The authors wish to acknowledge Dr. A. J. Hudspeth for funding and expertise in developing the preparations described in this paper. We also wish to thank Brian Fabella for creating and maintaining much of the custom equipment and software used in this protocol.
J. B. A. is supported by grant F30DC014215, J. D. S. is supported by grant F30DC013468, and both J. B. A. and J. D. S. are supported by grant T32GM07739 from the National Institutes of Health.

Ethics Statement: All procedures were approved by the Institutional Animal Care and Use Committee (IACUC) at The Rockefeller University.
1. Pre-experimental Prespanparation
<ol>
	<li>Solutions
	<ol>
		<li>Prepare the eugenol solution (2.5 g&#183;L-1&#183;kg-1 frog). Prepare the saline solutions (Table 1). 
		NOTE: For a one-chamber preparation, prepare artificial perilymph; for a two-chamber preparation, prepare both artificial perilymph and artificial endolymph.</li>
	</ol>
	</li>
</ol>
2. Experimental Tools
<ol>
	<li>Preparation of glass stimulation fiber
	<ol>
		<li>Narrow a borosilicate glass capillary with an electrode puller in a one-line pull with high heat and high velocity. Load the pulled capillary into a solenoid-driven puller.</li>
		<li>Bring the capillary&#39;s tip toward a filament with a glass bead melted onto it until the capillary contacts the glass bead.</li>
		<li>Turn on the filament to melt the tip of the capillary into the glass bead. Once a thin bridge of glass forms between the capillary and the bead, turn off the filament and around the same time activate the solenoid puller. 
		NOTE: This will pull the glass capillary at a right angle to its long axis and create a solid fiber.</li>
		<li>Ensure that the fiber&#39;s diameter is no more than 0.5 - 1 &#181;m and that its length does not exceed 100 - 300 &#181;m. If the fiber&#39;s length exceeds this dimension, trim it with iris scissors. Use scissors that are already dull to avoid damaging the dissection tools.</li>
		<li>To enhance optical contrast, sputter coat the fiber. Use a sputter coater with a gold-palladium source. Vertically load each fiber into a sputter coater with its tapered end towards the source. Bring the fiber&#39;s tip to a distance of 1 - 2 cm from the gold-palladium source.</li>
		<li>Close the sputter coater&#39;s chamber to form a seal and turn it on. Repeatedly flush out the ambient air with argon. After flushing the air, reduce the pressure within the chamber to 10 Pa (70 mTorr).</li>
		<li>Sputter coat in 10 s pulses with 10 s delays over a course of 120 s. 
		NOTE: The fiber&#39;s tip will darken over the duration of this protocol if the sputter coating was a success.</li>
	</ol>
	</li>
	<li>Preparation of sharp microelectrodes
	<ol>
		<li>Pull a glass capillary with an internal filament using a one-line high-heat pull. Electrodes should have a resistance of 100 - 300 M&#937; when filled with 3 M KCl. Fill each electrode with 3 M KCl.</li>
		<li>Bend each electrode&#39;s tip with the microforge to render it perpendicular to the hair cell apical surface when mounted in the amplifier headstage16.</li>
	</ol>
	</li>
	<li>Preparation of iontophoresis pipettes
	<ol>
		<li>Pull a glass capillary with an internal filament to a 50 M&#937; tip. Fill with concentrated solute (e.g., 500 mM gentamicin sulfate).</li>
	</ol>
	</li>
	<li>Preparation of aluminum mounting square
	<ol>
		<li>Cut a 1 cm x 1 cm square of aluminum foil. Perforate the foil in the center with the tip of a pithing rod or other sharp object. Shape the perforation so that it is circular and approximately 1 mm in diameter.</li>
	</ol>
	</li>
	<li>Preparation of vacuum grease-filled syringe.
	<ol>
		<li>Remove the plunger from a 5 mL syringe. Fill syringe from the back with vacuum grease. Replace the plunger.</li>
	</ol>
	</li>
	<li>Preparation of polytetrafluoroethylene glue applicator
	<ol>
		<li>Make a 2 mm long axial cut into the end of a wooden applicator stick. The slit should be in the center of the applicator when viewed end-on.</li>
		<li>With sharp scissors or a razor blade, cut a 2 mm x 4 mm rectangle of polytetrafluoroethylene from a sheet of 1 mm thickness.</li>
		<li>Use a razor blade to thin one of the 2 mm-long edges of the polytetrafluoroethylene rectangle. Starting in the middle of the rectangle, reduce the thickness of the rectangle by slicing outward to form a bevel at the tip of the rectangle.</li>
		<li>Insert the polytetrafluoroethylene rectangle into the wooden applicator slit such that the beveled edge faces away from the stick. Apply 5 min&#160;epoxy to the base of the polytetrafluoroethylene rectangle to secure it in place. Allow the glue applicator to cure for 1 h at room temperature.</li>
	</ol>
	</li>
	<li>Preparation of lower half of two-chamber mount
	<ol>
		<li>Machine or 3D print the lower half of the two-chamber mount (Supplemental file 2). Identify the 20 mm diameter recessed circle on the lower surface of the chamber.</li>
		<li>Apply a thin layer of epoxy to the lateral most 3 mm of the recessed circle. Apply one 18 mm circular glass coverslip on the epoxy. Allow epoxy to cure for 1 h.</li>
	</ol>
	</li>
</ol>
3. Extraction of Inner-ear Organs
<ol>
	<li>Anesthetize an American bullfrog by placing it in a small bucket containing eugenol anesthetic solution for 10 min. Adjust the volume of solution so that the frog&#39;s humero-scapular joint lies just above the liquid-air interface.</li>
	<li>Euthanize the anesthetized bullfrog by double pithing.
	<ol>
		<li>Grasp the anesthetized frog with one finger atop its nose and another below the jaw and rotate the frog&#39;s head forward.</li>
		<li>Rapidly plunge the pithing rod into the cranial vault through the foramen magnum, which is found at the midline between the frog&#39;s occipital processes.</li>
		<li>Slowly withdraw and rotate the rod until its tip departs from the foramen magnum. Force the rod caudally through the vertebral foramina to destroy the spinal cord.</li>
		<li>Confirm that the frog has been properly doubly pithed by observing that its lower extremities are extended.</li>
	</ol>
	</li>
	<li>Grasp the frog with a thumb atop the nose and first finger grasping the vomerine teeth for improved stability. Decapitate the frog by severing the temporomandibular joint bilaterally and subsequently cutting orthogonally to the rostrocaudal axis. To assure that the inner-ear organs within the skull remain intact, ensure that the cut is caudal to both tympana.</li>
	<li>Using a stereodissection microscope, execute a midline cut through the palatal tissue from the vomerine teeth to the most posterior extent of the tissue (Figure 1A).</li>
	<li>Sever and clear with horizontal cuts of the scalpel any muscle lying below the palatal tissue to reveal the posterior cartilage. After removing the muscle, observe the lollipop shape of the temporal cartilage forming the boundary of the otic capsule.</li>
	<li>Sever the columella at its point of contact with the otic capsule&#39;s cartilage.</li>
	<li>Repeatedly shave thin layers of this cartilage by making shallow horizontal cuts through it. Avoid deep cuts to prevent damage to the inner-ear organs. This opens the otic capsule found within the lollipop structure of the temporal cartilage (Figure 1B). Within the otic capsule are the frog&#39;s inner-ear organs (Figure 1C).</li>
	<li>Trim the posterior and lateral edges of the otic capsule, taking care not to damage the inner-ear organs. During the dissection, frequently flow saline over the inner ear organs to ensure that they remain submerged and hydrated.</li>
	<li>Locate the two circular openings in the temporal bone at the medial connection of the cartilage to the midline. Cut downward through the most medial opening to rupture the temporal bone.</li>
	<li>Make a second downward cut through the otic capsule at its postero-lateral edge. Remove the piece of cartilage between this and the previous cut to provide access to the inner-ear organs.</li>
	<li>Pry loose the cartilage between the cuts from steps 3.8 and 3.9 and cut it away from the otic capsule. This action severs the nearest semicircular canal, which can then be used as a handle for further manipulations.</li>
	<li>Taking care not to touch the sacculus, sever the VIIIth cranial nerve.</li>
	<li>Hold the ampulla of the nearest semicircular canal. Gently rotate the inner ear to expose the remaining two semicircular canals. Upon exposing each canal, sever it.</li>
	<li>Holding the nerve or a semicircular canal ampulla, extract the inner ear from the head and place it in a dish filled with chilled oxygenated artificial perilymph. Removal of the inner ear permits visualization of the passages within the temporal bone through which the semicircular canals once passed (Figure 1D). 
	NOTE: Repeat these steps to extract the second ear.</li>
</ol>
4. One-chamber Preparation
<ol>
	<li>Isolation of sacculus
	<ol>
		<li>Locate the sacculus by identifying its large white mass of otoconia and the saccular nerve that lies atop it (Figure 1E). Take care not to physically traumatize the sacculus during the following steps in order to maintain the integrity of saccular hair cells.</li>
		<li>Trim the semicircular canals to render the inner ear more maneuverable.</li>
		<li>Remove the perilymphatic cistern overlying the neural side of the sacculus. Make gentle cuts around the perimeter of the cistern. Next sever the small pillars of tissue that bridge the cistern&#39;s membrane to the neural side of the sacculus.</li>
		<li>Remove the lagena and its associated nerve.</li>
		<li>Holding the saccular nerve, gently lift the sacculus and cut through the thin membrane of the otoconial sac. As otoconia spill out of the sac, free the sacculus by cutting around its perimeter. 
		NOTE: After isolating the sacculus, the remaining inner-ear organs can be saved if desired. Removal of the sacculus eases identification of other structures, such as the amphibian and basilar papillae.</li>
		<li>Use scissors or forceps to carefully wipe away any otoconia remaining on the sacculus. Take care not to touch the sacculus during this process.</li>
		<li>Trim any remaining otoconial-sac membrane from the sacculus&#39; edge. This membrane tends to adhere to plastic and glass surfaces and its removal minimizes challenges in handling the tissue.</li>
		<li>Gently flow saline over the sacculus with a Pasteur pipette to remove any remaining otoconia (Figure 1F). 
		NOTE: Repeat this procedure for the second sacculus, which is a mirror image of the first.</li>
	</ol>
	</li>
	<li>Digestion and mounting
	<ol>
		<li>Using the back end of a Pasteur pipette with its tip broken, transfer the isolated sacculi to a Petri dish containing 3 mL of 67 mg&#8729;L-1 protease XXIV in artificial perilymph. 
		NOTE: This protease treatment digests the links tethering each kinociliary bulb to the otolithic membrane, enabling removal of the membrane without damaging the sensory hair bundles.</li>
		<li>Incubate the tissue for 30 min at 22 &#176;C (or 35 min at 21 &#176;C). Transfer each digested sacculus to an open-faced experimental chamber and secure the tissue with magnetic pins.</li>
		<li>The saccular macula lies directly below the otolithic membrane and is the portion of the sacculus that contains hair cells. Identify it and carefully remove its overlying otolithic membrane using a fine eyelash, taking care not to touch the saccular macula.</li>
	</ol>
	</li>
</ol>
5. Two-chamber Preparation
<ol>
	<li>Isolation of sacculus
	<ol>
		<li>Isolate the sacculus from the inner-ear organs as in Section 4.1.</li>
	</ol>
	</li>
	<li>Mounting and digestion
	<ol>
		<li>Fill the mounting block (Supplemental file 3) with artificial perilymph and place the perforated aluminum foil on one opening, using two spots of vacuum grease to hold it in place and form a weak seal.</li>
		<li>Transfer one sacculus to the foil and center it on top of the hole with the macula facing down and the nerve stump facing up.</li>
		<li>Remove the saline surrounding the sacculus with a piece of twisted tissue. Wick the saline to dry the surface of the aluminum square surrounding the sacculus.</li>
		<li>Use the polytetrafluoroethylene applicator to apply cyanoacrylate glue to form a tight seal along the boundary between the edge of the sacculus and the aluminum square. Ensure that the entire circumference of the sacculus is covered with glue. 
		NOTE: Proceeding too slowly allows the cyanoacrylate glue to creep over the neural side of the sacculus and eventually cover it. It is therefore imperative to complete this task rapidly.</li>
		<li>Place a drop of saline on top of the mounted tissue to cure the glue. A thin film of glue may form on top of the drop of saline; remove it with forceps.</li>
		<li>Carefully remove the foil from the mounting block. Flip the mounted tissue over so that the macular side of the sacculus faces upward and float it on an artificial perilymph-filled Petri dish.</li>
		<li>Add a drop of protease XXIV solution on top of the macula and incubate for 30 min at 22 &#176;C (or 35 min at 21 &#176;C). Fill the lower canal of the two-chamber apparatus (Supplemental file 2) with perilymph and place vacuum grease around the central chamber.</li>
		<li>Place the foil-mounted sacculus on the lower chamber with its nerve facing the chamber&#39;s surface. Add grease around the perimeter of the foil.</li>
		<li>Place the upper chamber (Supplemental file 1) of the preparation on the foil, taking care to form a complete seal with the vacuum grease. Fill the upper chamber with bubbled artificial endolymph and gently remove the otolithic membrane with an eyelash.</li>
	</ol>
	</li>
</ol>

