Isolating and Culturing Vestibular and Spiral Ganglion Somata from Neonatal Rodents for Patch-Clamp Recordings

Summary

Presented here are methods providing detailed instructions for dissecting, dissociating, culturing, and patch-clamp recording from vestibular ganglion and spiral ganglion neurons of the inner ear.

Abstract

The compact morphology of isolated and cultured inner ear ganglion neurons allows for detailed characterizations of the ion channels and neurotransmitter receptors that contribute to cell diversity across this population. This protocol outlines the steps necessary for successful dissecting, dissociating, and short-term culturing of the somata of inner ear bipolar neurons for the purpose of patch-clamp recordings. Detailed instructions for preparing vestibular ganglion neurons are provided with the necessary modifications needed for plating spiral ganglion neurons. The protocol includes instructions for performing whole-cell patch-clamp recordings in the perforated-patch configuration. Example results characterizing the voltage-clamp recordings of hyperpolarization-activated cyclic nucleotide-gated (HCN)-mediated currents highlight the stability of perforated-patch recording configuration in comparison to the more standard ruptured-patch configuration. The combination of these methods, isolated somata plus perforated-patch-clamp recordings, can be used to study cellular processes that require long, stable recordings and the preservation of intracellular milieu, such as signaling through G-protein coupled receptors.

Introduction

The bipolar neurons of the vestibulocochlear nerve connect the sensory hair cells of the inner ear to the brainstem. They are principal carriers of information about sound and head movements; damage to these important cells leads to deafness and balance disorders. The vestibular and auditory portions of the nerve are each comprised of distinct cell types that are morphologically and functionally diverse^1,2. In the vestibular system, two afferent subpopulations fire spontaneously at intervals that are either regular or irregular². Afferent spike timing is thought to reflect an underlying diversity in ion channel composition^3,4. In the auditory system, there are two main subpopulations of spiral ganglion neurons (SGNs); whereas Type I SGNs contact individual inner hair cells⁵, Type II SGNs contact multiple outer hair cells⁵. In vitro recordings from semi-intact and organotypic cultures suggest differences in the membrane properties of Type I and Type II SGNs^6,7.

Many ion channels and neurotransmitter receptors found at the terminals of these neurons are also found in their cell bodies. As such, cultures of the isolated vestibular and the spiral ganglion somata can be studied in vitro to understand how ion channels and neurotransmitter receptors contribute to the response of these neurons. The compact morphology of the isolated cell bodies allows for high-quality electrical recordings, suitable for detailed characterization of voltage-gated ion channels and neurotransmitter receptors. Easy access to a representative variety of neuron subtypes allows for high-throughput analysis of cell diversity.

This article presents a method for isolating and culturing dissociated ganglion cell bodies from the superior portion of the vestibular ganglion in rats at postnatal day (P)9 to P20. Suggestions are also provided for extending these methods to the spiral ganglion, in addition to the steps required for successfully extracting, dissociating, and plating the ganglion cells. These methods are an evolution of those devised in publications from various laboratories^8,9,10. Also included in this paper is guidance for selecting healthy cells for patch-clamp recordings.

Finally, the protocol outlines the procedure for patch-clamp recording using the perforated-patch configuration¹¹. Though the perforated-patch configuration is more time-consuming and more technically challenging than the more common ruptured-patch configuration, it is better for maintaining the cytoplasmic milieu that allows for long and stable recording sessions. The benefits of this recording configuration are illustrated here through the improved stability of hyperpolarization-activated cationic currents in perforated-patch relative to ruptured-patch recordings.

This protocol is organized into five sections. Sections 1-3 describe solutions and tools that can be prepared and stored ahead of time. Section 4 describes the steps for dissecting and plating the vestibular and SGNs. Section 5 describes the steps for recording from the neurons after a period in culture. In our hands, section 4 and section 5 are performed over a period of 2 consecutive days.

Protocol

All animal use described here has been approved by the Institutional Animal Care and Use Committee at the University of Southern California. Animals in this protocol are P3- to P25-aged Long Evans rats of both sexes obtained from Charles River Laboratories, but these methods can be applied to other rodent strains. A laboratory coat and gloves must be worn during all procedures, as well as splash-protective goggles when making solutions.

1. Preparations

NOTE: The solutions and tools described in this section can be made well ahead of time to be used on the day of dissection and recording.

