We thank Prof. Yuri Korchev (Imperial College, UK) for the long-term support and advice throughout all stages of the project. We also thank Drs. Pavel Novak and Andrew Shevchuk (Imperial College, UK) as well as Oleg Belov (National Research Centre for Audiology, Russia) for their help with software development. The study was supported by NIDCD/NIH (R01 DC008861 and R01 DC014658 to G.I.F.).

The study was performed in accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All animal procedures were approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Kentucky (protocol 00903M2005).
1.&#160;Manufacturing and testing the nanopipettes
<ol>
	<li>Create a program in the micropipette puller to obtain pipettes with a resistance between 200 and 400 M&#8486;, which corresponds to inner tip diameters of approximately 50-70 nm. The parameters will depend on the micropipette puller. To obtain short pipettes with non-flexible fine tips, check in the operational manual of the puller.</li>
	<li>Use borosilicate glass capillaries with outer/inner diameters of 1/0.58 mm and an inner filament to facilitate filling. The length of the pipette is crucial because it determines the frequency of the lateral mechanical resonance of the pipette. The longer is the pipette, the lower is the resonant frequency and the harder is to avoid this resonance. 
	NOTE: The user should try to manufacture the shortest pipette the holder can accept. The length of nanopipettes in this experiment is usually 15-25 mm (Figure 2A).</li>
	<li>Fill the nanopipette up to its middle point (Figure 2A) with a bath solution, either Leibovitz&#39;s L-15 or with Hank&#39;s Balanced Salt Solution (HBSS) supplemented with 20 mM D-glucose (to adjust osmolarity). To avoid potential artefacts, use the same solution that will be used in the bath for recordings.</li>
	<li>Using an optical microscope with a magnification of 10x, check for any bubbles at the tip of the pipette (Figure 2B). The bubbles would prevent the current flow. It is harder to remove bubbles in the pipettes that have been pulled several hours before the experiment. Therefore, it is recommended to pull new pipettes with every experiment.</li>
	<li>Once the pipette is free of bubbles, mount it in the HPICM pipette holder (Figure 2C).</li>
	<li>Place the sample (tissue or calibration standard) on the custom-built&#160;chamber and add 4 mL of the above-mentioned bath solution.</li>
	<li>Place the&#160;custom-built&#160;chamber on the HPICM stage and introduce the ground electrode in the solution.</li>
	<li>Make sure that the voltage being applied to the pipette by the patch clamp amplifier is zero.</li>
	<li>Move the pipette in Z until it touches the liquid.</li>
	<li>Set the amplifier offset to zero and then add +100 mV to check the pipette current.</li>
	<li>Calculate the resistance and the diameter of the pipette, based on Ohm&#39;s law: 
	R = V/I 
	where R is the resistance (MOhm), V is the voltage applied to the nanopipette (mV) and I is the current flowing through the nanopipette (nA).
	<ol>
		<li>Calculate the inner diameter of the pipette as described elsewhere according to the following formula12: 
		IDTip&#160;= 1000/ &#8730;R 
		NOTE: The ideal resistance value is between 200 and 400 M&#8486;. Pipettes with a resistance higher than 400 M&#8486; might lead to an unstable current due to their small size (&#60; 50 nm inner diameter). On the contrary, pipettes with resistances smaller than 200 M&#8486; are too large (&#62; 70 nm inner diameter) and would not resolve small features. Its recommended to start imaging with the pipettes of 200 M&#8486; resistance, as they are easier to manufacture and tend to provide less electrical noise.</li>
	</ol>
	</li>
</ol>
2.&#160;Minimizing sample drifts and vibrations
NOTE: To decrease the mechanical noise in the system during imaging, mount the samples on the custom-built chambers that utilize thick glass slides (~1.2 mm):
<ol>
	<li>Remove the glass portion off a 50 mm glass-bottom dish, leaving the plastic walls intact.</li>
	<li>Glue the plastic portion of the cell culture dish on top of the thick glass slide with silicon glue.</li>
	<li>Mount the calibration sample in the middle of the chamber (on top of the glass slide) using either silicon glue or thin double-sided tape.</li>
	<li>Firmly secure the chamber onto the HPICM stage using double-sided tape.</li>
	<li>During imaging, close the Faraday cage and cover it with a blanket to minimize electrical interference and thermal drift, correspondingly.</li>
</ol>
3.Testing the resolution with AFM calibration standards 
NOTE: It is strongly recommended to image AFM standards (see Table of Materials) before imaging live cells in order to troubleshoot the system and test its resolution in the X-Z-Y axes. The calibration standards have silicon dioxide pillars and holes of different shapes but fixed heights/depths (i.e., 20 or 100 nm) on a 5 x 5 mm silicon chip. Starting with the 100 nm calibration standard is recommended, to guarantee that the Z resolution is below 100 nm. After achieving a successful high-resolution image of the pillars or holes in this calibration sample (Figure 3A), move to the 20 nm standard. If the imaging of the latter standard is successful (Figure 3B), the resolution in the Z axis is guaranteed to be below 20 nm and appropriate for the imaging of the hair cell stereocilia bundles12. The following steps are used to image both calibration standards.
<ol>
	<li>Attach the calibration standard to the chamber with silicon glue.</li>
	<li>Add 4 mL of HBSS to the chamber to cover the calibration sample. Then, secure the chamber to the XY stage of the HPICM setup using double-sided tape.</li>
	<li>Clamp the magnetic holder of the ground electrode to the stage near the chamber and immerse the electrode into the bath solution (Figure 1B).</li>
	<li>Mount the nanopipette into the holder, immerse it into the bath solution, and set its current to ~1 nA following the steps described in Section 1.</li>
	<li>Position the nanopipette approximately above the center of the calibration standard using a course patch-clamp manipulator. Visual inspection is usually enough for this positioning, since the area covered by silicon dioxide structures is relatively large (1 x 1 mm). In contrast to the organ of Corti explants (see below), calibration standard is non-transparent and, therefore, a more precise positioning guided by optical imaging is not possible for this sample.</li>
	<li>Increase the setpoint while monitoring the signal from the sensor of the Z piezo actuator on an oscilloscope in real time. After establishing a stable repeatable Z approach cycle (as in Figure 1C, bottom), decrease the setpoint to the value that is just above the point of instability. This procedure would ensure the optimal setpoint for this particular nanopipette.</li>
	<li>Move the pipette down at a speed of ~5 &#181;m/s with a patch clamp micromanipulator until it reaches the sample. At this moment, the bottom level of the real-time Z positioning signal (Figure 1C) will increase, indicating that the nanopipette is withdrawn due to &#34;sensing&#34; the sample surface. Any further movement of the nanopipette will result in further positive shift of Z-positioning signal. 
	NOTE: Be careful not to exceed the upper limit of Z piezo actuator movement.</li>
	<li>Start imaging at low resolution (see Table 1). Due to uneven mounting of the AFM standard, the highest point of the area of interest may be unknown. Therefore, set the amplitude of pipette retraction (hop amplitude) to at least 200-500 nm.
	<ol>
		<li>Once the highest point of the sample in the imaging area is identified, decrease the hop amplitude. A smaller hop amplitude allows faster scanning, which is preferred for high resolution imaging due to diminished effects of the drifts and decreased vibration.</li>
	</ol>
	</li>
	<li>Before moving the pipette to a new X-Y location, retract it about 200 &#181;m in the Z axis to prevent any undesired collisions with the sample. 
	NOTE: In cases when the nanopipette is not aligned with the center of the calibration standard, it is possible that the scanning starts just off the area of surface features.</li>
	<li>After the area of interest is found, start imaging at a higher resolution (see&#160;Table 1).</li>
</ol>
4.&#160;Making custom-built chambers to secure the cochlear explants
NOTE: Mount the cochlear explants in the chambers with custom-built clamping systems that utilize either flexible glass pipettes (step 4.1) (Figure 4A) or dental floss (step 4.2) (Figure 4B). The glass pipette chamber could be sterilized and used for the cultured organs of Corti, while dental floss chamber provides a more secure holding of the sample and a control over stereocilia bundle orientation during mounting. These custom-built chambers need to be prepared in advance, but they can be cleaned and re-used in several imaging sessions.
<ol>
	<li>Make a chamber using flexible glass pipettes
	<ol>
		<li>Pull two thin and flexible glass fibers from glass capillaries using a pipette puller. Our pulled glass fibers typically measure 1 to 2 cm in length and are fairly flexible.</li>
