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¹ Department of Physiology and ² Department of Neuroscience, University of Texas Southwestern Medical Center at Dallas, Dallas, TX 75390

Correspondence to Donald Hilgemann: donald.hilgemann{at}utsouthwestern.edu

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Baby hamster kidney (BHK) fibroblasts increase their cell capacitance by 25–100% within 5 s upon activating maximal Ca influx via constitutively expressed cardiac Na/Ca exchangers (NCX1). Free Ca, measured with fluo-5N, transiently exceeds 0.2 mM with total Ca influx amounting to 5 mmol/liter cell volume. Capacitance responses are half-maximal when NCX1 promotes a free cytoplasmic Ca of 0.12 mM (Hill coefficient 2). Capacitance can return to baseline in 1–3 min, and responses can be repeated several times. The membrane tracer, FM 4-64, is taken up during recovery and can be released at a subsequent Ca influx episode. Given recent interest in signaling lipids in membrane fusion, we used green fluorescent protein (GFP) fusions with phosphatidylinositol 4,5-bisphosphate (PI(4,5)P₂) and diacylglycerol (DAG) binding domains to analyze phospholipid changes in relation to these responses. PI(4,5)P₂ is rapidly cleaved upon activating Ca influx and recovers within 2 min. However, PI(4,5)P₂ depletion by activation of overexpressed hM1 muscarinic receptors causes only little membrane fusion, and subsequent fusion in response to Ca influx remains massive. Two results suggest that DAG may be generated from sources other than PI(4,5)P in these protocols. First, acylglycerols are generated in response to elevated Ca, even when PI(4,5)P₂ is metabolically depleted. Second, DAG-binding C1A-GFP domains, which are brought to the cell surface by exogenous ligands, translocate rapidly back to the cytoplasm in response to Ca influx. Nevertheless, inhibitors of PLCs and cPLA2, PI(4,5)P₂-binding peptides, and PLD modification by butanol do not block membrane fusion. The cationic agents, FM 4-64 and heptalysine, bind profusely to the extracellular cell surface during membrane fusion. While this binding might reflect phosphatidylserine (PS) "scrambling" between monolayers, it is unaffected by a PS-binding protein, lactadherin, and by polylysine from the cytoplasmic side. Furthermore, the PS indicator, annexin-V, binds only slowly after fusion. Therefore, we suggest that the luminal surfaces of membrane vesicles that fuse to the plasmalemma may be rather anionic. In summary, our results provide no support for any regulatory or modulatory role of phospholipids in Ca-induced membrane fusion in fibroblasts.

Abbreviations used in this paper: AMP-PNP, adenosine 5'-(β,-imido)triphosphoate; BHK, baby hamster kidney; DAG, diacylglycerol; PI(4,5)P₂, phosphatidylinositol 4,5-bisphosphate; PS, phosphatidylserine.

© 2008 Yaradanakul et al. This article is distributed under the terms of an Attribution–Noncommercial–Share Alike–No Mirror Sites license for the first six months after the publication date (see http://www.jgp.org/misc/terms.shtml). After six months it is available under a Creative Commons License (Attribution–Noncommercial–Share Alike 3.0 Unported license, as described at http://creativecommons.org/licenses/by-nc-sa/3.0/).

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Eukaryotic cells use precisely orchestrated membrane fusion and fission events to perform multiple cell functions (Mayer, 2002; Bankaitis and Morris, 2003; Howell et al., 2006). Fusion events in the secretory pathway are under the control of protein–protein interactions of SNAREs and associated proteins that are the subject of intense ongoing investigation (Koumandou et al., 2007). Among fusion events, the release of neurotransmitters from neurons attracts the most attention owing to its fast triggering by Ca binding to synaptotagmins (Geppert et al., 1994) and to its fundamental role in neuronal signaling (Katz, 2003). It is less well appreciated that other cell types use Ca-activated fusion processes for other functions (Breitbart and Spungin, 1997; Gundersen et al., 2002; Steinhardt, 2005; Czibener et al., 2006), and that those mechanisms allow analysis of the membrane specificity of fusion processes, the physical basis of membrane mixing during fusion, and the nature of sensors that initiate fusion. Even in neurons, asynchronous neurotransmitter release appears to be controlled by Ca sensors that are different from those used in fusion events closely coupled to Ca influx (Maximov and Sudhof, 2005; Sun et al., 2007). Of most interest for this article, many eukaryotic cells employ Ca-triggered membrane fusion as part of a membrane repair reaction initiated by Ca influx through cell surface wounds (Togo et al., 2000; Reddy et al., 2001; Steinhardt, 2005). Standard cell culture fibroblasts, such as CHO cells, can rapidly expand their surface membranes via membrane fusion in response to a large increase of cytoplasmic Ca (Coorssen et al., 1996).

The cytoplasmic Ca concentrations needed to trigger the cell wound response are evidently higher than those needed for neurotransmitter release (Schneggenburger and Neher, 2005). In fibroblasts, fusion appears to be initiated in the range of 10–30 µM free Ca (Coorssen et al., 1996). In sea urchin eggs, the threshold Ca concentration is 3 mM (Terasaki et al., 1997), just a few fold less than the Ca concentration of sea water. This Ca sensitivity is low enough so that Ca binding by anionic phospholipids, such as phosphatidylserine (PS) (Wilschut et al., 1981;Papahadjopoulos et al., 1990) and PIP(4,5)P₂ (Toner et al., 1988), might in principle play a triggering role, as described for pure phospholipid vesicles (Fraley et al., 1980; Wilschut et al., 1981). As stressed by others, however, Mg does not substitute for Ca in the cell wound response (Steinhardt et al., 1994; Steinhardt, 2005), whereas Mg binds nearly as well to anionic membranes as Ca until the vesicles are brought into close proximity (Feigenson, 1986, 1989). Work in diverse cell types suggests, overall, that the cell wound response involves both SNAREs and synaptotagmins that are related to those underlying neurotransmitter release, albeit with a complication that the wound response can probably involve multiple membrane types, including endosomes and lysosomes (Krause et al., 1994; Steinhardt et al., 1994; Bement et al., 1999; Detrait et al., 2000; Rao et al., 2004; Andrews, 2005; Andrews and Chakrabarti, 2005). From the functional effects of synaptotagmin fragments (Rao et al., 2004; Andrews, 2005; Andrews and Chakrabarti, 2005) and from knockdown studies (Jaiswal et al., 2004), synaptotagmin VII has been proposed to be the Ca sensor in the cell wound response. Dysferlins are another group of Ca-binding proteins with C2 domains that are being considered for a role in cell membrane repair responses, in particular in muscle (Bansal and Campbell, 2004; Washington and Ward, 2006).

