Microelectrode Impalement Method to Record Membrane Potential from a Cannulated Middle Cerebral Artery

This protocol can be used to measure the membrane potential of normal and diseased blood vessels and to evaluate pharmacological agents that modulate membrane potential, vascular tone, and blood flow. The membrane potentials generated from this technique are close to the physiological range since vascular smooth muscles within a vessel are in a syncytium. Before beginning the procedure, place a dual channel differential electrometer amplifier close to the vessel chamber at the desired location.

Use a BNC-BNC cable to connect the output of the amplifier channel A or B to the channel input of the digitizer. Mount the probe in the micromanipulator, and turn the micromanipulator towards the microscope and the myograph on a vibration-free table. Place the knobs and switches on the front of the amplifier in the appropriate positions for the experiment, as described in the manual.

Then connect the bath ground to the circuit ground of the amplifier with an appropriate electrode, and ensure that the cage is grounded to the chassis of the amplifier. To prepare the borosilicate glass microelectrodes, use a standard puller to achieve a short, gradual, eight-to 10-millimeter taper with a less-than-one-micrometer diameter, looping twice for higher resistances and smaller tips. Use a microfiber syringe to fill the microelectrode with three-molar potassium chloride, slowly retracting the plunger during the injection to allow space for the fluid to fill and to prevent the formation of bubbles inside the microelectrode.

Place the fully loaded microelectrode onto the microelectrode holder, and carefully but firmly push the electrode shank into the holder through the bored hole. Remove any excess fluid with a lab tissue, and connect the electrode holder assembly to the amplifier probe. Conduct an electrode test to measure the electrode resistance.

Then open the recording software, assign a name to the file, and save the file for future analysis. Next, rinse the myograph chamber with distilled water several times before loading the chamber with five milliliters of normal physiological salt solution, or PSS. Using a five-or 10-milliliter syringe, fill both glass cannulas and the attached tubing with filtered normal PSS without introducing any air bubbles, and use blunt forceps to make a half-knot in two 10-0 monofilament nylon sutures.

Then, using dissection forceps under a dissection microscope, place the partially closed suture knots on both cannulas slightly away from the tips. For middle cerebral artery isolation, use spring scissors and forceps to identify and dissect out an unbranched segment of the middle cerebral artery, with an inner diameter of 100 to 200 micrometers, of the rat brain. Use fine forceps to mount the middle cerebral artery onto the glass cannulas, and tighten the sutures to secure the artery to the cannula.

Close off the distal cannula so that there will be no flow within the artery, and connect the inflow pipette to a reservoir of PSS. Visualize the cannulated middle cerebral artery using a charge-coupled device camera mounted on an inverted microscope and imaging software. Set the axial length of the MCA to an approximate length such that it appears neither rigid nor flaccid.

Equilibrate the bath solution with 95%oxygen and 5%carbon dioxide at 37 degrees Celsius. Immerse the ground of the amplifier in the PSS of the myograph. Illuminate the vessel chamber, and visualize the tip of the microelectrode in the bath solution through the microscope.

Using the micromanipulator, move the tip of the microelectrode close to the outer wall of the blood vessel, and begin the recording. Slowly move the tip of the microelectrode toward the vessel, aiming for the center of the vessel. When the tip comes in close proximity to the vessel, advance the electrode forward in one rapid motion to impale the membrane of the muscle.

Once the membrane has been penetrated, changes in the membrane potential may be observed. Do not touch the micromanipulator once the membrane has been impaled. When the membrane potential changes have been recorded, use the manipulator to remove the microelectrode in one rapid movement, and stop the recording before saving the data files.

Impalement is considered successful when there is a rapid deflection to negative values, the membrane potential is stable for a minimum of 30 seconds, and the voltage returns abruptly to zero millivolts upon removal of the electrode. After a successful impalement and membrane potential stabilization, drugs of interest can be perfused into the bath and changes in the membrane potential can be recorded. In this representative experiment, perfusion with potassium chloride depolarized the membrane by approximately six millivolts, while perfusion with a calcium-dependent activator of large conductance calcium-activated potassium channels hyperpolarized the membrane by nearly four millivolts.

The most important thing to remember is to pull highly resistant microelectrodes that have a short, gradual taper with a less-than-one-micrometer diameter.