		<li>Place a small drop of the silicone elastomer on top of a glass coverslip. Use coverslips of 2 cm in diameter.</li>
		<li>Place the ends of two glass fibers on the silicone drop and arrange the fibers to have a small degree of separation between them (Figure 4A).</li>
		<li>Place the coverslip on a hot plate to quickly cure the elastomer (1 to 3 min).</li>
		<li>Glue the coverslip onto the glass bottom of the chamber described in Section 2 using a small amount (1-3 &#181;L) of silicone elastomer and allowing it to cure overnight.</li>
	</ol>
	</li>
	<li>Making a chamber using dental floss:
	<ol>
		<li>Remove the glass portion of a 50 mm glass-bottom dish, leaving the plastic walls intact. Then glue the plastic portion of the cell culture dish on top of a 1.2 mm thick glass slide with silicon glue.</li>
		<li>Mount one plastic coverslip (6.5 x 6.5 mm) with the same glue to the center of the chamber. Then repeat the process with another coverslip on top of the previous one.</li>
		<li>Mount two small tungsten or gold-plated wires (12 mm of length and ~0.5 mm in diameter) with silicon glue, each one at opposites sides of the cover slides. Glue them far enough (&#62; 10-15 mm) from the cover slides (Figure 4B).</li>
		<li>Separate two dental floss strands and place them on top of the cover slides and secure them to the wires by making a knot. Leave a small gap between both strands (Figure 4B, short arrows).</li>
	</ol>
	</li>
	<li>Clean the chambers after every use
	<ol>
		<li>Gently remove the tissue from the chamber using fine tweezers and lightly scrape any tissue residues left behind.</li>
		<li>Rinse the chamber first with 70% ethanol and then with distilled water.</li>
		<li>Repeat the rinse cycle if needed.</li>
		<li>Place the chamber upside down on a filter paper to let it dry until next experiment. The chambers do not need to be sterilized, unless culturing of the organ of Corti is planned after the imaging.</li>
	</ol>
	</li>
</ol>
5.&#160;Dissecting the rodent organ of Corti
<ol>
	<li>Perform dissection of the young postnatal cochlear explants as described in detail elsewhere13.</li>
	<li>For HPICM imaging, dissect the organ of Corti from mice between postnatal days 3 and 6 (P3-6), and from rats between postnatal days 3 and 8 (P3-8). 
	NOTE: Older hair cells are more susceptible to the damage during dissection and, therefore, cannot be used for hours-long time lapse HPICM imaging.</li>
	<li>Do not forget to remove the tectorial membrane before HPICM imaging.</li>
	<li>Immediately after dissection, secure the tissue in one of the chambers described in Section 4, either placing it under the flexible glass pipettes or under the two dental floss strands (Figure 4). Pre-fill these chambers with 4 mL of the room temperature bath solution (to minimize bubble formation).</li>
</ol>
6.&#160;Imaging of the auditory hair cells
<ol>
	<li>Mount the chamber with a freshly isolated organ of Corti on the X-Y piezo stage using double-sided tape and make sure that it is firmly secured to minimize chamber drift in X and Y axes (Figure 1B).</li>
	<li>Follow steps in Section 1 to place a new nanopipette and check for the correct pipette resistance.</li>
	<li>Using patch clamp micromanipulator, position nanopipette over the hair cell region, while observing the organ of Corti explant in an inverted microscope.</li>
	<li>Check if the system is stable with a setpoint of 0.5% or lower by recording the real-time current and Z positioning signal on the oscilloscope (as in Figure 1C). If Z signal is not stable, try to decrease the cutoff frequency of the low-pass filter of the patch clamp amplifier. However, it cannot be lower than the response time of Z piezo actuator (to avoid pipette collision with the sample due to delayed current readings). 
	NOTE: In practice, 5 kHz setting of this filter is found to be optimal. It is better to replace the nanopipette, if Z-signal is still unstable.</li>
	<li>Once a stable recording is achieved, determine the optimal setpoint and approach the sample with HPICM pipette as described in the steps 3.6-3.7 above.</li>
	<li>First, perform low resolution imaging (see&#160;Table 1), using a hop amplitude of at least 6 to 8 &#181;m. To image tall structures such as the hair cell stereocilia bundles, make sure that the hop amplitude is enough to avoid collision with these structures. 
	NOTE: If the hop amplitude is not enough, the pipette will not be able to jump over a stereocilium and there will be an imminent collision. The collision of the HPICM probe with a stereocilium may damage the hair bundle. Therefore, in cases when the height of the stereocilia bundle is uncertain, use a bigger hop amplitude.</li>
	<li>Get familiar with the topography of the organ of Corti by first performing and/or studying images obtained with scanning electron microscopy (Figure 5). 
	NOTE: If the HPICM image is uniform with the heights smaller than 1 &#181;m in every imaging point, the pipette is likely scanning the glass bottom and not the tissue. Alternatively, the pipette may &#34;land&#34; on a different region of the cochlear explant, away from the hair cells.</li>
	<li>If the pipette needs to be moved to a new X-Y location, retract it about 500 nm to avoid collisions with any tall features within the tissue. Repeat low-resolution HPICM imaging until the region of interest with the hair cells is found.</li>
	<li>After the region of interest is found, start imaging at a higher resolution (see Table 1). Try to spend less than 15 min when imaging a whole hair bundle. 
	NOTE: The hair cell bundles in the live tissue are not still but may change their orientation, for example, due to shape changes in the underlying supporting cells. Therefore, the images may exhibit movement artefacts if the image acquisition is too slow.</li>
	<li>Once again, determine the tallest features in the low-resolution images before the decreasing the hop amplitude for high-resolution imaging. For a region of interest covering entire hair cell bundle, reduce the hop amplitude to 4-5 &#181;m, while for a relatively small and &#34;flat&#34; region within the bundle (e.g., 2 x 2 &#181;m) reduce the hop amplitude even further, down to less than 1 &#181;m, thereby increasing the speed and resolution of imaging.</li>
</ol>
7.&#160;Image processing
NOTE: Imaging artefacts are common in HPICM imaging. Some of them can be corrected by image acquisition parameters, while others require post-processing either with a specialized SICM viewer or with more general data-processing programs like ImageJ or MatLab. Here we describe the most common artefacts and how we fix them with SICM viewer.
<ol>
	<li>Perform slope correction 
	&#8203;NOTE: It is not obvious for a beginner, but human eye cannot resolve sub-micrometer size features at the surface of a cell, if the imaging area has an overall slope of equal or larger size (Figure 3A,B, left). Therefore, it is necessary to determine the average slope of an imaged area (by fitting HPICM 3D image data to a single plane) and subtract it from the HPICM image (Figure 3A,B, middle).
	<ol>
		<li>Click Open to open an image with SICM viewer.</li>
		<li>Select the Image Correction tab.</li>
		<li>Select the Correct Slope tab.</li>
		<li>Press the Correct Slope button for an automated slope correction.</li>
	</ol>
	</li>
	<li>Perform line alignment 
	&#8203;NOTE: As mentioned before, mechanical and/or thermal drifts as well as cell movement artefacts represent a significant problem in HPICM imaging. A small drift with a speed of less than a micrometer per minute is usually not noticeable in a regular patch clamp setup. Yet, it could produce artefacts of several tens of nanometers in HPICM imaging, which is significantly larger than the resolution of HPICM. Therefore, it is not uncommon to encounter sudden jumps in Z axis between two neighboring HPICM scan lines during imaging. This could be corrected by analyzing differences between starting (and/or ending) Z-values in these neighboring scan lines.
	<ol>
		<li>Click Open to open an image with SICM viewer.</li>
		<li>Select the Image Correction tab.</li>
		<li>Select the Correct Slope tab.</li>
		<li>Choose the width of lines to be aligned. Press on the button ButtonDestripeLineFit&#160;for an automated line alignment correction.</li>
	</ol>
	</li>
	<li>Perform noise reduction 
	&#8203;NOTE: While obtaining images with HPICM, small fluctuations in the nanopipette current can lead to the nanopipette stopping far from the surface of the sample, especially with low setpoints. It results in the appearance of small white dots in the image. In order to correct this imaging artefact, it is necessary to identify the imaging points with the Z-value significantly larger than that of the neighbors and replace this value with an average of the neighbors. This is be done by an adjustable median filter.
	<ol>
		<li>Click Open to open an image with SICM viewer.</li>
		<li>Select the Image Processing tab.</li>
		<li>Select the Noise reduction tab.</li>
		<li>Set the Threshold filter (&#181;m) for the pixels to be removed.</li>
	</ol>
	</li>
</ol>

