Levator Auris Longus Preparation for Examination of Mammalian Neuromuscular Transmission Under Voltage Clamp Conditions

All animal procedures were performed in accordance with the Animal Care and Use Committee of Wright State University.

1. Mouse Euthanasia

1. In a fume hood, place the mouse in an airtight glass anesthetizing chamber.
2. Expose the mouse via inhalation to a lethal dose of isoflurane (saturating, or ~25%). Leave the mouse in the chamber until no breathing can be observed.
3. Remove the mouse from the chamber and perform a cervical dislocation as a secondary method of euthanasia.

2. Removal of Hair from the Dorsal Surface of Head, Neck, and Back

1. Use an electric shaver to remove the majority of the hair on the dorsal surface of the head, neck, and back of the mouse. Also, remove the hair around and on the left ear.
   NOTE: Take care when shaving the skin around the ears, skin can get caught in the blades of the shaver and could damage underlying muscle tissue.
2. Apply hair removal cream with a cotton swab to the shaved area to remove the remaining hair. Wait approximately 1 min and rinse the hair removal cream with water using a squeeze wash bottle, then brush away any remaining cream or hair using a cotton swab and pat the skin dry.

3. Remove Skin to Expose the Levator Auris Longus Muscle

1. Under a stereo dissecting microscope, make a small incision (just deep enough to penetrate the skin) on the back of the mouse at the level of the scapulae. Cut the skin using micro dissection scissors, following the path shown in Figure 1A-B.
2. Using fine forceps (such as No. 5), pull up the skin along the cut near the scapulae. Using spring scissors, cut the connective tissue to separate the skin from the underlying muscle tissue while approaching the ear. Importantly, cut the connective tissue with the blades pointed into the skin to avoid accidental cutting of the exposed muscle tissue.
3. Periodically perfuse the exposed muscle with a physiological saline solution, such as the following Na⁺ external buffer that consists of (in mM): 144 NaCl, 4 KCl, 1.2 CaCl₂, 0.6 MgCl₂, 5 glucose, 1 NaH₂PO₄, pH 7.4 with NaOH, and has an osmolality of 300 ± 5 mmol/kg. Once the connective tissue has been cut up to the left ear, cut the skin around the ear to remove the skin completely from the mouse and discard it.
   NOTE: The LAL attaches to the base of the ear and can easily be damaged while removing the skin. It is good to leave around 1 mm of skin around the base of the ear to avoid cutting the LAL.

4. Removal of Levator Auris Longus Muscle and Surrounding Tissue

1. Using spring scissors, start by cutting muscles that are inferior to the LAL, which connect to the spinal column between the scapulae. Start at the right scapula, just to the right of the midline, and cut toward the rostral end of the mouse, while remaining on the right side of the midline. Continue cutting all the way through to the end of the muscle tissue on the cranium.
   NOTE: The LAL is the most superficial muscle connected to the medial side of the ear and the midline, as shown in Figure 2. For additional images, a book by E.C. Greene¹⁷ shows excellent, hand-drawn images of the LAL.
2. Use forceps to grab the cut tissue immediately to the right of the midline, then gently lift and begin cutting the tissue toward the left ear with the blades of the scissors pressed against the cranium to remove several layers of muscle that are inferior to the left LAL. Leave all of the layers attached temporarily to avoid accidentally cutting the LAL during removal.
3. Cut with blades parallel to and pressed against the cranium to avoid cutting the nerve that innervates the LAL, which wraps around the ear canal, and enters the muscles on the medial side of the ear. Continue cutting through the ear canal, keeping as much of the nerve attached as possible.
   NOTE: The nerve that supplies the LAL branches off of the facial nerve and should be preserved as it will be utilized later in the procedure.
4. Cut the fatty tissue that is just past the ear canal along the same path shown in Figure 1B on the ventrolateral portion of the ear. Also, cut along the left scapula as was done on the right side to remove the muscle completely from the mouse.