In this article, we extend the analysis of Ca-induced membrane fusion in fibroblasts with baby hamster kidney (BHK) cells, using cardiac Na/Ca exchangers to activate a large Ca influx and to manipulate cytoplasmic Ca. For multiple reasons, it appeared important to us to characterize the function of Ca-dependent phospholipases, especially PLCs in the protocols that evoke membrane fusion in fibroblasts. First, phosphatidylinositol 4,5-bisphosphate (PI(4,5)P₂), the substrate of PLCs, can bind to and modulate the function of synaptotagmins (Tucker et al., 2003; Bai et al., 2004) and an associated protein, CAPS (Grishanin et al., 2004), which control membrane fusion in the active zones of neurons (Geppert et al., 1994). In addition, there appear to be requirements for PI(4,5)P₂ in one or more processes leading up to membrane fusion (Milosevic et al., 2005). Second, diacylglycerol (DAG), the lipid product of PLC activity, can modulate proteins involved in membrane fusion, in particular the munc proteins (Madison et al., 2005; Speight and Silverman, 2005; Latham and Meunier, 2006), and a DAG-dependent PKC can evidently facilitate the cell wound response (Togo et al., 2003; Steinhardt, 2005). In Xenopus oocytes, the Ca-dependent exocytosis of cortical granules upon fertilization appears to be initiated by the activation of PKCs by DAG (Gundersen et al., 2002). Both DAGs and phorbol esters can mimic Ca in activating fusion in oocytes, and activation of the Ca-independent PKC- is implicated to be important (Gundersen et al., 2002). Thus, it is possible that in the oocyte a PLC is the actual Ca sensor for this type of fusion response. In yeast, a novel PLC activity has been suggested to play an important role in homotypic vacuole fusion with DAG serving potentially as a second messenger that facilitates fusion (Mayer et al., 2000; Jun et al., 2004). Finally, it is noteworthy that phospholipases appear to have a fusion-promoting (fusogenic) influence in multiple membrane fusion processes (Harrison and Roldan, 1990; Haslam and Coorssen, 1993; Goni and Alonso, 2000; Choi et al., 2002; Brown et al., 2003; Choi et al., 2006). As described in this article, Ca influx indeed strongly activates PLCs in BHK cells in the same Ca range and on the same time scale in which membrane fusion occurs. In addition, we identify another pronounced membrane change. Extracellular binding of multiple cationic agents suggests that the extracellular cell surface rapidly becomes anionic during and after membrane fusion. We describe a wide range of experiments to address the possible roles of these changes in membrane fusion, and the results suggest that membrane fusion and phospholipid changes are independent processes that are triggered independently by large cytoplasmic Ca transients.

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Cell Lines and Transfections
BHK cells expressing NCX1 (Linck et al., 1998) with and without expression of hM1 receptors were maintained as previously described (Yaradanakul et al., 2007). BHK cells were grown in Dulbecco's modified minimum essential medium (DMEM) supplemented with 10% FBS with periodic selection by 2.5 mg/ml methotrexate for NCX1 and continuous selection with 200 µg hygromycin for hM1 receptors. Lipofectamine 2000 was used for DNA transfections following the manufacturer's protocols. Cells were harvested with 0.25% trypsin and kept at 35°C for 20 min in culture medium before experimentation.

Patch Clamp
Patch clamp and capacitance recording were performed as described previously (Yaradanakul et al., 2007). In brief, giga seals were established with 4–6 µm inner diameter pipette tips that were coated with dental wax before cutting and polishing to reduce capacitance. The majority of capacitance recordings were performed with our own software, described in an accompanying article (Wang and Hilgemann, 2008). Sinusoidal voltage oscillation (20 mV, 0.5–2 kHz) was usually employed when input resistances were <2 M, and square wave oscillation (20 mV, 0.5 kHz) was usually employed when input resistances were 1 M or higher. In the former case, the optimal phase angle was selected digitally via small changes of the optimal capacitance compensation parameters. In the latter case, single exponentials were fit to the falling phase of current transients, and input resistance, cell conductance, and cell capacitance were calculated online. As described in the accompanying article, the routines were validated by simulating typical cell electrical properties with the perturbation patterns employed, whereby cell parameters and parameter changes could be retrieved with an accuracy of about 99.9%. A few recordings employed phase-lock amplifiers, as described previously (Yaradanakul et al., 2007), and we detected no qualitative or quantitative differences between capactiance recordings performed with the two methods.

For experiments without imaging, the voltage-clamped cells were moved rapidly between four parallel solution streams maintained at 35°C. For experiments with simultaneous confocal imaging, a commercial recording chamber (RC-26; Warner Instruments) was employed in conjunction with a Nikon TE2000-U microscope, and for extracellular solution changes four solution lines with on/off switches were merged to a single temperature-controlled outlet.