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B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+scanning+ion-conductance+microscope.">The scanning ion-conductance microscope.</a> Science. 243 (4891), 641-643 (1989).</li><li>Korchev, Y. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Specialized+scanning+ion-conductance+microscope+for+imaging+of+living+cells.">Specialized scanning ion-conductance microscope for imaging of living cells.</a> Journal of Microscopy. 188 (Pt 1), 17-23 (1997).</li><li>Shevchuk, A. I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+proteins+in+membranes+of+living+cells+by+high-resolution+scanning+ion+conductance+microscopy.">Imaging proteins in membranes of living cells by high-resolution scanning ion conductance microscopy.</a> Angewandte Chemie (International ed in English. 45 (14), 2212-2216 (2006).</li><li>Novak, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nanoscale+live-cell+imaging+using+hopping+probe+ion+conductance+microscopy.">Nanoscale live-cell imaging using hopping probe ion conductance microscopy.</a> Nature Methods. 6 (4), 279-281 (2009).</li><li>V&#233;lez-Ortega, A. C., Frolenkov, G. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualization+of+live+cochlear+stereocilia+at+a+nanoscale+resolution+using+hopping+probe+ion+conductance+microscopy.">Visualization of live cochlear stereocilia at a nanoscale resolution using hopping probe ion conductance microscopy.</a> Methods in Molecular Biology. 1427, 203-221 (2016).</li><li>Gu, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=High-resolution+scanning+patch-clamp:+new+insights+into+cell+function.">High-resolution scanning patch-clamp: new insights into cell function.</a> FASEB Journal Official Publication of the Federation of American Societies for Experimental Biology. 16 (7), 748-750 (2002).</li><li>Frolenkov, G. I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-channel+recordings+from+the+apical+surface+of+outer+hair+cells+with+a+scanning+ion+conductance+probe.">Single-channel recordings from the apical surface of outer hair cells with a scanning ion conductance probe.</a> Association for Research in Otolaryngology. Abs. 444, (2004).</li><li>S&#225;nchez, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Noncontact+measurement+of+the+local+mechanical+properties+of+living+cells+using+pressure+applied+via+a+pipette.">Noncontact measurement of the local mechanical properties of living cells using pressure applied via a pipette.</a> Biophysical Journal. 95 (6), 3017-3027 (2008).</li><li>Korchev, Y. E., Negulyaev, Y. A., Edwards, C. R., Vodyanoy, I., Lab, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+localization+of+single+active+ion+channels+on+the+surface+of+a+living+cell.">Functional localization of single active ion channels on the surface of a living cell.</a> Nature Cell Biology. 2 (9), 616-619 (2000).</li><li>Shevchuk, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Angular+approach+scanning+ion+conductance+microscopy.">Angular approach scanning ion conductance microscopy.</a> Biophysical Journal. 110 (10), 2252-2265 (2016).</li><li>Furness, D. N., Katori, Y., Nirmal Kumar, B., Hackney, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+dimensions+and+structural+attachments+of+tip+links+in+mammalian+cochlear+hair+cells+and+the+effects+of+exposure+to+different+levels+of+extracellular+calcium.">The dimensions and structural attachments of tip links in mammalian cochlear hair cells and the effects of exposure to different levels of extracellular calcium.</a> Neuroscience. 154 (1), 10-21 (2008).</li></ol>

The authors have no competing interests.

