We thank Vincent Schram from the NICHD microscopy and imaging core for assisting in the confocal image acquisition, and Tsg-Hui Chang for invaluable help with colony management and mice care. This research was supported by the Intramural Research Program of the NINDS, NIH, Bethesda, MD, to K.J.S. A.B. was supported by the Intramural Research Program of the NINDS, NIH, and by a Robert Wenthold Postdoctoral Fellowship from the intramural research program of the NIDCD.

The animal care and experimental procedures were performed following the guidelines for the Care and Use of Laboratory Animals, which were approved by the Animal Care and Use Committee of the National Institute of Neurological Disorders and Stroke (Animal protocol #1336 to KJS).
1. Mice
<ol>
	<li>Set a couple of breeding pairs of C57BL/6J wild-type to breed in the animal facility to control the date of birth of the litters and keep track of the age of the pups.</li>
</ol>
2. Cochleae Dissection
<ol>
	<li>Set a clean space close to a stereomicroscope to perform the dissections (Figure 1A). Use 70% ethanol to clean the space and surroundings and place a clean bench pad. A Medical Pathological Waste (MPW) plastic bag would be required to discard the animal carcasses.</li>
	<li>Prepare several 35 mm dishes with some Leibovitz&#39;s L15 media. 
	NOTE: Leibovitz&#39;s L-15 cell media contains 1&#8211;2 mM Ca2+, which is required to maintain the integrity of the tip-links, and contains essential amino acids, vitamins, and sodium pyruvate to improve cell health and survival. Serum was excluded to avoid experimental variability due to its poorly defined composition and potential interference with the dextran.</li>
	<li>Euthanize postnatal-day-6 (P6) mice by decapitation. 
	NOTE: 6 day old mice are somewhat resistant to inhalant anesthetics. Although isoflurane or prolonged CO2 exposures (up to 50 min) may be used for euthanasia, a secondary physical method is recommended to ensure death.</li>
	<li>Use surgical scissors to remove the skin of the skull by making a superficial cut from the anterior to the posterior end and across the external auditory canals.</li>
	<li>Fold the skin towards the nose to expose the cranium (Figure 2B).</li>
	<li>Make an incision from the back to the front of the skull and across the eye line (Figure 2B-C).</li>
	<li>Separate the skull in two halves and remove the brain with the use of a small spatula to expose the temporal bones (Figure 2C-D).</li>
	<li>With small scissors, cut around the temporal bones, and excise the tissue.</li>
	<li>Place both temporal bones in a 35 mm dish and make sure they are covered with L15 media (Figure 2E). 
	NOTE: The following steps are performed under the stereomicroscope. A black background usually helps to visualize the tissue during the fine dissection steps.</li>
	<li>Under a stereomicroscope equipped with a widefield eyepiece (a 10X magnification power (WF10X) and an external alternating current (AC) halogen light source), remove the surrounding cochleae tissue, semicircular canals, and vestibular organs with surgical forceps (Figure 2F).</li>
	<li>To allow the dextran and L15 media to enter the cochlear duct, perform two puncture bounds on the dissected cochleae, one on the round window and other at the apical cochlear region.</li>
	<li>Add 300 mL of Leibovitz&#39;s L15 media in each well from a 9-well glass depression plate.</li>
	<li>Place at least three dissected cochleae on each well.</li>
</ol>
3. Dextran Labeling
<ol>
	<li>Reconstitute the dextran in Hanks&#39; balanced salt solution without Ca2+ and Mg2+ (HBSS-CFM) at a final concentration of 10 mg/mL. This stock solution must be aliquoted in opaque black tubes (protected from light) and stored at -30 &#176;C until use. 
	NOTE: The use of lysine-fixable dextran is critical for a successful outcome of this protocol.</li>
	<li>Prepare each dextran at a final concentration of 2 mg/mL in 500 mL of Leibovitz&#39;s L15 media.</li>
	<li>Remove the media from the cochlea and add Leibovitz&#39;s L15 media containing the dextran of interest at a final concentration of 2 mg/mL. 
	NOTE: Although a proportion of the MET channels are open at rest41,42, the dextran incubation was performed with a gentle shaking of the explants to increase the open probability of the MET channel.</li>
	<li>Incubate at room temperature for 2 h with gentle shaking (25 rpm) by using a 3-dimensional shaker with a tilted angle of 25&#176;. 
	NOTE: Fluorescently labeled dextran must be protected from light when possible. To protect the dextran during the 2 h incubation, place the 9-well glass plate inside a cell culture P150 dish wrapped in aluminum foil.</li>
</ol>
4. Sample Preparation for Imaging
<ol>
	<li>After incubation with the dextran, wash the tissue for 2 min twice with media and once with HBSS.</li>
	<li>Incubate the tissue at room temperature for 30 min with 4% paraformaldehyde in Hanks&#39; balanced salt solution (HBSS). 
	CAUTION: Exposure to formaldehyde can be irritating to the eyes, nose, and upper respiratory tract. In certain individuals, repeated skin exposure to formaldehyde can cause sensitization that may result in allergic dermatitis. Formaldehyde is a known human carcinogen and a suspected reproductive hazard.</li>
	<li>Quickly and gently wash the fixed tissue twice with HBSS to remove the paraformaldehyde. 
	NOTE: Decalcification of the temporal bones is not needed at this developmental stage of the cochlea.</li>
	<li>Remove the spiral ligament and the tectorial membrane with fine tip forceps to dissect the organ of Corti (Figure 2G).</li>
	<li>Remove all the small pieces of tissue and wash the tissue with HBSS.</li>
	<li>Permeabilize the tissue in 0.5% Triton X-100 in PBS containing fluorescently-labeled phalloidin (conjugated to green or red when testing the uptake of TR- or FITC-labeled dextran, respectively) at a 1:200 dilution for 30 min to label F-actin and visualize the actin-based stereocilia.</li>
	<li>Wash the tissue 2&#8211;3 times for 2 min each time with HBSS buffer to remove the excess of triton and phalloidin, and once with PBS to remove the salts.</li>
	<li>Mount the organ of Corti tissues on a microscope slide and cover it with mounting media. 
	NOTE: When mounting the tissue, make sure that the side of the tissue containing the hair cell stereocilia is facing the coverslip.</li>
	<li>Remove any potential bubbles generated during the addition of the mounting media and prevent the generation of new bubbles during the placement of the coverslip. 
	NOTE: Aspirate with a pipette any bubble generated during the addition of the mounting media. To prevent air bubbles from being trapped under the coverslip, place an edge of the coverslip close to the sample and carefully and slowly lower the coverslip over the tissue using forceps or a pipette tip.</li>
	<li>Cover the tissue in mounting media with a glass coverslip (Figure 2H). 
	NOTE: Objectives with a numerical aperture above 0.4 are designed to use #1.5 coverslips (0.17 mm thickness). Using the wrong coverslip may have severe implications for the intensity and quality of the images.</li>
	<li>Incubate the mounted tissue overnight at room temperature to let the mounting media dry and store the slides at 4 &#176;C until imaging.</li>
</ol>
5. Image Acquisition and Image Processing
NOTE: The confocal images were taken with a LSM 880 confocal microscope equipped with a 32 channel Airyscan detector in the super-resolution (SR) mode43 and a 63x objective.
<ol>
	<li>Add a small drop of immersion oil on the objective.</li>
	<li>Place the microscope slide containing the mounted tissue sample in the microscope stage with the glass coverslip facing the immersion oil.</li>
	<li>Focus on the sample and set the imaging parameters using the image acquisition software.</li>
	<li>Use identical image acquisition settings and optimal parameters for x, y, and z resolution for each independent experiment. A piezo-driven focus system is required to quickly and precisely move the objective when acquiring the z-stack of images. 
	NOTE: To image the entire apical region of the hair cells, collect a z-stack of images from the stereocilia to the apical half of the hair cell body using the optimal settings. It is crucial to collect a large z-stack along the hair cell to assure the imaging of the vesicle-like particles.</li>
	<li>Use the image acquisition software to process the raw confocal images using the Airyscan 3D reconstruction algorithm with the automatic default deconvolution filter settings.</li>
	<li>Open the confocal images in an image processing software to adjust the brightness and contrast, add the scale bar, and export the images for the final figures.</li>
</ol>