5. Levator Auris Longus Muscle Isolation

1. Place the LAL, and surrounding tissue, into a container. Use a container in which the dissected tissue can be pinned to the bottom, such as a Petri dish with a silicone elastomer bottom. Bathe and frequently wash the tissue in a physiological saline solution.
2. Cut off the pinna of the ear along the base of the ear, leaving the cartilaginous part of the ear attached to the LAL. Flip the muscle so that the inferior side is facing up (LAL on the bottom of the dish).
3. Place a pin, such as an insect pin, through the ear canal to hold the prep in place. Then using smaller pins, pin the remaining tissue on the opposite side of the midline of the LAL. Using forceps, gently pull the skin on the lateral portion of the ear to stretch the muscle out and place a small pin through the skin. Repeat this step until the tissue is well-secured to the dish, as shown in Figure 3.
   NOTE: Small dissection pins can be made by cutting the tips of acupuncture needles to the desired length. These small pins minimize damage to the tissue and can be used with water-immersion microscope objectives with long working distances.
4. Begin removing the muscles (shown in Figure 4), that are covering the LAL (auricularis superior, abductor auris longus, and the interscutularis), and those bound to the LAL via connective tissue and are especially tight near the midline, using No. 5 forceps and spring scissors.
5. Using the forceps to pull up on the overlying muscle layer, cut the connective tissue with the blades oriented toward the muscle layer being pulled and take care to avoid cutting or nicking the LAL. Cut toward the midline and stop cutting about three quarters of the way to the midline. Then, cut parallel to the midline to remove each muscle layer. Keep removing muscle layers until only the LAL remains.
6. Remove some of the remaining connective tissue that is covering the LAL, which aids in impalement of the electrodes. Gently use No. 5 forceps to pull the connective tissue away from the LAL and cut it away using spring scissors. Only remove tissue that can be done so easily without risk of damaging the LAL in the process. Remove any large nerves that innervate the inferior muscle layers remaining on the inferior surface of the LAL that may obstruct the visualization of the neuromuscular junctions.

6. Isolation of Nerve

1. Identify the nerve that innervates the LAL using a nerve stimulator (for example, two platinum wires connected to a pulse generator); there will be several nerves running from the ear canal to the muscles. Touch the nerves with the nerve stimulator supplying some current (current will vary around 5 V). When the muscle contracts, the correct nerve has been identified.
2. Carefully grab the tissue near the nerve and use spring scissors to separate the nerve from the tissue surrounding the ear. To minimize damage, keep most of the nerve embedded in some surrounding tissue, which will be used later to secure the nerve to the recording dish.
   NOTE: The motor nerve that innervates the LAL is located on the medial side of the ear canal opening.
3. If using a bipolar stimulating electrode, remove the surrounding tissue only around a region of nerve, distal from the muscle. If using a suction electrode, remove surrounding tissue at the cut end of the nerve.
   NOTE: This is a good stopping point. This is a good stopping point. If a break is needed the dissected LAL prep can be maintained in a physiological solution or constant perfusion for a couple of hours to ensure that harmful metabolites do not accumulate. Use a large volume of solution or constant perfusion to ensure that harmful metabolites do not accumulate.
4. Next, unpin and transfer the muscle to a perfusion chamber for electrophysiology experiments under an upright compound microscope.
   NOTE: Use a perfusion chamber with a soft bottom to which the tissue can be securely pinned (Figure 5A).
5. Pin the muscle at the ends (ear and midline) and along the edge of the muscle. Position the nerve perpendicular to the muscle fibers and pin it to the bottom of the dish through some excess tissue that was left intact at the end of the nerve. Keep the tissue bathed in a physiological saline solution at all times.
   NOTE: It is also helpful to have rounded, elevated material directly under the LAL muscle fibers. This extra support aids in electrode impalement and can be made from a small section of silicone elastomer cast in a 50 mL conical tube lying on its side. The rounded section of the elastomer can be secured to the bottom of the perfusion chamber using a small amount of fresh silicone. A diagram of the perfusion chamber is shown in Figure 5B-C. The muscle fibers can adhere very tightly to newly cast silicone elastomer (which is very hydrophobic) and may become damaged. It is helpful to coat or block newly cast silicone with protein, similar to the blocking step in an immunoblot. To block it, we have incubated newly cast silicone in a concentrated solution of bovine serum albumin overnight.