To obtain successful HPICM images, users need to establish a low noise and low vibration system and manufacture appropriate pipettes. We strongly recommend the use of AFM calibration standards to test the stability of the system before attempting to perform any live cell imaging. Once the resolution of the system is tested, users can consider imaging fixed organ of Corti samples to get familiar with the imaging settings before attempting any live cell imaging.
The optimal setpoint for imaging ...

Despite the fact that stereocilia bundles in the auditory hair cells are big enough to be visualized by optical microscopy and deflected in live cells in a patch clamp experiment, the essential structural components of the transduction machinery such as tip links could be imaged only with the electron microscopy in dead cells. In the mammalian auditory hair cells, the transduction machinery is located at the lower ends of the tip links, i.e., at the tips of the shorter row stereocilia1 and regulated locally through the signaling at the tips of stereocilia2,3. Yet, label-free imaging of the surface structures at this location in live hair cells is not possible due to the small sizes of stereocilia.
The mammalian cochlea has two types of auditory sensory cells: inner and outer hair cells. In the inner hair cells, stereocilia are longer and thicker compared to those in the outer hair cells4. The first and second row of stereocilia have a diameter of 300-500 nm in mouse or rat inner hair cells. Due to the diffraction of light, the maximum resolution achievable with a label-free optical microscopy is approximately 200 nm. Hence, visualization of individual stereocilia within first and second rows of the inner hair cell bundle is relatively easy&#160;with optical microscopy. In contrast, shorter row stereocilia in the inner hair cells and all stereocilia of the outer hair cells have diameters around 100-200 nm and cannot be visualized with optical microscopy5. Despite recent progress in super-resolution imaging, this fundamental limitation persists in any optical label-free imaging. All current commercially available super-resolution techniques do require some sort of fluorescent molecules6, which limits their applications. In addition to limitations due to the need for specific fluorescently tagged molecules, exposure to intense light irradiation has been shown to induce cellular damage and might influence cellular processes, which is a great disadvantage when studying live cells7.
Our current knowledge of the ultrastructural details of hair cell stereocilia bundles has been obtained mostly with various electron microscopy (EM) techniques, such as scanning electron microscopy (SEM), transmission electron microscopy (TEM), freeze-fracture EM, and recently with 3D techniques such as serial sectioning with focused ion beam or cryo-EM tomography8,9,10,11,12,13,14,15,16. Unfortunately, all these EM techniques require chemical or cryofixation of the sample. Depending on the time scale of the phenomena, this requirement makes a study of the dynamic processes at the tips of stereocilia either impossible or very labor intensive.
Limited efforts have been made to image hair bundles of live hair cells with atomic force microscopy (AFM)17,18. Since AFM operates in physiological solutions, it could, in theory, visualize dynamic changes in the stereocilia bundles of live hair cells over time. The problem lies in the principles of high-resolution AFM, which implies certain physical contact between the AFM probe and the sample, even in the least damaging &#34;tapping&#34; mode19. When the AFM probe encounters a stereocilium, it usually crashes against it, damaging the structure of the hair bundle. As a result, this technique is not suitable for visualizing live, or even fixed, hair cells bundles17,18. The problem may be partially alleviated by using a large ball-shaped AFM probe that imposes only hydrodynamic forces to the surface of the sample20. However, even though such a probe is ideally suited to test mechanical properties of the sample21, it provides only a sub-micrometer resolution when imaging the organ of Corti22 and still applies to the sample a force that may be substantial for the highly sensitive stereocilia bundles.
Scanning ion conductance microscopy (SICM) is a version of scanning probe microscopy that uses a glass pipette probe filled with a conductive solution23. SICM detects the surface when the pipette approaches the cell and the electric current through the pipette decreases. Since this is happening well before touching the cell, the SICM is ideally suited for non-contact imaging of live cells in physiological solution24. The best resolution of SICM is on the order of single nanometers, which allows resolving individual protein complexes at the plasma membrane of a living cell25. However, similar to other scanning probe techniques, SICM is able to image only relatively flat surfaces. We overcame this limitation by inventing hopping probe ion conductance microscope (HPICM)26, in which the nanopipette approaches the sample at each imaging point (Figure 1A). Using HPICM, we were able to image stereocilia bundles in live auditory hair cells with nanoscale resolution27.
Another fundamental advantage of this technique is that HPICM is not only an imaging tool. In contrast to other scanning probe techniques, the HPICM/SICM probe is an electrode widely used in cell physiology for electrical recordings and local delivery of various stimuli. Ion channel activity does not usually interfere with HPICM imaging, because the total current through the HPICM probe is several orders of magnitude larger than the extracellular current generated by the largest ion channels25. However, HPICM allows precise positioning of the nanopipette over a structure of interest and subsequent single-channel patch-clamp recording from this structure28. This is how we obtained the first preliminary recordings of single channel activity at the tips of the outer hair cell stereocilia29. It is worth mentioning that even a large current through the nanopipette cannot produce significant changes of the potential across the plasma membrane due to enormous electrical shunt of the extracellular medium. However, individual ion channels can be activated mechanically by flow of liquid through the nanopipette30 or chemically by local application of an agonist31.
In HPICM, the image is generated when a nanopipette sequentially approaches the sample at one point, retracts, and then moves in lateral direction to repeat the approach (Figure 1A). A patch clamp amplifier constantly applies voltage to an AgCl wire in the pipette (Figure 1B) to generate a current of ~1 nA in the bath solution. The value of this current when the pipette is away from the surface of the cell is determined as a reference current (Iref, Figure 1C). Then, the pipette moves in the Z axis to approach the sample until the current is reduced by an amount predefined by the user (setpoint), usually 0.2%-1% of the Iref (Figure 1C, top trace). The system then saves Z value at this moment as the height of the sample, together with X and Y coordinates of this imaging point. Then, the pipette is retracted away from the surface (Figure 1C, bottom trace) at a speed defined by the user, usually 700-900 nm/ms. After retraction, the pipette (or, in our case, the sample - see Figure 1B) is moved laterally to the next imaging point, a new reference current value is obtained, and the pipette once again approaches the sample, repeating the process. The X-Y movement of the pipette is preferred in an upright microscope setup that is typically used for recordings of the hair cell mechanotransduction currents. In this setting, the HPICM probe approaches the hair cell bundles not from the top but at an angle32. However, the best resolution of HPICM imaging is achieved in an inverted microscope setup (Figure 1A,B), where the movement of the sample in X-Y directions is de-coupled from Z-movement of the nanopipette, thereby eliminating potential mechanical artefacts.
Using HPICM, we obtained topographic images of mouse and rat inner and outer hair cell stereocilia bundles, and even visualized the links between the stereocilia that are about 5 nm in diameter26,27. The success of hair cell bundle imaging with this technique relies on several factors. First, the noise (variance) of the nanopipette current should be as small as possible to allow the lowest possible setpoint for HPICM imaging. A low setpoint allows HPICM probe to &#8220;sense&#8221; stereocilia surface at a larger distance and at any angle to the probe approach and, surprisingly, improves X-Y resolution of HPICM imaging (see Discussion). Second, the vibrations and drifts in the system should be decreased to less than 10 nm, since they contribute directly to the imaging artefacts. Finally, even though the HPICM probe and the specimen stage are moved in Z and X-Y axes by the calibrated feedback-controlled piezo actuators that have an accuracy of a single nanometer or better, the diameter of the nanopipette tip determines the spread of the current (sensing volume) and hence the resolution (Figure 1A). Therefore, before imaging live hair cells, it is vital to pull the adequate pipettes, reach the desired resolution with calibration samples, and achieve low noise in the recording system.
For at least a couple of decades, the SICM technique has not been commercially available and it has been developing by only few labs in the world with the leading lab of Prof. Korchev in the Imperial College (UK). Recently, several SICM systems became commercially available (see Table of Materials), all of which are based on the original HPICM principles. However, imaging stereocilia bundles in the hair cells requires several custom modifications that are technically challenging (or even impossible) in the closed &#8220;ready-to-go&#8221; systems. Therefore, some component integration is needed. Since HPICM setup represents a patch clamp rig with more stringent vibration and drift requirements and a piezo-driven movement of the HPICM probe and the sample (Figure 1D), this integration is relatively easy for any researcher, who is proficient in patch clamping. However, a scientist without proper background would definitely need some training in electrophysiology first. Despite remaining challenges such as increasing the speed of imaging (see Discussion), we have been able to image stereocilia bundles in live hair cells with nanoscale resolution without damaging them.&#160;
This paper presents a detailed protocol to perform successful HPICM imaging of the live auditory hair cell bundles in young postnatal rat or mouse cochlear explants using our custom system. The integrated components are listed in the Table of Materials. The paper also describes common problems that can be encountered and how to troubleshoot them.