The authors have nothing to disclose.

This protocol describes how to perform uptake experiments in murine organ of Corti explants with 3 kDa dextran Texas Red. The goal of this method is to test whether molecules larger than others previously tested were also able to specifically label auditory hair cells and permeate through the MET channel. Similar experimental protocols have been previously used to evaluate the permeability of hair cells to other fluorescent dyes such as FM1-43 (0.56 kDa)12,19,20 and Texas Red-labeled aminoglycosides (1.29&#8211;1.43 kDa)13,52. The time needed for labeling hair cells using these fluorescent molecules varies with their size. The maximal labeling requires a few minutes for FM1-4313,53, 30-60 min for Texas Red-labeled aminoglycosides13, and 1.5 h for Texas Red-labeled 3 kDa dextran. Thus, long incubation times are crucial for this experiment as we saw only minimal labeling of the hair cell body at short incubation times (Figure 2C). We chose to study P6 mice for these experiments because, at this age, most apical and basal hair cells carry transduction currents and express both TMC1 and TMC2 proteins12,18,54,55. It should be possible to study younger animals as MET currents are present earlier than P6, in particular near the base of the cochlea12. Studying older animals may be more challenging because dissection of the bony cochlea becomes more difficult as this structure continues to ossify56.
There are several critical steps in this protocol. The most crucial one being the successful dissection and preparation of the organ of Corti tissue (Figure 1). The delicate structure of the organ of Corti and encasement in the boney cochlea presents a challenge for histological and biochemical analysis. Several detailed protocols for the successful dissection of this tissue have been previously published in this journal56,57,58 and others59. Disruption of the cell-cell junction and loss of the auditory sensory epithelium integrity may lead to the uptake of dextran of up to 40 kDa and the labeling of the affected cells39,40. In agreement with this, we observed unspecific labeling of hair cells and the surrounding supporting cells when the tissue was unintentionally damaged with the forceps during dissection before dextran incubation (Figure 2B and Figure 3B). There are no tricks or guidelines to reach proficiency at dissecting an intact organ of Corti other than practice and patience. However, the presence of 1&#8211;2 mM Ca2+ during tissue dissection and uptake experiments is crucial to maintain the integrity of the tip-link that transmits the mechanical force to the MET channel and thereby preserve MET channel function. Once the tissue is fixed with 4% PFA, the presence of calcium in the buffer or media becomes trivial.
One of the limitations of this study resides in the heterogeneous nature of the dextran molecules. The dextran elaborated by the Leuconostoc bacteria is a polymer of anhydroglucose composed of approximately 95% alpha-(1&#8211;6) D-linkages (Figure 2A). The remaining linkages account for the branching of dextran, which correlates with its length, so dextran of lower molecular weight are more rod-like and have a narrower size distribution while larger dextrans are polydisperse60. Therefore, the actual molecular weight of the molecules in each preparation of the 3 kDa dextran ranges from 1.5 to 3.0 kDa, including the fluorescent molecule61. Additionally, the fluorescently labeled dextran differ in the number of Texas Red molecules attached to the dextran (degree of labeling). It is advisable to use dextran with a degree of labeling of at least 0.4 since a lower degree of labeling will hinder the ability to detect the fluorescently labeled dextran by microscopy. The fluorescent dye coupled to the dextran and the lysine residues in the fixable versions, will determine the overall charge of the dextran. This is an important consideration since the cationic selectivity of the MET channel would preferably uptake cationic dextran27. Lastly, a lysine-fixable dextran is necessary to reach the resolution displayed in these experiments. A non-fixable dextran will continue to diffuse into the tissue hampering its accurate localization and visualization.
The LSM 880 confocal microscope equipped with an Airyscan detector used to image the organ of Corti samples provided us with twice the resolution, and better signal-to-noise ratio (SNR) compared to a conventional confocal microscope62. Therefore, this microscope allowed us not only to observe the robust dextran accumulation at the cell body but also to pinpoint the accumulation of the dextran at the stereocilia tips and the vesicle-like pattern at the cell body. A conventional confocal microscope would allow for the visualization and quantification of the accumulation of fluorescent dextran at the cell body, but it would difficult to distinguish the three rows of stereocilia and the precise localization of the dextran at the tips of the shorter stereocilia rows. A similar resolution to the one achieved with the LSM 880 Airyscan confocal could be reached using a conventional confocal microscope with oversampling and deconvolution, although the SNR would be lower than the one reached with the Airyscan detector.
In addition to the use of 3 kDa dextran-TR to assay for functional MET channels, this protocol could be further used to study endocytic processes in hair cells since all the dextrans tested reveal an intracellular vesicle-like pattern resembling endocytosis. This process was not the focus of our work, and additional studies would be required to fully characterize this phenomenon.