7. Electrophysiology Equipment Setup

1. Prepare or thaw aliquots of a dye to aid in visualizing the neuromuscular junction, such as the mitochondrial dye 4-(4-diethylaminostyryl)-1-methylpyridinium¹⁸, which is light sensitive and should be stored away from light. Also, thaw electrode solutions. Vortex all aliquots to ensure that all solutes are in solution. Prepare a physiological saline solution containing 1 µM µ-Conotoxin GIIIB (µ-CTX) and 80 µM BTS (optional).
2. Secure the perfusion chamber with the LAL to the microscope stage. Place the reference electrode into a cup filled with 3 M KCl, which is connected to the recording chamber via an agar bridge. Connect the reference electrode to the amplifier per manufacturer instructions.
3. At this point, position the nerve-stimulating electrode on the nerve. Use either a suction electrode or a small bipolar electrode, as they work well for nerve stimulation.
   1. Position the stimulating electrode as far from the muscle as possible to minimize the number of stimulation artifacts in the recordings.
      NOTE: The stimulating electrode should be controlled with a stimulus isolation unit that can control the voltage amplitude as necessary.
4. Slowly raise the voltage by turning the voltage knob on the stimulus isolation unit until contractions are observed. The point at which the contraction was first observed is the threshold. Once the threshold has been identified, set the voltage on the stimulus isolation unit at 1.5*threshold (or as defined by the experiment).
   NOTE: Figure 6 shows the muscle prep set up under an upright microscope and the small bipolar electrode positioned on the nerve using a ball-joint manipulator. It is favorable to have the lowest voltage for stimulation possible while still being sufficiently above threshold. A lower stimulus voltage reduces the number of stimulation artifacts during recordings. Also, placing the stimulating electrode away from the muscle will decrease the size of the stimulation artifact. Sometimes the end of the nerve will be damaged from the dissection, so it may be necessary to experiment with where along the nerve the stimulator should be placed.
5. Remove the bathing saline solution and replace with 5 µM 4-Di-2-Asp. Expose the LAL prep to the 4-Di-2-Asp for 10 min to achieve adequate fluorescence for neuromuscular junction visualization (Figure 7).
6. Prepare two electrode solutions, 3 M KCl and a K⁺ internal solution consisting of (in mM) 75 aspartate, 5 MgCl₂, 15 Ca(OH)₂, 5 ATP disodium, 5 phosphocreatine disodium, 5 glutathione, 20 MOPS, 30 EGTA, pH 7.2 with KOH.
   NOTE: The solutions should be prepared in advance; the K⁺ internal solution can be stored in aliquots at -20 °C.
7. Fill the pulled glass capillary for the voltage-sensing electrode with 3 M KCl. Use the K⁺ internal solution (prepared in step 7.6) for the current-passing electrode; use a narrow nonmetallic needle (~34 gauge) attached to a 1 mL syringe to easily fill the glass capillaries. Gently tap the capillaries to remove air bubbles; ensure that each filled electrode has a resistance of 10-15 MΩ.
   NOTE: Check and note the resistance of both electrodes using the data acquisition software.
8. After 10 min, exchange the 4-Di-Asp solution with the normal Ca²⁺ solution.

8. Identification of Neuromuscular Junction Using Fluorescence

1. Using an upright microscope with standard bright-field and fluorescence illumination, look for a bright band of fluorescent green neuromuscular junctions running perpendicular to the muscle fibers along the prep as shown in Figure 7A. Use a low magnification water-immersion objective (~10X) to identify neuromuscular junctions
   NOTE: The microscope should be equipped with a FITC cube (Ex: 480/40, Dichroic: 505LP, Em: 535/50), and an LED, laser, halogen lamp, or mercury lamp for fluorescence.To minimize photo bleaching, alternate between bright-field and fluorescence illumination.
   NOTE: This band is usually more proximal to the base of the ear than to the midline. The band is the region where the neuromuscular junctions are most concentrated.
2. Switch to a higher magnification water-immersion objective (~40X) and identify a neuromuscular junction on the top layer of muscle to examine with electrophysiology (Figure 7B).
3. Secure the filled pipettes into the pipette holders on the appropriate headstages, per manufacturer's instructions. Use micromanipulators to position the electrodes above the muscle membrane within 100 µm of the identified neuromuscular junction (Figure 7C). Position the electrodes using primarily bright-field microscopy; use fluorescence to confirm the location of the electrodes relative to the neuromuscular junction.
4. First, use a low magnification (~10X) objective to locate the electrodes and then switch to a higher magnification (~40X) objective for final positioning of the electrodes. Do not impale the electrodes into the muscle at this point, first tune the electrodes.