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F., J&#252;licher, F., Martin, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transduction+channels'+gating+can+control+friction+on+vibrating+hair-cell+bundles+in+the+ear.">Transduction channels' gating can control friction on vibrating hair-cell bundles in the ear.</a> Proceedings of the National Academy of Sciences of the United States of America. 111 (20), 7185-7190 (2014).</li><li>Crawford, A. C., Evans, M. G., Fettiplace, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+actions+of+calcium+on+the+mechano-electrical+transducer+current+of+turtle+hair+cells.">The actions of calcium on the mechano-electrical transducer current of turtle hair cells.</a> The Journal of physiology. , 369-398 (1991).</li><li>Kozlov, A. S., Risler, T., Hudspeth, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coherent+motion+of+stereocilia+assures+the+concerted+gating+of+hair-cell+transduction+channels.">Coherent motion of stereocilia assures the concerted gating of hair-cell transduction channels.</a> Nature Neuroscience. 10 (1), 87-92 (2007).</li><li>Lumpkin, E. A., Hudspeth, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+of+free+Ca2++concentration+in+hair-cell+stereocilia.">Regulation of free Ca2+ concentration in hair-cell stereocilia.</a> Journal of Neuroscience. 18 (16), 6300-6318 (1998).</li></ol>

The authors declare no competing interests.

