| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Department of Physiology, University of Cambridge, United Kingdom
Correspondence: Address reprint requests to Dr. Michael J. Mason, Dept. of Physiology, University of Cambridge, Downing St., Cambridge CB2 3EG, UK. Tel.: 44-1223-333899; E-mail: mjm39{at}cam.ac.uk.
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
|---|
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
|---|
; Hamill et al., 1981). The composition of the pipette solution is more critical in the whole-cell patch-clamp configuration than with sharp electrodes since patch electrodes are of lower resistance, resulting in dialysis of the cytosol by the composition of the patch pipette (Pusch and Neher, 1988). This is a particularly important issue since dialysis can result in the loss of important cytosolic constituents including soluble kinases and phosphatases and small-molecular weight proteins and substrates required for the normal cellular functions setting membrane potential. This problem has been overcome by the use of the perforated patch technique (Horn and Marty, 1988; Rae et al., 1991). Nystatin and amphotericin have been used to reduce patch resistance for the purpose of obtaining adequate voltage control of the cell under voltage clamp and accurate measurements of membrane potential under current clamp without washing out large-molecular weight compounds. However, these agents reduce access resistance by introducing exogenous channels with high permeability, thus requiring careful consideration of the ionic composition of the patch pipette and the impact of this ionic composition on membrane potential.
Noninvasive techniques for the measurement of membrane potential have been used in a variety of cell types. The Nernstian distribution of radiolabeled compounds has been used to estimate transmembrane potential. Although this technique is of limited value for the measurement of dynamic changes in potential, given the requirement of steady-state isotope distribution for calibration of potential, it has proved useful for estimates of steady-state potential (Catterall et al., 1976; Lichtshtein et al., 1979).
A variety of potentiometric fluorescent indicators have been employed to monitor membrane potential in a variety of cellular preparations (Waggoner, 1979; Rink et al., 1980; Grinstein et al., 1984; Ehrenberg et al.,1988; London et al., 1989; Zhang et al., 1998; Orbach et al., 1985; Mason and Grinstein, 1990; Rohr and Salzberg, 1994; Mason et al.,1999; Kao et al., 2001). Although potentiometric indicators are noninvasive, the influence of the fluorescent probe on cellular function must be considered (Simons, 1979; Montecucco et al., 1979; Rink et al., 1980). An additional concern with potentiometric indicators is converting their fluorescence or absorbance recordings into meaningful measurements of potential.
Indications of membrane potential changes across the membranes of intact bovine chromaffin cells have been recorded using the cell-attached voltage-clamp configuration (Fenwick et al., 1982). In these experiments current waveforms arising from intracellular action potentials could be recorded across the intact patch. More recently, using the cell-attached voltage-clamp configuration, Verheugen and colleagues have used changes in the reversal potential of the voltage-gated K+ channel to quantitatively estimate whole-cell membrane potential across an intact membrane patch (Verheugen et al., 1995). These data provided early indications that knowledge of the whole-cell membrane potential could be inferred from current recordings made across intact patches. To date the extent to which cell-attached current-clamp measurements reflect the whole-cell membrane potential has not been extensively investigated. In the experiments presented here, we demonstrate the feasibility of noninvasive measurements of both steady-state and dynamic changes in cell membrane potential across an intact membrane patch using the current-clamp mode of a patch-clamp amplifier.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
|---|
Cell culture
Rat basophilic leukemia cells of the RBL-1 clone were obtained from Dr. S. Ikeda (National Institutes of Health, Bethesda, MD) and were propagated in minimal essential medium with added Earles salts (Sigma-Aldrich). The media was supplemented with 10% fetal bovine serum (Sigma-Aldrich), 1% nonessential amino acids (Invitrogen, Paisley, UK), 100 U·ml–1 penicillin (Sigma-Aldrich), 50 µg·ml–1 streptomycin (Sigma-Aldrich), 2 µg·ml–1 amphotericin (Sigma-Aldrich) and 2 mM L-glutamine (Sigma-Aldrich). Cells were grown at 37°C in a humidified atmosphere of 95% air and 5% CO2. Suspension cells were used for propagation in all experiments as previously reported (Schofield and Mason, 1996). Cells were normally used 36–48 h after passaging. For experiments, 1-ml aliquots of media containing healthy floating RBL-1 cells were transferred to a 1.5-ml microcentrifuge tube. A small aliquot of this cell suspension was added directly to the experimental chamber for electrophysiological recording when required. All experiments were performed at room temperature.