1. Prepare Liebovitz media supplemented with 10 mM HEPES (L-15 solution). The following steps describe making 1 L of L-15 solution.
   1. Pour 990 mL of deionized H₂O into one beaker. Measure and add 2.386 g (10 mM) of HEPES. Add one bottle of 1 L of L-15 solution powder.
      NOTE: Powdered L-15 has a longer shelf life and takes up less storage space. Alternatively, purchase L-15 solution and supplement with HEPES. The latter option also has the option of using a phenol-free solution, which is useful for fluorescence imaging.
   2. Stir the solution on a stir plate. Using a pH meter, slowly add 1 N sodium hydroxide (NaOH) to obtain a pH between 7.34 and 7.36.
   3. Add deionized H₂O until the solution reaches a volume of 1,000 mL. Filter the solution using a 0.22 µm pore size membrane.
      NOTE: L-15 solution can be stored at 4 °C for around 1-2 weeks. If using L-15 made with phenol-red, the solution will turn pink if too basic (pH > 7.4) or orange if too acidic (pH < 7.34).
2. Prepare the culture medium for vestibular ganglion neurons (VGNs) (with modification for SGNs). Follow the steps below to make 50 mL of culture medium:
   1. Add 47.5 mL of GluMAX-I minimum essential medium to a clean glass beaker.
   2. Measure and add 0.1194 g (10 mM) of HEPES. This additional buffer resists pH changes while the cells settle, before being moved to the incubator.
   3. Add 2.5 mL of fetal bovine serum (FBS). This makes 5% by volume FBS in a total volume of 50 mL of medium. For SGN preparations, 2.5 mL of FBS is substituted with 0.5 mL of N2 and 1 mL of B27 solutions.
   4. Stir the solution on a stir plate. Using a pH meter, slowly add 1 N NaOH to obtain a pH between 7.38 and 7.4.
   5. Add 0.5 mL of penicillin-streptomycin. Filter the solution through a 0.22 µm sterile filter. Store the solution at 4 °C.
   6. At least 30 min before use, transfer the culture media to a tissue culture flask with a vented cap. Place the flask with culture media in an incubator with 5% CO₂ concentration at 37 °C.
3. Prepare the perforated-patch internal solution
   NOTE: Here, 100 mL of perforated-patch internal solution is used for whole-cell recording (recipe is summarized in Table 1). These solutions can be prepared well in advance and stored at -20 °C. Aliquots can be stored indefinitely at -20 °C if they don't undergo a repeated cycle of freezing and thawing.
   1. Up to 6 months ahead of time, make and store stock solutions of the following: 1 M KCl, 0.5 M Mg₂Cl (hexahydrate), and 0.5 M CaCl₂. Store in airtight bottles for up to 6 months at 4 °C.
      NOTE: It is a good idea to check the osmolality of the stock solutions (2,000 mOsm for the 1 M KCl stock solution; 1,500 mOsm for the Mg₂Cl and CaCl₂ solutions) to make sure the target concentration is reached. Do not use if the solution becomes cloudy or forms precipitates. This can be a possible pause point.
4. On the day of internal solution making, follow the below steps using the stock solutions from step 1.3.
   1. Measure 100 mL of milli-Q (MQ)H₂O. Draw out 40 mL from the 100 mL and set aside in a clean beaker.
   2. Add the remaining 60 mL of H₂O to a clean and sterile 100 mL beaker. Add 2.5 mL of 1 M KCl, 1 mL of 0.5 M Mg₂Cl-hexahydrate, and 0.02 mL of 0.5 M CaCl₂.
   3. Add a stir bar and place the beaker on a stir plate. Stir till completely dissolved. Add 1.307 g of K₂SO₄. Stir until the K₂SO₄ dissolves.
   4. Weigh 0.1192 g of HEPES (target 5 mM in a 100 mL final volume) and add to the solution. Stir until dissolved.
      NOTE: Ensure that all the components are fully dissolved, as sometimes it takes time for the K₂SO₄ to dissolve.
   5. Weigh 0.1902 g of ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid (EGTA) (target 5 mM in a 100 mL final volume) and add to the solution. As EGTA does not completely dissolve in an acidic solution, place the beaker on a stir plate and insert the pH probe.
   6. While stirring, slowly titrate 1 M KOH until the EGTA fully dissolves. Expect this to take place when the pH is between 6 and 7. Continue to slowly add KOH until the final pH is 7.35 (approximately 1.2 to 1.3 mL of KOH in total).
   7. Add MQH₂O to bring the solution to a final volume of 100 mL and stir. Check the osmolality of the solution (target 270 mOsm/kg for the perforated-patch solution) with an osmometer.
   8. Filter the perforated-patch internal solution using a 0.22 µm sterile filter. Store in 5 mL aliquots at -20 °C. Use a new aliquot for each new recording session.
      ​NOTE: This can be a possible pause point.

2. Fabricating trituration pipettes

NOTE: Trituration pipettes can be reused for multiple experiments. Flush the pipettes with ethanol, water, and L-15 solution before each use and clean them with water and ethanol after each use. Carefully store between uses.