Solutions, Chemicals, and Constructs
The standard extracellular solution for maximal outward Na/Ca exchange current contained (in mM) 110 NMG, 20 TEA-OH, 15 HEPES, 3 MgCl₂, and 0.5 EGTA, set to pH 7.0 with aspartate, with activation of current by substituting 2 MgCl₂ for 2 CaCl₂. The standard cytoplasmic solution contained (in mM) 80 NMG, 40 NaOH, 20 TEA-OH, 15 HEPES, 0.5 EGTA, 0.25 CaCl₂, 0.5 MgCl₂, 1.6 MgATP, 0.4 TRIS-ATP, and 0.2 MgGTP, set to pH 7.0 with aspartate. For Figs. 3 –5, 80 mM NMG was replaced in both solutions with 80 mM LiOH. In Fig. 3, 40 mM LiOH was replaced with 40 NaOH in the extracellular solution used to activate exchange current. In the experiments for Figs. 6 and 7, the cytoplasmic solution contained (in mM) 60 NaOH, 30 KOH, 30 TEA-OH, 0.5 MgCl₂, 0.5 EGTA, 0.25 CaCl₂, 30 HEPES and set to pH 6.9 with aspartate. For experiments with inward exchange current in Fig. 13 C, the extracellular solution contained (in mM) 120 NaOH (or LiOH), 1.5 EGTA, 26 TEA-OH, 20 HEPES, and 1 MgCl₂, set to pH 7.0 with aspartic acid. The cytoplasmic solution contained (in mM) 120 LiOH, 20 TEA-OH, 1 MgCl₂, 3.0 EGTA, CaCl₂ to achieve the desired free Ca, 15 HEPES, 2 ATP, and 0.2 GTP, set to pH 6.9 with aspartic acid. Rhodamine-conjugated hepta-lysine (rhodamine-KKKKKKK-amide; K7-Rhod) was prepared by Multiple Peptide Systems (NeoMPS, Inc.). Annexin-V Alexa Fluor 488 (#A13201) conjugate was from Invitrogen and was employed at 1:100 dilution. All other chemicals were from Sigma-Aldrich and were the highest purity available. The PLC1PH-GFP fusion protein construct was a gift of Tobias Meyer (Stanford University, Stanford, CA), and the construct for C1A-GFP fusion protein (Oancea et al., 1998) was provided by Mark Shapiro (UT San Antonio).

Imaging
Confocal imaging was as described (Yaradanakul et al., 2007) with a Nikon TE2000-U microscope and a Nikon 60x 1.45 NA oil immersion objective. A 40 mW Spectra Physics 163-CO2 laser was used for 488/514 nm excitation (i.e., for GFP constructs and FM dye); a 1.5 mW Melles Griot cylindrical HeNe Laser was used for 543.5 nm excitation (i.e., for rhodamine constructs). The time lapse images were recorded either at 160 x 160 or 256 x 256 resolution with an 400-ms total exposure time. The exposure interval was 3 s in all records presented. Lasers were operated at 1% power with a detector pinhole setting of 60 µm and an average linear gain of 7.00. Fluroescence intensity is quantified in most figures as percent of maximal fluorescence occurring during the recording (%).

Lipid Analysis
Anionic phospholipid mass measurements were performed as previously described (Nasuhoglu et al., 2002a). The total mono- and diacylglycerols were determined via a bacterial DAG kinase that phosphorylates both mono and diglycerides (Preiss et al., 1987). To do so, phospholipids were first extracted, washed, and dried by the same procedures used for other mass determinations. They were then dissolved in 50 µl of a solution containing 0.3 mM tetradecylsulfate and 0.6% Triton X-100. An equal amount of DAG kinase reaction buffer (2 mM ATP, 2 mM DTT, 100 mM imidazole, 100 mM NaCl, 25 mM MgCl₂, 2 mM EGTA, pH 6.6) was then added to separate samples with and without 10 µg/ml DAG kinase (Calbiochem, #266726). The aliquots were reacted to completion in 30 min at 30°C, phospholipids were again extracted and analyzed as previously described (Nasuhoglu et al., 2002a), and the total mono- and diacylglycerol was determined from the increase of glycerol phosphate in HPLC analysis of deacylated phospholipids treated versus not treated with DAG kinase.

Electron Microscopy
The BHK cell preparation for electron microscopy was performed largely as described for other cell types (Sugita et al., 2001). Cells were removed from dishes, as described above, and they were spun down to form loose pellets before fixing in 2% glutaraldehyde with 1% sucrose in 0.1 M sodium cacodylate buffer (pH 7.4) at 37°C for 2 h. The pellets were then carefully separated and rinsed twice in buffer and post-fixed in 0.5% OsO₄, 0.8% K-ferricyanide in the same buffer for 30 min at room temperature. After rinsing with distilled water, specimen were stained in bloc with 2% aqueous uranyl acetate for 15 min, dehydrated in ethanol and embedded in Poly/bed 812 for 24 h. Thin sections (65 nm) were post-stained with uranyl acetate and lead citrate, and they were viewed with an FEI Tecnai G2 Spirit Biotwin transmission electron microscope.

	RESULTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
BHK Cell Response to Maximal NCX1-mediated Ca Influx
Fig. 1 presents a typical electrophysiological recording from an NCX1-expressing BHK cell during repeated brief activation of the maximal outward exchange current when the cytoplasm was weakly Ca buffered. Whole-cell current, capacitance, and conductance are monitored as described in Materials and methods with a 20-mV sinusoidal voltage oscillation at 500 Hz. The cytoplasmic solution contains 2 mM ATP, 0.2 mM GTP, 40 mM Na, and 0.5 mM EGTA with 0.25 mM Ca (0.4 µM free Ca), and the extracellular solution is Na free. The outward current is activated by switching from an extracellular solution with 0.5 mM EGTA and no Ca to one with 2 mM Ca (i.e., 1.5 mM free Ca; see Materials and methods for further details). Panel A shows the entire experimental records over 6 min in which the exchange current was activated and deactivated four times. Panel B shows the records during each activation cycle at higher time resolution, together with the calculated capacitance derivative (dCap/dt).

View larger version (14K): [in this window] [in a new window] [Download PPT slide]   Figure 1. Typical records of membrane capacitance, conductance, and current from BHK cells consitutively expressing the cardiac Na/Ca exchanger (NCX1). Cell parameters are monitored via 20-mV sinusoidal membrane voltage perturbation at 0.5 kHz. Input resistance 1 M. Cytoplasmic solution contains 40 mM Na and 0.5 mM EGTA with 0.25 mM Ca (free Ca, 0.4 µM). Exchange current was activated four times for increasing durations of 1–6 s by applying and removing 2 mM extracellular Ca, as indicated below the records. (A) Complete signal records. (B) Expanded signal records for the time periods during and immediately after activation of exchange current. dCap/dt is the first derivative of capacitance. Two vertical dotted lines in each activation cycle mark the time when current was activated and the time at which the maximal rate of rise of capacitance occurred. Note that the rise of capacitance is preceded by a small negative capacitance phase. The time from current activation to peak rate of rise of capacitance increases as the peak exchange current decreases in this sequence.