Megakaryocyte isolation
Adult male Wistar rats were killed by exposure to a rising concentration of CO2 followed by cervical dislocation. Marrow containing megakaryocytes was isolated from the tibial and femoral bones of the hind limbs into saline containing (in mM) 145 NaCl, 5 KCl, 1 CaCl2, 1 MgCl2, 10 Hepes, 10 D-glucose, pH 7.35 with NaOH, and 0.32–0.64 U·ml–1 type VII apyrase (Sigma-Aldrich) as described previously (Mahaut-Smith et al., 1999). Apyrase was present during the isolation and storage of the marrow preparation to degrade spontaneously released adenosine nucleotides, but was absent during experiments. For electrophysiological recordings, a small aliquot of the marrow preparation was added directly to the experimental chamber with megakaryocytes being easily identifiable based upon their large size and polyploidic nucleus. All experiments were performed at room temperature.
Electrophysiological recording
Single electrode recording in RBL-1 cells
Tight-seal whole-cell and cell-attached patch-clamp recordings in voltage- and current-clamp mode were carried out using an Axopatch 200A amplifier (Axon Instruments, Union City, CA). In whole-cell voltage-clamp mode 70–75% series resistance compensation was achieved using the series resistance compensation feature of the 200A amplifier. Two different pipette solutions were used in whole-cell and cell-attached recordings. A KCl-based pipette solution had the following ionic composition (in mM): 150 KCl, 0.15 EGTA, 2 MgCl2, 10 Hepes, pH 7.3 with KOH. A K-glutamate-based pipette solution contained (in mM) 150 K-glutamate, 2 EGTA, 1 CaCl2, 10 Hepes, pH 7.3 with KOH. Experiments were performed in normal external solution of the following composition (in mM): 145 NaCl, 5 KCl, 1 CaCl2, 1 MgCl2, 10 Hepes, 10 D-glucose, pH 7.35 with NaOH. High-K+ solution was made by replacing 145 mM NaCl with KCl and titrating with KOH ([K+] = 154 mM). Voltage- and current-clamp recordings made with the K-glutamate internal are corrected for an experimentally determined +13 mV junction potential. The experimental chamber was earthed via an Ag/AgCl pellet placed directly in the chamber downstream of the cell.
Amplifier control and data acquisition were performed using Axograph 4.8 software (Axon Instruments) running on a Macintosh computer using a Digidata 1322A 16-bit data acquisition system (Axon Instruments). For identification of whole-cell currents under voltage-clamp, cells were held at –40 mV and 255-ms ramps from –140 to +60 mV were initiated every second. Currents were filtered at 1 kHz using an 8-pole low-pass Bessel filter (Frequency Devices, Haverhill, MA) and acquired to disk at 2 kHz. Single-channel events from cell-attached patches were recorded during 950-ms ramps from +140 to –60 mV pipette potential from a pipette holding potential of +40 mV. Ramps were applied every 3 s. Membrane voltage under current-clamp conditions in both the whole-cell and cell-attached modes was recorded at 2 kHz from either the unfiltered 10-Vm output of the Axopatch 200A amplifier or from the scaled output filtered at 1 kHz. In some experiments the 10-Vm output was filtered at 15 Hz before sampling at 2 kHz. Data were analyzed using Axograph software and IGOR Pro (Wavemetrics, Lake Oswego, OR).
Simultaneous whole-cell voltage-clamp and cell-attached current-clamp recordings in megakaryocytes
Simultaneous whole-cell voltage-clamp and cell-attached current-clamp recordings were performed using two patch-clamp amplifiers. Conventional tight seal whole-cell patch-clamp in voltage-clamp mode was performed using an Axopatch 200A amplifier with a ß = 0.1 headstage configuration, thus enabling whole-cell capacitance compensation of the large megakaryocyte membrane capacitance (Mahaut-Smith et al., 2003). An Axopatch 200A amplifier was used to record from a second electrode in the tight seal cell-attached current-clamp mode. Both electrodes were filled with a KCl-based pipette solution with the following ionic composition (in mM): 150 KCl, 0.1 EGTA, 0.05 K5-fura-2 salt, 0.05 Na2GTP, 2 MgCl2, 10 Hepes, pH 7.3 with KOH. Megakaryocyte experiments were carried out in an extracellular solution of the following composition (in mM): 145 NaCl, 5 KCl, 1 CaCl2, 1 MgCl2, 10 Hepes, 10 D-glucose, pH 7.35 with NaOH. Experiments were performed on a microscope equipped for simultaneous recording of single-cell fura-2 fluorescence as previously reported (Mahaut-Smith et al, 1999). Electrophysiological signals were acquired using hardware and software provided by Cairn Research (Faversham, UK). The current record from the whole-cell electrode under voltage-clamp (Axopatch 200B amplifier) was filtered at 30 Hz (Kemo Variable Filter, Beckenham, UK) and recorded by the Cairn software at 60 Hz. The whole-cell membrane voltage was recorded at 60 Hz from the unfiltered 10-Vm output of the 200A amplifier. Voltage steps were controlled by PClamp 6 (Axon Instruments) running on a second PC. Membrane voltage recorded from the second electrode in the tight seal cell-attached current-clamp configuration was filtered at 30 Hz and recorded at 60 Hz from the scaled output of the Axopatch 200A. All data were further averaged with software to give a 15 Hz acquisition rate for all channels.
