17-3-methoxyester-1,3,5(10)-trien-17-yl)amino)hexyl)-1H-pyrrole-2,5-dione (U73122
[GenBank]
), and amphotericin B were all obtained from Sigma-Aldrich. Calcein-AM, fura-2 AM, and BODIPY FL were obtained from Molecular Probes. The primary anti-NCS-1 antibody used in the immunocytochemistry was kindly provided by J. Weiss; the secondary antibody was obtained from Santa Cruz Biotechnology (Santa Cruz, CA).
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Results
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The response to muscarinic receptor stimulation
In the absence of receptor stimulation, the expressed GFP-PLC-PH construct was highly localized to the plasma membrane (Fig. 1A, control). The addition of the mAChR agonist Oxo-M (10 µM) to the bathing fluid produced a rapid and substantial translocation of GFP-PLC-PH into the cytosol. Thus, a confocal line scan showed a clear reduction in membrane fluorescence as well as an increase in cytosolic fluorescence (Fig. 1B). However, because of cell movement, line scans could not be routinely used to quantitate translocation; instead, we did this in the first instance by monitoring the time-dependent increase of fluorescence within a defined region of the cytoplasm, as shown in Fig. 1C.
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Figure 1. Agonist-induced translocation of GFP-PLC-PH. A, Scanning confocal microscope images (8-bit resolution; i.e., 256 gray scales) of GFP-PLC-PH fluorescence distribution in a sympathetic neuron before and during application of 10 µM Oxo-M. B, Plot of fluorescence intensity across a single line scan (A, white line) before and during the peak of the Oxo-M response. C, Continuous time plot of cytosolic fluorescence intensity recorded in a single region of interest (A, white box). D, Changes in CFI (12-bit resolution; i.e., 4096 gray scales) in a single neuron produced by successive applications of 10 µM Oxo-M and 100 nM BK. E, Mean changes in cytosolic fluorescence in seven neurons produced by 10 µM Oxo-M and 100 nM BK applied sequentially as in D. The change in fluorescence is given as  F/F0 x 100%, where F = F - F0, F is peak fluorescence after agonist application, and F0 is baseline fluorescence before agonist application.
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Figure 2 shows that, when simultaneously recorded, the time courses of the changes in cytosolic fluorescence (Fig. 2A), resting membrane current (Fig. 2B), M-current amplitude (Fig. 2C, deduced from deactivation tail-currents, B), and their recoveries after washout were closely matched. Likewise, the concentration dependence of the cytosolic fluorescence increase mirrored that for membrane current reduction when recorded simultaneously (Fig. 3A), and the mean increase in cytosolic fluorescence (F/F0) coincided very closely with the mean inhibition of the M current recorded separately (Fig. 3B). Thus, there is a good concordance between PIP2 hydrolysis, as monitored by GFP-PLC-PH translocation, and M-current inhibition for this agonist.
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Figure 3. Concentration dependence of oxotremorine-M-induced increases in cytosolic fluorescence and M-current inhibition. A, Simultaneous recording of CFI (arbitrary units) and membrane current (in picoamperes) during step-wise increments in the concentration of oxotremorine-M added to the bath-perfusion fluid. B, Mean concentration-response curves for percentage of inhibition of M current (filled squares and dashed line) and percentage increase in cytosolic fluorescence (F/F0 x 100%; open circles and dotted line) produced by increasing concentrations of oxotremorine-M (in log-molar units). Oxotremorine-M was added cumulatively as in A. Each point is the mean from measurements in 5-13 neurons; bars show unidirectional SEMs. Curves are drawn according to the equation y = ymax x x/(x + K). Values for ymax and K were (mean ± SEM) as follows: M-current inhibition, 61.0 ± 2.4%, 0.68 ± 0.13 µM; fluorescence, 58.2 ± 3.3%, 0.68 ± 0.94 µM.
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As reported previously (Suh and Hille, 2002), the aminosteroid phospholipase C inhibitor U73122
[GenBank]
(3 µM) produced a slow inhibition of the M current and substantially reduced the inhibitory action of 10 µM oxotremorine-M on the residual current, from 66 ± 2.5% (n = 6) to 23.5 ± 4.2% (n = 12). U73122
[GenBank]
did not change the resting cytosolic fluorescence intensity (CFI) in cells expressing GFP-PLC-PH but very strongly reduced the increment in fluorescence after application of 10 µM oxotremorine-M, from 72.2 ± 4.2% (n = 9) to 5.9 ± 1.9% (n = 7). This reinforces the view that the increased fluorescence induced by oxotremorine-M results from activation of phospholipase C and consequent hydrolysis of PIP2.