1. To prepare trituration pipettes for cell dissociation, hold a glass Pasteur pipette to the flame of a Bunsen burner. Once the glass tip begins to melt, stretch the glass out to the desired tip diameter. Pull one end of the pipette away from the flame to create a small bend.
2. Bring the bent pipette near to a microscope. Using a scoring tile, score and break the glass at the desired diameter.
3. Pass the tip of the pipette over the top of the Bunsen burner flame. This will quickly polish the rough edges near the tip. Check to ensure that the tip is not sealed by the flame.
4. Repeat the process until four or five pipettes have been prepared with varying sizes.

3. Fabricating patch pipettes

NOTE: Prepare a set of patch pipettes before the recording session, but after ganglion dissection and culture. Although electrodes that are made one at a time during the recording session are optimal in performance, reasonable success has been achieved when these are made as a batch the night before. Store the pipettes in a covered glass container to protect the tips from dust.

1. Use a vertical electrode puller and 1.5 mm outer diameter/1.17 mm inner diameter filamented borosilicate glass pipettes. Ensure that the glass is always kept free from dust and fingerprints.
2. Pull 8-10 recording pipettes using a two-step program, as recommended for patch-pipettes by the manufacturer.
3. Inspect and heat-polish every recording pipette tip with a microforge; heat-polishing promotes better sealing. The target pipette resistance is between 4 and 8 MΩ.
4. Store the polished pipettes in a covered container. This can be treated as a pause point in the experiment.
   NOTE: Optimizing the pipette diameter to reach the target resistance range requires some trial and error with steps 3.2 and 3.3. Most commercial patch-clamp systems have a built-in membrane test feature for measuring the electrode resistance in the bath solution. The resistance is computed as a ratio of the steady-state output current in response to an applied 5 mV step stimulus. The pull settings in step 3.1 and the polishing in step 3.2 must first be adjusted using trial pipettes, until the resistance of the trial pipettes falls within the 4 to 8 MΩ range.
5. On the day of recording, coat the pipette tips to reduce the pipette capacitance and recording noise. To do so, wrap a small strip of paraffin film on the shaft of the pipette. It is not necessary to wrap all the way to the tip.
   ​NOTE: An alternate strategy is to coat and heat set a silicone elastomer on the shaft of the pipette¹².