Patch electrodes
Fire-polished patch electrodes were constructed from filamented borosilicate glass (Harvard Apparatus, Edenbridge, UK) and had a resistance of 
5–8 M
(for use with RBL cells) or 3–6 M (for use with megakaryocytes) when filled with KCl-based internal. For some single-channel recordings electrode shanks were coated with Sylgard 184 silicone elastomer (Dow Corning, Wiesbaden, Germany) to reduce pipette capacitance.
Simultaneous whole-cell voltage-clamp and cell-attached current-clamp recordings in a model circuit
Simulated simultaneous whole-cell voltage-clamp and cell-attached current-clamp recordings in a single cell were made using an equivalent model circuit connected to two patch-clamp amplifiers. The equivalent model circuit used to approximate this experimental situation is presented in Fig. 1 A with the actual circuit presented in Fig. 1 B. Resistors X1 and X2 in this circuit diagram represent the seal resistance and the resistance of the cell-attached patch, respectively, and were altered to experimentally define the relationship between the ratio of these resistances and the fraction of the whole-cell membrane potential recorded across a cell-attached patch. This model circuit deviates from a true equivalent membrane circuit in three ways. First, the circuit does not include a low-value resistor to represent the resistance of the cell-attached electrode. This was neglected as the resistance of this component of the circuit is at least two orders of magnitude less than the patch resistance and thus has negligible impact upon the results. Second, we have designed the circuit to remove series resistance error in voltage-clamping the cell. Third, the whole-cell voltage-clamp component of the equivalent circuit does not contain a high-value resistor in parallel with the cell input resistance representing the seal resistance of the electrode. Since this resistance represents a pathway to earth for current flow it was omitted, indicative of an infinite seal resistance. This minimizes the required holding current passed by the amplifier to clamp the potential of the whole-cell component of the circuit, and its absence has no influence upon the results of the model circuit experiments.
|
Experimental chamber and solution exchange
Cell experiments were performed in a shallow perspex chamber of 0.5 ml, the bottom of which was formed by adhering a number 1 coverslip with silicone high-vacuum grease. Solution exchange in the experimental chamber was made by gravity fed bath superfusion with solution removal being achieved using a 100 millibar vacuum pump.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
|---|
 | (1) |
 | (2) |
 | (3) |
|
To investigate the usefulness of this technique, measurements of membrane potential were performed in RBL-1 cells. As a result of the small size of the RBL-1 cell it was not possible to reliably perform simultaneous whole-cell and cell-attached experiments. Rather, potential was recorded under current-clamp in cell-attached mode immediately before obtaining the whole-cell configuration and recording membrane potential again under current-clamp. Paired values for steady-state membrane potential recorded in the cell-attached and whole-cell configuration were obtained in 24 cells. Fourteen cells were depolarized (i.e., less negative) in both the cell-attached and whole-cell configuration whereas 10 cells showed a hyperpolarized potential when recorded in the cell-attached configuration. Of these 10 cells, eight were still hyperpolarized when potential was recorded in the whole-cell configuration. The two remaining cells were depolarized, possibly a result of alterations in the whole-cell currents or seal resistance during the transition to the whole-cell mode. Data from these two cells are excluded from the membrane potential values summarized in Table 1. These data demonstrate that in hyperpolarized cells 77% of the membrane potential recorded in the whole-cell mode is recorded across the cell-attached patch. In addition, no significant difference in the potential recorded by the two configurations was detected in depolarized cells.