9. Tuning and Impaling the Electrodes

1. Once the electrodes are positioned above the desired fiber, zero and tune the voltage-sensing electrode by bridge balancing (or an analogous approach) and neutralizing the electrode capacitance per the manufacturer's instructions. Also, zero the current-passing electrode.
2. Bring both electrodes down to the surface of the fiber. Slowly adjust the position of the electrodes on the way down to the fiber so that the electrodes remain oriented with respect to the neuromuscular junction, as described in steps 8.3 and 8.4.
3. After both electrodes have contacted the surface of the muscle fiber, impale the blunt current-passing electrode first. Use techniques such as buzz (brief pulses of excess capacitance compensation), brief current pulses, or gently tapping the table to facilitate electrode impalement.
   1. Monitor the signal, using an oscilloscope or oscilloscope protocol in the data acquisition software, until a negative membrane potential is identified, which indicates that the electrode has been impaled.
4. Depress the sharper voltage-sensing electrode onto the muscle to the level of the impaled current-passing electrode. Once both electrodes are in line with one another, impale the voltage-sensing electrode using the same techniques applied with the current-passing electrode.
5. Upon successful impalement, using the controller for the amplifier, apply a constant negative holding current with the current passing electrode to compensate for any membrane damage due to electrode impalement. As the cell starts to slowly charge toward -80 mV, increase the holding current as needed to bring the cell to the desired resting potential (i.e., -80 to -85 mV).
   NOTE: Avoid recording from fibers that require a holding current larger in absolute value than -25 nA in wild type muscle to minimize the chance of recording from unhealthy fibers.
6. Once the fiber is stable for a couple of minutes at the desired resting potential, proceed to recording muscle membrane properties or neuromuscular transmission.

10. Recording Postsynaptic Membrane Properties and Synaptic Transmission

1. Start with current-clamp recordings by applying an equal number of positive and negative current steps to measure the corresponding membrane voltage responses, which will be used to determine motor endplate R_in and τ_m. To ensure passive properties, use small voltage responses that reach a steady state and have a linear voltage-current relationship^16,19,20. Avoid recording from fibers with a damaged or unhealthy nerve, such fibers have a mEPP frequency of >3 Hz.
2. Next, enter two-electrode voltage-clamp per the manufacturer's instructions. Voltage-clamp fibers at a membrane potential of -85 mV. Voltage-clamp settings should acheive signal-to-noise ratios that allow detection of mEPCs.
   NOTE: Maintaining adequate voltage control can be an issue. We use two methods to minimize these issues. First, the fibers are impaled within 100 µm of the motor endplate as described previously, which is within the length constant of these fibers²¹. Second, we use the DC restore feature of the amplifier. This feature sets a very high voltage-clamp gain. Our previous work has described this method in detail and outlined a method for assessing the voltage-clamped endplate¹⁶.
3. Once in voltage-clamp, record synaptic currents (mEPCs and eEPCs) using various nerve-stimulation protocols, based on the requirements of the experiment.
   1. Record spontaneous mEPCs and EPCs using a low-frequency nerve stimulation protocol (≤0.5 Hz) which enables one to record EPCs without synaptic modulation (facilitation or modulation) to assess quantal content. To assess synaptic facilitation or depression, use a high-frequency nerve stimulation protocol (10 pulses at 50 Hz).
   2. Stimulate the nerve at the desired frequency via a stimulus isolation unit, which can be triggered using a pulse generator that is controlled using the data acquisition software.
4. Once the voltage-clamp recordings are done, return the amplifier to a current-clamp setting, turn off all currents and remove the electrodes from the fiber. Check that the electrodes did not become clogged or drift in baseline voltage during the recording.
   NOTE: Regarding drift, the electrode voltage should be 0 volts in the extracellular solution, as they were set before fiber impalement. For example, the recorded voltages would be uncertain if the voltage drifted during the experiment.