Muscle Resting Potential: Ionic Basis and Nernst Potentials

Introduction

If you have not already done so, read Appendix A, Crayfish Neuromuscular Preparation, for background. In this lab exercise, you will examine the ionic basis of the resting potential in the superficial flexor (SF) muscle of of the crayfish abdomen. In doing so, you will learn and practice intracellular recording techniques.

The resting potential (RP) of a cell is the difference in electrical potential between the inside of the cell and its surrounding medium. This potential is determined by the distribution of ions across the cell membrane. To help determine whether a given ion is involved in setting a cell’s resting potential, we can compare the calculated equilibrium potential of that ion with measured values of the resting potential (Wright, 2004; Purves et al., 2012).

The equilibrium potential (E_ion) of an ion is the electrical potential at which the electrical forces and chemical gradient forces acting on that ion to move it across a permeable membrane exactly offset each other. Thus, when the membrane potential is equal to E_ion, there is no net movement of that ion across the membrane. This equilibrium is described by the Nernst potential:

One way to show that a given ion can contribute to a voltage across a membrane is to change the concentration of that ion in the external fluid and measure the potential across the membrane (Hodgkin and Horowicz, 1959). If the membrane potential changes in the direction predicted by the Nernst equation, then that ion is involved in the potential. In special circumstances, one can even change the internal concentration of an ion to test its role in setting the membrane potential (Baker et al., 1962). You will use the first approach to examine the ionic basis of the resting potential of crayfish muscle; it can also be used to determine the ionic basis of action and synaptic potentials. Your experiments will test three hypotheses: (1) K⁺ contributes to the RP, (2) K⁺ is the only ion responsible for the RP, (3) Na⁺ contributes to the RP.

Dissection

Before starting, look at the methylene blue-stained specimen that was prepared in advance (it will resemble Figure A.2). This specimen shows nerve 3 leaving the ganglion, projecting to the superficial flexor (SF) muscle, and branching out over the surface of the muscle. If more than one segment has been dissected, note the different orientations of the superficial flexor muscle fibers in different segments. Use this specimen to get a feel for the depth of the muscle below the cuticle.

• Video 4.1, Muscle Dissection and Recording, shows the entire dissection and recording procedure. You may want to watch that version during the lab after reading the descriptions below.
• Video A.1, Preparing Crayfish Abdomen. Place a crayfish in the freezer or in ice and leave it until it has stopped moving. Cut off and keep the tail; place the remainder in the freezer. Pin the tail, ventral surface up, in a dissecting dish and cover it with normal crayfish saline.
• Video A.2, Removing Swimmerets. Remove and discard the swimmerets.
• Choose any intact tail segment for the dissection.
• Video 4.2, Exposing Superficial Flexor. Dissection to expose the muscle consists of three incisions. This crayfish, like many others, has a blue stripe on the ventral midline. It lies right over the ventral nerve cord. Before starting the dissection, find the muscle attachment point (the white line in Figure A.2). Take care not to cut through this line.
  1. Turn the dish so that the cut end of the tail faces away from you. Using a scalpel with a sharp (preferably new) blade, make an incision along the midline between the two sternites. This cut should go all the way through the cuticle. If your crayfish has a blue stripe, this cut will go along the center of the stripe.
  2. With forceps, lift the cuticle at the posterior end of the first incision and cut along the sternite toward the lateral edge. Although the video shows this dissection being done with a scalpel, you should use fine scissors if they are available. Hold the scissors so that the blades are flat and move parallel to the muscle during cutting. If the blades instead move up and down, they can damage the muscle. If using a scalpel, keep the blade shallow and parallel to the muscle, as shown in the video.
  3. Lift the flap of cuticle so that it stands up straight from the muscle insertion (the white line). Use scissors (preferred) or a scalpel to cut the flap away. Again, keep the blade flat and avoid touching the muscle.
• After the dissection, rinse all tools with fresh water and lay them aside where they will not be damaged.

Recording

Figure 4.1 shows the setup for intracellular recording; Video 4.3, Muscle Recording, shows the recording sequence. Before starting to record from the muscle, empty the saline out of the dissecting dish and replace it with fresh cold saline. You should replace the saline with fresh cold saline every half hour or so, but there's no need to interrupt a good recording to do this.