|
+2 mV when recorded under current clamp in the cell-attached configuration. This depolarization was fully reversible upon removal of the high-K+ saline. After the transition to the whole-cell configuration, the resting potential was 10 mV more hyperpolarized with the response to high-K+ exposure being virtually identical. It was consistently found that the high-frequency noise evident upon the cell-attached records was dramatically reduced upon transition to the whole-cell configuration. This observation was very useful in monitoring the integrity of the cell-attached configuration. Table 2 summarizes the membrane potential readings from unpaired cell-attached and whole-cell recording made immediately before and at the peak of exposure to high-K+ saline.
|
|
|
|
; McCloskey and Cahalan, 1990; Mukai et al., 1992, Mason et al., 1999). A close inspection of the null current value indicates that the membrane potential would be –21 mV, consistent with the depolarized nature of the cell when recorded under current clamp. This arises as a result of the lack of an outward current component to the I-V relationship at more negative potentials. The current-voltage relationships labeled 2 s, 5 s, 10 s, and 21 s were obtained at those times after the first detected change in the I-V relationship in response to changing to a solution containing 154 mM K+. During superfusion of high K+, there is an increase in the slope conductance of the inwardly rectifying K+ channel I-V curve at negative potentials and, importantly, an increase in the outward component of the I-V relationship, observations previously reported in response to increasing extracellular [K+] (Lindau and Fernandez, 1986; McCloskey and Qian, 1994; Kelly et al., 1992). The effect of the increase in outward component of the I-V relationship is to transiently shift the null current potential toward a hyperpolarized potential set by the K+ equilibrium potential. As discussed above, the extracellular [K+] does not change instantaneously, but rather, increases slowly as a result of mixing. A modest increase in extracellular [K+] is sufficient to increase the outward component of the I-V relationship but still results in a hyperpolarized K+ equilibrium potential. As the extracellular [K+] continues to rise, the null current potential depolarizes in accordance with the changing K+ equilibrium potential. Under current-clamp conditions this is expected to result in a rapid transient hyperpolarization followed by depolarization toward a potential set by the final extracellular [K+] of 154 mM, consistent with the membrane potential changes presented in Fig. 5. This would be expected to occur in reverse upon returning to normal saline. Therefore, the noninstantaneous exchange of solution in the chamber and the resultant transient changes in the outward current component of the inwardly rectifying K+ channel most likely underlie the anomalous changes in membrane potential recorded in Fig. 5. Importantly, these changes are faithfully recorded across the cell-attached patch, as is evident in Fig. 5 A.
|
; Yamada, 1957). This membrane is electrically coupled to the plasma membrane, suggesting that a conventional cell-attached patch might possibly be electrically coupled to far more membrane than predicted from the area of the patch (Mahaut-Smith et al., 2003). Theoretically, this increase in membrane would be expected to reduce the resistance of the patch, thereby making it an excellent candidate for further investigations into the feasibility of potential recordings across a cell-attached patch. In addition, the large size of the megakaryocyte facilitates two electrode patch-clamp experiments so that the two-electrode configuration denoted by the model circuit of Fig. 1 can be achieved in a biological sample. Experiments were undertaken to investigate the accuracy of cell-attached membrane potential measurements in these large cells. Simultaneous cell-attached and whole-cell recordings were obtained in four cells. In three of these cells a high-resistance seal was obtained with both electrodes whereas low-resistance seals were obtained in one cell. Fig. 7 shows a representative experiment with two high-resistance seals. Whole-cell potential was either measured under current-clamp or controlled by voltage-clamp while membrane potential was simultaneously recorded across an intact patch using a second electrode in current-clamp mode. The cell was initially held at –75 mV with 70% series resistance compensation and the cell-attached potential recorded was 2 mV more negative (–77 mV). A change in holding potential to 0 mV was accompanied by a rapid initial depolarization of the cell-attached recording followed by a secondary slower further increase to a value approaching 0 mV. This secondary slower change in potential mirrored the inactivation of the large outward voltage-gated K+ current and most likely represents the true whole-cell voltage as a result of uncompensated series resistance. The whole-cell electrode was placed in current-clamp mode and a spontaneous transient hyperpolarization was detected on both electrodes, consistent with a transient rise in [Ca2+]i and the activation of Ca2+-gated K+ channels (Uneyama et al., 1993a,b; Mason et al., 2000). The cell was returned to whole-cell voltage-clamp and a holding potential of –75 mV. Voltage steps, both depolarizing and hyperpolarizing, were made while recording the potential across the cell-attached patch. Changes in holding potential that did not activate large membrane currents were faithfully mirrored by the cell-attached electrode. Only when large membrane currents were activated by the voltage steps