4. Extraction of vestibular ganglion and plating of vestibular neurons

1. Preparation for dissection
   1. Prepare an enzyme mixture of L-15 solution with 0.05% collagenase and 0.25% trypsin. For example, add 0.001 g of collagenase and 0.005 g of trypsin to 2 mL of L-15 solution into a beaker, add a small magnetic stirrer, and stir until fully mixed. Set aside at room temperature.
   2. Prepare a commercially obtained guillotine by placing paper towels to the side and underneath the guillotine to minimize exposure of the benchtop and other surfaces to blood and other biological materials.
   3. Lay out large stainless-steel scissors, large forceps, and large spring scissors. Keep a large spray bottle with 70% ethanol handy for head cleaning.
   4. Fill a 100 mL beaker with L-15 solution and place on ice. Oxygenate the L-15 solution.
   5. Lay out the spring scissors, surgical scissors, forceps, scalpel, and transfer pipette (see Table of Materials).
   6. Prepare a 60 mm Petri dish (for the gross dissection) and two 35 mm Petri dishes (for the cleaning/fine dissection of the ganglia). Fill the 60 mm dish with oxygenated L-15 solution using a 30 mL syringe with a 0.22 µm filter tip.
2. Euthanasia, head-cleaning, and hemisection
   1. Weigh the pups to compute the appropriate dosage of fatal-plus dilution to administer intraperitoneally (~0.01 mL per 10 g).
   2. Once a deep plane of anesthesia is reached, as assessed by a lack of response to the toe pinch, decapitate using the guillotine.
   3. Rinse the head thoroughly with 70% ethanol and then thoroughly with L-15 solution.
   4. Completely remove the skin from the skull. Bisect the head using large spring scissors by starting at the entry point of the spinal cord to the brain and making two cuts, the first cut through the top of the skull and the second through the bottom of the jaw. Finish bisecting the head using surgical scissors.
   5. Place both halves of the head brain side down in a 60 mm Petri dish filled with L-15 solution. Place the fresh tissue not currently being dissected on ice.
3. Extracting the superior vestibular ganglion
   1. Scoop out the brain using closed surgical scissors. Sever and remove the cranial nerve to completely detach the brain from the skull.
      NOTE: At this point, the otic capsule (shaped like the number 8) should be visible at the back of the head.
   2. Using forceps and starting at the back of the head, pull clear membranous material off.
   3. Cut the top of the skull out using surgical scissors. Remove excess tissue from the back of the head and neck to make the dissection area and otic capsule cleaner and easier to access.
      NOTE: Avoid making the dish too cloudy by discarding excess tissue throughout the procedure.
   4. Transfer the tissue to a second Petri dish with fresh L-15 solution. Locate the otic capsule and the auditory, superior vestibular, and inferior vestibular nerves. Cut away the auditory nerve and separate the superior and inferior ganglia using small spring scissors.
      NOTE: Although the somata of the vestibular nerve are found in both the inferior (thinner) and superior (thicker) ganglia, cells extracted from the superior ganglion have better survival.
   5. Using a scalpel, gently shave off the bony ridge to weaken the bony area, under which the nerve dives into the bony capsule. Carefully remove debris with fine forceps, exposing the entire swollen portion of the ganglion.
   6. Use the fine spring scissors to cut and separate the ganglion from the peripheral nerve branch that is diving toward the utricle.
   7. Remove the superior ganglion using fine forceps, making sure not to pinch too close to the ganglionic tissue. Transfer to a 35 mm Petri dish with fresh L-15 solution.
   8. Before proceeding, pre-heat the enzyme solution: pour the enzyme mixture into a 35 mm Petri dish and place in a 37 °C incubator for 10-15 min.
   9. Clean the ganglion using fine forceps and small spring scissors by removing bone (which appears white and crystallized in structure), excess tissue, nerve fibers, and any other superfluous structures. Take care to minimize the removal of any ganglionic tissue.
4. Tissue dissociation and plating
   1. Transfer cleaned ganglia into the pre-heated enzyme solution and place it back in the incubator for 10 to 40 min. Enzyme treatment is complete once the tissue starts to break apart but remains in one piece. Overtreating the tissue with enzymes will result in complete dissolution of the ganglia before trituration.
      NOTE: The amount of time that the tissue undergoes enzymatic treatment for depends on the age of the animal. For example, ganglia from P9 rats are treated with enzyme for 25 min, and ganglia from P15 rats are treated with enzyme for 35 min.
   2. Transfer the ganglia to the 35 mm Petri dish with fresh L-15 solution and incubate for 2-3 min.
   3. Transfer the ganglia to another 35 mm Petri dish filled with filtered culture media.
   4. Using a 200 µL micropipette, pipette a ~150 µL drop of filtered culture media onto a coated glass-bottom dish. Do not add too much solution, as it is important to maintain surface tension to form a bubble of solution that remains over the glass coverslip.
   5. Transfer the desired number of ganglia (one to four) to the coverslip.
   6. Draw a small amount of culture medium from the culture dish to rinse the trituration pipette with medium to prevent the tissue from sticking to the sides of the glass pipette. Triturate by gently and repeatedly passing the tissue through the pipette until the ganglia are sufficiently dissociated.
      NOTE: Do not overwork the tissue to attempt single-cell suspensions or allow the cells to crash to the bottom of the dish using excessive positive pressure. Also, avoid forming air bubbles, as this will reduce cell survival. Gentle trituration is the key to achieving successful cell survival.
   7. Let the cells rest for 5 min. Check under a light microscope to see if the cells have settled on the coverslip.
   8. Carefully place a culture dish into a 37 °C incubator for 12-24 h. Again, make sure to keep the bubble of solution intact.
      ​NOTE: When culturing for longer than 24 h, refresh the culture media daily. This can be a possible pause point.
5. Plating SGNs
   1. Follow steps 4.2.1 to 4.2.5 of vestibular ganglion instructions to bisect the head.
   2. Locate the otic capsule (shaped like the number 8), and extract from the skull by chipping away at the edges using blunt forceps.
   3. Change the L-15 solution. The spiral portion of the otic capsule houses the cochlea with the organ of Corti. Chip away at the bone overlying the cochlear turns without damaging the underlying tissue, with fine forceps parallel to the curve of the bone.
   4. Once enough bone has been removed to reveal the full extent of the cochlear turns, use small spring scissors to sever the modiolus (thick, white, fibrous tissue containing central axons of SGNs) at the base of the cochlea to free it from the rest of the otic capsule.
   5. Remove the stria vascularis by pinching the stria at the base with fine forceps. Unwind around the spiral and up to the apex to peel it free from the organ of Corti.
   6. Cut the organ of Corti into two or three turns using small spring scissors such that it lays flat.
   7. Excise the modiolus from each turn with small spring scissors.
   8. Remove the spiral ganglion by cutting at the edge of where SGN fiber tracts project toward the hair cells.
   9. Clean the ganglion using fine forceps by removing bone (which appears white and crystallized in structure), the modiolus (which appears white and fibrous), and any other superfluous structures. Take care to minimize the removal of any ganglionic tissue.
   10. Follow the same procedure as for enzymatically treating the spiral ganglion as used for the vestibular ganglion in section 4.4. Enzymatic times are typically shorter in the spiral ganglion, which is more elongated than the vestibular ganglion. Use smaller diameter trituration pipettes for the spiral ganglion as compared to the vestibular ganglion.
   11. Triturate the spiral ganglion in culture medium supplemented with N2 and B27, indicated in the recipe for culture media.
   12. Place the culture dish containing the dissociated ganglia in a 37 °C, 5% CO₂ incubator for 12-24 h.
       ​NOTE: The protocol can be paused here.