It might be expected that the PLC-PH peptide could itself reduce M current (by sequestering PIP2) and/or reduce the inhibition of M current by agonist (by competing with PLC), as reported previously for thyrotropin-releasing hormone-induced inhibition of GIRK currents (Lei et al., 2001). No significant reduction of M current or oxotremorine-M-induced inhibition was apparent 1 d after GFP-PLC-PH cDNA injection (as used for the experiments described above). However, M current was substantially and significantly reduced after 3 d of expression when compared with control neurons cultured for 3 d simultaneously, from 526 ± 76 to 198 ± 16 pA (n = 7 for each).
The response to bradykinin
Bradykinin (10-100 nM) also induced translocation of the GFP-PLC-PH construct, similar to that produced by 10 µM oxotremorine-M (Fig. 1D,E). However, the effect of bradykinin differed in two respects from that of oxotremorine-M. First, recovery was much slower. Second, the effect showed strong desensitization, such that it was not possible to obtain two equivalent responses from the same cell. This accords with the slow recovery from, and desensitization of, M-current inhibition by bradykinin (Jones et al., 1995) and unfortunately precluded construction of accurate concentration-response curves.
PI5-K overexpression reduces the effects of both oxotremorine-M and bradykinin
Individual neurons were injected intranuclearly with plasmids encoding GFP-tagged versions of either the wild-type PI5-K or a mutant form PI5-K(K179M) devoid of kinase activity (Davis et al., 1997). In both cases, substantial fluorescence in association with the outer membrane was apparent after 2 d of expression but with additional patches of cytoplasmic fluorescence (perhaps in association with the Golgi/endoplasmic reticulum) (Fig. 4A).
M-current amplitudes (measured from deactivation tail-currents) (see Materials and Methods) in GFP-PI5-K-expressing cells and mutant PI5-K(K179M)-expressing cells were not significantly different from those in control cells. Mean amplitudes (in picoamperes) ± SEM were as follows: controls, 215 ± 16 (n = 15); PI5-K, 236 ± 27 (n = 19); PI5-K(K179M), 223 ± 35 (n = 11). Mean deactivation time constants also were not significantly different between the three groups, but time constants varied considerably in PI5-K-expressing neurons and, in some cells, were noticeably slow. This might suggest a PIP2-dependent effect on gating, as previously noted for KCNQ1/KCNE1 channels by Loussouarn et al. (2003). Because of the variability, however, we could not explore this in more detail.
M-current inhibition by both oxotremorine-M and bradykinin was strikingly reduced in wild-type PI5-K overexpressing neurons by 80 and 85%, respectively (Fig. 4B). This effect was not observed in cells overexpressing the kinase-deficient mutant PI5-K(K179M). It should be noted that the concentrations of oxotremorine-M and bradykinin used were maximally effective in normal neurons (Fig. 3) (Jones et al., 1995).
To assess the effect of PI5-K overexpression on GFP-PLC-PH translocation, we replaced the wild-type GFP-tagged PI5-K construct with a construct encoding the red fluorescent protein, pDSRed, and then recorded responses from cells that expressed both red and green fluorescence (Fig. 4A). The elevation of cytosolic GFP-PLC-PH fluorescence produced by both oxotremorine-M and bradykinin was strongly reduced by 88 and 55%, respectively (Fig. 4C). We interpret this to result from the apparent increased level of PIP2 caused by increased PI5-K activity and consequent competitive inhibition of cytosolic IP3 binding to GFP-PLC-PH (see below).
To assess whether IP3 production was modified by PI5-K overexpression, we recorded the elevation of intracellular Ca2+ produced by bradykinin using fura-2 AM. [Oxotremorine-M does not produce a detectable Ca2+ signal in these cells, but bradykinin produces a small rise (Cruzblanca et al., 1998; Delmas et al., 2002).] To avoid desensitization, we used brief pressure applications of 100 nM bradykinin. Ca2+ signals, although highly variable, were not clearly altered in PI5-K overexpressing cells (controls, 221 ± 120 nM, n = 5; PI5-K, 198 ± 108 nM, n = 6). This suggests that the total amount of IP3 production remained approximately the same, despite the apparent elevation of the initial membrane PIP2 concentration (see below), although we cannot exclude the possibility that the release mechanism might have been partly desensitized at the (presumed) higher resting IP3 concentration.
