[19] Electrophysiological recording from Xenopus oocytes

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PosttranscriptionalPolyadenylation Reaction 50 m M 10 m M 2.5 m M 250 m M 150 U 25 #g 50/tg 0.1 m M 1U

Tris-HC1, pH 8.0 MgC12 MnC12 NaCI RNase inhibitor RNA Nuclease-free bovine serum albumin ATP (containing a trace of [a-32P]ATP) Poly(A) polymerase

The reaction is carried out in a volume of 100 ~1, at 37 °, for 1 hr. The average number of moles of adenosine incorporated per mole of RNA [i.e., the average poly(A) tail length] is calculated from the fraction of [32p]ATP converted to an acid-insoluble form during the reaction. Typically, under the conditions given above, the average tail length is approximately 50- 80 residues. The polyadenylated transcripts are extracted with phenol, to remove the protein, ethanol precipitated, and dissolved in TE at about 1 mg/ml.

[ 19] E l e c t r o p h y s i o l o g i c a l R e c o r d i n g Oocytes

from Xenopus

B y W A L T E R ST13HMER

Introduction The oocytes from Xenopus laevis have proved to be a reliable expression system for ion channels and transport systems. Electrophysiologists from all fields are increasingly making use of the oocyte expression system for their studies. Many measurements can now be carried out successfully which would have been impossible without this new technique. This chapter describes methods for electrophysiological measurements as well as some applications and limitations. Methods used to maintain Xenopus laevis, to microinject mRNA, to remove the follicular cell layer, and to express ion channels in Xenopus oocytes are described elsewhere in this volume ([14]-[16]). Therefore I only briefly describe the methods mentioned above as a general review, emphasizing some variations used in our laboratory. Xenopus oocytes have been used successfully to translate messenger METHODS IN ENZYMOLOGY, VOL. 207

~ t © 1992by Acadc.mi¢Press, Inc. All fightsof reproductionin any form reserved.

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RNAs (mRNA) into the respective proteins including posttranslational modifications. 1,2 Various receptors and ion channels have been functionally expressed after injection of poly(A) mRNA (e.g., Refs. 3-6). The development of genetic engineering techniques has made it possible to clone cDNAs coding for various receptor and/or channel proteins, to transcribe RNA from the cDNAs (cRNA), to inject the cRNAs into oocytes, and to analyze the functionally expressed proteins (e.g., Ref. 7- 11). Although other translation systems (detailed in [26]- [29], this volume) have been developed, certain experiments are only feasible in oocytes. The expression of foreign proteins in Xenopus oocytes has many advantages for electrophysiological measurements. The large size which easily accommodates manipulations like mRNA injections and electrode penetration is certainly one aspect. However, the fact that it is possible to obtain cellattached patch clamp recordings~2 and, in particular, recordings from macropatches ~3is among the main advantages. These low noise recordings from a large number of channels would be difficult to obtain from other systems, and measurements, like gating current fluctuations, ~4 are presently limited to the oocyte expression system.

J. B. Gourdon, C. D. Lane, H. R. Woodland, and G. Marbaix, Nature (London) 233, 177 (1971). 2 C. D. Lane, Curr. Top. Dev. Biol. 18, 89 (1983). K. Sumikawa, M. Houghton, J. S. Emtage, B. M. Richards, and E. A. Barnard, Nature (London) 292, 862 (1981). 4 R. Miledi, I. Parker, and K. Sumikawa, Proc. R. Soc. London B 216, 509 (1982). E. A. Barnard, R. Miledi, and K. Sumikawa, Proc. R. Soc. London B 215, 241 (1982). 6 C. B. Gundersen, R. Miledi, and I. Parker, Proc. R. Soc. London B 220, 131 (1983). 7 M. Mishina, T. Kurosaki, T. Tobimatsu, Y. Morimoto, M. Noda, T. Yamamoto, M. Terao, J. Lindstrom, T. Takahashi, M. Kuno, and S. Numa, Nature (London) 307, 604 (1984). s M. Noda, I. Takayuki, H. Suzuki, H. Takeshima, T. Takahashi, M. Kuno, and S. Numa, Nature (London) 322, 826 (1986). 9 L. C. Timpe, T. L. Schwarz, B. L. Tempel, D. M. Papazian, Y. N. Jan, and L. Y. Jan, Nature (London) 331, 143 (1988). ~oA. Mikami, K. Imoto, T. Tanabe, T. Niidome, Y. Mori, H. Takeshima, S. Narumiya, and S. Numa, Nature (London) 340, 230 (I 989). H U. B. Kaupp, T. Niidome, T. Tanabe, S. Terada, W. BOnink, W. Stfihrner, N. J. Cook, K. Kangawa, H. Matsuo, T. Hirose, T. Miyata, and S. Numa, Nature (London) 342, 762 (1989). ~2C. Methfessel, V. Witzemann, T. Takahashi, M. Mishina, and S. Numa, Pfluegers Arch. 4107, 577 (1986). ~3W. StOhmer, C. Methfessel, B. Sakmann, M. Noda, and S. Numa, Eur. Biophys. J. 14, 131 (1987). ~4F. Conti and W. StOhmer, Eur. Biophys. J. 17, 53 (1989).