During Ca influx, the cell conductance increases by 20 nS within 700 ms and then decreases partially toward baseline. The irreversible component of this increase probably reflects a decrease of seal resistance. The transient part of the response may in part reflect exchange current activity, but several arguments suggest that it reflects mostly conductive properties of fusion pores (Lindau and Alvarez de Toledo, 2003; Neef et al., 2007). First, the transient conductance changes do not track exchange current. Conductance rather rises slowly after exchange current has been completely activated. Second, the outward NCX1 current under the conditions of these experiments (i.e., with 120 mM extracellular NMG and no other extracellular monovalent cations) is nearly voltage independent (Matsuoka and Hilgemann, 1992; Matsuoka and Hilgemann, 1994) and therefore is not expected to contribute substantial conductance changes. Third, the slow rise of conductance correlates roughly in time with the rate of rise of capacitance (i.e., dCap/dt), and the peaks of these signals occur at approximately the same time, as denoted by a second dotted vertical line in each response.

The peak exchange current typically decreased by at least 40% from one Ca pulse to the next with Ca pulse durations of 1–5 s. Also evident in these records, the rise of capacitance typically occurred with a longer delay at the second and later responses when the exchange current was decreased. Nevertheless, the second increase of capacitance was typically larger and occurred with a higher maximal rate than during the first response. This is consistent with reports that the cell wound response shows facilitation when induced multiple times (Togo et al., 2003). As in most experiments, the same magnitude of capacitance increase was achieved at the third and fourth Ca pulses when the Ca pulse period was increased to compensate for the decreased exchange current.

We stress that both the magnitudes of capacitance increase and the extent to which capacitance signals reversed to baseline were somewhat variable and showed statistically highly significant dependence on cell batch and growth conditions. The average increase of capacitance was found to vary by up to fivefold between cell batches with similar basal capacitance and exchange current density. For example, we verified in two datasets that removal of serum from cells for 24 h caused a highly significant decrease of the capacitance response by >50%, while the exchange current density was unchanged. In 50% of cell batches, cell capacitance did not reverse completely, as occurs in Fig. 1, and in 25% of cases the cell capacitance decreased to values lower than the initial cell capacitance before activating Ca influx. Rarely, reversal amounted to only a small fraction of the initial capacitance increase, and capacitance increments were then very small at the second to fourth Ca pulses. Finally, we mention that capacitance responses in CHO and HEK293 cells were typically smaller than those of BHK cells when equivalent NCX1 expression was achieved by transient or stable expression methods. Responses in CHO cells seldom exceeded 40% of basal cell capacitance, and responses in HEK293 cells were usually close to 15%.

Current and Ca Dependencies of Capacitance Responses
Using the protocol described in Fig. 1, we proceeded to analyze the exchange current and Ca dependencies of the capacitance responses. As evident in Fig. 1 B, capacitance increases smoothly during the same time period in which exchange current is deactivated by removing extracellular Ca. This observation indicates that Na/Ca exchangers do not generate a local Ca signal that is critical for this type of membrane fusion. To determine the mean free Ca concentrations occurring in cells, we employed the low affinity Ca indicator dye fluo-5N (Invitrogen) at a concentration of 3 µM in the pipette solution. Fig. 2 A shows a typical fluorescence record obtained during NCX1 activation under the same conditions as Fig. 1 (0.5 mM EGTA with 0.25 mM total Ca, 0.4 µM free Ca). After recording background fluorescence for 1.5 min, the cell was exposed to 2 mM extracellular Ca for 3 s, as is usual with substitution for 2 mM Mg, fluorescence signals were allowed to dissipate for 30 s. Then, the same cytoplasmic solution was perfused into the cell with the dye saturated by 1 mM free Ca (i.e., 1.5 mM total Ca with 0.5 mM EGTA) to determine the maximal Ca response of the dye in the cell. Peak free Ca induced by Ca influx was then estimated assuming a K_d for Ca of 90 µM with free Ca calculated as K_d/(F_max/F–1), where F_max is the cell fluorescence with Ca-saturated fluorophore and F is the peak fluorescence obtained in response to activating Ca influx.

View larger version (19K): [in this window] [in a new window] [Download PPT slide]   Figure 2. Measurement of cytoplasmic free Ca changes during activation of outward Na/Ca exchange currents in BHK cells. The pipette solution contains 0.5 mM EGTA and 0.25 mM Ca (i.e., 0.4 µM free Ca) with 3 µM Fluo 5N (Kd, 90 µM). Here, and in all figures, Ca concentrations given indicate the free Ca concentration. (A) Typical fluorescence and current records. When 2 mM Ca is applied for 4 s, current rises rapidly to a peak of 175 pA and decays to baseline within <1 s when Ca is removed. Peak fluorescence occurs at 3 s and begins to decay rapidly before current is deactivated. Upon perfusion of the pipette with 1 mM additional Ca, fluorescence rises to a steady "maximal" level with a time constant of 20 s. (B) Free and total Ca are calculated from the fluorescence and current records as described in the text. Peak free Ca is estimated to be 0.20 mM at a time point when total Ca influx amounts to 3.0 mmol/liter cell volume. (C) Peak free and total Ca influx values obtained 14 control cells and 5 cells perfused with 2 µM thapsigargin and 2 mM AMP-PNP to block Ca pumping. Results are not significantly different.