5. Recording

NOTE: In this procedure, patch-clamp recordings from the isolated ganglion are typically performed 12-24 h after the plating. Other labs have reported results from neurons after much longer periods in culture¹⁰.

1. Prepare the recording chamber
   1. Use the quick exchange recording chamber (or equivalent) that allows for patch-clamp recordings directly in the culture dish. Set up the chamber on the inverted microscope.
   2. Insert the glass-bottom culture dishes into the chamber.
   3. Feed the L-15 solution through a perfusion tubing system to the recording chamber. Stabilize the perfusion in-flow and out-flow in the chamber at a rate of 0.5-1 mL per minute.
2. Prepare the internal solution for loading electrodes
   1. Thaw an aliquot of the perforated-patch internal solution. Allocate 2.5 mL of the solution onto a 35 mm culture dish. Replace the lid on the culture dish and label as "Tip Dip". Set aside in a dust-free zone, as it is very important to keep this solution as clean as possible.
   2. Weigh 1 mg of amphotericin-B and add 20 µL of dimethyl sulfoxide (DMSO). Vortex and spin down the solution until all the amphotericin-B is in the solution. Add 10 µL of the DMSO/amphotericin-B solution to the remaining 2 mL of the defrosted perforated-patch internal solution. Withdraw and dispel the solution with a pipette two or three times to ensure that the DMSO/amphotericin-B has uniformly mixed into the internal solution. The solution will have a mild-yellowish tinge.
   3. Draw the amphotericin-B perforated-patch internal solution into a 3 mL syringe. Add a 34 G tip to the syringe. Wrap the syringe in aluminum foil and keep the syringe on ice.
      NOTE: This solution must be remade every 2 h to ensure the effectiveness of the amphotericin-B in perforating the cell membrane. Amphotericin-B is light sensitive, and it must be shielded from light using aluminum foil or other methods.
3. Patch-clamp recordings
   1. Visualize the neurons using a 10x or 20x objective. Adjust the illumination and optimize the optics to see the boundaries and shadows around the cell.
   2. Check the quality of the isolated neurons. Only attempt neurons with smooth and even surfaces that are not too strongly contrasted and have only small, dispersed craters. Also, make sure to avoid pairs of neurons, neurons surrounded by debris, or other cells.
   3. Under 100x magnification, identify a neuron or field of neurons to patch and line them up in the center of the field of view.
   4. Fill the tip of the pipette with a clean solution that does not contain amphotericin by dipping the tip (for ~20 s) into the clean solution that was placed in a culture dish. Capillary action will draw a small amount of solution into the tip.
      NOTE: Perform this step under a stereo dissection microscope, which allows for determining how well the tip holds the Tip Dip solution. If the solution does not hold for ~10-20 s, the shape of the electrode is not ideal. In this case, reconfigure the electrode-pulling program to produce a longer tip.
   5. Next, fill the back of the pipette with the internal solution containing amphotericin. The filament within the pipette will draw down the solution to meet the clean solution in the tip. Allow the filament to smoothly pull the solution down, and do not attempt to tap out bubbles, as this will force the amphotericin to the tip too fast.
   6. Use a syringe to draw out solution that may be at the very back of the electrode.
      NOTE: It is very important to work fast from this point to land and form a high-resistance seal on the desired cell.
   7. Insert the pipette into the pipette holder. Check that the pipette is snug in the holder to prevent pipette drift. If the pipette is not stable, switch out the sealing O-rings in the front and back of the pipette holder to minimize drift and maintain strong suction. Replace the pipette with one that is freshly filled.
   8. Lower the pipette into the bath of the recording chamber.
   9. Locate the pipette tip in the middle of the field of view. Ensure that the tip is free of air bubbles or other debris.
   10. Apply a voltage step (5 mV) in the membrane test to monitor the pipette resistance. Continue monitoring the input resistance through the entire process of forming a seal on the cell.
   11. Cancel the pipette offset potential, such that the pipette current reads zero in the membrane test mode of pClamp.
   12. Correct for the pipette capacitance using the fast capacitance compensation function on the patch-clamp amplifier (Cp fast dials in the Multiclamp Commander soft panel).
   13. Move the pipette down to the cell.
   14. Once the pipette is close enough to the neuron, switch to the higher magnification objective and send the image through a camera to a monitor (use a 40x objective, total 400x magnification). Adjust the pipette and re-center the neuron.
   15. Move the recording electrode close to the soma. Locate the center of the neuron on the monitor and position the recording electrode above the neuron.
   16. Approach the neuron from above, such that the electrode will land in the center of the spherical cell. Adjust the pipette offset to zero the constant DC potentials in the system.
   17. Position the pipette close to the membrane. Land firmly on the center of the cell.
       NOTE: When landing, a small dimpling on the surface of the neuron and a doubling/tripling of the input resistance will occur.
   18. Apply negative pressure (suction) using a syringe or mouth pipette. Turn on the holding potential of -60 mV. The seal resistance should increase until it passes a giga-ohm.
       NOTE: Work fast to reduce the time between filling an electrode and landing on a cell. There is a limited time before the amphotericin added to the perforated-patch internal solution in step 5.3.5 reaches the electrode tip. Once the tip is contaminated with amphotericin, it is difficult to form high-resistance seals, and the resistance often plateaus to ~200 megaohms.
   19. Once a giga-ohm seal forms, release the negative pressure. As soon as the seal forms, apply fast-capacitance compensation to reduce the amplitude of the pipette capacitance transients in membrane test mode as much as possible.
   20. As amphotericin begins to work, watch as the input resistance slowly decreases and the current flowing in response to the 5 mV voltage step progressively increases as the amphotericin enters the membrane. A sudden decrease in series resistance instead of gradual stabilization suggests that spontaneous rupture has occurred.
   21. Estimate the whole-cell capacitance and the series resistance using the amplifier's capacitive transient nulling function. Document and monitor the series resistance at the beginning of, and regularly during, the experiment.
       NOTE: The series resistance can take anywhere from 5-20 min to stabilize, depending on the size of the electrode and how much clean solution is drawn into the pipette. Voltage-clamp and current-clamp modes can be used once the series resistance has stabilized below 30 megaohms. Swelling or shrinking of the neurons during recording is sometimes observed, but rarely when the osmolarity and pH of the solutions are correctly adjusted.