<table><tbody><tr><td>Analog oscilloscope</td><td>B&amp;amp;K Precision</td><td>2160C</td><td>Analog oscilloscope for real-time monitoring of nanopipette current and Z-axis approach</td></tr><tr><td>AFM calibration standards</td><td>TED PELLA Inc</td><td>HS-100MG; HS-20MG</td><td>These 100 and 20 nm calibration standards are used to test the performance of HPICM system</td></tr><tr><td>Benchtop vibration Isolator</td><td>AMETEK/TMC</td><td>Everstill K-400</td><td>Active vibration isolation</td></tr><tr><td>Borosilicate glass capillaries</td><td>World Precision Instruments (WPI)</td><td>1B100F-4</td><td>Borosilicate glass capillaries for the nanopipettes</td></tr><tr><td>D-(+)-Glucose</td><td>Sigma-Aldrich</td><td>G8270</td><td>To be added to the bath solution to adjust osmolarity</td></tr><tr><td>Digitizer</td><td>National Instruments Corporation</td><td>PCI-6221</td><td>Multi-channel input/output digitizer</td></tr><tr><td>Fast analog Proportional-Integral-Derivative (PID) control for Z movement</td><td>Standford Research Systems</td><td>SIM900, SIM960, SIM980</td><td>Instrumentation modules integrated in an external PID controller for Z movement. It requires a fast response that is usually not implemented in commercial piezo amplifiers.</td></tr><tr><td>Faraday cage</td><td>AMETEK/TMC</td><td>Type II</td><td>Required to shield electromagnetic interference</td></tr><tr><td>Glass bottom dish</td><td>World Precision Instruments (WPI)</td><td>FD5040-100</td><td>Used as the dish for the chamber for the tissue</td></tr><tr><td>Hanks&#039; Balanced Salt Solution (HBSS)</td><td>Gibco, Thermo Fisher Scientific</td><td>14025092</td><td>Extracellular (bath) solution</td></tr><tr><td>Instrumentation amplifier</td><td>Brownlee Precision</td><td>Model 440</td><td>Instrumentation amplifier provides required offsets, filtering, and secondary magnification or attenuation</td></tr><tr><td>Laser-based micropipette puller</td><td>Sutter Instrument</td><td>P-2000/G</td><td>Micropipette puller to fabricate the nanopipettes. Laser is needed for sharp quartz pipettes.</td></tr><tr><td>Lebovitz&#039;s L-15, without phenol red</td><td>Gibco, Thermo Fisher Scientific</td><td>21083027</td><td>Extracellular (bath) solution</td></tr><tr><td>Micromanipulator</td><td>Scientifica</td><td>PatchStar</td><td>Used for &quot;course&quot; positioning of the Z piezo actuator</td></tr><tr><td>Microscope</td><td>Nikon</td><td>Eclipse TS100</td><td>Inverted optical microscope</td></tr><tr><td>Patch amplifier</td><td>Molecular Devices</td><td>Axopatch 200B</td><td>The patch clamp amplifier measures the current through the nanopipette</td></tr><tr><td>Piezo amplifier (XY axes)</td><td>Physik Instrumente (PI)</td><td>E-500.00, E-505.00, E-509.C2A</td><td>Amplification and PID control for XY piezo translation stage</td></tr><tr><td>Piezo amplifier (Z axis)</td><td>Piezosystem jena</td><td>ENT 400 &amp;amp; 800</td><td>Custom amplifier consisting of ENT 400 power supply and two ENT 800 amplifiers in parallel to achieve max current of 1.6 nA</td></tr><tr><td>Plastic Coverslips</td><td>TED PELLA Inc</td><td>26028</td><td>Used in the fabrication of the chambers for the tissue&amp;nbsp;</td></tr><tr><td>SICM controller &amp;amp; software*</td><td>Ionscope, UK (ionscope.com)</td><td>N/A</td><td>Custom controller based on SBC6711 digital signal processing board from Innovative Integration Ltd</td></tr><tr><td>Silicone elastomer (Sylgard)</td><td>World Precision Instruments (WPI)</td><td>SYLG184</td><td>Used to attach the flexible glass fibers to the chamber for the tissue</td></tr><tr><td>Silicon glue</td><td>The Dow Chemical Company</td><td>734</td><td>Used to glue the different parts of the chamber for the tissue</td></tr><tr><td>Tungsten rod</td><td>A-M Systems</td><td>717500</td><td>Used for holding the dental floss strands in the chamber for the tissue</td></tr><tr><td>XY piezo nanopositioner</td><td>Physik Instrumente (PI)</td><td>P-733.2DD</td><td>XY translation stage with capacitive sensors</td></tr><tr><td>Z piezo nanopositioner</td><td>Piezosystem jena</td><td>RA 12/24 SG</td><td>Ring piezoactuator with a strain gage sensor</td></tr><tr><td>*Ionscope does not sell separate SICM controllers anymore. There are few other commercial systems:&amp;nbsp; NX12-Bio and NX10 SICM,&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>Park Systems, Korea and SICM modules from ICAPPIC Limited, UK (icappic.com). All these systems are based on the original&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>HPICM principles. However, imaging stereocilia bundles in the hair cells requires several custom modifications that are technically&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>challenging (or even impossible) in the closed &amp;ldquo;ready-to-go&amp;rdquo; systems such as Ionscope or NX12-Bio/NX10. Currently, there is only one&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>modular system (ICAPPIC) that has the flexibility to suit any SICM/HPICM experiment but requires some component integration.&amp;nbsp;</td><td></td><td></td><td></td></tr></tbody></table>