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Preparation of Oocytes Details regarding the maintenance of Xenopus laevis and their oocytes have been described elsewhere in this volume (see [15]). Here, a brief review on the preparation and handling of RNA is given. A precondition for electrophysiological measurements on integral membrane proteins in oocytes is the injection of a sufficient amount of mRNA. This mRNA can be from various origins: either total RNA or poly(A) mRNA extracted from tissue samples, or cDNA-derived mRNA (cRNA). A tissue sample of about 1 g yields 10 to 100 #g of poly(A) mRNA. Extraction procedures have been described for total RNA 15.16 and poly(A) mRNA. 17 The main disadvantage of injecting total RNA or poly(A) mRNA is that all possible mRNAs are translated into proteins, diluting the desired mRNA with the consequence that the desired protein will be only a fraction of the expressed proteins. The relative abundance of the desired mRNA can be increased by size fractionation ~8,19 of poly(A) mRNA. The mRNA from cellular extracts should be injected at a concentration of 1 to 10/zg//zl, and the cRNA can be injected at a concentration of 0.2 to 0.8 pg/#l, both in aqueous solution. Standard procedures to avoid and inhibit RNases should be followed, such as using baked glassware (e.g., 250 ° for 6 hr), wearing sterile gloves at all stages, and using diethyl pyrocarbonate-treated, autoclaved doubledistilled water (DEPC-H20) for all solutions. Great care should be taken when handling the mRNA, not only to avoid contamination with RNases, but also to keep the solutions free of particles which could clog the injection pipettes. Thus the mRNA should be centrifuged to precipitate panicles suspended in solution. Silanized Eppendorf tubes (1 or 0.5 ml) are best suited as containers for mRNA. The mRNA should be stored at - 8 0 ° ; however, for brief periods, storage at - 20 ° is allowable. It is best to aliquot the mRNA into portions sufficient for the injections on a single day. For shipment and long-term storage it is best to store the mRNA precipitated in ethanol. The precipitation of mRNA can be carded out as follows: to 1 volume of aqueous mRNA solution 0.1 volume of 20% potassium acetate and 2.5 volumes of cold 100% ethanol are added. After is j. M. Chirgwin, A. E. Przybyla, R. J. MacDonnald, and W. J. Rutter, Biochemistry 18, 5924 (1979). 16 p. Dierks, A. van Ooyen, N. Mantel, and C. Weissmann, Proc. Natl. Acad. Sci. U.S.A. 78, 1411 (1981). 17 H. Aviv and P. Leder, Proc. Natl. Acad. Sci. U.S.A. 69, 1408 (1972). 18 p. Fourcroy, Electrophoresis 5, 73 (1984). 19 H. Ltibbert, B. J. Hoffman, T. P. Snutch, T. van Dyke, A. J. Levine, P. R. Hartig, H. A. Lester, and N. Davidson, Proc. Natl. Acad. Sci. U.S.A. 84, 4332 (1987).

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mixing by inversion of the tube, the mRNA is precipitated for at least 30 rain at - 7 0 °. The mRNA can be stored like this; however, it is more convenient to keep it in pure ethanol after decreasing the salt concentration. To do this the tube containing the mRNA is left at room temperature for about 5 min before centrifuging at 12,000 g for 15 rain at 4 ° in a cooled centrifuge (e.g., Hettich centrifuge, Eppendorf centrifuge, or Heraeus Biofuge A). Then the supernatant is carefully removed with a sterile suction pipette, and the pellet is washed with 2 volumes of 70% ethanol. A centrifugation of 10 min will pellet the mRNA again. After removing the saltcontaining supernatant the mRNA is stored in 100% ethanol. Care should be taken to avoid touching the mRNA pellet with the suction pipette and not to leave the tube open for long times to minimize contact with airborne RNases. To recover the precipitated mRNA the Eppendorf tube is brought to room temperature and centrifuged for 10 min at 12,000 g, the supernarant is removed, and the opening of the tube is covered with Parafilm into which three or four holes are poked with a sterile needle. The mRNA is then dried under reduced pressure in a desiccator for 4 - 10 rain. Then the mRNA can slowly be resuspended in D E P C - H 2 0 at the desired concentration for use. The best method for drying the RNA prior to resuspending it in water is to use a Speed-Vac or any other centrifugation under vacuum, so that the alcohol evaporates at the same time that the pellet is maintained at the bottom of the Eppendorf tube.

Injection of Messenger RNA Capillaries, such as transpipettor tubes (disposable micropipettes; Brand) are used for transferring the mRNA from the Eppendorftube to the injection pipette. The transpipettor tubes should be made hydrophobic by flushing them in a solution consisting of 50% ether and 50% silane (dimethyldichlorosilane, Fluka, Ronkonkoma, NY; caution: health hazard). After allowing the ether to evaporate under a hood, the tubes are baked at 180° for 2 hr to inactivate any remaining RNases. Tubes treated in this way can be stored under sterile conditions for several months. A micrometer controlled syringe, filled with mineral oil to reduce the air volume and evaporation of water from the mRNA solution, is used to load the transfer pipette with a few microliters of mRNA solution. Injection pipettes can be pulled using a standard pipette puller. They should have a long shank for estimating the volume of mRNA solution loaded and providing control that the oocytes are actually being injected. If pulled shortly before use, the injection pipettes will be RNase-free owing to the high temperature during the pulling process. The tips of the injection pipettes are broken under a microscope until they have an opening diame-