was a biphasic change in potential recorded by the cell-attached electrode. These data further support the conclusion that the biphasic potential change is a true indication of the whole-cell potential arising from the uncompensated component of the series resistance. It is important to note that under conditions where no large membrane currents are activated, the voltage excursions recorded across the cell-attached patch were virtually identical to the voltage step applied under voltage clamp. This high degree of accuracy of measurement of membrane potential by the cell-attached electrode was observed in all three cells in which high-resistance seals were obtained and is consistent with a large seal/patch resistance ratio. In the cell in which low-resistance seals were obtained, whole-cell current-clamp recordings revealed spontaneous oscillations in potential driven by oscillations in [Ca2+]i and the activation and inactivation of Ca2+-activated K+ channels, as discussed above. Despite the low-resistance seal, the oscillations, including the inherent variability in their magnitude, were faithfully recorded by the cell-attached electrode. As expected, both the absolute magnitude of the oscillations and the resting potential recorded with the cell-attached electrode were significantly less than that recorded by the whole-cell electrode, as predicted by the much lower seal/patch resistance ratio.
|
; Mason et al.,2000). A major concern of these experiments is that dialysis of the cell with the pipette solution is contributing to the oscillatory nature of the response. To address this question, experiments were undertaken to monitor membrane potential using a cell-attached electrode during application of ADP. In eight out of nine cells oscillations in membrane potential were recorded in response to ADP exposure. A representative experiment is shown in Fig. 8. Superfusion with 1 µM ADP resulted in oscillations in membrane potential between a resting potential of –40 mV and a value of –70 mV. Resting potential, the magnitude of the oscillations, and the kinetics of the oscillations were indistinguishable from oscillations recorded in the whole-cell configuration (Mahaut-Smith et al., 1999; Mason et al.,2000). Thus, cell dialysis does not underlie the oscillatory nature of the response. These experiments highlight the usefulness of cell-attached current-clamp recordings for noninvasive detection of dynamic changes in membrane potential.
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
|---|
When recording dynamic changes in potential using this technique it is important to consider the speed of the changes. Two issues arise. First, the design of many patch-clamp amplifiers, that enables them to monitor potential using the current-clamp mode, has limitations in the recording of high-frequency changes in potential (Magistretti et al, 1996,1998). This is a major problem for quantitative analysis of membrane potential in excitable cells where action potentials and burst-firing have high-frequency components to their signal. This is a general problem associated with current-clamp measurements using conventional patch-clamp amplifiers and is not confined to the cell-attached configuration. However, in nonexcitable cells there is a greatly reduced high-frequency component in the membrane potential signal making current-clamp measurements less prone to the artifacts present in measurements in excitable cells. Second, the capacitance elements of the cell-attached recording of potential effectively act as low-pass filters that attenuate high-frequency signal components. With the availability of patch-clamp amplifiers with true current-clamp circuits the concerns raised by point 1 are muted. However, concerns raised by the filtering influences of the capacitance elements of the cell-attached recording remain. We have calculated the step response of the equivalent circuit presented in Fig. 1 A. The time constant of the change in potential recorded by the cell-attached electrode in response to a step change in whole-cell potential is given by the following relationship:
 | (4) |
120 Hz if the product of the patch resistance and capacitance was 1 ms. It is important to bear in mind that even small headstage offset currents can result in dramatic errors in cell-attached measurements of potential as a result of the high resistance of the membrane patch. Headstage offset currents and leakage currents in current-clamp mode must be offset frequently to ensure that large errors are not introduced.
The high-resistance seals, which are required for good patch-clamp recordings, contributed to the relatively high seal/patch ratio of 3.3 estimated in the RBL-1 cell. Interestingly, all of our measurements of single-channel events in RBL-1 cells revealed multiple channels per patch with a high open probability as determined by the lack of complete closed channel events. Therefore, it seems likely that a low patch resistance in the RBL-1 cells is a contributing factor for the relatively efficient detection of the whole-cell potential. Interestingly, previous estimates in adrenal chromaffin cells by Fenwick and colleagues (1982) indicate that the membrane resistance is much lower than predicted from the ratio of patch area to membrane surface area. The authors propose that the process of forming a high-resistance seal may in fact be responsible for lowering the patch resistance, possibly via membrane damage. However, should membrane damage underlie the low resistance achieved in RBL cells it does so without compromising the ability to detect single-channel events, thus ensuring that the cell-attached configuration is intact.