Within the bullfrog&#39;s sacculus lie several thousand easily-accessible sensory hair cells. Here we demonstrate extraction and preparation of the sacculus for one- and two-chamber recordings. These two preparations permit both micromechanical and electrophysiological studies of hair cells and their associated bundles. Because the tissue can survive for several hours with frequent replacement of oxygenated saline, experiments may continue for long durations. Hair cells in these preparations typically remain viable for microelectrode recording for up to 6 h after dissection, while hair bundles oscillate spontaneously for up to 24 h after extraction.
Successful extraction and mounting of the sacculus hinges upon surmounting several common challenges. First, direct contact with the apical surface of the saccular macula should be avoided throughout the preparation procedure. The saccular nerve provides a convenient handle for safe manipulation of the sacculus. Once freed from the remainder of the inner-ear organs, the sacculus should be transferred using a large-bore pipette while remaining immersed in fluid to avoid mechanical damage to its sensory epithelium. The removal of otoconia from the macular surface must be completed without mechanical damage to hair cells. Because the otoconia lie directly atop the macula, hair cells can be damaged by accidental contact between dissection tools and the otolithic membrane while removing otoconia. To avoid damage, we recommend that the gelatinous mass of otoconia be held at a location far from the macula and removed as a single mass. This avoids fragmentation of the otoconial mass into numerous clusters, each of which would be individually extracted. If small clusters of otoconia remain they can be removed with gentle fluid pressure delivered by a Pasteur pipette. A final challenge involves the formation of a tight seal between the sacculus and aluminum mounting square in the two-chamber preparation. Employing a square with a perforation small enough to allow overlap of about 100 &#956;m between the sacculus and the surrounding aluminum permits complete sealing of the tissue. The glue should be brought into contact with approximately 100 &#956;m of saccular tissue around the macula&#39;s perimeter in order to form a tight seal.
The concentration of free Ca2+ is an important consideration in the study of hair cells. Ca2+ regulates both fast and slow adaptation, thus determining the kinetics of the mechanotransduction apparatus and the characteristics of the hair bundle&#39;s active-process phenomena, including spontaneous bundle motion8,23. Endolymphatic calcium in vivo is present at 250 &#956;M, therefore the most physiologically relevant kinetics are assessed at this concentration (Maunsell J. H. R., R. Jacobs, and A. J. Hudspeth. Unpublished observations16). However, microelectrode recordings from hair cells require an external calcium concentration exceeding 2 mM for proper sealing of the cellular membrane around the microelectrode. It is therefore imperative to use a high-calcium saline for these experiments. Finally, one may wish to study the effects of external calcium upon mechanotransduction using a variety of calcium concentrations. In these cases, it is important to remember that calcium concentrations below 1 &#956;M typically lead to tip-link rupture and irreversible loss of transduction28.
The two experimental preparations described here allow for a range of biophysical measurements on hair cells. However, additional measurements can be made with slight modifications to these preparations. In the folded saccular preparation, hair bundles are visualized laterally. Imaging hair-bundle motion from this vantage point reveals coherent motion of both short and tall stereocilia29. Here the saccular macula is first separated from its underlying tissue and subsequently folded along the axis defined by the saccular nerve such that hair bundles face outwards and are visualized laterally at the crease. A second modification, hair-cell dissociation, enables the study of both the hair cell&#39;s bundle and its soma. Hair cells are mechanically dissociated onto a glass slide for imaging and electrophysiological recording30. Finally, hair cells can be extruded from the epithelium by following a similar dissociation protocol but without the mechanical dissociation step. This treatment results in hair cells that gradually extrude out of the epithelium, providing basolateral access for electrophysiological recordings while minimizing mechanical damage. These preparations and their many modifications demonstrate the versatility of the frog sacculus as a model system for the biophysical study of sensory hair cells.

The acousticolateralis organs of mammals possess a complex architecture and lie within an anatomical niche that can be difficult to access. For example, the mammalian cochlea comprises a spiraling labyrinth and is embedded within the thick temporal bone. Isolation of the cochlea often causes mechanical damage to the sensory cells lying within it and has therefore proven to be a difficult task1. Neuroscientists have thus turned to model systems which are more readily extracted from the sanctum of the ear.
One of these model systems, the sacculus of the American bullfrog (Rana catesbeiana), has for decades yielded generalizable insight into the function of auditory and vestibular systems. The sacculus is a mixed-function organ with sensory roles in both low-frequency hearing and seismic sensation. The sensory cells of the sacculus are its hair cells, specialized transducers that convert mechanical energy into electrical signals within our auditory and vestibular organs. Projecting from the apical surface of each hair cell is a mechanosensitive hair bundle that comprises a graded tuft of enlarged microvilli called stereocilia. The tips of adjacent stereocilia are interconnected by filamentous tip-link proteins that mechanically gate ion channels in response to mechanical stimuli2,3. Although auditory and vestibular organs respond to different types of stimuli, they share a common detection mechanism. This commonality underlies the many insights gained into hair-cell mechanotransduction through studies of the bullfrog sacculus. For example, the hair cell&#39;s active process has been studied extensively in this organ4,5,6,7, and the hair bundle employs an energy-consuming process to produce mechanical work. Not only has it been shown that hair cells generate active work6, but distinct mechanisms underlying the active process and a hair cell&#39;s tuning characteristics have been unveiled through studies of bullfrog acousticolateralis organs. These include active hair-bundle motility8 and hair cell electrical resonance9,10,11 in the sacculus and frequency selectivity at the hair cell&#39;s ribbon synapse12 in the amphibian papilla.
The bullfrog&#39;s sacculus appeals to sensory neuroscientists for numerous reasons. Unlike the mammalian cochlea, this organ lies within the readily accessible otic capsule. Second, hair cells within this organ can remain healthy for several hours under appropriate conditions13,14. This permits experimentation on these cells over long timescales relative to their mammalian counterparts. Third, the organ bears little curvature, permitting easy manipulation. Fourth, each organ comprises a thousand or more hair cells15, providing both a high throughput and a high probability of locating an appropriate set of hair cells for a given experiment. Finally, the bullfrog&#39;s sacculus is easily visualized due to the thinness of this organ and large size of its hair cells.
These properties provide great versatility for the study of sensory cells within the bullfrog&#39;s sacculus. Depending on the question at hand, one of several experimental preparations can be obtained from the sacculus. The simplest of these is the one-chamber preparation. Here the sacculus is immobilized in a chamber filled with artificial perilymph, a sodium-rich and high-calcium saline. This preparation enables the study of hair cell currents and basic hair bundle mechanics. A second configuration, the two-chamber preparation, can be used to study spontaneous hair bundle movements. Here the apical side of hair cells is exposed to a potassium-rich and calcium-poor saline termed artificial endolymph, whereas the basolateral side is bathed in artificial perilymph. These two compartments mimic the in vivo arrangement of salines and provide an environment that allows hair bundles to oscillate spontaneously.
We describe in this paper the preparation of the bullfrog&#39;s sacculus for biophysical study of its sensory hair cells. We first provide a detailed depiction of the isolation of this organ from the frog&#39;s inner ear. We then describe both the one- and two-chamber experimental preparations and include representative results for each configuration.