From experiments in which we employed BHK cell lines with different NCX1 current densities, our impression was that the rate and extent of capacitance increase were nearly proportional to the current density. As described in Fig. 3, we analyzed quantitatively the relationships between peak current density and capacitance changes by varying extracellular Ca. By performing these experiments in the presence of 40 mM Na on both membrane sides, at 0 mV, the maximal free cytoplasmic Ca is thermodynamically limited to not exceed the free Ca applied to the extracellular side. Fig. 3 (A and B) shows representative results for applying 100 and 10 µM free Ca together with 40 mM extracellular Na. We quantified the rate of rise of capacitance as the fractional increase of cell capacitance per second (dCap/dt/Cap, i.e., dCap/dt divided by the basal cell capacitance). Fig. 3 C presents the dependence of capacitance change on the extracellular Ca concentrations employed (i.e., the maximal possible cytoplasmic free Ca), and Fig. 3 D presents the dependence on the peak current density. The maximal fractional rate of rise of capacitance is 0.3 per second (i.e., 30% per second). In both plots, the rate of rise of capacitance shows a sigmoidal dependence on current with a Hill coefficient of 2. Half-saturation occurs at 140 µM free extracellular Ca and 8 pA/pF, respectively. From these results and knowledge of the peak Ca/peak current density relationship from Fig. 2, we conclude that a free Ca concentration >10 µM is required to achieve 10% of the maximal fusion rate, and that a concentration >100 µM is required to achieve the half-maximal fusion rate. We mention that the capacitance delay, defined as the time from activation of current to the time at which capacitance crossed the zero line (see Fig. 3 B), decreased with increasing current density in an almost linear fashion (unpublished data).

View larger version (18K): [in this window] [in a new window] [Download PPT slide]   Figure 3. Dependence of BHK cell capacitance responses on the extracellular Ca concentration and exchange current density (0 mV, 35°C). (A) Typical current, capacitance, and conductance records during application of 0.1 mM free Ca together with 40 mM Na, thereby limiting the maximal free cytoplasmic Ca concentration to 0.1 mM. (B) Typical current, capacitance, and conductance records during application of 10 µM free Ca together with 40 mM Na, thereby limiting the maximal free cytoplasmic Ca concentration to 10 µM. (C) Maximal rate of capacitance increase as fractional increase of total cell capacitance per second (dCap/dt/Cap) in dependence on the applied extracellular free Ca. (D) Maximal fractional rate of capacitance rise in dependence on peak exchange current density.

View larger version (12K): [in this window] [in a new window] [Download PPT slide]   Figure 4. Capacitance and current record of a BHK cell expressing NCX1 in perforated patch whole cell configuration. Same solutions as previous figures with 40 µM β-escin in the pipette solution and capacitance recording via square wave perturbation. Error bars represent the results of 13 similar experiments.

View larger version (12K): [in this window] [in a new window] [Download PPT slide]   Figure 5. Effects of nonhydrolyzable ATP analogue and protein phosphatase inhibitors on the BHK cell capacitance response and exchange current rundown. (A) Capacitance, conductance, and current records from a BHK cell perfused with 2 mM AMP-PNP for 4 min before activating exchange current. (B) Capacitance changes and peak exchange current densities for control cells, cells perfused with 2 mM AMP-PNP, cells perfused with 2 mM AMP-PNP, and with 5 µM cyclosporin for 4 min before activating exchange current, and cells perfused with AMP-PNP with 9 mM pyrophosphate and 4 mM fluoride.

View larger version (53K): [in this window] [in a new window] [Download PPT slide]   Figure 6. PLC activation monitored via PLC1PH-GFP (panel I) and C1-GFP (panel II) protein fusions in BHK cells voltage clamped to 0 mV via large-diameter pipette tips. The continuous records give the average fluorescence in a central region of the cytoplasm. As evident in micrographs, shown above the cells, the large-diameter tips promote bleb formation. (Panel I, A) The PLC1PH-GFP domain, initially localized mostly to the surface membrane, rapidly translocates to the cytoplasm during carbachol (0.3 mM) application for 20 s. When agonist is removed, PH domains return to the cell membrane with a time constant of 32 s. (B) Similar PH domain responses recorded when cells are loaded with high Na (40 mM) and Ca influx via NCX1 is activated for 12 s by applying and removing 2 mM extracellular Ca. (C) Comparison of PH domain signals in a cell activated first by Ca influx, then by carbachol, and finally again by Ca. (Panel II, D) The C1 domain is initially cytoplasmic and then rapidly translocates to the membrane during carbachol (0.3 mM) application for 20 s. (E) Slower and smaller C1 domain response, typically obtained during activation of outward NCX1 current. (F) Typical C1 domain cytosolic fluorescence changes when carbachol (0.3 mM) is applied continuously and Ca (2 mM) is then applied for 15 s together with carbachol, followed by washout. C1-GFP domains dissociate rapidly from the plasma membrane during Ca influx and accumulate diffusely in the cytoplasm.

In response to carbachol (Fig. 6 A), cytoplasmic fluorescence rises by fourfold within 20 s as PLC1PH-GFP domains are lost from the cell surface. Upon removal of carbachol, the amount of cytoplasmic fluroescence begins to decrease within 5 s and returns to baseline in a nearly exponential fashion with a best-fit time constant of 32 s. In response to activation of outward NCX1 current for 12 s (Fig. 6 B), PLC1PH-GFP domain responses of similar magnitude and kinetics are recorded. As shown in Fig. 6 C, the magnitudes of responses to activation of NCX1 were typically at least as large as the responses to carbachol in the same cell, and responses could be repeated multiple times with only little loss of signal magnitude.

From these results, it is clear that Ca influx can activate large PLC responses in BHK cells. To determine more accurately the Ca range over which PLCs become activated, we analyzed PH domain distributions in dependence on the inward exchange current in the presence of different free Ca concentrations in the pipette solution. From previous experiments, it is clear that free cytoplasmic Ca decreases significantly during activation of inward current by extracellular Na, even in the presence of several millimolar EGTA buffering (Yaradanakul et al., 2007). Since the free cytoplasmic Ca cannot exceed the free Ca of the patch pipette solution, protocols with inward current can give a clear indication of the minimum Ca required to activate PLCs in the absence of receptor activation. To summarize briefly results from >20 experiments, removal of extracellular Na to deactivate inward current caused clear PLC activation when cells were perfused with solutions containing free Ca of 20 µM or higher. When cells were opened with free Ca of 8 µM or less, the activation and deactivation of inward exchange currents had no evident effect on the PH domain distribution in cells (unpublished data). Thus, we conclude that free Ca in the range of 10 µM is needed to significantly activate PLCs in the absence of GPCR activation.