If the effects of PI5-K overexpression on M-current inhibition result from increased levels of PIP2, these should be reduced or reversed by inhibiting the supply of the substrate phosphatidylinositol-4-phosphate (PIP). To do this, we used 15-20 µM wortmannin, which, at such concentrations, is an effective membrane-permeable inhibitor of PI4-kinase (Nakanishi et al., 1995). As reported previously (Suh and Hille, 2002; Ford et al., 2003), wortmannin itself produced a slow "rundown" of the resting M current (by 28.7 ± 4.7%; n = 7; after 10 min in 20 µM wortmannin) and greatly slowed recovery from oxotremorine-M-induced inhibition. It also slowed the recovery of the cytosolic GFP-PLC-PH signal; this effect was variable, but, in some neurons, several minutes were required for full recovery. (Attempts to test whether it affected recovery from bradykinin-induced inhibition were frustrated by the very variable duration of bradykinin action in the absence of wortmannin.) However, wortmannin (15 µM) did not significantly affect the fractional inhibition of the residual M current by either oxotremorine-M or bradykinin (Fig. 4D). In contrast, it enhanced the (small) inhibition observed in cells overexpressing PI5-K and, hence, partly reversed the effect of the PI5-K (Fig. 4D, filled bars). This provides evidence that the effects of PI5-K overexpression are indeed a result of the increased synthesis of PIP2.
NCS-1 overexpression selectively reduces the effects of bradykinin
NCS-1 is a myristoylated protein that enhances the activity of PI4-kinase (see the Introduction). In PC12 cells, overexpression of NCS-1 has been reported to increase cellular levels of PIP and PIP2 to enhance IP3 production and Ca2+ release by UTP and to increase Ca2+ release by bradykinin (Koizumi et al., 2002; Rajebhosale et al., 2003). In the present experiments, previous intranuclear injection of an NCS-1 cDNA plasmid increased levels of NCS-1 protein in sympathetic neurons as determined by immunofluorescent antibody staining 2 d later (Fig. 5A, B). Deconvolution imaging indicated that much of the increased staining was in, or near, the plasma membrane (Fig. 5Bii).
M-current amplitudes in NCS-1 overexpressing neurons were not significantly different from those in uninjected neurons or in sham (GFP-plasmid)-injected neurons. Mean amplitudes were as follows: controls, 129 ± 27 pA (n = 11); NCS-1, 105 ± 12 pA (n = 9); GFP, 122 ± 18 pA (n = 9). There was also no significant change in the inhibition of M current produced by 10 µM oxotremorine-M (Fig. 5C). However, the inhibition produced by 100 nM bradykinin was reduced from 43 ± 9.0% (n = 5) to 14 ± 4.4% (n = 6) (Fig. 5D). This effect was reversed by the PI4-K inhibitor wortmannin (20 µM) (Fig. 5E).
NCS-1 overexpression also did not affect the resting membrane:cytosol distribution of GFP-PLC-PH. However, as with M-current inhibition, NCS-1 selectively reduced the increase in cytosolic fluorescence produced by bradykinin (Fig. 5F).