stereocilia bundle imaging with nanoscale resolution in live mammalian auditory hair cells

Inner ear hair cells detect sound-induced displacements and transduce these stimuli into electrical signals in a hair bundle that consists of stereocilia that are arranged in rows of increasing height. When stereocilia are deflected, they tug on tiny (~5 nm in diameter) extracellular tip links interconnecting stereocilia, which convey forces to the mechanosensitive transduction channels. Although mechanotransduction has been studied in live hair cells for decades, the functionally important ultrastructural details of the mechanotransduction machinery at the tips of stereocilia (such as tip link dynamics or transduction-dependent stereocilia remodeling) can still be studied only in dead cells with electron microscopy. Theoretically, scanning probe techniques, such as atomic force microscopy, have enough resolution to visualize the surface of stereocilia. However, independent of imaging mode, even the slightest contact of the atomic force microscopy probe with the stereocilia bundle usually damages the bundle. Here we present a detailed protocol for the hopping probe ion conductance microscopy (HPICM) imaging of live rodent auditory hair cells. This non-contact scanning probe technique allows time lapse imaging of the surface of live cells with a complex topography, like hair cells, with single nanometers resolution and without making physical contact with the sample. The HPICM uses an electrical current passing through the glass nanopipette to detect the cell surface in close vicinity to the pipette, while a 3D-positioning piezoelectric system scans the surface and generates its image. With HPICM, we were able to image stereocilia bundles and the links interconnecting stereocilia in live auditory hair cells for several hours without noticeable damage. We anticipate that the use of HPICM will allow direct exploration of ultrastructural changes in the stereocilia of live hair cells for better understanding of their function.

The protocol presented in this paper can be used to visualize any live cells with complex topography. Following these steps, we routinely obtain images of live rat auditory hair cell bundles (Figure 6B,D). In spite of having lower X-Y resolution when compared to SEM images, our HPICM images can successfully resolve the different rows of stereocilia, the shape of the stereocilia tips, and even the small links (~5 nm in diameter) connecting adjacent stereocilia (<strong class=...

Watch this Scientific Journal Video about Stereocilia Bundle Imaging with Nanoscale Resolution in Live Mammalian Auditory Hair Cells at JoVE.com

Stereocilia Bundle Imaging with Nanoscale Resolution in Live Mammalian Auditory Hair Cells

Here we present a protocol for the Hopping Probe Ion Conductance Microscopy (HPICM), a non-contact scanning probe technique that allows nanoscale imaging of stereocilia bundles in live auditory hair cells.

Inner ear hair cells detect sound-induced displacements and transduce these stimuli into electrical signals in a hair bundle that consists of stereocilia ...

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3.1K Views.  University of Kentucky. Here we present a protocol for the Hopping Probe Ion Conductance Microscopy (HPICM), a non-contact scanning probe technique that allows nanoscale imaging of stereocilia bundles in live auditory hair cells.

Video: Stereocilia Bundle Imaging with Nanoscale Resolution in Live Mammalian Auditory Hair Cells

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 ...

Investigating Outer Hair Cell Motility with a Combination of External Alternating Electrical Field Stimulation and High-speed Image Analysis

OHCs are cylindrical sensorimotor cells located in the Organ of Corti, the auditory organ inside the mammalian inner ear. The name "hair cells" derives from their characteristic apical bundle of stereocilia, a critical element for detection and transduction of sound energy 1. OHCs are able to change shape &#x2014;elongate, shorten and bend&#x2014; in response to electrical, mechanical and chemical stimulation, a motor response considered crucial for cochlear amplification of acoustic signals 2.
OHC stimulation induces two different motile responses: i) electromotility, a.k.a fast motility, changes in length in the microsecond range derived from electrically-driven conformational changes in motor proteins densely packed in OHC plasma membrane, and ii) slow motility, shape changes in the millisecond to seconds range involving cytoskeletal reorganization 2, 3. OHC bending is associated with electromotility, and result either from an asymmetric distribution of motor proteins in the lateral plasma membrane, or asymmetric electrical stimulation of those motor proteins (e.g., with an electrical field perpendicular to the long axis of the cells) 4. Mechanical and chemical stimuli induce essentially slow motile responses, even though changes in the ionic conditions of the cells and/or their environment can also stimulate the plasma membrane-embedded motor proteins 5, 6. Since OHC motile responses are an essential component of the cochlear amplifier, the qualitative and quantitative analysis of these motile responses at acoustic frequencies (roughly from 20 Hz to 20 kHz in humans) is a very important matter in the field of hearing research 7.
The development of new imaging technology combining high-speed videocameras, LED-based illumination systems, and sophisticated image analysis software now provides the ability to perform reliable qualitative and quantitative studies of the motile response of isolated OHCs to an external alternating electrical field (EAEF) 8. This is a simple and non-invasive technique that circumvents most of the limitations of previous approaches 9-11. Moreover, the LED-based illumination system provides extreme brightness with insignificant thermal effects on the samples and, because of the use of video microscopy, optical resolution is at least 10-fold higher than with conventional light microscopy techniques 12. For instance, with the experimental setup described here, changes in cell length of about 20 nm can be routinely and reliably detected at frequencies of 10 kHz, and this resolution can be further improved at lower frequencies. 
We are confident that this experimental approach will help to extend our understanding of the cellular and molecular mechanisms underlying OHC motility.