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ter of about 10/zm. Usually this will leave a rough and uneven rim, which might lesion the oocytes when penetrating them. This can be avoided by fire polishing the rim and pulling a sharp tip. This is done by approaching the tip of the injection pipette with a microfilament that has a small drop of molten glass on the tip. A sharp, needlelike protrusion can be obtained by rapidly removing the glass covered filament after touching the injection pipette. Radiation heat during this process is sufficient to smooth the ragged rim. The injection pipettes are filled under stereomicroscopic control by suction applied through a conventional syringe. For this purpose, mRNA solution is extruded to form a droplet just outside of the transfer pipette by means of the micromanipulator-driven syringe. At this stage, the injection pipette is filled by application of suction with approximately 500 nl of mRNA solution, sufficient for injecting about 10 oocytes. After this, it is convenient to draw the remaining mRNA solution a few millimeters back into the transfer pipette by slight suction. This procedure decreases evaporation of water from the mRNA solution, which will be used for the next batch of oocytes to be injected. A hand-driven coarse manipulator is sufficient to maneuver the injection pipette. Grooves carved into a thin Perspex plate fixed to the bottom of the injection chamber support the oocytes during injection. Alternatively, a scratched petri dish or one covered with either silicone curing agent (RTV615, General Electric, Waterford, NY) or agar can be used to fix the oocytes. The fixation of the oocytes is achieved by reducing the amount of solution in the injection chamber, so that the surface tension holds the oocytes in the grooves. Manipulation of oocytes is performed with the aid of Pasteur pipettes whose tips have been broken to enlarge the opening and then fire polished. A slight kink in the pipette tips helps in handling the oocytes during transport. The mRNA is injected in aliquots of about 50 nl per oocyte. There are many procedures to achieve this, and very simple manual syringe systems or injection machines can be used. More sophisticated injection machines such as the Eppendorf microinjector or the Drummond oocyte injector are helpful when injecting large quantities of oocytes at a time (see also the description in [15], this volume). Incubation In most cases the large stage V and VI oocytesz° can be used. The oocytes are incubated in small petri dishes in Barth's medium [in mM: 20 j. N. Dumont, J.

Morphol. 136, 153 (1972).

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IX 9]

84 NaC1, 1 KC1, 2.4 NaHCO3, 0.82 MgSO4, 0.33 Ca(NO3)2, 0.41 CaC12, 7.5 Tris-HC1, pH 7.4] with additions of penicillin and streptomycin (100 U/ml and 100/tg/ml, respectively). Some protocols use gentamicin (50-100 ~tg/ml). The optimal incubation temperature is 19 °. Barth's medium should be changed every day using sterile pipettes. Oocytes having a blurred delimitation between the animal and vegetal pole should be removed.

Removal of Follicular Cell Layer The follicular cell layer can be removed either before or after injection of the mRNA. It can be removed by purely mechanical means, but partial or total digestion of the follicular cell layer helps to prevent mechanical damage of the oocytes. Mild digestion is achieved in Ca2+-free Barth's medium containing 1 mg/ml collagenase I (e.g., Sigma, St. Louis, MO, type I) for 1 hr at room temperature. The follicular cell layer is then removed mechanically using two pairs of forceps (No. 5, Dumont & Fils, Montignez, Switzerland). Alternatively, the ovaries can be incubated for 2 - 3 hr in the collagenase-containing solution under shaking until the oocytes are dispersed. It is important to wash (3-4 times in about 5 ml) the oocytes extensively in Barth's medium after collagenase treatment to stop the enzymatic reaction. There seems to be no ad hoc rule to anticipate the time after injection when the various proteins will be inserted in the oocyte membrane. Therefore it is necessary to assay for protein expression from day to day. In general this will entail two-electrode voltage clamping for ion channels and electrogenic pumps. If no patch clamp experiments are intended, the vitelline envelope does not need to be removed. However, the removal of this last barrier to the plasma membrane makes insertion of electrodes easier, improves access and washout of solutes, and fixes the oocytes to the bottom of the experimental chamber (we use new 35-mm petri dishes as experimental chambers). Methods for removing the vitelline layer are described below in the section on patch clamp recording. Electrophysiological Measurements For most electrophysiological recordings from oocytes, the membrane potential has to be under control. This is achieved by clamping the oocyte to predefined potentials using a conventional two-electrode voltage clamp. 2~,22For this, one intracellular electrode is used to record the actual 21 T. G. Smith, J. Lecar, S. J. Redmann, and P. W. Gage, eds., in "Voltage and Patch Clamping with Microelectrodes," American Physiological Society, Bethesda, Maryland,

(Williams& Wilkins,Baltimore,Maryland), 1985.