A component, at least, of the inhibition of M current by bradykinin appears to be mediated by Ca2+ release (see the Introduction). Because NCS-1 is a Ca2+-binding protein (Schaad et al., 1996), the reduction of bradykinin-induced inhibition in NCS-1 overexpressing neurons in Figure 5D might have resulted from increased buffering of intracellular Ca2+. This was difficult to assess directly because of the small amplitude and variability of the bradykinin-induced Ca2+ signals (see above). Instead, we tested for changes in buffering capacity by comparing the integral of the Ca2+ current evoked by 200 ms depolarizing voltage steps to 0 from -70 mV with the peak rise in intracellular free [Ca2+] recorded using fura-2 as described by Wanaverbecq et al. (2003). From this, the Ca2+-binding ratio may be calculated as 
= [Ca2+]bound/[Ca2+]i = ([Ca2+]total - [Ca2+]i)/[Ca2+]I, where [Ca2+]i is the increase in free cytosolic Ca2+ measured with fura-2, and [Ca2+]total is the total Ca2+ entry calculated from the clamp current. The baseline for the latter was set using 1 mM Co2+ to suppress the Ca2+ current. NCS-1-expressing cells were identified using pDSRed instead of GFP to prevent interference with the fura-2 signals. NCS-1 had no significant effect on the relationship between Ca2+ entry and fura-2 fluorescence or on the mean calculated binding ratio deduced therefrom (control, 650 ± 140, n = 6; NCS-1, 830 ± 180, n = 7). Also, NCS-1 had no effect on the resting levels of [Ca2+]i as measured by fura-2 fluorescence (controls, 231 ± 12 nM, n = 18; NCS-1, 210 ± 12 nM, n = 10; GFP, 224 ± 25 nM, n = 8). Thus, the effects of NCS-1 are unlikely to be the result of any appreciable increase in Ca2+ buffering.
Effect of PI5-K overexpression on inhibition of GIRK currents
GIRK currents were generated in these neurons by intranuclear coinjection of plasmids encoding GIRK1 and GIRK2 subunits (Kv3.1 and Kv3.2) along with a marker plasmid encoding GFP, as described previously (Ruiz-Velasco & Ikeda, 1998; Fernandez-Fernandez et al., 1999; Filippov et al., 2004). Currents were generated by voltage ramps (from -140 to -40 mV) and identified from their current-voltage relationship and inhibition by 100 µM to 1 mM Ba2+ (Filippov et al., 2004) (Fig. 6C).
Oxotremorine-M produced a biphasic effect: generation of an inward current, followed by a decline toward baseline (Fig. 6A, B). We attribute this to an initial activation of the GIRK current via M2-mAChRs (Fernandez-Fernandez et al., 1999) with subsequent inhibition via M1-mAChRs (Hill and Peralta, 2001). That the latter effect was attributable to inhibition of the M2-activated current (rather than desensitization) is confirmed by the fact that oxotremorine-M also inhibited GIRK currents preactivated by stimulating 2-adrenoceptors with 10 µM norepinephrine (data not shown) or by expressing G-protein 1
2-subunits (Fig. 6C) (Filippov et al., 2004). In the latter case, inhibition was measured in the presence of Pertussis toxin (to prevent M2-mAChR activation). Oxotremorine-M produced a concentration-dependent inhibition of the preactivated current of up to 60% of that inhibited by 1 mM Ba2+, with an IC50 value of 1.16 ± 0.14 µM (pooled data from four cells), quite comparable with that for M-current inhibition (compare with Fig. 3).
In PI5-K overexpressing neurons, the initial (M2) activation of the GIRK current noted in Figure 6 was unchanged, but the subsequent (M1) inhibitory effect was strongly reduced compared with that in control cells (Fig. 6A). When measured 30 s after the peak of the M2-activated current, inhibition was reduced by 57%, from 29.6 ± 3.3% (n = 12) to 12.8 ± 2.5% (n = 4) (Fig. 6B). We also tested the effect of bradykinin (100 nM), but this agonist failed to inhibit the GIRK current in any of six cells tested.
Quantification of membrane PIP2 levels from GFP-PLC-PH measurements
The PLC-PH peptide binds to both PIP2 and IP3 (Garcia et al., 1995; Lemmon et al., 1995). Hence, translocation from membrane to cytosol during PIP2 hydrolysis reflects (to differing extents) both the reduction of membrane [PIP2] and the rise in cytosolic [IP3] and indeed has been used as an index of both changes (Stauffer et al., 1998; Varnai and Balla, 1998; Hirose et al., 1999; Van der Wal et al., 2001; Nahorski et al., 2003). Notwithstanding, it is possible (in principle) to estimate individual changes in [PIP2] and [IP3] from translocation measurements (Xu et al., 2003). Because the inhibition of M current by oxotremorine-M is suggested to result from changes in [PIP2] rather than [IP3] (see the Introduction), we attempted to use our measurements of GFP-PLC-PH fluorescence to calculate both the resting concentrations of membrane [PIP2] and the changes in these concentrations after application of oxotremorine-M. For this, we used a different approach to the "model cell" approach used by Xu et al. (2003), based on (so far as possible) data obtained directly from our experiments on ganglion cells.