Representative Results

Running voltage-clamp protocols by applying families of voltage steps reveals the voltage-dependent activation of a variety of different families of currents. Representative examples of whole-cell currents evoked from a VGN and adapted from published recordings¹³ are shown in Figure 1A,B. Applying depolarizing voltages (Figure 1B) activates an inward current (negative by convention) that activates and inactivates very rapidly (Figure 1A). This is stereotypical for the voltage-gated properties of sodium channels^14,15, which principally drive the upstroke of action potentials^16,17. Depolarization also activates a long-lasting and relatively slow-activating outward current that drives the downstroke of an action potential. Pharmacology has revealed that these currents are largely carried by a variety of potassium channels in VGNs^{3,18,19,20,21}.

Deep, long-lasting hyperpolarizing voltages evoke a slow-activating inward current carried by hyperpolarization-activated cyclic nucleotide-gated (HCN) channels²² (Figure 1C). The activation of these currents can be studied using the tail-current protocol shown in Figure 1D. Here, the activation percentage of the current is probed by plotting the current that flows during the tail step (Itail) as a function of the conditioning voltage. The current-voltage activation curve has a sigmoidal shape (Figure 1E). Although this channel is gated by soluble second messengers such as cyclic AMP (cAMP), the activation curves measured using the perforated-patch configuration are stable throughout a long recording. In contrast, the size and voltage-activation range of the channel are altered (presumably due to the washout of cytosolic components) when the recording is made in the ruptured-patch configuration.

Neuronal diversity is illustrated by the range of firing patterns elicited when currents are injected into different VGNs and SGNs (Figure 2; top and bottom, respectively). Some neurons fire only at the onset of current injection, while others fire multiple times. This heterogeneity reflects a fundamental diversity in the composition of underlying sodium and potassium channels in both VGNs and SGNs^23,24,25.

Figure 1: Examples of voltage-clamp protocols for measuring diverse groups of ionic currents. (A,B) Example whole-cell currents (A) induced by a family of voltage steps (B). The net inward sodium currents (negative currents) are identified here by their transient activation and inactivation (labeled arrow, Na+). The net outward currents (labeled arrow, K+, long-lasting positive currents) are carried largely by potassium ions and have much slower activation and inactivation kinetics than sodium currents. (C,D) HCN currents are activated by a family of long-duration hyperpolarizing voltages. (E,F) Stability of ion channel characterization in the perforated-patch at different time points during recording. The voltage-dependent activation curves of HCN currents were measured in a perforated-patch configuration I. Activation curves of HCN currents during a ruptured-patch configuration (F). Images C-F have been modified from¹³. Please click here to view a larger version of this figure.

Figure 2: Diverse firing patterns are evoked by injecting steps of currents. (A) Firing patterns in five vestibular ganglion somata. (B) Firing patterns in five SGNs. (C) Corresponding current steps. Please click here to view a larger version of this figure.

