Introduction

Whole-cell voltage clamp of isolated heart cells has led to an explosion of information about ionic currents in the heart. This article briefly summarizes the use of this variant of the patch-clamp technique in isolated cardiac myocytes with an emphasis on practical suggestions. For an overview of the basics of the whole-cell voltage clamp technique in general, I would advise the beginner to read the classic article by Hamill et al. (1981). If the reader wishes to learn more about the details of whole-cell recording, I suggest the book by Sakman and Neher (1998) and the Axon Guide.

This description of the use of whole-cell technique in heart cells is divided into three sections: isolation of the cells, the actual experimental recording, and “tips and pitfalls”. Assuming that the investigator has adequate equipment and a basic understanding of the whole-cell technique, the quality of the isolated heart cell preparation is typically the single most important factor determining the success or failure of whole-cell recording experiments. Adult mammalian ventricular myocytes, the preparation discussed in this article, are extremely fragile and sensitive to seemingly minor alterations in isolation conditions or solutions. Thus, the importance of taking extreme care in the isolation of ventricular myocytes for whole-cell recording cannot be overstated.

Isolation of Adult Mammalian Myocytes for Whole-Cell Recording

My laboratory uses the isolation method described below for the recording of macroscopic currents (primarily calcium and potassium currents) from adult rat and guinea pig ventricular myocytes (e.g., Kawamura & Wahler, 1994). We have also applied this method, on occasion, to rats as young as 10-12 days and to adult mice and rabbits. Using somewhat different isolation procedures, we have also applied these whole-cell recording techniques to isolated (and sometimes cultured) neonatal rat and embryonic chick ventricular myocytes. However, we try to avoid using cultured cardiac myocytes, since we have found that the electrophysiology of cells is altered within hours of culturing. Similar methods for the isolation of adult ventricular myocytes for a variety of purposes are described in several other articles in this Help section. I will particularly emphasize those factors that are extremely important for whole-cell recording but may not be important (nor even appropriate) for the isolation of myocytes for other purposes (e.g., biochemical assays).

An adult rat is injected with heparin (200 units) approximately 30 min prior to removal of the heart. Following appropriate anesthesia, the heart is rapidly removed and dropped into a chilled (£ 4°C) nominally Ca-free Isolation Solution. Speed is of the essence. Any extraneous tissue (e.g., lung) can be removed after the heart is in the chilled solution. The heart is weighed in the chilled Isolation Solution and gently squeezed to remove blood. Then the heart is quickly mounted on a Langendorff perfusion apparatus (Baker Double Coil #AH 50-8382 and glass cannula #AH 59-7926, Harvard Apparatus, Inc. Holliston, MA). and retrogradely perfused for 5 min (at 37°C) with an oxygenated (95% O₂ - 5% CO₂) and humidified Isolation Solution to remove any remaining blood. We use a simple gravity system for perfusion of the coronary arteries (70 cm column of solution above the coronary ostia), though pump perfusion can also be used.

Following the 5 min initial perfusion period, the heart is perfused with the Enzyme Solution containing 45 U/ml of crude collagenase (Type II, Worthington Biochemical Corp. Lakewood, NJ) for 16.5 min/g heart weight at 37°C. Following the digestion period, the heart is taken down, the ventricles minced into approximately 1 X 1 mm chunks, and the tissue gently dissociated by repeated trituration over a period of 10 min in Storage Solution at 37°C. The wide-bore trituration pipettes are made by cutting the tapered portion of glass Pasteur pipettes at different points, fire-polishing the cut ends with a Bunsen burner and then silanizing the pipettes to minimize myocyte adherance. Subsequently, the cells are filtered (approx. 100 mM mesh), centrifuged at 60 X g for 3 min, and "cleaned" for 12 min at 37°C with the Cleaning Solution (Yazawa et al., 1990), centrifuged at 60 X g for 3 min, and resuspended in the Storage Solution. The cells are then centrifuged again at 60 x g for 3 min and then resuspended a final time in Storage Solution. The cells are kept at room temperature and used for whole-cell recording within 3-4 hrs of isolation.

Experimental Recording of L-type Calcium Current

The following description is our method for recording the macroscopic L-type calcium current (I_Ca) in rat ventricular myocytes. We have recorded other currents from these and other cells using different solutions and voltage protocols.