Resting membrane [PIP2] and the effect of PI5-K overexpression
We estimated resting membrane [PIP2] ([PIP2]Rest) using an intracellular IP3 competition assay. Starting from the situation in which most of the GFP-PLC-PH fluorescence is localized to the plasma membrane (where it is presumably bound to phosphatidylinositides, including PIP2), we patched neurons in open-tip configuration with pipettes containing different concentrations of IP3 and recorded the rise in cytosolic fluorescence (Fig. 7A). To "freeze" PIP2 concentrations at their prepatch level, we used an EGTA-buffered Ca2+-free intracellular solution, because Ca2+ is required for PLC function (Rhee, 2001).
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Figure 7. Displacement of membrane-associated GFP-PLC-PH by intracellular dialysis with IP3. A, Records showing the effect of breaking through with patch pipettes containing 0, 30, and 100 µM IP3 on cytosolic fluorescence in three sympathetic neurons expressing the GFP-PLC-PH construct. Initial fluorescence was measured over a cytoplasmic region of interest (Fig. 1) with the pipette in cell-attached configuration. Records show percentage change in this fluorescence after disrupting the patch (F/F0 x 100%, where F = F - F0, F is fluorescence after breakthrough, and F0 is baseline fluorescence before breakthrough). Inset, Images showing the distribution of fluorescence before and after pipette breakthrough. Note that, before breakthrough, fluorescence is localized primarily to the outer membrane but that, at the highest concentration of IP3, the fluorescence distributes uniformly with no clear membrane localization. B, Concentration dependence of IP3-induced change in cytosolic fluorescence F/F0 x 100% (A), where F is peak fluorescence after breakthrough. Each point represents a measurement in a separate neuron. C, Concentration-response curve for IP3-induced GFP-PLC-PH membrane-to-cytosol translocation. The mean fluorescence change in 0 [IP3](-40%) was subtracted from the individual values for F/F0 for each finite IP3 concentration in B and scaled to the mean value at 1 mM [IP3] (100%). Concentrations were corrected for phosphatase-induced IP3 breakdown as described in supplemental material 1 (available at www.jneurosci.org). Filled symbols indicate means and SEM of the transposed data points in B; open symbols indicate means and SEM of equivalent data at 30 and 100 µM [IP3] in cells preinjected with a cDNA plasmid encoding PI5-K. Curves are least-squares fits to the equation y = ymax · [IP3]/(IC50 + [IP3]). Mean values for IC50 were as follows: controls, 16.1 ± 1.24 µM; +PI5-K, 42.7 ± 9.4 µM.
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In the absence of intrapipette IP3, cytosolic fluorescence declined on breaking through with the pipette. This may be attributed to washout not only of resting cytosolic IP3 but also of any other diffusible molecules with polar groups that might bind GFP-PLC-PH, plus any free (unbound) GFP-PLC-PH. [There may also be some increase in the available "free space" (see below) as a result of disruption of cytoplasmic organelles by whole-cell patching.] With increasing concentrations of IP3 added to the pipette solution, there was a progressively increasing fluorescence above this zero-IP3 baseline (Fig. 7B). After baseline subtraction and scaling (Fig. 7C), the concentration-response curve yielded a mean EC50 value of 16 ± 1.2 µM. We then repeated these experiments in cells overexpressing PI5-K. For these tests, we used two concentrations of IP3, 30 and 100 µM. These concentrations clearly produced less translocation than in the control cells, giving an estimated EC50 value of 43 µM from the transposed concentrationresponse curve (Fig. 7C, filled circles).
With a small correction for intracellular IP3 breakdown (estimated at <5% at 3 µM intrapipette [IP3], <1% at >10 µM) (see supplemental material 1, available at www.jneurosci.org), the data in Figure 7C yielded the following values for the IP3 EC50 value (means, with lower and upper 95% confidence limits) (in µM): controls, 16.1 (11.9-23.5); +PI5-K, 42.7 (25.7-70.9). These results are broadly similar to those obtained by Van der Wal et al. (2001) in neuroblastoma cells using flash photolysis of caged IP3, in which IP3 concentrations between 10 and 100 µM were required to produce substantial translocation. The requirements for such high concentrations may seem surprising, because the dissociation equilibrium constant for IP3 binding to PLC-PH has been reported to be 0.1-0.2 µM, as opposed to 2 µM for binding to PIP2 (Lemmon et al., 1995; Hirose et al., 1999). The reason is that, clearly, the resting concentration of PIP2 (and other potential GFP-PLC-PH-binding phosphoinositides) in the membrane must be even higher.