A reliable method to investigate outer hair cell (OHC) motile responses, including electromotility, slow motility and bending, is described. OHC motility is elicited by stimulation with an external alternating electrical field, and the method takes advantage of high-speed image recording, LED-based illumination, and last generation image analysis software.

OHCs are cylindrical sensorimotor cells located in the Organ of Corti, the auditory organ inside the mammalian inner ear. The name "hair cells" derives ...

Evaluation of Planar-Cell-Polarity Phenotypes in Ciliopathy Mouse Mutant Cochlea

In recent years, primary cilia have emerged as key regulators in development and disease by influencing numerous signaling pathways. One of the earliest signaling pathways shown to be associated with ciliary function was the non-canonical Wnt signaling pathway, also referred to as planar cell polarity (PCP) signaling. One of the best places in which to study the effects of planar cell polarity (PCP) signaling during vertebrate development is the mammalian cochlea. PCP signaling disruption in the mouse cochlea disrupts cochlear outgrowth, cellular patterning and hair cell orientation, all of which are affected by cilia dysfunction. The goal of this protocol is to describe the analysis of PCP signaling in the developing mammalian cochlea via phenotypic analysis, immunohistochemistry and scanning electron microscopy. Defects in convergence and extension are manifested as a shortening of the cochlear duct and/or changes in cellular patterning, which can be quantified following dissection from developing mouse mutants. Changes in stereociliary bundle orientation and kinocilia length or positioning can be observed and quantitated using either immunofluorescence or scanning electron microscopy (SEM). A deeper insight into the role of ciliary proteins in cellular signaling pathways and other biological phenomena is crucial for our understanding of cellular and developmental biology, as well as for the development of targeted treatment strategies.

Primary cilia influence various signaling pathways. The mammalian cochlea is ideal for examining planar cell polarity (PCP) signaling. Cilia dysfunction affects cochlear outgrowth, cellular patterning and hair cell orientation, readouts of PCP. Our goal is to analyze PCP signaling in mouse cochlea via phenotypic analysis, immunohistochemistry and scanning electron microscopy.

In recent years, primary cilia have emerged as key regulators in development and disease by influencing numerous signaling pathways. One of the earliest ...

Medicine

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.

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.

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

Dextran Labeling and Uptake in Live and Functional Murine Cochlear Hair Cells

The hair cell mechanotransduction (MET) channel plays an important role in hearing. However, the molecular identity and structural information of MET remain unknown. Electrophysiological studies of hair cells revealed that the MET channel has a large conductance and is permeable to relatively large fluorescent cationic molecules, including some styryl dyes and Texas Red-labeled aminoglycoside antibiotics. In this protocol, we describe a method to visualize and evaluate the uptake of fluorescent dextrans in hair cells of the organ of Corti explants that can be used to assay for functional MET channels. We found that 3 kDa Texas Red-labeled dextran specifically labels functional auditory hair cells after 1&#8211;2 h incubation. In particular, 3 kDa dextran labels the two shorter stereocilia rows and accumulates in the cell body in a diffuse pattern when functional MET channels are present. An additional vesicle-like pattern of labeling was observed in the cell body of hair cells and surrounding supporting cells. Our data suggest that 3 kDa Texas-Red dextran can be used to visualize and study two pathways for cellular dye uptake; a hair cell-specific entry route through functional MET channels and endocytosis, a pattern also available to larger dextran.

Here, we present a method for visualizing the uptake of 3 kDa Texas Red-labeled dextran in auditory hair cells with functional mechanotransduction channels. In addition, dextrans of 3&#8211;10 kDa can be used to study endocytosis in hair and supporting cells of the organ of Corti.

The hair cell mechanotransduction (MET) channel plays an important role in hearing. However, the molecular identity and structural information of MET ...

Bioengineering

Auditory Brainstem Response and Outer Hair Cell Whole-cell Patch Clamp Recording in Postnatal Rats

The outer hair cell is one of the two types of sensory hair cells in the mammalian cochlea. They alter their cell length with the receptor potential to amplify the weak vibration of low-level sound signal. The morphology and electrophysiological property of outer hair cells (OHCs) develop in early postnatal ages. The maturation of outer hair cell may contribute to the development of the auditory system. However, the process of OHCs development is not well studied. This is partly because of the difficulty to measure their function by an electrophysiological approach. With the purpose of developing a simple method to address the above issue, here we describe a step-by-step protocol to study the function of OHCs in acutely dissociated cochlea from postnatal rats. With this method, we can evaluate the cochlear response to pure tone stimuli and examine the expression level and function of the motor protein prestin in OHCs. This method can also be used to investigate the inner hair cells (IHCs).

This protocol describes methods to record the auditory brainstem response from postnatal rat pups. To examine the functional development of outer hair cells, the experimental procedure of whole-cell patch clamp recording in isolated outer hair cells is described step-by-step.

The outer hair cell is one of the two types of sensory hair cells in the mammalian cochlea. They alter their cell length with the receptor potential to ...

In Vivo Calcium Imaging of Lateral-line Hair Cells in Larval Zebrafish

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory stimuli that mechanically deflect apical protrusions called hair bundles. Deflection opens mechanotransduction (MET) channels in hair bundles, leading to an influx of cations, including calcium. This cation influx depolarizes the cell and opens voltage-gated calcium channels located basally at the hair-cell presynapse. In mammals, hair cells are encased in bone, and it is challenging to functionally assess these activities in vivo. In contrast, larval zebrafish are transparent and possess an externally located lateral-line organ that contains hair cells. These hair cells are functionally and structurally similar to mammalian hair cells and can be functionally assessed in vivo. This article outlines a technique that utilizes a genetically encoded calcium indicator (GECI), GCaMP6s, to measure stimulus-evoked calcium signals in zebrafish lateral-line hair cells. GCaMP6s can be used, along with confocal imaging, to measure in vivo calcium signals at the apex and base of lateral-line hair cells. These signals provide a real-time, quantifiable readout of both mechanosensation- and presynapse-dependent calcium activities within these hair cells. These calcium signals also provide important functional information regarding how hair cells detect and transmit sensory stimuli. Overall, this technique generates useful data about relative changes in calcium activity in vivo. It is less well-suited for quantification of the absolute magnitude of calcium changes. This in vivo technique is sensitive to motion artifacts. A reasonable amount of practice and skill are required for proper positioning, immobilization, and stimulation of larvae. Ultimately, when properly executed, the protocol outlined in this article provides a powerful way to collect valuable information about the activity of hair-cells in their natural, fully integrated states within a live animal.