Opening and closing events in patches containing a low channel density will undergo large changes in patch resistance that will show up as rapid transitional changes in potential as the fractional accuracy of the recorded membrane potential is altered with changing resistance. The rapid fluctuations in potential recorded in partially depolarized cells in the cell-attached configuration (Fig. 5 A) were first thought to be as a result of such events. However, this does not appear to be the case as similar transitions were recorded in the whole-cell configuration (Fig. 5 B). These fluctuations in potential most likely arise from the inherent instability of the null current potential of depolarized cells. This instability arises as a result of 1), the small peak outward current between –80 and –40 mV; and 2), the decline in the outward current between –60 and –40 mV. As a result, small alterations in membrane current can result in pronounced shifts in potential to a new null current potential (Mason et al., 1999). The origin of the current changes responsible for the instability of the membrane potential observed in Fig. 5 may lie at the level of endogenous membrane currents or at the level of alterations in the seal resistance of the whole-cell or cell-attached electrode.
In the case of the megakaryocytes, membrane potential recorded across patches approached that set by whole-cell voltage-clamp. The presence of the demarcation membrane which is electrically coupled to the plasma membrane (Mahaut-Smith et al., 2003) may help explain this high degree of accuracy of the detection of potential. This increase in membrane may be significantly reducing the effective patch resistance and therefore increasing the seal/patch resistance ratio to a much higher value than that obtained in the absence of demarcation membrane. However, further experiments are required to define the role of this platelet precursor membrane system in the high degree of accuracy recorded by the cell-attached electrode. As a potential use of this technique we have exploited cell-attached current-clamp measurements to ensure that the dynamic changes in membrane potential induced by ADP are in fact not an artifact of whole-cell recordings. Although the high degree of accuracy of the membrane potential recorded may be a result of the presence of the demarcation membrane system, the accuracy of the dynamic changes is independent of the presence of a low-resistance patch. Even measurements with low-resistance cell-attached seals faithfully mirrored the dynamic changes recorded whole-cell (data not shown). Therefore, the ability to faithfully measure complex changes in potential is not confined to cells in which a very low-resistance patch is achieved. Thus, this technique will be useful for monitoring dynamic changes in potential under conditions where the accuracy of the absolute values of potential are less important.
As noted above, accurate measurement of membrane potential across a cell-attached patch requires a patch displaying a constant low resistance. Nystatin and amphotericin have been used to reduce patch resistance for the purpose of obtaining adequate voltage control of the cell under voltage clamp (Horn and Marty, 1988; Rae et al., 1991). Nystatin and amphotericin can also be used to improve the accuracy of the recorded potential in current-clamp mode. However, although these agents will reduce the resistance of the patch, they do so by introducing exogenous channels with high permeability, thus requiring careful consideration of the ionic composition of the patch pipette and the impact of this ionic composition on cell potential after reaching steady-state conditions across the patch. We have performed our experiments without exogenously altering the patch resistance with a surprisingly high degree of accuracy.
These experiments highlight the feasibility of using a cell-attached electrode in current-clamp to measure a large fraction of the whole-cell transmembrane potential noninvasively under steady-state and dynamic conditions. The usefulness and future application of this method will be very much dependent upon the membrane characteristics of the cell under investigation. Numerous factors need to be considered when applying this technique, including 1), the seal/patch resistance ratio under your experimental conditions; 2), the stability of the patch and seal resistances; and 3), the kinetics of the changes in potential. All of these factors will determine the accuracy of the recorded potential. In the RBL cell and the rat megakaryocyte this technique has already proved highly useful for investigations of dynamic changes in potential under noninvasive conditions.
|
ACKNOWLEDGEMENTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
|---|
This work was funded in part by grants from the Medical Research Council (G9901465 to M.P.M.-S. and M.J.M), the British Heart Foundation (BS10 to M.P.M.-S.), and the Royal Society (M.P.M.-S.).
Submitted on July 16, 2004; accepted for publication October 19, 2004.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
|---|
Catterall, W. A., R. Ray, and C. S. Morrow. 1976. Membrane potential dependent binding of scorpion toxin to action potential Na+ ionophore. Proc. Natl. Acad. Sci. USA. 73:2682–2686.