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COMMAND

FIG. 1. Schematic diagram of the main components of a two-electrode voltage clamp. The output of the left amplifier-comparator (Vm) is the difference in potential between the bath and the oocyte electrodes. Vmis compared with the command potential, and any difference is injected into the oocyte through the current electrode. This current flows to ground through the bath ground electrode. If I'm equals the command potential, no current flows to ground.

intracellular potential, and the second electrode is used to pass current in such a way as to maintain the desired potential. This is achieved using a feedback circuit, which is the main component of the voltage clamp. The current needed to maintain a given potential is the measured parameter. There are two ways of measuring this current: as current flowing to ground through the grounding electrode (using a virtual ground amphfier) or as the current flowing through the current electrode. The intracellular potential recording electrode has such a high impedance that no current flows through it. Most voltage clamp setups use also a potential recording electrode (reference electrode) for the bath solution. This avoids polarization errors which arise from the current passing through the ground electrode. In this configuration, the transmembrane potential is taken as the difference between the intracellular potential electrode and the reference electrode. Therefore, the small error introduced because the bath is not always exactly at ground potential is corrected for. A block diagram of the main components of a voltage clamp is given in Fig. 1.

22N. B. Standen, P. T. A. Green, and M. J. Whitaker, eds., "Microelectrode Techniques: The Plymouth Workshop Handbook." The Company of Biologists, Cambridge, 1987.

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In adition to the voltage clamp itself, a pulse generator is required for adjusting the voltage clamp amplifier and to make experiments which require measurements in response to changes in membrane potential. This pulse generator can be anything from a simple waveform generator to a computer-controlled pulse generator. If the voltage clamp does not provide a builtin filter, a Bessel low-pass filter is required to reduce the high frequency noise and as an antialiasing filter if the data are to be recorded digitally. The data recording can be a simple chart recorder (for slow events), an instrumentation tape recorder, or a computer-based data recording system. In principle, all the instrumentation will be quite similar to the one used for patch clamp recording, and details are given in [2] in this volume. The two-electrode voltage clamp in oocytes is performed in accordance with techniques used for smaller cells. 2~a2 The main differences reside in the larger size of the oocytes, which implies a higher membrane capacitance. This introduces no complication, if only slow or no changes in potential are required for the experiments. However, when measuring voltage-gated channels, the speed of the clamp and the possibility of compensating for capacitive transients are of prime importance. The time constant for charging the membrane and hence the speed of the voltage clamp depend critically on the electrode resistance through the relation z = R C , where R is the current electrode resistance and C the cell capacitance. For time-critical applications, the cell capacitance can be reduced by using smaller (stage III2°) oocytes. 23 Intracellular Electrodes

Standard intracellular electrodes, whose tips have been broken, have resistances below 1 Mfl and are quite appropriate for intracellular electrodes in oocytes. Electrode pipettes are made from capillary glass containing a thin filament which ensures that the electrode filling solution reaches the tip. We use glass from Clark (Reading, England), types GC120TF-10 (thin), GC 150F-10 (medium), and GC200F-15 (thick). Pulling is best done using a standard intracellular electrode puller (David Kopf Instruments, Tujunga, CA) or a programmable electrode puller (see [3] in this volume). To avoid creeping of the pipette filling solution up the back end of the electrode, a small amount of dental wax or sticky wax is applied to the end after moderate heating in a small flame. Melted wax goes into the pipette along the filament for a few millimeters during application. After this, the electrode pipettes can be backfilled either with 3 M KC1 or with 0.5 M 29 D. S. Krafte and H. A. Lester, this volume [20].

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potassium sulfate (or aspartate), containing at least 30 m M KC1. The 3 M KC1 will be useful for most applications. The sulfate or aspartate solutions are to reduce the C1- load to the oocytes, and the KC1 is needed for the silver/silver chloride electrodes. The electrode pipettes can be stored for several days sustained in a covered container which has some water at the bottom to ensure high humidity. The electrode tips are normally in the submicron range and need to be broken at the very tip to decrease the resistance to the megohm range. This is done either under a separate microscope or by simply jamming the electrode against the bottom of the recording chamber before an experiment. Typical tip diameters will be in the range of 1 to 5/zm, giving resistances in the range of 2 to 0.4 Mfl. The electrical contact to the electrode filling solution is made through a silver chloride electrode. The silver chloride electrodes can be made from chlorinated silver wire, from silver wire immersed in melted silver chloride, or from a silver/silver chloride pellet. The lifetimes of the electrodes mentioned as well as their diameters are given in ascending order. Most voltage clamp amplifiers incorporate an electrode resistance measurement. The one from Polder (NPI Electronics, Tamm, Germany) allows electrode resistance measurements even with the electrodes positioned inside the oocyte, with a direct readout in megohms. Alternatively, the electrode resistance can be measured by applying current pulses (AI) through the Ag/AgC1 electrode and measuring the potential jump (AV) induced. The electrode resistance will be A VIA L As long as the electrodes retain their low resistance, they can be reused for several oocytes. Clogged electrodes can be cleared by applying pneumatic back pressure with a syringe. This process can be monitored by the extrusion of the electrode filling solution (which has a different refraction index) under the microscope. A patch clamp pipette holder can be used as a electrode pipette holder. For long-term experiments, the electrode filling solution should be of such an amount that the hydrostatic pressure approximately compensates the pressure from the surface tension within the pipette. If the hydrostatic pressure in the electrode allows a large net outflow of solution, the oocytes will bulge around the site where the electrode is inserted as the oocytes accumulate KC1. If, on the contrary, there is a net inflow, the electrode resistance will increase as intracellular fluid enters the pipette. A very slight net outflow is the best choice, since this will stabilize diffusion potentials at the electrode tip. Owing to the large size of the oocytes, the electrodes can be positioned with simple, coarse manipulators. It is important, however, that the setup be free of vibrations because otherwise oscillations will cause damage to the membrane where the electrodes penetrate the oocyte. A vibration-isolated

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table is usually not necessary in buildings where the floor is sufficiently quiet. This can be tested by observing the electrode tip under the set-up microscope while bouncing on the floor. A Faraday cage is usually not necessary for normal two-voltage electrode clamping, provided that standard grounding techniques are used (see [2] in this volume). The microscope can be a very simple one, either inverted or noninverted. The low magnification (5 - 10× objective) provides a comfortable working distance.