Pull a pair of glass microelectrodes and place them, sharp end up, in a small (10 ml) beaker with a small amount of 3 M KCl. After a few minutes, the tip of the electrode should have filled with KCl (Figure C.5). Use a fine syringe to fill the rest of the electrode with KCl (Video C.2, Electrode Filling); then put it in the electrode holder. With the micromanipulator, place the tip of the electrode in the saline over the exposed muscle and measure the electrode resistance. (The procedure for measuring resistance varies with different amplifiers; ask your instructor for help.) A resistance of 10 to 20 MΩ is best. The higher the electrode resistance, the easier it will for the electrode to penetrate the muscle fiber. If the resistance is low (0 to 5 MΩ), then the electrode is broken and must be replaced. At first, this may happen frequently, even when you think you handled the electrode gently. Be sure to dispose of broken or used electrodes safely in a closed canister; it is most unpleasant to be jabbed by a forgotten electrode! If the resistance is greater than 20-30 MΩ, the electrode may be blocked and needs replacement. For more information on intracellular recording, see Appendix C, Recording Tips.

At high magnification, focus the microscope on the apparent tip of the electrode. (The actual tip is too small to be seen and is somewhat lower than what you are focused on.) Adjust lighting for best visibility of the electrode. Try placing a light source such that light comes in from the side of the preparation dish and highlights the muscle and electrode. (If your light source has two fiber-optic branches, place one branch at the side and one above the prep.) Now bring the electrode closer to the muscle, refocusing along the way, until the visible part of the electrode appears to be a short distance from the muscle surface. At this point, looking under the microscope is no longer useful because the electrode tip is so small, it is invisible. Instead, watch the oscilloscope and use the fine-adjust knob of the manipulator to advance the electrode. When the oscilloscope trace starts to wobble slightly, the electrode is just pressing against a muscle fiber. You can either advance another quarter-turn with the manipulator or use the Buzz button on the amplifier to get into the cell. When the electrode is in a cell, the oscilloscope should indicate a negative voltage, the RP. This sequence is shown in Video 4.3, Muscle Recording, for two muscle fibers. Be careful not to penetrate the muscle too deeply. This may break the electrode tip and damage the muscles, or it may cause you to record from deeper flexor muscles that do not experience good saline exchange.

Before penetrating a muscle fiber with the electrode, be sure that the amplifier output is zero. Adjust the DC Offset knob of the amplifier to achieve this. To get an accurate measurement of the RP, you must do this before each recording.

Experiments

Practice impaling muscle fibers until you can repeatedly record RPs of at least −50 mV from muscle fibers, then move to a new, undamaged part of the SF muscle (either a new part of this muscle or another muscle in a different segment or on the other side). Record the resting potentials of five fibers in normal crayfish saline. Now you are ready to change the external K⁺ concentration and observe the effect on RP. See Table 4.1, below, for composition of crayfish salines (van Harreveld, 1936; Wallin, 1967). You will record resting potentials in four solutions of differing K⁺ concentrations.

The first solution is normal crayfish saline, which contains 5.4 mM K⁺. There is also a stock solution of high K⁺ saline, with 60 mM K⁺. This will be the fourth solution. (To preserve the osmotic balance, NaCl has been reduced in this saline.) Make two intermediate solutions with known K⁺ concentrations by mixing the two stocks. For example, 85 ml of normal saline and 15 ml of high K⁺ saline make 100 ml of saline with 13.6 mM K⁺; 60 ml of normal saline and 40 ml of high K⁺ saline make 100 ml of saline with 27.3 mM K⁺(the preparation dish holds about 100 ml of saline). Consider: why do we suggest these concentrations instead of evenly spaced ones like 23.6 and 41.8 mM?

Change from normal saline to the lower-concentration intermediate K⁺ saline and let the tail sit for at least 5 min before recording. Record the resting potential from another five fibers. Repeat this procedure for the next-higher-concentration intermediate K⁺ saline and then for high K⁺ saline. When finished, return the tail to normal saline, rinsing several times, and repeat the measurement at the normal concentration. Why is this last repeated measurement important?