<table border="0" cellpadding="0" cellspacing="0" width="790">
	<tbody>
		<tr height="20">
			<td colspan="4" height="20" style="height:20px;width:789px;">Common to both preparations</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Stereo-dissection microscope</td>
			<td style="width:189px;">Leica</td>
			<td style="width:119px;">MZ6</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Tricaine methanesulfonate</td>
			<td style="width:189px;">Sigma</td>
			<td style="width:119px;">E10521</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Metal pithing rod</td>
			<td style="width:189px;">Fine Science Tools</td>
			<td style="width:119px;">10140-01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Vannas spring scissors</td>
			<td style="width:189px;">Fine Science Tools</td>
			<td style="width:119px;">15000-03</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Dumont #5 forceps</td>
			<td style="width:189px;">Fine Science Tools</td>
			<td style="width:119px;">11252-20</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Glass Pasteur pipette and bulb (x2)</td>
			<td style="width:189px;">Fisher Scientific</td>
			<td style="width:119px;">22-042816</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:299px;">Fine eyelash mounted on a hypodermic needle</td>
			<td style="width:189px;">Fisher Scientific</td>
			<td style="width:119px;">22-557-172</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Dow-corning vacuum grease</td>
			<td style="width:189px;">Fisher Scientific</td>
			<td style="width:119px;">14-635-5C</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Syringe for vacuum grease</td>
			<td style="width:189px;">Fisher Scientific</td>
			<td style="width:119px;">14-829-45</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">35 mm Petri dish (x2-3)</td>
			<td style="width:189px;">Fisher Scientific</td>
			<td style="width:119px;">08-772A</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Micropipette puller</td>
			<td style="width:189px;">Sutter</td>
			<td style="width:119px;">P-97 or P-2000</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">120 V Solenoid puller</td>
			<td style="width:189px;"></td>
			<td style="width:119px;"></td>
			<td>Home-made, see parts list</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Sputter coater</td>
			<td style="width:189px;">Anatech USA</td>
			<td style="width:119px;">Hummer 6.2</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Current source for iontophoresis</td>
			<td style="width:189px;">Axon Instruments</td>
			<td style="width:119px;">AxoClamp 2B</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Piezoelectric actuator</td>
			<td style="width:189px;">Piezosystem Jena</td>
			<td style="width:119px;">P-150-00</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Amplifier for piezoelectric actuator</td>
			<td style="width:189px;">Piezosystem Jena</td>
			<td style="width:119px;">ENV800</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:299px;">Borosilicate glass capillary</td>
			<td style="width:189px;">World Precision Instruments</td>
			<td style="width:119px;">1B120F-3</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Name</td>
			<td style="width:189px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr height="20">
			<td colspan="4" height="20" style="height:20px;width:789px;">For one-chamber preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Microelectrode amplifier</td>
			<td style="width:189px;">Axon Instruments</td>
			<td style="width:119px;">AxoClamp 2B</td>
			<td>Can be used for iontophoresis and microelectrode recordings simultaneously</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Magnetic pins (x2)</td>
			<td style="width:189px;"></td>
			<td style="width:119px;"></td>
			<td>Home-made, see parts list</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Open-top chamber with magnetic sheet</td>
			<td style="width:189px;"></td>
			<td style="width:119px;"></td>
			<td>Home-made, see parts list</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Name</td>
			<td style="width:189px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr height="20">
			<td colspan="4" height="20" style="height:20px;width:789px;">For two-chamber preparation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Upper chamber</td>
			<td style="width:189px;"></td>
			<td style="width:119px;"></td>
			<td>Supplementary file 1</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Troughed lower chamber</td>
			<td style="width:189px;"></td>
			<td style="width:119px;"></td>
			<td>Supplementary file 2</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Aluminum foil</td>
			<td style="width:189px;">Fisher Scientific</td>
			<td style="width:119px;">01-213-100</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Mounting block</td>
			<td style="width:189px;"></td>
			<td style="width:119px;"></td>
			<td>Supplementary file 3</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Wooden applicator sticks</td>
			<td style="width:189px;">Fisher Scientific</td>
			<td style="width:119px;">23-400-112</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Teflon sheet</td>
			<td style="width:189px;">McMaster-Carr</td>
			<td style="width:119px;">8545K12</td>
			<td>For teflon applicator</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Cyanoacrylate glue</td>
			<td style="width:189px;">3M</td>
			<td style="width:119px;">1469SB</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Lab tissues (Kimwipes)</td>
			<td style="width:189px;">Fisher Scientific</td>
			<td style="width:119px;">06-666A</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Gentamicin sulfate</td>
			<td style="width:189px;">Sigma-Aldrich</td>
			<td style="width:119px;">G1914</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Quick-setting epoxy</td>
			<td style="width:189px;">McMaster-Carr</td>
			<td style="width:119px;">7605A18</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">18 mm glass coverslips</td>
			<td style="width:189px;">Fisher Scientific</td>
			<td style="width:119px;">12-546</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Name</td>
			<td style="width:189px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Saline components</td>
			<td style="width:189px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">NaCl</td>
			<td style="width:189px;">Fisher Scientific</td>
			<td style="width:119px;">S271-3</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">KCl</td>
			<td style="width:189px;">Sigma-Aldrich</td>
			<td style="width:119px;">P4504-500G</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:299px;">CaCl2 &#8226; 2H2O</td>
			<td style="width:189px;">Fisher Scientific</td>
			<td style="width:119px;">10035-04-8</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">HEPES</td>
			<td style="width:189px;">Sigma-Aldrich</td>
			<td style="width:119px;">H3375-100G</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">D-(+)-glucose</td>
			<td style="width:189px;">Sigma-Aldrich</td>
			<td style="width:119px;">G7021</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Name</td>
			<td style="width:189px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td>Comments/Description</td>
		</tr>
		<tr height="20">
			<td colspan="4" height="20" style="height:20px;width:789px;">Parts lists for home-made equipment</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Solenoid puller</td>
			<td style="width:189px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Solenoid</td>
			<td style="width:189px;">Guardian Electric</td>
			<td style="width:119px;">A420-065426-00</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Foot-pedal switch</td>
			<td style="width:189px;">Linemaster</td>
			<td style="width:119px;">T-51-SC36</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:299px;">Pipette holder</td>
			<td style="width:189px;">World Precision Instruments</td>
			<td style="width:119px;">MEH900R</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Coarse manipulator</td>
			<td style="width:189px;">Narishige Group</td>
			<td style="width:119px;">MM-3</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Platinum wire</td>
			<td style="width:189px;">Alfa Aesar</td>
			<td style="width:119px;">25093</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Power supply</td>
			<td style="width:189px;">Leica</td>
			<td style="width:119px;">Z050-261</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Name</td>
			<td style="width:189px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td>Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Magnetic pins</td>
			<td style="width:189px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Epoxy</td>
			<td style="width:189px;">McMaster-Carr</td>
			<td style="width:119px;">7556A33</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">1 mm thickness aluminum</td>
			<td style="width:189px;">McMaster-Carr</td>
			<td style="width:119px;">89015K45</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Insect pins</td>
			<td style="width:189px;">Fine Science Tools</td>
			<td style="width:119px;">26000-40</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Name</td>
			<td style="width:189px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td>Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Open-top magnetic chamber</td>
			<td style="width:189px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">Flexible magnetic strip</td>
			<td style="width:189px;">McMaster-Carr</td>
			<td style="width:119px;">5759K75</td>
			<td>Other sources can be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:299px;">1 mm thickness aluminum</td>
			<td style="width:189px;">McMaster-Carr</td>
			<td style="width:119px;">89015K45</td>
			<td>Other sources can be used</td>
		</tr>
	</tbody>
</table>

physiological preparation of hair cells from the sacculus of the american bullfrog (rana catesbeiana)