The hair cells of the inner ear are the sensory cells that detect sound and covert the mechanically stimuli in electrical signals, which are ultimately interpreted by our brain. These cells have a staircase-shaped bundle of three rows of actin-based filaments, known as stereocilia, which protrude from their apical region1,2. The mechanical stimuli deflect the stereocilia filaments toward the longest row and trigger the opening of the mechanotransduction (MET) channels3. The opening of the MET channels leads to an influx of cations that depolarizes the cell and consequently signals the release of synapse vesicles at the basal region of the hair cell.
The biophysical properties of the MET channel essential for hearing have been extensively characterized. Among other properties, these channels are cationic selective and have a relatively large conductance (150&#8211;300 pS in low Ca2+)4,5,6,7,8,9,10. Remarkably, large fluorescent molecules such as FM1-43 and Texas Red-labeled aminoglycosides are permeant blockers of the MET channel, resulting in their accumulation in the hair cell body that can be visualized using fluorescence microscopy11,12,13,14. Conversely, the molecular identity and the structure of the MET channel and its permeation pathway have remained elusive. Increasing experimental evidence indicates that the transmembrane-like channel protein 1 (TMC1) is a component of the MET channel in mature hair cells15,16,17,18,19. Mutations in the transmembrane-like channel 1 (TMC1) alter the MET channel properties19,20,21,22 and cause deafness. In addition, TMC1 localizes to the site of the MET channel18,23 and interacts with the tip-link responsible for transmitting the mechanical force to the MET channel24,25. Furthermore, recent bioinformatics analysis has identified the TMC proteins as evolutionary related to the mechanosensitive channels TMEM63/OSCA proteins and the TMEM16 proteins, a family of calcium-activated chloride channels and lipid scramblases26,27,28. A structural model of TMC1 based on the relationship between these proteins revealed the presence of a large cavity at the protein-lipid interface27. This cavity harbors the two TMC1 mutations that cause autosomal dominant hearing loss (DFNA36)27,29,30,31,32, and selective modification of cysteine mutants for residues in the cavity alter MET channel properties28, indicating that it could function as the permeation pathway of the MET channel. The large size of this predicted cavity in TMC proteins could explain the ability of large molecules to permeate the MET channel. To test the prediction that the MET channel contains an unusually large permeation pathway and to push the limits of the size of the cavity observed in TMC1, we developed a protocol to perform uptake experiments in the organ of Corti explants with a larger molecule, 3 kDa dextran fluorescently labeled with Texas Red.
Dextran is a complex branched polysaccharide composed of many D-glucose molecules bound by alpha-1,6 glycosidic linkages. Its high solubility in water, low cell toxicity, and bioinertity make it a versatile tool to study several cellular processes. In addition, dextran is available in a wide range of sizes and fluorescently labeled with fluorophores of several colors. Fluorescently labeled dextrans are commonly used in cell and tissue permeability research33,34, to study endocytosis in multiple cellular systems35,36, and for neural tracing37,38. In the auditory field, dextran molecules have also been used to assess the disruption of the cell-cell junction and loss of the auditory sensory epithelium integrity after exposure to intense noise in the chinchilla organ of Corti39,40.
In this work, we exploited the properties of some of the smallest (3 and 10 kDa) fluorescent dextrans to perform uptake experiments in murine inner ear hair cells and explore the size of the permeation pathway of the inner ear hair cell MET channel. In addition, we used a laser-scanning confocal microscope (LSM) 880 equipped with an Airyscan detector to visualize and localize fluorescent dextran at the stereocilia and the cell body of auditory hair cells.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>#1.5 glass coverslips 18mm</td>
			<td>Warner Instruments</td>
			<td>64-0714</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 Phalloidin</td>
			<td>ThermoFisher</td>
			<td>A12379</td>
			<td></td>
		</tr>
		<tr>
			<td>Alexa Fluor 594 Phalloidin</td>
			<td>ThermoFisher</td>
			<td>A12381</td>
			<td></td>
		</tr>
		<tr>
			<td>alpha Plan-Apochromat 63X/1.4 Oil Corr M27 objective</td>
			<td>Carl Zeiss</td>
			<td>420780-9970-000</td>
			<td></td>
		</tr>
		<tr>
			<td>Amiloride hydrochloride</td>
			<td>EMD MILLIPORE</td>
			<td>129876</td>
			<td></td>
		</tr>
		<tr>
			<td>Benchwaver 3-dimensional Rocker</td>
			<td>Benchmarks scientific</td>
			<td>B3D5000</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6J wild-type mice</td>
			<td>strain 000664</td>
			<td>The Jackson Laboratory</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell impermeant BAPTA tetrapotassium salt</td>
			<td>ThermoFisher</td>
			<td>B1204</td>
			<td></td>
		</tr>
		<tr>
			<td>Dextran, Fluorescein, 10,000 MW, Anionic, Lysine Fixable</td>
			<td>ThermoFisher</td>
			<td>D1820</td>
			<td></td>
		</tr>
		<tr>
			<td>Dextran, Texas Red, 10,000 MW, Lysine Fixable</td>
			<td>ThermoFisher</td>
			<td>D1863</td>
			<td></td>
		</tr>
		<tr>
			<td>Dextran, Texas Red, 3000 MW, Lysine Fixable</td>
			<td>ThermoFisher</td>
			<td>D3328</td>
			<td></td>
		</tr>
		<tr>
			<td>Formaldehyde Aqueous Solution EM Grade</td>
			<td>Electron microscopy science</td>
			<td>15710</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS, calcium, magnesium, no phenol red</td>
			<td>ThermoFisher</td>
			<td>14025</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS, no calcium, no magnesium, no phenol red</td>
			<td>ThermoFisher</td>
			<td>14170</td>
			<td></td>
		</tr>
		<tr>
			<td>Image J or FIJI</td>
			<td>NIH</td>
			<td><a href="http://fiji.sc/" target="_blank">http://fiji.sc/</a></td>
			<td></td>
		</tr>
		<tr>
			<td>Immersol 518F oil immersion media</td>
			<td>Carl Zeiss</td>
			<td>444970-9000-000</td>
			<td></td>
		</tr>
		<tr>
			<td>Leibovitz&#39;s L-15 Medium, GlutaMAX Supplement</td>
			<td>ThermoFisher</td>
			<td>31415029</td>
			<td></td>
		</tr>
		<tr>
			<td>neomycin trisulfate salt hydrate</td>
			<td>Sigma</td>
			<td>N6386</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS (10X), pH 7.4</td>
			<td>ThermoFisher</td>
			<td>70011069</td>
			<td></td>
		</tr>
		<tr>
			<td>Phalloidin-CF405M</td>
			<td>Biotium</td>
			<td>00034</td>
			<td></td>
		</tr>
		<tr>
			<td>ProLong Diamond antifade mounting media</td>
			<td>ThermoFisher</td>
			<td>P36970</td>
			<td></td>
		</tr>
		<tr>
			<td>superfrost plus microscope slide</td>
			<td>Fisherbrand</td>
			<td>22-037-246</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma</td>
			<td>T8787</td>
			<td></td>
		</tr>
		<tr>
			<td>Zen Black 2.3 SP1 software</td>
			<td>Carl Zeiss</td>
			<td><a href="https://www.zeiss.com/microscopy/us/products/microscope-software/zen.html" target="_blank">https://www.zeiss.com/microscopy/us/products/microscope-software/zen.html</a></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