We pull our pipettes on a P-87 Flaming-Brown Puller (Sutter Instrument Co., Novato, CA) and fire-polished on a MF-9 Microforge (Narishige Co., Ltd. East Meadow, NY). The pipette glass we use is 1B150F borosilicate (Corning #7740) glass from World Precision Instruments, Sarasota, FL). While softer “patch clamp” glasses (e.g., Corning #7052 or #8161) can often give higher resistance seals than borosilicate glass, we avoid them due to the fact that they have a very high lead content. Coating the pipettes with a hydrophobic elastomer is not necessary for recording these relatively large currents in adult mammalian cells.

A suspension of myocytes is placed into a bath chamber on the microscope stage and allowed to settle for a few minutes in order to attach lightly to the glass bottom of the chamber. The inverted microscope (Diaphot, Nikon Instruments , Melville, NY) sits on a vibration control table (Newport Corporation , Irvine, CA) in order to minimize mechanical vibration which could disrupt the seals. The pipette is filled with Pipette Solution (see Table) and then attached to a pipette holder with side port. A length of flexibile tubing is attached the side port for subsequent suction. The pipette holder is attached to the amplifier headstage, which is in turn mounted on a hydraulic micromanipulator (model MO-303 Narishige Co., Ltd.).

Both the Bath Solution and Pipette Solution contain cesium in place of potassium (Table) The Pipette Solution and Isolation Solution, and Enzyme Solution are all filtered with 0.2 mM, low protein binding filters prior to use.

The pipette should be brought down onto the cell at a fairly steep angle (>45°). Since the myocytes are flat, a too shallow angle will reduce the probability of forming a tight membrane-glass seal. Once a gigohm seal is formed between the glass and cell membrane (cell-attached mode), capacitance transients are generated by applying 5 mV voltage pulses to the pipette. An Axopatch-1D amplifier, pClamp software and Digidata 1200B hardware (all from Axon Instruments , Union City, CA) are used to deliver the voltage protocols and acquire all the data, which are stored on a PC for subsequent analysis (by the pClamp software). In the cell-attached mode, the capacitance transients due to the charging of the small membrane patch are minimal.

The whole-cell configuration is subsequently attained by rupturing the patch of membrane within the pipette by either suction (generally applied by mouth or alternatively by a gas-tight syringe) or by “zapping” the membrane. This latter zapping is a feature of some amplifiers (including the Axopatch-1D) and consists of a very short duration, high voltage (> 1V) pulse, which will often cause the dielectric breakdown of the small patch of membrane in the pipette tip. Attainment of the whole-cell mode is evident as a sudden dramatic increase in the duration of capacitance transient, due to charging of the entire cell membrane. Once the investigator has attained the whole-cell recording configuration, much of the cell capacitance and series resistance can be compensated for electronically by controls on the amplifier. The breakthrough, i.e., the attainment of the whole-cell configuration, is evident with such large cells as a very large increase in the capacitance transient. Once access to the cell interior is obtained, several minutes are often allowed to pass to enable cell dialysis and current levels to stabilize. Once the currents are stable, the experiment can proceed.

In the absence of potassium, the holding current between pulses should be minimal. Calcium currents are elicited by repeated 200-250 ms pulses to 0 mV from a holding potential of –80 mV, following a brief prepulse to –40 mV, at a frequency of 0.2 Hz. The use of the prepulse inactivates the fast sodium current and the replacement of potassium with cesium eliminates the potassium currents. Current-voltage (I-V) curves may also be generated by pulsing in 10 mV increments over the range from –40 mV to +50 mV. The use of room temperature (22-25° C) results in a slower and smaller I_Ca, which makes adequate voltage control in these cells more likely. With adequate voltage control, the I-V curve should have a smooth gradual activation component over several tens of millivolts (see Kawamura & Wahler, 1994, for discussion). The use of room temperature, together with a low stimulation frequency, also minimizes the amount of spontaneous “rundown” of the current, a problem that often plagues recording of I_Ca.

Following acquisition of the current data, calcium currents are measured from the stored data (by pClamp software from Axon Instruments) as the peak inward current minus the current at the end of the pulse. The kinetics of the current may also be analyzed.

Tips and Pitfalls

It is very important to obtain a lot of crude collagenase which is optimal for your particular needs. We use different lots of collagenase for rat versus guinea pig ventricular myocytes. We generally also use different lots for biochemical assays (e.g., for cyclic nucleotide assays) in which yield is a very important factor, for cell contraction, and for patching of adult rat ventricular myocytes. Worthington Biochemical Corp has a “collagenase sampling program” in which they will send you samples of a few different lots to try, while keeping several grams of each lot on reserve. Even if a company does not provide free samples, one can always purchase small amounts of different lots to try. Once we have found a lot of collagenase that works well for a specific purpose, we purchase enough to last for at least two years. It may take a fair number of samples before you find a lot that is optimal. Many lots of collagenase will give good yields of rod-shaped cells that appear healthy, but will not be optimal for patching. High yields of healthy appearing cells, or Trypan blue exclusion, do not guarantee high resistance seals or stable currents. In fact, attempts to optimize yields are probably counter-productive for electrophysiology studies. That is, my experience has been that attempts to increase the percentage of good-looking cells by such interventions as gradually increasing the final calcium concentration often seem to keep “borderline” cells alive that are not optimal for whole-cell recording.