The other principal PLC-PH-binding phosphoinositide is phosphatidylinositol-4-phosphate, which has a binding constant of 7 µM (Garcia et al., 1995) and is present at 80% of the concentration of PIP2 in neuroblastoma cell membranes (Willars et al., 1998) and 60% in human erythrocyte membranes (Ferrell and Huestis, 1984). We ignored phosphatidylinositol (PI) because, even if present in excess over PIP2 (Willars et al., 1998; Xu et al., 2003) (but see Ferrell and Huestis, 1984), phosphatidylcholine/phosphatidylserine (PC/PS) liposomes doped with 1% PI show very little extra binding of PLC-PH over PC/PS liposomes alone [Garcia et al. (1995), their Fig. 2]. We can also ignore other double-phosphorylated and triple-phosphorylated PIs, because, even though these bind PLC-PH with comparable affinity to PI(4,5)P (Garcia et al., 1995), their sum concentration is <1% of that of PI(4,5)P2 (Vanhaesebroeck et al., 2001).
The proportion of GFP-PLC-PH bound to IP3 (PIP3) is then given by the following standard competitive binding equation:
 | (1) |
where CIP3 = [IP3]/KIP3 CPIP2 = [PIP2]/KPIP2, and CPIP = [PIP]/KPIP, and KIP3, KPIP2, and KPIP are the dissociation equilibrium constants for GFP-PLC-PH to IP3, PIP2, and PIP, respectively. At the EC50 value for the IP3-induced rise in cytosolic fluorescence (when P1P3 = 0.5), this simplifies to the following:
 | (2) |
This was solved for [PIP2] by setting KIP3 = 0.1 µM, KPIP2 = 2 µM, KPIP = 7 µM, and [PIP] = 0.8[PIP2] (Table 1). The EC50 values for IP3-induced GFP-PLC-PH displacement in Figure 7 then yielded the following estimates for [PIP2] (means with upper and lower 95% confidence limits): controls, 261 µM (192-381); +PI5-K, 693 µM (417-1153).
These estimates of the initial (resting) levels of PIP2 assume that the instillation of IP3 does not significantly change the level of PIP2 in the membrane during the time required for the fluorescence measurements. This seems a reasonable assumption, because the slope of the [IP3]/translocation curve in Figure 7C was near unity, whereas elevation of membrane PIP2 would be expected to produce a progressive reduction in translocation and hence flattening of the curve. The absolute values will also depend critically on the ratio of the binding constants, but the proportionate change on overexpressing PI5-K would be unchanged.
We also used the intracellular IP3 displacement assay to assess the effect of NCS-1 on membrane [PIP2]. For this, we used a single (30 µM) concentration of intracellular IP3. Because NCS-1 is a Ca2+-binding protein, we measured the IP3-induced rise in cytosolic fluorescence using intracellular solutions containing 0 and 200 nM buffered [Ca2+]. Mean values (±SEM) for F/F0 (%) from the 0 [IP3] baseline were as follows: 0 Ca2+ - NCS-1, 59.8 ± 3.8 (n = 5); 0 [Ca2+] plus NCS-1, 48.8 ± 1.8 (n = 6); 200 nM [Ca2+] - NCS-1, 57.3 ± 2.0 (n = 6); 200 nM [Ca2+] plus NCS-1, 50.3 ± 2.3. None of these values were significantly different from the control values at 30 µM in Fig. 7. Thus, although there might have been some reduction of IP3 displacement fluorescence in NCS-1 overexpressing cells (which was not dependent on the absence or presence of 200 nM [Ca2+]), the effect of NCS-1 was much less than that of PI5-K overexpression (in which the corresponding F/Fo at 30 µM [IP3] was reduced to 30.1 ± 3.8%). At best, we estimate a maximum increase in resting membrane [PIP2] of 100-150 µM, in line with the data of Koizumi et al. (2002) and Rajebhosale et al. (2003) suggesting rises of 50 and 25%, respectively.