We observed robust and specific labeling of hair cells after 2h incubation of organ of Corti explants from wild-type postnatal-day-6 (P6) mice with 3 kDa dextran fluorescently labeled with Texas Red (dextran-TR) (Figure 2A-B). Dextran labeling was observed in both inner and outer hair cells (IHC and OHC) at the basal, middle, and apical regions of the organ of Corti (Figure 2B).
Fluorescently labeled phalloidin was used to counterstain filamentous actin (F-actin) and visualize the actin-based hair cell stereocilia. We also performed similar experiments at shorter incubation times, and although we observed dextran-TR accumulation in the hair cell body, the signals were weaker and more variable than those at 2 h incubation (Figure 2C).
We next imaged both stereocilia and cell bodies of the hair cells and observed that only those cells that incorporated 3 kDa dextran-TR in their cell body showed fluorescent labeling of their stereocilia (Figure 3A). This relationship between stereocilia and cell body labeling was absent in hair cells from damaged tissue, which also presented unspecific labeling of dextran in several cell types of the sensory epithelium (yellow squares in Figure 2B, and Figure 3B). Importantly, we observed a uniform fluorescence signal along the stereocilia with enrichment at the tips of the shorter stereocilia rows where the MET channel is located18,23 (Figure 3C). Also, we noticed vesicle-like structures in the cell body of the hair cells and the neighboring supporting cells. The vesicle-like pattern of uptake in these cells suggests that dextran-TR can also be taken up by endocytosis (Figure 3C).
The protocol described here also allows for the examination of the uptake of larger dextrans and combinations of different dextrans. Larger dextran of 10 kDa labeled with Texas Red (dextran-TR) or fluorescein (dextran-FITC) also produced a vesicle-like pattern in the cell body of hair cells and supporting cells, in addition to accumulating around the hair cell membrane in a patchy pattern (Figure 4A,B). The 10 kDa dextran-TR also superficially labeled the three stereocilia rows of all the hair cells (Figure 4A, inset), probably due to the negatively charged surface of the hair cell plasma membrane44,45,46. We next examined the uptake of 3 kDa dextran-TR and 10 kDa dextran-FITC simultaneously in the organ of Corti explants. For the 3 kDa dextran-TR, we observed both a diffuse and a vesicle-like pattern. However, the 10 kDa dextran-FITC only displayed a vesicle-like pattern (Figure 4C). These data suggest that dextran uptake occurs by at least two distinct mechanisms that are dependent on the size of the dextran.
We next assessed whether functional MET channels were required for the uptake of 3 kDa dextran-TR. To do this, we tested dextran incorporation in hair cells from the organ of Corti explants in the presence of MET channel blockers (neomycin and amiloride)13,14,47 or the Ca2+ chelator BAPTA, which abolishes MET currents by breaking the tip-links and preventing the gating of the channel25,48,49. In these experiments, the tissue explants were previously incubated for 30 min with neomycin (500 mM), amiloride (150 mM), or with BAPTA (5mM) before the addition of 3 kDa dextran-TR in the presence of the corresponding MET blocker. The presence of BAPTA or the MET channel blockers prevented the stereocilia labeling (Figure 5A) and the uptake of 3 kDa dextran-TR in hair cells (Figure 5B). However, blockade of the MET channel preserved the vesicle-like pattern (Figure 5B), indicating that this pattern of uptake is independent of functional MET channels. Similar results have been observed in hair cells from TMC1/TMC2 double knock-out mice, which lack MET27. Intriguingly, these vesicle-like structures were not observable in the presence of amiloride, which is known to inhibit the Na+-H+ exchanger and thereby inhibit endocytosis50,51. These results indicate that 3kDa dextran-TR enters the hair cells through two different pathways, one common to larger dextrans involving vesicle-like structures and another that depends on functional MET channels.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60769/60769fig1.jpg" /> 
Figure 1: Steps of the dissection of a murine organ of Corti. (A) Clean area and materials required for the dissection. (B) Exposed mouse cranium. Forceps are holding the skin, which has folded towards the nose. Black lines indicate the two incisions that are required for the removal of the brain. (C) Incised and opened cranium to allow for removal of the brain with a spatula (on the right). (D) Cranium and temporal bones after brain removal. (E) Excised skull, including the temporal bones (dashed black squares) in a P35 plate covered with media. (F) Excised cochleae. (G) Two dissected organs of Corti on the well of a glass depression plate. (H) Mounted sample on a glass coverslip ready for imaging. <a href="https://www.jove.com/files/ftp_upload/60769/60769fig1alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/60769/60769fig2.jpg" /> 