Another very important factor is the quality of the water used for all solutions. The deionized water typically available in laboratories is not adequate for isolating adult mammalian ventricular myocytes for whole-cell recording. We use a Milli-Q System from the Millipore Corporation, Bedford MA. We use tissue culture grade chemicals wherever available. Additionally, we clean all glassware using a tissue culture cleaning solution and acid wash. It is wise to avoid glassware cleaners that contain chromium or other toxic substances. We also acid-wash our pipette glass prior to use, which appears to improve the seals.

One important factor that is often not considered in whole-cell recording experiments is the importance of the pipette [Ca²⁺]. Almost all whole-cell recording protocols include the calcium chelator EGTA in the pipette. In many instances, no additional Ca²⁺ is added to the pipette. We have empiriically determined that the measured free pipette [Ca²⁺] in such pipette solutions is less than 10^-9 M. Dialysis of the cell interior with such an unphysiologically low calcium concentration causes numerous alterations in both the electrophysiology of the cell and the regulatory mechanisms responsible for modulating channel activity. In some instances, these alterations are entirely unpredictable. For example, we have found that the use of low [Ca²⁺] in the pipettes reduces the I_Ca response to b-adrenergic stimulation, but enhances the I_Ca response to the phosphodiesterase inhibitor IBMX. This suggests that the unphysiologically low [Ca²⁺] in the typical pipette solution causes an alteration in the normal balance between cAMP production and degradation by the myocyte. Therefore, we include sufficient calcium in the pipette solution (together with EGTA) to maintain the free [Ca²⁺] at a physiological 10^-7M.

Table - Isolation and ICa Solutions:¹

	Isolation Solution²	Enzyme Solution²	Cleaning Solution	Storage Solution	ICa Bath Solution³	ICa Pipette Solution
Albumin⁴	1 mg/ml	1 mg/ml
CaCl₂		0.005			1.0	3.8
Collagenase⁵		45 U/ml
CsCl					4.0	120.0
DNAase I, Type IV			30 U/ml
EGTA			0.5	0.5		10.0
Glucose	11.1	11.1	10.0	10.0	11.1
Glutamic Acid			50.0	50.0
HEPES			10.0	10.0	10.0	10.0
KCl	2.6	2.6	40.0	40.0
KH₂PO₄	1.18	1.18	20.0	20.0
KOH			70.0	70.0
MgSO₄	1.18	1.18			1.2
MgCl₂			3.0	3.0		6.0
Na₂ATP						5.0
NaCl	118.5	118.5
NaHCO₃	14.5	14.5			135.5
Protease, Type XIV			0.1 mg/ml
Taurine			20.0	20.0

¹All concentrations given in mM, except where indicated. All chemicals obtained from (Sigma-Aldrich Co ., St. Louis, MO), except where indicated below. The pH of all solutions is 7.4, except the Pipette Solution which has a pH of 7.2.

²Oxygenate with 95% O₂ – 5% CO₂.

³This bath solution is specifically for rat cells. We use a slightly different bath solution with higher [Ca²⁺] for other species.

⁴Pentex^Ò Bovine Albumin, Fraction V, very low endotoxin, fatty acid-free (catalog # 82-047-5 from Serologicals Corporation , Norcross, GA.

⁵Crude Collagenase Type II, Worthington Biochemical Corp.

References:

Axon Guide (out of print, but available online)

http://www.axon.com/MR_Axon_Guide.html

Hamill OP, Marty A, Neher E, Sakmann B, Sigworth F. Improved patch-clamp techniques for high resolution current recording from cells and cell-free membrane patches. Pflügers Arch 391:85-100, 1981.

Kawamura A, Wahler GM. Perforated-patch recording does not enhance effect of 3-isobutyl-1-methylxanthine on cardiac calcium current. Am J Physiol 266:C1619-C1627, 1994.

Sakman B, Neher E. Single-channel recording, 2^nd edition. Plenum Press, New York, NY. 1995.

Yazawa K, Kaibara M, Ohara M, Kameyama M. An improved method for isolating myocytes useful for patch-clamp studies. Jpn J Physiol 40:157-163, 1990.