Move to a new segment and compare resting potentials in normal saline (205 mM Na⁺) and low sodium saline (2.3 mM Na⁺). (Choline chloride replaces NaCl, but there is still 2.3 mM NaHCO₃ as a buffer.)

Further Exploration

As the experiment is outlined above, you draw conclusions from the mean of recordings from several fibers. You could attempt a more statistically powerful but more difficult approach, recording the resting potential of the same muscle fiber at each concentration. This approach requires great care to change saline concentrations without disturbing your recording.

You can investigate active, energy-requiring contributions to the resting potential (e.g., the Na⁺/K⁺ ATPase ion pump) (Kerkut and Thomas, 1965; Jones, 1989) by increasing the temperature of the saline, which will increase the activity of this ion pump; by reducing external [K⁺]; or by adding Na⁺/K⁺ pump blockers such as ouabain (Wang and Huang, 2006).

Neuromodulators and neurotransmitters can also influence the resting potential by opening or shutting channels that move the membrane potential toward or away from the E_ion of a particular ion. For example, closing resting K⁺ channels with thyroid-releasing hormone depolarizes rat motor neurons (Nistri et al., 1990). You can examine this phenomenon with the inhibitory transmitter GABA, which opens Cl⁻ channels in crayfish muscle. Like K⁺, Cl⁻ has a negative Nernst potential. See Appendix D, Pharmacopeia, for suggested concentrations.

Lab Cleanup

During the lab, be sure to immediately discard used glass electrodes in the appropriate container. After the lab, remove your last electrode from its holder and discard it as well. Clean up any spilled saline and rinse the ground electrode with distilled water. Put the crayfish tail in the freezer along with the frozen heads and rinse the dissecting dish with fresh water.

Questions

1. Why is the resting potential important for neurons and muscles? Would you expect other types of cells, such as red blood cells, to have resting potentials? How do K⁺ leak channels differ from the K⁺ channels that help to repolarize the membrane potential during an action potential (Lesage and Lazdunski, 2000; Honoré, 2007)?
2. Calculate Nernst potentials of K⁺, Na⁺, Ca²⁺, and Cl⁻ in normal saline, using values from Table 4.1. Which ones come closest to your measured resting potential values in normal saline?
3. Is the resting potential in normal saline statistically different from the Nernst potential of K⁺ using the literature [K⁺]_i (use a 1-group t-test)? If so, why might this be?
4. Calculate the mean and standard deviation of the resting potentials recorded in each test condition. Are the differences in mean resting potentials you measured statistically significant? Use an analysis of variance (ANOVA) with post hoc t-tests. Make a graph of mean resting potential vs. log of external [K⁺] and calculate the slope of the relationship. What sort of curve does the Nernst potential predict these points should follow? What should be the slope of the curve?
5. How did changing the external Na⁺ concentration affect the resting potential? What does this suggest about the relative importance of K⁺ and Na⁺ in the resting potential? About the permeability of a resting membrane to K⁺ and Na⁺?
6. If the resting potential is less negative than the Nernst potential for K⁺, there must be a driving force to move K⁺ out of the cell, thus hyperpolarizing it. How can a stable resting potential be maintained then? Suppose that there is a small resting inward conductance to Na⁺ that creates a current equal and opposite to the K⁺ current. How could a few open Na⁺ channels provide a current that balances that generated by many open K⁺ channels?
7. List all the factors you can think of, besides resting or leak K⁺ channels, that could also contribute to the resting potential. What other mechanism of moving ions across membranes could contribute to the resting potential (see Further Exploration; McCormick, 2009)? What other types of channels, both leak and voltage- or ligand-gated, could affect the resting potential?
8. Explain why the Goldman-Hodgkin-Katz equation:
   

   Em =	RT	ln	PK[K+]o + PNa[Na+]o + PCl[Cl−]i
   	F		PK[K+]i + PNa[Na+]i + PCl[Cl−]o

   or the parallel conductance equation:
   

   Em =	gKEK + gNaENa + gClECl
   	gK + gNa + gCl

   predicts the resting potential (as well as action and synaptic potentials) better than the Nernst equation on its own.

References

• Aidley DJ (1998). The Physiology of Excitable Cells (Cambridge University Press, Cambridge), p. 21.
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