Discussion

The methods presented here are specific to recordings from isolated neurons; previous studies have focused on recordings from axon terminals in a semi-intact preparation. When compared to existing terminal recording techniques, isolated recordings offer superior space-clamp and iso-potential behavior. In addition, this protocol provides access to a broader sample of neurons, since only calyx-bearing subpopulations are accessible in semi-intact recordings of the vestibular epithelia. Finally, isolated recordings allow for the use of the perforated-patch technique, which prevents disruption of the intracellular milieu that is often interrupted by the dialysis between the intracellular solution and cytosol in ruptured-patch recordings.

Successful recordings rely first on the quality of the isolated and cultured somata. A critical step in cell survival is the force required during trituration to produce the isolated neurons. A gentle hand is vital to cell survival. Trituration pipettes should be placed high in the bubble of L-15 solution to prevent forcefully expelling neurons onto the bottom of the dish. If issues of cell survival arise, extra care should also be taken to prevent the formation of air bubbles or in breaking the bubble of solution, which prevents the dissociated cells from settling onto the coverslip. It is also best to have heat-polished trituration pipettes available in a variety of sizes, so that investigators have the flexibility to choose one with a diameter such that the ganglion experiences mild resistance as it passes through the pipette. Heat-polishing the pipettes reduces damage caused by passing the ganglion over rough edges of glass. Leaving a rough edge may initially appear to be more effective in breaking up the ganglia, but this approach damages the somata and reduces survival after the period in culture. A final piece of advice is to jealously safeguard a successful trituration pipette.

Another factor that is critical to cell survival is the duration of treatment with digesting enzymes. When determining enzyme time, investigators need to carefully adjust the time to ensure that the tissue breaks up well, but not so much as to impact cell survival. We find that the impact of enzymatic treatment on cell survival and ion channel properties is minimal, as data gathered from enzyme-treated cells appear to be consistent with other methods that do not rely on this approach^19,26. Although enzymatic treatment minimizes the force required to triturate, it does come with the limitation that enzymatic digestion (particularly based on trypsin alone or papain) is known to cause damage to ion channels²⁷. Thus, although we recommend some guidelines for enzyme timing dependent on the age of the animal, it is best to be prepared to adjust the time based on the outcomes. Finally, we encourage a 'less is more' approach to trituration. This means resisting the urge to form a single-cell suspension and stopping trituration after three or four passes, even if there are several large chunks of tissue remaining.

An advantage of culturing is that it appears to clean the somatic cell membrane and promotes shedding of the myelin-forming glial cells, which then allows for more successful patch-clamping from the neurons. Thus, the isolated and cultured ganglion preparation is especially useful for extending patch-clamp recordings past the 1st postnatal week, after which the cell bodies become progressively covered with myelin, impeding the patch-clamp electrode. Ion channel activity observed in patch-clamp recordings from isolated plus short-term cultured (<24 h) VGNs beyond the 2nd postnatal week are consistent with zonal and maturational changes in ion channel expression seen by immunohistochemistry and direct recordings from calyx terminals in vestibular epithelia^28,29. However, it should be noted that prolonged culturing aided by neurotrophic factors and antibiotics may also affect ion channels^{30,31,32,33,34}. Any application of these techniques must carefully consider the impact of these factors on the underlying biophysics of the neurons and their ion channels^{30,31,32,33,34}.

Overall, patch-clamp recordings from isolated and cultured somata are suitable for studying diversity in the ion channel properties of inner ear neurons. The long-term stability of the perforated-patch configuration is especially suitable for studying ion channels such as HCN, whose activation properties are subject to modulation via cytosolic second messengers.