Given that high free Ca can strongly activate PI(4,5)P₂ breakdown in BHK cells, it may be expected that PLC activity will contribute to NCX1 current rundown in the protocols with massive Ca influx. However, as just described, the PH domains redistribute reversibly in response to both carbachol and Ca changes on a rather fast time scale. By comparison, the suppression of outward exchange current in response to Ca influx episode is very long-lived and often does not reverse at all (Figs. 1 and 4). Thus, PI(4,5)P₂ depletion cannot be the sole or even the major mechanism of long-term exchange current inactivation and rundown in the experiments described.

Dual C1A Domain Responses to a Rise of Cytoplasmic Ca
In panel II of Fig. 6, and in Fig. 7, we describe responses of the DAG-binding C1A-GFP fusion proteins (Oancea et al., 1998). Fig. 6 (D and E) shows the usual signals observed for carbachol and NCX1 activation, respectively (>20 observations), and F illustrates a major complexity encountered. As shown in D and E, the C1 domain appears almost uniformly distributed throughout the cytoplasm in unstimulated, voltage-clamped BHK cells. When M1 receptors are overexpressed, as in all three cells depicted, application of carbachol (0.3 mM) results in the association of these domains with the membrane within 20 s. Upon removal of carbachol, the domains reequilibrate into the cytoplasm with somewhat larger time constants (90 s) than those just described for PH domains. Thus, metabolism of DAG by lipases and kinases may be somewhat slower than resynthesis of PI(4,5)P₂ (i.e., phosphorylation of PI and PIP). As apparent in Fig. 6 E, activation of Ca influx by NCX1 causes a qualitatively similar response to that of carbachol, as anticipated, but the extent of the responses and the rate of rise of the responses were consistently less than for carbachol. As shown in Fig. 6 F, the blunted response of C1A domains to Ca elevation reflects a second, strong opposing influence of Ca on the distribution of C1 domains. This opposing influence is vividly revealed when the C1 domains are first brought to the membrane by carbachol, and then extracellular Ca is applied to induce a rise of cytoplasmic Ca in the continued presence of carbachol. As shown in Fig. 6 F, Ca influx in the presence of carbachol results in a rapid return of C1 domain to the cytoplasm whereby a nearly uniform distribution develops with a time constant of just 15 s.

View larger version (28K): [in this window] [in a new window] [Download PPT slide]   Figure 7. Translocation of C1-GFP domains to the surface membrane of BHK cells by short-chain DAG8 (A), phorbol ester (B), and carbachol (C), followed by redistribution to the cytoplasm in all three cases by activation of Ca influx (i.e., outward NCX1 current). (A) Application of 20 µM C8 diacylglycerol brings the majority of C1 domain to the surface membrane with a time constant of 45 s. Upon activating reverse exchange current, domains return quantitatively to the cytoplasm with a time constant of 35 s. (B) Application of 0.2 µM phorbol ester (PMA) brings C1 domains to the cell surface with a time constant of 30 s. During activation of exchange current, C1 domains reach a substantially higher concentration in the cytoplasm than in the basal state with a time constant of 30 s, and C1 domains begin to move back to membrane immediately upon terminating Ca influx. (C) C1-GFP domain records together with cell capacitance and membrane current in a BHK cell expressing NCX1 and hM1 receptors. With application of carbachol (0.3 mM) cytoplasmic fluorescence decreases by 60% with a time constant of 20 s, and cell capacitance increases modestly. Upon activation of Ca influx, cell capacitance increases by 8 pF (30%) in less than 10 s, and C1 domains accumulate again in the cytoplasm with time constant of 10 s. Upon deactivating exchange current, C1-GFP domains move partially back to the surface membrane.

Phospholipid Changes in Response to Changes of Cytoplasmic Ca
We can suggest four mechanisms by which high cytoplasmic Ca may cause translocation of C1 domains from the cell surface to the cytoplasm, and we attempted to eliminate these one by one in appropriate experiments. First, DAG metabolism may be Ca dependent, a major mechanism being that some DAG kinases are activated by Ca (Jiang et al., 2000a; Luo et al., 2004; Topham, 2006). Second, C1 domains interact not only with DAG but also with negatively charged phospholipids, especially PS (Bittova et al., 2001; Ho et al., 2001), which can bind Ca (Wilschut et al., 1981; Papahadjopoulos et al., 1990) and thereby might cause release of C1 domains from the membrane. Third, DAG might be generated by PLCs on internal membranes by multiple Ca-dependent mechanisms, thereby favoring translocation of C1 domains to the cytoplasm. And fourth, DAG might translocate to the outside of the cell during membrane fusion, assuming that phospholipid mixing between bilayers occurs during fusion, thereby decreasing DAG on the inner membrane leaflette and releasing C1 domain back to the cytoplasm. In this light, we performed both fluorescence and biochemical studies to determine how a rise of cytoplasmic Ca affects phosphoinositides, DAG, and phosphatidic acid, and how those effects depend on cytoplasmic ATP.