Figure 2: Hair cells uptake 3 kDa dextran-TR. (A) Schematic representation of Texas Red-labeled dextran (dextran-TR) containing six molecules of glucose corresponding to a molecular weight of 1.08 kDa. A succinimidyl ester reaction links a Texas Red molecule (magenta) to a glucose monomer. (B) Representative confocal image showing specific labeling of sensory hair cells (HC) with 3 kDa dextran-TR across the whole organ of Corti from a 6 day-old mouse. The basal (BA), middle (MD), and apical (AP) regions of the organ are indicated. Yellow squares indicate tissue damaged regions. (C) 3 kDa dextran-TR fluorescence after 30, 60, 90, or 120 min incubation with the organ of Corti explants is shown in magenta merged with F-actin in green (top images) and independently in grayscale (bottom images). The image display range was linearly adjusted in each one of the gray images independently for visualization of the 3 kDa dextran-TR fluorescence. Scale bar = 20 mm. This figure has been modified from27. <a href="https://www.jove.com/files/ftp_upload/60769/60769fig2alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/60769/60769fig3a.jpg" /> 
Figure 3: Hair cell body and stereocilia labeling with 3 kDa dextran-TR. (A) Confocal images displaying 3 kDa dextran-TR fluorescence (magenta) at the cell body and stereocilia counterstained with phalloidin to label F-actin (green) to visualize hair cell boundaries and stereocilia. The three rows of outer hair cells (OHC) and one row of inner hair cells (IHC) are indicated. Scale bar = 20 mm. (B) Confocal images of a damaged tissue area displaying 3 kDa dextran-TR fluorescence (magenta) at the cell body and stereocilia. Yellow arrows indicate the labeling of non-sensory supporting cells. Scale bar = 20 mm. (C) Closer view of 3 kDa dextran-TR fluorescence at the cell body and stereocilia of outer hair cells (OHC, top) and inner hair cells (IHC, bottom). Scale bar = 5 mm. This figure has been modified from27. <a href="https://www.jove.com/files/ftp_upload/60769/60769fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/60769/60769fig4.jpg" /> 
Figure 4: Labeling of larger 10 kDa dextran and a combination of 3 and 10 kDa dextran. (A) Confocal image of hair cells at the hair cell body (top) and stereocilia (bottom) levels incubated with 10 kDa dextran-TR. The dextran fluorescence signal is shown in magenta merged with F-actin in green and independently in grayscale. A couple of IHC are shown at higher magnification in the inset to appreciate the superficial labeling of stereocilia observed with the 10 kDa dextran-TR. Scale bar = 5 mm. (B) Confocal image of hair cells incubated with 10 kDa dextran-FITC represented as in A. (C) Confocal image of hair cells at the hair cell body (top) and stereocilia (bottom) levels incubated simultaneously with 3 kDa dextran-TR and 10 kDa dextran-FITC and imaged using independent channels and depicted separately in gray. In the right panel, F-actin (blue), 10 kDa dextran-FITC (green), and 3 kDa dextran-TR (magenta) are shown together along with phalloidin (blue). Scale bar = 20 mm, and the vesicle-like pattern of uptake is indicated with yellow arrows. This figure has been modified from27. <a href="https://www.jove.com/files/ftp_upload/60769/60769fig4alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/60769/60769fig5.jpg" /> 
Figure 5: Functional MET channels are required for 3 kDa dextran-TR uptake. Representative confocal images of hair cells at the stereocilia (A) or cell body (B) level after incubation with 3 kDa dextran-TR in the absence (control) or presence of BAPTA or the MET channel blockers amiloride and neomycin. 3 kDa dextran-TR fluorescence is shown in magenta. Tissue was counterstained with phalloidin to label F-actin for visualization of the hair cell stereocilia and boundaries (green). White arrows point to vesicle-like structures. The three rows of outer hair cells (OHC) and one row of inner hair cells (IHC) are indicated, Scale bar = 20 mm. This figure has been modified from27. <a href="https://www.jove.com/files/ftp_upload/60769/60769fig5alarge.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Dextran Labeling and Uptake in Live and Functional Murine Cochlear Hair Cells at JoVE.com