The zebrafish is a model system that has many valuable features including optical clarity, rapid external development, and, of particular importance to the field of hearing and balance, externally located sensory hair cells. This article outlines how transgenic zebrafish can be used to assay both hair-cell mechanosensation and presynaptic function in toto.

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory ...

Cochlear Surface Preparation in the Adult Mouse

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous etiologies such as exposure to excessive noise, the use of ototoxic medications, bacterial or viral ear infections, head injuries, and the aging process. Loss of sensory hair cells is a common pathological feature of the varieties of acquired hearing loss. Additionally, the inner hair cell synapse can be damaged by mild insults. Therefore, surface preparations of cochlear epithelia, in combination with immunolabeling techniques and confocal imagery, are a very useful tool for the investigation of cochlear pathologies, including losses of ribbon synapses and sensory hair cells, changes in protein levels in hair cells and supporting cells, hair cell regeneration, and determination of report gene expression (i.e., GFP) for verification of successful transduction and identification of transduced cell types. The cochlea, a bony spiral-shaped structure in the inner ear, holds the auditory sensory end organ, the organ of Corti (OC). Sensory hair cells and surrounding supporting cells in the OC are contained in the cochlear duct and rest on the basilar membrane, organized in a tonotopic fashion with high-frequency detection occurring in the base and low-frequency in the apex. With the availability of molecular and genetic information and the ability to manipulate genes by knockout and knock-in techniques, mice have been widely used in biological research, including in hearing science. However, the adult mouse cochlea is miniscule, and the cochlear epithelium is encapsulated in a bony labyrinth, making microdissection difficult. Although dissection techniques have been developed and used in many laboratories, this modified microdissection method using cell and tissue adhesive is easier and more convenient. It can be used in all types of adult mouse cochleae following decalcification.

This article presents a modified cochlear surface preparation method that requires decalcification and use of a cell and tissue adhesive to adhere the pieces of cochlear epithelia to 10 mm round cover slips for immunohistochemistry in adult mouse cochleae.

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous ...

Whole Neonatal Cochlear Explants as an In vitro Model

Untreated hearing loss imposes significant costs on the global healthcare system and impairs individuals' quality of life. Sensorineural hearing loss is characterized by the cumulative and irreversible loss of sensory hair cells and auditory nerves in the cochlea. Entire and vital cochlear explants are one of the fundamental tools in hearing research to detect hair cell loss and to characterize the molecular mechanisms of the inner ear cells. Many years ago, a protocol for neonatal cochlear isolation was developed, and although it has been modified over time, it still holds potential for improvement. 
This paper presents an optimized protocol for isolating and culturing whole neonatal cochlear explants in multi-well culture chambers that enables the study of hair cells and spiral ganglion neuron cells along the entire length of the cochlea. The protocol was tested using cochlear explants from mice and rats. Healthy cochlear explants were obtained to study the interaction between hair cells, spiral ganglion neuron cells, and the surrounding supporting cells. 
One of the main advantages of this method is that it simplifies the organ culture steps without compromising the quality of the explants. All three turns of the organ of Corti are attached to the bottom of the chamber, which facilitates in vitro experiments and the comprehensive analysis of the explants. We provide some examples of cochlear images from different experiments with live and fixed explants, demonstrating that the explants retain their structure despite exposure to ototoxic drugs. This optimized protocol can be widely used for the integrative analysis of the mammalian cochlea.

Whole Neonatal Cochlear Explants as an In vitro Model

The current protocol updates previous protocols and incorporates relatively straightforward approaches for culturing high-quality cochlear explants. This provides reliable data acquisition and high-resolution imaging in live and fixed cells. This protocol supports the ongoing trend of studying inner ear cells.

Untreated hearing loss imposes significant costs on the global healthcare system and impairs individuals' quality of life. Sensorineural hearing loss is ...

Efficient In Vitro Cultivation of Mouse Cochlear Hair Cells

Isolation and Culture of Primary Cochlear Hair Cells from Neonatal Mice

Cochlear hair cells are the sensory receptors of the auditory system. These cells are located in the organ of Corti, the sensory organ responsible for hearing, within the osseous labyrinth of the inner ear. Cochlear hair cells consist of two anatomically and functionally distinct types: outer and inner hair cells. Damage to either of them results in hearing loss. Notably, as inner hair cells cannot regenerate, and damage to them is permanent. Hence, in vitro cultivation of primary hair cells is indispensable for investigating the protective or regenerative effects of cochlear hair cells. This study aimed to discover a method for isolating and cultivating mouse hair cells. 
After manual removal of the cochlear lateral wall, the auditory epithelium was meticulously dissected from the cochlear modiolus under a microscope, incubated in a mixture consisting of 0.25% trypsin-EDTA for 10 min at 37 &#176;C, and gently suspended in culture medium using a 200 &#181;L pipette tip. The cell suspension was passed through a cell filter, the filtrate was centrifuged, and cells were cultured in 24-well plates. Hair cells were identified based on their capacity to express a mechanotransduction complex, myosin-VIIa, which is involved in motor tensions, and via selective labeling of F-actin using phalloidin. Cells reached &gt;90% confluence after 4 d in culture. This method can enhance our understanding of the biological characteristics of in vitro cultured hair cells and demonstrate the efficiency of cochlear hair cell cultures, establishing a solid methodological foundation for further auditory research.

Author Spotlight: Advancements in Cultivating Mouse Hair Cells for Auditory Research 

Herein, we present a detailed protocol for isolating and culturing primary cochlear hair cells from mice. Initially, the organ of Corti was dissected from neonatal (aged 3-5 days) murine cochleae under a microscope. Subsequently, cells were enzymatically digested into a single-cell suspension and identified using immunofluorescence after several days in culture.

Cochlear hair cells are the sensory receptors of the auditory system. These cells are located in the organ of Corti, the sensory organ responsible for ...

Stereocilia Bundle Imaging with Nanoscale Resolution in Live Mammalian Auditory Hair Cells

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

Investigating Outer Hair Cell Motility with a Combination of External Alternating Electrical Field Stimulation and High-speed Image Analysis

Evaluation of Planar-Cell-Polarity Phenotypes in Ciliopathy Mouse Mutant Cochlea

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

Dextran Labeling and Uptake in Live and Functional Murine Cochlear Hair Cells

Auditory Brainstem Response and Outer Hair Cell Whole-cell Patch Clamp Recording in Postnatal Rats

In Vivo Calcium Imaging of Lateral-line Hair Cells in Larval Zebrafish

Cochlear Surface Preparation in the Adult Mouse

Whole Neonatal Cochlear Explants as an In vitro Model

Isolation and Culture of Primary Cochlear Hair Cells from Neonatal Mice