Materials

Name	Company	Catalog Number	Comments
Amphotericin	Sigma-Aldrich	A4888-100MG	For perforated patch recordings.
ATP di-sodium	Sigma-Aldrich	A7699	Additive to internal solution
B27 Supplement (50x), serum free	Thermo Fisher Scientific	17504044	additive to culture medium, for SGN
Beakers (1000, 100, 10) milliliter
bench-top centrifuge	USA Scientific	2641-0016
Bunsen burner
CaCl2	J.T. Baker	1311-01	Additive to internal solution
Collagenase	Sigma-Aldrich	C5318	one out of three enzyme to digest tissue
Coverglass, rectangular, #1 thickness, 22x40	Warner Instruments	64-0707
DMSO	Biotium	90082
Dnase I,from bovine pancreas	Sigma-Aldrich	11284932001	one out of three enzyme to digest tissue
Dumont #3 Forceps (Blunt)	Fine Science Tools	11231-30
Dumont #5 Forceps (Fine)	Fine Science Tools	11251-10
Dumont #55 Forceps (Fine)	Fine Science Tools	11255-20
EGTA	Sigma-Aldrich	E0396	Additive to internal solution
Electrode Puller	Narashige	PC-10
Epi-illumination light source	Zeiss	CL 1500 ECO
Ethanol	Decon Labs	2716	for cleaning head and around dissection bench
Filamented Borosilicate Capillaries for electrodes	Sutter Instruments	BF140-117-10
Fine-edged dissection blade	Fine Science Tools	10010-00
Glass Pasteur Pipettes	VWR	14673-010	to pull trituration pipettes
Heat-inactivated Fetal Bovine Serum	Thermo Fisher Scientific	16140063	additive to culture medium
HEPES	Sigma-Aldrich	H3375-100G	for pH buffering all solutions in protocol
Hot plate / magnetic stirrers	VWR	76549-914
Insulated bucket filled with ice			to keep all samples and solutions cool
K2SO4, Potassium Sulfate	Sigma Aldrich	P9458-250G	Additive to internal solution
KCl	Sigma-Aldrich	P93333	Additive to internal solution
KOH (1 M)	Honeywell	319376-500ML	To bring internal solution to desired pH.
Large Spring Scissors	Fine Science Tools	14133-13
Leibovitz medium	Sigma Aldrich	L4386	dissection and bath solutions
Low-profile-bath recording chamber for culture dishes	Warner Instruments	64-0236
luer-lok syringes, 30 ml	BD	302832	for drawing L-15/HEPES/HEPES solution.
MEM + Glutamax Supplement	Fisher Scientific	41-090-101	base of the culture medium
MgCl2-Hexahydrate	Sigma-Aldrich	M1028	Additive to internal solution
microFil needle for filling micropipettes - 34 gauge	World Precision Instruments	MF34G
Microforge	Narashige	MF-90	For electrode polishing.
N2 Supplement (100x)	Thermo Fisher Scientific	17502-048	additiive to culture medium, for SGN
NaCl	Sigma-Aldrich	S7653	Additive to internal solution
NaOH (1 M)	Thomas Scientific	319511-500ML	for titration pH
Osmometer	Advanced Instruments Inc.	3320
Oxygen, Medical grade, with adequate regulator and tubing	USC Material Management	MEDOX200 (Identifier: 00015)	for dissolving into dissection and bath solutions
Parafilm	Bemis	PM992
Pasteur pipette bulb (3 ml)	Fisher Scientific	03-448-25	bulb for trituration pipettes
Penicillin/Streptomycin	Thermo Fisher Scientific	15140122	additive to prevent contamination of culture medium
Pentobarbital based euthanasia solution (e.g., Fatal Plus. 50 – 60 mg/kg dosing)	MWI Animal Health	15199	for euthanasia
PES membrane filters ,  0.2 micrometer	Nalgene	566-0020	for filtering solutions
PES membrane sterile syringe filters, 0.22 um, 30 mm	CELLTREAT	229747	for filtering solutions drawn into syringes
Petri dishes, 35 x 10 mm	Genessee Scientific	32-103	for micro dissection and to hold Tip dip solution in perforated-patch configuration
Petri Dishes, 60 x 15 mm	Midland Scientific	P7455	for gross dissection
pH Meter	Mettler Toledo	Model S20
Pipettors (1000, 200, 10) microliter	USA Scientific
Poly-d-lysine coated glass bottomed culture dish	Mattek	P35GC-0-10-C	to plate neurons for culture
Quick change platform, heated base, for 35 mm culture dishes	Warner Instruments	64-0375
Reference Cell	World Precision Instruments	RC1T
Scalpel blade	Miltex	4-315
Scalpel Handle	Fine Science Tools	10003-12
Scientific Scale	Mettler Toledo	XS64
Serological Pipettes (10, 25) milliliter	Fisher Scientific
Silicone Grease Kit (for sealing coverglass and chamber)	Warner Instruments	64-0378
Small Animal Guillotine	Kent Scientific	DCAP
Small animal guillotine	Kent Scientific	DCAP	for decapitation if dissecting rats older than P15.
Stereo Dissection Microscope	Zeiss	Stemi 2000
Straight surgical scissors	Fine Science Tools	14060-09
Syringe (3, 10, 30) milliliter
Trypsin	Sigma Aldrich	T1426	one out of three enzyme to digest tissue
Tuberculin syringe	Covidien	8881500105	for delivering euthanasia solution by intraperitoneal injection
Vannas Spring Scissor, 2.5 mm Cutting Edge	Fine Science Tools	15000-08
Volumetric flask, 1000 milliliter
Vortex	VWR	945300
Water, sterile u ltrapure, R>18.18 megaOhms cm (e.g., filtered by a Millipore-Sigma water purification system)	Millipore-Sigma	CDUFBI001