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

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 ...

dextran-labeling-uptake-live-functional-murine-cochlear-hair

Universidad San Sebastian

Research

JoVE Journal

Bioengineering

7.3K Views.  National Institute of Neurological Disorders and Stroke, National Institutes of Health. 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 3–10 kDa can be used to study endocytosis in hair and supporting cells of the organ of Corti.

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

Transplantation of Induced Pluripotent Stem Cell-derived Mesoangioblast-like Myogenic Progenitors in Mouse Models of Muscle Regeneration

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells similar to pericyte-derived mesoangioblasts (iPSC-derived mesoangioblast-like stem/progenitor cells: IDEMs) can be established from iPSCs generated from patients affected by different forms of muscular dystrophy. Patient-specific IDEMs can be genetically corrected with different strategies (e.g. lentiviral vectors, human artificial chromosomes) and enhanced in their myogenic differentiation potential upon overexpression of the myogenesis regulator MyoD. This myogenic potential is then assessed in vitro with specific differentiation assays and analyzed by immunofluorescence. The regenerative potential of IDEMs is further evaluated in vivo, upon intramuscular and intra-arterial transplantation in two representative mouse models displaying acute and chronic muscle regeneration. The contribution of IDEMs to the host skeletal muscle is then confirmed by different functional tests in transplanted mice. In particular, the amelioration of the motor capacity of the animals is studied with treadmill tests. Cell engraftment and differentiation are then assessed by a number of histological and immunofluorescence assays on transplanted muscles. Overall, this paper describes the assays and tools currently utilized to evaluate the differentiation capacity of IDEMs, focusing on the transplantation methods and subsequent outcome measures to analyze the efficacy of cell transplantation.

Induced pluripotent stem cell (iPSC)-derived myogenic progenitors are promising candidates for cell therapy strategies to treat muscular dystrophies. This protocol describes transplantation and functional measurements required to evaluate the engraftment and differentiation of iPSC-derived mesoangioblasts (a type of muscle progenitors) in mouse models of acute and chronic muscle regeneration.

Patient-derived iPSCs could be an invaluable source of cells for future autologous cell therapy protocols. iPSC-derived myogenic stem/progenitor cells ...

Ordering Single Cells and Single Embryos in 3D Confinement: A New Device for High Content Screening

Biological cells are usually observed on flat (2D) surfaces. This condition is not physiological, and phenotypes and shapes are highly variable. Screening based on cells in such environments have therefore serious limitations: cell organelles show extreme phenotypes, cell morphologies and sizes are heterogeneous and/or specific cell organelles cannot be properly visualized. In addition, cells in vivo are located in a 3D environment; in this situation, cells show different phenotypes mainly because of their interaction with the surrounding extracellular matrix of the tissue. In order to standardize and generate order of single cells in a physiologically-relevant 3D environment for cell-based assays, we report here the microfabrication and applications of a device for in vitro 3D cell culture. This device consists of a 2D array of microcavities (typically 105 cavities/cm2), each filled with single cells or embryos. Cell position, shape, polarity and internal cell organization become then normalized showing a 3D architecture. We used replica molding to pattern an array of microcavities, &#8216;eggcups&#8217;, onto a thin polydimethylsiloxane (PDMS) layer adhered on a coverslip. Cavities were covered with fibronectin to facilitate adhesion. Cells were inserted by centrifugation. Filling percentage was optimized for each system allowing up to 80%. Cells and embryos viability was confirmed. We applied this methodology for the visualization of cellular organelles, such as nucleus and Golgi apparatus, and to study active processes, such as the closure of the cytokinetic ring during cell mitosis. This device allowed the identification of new features, such as periodic accumulations and inhomogeneities of myosin and actin during the cytokinetic ring closure and compacted phenotypes for Golgi and nucleus alignment. We characterized the method for mammalian cells, fission yeast, budding yeast, C. elegans with specific adaptation in each case. Finally, the characteristics of this device make it particularly interesting for drug screening assays and personalized medicine.

We report a device and a new method to study cells and embryos. Single cells are precisely ordered in microcavity arrays. Their 3D confinement is a step towards 3D environments encountered in physiological conditions and allows organelle orientation. By controlling cell shape, this setup minimizes variability reported in standard assays.

Biological cells are usually observed on flat (2D) surfaces. This condition is not physiological, and phenotypes and shapes are highly variable. Screening ...

Cell Labeling and Targeting with Superparamagnetic Iron Oxide Nanoparticles

Targeted delivery of cells and therapeutic agents would benefit a wide range of biomedical applications by concentrating the therapeutic effect at the target site while minimizing deleterious effects to off-target sites. Magnetic cell targeting is an efficient, safe, and straightforward delivery technique. Superparamagnetic iron oxide nanoparticles (SPION) are biodegradable, biocompatible, and can be endocytosed into cells to render them responsive to magnetic fields. The synthesis process involves creating magnetite (Fe3O4) nanoparticles followed by high-speed emulsification to form a poly(lactic-co-glycolic acid) (PLGA) coating. The PLGA-magnetite SPIONs are approximately 120 nm in diameter including the approximately 10 nm diameter magnetite core. When placed in culture medium, SPIONs are naturally endocytosed by cells and stored as small clusters within cytoplasmic endosomes. These particles impart sufficient magnetic mass to the cells to allow for targeting within magnetic fields. Numerous cell sorting and targeting applications are enabled by rendering various cell types responsive to magnetic fields. SPIONs have a variety of other biomedical applications as well including use as a medical imaging contrast agent, targeted drug or gene delivery, diagnostic assays, and generation of local hyperthermia for tumor therapy or tissue soldering.

Targeted cell delivery is useful in a variety of biomedical applications. The goal of this protocol is to use superparamagnetic iron oxide nanoparticles (SPION) to label cells and thereby enable magnetic cell targeting approaches for a high degree of control over cell delivery and localization.