Adjusting Voltage Clamp Once the electrodes are in solution, the offset of the potential electrode should be cancelled to within + 1 inV. It is advisable to test this adjustment after every experiment as well, to ensure that there was no drift and that the potentials to which the o0cyte had been clamped were correct. Then the electrodes are positioned with the manipulators as depicted in Fig. 2. The use of the following (or any equivalent) procedure is advisable owing to the

i/ Top view :

FIG.2. Sideviewand top viewofthe oocytewiththe intraceUularelectrodespositionedfor penetration.

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fact that the oocytes are opaque, so that the actual site of oocyte penetration is not visible, particularly when using an inverted microscope. The positioning is done as follows. First, the plane even with the bottom of the oocyte is brought into focus (there are always enough particles or scratches on the bottom of the chamber). Then the rim of the oocyte is brought into focus, and the distance a can be estimated by the amount of refocusing necessary. Then the focus is brought up by about the same distance (b) so that the focal plane should be even with the top rim of the oocyte. The electrodes are brought into the positions shown in the bottom part of Fig. 2, where the electrode tips should be in focus. Axial movement of the electrodes will now ensure that the electrodes will penetrate the oocyte correctly. Obviously, for this procedure to work, it is important to have the ability to move the electrodes axially under a tilt of about 45 °. To prevent oocytes from moving away during penetration, the oocyte can be supported by first advancing the current electrode until the oocyte starts to move away. Then the potential electrode is inserted. A rough bottom (polypropylene mesh, Fisher, Pittsburgh, PA, Cat. No. 08670-185, or just simple scratches on the bottom of the experimental chamber) is helpful in holding oocytes in place during electrode penetration. The whole problem of perforating the vitelline envelope with microelectrodes and fixation of oocytes to the experimental chamber can be overcome if the viteUine envelope is removed as described below for patch clamping. This also makes the membrane more accessible to changes in the extracellular solution. Penetration of the potential electrode can be monitored through the resting potential recorded. The resting potential of oocytes can be anywhere in the range o f - 20 to - 80 mV. Monitoring the penetration of the current electrode is more difficult. The oocyte voltage clamp from Polder allows the measurement of the potential through the current electrode in the current clamp mode. Therefore, insertion of the current electrode is monitored in an analogous manner to the potential electrode. The Polder amplifier also features an audio monitor for the potential measured by either electrode, greatly simplifying electrode penetration while looking through the microscope. With most other amphfiers penetration of the current electrode is monitored using the current damp mode. For this, a current pulse is passed through the current electrode, which will cause a change in potential within the oocyte as soon as the electrode penetrates the cell. This pulsed change in cell potential is monitored through the potential electrode which has to be inserted first. The amplitude of the potential response will depend on the input resistance of the oocyte, which can be obtained by evaluating A V/AI. It will be low for leaky oocytes or for

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oocytes expressing large quantities of exogenous channels having high probabilities of being open under the experimental conditions present. Now that the two electrodes are inserted, the actual feedback circuit for clamping the transmembrane potential can be closed. To monitor the adjustments necessary for this step, voltage steps of about 10 mV are given as command potentials to the voltage clamp, and the current response is observed on an oscilloscope. Before closing the clamp, the gain of the feedback loop is set to a minimal value, and the holding potential is set to the desired value. Closure of the clamp will cause the membrane potential to approach the setting of the holding potential. Holding currents needed to maintain a potential of - 100 mV are in the range of 100 to 200 nA, with larger currents being an indication of a leaky oocyte. The gain can now be increased carefully. Higher gain will speed up the time response of the voltage clamp, as seen from the decrease in the time constant of the current response. Too much gain, however, will cause oscillation and normally irreversible damage to the oocyte and current electrode. This can be avoided by either limiting the current output or by using an automatic oscillation shutoff feature. Both possibilities are provided in the Polder clamp, and many other commercially available clamps have provisions for current limiting.

Improving Frequency Response The price for improved frequency response is an increase in noise. Therefore, no more gain than necessary should be used. The frequency response can be improved by higher gain, using series resistance compensation (not routinely necessary) and compensating for the potential electrode capacitance (capacitance neutralization). Electrode capacitance can be minimized by using a minimal solution level, just sufficient to cover the oocyte. Capacitive coupling between the two recording electrodes should be avoided, but the laborious procedures used for small cells like silver paint coating and subsequent isolation of the current electrode (effectively providing a grounded shield down to the very tip, see Refs. 21 and 22) are normally not required. For critical wide bandwidth applications, a simple grounded metallic shield around the current electrode reaching as close to the solution surface as possible is usually sufficient to reduce capacitive coupling between the electrodes. Capacitance transients can be mostly compensated by adding the differentiated command voltage step with appropriate amplitude and time constant to the current trace. Two components are normally sufficient to compensate more than 90% of the capacitance transients. This compensation is only needed when recording from fast, voltage-activated channels and will in general decrease the signal-to-noise ratio.