We emphasize that these calculated PIP2 concentrations are the effective concentrations in the plasma membrane as "seen" by the GFP-PLC-PH tag. Assuming they refer only to PIP2 in the inner (5-nm-thick) leaflet of the membrane, they equate to a density of 783 molecules of PIP2/µm2 membrane area if arranged as a monolayer, rising to 2079/µm2 after overexpressing PI5-K. These values are substantially less than previous estimates for neuroblastoma cells (4000/µm2) (Xu et al., 2003) based on measurements of total membrane phospholipid. In these measurements, PIP2 comprised 0.43% of total membrane phospholipid, whereas our measurements imply correspondingly lower values of 0.08%, rising to 0.22% with PI5-K. However, note the following: (1) the calculations of Xu et al. (2003) do not appear to allow for the area occupied by membrane protein, and (2) GFP-PLC-PH only bound to the plasma membrane (Fig. 1 A) not the intracellular membranes included in biochemical assays.
Changes in fluorescence after PIP2 hydrolysis
Stimulation of muscarinic acetylcholine receptors accelerates PIP2 hydrolysis and produces a fall in membrane [PIP2], with an equivalent fall in [PIP], presumably through conversion to PIP2 (Willars et al., 1998). Knowing the initial membrane concentrations of PIP2 and PIP, one can calculate the amount of IP3 formed as it is hydrolyzed and its concentration in the cytosol from the cytosolic/membrane volume ratio (CMVR) as follows:
 | (3) |
where [PIP2] = [PIP2]Rest - [PIP2] (the multiplicand 1.8 reflects the additional formation of IP3 by PIP through conversion to PIP2). The proportions of GFP-PLC-PH remaining in the membrane bound to PIP2 and PIP as the hydrolysis proceeds is then given by the following:
 | (4) |
and
 | (5) |
Of the GFP-PLC-PH that accumulates in the cytosol, most is bound to IP3, but some is free. These proportions are given by the following:
 | (6) |
and
 | (7) |
To solve these equations, we need to determine the following: (1) the cell size and cell volume available to IP3 to calculate CMVR (Eq. 3) and (2) the relationship between the measured increase in cytosolic fluorescence (F/F0) and proportion of total GFP-PLC-PH in the cytosol (i.e., the fractional translocation of GFP-PLC-PH from membrane to cytosol).
Cell volume and cytoplasmic:membrane volume ratio
We estimated the total cell volume from three-dimensional reconstructions of calcein-loaded neurons (see Materials and Methods). In a sample of five cells, the mean volume was 4289 ± 382 fl (SEM). This would correspond to a spherical cell of 10.1 µm radius and would yield a CMVR of 330, assuming a membrane thickness of 10 nm. However, not all of the intracellular space will be available to IP3. To estimate the volume of distribution of IP3, we compared the mean overall cell fluorescence in neurons patched with pipettes containing a low molecular weight hydrophilic fluorescent marker (BODIPY) with that in equivalent-sized aqueous vesicles (Fig. 8A) (see Materials and Methods). Intracellular fluorescence was clearly less than that expected from the vesicular fluorescence, suggesting that the volume of distribution of BODIPY was 25% of the total intracellular volume. Assuming that the volume of distribution of IP3 is similar to that of BODIPY, the effective CMVR in Equation 1 is then 83. This implies that, in a normal neuron (with membrane [PIP2] of 261 µM and [PIP] of 209 µM; see above), cytosolic [IP3] would rise to 5.82 µM from an assumed baseline value of 0.16 µM (Fink et al., 1999) when all PIP2 and PIP (after conversion to PIP2) is hydrolyzed. Again, this is not unreasonable, because this does not allow for any hydrolysis of IP3. There have been no direct measurements in sympathetic neurons; however, Fink et al. (1999) estimated a value of 2 µM in neuroblastoma cells after strong stimulation with bradykinin. However, these cells are appreciably larger than sympathetic neurons (giving a larger CMVR), and bradykinin is unlikely to produce complete hydrolysis of PIP2 (see below).
Relationship between cytosolic fluorescence increase and membrane-to-cytosol GFP-PLC-PH translocation
We assume that the increase in cytosolic fluorescence F/F0 produced by saturating concentrations of intracellular IP3 reflects total membrane-to-cytosol translocation of GFP-PLC-PH. This value (+86% from the zero-IP3 basel
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Abstract
Introduction