Three types of experiments address whether Ca-activated DAG kinase activity might play a role in the C1 domains translocation to the cytoplasm in response to Ca influx. The first type of evidence is represented by the PMA experiment in Fig. 7. PMA is not phosphorylated by DAG kinases, and yet activation of Ca influx causes C1 domains to move to the cytoplasm after being brought to the cell surface by PMA. In this response, DAG kinase activity at the cell membrane cannot be the cause of the C1 domain movement. Second, we tested for inhibition of C1 translocation by the two available DAG kinase inhibitors at concentrations recommended for strong inhibition (DAG kinase inhibitors I and II [Calbiochem]; R59022 and R59949, each at 10 µM). We found no inhibition of the C1 translocation by either of these agents (unpublished data; two observations each) upon activating Ca influx. These results alone are not conclusive because not all DAG kinases are inhibited by these agents (Jiang et al., 2000b), and we report in this same connection that a substantial rise of phosphatidic acid that occurs in parallel with loss of phosphatidylinositol during M1 receptor activation (Li et al., 2005) was unaffected by the DAG kinase inhibitor, R59022, at a concentration of 20 µM. Third, we performed the protocols of Fig. 7 with 2 mM AMP-PNP and no ATP in the pipette solution, and the C1 domain still translocated rapidly to the cytoplasm when Ca influx was activated (two observations). On this basis, the activation of DAG kinase activity at the cytoplasmic cell surface by a rise of cytoplasmic Ca seems eliminated as a cause of the C1 domain translocation.

As described in Figs. 8 and 9, we measured phospholipid changes in response to increasing cytoplasmic Ca in two different approaches using BHK cells. In the first approach, we used NCX1 to mediate a large Ca influx. To do so, the ionophore, nystatin (25 µM), was applied to load cells with Na in Ca-free PBS (130 mM NaCl, 20 mM HEPES, 15 mM glucose, pH 7.4) for 10 min. Thereafter, 4 mM Ca was added to the extracellular medium for 1 min without addition of Na (130 mM TEA-Cl). Anionic phospholipids were then determined as described, and the relative mass of the four phospholipids of most interest (PIP, PI(4,5)P₂, PI, and PA) are shown in Fig. 8. Calibrations are given as percent of total anionic phospholipid, whereby the majority of anionic phospholipid is PS and cardiolipin. The nystatin treatment resulted in a 30% reduction of PI(4,5)P₂ in the absence of Ca, and there was little or no change of other phospholipids. The Ca addition caused PIP to fall by 70% and PI(4,5)P₂ to fall by 50%, as expected for PLC activation. During the same 1 min, PI fell from 46.3 to 44.3% of total anionic phospholipid, while PA increased from 12.8 to 17.4% of total phospholipid. As with receptor activation, the fact that PI mass decreases and PA mass increases by similar amounts, more than threefold greater than changes of PIP and PI(4,5)P₂, suggests that there is a substantial flux of phospholipid from PI to PIP to PI(4,5)P₂ to DAG and finally to PA in this protocol. In summary, these results for cytoplasmic Ca elevation are qualitatively very similar to, but quantitatively less drastic, than published results for M1 receptor activation in cells with constitutive receptor expression (Li et al., 2005).

View larger version (20K): [in this window] [in a new window] [Download PPT slide]   Figure 8. Phosphoinositide and phosphatidate changes in BHK cells in response to Ca influx via NCX1. The bar graphs represent the relative phospholipid concentrations, as percent of total anionic phospholipid, for cells under control conditions (control), cells treated with 25 µM nystatin in the absence of Ca for 10 min (nystatin – Ca), and nystatin-treated cells that were exposed to 4 mM extracellular Ca for 1 min (nystatin + Ca).

View larger version (22K): [in this window] [in a new window] [Download PPT slide]   Figure 9. Phosphoinositide, phosphatidate, and total mono/diacylglycerol (M/DAG) levels of permeabilized BHK cells in dependence on ATP, AMP-PNP, and Ca. (A) PIP and PI(4,5)P2 levels of BHK cells permeabilized with 40 µM β-escin. All results are the average of two to four measurements. Left bars give levels for control cells and permeabilized cells with 2 mM ATP and no Ca (0.5 mM EGTA). The set of three bars in the middle give PIP and PI(4,5)P2 levels with ATP, without ATP for 5 min, and with 2 mM AMP-PNP for 5 min. The right two bars give phospholipid levels of control cells and cells exposed to 0.4 mM free Ca for 2 min. (B) From left to right, the bar graphs give mono and diacylglycerol (M/DAG) and phosphatidate (PA) levels without and with 0.4 mM free Ca for 1 min (left four bars), without and with 0.3 mM carbachol for 3 min, without and with 0.4 mM free Ca together with 0.3 mM carbachol, and without and with 0.4 mM free Ca for 1 min in the presence of AMP-PNP and no ATP (right four bars).

Fig. 9 B shows our measurements of acylglycerols (M/DAG) and phosphatidic acid, using diacylglycerol kinase to generate phosphatidate and lysophosphatidate from acylglycerols (Preiss et al., 1987). In preliminary experiments, we established that the bacterial DAG kinase employed in this assay converted both 1-O-palmitylglycerol and dipalmitylglycerol, in amounts >3-fold greater than determined for cell lysates to phosphorylated metabolites that were recovered quantitatively as glycerol phosphate in our HPLC assay of anionic phospholipid metabolites. As shown in Fig. 9 B, the total di- and monocylglycerol content of BHK cells is similar to that of phosphatidate, namely 6–8% of total anionic phospholipid. For orientation, this is six to eight times more than the content of PI(4,5)P₂. As described in the left four bars, the addition of Ca to generate a free concentration of 0.4 mM in the extracellular medium, containing 2 mM MgATP, caused a 30% decrease of acylglycerol and a 15% increase of phosphatidate. These changes are consistent with diacylglycerol kinases being activated by Ca in these cells. Using cells with M1 receptors, we did not obtain significant effects of carbachol (0.3 mM) on acylglycerol levels, while phosphatidate increased by 15%. In response to the combined application of 0.4 mM free Ca and carbachol (0.3 mM), acylglycerol levels increased by 40% and phosphatidate levels increased by 24%. Finally, as shown in the right four bars of Fig. 9 B, incubation of cells with AMP-PNP (2 mM) instead of ATP caused a decrease of acylglycerol. In the presence of AMP-PNP, application of Ca (0.4 mM) for 1 min caused an increase of acylglycerol by 12% with no change of phosphatidate. While the increase is small, as a relative change, the absolute change is in fact larger than the entire PI(4,5)P₂ mass of the cells, and we point out that the increase of acylglycerol was accompanied by a significant decrease of phosphatidylinositol (unpublished data). Since PIP and PI(4,5)P₂ are depleted and cannot be synthesized in this condition, phospholipases evidently cleave significant amounts of phosphatidylinositol. In summary, these biochemical data demonstrate first that high cytoplasmic Ca indeed promotes generation of phosphatidic acid by activating DAG kinases, although this activity cannot explain the translocation of C1 domains into the cytoplasm described in Figs. 6 and 7. Second, they demonstrate that Ca might indeed support the generation of DAG on internal membranes from sources other than PI(4,5)P₂ in the circumstances of Figs. 6 and 7.