Ehrenberg, B., V. Montana, M. Wei, J. P. Wuskell, and L. M. Loew. 1988. Membrane potential can be determined in individual cells from the Nernstian distribution of cationic dyes. Biophys. J. 53:785–794.
Fenwick, E. M., A. Marty, and E. Neher. 1982. A patch-clamp study of bovine chromaffin cells and of their sensitivity to acetylcholine. J. Physiol. 331:577–597.
Grinstein, S., J. D. Goetz, and A. Rothstein. 1984. 22Na+ fluxes in thymic lymphocytes. I. Na+/Na+ and Na+/H+ exchange through an amiloride-insensitive pathway. J. Gen. Physiol. 84:565–584.
Hamill, O. P., A. Marty, E. Neher, B. Sakmann, and F. J. Sigworth. 1981. Improved patch clamp techniques for high resolution current recording from cells and cell-free membrane patches. Pflugers Arch. 391:85–100.[CrossRef][Medline]
Horn, R., and A. Marty. 1988. Muscarinic activation of ionic currents measured by a new whole-cell recording method. J. Gen. Physiol. 92:145–159.
Kao, W. Y., C. E. Davis, Y. I. Kim, and J. M. Beach. 2001. Fluorescence emission spectral shift measurements of membrane potential in single cells. Biophys. J. 81:1163–1170.
Kelly, M. E. M., S. J. Dixon, and S. M. Sims. 1992. Inwardly rectifying potassium current in rabbit osteoclasts: A whole-cell and single-channel study. J. Membr. Biol. 126:171–181.[Medline]
Lichtshtein, D., H. R. Kaback, and A. J. Blume. 1979. Use of a lipophilic cation for the determination of membrane potential in neuroblastoma-glioma hybrid cell suspensions. Proc. Natl. Acad. Sci. USA. 76:650–654.
Lindau, M., and J. M. Fernandez. 1986. A patch-clamp study of histamine-secreting cells. J. Gen. Physiol. 88:349–368.
London, J. A., L. B. Cohen, and Y. J. Wu. 1989. Optical recordings of the cortical response to whisker stimulation before and after addition of an epileptogenic agent. J. Neurosci. 9:2182–2190.[Abstract]
Magistretti, J., M. Mantegazza, M. de Curtis, and E. Wanke. 1998. Modalities of distortion of physiological voltage signals by patch-clamp amplifiers: A modeling study. Biophys. J. 74:831–842.
Magistretti, J., M. Mantegazza, E. Guatteo, and E. Wanke. 1996. Action potentials recorded with patch-clamp amplifiers: are they genuine? Trends Neurosci. 19:530–534.[CrossRef][Medline]
Mahaut-Smith, M. P., J. F. Hussain, and M. J. Mason. 1999. Depolarization-evoked Ca2+ release in a non-excitable cell, the rat megakaryocyte. J. Physiol. 515:385–390.
Mahaut-Smith, M. P., D. Thomas, A. B. Higham, J. A. Usher-Smith, J. F. Hussain, J. Martinez-Pinna, J. N. Skepper, and M. J. Mason. 2003. Properties of the demarcation membrane system in living rat megakaryocytes. Biophys. J. 84:2646–2654.
Mason, M. J., and S. Grinstein. 1990. Effect of cytoplasmic acidification on the membrane potential of T-lymphocytes: Role of trace metals. J. Membr. Biol. 116:139–148.[CrossRef][Medline]
Mason, M. J., J. Limberis, and G. G. Schofield. 1999. Transitional changes in membrane potential and intracellular [Ca2+] in rat basophilic leukemia cells. J. Membr. Biol. 170:79–87.[CrossRef][Medline]
Mason, M. J., J. F. Hussain, and M. P. Mahaut-Smith. 2000. A novel role for membrane potential in the modulation of intracellular Ca2+ oscillations in rat megakaryocytes. J. Physiol. 524:437–446.
McCloskey, M. A., and M. D. Cahalan. 1990. G protein control of potassium channel activity in a mast cell line. J. Gen. Physiol. 95:205–227.
McCloskey, M. A., and Y. X. Qian. 1994. Selective expression of potassium channels during mast cell differentiation. J. Biol. Chem. 269:14813–14819.