The study of hearing and balance rests upon insights drawn from biophysical studies of model systems. One such model, the sacculus of the American bullfrog, has become a mainstay of auditory and vestibular research. Studies of this organ have revealed how sensory cells hair can actively detect signals from the environment. Because of these studies, we now better understand the mechanical gating and localization of a hair cell&#39;s transduction channels, calcium&#39;s role in mechanical adaptation, and the identity of hair cell currents. This highly accessible organ continues to provide insight into the workings of hair cells. Here we describe the preparation of the bullfrog&#39;s sacculus for biophysical studies on its hair cells. We include the complete dissection procedure and provide specific protocols for the preparation of the sacculus in specific contexts. We additionally include representative results using this preparation, including the calculation of a hair bundle&#39;s instantaneous force-displacement relation and measurement of a bundle&#39;s spontaneous oscillation.

The sensory epithelium of the bullfrog&#39;s sacculus can be employed in various configurations to probe the physiology of hair cells. Because the tissue is relatively flat, it can be mounted in both one- and two-chamber preparations. The one-chamber configuration provides a simple setup for electrophysiological and micromechanical recordings of hair cells. The two-chamber preparation instead simulates both the endolymphatic and perilymphatic compartments on respectively the apical and basal sides of hair cells. These compartments together provide a physiologically-relevant environment for the study of mechanotransduction by hair cells.
The sensitivity and transduction characteristics of hair cells underlie their electrical response to mechanical stimulation. To probe these features, we simultaneously recorded from an individual hair cell the position of its bundle and the cell&#39;s receptor potential (Figure 2). We first attached a flexible glass fiber to the kinociliary bulb of a hair bundle to apply force pulses. We then measured the hair bundle&#39;s displacement using a dual photodiode system2 (Figure 2A). We concurrently acquired the hair cell&#39;s potential by impaling the cell with a sharp microelectrode. We obtained a displacement-response curve by plotting the peak voltage response elicited by each mechanical stimulus against the hair bundle&#39;s corresponding displacement (Figure 2B). The hair cell&#39;s electrical response saturates for both positive- and negative-displacement extrema. The reduction of membrane potential with negative displacement steps indicates the presence of a resting inward mechanotransduction current. This resting current is modulated by the action of Ca2+ on both fast and slow adaptation17,18,19,20,21.
A hair cell&#39;s behavior depends not only upon its electrical properties, but also upon the micromechanics of its sensory hair bundle. The two-chamber configuration mimics the separation of endolymph and perilymph in vivo, providing ideal conditions for the study of a hair bundle&#39;s mechanics. Under these conditions and with the otolithic membrane removed, hair bundles can oscillate spontaneously6. Here we employed the two-chamber preparation to assess the micromechanics of individual bundles. We recorded the spontaneous oscillations of a hair bundle by casting its shadow on a dual-photodiode displacement monitor (Figure 3). To assess the role of the aminoglycoside antibiotic gentamicin on mechanotransduction, we iontophoretically released gentamicin directly onto the hair bundle (Figure 3). The concentration of released gentamicin rises proportionally with the current passed through the micropipette. Gentamicin inhibits a hair bundle&#39;s oscillations and induces a static offset of the bundle towards its tall side. These effects reflect the role of gentamicin as an open-channel blocker that maintains the open state of mechanotransduction channels while blocking their permeation pore. Iontophoresis of charged chemicals permits localized and quantifiable release of chemicals at various concentrations in the absence of fluid flow-induced mechanical disruption and is thus ideally suited for the study of mechanosensitive organelles such as the hair bundle22.
A hair bundle&#39;s spontaneous motion arises from the interplay between adaptation and nonlinear bundle stiffness7,8,23,24. This spontaneous motion is a signature of a hair bundle&#39;s active process, which converts signal energy into mechanical work to overcome viscous drag. Hair bundles have been shown to exhibit nonlinear instantaneous stiffness in the vestibular2, auditory25, and lateral-line systems26.
We directly measured the instantaneous stiffness of an individual hair bundle from the bullfrog&#39;s sacculus (Figure 4). To achieve this, we coupled the tip of a flexible glass fiber to the hair bundle&#39;s kinociliary bulb (Figure 4A). We delivered forces to the hair bundle by displacing the fiber&#39;s base. The force exerted onto the hair bundle by the stimulus fiber corresponds to the difference between the displacements of the fiber&#39;s base and tip, multiplied by the fiber&#39;s stiffness2,27. Delivering pulses across a range of forces reveals a relationship between the force exerted onto the bundle and the bundle&#39;s ensuing displacement. The slope of this force-displacement relation corresponds to the hair bundle&#39;s instantaneous stiffness (Figure 4A).
This method allowed us to measure an individual bundle&#39;s instantaneous stiffness as a function of its deflection (Figure 4B). The instantaneous force-displacement curve displays a nonlinear relationship, revealing a nonlinear stiffness of the bundle over a range of about 20 nm around its resting position. Outside this range, the hair bundle behaves like a Hookean material, its stiffness is linear for large-magnitude deflections.
These results demonstrate the versatility of the bullfrog sacculus in the study of hair-cell physiology. Using these and other preparations, one can explore mechanotransduction at multiple stages in the transmission of information from the bundle toward the brain.
<img alt="Figure 1" src="/files/ftp_upload/55380/55380fig1.jpg" /> 
Figure 1: Dissection of the Bullfrog&#39;s Inner Ear. (A) Viewing the bullfrog&#39;s upper palate from its ventral side permits identification of the Eustachian tube (circle). Lateral reflection of the skin covering the right side of the upper palate reveals the location of the inner ear (dashed box). (B) Removal of the cartilage on the ventral side of the frog&#39;s temporal bone opens the otic capsule (dashed line). (C) Displayed is a higher magnification image of the otic capsule, in which the sacculus, lagena, CN VIII, and saccular nerve can be readily identified. (D) A view of the otic capsule after removal of the inner-ear organs reveals the locations of the semicircular canals. (E) After removal of the isolated inner ear, the sacculus, lagena, and VIIIth cranial nerve (CN VIII) can readily be identified. (F) The isolated sacculus possesses a short stump of the saccular nerve and an otolithic membrane lying atop its sensory epithelium. Labels correspond to the (i) sacculus, (ii) lagena, (iii) CN VIII, (iv) saccular nerve, and (v) otolithic membrane. Axis labels P and A correspond respectively to the posterior and anterior directions. Scale bars represent 1 cm (A, B), 1 mm (C, D, E), and 400 &#956;m (F). 
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<img alt="Figure 2" src="/files/ftp_upload/55380/55380fig2.jpg" /> 
Figure 2: Displacement-response Curve for a Single Hair Cell. (A) The tip of a glass stimulus fiber was coupled to the kinociliary bulb of a hair bundle and the fiber&#39;s base was subsequently displaced across nine discrete steps. The bundle&#39;s position was tracked on a dual-photodiode system, and its receptor potential was simultaneously measured using a microelectrode whose output was passed through an amplifier in bridge mode. The electrode&#39;s tip resistance was 95 M&#937;&#160;and the bundle&#39;s resting membrane potential was -47 mV. (B) A plot of the bundle&#39;s receptor potential as a function of its displacement reveals a nonlinear relationship between the bundle&#39;s response and its position. Each point corresponds to the mean potential and mean displacement over a 2.5 ms time window, beginning 2.5 ms after the onset of mechanical stimulation. Each color represents a set of time series corresponding to the same displacement pulse. 
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<img alt="Figure 3" src="/files/ftp_upload/55380/55380fig3.jpg" /> 
Figure 3: Effect of Gentamicin on Spontaneous Hair Bundle Oscillation. The spontaneous motion of a hair bundle in a two-chamber preparation was recorded using a dual photodiode system. In the absence of iontophoretic release of gentamicin (0 nA), the hair bundle displays symmetric oscillations. As the magnitude of current passed through an iontophoretic pipette filled with 500 mM gentamicin sulfate grows (10 nA, 20 nA), the frequency of hair-bundle excursions falls in a dose-dependent manner and the bundle is offset towards its tall edge for longer periods of time. 
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<img alt="Figure 4" src="/files/ftp_upload/55380/55380fig4.jpg" /> 
Figure 4. Calculating a Hair Bundle&#39;s Instantaneous&#160;Stiffness. (A) A stimulus fiber (red) of stiffness KF is coupled to the kinociliary bulb (brown) of an individual hair bundle (yellow). Displacing the base of the fiber a known distance XF causes the bundle to move a distance XB. The difference between the displacements of the fiber and the bundle is proportional to the force exerted onto the bundle by the stimulus fiber, FF. Repeating this across a range of forces yields an instantaneous force-displacement relation (right), the slope of which corresponds to the hair bundle&#39;s instantaneous stiffness. (B) An individual bundle was subjected to force pulses of increasing magnitude and its displacement within the first 50 ms after the pulse onset was measured (blue points). Here the hair bundle displays a nonlinear instantaneous stiffness over a range of approximately 20 nm around its resting position. The red curve corresponds to a fit to the relation F = k*X - 60*z*(1/(1+exp(-z*(X-X0)/(kB*T))) + F0, in which F is the force applied to the bundle, X is the bundle&#39;s displacement, k = 790 &#177; 51 &#181;N&#8729;m-1 is the bundle&#39;s constant stiffness when all channels are either closed or open, z = 0.43 &#177; 0.04 pN is the force of a single gating spring, X0 = 2 &#177; 1.9 nm is the bundle&#39;s position at which 50% of its channels are open, kB is Boltzmann&#39;s constant, T is the temperature, and F0 = 11.7 &#177; 1.3 pN is an offset force. The fit possesses a coefficient of determination of 0.98. The stimulus fiber had a stiffness of 107 &#181;N&#8729;m-1. 
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<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td rowspan="2">Solute</td>
			<td rowspan="2">Formula Weight (g/mol)</td>
			<td colspan="2">Add to 1 L</td>
		</tr>
		<tr>
			<td>Artificial perilymph</td>
			<td>Artificial endolymph</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>58.4</td>
			<td>6.54 g</td>
			<td>0.117 g</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>0.149</td>
			<td>0.149 g</td>
			<td>8.62 g</td>
		</tr>
		<tr>
			<td>CaCl2 &#10625; 2H2O</td>
			<td>147</td>
			<td>2 mL of 1 M CaCl2 stock</td>
			<td>250 &#181;L of 1 M CaCl2 stock</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>238.3</td>
			<td>1.19 g</td>
			<td>1.19 g</td>
		</tr>
		<tr>
			<td>D-(+)-glucose</td>
			<td>180.2</td>
			<td>0.541 g</td>
			<td>0. 541 g</td>
		</tr>
	</tbody>
</table>
Table 1. Solutions for Dissection and Experimental Preparation. Displayed in this table are the recipes for artificial perilymph and artificial endolymph solutions used in dissection and in one- or two-chamber preparations. The solutions should be brought to pH 7.2 - 7.4 with about 2 mL of NaOH (perilymph) or 2 mL KOH (endolymph). The osmotic strength should read approximately 230 mmol&#183;kg-1 due to incomplete ionic dissociation.