Further Evidence against Roles for PLC, PLD, and cPLA2 Activities, as well as other Ca-dependent Processes, in Membrane Fusion
Using the BHK cell line with high NCX1 expression, we tested a wide range of agents expected to change phosphatidylinositide metabolism, bind phosphatidylinositides, and/or inhibit phospholipases. All negative results reported here without figures reflect at least three and usually six or more control and treatment observations. We found no significant inhibition of membrane fusion or the rate of fusion by any of the following agents when included in the pipette solution and when fusion was quantified as a percent change of capacitance and related to the peak exchange current density: U73122 (10 µM), edelfosine (10 µM), nitrocoumarin (20 µM), neomycin (30 µM), heptalysine (40 µM), and 2000 MW polylysine (20 µM) to inhibit PLCs and/or bind PI(4,5)P₂ and other anionic phospholipids (Ben-Tal et al., 1996; Bucki et al., 2000), IP₃ (0.1 mM) to release PLC-d from the membrane (Cifuentes et al., 1994), wortmannin (4 µM) to inhibit PI3- and type III PI4-kinases (Nakanishi et al., 1995), adenosine (1 mM) to inhibit type II PI4-kinases (Barylko et al., 2001), phosphatidylinositol transfer protein from yeast (0.1 mg/ml) to remove phosphatidylinositol from cell membranes (Routt and Bankaitis, 2004), recombinant C1A domains (20 µM) to bind DAG (Colon-Gonzalez and Kazanietz, 2006), 0.6% butanol to stop DAG generation from phosphatidate (Choi et al., 2002), and a putatively specific phenylacrylamide cPLA2 inhibitor (Calbiochem #525143, 2 µM; Seno et al., 2000). The fact that exchange currents are not inhibited by agents that bind anionic phospholipids may appear surprising. The likely reason is that in these protocols there are no extracellular NCX1 ligands present until Ca is applied. Therefore, exchangers orient with ion transport sites open to the outside, and this configuration does not support the Na-dependent inactivation (Matsuoka and Hilgemann, 1994). We also tested for possible roles of other Ca-dependent processes. A possible role of Ca-activated microfilament depolymerization seems unlikely because phalloidin (5 µM) had no significant effect when perfused into cells for 5 min before activating Ca influx. A role of Ca-dependent proteolysis by calpain (Demarchi and Schneider, 2007) seems unlikely because high concentrations of a peptidic calpain inhibitor IV (20 µM, Calbiochem) had no significant effect when added to the cytoplasmic solution. A role of Ca-activated free radical generation seems unlikely because multiple free radical scavengers (TEMPO, 5 mM; sucrose, 40 mM; dithiothreitol, 2 mM; acetylcysteine, 2 mM; and ascorbate, 4 mM) were without significant effect in these protocols at concentrations suggested in the literature to suppress free radical signaling.

Failure to Correlate Capacitance Changes with Transporter or Channel Activity Changes
The source of the membrane that traffics in these experiments is not well defined. In this connection, we tested whether Na/Ca exchange activities or Na/K pump activities might increase with membrane fusion or decrease with the subsequent fall of capacitance. To do so, we activated the transport currents very briefly by application of extracellular Ca or K, respectively, to determine if maximal activity increases with membrane fusion and/or decreases during the period of membrane retrieval. We found no evidence that these transporter activities followed the changes of membrane area inferred from capacitance changes. In addition, we performed similar experiments in cells expressing ROMK-type potassium channels (Zeng et al., 2003), and results were similarly negative.

Evidence for Membrane Fusion as the Cause of Ca-induced Capacitance Increase
The fact that phospholipase inhibitors had little or no effect on membrane fusion tends to eliminate the possibility that large biochemical changes could be causing dielectric changes in these experiments. Nevertheless, the large magnitudes of capacitance responses, in some cases doubling cell capacitance, do raise questions about the nature of the membrane fusing. Therefore, we performed an ultrastructural study to analyze subplasmalemmal vesicles and membrane structures in BHK cells that might participate in fusion events within 2–5 s of activating Ca influx. Cells were trypsinized and removed from plates, as in preparation for electrophysiological experiments. Cells were then fixed under conditions established previously for ultrastructural studies of submembrane compartments in neurons (see Materials and methods). As shown in Fig. 10, electron micrographs indeed show the presence of numerous vesicles within short distances of the surface membrane, both coated and uncoated vesicles with diameters ranging from 45 to 90 nm. In >20 cells examined, the distributions of noncoated and coated vesicles were found to be similar in dependence on distance from the membrane. Assuming that sections are 65 nm thick, we counted the numbers of vesicles per linear micron of membrane within 500 nm of the membrane, and we projected an average number of vesicles per square micron of membrane. With an average diameter of 80 nm, fusion of at least 50 vesicles per square micron would be required to double membrane area. Our estimate of vesicular densities was only 10% of this value, indicating an important discrepancy between ultrastructural analysis and functional results. Some possible explanations are that (1) vesicles are lost during the procedures employed, (2) vesicles fuse during the fixation protocols employed, and that (3) the vesicle population that fuses is generated during procedures to establish whole-cell voltage clamp. We mention that pipette perfusion of the fixative solutions into cells did not cause membrane fusion, based on capacitance recording.

View larger version (79K): [in this window] [in a new window] [Download PPT slide]   Figure 10. Electron microscopic analysis of BHK cell vesicles in close proximity to the surface membrane. (A) Electron micrograph of submembrane vesicles in a BHK cell. (B) Analysis of noncoated and coated vesicle numbers identified at the given distances from the surface membrane of >20 cells.