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Improving Signal-to-Noise Ratio For slower applications, several procedures may be helpful in improving the signal-to-noise ratio. For example, the bandwidth of the feedback loop can be limited. Some amplifiers provide a feedback limited in bandwidth to 10 Hz; other units provide a setting having a high de feedback gain. Otherwise, the lowest gain setting can be used. This, however, has the disadvantage that the clamp error will increase. The optimal solution in this case is to use an integrating feedback, which will eliminate the feedback error. This feature can even be used for fast potential-gated channels if the rise time of the pulse is adjusted accordingly. The Polder amplifier provides all the features described here. Responses should be filtered using the lowest cutoff frequency possible. The limits of current resolution will depend on this filter setting. The range of measurable current amplitudes spans from several tens of nanoamperes for slow processes to several hundred nanoamperes for the fast voltageactivated channels. Another limitation of current resolution is the presence of endogeneous channels and carriers in the oocyte membrane. For instance, depolarization above + 30 mV leads to development of a slow outward current, mostly owing to the presence of CaZ+-activated C1- channels. Several other channel types are present to various degrees, and great care should be taken to avoid these currents from contaminating the desired signals.

Changes in Solution In general, the two-electrode voltage clamp is stable for several hours, and extracellular solution changes are well tolerated. The perfusion can be simply gravity driven, with a suction pipette used for level control. While changing solutions manually with a Pasteur pipette or a syringe, the clamp feedback gain should be reduced, since otherwise some amplifiers tend to oscillate, particularly when close to a critical setting. Changes in fluid level will cause changes in electrode shunt capacitance and require readjustment of the capacitance compensation and neutralization. In some cases we have observed a dependence of current magnitude on fluid level. The reason for this is not quite clear, but it could be due to a hydrostatic access resistance dependence on the microvilli present to various degrees (depending on developmental stage) on the oocyte membranes. For critical applications, where an exact control of fluid level is mandatory, a fluidlevel controller (MPCU, Adams and List, Westbury, NY) is very useful. The intraceUular medium can be modified by injection of concentrated solutions. Up to 50 nl of solution can be injected while under voltage clamp with injection pipettes similar to the ones described for mRNA injection, provided that the oocyte is bare of the viteUine envelope.

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Problems In most cases, problems in recording currents from oocytes damped with two electrodes arise from the electrodes themselves. Therefore, the possibility of measuring the electrode resistance during the experiment is very helpful. Any increase in electrode resistance deteriorates the clamp performance. This can often lead to oscillations. Also, increases (and decreases, when using capacitance neutralization) in electrode shunt capacitance can cause oscillations. This can be the case particularly when changing the solution levels. Most of the problems (and some cures) related to the electrodes and oocyte preparation have been dealt with above. But how can the performance of the voltage clamp itself be tested? Figure 3 is a schematic diagram of the most simple equivalent circuit of an oocyte including its electrodes. It assumes intracellular electrode resistances of l M£), and the membrane resistance (1 Mf~) and capacitance

1 MOhm

1

1KOhm

VBath

I Bath

FIG. 3. Simple equivalent circuit for the electrical impedance of an oocyte. The upper part corresponds to the intracellular side.

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(100 nF) correspond to average values encountered in oocytes. The 1-kfl resistances represent mostly the bath electrode resistances. It is convenient to incorporate the circuit of Fig. 3 into a small (grounded) box with connections that can easily substitute the ones leading to the electrodes used in a real experiment. The electrical equivalent of an oocyte is also very helpful in getting acquainted with the voltage clamp amplifier being used. The equivalent circuit does not emulate the diffusion potential of 50 to 60 mV from real electrodes filled with a 3 M KC1 solution. Patch Clamp Recording from Oocytes Prior to patch clamping on oocytes, the vitelline envelope needs to be removed. This is done mechanically after osmotic shrinkage in a solution of the following composition (in mM): 200 potassium aspartate, 20 KC1, 1 MgC12, 5 E G T A n K O H , 10 HEPES-KOH, pH 7.4. After 3 to l0 rain in this solution, the transparent vitelline envelope becomes visible and can be removed mechanically with two pairs of blunt-tipped forceps. Extreme care has to be taken to avoid any damage to the cellular membrane since the bare oocytes are very fragile; they are particularly sensitive to air exposure. After a brief wash in normal frog Ringer's solution (NFR, in mM: 115 NaC1, 2.5 KC1, 1.8 CaC12, l0 HEPES, pH 7.2), the oocytes are transferred to the final experimental chamber. The bare oocytes will attach to any clean surface within minutes. Thereafter, any movement of the oocyte will cause membrane damage. More details of the procedure are described by Methfessel et al. ~2 Patch Pipettes