Watch this Scientific Journal Video about Physiological Preparation of Hair Cells from the Sacculus of the American Bullfrog Rana catesbeiana at JoVE.com

Physiological Preparation of Hair Cells from the Sacculus of the American Bullfrog Rana catesbeiana

The American bullfrog&#39;s (Rana catesbeiana) sacculus permits direct examination of hair-cell physiology. Here the dissection and preparation of the bullfrog&#39;s sacculus for biophysical studies is described. We show representative experiments from these hair cells, including the calculation of a bundle&#39;s force-displacement relation and measurement of its unforced motion.

Physiological Preparation of Hair Cells from the Sacculus of the American Bullfrog (Rana catesbeiana)

The study of hearing and balance rests upon insights drawn from biophysical studies of model systems. One such model, the sacculus of the American ...

physiological-preparation-hair-cells-from-sacculus-american-bullfrog

Research

JoVE Journal

Neuroscience

16.7K Views.  The Rockefeller University. The American bullfrog's (Rana catesbeiana) sacculus permits direct examination of hair-cell physiology. Here the dissection and preparation of the bullfrog's sacculus for biophysical studies is described. We show representative experiments from these hair cells, including the calculation of a bundle's force-displacement relation and measurement of its unforced motion.

Video: Physiological Preparation of Hair Cells from the Sacculus of the American Bullfrog Rana catesbeiana

Postsynaptic Recordings at Afferent Dendrites Contacting Cochlear Inner Hair Cells: Monitoring Multivesicular Release at a Ribbon Synapse

The afferent synapse between the inner hair cell (IHC) and the auditory nerve fiber provides an electrophysiologically accessible site for recording the postsynaptic activity of a single ribbon synapse 1-4. Ribbon synapses of sensory cells release neurotransmitter continuously, the rate of which is modulated in response to graded changes in IHC membrane potential 5. Ribbon synapses have been shown to operate by multivesicular release, where multiple vesicles can be released simultaneously to evoke excitatory postsynaptic currents (EPSCs) of varying amplitudes 1, 4, 6-11. Neither the role of the presynaptic ribbon, nor the mechanism underlying multivesicular release is currently well understood.
The IHC is innervated by 10-20 auditory nerve fibers, and every fiber contacts the IHC with a unmyelinated single ending to form a single ribbon synapse. The small size of the afferent boutons contacting IHCs (approximately 1 &mu;m in diameter) enables recordings with exceptional temporal resolution to be made. Furthermore, the technique can be adapted to record from both pre- and postsynaptic cells simultaneously, allowing the transfer function at the synapse to be studied directly 2. This method therefore provides a means by which fundamental aspects of neurotransmission can be studied, from multivesicular release to the elusive function of the ribbon in sensory cells.

Whole-cell patch-clamp recordings from auditory nerve fiber dendrites at the inner hair cell ribbon synapse in the mammalian cochlea.

The afferent synapse between the inner hair cell (IHC) and the auditory nerve fiber provides an electrophysiologically accessible site for recording the ...