Montecucco, C., T. Pozzan, and T. Rink. 1979. Dicarbocyanine fluorescent probes of membrane potential block lymphocyte capping, deplete cellular ATP and inhibit respiration of isolated mitochondria. Biochim. Biophys. Acta. 552:552–557.[Medline]
Mukai, M., I. Kyogoku, and M. Kuno. 1992. Calcium-dependent inactivation of inwardly rectifying K+ channel in a tumor mast cell line. Am. J. Physiol. 262:C84–C90.[Medline]
Neher, E., and B. Sakmann. 1978. The extracellular patch clamp: a method for resolving currents through individual open channels in biological membranes. Pflugers Arch. 375:219–228.[CrossRef][Medline]
Orbach, H. S., L. B. Cohen, and A. Grinvald. 1985. Optical mapping of electrical activity in rat somatosensory and visual cortex. J. Neurosci. 5:1886–1895.[Abstract]
Pusch, M., and E. Neher. 1988. Rates of diffusional exchange between small cells and a measuring patch pipette. Pflugers Arch. 411:204–211.[CrossRef][Medline]
Rae, J., K. Cooper, P. Gates, and M. Watsky. 1991. Low access resistance perforated patch recordings using amphotericin B. J. Neurosci. Methods. 37:15–26.[CrossRef][Medline]
Rink, T. J., C. Montecucco, T. R. Hesketh, and R. T. Tsien. 1980. Lymphocyte membrane potential assessed with fluorescent probes. Biochim. Biophys. Acta. 595:15–30.[Medline]
Rohr, S., and B. M. Salzberg. 1994. Characterization of impulse propagation at the microscope level across geometrically defined expansions of excitable tissue: multiple site optical recording of transmembrane voltage (MSORTV) in patterned growth heart cell cultures. J. Gen. Physiol. 104:287–309.
Schofield, G. G., and M. J. Mason. 1996. A Ca2+ current activated by release of intracellular Ca2+ stores in rat basophilic leukemia cells (RBL-1). J. Membr. Biol. 153:217–231.[CrossRef][Medline]
Simons, T. J. 1979. Actions of carbocyanine dyes on the Ca2+-dependent K+ channel in human red cell ghosts. J. Physiol. 288:481–507.
Somasundaram, B., and M. P. Mahaut-Smith. 1995. A novel monovalent cation channel activated by inositol trisphosphate in the plasma membrane of the rat megakaryocyte. J. Biol. Chem. 270:16638–16644.
Uneyama, C., H. Uneyama, and N. Akaike. 1993a. Cytoplasmic Ca2+ oscillations in rat megakaryocytes evoked by a novel type of purinoceptor. J. Physiol. 470:731–749.
Uneyama, H., C. Uneyama, and N. Akaike. 1993b. Intracellular mechanisms of cytoplasmic Ca2+ oscillations in rat megakaryocytes. J. Biol. Chem. 268:168–174.
Verheugen, J. A. H., H. P. M. Vijverberg, M. Oortgiesen, and M. D. Cahalan. 1995. Voltage-gated and Ca2+-activated K+ channels in intact human T lymphocytes. J. Gen. Physiol. 105:765–794.
Waggoner, A. S. 1979. Dye indicators of membrane potential. Annu. Rev. Biophys. Bioeng. 8:47–68.[CrossRef][Medline]
Yamada, E. 1957. The fine structure of the megakaryocyte in the mouse spleen. Acta Anat. (Basel). 29:267–290.[Medline]
Zhang, J., R. M. Davidson, M. D. Wei, and L. M. Loew. 1998. Membrane electric properties by combined patch clamp and fluorescence ratio imaging in single neurons. Biophys. J. 74:48–52.
This article has been cited by other articles:
 |
|
 |
|
  C. Sevcencu, C. P. Pennisi, K. Yoshida, and H. Gregersen Simultaneous monitoring of cellular depolarization and contraction: a new method to investigate excitability and contractility in isolated smooth muscle cells Am J Physiol Gastrointest Liver Physiol, March 1, 2008; 294(3): G648 - G654. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. A. MacCannell, H. Bazzazi, L. Chilton, Y. Shibukawa, R. B. Clark, and W. R. Giles A Mathematical Model of Electrotonic Interactions between Ventricular Myocytes and Fibroblasts Biophys. J., June 1, 2007; 92(11): 4121 - 4132. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 E. W Newell and L. C Schlichter Integration of K+ and Cl- currents regulate steady-state and dynamic membrane potentials in cultured rat microglia J. Physiol., September 15, 2005; 567(3): 869 - 890. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||