Targeted delivery of cells and therapeutic agents would benefit a wide range of biomedical applications by concentrating the therapeutic effect at the ...

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and dampen loads in the spine. The IVD consists of a proteoglycan-rich nucleus pulposus (NP) and a collagen-rich annulus fibrosis (AF) sandwiched by cartilaginous end-plates. Together with the adjacent vertebrae, the vertebrae-IVD structure forms a functional spine unit (FSU). These microstructures contain unique cell types as well as unique extracellular matrices. Whole organ culture of the FSU preserves the native extracellular matrix, cell differentiation phenotypes, and cellular-matrix interactions. Thus, organ culture techniques are particularly useful for investigating the complex biological mechanisms of the IVD. Here, we describe a high-throughput approach for culturing whole lumbar mouse FSUs that provides an ideal platform for studying disease mechanisms and therapies for the IVD. Furthermore, we describe several applications that utilize this organ culture method to conduct further studies including contrast-enhanced microCT imaging and three-dimensional high-resolution finite element modeling of the IVD.

An In Vitro Organ Culture Model of the Murine Intervertebral Disc

Whole organ culture of the intervertebral disc (IVD) preserves the native extracellular matrix, cell phenotypes, and cellular-matrix interactions. Here we describe an IVD culture system using mouse lumbar and caudal IVDs in their functional spinal units and several applications utilizing this system.

Intervertebral disc (IVD) degeneration is a significant contributor to low back pain. The IVD is a fibrocartilaginous joint that serves to transmit and ...

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by their natural environment, root hair morphology and function are difficult to study and often excluded from plant research. In recent years, microfluidic platforms have offered a way to visualize root systems at high resolution without disturbing the roots during transfer to an imaging system. The microfluidic platform presented here builds on previous plant-on-a-chip research by incorporating a two-layer device to confine the Arabidopsis thaliana main root to the same optical plane as the root hairs. This design enables the quantification of root hairs on a cellular and organelle level and also prevents z-axis drifting during the addition of experimental treatments. We describe how to store the devices in a contained and hydrated environment, without the need for fluidic pumps, while maintaining a gnotobiotic environment for the seedling. After the optical imaging experiment, the device may be disassembled and used as a substrate for atomic force or scanning electron microscopy while keeping fine root structures intact.

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

This article demonstrates how to culture Arabidopsis thaliana seedlings in a two-layer microfluidic platform that confines the main root and root hairs to a single optical plane. This platform can be used for real-time optical imaging of fine root morphology as well as for high-resolution imaging by other means.

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by ...

Melt Electrospinning Writing of Three-dimensional Poly(&#949;-caprolactone) Scaffolds with Controllable Morphologies for Tissue Engineering Applications

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology with great potential for biomedical applications. The technique facilitates the direct deposition of biocompatible polymer fibers to fabricate well-ordered scaffolds in the sub-micron to micro scale range. The establishment of a stable, viscoelastic, polymer jet between a spinneret and a collector is achieved using an applied voltage and can be direct-written. A significant benefit of a typical porous scaffold is a high surface-to-volume ratio which provides increased effective adhesion sites for cell attachment and growth. Controlling the printing process by fine-tuning the system parameters enables high reproducibility in the quality of the printed scaffolds. It also provides a flexible manufacturing platform for users to tailor the morphological structures of the scaffolds to their specific requirements. For this purpose, we present a protocol to obtain different fiber diameters using melt electrospinning writing (MEW) with a guided amendment of the parameters, including flow rate, voltage and collection speed. Furthermore, we demonstrate how to optimize the jet, discuss often experienced technical challenges, explain troubleshooting techniques and showcase a wide range of printable scaffold architectures.

Melt Electrospinning Writing of Three-dimensional Poly(&amp;#949;-caprolactone) Scaffolds with Controllable Morphologies for Tissue Engineering Applications

This protocol serves as a comprehensive guideline to fabricate scaffolds via electrospinning with polymer melts in a direct writing mode. We systematically outline the process and define the appropriate parameter settings for achieving targeted scaffold architectures.

This tutorial reflects on the fundamental principles and guidelines for electrospinning writing with polymer melts, an additive manufacturing technology ...

The Organoid Reconstitution Assay (ORA) for the Functional Analysis of Intestinal Stem and Niche Cells

The intestinal epithelium is characterized by an extremely rapid turnover rate. In mammals, the entire epithelial lining is renewed within 4 - 5 days. Adult intestinal stem cells reside at the bottom of the crypts of Lieberk&#252;hn, are earmarked by expression of the Lgr5 gene, and preserve homeostasis through their characteristic high proliferative rate1. Throughout the small intestine, Lgr5+ stem cells are intermingled with specialized secretory cells called Paneth cells. Paneth cells secrete antibacterial compounds (i.e., lysozyme and cryptdins/defensins) and exert a controlling role on the intestinal flora. More recently, a novel function has been discovered for Paneth cells, namely their capacity to provide niche support to Lgr5+ stem cells through several key ligands as Wnt3, EGF, and Dll12.
When isolated ex vivo and cultured in the presence of specific growth factors and extracellular matrix components, whole intestinal crypts give rise to long-lived and self-renewing 3D structures called organoids that highly resemble the crypt-villus epithelial architecture of the adult small intestine3. Organoid cultures, when established from whole crypts, allow the study of self-renewal and differentiation of the intestinal stem cell niche, though without addressing the contribution of its individual components, namely the Lgr5+ and Paneth cells.
Here, we describe a novel approach to the organoid assay that takes advantage of the ability of Paneth and Lgr5+ cells to associate and form organoids when co-cultured. This approach, here referred to as &#34;organoid reconstitution assay&#34; (ORA), allows the genetic and biochemical modification of Paneth or Lgr5+ stem cells, followed by reconstitution into organoids. As such, it allows the functional analysis of the two main components of the intestinal stem cell niche.