Isolation and Culture of Neural Crest Stem Cells from Human Hair Follicles

Hair follicles undergo lifelong growth and hair cycle is a well-controlled process involving stem cell proliferation and quiescence. Hair bulge is a well-characterized niche for adult stem cells1. This segment of the outer root sheath contains a number of different types of stem cells, including epithelial stem cells2, melanocyte stem cells3 and neural crest like stem cells4-7. Hair follicles represent an accessible and rich source for different types of human stem cells. We and others have isolated neural crest stem cells (NCSCs) from human fetal and adult hair follicles4,5. These human stem cells are label-retaining cells and are capable of self-renewal through asymmetric cell division in vitro. They express immature neural crest cell markers but not differentiation markers. Our expression profiling study showed that they share a similar gene expression pattern with murine skin immature neural crest cells. They exhibit clonal multipotency that can give rise to myogenic, melanocytic, and neuronal cell lineages after in vitro clonal single cell culture. Differentiated cells not only acquire lineage-specific markers but also demonstrate appropriate functions in ex vivo conditions. In addition, these NCSCs show differentiation potential toward mesenchymal lineages. Differentiated neuronal cells can persist in mouse brain and retain neuronal differentiation markers. It has been shown that hair follicle derived NCSCs can help nerve regrowth, and they improve motor function in mice transplanted with these stem cells following transecting spinal cord injury8. Furthermore, peripheral nerves have been repaired with stem cell grafts9, and implantation of skin-derived precursor cells adjacent to crushed sciatic nerves has resulted in remyelination10. Therefore, the hair follicle/skin derived NCSCs have already shown promising results for regenerative therapy in preclinical models.
Somatic cell reprogramming to induced pluripotent stem (iPS) cells has shown enormous potential for regenerative medicine. However, there are still many issues with iPS cells, particularly the long term effect of oncogene/virus integration and potential tumorigenicity of pluripotent stem cells have not been adequately addressed. There are still many hurdles to be overcome before iPS cells can be used for regenerative medicine. Whereas the adult stem cells are known to be safe and they have been used clinically for many years, such as bone marrow transplant. Many patients have already benefited from the treatment. Autologous adult stem cells are still preferred cells for transplantation. Therefore, the readily accessible and expandable adult stem cells in human skin/hair follicles are a valuable source for regenerative medicine.

This article presents a robust protocol for isolation and culture of neural crest stem cells from human hair follicles.

Hair follicles undergo lifelong growth and hair cycle is a well-controlled process involving stem cell proliferation and quiescence. Hair bulge is a ...

Tissue Preparation and Immunostaining of Mouse Sensory Nerve Fibers Innervating Skin and Limb Bones

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in animals and humans. These peripheral sensory nerve fibers express a plethora of receptors and ion channel proteins which detect and initiate specific sensory stimuli. Methods are available to characterize the electrical properties of peripheral sensory nerve fibers innervating the skin, which can also be utilized to identify the functional expression of specific ion channel proteins in these fibers. However, similar electrophysiological methods are not available (and are also difficult to develop) for the detection of the functional expression of receptors and ion channel proteins in peripheral sensory nerve fibers innervating other visceral organs, including the most challenging tissues such as bone. Moreover, such electrophysiological methods cannot be utilized to determine the expression of non-excitable proteins in peripheral sensory nerve fibers. Therefore, immunostaining of peripheral/visceral tissue samples for sensory nerve fivers provides the best possible way to determine the expression of specific proteins of interest in these nerve fibers. So far, most of the protein expression studies in sensory neurons have utilized immunostaining procedures in sensory ganglia, where the information is limited to the expression of specific proteins in the cell body of specific types or subsets of sensory neurons. Here we report detailed methods/protocols for the preparation of peripheral/visceral tissue samples for immunostaining of peripheral sensory nerve fibers. We specifically detail methods for the preparation of skin or plantar punch biopsy and bone (femur) sections from mice for immunostaining of peripheral sensory nerve fibers. These methods are not only key to the qualitative determination of protein expression in peripheral sensory neurons, but also provide a quantitative assay method for determining changes in protein expression levels in specific types or subsets of sensory fibers, as well as for determining the morphological and/or anatomical changes in the number and density of sensory fibers during various pathological states. Further, these methods are not confined to the staining of only sensory nerve fibers, but can also be used for staining any types of nerve fibers in the skin, bones and other visceral tissue.

Immunocytochemical identification of peripheral sensory nerve fiber subtypes (and detection of protein expression therein) are key to the understanding of molecular mechanisms underlying peripheral sensation. Here we describe methods for preparation of peripheral/visceral tissue samples, such as skin and limb bones, for specific immunostaining of peripheral sensory nerve fibers.

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in ...

Odorant-induced Responses Recorded from Olfactory Receptor Neurons using the Suction Pipette Technique

Animals sample the odorous environment around them through the chemosensory systems located in the nasal cavity. Chemosensory signals affect complex behaviors such as food choice, predator, conspecific and mate recognition and other socially relevant cues. Olfactory receptor neurons (ORNs) are located in the dorsal part of the nasal cavity embedded in the olfactory epithelium. These bipolar neurons send an axon to the olfactory bulb (see Fig. 1, Reisert &amp; Zhao1, originally published in the Journal of General Physiology) and extend a single dendrite to the epithelial border from where cilia radiate into the mucus that covers the olfactory epithelium. The cilia contain the signal transduction machinery that ultimately leads to excitatory current influx through the ciliary transduction channels, a cyclic nucleotide-gated (CNG) channel and a Ca2+-activated Cl- channel (Fig. 1). The ensuing depolarization triggers action potential generation at the cell body2-4.
In this video we describe the use of the "suction pipette technique" to record odorant-induced responses from ORNs. This method was originally developed to record from rod photoreceptors5 and a variant of this method can be found at jove.com modified to record from mouse cone photoreceptors6. The suction pipette technique was later adapted to also record from ORNs7,8. Briefly, following dissociation of the olfactory epithelium and cell isolation, the entire cell body of an ORN is sucked into the tip of a recording pipette. The dendrite and the cilia remain exposed to the bath solution and thus accessible to solution changes to enable e.g. odorant or pharmacological blocker application. In this configuration, no access to the intracellular environment is gained (no whole-cell voltage clamp) and the intracellular voltage remains free to vary. This allows the simultaneous recording of the slow receptor current that originates at the cilia and fast action potentials fired by the cell body9. The difference in kinetics between these two signals allows them to be separated using different filter settings. This technique can be used on any wild type or knockout mouse or to record selectively from ORNs that also express GFP to label specific subsets of ORNs, e.g. expressing a given odorant receptor or ion channel.

Olfactory receptor neurons (ORNs) convert odor signals first into a receptor current that in turn triggers action potentials that are conveyed to second order neurons in the olfactory bulb. Here we describe the suction pipette technique to record simultaneously the odorant-induced receptor current and action potentials from mouse ORNs.

Animals sample the odorous environment around them through the chemosensory systems located in the nasal cavity. Chemosensory signals affect complex ...