Pipettes for single-channel recording are similar to the ones used for small cells. However, a thick coating of a silicone (RTV615) curing agent, high enough up the pipette, is necessary because the large size of the oocytes requires deeper immersion of the pipette in the solution. The pipettes for macropatches will have tip opening diameters of 2 to 8/lm, giving resistances of 0.6 to 2 M~, depending also on taper shape and pipette solution. The taper should be as short as possible to avoid unnecessary access resistance. A hard aluminum silicate glass such as the one supplied by Hilgenberg (Malsfeld, Germany), or the type 7052 (e.g., A-M Systems, Everett, WA), gives smooth rims for macropipettes and low noise. Seal Formation

The formation of a seal on oocytes is similar to seal formation in other cells for the small pipettes usually used for single-channel recording. ~2For

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macropatches, however, seal formation can be very slow (3-5 min). Also, the suction used to obtain seals is much smaller than when using small pipettes. It is convenient to be able to measure the pressure in the pipette. Positive pressures when entering the solution range between 10 and 40 m m H20; the suction during seal formation is in the range of 100 to 200 m m H20. The suction should be stopped when a seal resistance larger than 1 Gf2 is achieved. The MPCU fluid level controller provides an adjustable pressure outlet which can be used to measure and control the pipette pressure. A slight depolarization (10-20 mV) of the patch is sometimes helpful. For macropatches seal formation becomes particularly slow when the seal resistance is in the 10 to 20 Mr2 range. A few oocytes will form gigohm seals as quickly as in small cells. Seal resistances up to 100 Gf2 can be obtained with small and large pipettes, having less than 200 fA root mean square (rms) of current noise. To avoid suspended particles, the solutions used in the bath and in the pipette should be filtered. The most disturbing conductance in some oocytes is an endogenous stretch-activated channel with a conductance of about 40 pS. It can be easily identified by applying gentle suction to the patch pipette, which will cause the channel activity to increase. The open probability is also potential dependent, increasing with increasing transmembrane potential. Concentrations of 10 to 100 p M gadolinium will block these channels to a large extent. 24 These channels seem to be totally absent in more than 50% of the oocytes from the same animal. For long depolarizations to very positive potentials, a slow outward current develops. This current is partially carried by C1- through Ca2+-acti vated C1- channels and by a slowly activating potassium conductance. The density and number of these channels are highly variable from oocyte to oocyte. Chloride ions can be eliminated from the solution by substitution with methanesulfonic acid.~°,23Alternatively, the current can be blocked by 0.3 m M niflumic acid or by 0.5 m M flufenamic acid. 26 Injection of EGTA or BAPTA (50 nl/oocyte at 0.1 M) also reduces the early response to inflowing Ca 2+. The most stringent requirements regarding the absence of any ion conductance apply when measuring gating currents. A detailed description of methods used for these particular measurements is found in [22] in this volume.

24 X. C. Yang and F. Sachs, Biophys. J. 53, 412a (1988). 23 N. Dascal, T. P. Snutch, H. Ltabbert, N. Davidson, and H. A. Lester, Science 231, 1147 (1986). 26 j. p. Leonard and S. R. Kelso, Neuron 4, 53 (1990).

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Other Configurations Besides the cell-attached patch the only other easily viable configuration for macropatches is the inside-out configuration. This is obtained by abrupt withdrawal of the patch pipette from the oocyte after seal formation, in a solution normally containing high K + and no Ca 2+ [e.g., (in mM): 100 KC1, 10 EGTA, 10 HEPES, pH 7.4]. Oocytes will survive for several hours in this solution as external medium, and several patches can be obtained from the same oocyte. Some batches of oocytes, usually from the same animal, do not survive the excision procedure. On rare occasions vesicles are formed during patch excision. This is in most cases a consequence of a slow excision. These vesicles are difficult to detect but are indicated by the appearance of a slow capacitive transient after excision. Therefore, before patch excision, the capacitance transient should be compensated and observed carefully during excision. With small patches, it is possible to disrupt the vesicle by brief exposure to air. With macropatches, this procedure leads to destruction of the patch in most cases. A better way of disrupting the vesicle is to carefully approach the pipette to a small, freshly made silicone (RTV615) sphere until a change in the capacitive transient is seen. This leads to rupture of the outer membrane of the vesicle and leaves the inside-out membrane intact in more than 95% of cases. Most patches will tolerate extensive solution changes. The stability of even large patches may be due to the fact that the membrane forms an omega shape within the pipette tip. The tendency to form this omegashaped patch is increased in ooeytes after long incubation times (7 or more days after injection of the mRNA). Formation of an extensive omega patch can be detected during seal formation by the development of a slow capacitance transient, which cannot be compensated well, leaving a biphasic capacitive transient at best. These patches are not suitable for the study of kinetic effects, since solution changes as well as potential changes will not have unrestricted access to the membrane regions in close contact with the pipette wall. Also, ion accumulation effects should be considered in these cases.