The Organoid Reconstitution Assay ORA for the Functional Analysis of Intestinal Stem and Niche Cells

Intestinal organoid cultures are established from whole crypts and do not allow the analysis of self-renewal and differentiation in a cell-specific fashion. This protocol describes reconstitution of sorted stem (Lgr5+) and niche (Paneth) cells, which give rise to organoids while enabling their prior biochemical and genetic modification and functional analysis.

The intestinal epithelium is characterized by an extremely rapid turnover rate. In mammals, the entire epithelial lining is renewed within 4 - 5 days. ...

Uptake of New Lipid-coated Nanoparticles Containing Falcarindiol by Human Mesenchymal Stem Cells

Nanoparticles are the focus of an increased interest in drug delivery systems for cancer therapy. Lipid-coated nanoparticles are inspired in structure and size by low-density lipoproteins (LDLs) because cancer cells have an increased need for cholesterol to proliferate, and this has been exploited as a mechanism for delivering anticancer drugs to cancer cells. Moreover, depending on drug chemistry, encapsulating the drug can be advantageous to avoid degradation of the drug during circulation in vivo. Therefore, in this&#160;study, this design is used to fabricate lipid-coated nanoparticles of the anticancer drug falcarindiol, providing a potential new delivery system of falcarindiol in order to stabilize its chemical structure against degradation and improve its uptake by tumors. Falcarindiol nanoparticles, with a phospholipid and cholesterol monolayer encapsulating the purified drug core of the particle, were designed. The lipid monolayer coating consists of 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC), cholesterol (Chol), and 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000] (DSPE PEG 2000) along with the fluorescent label DiI (molar ratios of 43:50:5:2). The nanoparticles are fabricated using the rapid injection method, which is a fast and simple technique to precipitate nanoparticles by good-solvent for anti-solvent exchange. It consists of a rapid injection of an ethanol solution containing the nanoparticle components into an aqueous phase. The size of the fluorescent nanoparticles is measured using dynamic light scattering (DLS) at 74.1 &#177; 6.7 nm. The uptake of the nanoparticles is tested in human mesenchymal stem cells (hMSCs) and imaged using fluorescence and confocal microscopy. The uptake of the nanoparticles is observed in hMSCs, suggesting the potential for such a stable drug delivery system for falcarindiol.

This article describes the encapsulation of falcarindiol in lipid-coated 74 nm nanoparticles. The cellular uptake of the nanoparticles&#160;by human stem cells into lipid droplets is monitored by fluorescent and confocal imaging. Nanoparticles are fabricated by the rapid injection method of solvent shifting, and their size is measured with the dynamic light scattering technique.

Nanoparticles are the focus of an increased interest in drug delivery systems for cancer therapy. Lipid-coated nanoparticles are inspired in structure and ...

High-resolution Imaging of Nuclear Dynamics in Live Cells under Uniaxial Tensile Strain

Extracellular mechanical strain is known to elicit cell phenotypic responses and has physiological relevance in several tissue systems. To capture the effect of applied extracellular tensile strain on cell populations in vitro via biochemical assays, a device has previously been designed which can be fabricated simply and is small enough to fit inside tissue culture incubators, as well as on top of microscope stages. However, the previous design of the polydimethylsiloxane substratum did not allow high-resolution subcellular imaging via oil-immersion objectives. This work describes a redesigned geometry of the polydimethylsiloxane substratum and a customized imaging setup that together can facilitate high-resolution subcellular imaging of live cells while under applied strain. This substratum can be used with the same, earlier designed device and, hence, has the same advantages as listed above, in addition to allowing high-resolution optical imaging. The design of the polydimethylsiloxane substratum can be improved by incorporating a grid which will facilitate tracking the same cell before and after the application of strain. Representative results demonstrate high-resolution time-lapse imaging of fluorescently labeled nuclei within strained cells captured using the method described here. These nuclear dynamics data give insights into the mechanism by which applied tensile strain promotes differentiation of oligodendrocyte progenitor cells.&#160;

Using a previously designed device to apply mechanical strain to adherent cells, this paper describes a redesigned substratum geometry and a customized apparatus for high-resolution single-cell imaging of strained cells with a 100x oil immersion objective.

Extracellular mechanical strain is known to elicit cell phenotypic responses and has physiological relevance in several tissue systems. To capture the ...

Near Simultaneous Laser Scanning Confocal and Atomic Force Microscopy (Conpokal) on Live Cells

Techniques available for micro- and nano-scale mechanical characterization have exploded in the last few decades. From further development of the scanning and transmission electron microscope, to the invention of atomic force microscopy, and advances in fluorescent imaging, there have been substantial gains in technologies that enable the study of small materials. Conpokal is a portmanteau that combines confocal microscopy with atomic force microscopy (AFM), where a probe &#8220;pokes&#8221; the surface. Although each technique is extremely effective for the qualitative and/or quantitative image collection on their own, Conpokal provides the capability to test with blended fluorescence imaging and mechanical characterization. Designed for near simultaneous confocal imaging and atomic force probing, Conpokal facilitates experimentation on live microbiological samples. The added insight from paired instrumentation provides co-localization of measured mechanical properties (e.g., elastic modulus, adhesion, surface roughness) by AFM with subcellular components or activity observable through confocal microscopy. This work provides a step by step protocol for the operation of laser scanning confocal and atomic force microscopy, simultaneously, to achieve same cell, same region, confocal imaging, and mechanical characterization.

Near Simultaneous Laser Scanning Confocal and Atomic Force Microscopy Conpokal on Live Cells

Presented here is the protocol of the Conpokal technique, which combines confocal and atomic force microscopy into a single instrument platform. Conpokal provides same cell, same region, near simultaneous confocal imaging and mechanical characterization of live biological samples.

Techniques available for micro- and nano-scale mechanical characterization have exploded in the last few decades. From further development of the scanning ...