The Specification of Telencephalic Glutamatergic Neurons from Human Pluripotent Stem Cells

Here, a stepwise procedure for efficiently generating telencephalic glutamatergic neurons from human pluripotent stem cells (PSCs) has been described. The differentiation process is initiated by breaking the human PSCs into clumps which round up to form aggregates when the cells are placed in a suspension culture. The aggregates are then grown in hESC medium from days 1-4 to allow for spontaneous differentiation. During this time, the cells have the capacity to become any of the three germ layers. From days 5-8, the cells are placed in a neural induction medium to push them into the neural lineage. Around day 8, the cells are allowed to attach onto 6 well plates and differentiate during which time the neuroepithelial cells form. These neuroepithelial cells can be isolated at day 17. The cells can then be kept as neurospheres until they are ready to be plated onto coverslips. Using a basic medium without any caudalizing factors, neuroepithelial cells are specified into telencephalic precursors, which can then be further differentiated into dorsal telencephalic progenitors and glutamatergic neurons efficiently. Overall, our system provides a tool to generate human glutamatergic neurons for researchers to study the development of these neurons and the diseases which affect them.

This procedure yields telencephalic neurons by going through checkpoints which are similar to those observed during human development. The cells are allowed to spontaneously differentiate, are exposed to factors which push them towards the neural lineage, are isolated, and are plated onto coverslips to allow for terminal differentiation and maturation.

Here, a stepwise procedure for efficiently generating telencephalic glutamatergic neurons from human pluripotent stem cells (PSCs) has been described. The ...

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

Lineage-reprogramming of Pericyte-derived Cells of the Adult Human Brain into Induced Neurons

Direct lineage-reprogramming of non-neuronal cells into induced neurons (iNs) may provide insights into the molecular mechanisms underlying neurogenesis and enable new strategies for in vitro modeling or repairing the diseased brain. Identifying brain-resident non-neuronal cell types amenable to direct conversion into iNs might allow for launching such an approach in situ, i.e. within the damaged brain tissue. Here we describe a protocol developed in the attempt of identifying cells derived from the adult human brain that fulfill this premise. This protocol involves: (1) the culturing of human cells from the cerebral cortex obtained from adult human brain biopsies; (2) the in vitro expansion (approximately requiring 2-4 weeks) and characterization of the culture by immunocytochemistry and flow cytometry; (3) the enrichment by fluorescence-activated cell sorting (FACS) using anti-PDGF receptor-&#946; and anti-CD146 antibodies; (4) the retrovirus-mediated transduction with the neurogenic transcription factors sox2 and ascl1; (5) and finally the characterization of the resultant pericyte-derived induced neurons (PdiNs) by immunocytochemistry (14 days to 8 weeks following retroviral transduction). At this stage, iNs can be probed for their electrical properties by patch-clamp recording. This protocol provides a highly reproducible procedure for the in vitro lineage conversion of brain-resident pericytes into functional human iNs.

Targeting brain-resident cells for direct lineage-reprogramming offers new perspectives for brain repair. Here we describe a protocol of how to prepare cultures enriched for brain-resident pericytes from the adult human cerebral cortex and convert these into induced neurons by retrovirus-mediated expression of the transcription factors Sox2 and Ascl1.

Direct lineage-reprogramming of non-neuronal cells into induced neurons (iNs) may provide insights into the molecular mechanisms underlying neurogenesis ...

Feeder-free Derivation of Neural Crest Progenitor Cells from Human Pluripotent Stem Cells

Human pluripotent stem cells (hPSCs) have great potential for studying human embryonic development, for modeling human diseases in the dish and as a source of transplantable cells for regenerative applications after disease or accidents. Neural crest (NC) cells are the precursors for a large variety of adult somatic cells, such as cells from the peripheral nervous system and glia, melanocytes and mesenchymal cells. They are a valuable source of cells to study aspects of human embryonic development, including cell fate specification and migration. Further differentiation of NC progenitor cells into terminally differentiated cell types offers the possibility to model human diseases in vitro, investigate disease mechanisms and generate cells for regenerative medicine. This article presents the adaptation of a currently available in vitro differentiation protocol for the derivation of NC cells from hPSCs. This new protocol requires 18 days of differentiation, is feeder-free, easily scalable and highly reproducible among human embryonic stem cell (hESC) lines as well as human induced pluripotent stem cell (hiPSC) lines. Both old and new protocols yield NC cells of equal identity.

Neural crest (NC) cells derived from human pluripotent stem cells (hPSC) have great potential for modeling human development and disease and for cell replacement therapies. Here, a feeder-free adaptation of the currently widely used in vitro differentiation protocol for the derivation of NC cells from hPSCs is presented.

Human pluripotent stem cells (hPSCs) have great potential for studying human embryonic development, for modeling human diseases in the dish and as a ...

In-depth Physiological Analysis of Defined Cell Populations in Acute Tissue Slices of the Mouse Vomeronasal Organ

In most mammals, the vomeronasal organ (VNO) is a chemosensory structure that detects both hetero- and conspecific social cues. Vomeronasal sensory neurons (VSNs) express a specific type of G protein-coupled receptor (GPCR) from at least three different chemoreceptor gene families allowing sensitive and specific detection of chemosensory cues. These families comprise the V1r and V2r gene families as well as the formyl peptide receptor (FPR)-related sequence (Fpr-rs) family of putative chemoreceptor genes. In order to understand the physiology of vomeronasal receptor-ligand interactions and downstream signaling, it is essential to identify the biophysical properties inherent to each specific class of VSNs.
The physiological approach described here allows identification and in-depth analysis of a defined population of sensory neurons using a transgenic mouse line (Fpr-rs3-i-Venus). The use of this protocol, however, is not restricted to this specific line and thus can easily be extended to other genetically modified lines or wild type animals.

Here, we describe a physiological approach that allows identification and in-depth analysis of a defined population of sensory neurons in acute coronal tissue slices of the mouse vomeronasal organ using whole-cell patch-clamp recordings.

In most mammals, the vomeronasal organ (VNO) is a chemosensory structure that detects both hetero- and conspecific social cues. Vomeronasal sensory ...

Use of Synaptic Zinc Histochemistry to Reveal Different Regions and Laminae in the Developing and Adult Brain

Characterization of anatomical and functional brain organization and development requires accurate identification of distinct neural circuits and regions in the immature and adult brain. Here we describe a zinc histochemical staining procedure that reveals differences in staining patterns among different layers and brain regions. Others have utilized this procedure not only to reveal the distribution of zinc-containing neurons and circuits in the brain, but also to successfully delineate areal and laminar boundaries in the developing and adult brain in several species. Here we illustrate this staining procedure with images from developing and adult ferret brains. We reveal a zinc-staining pattern that serves as an anatomical marker of areas and layers, and can be reliably used to distinguish visual cortical areas in the developing and adult visual cortex. The main goal of this protocol is to present a histochemical method that allows the accurate identification of layers and regions in the developing and adult brain where other methods fail to do so. Secondarily, in conjunction with densitometric image analysis, this method allows one to assess the distribution of synaptic zinc to reveal potential changes throughout development. This protocol describes in detail the reagents, tools, and steps necessary to successively stain frozen brain sections. Although this protocol is described using ferret brain tissue, it can easily be adapted for use in rodents, cats, or monkeys as well as in other brain regions.

We describe a histochemical procedure that reveals characteristic laminar and areal zinc staining patterns in different brain regions. The zinc-staining pattern may be used in conjunction with other anatomical markers to reliably distinguish layers and regions in the developing and adult brain.

Characterization of anatomical and functional brain organization and development requires accurate identification of distinct neural circuits and regions ...