Distribution of Ion Channels and Seasonal Variations The distribution of channels is not uniform over the membrane of the oocyte. The magnitude of variation of the channel densities depends on the channel type. In our experience, sodium channels seem to be more uniformly distributed than potassium channels. Therefore, prediction of current size in a patch from the measured current size using a two-dec-

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trode voltage clamp is more accurate for the former channel type. Wholeoocyte currents of about 10/tA should yield patch currents of the order of 50 to 100 pA. When recording potassium currents from oocytes (expressing more than 10 gA of whole-cell K + current), some patches have practically no current whereas patches from other regions will have large currents. Therefore, several regions have to be explored with various patches in order to localize an area of high channel density. The regions of high channel density are relatively large, so that four measurements around the accessible portion of an oocyte are usually sufficient for localizing such a region. We have not found any correlation of regions of high current density with the animal or vegetal pole in oocytes, despite numerous rumors in that respect. There are, however, clear seasonal variations. In the northern hemisphere, ion channels tend to give better results during the winter and ionic

FIG.4. (A) "Tower"arrangementfor conventionalpatchclampmeasurements,whichcan also be usedto maketwo-electrodevoltageclampexperimentsin oocytes.

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ELECTROPHYSIOLOGICAL RECORDING FROM OOCYTES

Fzo. 4. (B) Close-up view of setup.

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pumps seem to work better during the summer. In general, oocytes are more amenable for patch formation during the winter. The best results obtained for voltage-dependent channels are during the period from November to April. This period surprisingly cannot be shifted by using Xenopus acutely imported from South Africa. Using Two-Electrode and Patch Clamp Amplifiers Simultaneously When Na + channels are expressed at high densities, in particular channels that have been modified and will show no or very slow inactivation, the oocytes will spontaneously depolarize or even tend toward the sodium reversal potential. This persistent depolarization can be avoided during patch clamp experiments by holding the cell at hyperpolarizing potentials with a two-electrode voltage clamp. A high holding potential will also reduce the current required to hyperpolarize the patch, particularly with low resistance seals. However, the two-electrode voltage clamp introduces noise into the system. This noise can be drastically reduced by limiting the gain and consequently the bandwidth of the feedback amplifier of the two-electrode amplifier. Some two-electrode amplifiers have a mode that reduces the bandwidth of the feedback loop to l0 Hz (e.g., Polder), a speed sufficient to compensate for slow drifts in membrane potential. Figure 4 gives the setup for both two-electrode voltage clamp and patch clamp experiments, which can also be used for conventional patch clamp experiments on small cells.

Comparison of Recordingsfrom Two-Electrode VoltageClamps andfrom Macropatches in Oocytes The two-electrode clamp of oocytes offers a series of advantages over patch clamp recording from macropatches. It is simpler, more stable, allows recording at lower channel densities, and the extracellular solution is easily changed. In addition, it is not so sensitive to varying oocyte conditions since the formation of gigaseals is not required. However, the kinetics of fast processes cannot be resolved owing to the high cell capacitance in combination with an upper limit of resistance of the current electrode. The temporal resolution is of the order of 200 to 1000/lsec. In addition, kinetics will depend on several parameters of the voltage-clamp amplifier (i.e., rise time limit or de gain) as well as on the morphology of the oocyte membrane, which is extensively invaginated (microvilli). For example, extreme discrepancies can be obtained when recording from inactivating Shaker potassium channels in the two-electrode clamp mode. In some batches of oocytes these currents will show practically no inactiva-

USEOF STAGEII-III Xenopus OOCYTES

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tion. Currents from macropatches or ensemble averages from single-channel currents from these same oocytes will have reproducible, fast inactivating kinetics. Therefore, extreme caution has to be taken when trying to analyze channel kinetics from two-electrode data. Macropatches achieve a high temporal resolution by electrically isolating a small area of membrane through pipettes with resistances of the order of 1 Mfl. Therefore, the temporal resolution is of the order of 50 to 200/~sec. The kinetics of fast processes are much more reproducible, probably because the microvillar structure is made more accessible in the membrane patch under the pipette. The solutions on both sides of the membrane are well defined, and exchange of the "intracellular" solution is possible in inside-out patches. Exchange of the extracellular solution is more difficult. This requires either the perfusion of the patch pipette or recording in the more difficult outside-out configuration. Recording from single channels naturally provides more information on the actual conformational changes between the open and closed states. Macropatches allow the characterization of ionic currents equivalent to the whole-cell configuration. Therefore, each of the various recording modes has its own advantages (and disadvantages), and a combination of several modes will in most cases allow a detailed characterization of the parameters under study.

[20] U s e o f S t a g e I I - I I I X e n o p u s O o c y t e s t o S t u d y Voltage-Dependent Ion Channels

By

D O U G L A S S. K R A F T E a n d H E N R Y A . LESTER

Introduction Pioneering studies on the use of Xenopus oocytes to express ion channels were first performed by Sumikawa et al.~ on acetylcholine receptors and later by Gundersen et aL2 on voltage-gated channels. Since this work, the Xenopus oocyte has become a convenient and robust system for expression of a variety of ion channels. Because the oocyte is able to express proteins from different species and tissues in a common membrane environment, thus removing one source of variability, an important use is to compare the detailed functional characteristics of channels induced by i K. Sumikawa, M. Houghton, J. S. Emtage, B. M. Richards, and E. A. Barnard, Nature (London) 292, 862 (1981). 2 C. B. Gundersen, R. Miledi, and I. Parker, Proc. R. Soc. London B 220, (1983). METHODS IN ENZYMOLOGY, VOL. 207

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